Encyclopedia of Virology, Volume 3: Viruses as Infectious Agents: Plant Viruses [4 ed.] 0128234075, 9780128234075

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ENCYCLOPEDIA OF VIROLOGY FOURTH EDITION

Volume 3

ENCYCLOPEDIA OF VIROLOGY FOURTH EDITION EDITORS IN CHIEF

Dennis H. Bamford Molecular and Integrative Biosciences Research Programme Faculty of Biological and Environmental Sciences University of Helsinki, Helsinki, Finland

Mark Zuckerman South London Specialist Virology Centre King’s College Hospital NHS Foundation Trust London, United Kingdom and Department of Infectious Diseases School of Immunology and Microbial Sciences, King’s College London Medical School London, United Kingdom

Volume 3

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EDITORS IN CHIEF

Dennis H. Bamford, PhD, is Professor Emeritus of Virology at the Faculty of Biological and Environmental Sciences, University of Helsinki, Finland. He obtained his PhD in 1980 from the Department of Genetics, University of Helsinki. During 1981–1982 he was an EMBO postdoctoral fellow at the Public Health Research Institute of the City of New York, United States, and during 1983–1992 he worked as a Senior Scientist at the Academy of Finland. In 1993 he was appointed Professor of General Microbiology at the University of Helsinki. He was awarded the esteemed Academy Professorship twice, in 2002–2007 and 2012–2016, and he also served twice as the Director of the Finnish Center of Excellence (in Structural Virology, 2000–2005, and in Virus Research, 2006–2011). Prof. Bamford has had continuous external research funding (e.g., from several European Union, Academy of Finland, TEKES and Jusélius Foundation funds, as well as the Human Frontier Science Program). He is an EMBO member and has held several positions of trust in scientific and administrative organizations. Prof. Bamford has published approx. 400 articles in international peer-reviewed journals in virology, microbiology, biochemistry, and molecular biology (36 of them in high impact journals). About half of the primary articles have been published with international collaborators showing high international integration. He has also been invited to give 56 keynote and plenary presentations in major international meetings. Prof. Bamford has supervised over 35 Master’s and over 40 PhD theses. Seven of his graduate students or post docs have obtained a professorship and a similar number have a principal investigator status. Prof. Bamford has studied virus evolution from a structure-centered perspective, showing that seemingly unrelated viruses, such as bacteriophage PRD1 and human adenovirus have similar virion architecture. When the corona virion architecture was gradually revealed, it was observed that its structural elements were close to those seen in RNA bacteriophage phi6 so that phi6 has been actively used as surrogate for pathogenic viruses - quite a surprise!

Dr. Mark Zuckerman is Head of Virology, Consultant Medical Virologist, and Honorary Senior Lecturer at South London Specialist Virology Centre, King’s College Hospital NHS Foundation Trust and King’s College London Medical School, Department of Infectious Diseases, School of Immunology and Microbial Sciences in London, United Kingdom. His interests include the clinical interface between developing molecular diagnostic tests relevant to the local population of patients, respiratory virus infections, herpesvirus infections in immunocompromised patients and blood-borne virus transmission incidents in the healthcare setting. He has chaired the UK Clinical Virology Network, Royal College of Pathologists Virology Specialty Advisory Committee and Virology Examiners Panel and is a member of the Specialty Advisory Committee on Transfusion Transmitted Viruses. He is a co-author on four editions of the “Mims’ Medical Microbiology” textbook, has written chapters in a number of other textbooks and has over 100 publications in international peer-reviewed journals and is an associate editor for two journals.

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EDITORIAL BOARD Editors in Chief Dennis H. Bamford Molecular and Integrative Biosciences Research Programme, Faculty of Biological and Environmental Sciences, University of Helsinki, Helsinki, Finland Mark Zuckerman South London Specialist Virology Centre, King’s College Hospital NHS Foundation Trust, London, United Kingdom and Department of Infectious Diseases, School of Immunology and Microbial Sciences, King’s College London Medical School, London, United Kingdom

Section Editors Claude M. Fauquet St Louis, MO, United States Michael Feiss Department of Microbiology and Immunology, Carver College of Medicine, University of Iowa, Iowa City, IA, United States Elizabeth E. Fry Department of Structural Biology, Nuffield Department of Medicine, University of Oxford, Oxford, United Kingdom Said A. Ghabrial† Department of Plant Pathology, University of Kentucky, Lexington, KY, United States Eric Hunter Department of Pathology and Laboratory Medicine, Emory University School of Medicine and Emory Vaccine Center, Emory University, Atlanta, GA, United States Ilkka Julkunen Institute of Biomedicine, University of Turku, Turku, Finland Peter J. Krell Department of Molecular and Cellular Biology, University of Guelph, Guelph, ON, Canada Mart Krupovic Archaeal Virology Unit, Institut Pasteur, Paris, France Maija Lappalainen HUS Diagnostic Center, HUSLAB, Clinical Microbiology, University of Helsinki and Helsinki University Hospital, Helsinki, Finland Hubert G.M. Niesters Department of Medical Microbiology and Infection Prevention, Division of Clinical Virology, University Medical Center Groningen, Groningen, The Netherlands Massimo Palmarini MRC-University of Glasgow Centre for Virus Research, Glasgow, United Kingdom David Prangishvili Institut Pasteur, Paris, France and Ivane Javakhishvili Tbilisi State University, Tbilisi, Georgia David I. Stuart Department of Structural Biology, Nuffield Department of Medicine, University of Oxford, Oxford, United Kingdom and Diamond Light Source, Didcot, United Kingdom Nobuhiro Suzuki Institute of Plant Stress and Resources (IPSR), Okayama University, Kurashiki, Japan

†

Deceased.

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SECTION EDITORS Claude Fauquet received his PhD in biochemistry from University Louis Pasteur in Strasburg, France in 1974. Dr. Fauquet joined the Institut de Recherche pour le Dévelopement (IRD) and worked there as a plant virologist for 28 years, and served in Ivory Coast, West Africa for 14 years. In 1991, he founded the International Laboratory for Tropical Agricultural Biotechnology (ILTAB) at The Scripps Research Institute, CA, United States. ILTAB was then hosted by the Donald Danforth Plant Science Center, St. Louis, MO, from 1999 to 2012. In 2003, he co-founded the Global Cassava Partnership for the 21st Century (GCP21), which he directed until 2019 and which goal is to improve the cassava crop worldwide. Dr. Fauquet is an international leader in plant virology including taxonomy, epidemiology, molecular virology, and in gene-silencing as an antiviral strategy. He was Secretary of the International Committee on Taxonomy of Viruses (ICTV) for 18 years and the editor of several ICTV Reports including the VIIIth ICTV Report in 2005. He has published more than 300 research papers in reviewed journals and books. He is a fellow of the American Association for the Advancement of Science, of the American Phytopathological Society and a member of the St. Louis Academy of Sciences. In 2007, Dr. Fauquet was knighted “Chevalier de l’Ordre des Palmes Académiques” by the French Minister of High Education and Research.

Dr. Michael Feiss is Professor Emeritus in the Department of Microbiology and Immunology of the Carver College of Medicine at the University of Iowa, IA, United States. Dr. Feiss received his PhD in Genetics at the University of Washington followed by a postdoctoral traineeship in the laboratory of Dr. Allan Campbell at Stanford. Dr. Feiss is a microbial geneticist who studies virus assembly with an emphasis on how a DNA virus, bacteriophage lambda, packages viral DNA into the empty prohead shell. The lab investigates how sites in the viral DNA orchestrate the initiation and termination of the DNA packaging process. This work includes comprehensive examination of the DNA recognition sites. A related interest is study of terminase, the viral DNA packaging enzyme, including the functional domains for protein–DNA and protein–protein interactions. A second focus has been the roles of the bacterial host’s IHF and DnaJ proteins in the lytic life cycle of the virus. More recent work has involved a genetic dissection of the role of terminase’s ATPase center that powers translocation of viral DNA into the prohead. This interest in the ATP hydrolysis-driven packaging motor involves a multidisciplinary collaboration examining the kinetics of DNA packaging during individual packaging events. Finally, recent studies have also looked at how the packaging process has diverged among several lambda-like phages, including phages 21, N15, and Gifsy-1.

Elizabeth E. Fry is a senior postdoctoral scientist in structural biology at the University of Oxford, Oxford, United Kingdom, where she received her DPhil. for studies relating to the structure determination of Foot-and-Mouth Disease Virus. Specializing in structural virology, Dr. Fry has studied many virus/viral protein structures but her primary focus is on picornavirus structure and function, in particular receptor interactions and virus uncoating. She is particularly interested in rationally designing virus-like-particles as next generation vaccines to reduce the inherent risks in handling live viruses.

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Section Editors Said A. Ghabrial† received his BSc in 1959 from Cairo University, Cairo, Egypt, and his PhD from Louisiana State University, Baton Rouge, LA, United States, in 1965. Dr. Ghabrial did postdoctoral research at the University of California, Davis, CA, United States, before returning to Cairo, where he served as a plant virologist in the Ministry of Agriculture. He returned to the United States in 1970 to do postdoctoral research at Purdue University, West Lafayette, IN. In 1972, he joined the Plant Pathology Department at the University of Kentucky, Lexington, KY, United States, where he rose to the rank of professor in 1986 and worked until 2013. Dr. Ghabrial has served as an associate and senior editor of Phytopathology. He served on the editorial boards of the Encyclopedia of Virology, 3rd edition and Encyclopedia of Plant Pathology, and edited a thematic issue of Advances in Virus Research on “Mycoviruses”. He was a member of the American Phytopathological Society (APS) and the American Society for Virology (ASV); in July 2002 he was elected as a Fellow of the American Phytopathological Society. He also acted as Chair of the ICTV Subcommittee on Fungal Viruses in 1987–1993 and 2011–2014. His long professional career allowed him to make many scientific achievements in phytopathology and virology. Among them are molecular dissection of a legume-infecting RNA virus, bean pod mottle virus (BPMV), development of BPMV-based vectors, discovery of a transmissible debilitation disease of the phytopathogenic ascomycete, Helminthosporium victoriae (Cochliobolus victoriae), establishment of a viral etiology of the H. victoriae disease, and advancement of structural biology of diverse fungal viruses.

Eric Hunter, PhD, is Professor of Pathology and Laboratory Medicine at Emory University, Atlanta, GA, United States. He serves as Co-Director of the Emory Center for AIDS Research and is a Georgia Research Alliance Eminent Scholar. Dr. Hunter’s research focus has been the molecular virology and pathogenesis of retroviruses, including human immunodeficiency virus. He has made significant contributions to the understanding of the role of retroviral glycoprotein structural features during viral entry and providing unique insights into the assembly and replication of this virus family. In recent years the emphasis of his research has been on HIV transmission and pathogenesis, defining the extreme genetic bottleneck and selection of viruses with unique traits during HIV heterosexual transmission. He has described the selection of fitter viruses at the target mucosa, a gender difference in the extent of selection bias, and a role for genital inflammation in reducing selection. His research has defined the impact of HIV adaptation to the cellular immune response on immune recognition and control of HIV after transmission, as well as on virus replicative fitness in vitro and in vivo. Recent work highlights the roles that virus replicative fitness and sex of the host play in defining disease progression in a newly infected individual. His bibliography includes over 300 peer-reviewed articles, reviews, and book chapters. He has also been the recipient of four NIH merit awards for his work on retrovirus and HIV molecular biology. Dr. Hunter served as the Editor in Chief of the journal AIDS Research and Human Retroviruses for 10 years. He was Chair of the AIDS Vaccine Research Subcommittee which is charged with providing advice and consultation on AIDS vaccine research to the National Institute of Allergy and Infectious Diseases and continues to serve on editorial boards for several academic journals and on external advisory committees for several government, academic, and commercial institutions.

Ilkka Julkunen graduated as an MD/PhD in 1984 from the Department of Virology, University of Helsinki, Helsinki, Finland. He worked as a postdoctoral research fellow at Memorial SloanKettering Cancer Center in New York, United States, in 1986–1989, followed by positions as a senior scientist, group leader and research professor at Finnish Institute for Health and Welfare in 1989–2013. In 2013 he became a Professor of Virology at the University of Turku, Turku, Finland. The research interests of Dr. Julkunen have concentrated on innate and adaptive humoral immunity in viral and microbial infections. He has studied intracellular signaling and RIG-I and TLR-mediated activation of interferon system in human macrophages and dendritic cells and stable cell lines in response to human and avian influenza, Sendai, Zika and coronavirus infections. In addition, he has analyzed the downregulation of innate immunity by viral regulatory proteins from influenza, HCV, flavi-, filo- and coronaviruses. He has expertise in vaccinology, biotechnology and development of methods to analyze antiviral immunity, he has also been actively involved in research training and collaborations with biotechnological industry.

†

Deceased.

Section Editors

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Peter Krell started his career in virology early as a summer high school student working for the Canadian Forestry Service studying the resistance of nuclear polyhedrosis viruses (now called baculoviruses) to environmental exposure with Dr. Fred T. Bird at the Insect Pathology Research Institute in Sault Ste. Marie, ON, Canada. He received his BSc and MSc in biology from Carleton University studying the iridovirus Tipula Iridescent Virus with Dr Peter Lee, in Ottawa, the Canadian capital. For his PhD he headed east to Dalhousie University in Halifax, Nova Scotia on the Atlantic coast. In addition to enjoying the salt sea air, fresh cod, lobster and mussels, he studied the molecular biology of polydnaviruses under the guidance of Dr Don Stoltz. Heading south to Texas A&M University in College Station, TX, United States, as a Postdoctoral Fellow he worked with Dr. Max Summer, of baculovirus fame, and Dr. Brad Vinson continuing to study polydnaviruses, but also became steeped in the early days of molecular baculovirology. He then accepted a faculty position in the Department of Microbiology and Immunology at the University of Guelph in Guelph, ON, Canada. There he switched to baculovirus research, which was more tractable, due in part to available cell cultures and focused on viral DNA replication and functional genomics, particularly on chitinase, cathepsin and ME53. In collaboration with Dr. Eva Nagy he studied molecular biology of different animal viruses, notably Fowl Avian adenoviruses and their development as vaccine vectors, but also on the birnavirus infectious pancreatic necrosis virus, the coronavirus porcine endemic diarrhea virus, fowlpox virus and the paramyxovirus Newcastle disease virus. He has been involved extensively with virus taxonomy, being active in the International Committee on Taxonomy of Viruses (ICTV) as member of the Polydnaviridae and Baculoviridae study groups, national representative of Canada on the ICTV, member of the Executive Committee for the ICTV and Chair of the ICTV Invertebrate Virus Subcommittee. In terms of governance, Peter Krell was President of the Canadian Society of Microbiology, Secretary and later President of the Society for Invertebrate Pathology, as well as being on the Editorial Boards of the Canadian Journal of Microbiology and the ASM Journal of Virology. While at the University of Guelph, he rose through the ranks to Professor and is currently University Professor Emeritus.

Mart Krupovic is the Head of the Archaeal Virology Unit in the Department of Microbiology at the Institut Pasteur of Paris, France. He received his MSc in Biochemistry in 2005 from the Vilnius University, Vilnius, Lithuania and PhD in 2010 in general microbiology from the University of Helsinki, Helsinki, Finland. His current research focuses on the diversity, origin, and evolution of viruses, as well as molecular mechanisms of virus–host interactions in archaea. He has published over 170 journal articles and serves as an editor or on the editorial boards of Biology Direct, Research in Microbiology, Scientific Reports, Virology, and Virus Evolution. He is also a member of the Executive Committee of the International Committee on Taxonomy of Viruses (ICTV) and chairs the Archaeal Viruses Subcommittee of the ICTV.

Maija Lappalainen, MD, PhD, Associate Professor of Clinical Microbiology, is the Head of Clinical Microbiology in the HUS Diagnostic Center, HUSLAB, University of Helsinki and Helsinki University Hospital, Helsinki, Finland. In her thesis during the years 1987–1992 she studied the incidence and diagnostics of congenital toxoplasmosis. After PhD, her research interest has been in diagnostic clinical virology, viral hepatitis, respiratory infections, viral infections in the immunocompromised patients and viral infections during pregnancy.

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Section Editors

Hubert G.M. Niesters (1958) studied biology and chemistry in Nijmegen, the Netherlands. After obtaining his PhD in Utrecht (Prof. dr. M. Horzinek and Prof. dr. B. van der Zeijst, 1987) on the molecular epidemiology of infectious bronchitis virus, he worked as a post-doctoral fellow with Prof. dr. Jim Strauss at the California Institute of Technology (Pasadena, United States) on the replication of Alphaviruses. He received a Niels Stensen fellowship (The Netherlands) and an E.S. Gosney fellowship (Caltech) during this period. After returning to the Netherlands (1989), he became a research associate in medical microbiology at the Diagnostic Medical Center (Delft) but moved back to clinical virology as a senior research associate in 1991 at the Erasmus University Medical Center Rotterdam (Head Prof. dr. Ab Osterhaus). From 1993 to 2007, he was responsible for the molecular diagnostics unit. During this period, he was involved in the discovery and characterization of several new viruses and variants. In 2007, he became full professor and director of the Laboratory of Clinical Virology within the Department of Medical Microbiology at the University Medical Center Groningen and University of Groningen. He has been actively involved in the implementation and development of new technologies like real-time amplification and automation within clinical virology. He has been focusing on molecular diagnostics and its use and the clinical value in a transplant setting, as well as in monitoring treatment of hepatitis viruses. Recently, his interest focuses on rapid regional epidemiology, automation including MiddleWare solutions for molecular diagnostics, as well as the cost–benefit of rapid point-ofimpact molecular testing. Special interest is focused on raising awareness for the detection of enteroviruses (enterovirus D68) and its relationship with acute flaccid myelitis (AFM). Since 2017, he is the Chair of the executive board of QCMD (Quality Control of Molecular Diagnostics, Glasgow). He is an auditor and team leader for the Dutch Council of Accreditation and Co-Editor in Chief of the Journal of Clinical Virology. He is an (co)-author of more than 250 peer-reviewed papers, chapters and reviews including emerging viruses, such as enterovirus D68 and hepatitis E virus (H-index 80). For his entire work, he received in 2016 the “Ed Nowakowski Senior Memorial Clinical Virology Award” from the Pan American Society for Clinical Virology.

Massimo Palmarini is the Director of the MRC-University of Glasgow Centre for Virus Research and Chair of Virology at the University of Glasgow, Glasgow, United Kingdom. A veterinarian by training, his research programs focus on the biology, evolution and pathogenesis of arboviruses and the mechanisms of virus cross-species transmission. His work is funded by the MRC and the Wellcome Trust. Massimo Palmarini has been elected Fellow of the Academy of Medical Sciences, of the Royal Society of Edinburgh and of the Royal Society of Biology and he was a Wolfson-Royal Society Research Merit Awardee. He is a Wellcome Trust Investigator.

David Prangishvili, PhD, Honorary Professor at the Institut Pasteur, Paris, France, and Professor at Tbilisi State University, Tbilisi, Georgia, is one of the pioneers in studies on the biology of Archaea and their viruses. His scientific career spans ex-USSR (Institute of Molecular Biology, Moscow; 1970–1976), Georgia (Georgian National Academy of Sciences, Tbilisi; 1976–1991), Germany (Max-Planck Institute for Biochemistry, Munich; University of Regensburg; 1991–2004) and France (Institut Pasteur, Paris, 2004–2020). In the research groups headed by him, several dozens of new species and eight new families of archaeal viruses have been discovered and characterized, which display remarkable diversity of unique morphotypes and exceptional genome contents. The results of his research contribute to the knowledge on viral diversity on our planet and change the field of prokaryotic virology, leading to the notion that viruses of hyperthermophilic Archaea form a particular group in the viral world, distinctive from viruses of Bacteria and Eukarya, and to the recognition of the virosphere of Archaea as one of the distinct features of this Domain of Life. David Prangishvili is a member of the Academia Europaea, the European Academy of Microbiology, and the Georgian National Academy of Sciences.

Section Editors

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David I. Stuart is MRC Professor of Structural Biology in the Nuffield Department of Medicine, Oxford University, Oxford, United Kingdom, Life Science Director at Diamond Light Source and Director of Instruct-ERIC (pan-European organisation providing shared access to infrastructure and methods for structural biology). He has diverse interests in structural virology from picornaviruses, double-stranded RNA viruses and enveloped RNA viruses. His drive to develop structural techniques led to the determination of the structure of Bluetongue virus (1995) and then the first membrane containing virus, PRD1. More recently, he has been at the fore-front of bringing Cryo-EM technology to bear on virus structure determination and its future role in visualizing virus function in cellulo. In addition to basic science he has a strong commitment to structural vaccinology and the development of antiviral drugs.

Dr. Nobuhiro Suzuki, PhD, received his MSc (1985) in phytopathology and PhD (1989) in virology from Tohoku University in Sendai, Japan. Dr. Suzuki currently serves as a full Professor of the Institute of Plant Stress and Resources, formerly Institute of Plant Sciences and Bioresouces at Okayama University and as an Editor of Virus Research, Frontiers in Virology, Journal of General Plant Pathology, Virology Journal, and Biology. He has also been Guest Editor to PLoS Pathogens, PNAS, and mBio, and an Editorial Board member of Virology and Journal of Virology. Suzuki Laboratory focuses on characterization of diverse viruses infecting phytopathogenic fungi and exploration of their interplays. Recent achievements include the discovery of a neo-virus lifestyle exhibited by a (+)ssRNA virus and an unrelated dsRNA virus in a plant pathogenic fungus and of multilayer antiviral defense in fungi involving Dicer. Prior to coming to Kurashiki, Okayama Prefecture, he was a visiting fellow of the Center for Agricultural Biotechnology at the University of Maryland Biotechnology Institute (UMBI), College Park, MA, United States, for 4 years (1997–2001) to study molecular biology of hypoviruses in the laboratory of Professor Donald L. Nuss. Before visiting UMBI, he served as an assistant professor and a lecturer of the Biotechnology Institute at the Akita Prefectural College of Agriculture, Japan, for 11 years (1988–1998) where he was engaged in a project on molecular characterization of rice dwarf phytoreovirus, a member of the family Reoviridae. He received awards from the Japanese Phytopathological Society of Japan and Japanese Society for Virology for his outstanding achievements in plant and fungal virology.

FOREWORD I am delighted to write the foreword to this wonderful Fourth Edition of the Encyclopedia of Virology. The Third Edition was published in 2008, how the world has changed in the intervening years. The release of the updated fourth edition could not be more timely or more prescient. It is superb and a huge tribute to the authors, Elsevier the publisher, and to the brilliant editors, Dennis Bamford and Mark Zuckerman. SARS-CoV-2 has dominated the world since it emerged in 2019 and affected every continent and every aspect of life. A reminder, if it were needed, of the impact of infectious diseases, the importance of virology and the vulnerability and interconnectivity of our world. There is no doubt that with rapidly changing ecology, urbanization, climate change, increased travel, and fragile public health systems, epidemics and pandemics will become more frequent, more complex and harder to prevent and contain. Most of these epidemics will be caused by viruses, those we know about and maybe able to predict and some we do not know of that will emerge from animals, plants or the environment. Our changing climate will change the epidemiology of viruses, their vectors and the infections they cause, hence the critical importance of this totally revised Fourth Edition of the Encyclopedia of Virology which brings together research and an understanding of viruses in animals, plants, bacteria and fungi, the environment, and among humans. Never has a holistic, one-health understanding been more important. That starts with an understanding of the fundamentals of virology, a field of science that has been transformed in the years since the Third Edition. An understanding transformed by embracing traditional fields of molecular and structural biology, genomics, and influenced by immunology, genetics, pharmacology and increasingly by epidemiology and mathematics. Events of 2020 and 2021 also show why it is so important to integrate within traditional virology an understanding of the animal and human health and behavior, of climate change and its impact on the ecology of viruses, plant sciences and vectors. And why we must understand the viruses we think we know well, and those viruses less extensively studied. Research is critical to this, research that pushes the boundaries of what we know, has the humility to seek answers to things we do not understand and shares that knowledge with the widest possible community. That research will be most exciting at the interface between disciplines, most impactful when dynamic, open, inclusive, global, and collaborative. This is what the Fourth Edition of the Encyclopedia of Virology, the largest reference source of research in virology sets out to achieve. It is a wonderful contribution to a critical field of knowledge. It contains new chapters, every chapter revised and updated by a dedicated global community who have come together to provide what is a brilliant and inspiring reference. It is an honor to contribute in a very small way to the timely release of the Fourth Edition of the Encyclopedia of Virology. Jeremy Farrar

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PREFACE The fourth edition of the Encyclopedia of Virology is encyclopedic, but we wanted to move away from an alphabetical list, apart from where it was more logical, to a vision that encompassed a different structure. Articles describing novel trends as well as original discoveries in specific subfields of virology have been distributed into a set of five volumes, namely Fundamentals of Virology, Human and Animal Viruses, Plant Viruses, Bacterial, Archaeal, Fungal, Algal and Invertebrate Viruses, and Diagnosis, Treatment and Prevention of Virus Infections. We had hoped that the new edition would ‘go viral’ but it was ironic that the time to publication 12 years after the previous edition had been made a bit longer due to a virus infection. The world encountered a devastating global pandemic, COVID-19, caused by a new type of a coronavirus, SARS-CoV-2. Scientists in many disciplines all over the world started immediate efforts to discover solutions as to how to mitigate and stop the spread of the pandemic. Virology moved from being a highly specialized subject to one in which everyone became a virologist, proving just how significant the different aspects of virology are in terms of understanding the nature of viral infection. Since the previous edition, the growth in the field of general virology has been enormous, including huge advances in basic science, identification of novel viruses, diagnostic methods, treatment and prevention. Taking this into account, the introduction of the articles within the Encyclopedia are very timely and crucial for providing a wealth of knowledge of the latest findings in the field of virology to a vast range of people, whether school students, undergraduates, postgraduates, teachers, scientists, researchers, journalists and others interested in infections and the conflict between the host and the pathogen. Pandemic viruses have become a serious public concern in the changing world. We can ask ourselves whether we have reached the point in which nature can no longer cope with the consequences of increased population density and human activities that are harmful to the environment. Although several pandemics have threatened mankind before, this COVID-19 pandemic has highlighted the massive adverse economic consequences towards the wellbeing of society and the importance of research in virology. We aimed to produce a Major Reference Work that differs in approach to others and binds all the virology disciplines together. Chapters have been included on origin, evolution and emergence of viruses, environmental virology and ecology, epidemiology, techniques for studying viruses, viral life cycles, structure, entry, genome and replication, assembly and packaging and taxonomy and viral–host interactions. Information has been included on all known species of viruses infecting bacteria, fungi, plants, vertebrates and invertebrates. Additional topics include antiviral classification and examples of their use in management of infection, diagnostic assays and vaccines, as well as the economic importance of viral diseases of crops and their control. This edition used viral classification according to the 9th Report of the International Committee on Taxonomy of Viruses published in 2012. Updating it to the 10th Report in 2020 was affected by the pandemic and can be found online at http://ictv.global/report/. We wish to acknowledge the hard work, interest, flexibility and patience, during such difficult times both socially and professionally, of everybody involved in the process of writing this edition of the Encyclopedia of Virology, especially Katarzyna Miklaszewska, Priscilla Braglia, Sam Crowe and colleagues at Elsevier. We sincerely thank all the authors and section editors for their excellent contributions to this edition.

Book Cover Image: Viruses are obligate parasites and all cells have their own viruses increasing the total number of viruses to the estimated astronomical number of 1031 that extends the number of stars in the universe. The viral string illustrates how pandemic viruses surround the globe. The original picture was created by Dr. Nina Atanasova (Finnish Meteorological Institute and University of Helsinki) and amended by Matthew Limbert at Elsevier. Dennis H. Bamford Mark Zuckerman

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HOW TO USE THE ENCYCLOPEDIA Structure of the Encyclopedia All articles in the encyclopedia are arranged thematically as a series of entries within subjects/sections, apart from volume 2 where there it was more logical to have articles arranged alphabetically. There are three features to help you easily find the topic you are interested in: a thematic contents list, a full subject index, and contributors. 1. Thematic contents list: The alphabetical contents list, which appears at the front of each volume, lists the entries in the order that they appear in the encyclopedia. 2. Index: The index appears at the end of volume 5 and includes page numbers for quick reference to the information you are looking for. The index entries differentiate between references to a whole entry, a part of an entry, and a table or figure. 3. Contributors: At the start of each volume there is a list of the authors who contributed to all volumes.

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LIST OF CONTRIBUTORS Stephen T. Abedon The Ohio State University, Mansfield, OH, United States Peter Abrahamian Agricultural Research Service, US Department of Agriculture, Beltsville, MD, United States Jônatas S. Abrahão Federal University of Minas Gerais, Belo Horizonte, Brazil Florence Abravanel Toulouse University Hospital, Toulouse, France and Toulouse University Paul Sabatier, Toulouse, France Nicola G.A. Abrescia Center for Cooperative Research in Biosciences, Basque Research and Technology Alliance, Derio, Spain; Ikerbasque, Basque Foundation for Science, Bilbao, Spain; and Center for Biomedical Research in the Liver and Digestive Diseases Network, Carlos III Health Institute, Madrid, Spain Gian Paolo Accotto Institute for Sustainable Plant Protection, National Research Council of Italy, Torino, Italy

Aleksandra Alimova The City University of New York (CUNY), School of Medicine, The City College of New York, New York, NY, United States Juan C. Alonso National Biotechnology Center–Spanish National Research Council, Madrid, Spain Imran Amin National Institute for Biotechnology and Genetic Engineering, Faisalabad, Pakistan Stephanie E. Ander University of Colorado School of Medicine, Aurora, CO, United States Danielle E. Anderson Duke-NUS Medical School, Singapore, Singapore Ida Bagus Andika Qingdao Agricultural University, Qingdao, China Ana C.d.S.P. Andrade Federal University of Minas Gerais, Belo Horizonte, Brazil Juana Angel Pontifical Javeriana University, Bogota, Colombia

Elisabeth Adderson St. Jude Children’s Research Hospital, Memphis, TN, United States and University of Tennessee Health Sciences Center, Memphis, TN, United States

Vanesa Anton-Vazquez King’s College Hospital, London, United Kingdom

Mustafa Adhab University of Baghdad, Baghdad, Iraq

Guido Antonelli Sapienza University of Rome, Rome, Italy

Alexey A. Agranovsky Lomonosov Moscow State University, Moscow, Russia Nasim Ahmed National Institute for Biotechnology and Genetic Engineering, Faisalabad, Pakistan Maher Al Rwahnih University of California, Davis, CA, United States Olufemi J. Alabi Texas A& M AgriLife Research and Extension Center, Weslaco, TX, United States Aurélie A. Albertini Institute for Integrative Biology of the Cell (I2BC), French Alternative Energies and Atomic Energy Commission, French National Center for Scientific Research, Paris-Sud University, University of Paris-Saclay, Gif-sur-Yvette, France

Josefa Antón University of Alicante, Alicante, Spain Nanako Aoki Tokyo University of Agriculture and Technology, Fuchu, Japan Timothy D. Appleby King’s College Hospital, London, United Kingdom Miguel Arenas Department of Biochemistry, Genetics and Immunology, University of Vigo, Vigo, Spain and CINBIO (Biomedical Research Center), University of Vigo, Vigo, Spain Basil Arif Laboratory for Molecular Virology, Great Lakes Forestry Centre, Sault Ste Marie, ON, Canada

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xxii

List of Contributors

Vicente Arnau Institute for Integrative Systems Biology (I2SysBio), University of Valencia–Spanish National Research Council, Valencia, Spain Gaurav Arya Duke University, Durham, NC, United States Leyla Asadi University of Alberta, Edmonton, AB, Canada Sassan Asgari The University of Queensland, Brisbane, QLD, Australia Nina S. Atanasova Finnish Meteorological Institute, Helsinki, Finland and University of Helsinki, Helsinki, Finland Houssam Attoui UMR1161 Virologie, INRAE – French National Research Institute for Agriculture, Food and Environment, ANSES, Ecole Nationale Vétérinaire d’Alfort, University of Paris-Est, Maisons-Alfort, France Silvia Ayora National Biotechnology Center–Spanish National Research Council, Madrid, Spain

Xiaoyong Bao The University of Texas Medical Branch, Galveston, TX, United States Yiming Bao Beijing Institute of Genomics, Chinese Academy of Sciences, University of Chinese Academy of Sciences, Beijing, China Alan D.T. Barrett The University of Texas Medical Branch, Galveston, TX, United States Diana P. Baquero Archaeal Virology Unit, Institut Pasteur, Paris, France and Sorbonne University, Paris, France Moshe Bar-Joseph Agricultural Research Organization, Volcani Center, Bet Dagan, Israel Rachael S. Barr Bristol Royal Hospital for Children, Bristol, United Kingdom Ralf Bartenschlager Heidelberg University, Heidelberg, Germany

Walid Azab Free University of Berlin, Berlin, Germany

David L.V. Bauer Francis Crick Institute, London, United Kingdom

Sasha R. Azar The University of Texas Medical Branch, Galveston, TX, United States

Oliver W. Bayfield University of York, York, United Kingdom

Fengwei Bai The University of Southern Mississippi, Hattiesburg, MS, United States Dalan Bailey The Pirbright Institute, Pirbright, United Kingdom S.C. Baker Loyola University of Chicago, Maywood, IL, United States Fausto Baldanti University of Pavia, Pavia, Italy and Scientific Institute for Research, Hospitalization and Healthcare, San Matteo Polyclinic Foundation, Pavia, Italy Logan Banadyga Public Health Agency of Canada, Winnipeg, MB, Canada Ashley C. Banyard Animal and Plant Health Agency, Addlestone, United Kingdom; University of West Sussex, Falmer, United Kingdom; and St. George's Medical School, University of London, London, United Kingdom

Sally A. Baylis Paul-Ehrlich-Institute, Langen, Germany Philippa M. Beard The Pirbright Institute, Pirbright, United Kingdom and The Roslin Institute, University of Edinburgh, United Kingdom Paul Becher University of Veterinary Medicine, Hannover, Germany Björn Becker Saarland University, Saarbrücken, Germany Karen L. Beemon Johns Hopkins University, Baltimore, MD, United States Martin Beer Friedrich-Loeffler-Institute, Insel Riems, Germany Jose Miguel Benito Health Research Institute of the Jiménez Díaz Foundation, Autonomous University of Madrid and Rey Juan Carlos University Hospital, Móstoles, Spain Mária Benko ˝ Institute for Veterinary Medical Research, Center for Agricultural Research, Budapest, Hungary

List of Contributors

Max Bergoin National Institute of Scientific Research – ArmandFrappier Health Research Centre, Laval, QC, Canada Sabrina Bertin Council for Agricultural Research and Economics, Research Center for Plant Protection and Certification, Rome, Italy Shweta Bhatt University of Copenhagen, Copenhagen, Denmark Dennis K. Bideshi California Baptist University, Riverside, CA, United States and University of California, Riverside, CA, United States Yves Bigot INRAE – French National Research Institute for Agriculture, Food and Environment, Nouzilly, France Richard J. Bingham University of York, York, United Kingdom

Maxime Boutier University of Liège, Liège, Belgium P.R. Bowser Cornell University, Ithaca, NY, United States Daniel Bradshaw Public Health England, London, United Kingdom Claude Bragard University of Louvain, Louvain-la-Neuve, Belgium Aaron C. Brault Centers for Disease Control and Prevention, Fort Collins, CO, United States Nicolas Bravo-Vasquez St. Jude Children’s Research Hospital, Memphis, TN, United States Rob W. Briddon University of Agriculture, Faisalabad, Pakistan Thomas Briese Columbia University, New York, NY, United States

Vera Bischoff Institute for Chemistry and Biology of the Marine Environment, Oldenburg, Germany

Paul Britton The Pirbright Institute, Pirbright, United Kingdom

Kate N. Bishop Francis Crick Institute, London, United Kingdom

Thomas J. Brouwers Athena Institute, VU Amsterdam, Amsterdam, The Netherlands

Lindsay W. Black The University of Maryland School of Medicine, Baltimore, MD, United States Romain Blanc-Mathieu Institute for Chemical Research, Kyoto University, Kyoto, Japan Soile Blomqvist National Institute for Health and Welfare, Helsinki, Finland Bryony C. Bonning University of Florida, Gainesville, FL, United States Lisa M. Bono Rutgers, The State University of New Jersey, New Brunswick, NJ, United States Alexia Bordigoni Aix-Marseille University, CNRS, IRD, Mediterranean Institute of Oceanography, Marseille, France and Aix-Marseille University, IRD257, Assistance-Publique des Hôpitauxde Marseille, UMR Microbes, Evolution, Phylogeny and Infections (MEPHI), IHU Méditerranée Infection, Marseille, France Mihnea Bostina University of Otago, Dunedin, New Zealand

xxiii

Kevin E. Brown Frimley Park Hospital, Frimley, United Kingdom and Immunisation and Countermeasures Division, Public Health England, London, United Kingdom Corina P.D. Brussaard NIOZ Royal Netherlands Institute for Sea Research, Den Burg, Texel, The Netherlands and Utrecht University, Utrecht, The Netherlands Harald Brüssow Laboratory of Gene Technology, Department of Biosystems, KU Leuven, Leuven, Belgium Joachim J. Bugert Bundeswehr Institute of Microbiology, Munich, Germany Jozef J. Bujarski Northern Illinois University, DeKalb, IL, United States and Polish Academy of Sciences, Poznan, Poland Laura Burga University of Otago, Dunedin, New Zealand Sara H. Burkhard University Hospital of Zurich, Zurich, Switzerland Cara C. Burns Centers for Disease Control and Prevention, Atlanta, GA, United States

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List of Contributors

Felicity Burt University of the Free State, Bloemfontein, South Africa Kerry S. Burton Leamington Spa, United Kingdom Sarah J. Butcher University of Helsinki, Helsinki, Finland Mathias Büttner Leipzig University, Leipzig, Germany Jesse Cahill Sandia National Labs, Albuquerque, NM, United States Marianna Calabretto Sapienza University of Rome, Rome, Italy Thierry Candresse The National Research Institute for Agriculture, Food and the Environment, University of Bordeaux, Villenave d′Ornon, France Alan J. Cann University of Leicester, Leicester, United Kingdom Lorenzo Capucci The Lombardy and Emilia Romagna Experimental Zootechnic Institute, Brescia, Italy Irene Carlon-Andres University of Oxford, Oxford, United Kingdom José M. Casasnovas National Center for Biotechnology, Spanish National Research Council (CSIC), Madrid, Spain J.W. Casey Cornell University, Ithaca, NY, United States R.N. Casey Cornell University, Ithaca, NY, United States Sherwood R. Casjens University of Utah, Salt Lake City, UT, United States Antonella Casola The University of Texas Medical Branch, Galveston, TX, United States José R. Castón National Center for Biotechnology, Spanish National Research Council, Madrid, Spain

Patrizia Cavadini The Lombardy and Emilia Romagna Experimental Zootechnic Institute, Brescia, Italy Supranee Chaiwatpongsakorn Nationwide Children’s Hospital, Columbus, OH, United States Supriya Chakraborty Jawaharlal Nehru University, New Delhi, India Yu-Chan Chao Institute of Molecular Biology, Academia Sinica, Taipei, Taiwan Tyler P. Chavers Centers for Disease Control and Prevention, Atlanta, GA, United States Keping Chen Jiangsu University, Zhenjiang, China Xiaorui Chen Genomics Research Center, Academia Sinica, Taipei, Taiwan Yanping Chen Bee Research Laboratory, Agricultural Research Service, US Department of Agriculture, Beltsville, MD, United States Dayna Cheng National Cheng Kung University, Tainan, Taiwan Quentin Chesnais University of Strasbourg, Colmar, France Sotaro Chiba Nagoya University, Nagoya, Japan Wah Chiu Stanford University, Stanford, CA, United States David Chmielewski Stanford University, Stanford, CA, United States Irma E. Cisneros The University of Texas Medical Branch, Galveston, TX, United States Lark L. Coffey University of California, Davis, CA, United states Alanna B. Cohen Rutgers University, New Brunswick, NJ, United States

Carlos E. Catalano University of Colorado Anschutz Medical Campus, Skaggs School of Pharmacy and Pharmaceutical Sciences, Aurora, CO, United States

Jeffrey I. Cohen Laboratory of Infectious Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, United States

Roberto Cattaneo Mayo Clinic, Rochester, MN, United States

Seth Coleman Rice University, Houston, TX, United States

List of Contributors

Miquel Coll Institute for Research in Biomedicine, Barcelona, Spain and Institute for Molecular Biology of Barcelona, Barcelona, Spain John Collinge UCL Institute of Prion Diseases, London, United Kingdom Carina Conceicao The Pirbright Institute, Pirbright, United Kingdom Gabriela N. Condezo National Center for Biotechnology, Spanish National Research Council, Madrid, Spain

xxv

Amy Davis St Jude Children’s Research Hospital, Memphis, TN, United States William O. Dawson Citrus Research and Education Center, Lake Alfred, FL, United States and University of Florida, Lake Alfred, FL, United States Erik De Clercq Rega Institute for Medical Research, KU Leuven, Leuven, Belgium Raoul J. de Groot Utrecht University, Utrecht, The Netherlands

Michaela J. Conley MRC-University of Glasgow Centre for Virus Research, Glasgow, United Kingdom

Juan C. de la Torre The Scripps Research Institute, La Jolla, CA, United States

Charles A Coomer University of Oxford, Oxford, United Kingdom

Marcelo De las Heras University of Zaragoza, Zaragoza, Spain

Anne K. Cordes Hannover Medical School, Institute of Virology, Hannover, Germany

Juliana Gabriela Silva de Lima Federal University of Rio Grande do Norte, Natal, Brazil

Mauricio Cortes Jr. Department of Chemistry, College of Arts and Sciences, Fort Wayne, IN, United States Robert H.A. Coutts University of Hertfordshire, Hatfield, United Kingdom Jeff A. Cowley CSIRO Livestock Industries, Brisbane, QLD, Australia Robert W. Cross The University of Texas Medical Branch, Galveston, TX, United States Henryk Czosnek The Hebrew University of Jerusalem, Rehovot, Israel Håkon Dahle Department of Biological Sciences, University of Bergen, Bergen, Norway Janet M. Daly University of Nottingham, Sutton Bonington, United Kingdom Subha Das Okayama University, Kurashiki, Japan

Athos S. de Oliveira University of Brasília, Brasília, Brazil Nicole T. de Stefano University of Florida, Gainesville, FL, United States Greg Deakin NIAB-EMR, East Malling, United Kingdom Philippe Delfosse University of Luxembourg, Esch-sur-Alzette, Luxembourg Natacha Delrez University of Liège, Liège, Belgium Tatiana A. Demina Molecular and Integrative Biosciences Research Program, Faculty of Biological and Environmental Sciences, University of Helsinki, Helsinki, Finland Ismail Demir Department of Biology, Karadeniz Technical University, Trabzon, Turkey Zihni Demirbağ Department of Biology, Karadeniz Technical University, Trabzon, Turkey

Indranil Dasgupta University of Delhi, New Delhi, India

X. Deng Loyola University of Chicago, Maywood, IL, United States

Sibnarayan Datta Defence Research Laboratory, Defence Research and Development Organisation (DRDO), Tezpur, Assam, India

Cécile Desbiez Plant Pathology Unit, INRAE – French National Research Institute for Agriculture, Food and Environment, Montfavet, France

xxvi

List of Contributors

Christelle Desnues Aix-Marseille University, CNRS, IRD, Mediterranean Institute of Oceanography, Marseille, France and Aix-Marseille University, IRD 257, Assistance-Publique des Hôpitaux de Marseille, UMR Microbes, Evolution, Phylogeny and Infections (MEPHI), IHU Méditerranée Infection, Marseille, France

Lucy Dorrell University of Oxford, Oxford, United Kingdom

Samantha J. DeWerff University of Illinois at Urbana-Champaign, Urbana, IL, United States

Andreas Dotzauer University of Bremen, Bremen, Germany

Daniele Di Carlo Sapienza University of Rome, Rome, Italy Arturo Diaz La Sierra University, Riverside, CA, United States Alfredo Diaz-Lara University of California, Davis, CA, United States Ralf G. Dietzgen The University of Queensland, St. Lucia, QLD, Australia Michele Digiaro International Center for Advanced Mediterranean Agronomic Studies (CIHEAM), Mediterranean Agronomic Institute of Bari, Valenzano, Italy Michael Dills Montana State University, Bozeman, MT, United States Wayne Dimech National Serology Reference Laboratory, Fitzroy, VIC, Australia Savithramma P. Dinesh-Kumar University of California, Davis, CA, United States Linda K. Dixon The Pirbright Institute, Pirbright, United Kingdom Valerian V. Dolja Oregon State University, Corvallis, OR, United States Aušra Domanska University of Helsinki, Helsinki, Finland Leslie L. Domier Agricultural Research Service, US Department of Agriculture, Urbana, IL, United States Pilar Domingo-Calap Institute for Integrative Systems Biology (I2SysBio), University of Valencia-CSIC, Valencia, Spain Tatiana Domitrovic Federal University of Rio de Janeiro, Rio de Janeiro, Brazil Sarah M. Doore Michigan State University, East Lansing, MI, United States

Rosemary A. Dorrington Rhodes University, Grahamstown, South Africa Andor Doszpoly Hungarian Academy of Sciences, Budapest, Hungary

Simon B. Drysdale St George’s University Hospitals NHS Foundation Trust, London, United Kingdom and St George’s, University of London, London, United Kingdom Robert L. Duda University of Pittsburgh, Pittsburgh, PA, United States Carol Duffy University of Alabama, Tuscaloosa, AL, United States Siobain Duffy Rutgers, The State University of New Jersey, New Brunswick, NJ, United States David D. Dunigan University of Nebraska–Lincoln, Lincoln, NE, United States Stéphane Duquerroy University of Paris-Saclay, Orsay, France and Institut Pasteur, Paris, France Bas E. Dutilh Utrecht University, Utrecht, The Netherlands and Radboud University Medical Center, Nijmegen, The Netherlands Michael Edelstein Faculty of Medicine, Bar Ilan University, Ramat Gan, Israel Herman K. Edskes National Institutes of Health, Bethesda, MD, United States Rosina Ehmann Bundeswehr Institute of Microbiology, Munich, Germany Toufic Elbeaino International Center for Advanced Mediterranean Agronomic Studies (CIHEAM), Mediterranean Agronomic Institute of Bari, Valenzano, Italy Joanne B. Emerson University of California, Davis, CA, United States Ann Emery University of North Carolina at Chapel Hill, Chapel Hill, NC, United States

List of Contributors

xxvii

Christine E. Engeland University Hospital Heidelberg and German Cancer Research Center, Heidelberg, Germany and Witten/Herdecke University, Witten, Germany

Elvira Fiallo-Olivé Institute for Mediterranean and Subtropical Horticulture “La Mayora”–Spanish National Research Council–University of Malaga, Algarrobo-Costa, Málaga, Spain

Luis Enjuanes National Center for Biotechnology – Spanish National Research Council (CNB-CSIC), Madrid, Spain

Andrew E. Firth University of Cambridge, Cambridge, United Kingdom

Katri Eskelin University of Helsinki, Helsinki, Finland Rosa Esteban Institute of Biology and Functional Genomics, CSIC/University of Salamanca, Salamanca, Spain Mary K. Estes Baylor College of Medicine, Houston, TX, United States Cassia F. Estofolete São José do Rio Preto School of Medicine, São José do Rio Preto, Brazil Alyssa B. Evans National Institutes of Health, Hamilton, MT, United States Øystein Evensen Norwegian University of Life Sciences, Oslo, Norway Alex Evilevitch Department of Experimental Medical Science, Lund University, Lund, Sweden Montserrat Fàbrega-Ferrer Institute for Research in Biomedicine, Barcelona, Spain and Institute for Molecular Biology of Barcelona, Barcelona, Spain Francesco Faggioli Council for Agricultural Research and Economics, Research Center for Plant Protection and Certification, Rome, Italy Bentley A. Fane University of Arizona, Tucson, AZ, United States Brian A. Federici University of California, Riverside, CA, United States F. Fenner Australian National University, Canberra, ACT, Australia Isabel Fernández de Castro Cell Structure Laboratory, National Center for Biotechnology – Spanish National Research Council (CNB-CSIC), Madrid, Spain Giovanni Ferrara University of Alberta, Edmonton, AB, Canada

Roland A. Fleck King’s College London, London, United Kingdom Ricardo Flores Polytechnic University of Valencia, Higher Council of Scientific Research, Valencia, Spain Ervin Fodor University of Oxford, Oxford, United Kingdom Anthony R. Fooks Animal and Plant Health Agency, Addlestone, United Kingdom; University of Liverpool, Liverpool, United Kingdom; and St. George's Medical School, University of London, London, United Kingdom Patrick Forterre Archeal Virology Unit, Institut Pasteur, Paris, France and French National Center for Scientific Research, Institute of Integrative Biology of the Cell, University of Paris-Saclay, Gif sur Yvette, France Rennos Fragkoudis University of Nottingham, Sutton Bonington, United Kingdom and University of Edinburgh, Edinburgh, United Kingdom Manuel A. Franco Pontifical Javeriana University, Bogota, Colombia Giovanni Franzo Department of Animal Medicine, Production and Health (MAPS), Padua University, Padua, Italy Graham L. Freimanis The Pirbright Institute, Pirbright, United Kingdom Juliana Freitas-Astúa Brazilian Agricultural Research Corporation (Embrapa) Cassava and Fruits, Cruz das Almas, Brazil Elizabeth E. Fry Department of Structural Biology, Nuffield Department of Medicine, University of Oxford, Oxford, United Kingdom Marc Fuchs Cornell University, Geneva, NY, United States Tsutomu Fujimura Institute of Biology and Functional Genomics, CSIC/University of Salamanca, Salamanca, Spain

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List of Contributors

Kuko Fuke Tokyo University of Agriculture and Technology, Fuchu, Japan

Said A. Ghabrial† Department of Plant Pathology, University of Kentucky, Lexington, KY, United States

Toshiyuki Fukuhara Tokyo University of Agriculture and Technology, Fuchu, Japan

Clément Gilbert Evolution, Genomes, Behavior and Ecology Laboratory, CNRS University of Paris-Sud UMR 9191, IRD UMR 247, Gif-sur-Yvette, France

To S. Fung South China Agricultural University, Guangzhou, China Yahya Z.A. Gaafar Julius Kuehn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany Toni Gabaldon Barcelona Supercomputing Center-National Center for Supercomputing, Institute of Research in Biomedicine, and Catalan Institution for Research and Advanced Studies, Barcelona, Spain Morgan Gaïa University of Paris-Saclay, Evry, France José Gallardo National Center for Biotechnology, Spanish National Research Council, Madrid, Spain Hernan Garcia-Ruiz University of Nebraska–Lincoln, Lincoln, NE, United States Juan A. García National Center for Biotechnology-Spanish National Research Council, Madrid, Spain Matteo P. Garofalo The University of Texas Medical Branch, Galveston, TX, United States Yves Gaudin Institute for Integrative Biology of the Cell (I2BC), French Alternative Energies and Atomic Energy Commission, French National Center for Scientific Research, Paris-Sud University, University of Paris-Saclay, Gif-sur-Yvette, France Andrew D.W. Geering The University of Queensland, St. Lucia, QLD, Australia Thomas W. Geisbert The University of Texas Medical Branch, Galveston, TX, United States Andrea Gentili Council for Agricultural Research and Economics, Research Center for Plant Protection and Certification, Rome, Italy Volker Gerdts University of Saskatchewan, Saskatoon, SK, Canada

Robert L. Gilbertson University of California, Davis, CA, United States Efstathios S. Giotis Imperial College London, London, United Kingdom and University of Essex, Colchester, United Kingdom Laurent Glais French Federation of Seed Potato Growers/Research, Development, Promotion of Seed Potato, Paris, France and Institute for Genetics, Environment and Plant Protection, Agrocampus West, French National Institute for Agriculture, Food and Environment, University of Rennes 1, Le Rheu, France Miroslav Glasa Biomedical Research Center, Slovak Academy of Sciences, Bratislava, Slovakia Ido Golding University of Illinois at Urbana-Champaign, Urbana, IL, United States Esperanza Gomez-Lucia Complutense University of Madrid, Madrid, Spain Zheng Gong Beijing Institute of Genomics, Chinese Academy of Sciences, Beijing, China Andrea González-González University of Florida, Gainesville, FL, United States Michael M. Goodin University of Kentucky, Lexington, KY, United States Alexander E. Gorbalenya Leiden University Medical Center, Leiden, The Netherlands Paul Gottlieb The City University of New York (CUNY), School of Medicine, The City College of New York, New York, NY, United States M.-A. Grandbastien INRAE – French National Research Institute for Agriculture, Food and Environment, Versailles, France †

Deceased.

List of Contributors

Meritxell Granell National Center for Biotechnology, Madrid, Spain and Institute of Chemical Research of Catalonia (ICIQ), Tarragona, Spain

Sébastien Halary National Museum of Natural History, UMR 7245 CNRS/MNHN Molécule de Communication et Adaptation des Micro-organismes, Paris, France

Patrick L. Green The Ohio State University, Columbus, OH, United States

Aron J. Hall Centers for Disease Control and Prevention, Atlanta, GA, United States

Sandra J. Greive University of York, York, United Kingdom

John Hammond Floral and Nursery Plants Research, Agricultural Research Service, US Department of Agriculture, Beltsville, MD, United States

Diane E. Griffin Johns Hopkins Bloomberg School of Public Health, Baltimore, MD, United States Jonathan M. Grimes University of Oxford, Oxford, United Kingdom Nigel Grimsley Integrative Biology of Marine Organisms Laboratory, Banuyls-sur-Mer, France and Sorbonne University, Banuyls-sur-Mer, France Bruno Gronenborn Institute for Integrative Biology of the Cell, CNRS, University of Paris-Sud, CEA, Gif sur Yvette, France Julianne H. Grose Brigham Young University, Provo, UT, United States Scott Grytdal Centers for Disease Control and Prevention, Atlanta, GA, United States

Rosemarie W. Hammond Agricultural Research Service, US Department of Agriculture, Beltsville, MD, United States Virginia Hargest St Jude Children’s Research Hospital, Memphis, TN, United States and University of Tennessee Health Science Center, Memphis, TN, United States Scott J. Harper Washington State University, Prosser, WA, United States Balázs Harrach Institute for Veterinary Medical Research, Center for Agricultural Research, Budapest, Hungary Masayoshi Hashimoto The University of Tokyo, Tokyo, Japan Muhammad Hassan University of Agriculture, Faisalabad, Pakistan

Duane J. Gubler Duke-NUS Medical School, Singapore, Singapore

Asma Hatoum-Aslan University of Alabama, Tuscaloosa, AL, United States

Peixuan Guo College of Pharmacy, The Ohio State University, Columbus, OH, United States

Philippa C. Hawes The Pirbright Institute, Pirbright, United Kingdom

Tongkun Guo Beijing Institute of Genomics, Chinese Academy of Sciences, Beijing, China Anne-Lise Haenni Institut Jacques Monod, French National Center for Scientific Research, Paris Diderot University, Paris, France Susan L. Hafenstein Pennsylvania State University, Hershey, PA, United States Ahmed Hafez Biotechvana, Valencia, Spain; Pompeu Fabra University, Barcelona, Spain; and Minia University, Minya, Egypt Marie Hagbom Linköping University, Linköping, Sweden

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Janelle A. Hayes University of Massachusetts Medical School, Worcester, MA, United States Guijuan He Virginia Tech, Blacksburg, VA, United States Klaus Hedman University of Helsinki, Helsinki, Finland and Helsinki University Hospital, Helsinki, Finland Albert Heim Hannover Medical School, Hanover, Germany Gary L. Hein University of Nebraska–Lincoln, Lincoln, NE, United States Manfred Heinlein IBMP-CNRS, University of Strasbourg, Strasbourg, France

xxx

List of Contributors

Mercedes Hernando-Pérez National Center for Biotechnology, Spanish National Research Council, Madrid, Spain Carmen Hernández Institute for Plant Molecular and Cell Biology (Spanish National Research Council–Polytechnic University of Valencia), Valencia, Spain Etienne Herrbach University of Strasbourg, Colmar, France Stephen Higgs Biosecurity Research Institute, Kansas State University, Manhattan, KS, United States Bradley I. Hillman Rutgers University, New Brunswick, NJ, United States Deborah M. Hinton National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, United States Judith Hirsch Plant Pathology Unit, INRAE – French National Research Institute for Agriculture, Food and Environment, Montfavet, France Jody Hobson-Peters Australian Infectious Diseases Research Centre, School of Chemistry and Molecular Biosciences, The University of Queensland, St Lucia, QLD, Australia

Elisabeth Huguet Research Institute on Insect Biology, French National Center for Scientific Research, University of Tours, Tours, France Roger Hull John Innes Centre, Norwich, United Kingdom Kiwamu Hyodo Okayama University, Kurashiki, Japan Eugénie Hébrard Interactions Plantes Microorganismes Environnement, Institut de Recherche pour le Développement, Centre de coopération internationale en recherche agronomique pour le développement, University of Montpellier, Montpellier, France Martin Hölzer University of Jena, Jena, Germany Tetsuro Ikegami The University of Texas Medical Branch at Galveston, Galveston, TX, United States Niina Ikonen Finnish Institute for Health and Welfare, Helsinki, Finland Cihan I˙nan Department of Molecular Biology and Genetics, Karadeniz Technical University, Trabzon, Turkey

Natalie M. Holste University of Nebraska–Lincoln, Lincoln, NE, United States

I˙kbal Agah I˙nce Department of Medical Microbiology, Acıbadem University School of Medicine, Istanbul, Turkey

Jin S. Hong Kangwon National University, Chunchon, South Korea

Katsuaki Inoue Diamond Light Source, Didcot, United Kingdom

Margaret J. Hosie MRC-University of Glasgow Centre for Virus Research, Glasgow, United Kingdom

Toru Iwanami Tokyo University of Agriculture, Tokyo, Japan

Olivia G. Howell University of Alabama, Tuscaloosa, AL, United States

Jacques Izopet Toulouse University Hospital, Toulouse, France and Toulouse University Paul Sabatier, Toulouse, France

Liya Hu Baylor College of Medicine, Houston, TX, United States Zhaoyang Hu Jiangsu University, Zhenjiang, China Kuan-Ying A. Huang Chang Gung Memorial Hospital, Taoyuan, Taiwan Yu Huang Peking University, Beijing, China Natalia B. Hubbs Hanover College, Hanover, IN, United States

Fauziah Mohd Jaafar UMR1161 Virologie, INRAE – French National Research Institute for Agriculture, Food and Environment, ANSES, Ecole Nationale Vétérinaire d’Alfort, University of Paris-Est, Maisons-Alfort, France Andrew O. Jackson China Agricultural University, Beijing, China Daral J. Jackwood The Ohio State University/OARDC, Wooster, OH, United States

List of Contributors

Jean-Rock Jacques Cellular and Molecular Epigenetics (GIGA), Liège, Belgium and Molecular Biology (TERRA), Gembloux, Belgium Tiffany Jenkins Nationwide Children’s Hospital, Columbus, OH, United States and The Ohio State University, Columbus, OH, United States Jeffrey D. Jensen Arizona State University, Tempe, AZ, United States Daohong Jiang Huazhong Agricultural University, Wuhan, China Zhihao Jiang China Agricultural University, Beijing, China

xxxi

Laura Kakkola University of Turku, Turku, Finland Hannimari Kallio-Kokko University of Helsinki and Helsinki University Hospital, Helsinki, Finland Nassim Kamar Toulouse University Hospital, Toulouse, France and Toulouse University Paul Sabatier, Toulouse, France Phyllis J. Kanki Harvard T.H. Chan School of Public Health, Boston, MA, United States Peter Karayiannis University of Nicosia, Nicosia, Cyprus

Allison R. Jilbert The University of Adelaide, Adelaide, SA, Australia

Henry M. Kariithi Kenya Agricultural and Livestock Research Organization, Nairobi, Kenya

Peng Jing Department of Chemistry, College of Arts and Sciences, Fort Wayne, IN, United States

Brian A. Kelch University of Massachusetts Medical School, Worcester, MA, United States

Xixi Jing Central South University, Changsha, China

Karen E. Keller Horticultural Crops Research Unit, Agricultural Research Service, US Department of Agriculture, Corvallis, OR, United States

Meesbah Jiwaji Rhodes University, Grahamstown, South Africa Kyle L. Johnson The University of Texas at El Paso, El Paso, TX, United States and CQuentia, Memphis, TN, United States Welkin E. Johnson Boston College, Chestnut Hill, MA, United States Ian M. Jones University of Reading, Reading, United Kingdom and London School of Hygiene and Tropical Medicine, London, United Kingdom Ramon Jordan Agricultural Research Service, US Department of Agriculture, Beltsville, MD, United States Thomas Joris Cellular and Molecular Epigenetics (GIGA), Liège, Belgium and Molecular Biology (TERRA), Gembloux, Belgium Ilkka Julkunen Institute of Biomedicine, University of Turku, Turku, Finland Sandra Junglen Charité - University Medicine Berlin, Berlin, Germany Masanori Kaido Kyoto University, Kyoto, Japan

Japhette E. Kembou-Ringert University of Tel Aviv, Tel Aviv, Israel Peter J. Kerr University of Sydney, Sydney, NSW, Australia and CSIRO Health and Biosecurity, Black Mountain Laboratories, Canberra, ACT, Australia Tiffany King Nationwide Children’s Hospital, Columbus, OH, United States and The Ohio State University College of Medicine, Columbus, OH, United States Andrea Kirmaier Boston College, Chestnut Hill, MA, United States Thomas Klose Purdue University, West Lafayette, IN, United States Barbara G. Klupp Friedrich-Loeffler-Institute, Greifswald-Insel Riems, Germany David M. Knipe Harvard Medical School, Boston, MA, United States Nick J. Knowles The Pirbright Institute, Pirbright, United Kingdom Guus Koch Wageningen Bioveterinary Research, Lelystad, The Netherlands

xxxii

List of Contributors

Renate Koenig Julius Kühn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany Susanne E. Kohalmi The University of Western Ontario, London, ON, Canada Hideki Kondo Okayama University, Kurashiki, Japan Jennifer L. Konopka-Anstadt Centers for Disease Control and Prevention, Atlanta, GA, United States Eugene V. Koonin National Center for Biotechnology Information, National Library of Medicine, Bethesda, MD, United States and National Institutes of Health, Bethesda, MD, United States Marion P.G. Koopmans Erasmus Medical Center, Rotterdam, The Netherlands Richard Kormelink Wageningen University and Research, Wageningen, The Netherlands Ioly Kotta-Loizou Imperial College London, London, United Kingdom Peter J. Krell Department of Molecular and Cellular Biology, University of Guelph, Guelph, ON, Canada Mart Krupovic Archaeal Virology Unit, Institut Pasteur, Paris, France

Manish Kumar Jawaharlal Nehru University, New Delhi, India Gael Kurath US Geological Survey, Western Fisheries Research Center, Seattle, WA, United States Satu Kurkela University of Helsinki and Helsinki University Hospital, Helsinki, Finland Wan-Chun Lai Chang Gung Memorial Hospital, Taoyuan, Taiwan Kevin Lamkiewicz University of Jena, Jena, Germany Rebecca K. Lane University of Texas Health Science Center at San Antonio, San Antonio, TX, United States Andrew S. Lang Memorial University of Newfoundland, St. John’s, NL, Canada Daniel Carlos Ferreira Lanza Federal University of Rio Grande do Norte, Natal, Brazil Maija Lappalainen HUS Diagnostic Center, HUSLAB, Clinical Microbiology, University of Helsinki and Helsinki University Hospital, Helsinki, Finland Katherine LaTourrette University of Nebraska–Lincoln, Lincoln, NE, United States

Andreas Kuhn University of Hohenheim, Stuttgart, Germany

Chris Lauber TWINCORE – Center for Experimental and Clinical Infection Research, Hannover, Germany

Jens H. Kuhn National Institutes of Health, Frederick, MD, United States

Antonio Lavazza The Lombardy and Emilia Romagna Experimental Zootechnic Institute, Brescia, Italy

Richard J. Kuhn Purdue University, West Lafayette, IN, United States

C. Martin Lawrence Montana State University, Bozeman, MT, United States

Suvi Kuivanen University of Helsinki, Helsinki, Finland

Hervé Lecoq Plant Pathology Unit, INRAE – French National Research Institute for Agriculture, Food and Environment, Montfavet, France

Ranjababu Kulasegaram Guy’s and St Thomas’ NHS Foundation Trust, London, United Kingdom Raghavendran Kulasegaran-Shylini Department of Pathogen Infection, Faculty of Infectious and Tropical Diseases, London School of Hygiene and Tropical Medicine, London, United Kingdom Gaurav Kumar University of Delhi, New Delhi, India

Young-Min Lee Utah State University, Logan, UT, United States Kristen N. LeGault University of California, Berkeley, CA, United States James Legg International Institute of Tropical Agriculture, Dar es Salaam, Tanzania

List of Contributors

xxxiii

Anne Legreve University of Louvain, Louvain-la-Neuve, Belgium

Walter Ian Lipkin Columbia University, New York, NY, United States

Petr G. Leiman The University of Texas Medical Branch, Galveston, TX, United States

Jan G. Lisby Copenhagen University Hospital Hvidovre, Hvidovre, Denmark

Stanley M. Lemon Lineberger Comprehensive Cancer Center, The University of North Carolina at Chapel Hill, NC, United States and Department of Microbiology and Immunology, The University of North Carolina at Chapel Hill, NC, United States

Ding X. Liu South China Agricultural University, Guangzhou, China

Sebastian Leptihn Zhejiang University-Edinburgh University Institute, Zhejiang University, Haining, China Dennis J. Lewandowski University of Florida, Lake Alfred, FL, United States Sébastien Lhomme Toulouse University Hospital, Toulouse, France and Toulouse University Paul Sabatier, Toulouse, France Dawei Li China Agricultural University, Beijing, China Guangdi Li Central South University, Changsha, China Guoqing Li Huazhong Agricultural University, Wuhan, China Yi Li Peking University, Beijing, China Zhefeng Li College of Pharmacy, The Ohio State University, Columbus, OH, United States Zhenghe Li Zhejang University, Hangzhou, China

Qiang Liu University of Saskatchewan, Saskatoon, SK, Canada Sijun Liu Iowa State University, Ames, IA, United States Carlos Llorens Biotechvana, Scientific Park University of Valencia, Valencia, Spain L. Sue Loesch-Fries Purdue University, West Lafayette, IN, United States George P. Lomonossoff John Innes Centre, Norwich, United Kingdom L. Letti Lopez The University of Texas at Austin, Austin, TX, United States Alan T. Loynachan University of Kentucky, Lexington, KY, United States Garry A. Luke University of St. Andrews, St. Andrews, United Kingdom M. Luo University of Alabama at Birmingham, Birmingham, AL, United States Juan J. López-Moya Center for Research in Agricultural Genomics and Spanish National Research Council, Barcelona, Spain

Jia Q. Liang South China Agricultural University, Guangzhou, China

Che Ma Genomics Research Center, Academia Sinica, Taipei, Taiwan

Sebastian Liebe Institute of Sugar Beet Research, Göttingen, Germany

Stuart A. MacFarlane The James Hutton Institute, Invergowrie, United Kingdom

João Paulo Matos Santos Lima Federal University of Rio Grande do Norte, Natal, Brazil

Saichetana Macherla J. Craig Venter Institute, La Jolla, CA, United States

Bruno Lina HCL Department of Virology, National Reference Center for Respiratory Viruses, Institute of Infectious Agents, Croix-Rousse Hospital, Lyon, France and Virpath Laboratory, International Center of Research in Infectiology (CIRI), INSERM U1111, CNRS—UMR 5308, École Normale Supérieure de Lyon, University Claude Bernard Lyon, Lyon University, Lyon, France

Kensaku Maejima The University of Tokyo, Tokyo, Japan Fabrizio Maggi University of Pisa, Pisa, Italy and University of Insubria, Varese, Italy Melissa S. Maginnis The University of Maine, Orono, ME, United States

xxxiv

List of Contributors

Edgar Maiss Leibniz University Hannover, Hannover, Germany

Chikara Masuta Hokkaido University, Sapporo, Japan

Kira S. Makarova National Center for Biotechnology Information, National Library of Medicine, Bethesda, MD, United States

Carlos P. Mata University of Leeds, Leeds, United Kingdom

Ariana Manglli Council for Agricultural Research and Economics, Research Center for Plant Protection and Certification, Rome, Italy

Jelle Matthijnssens Rega Institute for Medical Research, KU Leuven, Leuven, Belgium

Annette Mankertz Robert Koch-Institute, Berlin, Germany

Claire P. Mattison Centers for Disease Control and Prevention, Atlanta, GA, United States and Cherokee Nation Assurance, Arlington, VA, United States

Pilar Manrique The Ohio State University, Wexner Medical Center, Columbus, OH, United States

William McAllister Rowan University School of Osteopathic Medicine, Stratford, NJ, United States

Shahid Mansoor National Institute for Biotechnology and Genetic Engineering, Faisalabad, Pakistan

Alison A. McBride National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, United States

Marco Marklewitz Institute of Virology, Charité – University Medicine Berlin, Berlin, Germany Giovanni P. Martelli† University of Bari Aldo Moro, Bari, Italy Darren P. Martin University of Cape Town, Cape Town, South Africa Robert R. Martin Horticultural Crops Research Unit, Agricultural Research Service, US Department of Agriculture, Corvallis, OR, United States Manuel Martinez-Garcia University of Alicante, Alicante, Spain Francisco Martinez-Hernandez University of Alicante, Alicante, Spain Natalia Martín-González Autonomous University of Madrid, Madrid, Spain Joaquín Martínez Martínez Bigelow Laboratory for Ocean Sciences, East Boothbay, ME, United States Manja Marz University of Jena, Jena, Germany Andrea Marzi National Institute of Allergy and Infectious Diseases, National Institutes of Health, Hamilton, MT, United States Hema Masarapu Sri Venkateswara University, Tirupati, India †

Deceased.

Michael McChesney University of California, Davis, CA, United States Elaine McCulloch Quality Control for Molecular Diagnostics (QCMD), Glasgow, United Kingdom Andrew J. McMichael University of Oxford, Oxford, United Kingdom Alexander McPherson University of California, Irvine, CA, United States Irene K. Meki French National Center for Scientific Research, Montpellier, France Ulrich Melcher Oklaoma State University, Stillwater, OK, United States Tomas A Melgarejo University of California, Davis, CA, United States Michael J. Melzer Department of Plant and Environmental Protection Sciences, University of Hawaii, Honolulu, HI, United States Luiza Mendonça University of Oxford, Oxford, United Kingdom Xiang-Jin Meng Virginia Polytechnic Institute and State University, Blacksburg, VA, United States Peter P.C. Mertens University of Nottingham, Sutton Bonington, United Kingdom

List of Contributors

Thomas C. Mettenleiter Friedrich-Loeffler-Institute, Greifswald-Insel Riems, Germany Philip D. Minor St Albans, United Kingdom Ali Mirazimi National Veterinary Institute, Uppsala, Sweden and Karolinska Hospital University, Huddinge, Sweden Nischay Mishra Columbia University, New York, NY, United States Edward S. Mocarski Emory University School of Medicine, Atlanta, GA, United States Florian Mock University of Jena, Jena, Germany Volker Moennig University of Veterinary Medicine, Hannover, Germany Ian J. Molineux The University of Texas at Austin, Austin, TX, United States Aderito L. Monjane Norwegian Veterinary Institute, Oslo, Finland Jacen S. Moore University of Tennessee Health Science Center, Memphis, TN, United States Marc C. Morais The University of Texas Medical Branch, Galveston, TX, United States Cristina Moraru Institute for Chemistry and Biology of the Marine Environment, Oldenburg, Germany Hiromitsu Moriyama Tokyo University of Agriculture and Technology, Tokyo, Japan Sergey Y. Morozov Lomonosov Moscow State University, Moscow, Russia Thomas E. Morrison University of Colorado School of Medicine, Aurora, CO, United States Léa Morvan University of Liège, Liège, Belgium Benoît Moury Plant Pathology Unit, INRAE – French National Institute for Agriculture, Food and Environment, Montfavet, France

xxxv

Muhammad Mubin University of Agriculture, Faisalabad, Pakistan Nicolas J. Mueller University Hospital of Zurich, Zurich, Switzerland Emmanuelle Muller The French Agricultural Research Center for International Development, Joint Research Units–Biology and Genetics of Plant-Pathogen Interactions, Montpellier, France and Biology and Genetics of PlantPathogen Interactions, University of Montpellier, The French Agricultural Research Center for International Development, French National Institute for Agricultural Research, Montpellier SupAgro, Montpellier, France John S. Munday Massey University, Palmerston North, New Zealand Jacob H. Munson-McGee Montana State University, Bozeman, MT, United States and Bigelow Laboratory for Ocean Sciences, East Boothbay, ME, United States Hacer Muratoğlu Department of Molecular Biology and Genetics, Karadeniz Technical University, Trabzon, Turkey Kenan C. Murphy University of Massachusetts Medical School, Worcester, MA, United States Ugrappa Nagalakshmi University of California, Davis, CA, United States Keizo Nagasaki Kochi University, Nankoku, Japan Nazia Nahid GC University, Faisalabad, Pakistan and University of Agriculture, Faisalabad, Pakistan Venugopal Nair The Pirbright Institute, Pirbright, United Kingdom Remziye Nalçacıoğlu Department of Molecular Biology and Genetics, Karadeniz Technical University, Trabzon, Turkey Shigetou Namba The University of Tokyo, Tokyo, Japan Rubab Z. Naqvi National Institute for Biotechnology and Genetic Engineering, Faisalabad, Pakistan Rachel Nash The Pirbright Institute, Surrey, United Kingdom C.K. Navaratnarajah Purdue University, West Lafayette, IN, United States

xxxvi

List of Contributors

Maria A. Navarrete-Muñoz Biotechvana, Madrid, Spain; Institute of Health Research-Jiménez Díaz Foundation, Autonomous University of Madrid; and Rey Juan Carlos University Hospital, Móstoles, Spain Jesús Navas-Castillo Institute for Mediterranean and Subtropical Horticulture “La Mayora”–Spanish National Research Council– University of Malaga, Algarrobo-Costa, Málaga, Spain Muhammad S. Nawaz-ul-Rehman University of Agriculture, Faisalabad, Pakistan Christopher L. Netherton The Pirbright Institute, Pirbright, United Kingdom Thu V.P. Nguyen Baylor College of Medicine, Houston, TX, United States Annette Niehl Julius Kühn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany Hubert G.M. Niesters Department of Medical Microbiology and Infection Prevention, Division of Clinical Virology, University Medical Center Groningen, Groningen, The Netherlands Jozef I. Nissimov University of Waterloo, Waterloo, ON, Canada Norman Noah London School of Hygiene and Tropical Medicine, London, United Kingdom Mauricio L. Nogueira São José do Rio Preto School of Medicine, São José do Rio Preto, São Paulo, Brazil Johan Nordgren Linköping University, Linköping, Sweden C. Micha Nübling Paul-Ehrlich-Institute, Langen, Germany Visa Nurmi University of Helsinki, Helsinki, Finland

Hanna M. Oksanen Molecular and Integrative Biosciences Research Program, Faculty of Biological and Environmental Sciences, University of Helsinki, Helsinki, Finland Graziele P. Oliveira Federal University of Minas Gerais, Belo Horizonte, Brazil Francesco Origgi University of Bern, Bern, Switzerland Nikolaus Osterrieder Free University of Berlin, Berlin, Germany Robert A. Owens Beltsville Agricultural Research Center, Beltsville, MD 20705, United States Emine Özsahin University of Guelph, Guelph, ON, Canada Sergi Padilla-Parra University of Oxford, Oxford, United Kingdom; Department of Infectious Diseases, Faculty of Life Sciences and Medicine, King’s College London, London, United Kingdom; and Randall Division of Cell and Molecular Biophysics, King’s College London, London, United Kingdom Joshua Pajak Duke University, Durham, NC, United States Massimo Palmarini MRC-University of Glasgow Centre for Virus Research, Glasgow, United Kingdom Amanda R. Panfil The Ohio State University, Columbus, OH, United States Marcus Panning Institute of Virology, Freiburg University Medical Center, Faculty of Medicine, University of Freiburg, Freiburg, Germany

Donald L. Nuss University of Maryland, Rockville, MD, United States

Vitantonio Pantaleo National Research Council, Research Unit of Bari, Bari, Italy

M. Steven Oberste Centers for Disease Control and Prevention, Atlanta, GA, United States

Anna Papa Aristotle University of Thessaloniki, Thessaloniki, Greece

Hiroyuki Ogata Institute for Chemical Research, Kyoto University, Kyoto, Japan Ane Ogbe University of Oxford, Oxford, United Kingdom

Nikolaos Pappas Utrecht University, Utrecht, The Netherlands Hanu R. Pappu Washington State University, Pullman, WA, United States

List of Contributors

xxxvii

Kristin N. Parent Michigan State University, East Lansing, MI, United States

Jean-Marie Peron Toulouse University Hospital, Toulouse, France and Toulouse University Paul Sabatier, Toulouse, France

Colin R. Parrish Cornell University, Ithaca, NY, United States

Karin E. Peterson National Institutes of Health, Hamilton, MT, United States

A. Lorena Passarelli Kansas State University, Manhattan, KS, United States

Karel Petrzik Biology Center CAS, Institute of Plant Molecular Biology, České Budějovice, Czech Republic

Basavaprabhu L. Patil ICAR–Indian Institute of Horticultural Research, Bengaluru, India Jade Pattyn University of Antwerp, Antwerp, Belgium T.A. Paul Cornell University, Ithaca, NY, United States Lillian Pavlik Laboratory for Molecular Virology, Great Lakes Forestry Centre, Sault Ste Marie, ON, Canada Susan L. Payne Texas A& M University, College Station, TX, United States Michael N. Pearson The University of Auckland, Auckland, New Zealand Mark E. Peeples Nationwide Children’s Hospital, Columbus, OH, United States and The Ohio State University College of Medicine, Columbus, OH, United States Ben Peeters Wageningen Bioveterinary Research, Lelystad, The Netherlands Joseph S.M. Peiris The University of Hong Kong, Pok Fu Lam, Hong Kong Malik Peiris The University of Hong Kong, Pok Fu Lam, Hong Kong Judit J. Pénzes National Institute of Scientific Research – ArmandFrappier Health Research Centre, Laval, QC, Canada Miryam Pérez-Cañamás Institute for Plant Molecular and Cell Biology (Spanish National Research Council–Polytechnic University of Valencia), Valencia, Spain

Mahtab Peyambari Pennsylvania State University, State College, PA, United States Sujal Phadke J. Craig Venter Institute, La Jolla, CA, United States Hanh T. Pham National Institute of Scientific Research – ArmandFrappier Health Research Centre, Laval, QC, Canada Mauro Pistello University of Pisa, Pisa, Italy Daniel Ponndorf John Innes Centre, Norwich, United Kingdom Leo L.M. Poon The University of Hong Kong, Pok Fu Lam, Hong Kong Welkin H. Pope University of Pittsburgh, Pittsburgh, PA, United States Minna M. Poranen University of Helsinki, Helsinki, Finland Claudine Porta The Pirbright Institute, Pirbright, United Kingdom and University of Oxford, Oxford, United Kingdom Samuel S. Porter National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD, United States and University of Maryland, College Park, MD, United States Frank A. Post King's College Hospital NHS Foundation Trust, London, United Kingdom

Marta Pérez-Illana National Center for Biotechnology, Spanish National Research Council, Madrid, Spain

Nils Poulicard Interactions Plantes Microorganismes Environnement, Institut de Recherche pour le Développement, Centre de coopération internationale en recherche agronomique pour le développement, University of Montpellier, Montpellier, France

Jaume Pérez-Sánchez Institute of Aquaculture Torre de la Sal, Spanish National Research Council, Castellon, Spain

David Prangishvili Institut Pasteur, Paris, France and Ivane Javakhishvili Tbilisi State University, Tbilisi, Georgia

xxxviii

List of Contributors

B. V. Venkataram Prasad Baylor College of Medicine, Houston, TX, United States Lalita Priyamvada Centers for Disease Control and Prevention, Atlanta, GA, United States

Chris M. Rands University of Geneva Medical School and Swiss Institute of Bioinformatics, Geneva, Switzerland Venigalla B. Rao The Catholic University of America, Washington, DC, United States

Simone Prospero Swiss Federal Institute for Forest, Snow and Landscape Research WSL, Birmensdorf, Switzerland

Janne J. Ravantti University of Helsinki, Helsinki, Finland

Elisabeth Puchhammer-Stöckl Medical University of Vienna, Vienna, Austria

Mandy Ravensbergen Wageningen University and Research, Wageningen, The Netherlands

Jianming Qiu University of Kansas Medical Center, Kansas City, KS, United States

Georget Y. Reaiche-Miller The University of Adelaide, Adelaide, SA, Australia

S.L. Quackenbush Colorado State University, Fort Collins, CO, United States

D.V.R. Reddy International Crops Research Institute for the Semi-Arid Tropics, Hyderabad, India

Killian J. Quinn King’s College Hospital, London, United Kingdom

Vishwanatha R.A.P. Reddy The Pirbright Institute, Pirbright, United Kingdom

Diego F. Quito-Avila Department of Life Sciences, ESPOL Polytechnic University, Guayaquil, Ecuador

Juan Reguera Aix-Marseille University, French National Center for Scientific Research, Marseille, France and French National Institute of Health and Medical Research, Marseille, France

Frank Rabenstein Julius Kühn Institute, Quedlinburg, Germany Sheli R. Radoshitzky United States Army Medical Research Institute of Infectious Diseases, Frederick, MD, United States

William K. Reisen University of California, Davis, CA, United states Jingshan Ren University of Oxford, Oxford, United Kingdom

Saleem U. Rahman National Institute for Biotechnology and Genetic Engineering, Faisalabad, Pakistan

Renato O. Resende University of Brasilia, Brasilia, Brazil

Mbolarinosy Rakotomalala FOFIFA, Antananarivo, Madagascar

Peter A. Revill The Peter Doherty Institute of Infection and Immunity, Royal Melbourne Hospital, Melbourne, VIC, Australia

Norma Rallon Institute of Health Research-Jiménez Díaz Foundation, Autonomous University of Madrid and Rey Juan Carlos University Hospital, Móstoles, Spain

Félix A. Rey Institut Pasteur, Paris, France

Robert P. Rambo Diamond Light Source, Didcot, United Kingdom

Simone G. Ribeiro Embrapa Genetic Resources and Biotechnology, Brasília, Brazil

Bertha Cecilia Ramirez The Institute for Integrative Biology of the Cell, The French Alternative Energies and Atomic Energy Commission, French National Center for Scientific Research, University of Paris-Sud, University of Paris-Saclay, Gif-sur-Yvette, France María D. Ramos-Barbero University of Alicante, Alicante, Spain

Lara Rheinemann University of Utah, Salt Lake City, UT, United States

Daniel Rigling Swiss Federal Institute for Forest, Snow and Landscape Research (WSL), Birmensdorf, Switzerland Cristina Risco Cell Structure Laboratory, National Center for Biotechnology – Spanish National Research Council (CNB-CSIC), Madrid, Spain

List of Contributors

Efraín E. Rivera-Serrano Lineberger Comprehensive Cancer Center, The University of North Carolina at Chapel Hill, NC, United States and Department of Microbiology and Immunology, The University of North Carolina at Chapel Hill, NC, United States Cécile Robin INRAE – French National Research Institute for Agriculture, Food and Environment, UMR BIOGECO, Cestas, France Rodrigo A.L. Rodrigues Federal University of Minas Gerais, Belo Horizonte, Brazil Elina Roine University of Helsinki, Helsinki, Finland

xxxix

Polly Roy Department of Pathogen Infection, Faculty of Infectious and Tropical Diseases, London School of Hygiene and Tropical Medicine, London, United Kingdom and University of Reading, Reading, United Kingdom Aaron P. Roznowski The University of Texas at Austin, Austin, TX, United States and University of Arizona, Tucson, AZ, United States Luisa Rubino Institute for Sustainable Plant Protection, National Research Council, Bari, Italy Olli Ruuskanen Turku University Hospital, Turku, Finland

Maria R. Rojas University of California, Davis, CA, United States

Eugene V. Ryabov USDA, Agricultural Research Service, Beltsville, MD, United States

Marilyn J. Roossinck Pennsylvania State University, State College, PA, United States

Martin D. Ryan University of St. Andrews, St. Andrews, United Kingdom

Vera I.D. Ros Wageningen University and Research, Wageningen, The Netherlands

Ki H. Ryu Seoul Women’s University, Seoul, South Korea

Cristina Rosa Pennsylvania State University, University Park, PA, United States Hanna Rose Leibniz University Hannover, Hannover, Germany David A. Rosenbaum University of Florida, Gainesville, FL, United States Shannan L. Rossi The University of Texas Medical Branch, Galveston, TX, United States Michael G. Rossmann† Purdue University, West Lafayette, IN, United States

Hanns-Joachim Rziha Eberhard Karls University of Tübingen, Tübingen, Germany Sead Sabanadzovic Mississippi State University, Starkville, MS, United States Roghaiyeh Safari Cellular and Molecular Epigenetics (GIGA), Liège, Belgium and Molecular Biology (TERRA), Gembloux, Belgium Azeez Sait Sahul Hameed C. Abdul Hakeem College, Melvisharam, India

L. Roux University of Geneva Medical School, Geneva, Switzerland

Nicole Samies University of Alabama at Birmingham, Birmingham, AL, United States

Simon Roux US Department of Energy Joint Genome Institute, Walnut Creek, CA, United States

Carmen San Martín National Center for Biotechnology, Spanish National Research Council, Madrid, Spain

J. Rovnak Colorado State University, Fort Collins, CO, United States David J. Rowlands University of Leeds, Leeds, United Kingdom †

Deceased.

Ruth-Anne Sandaa Department of Biological Sciences, University of Bergen, Bergen, Norway Hélène Sanfaçon Agriculture and Agri-Food Canada, Summerland, BC, Canada

xl

List of Contributors

Rafael Sanjuán Institute for Integrative Systems Biology (I2SysBio), University of Valencia-CSIC, Valencia, Spain

Declan C. Schroeder University of Reading, Reading, United Kingdom and University of Minnesota, St. Paul, MN, United States

Neeraja Sankaran Utrecht University, Utrecht, The Netherlands

Stacey Schultz-Cherry St. Jude Children’s Research Hospital, Memphis, TN, United States

Fernando Santos University of Alicante, Alicante, Spain Cecilia Sarmiento Tallinn University of Technology, Tallinn, Estonia Takahide Sasaya National Agriculture and Food Research Organization, Fukuyama, Japan Preethi Sathanantham Virginia Tech, Blacksburg, VA, United States Panayampalli S. Satheshkumar Centers for Disease Control and Prevention, Atlanta, GA, United States Yukiyo Sato Okayama University, Kurashiki, Japan Andreas Sauerbrei Jena University Hospital, Jena, Germany Eugene I. Savenkov Swedish University of Agricultural Sciences, Uppsala, Sweden and Linnean Center for Plant Biology, Uppsala, Sweden Carita Savolainen-Kopra National Institute for Health and Welfare, Helsinki, Finland Kay Scheets Oklahoma State University, Stillwater, OK, United States Uffe V. Schenider Copenhagen University Hospital Hvidovre, Hvidovre, Denmark Richard H. Scheuermann J. Craig Venter Institute, La Jolla, CA, United States; University of California, San Diego, CA, United States; La Jolla Institute for Immunology, La Jolla, CA, United States; and Global Virus Network, Baltimore, MD, United States Manfred J. Schmitt Saarland University, Saarbrücken, Germany James E. Schoelz University of Missouri, Columbia, MO, United States Jason R. Schrad Michigan State University, East Lansing, MI, United States

Thomas F. Schulz Hannover Medical School, Institute of Virology, Hannover, Germany and German Center for Infection Research, Hannover-Braunschweig Site, Braunschweig, Germany Catherine A. Scougall The University of Adelaide, Adelaide, SA, Australia Kimberley D. Seed University of California, Berkeley, CA, United States Joaquim Segalés Departament of Animal Health and Anatomy, Faculty of Veterinary Medicine, Autonomous University of Barcelona, Barcelona, Spain; Animal Health Research Center (CReSA) – Institute of Agrifood Research and Technology (IRTA), Campus UAB, Barcelona, Spain; and OIE Collaborating Center for the Research and Control of Emerging and Re-emerging Swine Diseases in Europe (IRTA-CReSA), Barcelona, Spain Mateo Seoane-Blanco National Center for Biotechnology, Madrid, Spain Madhumati Sevvana Purdue University, West Lafayette, IN, United States Kazım Sezen Department of Biology, Karadeniz Technical University, Trabzon, Turkey Arvind Sharma Institut Pasteur, Paris, France Sumit Sharma Linköping University, Linköping, Sweden James M. Sharp University of Zaragoza, Zaragoza, Spain and Edinburgh, United Kingdom Qunxin She Shandong University, Qingdao, China Keith E. Shearwin The University of Adelaide, Adelaide, SA, Australia Hanako Shimura Hokkaido University, Sapporo, Japan Reina S. Sikkema Erasmus Medical Center, Rotterdam, The Netherlands

List of Contributors

Aaron Simkovich Agriculture and Agri-Food Canada, London, ON, Canada and The University of Western Ontario, London, ON, Canada Peter Simmonds University of Oxford, Oxford, United Kingdom Tarja Sironen University of Helsinki, Helsinki, Finland Susanna Sissonen Finnish Institute for Health and Welfare, Helsinki, Finland Michael A. Skinner Imperial College London, London, United Kingdom Douglas E. Smith University of California, San Diego, La Jolla, CA, United States Melvyn Smith Viapath Analytics, Specialist Virology Centre, King’s College NHS Foundation Trust, London, United Kingdom Thomas J. Smith The University of Texas Medical Branch, Galveston, TX, United States Teemu Smura Helsinki University Hospital and University of Helsinki, Helsinki, Finland Eric J. Snijder Leiden University Medical Center, Leiden, The Netherlands Gisela Soboll Hussey Michigan State University, East Lansing, MI, United States Maria Söderlund-Venermo University of Helsinki, Helsinki, Finland Merike Sõmera Tallinn University of Technology, Tallinn, Estonia Eun G. Song Seoul Women’s University, Seoul, South Korea Milan J. Sonneveld Erasmus University Medical Center, Rotterdam, The Netherlands Beatriz Soriano Biotechvana, Scientific Park University of Valencia and Institute for Integrative Systems Biology (I2SysBio), University of Valencia–Spanish National Research Council, Valencia, Spain

xli

Thomas E. Spencer University of Missouri, Columbia, MO, United States Pothur Sreenivasulu Sri Venkateswara University, Tirupati, India Ashley L. St. John Duke-NUS Medical School, Singapore, Singapore David K. Stammers University of Oxford, Oxford, United Kingdom John Stanley John Innes Centre, Colney, United Kingdom Glyn Stanway University of Essex, Colchester, United Kingdom Thilo Stehle University of Tuebingen, Tuebingen, Germany and Vanderbilt University School of Medicine, Nashville, TN, United States Gregory W. Stevenson Iowa State University, Ames, IA, United States Lucy Rae Stewart Agricultural Research Service, US Department of Agriculture, Wooster, OH, United States C.C.M.M. Stijger Wageningen University and Research Center, Bleiswijk, The Netherlands Peter G. Stockley University of Leeds, Leeds, United Kingdom David Stone Weymouth Laboratory, Centre for Environment, Fisheries and Aquaculture Science, Weymouth, United Kingdom Ashley E Strother The University of Texas Medical Branch, Galveston, TX, United States Sundharraman Subramanian Michigan State University, East Lansing, MI, United States William C. Summers Yale University, New Haven, CT, United States Liying Sun Northwest A& F University, Yangling, China Wesley I. Sundquist University of Utah, Salt Lake City, UT, United States Petri Susi University of Turku, Turku, Finland Curtis A. Suttle University of British Columbia, Vancouver, BC, Canada

xlii

List of Contributors

Nobuhiro Suzuki Institute of Plant Stress and Resources (IPSR), Okayama University, Kurashiki, Japan Lennart Svensson Linköping University, Linköping, Sweden and Karolinska Institute, Stockholm, Sweden Ronald Swanstrom University of North Carolina at Chapel Hill, Chapel Hill, NC, United States

Nicholas M.I. Taylor University of Copenhagen, Copenhagen, Denmark Xu Tengzhi University of California, Davis, CA, United States Raquel Tenorio Cell Structure Laboratory, National Center for Biotechnology – Spanish National Research Council (CNB-CSIC), Madrid, Spain

Daniele M. Swetnam University of California, Davis, CA, United states

Robert B. Tesh The University of Texas Medical Branch, Galveston, TX, United States

Moriah L. Szpara Pennsylvania State University, University Park, PA, United States

Vaskar Thapa Pennsylvania State University, State College, PA, United States

Keisuke Tabata Heidelberg University, Heidelberg, Germany

John E. Thomas The University of Queensland, Brisbane, QLD, Australia

Anna Taglienti Council for Agricultural Research and Economics, Research Center for Plant Protection and Certification, Rome, Italy

Julie A. Thomas Rochester Institute of Technology, Rochester, NY, United States

Naoki Takeshita Tokyo University of Agriculture and Technology, Fuchu, Japan Kana Takeshita Urayama Tokyo University of Agriculture and Technology, Fuchu, Japan Michael E. Taliansky The James Hutton Institute, Dundee, United Kingdom Pan Tao The Catholic University of America, Washington, DC, United States

Lynn C. Thomason Frederick National Laboratory for Cancer Research, Frederick, MD, United States Elizabeth Ashley Thompson The University of Southern Mississippi, Hattiesburg, MS, United States Jeremy R. Thompson Cornell University, Ithaca, NY, United States Antonio Tiberini Council for Agricultural Research and Economics, Research Center for Plant Protection and Certification, Rome, Italy

Jacqueline E. Tate Centers for Disease Control and Prevention, Atlanta, GA, United States

Peter Tijssen National Institute of Scientific Research – ArmandFrappier Health Research Centre, Microbiology and Immunology, Laval, QC, Canada

Satyanarayana Tatineni Agricultural Research Service, US Department of Agriculture, Lincoln, NE, United States and University of Nebraska–Lincoln, Lincoln, NE, United States

Yuji Tomaru Japan Fisheries Research and Education Agency, Kanagawa, Japan

Sisko Tauriainen University of Turku, Turku, Finland Norbert Tautz University of Luebeck, Luebeck, Germany Paulo Tavares Institute for Integrative Biology of the Cell, CEA, CNRS, University of Paris-Sud, University of Paris-Saclay, Gif-sur-Yvette, France

Laura Tomassoli Council for Agricultural Research and Economics, Research Center for Plant Protection and Certification, Rome, Italy Ruben Torres National Biotechnology Center–Spanish National Research Council, Madrid, Spain Jia Q. Truong The University of Adelaide, Adelaide, SA, Australia

List of Contributors

Erkki Truve† Tallinn University of Technology, Tallinn, Estonia Chih-Hsuan Tsai Institute of Molecular Biology, Academia Sinica, Taipei, Taiwan Roman Tuma University of Leeds, Leeds, United Kingdom and University of South Bohemia, České Budějovice, Czech Republic Topi Turunen Infectious Disease Unit, Espoo, Finland and Finnish Institute for Health and Welfare, Helsinki, Finland Reidun Twarock University of York, York, United Kingdom Ioannis E. Tzanetakis University of Arkansas, Fayetteville, United States Antti Vaheri University of Helsinki, Helsinki, Finland Eeva J. Vainio Natural Resources Institute Finland (Luke), Helsinki, Finland Anna M. Vaira Institute for Sustainable Plant Protection, National Research Council of Italy, Torino, Italy Steven M. Valles Center for Medical, Agricultural and Veterinary Entomology, Agricultural Research Service, US Department of Agriculture, Gainesville, FL, United States Adrián Valli National Center for Biotechnology-Spanish National Research Council, Madrid, Spain Rodrigo A. Valverde Louisiana State University Agricultural Center, Baton Rouge, United States Pierre Van Damme University of Antwerp, Antwerp, Belgium Rene A.A. van der Vlugt Wageningen University and Research Center, Wageningen, The Netherlands Bernard A.M. Van der Zeijst Leiden University Medical Center, Leiden, The Netherlands Koenraad Van Doorslaer University of Arizona, Tucson, AZ, United States †

Deceased.

xliii

James L. Van Etten University of Nebraska–Lincoln, Lincoln, NE, United States Suzanne van Meer University Medical Center Utrecht, Utrecht, The Netherlands Monique M. van Oers Wageningen University and Research, Wageningen, The Netherlands Mark J. van Raaij National Center for Biotechnology, Madrid, Spain Marc H.V. Van Regenmortel University of Strasbourg, Strasbourg, France Piet A. van Rijn Wageningen Bioveterinary Research, Lelystad, The Netherlands and North-West University, Potchefstroom, South Africa Alain Vanderplasschen University of Liège, Liège, Belgium Dana L. Vanlandingham College of Veterinary Medicine, Kansas State University, Manhattan, KS, United States Olli Vapalahti Helsinki University Hospital and University of Helsinki, Helsinki, Finland Mark Varrelmann Institute of Sugar Beet Research, Göttingen, Germany Nikos Vasilakis The University of Texas Medical Branch, Galveston, TX, United States Michael Veit Free University of Berlin, Berlin, Germany Česlovas Venclovas Vilnius University, Vilnius, Lithuania H. Josef Vetten Julius Kühn Institute, Braunschweig, Germany Marli Vlok University of British Columbia, Vancouver, BC, Canada Anne-Nathalie Volkoff Diversity, Genomes and Insects-Microorganisms Interactions, National Institute of Agricultural Research, University of Montpellier, Montpellier, France Ian E.H. Voorhees Cornell University, Ithaca, NY, United States Alex Vorsters University of Antwerp, Antwerp, Belgium

xliv

List of Contributors

Jonathan D.F. Wadsworth UCL Institute of Prion Diseases, London, United Kingdom

Kerstin Wernike Friedrich-Loeffler-Institute, Insel Riems, Germany

Peter J. Walker The University of Queensland, St. Lucia, QLD, Australia

Rachel J. Whitaker University of Illinois at Urbana-Champaign, Urbana, IL, United States

Paul Wallace Quality Control for Molecular Diagnostics (QCMD), Glasgow, United Kingdom

K. Andrew White York University, Toronto, ON, Canada

Aiming Wang Agriculture and Agri-Food Canada, London, ON, Canada

Anna E. Whitfield North Carolina State University, Raleigh, NC, United States

Jen-Ren Wang National Cheng Kung University, Tainan, Taiwan

Richard Whitley University of Alabama at Birmingham, Birmingham, AL, United States

Lin-Fa Wang Duke-NUS Medical School, Singapore, Singapore Nan Wang Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Xiangxi Wang Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Xiaofeng Wang Virginia Tech, Blacksburg, VA, United States Katherine N. Ward University College London, London, United Kingdom Matti Waris University of Turku, Turku, Finland Ranjit Warrier Purdue University, West Lafayette, IN, United States Daniel Watterson Australian Infectious Diseases Research Centre, School of Chemistry and Molecular Biosciences, The University of Queensland, St Lucia, QLD, Australia Marta L. Wayne University of Florida, Gainesville, FL, United States

Reed B. Wickner National Institute of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, MD, United States Luc Willems Cellular and Molecular Epigenetics (GIGA), Liège, Belgium and Molecular Biology (TERRA), Gembloux, Belgium Brian J. Willett MRC-University of Glasgow Centre for Virus Research, Glasgow, United Kindom Alexis Williams The University of Texas Medical Branch, Galveston, TX, United States Stephen A. Winchester Frimley Park Hospital, Frimley, United Kingdom and Immunisation and Countermeasures Division, Public Health England, London, United Kingdom Clayton W. Winkler National Institutes of Health, Hamilton, MT, United States

Friedemann Weber FB 10 – Institute for Virology, Justus Liebig University Giessen, Giessen, Germany

Stephan Winter Leibniz Institute – DSMZ – German Collection of Microorganisms and Cell Cultures GmbH, Braunschweig, Germany

Sung-Chan Wei Institute of Molecular Biology, Academia Sinica, Taipei, Taiwan

William M. Wintermantel Agricultural Research Service, US Department of Agriculture, Salinas, CA, United States

Robin A. Weiss University College London, London, United Kingdom

Jennifer Wirth Montana State University, Bozeman, MT, United States

Tao Weitao Southwest Baptist University, Bolivar, MO, United States

Yuri I. Wolf National Center for Biotechnology Information, National Library of Medicine, Bethesda, MD, United States

List of Contributors

xlv

Thorsten Wolff Robert Koch Institute, Berlin, Germany

Lawrence S. Young University of Warwick, Coventry, United Kingdom

Blaide Woodburn University of North Carolina at Chapel Hill, Chapel Hill, NC, United States

Mark J. Young Montana State University, Bozeman, MT, United States

Michael E. Woodson The University of Texas Medical Branch, Galveston, TX, United States Courtney Woolsey The University of Texas Medical Branch, Galveston, TX, United States Chien-Fu Wu Tokyo University of Agriculture and Technology, Fuchu, Japan Mingde Wu Huazhong Agricultural University, Wuhan, China Songsong Wu Huazhong Agricultural University, Wuhan, China Yan Xiang University of Texas Health Science Center at San Antonio, San Antonio, TX, United States Jiatao Xie Huazhong Agricultural University, Wuhan, China Zhuang Xiong Beijing Institute of Genomics, Chinese Academy of Sciences, Beijing, China Hajime. Yaegashi Institute of Fruit Tree and Tea Science, NARO, Morioka, Japan Mehtap Yakupoğlu Trabzon University, Trabzon, Turkey Yasuyuki Yamaji The University of Tokyo, Tokyo, Japan Meng Yang China Agricultural University, Beijing, China Teng-Chieh Yang Scarsdale, NY, United States Qin Yao Jiangsu University, Zhenjiang, China Tianyou Yao Baylor College of Medicine, Houston, TX, United States Nobuyuki Yoshikawa Iwate University, Morioka, Japan George R. Young Francis Crick Institute, London, United Kingdom

Ry Young Texas A& M University, College Station, TX, United States Isaac T. Younker University of Alabama, Tuscaloosa, AL, United States Qian Yu School of Life Sciences, Jiangsu University, Zhenjiang, China Sang-Im Yun Utah State University, Logan, UT, United States Fauzia Zarreen University of Delhi, New Delhi, India Francisco M. Zerbini Federal University of Viçosa, Viçosa, Brazil Dong-Xiu Zhang University of Maryland, Rockville, MD, United States Jianqiang Zhang Iowa State University, Ames, IA, United States Junjie Zhang Texas A& M University, College Station, TX, United States Long Zhang College of Pharmacy, The Ohio State University, Columbus, OH, United States Pan Zhang Central South University, Changsha, China Peijun Zhang University of Oxford, Oxford, United Kingdom and Electron Bio-Imaging Centre, Diamond Light Source, Didcot, United Kingdom Tao Zhang Beijing Institute of Genomics, Chinese Academy of Sciences, Beijing, China Yongliang Zhang China Agricultural University, Beijing, China Zhenlu Zhang Shandong Agricultural University, Tai’an, China Lixia Zhou College of Pharmacy, The Ohio State University, Columbus, OH, United States

xlvi

List of Contributors

Ling Zhu Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Heiko Ziebell Julius Kühn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany John Ziebuhr The Queen's University of Belfast, Belfast, United Kingdom

Jeffrey J. Zimmerman Iowa State University, Ames, IA, United States Falk Zucker Institute for Chemistry and Biology of the Marine Environment, Oldenburg, Germany

CONTENT OF ALL VOLUMES Editors in Chief

v

Editorial Board

vii

Section Editors

ix

Foreword

xv

Preface

xvii

Guide to Use

xix

List of Contributors

xxi

VOLUME 1 The Virus as a Concept – Fundamentals of Virology A Brief History of Virology David J Rowlands

3

The Origin of Viruses Patrick Forterre and Morgan Gaïa

14

The Virocell Concept Patrick Forterre

23

Virus Taxonomy Jens H Kuhn

28

The Greater Virus World and Its Evolution Eugene V Koonin and Valerian V Dolja

38

The Virus Species Concept Peter Simmonds

47

Genetic Diversity and Evolution of Viral Populations Rafael Sanjuán and Pilar Domingo-Calap

53

Mechanisms of RNA Virus Evolution Lisa M Bono and Siobain Duffy

62

Mechanisms of DNA Virus Evolution Moriah L Szpara and Koenraad Van Doorslaer

71

Paleovirology Clément Gilbert

79

Evolution Steered by Structure Nicola GA Abrescia

87

xlvii

xlviii

Content of all Volumes

Pairwise Sequence Comparison in Virology Tao Zhang, Zheng Gong, Tongkun Guo, Zhuang Xiong, and Yiming Bao

100

Computational Analysis of Recombination in Viral Nucleotide Sequences Miguel Arenas

108

Phylogeny of Viruses Alexander E Gorbalenya and Chris Lauber

116

Virus Bioinformatics Nikolaos Pappas, Simon Roux, Martin Hölzer, Kevin Lamkiewicz, Florian Mock, Manja Marz, and Bas E Dutilh

124

Metagenomics in Virology Simon Roux, Jelle Matthijnssens, and Bas E Dutilh

133

Database and Analytical Resources for Viral Research Community Sujal Phadke, Saichetana Macherla, and Richard H Scheuermann

141

Classification of the Viral World Based on Atomic Level Structures Janne J Ravantti and Nicola GA Abrescia

153

Isolating, Culturing, and Purifying Viruses With a Focus on Bacterial and Archaeal Viruses Katri Eskelin and Hanna M Oksanen

162

High Throughput Sequencing and Virology Graham L Freimanis and Nick J Knowles

175

Single-Virus Genomics: Studying Uncultured Viruses, One at a Time Manuel Martinez-Garcia, Francisco Martinez-Hernandez, and Joaquín Martínez Martínez

184

Biophysical Characterizations in the Solution State Robert P Rambo and Katsuaki Inoue

191

Virus Crystallography Jonathan M Grimes

199

Advanced Light and Correlative Microscopy in Virology Sergi Padilla-Parra, Charles A Coomer, and Irene Carlon-Andres

208

Atomic Force Microscopy (AFM) Investigation of Viruses Alexander McPherson

218

Cryo-Electron Microscopy (CEM) Structures of Viruses David Chmielewski and Wah Chiu

233

Analysis of Viruses in the Cellular Context by Electron Tomography Peijun Zhang and Luiza Mendonça

242

Mathematical Modeling of Virus Architecture Reidun Twarock

248

Principles of Virus Structure Madhumati Sevvana, Thomas Klose, and Michael G Rossmann†

257

Structures of Small Icosahedral Viruses Elizabeth E Fry, Jingshan Ren, and Claudine Porta

278

Structural Principles of the Flavivirus Particle Organization and of Its Conformational Changes Stéphane Duquerroy, Arvind Sharma, and Félix A Rey

290

Reoviruses (Reoviridae) and Their Structural Relatives Liya Hu, Mary K Estes, and B V Venkataram Prasad

303

†

Deceased.

Content of all Volumes

xlix

Structures of Tailed Phages and Herpesviruses (Herpesviridae) Montserrat Fàbrega-Ferrer and Miquel Coll

318

Adenoviruses (Adenoviridae) and Their Structural Relatives Gabriela N Condezo, Natalia Martín-González, Marta Pérez-Illana, Mercedes Hernando-Pérez, José Gallardo, and Carmen San Martín

329

Negative Single-Stranded RNA Viruses (Mononegavirales): A Structural View Juan Reguera

345

Structure of Retrovirus Particles (Retroviridae) David K Stammers and Jingshan Ren

352

Structure of Helical Viruses C Martin Lawrence

362

Giant Viruses and Their Virophage Parasites Rodrigo AL Rodrigues, Ana CdSP Andrade, Graziele P Oliveira, and Jônatas S Abrahão

372

Viral Replication Cycle AJ Cann

382

Viral Receptors José M Casasnovas and Thilo Stehle

388

Bacterial and Archeal Virus Entry Minna M Poranen and Aušra Domanska

402

Nonenveloped Eukaryotic Virus Entry Ian M Jones and Polly Roy

409

Enveloped Virus Membrane Fusion Aurélie A Albertini and Yves Gaudin

417

Genome Replication of Bacterial and Archaeal Viruses Česlovas Venclovas

429

Viral Transcription David LV Bauer and Ervin Fodor

439

Translation of Viral Proteins Martin D Ryan and Garry A Luke

444

Recombination Jozef J Bujarski

460

Assembly of Viruses: Enveloped Particles CK Navaratnarajah, R Warrier, and RJ Kuhn

468

Assembly of Viruses: Nonenveloped Particles M Luo

475

Virion Assembly: From Small Picornaviruses (Picornaviridae) to Large Herpesviruses (Herpesviridae) Ling Zhu, Nan Wang, and Xiangxi Wang

480

Genome Packaging Richard J Bingham, Reidun Twarock, Carlos P Mata, and Peter G Stockley

488

Virus Factories Isabel Fernández de Castro, Raquel Tenorio, and Cristina Risco

495

Release of Phages From Prokaryotic Cells Jesse Cahill and Ry Young

501

Virus Budding Lara Rheinemann and Wesley I Sundquist

519

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Content of all Volumes

Vesicle-Mediated Transcytosis and Export of Viruses Efraín E Rivera-Serrano and Stanley M Lemon

529

Vector Transmission of Animal Viruses Houssam Attoui, Fauziah Mohd Jaafar, Rennos Fragkoudis, and Peter PC Mertens

542

The Human Virome Alexia Bordigoni, Sébastien Halary, and Christelle Desnues

552

Epidemiology of Human and Animal Viral Diseases Michael Edelstein

559

Zoonosis, Emerging and Re-Emerging Viral Diseases Janet M Daly

569

Antiviral Innate Immunity: Introduction Friedemann Weber

577

Humoral and T Cell-Mediated Immunity to Viruses Ane Ogbe and Lucy Dorrell

584

Antigenicity and Antigenic Variation Kuan-Ying A Huang, Xiaorui Chen, Che Ma, Dayna Cheng, Jen-Ren Wang, and Wan-Chun Lai

597

Antigen Presentation Andrew J McMichael

601

Defense Against Viruses and Other Genetic Parasites in Prokaryotes Kira S Makarova, Yuri I Wolf, and Eugene V Koonin

606

Defective-Interfering Viruses L Roux

617

Ecology and Global Impacts of Viruses Joanne B Emerson

621

The Role of Retroviruses in Cellular Evolution Andrea Kirmaier and Welkin E Johnson

627

The Role of Bacteriophages in Bacterial Evolution Chris M Rands and Harald Brüssow

633

Viruses and Their Potential for Bioterrorism Dana L Vanlandingham and Stephen Higgs

644

The Use of Viral Promoters in Expression Vectors Ian M Jones

652

Oncolytic Viruses Laura Burga and Mihnea Bostina

658

Biotechnology Approaches to Modern Vaccine Design George P Lomonossoff and Daniel Ponndorf

662

Viruses: Impact on Science and Society Neeraja Sankaran and Robin A Weiss

671

VOLUME 2 Viruses as Infectious Agents: Human and Animal Viruses Adenoviruses (Adenoviridae) Balázs Harrach and Mária Benkő

3

Content of all Volumes

li

African Horse Sickness Virus (Reoviridae) Piet A van Rijn

17

African Swine Fever Virus (Asfarviridae) Linda K Dixon, Rachel Nash, Philippa C Hawes, and Christopher L Netherton

22

Akabane Virus and Schmallenberg Virus (Peribunyaviridae) Martin Beer and Kerstin Wernike

34

Alphaviruses Causing Encephalitis (Togaviridae) Diane E Griffin

40

Anelloviruses (Anelloviridae) Fabrizio Maggi and Mauro Pistello

48

Animal Lentiviruses (Retroviridae) Esperanza Gomez-Lucia

56

Animal Morbilliviruses (Paramyxoviridae) Carina Conceicao and Dalan Bailey

68

Animal Papillomaviruses (Papillomaviridae) John S Munday

79

Astroviruses (Astroviridae) Virginia Hargest, Amy Davis, and Stacey Schultz-Cherry

92

Avian Hepadnaviruses (Hepadnaviridae) Allison R Jilbert, Georget Y Reaiche-Miller, and Catherine A Scougall

100

Avian Herpesviruses (Herpesviridae) Vishwanatha RAP Reddy and Venugopal Nair

112

Avian Influenza Viruses (Orthomyxoviridae) Nicolas Bravo-Vasquez and Stacey Schultz-Cherry

117

Avian Leukosis and Sarcoma Viruses (Retroviridae) Karen L Beemon

122

Bluetongue Virus (Reoviridae) Raghavendran Kulasegaran-Shylini and Polly Roy

127

Borna Disease Virus and Related Bornaviruses (Bornaviridae) Susan L Payne

137

Bovine Leukemia Virus (Retroviridae) Thomas Joris, Roghaiyeh Safari, Jean-Rock Jacques, and Luc Willems

144

Bovine Viral Diarrhea, Border Disease, and Classical Swine Fever Viruses (Flaviviridae) Paul Becher, Volker Moennig, and Norbert Tautz

153

Capripoxviruses, Parapoxviruses, and Other Poxviruses of Ruminants (Poxviridae) Philippa M Beard

165

Chikungunya Virus (Togaviridae) Thomas E Morrison and Stephanie E Ander

173

Circoviruses (Circoviridae) Giovanni Franzo and Joaquim Segalés

182

Coronaviruses: General Features (Coronaviridae) Paul Britton

193

Coronaviruses: Molecular Biology (Coronaviridae) X Deng and SC Baker

198

lii

Content of all Volumes

Crimean-Congo Hemorrhagic Fever Virus and Nairoviruses of Medical Importance (Nairoviridae) Ali Mirazimi, Felicity Burt, and Anna Papa

208

Dengue Viruses (Flaviviridae) Ashley L St. John and Duane J Gubler

218

Ebola Virus (Filoviridae) Andrea Marzi and Logan Banadyga

232

Enteroviruses (Picornaviridae) Carita Savolainen-Kopra, Soile Blomqvist, and Petri Susi

245

Enveloped, Positive-Strand RNA Viruses (Nidovirales) L Enjuanes, AE Gorbalenya, RJ de Groot, JA Cowley, J Ziebuhr, and EJ Snijder

256

Epstein–Barr Virus (Herpesviridae) Lawrence S Young

267

Equine Herpesviruses (Herpesviridae) Gisela Soboll Hussey, Nikolaus Osterrieder, and Walid Azab

278

Equine, Canine, and Swine Influenza (Orthomyxoviridae) Janet M Daly and Japhette E Kembou-Ringert

287

Feline Calicivirus (Caliciviridae) Margaret J Hosie and Michaela J Conley

294

Feline Leukemia and Sarcoma Viruses (Retroviridae) Brian J Willett and Margaret J Hosie

300

Fish and Amphibian Alloherpesviruses (Herpesviridae) Maxime Boutier, Léa Morvan, Natacha Delrez, Francesco Origgi, Andor Doszpoly, and Alain Vanderplasschen

306

Fish Retroviruses (Retroviridae) TA Paul, RN Casey, PR Bowser, JW Casey, J Rovnak, and SL Quackenbush

316

Fish Rhabdoviruses (Rhabdoviridae) Gael Kurath and David Stone

324

Foot-and-Mouth Disease Viruses (Picornaviridae) David J Rowlands

332

Fowlpox Virus and Other Avipoxviruses (Poxviridae) Efstathios S Giotis and Michael A Skinner

343

Hantaviruses (Hantaviridae) Tarja Sironen and Antti Vaheri

349

Henipaviruses (Paramyxoviridae) Lin-Fa Wang and Danielle E Anderson

355

Hepatitis A Virus (Picornaviridae) Andreas Dotzauer

362

Hepatitis B Virus (Hepadnaviridae) Peter Karayiannis

373

Hepatitis C Virus (Flaviviridae) Ralf Bartenschlager and Keisuke Tabata

386

Hepeviruses (Hepeviridae) Xiang-Jin Meng

397

Herpes Simplex Virus 1 and 2 (Herpesviridae) David M Knipe and Richard Whitley

404

Content of all Volumes

liii

History of Virology: Vertebrate Viruses F Fenner

414

Human Boca- and Protoparvoviruses (Parvoviridae) Maria Söderlund-Venermo and Jianming Qiu

419

Human Coronavirus-229E, -OC43, -NL63, and -HKU1 (Coronaviridae) Ding X Liu, Jia Q Liang, and To S Fung

428

Human Cytomegalovirus (Herpesviridae) Edward S Mocarski

441

Human Immunodeficiency Virus (Retroviridae) Blaide Woodburn, Ann Emery, and Ronald Swanstrom

460

Human Metapneumovirus (Pneumoviridae) Antonella Casola, Matteo P Garofalo, and Xiaoyong Bao

475

Human Norovirus and Sapovirus (Caliciviridae) Sumit Sharma, Marie Hagbom, Lennart Svensson, and Johan Nordgren

483

Human Papillomaviruses (Papillomaviridae) Alison A McBride and Samuel S Porter

493

Human Parainfluenza Viruses (Paramyxoviridae) Elisabeth Adderson

502

Human Pathogenic Arenaviruses (Arenaviridae) Sheli R Radoshitzky and Juan C de la Torre

507

Human Polyomaviruses (Papillomaviridae) Melissa S Maginnis

518

Human T-Cell Leukemia Virus-1 and -2 (Retroviridae) Amanda R Panfil and Patrick L Green

528

Infectious Bursal Disease Virus (Birnaviridae) Daral J Jackwood

540

Infectious Pancreatic Necrosis Virus (Birnaviridae) Øystein Evensen

544

Influenza A Viruses (Orthomyxoviridae) Laura Kakkola, Niina Ikonen, and Ilkka Julkunen

551

Influenza B, C and D Viruses (Orthomyxoviridae) Thorsten Wolff and Michael Veit

561

Jaagsiekte Sheep Retrovirus (Retroviridae) James M Sharp, Marcelo De las Heras, Massimo Palmarini, and Thomas E Spencer

575

Japanese Encephalitis Virus (Flaviviridae) Sang-Im Yun and Young-Min Lee

583

Kaposi’s Sarcoma-Associated Herpesvirus (Herpesviridae) Anne K Cordes and Thomas F Schulz

598

Marburg and Ravn Viruses (Filoviridae) Courtney Woolsey, Thomas W Geisbert, and Robert W Cross

608

Measles Virus (Paramyxoviridae) Roberto Cattaneo and Michael McChesney

619

Molluscum Contagiosum Virus (Poxviridae) Joachim J Bugert and Rosina Ehmann

629

liv

Content of all Volumes

Mumps Virus (Paramyxoviridae) Stephen A Winchester and Kevin E Brown

634

Murine Leukemia and Sarcoma Viruses (Retroviridae) George R Young and Kate N Bishop

643

Newcastle Disease Virus (Paramyxoviridae) Ben Peeters and Guus Koch

648

Orthobunyaviruses (Peribunyaviridae) Alyssa B Evans, Clayton W Winkler, and Karin E Peterson

654

Parapoxviruses (Poxviridae) Hanns-Joachim Rziha and Mathias Büttner

666

Parechoviruses (Picornaviridae) Sisko Tauriainen and Glyn Stanway

675

Parvoviruses of Carnivores, and the Emergence of Canine Parvovirus (Parvoviridae) Colin R Parrish, Ian EH Voorhees, and Susan L Hafenstein

683

Polioviruses (Picornaviridae) Philip D Minor

688

Porcine Reproductive and Respiratory Syndrome Virus and Equine Arteritis Virus (Arteriviridae) Jianqiang Zhang, Alan T Loynachan, Gregory W Stevenson, and Jeffrey J Zimmerman

697

Prions of Vertebrates Jonathan DF Wadsworth and John Collinge

707

Pseudorabies Virus (Herpesviridae) Thomas C Mettenleiter and Barbara G Klupp

714

Rabbit Hemorrhagic Disease Virus and European Brown Hare Syndrome Virus (Caliciviridae) Lorenzo Capucci, Patrizia Cavadini, and Antonio Lavazza

724

Rabbit Myxoma Virus and the Fibroma Viruses (Poxviridae) Peter J Kerr

730

Rabies and Other Lyssaviruses (Rhabdoviridae) Ashley C Banyard and Anthony R Fooks

738

Respiratory Syncytial Virus (Pneumoviridae) Tiffany King, Tiffany Jenkins, Supranee Chaiwatpongsakorn, and Mark E Peeples

747

Rhinoviruses (Picornaviridae) Matti Waris and Olli Ruuskanen

757

Rift Valley Fever Virus and Other Phleboviruses (Phenuiviridae) Tetsuro Ikegami

765

Roseoloviruses: Human Herpesviruses 6A, 6B and 7 (Herpesviridae) Katherine N Ward

778

Rotaviruses (Reoviridae) Juana Angel and Manuel A Franco

789

Rubella Virus (Picornaviridae) Annette Mankertz

797

Saint Louis Encephalitis Virus (Flaviviridae) William K Reisen, Lark L Coffey, Daniele M Swetnam, and Aaron C Brault

805

Severe Acute Respiratory Syndrome (SARS) and Middle East Respiratory Syndrome (MERS) (Coronaviridae) Joseph SM Peiris and Leo LM Poon

814

Content of all Volumes

lv

Severe Acute Respiratory Syndrome Coronavirus 2 (SARS-CoV-2) (Coronaviridae) Malik Peiris

825

Simian Immunodeficiency Virus (SIV) and HIV-2 (Retroviridae) Phyllis J Kanki

827

Sindbis Virus (Togaviridae) Satu Kurkela

837

Tick-Borne Encephalitis Virus (Flaviviridae) Teemu Smura, Suvi Kuivanen, and Olli Vapalahti

843

Transmissible Gastroenteritis Virus of Pigs and Porcine Epidemic Diarrhea Virus (Coronaviridae) Qiang Liu and Volker Gerdts

850

Vaccinia Virus (Poxviridae) Yan Xiang and Rebecca K Lane

854

Varicella-Zoster Virus (Herpesviridae) Jeffrey I Cohen

860

Variola and Monkeypox Viruses (Poxviridae) Lalita Priyamvada and Panayampalli S Satheshkumar

868

Vesicular Stomatitis Virus and Bovine Ephemeral Fever Virus (Rhabdoviridae) Peter J Walker and Robert B Tesh

875

West Nile Virus (Flaviviridae) Fengwei Bai and Elizabeth Ashley Thompson

884

Yellow Fever Virus (Flaviviridae) Ashley E Strother and Alan DT Barrett

891

Zika Virus (Flaviviridae) Nikos Vasilakis, Shannan L Rossi, Sasha R Azar, Irma E Cisneros, Cassia F Estofolete, and Mauricio L Nogueira

899

VOLUME 3 Viruses as Infectious Agents: Plant Viruses An Introduction to Plant Viruses Roger Hull

3

Emerging and Re-Emerging Plant Viruses Sabrina Bertin, Francesco Faggioli, Andrea Gentili, Ariana Manglli, Anna Taglienti, Antonio Tiberini, and Laura Tomassoli

8

Emerging Geminiviruses (Geminiviridae) Muhammad S Nawaz-ul-Rehman, Nazia Nahid, and Muhammad Mubin

21

Movement of Viruses in Plants Manfred Heinlein

32

Plant Antiviral Defense: Gene-Silencing Pathways Vitantonio Pantaleo, Chikara Masuta, and Hanako Shimura

43

Plant Resistance to Viruses: Engineered Resistance Marc Fuchs

52

Plant Resistance to Viruses: Natural Resistance Associated With Dominant Genes Mandy Ravensbergen and Richard Kormelink

60

lvi

Content of all Volumes

Plant Resistance to Viruses: Natural Resistance Associated With Recessive Genes Masayoshi Hashimoto, Kensaku Maejima, Yasuyuki Yamaji, and Shigetou Namba

69

Plant Viral Diseases: Economic Implications Basavaprabhu L Patil

81

Retrotransposons of Plants M-A Grandbastien

98

Vector Transmission of Plant Viruses Etienne Herrbach and Quentin Chesnais

106

Viral Suppressors of Gene Silencing Hernan Garcia-Ruiz

116

Virus-Induced Gene Silencing (VIGS) Xu Tengzhi, Ugrappa Nagalakshmi, and Savithramma P Dinesh-Kumar

123

Alfalfa Mosaic Virus (Bromoviridae) L Sue Loesch-Fries

132

Alphaflexiviruses (Alphaflexiviridae) Sergey Y Morozov and Alexey A Agranovsky

140

Alphasatellites (Alphasatellitidae) Rob W Briddon and Muhammad S Nawaz-ul-Rehman

149

Amalgaviruses (Amalgaviridae) Ioannis E Tzanetakis, Sead Sabanadzovic, and Rodrigo A Valverde

154

Badnaviruses (Caulimoviridae) Andrew DW Geering

158

Banana Bunchy Top Virus (Nanoviridae) John E Thomas

169

Barley Yellow Dwarf Viruses (Luteoviridae) Leslie L Domier

176

Bean Common Mosaic Virus and Bean Common Mosaic Necrosis Virus (Potyviridae) Ramon Jordan and John Hammond

184

Bean Golden Mosaic Virus and Bean Golden Yellow Mosaic Virus (Geminiviridae) Francisco M Zerbini and Simone G Ribeiro

192

Beet Curly Top Virus (Geminiviridae) Robert L Gilbertson, Tomas A Melgarejo, Maria R Rojas, William M Wintermantel, and John Stanley

200

Beet Necrotic Yellow Vein Virus (Benyviridae) Sebastian Liebe, Annette Niehl, Renate Koenig, and Mark Varrelmann

213

Benyviruses (Benyviridae) Annette Niehl, Sebastian Liebe, Mark Varrelmann, and Renate Koenig

219

Betaflexiviruses (Betaflexiviridae) Nobuyuki Yoshikawa and Hajime Yaegashi

229

Betasatellites and Deltasatelliles (Tolecusatellitidae) Muhammad S Nawaz-ul-Rehman, Nazia Nahid, Muhammad Hassan, and Muhammad Mubin

239

Bluner-, Cile-, and Higreviruses (Kitaviridae) Diego F Quito-Avila, Juliana Freitas-Astúa, and Michael J Melzer

247

Brome Mosaic Virus (Bromoviridae) Guijuan He, Zhenlu Zhang, Preethi Sathanantham, Arturo Diaz, and Xiaofeng Wang

252

Content of all Volumes

lvii

Bromoviruses (Bromoviridae) Jozef J Bujarski

260

Bymoviruses (Potyviridae) Annette Niehl and Frank Rabenstein

268

Cacao Swollen Shoot Virus (Caulimoviridae) Emmanuelle Muller

274

Carmo-Like Viruses (Tombusviridae) Miryam Pérez-Cañamás and Carmen Hernández

285

Cassava Brown Streak Viruses (Potyviridae) Basavaprabhu L Patil

293

Cassava Mosaic Viruses (Geminiviridae) James Legg and Stephan Winter

301

Caulimoviruses (Caulimoviridae) James E Schoelz and Mustafa Adhab

313

Cheraviruses, Sadwaviruses and Torradoviruses (Secoviridae) Toru Iwanami and René AA van der Vlugt

322

Citrus Tristeza Virus (Closteroviridae) Moshe Bar-Joseph, Scott J Harper, and William O Dawson

327

Closteroviruses (Closteroviridae) Marc Fuchs

336

Comoviruses and Fabaviruses (Secoviridae) George P Lomonossoff

348

Cotton Leaf Curl Disease (Geminiviridae) Nasim Ahmed, Imran Amin, and Shahid Mansoor

355

Cowpea Mosaic Virus (Secoviridae) George P Lomonossoff

364

Cucumber Mosaic Virus (Bromoviridae) Judith Hirsch and Benoît Moury

371

Dianthovirus (Tombusviridae) Kiwamu Hyodo and Masanori Kaido

383

Endornaviruses (Endornaviridae) Toshiyuki Fukuhara

388

Fimoviruses (Fimoviridae) Toufic Elbeaino and Michele Digiaro

396

Furoviruses (Virgaviridae) Annette Niehl and Renate Koenig

405

Geminiviruses (Geminiviridae) Jesús Navas-Castillo and Elvira Fiallo-Olivé

411

Hordeiviruses (Virgaviridae) Zhihao Jiang, Meng Yang, Yongliang Zhang, Andrew O Jackson, and Dawei Li

420

Idaeoviruses (Mayoviridae) Robert R Martin and Karen E Keller

430

Ilarviruses (Bromoviridae) Aaron Simkovich, Susanne E Kohalmi, and Aiming Wang

439

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Content of all Volumes

Luteoviruses (Luteoviridae) Leslie L Domier

447

Machlomovirus and Panicoviruses (Tombusviridae) Kay Scheets

456

Maize Streak Virus (Geminiviridae) Darren P Martin and Aderito L Monjane

461

Nanoviruses (Nanoviridae) Bruno Gronenborn and H Josef Vetten

470

Necro-Like Viruses (Tombusviridae) Luisa Rubino and Giovanni P Martelli†

481

Nepoviruses (Secoviridae) Hélène Sanfaçon

486

Ophioviruses (Aspiviridae) Anna M Vaira and John Hammond

495

Orthotospoviruses (Tospoviridae) Renato O Resende and Hanu R Pappu

507

Ourmiaviruses (Botourmiaviridae) Gian Paolo Accotto and Cristina Rosa

516

Papaya Ringspot Virus (Potyviridae) Cécile Desbiez and Hervé Lecoq

520

Pecluviruses (Virgaviridae) Hema Masarapu, Pothur Sreenivasulu, Philippe Delfosse, Claude Bragard, Anne Legreve, and DVR Reddy

528

Pepino Mosaic Virus (Alphaflexiviridae) Rene AA van der Vlugt and CCMM Stijger

539

Plant Reoviruses (Reoviridae) Yu Huang and Yi Li

545

Plant Resistance to Geminiviruses Basavaprabhu L Patil, Supriya Chakraborty, Henryk Czosnek, Elvira Fiallo-Olivé, Robert L Gilbertson, James Legg, Shahid Mansoor, Jesús Navas-Castillo, Rubab Z Naqvi, Saleem U Rahman, and Francisco M Zerbini

554

Plant Rhabdoviruses (Rhabdoviridae) Ralf G Dietzgen, Michael M Goodin, and Zhenghe Li

567

Plant Satellite Viruses (Albetovirus, Aumaivirus, Papanivirus, Virtovirus) Mart Krupovic

581

Plum Pox Virus (Potyviridae) Miroslav Glasa and Thierry Candresse

586

Poleroviruses (Luteoviridae) Hernan Garcia-Ruiz, Natalie M Holste, and Katherine LaTourrette

594

Pomoviruses (Virgaviridae) Eugene I Savenkov

603

Potato Virus Y (Potyviridae) Laurent Glais and Benoît Moury

612

Potexviruses (Alphaflexiviridae) Ki H Ryu, Eun G Song, and Jin S Hong

623

†

Deceased.

Content of all Volumes

lix

Potyviruses (Potyviridae) Adrián Valli, Juan A García, and Juan J López-Moya

631

Quinviruses (Betaflexiviridae) Ki H Ryu and Eun G Song

642

Reverse-Transcribing Viruses (Belpaoviridae, Metaviridae, and Pseudoviridae) Carlos Llorens, Beatriz Soriano, Maria A Navarrete-Muñoz, Ahmed Hafez, Vicente Arnau, Jose Miguel Benito, Toni Gabaldon, Norma Rallon, Jaume Pérez-Sánchez, and Mart Krupovic

653

Rice Tungro Disease (Secoviridae, Caulimoviridae) Gaurav Kumar, Fauzia Zarreen, and Indranil Dasgupta

667

Rice Yellow Mottle Virus (Solemoviridae) Eugénie Hébrard, Nils Poulicard, and Mbolarinosy Rakotomalala

675

Satellite Nucleic Acids and Viruses Olufemi J Alabi, Alfredo Diaz-Lara, and Maher Al Rwahnih

681

Secoviruses (Secoviridae) Jeremy R Thompson

692

Sequiviruses and Waikaviruses (Secoviridae) Lucy Rae Stewart

703

Solemoviruses (Solemoviridae) Cecilia Sarmiento, Merike Sõmera, and Erkki Truve†

712

Tenuiviruses (Phenuiviridae) Bertha Cecilia Ramirez and Anne-Lise Haenni

719

Tobacco Mosaic Virus (Virgaviridae) Marc HV Van Regenmortel

727

Tobamoviruses (Virgaviridae) Ulrich Melcher, Dennis J Lewandowski, and William O Dawson

734

Tobraviruses (Virgaviridae) Stuart A MacFarlane

743

Tomato Leaf Curl New Delhi Virus (Geminiviridae) Supriya Chakraborty and Manish Kumar

749

Tomato Spotted Wilt Virus (Tospoviridae) Hanu R Pappu, Anna E Whitfield, and Athos S de Oliveira

761

Tomato Yellow Leaf Curl Viruses (Geminiviridae) Henryk Czosnek

768

Tombusvirus-Like Viruses (Tombusviridae) K Andrew White

778

Tombusviruses (Tombusviridae) Luisa Rubino and Kay Scheets

788

Tritimoviruses and Rymoviruses (Potyviridae) Satyanarayana Tatineni and Gary L Hein

797

Triviruses (Betaflexiviridae) Yahya ZA Gaafar and Heiko Ziebell

805

Tymoviruses (Tymoviridae) Rosemarie W Hammond and Peter Abrahamian

818

†

Deceased.

lx

Content of all Volumes

Umbraviruses (Tombusviridae) Eugene V Ryabov and Michael E Taliansky

827

Varicosaviruses (Rhabdoviridae) Takahide Sasaya

833

Virgaviruses (Virgaviridae) Eugene I Savenkov

839

Viroids (Pospiviroidae and Avsunviroidae) Ricardo Flores and Robert A Owens

852

Watermelon Mosaic Virus and Zucchini Yellow Mosaic Virus (Potyviridae) Cécile Desbiez and Hervé Lecoq

862

VOLUME 4 Viruses as Infectious Agents: Bacterial, Archaeal, Fungal, Algal, and Invertebrate Viruses Bacterial Viruses History of Virology: Bacteriophages William C Summers

3

Icosahedral Phages – Single-Stranded DNA (φX174) Bentley A Fane and Aaron P Roznowski

10

Single-Stranded RNA Bacterial Viruses Peter G Stockley and Junjie Zhang

21

Enveloped Icosahedral Phages – Double-Stranded RNA (φ6) Paul Gottlieb and Aleksandra Alimova

26

Membrane-Containing Icosahedral DNA Bacteriophages Roman Tuma, Sarah J Butcher, and Hanna M Oksanen

36

Tailed Double-Stranded DNA Phages Robert L Duda

45

Helical and Filamentous Phages Andreas Kuhn and Sebastian Leptihn

53

Replication of Bacillus Double-Stranded DNA Bacteriophages Silvia Ayora, Paulo Tavares, Ruben Torres, and Juan C Alonso

61

Lytic Transcription William McAllister and Deborah M Hinton

69

Lysogeny Keith E Shearwin and Jia Q Truong

77

Decision Making by Temperate Phages Ido Golding, Seth Coleman, Thu VP Nguyen, and Tianyou Yao

88

Mobilization of Phage Satellites Kristen N LeGault and Kimberley D Seed

98

Portal Vertex Peng Jing and Mauricio Cortes Jr.

105

Content of all Volumes

lxi

Prohead, the Head Shell Pre-Cursor Marc C Morais and Michael E Woodson

115

Enzymology of Viral DNA Packaging Machines Carlos E Catalano

124

DNA Packaging: DNA Recognition Sandra J Greive and Oliver W Bayfield

136

DNA Packaging: The Translocation Motor Janelle A Hayes and Brian A Kelch

148

Biophysics of DNA Packaging Joshua Pajak, Gaurav Arya, and Douglas E Smith

160

Energetics of the DNA-Filled Head Alex Evilevitch

167

Bacteriophage Receptor Proteins of Gram-Negative Bacteria Sarah M Doore, Kristin N Parent, Sundharraman Subramanian, Jason R Schrad, and Natalia B Hubbs

175

Tail Structure and Dynamics Shweta Bhatt, Petr G Leiman, and Nicholas MI Taylor

186

Bacteriophage Tail Fibres, Tailspikes, and Bacterial Receptor Interaction Mateo Seoane-Blanco, Mark J van Raaij, and Meritxell Granell

194

Phage Genome and Protein Ejection In Vivo Ian J Molineux, L Letti Lopez, and Aaron P Roznowski

206

Dealing With the Whole Head: Diversity and Function of Capsid Ejection Proteins in Tailed Phages Lindsay W Black and Julie A Thomas

219

Jumbo Phages Isaac T Younker and Carol Duffy

229

CRISPR-Cas Systems and Anti-CRISPR Proteins: Adaptive Defense and Counter-Defense in Prokaryotes and Their Viruses Asma Hatoum-Aslan and Olivia G Howell

242

Bacteriophage: Therapeutics and Diagnostics Development Teng-Chieh Yang

252

Bacteriophage Vaccines Pan Tao and Venigalla B Rao

259

Bacteriophage Diversity Julianne H Grose and Sherwood R Casjens

265

Genetic Mosaicism in the Tailed Double-Stranded DNA Phages Welkin H Pope

276

Bacteriophages of the Human Microbiome Pilar Manrique, Michael Dills, and Mark J Young

283

Bacteriophage: Red Recombination System and the Development of Recombineering Technologies Lynn C Thomason and Kenan C Murphy

291

Nanotechnology Application of Bacteriophage DNA Packaging Nanomotors Tao Weitao, Lixia Zhou, Zhefeng Li, Long Zhang, and Peixuan Guo

302

General Ecology of Bacteriophages Stephen T Abedon

314

Marine Bacteriophages Vera Bischoff, Falk Zucker, and Cristina Moraru

322

lxii

Content of all Volumes

Ecology of Phages in Extreme Environments Tatiana A Demina and Nina S Atanasova

342

Archaeal Viruses Diversity of Hyperthermophilic Archaeal Viruses David Prangishvili, Mart Krupovic, and Diana P Baquero

359

Euryarchaeal Viruses Tatiana A Demina and Hanna M Oksanen

368

Vesicle-Like Archaeal Viruses Elina Roine and Nina S Atanasova

380

Virus–Host Interactions in Archaea Diana P Baquero, David Prangishvili, and Mart Krupovic

387

Antiviral Defense Mechanisms in Archaea Qunxin She

400

Discovery of Archaeal Viruses in Hot Spring Environments Using Viral Metagenomics Jennifer Wirth, Jacob H Munson-McGee, and Mark J Young

407

Metagenomes of Archaeal Viruses in Hypersaline Environments Fernando Santos, María D Ramos-Barbero, and Josefa Antón

414

Extreme Environments as a Model System to Study How Virus–Host Interactions Evolve Along the Symbiosis Continuum Samantha J DeWerff and Rachel J Whitaker

419

Fungal Viruses An Introduction to Fungal Viruses Nobuhiro Suzuki

431

Cross-Kingdom Virus Infection Liying Sun, Hideki Kondo, and Ida Bagus Andika

443

Diversity of Mycoviruses in Aspergilli Ioly Kotta-Loizou

450

Evolution of Mycoviruses Mahtab Peyambari, Vaskar Thapa, and Marilyn J Roossinck

457

Mixed Infections of Mycoviruses in Phytopathogenic Fungus Sclerotinia sclerotiorum Jiatao Xie and Daohong Jiang

461

Mycovirus-Mediated Biological Control Daniel Rigling, Cécile Robin, and Simone Prospero

468

Mycoviruses With Filamentous Particles Michael N Pearson

478

Prions of Yeast and Fungi Reed B Wickner and Herman K Edskes

487

Single-Stranded DNA Mycoviruses Daohong Jiang

493

Structure of Double-Stranded RNA Mycoviruses José R Castón, Nobuhiro Suzuki, and Said A Ghabrial†

504

†

Deceased.

Content of all Volumes

lxiii

Ustilago maydis Viruses and Their Killer Toxins Alexis Williams and Thomas J Smith

513

Vegetative Incompatibility in Filamentous Fungi Songsong Wu, Daohong Jiang, and Jiatao Xie

520

Viral Diseases of Agaricus bisporus, the Button Mushroom Kerry S Burton and Greg Deakin

528

Viral Killer Toxins Manfred J Schmitt and Björn Becker

534

Alternaviruses (Unassigned) Hiromitsu Moriyama, Nanako Aoki, Kuko Fuke, Kana Takeshita Urayama, Naoki Takeshita, and Chien-Fu Wu

544

Barnaviruses (Barnaviridae) Peter A Revill

549

Botybirnaviruses (Botybirnavirus) Mingde Wu, Guoqing Li, Daohong Jiang, and Jiatao Xie

552

Chrysoviruses (Chrysoviridae) - General Features and Chrysovirus-Related Viruses Ioly Kotta-Loizou, Robert HA Coutts, José R Castón, Hiromitsu Moriyama, and Said A Ghabrial†

557

Fungal Partitiviruses (Partitiviridae) Eeva J Vainio

568

Fusariviruses (Unassigned) Sotaro Chiba

577

Giardiavirus (Totiviridae) Juliana Gabriela Silva de Lima, João Paulo Matos Santos Lima, and Daniel Carlos Ferreira Lanza

582

Hypoviruses (Hypoviridae) Dong-Xiu Zhang and Donald L Nuss

589

Megabirnaviruses (Megabirnaviridae) Yukiyo Sato and Nobuhiro Suzuki

594

Mitoviruses (Mitoviridae) Bradley I Hillman and Alanna B Cohen

601

Mycoreoviruses (Reoviridae) Bradley I Hillman and Alanna B Cohen

607

Mymonaviruses (Mymonaviridae) Daohong Jiang

615

Narnaviruses (Narnaviridae) Rosa Esteban and Tsutomu Fujimura

621

Phlegiviruses (Unassigned) Karel Petrzik

627

Plant and Protozoal Partitiviruses (Partitiviridae) Hanna Rose and Edgar Maiss

632

Quadriviruses (Quadriviridae) Hideki Kondo, José R Castón, and Nobuhiro Suzuki

642

Totiviruses (Totiviridae) Bradley I Hillman and Alanna B Cohen

648

†

Deceased.

lxiv

Content of all Volumes

Yado-kari Virus 1 and Yado-nushi Virus 1 (Unassigned) Subha Das and Nobuhiro Suzuki

658

Yeast L-A Virus (Totiviridae) Reed B Wickner, Tsutomu Fujimura, and Rosa Esteban

664

Algal Viruses Algal Marnaviruses (Marnaviridae) Marli Vlok, Curtis A Suttle, and Andrew S Lang

671

Algal Mimiviruses (Mimiviridae) Ruth-Anne Sandaa, Håkon Dahle, Corina PD Brussaard, Hiroyuki Ogata, and Romain Blanc-Mathieu

677

Miscellaneous Algal Viruses (Alvernaviridae, Bacilladnaviridae, Dinodnavirus, Reoviridae) Keizo Nagasaki, Yuji Tomaru, and Corina PD Brussaard

684

Phycodnaviruses (Phycodnaviridae) James L Van Etten, David D Dunigan, Keizo Nagasaki, Declan C Schroeder, Nigel Grimsley, Corina PD Brussaard, and Jozef I Nissimov

687

Invertebrate Viruses An Introduction to Viruses of Invertebrates Peter Krell

699

Ascoviruses (Ascoviridae) Sassan Asgari, Dennis K Bideshi, Yves Bigot, and Brian A Federici

724

Baculovirus–Host Interactions: Repurposing Host-Acquired Genes (Baculoviridae) A Lorena Passarelli

732

Baculoviruses: General Features (Baculoviridae) Vera ID Ros

739

Baculoviruses: Molecular Biology and Replication (Baculoviridae) Monique M van Oers

747

Bidensoviruses (Bidnaviridae) Qin Yao, Zhaoyang Hu, and Keping Chen

759

Bunyaviruses of Arthropods (Mypoviridae, Nairoviridae, Peribunyaviridae, Phasmaviridae, Phunuiviridae, Wupedeviridae) Sandra Junglen

764

Dicistroviruses (Dicistroviridae) Yanping Chen and Steven M Valles

768

Entomobirnaviruses (Birnaviridae) Marco Marklewitz

776

Hytrosaviruses (Hytrosaviridae) Henry M Kariithi and Irene K Meki

780

Iflaviruses (Iflaviridae) Bryony C Bonning and Sijun Liu

792

Iridoviruses of Invertebrates (Iridoviridae) İkbal Agah İnce

797

Mesoniviruses (Mesoniviridae) Jody Hobson-Peters and Daniel Watterson

804

Content of all Volumes

lxv

Nimaviruses (Nimaviridae) Peter Krell and Emine Ozsahin

808

Nodaviruses of Invertebrates and Fish (Nodaviridae) Kyle L Johnson and Jacen S Moore

819

Nudiviruses (Nudiviridae) Yu-Chan Chao, Chih-Hsuan Tsai, and Sung-Chan Wei

827

Parvoviruses of Invertebrates (Parvoviridae) Judit J Pénzes, Hanh T Pham, Qian Yu, Max Bergoin, and Peter Tijssen

835

Polydnaviruses (Polydnaviridae) Anne-Nathalie Volkoff and Elisabeth Huguet

849

Poxviruses of Insects (Poxviridae) Basil Arif, Lillian Pavlik, Remziye Nalçacıoğlu, Hacer Muratoğlu, Cihan İnan, Mehtap Yakupoğlu, Emine Özsahin, Ismail Demir, Kazım Sezen, and Zihni Demirbağ

858

Reoviruses of Invertebrates (Reoviridae) Peter Krell

867

Rhabdoviruses of Insects (Rhabdoviridae) Andrea González-González, Nicole T de Stefano, David A Rosenbaum, and Marta L Wayne

883

Sarthroviruses (Sarthroviridae) Azeez Sait Sahul Hameed

888

Solinviviruses (Solinviviridae) Steven M Valles and Andrew E Firth

892

Tetraviruses (Alphatetraviridae, Carmotetraviridae, Permutotetraviridae) Rosemary A Dorrington, Tatiana Domitrovic, and Meesbah Jiwaji

897

VOLUME 5 Diagnosis, Treatment and Prevention of Virus Infections Diagnosis Introduction to Virus Diagnosis and Treatment Maija Lappalainen and Hubert GM Niesters

3

Electron Microscopy for Viral Diagnosis Roland A Fleck

5

Serological Approaches for Viral Diagnosis Klaus Hedman and Visa Nurmi

15

A Brief History of the Development of Diagnostic Molecular-Based Assays Hubert GM Niesters

22

Sequencing Strategies Sibnarayan Datta

27

Validating Real-Time Polymerase Chain Reaction (PCR) Assays Melvyn Smith

35

Rapid Point-of-Care Assays Jan G Lisby and Uffe V Schenider

45

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Content of all Volumes

Standardization of Diagnostic Assays Sally A Baylis, C Micha Nübling, and Wayne Dimech

52

Quality Assurance in the Clinical Virology Laboratory Paul Wallace and Elaine McCulloch

64

Biosafety and Biosecurity in Diagnostic Laboratories Hannimari Kallio-Kokko and Susanna Sissonen

82

Screening for Viral Infections Walter Ian Lipkin, Nischay Mishra, and Thomas Briese

91

Clinical Diagnostic Virology Marcus Panning

98

Virus Diagnosis in Immunosuppressed Individuals Elisabeth Puchhammer-Stöckl and Fausto Baldanti

105

Diagnosis; Future Prospects on Direct Diagnosis Marianna Calabretto, Daniele Di Carlo, Fabrizio Maggi, and Guido Antonelli

112

Treatment Antiviral Classification Guangdi Li, Xixi Jing, Pan Zhang, and Erik De Clercq

121

Antiretroviral Therapy – Nucleoside/Nucleotide and Non-Nucleoside Reverse Transcriptase Inhibitors Timothy D Appleby and Killian J Quinn

131

Protease Inhibitors Vanesa Anton-Vazquez and Frank A Post

139

HIV Integrase Inhibitors and Entry Inhibitors Daniel Bradshaw and Ranjababu Kulasegaram

145

Management of Respiratory Syncytial Virus Infections (Pneumoviridae) Rachael S Barr and Simon B Drysdale

155

Management of Influenza Virus Infections (Orthomyxoviridae) Bruno Lina

160

Management of Herpes Simplex Virus Infections (Herpesviridae) Nicole Samies and Richard Whitley

175

Management of Varicella-Zoster Virus Infections (Herpesviridae) Andreas Sauerbrei

181

Treatment and Prevention of Herpesvirus Infections in the Immunocompromised Host Sara H Burkhard and Nicolas J Mueller

190

Management of Adenovirus Infections (Adenoviridae) Albert Heim

197

Management of Hepatitis A and E Virus Infection Sébastien Lhomme, Florence Abravanel, Jean-Marie Peron, Nassim Kamar, and Jacques Izopet

206

Management of Patients With Chronic Hepatitis B (Hepadnaviridae) and Chronic Hepatitis D Infection (Deltavirus) Milan J Sonneveld and Suzanne van Meer

217

Studying Population Genetic Processes in Viruses: From Drug-Resistance Evolution to Patient Infection Dynamics Jeffrey D Jensen

227

Content of all Volumes

Virus-Based Cancer Therapeutics Roberto Cattaneo and Christine E Engeland

lxvii

233

Prevention Surveillance of Infectious Diseases Norman Noah

247

Preparing for Emerging Zoonotic Viruses Reina S Sikkema and Marion PG Koopmans

256

Use of Immunoglobulins in the Prevention of Viral Infections Leyla Asadi and Giovanni Ferrara

267

Vaccine Production, Safety, and Efficacy Thomas J Brouwers and Bernard AM Van der Zeijst

281

Vaccines Against Viral Gastroenteritis Scott Grytdal, Tyler P Chavers, Claire P Mattison, Jacqueline E Tate, and Aron J Hall

289

Human Papillomavirus (HPV) Vaccines and Their Impact Jade Pattyn, Pierre Van Damme, and Alex Vorsters

295

Influenza Vaccination Topi Turunen

300

Polio Eradication M Steven Oberste, Cara C Burns, and Jennifer L Konopka-Anstadt

310

Subject Index

315

PLANT VIRUSES

An Introduction to Plant Viruses Roger Hull, John Innes Centre, Norwich, United Kingdom r 2021 Elsevier Ltd. All rights reserved. This is a reproduction of R. Hull, History of Virology: Plant Viruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00589-6.

Introduction History shows that the study of virus infections of plants has led the overall subject of virology in the development of several major concepts including that of the entity of viruses themselves. To obtain a historical perspective of plant virology, five major (overlapping) ages can be recognized.

Prehistory The earliest known written record describing what was almost certainly a plant virus disease is a poem in Japanese written by the Empress Koken in AD 752 and translated by T. Inouye as follows: In this village It looks as if frosting continuously For, the plant I saw In the field of summer The color of the leaves were yellowing The plant, identified as Eupatorium lindleyanum, has been found to be susceptible to tomato yellow leaf curl virus, which causes a yellowing disease. In Western Europe in the period from about 1600 to 1660, many paintings and drawings were made of tulips that demonstrate flower symptoms of virus disease. These are recorded in the Herbals of the time and in the still-life paintings of artists such as Johannes Bosschaert in 1610. During this period, blooms featuring such striped patterns were prized as special varieties leading to the phenomenon of ‘tulipomania’. The trade in infected tulip bulbs resulted in hyperinflation with bulbs exchanging hands for large amounts of money or goods (Table 1). In describing an experiment to demonstrate that sap flows in plants, Lawrence reported in 1714 the unintentonal transmission of a virus disease of jasmine by grafting. The following quotation from Blair in 1719 describes the procedure and demonstrates that even in this protoscientific stage, experimenters were already indulging in arguments about priorities of discovery. The inoculating of a strip’d Bud into a plain stock and the consequence that the Stripe or Variegation shall be seen in a few years after, all over the shrub above and below the graft, is a full demonstration of this Circulation of the Sap. This was first observed by Mr. Wats at Kensington, about 18 years ago: Mr. Fairchild performed it 9 years ago; Mr. Bradly says he observ’d it several years since; though Mr. Lawrence would insinuate as if he had first discovered it.

Recognition of Viral Entity In the latter part of the nineteenth century, the idea that infectious disease was caused by microbes was well established, and filters were available that would not allow the passage of known bacterial pathogens. Mayer in 1886 showed that a disease of tobacco (Mosaikkrankheit; now known to be caused by tobacco mosaic virus; TMV) could be transmitted to healthy plants by inoculation with extracts from diseased plants; Iwanowski demonstrated in 1892 that sap from such tobacco plants was still infective after it had been passed through a bacteria-proof filter candle. This work did not attract much attention until it was repeated by Beijerinck who in 1898 described the infectious agent as contagium vivum fluidum (Latin for contagious living fluid) to distinguish it from contagious corpuscular agents. Beijerinck's discovery is considered to be the birth of virology. In 1904 Baur showed that an infectious variegation of Abutilon could be transmitted by grafting, but not by mechanical inoculation. Beijerinck and Baur used the term ‘virus’ in describing the causative agents of these diseases, to contrast them with bacteria; this term had been used as more or less synonymous with bacteria by earlier workers. As more diseases of this sort were discovered, the unknown causative agents came to be called ‘filterable viruses’. Between 1900 and 1935, many plant diseases thought to be caused by filterable viruses were described, but considerable confusion arose because adequate methods for distinguishing one virus from another had not yet been developed. The original criterion of a virus was an infectious entity that could pass through a filter with a pore size small enough to hold back all known cellular agents of disease. However, diseases were soon found that had virus-like symptoms not associated with any pathogen visible in the light microscope, but that could not be transmitted by mechanical inoculation. With such diseases, the

Encyclopedia of Virology, 4th Edition, Volume 3

doi:10.1016/B978-0-12-814515-9.00589-0

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4

An Introduction to Plant Viruses

Table 1

Tulipomania: the goods exchanged for one bulb of Viceroy tulip

4 t of wheat 8 t of rye 4 fat oxen 8 fat pigs 12 fat sheep 2 hogsheads of wine

4 barrels of beer 2 barrels of butter 1000 lb cheese 1 bed with accessories 1 full dress suit 1 silver goblet

criterion of filterability could not be applied. Their infectious nature was established by graft transmission and sometimes by insect vectors. Thus, certain diseases of the yellows and witches’-broom type, such as aster yellows, came to be attributed to viruses on quite inadequate grounds. Many such diseases are now known to be caused by phytoplasma or spiroplasma, and a few by bacteria or rickettsia.

The Biological Age During most of the period between 1900 and 1935, attention was focused on the description of diseases using the macroscopic symptoms and cytological abnormalities as revealed by light microscopy, host ranges, and methods of transmission which were the only techniques available. The influence of various physical and chemical agents on virus infectivity was investigated, but methods for the assay of infective material were primitive. Holmes showed that the local lesions produced in some hosts following mechanical inoculation could be used for the rapid quantitative assay of infective virus. This technique enabled properties of viruses to be studied much more readily and paved the way for the isolation and purification of viruses a few years later. Until about 1930, there was serious confusion by most workers regarding the diseases produced by viruses and the viruses themselves. This was not surprising, since virtually nothing was known about the viruses except that they were very small. In 1931 Smith made an important contribution that helped to clarify this situation. Working with virus diseases in potato, he realized the necessity of using plant indicators, plant species other than potato, which would react differently to different viruses present in potatoes. Using several different and novel biological methods to separate the viruses, he was able to show that many potato virus diseases were caused by a combination of two viruses with different properties, which he named virus X (potato virus X, PVX) and virus Y (potato virus Y, PVY). As PVX was not transmitted by the aphid Myzus persicae, whereas PVY was, PVY could be separated from PVX. He obtained PVX free of PVY by needle inoculation of the mixture to Datura stramonium which does not support PVY. Furthermore, Smith observed that PVX from different sources fluctuated markedly in the severity of symptoms it produced in various hosts leading to the concept of strains. An important practical step forward was the recognition that some viruses could be transmitted from plant to plant by insects. Fukushi recorded the fact that in 1883 a Japanese rice grower transmitted what is now known to be rice dwarf virus, RDV) by the leafhopper Recelia dorsalis. However, this work was not published in any available form and so had little influence. In 1922 Kunkel first reported the transmission of a virus by a planthopper; within a decade, many insects were reported to be virus vectors leading to the recognition of specific virus–vector interactions. Since Fukushi first showed in 1940 that RDV could be passed through the egg of a leafhopper vector for many generations, there has been great interest in the possibility that some viruses may be able to replicate in both plants and insects. It is now well established that plant viruses in the families Rhabdoviridae and Reoviridae and the genera Tenuivirus, Tospovirus, and Marafivirus multiply in insects as well as in plants.

The Biochemical/Biophysical Age Beale's recognition in 1928 that plants infected with TMV contained a specific antigen opened the age in which the biochemical nature of viruses was elucidated. In the 1930s Gratia showed that plants infected with different viruses contained different specific antigens and Chester demonstrated that different strains of TMV and PVX could be distinguished serologically. The high concentration at which certain viruses occur in infected plants and their relative stability turned out to be of crucial importance in the first isolation and chemical characterization of viruses, because methods for extracting and purifying proteins were not highly developed. In the early 1930s, various attempts were made to isolate and purify plant viruses using methods similar to those that had just been developed for purifying enzymes. Following detailed chemical studies suggesting that the infectious agent of TMV might be a protein, Stanley announced in 1935 the isolation of this virus in an apparently crystalline state. At first Stanley considered that the virus was a globulin containing no phosphorus but in 1936 Bawden et al. described the isolation from TMV-infected plants of a liquid crystalline nucleoprotein containing nucleic acid of the pentose type. They showed that the particles were rod-shaped, thus confirming the earlier suggestion of Takahashi and Rawlins based on the observation that solutions containing TMV showed anisotropy of flow. Electron microscopy and X-ray crystallography were the major techniques used in early work to explore virus structure, and the importance of these methods has continued to the present day. Bernal and

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Fankuchen applying X-ray analysis to purified preparations of TMV obtained accurate estimates of the width of the rods. The isolation of other rod-shaped viruses, and spherical viruses that formed crystals, soon followed. All were shown to consist of protein and pentose nucleic acid. Early electron micrographs confirmed that TMV was rod-shaped and provided approximate dimensions, but they were not particularly revealing because of the lack of contrast between the virus particles and the supporting membrane. The application of shadow-casting with heavy metals greatly increased the usefulness of the method for determining the overall size and shape of virus particles but not structural detail. With the development of high-resolution microscopes and of negative staining in the 1950s electron microscopy became an important tool for studying virus substructure. From a comparative study of the physicochemical properties of the virus nucleoprotein and the empty viral protein shell found in TYMV preparations, Markham concluded in 1951 that the RNA of the virus must be held inside a shell of protein, a view that has since been amply confirmed for this and other viruses by X-ray crystallography. Crick and Watson suggested that the protein coats of small viruses are made up of numerous identical subunits arrayed either as helical rods or as a spherical shell with cubic symmetry. Subsequent X-ray crystallographic and chemical work has confirmed this view. Caspar and Klug formulated a general theory that delimited the possible numbers and arrangements of the protein subunits forming the shells of the smaller isodiametric viruses. Until about 1948, most attention was focused on the protein part of the viruses. Quantitatively, the protein made up the larger part of virus preparations. Enzymes that carried out important functions in cells were known to be proteins, and knowledge of pentose nucleic acids was rudimentary. No function was known for them in cells, and they generally were thought to be small molecules primarily because it was not recognized that RNA is very susceptible to hydrolysis by acid, by alkali, and by enzymes that commonly contaminate virus preparations. In 1949 Markham and Smith isolated turnip yellow mosaic virus (TYMV) and showed that purified preparations contained two classes of particles, one an infectious nucleoprotein with about 35% of RNA, and the other an apparently identical protein particle that contained no RNA and that was not infectious. This result clearly indicated that the RNA of the virus was important for biological activity. Analytical studies showed that the RNAs of different viruses have characteristically different base compositions while those of related viruses are similar. About this time, it came to be realized that viral RNAs might be considerably larger than had been thought. A synthetic analog of the normal base guanine, 8-azaguanine, when supplied to infected plants was incorporated into the RNA of TMV and TYMV, replacing some of the guanine. The fact that virus preparations containing the analog were less infectious than normal virus gave further experimental support to the idea that viral RNAs were important for infectivity. However, it was the classic experiments in the mid-1950s of Gierer and Schramm, and Fraenkel-Conrat and Williams that demonstrated the infectivity of naked TMV RNA and the protective role of the protein coat. These discoveries ushered in the era of modern plant virology. In the early 1950s Brakke developed density gradient centrifugation as a method for purifying viruses. Together with a better understanding of the chemical factors affecting the stability of viruses in extracts, this procedure has allowed the isolation and characterization of many viruses. The use of sucrose density gradient fractionation enabled Lister to discover the bipartite nature of the tobacco rattle virus genome. Since that time, density gradient and polyacrylamide gel fractionation techniques have allowed many viruses with multipartite genomes to be characterized. Their discovery, in turn, opened up the possibility of carrying out genetic reassortment experiments with plant viruses leading to the allocation of functions to many of the viral genes. Density gradient fractionation of purified preparations of some other viruses revealed noninfectious nucleoprotein particles containing subgenomic RNAs. Other viruses have been found to have associated with them satellite viruses or satellite RNAs that depend on the ‘helper’ virus for some function required during replication. Further developments in the 1970s included improved techniques related to X-ray crystallographic analysis and a growing knowledge of the amino acid sequences of the coat proteins allowed the three-dimensional structure of the protein shells of several plant viruses to be determined in molecular detail. For some decades, the study of plant virus replication had lagged far behind that of bacterial and vertebrate viruses mainly because there was no plant system in which all the cells could be infected simultaneously to provide the basis for synchronous ‘one-step growth’ experiments. However, following the initial experiments of Cocking in 1966, Takebe and colleagues developed protoplast systems for the study of plant virus replication. Although these systems had significant limitations, they greatly increased our understanding of the processes involved in plant virus replication. Another important technical development has been the use of in vitro protein-synthesizing systems such as that from wheat germ, in which many plant viral RNAs act as efficient messengers. Their use allowed the mapping of plant viral genomes by biochemical means to begin.

The Molecular Biology Age The molecular age opened in 1960 with the determination of the full sequence of 158 amino acids in the coat protein of TMV. The sequence of many naturally occurring strains and artificially induced mutants was also determined at about the same time. This work made an important contribution to establishing the universal nature of the genetic code and to our comprehension of the chemical basis of mutation. Our understanding of the genome organization and functioning of viruses has come from the development of procedures whereby the complete nucleotide sequence of viruses with RNA genomes can be determined. In 1982 the genomes of both the first

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plant RNA virus (TMV) and DNA virus cauliflower mosaic virus (CaMV) were sequenced. Since then, the genomes of representatives of all the plant virus genera have been sequenced and there are many sequences of virus species. The late 1980s and 1990s was a period when molecular biological techniques were applied to a wide range of aspects of plant virology. These included the ability to prepare in vitro infectious transcripts of RNA viruses derived from cloned viral cDNA allowing techniques such as site-directed mutagenesis to be applied to the study of genome function and reverse genetics being used to elucidate the functions of viral genes and control sequences. These approaches, together with others such as yeast systems for identifying interacting molecules, the expression of viral genes in transgenic plants, and labeling viral genomes in such a manner that their sites of function within the cell are known, were revealing the complexities of the interactions between viruses and their hosts. Nucleotide sequence information has had, and continues to have, a profound effect on our understanding of many aspects of plant virology, including (1) the location, number, and size of the genes in a viral genome; (2) the amino acid sequence of the known or putative gene products; (3) the molecular mechanisms whereby the gene products are transcribed; (4) the putative functions of a gene product, which can frequently be inferred from amino-acid-sequence similarities to products of known function encoded by other viruses; (5) the control and recognition sequences in the genome that modulate expression of viral genes and genome replication; (6) the understanding of the structure and replication of viroids and of the satellite RNAs found associated with some viruses; (7) the molecular basis for variability and evolution in viruses, including the recognition that recombination is a widespread phenomenon among RNA viruses and that viruses can acquire host nucleotide sequences as well as genes from other viruses; and (8) the beginning of a taxonomy for viruses that is based on evolutionary relationships. On the host side, advances in plant genome sequencing are identifying plant genes that confer resistance to viruses. During the 1980s, major advances were made on improved methods of diagnosis for virus diseases, centering on serological procedures and on methods based on nucleic acid hybridization. Since the work of Clark and Adams reported in 1977, the enzyme-linked immunosorbent assay (ELISA) technique has been developed with many variants for the sensitive assay and detection of plant viruses. Monoclonal antibodies against TMV lead to a very rapid growth in their use for many kinds of plant virus research and for diagnostic purposes. The late 1970s and the 1980s also saw the start of application of the powerful portfolio of molecular biological techniques to developing other approaches to virus diagnosis, to a great increase in our understanding of the organization and strategy of viral genomes, and to the development of techniques that promise novel methods for the control of some viral diseases. The use of nucleic acid hybridization procedures for sensitive assays of large numbers of samples and the polymerase chain reaction, also dependent on detailed knowledge of genome sequences, are being increasingly used in virus diagnosis. Most recently DNA chips are being developed for both virus diagnostics and studying virus infection. In the early 1980s, it seemed possible that some plant viruses, when suitably modified by the techniques of gene manipulation, might make useful vectors for the introduction of foreign genes into plants. Some plant viruses have been found to contain regulatory sequences that can be very useful in other gene vector systems, notably the widely used CaMV 35S promoter. Another practical application of molecular techniques to plant viruses has been the modification of viral genomes so that products of interest to industry and medicine can be produced in plants. This is being done either by the introduction of genes into the viral genome or by modification of the coat protein sequence to enable epitopes to be presented on the virus. Early attempts (early to mid-1900s) to control virus diseases in the field were often ineffective. They were mainly limited to attempts at general crop hygiene, roguing of obviously infected plants, and searching for genetically resistant lines. Developments since this period have improved the possibilities for control of some virus diseases. Heat treatments and meristem tip culture methods have been applied to an increasing range of vegetatively propagated plants to provide a nucleus of virus-free material that then can be multiplied under conditions that minimize reinfection. Such developments frequently have involved the introduction of certification schemes. Systemic insecticides, sometimes applied in pelleted form at the time of planting, provide significant protection against some viruses transmitted in a persistent manner by aphid vectors. It has become increasingly apparent that effective control of virus disease in a particular crop in a given area usually requires an integrated and continuing program involving more than one kind of control measure. However, such integrated programs are not yet in widespread use. Cross-protection (or mild-strain protection) is a phenomenon in which infection of a plant with a mild strain of a virus prevents or delays infection with a severe strain. The phenomenon has been used with varying success for the control of certain virus diseases, but the method has various difficulties and dangers. In 1986 Powell-Abel and co-workers considered that some of these problems might be overcome by the application of the concept of pathogen-derived resistance of Sandford and Johnston. Using recombinant DNA technology, they showed that transgenic tobacco plants expressing the TMV coat-protein gene either escaped infection following inoculation or showed a substantial delay in the development of systemic disease. These transgenic plants expressed TMV coat-protein mRNA as a nuclear event. Seedlings from self-fertilized transformed plants that expressed the coat protein showed delayed symptom development when inoculated with TMV. Thus, a new approach to the control of virus diseases emerged. However, this approach revealed some unexpected results which led to the recognition that plants have a defense system against ‘foreign’ RNA. This defense system, initially termed post-translational gene silencing and now called RNA silencing or RNA interference (RNAi) was first recognized in plants and had, and is still having, a great impact on molecular approaches as diverse as disease control and understanding gene functions. Among the tools that have arisen from understanding this new phenomenon is virus-induced gene silencing which is being used to determine the functions of genes in plants and animals by turning them off. However, the RNAi defense system in plants which targets double-stranded RNA, an intermediate RNA virus replication, raised the question of how RNA viruses replicated in plants. Studies on gene functions revealed that many plant viruses contain so-called ‘virulence’ genes. Many of these have been shown to suppress host RNA silencing, thus overcoming the defense system.

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Suppression of gene silencing is widespread among plant viruses and examples are being found in animal viruses. Thus, as noted at the beginning of this article, plant virology is still providing insights onto phenomena applicable to virology in general.

See also: Plant Antiviral Defense: Gene-Silencing Pathways. Plant Viral Diseases: Economic Implications. Tobacco Mosaic Virus (Virgaviridae). Vector Transmission of Plant Viruses. Viral Suppressors of Gene Silencing. Virus-Induced Gene Silencing (VIGS)

Further Reading Baulcombe, D.C., 1999. Viruses and gene silencing in plants. Archives of Virology 15 (supplement), 189–201. Caspar, D.L.D., Klug, A., 1962. Physical principles in the construction of regular viruses. Cold Spring Harbor Symposium on Quantitative Biology 27, 1–24. Clark, M.F., Adams, A.N., 1977. Characteristics of the microplate method of enzyme-linked immunosorbent assay for the detection of plant viruses. Journal of General Virology 34, 475–483. Fischer, R., Emans, N., 2000. Molecular farming of pharmaceutical proteins. Transgenic Research 9, 279–299. Hull, R., 2001. Matthews’ Plant Virology. San Diego: Academic Press. Li, F., Ding, S.W., 2006. Virus counterdefense: Diverse strategies for evading the RNA silencing mechanism. Annual Review of Microbiology 60, 507–531. Lindbo, J.A., Dougherty, W.G., 2005. Plant pathology and RNAi: A brief history. Annual Review of Phytopathology 43, 191–204. Pavord, A., 1999. The Tulip. London: Bloomsbury. Van der Want, J.P.H., Dijkstra, J., 2006. A history of plant virology. Archives of Virology 151, 1467–1498.

Emerging and Re-Emerging Plant Viruses Sabrina Bertin, Francesco Faggioli, Andrea Gentili, Ariana Manglli, Anna Taglienti, Antonio Tiberini, and Laura Tomassoli, Council for Agricultural Research and Economics, Research Center for Plant Protection and Certification, Rome, Italy r 2021 Elsevier Ltd. All rights reserved. This is an update of G.P. Martelli, D. Gallitelli, Emerging and Reemerging Virus Diseases of Plants, In Encyclopedia of Virology (Third Edition), Brian W.J. Mahy, Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00705-6.

Nomenclature

MP Movement protein NC Nucleocapsid NSm Non-structural movement protein NSS Non-structural RNA silencing suppressor nt Nucleotide(s) ORF Open reading frame RB Resistance breaking RdRp RNA-dependent RNA polymerase sgRNA Sub-genomic RNA ssRNA Single-stranded ribonucleic acid

aa Amino acid(s) asl Above sea level BSC Bundle sheath cells CP Coat protein or capsid protein HC-Pro Helper component-protease HR Hypersensitive response HTS High-throughput sequencing kb Kilobase kbp Kilobase pair kDa Kilo Dalton

Glossary Emergence An emergent virus is a virus that has adapted and/or emerged as a new disease/pathogenic strain, with attributes facilitating pathogenicity in an environment normally not associated with that virus. Isolate Unique sampling material out of which a given virus has been identified.

Pathosystem Subsystem of an ecosystem that is defined by the phenomenon of parasitism, in the present case, viruses. Strain Virus strains are viruses that belong to the same species and differ in having stable and heritable biological, serological, and/or molecular characters.

Introduction Plant viruses have been considered one of the most limiting biotic stresses, and their intrinsic and epidemiological features require specific containment measures aimed either to prevent (i.e., use of virus-free germplasm), eradicate (i.e., destruction of infected material) or control them (i.e., control of vectors and use of resistance genes). Several outbreaks are recorded every year worldwide mainly due to: (1) new viruses (i.e., first detected within the last five-ten years and persisting within restricted areas); (2) emerging viruses (i.e., whose incidence has rapidly increased within the last decade); (3) re-emerging viruses (i.e., viruses facing genetic evolution and host shift). The key factors driving the emergence of plant viruses include genetic variability of a viral population, changes in agricultural practices and/or distribution of insect vectors. Further, these factors are likely modified by the effects of climate change which, in recent years, is a major concern for plant pathologists proving to be a global problem of difficult resolution. According to the Food and Agriculture Organization (FAO), the climate change is among the driving factors of global hunger and malnutrition. It is reported that damage caused by plant pests is responsible for loss of 20%–40% of global food production. In view of this, it is easy to highlight how climate change, the emergence of new and more serious plant diseases and global malnutrition are closely related. In addition, agricultural globalization, international trade and long-distance transport of plant materials, often associated with people migration flow, are relevant for virus spreading, leading to the introduction of viruses and their vectors to new geographical regions with unpredictable consequences at local level. In fact, epidemiological characteristics of viral pathosystems are variables depending on different agro-ecological contexts, hence the control strategy of a well-known disease in a new area can be quite challenging. For example, the introduction of a virus in a new region where highly susceptible hosts are usually cropped or, conversely, the import of such susceptible hosts in a region where the virus is already spread, can lead to severe viral outbreaks. Moreover, in such cases, early response from scientists, plant protection services and policy makers is needed, as delayed diagnosis, prevention and control measures can be inefficient. For the above reasons, the list of new, emerging and re-emerging viruses (Table 1) needs to be often updated due to the quick environmental changes, global market and crop production. Some emergent viruses belong to important genera and families as well as the Begomovirus (Geminiviridae) within which some species suddenly increase in rate of spread, causing outbreaks and disease in new countries (i.e., tomato leaf curl New Delhi virus, tomato yellow leaf curl viruses, cotton leaf curl viruses, East African

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Table 1 Emerging and re-emerging viruses reviewed in the present Encyclopedia of Virology. Gray cells indicate other articles in the Encyclopedia Order

Family

Genus

Species

Bunyavirales Fimoviridae Emaravirus Rose rosette virus (RRV) – Tospoviridae Orthotospovirus Tomato spotted wilt virus (TSWV) Ortervirales Caulimoviridae Badnavirus Grapevine Roditis leaf discolorationassociated virus (GRLDaV) – – – Cacao swollen shoot disease (CSSD) Tymovirales Alphaflexividae Potexvirus Pepino mosaic virus (PepMV) – Betaflexiviridae Trichovirus Grapevine Pinot gris virus (GPGV) – Potyviridae Ipomovirus Cassava brown streak virus (CBSV) Unassigned Virgaviridae Tobamovirus Cucumber green mottle mosaic virus (CGMMV) – – – Tomato brown rugose fruit virus (ToBRFV) Unassigned Nanoviridae Babuvirus Banana bunchy top virus (BBTV) – Geminiviridae Begomovirus East African cassava mosaic virus (EACMV) – – – Squash leaf curl virus (SLCV) –

–

–

–

–

–

Geographical spread

Status

Canada, USA, India Worldwide Europe: Greece, Italy, Croatia, Turkey

Emerging Re-emerging Emerging

West Africa Worldwide Europe, Americas, Asia, Australia Africa Europe, USA, Canada, Australia

Re-emerging Emerging Emerging Emerging Emerging

Middle East, Europe, Mexico, USA, China

Emerging

Africa, Asia Africa, India, South East Asia

Emerging Emerging

North and Central America, Saudi Arabia, Middle East, Egypt India, Pakistan, Bangladesh, Iran, Thailand, Tomato leaf curl New Delhi virus Indonesia, Spain, Italy, Morocco (ToLCNDV) Tomato yellow leaf curl virus (TYLC) Europe, Asia, Africa, North and Central America, Australia

Emerging Emerging Re-emerging

cassava mosaic viruses, etc.) or have the capability to produce variants by recombinant events (i.e., tomato yellow leaf curl disease, East African cassava mosaic virus) causing enhancement of virulence and resistance breaking in plants. This article reports the main viral diseases, not treated in other articles, that are currently threatening farmers, scientific community, regulators, agricultural industry and even final consumers worldwide.

Vegetable Viruses Tobamoviruses Tobamovirus is a genus in the family Virgaviridae, reported to infect mainly tobacco, potato, tomato, and cucurbits. To date, thirtyseven virus species are included in this genus. According to molecular features and host range, four informal subgroups have been identified within this genus: brassicas, cucurbits, malvaceous and solanaceous group. Tobamovirus is considered one of the most important virus genera and the type species member, Tobacco mosaic virus has been the first virus ever discovered and characterized, leading to modern plant virology.

Cucumber Green Mottle Mosaic Virus Geographical distribution Cucumber green mottle mosaic virus (CGMMV) is an economically important pathogen infecting many cucurbit species. It was first described in cucumber plants in England in 1935. After a first period of slow diffusion, the virus has begun to cause disease more frequently after 1986. Since 2007, CGMMV represents a serious concern for the protected crops of cucumber and other cucurbits. In recent years, it rapidly has spread across European countries where it was already known and in new countries such as Bulgaria (2008) and Poland (2017). Outbreaks of CGMMV were also reported in Canada (2014), the USA (2014) and Australia (2015) with a rapid spread in different locations. To date, the virus occurs in almost all continents and over 40 countries, achieving a global distribution.

Causal agent and classification CGMMV has a 6.4 kb single-stranded, positive sense RNA ( þ ssRNA) genome encapsidated within 2000 molecules of the capsid protein (CP) resulting in a stiff, rod-shaped 300  18 nm particle. The genomic RNA contains four protein-coding open reading frames (ORFs). The first two putative ORFs encode a 129 kDa protein containing the methyl-transferase and helical domains, and a 186 kDa protein that contains the polymerase domain respectively. Both proteins are translated directly from the genomic RNA and are involved in RNA replication. ORF3 and ORF4 are translated from subgenomic (sg) RNAs and encode a 29 kDa protein involved in cell-to-cell movement (MP) and a 17.4 kDa CP respectively. Serological and molecular variability has been reported between European and Asian isolates.

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Disease symptoms and yield losses Symptoms caused by CGMMV vary depending upon cucurbit species and cultivars that are grown. In cucumber it causes leaf mottle mosaic, blistering, fruit discoloration, mottling and distortion. In watermelon, leaf mottling and mosaic are observed in the young leaves as well as brown necrotic lesion in the stems and peduncles (Fig. 1). Moreover, CGMMV causes a pulp deterioration called blood flesh disease and fruits lose their marketable value. In melon young leaves develop initial mottle and mosaic often disappearing in mature foliage. Fruits develop different degrees of malformation, mottling and surface netting. Pumpkin, squash and zucchini infected leaves can show leaf mottling, mosaic or may also be asymptomatic. Pumpkin fruits are always asymptomatic, while squash and zucchini fruits are externally symptomless but internally discolored and necrotic. In cucurbit seedlings symptoms are not visible but in severe infections from seed the cotyledons become yellow. Symptom development is influenced by environment, growing condition, plant growth stage at the time of infection or viral strain. Lower temperatures in early spring with low light intensity induce more severe symptoms than in summer. The damage caused by CGMMV consists in yield losses in the field and poor quality of the fruits. Serious damages are reported in cucumber and watermelon crops with yield losses up to 15% and 40%, respectively.

Epidemiology Like other species belonging to the genus Tobamovirus, CGMMV is very stable and contagious. It can remain infectious for a long period in the soil contaminated with infected plant debris, in contaminated equipment surface, or on farming tools. It is easily transmitted through direct contact, seed, and other propagation material including grafting. Seed transmission is known to occur in eight different cucurbit species: watermelon, melon, cucumber, pumpkin, zucchini, bottle gourd and snake gourd. The virus contaminates the coat of the seed from infected plants. Studies reported different rates of seed transmission reaching up to 8% in bottle gourd, cucumber and watermelon. The transmission rate may decline rapidly after a few months during seed storage. Contaminated seed stocks and infected seedlings can constitute the primary source of inoculum in a new area of production. Once introduced, the virus readily spreads mechanically during cultural operations, through contaminated instruments, hands and clothing, and through direct contact between the plants. CGMMV is transmitted also through irrigation. Insect transmission has been investigated but no insect vector is known to transmit the virus in a specific manner. CGMMV host range is mainly restricted to the family Cucurbitaceae. The natural host range includes crops as Benincasa hispida, Citrullus lanatus, Cucumis sativus, C. anguria, C. melo, Cucurbita maxima, C. moschata, C. pepo, Lagenaria siceraria, Luffa acutangula, L. cylindrica, Momordica charantia, Trichosanthes cucumerina, and weeds as Citrullus colocynthis, Ecballium elaterium and Mukia maderaspatana. In the recent years, it has been naturally reported in non-cucurbit species in the families Amaranthaceae, Apiaceae, Boraginaceae, Portulacaceae, Euphorbiaceae and Solanaceae, which can act as reservoirs.

Control Use of virus-free seeds and seedlings is essential to avoid virus introduction in new areas. As virus contaminates mainly the tegument, seed treatments based on dry heat (i.e., 701C for 3 days) or disinfectants can be used without adversely affecting seed germination. In the seedling nurseries it is very important to practice stringent hygiene measures. Early detection and removal of infected plants is crucial to avoid the virus spread. Once the virus becomes established in a farm, all organic and contaminated disposable material should be destroyed by incineration or deep burial. All contaminated surfaces and irrigation systems should be cleaned with antiviral detergent or chemical disinfectant. Considering the high stability and infectivity of the virus, crop rotation avoiding any cucurbits for a minimum period of at least 24 months is recommended. Resistance to CGMMV has been reported in melon and cucumber, even if commercial varieties mainly display partial resistance (tolerance) resulting in a reduced symptom expression.

Fig. 1 Leaf mosaic and blistering of CGMMV on cucumber.

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Tomato Brown Rugose Fruit Virus Geographical distribution Tomato brown rugose fruit virus (ToBRFV) was originally described in Jordan in 2015 and in Israel in 2014 (but reported only in 2017) on tomato plants harboring the tomato mosaic virus (ToMV) resistance Tm-22 gene. In 2018–19, ToBRFV outbreaks have been reported in countries on almost all the continents: Mexico, California (USA), China (Shandong district), Germany, Greece (Crete island), Italy, the Netherlands, Spain, Turkey and United Kingdom. In Mexico, Jordan and Italy ToBRFV was also reported on pepper plants not harboring L resistance to TMV. In view of its quick spreading and ability to overcome ToMV resistance, ToBRFV is considered an emerging pathogen, included in the EPPO Alert List in January 2019, potentially spreading worldwide, and it was suddenly regulated to contrast its introduction in free areas and spread where present. To date, ToRBFV was reported to be established in Mexico (53 provinces among 20 states), and in Israel and Jordan, where virus incidence can reach up to 100% in the greenhouse crop production.

Causal agent and classification Tomato brown rugose fruit virus has structural and genomic features that are typical for the members of the genus Tobamovirus. The genome, encapsidated in a 300 nm long rod-shaped rigid particle, is constituted by a positive ssRNA of B6400 nucleotides (nt) and consists of four ORFs. ORF1 and ORF2 encode two replication-related proteins of 126 and 183 kDa respectively, the latter being expressed by the partial suppression of the stop codon; ORF3 encodes the MP of 30 kDa, and ORF4 the CP of 17.5 kDa which is expressed via the 30 coterminal sgRNAs. Phylogenetic analysis showed that ToBRFV clustered distinctly from either ToMV or TMV clades. In addition, preliminary sequences analysis showed a low level of mutation rate among isolates of ToBRFV, and conserved nt sequences have been retrieved in isolates that have a different geographic origin.

Disease symptoms and yield losses On tomato, the symptomatology includes chlorosis, mosaic and mottling on foliage, often leaf narrowing (needles) (Fig. 2) or more severe malformation; yellow and brown spots, uneven ripening and rarely rugose patches are observed on fruits which dramatically decrease their commercial value. Asymptomatic fruits were reported in several European countries and yield reduction is recorded due to a reduced number of branches and fruits. The occurrence and severity of symptoms vary by variety and environmental conditions (temperature and light intensity). The disease may result in yield losses of up to 70%. Plants for planting are generally asymptomatic. On pepper, foliar symptoms include deformation, yellowing and mosaic; fruits are deformed, with yellow or brown areas or green stripes.

Epidemiology ToBRFV, as all tobamoviruses, is mechanically transmitted by contact. This transmission manner includes different patterns such as plant-to-plant contact, grafting, anthropic transmission by contaminated tools, hands and clothing during cultural practices (transplanting, pruning, staking, trellising, tying, spraying, and harvesting) as well as commercial packing and trade operations. ToBRFV was found in seed coats, and there is reason to suspect seed transmission early infecting emerging plantlets, but additional experimental trials should be carried out to confirm. Nevertheless, even if transmission rate from seed to seedling is low, further dissemination by contact (e.g., during transplantation of seedlings or regular handling of the crop) allows a rapid spread within a glasshouse. In addition, ToBRFV could be carried by bumblebees (Bombus terrestris) and transmitted to healthy tomato plants during pollination (mechanically by visiting flowers of healthy tomato plants). As reported for all tobamoviruses, ToBRFV is characterized by very stable particles. In fact, preliminary studies showed that ToBRFV can survive in crop debris, in soil, and on implements, stakes, trellis wires, fruit containers, greenhouse benches and seedling trays for a long period (several weeks to months). Tomato (Solanum lycopersicum) and pepper (Capsicum spp.) are the main hosts of ToBRFV. By experimental inoculations, symptoms were showed in Nicotiana benthamiana, N. glutinosa, N. sylvestris, N. tabacum, Petunia hybrida, Datura stramonium, Chenopodium spp. In addition, ToBRFV is reported to infect weeds such as Chenopodiastrum murale and Solanum nigrum, that can

Fig. 2 Symptoms of mosaic and needles of ToBRFV on tomato.

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therefore act as reservoirs. In experimental studies, aubergine (Solanum melongena) and potato (S. tuberosum) did not show symptoms after inoculation, and ToBRFV was not found when the plants were subsequently tested for latent infection detection.

Control Regulations have been introduced to prevent entries of ToBRFV through seeds and/or plants for planting of tomato and pepper (i.e. EU Commission Decision 2019/1615; WTOs – Emergency phytosanitary measures). Certification of seeds by molecular tests and only seedling from certified seeds are required for import/export. Strict sanitation practices aiming to prevent ToBRFV establishment indoor (treatment of seeds, sanitation of clothing, tools and equipment, removal and destruction of infected debris) and its spread (packaging area out of production sites, sanitation of boxes for harvesting) are mandatory.

Orthotospoviruses Orthotospovirus is a genus of the family Tospoviridae (order Bunyavirales). These thrips-transmitted viruses are amongst the most devastating plant viruses worldwide, causing severe economic damage to many vegetable crops, such as tomato and sweet pepper. The often wide and overlapping host ranges, emergence of resistance-breaking strains, circulative and propagative transmission by thrips, and difficulties in predicting their outbreaks cause high concern for their control.

Tomato Spotted Wilt Virus The evolution of new tomato spotted wilt virus (TSWV) variants by mutation poses high concern for the capacity to adapt to tomato and pepper varieties carrying the resistance genes Sw5 and Tsw respectively. For this reason, TSWV may be considered a re-emerging pathogen and included in the present article as a potential threat for the vegetable crops.

Geographical distribution Tomato spotted wilt virus (TSWV) is present in all countries under temperate, tropical and subtropical climate conditions, being a serious and endemic pathogen since almost the last 20 years. Since the first description of “spotted wilt” disease in Australia in 1915, the spreading of TSWV became uncontrolled with outbreaks occurring in vegetable crops, such as tomato, lettuce and pepper, tobacco and in ornamentals especially in southern Europe. A threatening resurgence of TSWV occurred since 1980 in North America, 1987 in Western Europe and 1993 in Australia, leading to a worldwide distribution.

Causal agent and classification TSWV, as other members of the Orthotospovirus genus, is enveloped including additional protein package on its RNA genome, which is segmented in three portions named S, M and L, according to the size of the RNA molecules, 2.9 kb, 4.8 kb, and 8.9 kb, respectively. The L segment encodes putative RNA-dependent RNA polymerases (RdRp), such as replicase, transcriptase, nuclease, helicase, cap-binding and NTPase proteins responsible for several enzymatic functions of TSWV. The M and S RNAs have special structures where genomes are ambisense, both including parts of positive and parts of negative polarities. In particular, the M segment produces envelope glycoproteins (Gn-Gc) and in antisense orientation non-structural movement protein (NSm). The S segment has two ORFs encoding nucleocapsid (NC) protein and non-structural RNA silencing suppressor (NSS), with antisense and sense orientation, respectively.

Disease symptoms and yield losses TSWV can induce a wide variety of symptoms on economically important ornamental plants, vegetables and industrial crops. The symptomatology expression is affected by plant species and cultivar, the developmental stage at the time of inoculation, the nutritional and environmental conditions (outdoor or greenhouse cultivations) as well as TSWV strain. Generally, symptoms on leaves include chlorotic or necrotic lesions, ring spots, concentric rings, line patterns together with crinkling, bronzing, distortion (curling) of leaves and a general plant disorder as wilting, stunting, mottling and necrosis. Symptoms on fruits usually could include irregular discoloration, yellow or brown flecks or rings, or necrotic lesions or rings on unripe fruits (Fig. 3). Affecting both yield and marketing quality, TSWV has been reported as the second most destructive viral pathogen in the list of economic damage caused by plant viruses. In fact, in tomato, it causes a yield and marketable value reduction up to 42% and about 95% respectively, with worldwide estimated losses exceeding one billion dollars annually. The economic impact of TSWV is great, due to its wide geographical distribution and to its broad host range. Crops in which important losses due to TSWV have been reported are tomato, pepper, lettuce, eggplant, papaya, French beans, celery and ornamental plants.

Epidemiology TSWV is naturally transmitted by at least eight species of thrips, in a circulative and propagative manner. Analysis of factor determining vector competence showed Frankliniella occidentalis (Pergande) as the most efficient vector. In fact, the western flower thrips, F. occidentalis, contributed to the spreading and worldwide occurrence of TSWV starting from 80s. TSWV is characterized by a wide host range, and this relative lack of host specificity along with highly polyphagous nature of its vector are the two main factors that make TSWV one of the most widely distributed plant viruses. A recent report included more than 1000 plants species

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Fig. 3 Fruit malformation and irregular discoloration in pepper infected by TSWV. Courtesy of M. Turina.

(exactly 1090) capable of being locally and/or systematically infected by TSWV. The species are from a single family of the class Pteropsida (Division X - Pteridophyta), 69 families of the class Dicotyledones (Division XII - Angiospermae) and 15 families of the class Monocotyledones (Division XII - Angiospermae). Most of the plants listed as hosts of TSWV belong to the family Asteraceae (247 species), Solanaceae (172 species) and Fabaceae (60 species).

Control Prevention and control of the spread of TSWV is very difficult due to its vector which needs a complex integrated program itself for management. As for other plant viruses, virus-free planting material is a prerequisite to prevent TSWV in non-resistant cultivars. Cultural practices play an important role in controlling disease spreading as removing weeds and infected plants, destruction or removal of old crops. Additionally, density row planting, cultivar selection, date of planting, reflective mulches could have powerful effects on vectors, although optimum levels differ crop by crop, and by conditions of field and climate. Direct vector control by insecticides is needed but establishing appropriate rotations and employing products from different mode of action groups as essential to prevent the development of insecticide resistance on the target insect populations. An alternative indirect strategy to mitigate natural thrips density and damage is the use of predatory insects such as the minute pirate bugs (Orius insidiosus) and big eyed bugs (Geocoris punctipes), mainly successful in protected crops. Host plant resistance is probably the most effective way to control this virus. Natural resistance to TSWV has been reported for different crops, including chrysanthemum, lettuce, pepper and tomato. Currently, two single dominant resistance genes against TSWV, Sw-5 in tomato and Tsw in pepper, are known and both provide a hypersensitive response (HR) type of resistance. This means that resistant plants react with an HR against virus infection resulting in necrotic local lesions.

Resistance breaking strains In recent years, increasing occurrence of TSWV strains characterized by a phenotype able to break resistance (RB strains) in resistant plants were recorded, even in natural conditions. Emergence of RB isolates of TSWV seems to be correlated with the extensive cultivation of resistant hybrids. From 2011 (Spain) resistant tomato plants seriously infected with TSWV were reported widely in most of the cultivation areas where TSWV is endemic and resistant varieties are largely used. In 2016, in California (USA) unusually early and severe symptoms of TSWV occurred in fields of TSWV-resistant (Sw-5) fresh market tomato cultivars. The occurrence of RB TSWV strains in California occurred 5–6 years after ever-increasing planting of Sw-5 cultivars in hot spot areas, included in the local integrated pest management strategy. The extreme variability of TSWV isolates, joined with the possibility of the exchange of genetic information through reassortment of genome segments, is suggested as the main cause for the apparent readiness of the virus to adapt and overcome both natural and pathogen-mediated resistance.

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Begomoviruses The genus Begomovirus of the family Geminiviridae is the largest group of DNA plant-infecting viruses comprising 409 recognized species. Begomoviruses are transmitted by the whitefly Bemisia tabaci (Gennadius) and cause disease in dicotyledonous plants. In the warmer parts of the world, these viruses are responsible of significant yield losses for economically important vegetable and fiber crops. This genus comprises viruses of monopartite and bipartite single-stranded DNA genome encapsidated in geminate particles of B 30  18 nm consisting of two incomplete icosahedra. Two groups of begomoviruses are reported: “New World” with bipartite genome and “Old World” comprising either bipartite or monopartite genomes.

Squash Leaf Curl Virus Two strains named squash leaf curl virus-extended (SLCV-E) and squash leaf curl virus-restricted (SLCV-R) based on their host range and symptom severity have been identified. According to molecular analysis, the two strains were successively recognized as separate (but closely related) species: SLCV-E remained SLCV, while SLCV-R was named squash mild leaf curl virus (SMLCV).

Geographical distribution Squash leaf curl disease (SCLD) was described for the first time in squash in California (1981) and in cultivated buffalo gourd in Arizona (1986). Since at least 1977, both SLCV and SMLCV were reported in the southwestern United States and northern Mexico and successively in Texas in watermelon plants. SLCV remains restricted in central and north America until 2000, when it was also found in Saudi Arabia and rapidly spread in the Middle East and other countries: Israel (2003), Egypt (2006), Jordan (2008), Palestine (2010), Lebanon (2012).

Causal agent and classification SLCV is a typical “New World” begomovirus with bipartite genome DNA-A and DNA-B, each approximately 2.6 kb in size with different sequences except for an intergenic region (RI) also called as common region (CR) involved in replication initiation. DNA-A is essential for replication and encapsidation. DNA-A encodes five ORFs, one (AV1-coat protein) in the viral sense and the other four (AC1-replication initiator, AC2-transcriptional activator, AC3-replication enhancer protein and AC4-within the AC1) in the complementary sense. DNA-B encodes two ORFs, BV1- nuclear shuttle protein in viral sense and BC1- movement protein in complementary sense. DNA-B is essential in systemic movement of the virus and symptoms production.

Disease symptoms and yield losses SCLV causes different symptoms according to the host, cultivar or growing stage. In the squash plants, it causes stunting, severe leaf curling, yellow mottling, blisters, and fruit deformation (Fig. 4). Similar symptoms can occur also in other cucurbits as melon and watermelon. In common beans it causes leaf curling, vein and stem necrosis. Leaf curling, yellowing and stunting of whole plants can occur also in Malva parviflora. In cucurbit crops, severe epidemics with almost 100% of virus incidence were reported in Israel. Higher incidence up to 95% was also reported from different growing areas in Jordan.

Epidemiology The virus is transmitted in the field by B. tabaci, especially MEAM1 (formerly named biotype B) in a persistent circulative and nonpropagative manner. Once acquired the virus is retained in the vector for a period of several weeks, often for the entire life during which the vector remains viruliferous to transmit the virus. Different weed species reported until now to be naturally infected by SLCV suggest that these weeds can serve as reservoir for the virus/vector during the year. In international trade the virus can be introduced through transplant movement and mainly by its vector B. tabaci on host plants, considering the wide host range of the vector and the virus persistence in the vector for a long time after acquisition (several weeks to whole life). The virus infects plants among the Cucurbitaceae as squash (Cucurbita pepo), melon (Cucumis melo), watermelon (Citrulus lanatus), cucumber (C. sativus), giant pumpkin (C. maxima), pumpkin (C. moschata), buffalo gourd (Cucurbita foetidissima) and wild cucurbits (i.e., Ecballium elaterium). In addition to cucurbits, the virus has been reported to infect common bean (Phaseolus vulgaris) and other weeds such as Chenopodium murale, Datura stramonium, Prosopis farcta, Convolvulus sp. and Malva sp.

Control Outbreaks of the virus disease are associated with high population of B. tabaci. The virus control is closely related with the control of its vector B. tabaci, by chemical and physical methods in order to reduce whitefly density. Systemic insecticides can be applied to seedlings before transplanting and during the growing season. Row covers and reflective mulches can be used to protect plants from whiteflies early in the season. Other agricultural practices as a free cucurbit period of at least two-three months and weeds control can remove the reservoir hosts for virus and vector. To prevent the virus introduction and its spread in new areas the use of virus-free and whitefly-free transplants is very important. In cucumber, melon, squash and for some strains of the virus in watermelon, host resistance to SLCV has been identified but this resistance has not been widely introduced into commercial varieties. Commercial zucchini varieties with intermediate resistance (tolerance) to SLCV are available.

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Fig. 4 Stunting and leaf malformation caused by SLCV in zucchini plant. Courtesy of G. Anfoka.

Grapevines Viruses Grapevine Roditis Leaf Discoloration-Associated Virus Geographical distribution Grapevine Roditis leaf discoloration (GRLD) disease was first described in Greece in 1989 on 4-years old grapevines (Vitis vinifera) cv. Roditis, a red berry cultivar, grafted on 110R rootstock. The etiology of the disease remained unknown for more than 25 years until 2015, when a High-Throughput Sequencing (HTS) platform was used for sequencing siRNAs from samples of vine cv. Roditis showing typical GRLD symptoms. HTS analysis detected a DNA virus which had not been described yet; further studies showed that it was closely related to GRLD disease, hence it was given the name of Grapevine Roditis leaf discoloration-associated virus (GRLDaV). Since then, GRLDaV has been detected in a limited number of symptomatic and asymptomatic samples of different cultivars in Greece, Italy, Croatia and Turkey, mainly by HTS techniques; only in the latter country, positive samples were detected by Sanger sequencing of PCR amplicons. Since October 2018, GRLDaV has been added to the EPPO Alert List.

Disease symptoms and yield losses Symptoms mostly appear on young leaves in late summer, consisting of yellow and/or reddish discolorations along the veins, on the interveinal areas or in wide or small surface area of the leaf blade (Fig. 5). Symptomatic leaves are also deformed, reduced in size and display abnormal venation and downward curling on the discolored sectors. Yield losses are associated with this disease, both in quantity and quality of grapes: in infected plants, bunches decrease in number and size and do not reach complete ripening and full color; a decrease in sugar content in yielded berries is also reported. In Italy (Apulia region) GRLDaV was detected in asymptomatic plants of ‘Bombino nero’.

Causal agent and classification GRLDaV is a member of the genus Badnavirus, family Caulimoviridae and displays the closest relationship with fig badnavirus 1. It is a circular dsDNA virus having non-enveloped bacilliform particles, B 130  30 nm, and containing a genome of 6988 nt, which is organized in 4 ORFs. ORF3 encodes a polyprotein characterized by motifs involved in replication, encapsidation and movement function of the virus, while the other three ORFs encode proteins whose role is yet unknown.

Epidemiology The only known natural host of GRLDaV is V. vinifera and graft transmission was confirmed; the virus is mechanically transmissible to grapevine and to some herbaceous test plants (Chenopodium quinoa, Gomphrena globosa and Nicotiana benthamiana). On

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Fig. 5 Yellow discoloration of grapevine leaves by GRLDaV. Courtesy of N. Katis.

the basis of the transmission manner of other badnaviruses and the natural spreading of the disease observed in Greece in late 80s, transmission by mealybug vectors (family Pseudococcidae) is likely, but remains still unconfirmed. Such studies also assessed the disease incidence at very low level (less than 1%). To date, GRLDaV-infected grapevine propagative material remains the main way of virus spreading over long distances. The actual impact of GRLDaV on grapevine cropping is difficult to assess, due to the limited number of positive samples detected worldwide and insufficient information on its biology, epidemiology and distribution; nonetheless, due to the major cultural and economic importance of grapevine, surveys for the presence of GRLDaV in grapevinegrowing countries are recommended.

Control The good health status of propagative material is the main preventive measure to mitigate impact of virus diseases. In viticulture, the implementation of compulsory certification programs and the use of certified virus-free propagative material (scionwood and rootstocks) are playing a key role in a healthy grapevine production reducing the inoculum potential for further spread of the disease. To date, GRLDaV is not included in such programs but, in order to avoid its further diffusion, grapevine-growing countries where GRLDaV is present should consider extending regulation also for this virus.

Grapevine Pinot Gris Virus Geographical distribution Grapevine Pinot gris virus (GPGV) was first identified and characterized in northern Italy in 2012 where symptoms were observed since 2003, particularly in ‘Pinot gris’. During the last years the disease was reported in many other European countries, in North and South America (Brazil, Canada, Chile, the United States), Asia (China, Georgia, Korea, Pakistan) and Oceania (Australia).

Disease symptoms and yield losses The disease appears with chlorotic mottling, puckering and deformation of the leaves and causes a general growth disorder of plants, with delayed budburst and shortened shoot internodes (Fig. 6). Often, infected plants show most visible symptoms at the beginning of the vegetative revival (spring) while during summer they produce symptomless shoots and leaves. Reduced yield due to low grapes weight and number has been reported. The association between the symptomatology and the presence of GPGV is unclear, indeed the virus is often detected in symptomless plants. Moreover, GPGV causes a higher titratable berry acidity, so that wine produced from infected grapes presents altered flavor and aroma.

Causal agent and classification GPGV is a positive-sense ssRNA virus belonging to the family Betaflexiviridae, genus Trichovirus. The viral genome is of 7259 nt and is organized in 3 ORFs: ORF1 encoding a polypeptide of 214 kDa including methyl-transferase, helicase and RdRp; ORF2 econding the MP of 46 kDa and ORF3 the 22 kDa CP. Viral particles are filamentous and localized in the bundle sheath cells (BSCs) of the phloem parenchyma.

Epidemiology The main way of GPGV spread seems to be related to the use of infected propagation material probably facilitated by using asymptomatic mother plants. GPGV is also transmitted by grafting and by the eriophyid mite Colomerus vitis, although this second pathway of diffusion seems to have a much lower incidence. Silene latifolia subsp. alba (Mill.) and Chenopodium album L. were found infected by the virus suggesting these weeds could act as reservoir plants.

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Fig. 6 Growth reduction and yellowing of a GPGV infected grapevine plant.

Control The unclear role of C. vitis, due to its low efficiency of transmission and limited mobility, does not suggest the control of the vector as a resolutive measure to contrast the diffusion of the disease. The use and trade of infected propagation material are the main ascertained causes of rapid and global dispersal of GPGV worldwide. Therefore, a good control strategy should be focused on developing GPGV-free certified germplasm. To date, GPGV is not subjected to phytosanitary regulation in Europe, whereas it is considered a quarantine pathogen in Australia.

Staple Crops: Cassava Brown Steak Disease Geographical distribution Cassava (Manihot esculenta Crantz, family Euphorbiaceae) is vulnerable to at least 20 different viruses, of which those causing cassava mosaic disease (CMD) and cassava brown streak disease (CBSD) are the most economically important. At the turn of the 21st century, CBSD re-emerged and rapidly spread at an alarming rate in both East and Central Africa, threatening the food security of millions of cassava farmers and attracting intense scientific interest. The first report of CBSD was from northern coastal areas of Tanzania in 1935. Next reports by 1950 noted that the affected areas were almost entirely restricted to coastal areas of East Africa from north eastern Kenya to northern Mozambique and it was believed that the disease did not spread at altitudes over 1000 m above sea level (asl). The first systematic countrywide assessment of CBSD was completed in 1994 and high CBSD incidences in areas of Tanzania, Mozambique and Malawi were reported. However, the view that CBSD is a lowland disease remained unchanged until 2004, when a significant spread of CBSD occurred in central and southern Uganda at altitudes above 1000 m asl. Significant increases in the incidence and distribution of the disease were recorded later from 2007 to 2011 in Uganda, western Kenya and the Lake Victoria zone of Tanzania. Additional reports have also been published in recent years from Rwanda (2011), Burundi (2011), eastern Democratic Republic of Congo (2012), Mayotte Island (2014) and southern Sudan (2017).

Causal agent and classification Two species of viruses are known to cause CBSD: Cassava brown streak virus (CBSV) and Ugandan cassava brown streak virus (UCBSV) which belong to the genus Ipomovirus, family Potyviridae. The viruses have characteristic pinwheel-like or cylindrical inclusions found in the phloem tissue and a positive-sense ssRNA genome of B9 kb. The genome of CBSV is translated as a large polyprotein and autocatalytically cleaved by virus-encoded protease into 10 mature protein products: the serine protease/silencing suppressor (P1); the third protein (P3); two 6 kDa proteins (6K1 and 6K2); the cylindrical inclusion protein (CI); the viral genome-linked protein (VPg); the main viral protease (NIaPro); the viral RdRp (NIb); a putative pyrophosphatase (Ham1); the coat protein (CP). An additional P3N-PIPO (N-terminal of P3 fused with the Pretty Interesting Potyviridae ORF) protein is produced through ribosomal frameshifting. Genomic analysis has revealed that CBSVs share unusual features. First, CBSVs are the first members within the family Potyviridae that encode a single P1 proteinase and lack the multi-functional helper-component proteinase protein (HCPro). The HCPro activities (silencing suppressor, vector transmission and long-distance movement in planta) appear to have been replaced by silencing suppressor activity of P1. Moreover, CBSV and UCBSV encode novel Ham1 proteins with conserved motifs. The functions of Ham1 proteins are yet to be elucidated, but they are likely to provide essential functions in the lifecycle and pathogenicity of CBSVs and have a role in determining the rate of virus replication and movement as well as the severity of symptoms. Sequence analysis showed that CBSV and UCBSV share 76%–78% nt and 87%–90% aa identity respectively. These virus species are not limited to agro-ecological areas and frequently occur in mixed infections but the potential interaction between them is not currently understood.

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Fig. 7 Spoilage of fresh roots in cassava by CBSD (left) and symptoms on old leaves of cassava (right). Courtesy of J. Legg.

Disease symptoms and yield losses The aerial symptoms of CBSD in cassava include feathery chlorosis along the veins of the leaves or sometimes circular patches of chlorosis between the primary veins (Fig. 7, right), brown necrotic streaks on the stem and stem die-back in severe cases. Symptoms in the tuberous roots consist of a brown, corky necrosis of the starchy tissue, occasional radial constrictions (Fig. 7, left) and a reduction in the content of starch and cyanide. The two causal agents produce distinct symptoms on both cassava and indicator hosts: CBSV causes more severe root necrosis and feathery chlorosis along vein margins, which develops into chlorotic blotches, whereas UCBSV is responsible for circular chlorotic blotches between veins as well as leaf curling and blistering in indicator plants. The viral symptoms also depend on the virus isolate, variety of cassava, age of plant and environmental conditions. Despite the distribution circumscribed to Africa, CBSD is reported by FAO as one of the most impacting diseases since cassava represents the main food intake in the whole continent. From some of the earliest studies of CBSD, it was noted that the disease can cause losses of crop production up to 70% due to reduced growth and spoilage of fresh roots in the most susceptible cassava cultivars. This lets to consider CBSD a re-emergent cause of serious cassava losses in East Africa. The further westward spread in Africa towards Nigeria, which is currently the world’s largest producer of cassava, is a great concern.

Epidemiology Cassava is the only known natural host of CBSVs, and no alternative crop or weed hosts have been reported. CBSV has been detected in the wild cassava relative Manihot glaziovii but the importance of this plant to CBSD epidemiology is not currently known. CBSVs can be mechanically transmitted to several herbaceous plant species that serve as indicator hosts, including Petunia hybrida, Datura stramonium, Nicotiana benthamiana, N. tabacum, N. rustica and N. glutinosa. CBSV and UCBSV are semi-persistently transmitted by the whiteflies Bemisia tabaci, Trialeurodes vaporariorium and Aleurodicus dispersus. Among these species, B. tabaci is considered the primary vector. Whiteflies acquire viruses in 5–10 min, retain them for up to 48 h and transmit them over relatively short distances. Even if the reasons of the sudden increase in CBSD incidence and geographical range in the last decade have not been clearly understood, it has been hypothesized that the disease is dispersed over long distances through the trade transportation of infected planting material or plants carrying viruliferous whiteflies. For instance, the primary causes of the increase of CSBD above 1000 m asl in the African Great Lakes region appears to be the concurrent dramatic increase of B. tabaci populations in that region.

Control There is no cassava cultivar with high levels of CBSD resistance available to farmers. Anyway, the use of cultivars tolerant to CBSD is the more effective approach in reducing the economic impact. Conventional breeding programs produced the M. esculenta M. glaziovii hybrid, known as ‘Namikonga’ in Tanzania or ‘Kaleso’ in Kenya, which reacts to CBSVs infection merely with mild disease symptoms and low incidence of root necrosis. However, the adoption of this hybrid that remains susceptible to CBSVs does not remove viral inoculum from the field. Genetic engineering approaches can offer promising potential for the control of CBSD, bypassing the appearance of undesirable traits in the hybrids and increasing the chances of CBSD and CMD combined resistances in cassava. The management of CBSD needs to pass through adequate phytosanitary measures. The production of “clean” stocks of planting material as well as its maintenance through rouging and selection of healthy stems for replanting are the major components of the CBSD control programs that are currently being implemented at a community level in the affected

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regions of Africa. Moreover, it is very important to establish and enforce strict quarantine measures on the exchange of cassava germplasm between countries, regions and continents (i.e., in Latin America) in order to prevent accidental introduction of CBSD to countries where it is currently absent. Finally, the control of whiteflies is critical to prevent the dissemination of cassava viruses over short distances.

Ornamental Viruses: Rose Rosette Virus Geographical distribution Rose rosette disease (RRD) was originally described in Canada in 1940 and in the United States in 1941, where it was first reported in California and Wyoming. During the last decade, a virus, therefore named rose rosette virus (RRV), was associated with this disease which rapidly spread to North-Central, South-Central and South-East USA with an exponential growth in incidence for the cultivated and garden roses (e.g., in South-Central USA). In 2017, a first detection of RRV was reported in India.

Disease symptoms and yield losses Several different symptoms have been observed in plants infected by RRV. The most recognized symptoms include fast elongation of new vertical and lateral shoots with intense red pigmentation followed by development of witches’ brooms or clustering of small branches (Fig. 8). In the witches’ broom, leaves are observed to be smaller, distorted, and may have a marked red pigmentation, although red pigmentation alone is not considered a consistent symptom. Other symptoms include flowering decrease, reduction and color alteration of petals, as well as petals and sepals switching to leaf-like tissue. Not all the rose varieties show all these symptoms and some of them (e.g., witches’ broom) look very similar to herbicide injury caused by glyphosate on roses. Infected rose plants often die within one to two years.

Causal agent and classification The causal agent of the disease is Rose rosette virus (RRV), belonging to the genus Emaravirus, family Fimoviridae. Like other emaraviruses, RRV has double membrane-bound particles of 120–150 nm in diameter, and has multipartite, negative-sense ssRNA genome. The genome structure of emaraviruses displays significant variability. RRV was reported as a quadra-segmented virus and more recently three novel genomic RNA segments were discovered, one of which is predicted to be bicistronic.

Fig. 8 Rose rosette virus symptoms on a garden rose ‘Firefighter’. Courtesy of F. Ochoa Corona.

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Epidemiology The disease is transmitted by the eriophyid mite Phyllocoptes fructiphilus and by grafting. RRV causes severe symptoms in Rosa multiflora, which can represent a common source of inoculum: rose plants cultivated downwind of infected multiflora are at risk for the vector dispersion by air currents from infected to healthy plants. The causal agent of rose rosette disease is not soil-borne; however, being a systemic pathogen, the virus may persist in RRV-infected root fragments that remain in the soil. The plants that regrow from these old root pieces, as in multiflora rose, can serve as an inoculum source for the vector and the virus can be transmitted to healthy plants.

Control No effective control is available for rose rosette disease to recover infected plants, and there is no evidence of resistant cultivars so far. Early detection of the disease is a key measure for controlling the virus introduction in new areas. Any suspected rose should be removed and destroyed immediately as soon as presence of RRD is ascertained. In some areas, burning is permitted and can be used to destroy the diseased plants. If burning is not allowed in the area, plants should be bagged and removed. In the infected areas, chemical treatments with miticides can reduce the vector population and the risk of virus spread. For new planting beds, strict measures are suggested: use of healthy planting material, avoiding dense plantations, use of no-host barriers to limit wind dispersal of infectious mites, disinfection of pruning tools, thorough removal of symptomatic plants and ensure that infected plants do not have chance to regrow from old, infected root fragments.

Further Reading Aramburu, J., Marti, M., 2003. The occurrence in north-east Spain of a variant of tomato spotted wilt virus (TSWV) that breaks resistance in tomato (Lycopersicum esculentum) containing the Sw-5 gene. Plant Pathology 52, 407. Babu, B., Washburn, B.K., Poduch, K., Knox, G.W., Paret, M.L., 2016. Identification and characterization of two novel genomic RNA segments RNA5 and RNA6 in rose rosette virus infecting roses. Acta Virologica 60, 156–165. doi:10.4149/av_2016_02_156. Bertazzon, N., Forte, V., Filippin, L., Borgo, M., Angelini, E., 2016. Evolution of the new grapevine disease of pinot gris and of grapevine pinot gris virus (GPGV). BIO Web of Conferences 7, 01042. doi:10.1051/bioconf/20160701042. Dombrovsky, A., Tran-Nguyen, L.T.T., Jones, R.A.C., 2017. Cucumber green mottle mosaic virus: Rapidly increasing global distribution, etiology, epidemiology, and management. Annual Review of Phytopathology 55 (1), 231–256. doi:10.1146/annurev-phyto-080516-035349. Laney, A.G., Keller, K.E., Martin, R.R., Tzanetakis, I.E., 2011. A discovery 70 years in the making: Characterization of the rose rosette virus. Journal of General Virology 92, 1727–1732. doi:10.1099/vir.0.031146-0. Luria, N., Smith, E., Reingold, V., et al., 2017. A new Israeli tobamovirus isolate infects tomato plants harboring Tm-22 resistance genes. PLoS One 12 (1), e0170429. doi:10.1371/journal.pone.0170429. Maliogka, V.I., Olmos, A., Pappi, P.G., et al., 2015. A novel grapevine badnavirus is associated with the Roditis leaf discoloration disease. Virus Research 203, 47–55. Martelli, G.P., Saldarelli, P., 2015. Phytosanitary challenges for the Mediterranean viticultural industry: Emerging grapevine viruses. Watch Letter 33, 4. https://www.ciheam.org/ publications/174/016_-_Martelli.pdf. Patil, B.L., Legg, J.P., Kanju, E., Fauquet, C.M., 2015. Cassava brown streak disease: A threat to food security in Africa. Journal of General Virology 96, 956–968. Saldarelli, P., Gualandri, V., Malossini, U., Glasa, M., 2017. Chapter 17 – Grapevine pinot gris virus. In: Meng, B., Martelli, G.P., Golino, D.A., Fuchs, M. (Eds.), Grapevine Viruses: Molecular Biology, Diagnostics and Management. Springer International Publishing. doi:10.1007/978-3-319-57706-7_17. Saleem, A.M., Rana, J., Eman, H., Omar, M., Salam, A., 2013. Squash leaf curl virus (SLCV): A serious disease threatening cucurbits production in Palestine. Virus genes 48. doi:10.1007/s11262-013-1012-1. Tomlinson, K.R., Bailey, A.M., Alicai, T.T., Seal, S., Foster, G.D., 2018. Cassava brown streak disease: Historical timeline, current knowledge and future prospects. Molecular Plant Pathology 19 (5), 1282–1294.

Relevant Websites https://www.cabi.org/isc/datasheet/16951 Cucumber green mottle mosaic virus. https://roserosette.org Rose Rosette. https://microbewiki.kenyon.edu/index.php/Rose_Rosette_Virus Rose Rosette Virus. https://www.cabi.org/isc/datasheet/15038 Squash leaf curl virus (leaf curl of squash). https://gd.eppo.int/taxon/tobrfv Tomato brown rugose fruit virus (TOBRFV).

Emerging Geminiviruses (Geminiviridae) Muhammad S Nawaz-ul-Rehman, University of Agriculture, Faisalabad, Pakistan Nazia Nahid, GC University, Faisalabad, Pakistan and University of Agriculture, Faisalabad, Pakistan Muhammad Mubin, University of Agriculture, Faisalabad, Pakistan r 2021 Elsevier Ltd. All rights reserved. This is an update of C.M. Fauquet, M.S. Nawaz-ul-Rehman, Emerging Geminiviruses, In Encyclopedia of Virology (Third Edition), Brian W.J. Mahy, Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00705-6.

Nomenclature

nt Nucleotide(s) SEA South East Asia ssDNA single-stranded deoxyribonucleic acid ToLCNDD Tomato leaf curl New Delhi disease TYLCD Tomato yellow leaf curl disease

CLCuD Cotton leaf curl disease CMD Cassava mosaic disease CMG Cassava mosaic geminivirus CP Coat protein or capsid protein

Glossary Geminivirus A class of plant viruses having circular single stranded DNA genome encapsidated in icosahedral geminate particle.

Pandemic The outbreak of disease at large scale, across several countries. Satellites A sub-viral agent composed of nucleic acids, which requires a helper virus for its key functions.

Introduction The single stranded DNA (ssDNA) viruses of the family Geminiviridae are serious pathogens of agricultural crops in the tropical and sub-tropical regions of the world. The geminiviruses can be classified into 9 different genera and more than 440 species based on their genome organization, sequence similarity, insect vectors and specific plant hosts. These genera include, Becurtovirus, Begomovirus, Capulavirus, Curtovirus, Eragrovirus, Grablovirus, Mastrevirus, Topocuvirus, and Turncurtovirus. The genus Eragrovirus has only two member species, while the genus Begomovirus comprises hundreds of species. If we consider the number of species of each genus, the begomoviruses contribute the most to the economic impact of the family Geminiviridae. In the Old World, begomoviruses are generally associated with satellites, known as alphasatellites and betasatellites. The betasatellites encode for a pathogenicity determinant protein, while alphasatellites apparently do not play any significant role in disease development. In the past three decades, the number of geminivirus species described has tremendously increased. As begomoviruses have a more devastating impact on world’s agriculture, they are therefore, the main focus of this article. The success of begomoviruses can be attributed to many factors such climatic changes, introduction of susceptible host genotypes and the diversification of insect vector whitefly (Bemisia tabaci). Although, whitefly is the only vector for begomoviruses transmission, but their spread through stem cuttings and grafted material have also played an important role in their global spread for vegetatively propagated plant hosts. Begomoviruses can be monopartite or bipartite. The monopartite begomoviruses contain one circular molecule (DNA-A), coding for six different genes necessary for replication, transcription, and other important regulatory functions. While, bipartite viruses have two circular genomes (DNA-A and DNA-B). Where DNA-A of bipartite viruses is equivalent to genomic component of monopartite viruses, DNA-B encodes two necessary proteins for movement of the virus in the host. Because there is a huge diversity of begomoviruses infecting many crops, we have focused our interest only on four major diseases caused by begomoviruses: the Tomato yellow leaf curl disease, the Tomato leaf curl New Delhi disease, the Cassava mosaic disease, and the cotton leaf curl disease.

The Tomato Yellow Leaf Curl Disease The tomato yellow leaf curl virus (TYLCV) was originally reported from the Jordan Valley in 1920. During the 1960s, the virus was reported from Israel infecting tomatoes. Since then, the tomato yellow leaf curl disease has been reported from all over the world throughout the tropical and sub-tropical regions. The tomato yellow leaf curl disease (TYLCD) is caused by several species of begomoviruses generally termed as Tomato yellow leaf curl virus-like (TYLCV-Like) viruses. There are at least thirteen different species of the TYLCV-like viruses around the globe. However, only the Tomato yellow leaf curl virus-Israel strain (TYLCV-IL) and tomato yellow leaf curl virus-Mild strain (TYLCV-Mld) have the broadest geographical distribution.

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Fig. 1 The global spread of tomato yellow leaf curl virus (TYLCV): Tomato crop is infected by the at least 13 different begomoviruses, which collectively manifest yellow leaf curl disease. The phylogenetic tree represents biodiversity of all 13 begomoviruses. They include, tomato yellow leaf curl virus (TYLCV-Israel, mild, Gezira and Kerman strains), Tomato yellow leaf curl Mali virus (TYLCMLV), tomato yellow leaf curl Malaga virus (TYLCMaV), Tomato yellow leaf curl Sardinia virus (TYLCSaV), Tomato yellow leaf curl Axarquia virus (TYLAxV), Tomato yellow leaf curl Yunnan virus (TYLCYnV), Tomato yellow leaf curl Vietnam virus (TYLCVV), Tomato yellow leaf curl Guangdong virus (TYLCGdV), Tomato yellow leaf curl Shuangbai virus (TYLCShV), Tomato yellow leaf curl Thailand virus (TYLCTHV), Tomato yellow leaf curl China virus (TYLCCNV), Tomato yellow leaf curl Kanchanaburi virus (TYLCKaV) and Tomato yellow leaf curl Indonesia virus (TYLCIDV). The geographical map on left side indicates the spread of TYLCV-Israel and mild strain. Each strain is differentiated by different colors.

Geographical Distribution TYLCD is reported from all the continents, in all tomato growing areas (Fig. 1). Initially, it was presumed that there were different species of begomoviruses infecting tomatoes in different parts of the world. But later, it was proved that TYLCV-IL strain has a worldwide distribution due to the modern trade between different countries in different continents. To date, the TYLCV-IL strain is found in Mediterranean Region, India, Japan, Australia, China, Cuba, Egypt, France, Greece, Korea, Iran, Italy, Mexico, Morocco, Puerto Rico, Sudan, Tunisia, and USA. The TYLCV-IL strain still continues to spread in the Old World, North and South America. Nearly 30% of the total world tomato production is produced in the region surrounding Israel, Jordan, and Iran therefore it is identified as the center for diversity of TYLCV-IL.

The Disease Symptoms, Host Range and Yield Losses TYLCD can cause severe economic losses by reducing the tomato production. The disease symptoms are manifested by a stunted growth, upward curling of leaves and yellowing of plants (Fig. 2). During the early stages of infection, the leaf size is drastically reduced and plants remain stunted with no fruit development. In the epidemic conditions, the disease can cause 100% yield losses. Other than tomatoes, the TYLCV has been reported from several other hosts, such as Solanum nigrum, peppers, tobacco, common bean, soybean, and different weeds.

Causal Agent and Taxonomy TYLCD is caused by mono- bipartite begomoviruses in the family Geminiviridae. There are several widely distributed tomatoinfecting begomoviruses which are both monopartite or bipartite in nature, but TYLCV-IL is a true monopartite begomovirus. In

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Fig. 2 Typical disease symptoms of Tomato yellow leaf curl virus on tomato in Jordan.

fact, all the TYLCV-Like viruses are monopartite begomoviruses, with the exception of tomato yellow leaf curl Thailand virus, which is a mono/bipartite virus. In some exceptional cases like the TYLCV-Oman strain and the Tomato yellow leaf curl China virus they are also associated with betasatellites. TYLCV has a typical genome of a monopartite DNA-A component with six different ORFs. TYLCV has evolved through inter and intra species recombination. There are six known strains of TYLCV in the Middle Eastern region, known as, TYLC-Israel, -Mild, -Oman, -Iran, -Kerman, and -Gezira. However, only the Israel strain spread all over the world and has the broadest geographical distribution and host range.

Epidemiology All the begomoviruses including TYLCV are transmitted through whitefly (Bemisia tabaci). The literature suggests that TYLCV can be transmitted through B or Q whitefly biotypes. The whitefly ingests TYLCV when feeding on phloem tissues of infected tomatoes. The tomato leaves and fruit sepals can act as a reservoir of the virus for the plant. It is believed that TYLCV spread through infected fruit or nursery seedlings from the Middle East to the Americas. After the ingestion of TYLCV by whitefly, it can be transferred to the host plant after 8 h, whereas symptoms can appear after 2–4 weeks. TYLCV is an exception among all begomoviruses, because it can be transmitted through seeds in soybean. The viral replication was also observed in soybean seeds. The TYLCV virions can be observed in the eggs and nymphs of whiteflies. However, not all the whitefly biotypes have the equal capacity of transovarial transmission of TYLCV. The reasons for the fast spread of TYLCV can be attributed to trade related to agriculture among Middle Eastern countries and the rest of the world. Once, the TYLCV is introduced in an area it can spread very aggressively compared to other viruses in this particular region. The invasive spread of TYLCV can be due to high and diverse whitefly population, availability of susceptible plants and higher recombination rate among strains.

Control TYLCD is very difficult to control. However, continuous breeding efforts have resulted in the introgression of resistant genes from wild tomato to cultivated tomatoes in the world. Until now, three different genes have been cloned from wild tomato sources. These include, Ty-1, Ty-2, and Ty-3. Ty-1 and Ty-3 are alleles of each other and encode an RNA dependent RNA polymerase, while Ty-2 encodes an NB-LRR gene. Through breeding programs all three genes have been introgressed into cultivated tomato varieties. The Ty-2 based resistance was overcome by TYLCSV, while, Ty-1 based resistance cannot sustain the high pressure of TYLCV infection. The resistance breakage ability of the virus poses a challenge to the plant breeders to continuously improve the germplasm to control TYLCV. Recently, transgenic plants employing the CRISPR/CAS technology have been developed, targeting Rep and coat protein (CP) genes of TYLCV. The results from transgenic plants are very promising. In the future, the natural and transgene based resistance may be combined to develop more durable resistant varieties.

The Tomato Leaf Curl New Delhi Disease Tomato leaf curl New Delhi virus (ToLCNDV) is another tomato infecting bipartite begomovirus from the Old World. The disease symptoms include leaf curling, stunted growth, slight yellowing of leaves, and leaf puckering. ToLCNDV is known from in the Indian sub-continent from the last 3 decades. Initially, it was limited to the tomato crop in New Delhi, India, but later it spread to other crops, like potato and cucurbit species. Earlier, the ToLCNDV was reported as a typical bipartite virus, but now it is known to be able to capture several different DNA-B components and betasatellites. ToLCNDV infects more than 40 dicotylednous species in the world. Currently, two different strains of ToLCNDV (namely, ToLCNDV-India and ToLCNDV-Spain) are dominant in the world.

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Geographical Distribution ToLCNDV infects different crops in Asia, Middle East, North Africa, and Europe. ToLCNDV was first identified from Northern India during 1990s. Later it was reported from Pakistan, Sri Lanka, Bangladesh, Malaysia, Iran, Taiwan, Thailand, and Indonesia. ToLCNDV is a fast spreading virus around the world. Recently, it was reported from Iran and after 2012 it spread westward to different European territories (Fig. 3). In 2012, it heavily infected cucurbits in the greenhouses and other field crops in Spain. Subsequently, it was reported from Tunisia and Italy. The isolates of Spain, Italy, and Tunisia are closely related to each other indicating their common origin. After 2017, the ToLCNDV has been reported from Algeria and Morocco. Right now (2018–2020) it continues to spread in Greece and Portugal.

The Disease Symptoms and Yield Losses ToLCNDV is a major constraint in tomato production in the Indian subcontinent. Its host range is extended to 43 different hosts. It poses serious threats to vegetable and fibre crops. The tomato plants infected with ToLCNDV exhibit stunted growth, shortening and slight leaf yellowing, puckering and blistering of leaves. Subsequently, the internodes of plants remain shorter with impaired fruit setting, which leads to complete yield loss. In cuccurbits, ToLCNDV induces blistering of leaves along with yellow mosaic pattern. The disease symptoms can be more sever with the adoption of betasatellites.

Epidemiology Similarly to other begomoviruses, the ToLCNDV spread occurs through whitefly B. tabaci. Several whitefly species like, Asia-I, Asia-II and Middle East-Asia Minor-I spread the virus in South Asia. While in Spain, the Q1 biotype causes the spread of the virus in tomato, zucchini and melon. The virus-vector interaction studies demonstrated that ToLCNDV is transmitted in a circulative persistent transmission manner. Evidence also suggest that female whiteflies possess higher transmission efficiency. Interestingly, ToLCNDV is often found in mixed infection with other begomoviruses, such as Chili leaf curl virus or Bhindi yellow vein mosaic virus. During routine surveys, ToLCNDV is found with Chili leaf curl betasatellite, thus enhancing the pathogenicity and host range of the virus.

Component Capturing and Exchange of Helper Components ToLCNDV is among the most unique bipartite viruses, which can co-exist in mixed infection and can exchange its DNA-B component with other begomoviruses of the region. For example, it is very common to find DNA-B of Tomato leaf curl Gujarat virus, Tomato leaf curl Palampur virus, Squash leaf curl China virus and Tomato leaf curl Ranchi virus in combination with the helper component of ToLCNDV DNA-A. It has also been shown that Croton yellow vein mosaic virus, Radish leaf curl virus, Tomato leaf curl Palampur virus, and Tomoto leaf Gujarat virus can trans-replicate the DNA-B of ToLCNDV. These evidences suggest that ToLCNDV can result in serious agricultural losses due to its flexibility in component capturing.

Control Fortunately, both breeding and genetic engineering approaches have led to the development of resistance against ToLCNDV in tomato and other important cucurbits. The breeders have developed the resistant lines through crossing of wild relatives of tomato with cultivated species. Just like TYLCV, the Ty loci have been exploited to develop resistance against ToLCNDV. The transgenic approaches included the use of AV1, AV2, and AC1 genes to target the ToLCNDV. However, recombination and component capturing can still result in serious outbreaks.

The African Cassava Mosaic Disease The cassava mosaic geminiviruses (CMGs) in Africa are among the emerging geminiviruses and caused a pandemic in sub-Saharan African countries. The CMGs are known to infect the cassava crop in Africa for more than 100 years, but some of them emerged as major pathogens in the last 30 years. The cassava mosaic disease (CMD) spread in pandemic manner in the early 90 s in Uganda, East Africa, and then spread to other parts of Africa, mostly East and central Africa. More recently it was demonstrated that CMD was imported from the Indian subcontinent to South East Asia (SEA), and a rapid pandemic began about 3 years ago, invading 4 countries already and compromising the second most important production of cassava in the world. The spread of cassava pandemic throughout Africa is mainly due to the man-mediated transfer of infected material and the whitefly vector, while in SEA it is the international regional trade of ingected cuttings that is the driving force for the rapid spread of the disease. This African pandemic was responsible for very severe cassava crop losses and resulted in one of the worst modern famines on this continent. In the last twenty years, both the Cassava brown streak viruses (CBSV and UCBSV; RNA viruses of the genus Ipomovirus, family Potyviridae) and CMGs had resulted in very low cassava production in East and Southern Africa. The CMGs and CBSVs are spreading from East Africa to central Africa. Interestingly, both the viruses are transmitted through whiteflies that have been super adundant on cassava in the last 3 decades.

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Fig. 3 Geographical distribution and phylogeny of ToLCNDV in the world. The distribution of Tomato leaf curl New Delhi virus (ToLCNDV) overtime. Countries are colored, according to the color index (top right), with respect to the first report of ToLCNDV in the countries. The phylogenetic tree at the bottom is based on ToLCDNV DNA-A sequences, and a spotlight on each cluster indicates the geographical region from which the isolates in that cluster originate. The photograph was previously published in Molecular plant pathology by Zaidi, S.S., Martin, D.P., Amin, I., Farooq, M., Mansoor, S., 2017. Tomato leaf curl New Delhi virus: A widepread bipartite begomovirus in the territory of monopartitebegomoviruses. Molecular Plant Pathology 18 (7), 901–911. doi:10.1111/mpp.12481.

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Fig. 4 Geographical distribution of cassava mosaic begomoviruses and cassava brown streak virus in Africa. The territory of each virus is marked with a unique colored box. The countries plagued by CBSV are shaded with yellow color. Cassava crop is infected by African cassava mosaic virus (ACMV), African cassava mosaic Burkina Faso virus (ACMBFV), East African cassava mosaic virus (EACMV), East African cassava mosaic Cameroon virus (EACMCV), East African cassava mosaic Kenya virus (EACMKV), East African cassava mosaic Malawi virus (EACMMV), East African cassava mosaic Zanzibar virus (EACMZV), South African cassava mosaic virus (SACMV), Cassava mosaic Madagascar virus (CMMGV), and Cassava brown streak virus (CBSV).

Geographical Distribution CMD was observed in East Africa (Tanzania) since, 1894, several decades before the emergence of Cassava brown streak disease (CBSD- caused by CBSV and UCBSV). In 1994, the CMD spread from Uganda to neighboring countries surrounding the lake Victoria. The disease swept through Central Africa to West African countries (Congo, Gabon and south of Cameroon) (Fig. 4). The earlier sequencing data revealed the presence of the African cassava mosaic virus (ACMV) in the infected cassava plants. But the 1990s epidemic revealed the presence of a more severe strain of a different species, namely East African cassava mosaic virus Ugandan strain (EACMV-UG). The disease is currently present in more than 20 countries (Fig. 4). The East and West Africa are mainly dominated by ACMV and EACMV-UG strains respectively. While Southern and South East Africa are mainly dominated by several other CMGs and also by the two CBSD viruses. Although, the presence of ACMV and EACMV dominates in the whole African continent, the Island of Madagascar contains the largest number of CMGs (Fig. 4). In 2007, the non-cultivated cotton samples collected from Pakistan showed the presence of DNA-A component of ACMV. The DNA-B component of ACMV is still not discovered from Pakistan. The discovery of ACMV from non-cultivated cotton is surprising, not only because of the geographical distance, but also due to the host change.

The Disease Symptoms and Yield Losses The CMGs can cause very severe damages to the cassava crop with annual losses of over US$1 billion losses. During the epidemic period, the CMD resulted in a reduction of about 90% of the cassava production in Uganda which induced a severe famine and

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Fig. 5 The disease symptoms of cassava plants infected with CMD. The health plant leaves do not show any mosaic pattern (a), The panel b represents the symptoms by EACMV and panel c represents the infection by ACMV. The synergism between both viruses results in candlestick syndrome in panel d.

thousands of people died due to hunger. The disease symptoms due to various strains can be easily identified. The symptoms due to EACMV-UG are more severe than ACMV (Fig. 5). The EACMV-UG infection results in reduction of leaf surface area, yellowing of lamina and downward curling of the leaves. The entire plant looks yellow and stunted in growth compared to healthy plants. The cassava plants infected with CBSD also show yellow symptoms on the old leaves but they never show the severity of EACMV-UG. Furthermore, the plants infected with CBSD have dry-brown necrosis on cassava tubers and are inedible. In the 90 s in Uganda, the cassava plants showed dual infection of ACMV and EACMV-UG with an extreme reduction of the growth of the plants and no root production. In the case of dual infected plants, the plant fails to produce any proper foliage; the stem turns into yellow-purple color and the infected plants showed a “candle stick syndrome”, and no root production whatsoever. In addition, cassava is propagated through stem cuttings and dually infected plants cannot be used for further propagation. The farmers in this part of the world, at that period in time, ran out of food and out of planting material!

Classification of Cassava Mosaic Begomoviruses The CMD is caused by bipartite begomoviruses. Unlike TYLCV, the CMDs have two circular genomes (bipartite) i.e., DNA-A and DNA-B. As recombination is the driving force for geminivirus evolution, therefore CMDs are no exceptions. Indeed, the EACMVUG strain is a recombinant for the CP region between EACMV and ACMV. The sequence analysis revealed that EACMV-UG strain gained B400–550 nt from the CP of ACMV through recombination. Over the years, the molecular sequencing revealed nine different species of begomoviruses occurring in single or mixed infection. These species includes, African cassava mosaic virus (ACMV), African cassava mosaic Burkina Faso virus (ACMBFV), East African cassava mosaic virus (EACMV), East African cassava mosaic Cameroon virus (EACMCV), East African cassava mosaic Kenya virus (EACMKV), South African cassava mosaic virus (SACMV), East African cassava mosaic Malawi virus (EACMMV), East African cassava mosaic Zanzibar virus (EACMZV), and cassava mosaic Madagascar virus (CMMGV). The co-infection of different CMDs can lead to synergism and new recombinants are produces with variable virulence can arise.

Epidemiology The literature suggests that CMGs mainly spread through infected cassava cuttings, however, whitefly (Bemisia tabaci) also played an important role in spreading the virus to other field crops. The transmission of CMGs occurs through whitefly in a circulative but non-propagative manner. It requires at least 8 h of circulation inside the whitefly before transmission into plants can occur. However, once the whitefly acquires the virus, it can remain viruliferous all its life, thus increasing the chance of viral spread for a long period of time and a longer distance. The East African cassava pandemic could be correlated to the synergism between ACMV and EACMV (Fig. 5) along with the explosion of “super abundant” whitefly populations. Recombination between different CMGs is extremely prevalent although it is not known how important this factor is in the disease spread. Recent research demonstrated that there are several whitefly biotypes infesting cassava in Africa and that these biotypes are geographically distributed.

Recent Outbreak of CMD in South East Asia In 2016, the first report of the CMD, was published, describing the unequivocal identification of the Sri Lankan cassava mosaic virus (SLCMV) as responsible for the disease symptoms in the north east of Cambodia. Since that date, several surveys have been done in Cambodia, China, Vietnam, and Thailand, and as of January 2019, the disease was present in 10 provinces of Cambodia, 14 in Vietnam, and 7 provinces in Thailand (Fig. 6). There is no CMD case yet reported in Laos or Myanmar.

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Fig. 6 Geographical distribution of cotton leaf curl geminiviruses. Cotton leaf curl disease is present in India, China, Pakistan and Philippines. The disease was first known from Pakistan and then spread to India, China and Philippines. The Chinese and Philippine’s isolates are identical to each other. The dashed arrows represent direction of movement for cotton leaf curl Multan virus.

The presence of the same virus has also been reported from several locations of cassava collections in China. In 3 years, the disease reports extended from a single plantation in Cambodia to an estimated 44% of the surfaces cultivated in the whole region. The disease is transmitted in two ways: by the natural whitefly vector (B. tabaci, Aleyrodidae) and by the cassava cuttings, there is no transmission through the roots or the seeds. There is a very active cassava stake trade between several provinces of Cambodia and between Cambodia and Vietnam or Thailand. Although there is clearly insect transmission occurring everywhere, it is believed that currently the disease is mostly spread through cuttings. Considering the importance of cassava in the region (455 million tons/year and 4$10 billion business) urgent action is needed to stop the spread and put CMD under control. To this effect the Global Cassava Partnership for the 21st Century - GCP21 and CIAT, organized a Regional Workshop in September 18–20, 2018, in Phnom Penh, Cambodia establishing a unique regional plan of control of CMD. This control plan lists 5 different topics; 1- Regional & National Coordination Mechanism, 2- CMD Awareness & Mitigation Extension, 3- Surveillance and Diagnostic, 4- Propagation and distribution of disease-free planting material, and 5- Varietal testing and breeding for CMD resistance. This recent emergence of CMD exemplifies the major role of international movement of infected material complemented by biological factors.

Control In general terms, CMD can be controlled by three different ways. (1) Planting disease-free material to minimize the spread of infected material and crop losses, (2) Control of whitefly populations to delay the infection process till later growth stages, (3) Eradication of infected material to avoid the spread of infected plants through cuttings. Although CMD is known for more than a century, still the disease causes very significant losses in Africa and India. The earlier cassava breeding programs in East Africa led to the development of highly resistant germplasm against CMGs (multigenic type of resistance CMD1). Subsequently breeding programs in IITA in West Africa produced numerous CMD resistant cassava varieties between 1970 and 1990. In the 90 s another source of resistance (CMD2) of monogenic nature was identified in cassava landraces in West Africa, conferring extremely high levels of durable resistance to CMGs. SNP markers have been developed for CMD2 allowing its easy transfer to many more

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genotypes of cassava. The introduction of resistant cassava germplasm in Uganda resulted in restoration of cassava yield in the late 90 s. However, due to their better processing and organoleptic properties, the farmers often prefer local susceptible landraces.

The Cotton Leaf Curl Disease in the Indian Sub-Continent Geographical Distribution The Cotton leaf curl disease (CLCuD) was first observed in the Multan region of Pakistan during late 1960s. The disease was limited to the surroundings of Multan until 1987. After 1987, the disease spread to the surrounding areas and appeared in epidemic form between 1992 and 1997. During early 2000, the disease spread to India, where it caused serious damage to cotton crops. Today, the disease is prevalent in Southeastern China, Philippines, Pakistan, and India. In Philippines, CLCuMuV infects hibiscus plants instead of cotton. Although there is a huge diversity among cotton leaf curl geminiviruses (CGs), the most devastating viruses are Cotton leaf curl Multan virus (CLCuMuV) and Cotton leaf curl Gezira virus (CLCuGeV-African origin). The CLCuMuV is prevalent in Pakistan, India, China, and Philippines, while CLCuGeV is present all over the Africa, wherever cotton and okra crops are grown. Indeed, CLCuGeV is now also present in Southern parts of Pakistan; however, its infection is limited on cotton crop which is dominated by Cotton leaf curl Multan virus.

The Disease Symptoms and Yield Losses Like all other begomoviruses, the CLCuD is transmitted by whiteflies. So far there is no evidence for the seed transmission of the CLCuD. Upward or downward leaf curling, vein thickening, leaf enations and stunted growth are the main symptoms of infected plants (Fig. 7). Once the plant acquires CLCuD through whitefly inoculation, the infection is life long for the host. The infected plants remain stunted and dark green in color. Very few flowers are produced and they cannot produce fertile seeds. Over the past three decades, Pakistan had lost billions of dollars due to this disease. During the epidemic (1992–1997) Pakistan lost US$5 billions. With the introduction of resistant varieties after the year 2000 the disease was fairly well controlled. However, a new out-break of the Burewala strain during 2004 resulted in the resistance breakdown. It was hypothesized that the Burewala strain is the most dominant virus on cotton field in Pakistan and India, however recent data showed that the Multan strain is re-emerging as a major pathogen on cotton.

Causal Agent and Disease Classification The CLCuD is mainly caused by satellite associated monopartite begomoviruses. These satellites are known as betasatellites and alphasatellites. Both the satellites are approximately half the size of viral component and are encapsidated by the viral CP coded by the helper virus. The prevalent betasatellite is known as Cotton leaf curl Multan betasatellite (CLCuMuB). The DNA-A component of Cotton leaf curl geminiviruses alone cannot produce symptoms, until betasatellites are added. The CLCuMuB encodes a pathogenicity determinant protein. The second category of satellites associated with cotton geminiviruses are alphasatellites. They encode an alpha-Rep responsible for replication of alphasatellites and which is also a suppressor of gene silencing of the host, the natural virus defense mechanism of host. The origin of alphasatellites is considered to be from the family Nanoviridae. Apparently, the alphasatellites do not play an important role in the etiology of CLCuD. However, their relatively large diversity suggests that they have an important evolutionary role in disease epidemics. CLCuD is mainly caused by monopartite begomoviruses, but recently some bipartite viruses have also been reported from Pakistan. According to literature, there are at least 14 different begomovirus and one mastrevirus species, which infect the cotton crop in the Old World. CLCuMuV, CLCuGeV, Cotton leaf curl Kokhran virus (CLCuKoV), Papaya leaf curl virus (PaLCuV), and Cotton leaf curl Alabad virus (CLCuAlV) are frequently associated with CLCuD. However, African cassava mosaic virus (ACMV), Okra leaf curl enation virus (OLCEnV), Tomato leaf curl New Delhi virus

Fig. 7 Typical disease symptoms of cotton leaf curl disease in Pakistan. The healthy cotton plants have straight leaves with no curling (A). The infected plants have upward or downward leaf curling. Panel-B presents cotton infected by cotton leaf curl Multan virus, cotton leaf curl Multan betasatellite and Gossypium darwinii symptomless alphasatellite. The abaxial side of the infected leaf shows vein thickening and leaf like enations (panel-C).

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Fig. 8 Phylogenetic tree of cotton leaf curl geminiviruses. Cotton plant has been shown infected by 14 different begomoviruses species and one mastrevirus species. These species include, Cotton leaf curl Gezira virus (CLCuGeV), African cassava mosaic virus (ACMV), Tomato leaf curl New Delhi virus (ToLCNDV), Tomato leaf curl Bangalore virus (ToLCBaV), Papaya leaf curl virus (PaLCV), Okra leaf curl enation virus (OLCEnV), Cotton leaf curl Bangalore virus (CLCuBaV), Cotton leaf curl Barasat virus (CLCuBarV), Cotton leaf curl Kokhran virus (CLCuKoV), Cotton leaf curl Multan virus (CLCuMuV), Cotton leaf curl Alabad virus (CLCuAaV), Tomato leaf curl Gujarat virus (ToLCuGuV) Cotton yellow mosaic virus, Squash leaf curl virus (SqLCV) and a mastrevirus, Chickpea chloratic dwarf virus (CpCDV). Shadadpur, Burewala and Rajasthan strains are recombinants of CLCuMuV and CLCuKoV.

(ToLCNDV), Tomato leaf curl Gujarat virus (ToLCuGuV), Squash leaf curl virus (SqLCV), Cotton yellow mosaic virus (CYMV), Cotton leaf curl Bangalore virus (CLCuBaV), Cotton leaf curl Barasat virus (CLCuBV), Tomato leaf curl Bangalore virus (ToLCBaV), and Chickpea chlorotic dwarf virus (CpCDV- a mastrevirus) are rarely found on cotton crop (Fig. 8).

Epidemiology Upland cotton (Gossypium hirsutum) was introduced in Pakistan by the East India Company from Southern Mexico. The CLCuD is not prevalent in Southern Mexico. The disease is also not seed transmissible, therefore it can be presumed that viruses causing leaf curl disease in cotton in Pakistan shifted from a local plant host to cotton. The major spread of the disease occurred when the exotic germplasm was cultivated during the late 1980s in Pakistan. Indeed, the variety S12 (imported from Texas) proved to be hyper-susceptible to the CLCuD and resulted in the explosion of the disease in a very short period of time. Even after removing the variety S12 from the field, the CLCuD epidemic continued actively. The S12 varietry acted as a trigger of the epidemic but it is whiteflies that spread the disease in the country. Due to year around presence of alternative hosts for cotton leaf curl geminiviruses; the threat of CLCuD remains permanent in Pakistan and India. As a matter of fact the begomoviruses from other crops like tomato, cassava, and chickpea are also invading cotton crop. After the year 2000 a recombinant Burewala strain dominated the cotton crop in Pakistan and India. But recently CLCuMuV has re-emerged on cotton in Pakistan.

Control CLCuD can only be controlled through the use of resistant varieties and the control of whiteflies. Recently, virus resistant cotton varieties obtained through conventional breeding were introduced. It was an immediate success but more recently the resistance was broken down by CLCuMV strains resulting from recombination with other local geminiviruses. The recombinant strains, Burewala, Shadadpur and Rajasthan played an important role in resistance breakdown. Efforts have been made to control the

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CLCuD through RNAi or CRISPR/CAS9 but to date there is still no data to support the use of transgene mediated resistance in cotton. The conventional breeding still remains the first and only option in controlling the CLCuD.

Conclusion This article is documenting the emergence of 4 major geminivirus diseases in the world in the last 30 years, namely the TYLCD and ToLCNDV on tomato, the CMD on cassava and the CLCuD on Cotton. All three examples illustrate the importance of human intervention to trigger a new epidemic, but evidently are also showing the great importance of some biological factors; the greater effectiveness of TYLCV to infect tomatoes, the capacity of ToLCNDV to capture other components and satellites, the combination of two cassava geminiviruses to synergize, and the role of a hyper susceptible cotton variety. In addition, it is also showing the major role of whiteflies in each example to carry on the epidemic to vast surfaces and for many years. The recent global spread of ToLCNDV on various hosts is also an alarming situation. The spread of ToLCNDV and TYLCV is a global concern and concrete efforts must be done to save crops from their worldwide epidemics. Another important concern for the world’s agriculture is the Squash leaf curl virus (SLCV) spread from California to Middle Eastern countries. If the other 4 examples in this article are all Old World geminiviruses that managed to invade the rest of the world, SLCV is a New World geminivirus that successfully invaded the Middle East region. Indeed, evidences of SLCV infection on cotton also exist in Pakistan. However, it still needs authenticated experiments to prove its level of infection on cotton. The emergence of such viruses is also a future concern. Currently it is unclear why some strains spread more rapidly than others at the global level. For example there are several strains of TYLCV, but TYLCV-Mld is more wide spread than some others. Similarly, the Uganda strain of the East African cassava mosaic virus (EACMV-UG) in Africa and the cotton leaf curl Multan virus (CLCuMuV) and its betasatellite in Asia are more wide spread than other strains. The world spread of TYLCV can be attributed to trade of tomato seedlings from the Middle East to different parts of the world. Once, the virus is introduced in a new environment, the local whiteflies can quickly acquire and spread the virus to other hosts. The CMD spread in Uganda which was triggered by the synergism between two geminiviruses and the presence of a hyper abundant whitefly, is also attributed to human migration in Africa (Rwanda in the 90 s), carrying infected cassava cuttings for further propagation over a high mountain range. The recent CMD outbreak in SEA was initiated by the import of SLCMV from the Indian subcontinent, and hugely distributed by the clonal propagation in the region. If the initial pandemic of CLCuD in Pakistan was clearly triggered by the import of a super susceptible cotton variety from the US, local whiteflies then took the relay to spread the disease in the sun-continent and to many local hosts. However, the spread of CLCuD from Pakistan to China and Philippines is more intriguing. It is plausible that Pakistan and India sharing borders, therefore whiteflies could easily transfer the disease to the neighboring countries. However, there is a mountainous region between Pakistan and China and crossing of whiteflies is not possible. The transfer of infected ornamental plants could be a reason for the spread of CLCuMuV from Pakistan to China and Philippines. The Multan strain in Philippines is identical to the one isolated in China, therefore it can be hypothesized that CLCuMuV spread from China to Philippines. In all these examples it is clear that humans, in combination with other biological factors, have played a vital role in the spread of highly pathogenic geminiviruses. In the future, such epidemics could be avoided by more strict quarantines measures and immediate proper agricultural policy control measures.

Further Reading Chen, W., Wosula, E.N., Hasegawa, D.K., et al., 2019. Genome of the African cassava whitefly Bemisia tabaci and distribution and genetic diversity of cassava-colonizing whiteflies in Africa. Insect Biochemistry and Molecular Biology 110, 112–120. Jacobson, A.L., Duffy, S., Sseruwagi, P., 2018. Whitefly-transmitted viruses threatening cassava production in Africa. Current Opinion in Virology 33, 167–176. Lefeuvre, P., Martin, D.P., Harkins, G., et al., 2010. The spread of tomato yellow leaf curl virus from the Middle East to the world. PLoS Pathogens 6, e1001164. Nawaz-ul-Rehman, M.S., Briddon, R.W., Fauquet, C.M., 2012. A melting pot of Old World begomoviruses and their satellites infecting a collection of Gossypium species in Pakistan. PLoS One 7, e40050. Saleem, H., Nahid, N., Shakir, S., et al., 2016. Diversity, mutation and recombination analysis of cotton leaf curl geminiviruses. PLoS One 11, e0151161. Varsani, A., Roumagnac, P., Fuchs, M., et al., 2017. Capulavirus and Grablovirus: Two new genera in the family Geminiviridae. Archives of Virology 162, 1819–1831. Zaidi, S.S., Martin, D.P., Amin, I., Farooq, M., Mansoor, S., 2017. Tomato leaf curl New Delhi virus: A widepread bipartite begomovirus in the territory of monopartite begomoviruses. Molecular Plant Pathology 18 (7), 901–911. doi:10.1111/mpp.12481.

Movement of Viruses in Plants Manfred Heinlein, IBMP-CNRS, University of Strasbourg, Strasbourg, France r 2021 Elsevier Ltd. All rights reserved. This is an update of P.A. Harries, R.S. Nelson, Movement of Viruses in Plants, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00728-7.

Glossary 50 CAP The five-prime cap is a modified nucleotide at the 50 end of RNA molecules. It prevents RNA degradation by exonucleases and promotes RNA translation. Actin filaments (microfilaments) A component of the cytoskeleton formed from polymerized G-actin monomers. Chaperones Cellular proteins that assist in the correct folding or unfolding of proteins and in the assembly or disassembly of protein complexes. COPI/COPII Cytoplasmic protein complexes (coatomers) that coat vesicles transporting proteins form ER to Golgi (COPII), and from Golgi to the ER (COPI). Endosomes Membrane-bound compartments that function in the sorting and delivery of material internalized from the cell surface or destined for transport from the Golgi to the lysosome or vacuole. ER exit sites Specialized zones of the ER engaged in COPII-mediated exit of newly synthesized proteins from the ER into vesicles for transport from ER to Golgi.

ESCRT The “endosomal sorting complexes required for transport (ESCRT)” machinery consists of cytoplasmic protein complexes involved in membrane remodeling. GPI anchor Short glycolipid (glycosylphosphatidylinositol or glycophosphatidylinositol) fused to the C-terminus of a protein during posttranslational modification and tethering the protein to the outer leaflet of the PM. Microtubules A component of the cytoskeleton composed of hollow tubes assembled from a-tubulin/b-tubulin dimers. Multivesicular bodies (MVB) Single membrane-bound organelles with intraluminal vesicles that mediate protein cargo transport form the trans-Golgi network (TGN) to vacuoles for degradation. Pattern-triggered immunity (PTI) Innate immunity based on the recognition of conserved microbial signatures (patterns) by pathogen recognition receptors at the PM or in the cytoplasm. Reticulons Proteins residing at the ER and playing a role in promoting membrane curvature.

Introduction In order for a plant virus to infect its host systemically, it must be capable of hijacking the host's cellular machinery to replicate and move from the initially infected cell. Plant viruses require wounding, usually by insect or fungal vectors or mechanical abrasion, for an infection to begin. Once inside a cell, virus particles have to disassemble to release the viral genome which, then, can be transcribed (DNA viruses) and translated (DNA and RNA viruses) to initiate its replication and movement. Some of the viral products are required for virus movement and interact with host factors (proteins or membranes) to carry out this function. Virus movement in plants is tightly linked to virus replication and can be broken down into three distinct steps: (1) intracellular movement, (2) intercellular movement and (3) systemic movement. Intracellular movement refers to processes that allow the virus to reach plasmodesmata (PD), tiny (50 mm in diameter) channels in the plant cell walls that connect neighboring cells, whereas intercellular movement refers to the process of virus movement through PD allowing the spread of infection between cells. By intercellular movement from-cell-to-cell and by reiteration of infection and viral genome replication and the targeting of PD in each cell, the virus finally reaches cells of the vascular system (Fig. 1). Once the virus has entered the sugar-transporting phloem sieve elements, the virus moves systemically with the flow of photoassimilates, which directs the infection efficiently to the young, sugar-importing sink tissues before also source tissues are slowly infected. Upon delivery by the phloem to a tissue distant from the original infection site, the virus exits the vasculature and resumes cell-to-cell movement via PD in the new tissue. Although virus movement generally occurs by viral exploitation of the symplasmic intercellular communication network established by PD and the connected phloem (Fig. 1), it is important to realize that some viruses, like Turnip mosaic virus (TuMV) and Rice yellow mottle virus (RYMV), also exploit the water-transporting xylem vessels, but this will not be further elaborated here. The mechanism of virus movement in plants has been studied with a wide range of virus genera, including, but not limited to, tobamoviruses, potexviruses, hordeiviruses, comoviruses, nepoviruses, potyviruses, tombusviruses, tospoviruses, and geminiviruses. Although viruses generally move between cells with support of their Movement Proteins (MPs) (Fig. 2), the cellular pathways and host factors (proteins and membranes) used by viruses for movement show strong differences (Fig. 3). Several studies using GFP and other fluorescent proteins as visible markers have provided important insights into the dynamic changes in the subcellular distribution, accumulation, and movement patterns of viral and host proteins and helped to decipher the different processes occurring during virus infection in plant cells. The resulting observations are summarized in numerous review articles and highlight the roles of membranes and the associated cytoskeleton during virus replication and movement. This article will describe the three steps of virus movement in plants and will focus on model viruses within genera that provide the most

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Fig. 1 Local and systemic virus movement in infected plants. In the inoculated leaf, viruses move cell-to cell (i) and expand the local sites of infection (a) until they reach the phloem sieve elements (SE) in the leaf vein vasculature by which they move with the flow of photoassimilates over long distance to sink tissues (ii). Upon unloading from the SE, viruses move again cell-to-cell (iii) to spread infection throughout the newly infected systemic leaves (b). Successful infection depends on compatible interactions between the virus and its host (arrows) to allow the virus to replicate its genome and to target PD (curved arrow, intracellular movement) in each cell it enters. (a) Local sites of infection by TMV tagged with green fluorescent protein (TMV-GFP). (b) a systemic leaf showing TMV-GFP as it spreads infection from the veins into the surrounding cells. BS, bundle sheath; PP/CC, phloem parenchyma/companion cell.

information on the subject. This article also considers the virus-triggered defense responses of the host that the virus must overcome to ensure its efficient cell-to-cell spread.

Intracellular Movement Although the process of virus movement can be complex and requires support by the coordinated activity of several virus-and host-encoded proteins, many viruses achieve their movement with the help of virus-encoded ‘movement proteins (MP)’. Pioneering studies with the MP of TMV and later with other MPs demonstrated that these proteins bind single-stranded RNA and modify the size-exclusion limit (SEL) of PD (Fig. 2). Moreover, upon introduction into cells by microinjection, or by transient expression using microparticle bombardment or agroinfiltration, MPs interact with PD, move between cells and may co-transport co-injected nucleic acids, thus indicating that their function is independent of infection. The ability of MP to alter PD aperture is usually correlated with the accumulation of the protein at PD. Nevertheless, plants that express the MP and are able to complement the movement function of movement-deficient TMV mutants do not show obvious growth defects, suggesting the existence of mechanisms that tightly control the ability of MP to modify PD aperture and thus prevent the continuous trafficking of signaling macromolecules between cells. Microinjection studies indeed demonstrated that the MP of TMV modifies the SEL of PD only transiently in cells at the spreading front of infection, thus confirming that MP is tightly regulated. The MP of TMV has several amino acids that are phosphorylated in vivo and that may play a role in regulating MP functions. Consistent with this type of posttranslational control, PD are associated with several kinases and a specific kinase, plasmodesmal-associated kinase (PAPK), was shown to specifically phosphorylate the C-terminal phosphorylation sites of MP in vitro and also to be active on a subset of other MPs and non-cell-autonomous plant transcription factors. The ability of MPs to bind nucleic acids is sequence-independent, which raises the important question of how these proteins find their viral RNA cargo in infected cells. Since the MPs are produced by translation of the viral genome, they may simply attach co-translationally to viral genomes in their vicinity. Consistently, the MPs of several RNA viruses, including Tobacco mosaic virus (TMV), Potato virus X (PVX), Brome mosaic virus (BMV) or Red clover necrotic mosaic virus (RCNMV) were shown to colocalize with membrane-associated inclusions that harbor their viral replication complexes (VRC). VRCs can be highly organized structures involving membranes and cytoskeletal elements, and in which the binding of MP to viral RNA involves specific mechanisms, for

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example interaction with other viral proteins, as exemplified by VRCs formed by RCNMV. Another important question is whether MPs transport viral genomes as part of subcomplexes that are derived from anchored VRCs or whether infection moves between cells in the form of mobile VRCs. Studies with TMV and other viruses have shown that VRCs develop at multiple sites in the infected cell. Moreover, VRCs can be stationary or mobile, and their subcellular formation and movement is associated with the spread of the virus. Studies with PVX have shown that VRCs can associate with PD and load replicated viral genome into PD for movement, while observations with TMV suggested that TMV spread infection by movement through PD as a wholly intact VRC. Thus, it appears that viruses develop VRCs for replication as well as for movement and that virus replication and virus movement are tightly linked processes. Movement to PD or even between cells as a VRC ensures the vicinity of MPs to the viral genome and may explain the apparent lack of evolution of any sequence specificity in the binding of MPs to nucleic acids. Thus, how do viruses develop their VRCs and transport them to PD? TMV infection is initiated by attaching the viral RNA that entered the cell with its 50 CAP to the ER membrane. The RNA is then replicated before the virus can spread its RNA genome into adjacent cells. Time-resolved in vivo studies showed that TMV replication and, thus, the formation of VRCs occurs at distinct sites of the cortical ER. While some of the VRCs remain anchored at these sites and are destined for the development of ER-associated, virion-producing viral factories, other VRCs detach from their anchorage sites soon after formation and act as mobile RNA particles for intra-and intercellular transport (Fig. 3(a)). The formation of VRCs at distinct ER sites probably reflects the presence of localized mechanisms by which the virus recruits host factors and membranes for VRC formation. In vivo studies have shown that the membrane-associated mechanisms of VRC formation and movement are tightly associated with the cytoskeleton, thus reflecting the viral exploitation of the microtubule- and actin-dependent functions in plant endomembrane organization. The distinct sites of VRC formation by TMV overlap with sites of immobile ER at which the cortical ER-actin network intersects with microtubules. The literature suggests that such sites are attached to the plasma membrane (PM), thus potentially constituting ER-PM contact sites. In providing vertical vicinity between the PM and the ER and lateral vicinity between the ER and the associated cytoskeleton these sites may function as a cortical nexus at which actin- and microtubule-driven transport pathways for the trafficking of membranes, protein complexes and organelles converge. TMV, and potentially other viruses, may have evolved to target these junctions as sites at which membranes and host factors required for VRC formation can be recruited, concentrated, and organized. The MP of TMV has strong affinity for binding to the surfaces of both membranes and microtubules, and also interacts with microtubule-associated proteins such as tubulin dimers, g-tubulin, and end-binding protein 1 (EB1). While microtubules may support VRC attachment and the structural stabilization of the growing VRC, the ER-associated actin network supports the recruitment of membranes and host factors to these sites with support of myosin motor proteins. Many viruses associate with the ER for replication like TMV, but several other viruses use different locations to establish their VRCs. For example, Carnation Italian ringspot virus (CIRV) and Melon necrotic spot virus (MNSV) form factories from the outer membrane of mitochondria, whereas the tonoplast is implied to play a role in the replication of Alfalfa mosaic virus (AlMV) and Tomato mosaic virus (ToMV). Tomato bushy stunt virus (TBSV) induces the formation of multivesicular bodies (MVB) from peroxisomes and several other viruses show associations with chloroplasts. However, given that Golgi membranes, chloroplasts, peroxisomes, and also the tonoplast are tightly associated with the ER-actin network, ER membranes and associated vesicle trafficking as well as the above-described roles of the cytoskeleton in supporting membrane trafficking and anchorage may represent basic principles of membrane and host factor recruitment playing a role also in the association of viruses with these organelles. Tight linkage between virus replication and movement is particularly important if more than a single viral RNA must be moved between cells to spread infection. Unlike TMV, RCNMV has a bipartite genome but also forms ER-associated VRCs and encodes two N-terminally overlapping replicase subunits similar to TMV. To ensure replication of RNA2 together with RNA1, the smaller replicase subunit (p27) of the replicase complex, which is associated with RNA 1, binds to RNA2 through Fig. 2 Regulation of PD and modification by viruses. (a) PD structure and regulation. (i) PD are channels (ca. 50 mm in diameter) in the plant cell walls that provide cytoplasmic and membrane continuity between adjacent cells. Structurally they are concentric cylinders with the PM lining the cell wall and the ER forming a narrow rod called ‘desmotubule’ in the center. The desmotubule is linked to the PM by spoke-like extensions from all its sides. The cell wall regions surrounding the PD orifices at the PD neck regions contain deposits of callose (b-1,3 glucan). (ii) and (iii) Transport through PD occurs through the cytoplasmic space delimited by the PM and the desmotubule. This size of this space determines the size exclusion limit (SEL), thus the maximal size of the molecules that are allowed to pass from one cell to another. The SEL is regulated by two potential mechanisms. The first mechanism (ii) relies on the regulation of the callose deposits. Callose synthesis by callose synthases increases the size of the callose deposits in the cell wall, thereby narrowing the space between the PM and the desmotubule and, thus, reducing the SEL. Degradation of callose by b-1,3-glucanase has the opposite effect leading to an SEL increase. The second mechanism is based on the hypothesis that the spoke-like extensions between the desmotubule and the PM are actually formed by tether proteins that regulate their tether length in response to Ca2 þ . PD are indeed regulated by Ca2 þ . Elevated and reduced cytosolic Ca2 þ levels were correlated with a decrease and increase in PD SEL, respectively. However, callose synthase holoenzyme complexes are also regulated by Ca2 þ . Thus, further studies are needed to substantiate this model. (b) Modification of PD during viral infection. PD are associated with signaling complexes that sense the presence of pathogens and cause callose accumulation and PD closure (i). Virus-encoded MPs modify PD and allow virus movement irrespective of the physiological status of the cells by two different basic mechanisms. MPs that support virus movement in the form of a VRC/vRNP, as exemplified by the MP of TMV, modify the SEL by inducing the degradation of callose deposits, presumably by recruitment or activation of b-1,3, glucanases (ii). In contrast, tubule-forming viruses like CaMV, GFLV, or CPMV have MPs with the capacity to form a tubule-like structure within the PD channel allowing whole virion particles to move between cells. MP tubule formation depends on the interaction of MP with members of the Plasmodesmata Located Protein (PDLP) family and leads to the displacement of the desmotubule and, thus, to the disruption of the ER-connectivity between cells (iii).

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recognition of a Y-shaped RNA element (YRE) and also binds the complex to the membrane. Unlike the MP of TMV, the MP of RCNMV has no affinity for binding to the ER. Instead, the localization of this MP to the VRC rather depends on replicated RNA1. Interestingly, translation of MP from RNA2 depends on replicated RNA2. This way, virus movement can occur only after a functional VRC has been assembled and both viral RNAs have been replicated. The binding of the MP to the VRC is supported by the chloroplastic protein glyceraldehyde-3-phosphate dehydrogenase (GAPDH), which also binds to the smaller replicase subunit (p27). The role of chloroplastic proteins in virus replication and movement has also been noted for other viruses like TMV and Bamboo mosaic virus (BaMV). As chloroplastic proteins and salicylic acid (SA) produced in chloroplasts are implicated in the regulation of PD, it may not be surprising if chloroplasts play a yet uncharacterized role in coordinating virus replication and intercellular transport through PD. The interaction of the small subunit replicase of RCNMV with the ER and the resulting membrane perturbations involved in the formation of the VRCs are linked to interactions of p27 with components of the secretory pathway, such as ADP ribosylation factor 1 (ARF1), the target of the secretory pathway inhibitor Brefeldin A (BFA). Consistently, virus replication is inhibited by treatment of infected cells with BFA or by inhibition of the secretory pathway protein SAR1, indicating that RCNMV replication and, thus, also cell-to-cell movement depend on membrane trafficking within the COPI/COPII-dependent secretory pathway. These and many other studies indicate that the deformation and recruitment of membranes for virus replication is a rather complex process that generally involves viral interference with lipid metabolism and protein regulation, targeting and transport. Studies using heterologous expression in yeast have shown that the formation of replication vesicles by tombus-and bromovirus replicase proteins involves reticulons as well as ESCRT (Endosomal Sorting Complexes Required for Transport) and MVB (multivesicular endosome) proteins, thus implying a role of endosomal pathways and of proteins shaping the structure and curvature of membranes. RCNMV exemplifies a relatively simple system in which only two parts of the viral genome must be replicated and moved between cells to spread infection. A more extreme case is represented by the multipartite nanoviruses, for example Faba bean necrotic stunt virus (FBNSV), which consists of eight genome segments. Recent studies to determine how the replication and cell-to-cell movement of eight genome segments can be coordinated led to the finding that the different segments of the virus do not occur in the same cells but in different cells. This exciting observation challenges the current view that virus replication must rely on single cells. Rather, nanoviruses, and potentially also other viruses with segmented genomes, may hijack groups of plant cells for replication and use intercellular trafficking of the virus-encoded products for functional complementation between segments and their coordinated replication and movement. Given that virus replication and movement for many viruses are tightly connected, there must be a mechanism to determine which of the viral genomes stay in the cell to form virion progeny and which of the viral genomes are transported to PD to spread infection into adjacent cells. TMV does not require its coat protein (CP) for cell-to-cell movement and, during the time course of infection, the CP is expressed later than MP, thereby ensuring that virions are formed after the virus already spread infection between cells. The above-described studies with TMV but also studies involving other viruses like PVX and TuMV have shown that the VRC-containing membrane compartments occurring during infection are divided into small, mobile compartments associated with virus movement and larger, sessile compartments that occur later and function as viral factories that produce virion progeny. Although the two different compartments can often be observed in the same cells and may communicate with each other, the larger virion-producing compartments of TMV and also those of RCNMV and PVX rather mature after virus movement has occurred. The mechanisms by which viruses transport their mobile VRCs to PD differ (Fig. 3) and many of the differences correlate with the mechanism by which the respective viruses modify PD to facilitate intercellular movement (see below). Because the ER membrane is continuous through PD between adjacent cells, many viruses target PD simply by transporting their VRCs along the ER membrane. However, several viruses, particularly the ‘tubule-forming’ viruses disrupt the ER continuity between cells (Fig. 2) and, therefore, depend on other, or additional, pathways to target the PD. Non-tubule-forming viruses can utilize the ER membrane because it is a fluid structure allowing embedded proteins and protein complexes to move by Fig. 3 Examples of different PD targeting pathways used by viruses. (a) TMV forms VRCs at microtubule-associated sites of the cortical ER-actin network and targets the PD by diffusion along the ER. The targeting of the MP to PD and the spread of infection depends on class XI and class VIII myosin motors. While some early VRCs undergo movement and spread infection to new cells, other “early VRCs” stay behind and become virus factories that produce virion progeny. When class VIII myosins are inhibited, the MP accumulates in the PM and cannot enter PD. The concentration of MP within PM rafts at PD may involve endosomal recycling. (b) TuMV forms replication vesicles induced by the viral 6K2 protein that bud off from the ER at ER exit sites in a COPII-COPI-dependent manner and then move along actin filaments to reach PD. Cylindrical inclusions formed by the cylindrical inclusion protein (CI) acts as a docking point for 6K2 and for a viral protein involved in TuMV movement, called P3N-PIPO. The P3N-PIPO has the capacity to modify the SEL of PD. Thus, following docking, the cylindrical inclusion my guide P3N-PIPO and the 6K2-associated VRCs to the PD channel. (c) Tubule-forming viruses like GFLV, CaMV, and CPMV have MPs that assemble to hollow tubules that allow the cell-to-cell transport of whole virion particles. Tubule formation disrupts the ER connectivity between cells. PD targeting, tubule assembly, and virus movement depend on interaction of the MPs with members of the PDLP family at PD. The targeting of PDLP to PD depends on the ER-Golgi secretory pathway and, thus, virus movement also depends on this pathway. The cellular pathway by which GFLV targets its MP and virions to PD is not known. More is known for CaMV (but not shown here), which forms stationary and motile virion factories. The motile virion factories are transported to PD along actin filaments and deliver virion particles to the PD channel in a process involving viral proteins present in the motile complex and the MP at PD. Transport of virions through the tubules formed by the tubule-forming viruses depends on the interaction between the viral coat protein subunits of the virion and the MP within the tubule. The mechanism that drives virion transport though the tubule and whether this process may involve non-viral proteins remains to be elucidated.

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diffusion. Moreover, similar to the ER-associated movements of organelles, such as Golgi complexes, mitochondria and peroxisomes, also viruses enhance the efficiency and directionality of their transport with the help of myosin motor proteins. By blocking the pathway of myosin, achieved by expression of actin-binding protein or by blocking specific myosin motors using myosin RNA silencing or expression of dominantly inhibiting myosin mutants, it was demonstrated that TMV depends on class XI and class VIII myosins for transport. Whereas class XI myosins support the transport of the TMV VRCs along the ER-actin network, class VIII myosins play a role at the PM and in the interaction of the MP with PD (Fig. 3(a)). ER-mediated targeting to PD has also been demonstrated for many other viruses. However, while the mechanism of ER-mediated transport is rather simple for TMV, which encodes only one MP, the mechanism is more complex for viruses that encode more than one MP. For example, the rod-shaped hordei-like (hordei-, pomo-, peclu-, and beny-) viruses and potexviruses encode three MPs in overlapping open reading frames, the triple gene block (TGB). The general model for movement of these viruses involves the binding of viral RNA by TGB1 and the targeting of the TGB1:RNA complex to PD with the help of TGB2 and TGB3, transmembrane proteins that are localized to the ER. Studies using hordeiviruses, pomoviruses, and potexviruses as models demonstrated that TGB3 contains the signals for targeting specific sites on the ER, for the recruitment of TGB2, for transporting the TGB2/TGB3 complex to ER sites in proximity to the PD, and for recruitment of TGB1 to these sites. Several observations indicated that the TGB2/TGB3 complex is transported by ER-derived, motile complexes along actin filaments. Similar to the motile TMV VRCs, also these motile TGB complexes may represent VRCs. PVX, for example, replicates in association with the ER and the motile complexes contain the viral replicase as well as ribosomes and virions. At later stages of infection, the TGB proteins also colocalize with viral RNA to ER-associated replication factories that produce virions. Several lines of evidence suggest a role of microtubules in the localized formation and transport of the TGB VRCs. Thus, although many details remain to be explored, the mechanism of TGB virus movement using ER-guided movement of motile VRCs, potentially microtubules, and stationary VRCs for the formation of viral factories, appears to be similar to the ER-guided mechanism used by TMV. However, it is not always clear whether motile complexes described as “ER-derived” are still part of the ER or whether these rather represent ER vesicles that bud off from the ER. The TGB3 of Poa semilatent virus (PSV, an hordeivirus) forms motile complexes upon inhibition of the COPII-dependent ER-Golgi pathway thus indicating that PSV and potentially other TGB viruses target PD by an ER-mediated route that does not require exit from the ER. However, this is different, for example, for Turnip mosaic virus (TuMV, a potyvirus), which forms replication vesicles induced by the viral 6K2 protein that bud off from the ER at ER exit sites in a COPII-COPI-dependent manner and then move along actin filaments to reach PD (Fig. 3(b)). The requirement of exit from the ER for this virus may be linked to the specialized mechanism by which this virus targets and modifies PD for movement and which may disturb the normal ER continuity and PD function between cells. Indeed, this virus encodes a cylindrical inclusion protein (CI) that forms a highly elaborated virus-induced transport structure at PD. The CI protein acts as a docking point for 6K2 and for a viral protein involved in TuMV movement, called P3N-PIPO. P3N-PIPO has the capacity to modify the SEL of PD. Thus, following docking, the cylindrical inclusion my guide P3N-PIPO and the 6K2-associated VRCs to the PD channel. Alternative routes for the targeting of the virus to PD are indeed required for the already mentioned “tubule-forming viruses“, which alter the inner structure of PD by forming a transport tubule assembled by MP and disrupt the ER continuity between cells (Fig. 3(c)). Viruses encoding tubule-forming MPs can be found in several virus families, such as the Secoviridae, Caulimoviridae, Badnaviridae, Tospoviridae, Ilarviridae, and Bromoviridae. One example is Cauliflower mosaic virus (CaMV), a pararetrovirus which replicates its dsDNA genome by reverse transcription of a terminally redundant RNA template. Infected cells form electron dense inclusion bodies in which the P6 protein of the virus facilitates the translation of the polycistronic viral RNA, and viral and host proteins are recruited for replication and virion assembly. In a manner similar to the virus factories formed by TMV, the inclusion bodies form in association with the ER and microtubules and can either be large, functioning as stationary “virion factories’, or small, functioning as motile complexes that target the PD. To reach PD, the motile complexes are transported along actin filaments facilitated by a protein called CHUP1 (Chloroplast Unusual Positioning Protein) that usually anchors chloroplasts to actin microfilaments and is required for the movements of chloroplasts in response to light. At the PD, the inclusion body delivers virions to the PD channel, in a process potentially involving interaction of P6 with the PD-localized MP (P1) and another P6-interacting protein called AtSRC2.2. Interestingly, the MP of the virus (P1) targets PD independently of the P6 inclusion bodies. As shown for CaMV and also for Grapevine fanleaf virus (GFLV) (Fig. 3(c)), the PD targeting by MP, efficient tubule assembly by MP and virus infection supported by MP depend on interaction of the tubule-forming MP with PD-localized proteins (PDLP), a multigene protein family that localizes to PD via the ER-Golgi secretory pathway. The secretory pathway consists of interacting and highly dynamic organelles that move intracellularly with support of the actin cytoskeleton. Consistently, the PD targeting of PDLP and, consequently, that of the tubule-forming MP is reduced by inhibition of the secretory pathway (e.g., with the chemical inhibitor Brefeldin A, BFA), of actin filaments (e.g., with the chemical inhibitor Latrunculin A, Lat A), or of myosins (e.g., with the chemical inhibitor 2,3 butane-dione-monoxime, BDM). Moreover, PDLP and MP targeting to PD as well as GFLV movement and tubule formation by the GFLV MP depend on class XI myosins. Microtubules also play a role in this pathway, since GFLV MP tubules were shown to form at ectopic sites rather than at PD when microtubule inhibitors were present. So far, the described mechanisms of intracellular transport involve membranes of the ER and the secretory system. However, several viruses also interact with endosomal pathways for movement. For example, TGB2 and TGB3 proteins do not move with viral RNA into adjacent cells but are recycled by endosomal membrane trafficking for further rounds of transport. Endosomal recycling may also play a role for the MP of TMV (Fig. 3(a)) and other MPs that interact with synaptotagmin A (SYTA, SYT1), one of five SYTs in Arabidopsis. SYTA was shown to localize to PM-derived endosomes at discrete areas at the PM, and to play an important role in endocytosis as well as in MP-mediated cell-to-cell transport. TMV movement may indeed involve a PM-associated function, since the MP showed strong accumulation at the PM when its targeting to PD was blocked by inhibition

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of class VIII myosins. This observation may be related to the notion that PD contain PM rafts and that the MP of TMV and also the MPs of other viruses may be part of such rafts when present in PD. Thus, it appears conceivable that SYTA-associated endosomal recycling maintains the association of MP with membrane rafts and that myosin VIII is required to anchor the rafts at PD. Several MPs contain specific sorting motifs that are required for sorting into clathrin-coated endosomal vesicles. Mutation of such motif within the P1 MP of CaMV allowed the protein to accumulate at the PM but abolished its recruitment to PD thus again indicating that an endosomal recapture pathway plays a role in the recruitment of MPs from the PM to PD.

Intercellular Transport PD are cytoplasmic communication channels in the cell walls between adjoining cells and play pivotal roles during plant development and in the interaction of plants with biotic and abiotic factors by allowing the cell-to-cell transport of water, metabolites, nutrients and hormones as well as of informational macromolecules such as RNAs, proteins and protein-RNA complexes. Structurally, they are concentric structures in which a cytoplasmic sleeve is delimited by the centrally located rod of the ER (the desmotubule) on one side and the PM that lines the outer diameter of the channel on the other. PD are associated with signaling components such as receptor proteins and receptor protein kinases as well as signal relaying components (e.g., the already mentioned PDLPs) by which the PD constantly adjust their permeability in response to developmental and environmental cues. The PM of PD contains rafts or other types of microdomains enriched in specific lipids and which bind and concentrate proteins carrying lipophilic modifications (e.g., a glycolsyl-phosphatidyl-inositol (GPI) “anchor”) that are enriched in PD. The SEL of PD, thus the upper limit of the size of molecules transported by PD, is determined by the degree of callose deposition in the cell wall surrounding the PD neck regions (Fig. 2(a)). Callose synthesis and accumulation by callose synthases in the cell wall leads to closure of the cytoplasmic compartment by forcing the PM against the desmotubule, whereas the degradation of callose by beta1,3-glucanase opens the cytoplasmic space for intercellular transport. It has been proposed that the SEL of PD may also be regulated by proteins that tether the desmotubule to the PM and are able to regulate their tether length, and, thus, the size of the cytoplasmic space. The SEL of PD is regulated in response to Ca2 þ waves or the sensing of pathogens. For example, one protein of the PDLP family, PDLP5, was shown to regulate the SEL of PD by stimulation of callose synthesis through callose synthase 1 (CalS1) and CalS8. Moreover, PDLP5 is required for regulating PD permeability induced by bacterial pathogen or by accumulation of salicylic acid (SA), a hormone playing central roles in stimulating immunity against biotrophic pathogens, including viruses. To overcome the gating of PD, the virus-encoded MPs modify PD in different ways (Fig. 2(b)). Whereas the MP of TMV and the MPs of probably most other viruses cause a transient modification of the PD SEL, the MPs of ‘tubule forming viruses’ assemble a hollow tubule within PD and thereby drastically alter the internal structure and function of the PD channel. The tubules formed within PD allow the cell-to-cell transport of whole virion particles, whereas viruses that modify the PD SEL move between cells rather in a non-virion form (e.g., as VRCs) and may even be independent of CP, as in the case of TMV. Virion entry into tubules depends on direct or indirect interactions between the CP of the capsid and MP residues exposed in the inner tubule wall. Whereas the CPs and MPs of Cowpea mosaic virus (CPMV) and GFLV interact directly, the binding of the CP of CaMV to the MP (P1) is mediated through another viral protein (P3). The CaMV CP also binds to P6. Thus, both P6 and P3 may bind to the virion surface to assist in the transfer of virions from P6 inclusion bodies to MP (P1) within the tubule. Given that tubules are assembled by MP produced in the infected cells and have a given length, it seems likely that the tubules are dynamic structures in which MP units and associated virions that have docked to the tubule treadmill through the tubule length and are released at the distal end. However, whether tubules are indeed dynamic or rather static structures remains to be demonstrated. The mechanism by which other MPs like that of TMV modify the PD SEL is not known. Although electron micrographs indicated that the TMV MP forms fibrillar substructures within PD cavities, it remains unknown whether these structures are functional. Other studies indicate that viruses modify the PD SEL by direct or indirect interaction with one or more beta-1,3-glucanases (GLU) and thereby direct the degradation of callose. The expression of specific GLU genes was shown to increase during infection and the efficiency of TMV movement was affected when the level of GLU expression was altered, thus confirming the hypothesis that a callose-degrading enzyme activity may regulate virus movement. Consistently, a specific glucanase, AtBG_pap, which localizes to PD, was identified. However, although an atbg_pap mutant exhibited reduced virus movement associated with increased PD-localized callose levels, it still remains to be clarified whether viruses like TMV indeed operate GLU activities for virus movement or whether they rather act by preventing infection-triggered callose synthesis. The MP of TMV and the TBG1 protein of PVX were shown to bind specific ankyrin repeat-containing proteins (TIP1-3, ANK). The tobacco homolog ANK was shown to facilitate virus movement and its co-expression with MP resulted in its recruitment from the cytoplasm to PD, in reduced callose levels at PD and increased intercellular diffusion of MP. Thus, ANK and related factors may be recruited by MP to facilitate virus movement. However, the exact mechanism through which these proteins act in virus movement remains to be further studied. More recent studies indicated that the MP harbors a specific PD-localization signal (PLS) in its N-terminus. Interestingly, this domain is recognized by the MP-interacting protein SYTA/SYT1, which was already mentioned as a host protein required for the cell-to-cell movement of TMV and of several other viruses. Although the SYT proteins may play a role in Ca2 þ -sensitive membrane trafficking, their similarity to mammalian synaptotagmin (E-Syt) and yeast tricalbin (Tcb) families suggests that they (also) act as membrane tethers. Thus, in addition to its function in endosomal trafficking, SYTA is now also recognized as an ER-anchored protein that tethers the ER to the PM at ER:PM contact sites and has critical functions in stabilizing the ER network. In binding MP, SYTA may play overlapping roles both in endosomal recycling of MP to maintain its enrichment at the cell periphery and also in

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anchoring the MP-containing TMV VRCs to the microtubule-associated ER:PM contact sites mentioned above. Moreover, given the close proximity between the desmotubular ER and the PM within PD, it has been proposed that also the PD may represent ER:PM contact sites. In this case, SYTA may take part in the tethering of the desmotubule to the PM and thereby controlling the cytoplasmic space and, thus, the PD SEL by changing its tether length in response to Ca2 þ (Fig. 2(a)). SYTA and other markers of ER:PM contact sites (e.g., VAP27) were found to localize to PD and to interact with PD-localized reticulons, which is consistent with this hypothesis. The finding that SYTA binds the PLS of MP and is required for the MP targeting to PD suggests that SYTA recruits or binds MP at PD. The association of MP with SYTA at PD may also imply the possibility that MP modifies PD SEL through regulating the length of SYTA tethers. Several studies indicate that the PD SEL may also be directly or indirectly controlled by actin filaments. Consistent with a role of actin in controlling PD SEL, the MPs of TMV and of Cucumber mosaic virus (CMV) were shown to exhibit actin severing activity in vitro. Moreover, the stabilization of actin filaments by treatment with phalloidin prevented the ability of MP to increase the PD SEL in vivo. Although these observations suggest that MPs manipulate actin to control the PD permeability, it remains to be shown whether these MPs indeed interact with actin in vivo and, if so, whether the actin severing activity of these MPs occurs at PD or at other locations in the cell. Nevertheless, class VIII myosins were localized to PD, and such myosins were shown to be required for the interaction of the TMV MP with PD. Future studies may reveal whether or not PD indeed contain an actomyosin system and whether this system has a role in controlling the aperture of PD. Virus movement through PD likely requires the activity of virus-encoded chaperones. This is illustrated by the virally encoded Hsp70 chaperone homolog (Hsp70h) of Beet yellows virus (BYV). This virus has a particularly large RNA genome and forms very long and flexible virions. Hsp70h is one of five virus-encoded proteins required for transport. This protein targets PD and is also found in actin-associated motile granules. Consistent with a role of the granules in intracellular transport, the targeting of Hsp70h to PD depends on intact actin filaments and class VIII myosins. The Hsp70h is a component of the filamentous capsid and its ATPase activity is required for virus cell-to-cell movement. These findings led to a model where Hsp70h mediates virion assembly and, once localized to the PD, actively translocates the virion from cell to cell via an ATP-dependent process. A role of viral proteins in plasmodesmal translocation of virus is also supported by the requirement of NTPase activities in other viral proteins for virus movement, such as in the TGBp1 helicase of PMTV and the CP of PVX. Moreover, also the helicase domains within the N-terminally overlapping replicase subunits of TMV are required for cell-to-cell movement. It seems plausible that the helicase activity found in several viruses is necessary for the remodeling of viral RNA during its passage through the PD. As has been already mentioned, the process of virus movement through PD may also be regulated by posttranslational modifications of MP. In addition to its control by phosphorylation, the MP of TMV is also controlled by ubiquitinylation and degradation by the 26S proteasome. ER stress caused by viral replication and MP production leads to the activation of the AAA-ATPase CDC48, which binds and extracts the ubiquitinylated MP from the ER and guides it into the cytoplasm for degradation. While this CDC48-dependent process of removing MP from the ER transport pathway may represent a plant defense response, this mechanism may also be part of a viral strategy to inhibit further virus movement and allow the virus to switch infection from an early phase focused on virus movement to a late phase focused on virus replication and virion production.

Systemic Transport To achieve systemic movement, the infecting virus needs to pass through different cell types of the vascular veins until it reaches the phloem sieve elements for long-distance movement into other parts of the plant (Fig. 1). The phloem transports and distributes photosynthetic assimilates from source regions in mature leaves to sink regions that require carbon for growth and development. In addition to sugars the phloem allows the long-distance transport of many solutes, hormones, proteins and RNAs. Thus, once in the sieve element, the virus can exploit this mass flow from source to sink for its transport. The vascular veins are defined as major and minor, depending on their structure, location, branching patterns, and function. The vascular tissues of each vein, for example in tobacco, consist of cell types of the phloem, thus the sieve elements and adjacent companion cells together with phloem parenchyma cells, and cell types of the xylem, thus the xylem vessels and xylem parenchyma cells. All these cell-types are surrounded by cells of the bundle sheath which delimits the vascular tissue against the mesophyll tissue. Plant species differ in their strategy of loading sugars from the mesophyll into the phloem and this difference is reflected by the number of PD that connect the bundle sheath cells and the companion cells. In symplastic loaders, thus plant species that load sugars through PD, many PD are located at this interface, whereas in apoplastic loaders, thus plant species in which sugars are transported across the cell wall by specific transporter proteins located in the PM, only few PD are present at this interface. Since tobacco is an apoplastic loader, TMV is apparently able to find these few PD to enter the vascular tissue and cause systemic infection in this species. Moreover, whereas all species examined to date have PD between companion cells and sieve elements, PD are rather infrequent between the sieve element and either phloem parenchyma or bundle sheath cells. Thus, the sieve element is isolated from the rest of the leaf and the majority of molecules must enter the sieve element through companion cells, which therefore act as a major gatekeeper for phloem entry. Viruses use both minor or major veins to enter and exit the phloem in source and sink tissues, respectively. To reach the phloem, viruses recapitulate mechanisms of cell-to-cell movement. However, the ability to move longdistance depends on additional mechanisms. TMV, for example, moves cell-to-cell without CP but requires CP for systemic movement. Without the CP, the virus can reach bundle sheath and vascular parenchyma cells of the minor veins by cell-to-cell movement, but shows reduces accumulation in the companion cells compared to the wild type virus. This requirement of CP may

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be consistent with several other findings in the literature indicating a requirement of specific interactions between virus and host factors at the boundary between different cell types. Moreover, because the phloem sieve elements are devoid of ribosomes and therefore lack the machinery to support virus replication, the viral genome may also require specific protection to reach the distant tissues. Virion particles of several viruses have been detected in the phloem, thus suggesting that virions are the most prevalent transport form of viruses in the phloem. For viruses that move cell-to-cell in a non-virion form like TMV, the exact site of virion formation is not known. While virion particles of several viruses have been detected in the PD that connect companion cells with sieve elements, the virions of CMV were only found in sieve elements thus suggesting that the virions are formed in the parietal, ER membrane-containing layer of the sieve element. Interestingly, MPs can enter the sieve element and may also be translocated to distant sites to exert an influence on PD in these regions. PD in sink tissues have larger SELs than PD in source tissues which facilitates phloem unloading. Thus, MPs that have entered the phloem stream could spread into and between cells of sink tissues and thereby prime these tissues for the subsequent export and transport of the virus. Although long-distance movement appears to require encapsidation of the virus genome, umbraviruses like Groundnut rosette virus (GRV) do not encode a CP. Systemic movement of GRV depends on its ORF3 protein, which targets Cajal bodies in the nucleolus to recruit the nucleolar protein fibrillarin and to assemble complexes with viral RNA able to move systemically. Viroids are single-stranded, circular RNA molecules that even do not encode any protein but can nevertheless cause systemic infection. Thus, all proteins required for viroid replication and transport must be of plant host origin. The RNA is folded into a series of loops and bulges and studies on Potato spindle tuber viroid (PSTVd) and mutational analysis demonstrated that several of them are required for cell-to-cell and/or phloem entry and systemic transport. Certain phloem proteins that bind to viroids have been identified but their functional significance in viroid movement remains to be investigated. Several viruses, for example, viruses belonging to the aphid-transmitted Luteoviridae family, are phloem-limited because they can enter the phloem but fail to be unloaded. The restriction of virus movement of these phloem-limited viruses usually occurs at some point outside the sieve element-companion cell (SE-CC) complex, suggesting that the limiting step in tissue invasion lies in the cell-to-cell post-phloem movement rather than in the ability to exit from the SE-CC.

Virus Movement and Plant Defense Responses Successful systemic infection critically depends on the ability of the infecting virus to suppress the defense reactions of the plant. Replicating viruses trigger RNA silencing that targets viral RNA for cleavage and/or translational repression. RNA silencing is initiated by dsRNA produced during replication. This dsRNA is cleaved to small-interfering RNAs (siRNAs) that guide Argonaute protein-containing RNA-Induced Silencing Complexes (RISC) to further cleave viral RNA in an RNA sequence-dependent manner. Viruses encode a Viral Suppressor of RNA silencing (VSR) that binds small RNAs or interferes with another step of the RNA silencing pathway. Because plants use small RNAs and RNA silencing pathways to control gene expression, VSR activities can interfere with normal plant development and, thereby, play an important role virus-induced pathogenicity. Virus-derived and other small RNAs are mobile and may move ahead of the virus to immunize cells ahead of infection. The level and systemic spread of antiviral siRNAs is enhanced by an amplifying pathway involving RNA-dependent RNA polymerase 6 (RDR6). Enhanced production of antiviral siRNAs by RDR6 and their spread into sink tissues plays an important role in immune surveillance in meristems and derived young tissues. Viruses with weak or deleted VSR functions that are unable to suppress this amplified antiviral RNA silencing may fail to cause a sustained systemic infection. More recent studies have shown that in addition to triggering RNA silencing, dsRNAs also act as potent elicitors of Pattern-Triggered immunity (PTI), which inhibits virus propagation as well. Future studies may show how the two types of dsRNA-triggered plant host responses are coordinated during infection and whether they play synergistic roles in antiviral defense. A third layer of antiviral defense involves resistance (R) genes that cause Effector-triggered immunity (ETI) against specific viruses. ETI can lead to a hypersensitive response (HR) involving cell death, high levels of the immunity hormone SA, and the establishment of systemic acquired resistance (SAR). An important example is the tobacco N gene, which triggers an HR upon recognition of the VSR effector of TMV (the small subunits of the TMV replicase). An example for ETI without induction of an HR is provided by the RTM1 gene in Arabidopsis. The lectin-like RTM1 protein acts specifically in the phloem to restrict the long-distance movement of potyviruses, such as Tobacco etch virus (TEV), Lettuce mosaic virus (LMV), and Plum pox virus (PPV). The protein forms a complex with proteins encoded by other RTM resistance genes and likely targets the potyviral particle or a CP-containing ribonucleoprotein complex.

Further Reading Epel, B.L., 2009. Plant viruses spread by diffusion on ER-associated movement-protein-rafts through plasmodesmata gated by viral induced host beta-1,3-glucanases. Seminars in Cell and Developmental Biology 20, 1074–1081. Ghoshal, B., Sanfacon, H., 2015. Symptom recovery in virus-infected plants: Revisiting the role of RNA silencing mechanisms. Virology 497–480, 167–179. Gouveia, B.C., Calil, P., Machado, J.P., Santos, A.A., Fontes, E.P., 2017. Immune receptors and co-receptors in antiviral innate immunity in plants. Frontiers in Microbiology 7, 2139. Heinlein, M., 2015. Plasmodesmata: Channels for viruses on the move. Methods in Molecular Biology 1217, 25–52. Heinlein, M., 2015. Plant virus replication and movement. Virology 479–480, 657–671. Kleinow, T., 2016. Plant-virus interactions: Molecular biology, intra-and intercellular transport. Kleinow, T. (Ed.), Heidelberg: Springer. Niehl, A., Heinlein, M., 2011. Cellular pathways for viral transport through plasmodesmata. Protoplasma 248, 75–99.

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Niehl, A., Peña, E.J., Amari, K., Heinlein, M., 2013. Microtubules in viral replication and transport. Plant Journal 75, 290–308. Niehl, A., Wyrsch, I., Boller, T., Heinlein, M., 2016. Double-stranded RNAs induce a pattern-triggered immune signaling pathway in plants. New Phytologist 211, 1008–1019. Peña, E.J., Heinlein, M., 2013. Cortical microtubule-associated ER sites: Organization centers of cell polarity and communication. Current Opinion in Plant Biology 16, 764–773. Pitzalis, N., Heinlein, M., 2018. The roles of membranes and associated cytoskeleton in plant virus replication and cell-to-cell movement. Journal of Experimental Botany 69, 117–132. Reagan, B.C., Ganusova, E.E., Fernandez, J.C., McCray, T.N., Burch-Smith, T.M., 2018. RNA on the move: The plasmodesmata perspective. Plant Science 275, 1–10. Sambade, A., Brandner, K., Hofmann, C., et al., 2008. Transport of TMV movement protein particles associated with the targeting of RNA to plasmodesmata. Traffic 9, 2073–2088. Schoelz, J.E., Angel, C.A., Nelson, R.S., Leisner, S.M., 2016. A model for intracellular movement of Cauliflower mosaic virus: The concept of the mobile virion factory. Journal of Experimental Botany 67, 2039–2048. Sevilem, I., Yadav, S.R., Helariutta, Y., 2017. Plasmodesmata: Channels for intercellular signaling during plant growth and development. Methods in Molecular Biology 1217, 3–27. Sicard, A., Pirolles, E., Gallet, R., et al., 2019. A multicellular way of life for a multipartite virus. Elife 8, e43599. Tilsner, J., Linnik, O., Louveaux, M., et al., 2013. Replication and trafficking of a plant virus are coupled at the entrances of plasmodesmata. Journal of Cell Biology 201, 981–995. Tilsner, J., Nicolas, W., Rosado, A., Bayer, E.M., 2016. Staying tight: Plasmodesmal membrane contact sites and the control of cell-to-cell connectivity in plants. Annual Review in Plant Biology 67, 337–364. Waigmann, E., Heinlein, M., 2007. Viral Transport in Plants. Plant Cell Monographs. vol. 7. Heidelberg: Springer. Wan, J., Cabanilla, D.G., Zheng, H., Laliberté, J.F., 2015. Turnip mosaic virus moves systemically through both phloem and xylem as membrane-associated complexes. Plant Physiology 167, 1374–1388. Wang, A., 2015. Dissecting the molecular network of virus-plant interactions: The complex roles of host factors. Annual Review of Phytopathology 53, 45–66.

Plant Antiviral Defense: Gene-Silencing Pathways Vitantonio Pantaleo, National Research Council, Research Unit of Bari, Bari, Italy Chikara Masuta and Hanako Shimura, Hokkaido University, Sapporo, Japan r 2021 Elsevier Ltd. All rights reserved. This is an update of G. Szittya, T. Dalmay, J. Burgyan, Plant Antiviral Defense: Gene Silencing Pathway, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00733-0.

Glossary AGOs Argonaute (AGO) proteins bind to siRNAs and cleave target RNAs in plants as part of the RNA-induced silencing complex (RISC). Arabidopsis thaliana has ten AGOs with specific functions, which are summarized as follows: AGO1, antiviral silencing; AGO2, DNA methylation, antiviral silencing; AGO3, antiviral silencing; AGO4, DNA methylation; AGO5, antiviral silencing; AGO6, DNA methylation; AGO7, antiviral silencing; AGO8, induction of denfeses against herbivores (insects); AGO9, DNA methylation; and AGO10, antiviral silencing, but also a singular pro-viral role. DCLs Dicer-like proteins (DCLs, RNase III-like class of proteins) cleave double-stranded RNAs (dsRNA) to generate mostly 21- to 24-nt siRNAs or miRNAs. A. thaliana has four DCLs with specific functions that act synergistically: DCL1, biogenesis of 21-nt miRNAs, 21-nt nat-siRNAs; DCL2, biogenesis of 22-nt siRNAs for antiviral silencing, 24-nt nat-siRNAs; DCL3, biogenesis of 24-nt siRNAs (het-siRNAs) for RdDM; and DCL4, biogenesis of 21-nt siRNAs (24-nt in rice) for antiviral silencing. RNA drug Viruses can be controlled through cellular antiviral RNA silencing (av-RNAi) against target viral genes by spraying double-stranded RNAs (dsRNAs), which are

processed into viral siRNAs (vsiRNAs) in the applied plant cells. siRNAs Short double-stranded RNAs ranging in size from 21- to 24-nt long generated by Dicer-like proteins (DCLs). siRNAs include: vasiRNA, virus-activated siRNAs (21- and 22-nt) derived from endogenous RNAs; vsiRNA, viral smallinterfering RNAs derived from viral RNAs; nat-siRNA, natural antisense transcript-derived siRNAs (21- or 24-nt); ta-siRNA, trans-acting siRNAs targeting mostly transcription factor genes; pha-siRNA, phased siRNAs (21- to 24-nt) from endogenous RNAs triggered by a particular miRNA; piRNA, piwi-interacting RNAs (21-nt) that suppress transposable elements (TEs) in the germline (to date not found in plants); het-siRNA, heterochromatic siRNAs (24-nt) generated by DCL3; easiRNA, epigenetically associated siRNA (21-nt), derived from pericentromeric TEs due to the transient loss of RNA-dependent DNA methylation (RdDM) in sperm. Viral silencing suppressors (VSRs) Most plant viruses encode proteins that interfere with various steps in the host RNA silencing machinery. The main mechanism can be explained by their ability to bind to dsRNAs and siRNAs. Some VSRs can target the host DNA methylation machinery to inhibit host resistance modulated by epigenetics.

Introduction RNA silencing or RNA interference (RNAi) refers to a conserved gene silencing process mediated by short RNAs, such as shortinterfering RNAs (siRNAs) and micro RNAs (miRNAs). RNAi is one of the most important mechanisms in regulating gene expression and repressing transposable elements (TEs) at the transcriptional or posttranscriptional level. Across the eukaryotic kingdom, with the exception of some yeast species including Saccharomyces cerevisiae, the importance of RNAi-based gene regulation has been recognized as implicated in central developmental processes, maintaining genome integrity, and resilience to biotic and abiotic environmental inputs. Furthermore, plants synthesize viral siRNAs (vsiRNAs) and viral-activated siRNAs (vasiRNAs) as a means of defense in response to viral invasion. RNAi was first described in the free-living nematode model Caenorhabditis elegans. It has been considered one of the most striking biological discoveries of the 1990s. Studies on RNAi have been readily and markedly supported by the genomic and Next Generation Sequencing (NGS) era in model plants; thus, studies in the field of small noncoding RNAs have advanced at a dramatic pace. It is now clear that RNAi is diverse, and in the last decade scientific efforts have been dedicated to extending the acquired knowledge from model plants to crops. Moreover, progress is being made to combine knowledge to the mechanisms and functions or to the applications of RNAi for sustainable plant protection.

Diverse Gene-Silencing Pathways in Plants After 30 years since the discovery of RNAi in plants, most of the related knowledge is still rooted in analyzes of the model plant Arabidopsis thaliana. RNAi pathways in Arabidopsis show considerable conservation of the molecular components: (1) long perfect or imperfect doublestranded RNAs (dsRNAs) are processed by DCLs into 21- to 24-nucleotide (nt) siRNAs; (2) siRNAs can be incorporated into members of

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Argonaute (AGO), the core proteins of RNA-induced silencing complex (RISC); (3) sRNAs guide RISC to target nucleic acids in a sequencespecific manner and exert their function in transcriptional or post-transcriptional gene silencing (TGS and PTGS, respectively). Across eukaryotes, plants have the most complex RNA-based control of gene expression. PTGS is usually directed against endogenous mRNAs and includes miRNA, trans-acting siRNA (ta-siRNA), natural antisense transcript-derived siRNA (nat-siRNA), and secondary phased siRNA (pha-siRNA) pathways. Instead, TGS is directed against components of the genome, including genes, repetitive elements and TEs. The microRNA (miRNA) pathway. miRNAs derive from specific noncoding genes denoted as MIR genes, which are transcribed by RNA polymerase II (Pol II) into long single-stranded primary transcripts (pri-miRNA). pri-miRNAs exhibit typical Pol II cap structures at their 50 termini and poly(A) tails at their 30 termini and often contain introns. pri-miRNA transcripts adopt a fold-back stem-loop structure that is processed into a mature miRNA duplex by DCL1. The processing of pri-miRNA into the miRNA duplex has been described in detail and requires: ● ● ● ● ● ● ●

Cap-binding protein 20 (CBP20) and CBP80; zinc finger protein SERRATE; dsRNA-binding protein/HYPONASTIC LEAVES 1 (DRB1/HYL1); Forkhead-associated (FHA) domain-containing protein DAWDLE (DDL); TOUGH protein (TGH); Proline-rich protein SICKLE (SIC); and RNA-binding protein MODIFIER OF SNC1, 2 (MOS2).

miRNA duplexes in nuclei undergo a maturation process. Both strands of an miRNA duplex are methylated at their 30 terminal nucleotide by the RNA methyltransferase HEN1, an S-adenosyl methionine (SAMe)-binding methyltransferase; the methyl group protects miRNAs from polyuridylation and degradation at the 30 termini. Most methylated miRNA duplexes are exported to the cytoplasm by the exportin-5 homolog HASTY. One strand of the miRNA duplex acts as a guide strand and is selectively loaded onto an AGO-containing RISC, whereas the other strand, the passenger strand (miRNA*), is discarded from the complex and rapidly degraded. Most miRNAs associate with AGO1, but specific associations with AGO2, AGO7 and AGO10 have also been reported. miRNAs are mainly 21-nt long, and mRNA cleavage is assumed as the common approach for miRNA-mediated gene regulation in plants. In addition to regulating RNA degradation, miRNAs occasionally direct DNA methylation or inhibit translation. miRNAs regulate endogenous gene expression during development and environmental adaptation. Thousands of miRNAs have been identified in Arabidopsis and other plant species. Many of them are conserved and species-specific miRNAs. The list of currently known plant miRNAs can be found in miRbase (see “Relevant Websites section”), which has been recently updated. The trans-acting siRNA pathway. ta-siRNAs derive from TRANS ACTING siRNA (TAS) genes, which are transcribed by Pol II into long non-coding, single-stranded (ss)RNAs that contain one or two specific miRNA target sites. TAS RNAs are targeted by miRNAcontaining AGO1 (or AGO7)-RISC, and the cleaved TAS noncoding RNA is converted into dsRNA by the suppressor of gene silencing-3 (SGS3) and host RNA-directed RNA polimerase 6 (RDR6). Subsequently, DCL4 assisted by dsRNA-binding protein 4 (DRB4) processes the dsRNA to generate a population of 21-nt ta-siRNAs in phase with the miRNA-guided cleavage site. Similar to most miRNAs, ta-siRNA duplexes are methylated by HEN1. One strand of the ta-siRNA duplex associates with RISC to guide the cleavage of target mRNAs. Some ta-siRNAs regulate juvenile-to-adult transition or, more generally, transcriptional factors involved in plant response to environmental stimuli. The natural cis-Antisense transcript-derived siRNA pathway. In plant genomes there are many genes that can be transcribed from complementary DNA strands at the same locus. If these genes are co-expressed, they produce overlapping sense/antisense transcripts. The overlapping region initiates the biogenesis of primary 24-nt nat-siRNAs through the action of a DCL (i.e., DCL2). Primary nat-siRNAs are loaded onto a yet unidentified AGO protein to direct the cleavage of the constitutively expressed complementary transcript. In a second step, the cleaved transcript is converted into dsRNA in a RDR6- and SGS3-dependent manner. This RNA amplification step may extend beyond the overlapping region to form outer siRNAs. Further DCL1-dependent processing of the newly synthesized dsRNA will generate 21-nt nat-siRNAs in phase with the primary cleavage. Similar to miRNAs and ta-siRNAs, nat-siRNA duplexes are methylated by HEN1. nat-siRNAs were discovered in A. thaliana grown under salt stress conditions. However, recent genome-wide analyzes have shown the widespread existence of overlapping sense/antisense transcripts, which raises the possibility that natsiRNAs could be major effectors of gene regulation in the plant-adaptive protection mechanism in response to either abiotic or biotic stress. Phased-siRNAs. The ability of miRNAs to indirectly regulate gene expression through the production of secondary siRNAs is an aspect that has become more evident only recently. Indeed, mRNAs cleaved by miRNA-loaded RISC can result in dsRNA synthesis, having the targeted transcript as template (like TAS ncRNAs). The newly synthesized dsRNA molecule is primarily the substrate for DCL4 to generate a new population of “secondary siRNAs”; these have a phased pattern similar to the ta-siRNA population, with siRNAs being produced at intervals of 21-nt (or 24-nt in rice) from the miRNA cleavage site. Secondary siRNAs can have a dramatic impact on gene regulation. They can act in cis, amplifying the silencing effect on their own target, or in trans, promoting a regulatory cascade if they derive from a member of large gene families. Nucleotide-binding site–leucine-rich repeat (NBS-LRR) clusters of pathogen resistance R-genes and pentatricopeptide repeats have been found subject to such regulatory cascades.

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Heterochromatic and epigenetically activated siRNA Pathways. The majority of endogenous plant siRNAs consist in 24-nt siRNAs derived from heterochromatin, i.e. chromosome regions known to be extremely poor in genes and rich in repeated sequences, DNA transposons and silent retroviruses (retro-transposons), collectively named TEs. The biogenesis of heterochromatic (het)-siRNAs depends on plant DNA-dependent RNA polymerases Pol IV (p4) and Pol V (p5) and, therefore, het-siRNAs are also known as p4/p5-siRNAs. It has been proposed that p4/p5-dependent ssRNA transcripts from heterochromatic DNA loci generate dsRNAs through the action of RDR2, in partnership with other host factors. The long dsRNAs are processed by DCL3 into 24-nt–long het-siRNAs that are, in turn, methylated by HEN1. het-siRNAs associate with AGO4, AGO6, and AGO9, which then trigger TGS at RNA-dependent DNA methylation (RdDM) targets. This results in a silent heterochromatin, the prevention of TE evasion and preservation of genome stability. In Arabidopsis, pericentromeric retrotransposoms and thousands of DNA transposons lose RDR2-directed RdDM during pollen grain formation. Epigenetically reactivated TEs are preferentially targeted by more than 50 miRNAs bound to AGO1. TE transcripts engage RDR6 to form dsRNAs allowing DCL4 to produce 21-nt epigenetically activated (ea) siRNAs. easiRNAs trigger cascade PTGS of TE transcripts during the reprogramming of germlines, a critical window for the plant’s life cycle. However, the transient loss of RdDM in sperm is restored after fertilization by maternal 24-nt het-siRNAs; therefore, easiRNAs are recognized as playing a role in fertility and male/female compatibility. Another class of het-siRNAs is “Needed for RDR2-independent DNA methylation” (NERD) siRNAs. NERD is a class of proteins unique to plants that contain GW motifs known to bind AGO proteins. NERD has been shown to mediate TGS by recruiting AGO2-bound 21- to 22-nt siRNAs at specific genomic sites (i.e., those unmethylated at H3 lysine 9). This alternative TGS pathway is AGO4/RDR2 independent but RDR6/AGO2 dependent.

Antiviral RNA Silencing Viral infection in plants requires replication in the infected cells, cell-to-cell movement, and long-distance spread of the virus. To defend themselves against viral infections plants use RNAi, a mechanism that fights parasitic nucleic acids. Invasive viruses and subviral infectious entities (satellites and viroids) are triggers of antiviral (av)-RNAi, which in turn targets viral genomes or transcripts for functional inactivation. av-RNAi results in a potent response of viral restriction using components of endogenous RNA silencing pathways. The av-RNAi mechanism in plants is best understood by examining the model plant A. thaliana, where the main players of RNA-silencing pathways have first been identified. The four DCLs (DCL1 to 4), ten AGO proteins (AGO1 to 10) and six RDR proteins (RDR1 to 6) are involved in av-RNAi depending on the type of plant virus. Different nucleic-acid content, genome organization and replication strategies influence the av-RNAi activated by plants. From viral long dsRNAs to viral siRNAs. Viral dsRNA. The av-RNAi machinery is first triggered by viral dsRNAs at the level of infected plant cells. The origin of viral dsRNA is diverse. Av-RNAi is differentially activated depending on viral species (Fig. 1). Endornaviruses and reoviruses possess dsRNA genomes that trigger the RNAi response. In geminiviruses, dsRNA arises from the bidirectional transcription of overlapping genes. It is a common belief that most viruses with a positive ( þ ) or negative (  ) single-stranded ssRNA genome (approximately 75% of known plant viruses) also produce dsRNAs when replicating in infected cells. However, replication occurs in specific and protected membrane niches in the plant cell that releases solely viral ssRNA. This indicates that dsRNA replication intermediates (which have not been detected in vivo so far) are not essential to activate av-RNAi. Conversely, viral ssRNAs are potent triggers of av-RNAi. dsRNA signals of viral ssRNAs (viral genomes, sub-genomes, and transcripts) can anneal and form dsRNA motifs, also referred to as dsRNA-like structured ssRNAs. DCLs. Primary viral dsRNA signals, either long perfect or imperfect, are recognized and processed by DCLs associated with DRBs into 21- to 24-nt RNA duplexes called primary viral small-interfering (vsi)RNAs. A. thaliana DCLs (DCL1 to 4) are all involved in primary vsiRNAs. Indeed, the dsRNAs derived from cytoplasmic RNA viruses are diced by DCLs 4 and 2, while those derived from viruses establishing minichromosomes, i.e., geminiviruses and caulimoviruses, are processed by DCLs 1 and 3. RDRs. DCLs 2 and 4 are also deputed to process RDR-dependent viral dsRNAs into secondary vsiRNAs. Indeed, combinations of RDR activities contribute to the formation of secondary dsRNAs of viral origin. Viral aberrant ssRNAs (i.e., bona fide truncated viral RNAs lacking 30 -polyA or 50 -MG 7-CAP) are recruited by RDRs and converted into long perfect dsRNAs. RDR-dependent viral dsRNAs are processed by DCLs into 21- to 24-nt secondary vsiRNAs, which are structurally indistinguishable from primary vsiRNAs. vsiRNAs (viral small-interfering RNAs). 21-nt vsiRNAs are predominantly associated with positive ( þ ) strand RNA viruses and made by DCL4 (which also produces endogenous ta-siRNAs, see above). 22-nt vsiRNAs produced by DCL2 (also involved in natsiRNAs, see above) in the presence of DCL4 often constitute less than 20% of the total vsiRNA population in infected tissues. Infection with DNA viruses induces a more abundant production of DCL3-derived, 24-nt vsiRNAs than 21- and 22-nt vsiRNAs in A. thaliana. DCL1 is active in viruses with DNA genomes and only in the absence of other DCLs. Primary and secondary vsiRNA duplexes are methylated by HEN1 at the 30 -termini of 20 -OH groups, analogously to endogenous functional siRNAs (see above). RISCs in av-RNAi. AGOs are the core components of as yet incompletely characterized RISCs (see above). AGO/RISC-mediated av-RNAi is one of the most important components of the plant's immune response against viruses. In the last decade, major efforts were dedicated to accurately describe the antiviral RISC in plants. Besides in vivo studies, mainly based on reverse genetic approaches, in vitro systems have been recently launched and appeared to be valuable tools to define and characterize AGOs with

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1. dsDNA viruses

Nucleus Pol II

Viral dsDNA

dsDNA

RT pgRNA

ssDNA

dsDNA

(+)RNA

Translation

Particles

2. ssDNA viruses Nucleus Rep+Host DNA Pol Viral ssDNA

Host DNA Pol ssDNA dsDNA

ssDNA

ssDNA unit-length

Pol II RNA

Particles

(+)RNA

Translation

3. dsRNA viruses Viral replicase dsRNA

(+)ssRNA

Particles

dsRNA

Translation

4. ssRNA viruses Viral replicase

ssRNA

ssRNA

Particles

(+)RNA Translation

Viral replicase

Subgenomic RNA

(+)RNA

Fig. 1 Viral RNA species targeted by av-RNA silencing in host cells. Plant viruses, including DNA viruses, have RNA states in their transcription and replication cycles. Such RNAs are mainly cleaved by Argonaute (AGO) in RNA-induced silencing complex (RISC) or subjected to translational inhibition, though rarely. Double-stranded RNA (dsRNA) viruses may be subjected directly to Dicer cleavage, but their RNA transcripts (ssRNAs) seem to be a major target of AGO because dsRNAs are suspected to be formed in viral particles. Although dsDNA and ssDNA viruses use their RNA transcripts for replication in the nucleus, such RNAs become a target of RNA silencing when translated in cytoplasm.

antiviral functions. For instance, cytoplasmic extracts of evacuolated Nicotiana tabacum BY2 protoplasts enabled the reconstitution of av-RNAi with a defined AGO protein and (an) exogenous siRNA(s) of choice. Both in vivo and in vitro studies have clarified that of the ten AGO proteins that were first identified in A. thaliana, AGO1,  2,  3,  5,  7, and  10 have antiviral RNA-silencing activity. Upon incorporation of 21-nt or 22-nt–long vsiRNAs, these AGOs exhibited cleavage activity of targeted RNAs of viral origin and the capacity to inhibit viral replication. The most highly characterized antivirally acting AGO1 and AGO2 proteins contribute to the removal of viral or subviral RNA entities by endonucleolytic cleavage of the target RNA in a vsiRNA-directed, sequence-specific manner. AGO4 has also been shown to have antiviral activity against ssDNA viruses, such as geminiviruses, through RdDM. Indeed, geminiviral mRNAs from bi-directional transcription are subject to av-RNAi similarly to RNA viruses. In addition, upon incorporation of 24-nt–long vsiRNAs, AGO4 promotes RdDM of viral chromatin leading to transcriptional gene silencing as a potent antiviral defense. Either length or the 50 -terminal nucleotide has a strong impact on vsiRNA sorting into specific antiviral AGOs. For example, vsiRNAs of 21- and 22-nt have a 50 -terminal U co-immunoprecipitate with AGO1, whereas those that have a 50 -terminal A

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co-immunoprecipitate with AGO2. Accordingly, vsiRNAs that are effective with AGO1 contain a U at the 50 -termini, whereas those effective with AGO2 contain an A at the 50 -termini. As for endogenous regulatory sRNAs (see above), after the incorporation of a vsiRNA duplex into an AGO protein, the passenger strand is removed while the remaining guide(g) strand directs the RISC to the cognate viral RNA. Function of vsiRNAs as a means of symptom development. In plants infected with viruses or viroids, vsiRNAs are abundantly produced. vsiRNAs induce av-RNAi and, in some instances, symptoms in infected plants. If vsiRNAs share high degrees of sequence similarity (or complementarity) with host genes, they can oftentimes induce specific symptoms through the host RNAi pathway. One prime example of vsiRNA-mediated gene silencing is the bright yellowing symptom on tobacco infected with a satellite RNA (Y-sat) of cucumber mosaic virus (CMV). Y-sat–derived siRNAs can induce silencing of the chlorophyll biosynthetic gene (ChlI), leading to yellow chlorosis. Viroid vsiRNAs have also been implicated for a long time in specific symptom induction, yet their molecular mechanisms remain unknown. Viroid-induced symptoms seem to be a consequence of complex physiological disorder by viroid infection where viroids interact with many host factors. However, some target gene candidates of viroid vsiRNAs have been demonstrated to be associated with symptom expression; these include, for example, the chloroplast heat-shock protein 90 (cHSP90) gene by peach latent mosaic viroid (PLMVd), the chloride channel protein-b-like gene (CLC-b-like) and 40S ribosomal protein S3a-like gene (RPS3a-like) by potato spindle tuber viroid (PSTVd). Recent advances in NGS and computational analyses have enabled the identification of vsiRNAs in vivo (denoted as degradome analysis) or in silico, which target host genes as well as the corresponding viral RNAs. By virtue of this technical development, vsiRNA-mediated host gene silencing leading to cleavage of the target transcripts has been proposed in many virus–host interactions, e.g., tobacco mosaic virus (TMV)-Arabidopsis, cauliflower mosaic virus (CaMV)-Arabidopsis, and sugarcane mosaic virus (SCMV)-maize, as well as in grapevine with multiple viral infections. Widespread silencing of host genes and other regulatory cascades induced by viral infections. In addition to vsiRNAs, plants produce a newly discovered class of endogenous siRNAs, the 21- and 22-nt virus-activated siRNAs (vasiRNAs). The production of vasiRNAs from thousands of Arabidopsis-encoding transcripts is induced by at least two distinct species of RNA viruses, i.e., CMV and turnip mosaic virus (TuMV). The genetic requirements of vasiRNAs are distinct from those of miRNAs, vsiRNAs, ta-siRNAs and pha-siRNAs; although they are DCL4-dependent, they require RDR1 for biogenesis. Upon incorporation into AGO2, vasiRNAs direct the widespread silencing of host genes in cis. The observations performed to date suggest that the widespread silencing of host genes (that largely include factors responsive to biotic and abiotic stimuli) directed by vasiRNAs plays a role in the virus resistance activated during antiviral silencing in a manner that is independent of the antiviral activity of vsiRNAs. Unlike the unique symptom induced by Y-sat, typical symptoms such as leaf mosaics, plant stunting and abnormal leaf development may not be simply explained by a specific vsiRNA, but rather by the result of all combined molecular effects mediated by miRNAs, vsiRNAs and vasiRNAs.

Viral and Plant RNA-Silencing Suppressors Plant virus genomes embed genetic information that is surprisingly restricted but sufficient to coordinate the entire viral life cycle including cellular infection, replication processes, invasion of the host and spread into the environment. Additionally, viral structural or non-structural proteins are programmed to protect viral genomes against av-RNAi. Such proteins are widely known as viral silencing suppressors (VSRs). Initially, sRNA binding has been shown to be the prevalent and unifying mechanism of suppression activity at least in model systems such as p19 of tombusviruses, HC-Pro of potyviruses, and p21 of closteroviruses. In the last decade, the attention devoted by the scientific community to VSRs presented an extremely rich and diverse scenario: each step or player of the plant RNAi machinery emerged as a putative target of VSRs. It was also revealed that certain VSRs have multiple targets. The VSRs and silencing players in host immunity are summarized in Table 1. Many VSRs have been described as pathogenicity determinants and, indeed, the transient or constitutive overexpression of VSRs resulted in developmental abnormalities in plants. These capacities were attributed to the binding affinities of VSRs to the respective miRNAs and, accordingly, to the alteration of miRNA accumulation and functions. However, because VSRs differentially bind to miRNAs and the important silencing players such as DCL1, AGO1 and AGO2 are regulated by miRNAs, viral symptom development is not simply explained through miRNA sequestration by VSRs. In this perspective, the influence of VSRs may have a significant impact on endogenous RNAi during the early stages of infection. Many endogenous RNA silencing suppressors (endoRSSs) have been identified in plants. It has been well documented that stress response, differentiation and development in plants are controlled by tuning the gene expression at the right time using miRNAs. In contrast to VSRs, which are produced to destroy host RNA silencing, endoRSSs probably play a roll in breaking miRNA-mediated gene regulation to remove excessively generated miRNAs. For example, it has been demonstrated that RNA silencing suppressed by FRY1 and XRN4 would be important for shaping root architecture.

Antiviral Silencing in Crops In some virus-host couples (woody plants, agro-ecosystems, forests) that have cohabited for long periods, av-RNAi should be more properly considered as a mechanism of surveillance ensuring the coexistence of both partners and the absence of symptoms, with

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Table 1

Target sites of VSRs to promote viral infection

Target sites Direct PTGS suppression 1. Generation of siRNA/miRNA (1) dsRNA binding Long dsRNA ds-siRNA

2.

3.

4. 5.

ds-miRNA (2) ssRNA binding ss-siRNA/miRNA binding (3) DCL downregulation DCL3, DCL4 DCL1 (4) DCL binding DRB4 (DCL4 cofactor) siRNA/miRNA methylation (1) HEN1 binding (2) No SAM supply to HEN1 Target RNA cleavage (1) AGO binding AGO1 AGO2 AGO4 (2) AGO downregulation AGO1 (3) AGO degradation AGO1, AGO2, AGO4 mRNA Translation (1) Ribosome binding Secondary siRNA amplification (1) RDR6 binding SGS3 (RDR6 cofactor) (2) RDR6 downregulation

VSRs examples

Remarks

NSs (TSWV), 2b (CMV), P38 (TCV-CP) HC-Pro (poty), 2b, P19 (TBSV), 126K (TMV) V2 (begomo) 2b, 126K AC4 (begomo), NS3 (RSV), P10 (GVA), V2

V2 binds only 24-nt ss-siRNA

P38 2b

Antagonistic effect of DCL1 upregulation

P6 (CaMV) HC-Pro HC-Pro, bC1

bC1 is produced by bsatellite of TYLCV

HC-Pro, 2b, p38, AC2 (begomo) p38 2b HC-Pro, 2b, 126K, P38

Downregulated by miR168 to control AGO1

P25 (potex), P0 (polero) HC-Pro, P6

P25, V2, P2 (RSV), CMV-CP HC-Pro

Indirect PTGS suppression mediated by host factors VPg (poty), HC-Pro XRN4 (exoribonuclease 4) binding WEL1 activation AC2/AL2 rgsCaM binding HC-Pro VPg, HC-Pro DCP2 (decapping protein 2) binding vasiRNAs suppression 2b

Reduce antiviral function of XRN4 Enhance WEL1 exonuclease activity Stimulate endogenous RSS Reduce antiviral function of DCP2 Reduce RDR1-dependent production of vasiRNAs

TGS suppression 1. DNA methylation (1) ADK binding (2) SAHH binding (3) SAMS binding (4) SAM decarboxylase I binding

ADK, adenosine kinase SAHH, S-adnocyl-L-homocystein hydrolase SAMS,S-adenocyl-L-methione syntase SAM, S-adenocyl-L-methionine

AL2/L2 (begomo) HC-Pro, bC1 HC-Pro C2 (begomo)

periodical outbreaks. The study of RNA silencing and av-RNAi mechanisms in the Arabidopsis model system, in which genetic dissection is possible, has helped the scientific community to accelerate the identification of biological gene functions. Upfront basic research has demonstrated that the av-RNAi process holds the key to future technological applications for the sustainable production of food (see also below, in this section). Indeed, av-RNAi is also effective in crop plants, which are important from an economic standpoint. Genome sequencing projects involve all crop types, including cereals and legumes, and are being extended to minor crops, ancient varieties or to wild species as a source of resistance to plant pathogens. In addition, image-based phenotyping is extremely effective to quantify the progression of biotic impacts and to associate it to genetic traits. The time is ripe to focus on av-RNAi in crop systems allowing us to contrast viruses (as well as other pathogens and pests) with the situation in crop plants. In Arabidopsis, the existence of multiple paralogs of RNA-silencing–associated proteins (e.g., ten AGOs, six RDRs and four DCLs) may explain the diversification of siRNAs and gene-silencing pathways; in crops the layer of complexity is more pronounced. The majority of crop plants possesses an allopolyploid genome, as they often undergo whole-genome duplication through the recombination of two or more genomes from progenitors. Paralogs of DCL and AGO genes are often present in crops,

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along with the recruitment of different accessory proteins and co-factors. This has led to functional diversification or specialization in different crops, adding layers of complexity. In rice (Oryza sativa L.), only AGO1 and AGO18 expressed activity against viruses. Rice AGO1 binds vsiRNA to form a RISC for resistance against rice stripe virus (RSV) and rice dwarf virus. AGO18 expression is almost undetectable, but is highly induced upon RSV infection. Overexpression of AGO18 leads to increased expression of AGO1, which together confer broad-spectrum resistance against viruses in rice. Nicotiana benthamiana (Domin) is considered either a model plant, since it is a permissive host for many plant viruses, or a typical case of alloploydization which is common in the Solanaceae family (e.g., tomato, potato, eggplant, pepper). The NbAGO1 gene is duplicated, and both homeologs (generically called “duplicates” or “isoforms”) retain the capacity to be transcribed into mRNAs and mainly differ in one 18-nucleotide insertion/deletion (indel). The indel is located eight nucleotides upstream of the target site of miR168, which is an important modulator of AGO1 expression. During infection with cymbidium ringspot virus, miR168 is up-regulated; however, the indel affects miR168-guided regulation and AGO1 isoforms, both of which appear to have specific involvement in symptom development. Similarly to N. benthamiana, tomato (Solanum lycopersicum L.) has AGO1 isoforms that are differently regulated in the case of tomato yellow leaf curl Sardinia virus (TYLCSV) infection. Moreover, functional indels affect the miRNA regulation of several transcriptional factors and NBS-LRR genes during viral infection. In tomato a large family of disease-resistance proteins with NBS-LRR has been detected, and siRNA datasets revealed the presence of a regulatory cascade affecting NBS-LRRs. The initiator of the cascade is miR482, which targets mRNAs for NBS-LRR and triggers mRNA decay as well as the production of secondary siRNAs in an RDR6-dependent manner. These secondary siRNAs target other NBS-LRR mRNAs and continue the regulatory cascade. The miR482-mediated silencing cascade is suppressed by VSRs in tomato infected with viruses, increasing expression of the mRNAs targeted by miR482 or secondary siRNAs. This process allows pathogen-inducible expression of NBS-LRR proteins and contributes to a novel layer of defense against pathogen attack and a nonrace-specific plant immunity induced by viral infection. Certain secondary siRNAs from NBS-LRR upon TYLCSV infection have been recognized to target and silence tomato genes involved in the photosynthetic pathway, partially explaining the phenotype plasticity in plants as well as the appearance of yellowing symptoms in the viral pathosystem.

Viral siRNA-Mediated Antiviral RNAi in Mammalian Cells Paralogs of Dicer and AGO genes have emerged via duplication during the evolution of all eukaryotes, including mammals. Along with the recruitment of different accessory proteins and co-factors, this process has led to functional diversification or specialization in different organisms. For example, insects encode two DCL genes – one mediating miRNA biogenesis and the other being responsible for siRNA (and vsiRNA) biogenesis. Conversely, mammals only encode a single DCL gene that generates both miRNAs and siRNAs. AGO2 is responsible for siRNA-mediated target RNA cleavage in insects, whereas AGO1 mediates miRNA-dependent gene silencing. Mammals, on the other hand, encode four AGO genes, all of which are involved in miRNA-guided gene silencing; only AGO2 is capable of cleaving target RNAs to mediate siRNA-dependent RNAi. In plants, vsiRNAs mainly induce the destruction of invading RNAs, leading to antiviral immunity. This is also the case in fungi and invertebrate cells. On the other hand, because mammalian antiviral defense heavily relies on interferon-mediated production of antiviral proteins, whether RNAi can indeed induce antiviral immunity in mammalian cells (especially somatic cells) has long been a matter of debate. Although numerous studies have reported the existence of siRNA-mediated av-RNAi in mammalian cells (not necessarily somatic cells), some questions still remain unanswered. It is clear that RNAi is involved in regulating mammalian viruses by host cellular miRNAs and that TEs have been found to be regulated by TE-derived siRNAs as well as by piwi-interacting RNAs (piRNAs) in both undifferentiated and somatic cells. However, to highlight the importance of av-RNAi in mammalian cells several factors must be carefully considered; for example: (1) whether VSRs is strong enough to reduce viral siRNA levels; (2) whether VSR can suppress other types of antiviral immune response; (3) whether the components in the RNAi pathway are well expressed in somatic cells; and (4) how cellular miRNAs contribute to the observed antiviral response.

View of Applications of av-RNAi RNA drugs: novel non-transgenic and pesticide-free approaches for plant protection. Recovery phenotype in plants was described almost 80 years ago. In brief, primary virus-infected leaves manifested severe viral symptoms, whereas the upper leaves showed attenuated symptoms and became immune to secondary infection of the same or similar viruses in terms of viral sequence. av-RNAi in plants explained the recovery phenotype and supported the notion of searching for applicative solutions to boost natural av-RNAi. Thus, plant viruses became tools to examine RNA vaccination. RNA vaccination is attractive for several reasons: (a) RNA molecules can be applied transiently, i.e., not in the context of transgenic plants; (b) sRNAs are naturally produced in plants and other eukaryotes to regulate endogenous gene expression and mediate defense responses and crosstalk among organisms, with the potential of being extended to other pathogens including fungi, pests and insects; and (c) RNA vaccination enables rapid and flexible solutions against emerging pathogen or pest resistance by simple alterations of a target sequence.

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Fig. 2 Mechanism through which exogenous viral double-stranded RNAs (dsRNAs) sprayed onto leaves induce antiviral RNA silencing (from A to D). (A) dsRNA is sprayed onto leaves. (B) The applied dsRNA penetrates into cells through wound cells. (C) The dsRNA is then cleaved into short-interfering RNAs (siRNAs) by cellular Dicer-like proteins (DCLs). The siRNAs generated bind to Argonaute (AGO), which is a component of RNA-induced silencing complex (RISC), to guide AGO in targeting viral RNAs. Secondary siRNAs amplified by RNA-directed RNA polimerase (RDR) 6/DCL4 spread to neighboring cells. The transport of original dsRNA is not clear. (D) siRNAs (or dsRNAs) can be transported through phloem to induce systemic silencing in upper non-sprayed leaves.

dsRNA-based vaccination has been successfully applied against RNA viruses such as pepper mild mottle virus, tobacco etch virus, alfalfa mosaic virus, CMV, TMV, and zucchini yellow mosaic virus. dsRNAs can be easily applied to plants by spraying (Fig. 2). Recognition of viral dsRNA by DCLs in av-RNAi produces vsiRNAs to direct virus-specific resistance. In addition, viral dsRNA is being considered as a broadly conserved pathogen-associated molecular pattern (PAMP) by DCLs. PAMP involves the activation of

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transcriptional signaling cascades (see vasiRNAs above and other endogenous secondary siRNAs) and could be used to confer broad-spectrum, non-specific pathogen resistance. However, central to this application is the assumption that exogenous dsRNA does not show any non-specific side effects relating to mismatched off-target hybridization, protein binding to nucleic acids or activation of non-controlled cascades. A direct application of siRNAs ensures more specificity; the experience to date is positive and promising. Research, however, is currently focusing on aspects of stability, safety, efficiency and costs, which taken together will influence the diffusion of RNA drugs in agriculture. Virus-induced transgenerational memory. Plant viruses can induce transient, stable, and heritable changes in gene expression without changes in genome sequences. For instance, plant–virus systems have led to the discovery and investigation of many aspects of plant epigenetics. av-RNAi can, indeed, involve a transgenerationally inherited epigenetic modification. Thus, av-RNAi infection may induce heritable phenotypic variation that influences plant fitness in response to recurring pathogen invasions. For application purposes, its epigenetic mechanisms are far from being completely understood. However, science is not far from developing specific viral vectors that can be used to infect progenitors, conferring an inherited phenotype to the next generation. This would open the window to non-transgenic, virus-free engineered seeds. vsiRNAs for viral metagenomics: from diagnostics to environmental studies. Given the RNAi-based plant antiviral and viral counterdefense strategies, av-RNAi often appears as a mechanism of surveillance ensuring the co-existence of both partners and the absence of symptoms. Indeed, thanks to the support of far-reaching NGS technology, we can detect new vsiRNAs as a result of av-RNAi even in non-symptomatic plants. Viruses are incisively abundant on Earth and recognized as drivers of global processes. They also represent vast genetic and biological diversity on the planet. Viruses are obligate entities that infect the cells of living organisms, co-evolve with them and migrate from one host species to another. In plants, viruses or subviral entities can cause diseases that have serious bio-economical impacts, leading to the adoption of measures of prevention or resistance. However, in most cases, plant viruses are silent or asymptomatic in their hosts and continue to be regulators of plant gene expression, often through RNAi. Recent findings, for example, reinforce the notion that viruses are beneficial in complex ecosystems when used for specific purposes and when they do not cause diseases in agro-ecosystems. Indeed, plant viruses have been used to enhance the beauty of ornamental plants. They confer resilience under conditions of drought or cold temperatures. Genome-integrated viral forms can confer cross-protection against severe strains, support evolution by intra- or inter-specific horizontal gene transfer (plant-to-plant through virus), and can raise the fitness level of a crop in terms of seed production by attracting pollinators. The great potential that plant viruses have in the agriculture and forestry sectors requires: (1) a greater focus on our biased notions of viruses as pathogens; (2) a deeper understanding of the mechanisms of how plant viruses regulate gene expression; and (3) closing the gap and providing insights into the composition and structure of environmental viral communities. av-RNAi includes steps of host-dependent amplification vsiRNAs, which in turn are molecular markers of viral infections. This unique feature allowed the indentification of otherwise undetectable novel infectious entities in plants, present at a very low level of replication, using homology-independent approaches.

Further Reading Baulcombe, D.C., Dean, C., 2014. Epigenetic regulation in plant responses to the environment. Cold Spring Harbor Perspectives in Biology 6, a019471. Guo, Z., Li, Y., Ding, S.W., 2019. Small RNA-based antimicrobial immunity. Nature Reviews Immunology 19 (1), 31–44. Maillard, P.V., Ciaudo, C., Marchais, A., et al., 2013. Antiviral RNA interference in mammalian cells. Science 342, 235–238. Martinez de Alba, A.E., Elvira-Matelot, E., Vaucheret, H., 2013. Gene silencing in plants: A diversity of pathways. Biochimica et Biophysica Acta 1829, 1300–1308. Mitter, N., Worrall, E.A., Robinson, K.E., Xu, Z.P., Carroll, B.J., 2017. Induction of virus resistance by exogenous application of double-stranded RNA. Current Opinion in Virology. 49–55. doi:10.1016/j.coviro.2017.07.009. Roossinck, M.J., 2015. A new look at plant viruses and their potential beneficial roles in crops. Molecular Plant Pathology 16, 331–333. Rosa, C., Kuo, Y.W., Wuriyanghan, H., Falk, B.W., 2018. RNA interference mechanisms and applications in plant pathology. Annual Review of Phytopathology 56, 581–610. Schuck, J., Gursinsky, T., Pantaleo, V., Burgyan, J., Behrens, S.E., 2013. AGO/RISC-mediated antiviral RNA silencing in a plant in vitro system. Nucleic Acids Research 41, 5090–5103.

Relevant Websites http://www.mirbase.org miRBase.

Plant Resistance to Viruses: Engineered Resistance Marc Fuchs, Cornell University, Geneva, NY, United States r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

mRNA Messenger RNA nt Nucleotide PTGS Post-transcriptional gene silencing RdRp RNA-dependent RNA polymerase RISC RNA-induced silencing complex ssRNA Single-stranded RNA RNA Ribonucleic acid RNAi RNA interference siRNA Small interfering RNA ta-siRNA Trans-acting siRNA VSR Viral suppressor of RNA silencing

AGO Argonaute amiRNA Artificial microRNA CP Coat protein or capsid protein CRISPR Clustered regularly interspaced short palindromic repeats CAS CRISPR-associated proteins DCL Dicer-like proteins with a dsRNA-specific endoribonuclease activity DNA Deoxyribonucleic acid dsRNA Double-stranded RNA miRNA MicroRNAs

Glossary Argonaute is a family of proteins that bind small noncoding RNAs, including miRNAs and siRNAs, and are guided to RNA targets through sequence complementarity, leading to the cleavage or translation inhibition of mRNAs, including viral RNAs. CRISPR-Cas Clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated proteins (Cas) is a prokaryotic adaptive immune system that can be leveraged for genetic material to be added, removed, or altered at particular locations in the plant genome, or to directly target viral DNA or RNA molecules. MicroRNAs are a class of small non-coding B21–24 nt RNAs that are involved in the regulation of gene expression at the post-transcriptional level by degrading target

mRNAs, including viral RNAs, and/or inhibiting their translation. RNA interference is a biological process in which RNA molecules inhibit gene expression or translation by neutralizing target mRNA molecules, including viral RNAs. Small interfering RNA is a class of 20–25 nt non-coding dsRNA molecules. The RNA-induced silencing complex is a ribonucleoprotein complex that incorporates one strand of a miRNA or a siRNA duplex for RNAi. Trans-acting siRNA are siRNA that repress plant gene expression through post-transcriptional gene silencing. Transgenic plant is a plant with a genome modified by genetic engineering techniques to insert, remove or knockout genetic elements.

Introduction Plant viruses can cause severe damage to crops by substantially reducing vigor, yield and product quality. A number of approaches are strategically implemented to mitigate the impacts of plant viruses. For example, quarantine measures and certification of seeds and propagative planting units are used to limit the introduction of virus diseases in production fields. Cultural practices, rogueing, control of vectors and cross-protection based on mild virus strains or benign satellite RNA are selected to reduce secondary spread of viruses in areas where epidemics can be problematic. However, the ideal and most effective approach to control viruses relies on the use of resistant crop cultivars. Host resistance genes have been extensively exploited by traditional breeding techniques although only a limited number of commercial crop cultivars exhibit practical resistance to viruses. The advent of biotechnology through plant transformation, the application of the concept of pathogen-derived resistance, and the exploitation of the antiviral pathways of RNAi have opened new avenues for the development of virus-resistant crop cultivars, in particular when resistant material with desired horticultural characteristics has not been developed through conventional breeding or when no host resistance sources are known. RNAi is a conserved and RNA-dependent gene silencing process that is induced by dsRNA expression to target the cleavage of viral RNA molecules and confer resistance to virus infection via PTGS. In addition to the conventional expression of RNAi in transgenic plants to achieve virus resistance, RNAi can be triggered by the exogenous application of in vitro-produced dsRNA derived from viral sequences. More recently, genome editing via CRISPR/Cas has proven valuable to disrupt host susceptibility genes for the improvement of crop cultivars or directly target viral DNA or RNA for cleavage. The past three decades have witnessed the development of numerous virus-resistant transgenic plants. Some crop cultivars engineered for virus resistance have been successfully tested in the field but only a few have been commercialized, including summer

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squash and papaya which have been adopted for more than 20 years in the United States. Crop plants engineered for virus resistance offer numerous benefits to agriculture and cause extraordinarily limited, if any, hazard to the environment and human health.

A Historical Perspective on Engineered Resistance Against Plant Viruses Genetic engineering expanded the scope of innovative approaches for virus control in crops by providing new tools to achieve resistance in otherwise susceptible plants. The first approach to confer resistance to viruses in plants resulted in the mid-1980s from the development of efficient protocols for plant transformation and the application of the concept of pathogen-derived resistance. This concept was initially conceived to achieve resistance by transferring and expressing a dysfunctional pathogen-specific molecule into the host genome to inhibit the pathogen. As a potential application of the concept, a segment of the virus’ own genetic material was hypothesized to somehow protect a plant against virus infection by acting against the virus itself. The first demonstration of pathogen-derived resistance against a plant virus was with Tobacco mosaic virus (TMV) in 1986. This seminal work revealed that transgenic tobacco plants expressing the TMV CP gene showed a substantial delay in symptom development or did not display symptoms following inoculation with TMV particles. This type of resistance was known as CP-mediated resistance. It was successfully applied against numerous plant viruses. In the early 1990s, RdRp, and then movement protein, protease, satellite RNA, defective interfering RNAs, or non-coding viral sequences were described to confer virus resistance in plants. Virus-derived transgene constructs included full-length, untranslated, and truncated coding and non-coding fragments, in sense or anti-sense orientation. It became soon apparent that more or less any viral sequence could provide some level of resistance to virus infection in transgenic plants. Remarkably, transcription of the virusderived transgene rather than expression of a protein product was shown to be required for resistance. This pioneering work progressively led to the discovery of RNAi as a key antiviral defense of plants against viruses. Summer squash was the first virus resistant RNAi crop to be commercially released in 1996. Resistance to multiple viruses was achieved in summer squash cultivars by pyramiding several RNAi constructs derived from individual viruses. Antiviral factors other than sequences derived from viral genomes have also been used to achieve resistance to viruses in plants. Production of antibodies, 20 -50 oligoadenylate synthase, ribosome inactivating proteins, dsRNA-specific RNases, dsRNA-dependent protein kinases, cysteine protease inhibitors, nucleoprotein-binding interfering aptamer peptides, and pathogenesis-related proteins has been reported to protect plants against virus infection. Over the past three decades, resistance was engineered against numerous viruses with diverse taxonomic affiliations and various modes of transmission by insect vectors in many plant species and crop plants, essentially via RNAi. Further refinements of RNAi for virus resistance consisted of engineering amiRNAs to target a virus sequence and protect plants from virus infection. More recently, genome editing via CRISPR/Cas has proven valuable for engineered virus resistance by modifying host recessive resistance genes for crop improvement or directly degrading viral DNA or RNA.

Molecular Underpinnings of Engineered Resistant to Plant Viruses Several mechanisms underlying engineered virus resistance have been described. Early studies suggested that expression of a transgene protein product seemed to be required for resistance. For example, resistance to TMV in tobacco plants was initially related to the expression level of the viral CP. Interference with an early step in the virus infection cycle, i.e., disassembly of TMV particles, was hypothesized to explain CP-mediated resistance, maybe by inhibition of viral uncoating or reduction of proteinprotein interactions between transgene-expressed CP and challenge virus. Resistance could be overcome by high doses of inoculum and was not very effective against virus RNA inoculation. Next, resistance was shown to be RNA- rather than protein-mediated because untranslatable virus CP transgenes protected plants against virus infection. In addition, resistance was correlated to actively transcribed transgenes but low steady-state levels of transgene mRNA, indicating a sequence-specific post-transcriptional RNA-degradation system. RNA degradation was shown to target transgene transcripts and viral RNA with high nucleotide sequence identity. This was the first indication of the occurrence of PTGS as a manifestation of RNAi, a mechanism that inhibits gene expression in a sequence specific manner. RNAi is a conserved eukaryotic mechanism that is critical for host defense against viruses and has become an approach of choice to engineer resistance to viruses in transgenic crops. A conserved feature of RNAi in plants is the processing of dsRNA into siRNA by DCL enzymes. Briefly, RNAi is initiated by dsRNA molecules and controlled by RISC. Long dsRNA molecules, for instance dsRNA corresponding to viral sequences, are cleaved into short dsRNA fragments of 20–25 nt siRNAs duplexes. The passenger strand of the siRNA duplexes is degraded and the guide strand is incorporated into RISC by DCL endoribonuclease proteins. Hybridization of the guide siRNA strand with a complementary RNA sequence such as a viral RNA molecule induces cleavage by AGO2, the catalytic component of RISC. This nucleotide sequence-based cleavage results in the degradation of the viral RNA target, hence resistance. Duplexes of siRNAs result from dsRNA derived from RNA virus replication, synthesis by plant RdRp, transcripts and hairpin-loop structures of ssRNAs. Plants appear to have a set of four DCL proteins to process and generate siRNAs. DCL1 mainly processes imperfectly basepaired fold-back precursors to produce miRNAs. DCL2 generates 22 nt siRNAs and can produce secondary viral siRNAs. DCL3 generates 24 nt siRNAs that are associated with silencing of transposons and repetitive elements, and are involved in defense against DNA viruses. DCL4 generates 21 nt siRNAs that have major roles in antiviral defense against RNA viruses, and is involved in the generation of ta-siRNAs that regulate gene expression.

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As an alternative to RNAi transgenic plants. exogenous application of in vitro-produced dsRNA to initiate RNAi via sprays or use of carrier compounds, including layered double hydroxide clay nanosheets, among other means of deliver, confers resistance to virus infection. This strategy was even shown to inhibit insect-mediated virus transmission. To counter the RNAi plant defense, viruses have evolved RNAi-suppressor proteins called VSR. VSRs interfere at different steps of the antiviral RNAi pathways by sequestering siRNA, protecting dsRNA from degradation or interacting directly with host enzymes involved in RNAi. As a result of VSR expression, RNAi-mediated resistance is impaired. RNAi can sometimes be unpredictable with regard to the degree of virus resistance. Different factors may be involved in deceiving levels of resistance or even loss of resistance. Nucleotide sequence divergence between a dsRNA transgene construct and cognate sequence in a virus population, the emergence of new virus strains in terms of the evolutionary dynamic of virus populations, VSRs of cognate or heterologous viruses, environmental conditions with fluctuating growing temperature during a cropping season can all affect the efficacy of RNAi at protecting plants from virus infection. Another class of non-coding regulatory RNAs, miRNAs, has been exploited to engineer virus resistance in plants. Expression of modified miRNA precursors for which intrinsic plant specific nucleotides are replaced with virus-specific nucleotides results in the production of amiRNAs that target viral RNA sequences, resulting in resistance via RNAi. Although amiRNAs have been used for engineering resistance to a wide range of plant viruses, this strategy awaits field evaluation. Engineered plant virus resistance can also be achieved by using CRISP-Cas, a prokaryotic adaptive immune system which can be reprogrammed into a gene targeting technology. An RNA-guided Cas nuclease protein cleaves the substrate viral DNA or RNA at specific target sites, leading to their degradation. The specificity of the cleavage is governed by base complementarity between the CRISPR RNA and the target DNA or RNA molecule. A number of Cas proteins with sequencespecific nuclease activity have been identified to target DNA, RNA or both. Different CRISPR-Cas platforms have been successfully leveraged to engineer resistance to DNA viruses or RNA viruses in planta. CRISPR-Cas has also been used to knockout the host recessive eIF4E gene in cucumber and confer broad virus resistance. It will be interesting to see how this promising technology performs under field conditions.

Field Resistance and Commercial Adoption Summer squash line ZW-20 expressing the CP gene of Zucchini yellow mosaic virus (ZYMV) and Watermelon mosaic virus (WMV), and summer squash line CZW-3 expressing the CP gene of ZYMV, WMV and Cucumber mosaic virus (CMV) were the first disease resistant transgenic plant events commercially released. Summer squash CZW-3 plants are highly resistant to mechanical inoculation of viruses in the greenhouse (Fig. 1(A)) or to aphid-mediated virus transmission in the field (Fig. 1(D)). Summer squash ZW-20 plants are highly resistant to mechanical inoculation in the field (Fig. 1(C)) or aphid-mediated virus transmission under field conditions in which no insecticides were applied to control aphid vector populations (Fig. 1(F)). Transgenic summer squash plants produce fruits of marketable quality (Fig. 1(E) and (H)) whereas most fruits of control plants are malformed and discolored (Fig. 1(E) and (G)). Numerous transgenic crop plants such as cereal, vegetable, fruit, legume, flower, and forage crops, among others, that express RNAi constructs, virus-derived gene constructs or other antiviral factors have been developed and tested under field conditions (Table 1). A few transgenic crops engineered for virus resistance have been commercially released in the United States and the People’s Republic of China (Table 1). In the United States, summer squash lines ZW-20 and CZW-3 have been commercialized in 1996. Several squash types and cultivars have been developed by crosses and back crosses with the two initially deregulated lines. Papaya expressing the CP gene of Papaya ringspot virus (PRSV) has been deregulated and commercialized in Hawaii in 1998. PRSV-resistant papaya was the first transgenic fruit crop to be commercially released in the United States. PRSV-resistant transgenic papaya cultivars are also commercialized in the People’s Republic of China. Several potato expressing the RdRp gene of Potato leafroll virus or the CP gene of Potato virus Y were deregulated in 1998 and 2000. However, soon after their release, these potato lines were withdrawn from the market.

Benefits of Virus-Resistant Transgenic Plants to Agriculture Virus-resistant transgenic plants offer numerous benefits to agriculture, particularly in cases where no genetic source of resistance has been identified or host resistance is challenging to transfer into elite cultivars by traditional breeding approaches due to genetic incompatibility or linkage to undesired traits. In such cases, engineered resistance may be the only viable option to develop virusresistant cultivars. Engineered resistance may also be the only approach to develop cultivars with multiple sources of resistance by pyramiding several RNAi constructs. Benefits of virus-resistant transgenic plants are of economic importance because yields are increased and quality of crop products improved. For example, the adoption of PRSV-resistant transgenic papaya cultivars resulted in a similar production level than before PRSV became epidemic in Hawaii, providing a steady source of income to growers. Benefits are also of epidemiological importance because virus-resistant transgenic plants limit virus infection rates by restricting challenge viruses, reducing their titers, or inhibiting their replication and/or cell-to-cell or systemic movement. Therefore, lower virus levels reduce the frequency of acquisition by vectors and subsequent transmission within and between field sites. Consequently, virus epidemics are substantially limited. In addition, benefits of virus-resistant transgenic plants are of environmental importance because chemicals directed to

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Fig. 1 Reaction of transgenic summer squash ZW-20 expressing the CP gene of Zucchini yellow mosaic virus (ZYMV) and Watermelon mosaic virus (WMV), and of transgenic summer squash CZW-3 expressing the CP gene of ZYMV, WMV and Cucumber mosaic virus (CMV) to virus infection. (A) Resistance of transgenic CWZ-3 to infection by CMV, ZYMV and WMV (upper right), and susceptibility of conventional plants to single infection by CMV (lower right), ZYMV (upper left) and WMV (lower left); (B) Fields of transgenic ZW-20 (foreground) and conventional summer squash plants (background) surrounded by a border row of mechanically-inoculated non-transgenic plants; (C) Transgenic ZW-20 (right) and conventional (left) plants subjected to mechanical inoculation of ZYMV and WMV; (D) Transgenic CZW-3 (right) and conventional (left) plants mechanically inoculated with ZYMV, WMV and CMV; (E) Close-up of a fruit of transgenic ZW-20 (extreme left) and fruits of conventional summer squash plants; (F) Transgenic CZW-3 (middle and right rows) and conventional (left row) summer squash subjected to aphid-mediated inoculation by ZYMV, WMV and CMV; (G) Fruit production of conventional summer squash under conditions of infection by ZYMV and WMV; and, (H) Fruit production of transgenic summer squash ZW-20 under conditions of infection by ZYMV and WMV.

control virus vector populations are reduced or not necessary. Restricting the reliance on chemicals directed to arthropod, fungal and nematode vectors of plant viruses is important for sustainable agriculture. Finally, benefits of virus-resistant transgenic plants are of social importance. In Hawaii, growing papaya was not viable anymore prior to the adoption of PRSV-resistant transgenic papaya despite huge efforts to eradicate infected trees in order to limit the propagation of the virus. The impact of PRSV was so severe that some growers abandoned their farms and had to find new jobs to secure an income. PRSV-resistant transgenic papaya saved the papaya industry and strengthened the social welfare of local communities in Hawaii.

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Table 1

Examples of virus-resistant transgenic crops that have been tested in the field or commercially released. This list is not exhaustive

Crop category Crop common name

Crop scientific name

Resistance to virus

Genus

Family

Cereals Barley

Hordeum vulgare

Barley yellow dwarf virus Wheat dwarf virus Barley stripe mosaic virus Turnip mosaic virus Maize dwarf mosaic virus Maize chlorotic dwarf virus Maize chlorotic mottle virus Sugarcane mosaic virus Barley yellow dwarf virus Rice stripe virus Rice yellow mottle virus Rice hoja blanca virus Rice black-streaked dwarf virus Rice dwarf virus Rice gall dwarf virus Rice black streaked dwarf virus Rice grassy stunt virus Rice ragged stunt virus Rice tungro bacilliform virus Barley yellow dwarf virus Wheat streak mosaic virus Wheat yellow mosaic virus Triticum mosaic virus Barley stripe mosaic virus

Luteovirus Mastrevirus Hordeivirus Potyvirus Potyvirus Waikavirus Sobemovirus Potyvirus Luteovirus Tenuivirus Sobemovirus Tenuivirus Fjivirus Phytoreovirus Phytoreovirus Fijivirus Tenuivirus Oryzavirus Tungrovirus Luteovirus Tritimovirus Bymovirus Poacevirus Hordeivirus

Luteoviridae Geminiviridae Virgaviridae Potyviridae Potyviridae Secoviridae Solemoviridae Potyviridae Luteoviridae Phenuiviridae Solemoviridae Phenuiviridae Reoviridae Reoviridae Reoviridae Reoviridae Phenuiviridae Reoviridae Caulimoviridae Luteoviridae Potyviridae Potyviridae Potyviridae Virgaviridae

Cucumber mosaic virus Papaya ringspot virus Squash mosaic virus Watermelon mosaic virus Zucchini yellow mosaic virus Cucumber vein yellowing virus Lettuce mosaic virus Lettuce necrotic yellows virus Mirafiori lettuce big-vein virus Cucumber mosaic virus Tobacco etch virus Tomato mosaic virus Potato virus Y Pepper mild yellow mottle virus Cucumber mosaic virusa Papaya ringspot virus Squash mosaic virus Watermelon mosaic virusa Zucchini yellow mosaic virusa Beet necrotic yellow vein virus Beet western yellows virus Beet curly top virus Cucumber mosaic virus Tobacco mosaic virus Tomato mosaic virus Impatiens necrotic spot virus Tomato spotted wilt virus Ageratum yellow vein Malaysia virus Tomato yellow leaf curl virus

Cucumovirus Potyvirus Comovirus Potyvirus Potyvirus Ipomovirus Potyvirus Tenuivirus Ophiovirus Cucumovirus Potyvirus Tobamovirus Potyvirus Tobamovirus Cucumovirus Potyvirus Comovirus Potyvirus Potyvirus Benyvirus Polerovirus Curtovirus Cucumovirus Tobamovirus Tobamovirus Orthotospovirus Orthotospovirus Begomovirus Begomovirus

Bromoviridae Potyviridae Secoviridae Potyviridae Potyviridae Potyviridae Potyviridae Phenuiviridae Aspiviridae Bromoviridae Potyviridae Virgaviridae Potyviridae Virgaviridae Bromoviridae Potyviridae Secoviridae Potyviridae Potyviridae Benyviridae Luteoviridae Geminiviridae Bromoviridae Virgaviridae Virgaviridae Tospoviridae Tospoviridae Geminiviridae Geminiviridae

Citrus tristeza virus Banana bunchy top virus Citrus tristeza virus

Closterovirus Babuvirus Closterovirus

Closteroviridae Nanoviridae Closteroviridae

Canola Maize

Brassica napus Zea mays

Oat Rice

Avena sativa Oryza sativa

Wheat

Triticum aestivum

Vegetables Cucumber

Cucumis sativus

Lettuce

Lactuca sativa

Pepper

Capsicum

Squasha

Cucurbita pepo

Sugar beet

Beta vulgaris

Tomato

Solanum lycopersicum

Fruits Alemow Banana Grapefruit

Citrus macrophylla Musa sp. Citrus paradisi

Plant Resistance to Viruses: Engineered Resistance

Table 1

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Continued

Crop category Crop common name

Crop scientific name

Resistance to virus

Genus

Family

Grapevine Melon

Vitis sp. Cucumis melo

Mexican lime

Citrus aurantifolia

Passion fruit

Passiflora edulis

Papayaa,b Pineapple Plum Raspberry

Carica papaya Ananas comosus Prunus domestica Rubus idaeus

Strawberry Sweet orange

Fragaria sp. Citrus sinensis

Tamarillo Walnut Watermelon

Cyphomandra betacea Juglans regia Citrullus lanatus

Grapevine fanleaf virus Cucumber mosaic virus Papaya ringspot virus Squash mosaic virus Watermelon mosaic virus Zucchini yellow mosaic virus Cucurbit yellow stunt disorder virus Watermelon silver mottle virus Citrus tristeza virus Citrus psorosis virus Cowpea aphid borne mosaic virus Passion fruit woodiness virus Papaya ringspot virusa,b Pineapple wilt-associated virus Plum pox virus Raspberry bushy dwarf Tomato ringspot virus Strawberry mild yellow edge virus Citrus tristeza virus Citrus psorosis virus Tamarillo mosaic virus Cherry leafroll virus Cucumber mosaic virus Cucumber green mottle mosaic virus Watermelon mosaic virus Zucchini yellow mosaic virus Papaya ringspot virus

Nepovirus Cucumovirus Potyvirus Comovirus Potyvirus Potyvirus Crinivirus Orthotospovirus Closterovirus Ophiovirus Potyvirus Potyvirus Potyvirus Ampelovirus Potyvirus Idaeovirus Nepovirus Potexvirus Closterovirus Ophiovirus Potyvirus Nepovirus Cucumovirus Tobamovirus Potyvirus Potyvirus Potyvirus

Secoviridae Bromoviridae Potyviridae Secoviridae Potyviridae Potyviridae Closteroviridae Tospoviridae Closteroviridae Aspiviridae Potyviridae Potyviridae Potyviridae Closteroviridae Potyviridae Unassigned Secoviridae Alphaflexiviridae Closteroviridae Aspiviridae Potyviridae Secoviridae Bromoviridae Virgaviridae Potyviridae Potyviridae Potyviridae

Bean golden mosaic virus Alfalfa mosaic virus White clover mosaic virus Mungbean yellow mosaic India virus Cowpea severe mosaic virus Cowpea aphid-borne mosaic virus Peanut clump virus Groundnut rosette virus Alfalfa mosaic virus Bean leafroll virus Bean yellow mosaic virus Pea enation mosaic virus Pea seed-borne mosaic virus Pea streak virus Tomato spotted wilt virus Groundnut rosette assistor virus Peanut stripe virus Tobacco streak virus Alfalfa mosaic virus Soybean mosaic virus Bean pod mottle virus Southern bean mosaic virus Soybean dwarf virus Bean yellow mosaic virus

Begomovirus Alfamovirus Potexvirus Begomovirus Comovirus Potyvirus Pecluvirus Umbravirus Alfamovirus Luteovirus Potyvirus Umbravirus Potyvirus Carlavirus Orthotospovirus unassigned Potyvirus Ilarvirus Alfamovirus Potyvirus Comovirus Sobemovirus Luteovirus Potyvirus

Geminiviridae Bromoviridae Alphaflexiviridae Geminiviridae Secoviridae Potyviridae Virgaviridae Tombusviridae Bromoviridae Luteoviridae Potyviridae Tombusviridae Potyviridae Betaflexiviridae Tospoviridae Luteoviridae Potyviridae Bromoviridae Bromoviridae Potyviridae Secoviridae Solemoviridae Luteoviridae Potyviridae

Tomato spotted wilt virus Chysanthemum virus B Cymbidium mosaic virus Bean yellow mosaic virus Impatiens necrotic spot virus Poinsettia mosaic virus

Orthotospovirus Carlavirus Potexvirus Potyvirus Orthotospovirus Tymovirus

Tospoviridae Betaflexiviridae Alphaflexiviridae Potyviridae Tospoviridae Tymoviridae

Legumes Bean Clover

Phaseolus vulgaris Trifolium repens

Cowpea

Vigna unguiculata

Groundnut

Arachis hypogaea

Pea

Pisum sativum

Peanut

Arachis hypogaea

Soybean

Glycine max

Flowers Chrysanthemum Dendrobium Gladiolus Impatiens Poinsettia

Chrysanthemum indicum Encyclia cochleata Gladiolus sp. Impatiens walleriana Euphorbia pulcherrima

(Continued )

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Table 1

Continued

Crop category Crop common name

Crop scientific name

Resistance to virus

Genus

Family

Forage Alfalfa

Medicago sativa

Alfalfa mosaic virus

Alfamovirus

Bromoviridae

Grass Sugarcane

Saccharum sp.

Sugarcane mosaic virus Sugarcane yellow leaf virus Sorghum mosaic virus

Potyvirus Polerovirus Potyvirus

Potyviridae Luteoviridae Potyviridae

Tuberous root Cassava

Manihot esculenta

African cassava mosaic virus East African cassava mosaic virus Cassava brown streak virus Sri Lankan cassava mosaic virus Uganda cassava brown streak virus Potato virus A Potato virus X Potato virus Y Potato virus S Potato leafroll virus Tobacco rattle virus Tobacco vein mottling virus Sweet potato feathery mottle virus Sweet potato chlorotic stunt virus Sweet potato virus G Sweet potato mild mottle virus

Begomovirus Begomovirus Ipomovirus Begomovirus Ipomovirus Potyvirus Potexvirus Potyvirus Carlavirus Polerovirus Tobravirus Potyvirus Potyvirus Crinivirus Potyvirus Potyvirus

Geminiviridae Geminiviridae Potyviridae Geminiviridae Potyviridae Potyviridae Alphaflexiviridae Potyviridae Betaflexiviridae Luteoviridae Virgaviridae Potyviridae Potyviridae Closteroviridae Potyviridae Potyviridae

Cotton leaf curl Kokhran virus Cotton leaf curl Multan virus Cotton leaf curl Multan betasatellite

Begomovirus Begomovirus Betasatellite

Geminiviridae Geminiviridae Telocusatellidae

Potato

Solanum tuberosum

Sweet potato

Ipomea batatas

Fiber Cotton

Gossypium hirsutum

a

Commercially released in the USA. Commercially released in the People’s Republic of China.

b

Environmental and Human Health Safety Issues The insertion and expression of virus-derived genes or gene fragments in plants has raised concerns on their environmental and human health safety. Potential risks relate primarily to the occurrence and outcomes of heterologous encapsidation, recombination, gene flow, toxicity and allerginicity. It is important to keep in perspective that these three phenomena did not arise with transgenic plants expressing viral genes or gene fragments; they occur in conventional plants too. Therefore, it is critical to determine if they occur in transgenic plants beyond baseline events in conventional plants. Heterologous encapsidation refers to the encapsidation of the genome of a challenge virus by CP subunits expressed in a transgenic plant containing a viral CP gene. Since the CP carries determinants for vector specificity, among other features, the properties of field viruses may change. Therefore, it is conceivable that an otherwise vector non-transmissible virus could become transmissible through heterologous encapsidation and infect otherwise non-host plants. Extensive work has shown that heterologous encapsidation in transgenic plants expressing virus CP genes should be considered negligible in regard to adverse environmental effects, given this phenomenon is known to occur in conventional plants subjected to mixed virus infection. It should also be kept in perspective that any change in vector specific is a single generation rather than a permanent event because they are not perpetuated in the virus genome progeny. Recombination refers to template switching between viral transgene transcripts and the genome of a challenge virus. Resulting recombinant viruses may have a chimeric genome consisting of a segment from the challenge viral genome and another segment from viral transgene transcripts. It is argued that recombinant viruses may have identical biological properties as their parental lineages or new biological properties such as changes in vector specificity, expanded host range or increased pathogenicity. Since recombination alters the genome of challenge viruses, new properties of chimeric viruses will be stably transmitted to and perpetuated within the virus progeny. Comprehensive laboratory studies documented the occurrence of recombination in transgenic plants expressing viral genes. However, extensive field studies revealed a limited significance of recombination in transgenic plants expressing viral genes in regard to adverse environmental effects. This is because the stringency of selective pressure applied to the challenge virus is a critical factor in the recovery of recombinant viruses. Conditions of high selective pressure enhance the creation of recombinant viruses. In contrast, limited, if any, recombinant viruses are found to detectable level

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under conditions of low or no selective pressure. The latter conditions prevail under field conditions in which plants are infected by functional, not defective, viruses. Gene flow refers to the pollen-driven movement of transgenes from a virus-resistant transgenic plant into a non-transgenic compatible recipient plant, for example, a free-living relative. Hybrids resulting from gene flow can acquire and express virus-derived transgenes, and become resistant to the corresponding viruses. Subsequently, plants acquiring viral resistance traits can have a competitive advantage, exhibit increased fitness and eventually become more invasive. Movement of viral transgene constructs through pollen flow has been documented from a few virus-resistant transgenic crops, including summer squash, into free-living relatives under experimental field conditions. Hybrids between transgenic and free-living plants exhibited increased fitness under conditions of intense disease pressure. In contrast, under conditions of low disease pressure, no difference was observed between hybrids and free-living plant in terms of growth and reproductive potential. It is important to keep in perspective that gene flow with virus-resistant transgenic plants should not be perceived riskier than the equivalent situation with virus-resistant conventional plants. Safety assessment studies on human health have documented no evidence of toxicity and allergenicity in crop cultivars engineered for virus resistance or food derived thereof. To fully grasp the significance of these findings, it is critical to recognize that plant virus proteins or sequences are not known to code for toxins, create new allergens or change the expression of endogenous proteins to produce allergens. In addition, foods derived from virus-resistant transgenic summer squash and papaya have been consumed by millions of people for more than 20 years, with no reported ill effects or legal cases related to human health. So far, there is no compelling evidence to indicate that transgenic plants expressing virus genes or gene fragments increase the frequency of heterologous encapsidation, recombination, or food toxicity and allergenicity beyond background events. Similarly, there is little evidence, if any, to infer that transgenic plants expressing virus genes or gene fragments alter the properties of existing virus populations or create new viruses that could not arise naturally in conventional plants subjected to multiple virus infection. Finally, consequences of gene flow should be assessed on a case-by-case approach to make sound decisions on the safe release of crop cultivars engineered for virus resistance.

Conclusions Virus diseases remain a huge problem for a sustainable and efficient agriculture. Engineered resistance via RNAi and genome editing has expanded the scope of innovative approaches for the management of virus diseases by providing new tools to develop resistant crop cultivars and increasing opportunities to implement effective and sustainable management strategies. The past three decades have witnessed an explosion in the development of virus-resistant transgenic plants. Lessons from field experiments and commercial release of virus-resistant summer squash, papaya, and potato have demonstrated that benefits outweigh by far risks to the environment and human health. Nonetheless, despite remarkable progress, less than a handful of virus-resistant transgenic crops have been commercially adopted. A timely release of new virus-resistant transgenic crop cultivars would be desirable because viruses continue to be devastating.

Further Reading Baulcombe, D., 2019. How virus resistance provided a mechanistic foundation for RNA silencing. The Plant Cell 31, 1395–1396. Dalakouras, A., Wassenegger, M., Dadami, E., et al., 2019. GMO-free RNAi: Exogenous application of RNA molecules in plants. Plant Physiology. doi:10.1104/pp.19.00570. Dong, O.X., Ronald, P.C., 2019. Genetic engineering for disease resistance in plants: Recent progress and future perspectives. Plant Physiology 180, 26–38. Fuchs, M., Gonsalves, D., 2007. Safety of virus-resistant transgenic plants two decades after their introduction: Lessons from realistic field risk assessment studies. Annual Review of Phytopathology 45, 173–202. Gaffar, F.Y., Koch, A., 2019. Catch me if you can! RNA silencing-based improvement of antiviral plant immunity. Viruses 11, 673. doi:10.3390/v11070673. Gottula, J., Fuchs, M., 2009. Toward a quarter century of pathogen-derived resistance and practical approaches to engineered virus resistance in crops. In: Loebenstein, G., Carr, J.P. (Eds.), Natural and Engineered Resistance to Plant Viruses. Advances in Virus Research 75. Elsevier, pp. 161–183. Kalinina, N.O., Khromow, A., Love, A.J., Taliansky, M.E., 2019. CRISPR applications in plant virology: Virus resistance and beyond. Phytopathology. doi:10.1094/PHYTO-07-19-0267-IA. Kalid, A., Zang, A., Yasir, M., Li, F., 2017. Small RNA-based genetic engineering for plant viral resistance: Application in crop protection. Frontiers in Microbiology 8, 43. Lindbo, J.A., Falk, B.W., 2017. The impact of “coat protein-mediated virus resistance” in applied plant pathology and basic research. Phytopathology 107, 624–634. National Academies of Sciences, Engineering, and Medicine, Engineering, and Medicine, 2016. Genetically engineered crops: Experiences and prospects. Washington, DC: The National Academies Press, doi:10.17226/23395. Powell Abel, P., Nelson, R.S., De, B., et al., 1986. Delay of disease development in transgenic plants that express the tobacco mosaic virus coat protein gene. Science 232, 738–743. Pooggin, M.M., 2017. RNAi-mediated resistance to viruses: A critical assessment of methodologies. Current Opinion in Virology 26, 28–35. Rosa, C., Kuo, Y.-W., Wuriyanghan, H., Falk, B.W., 2018. RNA interference mechanisms and applications in plant pathology. Annual Review of Phytopathology 56, 581–610. Schmitt-Keichinger, C., 2019. Manipulating cellular factors to combat viruses: A case study from the plant eukaryotic translation initiation factors eIF4. Frontiers in Microbiology 10. doi:10.3389/fmicb.2019.00017. Tricoli, D.M., Carney, K.J., Russell, P.F., et al., 1995. Field evaluation of transgenic squash containing single or multiple virus coat protein gene constructs for resistance to cucumber mosaic virus, watermelon mosaic virus 2, and zucchini yellow mosaic virus. Nature Biotechnology 13, 1458–1465.

Relevant Websites http://www.isaaa.org/ ISAAA.

Plant Resistance to Viruses: Natural Resistance Associated With Dominant Genes Mandy Ravensbergen and Richard Kormelink, Wageningen University and Research, Wageningen, The Netherlands r 2021 Elsevier Ltd. All rights reserved. This is an update of P. Moffett, D.F. Klessig, Plant Resistance to Viruses: Natural Resistance Associated with Dominant Genes, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00469-6.

Nomenclature

NLRC Nod-like receptor nt Nucleotide(s) PAD Phytoalexin deficient R Dominant resistance RDR RNA-dependent RNA polymerase RISC RNA-induced silencing complex RITS RNA-induced transcriptional silencing RNAi RNA interference TIR Toll and interleukin receptor VIGS Virus-induced gene silencing VSR Viral suppressor of RNA silencing ZAR Hopz-activated resistance

AGO Argonaute protein Avr Avirulence CC Coiled-coil structure DCL Dicer-like dsRNA Double-stranded RNA EDS Enhanced disease susceptibility HR Hypersensitive response LRR Leucine-rich repeat MP Movement protein mRNA Messenger RNA NB-ARC Nucleotide binding adaptor

Glossary Apoptosome An oligomerized form of animal R proteins, for example of APAF1 and CED4, leading to activation of apoptotic initiator caspases. Avirulence gene Any pathogen gene whose protein product induces a resistance response in a plant possessing a matching resistance gene, thereby rendering the pathogen unable to establish an infection. Gene-for-gene resistance A genetic paradigm whereby the outcome of a plant–pathogen interaction is determined by the genotype of both organisms. Plant disease resistance proteins confer resistance to pathogens possessing a corresponding avirulence gene. Hypersensitive response A type of programmed cell death induced by a resistant plant in response to pathogen infection. Cell death is normally contained to infected cells. Inflammasome An oligomerized form of animal R proteins, for example of NLRC4, leading to activation of inflammatory caspases.

NB-LRR protein Plant disease resistance proteins containing a central nucleotide-binding domain (NB) and C-terminal leucine-rich repeat (LRR) domain. Also known as NBS-LRR, NB-ARC-LRR or NLR proteins. Resistance gene Plant genes which confer resistance to specific pathogens. Resistance genes confer resistance to pathogens possessing a corresponding avirulence gene. Resistosome An oligomerized form of activated plant NB-LRRs, recently identified for ZAR1, bearing structural resemblance to animal apoptosomes and inflammasomes. RNA interference A broad antiviral defense system used by plants, animals, and fungi. It recognizes double-stranded RNA as a cue of viral presence and induces degradation or silencing of viral transcripts or genomes. Virulence gene Any pathogen gene whose function contributes to increased pathogen proliferation or disease symptoms on an infected plant.

Introduction Host plants possess polymorphic resistance to different strains or races of pathogens, which is controlled by a single gene. Disease resistance controlled by recessive genes is often a passive form of resistance wherein the pathogen is unable to utilize the host-cell machinery. Dominant resistance (R) genes in plants appear to actively recognize the presence of specific pathogens and initiate responses to counteract infection. Usually, this involves a type of programmed cell death termed hypersensitive response (HR). However, this resistance is dependent on the pathogen possessing a corresponding gene that matches the R gene. These pathogen avirulence (Avr) genes are also dominant over their cognate virulent (avr) alleles. Since the outcome of an attempted infection is dependent on the genotype of both host and pathogen, this phenomenon is known as gene-for-gene resistance. Gene-for-gene resistance was first described over 60 years ago, with the discovery of various R genes against specific races of flax rust, caused by the fungus Melampsora lini, among different flax genotypes. Ever since, gene-for-gene resistance has been observed for many plant pathogens including fungi, oomycetes, bacteria, insects, nematodes, and viruses.

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R Gene Products R genes have been genetically characterized for many years and have been incorporated into breeding programs of multiple crops. To study the structure and function of the R gene encoded proteins, numerous R genes have been cloned. These proteins belong to a small number of protein classes including transmembrane proteins possessing extracellular leucine-rich repeat (LRR) domains as well as a small number of intracellular serine/threonine kinases. The most abundant class of R genes encodes intracellular proteins containing a central NB-ARC (nucleotide binding adaptor shared by APAF-1, certain R gene products, and CED-4) domain as well as a C-terminal LRR domain. Additionally, these proteins contain an N-terminal signaling domain, either displaying homology to the Toll and interleukin receptors (TIR domain) of animals or containing a coiled-coil structure (CC). These proteins are often denoted as CC-NB-LRR or CNL, and TIR-NB-LRR or TNL, collectively known as NB-LRR or NLR. Here, CC-NB-LRR and TIR-NB-LRR, collectively NB-LRR, will be used. Sequencing projects have revealed that plant genomes contain large numbers of genes encoding NB-LRR proteins. For example, the Arabidopsis genome contains 149 such genes, whereas the cassava genome encodes 327, 99 of which are lacking an LRR domain. The polymorphic nature of host plant resistance is attributed to the polymorphic NB-LRR gene loci, with different alleles recognizing different pathogens or pathogen races. Furthermore, the proteins encoded are highly divergent both within and between species, presenting a large potential for recognition of pathogen diversity. R genes recognizing a specific pathogen could be identified by testing many genotypes of a certain species. For crop species, resistance sources are often identified in, and introgressed from, wild relatives. Molecular studies have shown that recognition specificity is in most cases determined by the LRR domain of NB-LRR proteins. In certain cases, this could be demonstrated through swapping of LRR domains between two similar NB-LRR proteins. Correspondingly, the LRR domain is the most polymorphic region of NB–LRR proteins. Bioinformatics studies have suggested that the LRR domains are under diversifying selection, suggesting that this class of R genes is under selective pressure. There do not appear to be any obvious features that distinguish between those R genes that recognize, for example, bacteria versus those that recognize viruses. Within Arabidopsis, for example, the HRT, RCY1 (Resistance to CMV (Y)), and RPP8 genes are alleles of the same locus, but confer resistance to Turnip crinkle virus (TCV), Cucumber mosaic virus (CMV), and the oomycete Hyaloperonospora parasitica, respectively. Similarly, in potato, gene-duplication events have led to highly similar gene paralogues, Rx1 and Gpa2, which confer resistance to Potato virus X (PVX) and to the nematode Globodera pallida, respectively. Additionally, PVX can be engineered to express a bacterial Avr gene and a plant expressing the corresponding NB-LRR gene will be resistant to the recombinant virus. Thus, most NB-LRR genes probably have the potential to evolve specificity to a variety of different pathogen types, allowing for maximum adaptability. As such, the mechanisms of recognition and initiation of disease-resistance responses are likely to be similar between R proteins that recognize viruses and R proteins that recognize other types of pathogens.

R Gene Responses to Viruses Many different types of pathogen proteins are either synthesized in the host cell by intracellular pathogens (viruses) or delivered into the cytoplasm by the various secretion systems of bacteria, fungi, or oomycetes. Here, they can be recognized by NB-LRR proteins. In absence of a corresponding R gene, Avr proteins generally act as virulence factors, protect the pathogen from host defense responses, actively inhibit host defenses, or are involved in replication of the pathogen. Pathogen avr alleles can overcome R-gene-mediated resistance either by mutation or by elimination of the appropriate Avr gene. However, this often leads to a loss of fitness. Given their small genomes, this is particularly acute for viruses as most viral genes are crucial for viral fitness and often have multiple functions. An overview of cloned NB-LRR genes of which the Avr determinant has been identified is presented in Table 1. Anti-viral NB-LRR genes that have not been cloned yet or have an unknown elicitor can be found in recent extensive reviews, for example by de Ronde et al. One of the most studied antiviral R genes is the N gene. It was introduced into commercial tobacco cultivars from Nicotiana glutinosa via interspecific crosses as a means of conferring resistance to tobamoviruses, primarily Tobacco mosaic virus (TMV). The N gene was among the first R genes to be cloned and encodes a typical TIR-NB-LRR protein. Like several TIR-NB-LRR-encoding genes, the N gene produces two different transcripts via alternative splicing, one encoding the full protein and a shorter version encoding a protein lacking the LRR domain. The role of these truncated proteins is unclear; plants transgenic for versions of the N gene that cannot produce both splice forms are compromised in their resistance to TMV. The N gene confers resistance against a wide range of tobamoviruses, with the exception of a virus originally designated TMV-ob and renamed Obuda pepper virus (ObPV). The Avr effector of the N gene resides in the TMV viral replicase protein, as demonstrated through gene swaps between TMV and ObPV. Expression of a helicase domain (p50) of the replicase is sufficient to induce HR in N plants. The HR induced by N in response to TMV is visible as small necrotic lesions at the site of infection. In this case, the virus is able to replicate to a limited degree, spreading to cells surrounding the initially infected cell before the onset of cell death. Initiation of HR requires only the presence of a matching pair of R and Avr proteins. In the past decades it has become evident that cell death caused by HR does not equate the resistance response. Follow-up studies indicated that the N gene based antiviral response leads to a translational arrest of viral transcripts by a process involving Argonaute 4 (AGO4), ultimately preventing virus accumulation and spread. Additionally, in cases of HR-induction with Rx1, this response can be knocked out without affecting Rx1-mediated resistance against PVX. Similar observations have been made by others and suggest that the actual resistance response is different from an HR, although often both are triggered. It follows that HR continues to serve as a valuable indication of NB-LRR mediated resistance.

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Table 1 Overview of cloned and characterized antiviral NB-LRR genes. Columns list the plant species in which the R gene has been introgressed (plant), the domain facilitating oligomerization, the viral Avr determinant and the resistance response R gene

Encoded protein

Plant

Homo-oligomerization Virus domain

Avr determinant

Resistance responsea

N Rx1 Rx2 Sw-5b HRT RCY-1 Tm2 Tm22

TIR-NB-LRR CC-NB-LRR CC-NB-LRR CC-NB-LRR CC-NB-LRR CC-NB-LRR CC-NB-LRR CC-NB-LRR

Tobacco Potato Potato Tomato Arabidopsis Arabidopsis Tomato Tomato

NB-ARC –b ND NB-ARC ND ND ND ND

Helicase/p50 subunit CP CP NSm CP CP 30 kDa MP 30 kDa MP

HR ER ER ER, HR HR HR ER ER

Tobamoviruses PVX PVX Tospoviruses (TSWV, GRSV, TCSV) TCV CMV Tobamoviruses Tobamoviruses

a

Certain resistance genes condition more than one type of response depending on viral isolate, genetic background, or environmental conditions. Rx1 was found to heterodimerize with RanGAP2, there is no evidence for homodimerization. Note: ND, not determined.

b

For some plant-virus interactions, HR is observed although a strict gene-for-gene relationship has not been defined due to a lack of variability in either the host or the pathogen. For example, eggplant undergoes a typical HR response when inoculated with tobamoviruses. A compatible interaction was not found either because the corresponding R gene is present in all cultivars of eggplant, or because not enough genetic material has been tested. This raises the possibility that some cases of non-host resistance, where all accessions are resistant to all isolates of a pathogen, may in fact be controlled by the same mechanisms as gene-for-gene resistance. Aside from the typical HR, an interaction between an Avr determinant and an R gene encoded protein can result in two other comparable responses: extreme resistance (ER) and systemic necrosis (SN). ER is broadly defined as a lack of visible response to virus infection. In an ER response there is no necrosis and little or no virus accumulation can be detected. SN is caused by the capability of the virus to evade the initial resistance response and spread to systemic tissues. However, the accumulation of virus here eventually leads to the initiation of cell death. This response is not sufficient to contain the virus, causing a trailing HR that eventually leads to collapse of the plant. The differences between these three responses, HR, ER, and SN, are suggested to be quantitative rather than qualitative, as illustrated by the following examples. The Rx1 and Rx2 CC-NB-LRR genes in potato confer an ER response to PVX, upon recognition of the same domain of the coat protein (CP). However, co-expression with the PVX CP results in an HR. The ER responses induced by Rx1 and Rx2 are thus not qualitatively different from HR, but rather eliminate the source of the Avr protein before it accumulates to HR inducing levels. Varying recognition efficiency of an NB-LRR protein between Avr effectors of different viral strains can result in a differential response, as in the case of Sw-5b. This tomato CC-NB-LRR recognizes the NSm protein of Tomato spotted wilt virus (TSWV), Tomato chlorotic spot virus (TCSV), and Groundnut ringspot virus (GRSV). Depending on the strain used, either HR or ER responses can be seen. The Arabidopsis CC-NB-LRR gene HRT (HR to TCV) recognizes the TCV CP, but in roughly 10% of cases the virus escapes the initial HR and replicates in systemic tissue, leading to SN. In absence of a second recessive gene, rrt (regulates resistance to TCV), this effect is much more pronounced. Although as of yet unidentified, this gene is expected to elevate HRT transcript levels in the highly resistant line Di-17. Additionally, the Rsv1 gene of soybean recognizes the viral Avr determinant of different Soybean mosaic virus (SMV) strains with varying efficiency, leading to ER, systemic infection, or SN, depending on the viral isolate. In the case of the I TIR-NB-LRR gene in bean, defense responses range from ER to HR to SN depending on the potyvirus isolate. The defense response is both dependent on temperature and has a gene dosage effect. At 231C plants homozygous for the I gene (I/I) confer ER to Bean common mosaic virus (BCMV), whereas heterozygous plants (I/i) confer an HR-type of resistance. At 341C, SN is seen on I/i plants, but to a lesser degree on I/I plants. Thus, the different responses to viruses by resistant plants likely represent a continuum of quantitatively different responses. The degree of response can be modulated by the relative efficiency of recognition of the Avr determinant, R gene dosage, temperature, and genetic background.

Mechanisms of Recognition Given the relatively straightforward genetic relationship between R and Avr genes, it was originally proposed that this might represent a receptor–ligand interaction. Since LRR domains often act as protein–protein interaction domains, it was proposed that the variable LRR domains of NB–LRR proteins would bind to different Avr proteins and this binding would activate defense responses. However, initial difficulties in demonstrating such interactions led to the formulation of indirect models of recognition. The guard hypothesis states that, since many Avr genes are also virulence genes, R proteins would ‘guard’ their cellular targets. The pathogen effector would modify cellular proteins, which is in turn detected by R proteins. Several examples of indirect recognition have been studied. For example, when expressed in Pseudomonas syringae, the Avr proteins AvrB and AvrRpm1 are both recognized

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by the Arabidopsis CC-NB-LRR protein resistance to P. syringae pv maculicola 1 (RPM1). RPM1 interacts with rpm1 interacting protein 4 (RIN4), a cellular protein which in turn can interact with both AvrB and AvrRpm1. This latter interaction is thought to activate the RPM1 protein. The Avr protein AvrRpt2 from P. syringae is a cysteine protease and is recognized by the Arabidopsis CC-NB-LRR protein resistant to P. syringae 2 (RPS2). RPS2 also binds RIN4 and becomes activated when RIN4 is cleaved by AvrRpt2. In the absence of RIN4, for example through genetic ablation, RPS2 is constitutively activated. In this case, RIN4 thus appears to function as a negative regulatory protein. At the same time, however, there is some indication that certain Avr/R protein pairs may interact directly. Yeast two-hybrid studies have shown Avr proteins interacting with the rice Pi-Ta, Arabidopsis RRS1 (resistant to Ralstonia solanacearum 1), and the flax L NB-LRR proteins which confer resistance to rice blast fungus, R. solanacearum, and flax rust, respectively. Direct interactions have also been reported between the viral R protein N and its cognate elicitor p50, surprisingly mediated by the TIR domain of N. However, a great deal about these interactions remains unclear, such as whether the other Avr proteins interact with the LRR domain, another part of the protein, or a combination thereof. Two traits desired by plant breeders related to recognition are broad-spectrum resistance and durability. Often, R genes are deployed that provide good resistance but are quickly overcome by new strains of a pathogen. The durability of an R gene may be affected by different aspects of the pathogen such as dispersal rates and mechanisms, occurrence of sexual reproduction, and variable repertoires of virulence factors. However, since viruses cannot dispense with any of their limited number of genes, overcoming resistance must occur through sequence changes. Several cases have been studied where the virus Avr gene has mutated to a point it is no longer recognized by the corresponding R gene. However, this often results in a loss of viral fitness due to the detrimental effect of these mutations, as illustrated by the examples below. The CC-NB-LRR encoding genes Tm-2 and Tm-22 have been introgressed from different Lycopersicum peruvianum accessions into tomato to control TMV and Tomato mosaic virus (ToMV). These genes confer resistance to nearly all strains of tobamoviruses through recognition of the viral movement protein (MP). Molecular cloning has revealed that Tm-2 and Tm-22 are different alleles of the same gene. They differ by only four aa, two of which are in the LRR domain. Despite this similarity, Tm-22 has proven to be more durable than Tm-2. Strains of ToMV that overcome Tm-2 or Tm-22 have mutations in different regions of the MP. The Tm-22 gene seems more durable because the resistance-breaking mutations have a very negative impact on virus fitness, resulting in reduced viral load in plants infected with these strains. The Tm-22-breaking strains cause very severe symptoms on Tm-22 plants so that infected plants can be easily recognized and eliminated from the field or greenhouse. Other R genes that are very durable are N, Rx1, and Ry. There are no strains of Potato virus Y (PVY) that overcome Ry and although ObPV can overcome N, it is not widespread. Likewise, Rx1-breaking PVX strains have reduced virulence and are found only in South America. Less durable genes include Nx and Nb, where PVX strains that overcome this resistance are more widespread. Additionally, TSWV strains breaking Sw-5b resistance have been reported in several tomato-growing areas. Not all mutations lead to a loss of fitness for the virus, for example in the case of differential recognition by the N0 gene of tobacco. The N0 gene confers resistance to several tobamoviruses including ToMV, but not TMV. Although N0 has not been cloned, the Avr determinant has been shown to be the viral CP. Curiously, mutations can be introduced into the TMV CP that result in a gain of recognition by N0 , but which appear to compromise CP function. These mutations are hypothesized to interfere with the tertiary or quaternary structure of the CP, exposing an N0 recognition site previously masked. Differential recognition of viral Avr proteins by R proteins can also be seen for the products encoded by the allelic pepper genes L1-L4, also recognizing the CP of tobamoviruses. Certain virus strains are able to evade recognition of L1, and thus L1-mediated resistance, some overcome L1 and L2, and some overcome L1, L2, and L3. However, the L4 gene is very broad spectrum, controlling all known tobamoviruses.

Signaling Mechanisms Studies with Rx1, HRT, Sw-5b and N have shown that the different domains of NB-LRR proteins undergo physical intramolecular interactions with the LRR and N-terminal domains interacting with the NB domain. These interactions appear to condition an auto-repressed state and alterations thereof are associated with activation of the protein. In the presence of PVX CP, the interdomain interactions of Rx1 are disrupted and the N protein has been shown to undergo oligomerization in the presence of p50. Both N and Rx1 shuttle between the nucleus and the cytoplasm, and both depend on their nuclear localization in order to induce a defense response. Interestingly, p50 is not required in the nucleus to elicit the N mediated defense response, but does so even when restricted to the cytoplasm, and Rx1 can only be activated in the cytoplasm. Similarly, the barley NB-LRR MLA10, and the Arabidopsis NB-LRRs RRS1, RPS4, and SNC1 (suppressor of npr1–1), are all at least to some extent localized to the nucleus. Of these, MLA10, RPS4, and SNC1 show a reduced immune response upon induced relocation from the nucleus to the cytoplasm. Taken together, these NB-LRRs provide evidence towards a nuclear signaling component, concurrent with one of the first steps of the immune response pathway being modification of transcription. In vitro studies have shown that Rx1 binds unselectively to DNA after activation through association with the PVX CP. It induces melting and bending of DNA, leading to structures resembling transcription initiation complexes. Rx1 most likely interacts with a Golden2-like (Glk) transcription factor, as demonstrated by its binding to Nicotiana benthamiana Glk1 in a transient transformation assay. In Arabidopsis, Glk1 and Glk2 are involved in a defense response against CMV, possibly in a redundant manner. In a double mutant, induction of defense related genes is hampered. In the current Rx1 resistance model, Rx1 and Glk1 associate in a

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complex and bind to DNA through Rx1 mediated DNA binding activity. Upon infection, the CP induces conformational changes in the Rx1-Glk1 complex, exposing the Glk1 DNA binding domain, previously blocked by the Rx1 CC domain. Glk1 binds stronger to its consensus DNA binding motifs, providing specificity to the induced transcriptional changes in the defense response. Concomitant with the role of Glk1 as an immune activating transcription factor, overexpression of Glk1 without Rx1 is sufficient to induce ER to PVX. It is presumed that other R proteins induce a signal transduction cascade leading to disease resistance by interacting with signal adaptor molecules as well. Although these molecules have yet to be identified, a number of proteins required for NB-LRR function are known. TIR-NB-LRR proteins, including N, have a general requirement for the EDS1 (enhanced disease susceptibility 1) protein and often also require the EDS1-interacting protein PAD4 (phytoalexin deficient 4). Both of these proteins have lipase signature domains, of which no exact function is known. Given that the N gene is not effective above 321C, it is interesting to note that EDS1 expression levels are greatly reduced at higher temperatures in Arabidopsis. Likewise, the barley MLA NB-LRR proteins accumulate to greatly reduced levels at high temperatures. These phenomena may at least partially explain the temperature sensitivity of some R genes. A number of studies have shown that during the resistance response, kinase cascades are activated, and transcription is reprogrammed substantially. Studies using virus-induced gene silencing (VIGS) have implicated several protein kinases, including MEK1, MEK2, NTF6, SIPK, WIPK, and NPK1, in N gene-mediated resistance. Additionally, successful N mediated resistance to TMV requires Beclin 1, Atg3 (autophagy related 3), Atg7, and Pi3k/Vps34 (class III phosphoinositide 3-kinase). These proteins are involved in the autophagy pathway, one of the major cellular degradation pathways in eukaryotes. Silencing these genes leads to programmed cell death beyond the infected tissue, thus for N mediated resistance the autophagy pathway appears involved in limiting the HR to the site of infection. Autophagy has been implicated in many plant processes, including innate immunity. Furthermore, the mammalian nod-like receptor 2 (NLRC2) activates antiviral innate immune signaling, using autophagy to regulate inflammatory responses. A requirement was also shown for the TGA and WRKY families of transcription factors as well as the individual transcription factors MYB1 (myb domain protein 1) and the TGA-interacting protein NPR1 (nonexpresser of pr genes 1). These proteins and protein families have also been implicated in gene-for-gene resistance to other types of pathogens. Gene-for-gene resistance may also involve targeted proteolysis since silencing the expression of certain proteins involved in regulated protein degradation, including a subunit of the COP9 (constitutive photomorphogenesis 9) signalosome, SKP1 (S phase kinase-associated protein 1 phase kinase-associated protein 1) and COI1 (coronatine insensitive 1), also compromises N-mediated resistance. To date, the only genes whose silencing has been shown to break Rx1-mediated resistance are Hsp90 (heat shock protein 90) and Sgt1 (suppressor of the G2 allele of skp1). Along with the Sgt1 binding protein Rar1 (required for MLA12 resistance 1), these proteins are required for the function of a number of NB-LRR proteins. The Hsp90, Sgt1, and Rar1 proteins are likely chaperoning proteins in order to allow proper protein folding. The plant hormone ethylene plays a role in R-gene-mediated antiviral responses. RCY1 functions at reduced efficiency in Arabidopsis containing the ethylene-desensitizing mutations ein3 and etr1. Salicylic acid (SA) also plays a role in R-gene-mediated resistance. The involvement of SA is often studied using SA biosynthetic mutants in Arabidopsis (such as eds5) or by using plants transgenically expressing the bacterial nahG gene, which encodes a salicylate hydroxylase that degrades SA. The eds5 mutation and nahG transgene both decrease the efficiency of RCY1-mediated resistance and completely abrogate HRT-mediated resistance. When tobacco plants expressing N and nahG are infected with TMV, the resulting HR lesions are larger, and necrosis can spread beyond the infected leaf. Similarly, in nahG transgenic tomato the normal ER response of Tm-22 becomes a spreading HR phenotype in the inoculated leaf, although the virus does not spread systemically. The Rx1-mediated response, however, is not affected by the nahG transgene. SA may enhance R gene responses rather than directly targeting the virus, and its requirement for resistance may depend on the strength of the R-gene-mediated response. In agreement with this, SA application results in reduced lesion size in the N-mediated response to TMV and an enhancement of HRT-mediated resistance. Cell death during the HR induces increased levels of SA. Thus, cell death may indirectly prime and/or augment R-gene-mediated antiviral responses. It is clear that cell death itself is not the only mechanism by which virus is eliminated as ER does not involve cell death, and in the N–TMV interaction, TMV can be detected outside the area of necrosis. Furthermore, in the interaction between Cauliflower mosaic virus (CaMV) and Nicotiana spp., resistance is controlled by a single dominant gene. However, whether or not this resistance is accompanied by an HR is controlled by a separate gene. A clear understanding of how viruses are cleared from resistant plant cells requires further study.

Resistosome Although multiple individual NB-LRR genes and their corresponding Avr effectors have been both identified and studied, the pathway of defense response induction in plants remains elusive. In animal systems, these mechanisms are often elucidated to a higher extent. Translating this information to plant systems could provide a valuable resource. In animal systems, one of the key factors required for resistance mediated by several R genes was found to be oligomerization. The highly structured complexes are referred to as either an inflammasome or apoptosome. Three well-studied examples include the human R protein apoptotic protease-activating factor 1 (APAF1), the nematode R protein cell death protein 4 (CED-4), and the human protein NLRC4. Within plants, although a similar resistance mechanism was hypothesized in 2010, only one R complex has been identified, as recent as 2019. This work involved the CC-NB-LRR protein hopz-activated resistance 1 (ZAR1), conferring resistance within Arabidopsis and

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N. benthamiana to disease caused by the bacteria Xanthomonas campestris. The formation of the ZAR1 complex, termed resistosome, is essential for resistance induced by the X. campestris effector AvrAC. The resistosome structurally resembles the inflammasome and the apoptosome. All three consist of a wheel-like basis, and the NB-ARC domain is similarly positioned between ZAR1, APAF1, and NLRC4 to facilitate oligomerization (Fig. 1). Furthermore, although ZAR1 oligomerization requires certain co-factors, these are not part of the interacting domains between the sub-units, similar to the co-factor cytochrome C in the Apaf1 apoptosome. Important differences include the organization of the N-terminal domains. The Apaf1 apoptosome and the NLRC4 inflammasome contain disorganized caspase recruitment domains (CARDs), in the absence of certain caspases. These caspases are interacting partners required for downstream processing of the defense signal. However, the a helices in the CC domain of the ZAR1 resistosome are highly structured. Furthermore, the number of sub-units contributing to the complex differs from five in the ZAR1 resistosome, to seven for the Apaf1 apoptosome, nine for the CED4 apoptosome, and eleven for the NLRC4 inflammasome. Assembly of the ZAR1 resistosome requires the presence of two other proteins: resistance-related kinase 1 (RKS1) and PBS1 like protein 2 (PBL2), as well as the exchange of an ADP for ATP. Current evidence suggests ZAR1 and RKS1 form a dimer within the cytoplasm of the plant cell. Upon infection by the X. campestris bacteria, the AvrAc protein modifies the host protein

Fig. 1 Protein structure of the ZAR1 resistosome (A, D, G), the APAF1 apoptosome (B, E), the NLRC4 inflammasome (C, F), and the actinoporin FraC (H). Colors indicate the transition from N (blue) to C terminal (red) (A–C) or the different subunits in the complex (D–H). A–C show top views of the complexes, and D–H show side views. G shows a detail of the a helical bundle of the ZAR1 complex in D, for comparison to the a helical bundle of FraC in H.

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PBL2 by the addition of a uridine monophosphate. This modified PBL2 binds to the ZAR1-RKS1 dimer, promoting the release of ADP from this complex. The subsequent binding of ATP leads to the oligomerization of this complex in a resistosome consisting of five ZAR1-RKS1-PBL2 subunits. Interestingly, the oligomerization towards the resistosome seems to differ from the activation and assembly of the human NLRP3 inflammasome, where a potassium efflux seems required in almost all cases. Although the ZAR1-RKS1 dimer is only to a very limited degree localized in the plasma membrane, resistance mediated by the resistosome complex requires its presence in the plasma membrane. To understand the translocation of the resistosome, a closer look must be taken to the rearrangements of the different ZAR1 domains during activation. The NB-ARC domain of ZAR1 can be further divided in a NB domain, HD1 (helicase domain 1), and WHD (winged helix domain). Rearrangements within the CC domain lead to the exposure of the a1 helix, from a position buried and contacting the LRR and WHD domain, to a position protruding from the rest of the protein. Upon oligomerization, the a1 helices form a highly structured funnel shape. This most likely facilitates the association to the plasma membrane. Mutation of several aa in this a1 helix, as well as adding a FLAG peptide, leads to a loss of plasma membrane association. Although these perturbations of the a1 helix lead to loss of ZAR1 resistance, they do not affect the formation of the ZAR1 resistosome. Interestingly, this funnel shaped structure in the ZAR1 resistosome resembles that of the actinoporin fragaceatoxin C (FraC), although the proteins bear little homology. Actinoporins like FraC are pore-forming complexes found in sea anemone species, used both in defense and as a hunting strategy. FraC attaches to the plasma membrane, and oligomerizes into a complex consisting of nine subunits. This probably changes the protein conformation leading to the detachment of the N terminal a helix, and subsequent insertion of the a helices into the membrane, forming a highly structured a helical bundle. After the transition from a prepore to a pore, the membrane is no longer capable of maintaining ion balance. The presence of the nonselective hole leads to an efflux of ions and small molecules, depending on the pore size, and ultimately leads to cell injury or death. Aside from the similarities in structure between the a helical funnels of the ZAR1 resistosome and the FraC actinoporin complex, their size is also comparable, at 33 and 35 ångström, respectively. This could indicate that the ZAR1 funnel is also membrane spanning, rather than membrane inserted. Following this hypothesis, the ZAR1 resistosome could function through disturbance of the membrane integrity in a similar manner as the FraC complex, leading to localized cell death and limitation of pathogen spread. This is supported by the observation that mutating the inner surface of the ZAR1 resistosome funnel abolishes disease resistance. In animal cells, a similar route of using pore complexes to mediate a cell death response exists. For example, in humans and mice, inflammatory caspases oligomerize upon activation into an inflammasome. One such example is caspase 11, which cleaves gasdermin D (GSDMD), in a C-terminal and N-terminal part. After cleavage, GSDMD-NT oligomerizes within the cell membrane and forms pores, leading to cell death. For now, the exact resistance mechanism of ZAR1 remains unidentified, and future research will have to show whether ZAR1 functions as a pore complex as hypothesized, or through a different mechanism.

Non-NB-lRR Resistance Although by far most identified R genes belong to the NB-LRR class, other classes of R genes should not be ignored or overlooked. In fact, breeding for resistance mediated by NB-LRR genes may prove less durable than breeding for resistance conferred by nonNB-LRR genes. This inherent disadvantage attributed to NB-LRRs is caused by the gene-for-gene relationship NB-LRRs have with pathogen Avr determinants. This puts a relatively large selection pressure on the pathogen. Any mutation in the pathogen effector leading to less recognition by the NB-LRR increases the pathogens fitness considerably, as it is now able to infect a new host genotype. The likelihood of the frequency of this mutation rising within the population is therefore considerable, and the breeder would have to start over again by re-introducing resistance within the crop plant. Non-NB-LRR R genes could help alleviate this disadvantage, as often the principle of gene-for-gene resistance does not apply here. Instead, the underlying mechanism of resistance involves a more complex interplay of the pathogen and host, or the perception of more generic signals of the pathogen. Although less emphasis is placed on non-NB-LRRs R genes, they represent interesting sources to confer more durable (and sometimes broader) crop resistance. One such example can be found within the major plant anti-viral RNA interference (RNAi) defense mechanism. RNAi is triggered by a highly conserved cue of viral presence in the plant cell: double stranded RNA (dsRNA). These dsRNAs are formed during viral replication, either through overlapping open reading frames (ORFs), from highly structured viral mRNAs, or as a replication intermediate of RNA viruses. During the life cycle of a healthy plant cell, dsRNA is rarely formed, rendering this form of RNA a distinctive signal to the plant cell of pathogen presence. Dicer-like (DCL) enzymes cleave these RNA duplex molecules in small fragments, typically 20–25 nt in size. One strand of these small interfering RNAs (siRNAs) is then loaded into an AGO containing RNA-induced silencing complex (RISC). Any RNA complementary to this strand is targeted for degradation, giving rise to aberrant RNA molecules, or translational arrest. During an amplification cycle, a type RNA dependent RNA polymerases (RDRs) synthesize a full-length dsRNA molecule using the single-stranded aberrant RNA molecule as a template. This full-length dsRNA is cleaved by one of the DCLs, generating a population of secondary siRNAs. These can either be used again to target complementary RNAs for silencing or spread through the plasmodesmata between plant cells, leading to immunization beyond the infected cell. Upon infection by a DNA virus, siRNAs are alternatively loaded on an RNA-induced transcriptional silencing (RITS) complex, containing a distinct AGO protein. This complex targets complementary DNA for methylation, leading to a block of transcription of viral DNA.

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Table 2 Overview of cloned antiviral non-NB-LRR genes. The resistance mechanism is indicated, as well as the species in which the R gene is used (plant), and the viral elicitor Resistance mechanism

R gene

Encoded protein

Plant

Virus

Elicitor

RNAi Ty-1 RDRg Restricted systemic RTM1–5a Jacalin, HSP20, MATH, ND, ND movement Lectin-mediated resistance JAX1 Jacalin

Tomato Arabidopsis

TYLCV Potyviruses (TEV, PPV, LMV)

dsRNA ND

Arabidopsis

Replicase

Inhibition RNA replication Tm-1

Tomato

Potexviruses (PlAMV, PVX, Asparagus virus 3, White clover mosaic virus) ToMV

TIM-barrel-like domain

130K and 180K replication proteins

a

RTM mediated resistance requires the presence of all 5 genes.

Recently, Ty-1, one of the major dominant resistance genes against the geminivirus TYLCV (Tomato yellow leaf-curl virus) in tomato, has been cloned, and was shown to encode a member of the g subclass of RDRs (Table 2). Previously, members of this class had not been assigned any function. The main difference between the well characterized a type and the g type lies within the catalytic core domain, the a type containing a DLDGD motif and the g type a DFDGD motif. Ty-1 has been shown to confer resistance, still allowing very low levels of viral DNA replication, by enhancing the RNAi response. Specifically, it strengthens transcriptional gene silencing of viral DNA. As opposed to an NB-LRR recognizing one pathogen Avr effector, Ty-1 potentially entails a more durable R gene and confers resistance to a wider range of mono-/bipartite geminiviruses, besides TYLCV. The achilles heel of Ty-1-mediated resistance lies in co-infecting plant viruses with the ability to interfere with antiviral RNAi using proteins called viral suppressor of RNAi (VSR). Another interesting example can be found in the Arabidopsis R genes Restricted tobacco etch potyvirus (RTM) and Sieve element lining chaperone 1 (SLI1). RTM mediated resistance requires the presence of five genes, termed RTM1–5. Together, these genes limit the spread of TEV, Plum pox potyvirus (PPV), and Lettuce mosaic virus (LMV) through the plant to systemic tissues. RTM1 encodes the lectin protein jacalin. Several members of this gene family have been shown to provide resistance to insects, fungi, and viruses. RTM2 is an unusual small heat shock protein, containing a C-terminal transmembrane domain. It is neither inducible by heat treatment, nor providing tolerance towards high temperatures. RTM3 belongs to a gene family of unknown function, whose members contain a meprin and TRAF homology (MATH) domain and a C-terminal CC domain. RTM4 and RTM5 remain uncharacterized. Although the exact resistance mechanism is unknown, resistance breaking strains rely on changes in the N-terminal region of the CP. No interaction has been shown between the CP and RTM1–3. Both RTM1 and RTM2 are shown to localize to sieve tubes, rendering it likely that the resistance acts in phloem tissues. Oligomerization of several different factors seems required, as demonstrated by both the direct interaction of RTM1 and RTM3, and the presence of several domains involved in protein-protein interactions in RTM1–3. An alternative method of introducing resistance against viruses is by combatting vectors transmitting these viruses between plants. SLI1 is an example of an R gene that could be used in such a manner. It has been recently identified and confers resistance in Arabidopsis against the generalist aphid Myzus persicae through restriction of both feeding time and phloem ingestion rate. SLI1 limits the growth of the aphid population on the host plant, and in this manner SLI1 could aid in limiting virus spread throughout the plant population. As opposed to RTM2, SLI1 confers tolerance to higher temperatures. SLI1 is lined around the entire sieve elements, including the sieve plate, and specifically around mitochondria in a peripheral strip of cytoplasm. Interestingly, RTM2 and SLI1 are homologs belonging to the same superfamily of heat shock protein 20 (HSP20)-like chaperones.

Outlook With the recent discovery and elucidation of the ZAR1 resistosome a new chapter has opened on the functioning of NB-LRR resistance proteins. Whereas earlier research has indicated that the actual resistance response is different from HR, findings on the ZAR1 resistosome, most likely functioning as a membrane pore, implies that cell death is an essential and final stage of the resistance response/mechanism. This (again) forces us to re-visit our views on the resistance mechanism. If the hypothesized function of ZAR1 holds true, how can HR and the so-called actual resistance response be uncoupled? Maybe on this point the resistance mechanism consists of sequential layers, with the Avr being the initial trigger to all. The first layer(s) could be pathogenspecific and the formation of a resistosome represents the final stage that is generic and always leads to cell death, irrespective of the type of pathogen. This also would explain why in most cases HR occurs concomitant with the induction of NB-LRR-gene mediated resistance. The lack of HR in several cases could then be a quantitative rather than qualitative matter, similar to the differences observed between HR, ER, and SN responses. Alternatively, this could result from mutations that prevent the R protein from oligomerization but not compromise the earlier, upstream steps of the resistance response. Although speculative, the formation of different oligomers as observed with animal R proteins, like inflammasomes and apoptosomes, also raises the question on the existence of different resistosomes in plants. In light of this it is interesting to remark

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on differences between plant NB-LRRs. For ZAR1 the CC domain is not only essential for oligomerization, but its overexpression also leads to HR. This is compromised by N-terminal fusions or -deletions at the CC domain. In contrast, for several other NB-LRR genes, not CC but overexpression of NB-ARC (e.g., Sw-5b) or NB (e.g., Rx1) triggers HR. In addition, the folding structure of the ZAR1 CC domain is different from those of stem rust resistance 33 (Sr33), Rx1 and MLA10. This indicates a possible classification of NB-LRRs into subgroups. On this point, a recent study has shown that several NB-LRRs, termed sensors, require the presence of a second NB-LRR, termed helper. These helpers can be divided in three classes: NRC (NB-LRR required for cell-death), ADR1 (Activated disease resistance 1), and NRG1 (N required gene 1). Interestingly, these helpers can be (partially) redundant, as in the case of NRC2, 3, and 4, while several sensors can share the same helper. Silencing all three NRCs affects, among others, resistance mediated by Sw-5b and Rx1. Subsequent challenge of NRC silenced plants carrying Rx1 with PVX results in SN, instead of the usual ER response. This suggests NB-LRRs can be divided into at least two sub-groups, those that act without helper and those that require a helper. This could also explain why domains from different NB-LRR proteins are not all functionally equivalent in signal transduction. Building on this, a recent study revealed an N-terminal 21 aa motif, termed MADA, that is conserved among the NRC helper proteins and folds into an a helix similar to the a1 helix of ZAR1. Additionally, this motif is found in approximately 20% of the 988 CC-NB-LRRs tested. The MADA motif is essential for the HR response, and upon switching with the corresponding region in ZAR1 the helper NB-LRR NRC4 is still capable of conferring cell-death. Whether NB-LRRs that need a helper differ from ZAR1 in the formation of a distinct resistosome remains an intriguing question to be solved. Despite a strong interest for dominant R genes, interest in non-NB-LRR genes lags behind. Their identification being more difficult and elaborate than that of NB-LRR genes has so far limited the number of antiviral non-NB-LRR genes characterized. Nevertheless, non-NB-LRR R genes hold a strong promise towards the future, due to their more durable nature and broader resistance spectrum.

Further Reading Adachi, H., Contreras, M., Harant, A., et al., 2019. An N-terminal motif in NLR immune receptors is functionally conserved across distantly related plant species. eLife 8, e49956. Bentham, A., Burdett, H., Anderson, P.A., Williams, S.J., Kobe, B., 2016. Animal NLRs provide structural insights into plant NLR function. Annals of Botany 119 (5), 698–702. Carr, J.P., Murphy, A.M., Tungadi, T., Yoon, J.Y., 2019. Plant defense signals: Players and pawns in plant-virus-vector interactions. Plant Science 279, 87–95. de Ronde, D., Butterbach, P., Kormelink, R., 2014. Dominant resistance against plant viruses. Frontiers in Plant Science 5, 307. Jubic, L.M., Saile, S., Furzer, O.J., El Kasmi, F., Dangl, J.L., 2019. Help wanted: Helper NLRs and plant immune responses. Current Opinion in Plant Biology 50, 82–94. Qu, F., 2010. Plant viruses versus RNAi: Simple pathogens reveal complex insights on plant antimicrobial defense. Wiley Interdisciplinary Reviews: RNA 1 (1), 22–33. Wang, J., Hu, M., Wang, J., et al., 2019. Reconstitution and structure of a plant NLR resistosome conferring immunity. Science 364 (6435), eaav5870. Wu, C.H., Abd-El-Haliem, A., Bozkurt, T.O., et al., 2017. NLR network mediates immunity to diverse plant pathogens. Proceedings of the National Academy of Sciences of the United States of America 114 (30), 8113–8118. Zhang, L., Chen, S., Ruan, J., et al., 2015. Cryo-EM structure of the activated NAIP2-NLRC4 inflammasome reveals nucleated polymerization. Science 350 (6259), 404–409. Zhou, M., Li, Y., Hu, Q., et al., 2015. Atomic structure of the apoptosome: Mechanism of cytochrome c-and dATP-mediated activation of Apaf-1. Genes & Development 29 (22), 2349–2361.

Plant Resistance to Viruses: Natural Resistance Associated With Recessive Genes Masayoshi Hashimoto, Kensaku Maejima, Yasuyuki Yamaji, and Shigetou Namba, The University of Tokyo, Tokyo, Japan r 2021 Elsevier Ltd. All rights reserved. This is an update of C. Caranta, C. Dogimont, Plant Resistance to Viruses: Natural Resistance Associated with Recessive Genes, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00735-4.

Nomenclature aa amino acid(s) Avr avirulence Cas CRISPR-associated proteins CI cylindrical inclusion CITE cap-independent translation enhancer cPGK2 chloroplast phosphoglycerate kinase 2 CRISPR Clustered regularly interspaced short palindromic repeats dsRNA double-stranded RNA EMS ethyl methane sulfonate ER endoplasmic reticulum GM genetically modified gRNA genomic RNA HR hypersensitive response IRES internal ribosomal entry site KO knockout mRNA messenger RNA nCBP novel cap-binding protein

Glossary Allele Allele is one of the distinct forms of a gene at a single locus. Show different effect on the phenotype from the other forms. Avirulence Genetic traits for a pathogen to be impaired by genetically determined host resistance and to cause an incompatible (no disease) interaction. Dominant allele An allele expressing its phenotypic effect even when heterozygous with a recessive allele. Genome editing Genome editing is a technology for the modification of genomic DNA sequence (deletion, substitution, and insertion) using sequence-specific nucleases. Paralog Paralogs are a group of genes generated by duplication events within a genome. Orthologs, which are evolved from a common ancestral gene in different species, retain evolutionally conserved functions, whereas paralogs generally evolve new functions.

NGS next generation sequencing nt nucleotide(s) ORF open reading frame PABP poly(A)-binding protein PDIL5-1 protein disulfide isomerase like 5-1 Poly(A) polyadenylated QTL quantitative trait locus RdRp RNA-dependent RNA polymerase RIL recombinant inbred line SNP single-nucleotide polymorphism ssRNA single-stranded RNA TALEN transcription activator-like effector nuclease TILLING targeting induced local lesions in genomes tRNA transfer RNA UPR unfolded protein response Vpg viral protein genome-linked VPS41 vacuolar protein sorting 41 ZFN zinc-finger nuclease

Pleiotropic mutation A mutation showing multiple distinct effects. Positional cloning Determining the location of a gene-ofinterest on the chromosome without any idea of the biological function of that gene. Quantitative trait loci Correspond to genomic regions associated with the phenotypic variation of a quantitative trait. Recessive allele An allele whose phenotypic effect is not expressed in a heterozygote. TILLING A reverse genetics approach that relies on the ability of a special enzyme to detect mismatches in normal and mutant DNA strands when they are annealed. It can therefore detect single point mutations of the type induced by chemicals conventionally used in mutation breeding programs.

Introduction To defend against plant pathogenic microbes, plants deploy resistance machinery, which can be inherited genetically. Genetic resistance, also referred to as host or cultivar resistance, is determined by a specific genotype in the resistant host variety, relative to other varieties possessing distinct genotypes that confer susceptibility. Employing host resistance machinery through the conventional breeding has long been one of the most effective and sustainable ways to control diseases caused by plant pathogenic microbes including viruses.

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In terms of genetics, host resistance can be classified as monogenic, digenic, or polygenic ones, governed by one, two, or several genes, respectively. Most monogenic and digenic types of resistance are characterized by qualitative resistance phenotypes, such as the complete absence of pathogen proliferation. In contrast, most polygenic resistance is phenotypically quantitative, underlined by several resistance genes generally called quantitative trait loci (QTLs). QTL mapping is the main technique used to identify genetic markers statistically linked to the resistance phenotype, in which the recombinant inbred lines (RILs) derived from the crosses between parents with different resistance traits are used to collect the phenotypic data. In practice, the development of genetic markers tightly linked to resistance traits has been quite valuable for the conventional breeding of resistant plant varieties. Host resistance can be inherited in a dominant or recessive manner. Almost half of the host resistance against plant viruses that have been characterized in natural plant ecotypes are inherited recessively. This high ratio of recessive inheritance is specific to plant viruses and has not been observed for other plant pathogens, such as oomycetes, fungi, and bacteria: this feature is closely related to the distinct infection cycles of plant viruses, as will be discussed later. The degree of recessively inherited resistance ranges from partial to complete inhibition of pathogen invasion. Some plants with partial resistance show “tolerance,” in which symptom attenuation and/or no crop yield loss is observed, even under complete systemic infection by the pathogens. More phenotypic variation is observed for recessive resistance, than for dominant resistance, which mainly involves pathogen confinement at the initial infection site and is frequently associated with localized cell death, also known as hypersensitive response (HR). Recessive resistance can interrupt viral infection at any stage of the infection cycle, including during viral protein translation, viral genome replication, and cell-to-cell and systemic movement. Resistance breakdown by the emergence of a virus strain with genome mutations frequently accompanies recessive resistance in agricultural fields. Comparative genetic analysis using resistance-breaking viral strains can be used to identify the viral avirulence (Avr) factors that determine recessive resistance. Functional analysis of the interaction between the viral Avr factor and the plant resistance gene product promotes our understanding of the molecular dialog between viruses and their host plants, and contributes to the development of new breeding strategies based on natural recessive resistance to avoid resistance breaking. In this article, we demonstrate the phenotypic and molecular characteristics of natural virus recessive resistance in plants, and describe recent advances in this field.

Overview of Naturally Occurring Recessive Resistance Recessive resistance is a highly relevant trait for virus-resistant crop breeding. The ratio of recessive to dominant resistance to plant viruses is much higher than those of resistance to pathogens from other kingdoms, including oomycetes, fungi, and bacteria. Examples of recessive resistance found in nature, drawn from a survey of the literature, are shown in Table 1. Recessive resistance to viruses has been observed in dicots and monocots. It has been studied most intensively in the dicots families Fabaceae, Cucurbitaceae, and Solanaceae, and in the monocots barley and rice. Recessive resistance is rarely found in polyploid plants, likely due to the presence of dominant gene copies with redundant function. However, one allopolyploid plant, Brassica juncea (mustard), encodes recessive resistance gene retr03, which has been found at a higher than expected ratio in 7 resistant landraces among 35 tested. Inherently recessive resistance has been characterized for viruses of diverse taxa, with various types of RNA- or DNA-based genome structure. Many of these viruses belong to the genus Potyvirus. Potyviruses are single-stranded RNA viruses that account for about 70% of the recessive resistance in a wide range of plant species, together with other related viruses encoding the viral protein genome–linked (VPg) protein, which binds to the 50 end of their genomic RNA as a cap analog. Potyvirus is the largest genus of phytopathogenic viruses; it contains many virus species that cause severe crop yield losses worldwide. The frequency of recessive resistance varies among virus families and genera: it is almost 50% in the genus Potyvirus, but rare in other virus genera, such as Tobamovirus. This inconsistency is likely to relate to virus infection strategies and selection pressure due to viral infection in natural plant habitats. Thus, the frequency of recessive resistance for a specific plant–virus interaction may be determined by a trade-off between the fitness costs of recessive mutations of resistance genes and the costs of viral infection to healthy plant growth. Several resistance loci control distinct strains of single virus species, or two or more distinct viral species. In barley (Hordeum vulgare), two distinct alleles, rym4, and rym5, at a single recessive locus confer resistance to several bymoviruses, including Barley mild mosaic virus (BaMMV), Barley yellow mosaic virus (BaYMV), and Barley yellow mosaic virus-2 (BaYMV-2). Alleles at the pvr2 locus in pepper (Capsicum spp.) are responsible for resistance to several potyviruses, including Potato virus Y (PVY), Tobacco etch virus (TEV), and Pepper mottle virus (PepMoV). Another example is pea (Pisum sativum) with two well-documented recessive resistance loci against several potyviruses. At the first locus are bcm against Bean common mosaic virus (BCMV), cyv1 against Clover yellow vein virus (ClYVV), mo against Bean yellow mosaic virus (BYMV) and Watermelon mosaic virus (WMV), sbm-2 against Pea seed-borne mosaic virus (PSbMV), and pmv against Pea mosaic virus (PMV). At the second locus are sbm-1, sbm-3, and sbm-4, conferring resistance to three PSbMV strains, cyv2 against ClYVV, wlv against BYMV, and mo11 and mo12 against Lettuce mosaic virus (LMV). With recent advances in the molecular understanding of natural recessive resistance, discussed below, the broad resistance spectrum of these loci has been attributed to the action of a single gene with a pleiotropic effect. Although recessive resistance is usually governed by a single gene, it is controlled by two distinct loci in several plant–virus pairs. In common bean (Phaseolus vulgaris), resistance against BCMV pathotypes requires two complementary genes at the bc-u locus and at one of the three strain-specific loci (bc-1, bc-2, and bc-3). Interestingly, in pepper, only the pvr2 locus controls resistance against potyviruses including PVY, whereas the pvr6 and pvr2 loci are required for recessive resistance to another

Plant Resistance to Viruses: Natural Resistance Associated With Recessive Genes

Table 1

71

Virus recessive resistance found in hosts

Genus – Family/Virus (acronym)

Host plant

RNA viruses with a 50 VPg and 30 poly(A) tail Bymovirus - Potyviridae Hordeum vulgare (barley) Barley mild mosaic virus (BaMMV) Barley yellow mosaic virus (BaYMV)

Hordeum vulgare (barley)

Barley yellow mosaic virus-2 (BaYMV-2) Hordeum vulgare (barley) Potyvirus - Potyviridae Bean common mosaic virus (BCMV)

Pisum sativum (pea) Phaseolus vulgaris (common bean) Phaseolus vulgaris (bean) Bean yellow mosaic virus (BYMV) Pisum sativum (pea) Vicia mungo (mungbean) Pisum sativum (pea) Bean yellow mosaic virus-W (BYMV) Lactuca sativa (lettuce) Bidens mottle virus (BiMV) Blackeye cowpea mosaic virus (BlCMV) Vigna unguiculata (cowpea) Apium graveolens (celery) Celery mosaic virus (CeMV) Phaseolus vulgaris (bean) Clover yellow vein virus (ClYVV) Pisum sativum (pea)

Lettuce mosaic virus (LMV) Moroccan watermelon mosaic virus (MWMV) Papaya ringspot virus-W (PRSV)

Passionfruit woodiness virus (PWV) Pea seed-borne mosaic virus (PSbMV)

Peru tomato virus (PTV) Pepper mottle virus (PepMoV) Pepper veinal mottle virus (PVMV) Plum pox virus (PPV) Potato virus A (PVA) Potato virus Y (PVY)

Tobacco etch virus (TEV)

Tobacco vein mottling virus (TVMV) Turnip mosaic virus (TuMV)

Glycine max (soybean) Lactuca sativa (lettuce) Cucumis sativus (cucumber) Cucurbita ecuadorensis Cucumis sativus (cucumber)

Resistance gene(s) or locia

Resistance phenotypea

rym4, rym5 715 rym on 5chr. rym4, rym5 715 rym on 5chr. rym5 rym1, rym11

CR PR-CR CR PR-CR CR

bcm bc-1, bc-12, bc-2, bc-22, bc-3, bc-u (a locus) cyv mo 2 genes wlv bi bcm cmv desc cyv1 cyv2 (a locus) d-cv mo11, mo12 mwm

No SI

2 genes prsv-1 prsv2245 Cucurbita moschata (squash) prv Citrullus amarus (watermelon) prv Pisum sativum (pea) pwv Lens culinaris (lentil bean) sbv sbm1, sbm3, sbm4 Pisum sativum (pea) sbm2 (a locus) Solanum lycopersicum Monogenic (tomato) Capsicum annuum (pepper) pvr3 Capsicum chinense (pepper) pvr1 Capsicum annuum (pepper) pvr22 þ pvr6 (digenic) Arabidopsis thaliana rpv1 Solanum tuberosum subsp. raadg andigena Capsicum annuum (pepper) pvr21, pvr22 pvr3 (a locus) Several QTLs Capsicum chinense (pepper) pvr1 (allelic to pvr2) pot-1 Solanum habrochaites (tomato) Nicotiana tabacum (tobacco) va Capsicum annuum (pepper) pvr22 Capsicum chinense (pepper) pvr1 pot-1 Solanum habrochaites (tomato) Nicotiana tabacum (tobacco) 2 genes linked Nicotiana tabacum (tobacco) va Brassica campestris 2 genes (cabbage) 1 to 3 genes Brassica rapa (turnip) trs retr01 retr02 retr03 Brassica juncea (mustard)

No SI SL No local or SI CR Cell R Cell-to-cell Mvt Red Acc Red Acc SL Red Acc Red Acc Red Acc SL CR Red Acc Cell R

Long-distance Mvt Cell R CR Long-distance Mvt Vascular Mvt Cell R CR PR Cell R CR Cell-to-cell Mvt Cell R Cell R CR Long-distance Mvt Cell-to-cell Mvt No SI No SI Red Acc

(Continued )

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Table 1

Plant Resistance to Viruses: Natural Resistance Associated With Recessive Genes

Continued

Genus – Family/Virus (acronym) Watermelon mosaic virus (WMV)

White lupin mosaic virus (WLMV) Zucchini yellow mosaic virus (ZYMV)

Zucchini yellow fleck virus (ZYFV) Comovirus - Secoviridae Cowpea severe mosaic virus (CPSMV) Waikavirus - Secoviridae Rice tungro spherical virus (RTSV)

Host plant Cucumis melo (melon)

No SI No SI

Vigna unguiculata (cowpea)

Oligogenic (3 genes)

CR

Oryza sativa (rice)

tsv-1 tsv-2 (a loci)

No SI

lr lrv

SL SL

Lactuca sativa (lettuce) Cucumis melo (melon)

bwy cab-1 þ cab-2 (digenic)

Red Acc CR

Oryza sativa (rice)

rymv1 Several QTLs rymv1 rymv2

CR PR CR

Oryza glaberrima (rice) RNA viruses with capped 50 end and non-polyadenylated 30 end Alfamovirus - Bromoviridae Medicago sativa (alfalfa) Alfalfa mosaic virus (AMV) Bromovirus - Bromoviridae Cowpea chlorotic mottle virus (CCMV) Glycine max (soybean) Cucumovirus - Bromoviridae Cucumis melo (melon) Cucumber mosaic virus (CMV) Cucurbita pepo (pumpkin) Capsicum frutescens (pepper) Capsicum annuum (pepper) Tobamovirus - Tobamoviridae Cucumis melo (melon) Cucumber green mottle mosaic virus (CGMMV)

Umbravirus - Tombusviridae Groundnut rosette virus (GRV) Tospovirus - Bunyaviridae Tomato spotted wilt virus (TSWV)

Resistance phenotypea

monogenic wmv1551 Cucumis sativus (cucumber) wmv-2 wmv-3 þ wmv-4 (digenic) mo Pisum sativum (pea) Arabidopsis thaliana rmv1 Cucumis sativus (cucumber) wmv02245 Citrullus lanatus (watermelon) 2 genes Pisum sativum (pea) wlv Cucumis sativus (cucumber) zym Citrullus lanatus (watermelon) zym-CH zym-FL Citrullus mucosospermus (watermelon) Citrullus lanatus (watermelon) monogenic Cucumis sativus (cucumber) zyf

RNA viruses with a 50 VPg and non-polyadenylated 30 end Enamovirus - Luteoviridae Pisum sativum (pea) Bean leafroll virus (BLRV) Polerovirus - Luteoviridae Beet western yellows virus (BWYV) Cucurbit aphid-borne yellows virus (CABYV) Sobemovirus - Solemoviridae Rice yellow mottle virus (RYMV)

Resistance gene(s) or locia

Red Acc; Long-distance Mvt To SL cotyledons SL No SI

PR Red Acc No SI

am-1 2 genes

Long-distance Mvt

Polygenic with a major QTL (cmv1) 2 unlinked genes Oligogenic (2–3 genes) cmr2

SL SL No SI

Polygenic

SL

cgmmv-1 cgmmv-2 Arachis hypogaea (peanut)

Solanum lycopersicum (tomato) RNA viruses with unmodified 50 and 30 ends Carmovirus - Tombusviridae Arabidopsis thaliana Turnip crinkle virus (TCV) Cucumis melo (melon) Melon necrotic spot virus (MNSV) Nicotiana benthamiana Luteovirus - Luteoviridae Hordeum vulgare (barley) Barley yellow dwarf virus (BYDV) Idaeovirus Rubus idaeus (raspberry) Raspberry bushy dwarf virus (RBDV) dsRNA virus

2 genes

SL

sw2, sw3, sw4

PR

rrt þ HRT (digenic) nsv (no locus name)

HR Cell R Non-host R

ryd1

PR

Monogenic

CR

Plant Resistance to Viruses: Natural Resistance Associated With Recessive Genes

Table 1

73

Continued

Genus – Family/Virus (acronym) Fijivirus - Reoviridae Maize rough dwarf virus (MRDV) Mal de Rio Cuarto virus (MRCV) Rice black-streaked dwarf virus (RBSDV) DNA viruses Begomovirus - Geminiviridae Bean golden yellow mosaic virus (BGYMV)

Host plant

Resistance gene(s) or locia

Zea mays (maize) Zea mays (maize) Zea mays (maize)

Polygenic with a major QTL (qMrdd1) Polygenic with a major QTL (qMrdd1) Polygenic with a major QTL (qMrdd1)

Phaseolus vulgaris (common bgm-1 bean) bgm-2 (a loci) Phaseolus coccineus (scarlet bgm-3 (a loci) runner bean) Solanum pimpinellifolium Potato yellow mosaic virus (PYMV) Digenic Tomato yellow leaf curl virus (TYLCV) Solanum habrochaites Oligogenic A major QTL, ty-5, and four minor QTLs Solanum lycopersicum (tomato) Solanum chilense Tomato leaf curl virus (ToLCV) tgr-1 tcm-1 Tomato chlorotic mottle virus (ToCMoV) Solanum lycopersicum (tomato) Arabidopsis thaliana Cabbage leaf curl virus (CaLCuV) gip-1 (Pla-1 ecotype)

Resistance phenotypea

Red S Red S Red S No SI CR Red S Cell-to-cell Mvt No SI No Acc Inoc leaves; R to BCTV and TYLCV

Curtovirus - Genimiviridae Beet curly top virus (BCTV)

Arabidopsis thaliana

Monogenic R loci in three ecotypes, Ms-0, To or No SI Pr-0, and Cen-O

Tungrovirus (Caulimoviridae) Rice tungro bacilliform virus (RTBV)

Oryza sativa (rice)

Polygenic

To, Red Acc

Abbreviations: CR, complete resistance; chr, chromosomes; PR, partial resistance; a, independent; SL, symptomless; Cell R, cellular resistance (resistance at a single cell level); To, tolerance; SI, systemic infection; Mvt, movement; Red, reduced; Acc, accumulation; Inoc, inoculated; HR, hypersensitive response; S, symptoms; R, resistance.

a

potyvirus, Pepper veinal mottle virus (PVMV). These phenomena imply that the pyramiding of independent resistance loci in a single cultivar, if possible, would broaden the spectrum of recessive resistance. Among a number of studies performing the genetic analysis on resistances governed by QTLs, some of them have been proved to be recessive. For example, quantitative recessive resistance in a melon cultivar to Cucumber mosaic virus (CMV; Cucumovirus, family Bromoviridae) is conferred by polygenes including the major QTL cmv1. In melon, the major QTL wmv1551, together with three minor QTLs on different chromosomes, regulates recessive resistance against WMV. The major QTL qMrdd1, which confers recessive resistance against maize rough dwarf disease caused by a group of viruses in the genus Fijivirus, family Reoviridae, has also been mapped in maize. The major QTL rymv-QTL1 in rice modulates quantitative recessive resistance to Rice yellow mottle virus (RYMV; Sobemovirus, family Solemoviridae). Interestingly, rymv-QTL1 was mapped to the same locus as the monogenic recessive resistance gene rymv2, indicating that the same gene can be involved in quantitative and qualitative resistance.

Cloned Recessive Resistance Genes Plant viruses rely heavily on a number of host factors, also referred to as susceptibility factors, in host cells to replicate their genomes and spread into healthy tissues. In host cells, viruses must simultaneously deal with multiple defense responses by plant innate immune systems. Due to such inherent nature of virus infection strategies in host cells, recessive resistance is hypothesized to be genetically explained by loss-of-function (null) or point mutations of a gene encoding a host factor involved in viral infection, and of a negative regulator of defense responses. Both parts of this hypothesis appear to be supported by recent developments in the cloning and the characterization of recessive resistance genes found in natural plant germplasm (Table 2). Eukaryotic translation initiation factor 4E (eIF4E)-mediated recessive resistance has been observed in a wide range of plant species, including dicots and monocots. The possible involvement of eIF4E in plant viral infection was first documented by the isolation of an isoform of eIF4E (eIFiso4E) as an interactor of the VPg protein of Turnip mosaic virus (TuMV) in a yeast two-hybrid assay. eIF4E-mediated resistance has been discovered in various crop species via the candidate gene approach. Additionally, there are also distinct classes of the recessive resistance genes in terms of their intrinsic functions in host plants, including both dicots and monocots, against a wide range of viruses. Elucidation of the characteristics and mechanisms of the natural recessive resistance would create a foundation for the development of the new plant breeding strategies for virus resistance. Recent findings imply the existence of an unexpected regulatory mechanism for natural recessive resistance, rather than a simple working hypothesis

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Table 2

Plant Resistance to Viruses: Natural Resistance Associated With Recessive Genes

Cloned recessive resistance genes against plant viruses Locus or allele

Host plant

Resistance Gene

Avr gene

wlv cyv2 desc (¼ bc  3 & cyv) mo11 mo12 sbm1 ( ¼ wlv & cyv2) pvr6 ( þ pvr2) rpv1 pvr21 pvr22 pot  1 pvr1

Pisum sativum Pisum sativum Phaseolus vulgaris Lactuca sativa Lactuca sativa Pisum sativum Capsicum annuum Arabidopsis thaliana Capsicum annuum Capsicum annuum Solanum habrochaites Capsicum chinense

eIF4E eIF4E eIF4E eIF4E eIF4E eIF4E eIFiso4E cPGK2 eIF4E eIF4E eIF4E eIF4E

VPg P1, P3N-PIPO

retr01, retr02 retr03 rwm1

Brassica rapa Brassica rapa Arabidopsis thaliana

eIFiso4E eIF2Bß cPGK2

VPg

rym4

Hordeum vulgare

eIF4E

VPg

rym5

Hordeum vulgare

eIF4E

VPg

rym11

Hordeum vulgare

PDIL5–1

Cucumovirus - Bromoviridae Cucumber mosaic virus (CMV)

cmv1

Cucumis melo

VPS41

MP

Sobemovirus - Solemoviridae Rice yellow mottle virus (RYMV) Rice yellow mottle virus (RYMV)

rymv1 rymv2

Oryza sativa Oryza glaberrima

eIFiso4G CPR5

VPg

Carmovirus - Tombusviridae Melon necrotic spot virus (MNSV) Melon necrotic spot virus (MNSV) Melon necrotic spot virus (MNSV)

nsv – (EcoTILLING)

Cucumis melo Nicotiana benthamiana Cucumis zeyheri

eIF4E eIF4E eIF4E

30 UTR 30 UTR

Waikavirus - Secoviridae Rice tungro spherical virus (RTSV)

tsv1

Oryza sativa

eIF4G

Begomovirus Geminiviridae Tomato yellow leaf curl virus (TYLCV)

ty  5

Solanum lycopersicum

Pelota

Genus – Family/Virus (acronym) Potyvirus - Potyviridae Bean yellow mosaic virus (BYMV) Clover yellow vein virus (ClYVV) Clover yellow vein virus (ClYVV) Lettuce mosaic virus (LMV) Lettuce mosaic virus (LMV) Pea seed-borne mosaic virus (PSbMV) Pepper veinal mottle virus (PVMV) Plum pox virus (PPV) Potato virus Y (PVY) Potato virus Y (PVY); Tobacco etch virus (TEV) Potato virus Y (PVY); Tobacco etch virus (TEV) Potato virus Y (PVY); Tobacco etch virus (TEV); Pepper mottle virus (PepMoV) Turnip mosaic virus (TuMV) Turnip mosaic virus (TuMV) Watermelon mosaic virus (WMV) Bymovirus - Potyviridae Barley mild mosaic virus (BaMMV); Barley yellow mosaic virus (BaYMV) BaMMV; BaYMV; Barley yellow mosaic virus  2 (BaYMV  2) BaMMV; BaYMV; Barley yellow mosaic virus  2 (BaYMV  2)

CI, VPg, NIa CI, VPg, NIa VPg

VPg VPg VPg

explained by null mutation of a single susceptibility factor. A deeper understanding of recessive resistance in natural germplasm will be valuable for durable resistance applications in crop species.

Characteristics of eIF4E-Mediated Recessive Resistance The first molecular cloning of a naturally occurring recessive resistance gene revealed that eIF4E gene is responsible for pvr2-mediated resistance against PVY and TEV in pepper (Capsicum annuum). Although a number of susceptibility factors has been identified from naturally resistant accessions and artificially induced mutants, and as interactors of viral proteins in various plant species, the most-studied naturally recessive resistance genes remain eIF4E, its interacting partner eIF4G, and their isoforms (hereafter eIF4Es). The protein eIF4E is commonly involved in canonical translation of mRNA in eukaryotic cells. eIF4E physically interacts with eIF4G scaffold proteins to form the eIF4F translation initiation binary complex. This complex initially binds to the m7G cap structure at the 50 end of mRNA through the eIF4E subunit. Subsequently, eIF4G recruits several other initiation factors, such as poly(A)-binding protein (PABP); eIF4A, a helicase for the unwinding of RNA secondary structures; and eIF3, a multi-subunit complex that binds to the 40S ribosomal subunit (Fig. 1). The eIF4F complex induces the close interaction of both termini within the mRNA molecules in a process termed circularization, which promotes efficient translation.

Plant Resistance to Viruses: Natural Resistance Associated With Recessive Genes

75

Fig. 1 The eukaryotic translation initiation complex. eIF, eukaryotic initiation factor; PABP, poly(A)-binding protein; 40S, 40S ribosomal subunit. Reproduced from Robaglia, C., Caranta, C., 2006. Translation initiation factors: A weak link in plant RNA virus infection. Trends in Plant Science 11(1), 40–45.

Plants possess two isoforms of eIF4E: eIFiso4E, and novel cap-binding protein (nCBP). eIF4E and eIFiso4E form the distinct protein complexes eIF4F and eIFiso4F with eIF4G and eIFiso4G, respectively. Although these complexes can have partially redundant functions in the protein translation machinery, each complex selectively recruits specific mRNA species for translation. Genes encoding the components of the eIF4F complexes constitute a small gene families in plants. For example, in the model plant Arabidopsis, three genes (eIF4E1, eIF4E2, and eIF4E3) belongs to the eIF4E subfamily, one gene belongs to eIFiso4E; one gene belongs to eIF4G; two genes (eIFiso4G1 and eIFiso4G2) belongs to the eIFiso4G subfamily. Thus, the paralogous genes in each subfamily showing high degrees of sequence similarity are predicted to be functionally redundant in the translation machinery. The protein eIF4Es may have biological functions other than translation initiation. eIF4E and/or eIFiso4E have been deduced to be involved in the nuclear export of a subset of mRNA molecules. The eIFiso4F complex may regulate microtubule dynamics through the physical interaction of eIFiso4G with microtubules. In addition, each component of the two eIF4F complexes had been demonstrated to be a susceptibility factor for some plant viruses, especially those encoding VPg. In contrast, although nCBP has a sequence motif related to eIF4E and less cap-binding activity than eIF4E and eIFiso4E, its intrinsic function in the plant translation machinery and interacting partners remain unknown. However, nCBP has been recently reported to be a susceptibility factor for several potexviruses (family Alphaflexiviridae) in Arabidopsis, and of Cassava brown streak virus (CBSV) and Ugandan cassava brown streak virus (UCBSV; genus Ipomovirus, family Potyviridae) in cassava (Manihot esculenta). As mentioned above, more than half of recessively inherited resistance alleles are associated with potyviruses. Similarly, the majority of viruses influenced by eIF4E-mediated resistance are potyviruses. This dominance of potyviruses may be consistent with the relevance of this large virus group in agricultural production. Given the essential role of eIF4Es in translation and the lack of their own translational machinery for plant viruses, all plant viruses very likely utilize the eIF4Es for the translation of viral factors. Nevertheless, a limited range of virus species is impaired by mutation of one of these translation initiation factors, suggesting that some potyviruses have a fitness advantage in their infection cycles characterized by specific interaction with one of the eIF4Es possessing redundant functions. In contrast, as mentioned above, resistance to PVMV in pepper requires both pvr6 and pvr2-eIF4E. pvr6 encodes a loss-offunction variant of the eIFiso4E gene, suggesting that PVMV can recruit both eIF4E and eIFiso4E to establish infection. Except for the pvr6 resistance gene in pepper, almost all natural recessive resistance alleles related to eIF4Es possess only several point mutations in their coding regions, which do not interfere with the intrinsic function of eIF4E variants in plant cells. To date, several surveys of natural variation in eIF4E alleles in various crop cultivars indicated that the nonsynonymous substitutions correlated with the resistance phenotype are predominantly found at the non-conserved aa residues near the cap-binding pocket; these substitutions occurs in two adjacent regions on the surface of the predicted three-dimensional eIF4E structure. Functional resistance alleles with subtle aa substitutions are also known in other viruses affected by eIF4E-mediated recessive resistance, such as bymoviruses, RYMV, and Melon necrotic spot virus (MNSV). In pepper, lettuce, and pea, several allelic series show distinct resistance specificity against different virus species and pathotypes. These series represent multiple combinations of point mutations at a small number of residues, enabling multiple distinct resistance specificities. Another common feature of eIF4E-mediated resistance is that VPg, a viral protein covalently attached to the 50 end of viral RNA, has been annotated as an Avr factor of bymoviruses and sobemoviruses, and potyviruses. A few other viral factors, including the P1 cistron of ClYVV in pea and the cylindrical inclusion (CI) protein of LMV in lettuce, have been characterized as Avr proteins. Remarkably, the Avr determinant of MNSV for nsv-eIF4E–mediated recessive resistance in melon can be mapped to 30 capindependent translation enhancer (30 -CITE) at 30 untranslated region of the viral genome, but not to viral proteins. Interestingly, the incompatibility of the MNSV 30 -CITE with eIF4E may also explain non-host resistance against MNSV in Nicotiana benthamiana. The roles of eIF4Es in plant–virus interactions have long been studied using Arabidopsis. Similarly, Arabidopsis mutants of eIF4Es introduced by ethyl methane sulfonate (EMS) treatment or transposon knockout (KO) show phenotypes characterized by loss of

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Plant Resistance to Viruses: Natural Resistance Associated With Recessive Genes

susceptibility to several plant RNA viruses, including potyviruses, CMV, and Turnip crinkle virus (TCV; Carmovirus, family Tombusviridae). Such phenotypes are genetically equivalent to natural recessive resistance found among crop cultivars. However, no natural Arabidopsis ecotype deploying eIF4Es-mediated recessive resistance has been identified to date, implying that Arabidopsis has a greater fitness advantage during probable interactions with viruses relying on eIF4Es than conferred by recessive resistance mutation in eIF4Es. Functional specificities to eIF4Es of particular viral strains appear to be distinct between Arabidopsis and crop species. For instance, whereas TEV and LMV rely on eIFiso4E to infect Arabidopsis, they rely on eIF4E to infect pepper, tomato, and lettuce. These results highlight the significance of the basic research to reveal specific interactions among eIF4Es and viruses in crop species and model plants for the development of recessive resistance cultivars.

Molecular Mechanisms of eIF4E-Mediated Recessive Resistance Due to the early cloning of eIF4Es as recessive resistance genes, which have the most dominance among cloned resistance genes, the molecular mechanism of recessive resistance governed by eIF4Es has been the most intensively studied so far. The two main topics of this research area have been the molecular characterization of viral Avr factors, mainly the potyvirus VPg protein, and the functional analysis of natural allelic variation in eIF4Es in crops and wild relatives. These studies have provided profound insight into the principles and diversity of molecular mechanisms of eIF4E-mediated recessive resistance. In most cases, it is well known that eIF4E resistance alleles fails to physically interact with the potyvirus VPg protein (Fig. 2). Given that point mutations in eIF4E resistance alleles are mainly found near the cap-binding pocket, eIF4E-mediated recessive resistance against potyviruses is plausibly due to the disruption of eIF4E–VPg interaction. Point mutations of eIF4Es may abolish the recruitment by VPg, instead of the 50 cap structure of intrinsic mRNA for translation of the potyvirus genome. This hypothesis is consistent with the finding that eIF4G, as well as eIF4E, is a susceptibility factor for ClYVV in Arabidopsis, suggesting that the eIF4F complex is involved in potyvirus infection. However, a few inconsistencies were found in the correlation of eIF4E–VPg interactions with resistance phenotypes, implying the existence of another distinct mechanism for resistance against at least a few potyviruses that is functionally independent of such interaction. Alternatively, it is possible the methods used to detect eIF4E–VPg interactions, such as the yeast two-hybrid assay, may not perfectly represent these biological interactions in host cells. Intriguingly, each plant–virus pair shows diversity of infection steps and the extent to which viral infection is impaired in eIF4E-mediated resistance against potyviruses (Table 1). Different levels of recessive resistance against a virus are frequently observed on distinct alleles at a single locus in a host. This may also support the hypothesized diversity of potyvirus infection strategies, some of which are independent of eIF4E–VPg interaction. Alternatively, eIF4E may be involved in multiple steps of the potyvirus infection cycle. Thus, distinct eIF4E alleles may differentially change the eIF4E function at each step in a qualitative or quantitative manner, generating diversity within the resistance phenotype. The translation of viral proteins from genomic RNAs of PVY and TuMV had been proposed to depend on the internal ribosomal entry site (IRES), suggesting an alternative hypothesis that the eIF4E–VPg interaction does not directly control genomic RNA translation. Rather, VPg may cause host translation shutoff via interaction with eIF4E. Such activity promotes the release of translation initiation factors and/or ribosomes from host mRNA translation, thereby directing eIF4E to translate viral proteins. Since replication is tightly linked to translation during the plant viral infection cycle, eIF4E–VPg interaction had also been proposed to control replication, rather than translation. Indeed, the VPg region of the 6K2-VPg-Pro polyprotein has been demonstrated to be recruited into the viral replication complex (VRC) in TuMV-infected cells. A few exceptions to the role of VPg as an Avr determinant of recessive resistance have been found. For example, the breaking of mo11-mediated resistance is attributed to a single aa mutation of the LMV CI, which is involved in potyviral replication and systemic movement. Recessive resistance mediated by cyv2 is also overcome by an aa substitution of the ClYVV P1 cistron. In the sobemovirus RYMV, virulent VPg protein has been shown to interact with eIFiso4G, but not with eIF4E. These results indicate

Fig. 2 Predicted three-dimensional structural model of the PVY VPg (green) and eIF4E (blue) complex. W56 in eIF4E protein indicates the tryptophan residue that interacts with the cap structure. Nt, N-terminus; Ct, C-terminus. Reproduced from de Oliveira, L.C., Volpon, L., Rahardjo, A.K., et al., 2019. Structural studies of the eIF4E-VPg complex reveal a direct competition for capped RNA: Implications for translation. Proceedings of National Academy of Science of the United States of America 116, 24056–24065.

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qualitative differentiation in the mechanism of resistance mediated by eIF4E, even between closely related viruses. VPg may not be the sole interactor with eIF4Es; other viral factors together with VPg, including the polyprotein intermediates containing the VPg domain, may also play essential roles in this interaction. The eIF4Es-mediated recessive resistance has also been found to be functional against viruses with distinct genomic structures, such as the lack of VPg, clearly suggesting a resistance mechanism distinct from the one mediated by eIF4Es against VPg-containing viruses. As mentioned above, nsv-eIF4E–mediated resistance against MNSV is attributed to the 30 -CITE sequence, which is an Avr factor. A resistance-breaking strain of MNSV had been shown to gain the ability to interact with eIF4E from resistant melon. Although direct evidence for the molecular resistance mechanism remains lacking, eIF4E may facilitate circularization of MNSV genomic RNAs via interaction with 30 -CITE. Large-scale surveys of natural polymorphism of eIF4E alleles in several crops and their wild relatives have been conducted to explore the genetic variation and evolutionary patterns of eIF4E recessive resistance loci. KO mutations of eIF4E has not been found among natural accessions in these surveys. Although artificial development of an eIF4E KO mutant in Arabidopsis is possible, this may be due to the moderate greenhouse conditions normally used in Arabidopsis research. In a survey using the wild pepper chiltepin (Capsicum annuum var. glabriusculum), which is undergoing early domestication, eIF4E resistance alleles were frequently found in cultivated chiltepin populations. The same trend has also been reported for bymovirus resistance in barley, where possible eIF4E resistance alleles are predominantly encoded in domesticated barley accessions. In contrast, a survey of melon wild relatives successfully found several natural alleles of eIF4E genes, but no correlation of these alleles with the resistance phenotype. These findings indicate that human management of plant populations exerts profoundly strong selection pressures on the diversity and evolution of recessive resistance genes. Each potyvirus species has evolved to specifically recruit one eIF4E for viral infection, despite functional overlap among eIF4Es in the host translational machinery. Therefore, if possible, simultaneous KO of eIF4Es is desirable for the engineering of broadspectrum resistance against viruses. However, several lines of evidence suggest that double mutation of eIF4Es is harmful to plant development. In Arabidopsis, the eif4e/eifiso4e double-KO mutant shows a lethal phenotype; in tomato, functional repression of two eIF4E paralogues, eIF4E1 and eIF4E2, by RNA silencing induces severe dwarf phenotypes. Given the functional relevance of eIF4Es in host translation, it is plausible that simultaneous mutation of two eIF4Es cause severe defects in plant growth. Despite these results, recent studies have suggested that natural resistance alleles would solve the problem attributed to the intrinsic function of eIF4Es to create a broad resistance spectrum. Tomato pot-1–eIF4E1–mediated resistance, which is attributed to a small number of eIF4E1 aa mutations, impairs infection by PVY and TEV strains. In contrast, KO mutation of eIF4E1 in tomato yields a much narrower spectrum of resistance. Furthermore, the broad spectrum of resistance found in natural alleles can be reconstituted by artificially combining KO mutations of eIF4E1 and eIF4E2 in tomato with severe growth defects, indicating that eIF4E2 also has the capacity to participate in potyvirus infection, and that pot-1–eIF4E negatively regulates eIF4E2 in the potyvirus infection cycle without no detriment to plant growth. Consistently, overexpression of the eIF4E1 resistance allele in tomato provides dominant resistance to potyviruses, even in the presence of the intrinsic eIF4E1 susceptible allele. These results suggest that eIF4E1 resistance alleles negatively control eIF4E2 with redundant function to obtain a broad spectrum of resistance. This may coincide with the fact almost all eIF4E resistance alleles identified to date sustain only several point mutations, but not null mutations.

New Recessive Resistance Genes Found in Natural Diversity In the past decade, several distinct classes of recessive resistance genes other than eIF4Es have been discovered from a wide range of plant species. A few of these genes likely have functions similar to those of eIF4Es in recessive resistance, due to their known intrinsic functions in the translation machinery. Others appear to have a distinct recessive resistance mechanism, involving a negative defense regulator in recessive resistance. These newly identified recessive resistance genes show different specificities of resistance to plant viruses than those mediated by eIF4Es. Given candidate gene approaches have helped identify eIF4E resistance alleles in various crops, recent progress on the new recessive resistance genes will further promote to identify the resistance genes responsible for other plant–virus interactions, and will increase the potential of recessive resistance (in terms of the resistance spectrum and the gene repertoire) in the development of resistant cultivars in a wide range of crops. In barley, although the recessive resistance genes rym4/rym5, which are alleles of eIF4E, are frequently overcome by BaMMV and BaYMV resistance-breaking isolates, cultivars with the rym11 locus are highly resistant against both virus isolates. The rym11 locus has been demonstrated to encode an allele of protein disulfide isomerase like 5-1 (PDIL5-1), a distinct type of recessive resistance gene. A natural polymorphism survey of PDIL5-1 genes suggested that most rym11 resistance alleles are likely the results of frequent interactions with highly divergent forms of BaMMV and BaYMV in eastern Asia. PDIL5-1 is an endoplasmic reticulum (ER)–localized chaperone in plants and animals; it plays an essential role in the unfolded protein response (UPR), a cellular machinery that alleviates unfolded protein overload in ER. Although the mechanism of rym11-PDIL5-1–mediated recessive resistance remains unknown, downregulation of other UPR components commonly impairs efficient viral infection in several plant–virus pathosystems, implying that they are promising targets for recessive resistance to a wide range of viruses. Distinct classes of recessive resistance genes against potyviruses have also been cloned. Single-point mutation of eukaryotic translation initiation factor 2Bβ (eIF2Bβ) is responsible for the retr03 resistance locus against TuMV in allopolyploid mustard. eIF2Bß is a subunit of the eIF2B complex, which mediates guanine nucleotide exchange in eIF2 in the later step of eIF4E in translation initiation. The role of eIF2Bβ in TuMV infection, although still unknown, may be as a host factor because of the

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intrinsic function related to eIF4E in translation initiation. Similarly, a susceptibility factor EXA1 for potexviruses, identified from loss-of-susceptibility mutant of Arabidopsis, contains a putative eIF4E-binding motif. These findings imply that the factors functionally related to eIF4E are promising targets for recessive resistance. eIF2Bβ recessive resistance alleles are frequently found in several mustard cultivars, albeit those with allopolyploid genomes, suggesting the relevance of this gene in mustard–TuMV interaction. Another distinct type of recessive potyvirus resistance gene is nuclear-encoded chloroplast phosphoglycerate kinase 2 (cPGK2), validated in Arabidopsis ecotypes, independently designated WMV resistance loci rwm1, and rpv1 against Plum pox virus (PPV). cPGK2 gene homologs are found in a diverse range of plants, including dicots and monocots. Consistently, knockdown of cPGK genes in N. benthamiana compromises efficient accumulation of Bamboo mosaic virus (BaMV; Potexvirus, family Alphaflexiviridae). In addition, cPGK2 is predicted to recruit BaMV genomic RNA to chloroplasts for BaMV replication. Although the exact molecular mechanism of cPGK2-mediated resistance in Arabidopsis ecotypes remains uncertain, cPGK2 may be a general susceptibility factor for at least a subset of potyviruses and other distantly related viruses. A mutation in vacuolar protein sorting 41 (VPS41) gene has been demonstrated to be involved in resistance against CMV, mediated by the recessive allele cmv1 in melon. VPS41 plays a role in membranous vesicle trafficking from the late Golgi to the vacuole in eukaryotic cells. Intriguingly, in cucumber, the gene encoding another vesicle trafficking component, VPS4, has been proposed as a strong candidate of the recessive resistance locus zym against Zucchini yellow mosaic virus (ZYMV; Potyvirus, family Potyviridae). Although evidence for a specific molecular mechanism for cmv1-VPS41–mediated resistance is insufficient, several viruses appear to commonly recruit components of the vesicle trafficking machinery for efficient infection. In an African rice (Oryza glaberrima) accession that is resistant against Rice yellow mottle virus (RYMV; Sobemovirus, family Solemoviridae), the rymv2 resistance gene encodes an allele of the rice homolog of the Arabidopsis CPR5 gene, whose dominant form plays a negative role in the plant defense response. Seven other African rice accessions also encode rymv2 resistance alleles. Because of the CPR5 function in Arabidopsis defense responses, rymv2-CPR5–mediated resistance is highly likely the first example of the activation of defense responses by recessive mutation of a negative defense regulator gene. Remarkably, the recessive resistance locus ty-5 in tomato has also been identified to confer resistance to a plant DNA virus, Tomato yellow leaf curl virus (TYLCV; Begomovirus, family Geminiviridae). The ty-5 locus has been reported to encode an allele of Pelota, whose protein product is implicated in the recycling phase of translation, the latest step of protein biosynthesis. Although the resistance mechanism mediated by Pelota remains unclear, this result reinforces the relevance of the host translational machinery in recessive resistance despite significant differences in infection strategies among DNA and RNA viruses. Collectively, the identification of these distinct classes of recessive resistance genes will broaden the potential for recessive resistance application to the plant–virus pathosystem, for which this type of resistance has never been described.

Toward the Application of Recessive Resistance in Agriculture In the last ten years, due to the increase in the acquisition of genomic information and the development of techniques for the analysis of gene functions, greater numbers of studies have identified several classes of recessive resistance genes and provided mechanistic insight on recessive resistance. Loss of susceptibility to viral infection is genetically equivalent to natural recessive resistance. Therefore, random mutagenesis using crops and model plants would extend the potential to identify unknown susceptibility factors and find new ways to utilize host factors beyond the genetic variations found in naturally occurring cultivars. Model plants further facilitate the isolation of loss-of-susceptibility mutants and subsequent identification of responsible genes, due to the availability of high-quality whole-genome sequence, simple genetic approaches, and many other analytical tools. However, it should be noted that the generation of polyploid plant mutants with desired phenotypes by random mutagenesis is often difficult due to the presence of redundant genes. Conventional breeding techniques based on crossbreeding and molecular marker selection has long been used to introduce the recessive resistance phenotypes from one cultivar to another elite cultivar. This method has contributed to the alleviation of crop yield losses caused by devastating viruses. However, the stable introduction of resistant traits to establish resistant cultivars through conventional techniques or by the targeting of a specific gene using random mutagenesis and screening are substantially technically challenging and time consuming. Targeting induced local lesions in genomes (TILLING) is a technology applied to rapidly identify a mutant plant with genetic modification in a target gene by combining random mutagenesis by ethyl methanesulfonate (EMS) and single-nucleotide polymorphism (SNP) detection in a specific gene. TILLING has been successfully applied to establish a null allele of eIF4E gene in tomato showing loss of susceptibility to potyviruses. As discussed above, the range of resistance conferred by the null eIF4E allele is narrower than that of the natural functional allele pot1. Thus, at least when eIF4E is the gene of interest, functional resistance alleles are preferable to a null allele for obtaining a broad spectrum of resistance. EcoTILLING is a derivative of TILLING that is used to isolate the genetic variation of a target gene from that in natural cultivars. Using the EcoTILLING approach, several natural allelic variations of eIF4E gene have been found in wild relatives of melon; however, they do not correlate with potyvirus resistance phenotypes. Considering the results of other surveys of natural allelic polymorphism, EcoTILLING for natural resistance alleles may be performed more effectively with collections of crop cultivars, rather than their wild relatives. Significant technological and conceptual progress is currently being made in further promoting the application of recessive resistance and loss of susceptibility for resistant crop breeding. Sometimes, the elimination of an undesired trait that is tightly linked to the resistance phenotype using conventional breeding techniques is difficult. However, the recent emergence of

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next-generation sequencing (NGS) technology and genome editing could pave the way for the rapid introduction of recessive resistance phenotypes to crop cultivars as intended. NGS technology promotes access to the whole-genome sequences of all organisms. Genomics-based crop breeding combined with NGS is expected to enable the rapid establishment of new crop cultivars with desirable traits. NGS technology, which have rarely been applied to antiviral crop breeding using natural variants, would be quite useful for the identification of loci including major resistant QTLs (e.g., QTL-seq) in naturally resistant variants, as well as for crop breeding, to introduce resistant loci into elite cultivars. Genome editing technology is based on sequence-specific nucleases, including zinc-finger nuclease (ZFN), transcription activator-like effector nuclease (TALEN), and clustered regularly interspaced short palindromic repeat (CRISPR)-associated protein 9 (Cas9) in the system referred to as CRISPR/Cas. These technologies, which have emerged during the last two decades to enable mutagenesis and insertion at specific genomic sites in eukaryotic genomes, are now widely used in diverse crop species, as well as model plants. Targeted mutagenesis, made possible by genome editing, can overcome potential problems caused by existing technologies, such as the timeconsuming aspect of random mutagenesis and potential tight linkage of undesired traits in crossbreeding. One advantage of genome editing for crop breeding is that the original transgenes can be removed after editing via segregation. This feature permits the removal of bottlenecks for social acceptance of traditional genetically modified (GM) crops. The CRISPR/ Cas9 system was applied to modify eIF4E genes in Arabidopsis, cucumber, and rice. As expected, the resulting mutant plants, which were free of transgenes, exhibited recessive resistance to potyviruses and waikaviruses. Multiple gene editing has been demonstrated in model plants Arabidopsis and rice, and is thus possible in polyploid plants. Genome editing technologies can be also applied to develop loss-of-susceptibility mutants by modifying host factors identified through basic research on plant–virus interactions. In this sense, genes encoding host factors with loss-of-susceptibility mutations in Arabidopsis and other plant species can be regarded as potential recessive resistance genes. As a proof of concept, nCBP, known as an eIF4E isoform, was isolated in cassava as an interactor with VPg proteins of two ipomoviruses, CBSV and UCBSV. CRISPR-Cas9–mediated editing of two cassava nCBP genes successfully generated a resistant phenotype. Considering that nCBP is a susceptibility factor for potexviruses in Arabidopsis, nCBP gene is a promising target of genome editing to obtain recessive resistance with distinct spectrum from eIF4Emediated resistance. Furthermore, genome editing technologies are evidently compatible with the application of current knowledge of recessive resistance in antiviral resistance breeding of various crop species, including polyploids. Derived from the original CRISPR/Cas9 system, precise base editing has been accomplished using a new technology termed Target-AID, in which a nucleasedeficient Cas9 mutant is fused with activation-induced deaminase (AID), which catalyzes C to T mutations. This technology has been utilized to confirm that base editing of the eIF4E gene mimicking the pea sbm1 allele achieved recessive resistance against ClYVV in Arabidopsis, where no natural eIF4E resistance allele has been found. Conventionally, recessive resistance traits must be introduced into new crop species from closely related species that exhibit recessive resistance and can be crossbred. Since some crop species lack recessive resistance traits, random mutagenesis has been the only method for establishment of a resistant mutant in which a susceptibility gene is knocked out. However, in tomato, the eIF4E1 KO mutant shows a much narrower potyvirus resistance spectrum than does the natural pot1-eIF4E1 resistance allele. Considering this finding together with other results, researchers have hypothesized that the pot1 resistance allele, which maintains function in translation, negatively regulates eIF4E2 with redundant function in viral infections. Based on this insight, the researchers tested whether introgression of synthetic eIF4E (eIF4ER) alleles, rationally-designed from the pea sbm1 gene in trans-species manner, produces a broad resistance spectrum in the background of double KO of eIF4E/eIFiso4E (eif4eKO/eifiso4eKO) in Arabidopsis. The Arabidopsis genotype eIF4ER/eif4eKO/ eifiso4eKO confers no growth defect and shows a broad spectrum of resistance to several potyviruses, and more significantly, to WMV and one strain of Beet western yellows virus (BWYV; genus Polerovirus, family Luteoviridae), to both of which no recessive resistance has been documented in either KO mutant, eif4eKO and eifiso4eKO, in Arabidopsis. These results suggest the new crop breeding concept of developing a synthetic eIF4E allele that mimics natural resistance alleles from different species will retain intrinsic function in translation and shows a broad resistance spectrum in crops in which natural recessive resistance cannot be obtained by conventional breeding. However, it should be noted that the Arabidopsis genotype eIF4ER/eif4eKO/eifiso4eKO does not show recessive resistance against all viruses tested. Thus, to control the viruses for which recessive resistance alleles have not been found yet, the identification of natural resistance alleles for these viruses may be necessary.

Conclusions In this article, we addressed the characteristics of natural recessive resistance found in diverse plant species, the isolation and functional features of recessive resistance genes, and recent advances toward the application of recessive resistance in a wide range of crop species. Indeed, some of the recessive resistance genes summarized in Table 1 are currently utilized to control plant virus diseases in agriculture. Table 1 shows a strong bias toward recessive resistance against RNA viruses, perhaps due partly to research focus. Recent progress in the isolation of recessive resistance genes suggests the continuing relevance of eIF4Es as highly dominant recessive resistance genes, as well as the functional diversity of recessive resistance. Further research efforts should explore more diverse classes of recessive resistance genes in the near future. An unexpected regulatory mechanism of natural eIF4E-mediated recessive resistance has been proposed, such that a functional eIF4E1 resistance allele likely negatively regulates eIF4E2 with redundant functions. Recently emerged genome editing technologies are anticipated to be applied to the molecular breeding of recessive resistance in crops. In particular, the target-AID method will facilitate precise base editing, and is highly suitable for the design of a synthetic resistance allele mimicking natural resistance alleles in a trans-species manner.

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As also discussed in this article, genome editing technologies are clearly promising tools for the application of recessive resistance to various crop species. However, social acceptance of genome-edited crops is uncertain in many parts of the world, partly because evidence supporting the safety of genome-edited crops seems to be insufficient due to still developing technologies. Indeed, although some genome-edited crops have been approved without restriction in the USA, the Court of Justice of the European Union decided to make genome-edited crops subject to the same regulations as conventional GM crops. Further basic research is essential to enhance the utility of recessive resistance and support the safety of genome editing technologies for recessive resistance–based crop breeding.

Further Reading Bastet, A., Robaglia, C., Gallois, J.L., 2017. eIF4E resistance: Natural variation should guide gene editing. Trends in Plant Science 22, 411–419. Bastet, A., Lederer, B., Giovinazzo, N., et al., 2018. Trans-species synthetic gene design allows resistance pyramiding and broad-spectrum engineering of virus resistance in plants. Plant Biotechnology Journal 16, 1569–1581. de Oliveira, L.C., Volpon, L., Rahardjo, A.K., et al., 2019. Structural studies of the eIF4E-VPg complex reveal a direct competition for capped RNA: Implications for translation. Proceedings of National Academy of Science of the United State of America 116, 24056–24065. Fraser, R.S.S., 1990. The genetics of resistance to plant viruses. Annual Review of Phytopathology 28, 179–200. Hashimoto, M., Neriya, Y., Yamaji, Y., Namba, S., 2016. Recessive resistance to plant viruses: Potential resistance genes beyond translation initiation factors. Frontiers in Microbiology 7, 1695. Kang, B.C., Yeam, I., Jahn, M.M., 2005. Genetics of plant virus resistance. Annual Review of Phytopathology 43, 581–621. Robaglia, C., Caranta, C., 2006. Translation initiation factors: A weak link in plant RNA virus infection. Trends in Plant Science 11, 40–45. Sanfaçon, H., 2015. Plant translation factors and virus resistance. Viruses 7, 3392–3419. Truniger, V., Aranda, M.A., 2009. Recessive resistance to plant viruses. Advances in Virus Research 75, 119–159. Wang, A., Krishnaswamy, S., 2012. Eukaryotic translation initiation factor 4E-mediated recessive resistance to plant viruses and its utility in crop improvement. Molecular Plant Pathology 13, 795–803.

Plant Viral Diseases: Economic Implications Basavaprabhu L Patil, ICAR–Indian Institute of Horticultural Research, Bengaluru, India r 2021 Elsevier Ltd. All rights reserved. This is an update of G. Loebenstein, Plant Virus Diseases: Economic Aspects, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, https://doi.org/10.1016/B978-012374410-4.00470-2.

Nomenclature

ssRNA Single strand RNA SY Seedling yellows t Tonnes t/Ha Tonnes per hectare USA United States of America WTGs Whitefly-transmitted geminiviruses

Ha Hectares mHa Million hectares mt Million tonnes SP Stem pitting SSA Sub-Saharan Africa ssDNA Single strand DNA

Glossary Non-persistent manner A mode of virus transmission characterized by short acquisition and inoculation times (from seconds to minutes). Non-propagative manner A mode of virus transmission in which the virus is not internalized by the vector: the virus is carried externally by attachment to the mouthparts or foregut of the vector.

Pseudo-recombination Component exchange between two geminiviruses. The usual mechanism of pseudorecombination is by a process known as ‘regulon grafting’: the A component of a virus donates its common region by recombination to the B component of a second virus being captured. This results in a new dependent interaction between two components.

Introduction Estimates of losses as a result of pest or pathogen attack in crop plants are mostly lacking. Whatever meager amount of data is available is not necessarily accurate nor up-to-date. Such data from different countries assessed by different methods should be taken as only a rough estimate. Nevertheless, the available data gives an approximate idea on the tremendous losses to the food and fiber production for the increasing global population. During 1988–1990 a team of Scientists from Germany, supported by the European Crop Protection Association, published a comprehensive report on pest-induced crop losses on major food and cash crops. In the four principal cereal crops, rice, wheat, barley, and maize, the yield losses due to diseases were in the range of 10%–16% which translates to approx. $64 billion. When four more crops (potatoes, soybeans, cotton, and coffee) were taken into account the monetary estimate of yield loss increased to $84 billion, with an estimated harvests worth $300 billion in 1988–1990. However, this analysis did not cover some important food crops, such as cassava, millet, sorghum, and the horticultural crops, cultivated in developing countries, which are also significantly affected by viral diseases. It was also revealed that the pests and diseases accounted for pre-harvest losses of 42% of the potential value of output during 1988–1990, with 15% due to insect damage and 13% each to weeds and pathogens. An additional 10% of the potential yield was lost post-harvest. Among different plant pathogens, the fungal pathogens cause maximum damage with an estimated loss of 40%–60%, followed by the viral diseases causing a loss of 10%–15% and the rest of the losses are caused by bacterial pathogens, phytoplasma, and nematodes. The above data should be cautiously interpreted, as such survey data are mere rough estimates and probably the selection of crops was different and fewer viral diseases were considered. In several crops the yield losses caused by different viral diseases can range from 0% to 100% (Table 1). For example, in one of the surveys conducted in eight African countries, the yield losses because of cassava mosaic disease (CMD) were estimated to be in the range of 30%–40%. Barley yellow dwarf viruses (BYDVs) has a global distribution and infect over 150 species of the family Poaceae, including staple cereals such as wheat, barley, oats, rye, rice, and maize. A number of studies have demonstrated the devastating effect these viruses may have on cereals and the seed yield reductions are in the range of 15%–40% depending on the virus isolate and the cereal crop species. The economic impact caused by viral diseases is not reduction in yield alone, but also deterioration in the quality of the plant produce, such as deformations in peaches affected by Plum pox virus (PPV) or squash with Zucchini yellow mosaic virus (ZYMV) infection. In addition, major economic aspects to be considered are the expenses incurred in production of virus-indexed propagation material, eradication programs, control of insect vectors, and breeding for resistance to viruses. The viral epidemics and the subsequent economic impact are more prominent in tropical and subtropical regions with continuous vegetation round the year, which enables persistence and proliferation of the viruses and their vectors. In contrast, continuous herbaceous vegetation is interrupted during winters

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Table 1 Crop

Estimated yield or monetary losses due to viral diseases in different crop plants Country/Region

Cereals Barley, oat, Worldwide and wheat Worldwide Maize Africa USA Rice South and South East Asia

Wheat

Africa South America USA

Root and Tuber Crops Cassava Africa & Indian Subcontinent East Africa Potato Worldwide UK

Sweet potato

Worldwide Eastern Europe Eastern Europe Worldwide, East Africa in particular

Viral name (Acronym)

Virus taxonomy (Genus/Family)

Monetary/Yield losses

Barley yellow dwarf viruses (BYDVs) Cereal yellow dwarf viruses (CYDVs) Maize streak virus (MSV) Maize dwarf mosaic virus (MDMV) Rice tungro bacilliform virus (RTBV) and Rice tungro spherical virus (RTSV)

Luteovirus/Luteoviridae Polerovirus/Luteoviridae Mastrevirus/Geminiviridae Potyvirus/Potyviridae Tungrovirus/Caulimoviridae

Up to 50% yield loss 11%–12% yield loss 17%–71% yield loss 42%–75% yield loss US$1.5 billion

Rice yellow mottle virus (RYMV) Rice hoja blanca virus (RHBV) Wheat streak mosaic virus (WSMV) African cassava mosaic viruses (ACMVs), and East African cassava mosaic viruses (EACMVs) Cassava brown streak viruses (CBSVs) Potato leafroll virus (PLRV) Potato virus X (PVX), Potato virus Y PVY), and Potato leafroll virus (PLRV)

Potato virus X (PVX) Potato virus S (PVS) Potato virus M (PVM) Sweet potato feathery mottle virus (SPFMoV) and Sweet potato chlorotic stunt virus (SPChSV)

Waikavirus/Secoviridae Sobemovirus/Solemoviridae 84%–97% yield loss Tenuivirus/Phenuiviridae 25%–50% yield loss Tritimovirus/Potyviridae 27%–75% yield loss Begomovirus/Geminiviridae

US$1.2–2.3 billion per year

Ipomovirus/Potyviridae Polerovirus/Luteoviridae Potexvirus/Alphaflexiviridae

US$75 million 50%–60% yield loss US$5.5 million

Potyvirus/Potyviridae Polerovirus/Luteoviridae Potexvirus/Alphaflexiviridae Carlavirus/Betaflexiviridae Carlavirus/Betaflexiviridae Potyvirus/Potyviridae

Up to 80% Yield loss 5%–15% yield loss 3%–20% yield loss 40%–75% yield loss 50%–90% in combined infections

Crinivirus/Closteroviridae Vegetables Cucurbits Tomato

Fruits Banana

Cacao Citrus

Worldwide Worldwide Worldwide

Cucumber mosaic virus (CMV) Zuchini yellow mosaic virus (ZYMV) Tomato yellow leaf curl viruses (TYLCVs)

Cucumovirus/Bromoviridae Potyvirus/Potyviridae Begomovirus/Geminiviridae

Worldwide

Tomato leaf curl New Delhi virus (ToLCNDV)

Begomovirus/Geminiviridae

Worldwide

Tomato spotted wilt virus (TSWV)

Orthotospovirus/ Tospoviridae

Worldwide Worldwide

Banana streak viruses (BSVs) Banana bunchy top virus (BBTV)

Badnavirus/Caulimoviridae Babuvirus/Nanoviridae

Asia

Banana bract mosaic virus (BBrMV)

Africa Worldwide

Cacao swollen shoot viruses (CSSVs) Citrus tristeza virus (CTV)

Grape France Papaya Worldwide Stone fruits Worldwide Legumes Groundnut

Legumes Pigeonpea Soybean

Africa

Grapevine fanleaf virus (GFLV) Papaya ring spot virus (PRSV) Plum pox virus (PPV)

India

Groundnut rosette virus (GRV) Groundnut rosette assistor virus (GRaV) Groundnut bud necrosis virus (GBNV)

India/West Africa Worldwide India Worldwide

Peanut clump viruses (PCVs) Bean yellow mosaic virus (BYMV) Pigeonpea sterility mosaic viruses (PPSMVs) Soybean mosaic virus (SbMV)

10%–20% yield loss Up to 100% yield loss 93%–100% yield loss (in Jordan) 93%–100% yield loss (in Jordan) US$1 billion in 1994

Up to 82% yield loss US$50 million in India during 2007–2010 Potyvirus/Potyviridae 30%–70% in India & Philippines Badnavirus/Caulimoviridae up to 100% yield loss Closterovirus/Closteroviridae Since 1919, 80 million citrus trees eradicated Nepovirus/Secoviridae US$100 million Potyvirus/Potyviridae Up to 70% yield loss Potyvirus/Potyviridae US$10 billion during 1976–2006 Umbravirus/Tombusviridae Unassigned/Luteoviridae Orthotospovirus/ Tospoviridae Pecluvirus/Virgaviridae Potyvirus/Potyviridae Emaravirus/Fimoviridae Potyvirus/Potyviridae

US$100 million US$89 million US$38 million US$300 million (in India) US$300 million 8%–25% yield loss

Plant Viral Diseases: Economic Implications

Table 1

Continued Viral name (Acronym)

Virus taxonomy (Genus/Family)

Monetary/Yield losses

Cash Crops Cardamom India Cotton India/Pakistan

Cardamom mosaic virus (CarMV) Cotton leaf curl viruses (CLCVs)

Macluravirus/Potyviridae Begomovirus/Geminiviridae

Sugarcane

Sugarcane mosaic virus (SCMV)

Potyvirus/Potyviridae

70%–100% yield loss 68%–71% yield loss US$5 billion in Pakistan, during 1992–1997 30%–80% yield loss

Tobacco mosaic virus (TMV)

Tobamovirus/Virgaviridae

Crop

Tobacco/ Tomato

83

Country/Region

Worldwide except Guyana and Mauritius Worldwide

B1% yield losses in Tobacco and Up to 20% in Tomato

in the temperate regions and also in arid regions with dry hot summers, impacting negatively the vector population and subsequent virus transmission is significantly reduced. Furthermore the spread and prevalence of viral diseases is much higher in plants that are vegetatively propagated, such as cassava, potato, and citrus, than in those that are propagated through seeds. With a few exceptions, a majority of viruses are not seed transmitted, because of lack of vascular or the plasmodesmatal connection between the embryo and its parent plant. In vegetatively propagated plants, when the seed tubers, cuttings, bud wood, bulbs, runners, etc., are derived from infected plants viruses are transmitted to the subsequent generations. This is in addition to spread of the virus by vectors, which may occur in both cases. In the last two decades the whitefly transmitted viruses have been on the raise in several crops, including vegetables, often becoming a major constraint of production. In this article, it is not possible to cover the economic impact of all plant viral diseases, however we have made an effort to highlight the impact of most important virus diseases on economically important crops (Table 1).

Cereal Viral Diseases Globally wheat is cultivated in an area of 218 mHa with a total production of 772 mt. Diseases caused by viral pathogens are the major threat to early planted wheat in the southern Great Plains of United States (USA). The Wheat streak mosaic virus (WSMV, genus Tritimovirus, family Potyviridae) transmitted by wheat curl mite causes grain and forage yield losses that significantly affect profits from wheat production. The yield losses in wheat crop due to WSMV ranged from 27% to 75%. WSMV shows synergistic interactions with other viruses (specifically Triticum mosaic virus and High plains virus), further contributing to yield reductions. WSMV infects some of the most agriculturally important members of the family Poaceae, including wheat, corn, rye, oats, barley, sorghum, millets, and the grassy weeds that may act as alternate hosts. Barley yellow dwarf virus (BYDV; genus Luteovirus, family Luteoviridae), transmitted by aphids, is another important virus known to infect economically important crops such as barley, oats, wheat, maize, triticale, and rice. The yield losses caused by BYDV infection can vary widely depending on the viral strain, time of infection and rate of spread. Maximum yield losses up to 50% occur from early infections. Brome mosaic virus (BMV, genus Bromovirus, family Bromoviridae) is another economically important virus infecting barley and other cereal crops. BMV is also known to co-infect triticale along with WSMV. As per a study conducted in Ohio (USA), BMV could reduce the yield by as much as 61% in soft red winter wheat. Corn cultivated in a global area of 197 mHa, is also affected by more than 25 viral diseases. The total estimated corn loss (bushels) by different viral diseases in the United States and Canada, during 2012–2015 was estimated to be 23.6 million bushels and of this B46% was by Maize dwarf mosaic virus (MDMV, genus Luteovirus, family Luteoviridae) alone. Maize streak virus (MSV; genus Mastrevirus, family Geminiviridae) transmitted by six leafhopper species, is one of the most devastating viruses that occurs throughout Africa on corn. MSV epidemiology is controlled by environmental conditions on its vector, resulting in epidemics every 3–10 years. During the epidemic years, disease incidence can vary from a few infected corn plants per field, with little associated yield loss, to 100% infection rates and complete yield loss (Fig. 1).

Root and Tuber Crops Viral Diseases Cassava Viral Diseases Cassava is an important food security crop for small holder and marginal farmers of developing countries. Cassava is a climate resilient crop with exceptional drought tolerance and is a staple food for close to one billion people in the tropics. Globally, cassava is cultivated in an area of 26 mHa, with a total production of 292 mt and of this total area, 76% area is in African continent itself accounting for a total production of 178 mt. However, because of its vegetative propagation, cassava is affected by at least 20 different viruses worldwide. In sub-Saharan Africa, these viruses are persistently posing a big threat to food security, and resulting in a monetary loss of more than $1 billion annually. In particular, the two groups of whitefly transmitting cassava infecting viruses, namely begomoviruses (genus Begomovirus, family Geminivirdae) with single strand DNA (ssDNA) as their genome and Cassava brown streak viruses (genus Ipomovirus, family Potyviridae) with single strand RNA (ssRNA) as their genome,

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Fig. 1 Characteristic symptoms of important cereal viral diseases: (A) Wheat streak mosaic virus (WSMV) symptoms on wheat, (B) Barley yellow dwarf virus (BYDV) symptoms on barley, (C) Corn plant infected with a severe strain of Maize dwarf mosaic virus (MDMV), (D) Symptoms of Maize dwarf mosaic virus (MDMV) on corn leaf, (E) Corn infected with Maize streak virus (MSV), (F) Symptoms of Maize streak virus (MSV) on a leaf corn, (G) Rice tungro disease (RTD) on rice. (B) Source: Brian Olson, Oklahoma State University, Available at: Bugwood.org. (C) Source: https://krex.k-state.edu/dspace/bitstream/handle/2097/20917/KSUL0006Claflini37262.jpg?sequence=1&isAllowed=y. (E) Source: https://alchetron. com/Maize-streak-virus. (G) Source: IRRI.

have been responsible for the severe pandemics in Africa. The cassava infecting geminiviruses cause cassava mosaic disease (CMD) and the cassava infecting ipomoviruses cause cassava brown streak disease (CBSD). Currently, eleven distinct species of begomoviruses are involved in CMD, of these 9 are reported from Africa and 2 from the Indian subcontinent. In Africa, these 9 begomoviruses can be broadly categorized into African cassava mosaic viruses (ACMVs) and East African cassava mosaic viruses (EACMVs). ACMV and EACMV-like viruses can either occur singly or as mixed infections causing more severe symptoms. The yield losses in different cassava cultivars, in different African countries range from 20% to 95%. Estimates done in mid-1990s, revealed that the yield losses in the entire African continent was 12–23 mt of fresh edible roots per year, worth about $1.2–2.3 billion. Starting from 1988 till date, an unusually severe form of CMD pandemic is spreading across East and Central Africa, affecting food supplies and creating a famine in Uganda in the late 90s. In 1999, this CMD pandemic had affected at least nine countries in East and Central Africa, encompassing an area of 2.6 million square kilometers and causing an estimated loss of $1.9–2.7 billion per year. This severe form of CMD has been attributed at first to a recombinant strain of cassava mosaic virus. Later studies revealed that synergistic interactions between ACMV and a Ugandan strain of EACMV on mixed infection was mainly responsible for this severe form of CMD. The local cassava cultivars were highly susceptible to this recombinant strain and due to heavy losses many farmers abandoned cassava cultivation, resulting in destabilization of food security in Uganda. It was estimated that each year an area of c. 60,000 ha of cassava yielding 600,000 t of edible roots worth $60 million was lost due to CMD. Thus CMD was considered as the most damaging plant viral disease in the world, causing famine and death of thousands of people in Uganda which lost 90% of its production by 1998. In 2010, the Science journal published a report on seven most dangerous plant diseases in the world that can significantly affect the food security. Of all the plant diseases, CBSD was the only viral disease to be listed in this report. Although the first report of CBSD was made in 1930s from coastal region of Tanzania, in contrast to CMD, CBSD received much less attention, since it was restricted to the coastal low lying areas of East Africa, it appeared sporadically and did not cause any significant loss. However, post 2004, CBSD has been spreading across East and Central Africa at an alarming rate, with reports of CBSD incidence in highland areas too and hence risking the food security of millions of farmers in Africa. One of the estimates on incidence of CBSD, by National Agricultural Research Organization (NARO, Uganda), put the overall incidence at 16% in 2008 and 29% in 2009, clearly indicating the rapid expansion of the new CBSD epidemic. Shortly after the reports of spread in Uganda, similar observations were made in western Kenya and the Lake Victoria zone of Tanzania. A survey in 19 districts of Tanzania revealed a steep increase in CBSD incidence from 6% in 2006 to 12% in 2007 and in 2008 it was 32%.

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Most of the increase in disease incidence was attributed to greater levels of disease in districts in which CBSD was already present by 2006 (12 of 19 districts). In addition to the reduced growth of the cassava plants as a result of CBSD infection, significant losses occur due to spoilage of harvested roots due to necrotic rots. There have been very few quantitative assessments of yield losses and one such assessment done in southern coastal Tanzania revealed that the losses could occur to an extent of 70% in susceptible cassava cultivars. As the CBSD infected cassava plants mature, the root symptoms can further increase and hence farmers opt for early harvest to prevent further damage of roots. The effects on roots could be variable, such as: reduction in the quality of roots caused by pitting, constrictions and root necrosis, as well as yield reduction through reduction in the number and weight of tuberous roots. An estimate done in 2001, indicated an overall loss of 20%–25%, equivalent to a monetary value of $6–7 million. An effort was made to estimate the economic impact due to CBSD in Africa through development of a framework. This revealed that there are some of the undocumented elements of CBSD yield loss, such as additional labor cost incurred to separate necrotic from non-necrotic portions of affected roots and the deleterious effects on starch quality of non-necrotic portions of affected roots. A total loss of $75 million was estimated for all the eight countries of East and Central Africa affected by CBSD. For control of both CMD & CBSD, the main emphasis has been on the development of virus resistant cassava cultivars, phytosanitary practices such as using cuttings from healthy plants, elimination of infected plants, and production of virus-free tissue-cultured plant propagules. CMD can also be controlled by introgressing one of these three sources of resistance: polygenic recessive resistance, termed CMD1, the dominant monolocus type, named CMD2, and the recently identified CMD3 locus. Application of pesticides to stop the spread of CMD by controlling the whitefly vectors has not been successful.

Potato Viral Diseases Globally, potato is one of the major crops cultivated in an area of 19 mHa with a total production of 388 mt per year. Since potato is mainly vegetatively propagated, viral diseases are major constraint for cultivation of potato and the yields are reduced beyond 50%. Since potato is an economically important crop maximum research was devoted to it in the first half of the 20th century. Several new tools and techniques such as, serology, meristem cultures, and advanced microscopy were developed for management of potato infecting viruses. More than 37 viruses are known to naturally infect potato, with Potato virus Y (PVY; genus Potyvirus, family Potyviridae), Potato virus X (PVX; genus Potexvirus, family Alphaflexiviridae), Potato leafroll virus (PLRV; genus Polerovirus, family Luteoviridae), Potato virus S (PVS; genus Carlavirus, family Betaflexiviridae), Potato virus M (PVM; genus Carlavirus, family Betaflexiviridae), Potato aucuba mosaic virus (PAMV; genus Potexvirus, family Alphaflexiviridae), and Potato mop-top virus (PMTV; genus Pomovirus, family Virgaviridae) being the most economically important and widely spread. Additionally, Potato spindle tuber viroid (PSTVd; genus Pospiviroid, family Pospiviroidae) also cause major damages. PLRV, present across the globe, is reported to cause 50%–60% yield losses in potato and this virus is tuber borne and mainly vectored by aphids in a circulative and non-propagative manner. Whenever potato plants are co-infected with PLRV and PVX or PVY, yields are reduced to an extent of 70%. PVY first described in 1930s, has a global distribution and has been known for several decades as a threat to seed, ware and processed potatoes. In addition to potato it is also known to infect several other host plants in Solanaceae family, such as tobacco, tomato, and pepper, causing yield losses up to 80%. PVY is transmitted by more than 40 aphid species in a non-persistent manner. The isolates of PVY are highly variable at the biological, serological, and molecular levels. Due to lack of source of resistance to PVY and its plant-to-plant transmission through daughter tubers, the main control strategy used is primarily based on certification of seed production. A severe group of PVY strains named as PVYNTN are reported from Europe, Japan, and the US, and cause a severe form of disease in potato. The infected tubers develop superficial rings that later are sunken and necrotic. During storage these necrotic damages often become more conspicuous, and can affect 90% of tubers in susceptible cultivars. In spite of the latest improvements in diagnostic methods, routinely applied methods are unable to accurately characterize isolates responsible for tuber necrosis. Currently there is no serological assay, which can distinguish between PVYNTN and other PVY strains. Hence, there is no efficient means to manage the risks of epidemics caused by the emerging necrotic variants of PVY. The global potato market is worth several billion dollars and highly competitive, with poor understanding of the PVY-host interactions involved in induction of necrotic symptoms and inefficient diagnostic tools have led to a situation in which necrotic strains of PVY can potentially cause huge agronomic and economic losses. Furthermore, when PVY occurs along with PVX the yield losses are much more drastic. Crop yields can be further reduced by 5%–15% whenever plants are secondarily infected by PVX and PVS. PVM and PVS are common in countries of Eastern Europe. PVM can result in 40%–75% loss in tuber yield and it generally occurs in complex with other viruses (especially PVS). While with infection by PVS isolates tuber yield may be reduced by 3%–20%. Plants propagated from infected stocks, for several generations, may suffer more severe losses. Thus, for example, during the years 1993–1995, potato yields in Kazakhstan were extremely low, averaging 9 t/Ha. In a pilot project funded by USAID, micro-tubers obtained from meristems of potatoes screened for virus, were used, which resulted in a low infection rate in the elite seed tubers. These elite seeds enhance the yields by about 90% when compared with tuber yields from commercial fields in the neighborhood. Potato yields in erstwhile Soviet Union countries were low, averaging 11 t/Ha during 1993–1995 as there was no reliable system for providing certified virus-free tuber seeds, and farmers used seeds from their own farms. Reduced yields are mostly due to virus diseases carried over in the planting material. Thus, Loshitsky and Belorusky 3N cultivars developed by the Belarusian Research Institute for Potato Growing (BRIP) yielded 30–50 t/Ha in the experimental fields, while Belarus’ average potato yields

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were in the range of 12.2 and 16.3 tonnes per hectare. Assuming that the viral diseases in potato cause 20% damage, that would be translated to a total loss of 18 mt of potato in former USSR with 6 mHa under potato cultivation, with a total production of 70 mt. For any estimate for monetary implication, it is also important to consider the expenses involved in seed potato certification schemes. The main objective of such schemes is to produce true-to-type disease-free (mainly viruses) potato seed tubers. Such schemes mostly include in vitro produced plantlets for the initial increase of pathogen-tested clonal selections, including production of mini-tubers. Price of certified tuber seeds are almost double the price for ware potatoes.

Sweet Potato Viral Diseases Sweet potato (Ipomoea batatas) is cultivated in both tropical and warm temperate regions and is the seventh most important food crop in terms of production. In 2017, globally sweet potato was cultivated in an area of 9 mHa, with a total production of 113 mt, with China as the leading producer. Sweet potato is called ‘poor man’s crop’, since most of the production is done on a small or subsistence level and can be cultivated without much inputs such as fertilizers and irrigation, and is one of the crops that can survive the famines. Sweet potato produces more biomass and nutrients per hectare than any other food crop in the world. Sweet potato is a major crop in East Africa and thousands of villages depend on sweet potato for food supply and the Japanese use it when typhoons demolish their paddy fields. Yields vary greatly in different areas or even fields in the same region. The average yields in African countries is about 4.7 t/Ha, while the yields in Asia are significantly higher, averaging to 18.5 t/Ha. Such large variation in yields is attributed to variation in quality of the propagation material. Sweet potatoes are vegetatively propagated from vines, root slips (sprouts) or tubers, and generally the farmers in African countries use vines from their own fields for propagation year after year. Thus, use of virus infected material for propagation results in reduced yield and often these fields are infected with several viruses, thereby compounding the effect on yields. Some of the viruses that are reported to infect sweet potato are Sweet potato feathery mottle virus (SPFMV; genus Potyvirus, family Potyviridae), Sweet potato mild mottle virus (SPMMV; genus Ipomovirus, family Potyviridae), Sweet potato latent virus (SPLV; genus Potyvirus, family Potyviridae), and Sweet potato chlorotic stunt virus (SPCSV; genus Crinivirus, family Closteroviridae). Infection by both SPFMV and SPCSV causes the very severe sweet potato virus disease (SPVD) and it is the most important disease of sweet potato in Africa. SPVD can cause 50%–80% yield losses in Uganda and Kenya. In a 3-years field study in Cameroon, SPVD incidence resulted in a yield loss of 56%–90%, especially in the susceptible cultivars. Similarly, 78% yield reductions have been reported from field trials in Nigeria due to SPVD. Yield reduction of 50% were reported from a 2-year field experiment in Israel, in plots planted with SPVD-infected cuttings, while infection by SPFMV or SPCSV alone did not have any significant effect on yield. In China, the average yield losses due to sweet potato viral diseases are of over 20% and the infection rate in the Shandong province can range from 5% to 41%. In countries where virus-free planting material is provided, for instance in the USA and Israel, markedly higher yields of 16.3 and 30 t/Ha respectively, are obtained. Cucumber mosaic virus (CMV) alone does not infect sweet potato plants that are healthy and virus free, however, superinfection by CMV can cause a complete crop failure when sweet potatoes are carrying SPCSV. Most effective way to control viral diseases of sweet potato is through supply of virus-indexed healthy planting material, which is being practiced in Israel and in the Shandong province of China. Planting virus-tested material increased yields by at least by 100% in Israel, while in China the yield increase ranged from 22% to 92%. In Israel use of certified material costing 3 US cents per cutting, prepared by special nurseries, is a common practice. Initially, farmers buy about 30% of planting material required to plant their fields and later use the cuttings from this to fill in the rest. Yields are high and stable when virus-free material is used. In African countries virus-indexing programs operate only on a limited scale, since sweet potatoes are mainly grown as a food security crop, and not as a commercial one. Resistance breeding programs might be an answer for introgressing SPVD resistance and such programs are in operation in Uganda. One has to see if these improved sweet potato cultivars have stable, durable and broad spectrum resistance. An example for unstable resistance was reported when several clones resistant to SPFMV from the International Potato Center (CIP, Peru) were found to be susceptible when Israeli and Ugandan isolates of SPFMV were used for screening (Fig. 2).

Vegetable Viral Diseases Chilli Viral Diseases Chilli or hot pepper (Capsicum annuum L., family Solanaceae) is economically most important and valuable spice and vegetable crop throughout the world. Chilli is native to New Mexico and the Portuguese introduced chillies in Indo-Pak subcontinent from Brazil in 16th century. India is the world’s largest producer, consumer and exporter of chillies in the world, contributing 40% of the total world production. Other major chillies producing countries are China, Bangladesh, and Peru. Chilli is vulnerable to several pathogens, including the viral diseases and till date 65 viruses are reported to infect chilli resulting in heavy yield losses. Mostly, the viruses are spread or transmitted by infected seeds, insects and or by the mechanical tools. Most of the viral diseases are difficult to diagnose due to significant overlap of symptomatology. Symptoms produced by different viruses are mosaic pattern on leaves, yellowing, ring spots, leaf deformation, curling of leaves, and stunting of plants. Common viral diseases are; Alfalfa mosaic virus, Beet curly top virus, Cucumber mosaic virus, Pepper mottle virus, Tobacco mosaic virus, Chilli leaf curl virus, and Pepper mosaic virus, which are transmitted by aphids and whitefly vectors. Chilli leaf curl viral disease

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Fig. 2 Characteristic symptoms of important root and tuber crops viral diseases: (A) African cassava mosaic virus (ACMV), on cassava, (B) East African cassava mosaic virus (EACMV), on cassava leaf (Courtesy of James Legg), (C) Cassava brown streak virus (CBSV), on an old cassava leaf, (D) Cut roots of cassava infected with Cassava brown streak virus (CBSV), (E) Potato virus Y (PVY); symptoms on potato leaves, (F) Potato virus Y (PVY); symptoms on potato tuber, (G) Necrosis induced by Potato virus Y (PVY); on potato leaf, (H) Sweet potato virus disease (SPVD) on sweet potato (in the red circle) and healthy plants. (A) Source: https://www.labmanager.com/news/2019/04/gene-editing-technology-may-produceresistant-virus-in-cassava-plant#.XgOqPy2ZPyk. (B) Courtesy of James Legg. (C) Courtesy of Claude Fauquet. (E) Source: https://cpb-us-e1. wpmucdn.com/blogs.cornell.edu/dist/5/6962/files/2018/02/Severe-mosaic-symptoms-2g3v8aq.jpg. (F) Source: Bruce Watt, University of Maine, Available at: Bugwood.org. (G) Source: Bruce Watt, University of Maine, Available at: Bugwood.org. (H) Source: http://biosafetykenya.go.ke/images/ download/conferencematerials/6thannualpresentations/Proposal-presentaion-GMO-NBA-presentation.pdf.

(ChiLCVD, genus Begomovirus, family Geminiviridae) persistently transmitted by whiteflies, is the most destructive virus disease in terms of incidence and yield loss, causing a yield loss of 53% or more. The disease can be identified by typical upward leaf curling, crinkling, puckering and reduction in leaf area along with stunting of whole plants. It is transmitted by the whitefly Bemisia tabaci in persistent manner. The first report of Chilli leaf curl disease virus in India was in 1954, which was later reported in frequent intervals. However, after 2005 the virus complex has been emerging rapidly across the Indian subcontinent.

Cucurbit Viral Diseases Worldwide, cucurbits are known to be infected by large number of viruses causing significant economic losses. Cucurbit viruses represent a complex and changing pathosystem and cucumber alone is affected by at least 153 viruses. Virus infection in cucurbits generally result in three broad categories of symptoms: (1) Mosaic in leaves, often associated with deformations: leaf size reduction, rugosity, and enations; (2) Yellows, generally affecting the older leaves and later spreading to younger leaves, often accompanied by leaf thickening and reduction in fruit production, but not the fruit quality; (3) Necrosis as necrotic spots on leaves or as generalized necrosis resulting in death of plant, that occur in certain virus/strain and host/cultivar combinations, particularly with potyviruses. Fruits generally don’t mature and may develop necrotic symptoms. Often in the field there are mixed infections of two or more viruses, that lead to combination of above three symptom types. Cucumber mosaic virus (CMV, genus Cucumovirus, family Bromoviridae), transmitted by aphids, known since beginning of 20th century, is the first mosaic disease reported in cucurbits in the U.S. CMV causes most significant economic losses in melon, cucumber and squash, but is rarely found in watermelon. CMV has a broad host range, that infects more than 775 different plant species representing more than 86 different families. Two major CMV subgroups have been identified based on the serological and molecular properties. A diversity of CMV strains are also reported that differ in symptomatology, host range, virulence towards resistance genes, and aphid transmissibility. Epidemics of CMV were found to be very rapid in Southern Europe and generalized in melons, while in zucchini and squash were often slow to develop symptoms and limited to a few plants.

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Epidemics are occasionally reported from temperate regions such as north-eastern U.S. or central Europe. Several distinct species of the Potyvirus genus (family Potyviridae) infect cultivated cucurbits. Three of them, Papaya ringspot virus (PRSV), Watermelon mosaic virus 2 (WMV2), and Zucchini yellow mosaic virus (ZYMV) have a global spread, and cause significant yield reductions in melon, cucumber, squash, and watermelon. Epidemics of PRSV are common in tropical and subtropical regions where the wild cucurbit species act as reservoirs for the virus. While, WMV2 has been reported more than 50 years ago from across the world. Zucchini yellow mosaic virus (ZYMV) is one of the first cucurbit 'emerging' viruses that threatened the cucurbit industries across the globe in the early 1980s. In addition to mosaic and deformations on leaves and fruits of all cultivated cucurbits, various "bump" shapes are observed on mature fruits, making them unmarketable. First reported from Italy and France, within a decade’s time ZYMV has spread to the major cucurbit producing regions of the world. ZYMV has also colonized new regions or islands. Economic effect of the virus can be dramatic, early infections can result in complete yield loss. LIYV is another major virus prevalent in cucurbits, lettuce, and sugarbeets. Despite the atypical shift in vectors populations, LIYV caused major economic losses due to its prevalence in lettuce, sugarbeets, and melons. In 1981 alone, the estimated losses were US$20 million. Squash leaf curl virus (SCLV) and Watermelon chlorotic stunt virus (WmCSV) are transmitted by B. tabaci in a persistent manner are two other important viruses that cause significant damage to cucurbits.

Okra Viral Diseases Okra or bhendi or ladies finger (Abelmoschus esculentus) is an economically important vegetable crop grown in tropical and subtropical parts of the world and India is the leading producers of okra in the world. However, cultivation of okra is constrained by very high incidence of Bendhi yellow vein mosaic virus (BYVMV) disease, transmitted through whitefly. BYVMV infects the crop at all growth stages and causes 50%–100% yield loss, as well as reduction in quality, if the plants get infected within 20 days after germination. In India, Bhendi yellow vein mosaic disease (BYVMD) is one of the most economically important diseases of bhendi/okra and is caused by a monopartite begomovirus Bhendi yellow vein mosaic virus (BYVMV; genus Begomovirus, family Geminiviridae) associated with Bhendi yellow vein betasatellite (BYVb). The first report of BYVMV was from Mumbai in 1924.

Tomato Viral Diseases Globally, tomato (Solanum lycopersicum) is cultivated in an area of 4.8 mHa, with a total production of 182 mt and with an estimated value of over $40 billion in 2017. In terms of economic value, tomato shares approx. 70% of the total value of fresh vegetables produced worldwide. At least 136 different viruses are known to infect tomato, whereas this number is significantly lower for other vegetable crops, for example, 49 viruses are reported to infect pepper (Capsicum annuum), 46 infect melon (Cucumis melo), 53 infect lettuce (Lactuca sativa), 54 infect potato (S. tuberosum), and 44 infect eggplant (S. melongena). Only cucumber (C. sativus) is infected by more viruses than tomato (153). Years of intensive breeding for yield enhancement in tomato, ignoring the resistance traits has reduced the genetic base for virus resistance in cultivated tomato. Further intensified cultivation of tomato, with large areas under tomato mono-cropping might have favored proliferation of tomato viruses and their vectors. Begomoviruses and other whitefly-transmitted viruses are invading into new regions, and other emerging viruses such as new tospoviruses and Tomato torrado virus (ToTV; genus Torradovirus, family Secoviridae) are rapidly spreading over larger geographic areas. In this article, we will restrict to economic impact caused by tomato leaf curl begomoviruses and Tomato spotted wilt virus (TSWV). The spotted wilt disease of tomatoes was first reported from Australia in 1915 and later demonstrated to be caused by a virus and transmitted by thrips. The causative agent, was named as Tomato spotted wilt virus (TSWV; genus Tospovirus, family Bunyaviridae). TSWV causes a range of symptoms, such as chlorotic/necrotic rings and flecking on leaves, stems and fruits. Early infection of TSWV can lead to one-sided growth, drooping of leaves reminiscent of vascular wilt, and stunting or death of the tomato plant. Subsequently, these infections produce unmarketable fruits with conspicuous chlorotic/necrotic ringspots. Control of the TSWV vector thrips is almost impossible due to their high fecundity, propensity to develop insecticide resistance and due to their vast host range. TSWV can replicate within the thrips vectors, which is a characteristic feature of many membrane-bound bunyaviruses. The worldwide dispersal of the TSWV vector Western flower thrips (Frankliniella occidentalis) resulted in re-emergence of TSWV as a major agricultural pathogen in 1980s. In 1994, the monetary losses by menace of TSWV, across the globe was estimated to be over $1 billion annually. Till date TSWV is considered to be economically the most important virus due to its global presence and a broad host range with ability to infect more than 800 plant species, including tomato, pepper, lettuce, peanut, and chrysanthemum. Tomato yellow leaf curl disease (TYLCD) is another major constraint for cultivation of tomato. TYLCD is widely distributed in Asia, sub-Saharan Africa, the Caribbean Islands, Australia, and the USA. More than 57 different geminivirus species generally called as Tomato yellow leaf curl virus (TYLCV, genus Begomovirus, family Geminivirdae) are reported to infect tomato. In the United States the whitefly-transmitted geminiviruses (WTGs) appeared first in early 1990s, accounting for 20% estimated yield loss. Whereas in the Latin American countries, namely, Brazil, Cuba, Guatemala, Mexico, and Costa Rica, yield losses were in the range

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Fig. 3 Characteristic symptoms of important vegetable viral diseases: (A) Tomato yellow leaf curl virus (TYLCV) on tomato, (B) Tomato spotted wilt virus (TSWV) on tomato fruits, (C) Tomato spotted wilt virus (TSWV) on tomato leaf, (D) Okra yellow vein mosaic virus (OYVMV) on okra, (E) Chilli leaf curl disease (ChiLCD) on chili, (F) Zucchini yellow mosaic virus (ZYMV) on Cucurbita pepo, fruits (left) and leaf (right). (A) Source: https://plantpath.ifas.ufl.edu/plant-virus-profiles/images/Picture1.jpg. (B) Source: Whitney Cranshaw, Colorado State University, Available at: Bugwood.org. (C) Source: Whitney Cranshaw, Colorado State University, Available at: Bugwood.org. (F) Source: http://www.pestnet.org/fact_sheets/ zucchini_yellow_mosaic_202.htm.

of 30%–100%. The estimated losses during 1989–1995 in Dominican Republic were to the level of $50 million. Another study has revealed that the Tomato leaf curl viruses (ToLCVs) cause an annual yield loss of $140 million in Florida, USA. Although it is possible to chemically control the whiteflies, but this may not be possible when there are large population sizes of whiteflies. However, under protected cultivation, physical barriers such as greenhouses enveloped by net screens are effective, while ‘floating barriers’ of perforated polyethylene sheets stretched over tomato fields prevent the landing of whiteflies. Such measures, increase the costs of cultivation and it may be difficult for small and marginal farmers to take up such protected cultivation. Hence, cultivation of virus resistant/tolerant tomato varieties is economically more feasible and breeding programs for introgression of virus resistant traits are in progress. Tomato leaf curl New Delhi virus (ToLCNDV) (genus Begomovirus, family Geminiviridae) is a major constraint in tomato production and is economically most important disease affecting tomato in Indian subcontinent. The first report of association of ToLCNDV with tomato leaf curl disease on solanaceous crops was from India in 1995. However, in the last few years ToLCNDV has significantly widened its host range and has moved to new geographical areas, including the Middle East, the western Mediterranean Basin, and some of the European countries. Recombination and pseudorecombination are the major factors for diversification and adaptive evolution of ToLCNDV. Emergence and epidemics of ToLCNDV has resulted in increased yield losses in several important crops, including tomato. Recent reports from Tunisia, Italy, and Spain indicate infection of ToLCNDV isolates in several cucurbit species. Sequence analysis revealed that a novel strain of ToLCNDV (ToLCNDV-ES) is spreading in the western Mediterranean region, which probably evolved through recombination from a highly diverse population of ToLCNDV. Several reports have demonstrated the involvement of monopartite begomoviruses in the major disease complexes affecting various crops in the “Old World”. However, ToLCNDV a bipartite begomovirus is the only exception to this, which causes diseases in a number of plant species of the families Solanaceae, Cucurbitaceae, Euphorbiaceae, Fabaceae, and Malvaceae. Furthermore in addition to vegetable crops, ToLCNDV has been reported to cause damage to fiber crop species and was also found to infect a number of weeds. Currently, ToLCNDV is known to infect 43 different plant species in Bangladesh, India, Indonesia, Iran, Italy, Malaysia, Pakistan, Sri Lanka, Spain, Taiwan, Thailand, and Tunisia. Co-existence of satellite DNAs (betasatellites) along with ToLCNDV is also reported from natural infections and are known to modulate its pathogenesis, including host range (Fig. 3).

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Fruit Virus Diseases Banana Viral Diseases Banana is an important source of carbohydrate and is cultivated in more than 130 countries. Banana is also a major source of income for millions of farmers from tropical regions of the world. In 2017, total global area under banana cultivation was 5.6 mHa, with a total production of 114 mt. The Asian continent has the largest area under banana cultivation with more than 50% share in total world area. In 2011, the total export value of banana was estimated to be $895 million. Banana is propagated vegetatively through the use of suckers and this practice has been one of the main cause for outbreaks of several diseases and pests around the world, especially the viruses. At least 20 different viral species belonging to five different families are known to infect banana globally. The economically most important viruses are: Banana streak viruses (BSVs; genus Badnavirus, family Caulimoviridae), Banana bunchy top virus (BBTV; genus Babuvirus, family Nanoviridae), Banana bract mosaic virus (BBrMV; genus Potyvirus, family Potyviridae), and Cucumber mosaic virus (CMV, genus Cucumovirus, family Bromoviridae). Worldwide, Banana bunchy top disease (BBTD) is the most devastating disease caused by BBTV and transmitted by the aphid vector Pentalonia nigronervosa. This disease was first reported in 1889 from Fiji and an outbreak of BBTD occurred in 1953 resulting in loss of 641,000 bunches. Later, BBTD has spread to more than 36 countries in South Pacific, Asia, and Africa. However, there are no records of BBTD in the New World except in Hawaii (USA). In the 1920s, due to the bunchy top disease epidemics, the Australian banana industry in New South Wales had collapsed, and the total area under banana cultivation was reduced by 90%. Similarly, in Southern Queensland banana production was reduced by more than 95%. In India, BBTD is known to cause heavy losses. One of the most significant BBTD outbreaks was in highly prized hill banana cv. Virupakshi, in the state of Tamil Nadu (India), which reduced the cultivation area from 18,000 to 2,000 Ha. Several epidemics of BBTD during 2007–2010 in Koduru, Andhra Pradesh, and Jalgaon, Maharashtra, India, led to an annual loss of $50 million. An outbreak of BBTD in Pakistan led to huge reduction in area under banana cultivation from 60,000 to 26,000 Ha. None of the farmer’s cultivated banana varieties are resistant to BBTV and production of disease-resistant bananas by conventional breeding is very difficult. Hence, transgenic development of BBTV resistant bananas is the most viable option to achieve resistance to BBTD. Banana streak disease caused by Banana streak virus (BSV) is also one of the important viral disease of banana which has a wider presence in banana plantations throughout the world. The streak disease was first observed in the Nieky Valley on the Ivory Coast in 1958 and later it was reported from Morocco where almost every established plantation was affected. Banana streak disease is now reported to occur in over 43 countries of Africa, Asia, Australia, Europe, Oceania, and tropical America. Another viral disease called Banana bract mosaic disease, caused by the Banana bract mosaic virus (BBrMV), was first reported in most of the banana cultivars, from Philippines (island of Mindanao) in 1979 and thought to be different from all other recognized viruses of banana. Later, it was found widespread in entire Philippines. Subsequently the banana bract disease was discovered in other Asian countries including India, Sri Lanka, Samoa, Thailand, and Vietnam. In Latin America, BBrMV was first reported from Colombia. However, there are limited reports available on the economic impact of the disease. In 1997, up to 40% yield losses were reported from Mindanao island of the Philippines. Yield losses in the range of 30%–70% are also reported from India and the Philippines.

Cacao Swollen Shoot Disease Cacao swollen shoot disease (CSSD) caused by Cacao swollen shoot virus (CSSV; genus Badnavirus, family Caulimoviridae), was first reported from Ghana in 1936 and is endemic to West Africa covering Togo, Ivory Coast and Nigeria. CSSD is semi-persistently transmitted by mealy bug (Planococcoides njalensis) vectors and this disease reduces cacao yield in first year of infection by almost 25%, in second year the yields are reduced by 50%, and in next 3–4 years the tree dies from die back. CSSD economically most important disease, since cacao has high industrial value and this disease has resulted in drastic reduction in cocoa production in Ghana. Since its first report in 1936, more than 200 million cacao trees have been eradicated from about 130,000 Ha. Yield of cacao bean were reduced by 86% during 1945–1950. In 1953, the 50 million diseased trees represented a capital depreciation of d25 million since the report of CSSD. During 2006–2010, over 28 million infected trees were removed and globally today CSSV is responsible for 15% of total cacao crop loss. This disease caused much rural disconnect and political uproar in Ghana. In response to the menace caused by CSSD, Ghana has launched one of the most ambitious eradication program with high investments. In the late 1940s, the British colonial authorities in Ghana enforced an eradication policy for removal of infected cacao trees. This met with a serious opposition, resulted in the rise of a political party led by Kwame Nkruma, who accused the authorities of attempting to destroy farmers’ income. Some of the strategies employed to control this disease are: use of insecticides, biological control of the mealybugs, breeding for resistant cocoa cultivars, and cross-protection with mild strains of CSSV, but the main practice is to rogue the infected and adjacent trees. CSSV has several alternate hosts, such as Cola gigantean, and mere removal of infected trees would not solve the CSSD menace and it is important to remove the alternate hosts too.

Citrus Tristeza Virus Citrus tristeza virus (CTV) (genus Closterovirus, family Closteroviridae) is the causal agent of devastating epidemics that changed the course of the citrus industry. There are mainly three distinct syndromes caused by CTV, named as tristeza, stem pitting (SP), and

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seedling yellows (SY), which is due to differential interaction of CTV with different scions and rootstocks from different regions. Tristeza disease is a decline syndrome caused by infection of CTV on different citrus species such as sweet oranges, mandarins, grapefruits and limes, which are propagated on rootstocks derived from sour orange or lemon. During 1940–1960, about 30 million citrus trees grafted on sour orange rootstocks were lost in Argentina and Brazil. In 1980s, about 6.6 million trees were lost in Venezuela, and B10 million trees were lost in Florida and other Caribbean countries. In these regions CTV is mainly transmitted by the brown citrus aphid, Toxoptera citricida. In Spain alone CTV took a toll of 10 million citrus trees and several million more were lost in California (USA), Israel, and other areas. In these areas CTV is being spread by the melon aphid, Aphis gossypii. In the presence of both the aphid vectors, the spread of CTV is greatly enhanced, as in the case of Florida (USA). Significant yield losses have been documented from several regions and the monetary losses have been estimated. However, these estimates do not include the expenses incurred on removal of dead trees and replanting with healthy plants. Despite the loss of millions of citrus trees in Brazil and Spain, in the long run the citrus production was not affected. In Brazil, in a span of about 40 years, the citrus production increased from 1.7 to 20 mt in 1990s. This would not have been possible without replanting new orchards on tolerant rootstocks, along with budwood control. Some of the strategies employed for control of CTV damage are quarantine and budwood certification programs, rouging of infected trees, use of tristeza-tolerant rootstocks, or cross protection with mild isolates. The choice of the management practice mostly depends on CTV incidence, the virus strains present and host varieties cultivated in an area. Introgression of virus resistance genes into cultivated genes through conventional breeding is not feasible, while genetic engineering for virus resistance has given variable results. Long distance spread of CTV occurs through movement of infected planting material and then short distance local transport is facilitated by different aphid species and is not known to be seed transmitted. Infection of CTV is mostly restricted to species of two genera of Rutaceae family and the virus is then localized to the phloem cells. The most devastating epidemics of CTV occurred in Argentina (1930), Brazil (1937), California (1939), Florida (1951), Spain (1957), Israel (1970), and Venezuela (1980). Outbreaks of tristeza have also been reported from Cyprus (1989), Cuba (1992), Mexico (1995), the Dominican Republic (1996), and Italy (2002). In addition to loss of trees, CTV is also indirectly associated with loss of sour orange root stocks with very good agronomic traits. Further, the use of tristeza-tolerant rootstocks has led to the emergence of new problems such as damage from soil salinity or alkalinity, waterlogging, or vulnerability to soil fungi or graft-transmissible pathogens such as citrus blight, Citrus sudden death-associated virus or Citrus tatter leaf virus. Regardless of the rootstock used, the affected trees showed stunting, SP, low yield and poor quality of fruits. Isolates of CTV causing SP, that were initially restricted to regions of Asia, Australia, South Africa, and South America, were subsequently reported from California, Florida, and the Mediterranean region, although at lower frequency. Implementation of mandatory certification program for maintenance and distribution of virus-free material for commercial use is the best way to control CTV or any other graft transmissible disease. One such publicly operated self-supportive program called ‘clean stock program’ in Florida charge a fee of $2 for registration of validated or parent trees. Cross-protection is widely practiced in Brazil, Australia, and South Africa. In particular, this has been highly successful in Brazil for CTV strains causing SP on sweet orange and grapefruit. By 1980, over 8 million trees of Pera sweet orange were cross-protected in Brazil and this is still practiced. In California, an eradication program for tristeza is in place and during 2000–2001 the Central California Tristeza Eradication Agency surveyed approximately 31,400 acres of citrus orchards and carried ELISA tests for about 375,000 individual plants. Undoubtedly this program deferred the spread of tristeza, but huge costs were incurred to remove the sick trees. In Israel, during 1970–1977, a tristeza suppression program was in place, wherein about 300,000 tests were made using indicator plants and electron microscopy and eventually 1700 trees diagnosed with CTV infection were removed. Later in 1979–1980, indexing of 1.25 million of citrus trees by ELISA revealed CTV infection in only 0.13% of the tested trees. Hence such eradication programs delayed the outbreak of a citrus tristeza pandemic in Israel by 15–20 years. Eighty-five years of epidemics of CTV have made a devastating economic impact on the citrus farms and the industries dependent on it. Worldwide, CTV has taken a toll of more than 80 million citrus trees, mostly in South Africa since 1910, Argentina (10 million) and Brazil (6 million) since 1970, and the US. (3 million) since 1950. Dispersal of CTV is a continuous process and it is feared that the future losses from tristeza may be much more than the losses that have occurred till date. There is also a rapid dispersal of the CTV vector T. citricida in Mexico and the Mediterranean basin, with large area under citrus cultivation using the sour orange rootstock, could facilitate the spread of tristeza. The replacement of declining citrus trees with new trees on tristeza tolerant rootstocks can only restore the production of citrus. In the last 40 years, new isolates of SP are being periodically reported in new regions and is going to be a more enduring problem. Cross-border movement of new citrus varieties has resulted in invisible spread of severe SP isolates and is likely to continue despite the quarantine measures.

Papaya Ring Spot Virus Worldwide papaya is cultivated in an area of 0.44 mHa with a total production of 13.2 mt and 45% of the total production is contributed by India alone. Several pests and pathogens are known to affect papaya (Carica papaya) plants, of which viral diseases are the most damaging ones. Of the several viral diseases Papaya ringspot virus (PRSV, genus Potyvirus, family Potyviridae) inflicts maximum damage to papaya and has a worldwide presence. Other viruses that are known to infect papaya are geminiviruses causing leaf curl symptoms, Papaya meleira virus (PMeV; Unassigned in the family Totiviridae), Papaya mosaic virus (PapMV;

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genus Potexvirus, family Alphaflexiviridae), and Papaya lethal yellowing virus (PLYV; genus Sobemovirus, family Solemoviridae). PRSV is transmitted by several different aphid species in a non-persistent manner. There are no known resistance genes against PRSV in papaya, but tolerance to PRSV is conferred in a quantitative manner. In 1950s, Oahu island in Hawaii (USA) was the main papaya-growing area, however because of the heavy PRSV infection, papaya cultivation was shifted to fertile lands of Puna district (Hawaii, USA) where PRSV was absent. By 1970s, 95% of Hawaiian papaya were cultivated in Puna, but a potential threat of PRSV outbreak remained, since Hilo was just 30 km away. In 1992, for the first time PRSV was reported from commercial plantings of papaya in Puna district on Hawaii island. Despite all the efforts to minimize the spread of PRSV, more than 50% of Puna was severely affected by PRSV by 1994. Subsequently the Hawaii Department of Agriculture (HDOA) abandoned all the eradication programs. By 1998, most of the papaya plants in Puna were PRSV infected and papaya production was reduced to about half of the 1992 level (12,000 vs 24,000 t of fruits). In 1992, Puna produced 95% (24,000 t) of Hawaii’s papaya; but after the entry of PRSV in that region, the production decreased significantly and by 1998 Puna produced only 75% (12,000 t) of Hawaii’s production. Papaya production in Puna was reduced to half in 1998, compared to 1992. Clearly, PRSV had a devastating effect on papaya production in 1998. To maintain its markets, especially in mainland of USA and Japan, Hawaii had to keep up production as much as possible. In addition to significant yield losses, PRSV infection severely affected the quality of the produce and the papaya fruits harvested were of inferior quality. Transgenic papaya overexpressing the coat protein sequence of PRSV were developed by researchers from University of Hawaii (USA) and released as papaya varieties “SunUp” and “Rainbow”. These varieties are now widely cultivated in the Hawaii islands of United States and is the most successful field application of transgenic technology. Since the release of PRSV resistant transgenic papaya, the papaya production in Puna showed an increasing trend, starting from 15,400 t in year 2000 to 18,300 t in 2001. Furthermore, papaya production in Puna accounted for 84% of the production in 2002, compared to the low of 65% in 1999. In 2004, Rainbow accounted for 88% of the total area of papaya in Puna (Hawaii, USA).

Plum Pox Virus (Sharka) Disease of Stone Fruits Plum pox disease, also called as “sharka” is one of the most devastating viral diseases of stone fruits resulting in huge yield losses and a major economic impact. This disease is caused by Plum pox virus (PPV), a positive ssRNA virus and belonging to the genus Potyvirus and family Potyviridae. In addition to plums (Prunus domestica), PPV also infects other stone fruits such as, peaches (Prunus persica), apricots (Prunus armeniaca), nectarines (Prunus persica var. nucipersica), almonds (Prunus dulcis), sweet cherries (Prunus avium) and tart cherries (Prunus cerasus). PPV also infects important ornamental and wild Prunus species, including those used in traditional medicine: myrobalan, American plum, dwarf flowering almond, and blackthorn. Further, approx. 60 plant species representing eight families are recognized as experimental host plants of PPV. Nevertheless, PPV also infects plants that are not members of genus Prunus. The first report of sharka disease was from plum orchards in Bulgaria during 1915–1918. During 1932–1960, the disease disseminated to north and east from Bulgaria into Yugoslavia, Hungary, Romania, Albania, Czechoslovakia, Germany, and Russia. Initially, sharka disease was mainly found in plums and apricots and since the 1980s has been reported in peaches. About a decade ago, about 100 million stone fruit trees in Europe were reported to be infected. The susceptible varieties could result in 80%–100% yield losses. In Eastern and Central Europe, susceptible plum cultivars exhibit premature fruit drop and bark splitting, and sweet cherry fruits develop chlorotic and necrotic rings and fruit drop occurs prematurely. Although the fruit trees don’t die from PPV infection, but the fruits drop prematurely and are unfit for consumption and marketing due to their bitter taste. In 1968, the estimated yield losses due to sharka disease in Bulgaria was 30,000 t. In last several decades, the fruit drop production was to an extent of 80% in Czech Republic, and there was a drastic reduction in number of plum trees from 18 million to 4 million. During 1998–2002, due to PPV infection, about 69,000 stone fruit trees were removed in Emilia–Romagna region of Italy; and about $450,000 is spent annually for the removal and replanting of trees. An expenditure of $17 million was incurred for removal of B1.5 million trees in Spain during 1988–2008. PPV is predominantly present in south east of France and all the symptomatic trees are eliminated and if infection levels are more than 10%, the entire orchard is destroyed. Such operations have resulted in removal of about 27,000 trees in 1992 and 100 ha of plantations (mainly peach) were eliminated in 1993. During 1973–1990, about 91,000 trees were estimated to be destroyed in France. PPV is transmitted by aphids in a non-persistent manner, however the spread in Europe is mainly due to movement of infected nursery material from the Balkans. Vegetative propagation of host plants is the main cause for efficient dissemination of PPV both locally and globally. General susceptibility of hosts, with scarce resistance sources identified, has largely frustrated international resistance breeding efforts. For most of the 20th century PPV was limited to Europe, however in the past 20 years PPV infection has been reported from Africa, South & North America, and Asia. Currently almost all the major plum production areas are now affected by PPV at varying degrees. Huge efforts and costs are involved in the control and eradication of PPV in several countries. Though successful in a few cases, such efforts have generally slowed the progression of PPV onslaught, but certainly has not stopped it. In 2006, the combined costs for control of Sharka disease were estimated to be $10 billion over the past 30 years worldwide. Large number of efforts are made for genetically engineering PPV resistance in plums, resulting in successful field trial of the PPV resistant HoneySweet transgenic plum. Deregulation and cultivation of HoneySweet transgenic plum in USA, is one of the few example of virus-resistant transgenic crops developed to marketability (Fig. 4).

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Fig. 4 Characteristic symptoms of important fruit viral diseases: (A) Cocoa swollen shoot virus (CSSV) on young cocoa tree branches and leaf, (B) Citrus tristeza virus (CTV) on citrus tree, (C) Stem pitting symptom induced by Citrus tristeza virus (CTV) on a citrus branch, (D) Banana bunchy top virus (BBTV) on banana, (E) Banana streak virus (BSV), on banana leaf, (F) Papaya ring spot virus (PRSV), symptoms on a tree, (G) Papaya ring spot virus (PRSV), symptoms on a leaf, (H) Papaya ring spot virus (PRSV), symptoms on a fruit, (I) Plum pox virus (PPV) on several fruits and leaves of plum, apricot and peach. (A) Source: wikimedia. (B) Source: L. Navarro, Instituto Valenciano de Investigaciones Agrarias, Available at: Bugwood.org. (D) Source: Rui map Zheng, Available at: Bugwood.org. (H) Source: https://caes.ucdavis.edu/news/articles/ 2014/01/science-provides-facts-for-hawaii-gmo-debate. (I) Source: European and Mediterranean Plant Protection Organization, Available at: Bugwood.org.

Legume Viral Diseases A wide range of legumes (Fabaceae: Papilionoideae) are cultivated across the globe as an important source of proteins. These legume crops are severely affected by a range of viral diseases that result in reduced grain yield, poor quality seeds and increased cost of cultivation for phytosanitary activities and disease control, thus making significant impact on economic losses. The majority of the legume infecting viruses are transmitted by insect vectors, and many of them are also seed transmitted. Viral diseases are the major biotic constraints to legumes production, especially in the tropical and subtropical regions. Cultivated food legumes are vulnerable to infection by at least 150 viruses, belonging to different plant viral genera/families. Begomoviral diseases alone in legumes is known to cause an annual estimated yield losses of $300 million in India. The economic impact of viral diseases in most important legumes/pulse crops, such as soybean, groundnut, cowpea, pigeonpea, and mungbean are listed in Table 1. Worldwide, about 70 different viral species are reported to infect soybean, of which, Soybean mosaic virus (SbMV; genus Potyvirus, family Potyviridae), Tobacco ring spot virus (TRSV; genus Nepovirus, family Secoviridae), Peanut bud necrosis virus (PBNV; genus Tospovirus, family Bunyaviridae), Tobacco streak virus (TSV; genus Ilarvirus, family Bromoviridae), Soybean dwarf virus (SbDV; genus Luteovirus, family Luteoviridae), and some begomovirus species are considered to be economically important. SbMV is reported to cause a yield loss in the range of 8%–50%, and can reach up to 100% in severe outbreaks and when coinfected with Bean pod mottle virus (BPMV; genus Comovirus, family Secoviridae), Cowpea mosaic virus (CPMV; genus Comovirus, family Secoviridae), Abutilon mosaic virus (AMV; Begomovirus), and TRSV. Several different begomovirus species are associated with the soybean yellow mosaic disease (SYMD). In India alone, the combined yield losses due to SYMD were estimated to exceed $300 m in mungbean, black gram, and soybean.

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At least 32 viruses are reported to naturally infect groundnut, of which TSWV, PBNV, TSV, Peanut clump virus (PCV; genus Pecluvirus, family Virgaviridae), Indian peanut clump virus (IPCV; genus Pecluvirus, family Virgaviridae), PeMoV, PStV, CMV, and Groundnut rosette disease (GRD) caused by a complex of 3 viral agents, Groundnut rosette assistor virus (GRAV), Groundnut rosette virus (GRV, genus Umbravirus, family Tombusviridae), and a satellite RNA, are considered to be economically important. GRD is the most damaging disease in sub-Saharan Africa (SSA) and many GRD epidemics were reported from Africa, which has caused enormous crop losses. Spotted wilt disease of groundnut caused by TSWV, can lead to 100% yield loss and this disease is reported from North and South Americas, several African countries, and Australia. In 1998, a similar disease has been reported from Asia, but was shown to be caused by PBNV, a distinct tospovirus. Groundnut bud necrosis disease (GBND) first reported from India, is also prevalent in Sri Lanka, Nepal, Myanmar, Thailand, and parts of China. GBND is caused by a distinct tospovirus, PBNV (synonym Groundnut bud necrosis virus, GBNV). PBNV also infects mungbean, urdbean, soybean, cowpea, pea, and lablab bean under field conditions. Stem necrosis disease (SND), previously assumed to be caused by PBNV, was later found to be caused by TSV in India. TSV is known to occur on groundnut in India, Pakistan, South Africa, and Brazil and has a broad host range that includes sunflower (Helianthus annuus), soybean, mungbean, black gram, cowpea, and sunnhemp (Crotalaria juncea). TSV continues to expand its host range. In year 2000, SND was prevalent in nearly 225,000 Ha of erstwhile Andhra Pradesh state of India, which resulted in crop losses exceeding $65 million. Peanut clump disease (PCD) is prevalent in India, Pakistan, and West Africa, and the causal virus is referred as IPCV in Indian subcontinent and PCV in Africa, both of which cause indistinguishable symptoms. In 2003, worldwide the economic losses due to PCD was estimated to exceed $38 million. Both IPCV and PCV, infect pigeonpea and other economically important cereal crops. Since the casual viruses are seed transmissible in groundnut, cereals, and millets, PCD has quarantine importance. Cowpea is most widely cultivated legume in SSA, and is the second most important food legume after groundnut and more than 80% of cowpea production is from West Africa. More than 140 viruses are reported to infect cowpea and of these 20 viruses are known to be economically important. Except CGMV (Cowpea golden mosaic virus) and CCMV (Cowpea chlorotic mottle virus), all the other cowpea viruses are known to be seed transmitted. Pigeonpea is another important grain legume crops, predominately cultivated in the Indian subcontinent, and also in Southern and Eastern Africa, the Caribbean, and China. About 15 viruses are reported to infect pigeonpea, of these, Pigeonpea sterility mosaic emaravirus (PPSMV, genus Emaravirus, family Fimoviridae) is most important, followed by the whitefly-transmitted begomoviruses. Sterility mosaic disease (SMD) caused by PPSMV is a serious constraint for pigeonpea cultivation in South Asia, with an estimated annual loss of over $300 million in India alone. Of several viruses infecting mungbean and urdbean, yellow mosaic caused by begomoviruses, leaf curl caused by PBNV, and leaf crinkle caused by Urdbean leaf crinkle virus (ULCV) are economically most important. In India, both Mungbean yellow mosaic Indian virus (MYMIV) and ULCV are prevalent in both mungbean and urdbean, and are often synergistic leading to yield losses over 90%. Viruses that are transmitted through legume seeds serve as primary source of virus in virus ecology and disease epidemiology in cultivated legumes and have quarantine importance. Further the multiplication and spread of insect vectors is facilitated by intensive cropping and changes to cropping patterns as a result of increased irrigation and overuse of pesticides. Occurrence and the drastic expansion of the host range of tospoviruses and TSV in particular are major concerns. Some of the strategies employed for management of legume viruses are: selection and planting of virus-free seeds, modification of agronomic practices, chemical, physical, and biological control of viral vectors, and planting of virus-resistant crop varieties developed through conventional or non-conventional crop improvement strategies (Fig. 5).

Cash Crops Viral Diseases Sugarcane Viral Diseases Sugarcane is the largest crop production in the world with more than 1800 mt in 2017 on a surface of nearly 26 mt in the world. Worldwide, sugarcane is affected by diverse number of viruses, mainly because of its vegetative propagation. Sugarcane mosaic disease caused by Sugarcane mosaic virus (SCMV; genus Potyvirus, family Potyviridae) is economically the most important of all and is seen in almost all the major sugarcane-growing countries. The first report of sugarcane mosaic was in 1892, and was subsequently reported from other countries growing sugarcane, maize, and sorghum, namely, Bangladesh, Indonesia, Thailand, Malaysia, China, the United States, Pakistan, Argentina, Australia, Brazil, Cameroon, Colombia, Congo, Cuba, Denmark, Ethiopia, Egypt, France, Germany, Iran, Israel, Mexico, the Netherlands, and Vietnam. The first report from India was in 1921, since then, it invariably infected all the sugarcane cultivars. During 1914–1916, a severe epidemic of sugarcane mosaic occurred in Argentina, resulting in up to 80% losses in sugar production. Later, this crisis was overcome by replacing susceptible varieties of sugarcane with the resistant ones (POJ varieties) from Indonesia (Java). In mid-1920s, epidemic of sugarcane mosaic threatened the sugar industry in both Brazil and the United States, resulting in severe economic crisis in the sugar industry. Subsequently, the disease was controlled by replacing the susceptible varieties with mosaic resistant hybrids. However, later in 1971–1972, it was revealed in Brazil that the mosaic was found in tolerant hybrids with 100% infection, causing a loss of 18% in susceptible clones. Despite the report of sugarcane mosaic in India, it was ignored initially, however later, it was reported to have severe impact on popular sugarcane varieties in the field. Studies showed that the virus infection significantly reduced the number of millable canes, net CO2

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Fig. 5 Characteristic symptoms of important legume viral diseases: (A) Groundnut rosette disease (GRD) on groundnut (center) and healthy plants (left and right), (B) Bean com mon mosaic virus (BCMV) on bean leaf, (C) Bean yellow mosaic virus (BYMV) on bean leaf, (D) Bean golden mosaic virus (BGMV) on bean, (E) Mungbean yellow mosaic disease (MYMD) on mungbean, (F) Soybean mosaic virus (SbMV) on soybean. (A) Source: http://www.dpvweb.net/dpv/showfig.php?dpvno=345&figno=03. (B) Source: https://www.invasive.org/browse/detail.cfm? imgnum=5364007. (C) Source: Dr. Parthasarathy Seethapathy, Tamil Nadu Agricultural University, Available at: Bugwood.org.

assimilation rate, cane growth parameters and other post-harvest juice quality parameters. Extensive surveys conducted at different sugar factories in Indonesia, revealed that about 30% of sugarcane fields were affected by the mosaic disease. Sugarcane bacilliform virus (SCBV; genus Badnavirus, family Caulimoviridae) causing leaf fleck disease in sugarcane was first reported from Morocco and now it shows its presence in all the major sugarcane-growing countries. Studies revealed that the S. officinarum clones recorded more than 90% SCBV incidence, and other Saccharum species also showed the presence of SCBV. SCBV infection is presumed to have originated from Papua New Guinea (Southeast Asia), the center of sugarcane origin, and from there, it might have spread to other areas through the germplasm movement. SCBV is distributed throughout the world in both tropical and subtropical regions and can result in economic losses in the range of 10%–90% in severely infected crops. Sugarcane yellow leaf virus (SCYLV; family Luteoviridae) causing yellow leaf disease (YLD) is also one of the major threat to sugarcane cultivation worldwide and the commercial sugarcane cultivars of major sugarcane-growing countries are reported with 100% yield incidence in Florida, Brazil, India, etc.

Ornamentals and Orchids Viral Diseases In 2004, the global floricultural produce was estimated to be $75 billion, for which Netherlands, USA and Japan were the major contributors. Virus infections in ornamental plants have enhanced the esthetic value in certain cases as well as have resulted in significant monetary losses in several other cases. One of the first uses of esthetic effects of plant virus infection was the production and marketing of highly valued variegated tulips, infected with Tulip breaking virus. Other popular examples are the attractive mosaic patterns on the leaves of Abutilon infected with Abutilon mosaic virus, the variegated flowers of camellia infected with Camellia yellow mottle virus, and the yellow-veined Japanese honeysuckle infected with Honeysuckle yellow vein mosaic virus. In contrast, there are also hundreds of viruses that are known to infect the ornamental plants and cause severe economic losses. Viral diseases reduce the esthetic value and marketability of ornamental plants. Large scale vegetative propagation through virus infected material such as cuttings, bulbs, and corms has significantly contributed for spread of viral diseases in ornamentals. Tospoviruses transmitted by the thrips are known to cause maximum damage to the ornamental plants and their incidence is increasing year by

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Fig. 6 Characteristic symptoms of important cash crops viral diseases: (A) Cotton leaf curl disease (CLCuD) on cotton, (B) Sugarcane mosaic virus (SCMV) on sugarcane leaf, (C) Tobacco mosaic virus (TMV) on tobacco plant, (D) Cymbidium mosaic virus (CymMV) on orchid plant, (E) Dendrobium mosaic virus (DMV) on orchid leaf, (F) Odontoplossum ring spot virus (ORSV) on orchid leaf. (A) Courtesy of Suresh Kunkalikar. (C) Source: https://burleytobaccoextension.ca.uky.edu-content-tobacco-mosaic-virus-tmv. (D) Source: http://www.orchidboard.com/community/ pests-and-diseases/92602-cymbidium-mosaic-virus.html. (E) Source: flicker. (F) Source: https://www.flickr.com/photos/scotnelson/33983505945.

year mainly because of international trade and movement of propagative material. TSWV and INSV (Impatiens necrotic spot virus) are the two most important tospoviruses infecting the ornamental plants. In India, Chrysanthemum diseases caused by Chrysanthemum virus B (CVB; genus Carlavirus, family Betaflexiviridae), Cucumber mosaic virus, Tomato aspermy virus and Gladiolus diseases caused by Cucumber mosaic virus (all three viruses; genus Cucumovirus, family Bromoviridae) are of economic importance. More than 30 viruses are reported to infect different orchids in various geographical locations. Orchid viruses generally produce mosaics, ringspots, chlorosis, chlorotic and necrotic sunken patches on the leaves, deformation and a drastic reduction in the size of flowers and overall stunting of the plant. The orchid plants, Phalaenopsis and Dendrobium in particular, are commercially valuable ornamental plants with a global market. Unfortunately, orchid plants are highly susceptible to viruses such as Cymbidium mosaic virus (CymMV; genus Potexvirus, family Alphaflexiviridae) and Odontoglossum ringspot virus (ORSV; genus Tobamovirus, family Virgaviridae), posing a major threat and serious economic loss to the orchid industry worldwide. Among the orchid viruses, CymMV and ORSV are considered most important due to their worldwide occurrence and severity of the symptoms in several orchid genera. They reduce the general vigor of the plant and affect the flower quality, thereby reducing the marketability and incurring severe economic losses. These two viruses also occur naturally as mixed infections in several orchid species (Fig. 6).

Further Reading Gonsalves, D., 1998. Control of papaya ringspot virus in papaya: A case study. Annual Review of Phytopathology 36, 415–437. Legg, J.P., Kumar, P.L., Makeshkumar, T., et al., 2015. Cassava virus diseases: Biology, epidemiology, and management. Advances in Virus Research 91, 85–142. Loebenstein, G., Thottappilly, G., 2003. Virus and Virus-Like Diseases of Major Crops in Developing Countries. Amsterdam: Elsevier/Kluwer Academic Publishers. Loebenstein, G., Katis, N., 2014. Control of plant virus diseases: Seed-propagated crops. Advances in Virus Research 90, 530. Loebenstein, G., Katis, N., 2015. Control of plant virus diseases: Vegetatively-propagated crops. In: Advances in Virus Research, first ed., 91. Elsevier, p. 332. Orke, E.C., Dehne, H.W., Schonbeck, F., Weber, A., 1994. Crop Production and Crop Protection: Estimated Losses in Major Food and Cash Crops. Elsevier. Patil, B.L., Legg, J.P., Kanju, E., Fauquet, C.M., 2015. Cassava brown streak disease: A threat to food security in Africa. Journal of General Virology 96, 956–968.

Plant Viral Diseases: Economic Implications

Patil, B.L., 2018. Genes, Genetics and Transgenics for Virus Resistance in Plants. Norfolk: Caister Academic Press, doi:10.21775/9781910190814. Scholthof, K.B., Adkins, S., Czosnek, H., et al., 2011. Top 10 plant viruses in molecular plant pathology. Molecular Plant Pathology 12 (9), 938–954. van der Zaag, D.E., 1987. Yield reduction in relation to virus infection. In: de Bolas, J.A., van der Want, J.P.H. (Eds.), Viruses of Potatoes and Seed-Potato Production. Wageningen: Pudoc, pp. 146–150. Waterworth, H.E., Hadidi, A., 1998. Economic losses due to plant diseases. In: Hadidi, A., Khetarpal, R.H., Koganezawa, H. (Eds.), Plant Virus Disease Control. St. Paul: American Phytopathological Society Press, pp. 1–13.

Relevant Websites http://faostat.fao.org FAOSTAT.

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Retrotransposons of Plants M-A Grandbastien, INRAE – French National Research Institute for Agriculture, Food and Environment, Versailles, France r 2008 Elsevier Ltd. All rights reserved. This is a reproduction of M.-A. Grandbastien, Retrotransposons of Plants, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00544-6.

Glossary Env Envelope protein of retroviruses and errantiviruses, containing a transmembrane domain and a host receptorbinding domain, and mediating formation of infectious virions and targeting of host receptor cells. Env-like Coding domains possessing some features similar to those of the env domains of retroviruses (such as a transmembrane domain), but for which no similar function has been shown as yet; env-like domains are found in addition to the gag-pol domain of many plant LTR retrotransposons. Gag (group-specific antigen) Nucleic acid-binding protein forming the structural core of the retrovirus virion core and of the cytoplasmic VLP of retrotransposons. Int Integrase function ensuring the insertion of the double-stranded DNA daughter copy in the genome; also sometimes referred to as endonuclease (endo). LTR (long terminal repeat) Long sequence found repeated in the same orientation at both ends of retroviruses and LTR retrotransposons and containing promoter and regulatory sequences involved in transcription, that starts in the 50 LTR and terminates in the 30 LTR.

PBS (primer binding site) Short sequence found downstream to the 50 LTR, and involved in the priming of the (  )-DNA strand synthesis during reverse transcription. PPT (polypurine tract) Short AG-rich sequence found upstream to the 30 LTR, and involved in the priming of the ( þ )-DNA strand synthesis during reverse transcription Prot Protease protein involved in the cleavage of the functional proteins units from the (gag) pol polyprotein produced by the retrovirus or the retrotransposon. RT Reverse transcriptase protein ensuring the synthesis of the double-stranded daughter DNA copy from the RNA template produced by the integrated provirus or retrotransposon; an associate ribonucleaseH (RNaseH) function ensures the concomitant degradation of the RNA template. VLP (virus-like particle) Cytoplasmic intermediate analogous to the virion core of retroviruses; the VLP is constituted by structural gag proteins associated to two copies of the RNA template, together with enzymatic proteins (RT, int); the reverse transcription of the RNA template is thought to occur within the VLP.

Introduction and General Classification Transposable elements are ubiquitous mobile DNA sequences found in both prokaryotic and eukaryotic genomes. For a long time, they were considered as ‘parasite’ DNA, but have now been shown to be major components of plant genomes, where they can represent up to 80% of the bulk of large cereal genomes. Transposable elements, with a few notable exceptions, encode the functions involved in their mobility and are classified into two main classes with radically different transposition mechanisms. Class I elements or retrotransposons transpose via an RNA intermediate that is reverse-transcribed into a daughter copy DNA and re-inserted into the genome, while class II elements or DNA transposons move directly from DNA to DNA. Retrotransposons thus belong to the large class of reverse-transcribing elements (or retroelements) that multiply by transferring their genetic information from RNA to DNA, and are close relatives to viral entities such as animal retroviruses or pararetroviruses infecting both animal and plant hosts. However, in contrast to true infectious viruses, which propagate only functional genomes, retrotransposons are vertically transmitted intrinsic components of host genomes. Their integrated genomes are thus submitted to evolutionary drift, with various mutations and restructurations destroying their functionality, and defective copies can still be amplified, sometimes at surprisingly high levels, via functional related copies. Plant retrotransposons are thus found in a tremendous variety of structural variants, including highly defective and deleted versions, making their classification a difficult task. In addition, retrotransposons are composed of modules that frequently show incongruent phylogenies, and this intrinsic chimerical nature often obscures classifications based on phylogenies. There are two main categories of retrotransposons: LTR retrotransposons that contain two long terminal repeats (LTRs), and non-LTR retrotransposons often referred to as retroposons. All functional copies encode as basic modules the structural and functional proteins required for the retrotransposition cycle (summarized in Fig. 1). These include a structural RNA-binding protein (gag or gag-like), and a pol domain encoding the reverse transcriptase (RT) ensuring the synthesis of the double-stranded daughter DNA copy from the RNA template, as well as in most cases a protein ensuring insertion of the double-stranded DNA daughter copy in the genome (referred to as integrase for most elements). A number of plant LTR retrotransposons contain an additional coding domain currently termed env-like, although, as discussed below, functional and structural analogies to retroviral env proteins remain to be demonstrated. LTR retrotransposons have been further classified in Ty1/copia-type elements and Ty3/gypsy-type depending on the order of the coding domains. Non-LTR retrotransposons include LINEs (long interspersed nuclear elements), which are elements carrying

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Suborder Orthoretrovirineae (retroviruses) PBS

Retroviridae

PPT

gag

5 LTR

3 LTR

env

prot RT int pol

Suborder Retrotransposineae (LTR retrotransposons) PBS

gag

5 LTR Pseudoviridae

PPT

prot int RT 3 LTR pol

PBS

gag

5 LTR

Pseudovirus

PPT

prot int RT

env-l

Sirevirus

3 LTR

pol PPT

PBS

5 LTR

Metaviridae

gag

pol

PBS

5 LTR

Metavirus

prot RT int 3 LTR

gag

PPT

prot RT int

env-l 3 LTR

? Chromovirus

pol

PBS PPT 5 LTR

Unclassified

TRIMs

3 LTR

PBS

PPT

5 LTR

3 LTR

LARDs

Suborder Retroposineae (non-LTR retrotransposons, or retroposons)

gag-l

A(n)

pol

A(n)

LINEs

SINEs

Fig. 1 Structural diversity of retrotransposons found in plant genomes. The structure of a retrovirus has been provided for reference. The gag and pol domains are encoded in one or two frames, depending on the element, and have been represented here as a single ORF for simplification. Family names are indicated on the left and genus names on the right. LTR, long terminal repeat; gag, nucleic acid-binding protein forming the structural core of the retrovirus virion core and of the cytoplasmic VLP of retrotransposons; prot, protease protein involved in the cleavage of the functional proteins units from the (gag) pol polyprotein; int, integrase function ensuring the insertion of the double-stranded DNA copy in the genome; RT, reverse transcriptase ensuring the synthesis of the double-stranded DNA copy from the RNA template; env, envelope protein ensuring infectiosity of the retrovirus; env-l, env-like coding domain possessing structural features similar to env, but for which no similar function has been shown yet (a dotted box framing the env-like domain indicates that elements within the genus may or may not contain the domain); PBS, primer binding site; PPT, polypurine tract. Arrows flanking elements represent the target site duplication. The question mark linking the genera Metavirus and Chromovirus indicates that the definition of Chromovirus as an additional genus within the family Metaviridae has not been included in the official classification, with the result that many chromoviruses are still considered to be metaviruses.

coding sequences, and SINEs (short interspersed nuclear elements), small noncoding elements of a few hundred base pairs that exploit the transposition machinery of LINEs to ensure their amplification. LTR retrotransposons are by far the most abundant transposable elements in plant genomes, while LINEs appearsomewhat less represented, in contrast to mammalian genomes. SINEs are also abundant in plant genomes, although their small size prevents them from playing a large structural role. The continuous discovery of new structural variants of these major categories has made it necessary to revise and refine previous classifications. Retrotransposons have been included in a recent classification proposed for all reverse-transcribing elements, and based on viral nomenclature with type species forms defined for some genera (Table 1). All retroelements have been defined as belonging to class Retroelementopsida, presently divided in two orders, order Retrovirales that encompasses retroviruses, pararetroviruses, and LTR

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Classification of retroelements Type species

Order Retrovirales Suborder Orthoretrovirineae Family Retroviridae (retrovirus) Suborder Pararetrovirineae (pararetrovirus) Family Hepadnaviridae Family Caulimoviridae Suborder Retrotransposineae (LTR retrotransposons) Family Pseudoviridae (Ty1/copia retrotransposons) Genus Pseudovirus

Genus Hemivirus Genus Sirevirus Family Metaviridae (Ty3/gypsy retrotransposons) Genus Errantivirus Genus Metavirus

Genus Semotivirus Genus ? Unclassified

A few examples in Viridiplantaea

Not yet

Not yet CaMV, BSV, integrated EPRVs (banana BSV; petunia PVCV, tobacco TVCV, Nicotiana NtoEPRV, and Ns EPRV)

Ty1

Copia SIRE1

gypsy Ty3

BEL DIRS

Ta1 (Arabidopsis X1329), Tnt1 (tobacco X13777), Tst1 (potato X52387), Tto1 (tobacco D83003), WIS2 (wheat X63184), BARE1 (barley Z17327), Hopscotch (maize U12626), Tto1 (tobacco D83003), Tos17 (rice D85876), RIRE1 (rice D85597), Art1 (arabidopsis Y08010), Retrofit (rice U72726), Melmoth (kale Y12321), Tgmr (soybean U96748), Stonor (maize AF082134), Panzee (pigeon pea AJ000893), Tpv2 (bean AJ005762), LERE1 (tomato AF275345), Retrolyc1 (S. peruvianum AF228701), AtRE1 (Arabidopsis AB021265), TLC1 (S. chilense AF279585), Toto1 (tomato AF220602), Angela (wheat DQ666286) Osser (Volvox X69552) SIRE1 (soybean AY205608), ToRTL1 (tomato U68072), Endovir (Arabidopsis AY016208) Opie-2 (maize U68408), PREM2 (maize U41000), Osr7 (rice AP002538), Osr8 (rice AC021891)

Not yet Athila (Arabidopsis X81801), Cyclops (pea AJ000640), RIRE8 (rice AB014740), RIRE2 (rice AB030283), BAGY2 (barley AJ279072), Cinful (maize AF049110), Grande1 (maize X97604), Piggy1 (pea AY299398), Ogre (pea AY299398), Diaspora (soybean AF095730) Genus Chromovirus?: del1 (lilium X13886), IFG7 (pine AJ004945), Reina (maize U69258), RIRE3 (rice AB014738), RIRE7 (rice AB033235), Galadriel (tomato AF119040), tekay (maize AF448416), Beetle1 (beet AJ539424) Not yet Not yet LARDs: Sukkula (barley AY054378), Squiq (rice AY355293), Spip (rice AY355292), Dasheng (rice) TRIMs: (potato AJ276865), Katydid (Arabidopsis), mini-Toto1 (tomato X58273)

Order Retrales Suborder Retroposineae (non-LTR retrotransposons) L1 Alu

Cin4 (maize Y00086), del2 (lilium Z17425), Isabelle (maize AF326781), Karma (rice AB081316), Bali1A (rapeseed AF525305) TS (tobacco D17453), S1 (rapeseed L76840), Au (Aegilops AB046134)

Suborder Retronineae (bacterial retrons) a Elements have been classified according to year of publication or sequence submission to Genbank databases. Note that in several cases, only partial sequences are available. Extensive data on various plant elements can be found at: Repbase, a database of repetitive DNA elements of all organisms, the TIGR Plant Repeat Database and the Triticeae-specific TREP database. Retroelements were classified according to Hull R (2001) Classifying reverse transcribing elements: A proposal and challenge to the ICTV. Archives of Virology 146: 2255–2261; and Boeke JD, Eickbush T, Sandmeyer SB, and Voytas DF (2004) Pseudoviridae/Metaviridae In: Fauquet CM, Mayo MA, Maniloff J, Desselberger U, and Ball LA (eds.) Virus Taxonomy: Eighth Report of the International Committee on Taxonomy of Viruses, pp. 397–420. San Diego, CA: Elsevier Academic Press.

retrotransposons, and order Retrales that contains non-LTR retrotransposons and bacterial retrons. While order Retrales has not been further organized at the present time, order Retrovirales has been divided in several suborders, families, and genera. Suborder Orthoretrovirinae contains the family Retroviridae, or true retroviruses, of which no example is known in plants at the present time. Suborder Pararetroviridae contains two families of pararetroviruses, one of which (Caulimoviridae) is represented in plants. Pararetroviruses represent sensibly different lineages of retroelements, and are thought to derive from preexisting viruses having acquired a reverse transcription function, presumably from retrotransposons of the family Metaviridae. Many plant genomes also contain vertically transmitted integrated endogenous pararetroviral sequences (EPRVs), which represent integrated derivatives of pararetroviruses, although the infectious counterpart has not always been characterized. Plant pararetroviruses are not discussed here.

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Fig. 2 The amplification cycle of LTR retrotransposons. Transcription begins in the 50 LTR and ends in the 30 LTR, and produces both the RNA template used for reverse transcription and mRNAs translated into proteins. The gag protein binds to the RNA template and forms a cytoplasmic virus-like particle (VLP). Together with the RNA template, VLPs encapsulate the RT that will produce the double-stranded DNA copy, and the integrase (int) involved in transfer of the linear DNA copy to the nucleus and its insertion in the genome.

Suborder Retrotransposineae contains LTR retrotransposons, and has been divided in family Pseudoviridae (Ty1/copia retrotransposons) and family Metaviridae (Ty3/gypsy retrotransposons), based on the organization of the pol domain.

The Life Cycle of LTR Retrotransposons High Similarities with Intracellular Steps of the Retroviral Cycle LTR retrotransposons represent by far the most abundant plant retrotransposons. They share striking similarities with animal retroviruses, both in their structure (Fig. 1) and in their replication cycle (Fig. 2). Like integrated retroviral forms (termed proviruses), LTR retrotransposons are terminated by two LTR sequences. As the reverse transcription process generates two identical LTRs, with mutations accumulating subsequently, the rate of divergences between LTRs of a given copy is often used as a molecular clock to date its insertion. The gag and pol coding domains are found between the LTRs. Other typical features include short sequences known as PBS (primer binding site) and PPT (polypurine tract). The PBS is found immediately downstream the 50 LTR, and is involved in priming of the (  )-DNA strand synthesis during reverse transcription. It is generally complementary to the 30 end of a tRNA recruited for the priming of DNA synthesis (tRNAmet for most Pseudoviridae, more variable for Metaviridae). The PPT is a short AG-rich sequence found immediately upstream to the 30 LTR, and involved in the priming of the ( þ )-DNA strand synthesis. Inserted copies are typically bounded by short direct repeat of the target host site, generated upon insertion, and usually 5 bp long for plant LTR retrotransposons. Transcription begins in the 50 LTR, ends in the 30 LTR, and produces both the RNA template used for reverse transcription and the mRNA(s) that will be translated into proteins, after cleavage of the gag-pol polyprotein by the protease (prot) function. The gag protein binds to the RNA template and forms a cytoplasmic virus-like particle (VLP) analogous to the retroviral virion core. Together with the RNA template, VLPs encapsulate the RT that will produce the double-stranded DNA copy, and the integrase (int) that will be involved in transfer of the linear DNA copy to the nucleus and its insertion in the genome. This amplification cycle includes multiple steps of control, with transcription being absolutely crucial for retrotransposition. As a consequence, the mode of amplification of a particular element will be conditioned by its transcriptional regulation. The amplification cycle of LTR retrotransposons is thus closely related to the intracellular steps of the retroviral cycle, with strong similarities in the reverse transcription and integration processes, as well as similar involvement of a cytoplasmic encapsulated intermediate. It has to be noticed that the amplification of non-LTR retrotransposons is sensibly different, with reverse transcription occurring at the integration site. The reality of the retroviral-type cycle of LTR retrotransposons has been demonstrated in yeast, but not yet in plants. However, VLPs have been detected for the barley BARE1 element. In addition, it has been demonstrated very recently that plant elements of the genus Sirevirus (see below) encode unusually large Gag domains extended in their C-terminus, and that this Gag extension binds to LC8/LC6 dynein proteins. Although the biological significance of this interaction remains to be determined, it is very similar to the binding of some human retroviruses’ Gag protein to LC8 proteins to

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ensure intracellular movement of virion cores along microtubules. Such observations reinforce the strong similarities between the life cycle of retroviruses and LTR retrotransposons. The major difference between retroviruses and LTR retrotransposons resides in the infectious potentialities of the former, mediated by the env function that allows them to produce extracellular virions and infect host receptor cells. Bona fide retrotransposons, on the other hand, are not infectious and their amplification cycle is exclusively intracellular. However, we shall see later that this assumption should be treated with care and that the delimitation between retroviruses and retrotransposons is increasingly fuzzy, possibly even in plants. It is now considered as certain that retroviruses are derivatives of some LTR retrotransposon Metaviridae lineages. LTR retrotransposons are themselves derived, with LINE-like ancestors, from the assembly of various modules together with an RT-like function derived from ancestral cellular functions used for the transition of genetic information from RNA to DNA at the dawn of life. Retroviruses have evolved from LTR retrotransposons through transduction of env domains ensuring their ability to exit the cell. Although the process can be reversed (e.g., with vertebrate endogenous retroviruses having lost their env functionality and reversed to a restricted intracellular life cycle), retroviruses can be seen as the ultimate evolutionary step in freeing of cellular genetic information outside of the cell, and they basically represent LTR retrotransposons that have succeeded in life. It is not clear whether this process has occurred only once during the evolution of Retroviridae; however, it is now increasingly clear that a number of their LTR retrotransposons’ left-behind cousins have repeatedly attempted to join them in acquiring an extracellular life and that some of them have truly succeeded, albeit not yet in plants.

Evolution as Quasispecies-Like Populations Other similarities of retrotransposons with retroviral characteristics include their high level of sequence variability. This has been best illustrated for elements related to the tobacco Tnt1 element, which are found in many species of the family Solanaceae, and evolve via a high variability of the LTR region carrying regulatory features (U3 region). Species of the genus Nicotiana contain Tnt1 elements that are classified into several groups (termed subfamilies) which are closely related, except in their U3 regions that differ strongly. Members of each subfamily are detected within each species, and their U3 divergence thus predates the Nicotiana radiation. However, the respective proportions of each subfamily differ in each species, indicating that each host species has preferentially amplified specific subfamilies. Similar evolutionary patterns have been detected in other Solanaceae species, such as tomato, pepper, or aubergine, which contain elements closely related to Nicotiana elements, except in the U3 region, and which can also be further subdivided in subfamilies based on the nature of the U3 region only. Interestingly, the different tobacco Tnt1 subfamilies are each expressed in response to different stimuli, within the same host genome. The very ancient Tnt1 element has thus evolved in Solanaceae a large range of highly related progeny populations that have gained new regulatory sequences and new expression patterns. This strategy is highly similar to the evolutionary patterns of RNA viruses and retroviruses, generating populations of closely related but different genomes referred to as quasispecies. This variability has been attributed to the high-error-prone process of reverse transcription, due the lack of proofreading repair activity of RNA polymerases and reverse transcriptases. This allows viral populations to evolve very rapidly when environmental conditions change and endow retroviruses with high adaptative capacity. Thus, LTR retrotransposons similarly evolve continuums of closely related sequences, which can be defined as quasispecies-like. This evolutionary pattern allows them to vary their amplification conditions, as regulatory controls in the U3 will determine the conditions in which the element will be transcribed and amplified. The preferential amplification of specific subfamilies in each host species suggests that optimal patterns of amplification may have been selected and maintained in each host genome, possibly depending on its own evolutionary, reproductive, and environmental history.

The Different Genera of LTR Retrotransposons The Family Pseudoviridae (Ty1/copia Retrotransposons) Pseudoviridae are differentiated from all other LTR retrotransposons and from retroviruses by an unusual organization of the pol domain, with the int domain placed before the RT domain (Fig. 1). This confirms that the different LTR retrotransposon families have evolved by independent acquisition of modular functions. Historically, Ty1/copia retrotransposons have been the first LTR retrotransposons characterized in higher plants, and the best-known plant LTR retrotransposons belong to this family, including the tiny handful of elements for which mobility has been demonstrated. Pseudoviridae have been classified in the genera Pseudovirus (type species Ty1 of yeast), Hemivirus (type species copia of Drosophila), and the recent genus Sirevirus (type species, SIRE1 of soybean). Pseudoviruses and hemiviruses differ by the fact that the former use a full tRNAmeti to prime synthesis of the (  )-DNA strand, while the latter use a half tRNAmeti. So far, no hemivirus has been referenced in plants, and all LTR retrotransposons usually – and incorrectly – referred to as copia-type retrotransposons in fact classify in the genus Pseudovirus. Sireviruses have so far been identified only in plants. They are phylogenetically well separated from the two other genera, and display some very interesting characteristics. In particular, several members of the genus have acquired additional ORFs carrying additional functions that present some

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similarities to the env domain of retroviruses, such as a transmembrane domain (TMD). This is, for instance, the case with SIRE1, ToRTL1, and Endovir (Table 1). A number of other elements classified as sireviruses, such as Opie-2, PREM-2, Osr7, or Osr8 (Table 1), do not carry additional ORFs with typical env-like features. However, all of them carry a significant amount of noncoding DNA after the end of the pol domain, from a few hundred base pairs to over 1 kbp, while members of the genera Pseudovirus and Hemivirus, such as Tnt1, Ty1, or copia, separate the pol domain from the 30 LTR by a few dozen nucleotides or less. This suggests that the env-less sireviruses may also have contained similar additional ORFs that have decayed over evolutionary times.

The Family Metaviridae (Ty3/gypsy Retrotransposons) The family Metaviridae is characterized by an organization of the pol domain identical to that of retroviruses, with the int protein encoded at the end of pol, after the RT domain (Fig. 1). Metaviruses appear to be basal to both retroviruses and pseudoviruses, and the family Metaviridae therefore encompasses a large variety of members, whose classification is probably not at its final stage yet. Although metaviruses have been characterized in plants more recently than pseudoviruses, they have been found to be more abundant in several plant genomes. The family Metaviridae comprises the genera Errantivirus (type species gypsy of drosophila), Metavirus (type species Ty3 of yeast), and Semotivirus (type species BEL of drosophila) (Table 1). To these three genera are often added the DIRS retrotransposons, originally described from Dictyostelium discoidum, that show differences with other metavirus elements. DIRS do not carry typical LTRs, that is, direct repeats in the same orientation, and do not encode an int. Instead, they are bounded by inverted long repeats and integration in the host genome is ensured by a tyrosine-recombinase. In spite of these differences, they are more related to metaviruses, both in terms of RT sequence and organization of the pol domain, with the tyrosine-recombinase placed downstream to the RT domain. They could represent chimerical elements having acquired an RT domain from some metaviruses, and are placed by several authors as a putative additional genus (still unnamed) in this family. No DIRS, semotivirus, or errantivirus has yet been characterized in plants, with the totality of so-called gypsy-like retrotransposons of plants classified in the genus Metavirus (Table 1). Errantiviruses are grouped together on the basis of the presence of an additional env-like domain, which ensures a real envelope function for the two infectious gypsy and ZAM elements of drosophila. Gypsy and ZAM are the first examples of true retroviruses outside of the vertebrate subphylum. However, their phylogenies place them quite apart from retroviruses, and they are more closely related to other LTR retrotransposons of the family Metaviridae. They do presumably represent recent progresses of LTR retrotransposons toward an extracellular life cycle. The separation of errantiviruses and metaviruses has been interpreted by many authors as based on the lack of an env or env-like domain in metaviruses. However, the situation is more complex. Many plant elements presently classified in the genus metavirus indeed do not encode for additional ORFs after the pol domain (such as del1, IFG7, Reina, RIRE3, RIRE7, Galadriel, tekay, and Beetle-1; Table 1). In contrast to env-less sireviruses mentioned above, these elements do not show significant stretches of additional sequences between the pol domain and the 30 LTR (only a few dozens at the most, often very few), indicating that they may never have contained any additional ORF. However, a large number of plant elements presently classified in the genus Metavirus do contain an additional ORF with typical env-like features (such as Athila, Cyclops, RIRE8, RIRE2, BAGY2, Cinful, Grande1, Piggy1, or Ogre; Table 1). The two groups of elements cluster separately in most phylogenetic studies, indicating that they could be considered as two different genera of the family Metaviridae. An additional genus, Chromovirus, has recently been proposed within the family Metaviridae, based on the presence of a chromodomain in the integrase. However, it has not been included in the current official classification. Interestingly, all plant metaviruses devoid of additional coding domains seem to be chromoviruses. It is therefore likely that the classification of Metaviridae will be reconsidered in future, to separate the two groups of metaviruses and recognize the existence of the chromodomain. Interestingly, the soybean Diaspora element is devoid of any additional DNA between pol and the 30 LTR; however it clusters with env-containing elements, suggesting that its original env-like domain has been deleted. Therefore, the presence/absence of an env-like domain would not be a relevant criterion in the redefinition of genera within the family Metaviridae.

Unclassified LTR Retrotransposons A number of LTR retrotransposons have recently been discovered that cannot be classified into any of the above genera due to their lack of coding sequences. The TRIMs (terminal repeat retrotransposons in miniature) consist of small LTRs framing a short central domain of noncoding sequences, while the LARDs (large retrotransposon derivatives) are larger defective derivatives that contain a large internal domain without significant similarities to retrotransposon coding domains. Both TRIMs and LARDs share many of the structural characteristics of LTR retrotransposons, such as RT priming sites PBS and PPT, and are highly amplified, suggesting efficient transactivation by functional retrotransposons. For several LARDs, sequence identities have been found with LTRs of canonical LTR retrotransposons (e.g., Dasheng with RIRE2, Squip with RIRE8, Spip with RIRE3), suggesting that they have been derived from these elements, and may eventually be classified together. Similarly, some TRIM-like small elements are derived from canonical LTR retrotransposons, such as the tomato mini-Toto1 element, derived from the Toto1 pseudovirus. In addition, a number of intermediate situations are frequently found, such as TRIM derivatives containing larger additional unknown DNA, a

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possible intermediate situation between TRIMs and LARDs. The existence of TRIMs and LARDs and various intermediates indicates that plant genomes contain a large range of retrotransposons with structural features much more diverse and complex than previously thought. As long as the features required in cis for amplification are conserved (e.g., LTRs, PBS, and PPT, as well as a few internal sequences involved in packaging), and as long as the LTR promoter remains transcriptionally active to ensure the production of an RNA template, these defective or atypical elements can be amplified to high levels using the structural and enzymatic retroviral functions provided by their original functional parent, or possibly by other related LTR retrotransposons.

Env-Like Functions in Plants? Although plant retrotransposons with additional ORFs of the env type have been detected, the potential role of such a function in infectivity remains very hypothetical in plants. Retroviral env glycoproteins carry a number of typical structures, such as a TMD mediating formation of infectious virions and a host receptor-binding domain, mediating targeting of host receptor cells. No significant homology to any retroviral env gene has been shown for any plant element; however, such identification is highly unlikely due to the intrinsic high variability shown by animal retroviral env proteins. As the definition of env-like functions in plant retrotransposons is mostly based on the presence of TMDS, the issue of the naming of these additional ORFs has been hotly debated. TMDs can also be found in various proteins, and the env denomination has a specific meaning related to anchoring of the protein in the membrane envelope of extracellular virion particles and to recognition of receptors on target cells to ensure viral infection after membrane fusion. Quite likely, such mechanisms are rarely used by plant viruses for host-to-host transmission, due to the cell wall barrier. Plant retrotransposons carrying additional ORFs related to functions used by most plant viruses to ensure host-to-host transmission, for example, via vector insects, have not yet been found. However, examples of enveloped viruses do exist in plants, and retrotransposon-encoded env-like proteins could be involved in anchoring of VLPs to cellular membranes, playing a role in cell-to-cell transmission via plasmodesmata and possibly in transmission by insects. Furthermore, conservation of env-like domains has been observed between elements from different plant genera within both the Metaviridae and the Pseudoviridae (SIRE1 of soybean and Endovir of Arabidopsis). In addition, conserved splice acceptor sequences are detected 50 of env-like domains of several plant metaviruses, and production of an env-like spliced subgenomic RNA has been demonstrated for the barley BAGY-2 element, a mechanism similar to those used by retroviruses to ensure production of the env protein. In contrast, sireviruses such as SIRE1 use stop codon suppression to express env-like protein. The existence of such specific expression mechanisms strongly suggests that env-like domains are not mere incidental transduction of cellular functions, as shown for the maize Bs1 element, and that acquisition of additional coding domains with TMDs, whether in a quest or not for extracellular life, may play an important role in the plant LTR retrotransposon life cycle. In the search of a better definition, and in spite of lack of evidence for any retroviral env-type function, it has been proposed to maintain the env denomination due to its vernacular long-term use: for example, the nucleocapsid protein used to build up the VLP core particle is commonly termed gag in all retrotransposons, by analogy to the gag ‘group-specific antigen’ region of retroviruses, although such functional denomination does not make sense in plants. A striking observation is that LTR retrotransposons generally have been very efficient in acquiring env-like domains, and have done so many times. For instance, additional ORFs with env-like features have been observed both for Ty1/copia and Ty3/gypsy retrotransposons, suggesting that they have been acquired independently during evolution. In addition, the Metaviridae contain clear examples of acquisition of env function from different sources, for example, from baculoviruses for gypsy of drosophila, and from phleboviruses and herpesviruses for the semotiviruses Cer of Caenorhabditis elegans and Tas of Ascaris lumbrocoides, respectively. This indicates that LTR retrotransposons have repeatedly been able to hijack viral functions leading to extracellular autonomy, and there is no conceivable reason for plants to be an exception.

The Nomenclature Issue Rules for naming individual elements also continue to be hotly debated. Earlier retrotransposons have been named according to authors’ personal fancies (e.g., Athila, Cyclops, Opie, Ogre) or based on the plant species they were isolated from, but following no defined rule (e.g., Tnt1 for Transposon of Nicotiana Tabacum, BARE-1 for Barley Retrotransposon, WIS-2 for Wheat Insertion Sequence). An official nomenclature has recently been proposed for all retroelements, with three letters used for the species, up to three (exceptionally four) for the element and ‘V’ for virus. In this nomenclature, the yeast Ty1 and drosophila copia elements have been renamed SceTy1V and DmeCopV, respectively, and Tnt1 and BARE1 have been renamed NtaTnt1V and HvuBV, respectively. However, this nomenclature has so far not been implemented by the plant scientific community, most likely because it does not solve the major problem arising from the presence of highly related elements in different plant species or genera, which is a general rule in plants. In addition, most transposable element populations, and especially retrotransposons, are generally composed of populations of closely related sequences. Defining whether a particular copy is related to an already known element, or is different, is thus often a difficult task. A consensus position has been to consider that copies that show above 75% identity over most of their length (not taking into account deletions or insertions of unrelated material) are variants of the same element. However, many elements have been discovered and named independently. For instance, the wheat WIS2 and Angela elements are quite similar to the barley BARE1 element, with internal nucleotidic sequences 75–80% identical between BARE1 and WIS2 and over

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80% identical between BARE1 and Angela. Similarly, Solanum subsection Lycopersicon counterparts of the tobacco Tnt1 element have been named Retrolyc1 (first identified in S. peruvianum) and TLC1 (identified in S. chilense), although Retrolyc1 and Tnt1 nucleotide sequences are over 85% identical and TLC1 and Retrolyc1 are 93% identical. This highlights the difficulties inherent to a nomenclature based on host species names. A proposition was made to replace species-based names by first names, a trend that seems to be developing at present, at least in the plant field. More specifically, it has been proposed that female first names should be used for retroelements and male first names for class II DNA transposons, although this particular proposal is not really implemented. Such type of nomenclature should alleviate the problem of the presence of related elements in different hosts, provided authors make sure their favorite first name has not been used yet. Attempts to unify the nomenclature (as well as the general classification) of transposable elements of Triticeae are currently coordinated by Dr. Thomas Wicker, who curates the TREP database for Triticeae repeats. This should lead to the redefinition of consensus guidelines that could be fruitfully followed by the entire scientific community for transposable elements of all plant genera.

See also: Caulimoviruses (Caulimoviridae). Movement of Viruses in Plants. Plant Reoviruses (Reoviridae). Vector Transmission of Plant Viruses

Further Reading Boeke, J.D., Eickbush, T., Sandmeyer, S.B., Voytas, D.F., 2004. Pseudoviridae/Metaviridae. In: Fauquet, C.M., Mayo, M.A., Maniloff, J., Desselberger, U., Ball, L.A. (Eds.), Virus Taxonomy: Eighth Report of the International Committee on Taxonomy of Viruses. San Diego, CA: Elsevier Academic Press, pp. 397–420. Casacuberta, J.M., Vernhettes, S., Audéon, C., Grandbastien, M.-A., 1997. Quasispecies in retrotransposons: A role for sequence variability in Tnt1 evolution. Genetica 100, 109–117. Grandbastien, M.-A., Audeon, C., Bonnivard, E., et al., 2005. Stress activation and genomic impact of Tnt1 retrotransposons in Solanaceae. Special Issue: Retrotransposable Elements and Genome Evolution. Cytogenetic and Genome Research 110, 229–241. Havecker, E.R., Gao, X., Voyas, D.F., 2004. The diversity of LTR retrotransposons. Genome Biology 5, 225. Havecker, E.R., Gao, X., Voytas, D.F., 2005. The sireviruses, a plant-specific lineage of the Ty1/copia retrotransposons, interact with a family of proteins related to dynein light chain 8. Plant Physiology 139, 857–868. Hull, R., 2001. Classifying reverse transcribing elements: A proposal and challenge to the ICTV. Archives of Virology 146, 2255–2261. Kalendar, R., Vicient, C.M., Peleg, O., Anamthawat-Jonsson, K., Bolshoy, A., Schulman, A.H., 2004. Large retrotransposon derivatives: Abundant, conserved but nonautonomous retroelements of barley and related genomes. Genetics 166, 1437–1450. Kordis, D., 2005. A genomic perspective on the chromodomain-containing retrotransposons: Chromoviruses. Gene 247, 161–173. Lucas, H., Yot, P., Lockhart, B.E.L., Capy, P., 2004. Taxa of viruses, virus-like and subviral agents infecting Poaceae, class ‘Retroelementopsida’. In: Lapierre, H., Signoret, P.A. (Eds.), Virus and Virus Diseases of Poaceae (Graminae). Paris: INRA Editions, pp. 279–303. Malik, H.S., Henikoff, S., Eickbush, T.H., 2000. Poised for contagion: Evolutionary origins of the infectious ability of invertebrate retroviruses. Genome Research 10, 1307–1318. Peterson-Burch, B.D., Wright, D.A., Laten, H.L., Voytas, D.F., 2000. Retroviruses in plants? Trends in Genetics 16, 151–152. Vicient, C.M., Kalendar, R., SchulmanEnvelope-class, A.H., 2001. Envelope-class retrovirus-like elements are widespread, transcribed and spliced, and insertionally polymorphic in plants. Genome Research 11, 2041–2049. Wicker, T., Sabot, F., Hua-Van, A., et al., 2007. A unified classification system for eukaryotic transposable elements. Nature Previews Genetics 8, 973–982. Witte, C.P., Le, Q.H., Bureau, T., Kumar, A., 2001. Terminal-repeat retrotransposons in miniature (TRIM) are involved in restructuring plant genomes. Proceedings of the National Academy of Sciences, USA 98, 13778–13783. Wright, D.A., Daniel, F., Voytas, D.F., 2001. Athila4 of Arabidopsis and Calypso of soybean define a lineage of endogenous plant retroviruses. Genome Research 12, 122–131.

Relevant Websites http://www.girinst.org Genetic Information Research Institute (GIRI). http://www.tigr.org The TIGR Plant Repeat Databases, J. Craig Venter Institute (JCVI). http://wheat.pw.usda.gov The Triticeae Repeat Sequence Database (TREP), GrainGenes, Agricultural Research Service, USDA.

Vector Transmission of Plant Viruses Etienne Herrbach and Quentin Chesnais, University of Strasbourg, Colmar, France r 2021 Elsevier Ltd. All rights reserved. This is an update of S. Blanc, Vector Transmission of Plant Viruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00433-7.

Nomenclature

kDa Kilo Dalton ORF Open reading frame

HC Helper component HC-Pro Helper component-protease

Glossary Helper component Virus-encoded nonstructural protein required for vector transmission of a virus. Hemolymph Physiological fluid that circulates in the body cavity, or hemocoel, of arthropods in direct contact with their inner organs. Horizontal transmission Transmission of a pathogen from one host to another within the same generation, as opposed to vertical transmission. Phytoviruses Viruses infecting plants. Piercing-sucking insects Insects adapted to sap or blood feeding, with the mouthparts in form of long chitin stylets

able to pierce and penetrate tissues and cells, and allow ingestion of their content. Protist A unicellular eukaryotic organism. Readthrough protein A protein expressed by two open reading frames separated by a stop codon that is bypassed during translation. Vector Mobile organism acquiring a pathogen on an infected host and inoculating it in a healthy one. Vertical transmission Transmission of a pathogen from the parent(s) to the offspring, usually through the germline.

Introduction Plant-infecting viruses, here referred to as phytoviruses, are known since the last two decades of the 19th century. To date, over 900 species of phytoviruses have been described and their number is steadily increasing as a result of latest virological technologies. Unlike animal viruses, phytoviruses infect immobile hosts; moreover, they must cross the plant cuticula and the cell wall in order to infect living plant cells. The majority of phytoviruses have evolved a close relationship with mobile organisms, called ‘vectors’, which help them in their plant-to-plant propagation; this biological phenomenon is coined ‘horizontal transmission’. However, some very stable viruses, such as tobamoviruses, are able to infect plants by leaf contact, or by wounds, and require no vector. Many viruses are transmissible through seed and/or pollen, and the majority can be transferred through vegetative multiplication; these routes are referred to as ‘vertical transmission’. Many virus species can use both horizontal and vertical transmissions. Finally, for many viruses (e.g., in genera Velarivirus, Foveavirus, or Hordeivirus), the natural vector, if any, remains unknown. The present article aims at summarizing the current knowledge on the various strategies used for vector transmission of plant viruses.

Plant Virus Vectors The vast majority of vector organisms belong to phytophagous insect taxa, mainly sap feeding members of the order Hemiptera (aphids, leaf- and planthoppers, whiteflies, coccoids). Several chewing insects in the order Coleoptera also are phytovirus vectors. Other important vectors belong to phytophagous Eriophyid and Tenuipalpid mites, to rootectoparasitic Longidorid and Trichodorid nematodes, and to soil-dwelling plant-infecting Chytrid fungi and Plasmidiophorid protists. The transmission process includes several successive steps. ‘Acquisition’ refers to the uptake of virions while the vector feeds on an infected plant, whereas ‘inoculation’ occurs during a subsequent feeding on another plant. ‘Latency’ is the time lapse between virus acquisition and time when a viruliferous (i.e., virus bearing) vector becomes infective (i.e., able to inoculate the virus). ‘Retention’ time refers to the time period during which the vector remains infective. These criteria served to define the typology of the strategies involved, i.e., the transmission modes as described below.

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Transmission of Plant Viruses by Insects History of the Typology of the Transmission Modes Vector transmission of a plant virus was first demonstrated in Japan in 1893 for the leafhopper-transmitted Rice dwarf virus (Phytoreovirus, Reoviridae). Interestingly, this discovery occurred shortly after that of insects as vectors of animal viruses. Pioneer researchers soon noticed that all virus–vector associations do not follow the same pattern, due to the specific biology of each of the three partners, i.e., plant, virus, and the vector. The typology of these ‘transmission modes’ was then defined for aphid vectors; however, it applies quite well to other vectors with a piercing-sucking apparatus, but not to beetles, equipped with biting-chewing mouthparts. The first typology, proposed in 1939, was based on the optimal times for acquisition and inoculation and defined the ‘nonpersistent’ (seconds to minutes) and the ‘persistent’ (hours to days) modes. This typology was completed in 1956 with the semi-persistent mode, with intermediary features (i.e., minutes to hours). In 1962, the second typology coined the terms circulative and stylet-borne viruses, according to whether the infectivity, and therefore the virus, is maintained or lost through molting (trans-stadial passage) and to whether the virus is detected or not in the insect hemolymph. In practice, circulative corresponds to persistent, and stylet-borne to non-persistent; however, the semi-persistent mode does not fit in this typology. In 1977, it was proposed to unify these classifications on the basis of the retention site of the virions. Thus, in the non-circulative (or externally-borne) mode, the virions are retained at the cuticular lining of stylets and/or foregut, without being transported across any epithelium, and therefore lost during molt; this includes the semipersistent mode, also named ‘foregut-borne’ according to the retention site. In the ‘circulative’ (or internally-borne) mode, the virions are internalized across the gut epithelium into the hemolymph, and from whence across the salivary gland epithelium, the circulation within the hemolymph occurring as virions. Circulative viruses are further classified as propagative or non-propagative, according to their ability to replicate or not in the vector’s organs. Furthermore, a few propagative circulative viruses are capable of a mother-to-progeny transovarial transmission. The correspondence between feeding behavior of aphids and other hemipteran vectors, and transmission modes is well established. Aphid feeding behavior is characterized by two kinds of stylet insertion into plants. They first proceed to one or usually several brief probes (i.e., within seconds) in epidermal and mesophyll cells, during which cell content is sampled and gustated (host selection behavior) and saliva is expelled into these cells; after this phase the aphid may settle or leave the plant. If it settles, the stylets continue a deeper path into plant tissues, mainly intercellularly, to finally reach the phloem vessels, where the sap ingestion starts. Non-circulative viruses are acquired and inoculated during short probes in epidermal cells or phloem cells, in respectively non- and semi-persistent modes, whereas circulative viruses are generally acquired and inoculated during ingestion from, and salivation into phloem sieve elements. The feeding behavior of other hemipteran vectors differs somewhat from that of aphids. For example, coccoids (scale insects) make no epidermal probes, therefore transmit viruses in the semipersistent mode only. Moreover, the stylet pathway of whiteflies to the phloem is mainly intercellular, whereas that of leafhoppers, with less flexible stylets, pierce straight through mesophyll cells on hosts and non-hosts. For all hemipterans, patterns of feeding behavior vary according to host plant suitability. Many hemipterans can also ingest xylem sap, though this behavior is irrelevant to virus transmission, as xylem-located viruses are unknown.

Circulative Transmission Circulative Non-Propagative Transmission This strategy is very specific of phytoviruses, as it has ever been reported for the transmission of neither arthropod-transmitted animal virus (arboviruses), nor non-viral plant pathogens. The best studied pathosystems following this mode are aphids–luteovirids, leafhoppers–mastreviruses, and whiteflies–begomoviruses. Other pathosystems using this mode include aphids–nanovirids and –capulaviruses, and probably many of those involving fungus and protist vectors. Case of Aphid-Transmitted Luteovirids These viruses of the family Luteoviridae, that are causal agents of several economically important diseases in arable and vegetable crops worldwide, share the property of being phloem-restricted in their host-plants and transmitted by aphids (family Aphididae) during phloem feeding (Fig. 1(A)). The route of virions inside the vector has been extensively studied using cellular and molecular biology approaches. The widely accepted model is as follows. Virions are acquired into the vector along with ingested phloem sap. Then, in compatible aphid species–virus species combinations, the particles are retained at putative receptors borne by the luminal side of gut epithelium, then invaginated into gut cells and released at the basal side of these cells (‘transcytosis’) into the hemolymph. From there they diffuse, probably passively, until they reach accessory salivary glands, whose cells transport them by transcytosis from basal to apical side into the saliva (Fig. 2). They can then be inoculated during salivation into the phloem tissue of a new host. No virus particle has ever been observed in any other organ than the gut or the accessory salivary glands, nor has any viral replication ever been reported. During transcytosis in epithelial cell of the gut and the salivary glands, virions observed using microscopy are always seen inside membranous vesicles, either alone, or grouped in clusters or tubular arrays, suggesting that the particles follow intracellular trafficking routes. In the gut, the transcytosis occurs at either the midgut, the hindgut, or both, depending on the aphid and virus species concerned. The gut barrier seems less specific than the salivary one, since viral particles are detected in the hemolymph of some non-vector species. Specificity is thought to rely on viral ligands that interact with receptors borne on the plasmalemma of

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Fig. 1 Main insect vectors of plant viruses. (A) Acyrthosiphon pisum (Aphididae); (B) Bemisia tabaci (Aleyrodidae); (C) Parthenolecanium persicae (Coccidae); (D) Psammotettix alienus (Cicadellidae); (E) Peregrinus maidis (Delphacidae); (F) Spissistilus festinus (Membracidae); (G) Frankliniella occidentalis (Thripidae); (H) Epilachna varivestis (Coccinellidae). Photographs: (A)–(B) Courtesy of Ian Wright, UC Riverside USA; (C) Courtesy of Gérard Hommay, INRAE Colmar France; (D) Courtesy of Emmanuel Jacquot, INRAE Montpellier France; (E) Courtesy of Antoine Franck, CIRAD France; (F) Courtesy of Ken Childs, USA; (G) Courtesy of Dominique Blancard, INRAE Bordeaux France; (H) Courtesy of Bryan Cassone, Brandon University Canada.

traversed cells of the gut then the salivary gland. Luteovirids have two structural proteins: the major coat protein and a minor readthrough protein composed of the coat protein at the N-terminus followed by the readthrough domain which is extended outside the particle and indispensable for the interaction with putative receptors. Recently, alanyl aminopeptidase N and ephrin receptor proteins have been shown to act as a virion receptor at the gut level, whereas other proteins are suspected to do so. During virion diffusion in the hemolymph, the readthrough protein of at least some species interacts with a chaperone protein (homolog to Escherichia coli GroEL) secreted by endosymbiotic bacteria; this interaction is believed to protect the particle from degradation and/or assist its transfer to the salivary glands. Case of Leafhopper-Transmitted Mastreviruses Unlike luteovirids, mastreviruses (Geminiviridae) infect mesophyll cells and are transmitted by mesophyll- and phloemfeeding leafhoppers (Cicadellidae) (Fig. 1(D)). Acquired virions mainly accumulate in the epithelia of the filter chamber and of the midgut, though without replication, from whence they reach the salivary glands. Unlike luteovirids, the gut provides the main selective barrier, as a non-vector can become infective upon microinjection of virions into its hemolymph, as well as upon a simple puncture of the gut wall with a needle. A probing time of less than a minute may be enough for the vector to acquire the virus, indicating that acquisition occurs in the mesophyll while puncturing cells. Conversely, inoculation, requiring longer times, occurs in phloem. Virion entry into the gut cells, mainly at the filter chamber and the anterior part of the midgut, is thought to use lipid-raft-mediated endocytosis, and the viral capsid protein is thought to bear the transmission determinants. In the same family, the genera Curtovirus and Topocuvirus are also transmitted according to the circulative non-propagative mode, by respectively leafhoppers and treehoppers (Membracidae) (Fig. 1(F)). A treehopper has been reported as a vector of a Grablovirus member, with a still undetermined transmission mode. Finally, the recently defined genus Capulavirus is transmitted by aphids. Case of Whitefly-Transmitted Begomoviruses The genus Begomovirus is the genus with the largest number of species in the family Geminiviridae, with over 400 species described, mainly in the tropics and subtropics where they cause severe diseases to crops. The vector of all these species is exclusively the whitefly Bemisia tabaci (Aleyrodidae) (Fig. 1(B)). However, B. tabaci now appears as a vast complex of biotypes and ‘cryptic’ species, with a great invasive potential worldwide; moreover, this complexity is associated to a wide viral genetic diversity and a strong proneness to viral recombination. Like all geminivirids, begomoviruses are reported to follow the circulative non-propagative transmission mode. However, a propagative and a transovarial transmission of Tomato yellow leaf curl virus (TYLCV) DNA by B. tabaci have been reported, though this

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Table 1

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Families and genera of plant viruses and their vectors. Virus taxa with no known vector are not mentioned

Family

Genus

Vector

Alphaflexiviridae

Allexivirus Potexvirus Ophiovirus

Mite Aphida, chytrid?, cycloraphan flyb Chytrid

Carlavirus Trichovirus Vitivirus

Aphid, leafhopper, whiteflyb Mite Coccoid, aphidc

Benyviridae

Benyvirus

Plasmidiophorid

Bromoviridae

Alfamovirus Anulavirus Bromovirus Cucumovirus Ilarvirus

Aphid Thripsd Beetle, aphide Aphid Thripsd

Caulimoviridae

Badnavirus Caulimovirus Tungrovirus

Coccoid, aphid, lacebugb Aphid Leafhopper

Non-circulative Non-circulative helper strategy Circulative propagative

Closteroviridae

Ampelovirus Closterovirus Crinivirus

Coccoid Aphids Whitefly

Non-circulative Non-circulative Non-circulative capsid strategy

Fimoviridae

Emaravirus

Mite

Geminiviridae

Becurtovirus Begomovirus Capulavirus Curtovirus Grablovirus Mastrevirus Topocuvirus Turncurtovirus

Leafhopper Whitefly Aphid Leafhopper Treehopper Leafhopper Treehopper Leafhopper

Circulative non-propagative Circulative non-propagative Circulative non-propagative

Kitaviridae

Cilevirus

Mite

Circulative (propagative?)

Luteoviridae

Enamovirus Luteovirus Polerovirus

Aphid Aphid Aphid, whiteflyb

Circulative non-propagative Circulative non-propagative Circulative non-propagative

Nanoviridae

Babuvirus Nanovirus

Aphid Aphid

Circulative non-propagative Circulative non-propagative (with helper factor)

Peribunyaviridae

Tospovirus

Thrips

Circulative propagative

Phenuiviridae

Tenuivirus

Planthopper

Circulative propagative

Potyviridae

Bymovirus Ipomovirus Macluravirus Poacevirus Potyvirus Roymovirus Rymovirus Tritimovirus

Plasmidiophorid Whitefly Aphid Mite Aphid Mite? Mite Mite

Circulative Non-circulative Non-circulative

Reoviridae

Fijivirus Oryzavirus Phytoreovirus

Planthopper Planthopper Leafhopper

Circulative propagative Circulative propagative Circulative propagative

Rhabdoviridae

Cytorhabdovirus Dichorhabdovirus Nucleorhabdovirus Varicosavirus

Aphid, planthopper, leafhopper Mite Leafhopper, planthopper, aphid Chytrid

Circulative propagative Circulative (propagative?) Circulative propagative Circulative

Aspiviridae Betaflexiviridae

Transmission mode

Non-circulative Non-circulative

Non-circulative capsid strategy

Non-circulative capsid strategy

Circulative Circulative Circulative Circulative

non-propagative non-propagativef non-propagative non-propagative

Non-circulative helper strategy

(Continued )

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Table 1

Continued

Family

Genus

Vector

Transmission mode

Secoviridae

Cheravirus Comovirus Fabavirus Nepovirus Sequivirus Torradovirus Waikavirus

Nematode Beetle Aphid Nematode, mite Aphid Whitefly Aphid, leafhopper

Non-circulative Non-circulative capsid strategy Non-circulative Non-circulative Non-circulative

Solemoviridae

Sobemovirus

Beetle, aphid, mirid, (thrips, diptera)g

Non-circulative?

Tombusviridae

Alphanecrovirus Aureusvirus Avenavirus Betacarmovirus Betanecrovirus Dianthovirus Gammacarmovirus Machlomovirus Tombusvirus Umbravirus Zeavirus

Chytrid Chytrid, plasmidiophorid?b Chytrid Beetle Chytrid Chytridb Chytrid Beetle, thripsb Chytrid Aphid Chytrid

Tymoviridae

Marafivirus Tymovirus

Leafhopper Beetle

Virgaviridae

Furovirus Pecluvirus Pomovirus Tobravirus

Plasmidiophorid Plasmidiophorid Plasmidiophorid Nematode

Non-circulative capsid strategy

Circulative propagative

a

Only one viral species when assisted by the HC of a potyvirus. Only one viral species. c Two viral species when assisted by a helper virus. d Virus transported along with pollen. e Only one viral species, non-circulative mode, capsid strategy. f Circulative propagative transmission seems to occur in some specific virus-vector combinations. g Rice yellow mottle virus (RYMV) can also be transmitted by orthopterans, hemi- and heteropterans, probably mechanically. Sources: ICTV 9th and 10th Reports (https://talk.ictvonline.org/). Note: Bragard, C., Caciagli, P., Lemaire, O., et al., 2013. Status and prospects of plant virus control through interference with vector transmission. Annual Review of Phytopathology 51, 177–201. Brown, J.K., 2016. Vector-mediated Transmission of Plant Pathogens. St Paul, MN: American Phytopathological Society Press. b

DNA is probably not infective. Moreover, the report of a venereal transmission of a specific TYLCV isolate in certain B. tabaci biotypes/ species adds a further originality, unique among arthropod-transmitted phytoviruses. Globally, the begomoviruses–whiteflies interaction and its efficiency appear complex and diverse, possibly linked to a co-evolution between vector species or subspecies and a great diversity of viruses. Immuno-microscopic studies on several begomovirus species revealed para-crystalline structures, thought to be virions, in the filter chamber and salivary glands of the vector. Like for luteovirids, the participation of an endosymbiont-secreted chaperone protein during virion diffusion in the whitefly’s hemolymph has been shown. The coat protein of begomoviruses, whose sequence is only slightly variable, determines the transmission specificity and its N-terminus is required for receptor-mediated endocytosis in the gut and salivary glands. Whitefly proteins, such as cyclophilin B, are known to be involved in TYLCV transmission.

Circulative Propagative Transmission While circulative propagative mode is almost the general rule for arthropod-transmitted animal viruses (arboviruses), this strategy is also well known for many phytoviruses, such as the genera Cytorhabdovirus and Nucleorhabdovirus (Rhabdoviridae) (transmitted by leaf-, planthoppers, or aphids), Fijivirus, Oryzavirus (Reoviridae) and Tenuivirus (Phenuiviridae) (planthoppers), Marafivirus (Tymoviridae) and Phytoreovirus (Reoviridae) (leafhoppers), and Tospovirus (Tospoviridae) (thrips) (Fig. 1(G)). In these cases, virions infect gut cells, are then released into the hemolymph and infect various tissues and organs, including salivary glands. Rhabdoviruses infect the central nervous system, from whence they spread to other organs. The infection of the vector is very similar to the infection of an insect by an entomopathogenic virus, and the vector can therefore also be seen as a virus host, even though such vectors are usually much less impacted by the infection in terms of survival and fecundity. Remarkably, propagative phytoviruses belong (apart from marafiviruses) to families comprising also animal-infecting viruses, and the question of whether they may have evolved from insect viruses has been raised.

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Non-Circulative Transmission The non-circulative transmission is specific to phytoviruses, for which it is the most frequent case (4 50% of viruses), and unknown for vertebrate arboviruses. The virions are adsorbed at the luminal side of cuticula-lined tissues, either stylet food canal or/and foregut, without being internalized, and are therefore also called ‘externally-borne’. This accounts for the loss of infectivity of the vector during molting and the shorter periods needed for efficient acquisition and inoculation (Table 2). As mentioned above, classical authors defined (1) the non-persistent mode, where the virions are acquired and inoculated from epidermal and mesophyll cells retained at the stylet level, and (2) the semi-persistent mode, where virions are acquired and inoculated from deeper cells, even in phloem, and/or retained at the foregut level (Fig. 2). However, it is sometimes uneasy to verify these features experimentally and this often vague distinction is less used today. Two hypotheses, though not mutually exclusive, have been put forward to explain the inoculation phase in this transmission mode. On one hand, the ingestion–egestion hypothesis states that the retained virions are regurgitated by the vector with the sampled plant sap during probing and feeding. On the other hand, the ingestion–salivation hypothesis, which is favored in many cases, postulates that the salivation into plant tissues is responsible to detach the retained virions and inoculate them into the host; this implies that, in the case of aphids at least, the retention site lies in the distal part of the stylets where the food and the salivary canals merge together. Table 2

Characteristics of the different modes of vector transmission of plant viruses

Viruses

Non-circulative

Mode

Non-persistent

Acquisition time Acquisition and inoculation sites Latency time Retention time Inoculation time Passage through molt Replication in vector Transovarial passage Vector specificity

Secondes to minutes Epidermis None Minutes to hours Secondes to minutes No No No Large

Circulative

Circulative propagative

Semi-persistent

Persistent non-propagative

Persistent propagative

Minutes to hours Phloem None Hours to days Minutes to hours No No No Medium

Hours to days Phloem Hours to days Several days Hours to days Yes No No Narrow

Hours to days Phloem Days Life long Hours to days Yes Yes Possible for certain viruses Narrow

Fig. 2 Schematic representations of plant virus transmission mechanisms. Retention sites for each virus are indicated by a red star adjacent to the labeled area. C-P-Prop: circulative-persistent propagative viruses; C-P-NProp: circulative-persistent non-propagative viruses; NC-SPer: non-circulative semipersistent viruses; NC-NPer: non-circulative nonpersistent viruses. With the permission of Mauck et al. (2018). Evolutionary determinants of host and vector manipulation by plant viruses. Advances in Virus Research 101, 189–250.

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Modern authors characterize the non-circulative mode according to the retention strategy of virus particles in the vector. In the ‘capsid strategy’ the virion is directly linked to cuticular sites in the vector, whereas the ‘helper strategy’ requires one or two extracapsid peptides building a ‘molecular bridge’ between cuticular sites and virions. For both strategies, the existence of virus receptors in the vector, explaining the virus–vector specificity, has been postulated and in a few cases demonstrated. Remarkably, a helper factor has also been shown to be required in two circulative viruses, Faba bean necrotic stunt virus (FBNSV, Nanovirus, Nanoviridae) for its transfer within the aphid’s hemocoel, and Rice stripe virus (RSV, Tenuivirus, Phenuiviridae) for its endocytosis into gut cells.

Capsid Strategy Case of Aphid-Transmitted Cucumoviruses This strategy is illustrated by the interaction between cucumoviruses and aphid vectors. The type-member Cucumber mosaic virus (CMV, Cucumovirus, Bromoviridae) is easily transmitted by aphids fed on purified virions through a stretched Parafilm™ membrane. This clearly demonstrates that virions alone are transmissible, without need for any other compound, indicating that the virion structure bears the determinant required for attachment to the cuticula. The viral retention site is likely in the stylet alimentary and/or common canals. Moreover, the three RNAs and the coat protein can be separated in vitro, then reassembled; thus, exchanging the components of transmissible and non-transmissible isolates confirmed the implication of the sole coat protein in transmission. Sequence modifications in this protein have been reported to modify vector specificity. Moreover, the vector-less Tobacco mosaic virus (Tobamovirus, Virgaviridae) can be aphid-transmitted when its RNA molecule is encapsidated within the capsid of a cucumovirus. The same strategy operates in the aphid transmission of alfamoviruses and carlaviruses, as well as probably for whiteflytransmitted criniviruses; however, very few is known so far for other closterovirids.

Helper Strategy Case of Aphid-Transmitted Potyviruses This strategy was first described for members of the genus Potyvirus (Potyviridae). In this species-rich genus, most species are aphidtransmitted, many of them are also transmissible mechanically and some by seed. In terms of acquisition, retention, and inoculation times, potyviruses are transmitted similarly to cucumoviruses. However, purified potyvirus particles are not transmissible by aphids fed through a Parafilm™ membrane, albeit they are mechanically transmitted, showing that vector transmission requires an additional factor which is lost during purification process. Subsequent experiments proved that this factor is a virus-encoded nonstructural peptide, called helper factor or helper component (HC), released in infected cells and mediating the attachment of virions to cuticular sites during acquisition. For an efficient transmission, the HC must be taken up by the vector before or during virus acquisition, but not after. Comparison of transmissible and non-transmissible isolates or species, and mutagenesis and microscopy experiments revealed the role of the peptidic motifs, borne by the coat protein and the HC, in building a ‘molecular bridge’ between virion, HC and stylet cuticula. The HC is commonly designated as ‘HCPro’ in literature, since it also has a protease function. The precise retention site of the HC–virion complex in the vector’s stylets likely comprises one or more cuticular protein(s). Trans-complementation is another feature of the helper strategy of potyviruses. The HC of a potyvirus species can interact with the capsid of a related species acquired on the same plant or another one, due to sequence homologies, and thus assist its transmission; thus, this common phenomenon advantages the population genetics and evolution of potyvirus species. Case of Aphid-Transmitted Caulimoviruses Caulimoviruses (Caulimoviridae) are transmitted by aphids according to a particular helper strategy, that is best studied for Cauliflower mosaic virus (CaMV). Like for potyviruses, purified CaMV particles are not transmissible by artificially-fed aphids, indicating the requirement of an additional factor, later demonstrated to be the nonstructural transmission helper protein P2 expressed from the open reading frame (ORF) II of viral DNA. Moreover, a second helper peptide, the product P3 expressed from the ORF III and associated to the viral capsid, was proven indispensable in the transmission process. During cell sap ingestion, virus receptor(s) harbored on the stylets’ inner cuticula retain either the complete P2–P3–virion complex simultaneously, or sequentially with P2 binding first to the stylets, followed by subsequent attachment of the P3–virion complex. In infected plant cells, CaMV virions accumulate and concentrate in specific inclusions called virus factories, whereas P2 and P3 accumulate in separate inclusions named transmission bodies. When the aphid stylets enter such cells, the transmission bodies are instantly (r 5 s) disrupted, and the freed P2 relocalizes to the cellular microtubules. Simultaneously, the virus factories release virions that join P2 on the microtubules and form transmissible virions that can be acquired efficiently by the vector. This remarkable phenomenon, coined ‘transmission activation’, is reversible after the stylet withdrawal. A similar reaction, i.e., vector-induced formation of transmissible virions, has recently been described for Turnip mosaic virus, a potyvirus, whose transmission also relies on a helper strategy. Microscopy studies demonstrated that CaMV virions are retained at the tip of the aphid’s stylet bundle, i.e., the common duct where the food and salivary canals converge, and more precisely at a particular part in the inner side of the maxillary stylets called acrostyle. Among cuticular proteins embedded in this chitinous organ, two proteins called stylin-01 and stylin-02, largely conserved among aphid species, have recently been shown to interact with either P2 or the transmissible complex P2–P3–virion, and are thus good candidate receptors for CaMV and possibly other aphid-transmitted non-circulative phytoviruses.

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Transmission by Beetles Vectors of plant viruses are known in phytophagous beetles within the Chrysomelid and Coccinellid families (Fig. 1(H)), and for members of the genera Bromovirus (Bromoviridae), Comovirus (Secoviridae) Machlomovirus (Tombusviridae), Sobemovirus (Solemoviridae), and Tymovirus (Tymoviridae). These viruses share isometric, stable and also mechanically transmissible particles. Virions are rapidly acquired when the vector’s biting-chewing mouthparts damage plant tissues. The interaction, still poorly described, is likely ingestion–regurgitation and specific. Phytophagous beetles are devoid of salivary glands and coat their mandibles with an RNase-rich food regurgitate. The viruses transmitted display a certain resistance to RNases that may account for their vector specificity, and they are thought to translocate rapidly to unwounded cells and/or vascular tissues. In some cases, virions are detectable in the insect hemolymph within seconds, though whether this feature refers to a circulative mode or not is unknown. Some vectors can efficiently infect a plant after ‘drinking’ a purified virion suspension, indicating that the mere virion is required for transmission.

Transmission by Non-Insect Vectors The transmission of several plant viruses relies on non-insect vectors (Table 1), such as mites, nematodes, and ‘fungus-like’ organisms (i.e., true fungi and protists).

Transmission by Mites Two families of mites (Acari), Eriophyidae (gall mites), and Tenuipalpidae (false spider mites, flat mites), contain vectors of members of several virus genera (Table 1, Fig. 3(A)). However, these minute vectors have been less studied than insects, and the underlying mechanisms are poorly known, and only in some cases. For feeding, phytophagous mites empty plant epidermal cells, using a short stylet bundle. For Brevipalpus spp. (Tenuipalpidae) and Citrus leprosis virus (Cilevirus, Kitaviridae), virions were detected in midgut cells. Virions of the eriophyid (Aceria tosichella)-transmitted Wheat streak mosaic virus (WSMV, Tritimovirus, Potyviridae) were also detected in hemolymph and salivary glands. Whereas the transmission mode was long considered as non-circulative and semi-persistent, the observations mentioned above rather suggest a circulative (propagative?) interaction, but experimental proofs are lacking so far. Considering that tritimoviruses belong to the same family as aphid-transmitted potyviruses and possess an HCPro homolog ORF, experiments showed that HCPro of WSMV bears motifs required for transmissibility, even though the transmission modes (non-circulative for potyviruses) likely differ substantially between the two genera.

Transmission by Nematodes Members of Nepovirus and Cheravirus in the family Secoviridae, and of Tobravirus (Virgaviridae) are transmitted by, respectively, longidorid and trichodorid nematodes. These nematodes feed on root cells using stylet-like organs called odontostyle and onchiostyle, respectively. The mode of transmission is described to be non-circulative, as virions are mainly adsorbed at the inner lining of the esophagus, at still unknown receptors. Despite the loss of virions during molting, Xiphinema spp. longidorids (Fig. 3(B)) harboring Grapevine fanleaf virus (GFLV, Nepovirus, Secoviridae) can remain viruliferous after at least 4 years rest in soil; thus, these slow-moving vectors provide the virus with a travel in time, rather than in space. The specific transmission of GFLV and probably other nepoviruses requires the sole capsid protein (capsid strategy), whereas that of tobraviruses needs one or two nonstructural proteins (helper strategy). The viral determinants of GFLV are located around a canyon on the capsid surface and account for its vector specificity.

Transmission by Fungi and Protists These soil-dwelling unicellular organisms, either chytrids (zoosporic Fungi or plasmidiophorid protists (Cercozoa) (Figs. 3(C) and (D)), have flagellate forms, called zoospores, that can enter the cytoplasm of root cells and from there colonize the plant. While doing so, these species may inoculate viruses of several genera (Table 1). Several mechanisms seemingly exist. The best documented associations are the chytrids Olpidium spp. transmitting tombusvirids and ophioviruses, and the plasmidiophorids Polymyxa spp. vectors of benyviruses and furoviruses. In the first association, during zoospore formation in infected host plants, virions bind to the zoospore external coat using fungal glycoproteins and specific regions in the capsid protein (homologous to non-circulative capsid strategy), leading to a swelling of virions. They are then released into the newly infested host cell upon cell wall digestion and fungal penetration. In the case of the protists Polymyxa spp. (Fig. 3(C)), the virions, or at least viral RNA-protein complexes, are endocytosed in the zoospore, presumably without replication (homologous to circulative non-propagative mode) and then inoculated into root cells with the zoospore cytoplasm. A 32 kDa viral protein was shown to be required for the transmission of a benyvirus by Polymyxa sp. Precise mechanisms of acquisition and inoculation are poorly known.

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Fig. 3 Main non-insect vectors of plant viruses. (A) Aceria tosichella (Acari Eriophyidae); (B) Xiphinema index (Nematoda Longidoridae); (C) Polymyxa betae (Cercozoa Plasmidiophorida); (D) Olpidium sp. (Fungi Chytridiomycota). Photographs: (A) Courtesy of Anna Skoracka, Wielkopolska Centre for Advanced Technologies, Adam Mickiewicz University, Poznan, Poland; (B) Courtesy of Gérard Demangeat, INRAE Colmar France; (C) Courtesy of Olivier Lemaire, INRAE Colmar France; (D) Courtesy of Dominique Blancard, INRAE Bordeaux France.

Host and Vector ‘Manipulation’ by Plant Viruses Plant virus infection can drastically alter host’s phenotype, including size, color, volatiles emission, primary and secondary metabolites composition. Historically, most studies have focused on exploring the effects of viruses on plants from an agronomic point of view, such as impacts on crop yield or quality. In recent years, it has been highlighted that virus-induced changes in plant physiology may also influence host–vector interactions and, thereby, the probability of virus transmission. Given that this crucial step of transmission obligately depends on vectors for many plant viruses, it has been proposed that vector-borne viruses evolved adaptive traits that induce host phenotypes, and effects on vectors that promote their own spread. Since then, numerous studies showed that virus infection in host plants modifies vector behavior, such as orientation and settling preferences, but also reproduction and survival, in ways that are generally conducive to transmission. These effects fall into the category of putative ‘indirect manipulations’, because, in these cases, the viruses modify the behavior and/or performance of their insect vector through phenotypic alterations of the shared host. Another category of effects, qualified as ‘direct’, corresponds in changes within the viruliferous vector’s orientation and feeding preferences. In the early 1950s, it was reported for the first time a positive effect of plant infection on the fitness of an aphid vector. Thereafter, indirect effects of plant viruses on vectors, linked to induced volatiles and to changes in leaf attractiveness and suitability for the vector, have been reported with aphid vectors, and were shown to vary according to the transmission mode of the virus. Over the last two decades, these observations were substantially confirmed and documented by many published studies with aphids and other insect vectors (whiteflies, leafhoppers, thrips, etc.). Indeed, vector-borne viruses can be classified in different categories based on their associations with the host plant and the vector (Table 1). Some viruses are restricted to the phloem tissue of their host plant, virions are thus only acquired and inoculated by vectors achieving sustained phloem sap ingestion (i.e., during hours). In contrast, viruses that are not restricted to the phloem are acquired and inoculated most efficiently during brief epidermal or mesophyll intracellular probes and are generally retained within the vector only for seconds or minutes. In this case, if the vector initiates ingestion of phloem sap prior dispersal, virus particles might be lost, and viral transmission impeded. Consequently, short probing followed by rapid dispersal is the most-conducive vector behavior for nonphloem restricted virus transmission. From this classification, it was predicted that viruses sharing the same transmission modes – and thus, benefiting from a similar sequence of attraction, vector probing, feeding, and dispersal behavior – should exhibit convergent indirect host-mediated effects on vectors. Indeed, most phloem-restricted viruses improve the nutritive quality of the host plant for vectors (i.e., improved survival and/or fecundity), which is expected in turn to lead to crowding and subsequent dispersal of viruliferous vectors. These viruses also increase the palatability of the infected host plant compared to healthy ones, which results in improved vector feeding behavior and retention, facilitating virion acquisition from the phloem tissue. For the non-phloem restricted viruses, infections often reduce plant quality for vectors, and tend to reduce the retention of vectors on infected plants, thus potentially promoting

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the rapid dispersion of viruliferous vectors to healthy plants. Regardless of the type of virus, the vectors are preferentially attracted by the visual and volatile cues of infected plants over healthy plants, which is expected to promote virus spread. Only a few studies have documented cases of direct effects of plant viruses on vector behavior and, for the moment, the mechanisms are poorly characterized. Phloem-restricted viruses with persistent-circulative transmission mode need to perform a route from the gut to the salivary glands of their vector before they can be inoculated into a new host. These intimate associations offer strong opportunities for direct interactions of viruses on the physiology of their vector. For this kind of virus, vector dispersal is favored only after sufficient virions have been acquired by the insect to become infective. Therefore, it is predicted that these viruses may have evolved mechanisms to encourage dispersal following sustained phloem sap ingestion on infected hosts, and to discourage viruliferous vectors from visiting infected hosts; these changes in vector preference are named ‘conditional vector preferences’. This hypothesis was verified on aphids carrying two luteovirids, settling preferentially on uninfected plants, while non-viruliferous aphids preferred the infected plant. The same effect was observed with aphids having acquired virions from an artificial medium (thus overcoming any potential indirect plant-mediated effect). Some examples have also documented changes in viruliferous vectors feeding behavior, such as alteration of phases related to virus transmission (i.e., salivation, phloem sap intake, sustained ingestion of phloem sap), most of which are expected to enhance the probability of virus transmission.

Future Prospects From a mechanistic point of view, basic information on the virus-vector interaction process is increasingly documented in several cases, but is lacking in many other cases. Further studies in various transmission modes will provide a better understanding of the complex virus–vector–plant interactions, including the viral ‘manipulation’ of the two other partners favoring disease spread, and the localization and identification of vector receptors used by the virus for its transmission. Novel knowledge is based on ongoing progress in high-throughput data analysis and microscopic technologies. From an evolutionary point of view, the vector provides an obvious bottleneck for the virus, since only some genome variants encapsidated in transmissible virions will be transferred into a new host plant. This issue has started to be documented but requires more information to fully understand its implications. From an agricultural point of view, vector-borne viruses represent a burden for most crops worldwide. Since virus-infected plants remain incurable, phyto-protection against viruses mainly relies on (1) use of resistant varieties, provided resistant genes are available, although resistance-breaking viral strains are susceptible to be selected, and (2) insecticide, or acaricide and nematicide, treatments directed against the vector, which have potential environmental drawbacks and can also lead to the selection of resistant vector populations. In case of perennial crops or seed-transmitted viruses, using virus-free material is also important, even though it does not prevent further contamination. Deeper knowledge of the transmission process and the viral epidemiology is expected to generate new strategies to protect plants from viruses using vector-targeted, and more environment-friendly and integrated practices. This includes the use of molecular and/or behavioral disruption of the virus–vector–plant interaction, and the adaptation of crop management to climate change.

See also: Betaflexiviruses (Betaflexiviridae). Bluner-, Cile-, and Higreviruses (Kitaviridae). Bromoviruses (Bromoviridae). Caulimoviruses (Caulimoviridae). Closteroviruses (Closteroviridae). Cucumber Mosaic Virus (Bromoviridae). Geminiviruses (Geminiviridae). Luteoviruses (Luteoviridae). Nanoviruses (Nanoviridae). Orthotospoviruses (Tospoviridae). Plant Reoviruses (Reoviridae). Plant Rhabdoviruses (Rhabdoviridae). Potyviruses (Potyviridae). Secoviruses (Secoviridae). Tenuiviruses (Phenuiviridae). Tobamoviruses (Virgaviridae). Tombusviruses (Tombusviridae). Tymoviruses (Tymoviridae). Virgaviruses (Virgaviridae)

Further Reading Blanc, S., Drucker, M., Uzest, M., 2014. Localizing viruses in their insect vectors. Annual Review of Phytopathology 52, 403–425. Bragard, C., Caciagli, P., Lemaire, O., et al., 2013. Status and prospects of plant virus control through interference with vector transmission. Annual Review of Phytopathology 51, 177–201. Brown, J.K., 2016. Vector-mediated Transmission of Plant Pathogens. St Paul, MN: American Phytopathological Society Press. Eigenbrode, S.D., Bosque-Pérez, N.A., Davis, T.S., 2018. Insect-borne plant pathogens and their vectors: Ecology, evolution, and complex interactions. Annual Review of Entomology 63, 169–191. Hogenhout, S.A., Ammar, E.D., Whitfield, A.E., Redinbaugh, M.G., 2008. Insect vector interactions with persistently transmitted viruses. Annual Review of Phytopathology 46, 327–359. Hull, R., 2014. Plant to plant movement. In: Hull, R. (Ed.), Plant Virology, fifth ed. Amsterdam: Academic Press, pp. 669–751. Mauck, K.E., Chesnais, Q., Shapiro, L.R., 2018. Evolutionary determinants of host and vector manipulation by plant viruses. Advances in Virus Research 101, 189–250. Wilson, J.R., DeBlasio, S.L., Alexander, M.M., Heck, M., 2019. Looking through the lens of 'Omics technologies: Insights into the transmission of insect vector-borne plant viruses. In: Bonning, B.C. (Ed.), Insect Molecular Virology: Advances and Emerging Trends. Poole: Caister Academic Press, pp. 113–144. Zhou, J.S., Drucker, M., Ng, J.C.K., 2018. Direct and indirect influences of virus–insect vector–plant interactions on non-circulative, semi-persistent virus transmission. Current Opinion in Virology 33, 129–136.

Viral Suppressors of Gene Silencing Hernan Garcia-Ruiz, University of Nebraska–Lincoln, Lincoln, NE, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of J. Verchot-Lubicz, J.P. Carr, Viral Suppressors of Gene Silencing, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00718-4.

Glossary Antiviral immunity Is a group of mechanisms that protect cells from virus infection. They target viral proteins or nucleic acids. Antiviral gene silencing Is the downregulation of viral RNA accumulation. It targets viral RNA for degradation or translational repression. Additionally, in DNA viruses, it prevents transcription. Gene silencing Is a sequence-specific gene inactivation system mediated by small interfering RNAs (siRNA). It downregulates RNA accumulation without affecting DNA sequence. It is also known as RNA interference. Pathogenicity determinants Are viral components, usually proteins, that promote virus infection in cells and cause developmental defects in infected organisms. Post-transcriptional gene silencing Is the degradation or translational repression of mRNA. It targets both cellular and viral RNA.

Small interfering RNAs (siRNAs) Are double-strand RNAs that measure 21–24 nt in length. SiRNAs are the specificity determinant of gene silencing and program argonaute proteins for specific degradation of RNA with sequence complementarity to the siRNA. Suppressors of gene silencing Are proteins encoded by viruses that interfere with gene silencing. Their activity prevents degradation of viral RNA, enhance transcription of viral RNA, and alters silencing of cellular genes. These effects are in part responsible for developmental defects in virus infected organisms. Transcriptional gene silencing Prevent inhibits transcription from chromosomes, plasmids or viral DNA mini chromosomes. It is directed by siRNAs guiding enzymatic DNA methylation. Virus-induced gene silencing Virus infection induces silencing of cellular genes with sequence similarity to viral RNA.

Introduction Gene silencing is a sequence-specific gene inactivation system mediated by small interfering RNAs (siRNA) that measure 21–24 nt. Gene silencing downregulates RNA accumulation without affecting DNA sequence and is also known as RNA interference. Gene silencing regulates gene expression at the transcriptional or post-transcriptional levels. Transcriptional gene silencing is directed by siRNAs guiding enzymatic DNA methylation and inhibits transcription from chromosomes, plasmids or viral DNA mini chromosomes. Post-transcriptional gene silencing is the nucleolytic cleavage and degradation or translational repression of mRNA (Fig. 1). The pathway and biological roles of gene silencing are conserved in eukaryotes. Gene silencing regulates gene expression in a temporal and tissue-specific manner to control development, response to stress, and to preserve genome integrity. Additionally, in plants, nematodes, and insects, gene silencing is an essential component of antiviral immunity. In these organisms, plant viruses are inducers and suppressors of gene silencing. Our mechanistic understanding of gene silencing and virus-encoded gene silencing suppressors is derived mainly from the study of plant viruses. Accordingly, this article is focused on suppressors of gene silencing encoded by plant viruses. Viruses use RNA to replicate, express their genes, or both. The RNA might be perceived by the cell and activates antiviral defense, including gene silencing and mRNA decay. DNA viruses are targeted by both transcriptional and post-transcriptional gene silencing. RNA viruses, viroids and satellite RNAs are targeted by post-transcriptional gene silencing. To overcome gene silencing and promote susceptibility, viruses encode suppressors of gene silencing that are essential for viruses to replicate and spread. Virus-encoded gene silencing suppressors interfere with antiviral defense, and have been identified in plant viruses, insect viruses, fungal viruses and in some viruses infecting humans. However, the antiviral role of gene silencing in mammals is only beginning to be elucidated. In plant-virus interactions, the establishment of infection is determined by the balance between antiviral gene silencing versus virus evasion or suppression of antiviral defense. Endogenous and antiviral gene silencing have conserved and overlapping pathways. Accordingly, virus-encoded silencing suppressors affect both endogenous and antiviral gene silencing and their effects are in part responsible for symptom development in infected plants.

Antiviral Gene Silencing Pathway Antiviral gene silencing is non-cell autonomous, initiates at the single cell level and spreads cell-to-cell and long distance through plasmodesmata and the plant vascular system. The pathway consists of four parts: Initiation, targeting, amplification, and systemic spread (Fig. 2). Upon initial detection, Dicer-like (DCL) proteins process viral dsRNA into virus-derived siRNA that associate with

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DNA DNA viruses

Transcriptional gene silencing

Transcription RNA

RNA viruses Viroids Satellite RNA viruses Satellite RNAs

Non-coding RNA

Protein-coding RNA (mRNA)

Post-transcriptional gene silencing

Translation

Proteins

lncRNA

siRNA miRNA tasiRNA hc-siRNA nat-siRNA long siRNA

Virus-infected plants

Virus-activaded siRNAs Virus-derived siRNAs RNA-Induced silecing complex

Fig. 1 Flow of genetic information and gene silencing in plants. RNA-induced silencing complexes are formed between effector proteins and small interfering RNAs (siRNAs) derived from non-coding transcripts. Gene silencing is a block in the flow of genetic information by preventing transcription (transcriptional gene silencing) or by inducing degradation or translational repression of protein-coding RNAs (post-transcriptional gene silencing). In addition to endogenous long non-coding RNAs and siRNAs, virus-infected plants accumulate virus-induced siRNAs and virus-de-rived siRNAs, which enter the RNA-induced silencing complex and program degradation of host or viral mRNA. Gene silencing is essential for normal development and antiviral immunity. It targets viruses at the transcriptional or post-transcriptional level. Adapted from Garcia-Ruiz, H., Ruiz, M.T.G., Peralta, S.M.G., Gabriel, C.B.M., El-Mounadi, K., 2016. Mechanisms, applications, and perspectives of antiviral RNA silencing in plants. Mexican Journal of Phytopathology 34, 286–307.

argonaute (AGO) proteins to form RNA-induced silencing complexes (RISC) that target viral mRNA, and potentially host mRNAs, for endonucleolytic cleavage or translational repression. Initial recognition of viral RNA is necessary but not sufficient to restrict plant virus infection. Restriction of infection requires silencing amplification by cellular RNA-dependent RNA polymerases (RDRs). After moving out of the infected cell, in recipient cells the silencing response is amplified. Cellular RDRs synthesize new viral dsRNA that is the substrate for the formation of secondary virus-derived siRNAs that move systemically ahead of the virus to establish an antiviral state. DNA viruses express their genes through mRNA. Silencing is activated at the post-transcriptional level, resulting in the formation of virus-derived siRNAs that guide targeting of viral RNA. Furthermore, virus-derived siRNAs in association of AGO and RDR proteins guide methylation of viral DNA to prevent mRNA transcription (transcriptional gene silencing). Combined, the effects of both transcriptional and post-transcriptional gene silencing create an antiviral state that prevents virus replication and movement.

Small Interfering RNAs Direct Antiviral Immunity Through their association with AGO proteins, virus-derived siRNAs are the specificity determinant guiding translational repression or cleavage of viral RNA with sequence complementarity. In a variation of the pathway targeting DNA viruses, silencing complexes formed by virus-derived siRNAs and AGO proteins direct methylation of viral DNA to prevent transcription. In both cases, the end result is the inhibition of virus replication and movement. In plant-virus interactions, several phenomena are mediated by gene silencing such as cross protection, symptom recovery, synergism, and non-host resistance. Cross protection occurs when infection by a mild virus strain prevents subsequent infection by a second virus of the same or closely related species. siRNAs-derived from the mild virus strain direct targeting of viruses with sequence similarity. However, not all cross-protection cases are explained by gene silencing. In Turnip crinkle virus, and Wheat streak mosaic virus, supper infection exclusion is a form of cross protection mediated by viral proteins. Symptom recovery has been described for several plant-virus combinations, such as Arabidopsis thaliana infected by Oilseed rape mosaic virus or Tobacco rattle virus. Infected plants accumulate high virus titers and develop visible symptoms. However, the upper leaves of those plants contain low virus titters and do not develop symptoms. Virus-derived siRNAs formed in the lower part of the plant move systemically to the meristem to direct viral RNA targeting and prevent virus invasion of the meristem. Leaves formed after that harbor virus-derived siRNAs that mediate virus downregulation. Interestingly, in recovered A. thaliana tissue, the suppressor from Oilseed rape mosaic virus lost its activity.

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NonInfected cell

Infected cell Virus entry

Virus cell-to-cell and systemic movement

Disassembly Virion formation

Replication and transcription

Virus recognition RNA replication intermediate

Primary virus-derived siRNAs DCL

Foldback structure on ssRNA

Transcription DCL

Viral DNA Methylation and transcriptional silencing

Endogenous siRNAs

siRNA systemic movement and immunization

AGO-siRNA association

Target slicing or translational repression

Secondary virus-derived siRNAs

Viral dsRNA DCL

AGO

Host RNA

Viral RNA RDR DRB

AGO

AGO

SGS Silencing amplification

Plant development

Antiviral defense

Fig. 2 Antiviral gene silencing and basic components in plants. In infected cells, viruses are recognized by dsRNA replication intermediates or self-complementary sequences in genomic RNA or mRNA. Dicer-like (DCL) proteins cut viral dsRNA to form primary virus-derived small interfering RNAs (siRNAs), that associate with argonaute (AGO) proteins and program endonucleolytic slicing and degradation or translational repression of viral RNA. Host transcripts with sequence complementarity to virus-derived siRNAs might be targeted. Slicing of viral RNA triggers amplification of antiviral RNA silencing that results in the formation of viral dsRNA by cellular RNA-dependent RNA polymerases (RDR) and processing by DCL to form secondary-virus-derived siRNAs. Silencing amplification triggers methylation of viral DNA which results in transcriptions silencing. Virus-derived siRNA move cell-to-cell and systemically to establish a state of antiviral immunity away from the initial infection site. Adapted from Garcia-Ruiz, H., Ruiz, M.T.G., Peralta, S.M.G., Gabriel, C.B.M., El-Mounadi, K., 2016. Mechanisms, applications, and perspectives of antiviral RNA silencing in plants. Mexican Journal of Phytopathology 34, 286–307.

Several mechanisms explain non-host resistance, such as the absence of pro-viral factors needed for replication or movement. However, antiviral gene silencing early in infection guided by endogenous siRNAs or virus activated siRNAs might target viral RNA to prevent the establishment of infection and plants show the phenotype of a non-host.

Viral Suppressors of Gene Silencing Promote Infection Gene silencing is an essential component of antiviral immunity in plants, insects, and nematodes. To establish infection, viruses must protect themselves from antiviral defense responses such as RNA decay, gene silencing and protein-based

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mechanism such as autophagy or ubiquitination. At least two mechanisms have been proposed to explain gene silencing: evasion and suppression. Positive-strand RNA viruses replicate in membrane bound compartments that sequester replication intermediates. Negative-strand RNA viruses and dsRNA viruses replicate in enveloped vesicles. These structures might prevent access to viral dsRNA by the gene silencing machinery. In an interesting variation of gene silencing, a non-coding region in Cauliflower mosaic virus produces large amounts of siRNAs that saturate the silencing machinery. This decoy role prevents targeting of viral RNA.

Mechanisms of Gene Silencing Suppression To protect themselves from gene silencing and promote infection, most plant and some insect viruses encode silencing suppressors that inactivate gene silencing by multiple mechanisms (Table 1) discussed below. Table 1

Representative virus-encoded gene silencing suppressors

Virus

Suppressor

Silencing target

Other functions

siRNA binding Potyviruses TBSV BYV TSWV WDV RGDV ACMV

HC-Pro P19 P21 NSs Rep PNS11 AC4

siRNA monomers and dimers siRNA dimers siRNA dimers siRNAs siRNAs siRNAs Single strand siRNAs

Virus movement, symptom development, pathogenicity determinant Virus movement, symptom development pathogenicity determinant Required for replication Pathogenicity determinant Viral DNA replication Pathogenicity determinant Viral synergism

Long dsRNA binding PoLV P14 TCV P38 FHV B2

dsRNA and siRNAs dsRNA dsRNA

Symptom determinant Virion formation Virus movement NA

AGO degradation or inhibition CMV 2b PVX P25 ToRSV CP Polerovirus P0 TCV P38

AGO1 AGO4 AGO1 AGO1 AGO1 AGO1 AGO2

Virus movement Virus movement Virion formation Pathogenicity determinant Virion formation Virus movement

DCL inhibition RYMV P1 TCV P38

DCL1 upregulation, DCL4 DCL1 upregulation, DCL4

Pathogenesis, symptom development Virion formation Virus movement

Silencing amplification inhibition RDV PNS10 TuMV PVA VPg CaMV P6 (TAV) RGDV PNS12

RDR6 downregulation ND RDR6 and SGS3 downregulation RNA translation DRB4 inactivation Cell-to-cell movement, NPR1 downregulation, translational activator NA

Interference with siRNA methylation TMV 129K Tombusvirus P19

HEN1 siRNAs

RNA replication Virus movement, symptom development

NA Adenosine kinase inhibition

Symptom determinant Pathogenesis

Cellular transcription Cellular transcription

Viral synergism Transcription activation

DNA methylation interference TYMV P69 TGMV BCTV AL2 L2 CaLCuV AL2 Activation of cellular transcription EACMV AC2 CaLCuV AL2 Decoy RNA CaMV

35S RNA translational leader Overloading DCL

Translation, reverse transcription template

Viruses: African cassava mosaic virus (ACMV), East African cassava mosaic virus (EACMV), Beet curly top virus (BCTV), Beet yellows virus (BYV), Cabbage leaf curl virus (CaLCuV), Cauliflower mosaic virus (CaMV), Cucumber mosaic virus (CMV), Flock house virus (FHV), Potato virus A (PVA), Potato virus X (PVX), Pothos latent virus (PoLV), Rice gall dwarf virus (RGDV), Rice dwarf phytoreovirus (RDV), Rice yellow mottle virus (RYMV), Tobacco mosaic virus (TMV), Tomato bushy stunt virus (TBSV), Tomato golden mosaic virus (TGMV), Tomato ringspot virus (ToRSV), Tomato spotted wilt virus (TSWV), Turnip crinkle virus (TCV), Turnip mosaic virus (TuMV), Turnip yellow mosaic virus (TYMV), Wheat dwarf virus (WDV).

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Viral Suppressors of Gene Silencing

Small RNA Binding Potyviral HC-Pro, tombusviral p19, and others, bind siRNAs of viral and cellular origin including microRNAs and secondary siRNAs. Binding by silencing suppressors prevents association of siRNAs with AGO proteins. Consequently, RISC complexes are not formed and viral RNA, and host mRNAs, are not targeted. As a consequence, host genes normally downregulated by siRNAs, accumulate to levels higher than normal and plants display malformation that resemble symptoms caused by virus infection.

Inhibition of siRNA Biogenesis Several virus-encoded gene silencing suppressors interfere with the biogenesis of siRNA by at least two mechanisms: binding long dsRNA and altering homeostasis of core genetic components of the pathway (Table 1). Binding long dsRNA prevents access to DCL proteins and inhibits siRNA formation. Some suppressors induce degradation of cellular components needed for amplification of gene silencing, such as RDR6, SGS3 or DRB4. In both cases, the end result is a reduction in the biogenesis of virus-derived and endogenous siRNAs.

Homeostasis of Gene Silencing Components Core components of gene silencing (DCL, AGO, RDR, and DRB proteins) are targeted for degradation by some virus-encoded gene silencing suppressors (Table 1). Cucumber mosaic virus 2b and polerovirus P0 induced degradation of AGO1. AGO1 mainly associate with microRNAs and is essential for normal plant development. AGO2 and AGO4, are also needed for plant development and are targeted for degradation by silencing suppressors. Turnip crinkle virus P38 inhibits activity of DCL4, the first line of defense against viral RNA, and prevents biogenesis of DCL4-dependent siRNAs. Furthermore, DCL1 downregulates expression of DCL4 and DCL2, which have essential roles in siRNA biogenesis in both endogenous and antiviral gene silencing. Several viruses upregulate DCL1, consequently downregulating DCL4 and DCL2 to protect viral RNA from silencing.

Inhibition of Assembly or Activity of RNA-Induced Silencing Complexes The catalytic components of RNA-induced silencing complexes (RISC) are binary complexes formed between AGO proteins and siRNAs. Several virus-encoded silencing suppressors inhibit assembly or activity of RISC complexes by binding siRNAs or targeting AGO proteins for degradation (Table 1). Interestingly, in a complementary mechanism 2b, the silencing suppressor encoded by Cucumber mosaic virus and related species, binds to AGO1 and prevents cleavage of target mRNA.

Inhibition of Silencing Amplification A critical step in antiviral gene silencing is the systemic spread and amplification of silencing signals to establish a state of antiviral immunity away from the initially infected tissues (Fig. 2). The silencing signal consists of siRNAs, very likely in endosomes. After moving out of the initially infected cell, silencing is amplification by cellular RNA-dependent RNA polymerases (RDRs) and auxiliary proteins such as suppressor of gene silencing (SGS3) and double stranded RNA binding protein 4 (DRB4). Silencing amplification consists on the synthesis of new viral dsRNA that is processed by DCL proteins into secondary virus-derived siRNAs that move ahead of the virus to establish an antiviral state. Some virus-encoded gene silencing suppressors prevent spread or amplification of the gene silencing signal (Table 1). The mechanisms are dependent on siRNA sequestration, or induction of degradation of key components of secondary siRNA biogenesis, such as RDR6, SGS3 or DRB4.

Interference With DNA Methylation Plants silence geminiviruses by methylating their mini chromoses through transcriptional silencing. As counter-defense, some geminiviruses encode suppressors that interfere with DNA methylation (Table 1). AL2, encoded by Cabbage leaf curl virus, and L2 encoded by Beet curly top virus, interact with and inactivate adenosine kinase (ADK). ADK is a methyl cycle-associated enzyme required for the efficient production of S-adenosyl methionine (SAM), an essential methyltransferase co-factor. These viral suppressors interfere with methylation of both viral mini chromosomes and host chromosomes. The former promotes viral infection. The latter is associated to symptom development.

Activation of Cellular Transcription AL2, encoded by Cabbage leaf curl virus, and AC2 encoded by East African cassava mosaic virus are a transcription factor that suppresses post-transcriptional gene silencing by upregulating plants genes that negatively regulate gene silencing: Werner exonuclease-like 1 (WEL1), and regulator of gene silencing-calmodulin-like protein (rgsCam). In the presence of WEL1 and rgsCam gene silencing is less effective, thus favoring virus infection.

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Assays to Identify and Characterize Gene Silencing Suppressors Viral suppressors of gene silencing have been identified and characterized using a standard transient assay in Nicotiana benthamiana. The assay consists in providing a transgene encoding green fluorescent protein (GFP) as a reporter gene. Accumulation of GFP is initially evaluated using a hand-held UV lamp. The GFP transgene is normally silenced by the plant in two days. However, when a silencing suppressor is provided, GFP expression remains active for 4–5 days. Both the GFP transgene and genes being tested are delivered using Agrobacterium tumefaciens carrying binary vectors. This assay measures suppression of transgene silencing in infiltrated leaves (local silencing), and N. benthamiana can be wild type or constitutively express a GFP transgene (16C). The 16C line was developed by Dr. David Baulcombe’s group in 1998 and has been widely used to visualize local and systemic gene silencing. In 16C N. benthamiana plants, GFP is silenced in the infiltrated leaves and the silencing signal moves systemically. After 10–15 days of triggering local silencing, GFP silencing is evident in young laves, as a result of systemic gene silencing. Assays using 16C N. benthamiana have been used to identify suppressors that interfere with movement or perception of the gene silencing signal. In a variation of the assay, suppressor-deficient Turnip crinkle virus (TCV) tagged with GFP (TCV-GFP) is used instead of a single- or double-stranded GFP. TCV-GFP is unable to move cell-to-cell in N. benthamiana and is co-expressed with a gene of interest using A. tumefaciens carrying binary vectors. Silencing suppressors restore cell-to-cell movement to TCV-GFP and local infection foci visible to the naked eye are formed. After initial identification in transient assays using single- or double-stranded GFP or TCV-GFP as reporter genes, silencing suppressors are further tested in a heterologous virus, most commonly Potato virus X (PVX). PVX expressing a heterologous silencing suppressor normally cause more severe symptoms the wt PVX. In a complementary approach, silencing suppressors have been expressed constitutively expressed in model plants such as A. thaliana or N. benthamiana. Examples are potyviral HC-Pro, CMV 2b, and polerovirus P0. Interestingly, plants expressing one of these silencing suppressors, display development defects that resemble AGO mutants or symptoms characteristic of virus infected plants. These observations support the model that silencing suppressors interfere with endogenous gene silencing and cause the developmental malformations perceived as symptoms in virus infected plants. Using these assays, a large number of silencing suppressors have been identified in plant and insect viruses. Interestingly, proteins encoded by several human viruses have suppression activity in these assays. Vaccinia virus E3L and Influenza virus NS1 have gene silencing suppression activity. However, the biological role of this activity remains to be determined. Furthermore, Dengue virus proteins NS3 and NS4 are silencing suppressors. Since Dengue virus is transmitted by mosquitos these proteins likely protect the virus from silencing in the mosquito.

Viral Suppressors of Gene Silencing Need Cellular Factors Virus-encoded gene silencing suppressors are essential pathogenicity determinants necessary for viruses to interfere with gene silencing. Interestingly, some virus-encoded gene silencing suppressors interact with and need host factors to function. For example, RAV2, an ethylene-inducible transcription factor, is required for suppression of gene silencing by potyviral HC-Pro and TCV P38. In N. benthamiana, the calmodulin-like protein (Nbrgs-CaM) is an endogenous plant regulator of gene silencing that functions by repressing expression of RDR6. RDR6 and SGS3 participate in the biogenesis of secondary siRNAs necessary to amplify endogenous and antiviral gene-silencing signals against RNA viruses and geminiviruses. Infection of N. benthamiana by Tomato yellow leaf curl China virus and the associated ßC1 DNA satellite induces expression of Nbrgs-CaM that in turn downregulates RDR6 expression, thus reducing antiviral defense mediated by gene silencing.

Evolution of Viral Suppressors of Gene Silencing Most silencing suppressors are multifunctional and perform activities necessary for completion of the infection cycle, such as viral RNA replication, virus movement, virion formation or vector transmission (Table 1). Accordingly, virus-encoded silencing suppressors act through diverse mechanisms and are structurally diverse. Furthermore, even for suppressors with the same mechanism, their amino acid sequence is generally diverse. These features have limited the use of computational approaches to identify and characterize gene silencing suppressors and supports the model that silencing suppression has evolved multiple times, independently, through convergent evolution due to host pressure imposed through gene silencing as an antiviral mechanism.

Effect of Gene Silencing Suppressors on Plant Development A wide range of biological processes are regulated through silencing mechanisms, including plant development, organ formation, and stress responses in eukaryotes. Thus, perturbation of silencing pathways leads to changes in host gene expression that alter normal development. This effect is, at least in part, responsible for symptom development in virus infected plants. As an example, the abaxial-adaxial leaf polarity is regulated by siRNA gradients moving between cells. AGO7 and AGO10 are expressed specifically

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in the adaxial leaf primordial and in vascular tissues. AGO7 restricts activity of miR390 and directs a gradient of trans-acting small interfering RNAs from the adaxial to the abaxial side of developing leaves. Virus-encoded silencing suppressors such as potyviral HC-Pro, orthotospoviral NSs and tombusviral P19 bind cellular miRNAs, and secondary siRNAs. Additionally, potyviral VPg induces degradation of SGS3 and RDR6. TCV P38 inactivates DCL4, while polerovirus P0 and CMV 2b induce AGO1 degradation (Table 1). Collectively, these effects prevent activity of cellular microRNAs and the biogenesis or activity of trans-acting secondary siRNAs. Normal activity of cellular miRNAs and trans-acting secondary siRNAs is essential for plant development. As a consequence of silencing suppression, host genes normally downregulated by siRNAs, accumulate to levels higher than normal and plants display malformation that resemble symptoms caused by virus infection.

Silencing Suppressors Mediate Viral Synergism Viral synergism occurs when co-infection by two viruses results in disease with more severe symptoms than single virus infection. Most documented cases of viral synergism involve viruses in the family Potyviridae and depend on silencing suppressors HC-Pro or P1. Potyviral HC-Pro binds both endogenous and virus-derived siRNAs, their own and those derived from co-infecting viruses. SiRNA binding by HC-Pro prevents targeting of viral RNA and results in higher accumulation of the non-potyvirus and symptoms are more severe than those of single infection. An example of economic importance is maize lethal necrosis, caused the coinfection of Maize chlorotic mottle virus and Sugarcane mosaic virus. Maize lethal necrosis is a re-emerging disease that has reached epidemic proportion and is threatening food security in sub-Saharan Africa. In potato, co-infection by Potato virus X and Potato virus Y causes synergism. In pepper, co-infection by Cucumber mosaic virus and Pepper mottle virus causes synergism. Similarly, in papaya co-infection by Papaya mosaic and Papaya ringspot virus causes synergism. In all these cases, the non-potyvirus accumulates to higher levels in plants double infected, compared to plants single infected. Interestingly, accumulation of the potyvirus is similar in single or double-infected plants. The genetic determinant of synergism is potyviral silencing suppressor HC-Pro. There is one documented case of synergism in DNA viruses. In cassava, a synergistic disease is caused by the co-infection of African cassava mosaic virus (ACMV) and East African cassava mosaic virus (EACMV). ACMV and EACMV encode AC4 and AC2, respectively, that suppress gene silencing by different mechanisms. Furthermore, it is the only documented case of synergism between viruses of the same genus.

Viral Suppressors in Support of Biotechnology Several pharmaceutical products, including vaccines against human pathogens such as influenza viruses are produced in plants. The corresponding coding sequence is provided as a transgene, which often is turned off by gene silencing. In transient assays, this limitation has been resolved using virus-encoded gene silencing suppressors, which in enhance gene expression and bust yield of the pharmaceutical product of interest.

Further Reading Burgyan, J., Havelda, Z., 2011. Viral suppressors of RNA silencing. Trends in Plant Science 16, 265–272. Chapman, E.J., Prokhnevsky, A.I., Gopinath, K., Dolja, V.V., Carrington, J.C., 2004. Viral RNA silencing suppressors inhibit the microRNA pathway at an intermediate step. Genes & Development 18, 1179–1186. Csorba, T., Kontra, L., Burgyan, J., 2015. Viral silencing suppressors: Tools forged to fine-tune host-pathogen coexistence. Virology 479–480, 85–103. Deleris, A., Gallego-Bartolome, J., Bao, J., et al., 2006. Hierarchical action and inhibition of plant dicer-like proteins in antiviral defense. Science 313, 68–71. Diaz-Pendon, J.A., Ding, S.W., 2008. Direct and indirect roles of viral suppressors of RNA silencing in pathogenesis. Annual Review of Phytopathology 46, 303–326. Ding, S.W., 2010. RNA-based antiviral immunity. Nature Reviews Immunology 10, 632–644. Ding, S.W., Voinnet, O., 2007. Antiviral immunity directed by small RNAs. Cell 130, 413–426. Garcia-Ruiz, H., Carbonell, A., Hoyer, J.S., et al., 2015. Roles and programming of Arabidopsis ARGONAUTE proteins during Turnip mosaic Virus Infection. PLOS Pathogens 11, e1004755. Incarbone, M., Dunoyer, P., 2013. RNA silencing and its suppression: Novel insights from in planta analyses. Trends in Plant Science 18, 382–392. Johansen, L.K., Carrington, J.C., 2001. Silencing on the spot. induction and suppression of RNA silencing in the Agrobacterium-mediated transient expression system. Plant Physiology 126, 930–938. Korner, C.J., Pitzalis, N., Pena, E.J., et al., 2018. Crosstalk between PTGS and TGS pathways in natural antiviral immunity and disease recovery. Nature Plants 4, 157–164. Mlotshwa, S., Pruss, G.J., Vance, V., 2008. Small RNAs in viral infection and host defense. Trends in Plant Science 13, 375–382. Muhammad, T., Zhang, F., Zhang, Y., Liang, Y., 2019. RNA interference: A natural immune system of plants to counteract biotic stressors. Cells 8. Vanitharani, R., Chellappan, P., Pita, J.S., Fauquet, C.M., 2004. Differential roles of AC2 and AC4 of cassava geminiviruses in mediating synergism and suppression of posttranscriptional gene silencing. Journal of Virology 78, 9487–9498. Zhang, X., Yuan, Y.R., Pei, Y., et al., 2006. Cucumber mosaic virus-encoded 2b suppressor inhibits Arabidopsis Argonaute1 cleavage activity to counter plant defense. Genes & Development 20, 3255–3268. Zhao, J.H., Hua, C.L., Fang, Y.Y., Guo, H.S., 2016. The dual edge of RNA silencing suppressors in the virus-host interactions. Current Opinion in Virology 17, 39–44.

Virus-Induced Gene Silencing (VIGS) Xu Tengzhi, Ugrappa Nagalakshmi, and Savithramma P Dinesh-Kumar, University of California, Davis, CA, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of M.S. Padmanabhan, S.P. Dinesh-Kumar, Virus-Induced Gene Silencing (VIGS), In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00578-1.

Nomenclature ABA Abscisic acid AGO Argonaute CHS Chalcone synthase CP coat protein or capsid protein dsRNA Double stranded RNA HSP90 Heat shock protein 90 LIC Ligation-independent cloning mRNA Messenger RNA NLR Nucleotide-binding leucine rich repeat NOS Nopaline synthase

Glossary Agrobacterium-mediated genetic transformation Agrobacterium genetically transforms its host by transferring a well-defined DNA segment from its tumor-inducing (Ti) plasmid to the host-cell genome. Agro-infiltration It is a method used in plant biology and especially lately in plant biotechnology to induce transient expression of genes in a plant, or isolated leaves from a plant, or even in cultures of plant cells, in order to produce a desired protein. In the method a suspension of Agrobacterium tumefaciens is introduced into a plant leaf by direct injection or by vacuum infiltration, where after the bacteria transfer the desired gene into the plant cells via transfer of T-DNA. Argonaute It is a family of proteins that bind small noncoding RNAs, including miRNAs and siRNAs, and are guided to RNA targets through sequence complementarity, leading to the cleavage or translation inhibition of mRNAs, including viral RNAs. Co-suppression phenomenon understood as posttranscriptional gene silencing, which can be triggered by introducing transgenes or a certain viruses expressing the target gene. CRISPR-Cas Clustered regularly interspaced short palindromic repeats (CRISPR) and CRISPR-associated proteins (Cas) is a prokaryotic adaptive immune system that can be leveraged for genetic material to be added, removed, or altered at particular locations in the plant genome, or to directly target viral DNA or RNA molecules. Dicer It is a multidomain ribonuclease that processes double-stranded RNAs to 21-nt small interfering RNAs during RNA interference and excises microRNAs from precursor hairpins.

Encyclopedia of Virology, 4th Edition, Volume 3

ORF Open reading frame(s) PDS Phytoene desaturase PTGS Post-transcriptional gene silencing RdRp RNA-dependent RNA polymerase RISC RNA-induced silence complex RNAi RNA interference SGN Solanaceae genomics network UTR Untranslated region VIGS Virus-induced gene silencing VPGD VIGS phenomics and functional genomics database VPI Vascular puncture inoculation

Forward genetics Approaches used to identify directly genes responsible for a particular phenotype of an organism (DNA). Functional genomics The study of genes with respect to the role they play within biological processes. Knockdown Reduction in the expression of a gene. Knockout Complete inhibition of gene expression. MicroRNAs These are a class of small non-coding B21–24 nt RNAs that are involved in the regulation of gene expression at the post-transcriptional level by degrading target mRNAs, including viral RNAs, and/or inhibiting their translation. Post-transcriptional gene silencing Silencing of an endogenous gene caused by the introduction of a homologous dsRNA, transgene or virus. In PTGS, the transcript of the silenced gene is synthesized but does not accumulate because it is rapidly degraded. Reverse genetics Approaches used to define the function of a gene or sequence of DNA within the context of the organism (RNA, proteins). RNA interference It is a ribonucleoprotein complex that incorporates one strand of a miRNA or a siRNA duplex for RNAi. RNA-induced silencing complex It is a ribonucleoprotein complex that incorporates one strand of a miRNA or a siRNA duplex for RNAi. Small interfering RNA It is a class of 20–25 nt non-coding dsRNA molecules. Trans-acting siRNA These are siRNA that repress plant gene expression through post-transcriptional gene silencing.

doi:10.1016/B978-0-12-809633-8.21530-6

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Introduction Virus-induced gene silencing (VIGS) is an effective forward and reverse genetics approach in plants. VIGS uses the antiviral defense mechanism in plants to transiently downregulate specific transcripts through a recombinant vector. The VIGS vector, which carries a partial sequence from the gene to be silenced, is introduced into the plant cell via Agrobacterium tumefaciens-mediated transient transformation or using in vitro-generated transcripts. Double stranded RNA (dsRNA) molecules generated during viral infection trigger the RNA interference (RNAi) pathway; leading to the degradation of messenger RNAs (mRNAs) homologous to the dsRNA and subsequent downregulation of the corresponding gene. The first VIGS vector, based on Tobacco mosaic virus (TMV), was developed in 1995. The last two decades have seen a significant increase in the number of viruses that have been adapted for VIGS to study gene function in plants. VIGS has major advantages over the labor-intensive transgenic approaches including speed, efficacy and cost. These attributes have made VIGS a versatile functional genomic tool in a wide range of plant species to unravel gene functions involved in a variety of plant biological processes.

PTGS and VIGS The term “cross-protection” was first described in the 1920s as a phenomenon of induced resistance in a host plant against one virus or viroid strain infection if the host plant had been previously infected with another milder strain of the same virus or viroid. However, the mechanistic basis of this phenomenon was not understood. In 1990, researchers attempted to develop an enhanced purple colored petunia flower by overexpressing a pigmentation gene chalcone synthase (CHS). Unexpectedly, the flowers of CHS overexpressing plants were either entirely white or a mottled purple/white. This phenomenon was named “co-suppression” and considered to be an oddity of petunia. In the following few years, the same phenomenon was reported in many other plant species. The “cross-protection” and “co-suppression” phenomenon were soon understood as post-transcriptional gene silencing (PTGS), which could be triggered by introducing transgenes or using viruses as gene carrier. Around the same time, a similar phenomenon was described in the fungus Neurospora crassa and named as “quelling”. This phenomenon has since been observed in many organisms, including nematodes, flies, and mammals; coming to be known more widely as RNA interference (RNAi). These cross-kingdom PTGS/quelling/RNAi are evolutionary conserved and induced by dsRNA. The term VIGS was coined in 1997 and used to describe the symptoms recovery phenomenon of plants upon viral infection. As mentioned above, viral infection activates the plant RNAi-mediated antiviral defense mechanism, PTGS. The machinery involved in PTGS targets the viral genome as an antiviral defense. This was exploited as the basis of VIGS. Now, VIGS specifically refers to the reverse genetics technique for down-regulation of target gene expression in plants by recombinant viruses which carry a partial sequence of the target gene. PTGS is activated upon virus infection in plants. During the viral replication process, dsRNA intermediates are generated. Synthesis of dsRNA is mediated by RNA-dependent RNA polymerase (RdRp). DNA viruses employ the host RdRp for the dsRNA synthesis, while RNA viruses encode their own RdRp for the dsRNA production. Viral dsRNAs are recognized as host aberrant RNAs and cleaved by the host plant’s Dicer-like ribonuclease, producing short interfering RNA (siRNA) duplex molecules which are of 21–24 nt in length. The siRNA duplex can then be recognized and unpaired by Argonaute (AGO) protein into a guide strand and a passenger strand in an ATP-dependent manner. The Passenger strand is degraded while the Guide strand is loaded into a multiprotein complex known as the RNA-induced silence complex (RISC). The RISC complex, harboring siRNA, scans the mRNA population inside the cell and specifically targets transcripts that are complementary to siRNA, resulting in cleavage of target transcript. As a result, the mRNA level of the target gene is reduced. The silencing signal can be further amplified by plant RNAdependent RNA polymerase 6 (RDR6) and spread systemically throughout the plant, causing systemic gene silencing.

Establishment of a VIGS System To develop a virus vector into a more efficient VIGS tool, the selected virus should have a broad host range and should cause only mild disease symptoms. In this way, the VIGS-vector-induced silencing phenotype will not be obscured by disease symptoms. The best VIGS vectors are characterized by high silencing efficiency as well as persistent silencing phenotype. Furthermore, uniform expression of a silencing phenotype, in a tissue-independent manner, is also an important factor that should be considered when choosing a virus for VIGS vector development. In the last two decades, many plant viruses have been used to develop VIGS vectors. Most VIGS vectors are derived from RNA viruses, with a few DNA viruses and satellites having been used as well (Table 1). For development of a VIGS vector, the viral genome is cloned under the control of a constitutive promoter such as Cauliflower mosaic virus (CaMV) 35S promoter and a terminator such as nopaline synthase (NOS) terminator in a T-DNA binary vector. The VIGS T-DNA vector, containing the target gene insert, is transferred into plants via Agrobacterium tumefaciens-mediated transient plant transformation system or DNA of VIGS vector could be delivered by biolistic bombardment. In addition, transcripts of T7 promoter-based VIGS vector generated by in vitro transcription could be used for rub-inoculation onto plants. Agroinfiltration is the most widely used method for VIGS because it is inexpensive and highly reproducible. Syringe infiltration is commonly used to introduce the agrobacterium into the

Virus-Induced Gene Silencing (VIGS)

Table 1

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List of plant viruses used as VIGS vectors for gene silencing and their experimental plant hosts

Virus

Genus/Family

Host species

Apple latent spherical virus (ALSV)

Cheravirus Secoviridae

Bamboo mosaic virus (BaMV) Barley stripe mosaic virus (BSMV) Bean pod mottle virus (BPMV) Brome mosaic virus (BMV) Cucumber mosaic virus (CMV) Cymbidium mosaic virus (CymMV) Foxtail mosaic virus (FoMV) Pea early browning virus (PEBV) Potato virus X (PVX) Tobacco rattle virus (TRV)

Potexvirus Alphaflexiviridae Hordevirus Virgaviridae Comovirus Secoviridae Bromovirus Bromoviridae Cucumovirus Bromoviridae Potexvirus Alphaflexiviridae Potexvirus Alphaflexiviridae Tobravirus Virgaviridae Potexvirus Alphaflexiviridae Tobravirus Virgaviridae

Turnip yellow mosaic virus (TYMV)

Tymovirus Tymoviridae

Chenopodium quinoa, soybean, apple and Nicotiana benthamiana Brachypodium distachyon Wheat, barley Soybean Maize Nicotiana benthamiana and maize Orchid Barley, wheat, foxtail millet, and maize Medicago truncatula and pea Nicotiana benthamiana Nicotiana benthamiana, tobacco, tomato, potato, pepper, Arabidopsis, cotton, poppy and others Chinese cabbage and Arabidopsis

DNA virus

African cassava mosaic virus (ACMV) Cabbage leaf curl virus (CaLCuV) Cotton leaf crumple virus (CLCrV) Pepper huasteco yellow vein virus (PHYVV) Rice tungro bacilliform virus (RTBV) Tomato golden mosaic virus (TGMV)

Begomovirus Geminiviridae Begomovirus Geminiviridae Begomovirus Geminiviridae Begomovirus Geminiviridae Tungrovirus Caulimoviridae Begomovirus Geminiviridae

Cassava Arabidopsis Cotton Pepper Rice Nicotiana benthamiana

Satellite virus

Satellite Bamboo mosaic virus (satBaMV) Tomato yellow leaf curl China betasatellite (TYLCCNV)

Potexvirusa Begomovirusa Geminiviridae

Brachypodium distachyon Tomato

RNA virus

a

Genus of the helper virus.

Fig. 1 Silencing phenotype of the phytoene desaturase (PDS) gene in Nicotiana benthamiana. Four week old N. benthamiana plants infected with TRV-NbPDS show photobleaching phenotype (right panel) compared to the TRV-VIGS vector (left panel) infected plant. Pictures were taken two weeks post VIGS vector inoculation.

plants. However, other delivery routes such as spraying, vacuum infiltration and agro-drench have also been used to deliver VIGS vector containing T-DNA vectors in plants. Virus-encoded silencing suppressor genes can reduce VIGS silencing efficiency and should be removed or modified in the VIGS vector. Some non-essential genes, for example, those that are not necessary for viral replication, cell-to-cell movement and systemic movement, could be removed from the viral genome while constructing a VIGS vector. To clone a target gene fragment into a VIGS vector, an appropriate position in the viral genome should be selected in such a way that it does not interfere with viral lifecycle. In some cases, a sub-genomic promoter from a related virus has been used in a VIGS vector to drive the expression of a target gene for silencing. To clone the target gene fragment into the VIGS vector, the restriction/ligation, gateway cloning, or ligation-independent cloning (LIC) strategies have been used. The optimal gene fragment length for efficient silencing of a target gene is between 200 and 500 bp. However, gene fragments as short as 23 nt are sufficient to knockdown target gene expression. The silencing

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Fig. 2 VIGS assay applied to tomato fruit. (A) Phenotype of Nor-silenced fruit. Vector only (pTRV2–00) and PDS-silenced (pTRV2-PDS) fruits were used as the control. (B) Flow chart of the VIGS assay for tomato fruit. To construct the pTRV2 vector, a fragment of the target gene (FTG) was inserted into the multiple cloning site (MCS). pTRV1 and pTRV2 plasmids were independently transferred to A. tumefaciens (GV3101). The cells in the LB cultures containing pTRV1 and pTRV2 were harvested and mixed at a ratio of 1:1 after resuspension in Agrobacterium infiltration buffer (AI buffer); the mixture was used to infiltrate the carpopodium of the tomato fruit attached to the plant at 7–10 DPA. (C) The silencing efficiency of the Nor gene in AC tomato fruit at the red-ripe stage (RR). CaMV 35S promoter (35S), nopaline synthase terminator (NOSt), coat protein (CP). Luria-Bertani medium (LB). Asterisks indicate a significant difference as determined using Student’s t-test (**, P o 0.01). Reproduced Fig. 1 from Yuan, X.Y., Wang, R.H., Zhao, X.D., Luo, Y.B., Fu, D.Q., 2016. Role of the tomato non-ripening mutation in regulating fruit quality elucidated using iTRAQ protein profile analysis PLOS ONE 11, e0164335. Available at: https://doi.org/10.1371/journal.pone.0164335.

efficiency varies depending on the target region selected. Therefore, multiple fragments targeting different regions of the target gene of interest should be tested. In view of this, Solanaceae Genomics Network (SGN) VIGS Tool (See Relevant Websites Section) could be used to select an optimal target gene fragment for cloning into a VIGS vector.

VIGS Vectors Many VIGS vectors developed to-date have been used for gene silencing in dicot plants. Since Nicotiana benthamiana is susceptible to a wide range of plant viruses and amenable to agro-infiltration it is widely used to test the efficacy of a newly developed VIGS vector. The

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Fig. 3 The RLKs and WRKYs are required for resistance against V. dahliae in cotton via VIGS analysis. (A) Silencing of CLA1 gene in G. hirsutum variety Zhongzhimian KV3. Left, plant infiltrated with CLCrV-based empty vector (CLCrV-00); right, plant infiltrated with CLCrV-CLA1. (B) Relative expression levels of candidate gene in the silenced and non-silenced cotton plants at 20 dpi were determined through qRT-PCR. Asterisk represents the data point that was statistically different from the non-silenced (p o 0.05) analyzed by one-way ANOVA, using the SAS 8.1. The vertical bars indicate standard Deviation. (C) a, Negative control-silenced upland cotton (Zhongzhimian KV3, CLCrV-00); b, GhGsSRKsilenced Zhongzhimian KV3 plant; c, GhWRKY2-silenced Zhongzhimian KV3 plant; d, GhWRKY29-silenced Zhongzhimian KV3 plant; e, GhFLS2silenced Zhongzhimian KV3 plant; f, Negative control-silenced upland cotton (86–1, CLCrV-00); g, GhWRKY13-silenced 86–1plant. (D) The disease index (DI) after silencing different genes. The results are presented as mean 7standard deviation (SD) from three replicates with at least 20 plants per replicate. Black represents candidate gene silenced in Zhongzhimian KV3, and blue represents candidate gene silenced in 86–1. Asterisk represents the data point that was statistically different from wild-type and CLCrV-00 plants (p o 0.05) analyzed by one-way ANOVA, using the SAS 8.1. The vertical bars indicate standard Deviation. Reproduced Fig. 3 from Zhang, W., Zhang, H., Liu, K., et al., 2017. Large-scale identification of Gossypium hirsutum genes associated with Verticillium dahliae by comparative transcriptomic and reverse genetics analysis. PLOS ONE 12 (8), e0181609. Available at: https://doi.org/10.1371/journal.pone.0181609.

Tobacco rattle virus (TRV)-VIGS vector is by far the most widely used VIGS vector due to its broad host range and high silencing efficiency in addition to its very mild disease symptoms. TRV is a bipartite, positive-strand RNA virus consisting of RNA1 and RNA2 genomes. Since the RdRp, movement protein, and 16K protein encoded by RNA1 are essential for TRV infection, the RNA1 is unmodified in the TRV-VIGS system. The cDNA corresponding to RNA1 is cloned under the control of a 35S promoter and a NOS terminator (TRV1). In RNA2, the coding region required for vector transmission has been removed and the cDNA corresponding to the remaining parts are cloned between a 35S promoter and a NOS terminator. In RNA2 vector (TRV2), a multiple cloning site, gateway cloning cassette, or ligation-independent cloning cassette has been introduced downstream of intergenic region after the coat protein (CP) open reading frame (ORF). TRV-VIGS has been used successfully for gene function studies in more than 25 plant species including a large number of plants in Solanaceae (Figs. 1 and 2), as well as in Malvaceae (Fig. 3), Brassicaceae, Euphorbiaceae, Papaveraceae and Rosaceae. In legumes, Bean pod mottle virus (BPMV) and Pea early browning virus (PEBV) have been used successfully as VIGS vectors.

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Fig. 4 PDS and magnesium chelatase subunit H (ChlH) gene silencing phenotypes in barley, wheat and B. distachyon. (A) Barley showing phenotypes typical of suppression of ChlH and PDS by BSMV:HvChlH or BSMV:HvPDS inserts of the indicated lengths. (B) Upper panel: Wheat inoculated with BSMV:TaChlH250 showing more than 90% of inoculated plants developing the chlorotic phenotype associated with suppression of the ChlH gene. Middle panel: PDS and ChlH gene silencing phenotypes on wheat leaves infected with BSMV:TaChlH and BSMV:PDS derivatives with different length inserts. Bottom panel: PCR amplification of transcripts from leaves shown in the middle panel. (C) B. distachyon leaves showing effects of PDS suppression after infection with BSMV:BdPDS inserts. In these experiments, plants were inoculated at the two-leaf stage with infected N. benthamiana sap harboring BSMV derivatives targeting PDS and ChlH cognate genes from each species. Leaf photographs were taken at 14 dpi, and fragment lengths from each source are indicated as subscripts above each leaf. Relative transcript levels of PDS and ChlH genes in the leaves infected with the different BSMV derivatives are shown under the leaf photographs. RNA extracted from the leaves was subjected to semi-quantitative RT-PCR amplification (28 cycles for wheat and barley, 31 cycles for B. distachyon) with the gene-specific oligonucleotide primers. Amplified species-specific 18S rRNA served as internal controls for each species. Reproduced Fig. 6 from Yuan, C., Li, C., Yan, L., et al. 2011. A high throughput Barley stripe mosaic virus vector for virus induced gene silencing in monocots and dicots. PLOS ONE 6, e26468. Available at: https://doi.org/10.1371/journal.pone.0026468.

Very few vectors have been developed for gene silencing in monocot plants (Table 1). Monocots are generally recalcitrant to agro-infiltration. As such, mechanical inoculation methods such as rub-inoculation of crude sap from infected N. benthamiana plants, in vitro generated VIGS vector transcripts, or vascular puncture inoculation (VPI) of in vitro transcripts or virions has been used to introduce VIGS vectors into monocots. Brome mosaic virus (BMV) and Barley stripe mosaic virus (BSMV; Fig. 4) VIGS vectors have been used to silence genes in maize, wheat and barley. Recently, Foxtail mosaic virus (FoMV)-based VIGS vector has been used to silence genes in barley, wheat, foxtail millet, and maize and Cucumber mosaic virus (CMV)-based vector has been used to silence genes in maize. Generally, Phytoene desaturase (PDS), a gene involved in carotenoid biosynthesis, has been used as a target to test the efficacy of newly developed VIGS vectors. Since carotenoid synthesis is important for photosynthesis and photoprotection, low levels of carotenoids result in a degradation of chlorophyll. This is manifested in a visible photobleaching phenotype in PDS silenced plants (Fig. 1). This visible feature makes PDS an excellent positive control for monitoring the silencing efficiency.

VIGS Application VIGS vectors have been used widely as a reverse and forward genetic tool to unravel the functions of genes involved in various biological processes such as disease resistance, cell death, abiotic stress, cell signaling, flower development (Fig. 5), fruit development (Fig. 2), secondary metabolite biosynthesis, and evolutionary developmental biology. The role of Heat Shock Protein 90 (HSP90) in Rx1 nucleotide-binding leucine rich repeat (NLR) class receptor-mediated immunity was uncovered in a forward genetics by screening nearly 5000 cDNAs in TRV-VIGS vector in N. benthamiana plants. The role of autophagy in immunity and cell death in plants was uncovered by screening TRV-VIGS library. Similar TRV-VIGS library was used to identify the role for ribosomal proteins, RPL12 and RPL19, in non-host disease resistance against bacterial pathogens in N. benthamiana. Numerous studies have used VIGS as a reverse genetics tool to study gene function in the literature. NCBI PubMed database using the term “VIGS” lists 828 publications. Majority of these publications describe using VIGS to discover gene function in various plant biological processes. By far the most application of VIGS is in the disease resistance area (168 publications in NCBI). VIGS has been used to study function of genes involved in disease resistance in various plants including N. benthamiana, tomato, pepper, potato, soybean, wheat, barley, maize, cotton, cassava, and other plants. In addition to biotic interaction, VIGS has been used to study abiotic stress responses. VIGS vectors based on TRV, BSMV, BPMV, PEBV, and Tomato yellow leaf curl China virus (TYLCCNV) have been used successfully to study genes function in abiotic

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Fig. 5 Virus-induced gene silencing of Thalictrum clavatum AGAMOUS ortholog TcAG-1 results in homeotic floral phenotypes. (A) Flower silencing phenotypes of TcAG-1, relative to controls. Ai, Untreated flower of T. clavatum showing sepals (se), stamens (st) and carpels (ca); Aii: strongly silenced flower in TRV2-TAG-1 treated plant, showing an array of sepals and no stamens nor carpels, all reproductive organs have been homeotically converted to sepals; Aiii: intermediate phenotype with partial conversion of organs and some normal ones; Aiv: detail of dissected organs in an untreated flower (sepal, stamen, carpel, from left to right); Av: detail of all sepaloid dissected organs from a strong TcAG-1 silencing phenotype (from the outside to the inside of the flower, left to right); Avi: detail of sample chimeric organs, arrows point to anther tissue on the edges of an internal “sepal”. Scale bar ¼ 1 mm. (B) Gene expression by Reverse Transcriptase (RT)-PCR in TcAG-1 silenced plants compared to controls. Untreated and mock-treated (empty TRV2) plants are compared to TRV2-TAG-1 treated plants showing strong homeotic conversions (Aii, Av). RT-PCR was performed with locus-specific primers to the housekeeping gene ACTIN (loading control); to the MADS box gene TcAG-1 and to the viral transcripts TRV1/TRV2. For TRV2: larger bands result from the presence of insert, the smaller band from an empty TRV2 (mock control). Reproduced Fig. 4 from Di Stilio, V.S., Kumar, R.A., Oddone, A.M., et al., 2010. Virus-induced gene silencing as a tool for comparative functional studies in Thalictrum. PLOS ONE 5, e12064. Available at: https://doi.org/10.1371/journal.pone.0012064.

stress responses including drought, highlight, iron deficiency, oxidative stress, abscisic acid (ABA) response, salt stress, osmatic stress, dehydration and nutrient deficiency in crop plants such as tomato, pepper, pea, soybean, wheat, and barley. Many groups have also used VIGS to study genes involved in various plant developmental pathways including flower and fruit development. Genes involved in flower development has not only been studied in model plants such as N. benthamiana and Arabidopsis but also in crop plants such as tomato, petunia, soybean, and cherry. In addition, TRV-based VIGS has been used in comparative studies involving flower development genes in basal eudicots such as Aquilegia, Cysticapnos vesicaria, Papaver somniferum (opium poppy) and Eschscholzia californica (California poppy); Gerbera hybrida, Phalaenopsis orchids, and rose. TRV-based VIGS has been used successfully to uncover the role of genes involved in fruit development in tomato, pepper, peach, and strawberry. Recently, a database describing phenotypes resulting from silencing of approximately 1300 genes in N. benthamiana has been developed, VIGS phenomics and functional genomics database (VPGD; See Relevant Websites Section). Since genes in N. benthamiana

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have homologs in other Solanaceae plants such as tomato, potato and pepper; one could infer the function of the corresponding genes in these crop plants.

Advantages and Limitations VIGS is an efficient forward genetic approach to identify loss-of-function phenotypes of specific genes. TRV-VIGS requires less than 2 weeks to knockdown target gene expression in N. benthamiana plants. However, it usually takes 3–4 months to generate transgenic N. benthamiana with the traditional Agrobacterium-mediated genetic transformation; homozygous lines require additional time. Additionally, the transformation process is labor-intensive and expensive. The VIGS could be used to silence a single gene or even multiple genes. Since VIGS has been applied in a broad range of plant species, it is possible to conduct comparative studies using homologous genes in different plant species. Many plants are highly recalcitrant to genetic transformation. For some genes in sexually propagated plants, the mutation or antisense-mediated knockdown is lethal. In these cases, VIGS can serve as a valuable alternative for gene function studies as it eliminates the need to establish a stable genetic transformation method. Silencing of a specific gene within a multi-gene family and silencing multi-copy genes in polyploid plants is also feasible by VIGS method by choosing the targeting sequence derived from the most conserved region among gene family members. Similarly, one could use 30 untranslated region (UTR) to specifically silence a single gene within a gene family. VIGS, like any technique, has limitations. The silencing effect of VIGS is transient. However, silencing by TRV-VIGS has been shown to last for longer time. Another limitation is that VIGS fails to completely knockdown target gene expression. In some cases this imperfect knockdown may not manifest in a phenotypic change compared to the control plants. Since the VIGS vector is essentially an infectious plant pathogen, the silencing process along with virus infection may interfere with the plant metabolism which could affect the results of some studies. Therefore, it is always important to use negative control such as VIGS vector alone (Fig. 1) without any target insert in VIGS experiments to make sure that the observed phenotype is specifically due to the knockdown of a target gene. VIGS is an ideal method for preliminary identification of function of genes due to its quick and easy features. Following preliminary identification with VIGS, the most promising candidate genes could be further validated with other approaches.

Conclusion and Prospects VIGS is an efficient forward and reverse genetic technique that exploits the plant’s PTGS mechanism. It facilitates the downregulation of single genes or gene families. A large number of plant viruses have been engineered as VIGS tools for gene function studies in a variety of plant species. In the last 20 years, VIGS has made a significant contribution to gene function studies in various plant processes. With the availability of more plant genome sequences in the post-genomic era, the improvement of existing VIGS vectors as well as the development of new VIGS vectors for additional plant species is required for VIGS-based functional genomic studies to achieve its full potential in plants.

Further Reading Bachan, S., Dinesh-Kumar, S.P., 2012. Tobacco rattle virus (TRV)-based virus-induced gene silencing. Methods Molecular Biology 894, 83–92. Burch-Smith, T.M., Anderson, J.C., Martin, G.B., Dinesh-Kumar, S.P., 2004. Applications and advantages of virus-induced gene silencing for gene function studies in plants. Plant Journal 39, 734–746. Ding, X.S., Mannas, S.W., Bishop, B.A., et al., 2018. An improved Brome mosaic virus silencing vector: Greater insert stability and more extensive VIGS. Plant Physiology 176, 496–510. Di Stilio, V.S., Kumar, R.A., Oddone, A.M., et al., 2010. Virus-induced gene silencing as a tool for comparative functional studies in Thalictrum. PLOS ONE 5, e12064. Available at: https://doi.org/10.1371/journal.pone.0012064. Dommes, A., Gross, T., Herbert, D.B., Kivivirta, K.I., Becker, A., 2019. Virus-induced gene silencing: empowering genetics in non-model organisms. Journal of Experimental Botany 70, 757–770. Fu, D.Q., Zhu, B.Z., Zhu, H.L., Jiang, W.B., Luo, Y.B., 2005. Virus-induced gene silencing in tomato fruit. Plant Journal 43, 299–308. Jiang, C.Z., Chen, J.C., Reid, M., 2011. Virus-induced gene silencing in ornamental plants. Methods in Molecular Biology 744, 81–96. Kant, R., Dasgupta, I., 2019. Gene silencing approaches through virus-based vectors: speeding up functional genomics in monocots. Plant Molecular Biology 100, 3–18. Kumagai, M., Donson, J., Della-Cioppa, G., et al., 1995. Cytoplasmic inhibition of carotenoid biosynthesis with virus-derived RNA. Proceedings of the National Academy of Sciences of the United States of America 92, 1679–1683. Lacomme, C., 2015. Strategies for altering plant traits using virus-induced gene silencing technologies. Methods in Molecular Biology 1287, 25–41. Liu, Y., Schiff, M., Czymmek, K., et al., 2005. Autophagyregulates programmed cell death during the plant innate immune response. Cell 121, 567–577. Liu, N., Xie, K., Jia, Q., et al., 2016. Foxtail mosaic virus-induced gene silencing in monocot plants. Plant Physiology 171, 1801–1807. Lu, R., Malcuit, I., Moffett, P., et al., 2003. High throughput virus-induced gene silencing implicates heat shock protein 90 in plant disease resistance. EMBO Journal 22, 5690–5699. Mei, Y., Zhang, C., Kernodle, B.M., Hill, J.H., Whitham, S.A., 2016. A Foxtail mosaic virus vector for virus-induced gene silencing in maize. Plant Physiology 171, 760–772. Nagaraj, S., Senthil-Kumar, M., Ramu, V.S., Wang, K., Mysore, K.S., 2016. Plant ribosomal proteins, RPL12 and RPL19, play a role in non-host disease resistance against bacterial pathogens. Frontiers in Plant Science 6, 1192.

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Ramegowda, V., Mysore, K.S., Senthil-Kumar, M., 2014. Virus-induced gene silencing is a versatile tool for unraveling the functional relevance of multiple abiotic-stressresponsive genes in crop plants. Frontiers in Plant Science 5, 323. Rosa, C., Kuo, Y.W., Wuriyanghan, H., Falk, B.W., 2018. RNA interference mechanisms and applications in plant pathology. Annual Review Phytopathology 56, 581–610. Senthil-Kumar, M., Mysore, K.S., 2011. New dimensions for VIGS in plant functional genomics. Trends in Plant Science 16, 656–665. Senthil-Kumar, M., Wang, M., Chang, J., et al., 2018. Virus-induced gene silencing database for phenomics and functional genomics in Nicotiana benthamiana. Plant Direct 2, e00055. Wang, R., Yang, X., Wang, N., et al., 2016. An efficient virus-induced gene silencing vector for maize functional genomics research. Plant Journal 86, 102–115. Yuan, C., Li, C., Yan, L., et al., 2011. A high throughput Barley stripe mosaic virus vector for virus induced gene silencing in monocots and dicots. PLOS ONE 6, e26468. Available at: https://doi.org/10.1371/journal.pone.0026468. Yuan, X.Y., Wang, R.H., Zhao, X.D., Luo, Y.B., Fu, D.Q., 2016. Role of the tomato non-ripening mutation in regulating fruit quality elucidated using iTRAQ protein profile analysis. PLOS ONE 11, e0164335. Available at: https://doi.org/10.1371/journal.pone.0164335. Zhang, W., Zhang, H., Liu, K., et al., 2017. Large-scale identification of Gossypium hirsutum genes associated with Verticillium dahliae by comparative transcriptomic and reverse genetics analysis. PLOS ONE 12 (8), e0181609. Available at: https://doi.org/10.1371/journal.pone.0181609.

Relevant Websites http://vigs.solgenomics.net/ SGN-VIGS Sol Genomics Network. http://vigs.noble.org VIGS Database Noble Research Institute.

Alfalfa Mosaic Virus (Bromoviridae) L Sue Loesch-Fries, Purdue University, West Lafayette, IN, United States r 2021 Published by Elsevier Ltd. This is an update of J.F. Bol, Alfalfa Mosaic Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00635-X.

Nomenclature aa Amino acid(s) CP Capsid or coat protein kb Kilobases; the size of a ssDNA or ssRNA molecule kDa Kilodaltons; the size of a protein MP Movement protein MVBs Multi-vesicular bodies

Glossary Multi-vesicular bodies Organelles bound by a membrane that contain intraluminal vesicles that mediate transport between the trans-Golgi network and vacuoles. Poly(A)-binding protein Regulatory proteins that bind to the poly(A) tail of mRNAs.

nm Nanometer(s) nt Nucleotide(s) PABP Poly(A)-binding protein Poly(A) Polyadenylated TLS Transfer RNA-like structure tRNA transfer RNA UTR Untranslated region

T-DNA vector Plasmid with the T-DNA (transferred DNA) sequence of the Ti plasmid of Agrobacterium tumefaciens, which is transferred to plant cells where it can be transiently expressed or become integrated in the plant genome. Transgenic P12 tobacco Tobacco plants that express P1 and P2 replicase proteins from nuclear genes inserted via a T-DNA vector.

History Alfalfa mosaic virus (AMV) was identified in 1931 as the causal agent of an economically important disease in alfalfa. Purification of AMV around 1960 showed that virus preparations contained bacilliform particles of different length. Fractionation of these particles by sucrose-gradient centrifugation revealed that the four major components each contained a specific type of RNA, termed RNAs 1, 2, 3, and 4. Initially, an analysis of the biological activity of the viral nucleoproteins and RNAs resulted in a puzzle. A mixture of the three largest viral particles was fully infectious, but a mixture of RNAs 1, 2, and 3, purified from these particles, was not. At the RNA level, RNA4 was required in the inoculum to initiate infection. In 1971, it became clear that the AMV genome consisted of RNAs 1, 2, and 3, and that a mixture of these genomic RNAs became infectious only after addition of capsid or coat protein (CP) or its sub-genomic messenger, RNA4. High-affinity binding sites for CP were identified in the AMV RNAs in 1972 and could be localized to the 30 termini of the RNAs in 1978. From 1975 onwards it became clear that, similar to AMV, the ilarviruses required the CP to initiate infection. Moreover, the CPs of AMV and the ilarviruses could bind to the 30 termini of each other’s RNAs, and could be freely exchanged in the initiation of infection. Currently, AMV is the type species of the genus Alfamovirus and is classified together with the genus Ilarvirus in the family Bromoviridae.

Taxonomy and Classification In addition to the genera Alfamovirus and Ilarvirus, the family Bromoviridae contains the genera Bromovirus, Cucumovirus, Anulavirus, and Oleavirus. These viruses all have three positive-sense single-stranded genomic RNAs packaged into separate particles with sub-genomic, defective, or satellite RNAs. The particles are bacilliform or spherical. AMV has been placed into a separate genus from the other viruses because of its mode of transmission (aphids, pollen, and seed), its shape (bacilliform), and its very wide host range. In addition, AMV requires CP for the initiation of infection (termed ‘genome activation’), whereas other members of the family do not, except for the ilarviruses. It has been suggested that the alfamoviruses and the ilarviruses be combined into a single genus, but this has yet to be resolved. The ilarviruses differ from AMV, however, by mode of transmission (thrips/pollen) and shape (irregular spheres). Many isolates of AMV are known which are closely related serologically and by nt sequence similarity. The complete nt sequences of AMV isolate 425 (Leiden [L] and Madison [M] isolates) were the first determined; GenBank now contains 11,602 full and partial sequences of many additional strains and isolates. The genetic diversity of the sequences is generally low with agro-ecological factors affecting their distribution.

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Fig. 1 Electron micrograph of Alfalfa mosaic virus (Alfamovirus, Bromoviridae). A. Bacilliform bottom particle, B, middle particle, M, and top particle Tb. Scale bar is 100 nm. (Courtesy of J. Bol). B. Cryo-electron micrograph of a T¼1 AMV particle. Reproduced from http://viperdb. scripps.edu; Kumar, A., Reddy, V.S., Yusibov, V., et al., 1997. The structure of Alfalfa mosaic virus capsid protein assembled as a T¼1 icosahedral particle at 4.0-Å resolution. Journal of Virology 71, 7911–7916.

Particle Structure and Composition Fig. 1 shows an electron micrograph of AMV. The four major classes of particles in AMV preparations are called bottom component (B), middle component (M), top component b (Tb), and top component a (Ta). B, M, and Tb are bacilliform and contain the genomic RNAs 1, 2, and 3, respectively. Ta contains two molecules of the sub-genomic RNA4 and can be subdivided into bacilliform Ta-b and spheroidal Ta-t particles. The bacilliform particles are all 19 nm wide and have lengths of 56 nm (B), 43 nm (M), 35 nm (Tb), and 30 nm (Ta-b) (Fig. 1(A)). The RNAs are encapsidated by a single type of CP, which in the case of strain AMV-L has a length of 220 aa (Mr 24.3 kDa). In solution, AMV CP occurs as dimers, which under appropriate conditions of pH and ionic strength form a T¼ 1 icosahedral structures built from 30 dimers. This structure can be crystallized and has been studied by X-ray diffraction, cryoelectron microscopy, and image reconstruction methods (Fig. 1(B)). The CP was found to have the canonical eight-stranded bbarrel fold with the N- and C-terminal arms as extended chains. Dimer formation in the T¼1 particle is based on the clamping of the C-terminal arms of the subunits. From particle weight measurements and analysis of electron microscopic images, it was determined that the number of CP monomers in the major viral components is 60 þ (n  18), n being 10 (B, 240 subunits), 7 (M, 186 subunits), 5 (Tb, 150 subunits), or 4 (Ta, 132 subunits). By gel electrophoresis, at least 13 minor components have been resolved which probably represent other n values and contain monomers of genomic RNAs, multimers of genomic RNAs or RNA4, or specific degradation products of RNA3. Although details of the arrangement of the protein monomers have not been established, electron microscopical studies indicate that the cylindrical parts of the bacilliform particles have a hexagonal surface lattice with dimers of CP associated with the twofold symmetry axes. The cylindrical portion of the particle is thought to be capped by two halves of an icosahedron by changing the axes from sixfold symmetry in the cylinder into axes of fivefold symmetry. Neutron scattering data suggested that the capsid structure of spheroidal Ta-t particles is represented by a deltahedron with 52-point group symmetry built from 120 subunits. The percentage of RNA in the particles decreases from 16.3 in B to 15.2 in Ta-b. The buoyant density in CsCl of the major components fixed with formaldehyde varies from 1.366 (Ta) to 1.372 (B) g cm3. The protein shell has an inner radius of 6.5 nm and an outside radius of 9.4 nm. The RNA is uniformly packed within the 6.5 nm radial limit, occupies about 20% of the interior volume available, and slightly penetrates the protein shell to stabilize the particles by protein–RNA interactions. The RNA is easily accessible to ribonucleases A and T1 through holes in the protein shell.

Genome Structure The genome structure of AMV strain 425 is shown in Fig. 2. RNAs 1 and 2 encode the replicase proteins P1 and P2, respectively. P1 contains an N-terminal methyl-transferase-like domain and a C-terminal helicase-like domain, whereas P2 contains a polymerase-like domain. RNA3 is dicistronic and encodes the 50 proximal movement protein (MP) gene and the 30 proximal CP gene. MP and CP are both required for cell-to-cell transport of the virus. CP is translated from RNA4, which is identical in sequence to the 30 terminal 881 nt of RNA3. The length of the intercistronic region in RNA3 is 52 nt including the leader sequence of RNA4 of 36 nt. At the 51 end, all four AMV RNAs are capped. The organization of the leader sequence of RNA3 varies between strains. For strains M, S, L, and Y the length of this leader sequence is 240, 313, 345, and 391 nt, respectively. The increased length of the last three strains is due to the presence of direct repeats of 56 (S), 75 (L), or 149 (Y) nt. At their 30 termini, the viral RNAs contain a homologous sequence of 145 nt. The 30 112 nt of this sequence may adopt two mutually exclusive conformations, one representing a strong CP binding site (CPB) and the other representing a transfer RNAlike structure (TLS) resembling the TLS of bromo- and cucumoviruses. The 112 nt CPB structure, consists of four hairpins

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Fig. 2 Structure of the AMV genome. RNAs 1 and 2 are monocistronic and encode the replicase proteins P1 of B110 kDa, and P2 of B85 kDa. RNA3 is dicistronic and encodes a movement protein of B34 kDa and coat protein of B22 kDa. CP is translated from sub-genomic RNA4 (not shown). The gray squares represent the 30 terminal 112 nt of the RNAs, which can adopt either a tRNA-like structure or a structure with high affinity for CP. From: Bujarski et al., 2019. Journal of General Virology 100, 1206–1207.

(designated A, B, C, and D from 30 to 50 end) flanked by tetranucleotide sequences AUGC (or a UUGC derivative). Basepairing between a four nt sequence in the loop of the 50 proximal hairpin D and a four nt sequence in the stem of the 30 proximal hairpin A results in a pseudoknot interaction that generates the TLS structure. Structures similar to the AMV CPB and TLS conformations have been identified at the 30 termini of the RNAs of the ilarviruses. The TLS of AMV and ilarviruses cannot be charged with an aa as can the TLS of bromo- and cucumoviruses, but it can be recognized by the host enzyme that adds CCA to the 30 termini of cellular tRNAs (CTP/ATP: tRNA nucleotidyl transferase).

Interaction Between Viral RNA and Coat Protein In vitro, the 30 untranslated region (UTR) of AMV RNAs can bind several dimers of CP. A minimal CP binding site consists of the 30 -terminal 39 nt of the RNAs with the structure 51-AUGC-[hairpin B]-AUGC-[hairpin A]-AUGC-30 . This structure contains two overlapping binding sites for the N-terminus of the two subunits of a CP dimer. The consensus binding site in the RNA is UGC-[hairpin]-RAUGC (in which R is a purine). In addition to dimers of native CP, N-terminal peptides of CP can bind the 39 nt RNA fragment in a 2:1 ratio. The N-terminus of CP contains basic residues at positions 5, 6, 10, 13, 16, 17, 25, and 26, but only arginine-17 appears to be critical for binding of CP to RNA. This Arg residue is part of a Pro-Thr-x-Arg-Ser-x-x-Tyr (PTxRSxxY) RNA binding domain conserved among AMV and ilarvirus CPs. A complex of the 30 terminal 39 nt RNA fragment and peptides corresponding to the N-terminal 26 aa of CP has been crystallized and its structure was solved to 3 A1 resolution. Co-folding altered the structure of both peptide and RNA (Fig. 3). In the co-crystal, hairpins A and B are oriented at approximately right angles and each hairpin is extended by 2 bp formed between nt from adjacent AUGC sequences. If the AUGC motifs in the 39 nt fragment are numbered 1–3, starting from the 30 end of the RNA, hairpin B is extended by a duplex formed by base-pairing between the U and C residue of motif 3 and the A and G residue of motif 2. Similarly, hairpin A is extended by a duplex formed by base-pairing between the U and C residue of motif 2 and the A and G residue of motif 1. The two peptides in the co-crystal each form a helix with residues 12–26 ordered in peptide 1 and residues 9–26 ordered in peptide 2. The data has provided insight into the role of the PTxRSxxY-motif in RNA binding. In vitro selection of RNA fragments with high affinity for full-length AMV CP from a pool of randomized RNAs yielded fragments that maintained the unusual inter-AUGC base pairs observed in the crystal structure, but the primary sequences diverged from the wild-type RNA.

Translation of Viral RNA Extension of the 30 end of AMV genomic RNAs with an artificial poly(A)-tail increased the basal level of infectivity of these RNAs 50-fold, compared to a 1000-fold increase caused by binding of CP to the RNAs. A role of CP in translation of viral RNAs was investigated by extension of the 30 end of a luciferase reporter RNA with the AMV 30 UTR [Luc-AMV]. CP stimulated translational efficiency of Luc-AMV 40-fold without affecting the half-life of Luc-AMV. GST-pull-down assays and Far Western blotting revealed that AMV CP specifically interacted with the eIF4G-subunit in the eIF4F initiation factor complex from wheat germ and with the eIFiso4G-subunit from the eIFiso4F complex in plants This interaction allows the formation of a closed-loop structure for AMV RNAs similar to that formed by the interaction of poly(A) binding protein (PABP), the poly(A) tail of eukaryotic mRNAs, and the eIF4G-subunit during translation (Fig. 4). Therefore, AMV CP stimulated translation of viral RNAs by mimicking the PABP. Protoplasts transfected with wild-type AMV RNA4 accumulated CP at a detectable level, although accumulation was 100-fold lower than in productively infected protoplasts. However, translation of an RNA4 mutant, R17A, which encodes a CP with

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Fig. 3 Proposed structure of the entire 30 end of AMV RNA. A. Diagram of the 30 untranslated region (UTR) with color-coded U-C-hairpin-A-G CP-binding sites. B. End-on view of the simulated 30 UTR. C. View of the 30 UTR along its length. RNA is colored green; peptide is colored yellow. From Guogas, L.M., Filman, D.J., Hogle, J.M., Gehrke, L., 2004. Cofolding organizes alfalfa mosaic virus RNA and coat protein for replication. Science 306, 2108–2111.

Fig. 4 Model for the role of coat protein in translation of AMV RNAs. Translation of cellular mRNAs is strongly enhanced by the formation of a closed-loop structure by interactions of the poly(A) binding protein (PABP) with the 30 poly(A) tail and with the eIF4G subunit of the eIF4F complex of initiation factors bound to the 50 cap structure (upper panel). AMV coat protein (CP) enhances translational efficiency of viral RNAs 40-fold by binding to the CP binding site (CPB) at the 30 end of the viral RNAs. The finding that CP also interacts with eIF4G indicates that CP mimics the function of PABP in formation of the closed-loop structure (lower panel).

arginine-17 replaced by alanine that is unable to bind to the 30 end of AMV RNAs, was below the detection level. Translation of mutant R17A could be rescued to wild-type levels by expression in trans of a CP that was functional in RNA binding. Also, translation of this mutant was rescued to wild-type levels by replacing its 30 UTR by the 30 UTR of brome mosaic virus (BMV) RNA4. BMV is the type species of the genus Bromovirus in the family Bromoviridae and its 30 UTR stimulates translation independently of CP, possibly by the binding of host factors.

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Replication of Viral RNA AMV replication complexes are associated with vacuolar membranes. Replicase protein P1 recruits P2 to multivesicular bodies, also called prevacuolar compartments. P1 and P2 localize to the tonoplast in the presence of full-length AMV RNA suggesting that functional replicase complexes assemble on multivesicular bodies (MVBs) then traffic to the tonoplast. Template-dependent replicase preparations were purified from infected normal and transgenic plants that constitutively express the replicase proteins. The purified replicase preparations, which specifically accept exogenous AMV RNA as template, were used to study viral minusstrand and plus-strand RNA synthesis in vitro. The identification in 1978 of high-affinity binding sites for CP at the 30 termini of AMV RNAs led to the hypothesis that binding of CP was required to permit initiation of minus-strand RNA synthesis. However, all experiments that were done with replicase preparations to test this hypothesis showed that CP was not required for the synthesis of AMV minus-strand RNA in vivo or in vitro. This led to the conformational switch model of AMV RNA replication. An analysis of sequences of AMV RNAs, which direct minus-strand RNA synthesis by the purified viral replicase in vitro, showed that the entire 30 terminal homologous sequence of 145 nt is required for minus-strand promoter activity. This promoter consists of the TLS structure formed by the 30 -terminal 112 nt and a hairpin structure, termed hairpin E, between nt 112 and 145 from the 30 end of the RNAs. Hairpin E may be the primary element recognized by the viral replicase, and the TLS could serve to direct the replicase to the very 30 end of the template. If the TLS structure is disrupted by mutations affecting the pseudoknot or if the TLS is completely deleted, hairpin E directs initiation of minus-strand synthesis to a position located 50 from the hairpin. In the absence of the TLS, the mechanism of RNA synthesis directed by hairpin E is very similar to that directed by the sub-genomic promoter hairpin located in minus-strand RNA3, which directs plus-strand RNA4 synthesis. The finding that the sub-genomic promoter hairpin could be replaced by hairpin E without loss of infectivity illustrates the functional equivalence between the two hairpins. Although a knockout mutation of the CP gene does not affect AMV minus-strand RNA synthesis in infected protoplasts or in leaves expressing AMV replicase proteins, it results in a 100-fold drop in the accumulation of viral plus-strand RNAs. The expression of CP in leaves along with the replicase proteins not only enhanced the accumulation of replication-competent RNAs, but also the accumulation of replication-defective viral RNAs. Thus, it is possible that the stimulation of the accumulation of plus-strand RNAs by CP in infected cells reflects a role of CP in protection of the RNAs from degradation. RNA3 can be replicated in trans by P1 and P2 proteins expressed from replication-defective cDNAs RNAs 1 and 2 in tobacco leaves or by P1 and P2 expressed in transgenic P12 tobacco. However, RNAs 1 and 2 are unable to make use of these expressed replicase proteins and are dependent for replication on their encoded proteins in cis. This requirement in cis explains why RNA3 can replicate in protoplasts of transgenic P12 plants without a requirement for CP, whereas replication of RNA1 or RNA2 in this system requires CP to enable translation of these RNAs. Moreover, replication of RNAs 1 and 2 is strictly coordinated. Mutations in either P1 or P2 ORF that is lethal to the function of the encoded proteins block the replication of both mutant and wild-type RNA1 or 2, but does not affect the replication of RNA3 by the expressed replicase. Apparently, replication of RNA1 controls replication of RNA2 and vice versa.

Virus Encapsidation and Movement The high-affinity binding sites for CP at the end of RNAs 1 and 2 are dispensable for encapsidation of the RNAs. This was found by the expression of mutant RNAs 1 and 2 without their 30 UTRs along with wild-type RNA3 in leaves expressing replicase proteins. RNAs 1, 2, and 3 were all encapsidated into particles. Possibly, assembly of particles initiates on the identified internal CP binding sites in AMV RNAs. The RNA3 encoded MP and CP are both required for cell-to-cell movement of AMV in infected plants. The MP is phosphorylated and forms tubular structures on the surface of infected protoplasts, suggesting that virus movement in plants involves transport of virus particles through tubules, which traverse the cell wall through modified plasmodesmata. Such a mechanism has been found for viruses of the families Bromoviridae, Comoviridae, Caulimoviridae, and Bunyaviridae. However, tubular structures filled with virus particles have not been observed in AMV infected leaf tissue. Moreover, a CP mutant defective in the formation of virions is able to move cell-to-cell at a reduced level. This suggests, along with information about other members of the Bromoviridae, that the transport of AMV moves cell-to-cell as viral ribonucleoprotein complexes that structurally differ from virions. AMV MP is functional in the cell-to-cell transport of Tobacco mosaic virus, Brome mosaic virus, Prunus necrotic ringspot virus, Cucumber mosaic virus, and Cowpea mosaic virus. Conversely, the MPs of these viruses are competent in systemic transport of AMV as long as the MPs contain the last 44 CP-interacting aa of AMV MP at their C-termini. Analysis of AMV MP and those of other viruses in the family Bromoviridae suggests that the C-termini of these MPs enable specificity in virus transport by binding specific virus CPs and that the N-terminal portions of the MPs function in transport. AMV MP interacts with Arabidopsis Patellin 3 (atPATL3) and Patellin 6 (atPATL6), which are implicated in membrane trafficking. The interaction interferes with the plasmodesmatal targeting of the MP and reduces AMV cell-to-cell movement. AMV replicated to higher levels in Patellin knockout mutants of Arabidopsis than in normal plants where replication is limited to fewer cells due to inhibition of cell-to-cell movement.

Role of Coat Protein in the AMV Replication Cycle Natural infections by AMV are initiated by viral particles containing the genomic RNAs 1, 2, and 3, and sub-genomic RNA4. However, inoculation of plants in the laboratory with a mixture of the three genomic RNAs results in barely detectable levels of

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AMV, but the infectivity of this mixture is increased 1000-fold by the addition of four to ten molecules of CP per RNA molecule. Alternatively, infectivity can be increased by including RNA4, the messenger for CP. It is assumed that AMV infection starts with the RNAs associated with one or more dimers of the viral CP. These RNA/CP complexes can be generated by partial disassembly of the particles in inoculated cells, by mixing purified viral RNAs and CP in vitro, or by translation of RNA4 in the inoculated cells and subsequent binding of de novo synthesized CP to the viral RNAs. The requirement of CP for translation of AMV RNAs was explained by the finding that AMV CP mimicks the function of the host cell poly(A) binding protein for efficient translation. Two alternative models have been proposed for the role of CP in the replication of AMV RNA: a conformational switch model and a 30 organization model. The conformational switch model suggests that AMV RNA 30 ends form two mutually exclusive conformations, one that is extended and binds AMV CP and a second CP-free TLS pseudoknot conformation. This model proposes that CP enhances the translation of RNA 1 and 2 to make the replicase proteins and that the switch from translation to replication requires dissociation of CP from the 30 end of the RNAs to allow the formation of the TLS structure for minus-strand promoter activity. This model is based upon many experiments done in vivo and in vitro with mutant RNAs or replicase preparations, which found that CP was not needed for minus-strand synthesis. In contrast, the 30 organization model suggests that the CP organizes the 30 termini into compact structures for activation of early RNA replication. This was proposed based on the imperfect fit and uncertain functionality of the reported TLS at the 31 end of AMV RNAs and the crystal structure of the 30 terminal 39 nt of AMV RNA in complex with the RNA binding domain of the CP. This model proposes that a structurally organized RNA end is important for replication for all members of the family Bromoviridae. The compact structure is formed at the 30 end of some virus RNAs, such as those of BMV, by a TLS structure, while others, such as the 30 end of AMV and the ilarviruses, require CP binding to fold into the active structure. Replicase proteins translated from inoculum RNAs 1 and 2 target the viral RNAs to vacuolar membranes where RNA synthesis takes place. The mechanism of the switch from minus-strand to plus-strand RNA synthesis is not yet clear. Although the promoter for plus-strand sub-genomic RNA4 synthesis is structurally similar to hairpin E in the minus-strand promoter, the promoters for plus-strand genomic RNA synthesis have not yet been characterized. The requirement of CP for efficient accumulation of plusstrand RNA in infected cells may reflect a role of CP in protection of the RNAs from degradation, but a role in plus-strand RNA synthesis has not yet been ruled out. There is growing evidence for AMV and other viruses in the family Bromoviridae that the many steps in the viral replication cycle are tightly linked.

Virus-Host Relationships The AMV group is a large conglomerate of strains infecting a large number of susceptible hosts. This accounts for the tremendous range of symptoms displayed by AMV-infected plants. Mutations in the coat protein gene and 50 UTR of RNA3 have been shown to affect symptom formation in tobacco. Cytological modifications in AMV-infected plants occur only in cells of organs showing symptoms. In these cells, fragmentation of the ground cytoplasm and an increased accumulation of membrane-bound vesicles has been observed. Sometimes the lamellar system of chloroplasts is affected and invaginations of the nuclear membrane have been reported. The MP protein localizes to plasmodesmata. It interacts with atPATL3 and atPATL6, which are normally located at the cell periphery, to form complexes that were at the plasmodesmata and also at the cell periphery. Overexpression of atPATL3 and atPATL6 reduces the plasmadesmatal localization of AMV MP thereby reducing the spread of infection. When expressed alone in protoplasts, the P1 protein is found on MVBs, while the P2 protein is throughout the cytosol. When P1 and P2 are expressed together, both are found on MVBs; when both proteins are expressed with a functional AMV RNA, the proteins localize to the vacuolar membrane, the tonoplast, where replication takes place. Following encapsidation, virus particles are found throughout the cytoplasm with a few records of particles in the nucleus. Depending on the strain, the particles form different types of intracellular aggregates. Recent studies have shown that the CP of AMV is multifunctional and interacts with a number of host proteins. Chloroplast proteins are increasingly found to be associated with virus proteins; Arabidopsis PsbP (atPsbP) protein is one example. This nuclear-encoded component of the oxygen-evolving complex of photosystem II was identified in a yeast two-hybrid screen to interact with AMV CP. Newly synthesized AtPsbP was sequestered in the cytosol during virus infection, whereas it localizes to the chloroplasts in healthy cells. Overexpression of atPsbP inhibited virus replication suggesting that atPsbP has a role in plant defenses. AMV CP also interacts with the transcription factor, ILR3, in both tobacco and Arabidopsis. This interaction takes place in the nucleus where CP-interaction moves ILR3 to the nucleolus in which CP had previously been found. CP has a nucleolar localization signal in its N-terminus and interacts with importin-a to gain access to the nucleus. It is suggested that ILR3-CP interaction leads to a down-regulation of a host factor that is involved in host defense modulation. AMV CP also interacts with atALKBH9B, recently identified as the first demethylase described in plants. Such enzymes have broad regulatory roles in RNA biology and can remove m6A in virus RNAs to modulate infectivity of the virus. atALKBH9B also interacts with AMV RNA, whose sequence contains three of the common m6A consensus motifs. Downregulation of atALKBH9B in infected plants increased the abundance of m6A in AMV RNA and decreased infection.

Host Range and Economic Significance AMV occurs worldwide. Strains of this virus have been found in natural infections of about 305 plant species representing 22 families. The experimental and natural host ranges include over 600 species in 70 families. Recently, 68 Arabidopsis thaliana ecotypes were analyzed for their susceptibility to AMV infection. In total 39, ecotypes supported both local and systemic infection, 26 ecotypes supported only local infection, and three ecotypes could not be infected.

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Fig. 5 Symptoms caused by AMV infection. (A) Interveinal chlorosis on alfalfa plants; (B) Mottle on potato; (C) Vein clearing on Burley tobacco; (D) Mosaic on pepper; (E) Mosaic on soybean; (F) Chlorosis on viburnum; (G) Interveinal chlorosis on white clover; (H) Tip necrosis on lentil. From: (A) South Dakota State University, extension.sdstate.edu; (B) Howard F. Schwartz, Colorado State University, Bugwood.org; (C) Kenneth Seebold, Univeristy of Kentucky, burleytobaccoextension.ca.uky.edu; (D) Whitney Cranshaw, Colorado State University, Bugwood.org; (E) Craig Grau, University of Wisconsin, cropprotectionnetwork.org; (F) Tom Creswell, Purdue University; (G) Barenbrug agriseeds, agriseeds.co.nz; (H) agriculture.vic.gov.au.

AMV causes significant losses in forage crops, can reduced winter survival, and increase the susceptibility to other pathogens. AMV is an economically important pathogen in temperate pulses (chickpeas, faba beans, field peas, and lentils), pasture legumes (alfalfa, and clovers), and may infect pepper, tobacco, tomato, soybean, potato, celery, weeds, and ornamental plants. In alfalfa, losses have been reported up to 30% and seed production up to 69%. In soybean, AMV infection along with soybean mosaic virus causes disease synergism with the development of severe symptoms. Such co-infection is of concern in areas where the soybean

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aphid is present. Time of infection is important in yield loss caused by AMV. For example, late infection of faba bean can cause up to 45% of seed loss. Symptoms caused by AMV vary depending upon the virus strain and host, but include: yellow mosaic and mottle, necrosis, stunting, leaf distortion, leaf rolling, vein-clearing, chlorotic spots, line patterns, and ringspots. The environment plays a role in symptom production. Infected field plants are more symptomatic in cool spring and autumn weather and may be symptomless in warm/hot summers. Fig. 5 shows the symptoms for a variety of infected plants.

Transmission AMV is easily mechanically transmissible. However, field spread occurs predominantly by aphid transmission. At least 15 aphid species are known to transmit the virus in a stylet-borne or non-persistent manner. Acquisition of the virus occurs within 10–30 s of feeding and is followed by immediate transmission without a latent period. The ability of the aphid to continue to transmit is lost within 1 h. The variability of individual aphid species in their capacity to transmit different AMV strains suggests a specific virus–vector relationship which is probably governed by the structural properties of the CP. Seed transmission of AMV has been reported for alfalfa and other plant species with rates of transmission varying from 0.1% to 50%. Transmission of the virus between plants by parasitic dodder has been observed with five Cuscuta species.

Epidemiology and Control Genes conferring resistance to AMV have been reported in alfalfa (amv-1 gene), soybean (rav1 gene), and tomato (am gene). These, however, have not been deployed in commercial crops. Control of AMV in alfalfa is primarily achieved by using virus-free seed and avoiding reservoir hosts of the virus. Because AMV infection occurs naturally in many different plant species, control is difficult. Transgenic tobacco, alfalfa, and white clover plants expressing the CP gene of AMV were found to be highly resistant to the virus when mechanically inoculated. The resistance was clearly protein-mediated because plants with the highest levels of CP accumulation were the most resistant. A mutation in the transgene that affected the N-terminal sequence of the encoded CP destroyed resistance to the wild-type virus but the mutant transgene conferred resistance to virus expressing the mutant CP. As yet, transgenic resistance to AMV has not been commercialized. AMV is primarily spread by aphids. The incidence of AMV in alfalfa pastures can be as high as 50%–98%. Susceptible crops growing outdoors or in greenhouses close to alfalfa fields are at risk of infection. Because of AMV’s wide host range, many weed species and ornamental plants are infected, which provides reservoirs for the aphids to acquire the virus. AMV is also seed-born. It has been spread throughout the world by importation of infected alfalfa seed lots. AMV is also mechanically transmitted. It can be transmitted to healthy plants by handling infected plants or seedlings. Control of AMV in alfalfa is primarily achieved by using virus-free seed and transplants, managing weeds, and avoiding reservoir hosts of the virus. Insecticides are largely ineffective in field management of AMV because aphids do not have to colonize a plant to acquire or transmit the virus; test feedings are sufficient. However, control of aphids early in the season may avoid buildup of populations. Various cultivars of crop plants may vary in susceptibility; however, there are no AMV-resistant commercial varieties.

See also: Brome Mosaic Virus (Bromoviridae). Bromoviruses (Bromoviridae). Cucumber Mosaic Virus (Bromoviridae)

Further Reading Balasubramaniam, M., Ibrahim, A., Kim, B.-S., Loesch-Fries, L.S., 2006. Arabidopsis thaliana is an asymptomatic host of Alfalfa mosaic virus. Virus Research 121, 215–219. Balasubramaniam, M., Kim, B.-S., Hutchens-Williams, H.M., Loesch-Fries, L.S., 2014. The photosystem II oxygen-evolving complex protein PsbP interacts with the coat protein of Alfalfa mosaic virus and inhibits infection. Molecular Plant Microbe Interactions 27, 1107–1118. Bol, J.F., 2005. Replication of alfamo- and ilarviruses: Role of the coat protein. Annual Review of Phytopathology 43, 39–62. Guogas, L.M., Filman, D.J., Hogle, J.M., Gehrke, L., 2004. Cofolding organizes Alfalfa mosaic virus RNA and coat protein for replication. Science 306, 2108–2111. Hull, R., 1969. Alfalfa mosaic virus. Advances in Virus Research 15, 365–433. Jones, R.A.C., 2012. Virus diseases of annual pasture legumes: Incidences, losses, epidemiology, and management. Crop & Pasture Science 63, 399–418. Krab, I.M., Caldwell, C., Gallie, D.R., Bol, J.F., 2005. Coat protein enhances translational efficiency of Alfalfa mosaic virus RNAs and interacts with the eIF4G component of initiation factor eIF4F. Journal of General Virology 86, 1841–1849. Kumar, A., Reddy, V.S., Yusibov, V., et al., 1997. The structure of Alfalfa mosaic virus capsid protein assembled as a T¼ 1 icosahedral particle at 4.0-Å resolution. Journal of Virology 71, 7911–7916. Martinez-Pérez, M., Aparicio, F., López-Gresa, M.P., et al., 2017. Arabidopsis m6A demethylase activity modulates viral infection of a plant virus and the m6A abundance in its genomic RNAs. Proceedings of the National Academy of Sciences of the United States of America 114, 10755–10760. Navarro, J.A., Sanchez-Navarro, J.A., Pallas, V., 2019. Key checkpoints in the movement of plant viruses through the host. Advances in Virus Research 104, 2–63. Olsthoorn, R.C.L., Mertens, S., Brederode, F.T., Bol, J.F., 1999. A conformational switch at the 30 end of a plant virus RNA regulates viral replication. EMBO Journal 18, 4856–4864. Pallas, V., Apariio, F., Herranz, M.C., Sanchez-Navarro, J.A., Scott, S.W., 2013. The molecular biology of Ilarviruses. Advances in Virus Research 85, 140–181. Petrillo, J.E., Rocheleau, G., Kelley-Clarke, B., Gehrke, L., 2005. Evaluation of the conformational switch model for Alfalfa mosaic virus RNA replication. Journal of Virology 79, 5743–5751.

Alphaflexiviruses (Alphaflexiviridae) Sergey Y Morozov and Alexey A Agranovsky, Lomonosov Moscow State University, Moscow, Russia r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) Avr Avirulence CP Coat protein DCL Dicer-like ER Endoplasmic reticulum HEL RNA helicase domain kb Kilobases kDa Kilo Daltons MP Movement protein MTR Methyl-transferase domain nm Nanometer(s) NMD Nonsense-mediated degradation pathways of mRNA decay

Glossary 30 -co-terminal sub-genomic RNA Virus-specific RNA corresponding to the 30 -terminal portion of virus RNA genome. Cap m7GTP moiety at the 50 end of the genomic virus RNA. Cell-to-cell movement Substance movement between adjacent cells. Coat protein Virion protein directly covering the virus genomic nucleic acid in the particle. Endoplasmic reticulum Cell network of endomembrane tubules.

nt Nucleotide(s) ORF Open reading frame PAMP Pathogen-associated molecular pattern PCNA Proliferating cell nuclear antigen PD Plasmodesmata POL RNA-dependent RNA polymerase domain ROS Reactive oxygen species satRNA Satellite RNA SFI helicase Superfamily I helicase sgRNA Sub-genomic RNA TGB Triple gene block VSR Viral suppressor of RNA silencing

Long-distance movement Substance virus movement through the plant phloem system. Movement protein Virus protein responsible for cell-tocell movement. Open reading frame Sequence of codons (usually not less than 15–20 codons) containing no termination triplets. Triple gene block The module of three overlapping genes involved into cell-to-cell and long distance movement of the virus.

Introduction The family Alphaflexiviridae (order Tymovirales) contains seven genera (Allexivirus, Botrexvirus, Lolavirus, Mandarivirus, Platypuvirus, Potexvirus, and Sclerodarnavirus) and five species unassigned to a genus (Tables 1 and 2). These viruses have flexuous filamentous particles (Fig. 1) and infect higher plants or fungi. The 50 -capped and 30 - polyadenylated monopartite positive-stranded RNA genomes of the Alphaflexiviridae members are 5.9–9.0 kb in length. The 50 -terminal open reading frame (ORF) encodes the replicase with conserved methyltransferase (MTR), helicase (HEL), and RNA-dependent RNA polymerase (POL) domains, a configuration typical of the alphavirus-like superfamily of plant and animal viruses (Fig. 2). Most plant-infecting alphaflexivirus genomes encompass the module of three overlapping ORFs (Triple Gene Block, TGB) that are necessary for the cell-to-cell movement. Coat protein (CP) is encoded by the gene located at or close to the 30 -end of the genome (Fig. 2). Alphaflexiviruses produce 30 -coterminal sub-genomic RNAs that allow expression of the CP and other accessory proteins. Some alphaflexiviruses amplify to high yields in host plants, and are transmitted mechanically, whereas others have mites and insect vectors. Plantinfecting family members have been found in a wide range of mono- and dicotyledonous plants species, and the majority of them have only mild effects on their host.

Taxonomy, Phylogeny of Family Members, and Evolution The genera of Alphaflexiviridae show close similarity of the encoded proteins (mostly the replicases) and share common features of the virion structure and genome organization. These properties are shown in Fig. 2 and Table 1. The affiliation with Alphaflexiviridae is based on the replicase amino acid sequence, and viruses from different genera usually have less than about 45% nt identity in the respective replicase genes.

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Some properties of the viruses in the genera of the family Alphaflexiviridae

Genus

Host

Allexivirus Botrexvirus Lolavirus Mandarivirus Platipuvirus Potexvirus Sclerodarnavirus

Higher Fungi Higher Higher Higher Higher Fungi

plants plants plants plants plants

Virion length

Number of ORFs

Replicase size (kDa)

Coat protein size (kDa)

800 nm 720 nm 640 nm 650 nm 800 nm 470–580 nm No virions

6 5 6 6 7 5 1

170–195 160 196 187 157 150–195 193

25–29 43 32 34 22 18–27 No CP

Fig. 1 Electron micrograph of negatively-stained virions of Alternanthera mosaic potexvirus. The bar ¼ 500 nm. Courtesy of O.V. Karpova, Lomonosov Moscow State University. Department of Virology.

Among the plant-infecting Alphaflexiviridae five genera are currently recognized (Potexvirus, Allexivirus, Lolavirus, Mandarivirus, and Platypuvirus; Table 2). The most populated genus Potexvirus includes 38 members (Table 2). The next most populated genus Allexivirus contains 12 approved members. The genus Lolavirus contains two members, whereas each of the genera Mandarivirus and Platypuvirus includes only a single species. Two remaining fungus-infecting genera also include a single species (Table 2). The single member of the genus Botrexvirus (Botrytis virus X) infects a filamentous fungus. The BVX genome lacks the movement protein genes, but encodes the CP and several unique proteins of unknown function. Another fungus-infecting genus Sclerodarnavirus is also represented by a single species whose genome contains only the replicase ORF (Fig. 2 and Table 2). Phylogenetic trees based on the nucleotide and protein sequences depended on the genome region used for analysis. In a phylogenetic analysis of the whole replication protein, Botrexvirus and Allexivirus genera fall on a well-supported separated branch (Fig. 3). Potexvirus, Mandarivirus and Lolavirus genera form another branch in replicase phylogenetic tree, whereas Platipuvirus (DOSV) forms the most basal branch in this dendrogram (Fig. 3). If the genome nucleotide sequence is considered, the genomic RNAs of Alphaflexiviridae are most closely related to those of the other families in the order Tymovirales, namely Betaflexiviridae, Gammaflexiviridae and Tymoviridae. Strikingly, in this genome-based tree two genera of betaflexiviruses, namely, Vitivirus and Tepovirus, show common root with the Alphaflexiviridae branch but not with the other members of Betaflexiviridae (Fig. 4). This is in line with replicase protein tree where Vitivirus and Tepovirus proteins form clear distinct brunch separated from the rest of betaflexiviruses (Fig. 3). The TGB proteins are related to the equivalent proteins encoded by some members of the family Betaflexiviridae and, more distantly, to those of the rod-shaped viruses in the family Virgaviridae (genera Hordeivirus, Pecluvirus, and Pomovirus).

Virion Structure Typical alphaflexiviruses have flexuous filamentous particles (12–13 nm in diameter and up to 800 nm in length) with helical symmetry (pitch of ca. 3.4 nm) (Fig. 1). No lipids have been reported in the virions of alphaflexiviruses. Usually no carbohydrates are associated with the virions. The potexvirus and lolavirus CPs are reportedly glycosylated. The viral capsid of most members of the family (except lolaviruses) is composed of identical CP subunits ranging in size from 18 to 43 kDa. In allexiviruses, the 42 kDa protein (encoded in the third ORF from the 30 end; Fig. 2) is a minor virion component non-uniformly distributed along the particle. The particles of lolaviruses are composed of equimolar amounts the larger and smaller versions of the CP (32 kDa and

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Fig. 2 Schematic representation of the genome structure of the Alphaflexiviridae (Potexvirus, Potato virus X; Alexvirus, Shallot virus X; Mandarivirus, Citrus yellow vein clearing virus; Botrexvirus, Botrytis virus X; Sclerodarnavirus, Sclerotinia sclerotiorum debilitation-associated virus; Lolavirus, Lolium latent virus; Platipuvirus, Donkey orchid symptomless virus). Arrowheads denote the 30 -termini of genomic RNAs; broken arrows – the start and direction of the potexvirus sub-genomic RNAs. Open reading frames (ORFs) are shown by boxes. MTR, HEL, and POL stand for the conserved methyl-transferase, NTPase/helicase, and RNA polymerase domains, respectively. TGB (triple gene block), CP (coat protein gene), and the genes for RNA-binding proteins are shown as boxes with different fills.

28 kDa, respectively). The 28 kDa CP seemingly originates from internal translation initiation of the coat protein gene. It should be noted that the sclerodarnavirus exists only as a capsid-less single-stranded and double-stranded RNA, making an intriguing parallel with capsid-less RNA agents associated with chestnut blight hypovirulence (family Hypoviridae) that code for the replicase orthologs of plant picorna-like viruses of the family Potyviridae (Fig. 2).

Genome Organization Genomic RNAs of plant alphaflexiviruses typically have five or six ORFs. The 50 terminal ORF1 encodes the viral replicase (Fig. 2). The 30 proximal ORFs encode the movement proteins (TGB proteins), the CP, and an accessory zinc-binding and RNA-binding protein (Fig. 2). The last protein is encoded only in genera Allexivirus and Mandarivirus. ORF2 to ORF5 are expressed via the 30 co-terminal sgRNAs (Fig. 2).

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Table 2 List of genera, virus species, acronyms, NCBI accession # and Genome length (nt) in the family Alphaflexiviridae. Type species are written in bold Genus/Species name

Acronym

Accession #

Length (nt)

Potexvirus Actinidia virus X Allium virus X Alstroemeria virus X Alternanthera mosaic virus Asparagus virus 3 Babaco mosaic virus Bamboo mosaic virus Cactus virus X Cassava common mosaic virus Cassava virus X Clover yellow mosaic virus Cymbidium mosaic virus Foxtail mosaic virus Hosta virus X Hydrangea ringspot virus Lettuce virus X Lily virus X Malva mosaic virus Mint virus X Narcissus mosaic virus Nerine virus X Opuntia virus X Papaya mosaic virus Pepino mosaic virus Phaius virus X Pitaya virus X Plantago asiatica mosaic virus Potato aucuba mosaic virus

AVX AlVX AlsVX AltMV AV3 BabMV BaMV CVX CsCMV CsVX CYMV CymMV FoMV HVX HdRSV LeVX LVX MaMV MVX NMV NVX OpVX PapMV PepMV PhVX PiVX PlaMV PAMV

NC028649 NC012211 NC007408 NC007731 NC003400 NC036587 NC001642 NC002815 NC001658 NC034375 NC001753 NC001812 NC001483 NC011544 NC006943 NC010832 NC007192 NC008251 NC006948 NC001441 NC007679 NC006060 NC001748 NC004067 NC010295 NC024458 NC003849 NC003632

6888 7118 7009 6607 6985 6692 6366 6614 6376 5879 7015 6227 6151 6528 6185 7212 5823 6858 5914 6955 6582 6653 6656 6450 5816 6677 6128 7059

Potato virus X Schlumbergera virus X Senna mosaic virus Strawberry mild yellow edge virus Tamus red mosaic virus Tulip virus X Turtle grass virus X Vanilla virus X White clover mosaic virus Yam virus X Zygocactus virus X

PVX SchVX SenMV SMYEV TRMV TVX TGVX VVX WClMV YVX ZyVX

NC011620 NC011659 NC030746 NC003794 NC016003 NC004322 NC040644 NC035205 NC003820 NC025252 NC006059

6435 6633 6775 5966 6495 6056 6278 6295 5845 6158 6624

Allexivirus Alfalfa virus S Arachis pintoi virus Blackberry virus E Garlic mite-borne filamentous virus Garlic virus A Garlic virus B Garlic virus C Garlic virus D Garlic virus E Garlic virus X

AVS ApV BVE GarMbFV GarV-A GarV-B GarV-C GarV-D GarV-E GarV-X

NC034622 NC032104 NC015706 NC038864 NC003375 NC025789 NC003376 NC022961 NC004012 NC001800

8349 7599 7718 762 a 8660 8336 8405 8424 8451 8106

Shallot virus X Vanilla latent virus

ShVX VLV

NC003795 NC035204

8832 7462

Lolavirus Lolium latent virus

LoLV

NC010434

7674

Mandarivirus Citrus yellow vein clearing virus Indian citrus ringspot virus

CYVCV ICRSV

NC026592 NC003093

7531 7560 (Continued )

144

Table 2

Alphaflexiviruses (Alphaflexiviridae)

Continued

Genus/Species name

Acronym

Accession #

Length (nt)

Platypuvirus Donkey orchid symptomless virus

DOSV

NC022894

7838

Botrexvirus Botrytis virus X

BVX

NC005132

6966

Sclerodarnavirus Sclerotinia sclerotiorum debilitation-associated RNA virus

SSDaRV

NC007415

5470

Unassigned species Insect-associated alphaflexivirus 3 Euonymus yellow vein virus Potexvirus sp.

IaAlV-3 EuYVV –

MN203145 NC035190 NC040842

796a 7279 5839

IaAlV-1

MN203143 MN203144

4165 868a

Insect-associated alphaflexivirus 1 Insect-associated alphaflexivirus 2 IaAlV-2 a

partial sequence.

Fig. 3 Maximum-likelihood tree derived from the complete amino acid sequences of the replicases of representative members of the families Betaflexiviridae and Alphaflexiviridae. Names of viruses and GenBank accession numbers are shown in the tree. Bootstrap analysis was done using 1000 replicates in MEGA7. The newly identified virus in this study is indicated by a red star. The scale bar represents the genetic distance. Accession number for Loquat virus A genome is MK936045. After Liu, Q., Yang, L., Xuan, Z., et al., 2020. Complete nucleotide sequence of Loquat virus A, a member of the family Betaflexiviridae with a novel genome organization. Archives of Virology 165, 223–226.

Notable specific feature of Allexivirus members is the absence of TGB3 gene and the presence, in place of TGB3, of the gene for a large protein of unknown function (Fig. 2). Even more distinctive features in genome structure have been found in an unusual Platipuvirus, Donkey orchid symptomless virus (DOSV) (Fig. 2). The ORF1 that overlaps the replicase gene has marginal similarity with MPs of plant tymoviruses (family Tymoviridae). The replicase gene (ORF2) shares domain organization of the encoded protein with other members of the family Alphaflexiviridae. A 44 kDa protein encoded by ORF3 shows low level of identity with myosin. The putative 25 kDa movement protein (MP) (ORF7) shares low level of identity with the 3A-like MPs of some members of the family Tombusviridae (Dianthovirus).

Alphaflexiviruses (Alphaflexiviridae)

145

Fig. 4 Molecular Phylogenetic analysis of full-genome tymoviral reference sequences by Maximum Likelihood method based on the General Time Reversible model with a discrete Gamma distribution. The scale bar represents the genetic distance. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. After Çağlayan, K., Roumi, V., Gaze, L.M., et al., 2019. Identification and characterization of a novel Robigovirus species from sweet cherry in Turkey. Pathogens 8, 57.

In genus Potexvirus, three 30 co-terminal sgRNAs are produced in infected plant cells (Fig. 2). The 2.1 kb species serves as mRNA for the TGB1 protein, whereas the 1.2-rb sgRNA directs expression of the TGB2 and TGB3 via leaky scanning. The 1.0-kb sgRNA encodes the coat protein. The genome organization of the TGB-containing alphaflexiviruses is quite similar to the TGB-containing betaflexiviruses belonging to the subfamily Quinvirinae (genera Carlavirus, Robigovirus and Foveavirus). Fungal alphaflexiviruses (genera Botrexvirus and Sclerodarnavirus) are drastically different with respect to their genome architecture. The capsid-less sclerodarnavirus encodes the replicase gene only, whereas Botrytis virus X contains six genes encoding replicase, CP and four proteins with unknown function (Fig. 2).

Properties and Functions of Gene Products The ORF1 product contains the invariant array of protein domains involved in RNA capping (MTR), duplex unwinding (HEL), and RNA synthesis (POL) (Fig. 2). The POL domain of potexviruses is vital for the functioning of the viral replication complex. It contains the typical signature of plus-RNA viral RdRp – tripeptide GDD. Mutational analysis of potexvirus genomes unambiguously showed that during the catalysis of polymerization reaction this motif has the essential role. The POL domain of potexviruses also recognizes the cis-signals in the 30 -terminal region of both the positive and negative strands of RNA genome (see also below). These specific protein–RNA interactions allow correct initiation of the replication. The replicative HEL domain of potexviruses locates in the middle of replicase protein and belongs to SFI helicase family. The purified HEL is able to unwind dsRNAs and to remove the g-phosphate from nucleoside triphosphates. Both of these activities depend on Mg2 þ or Mn2 þ cations. Mutations of the SFI conserved motifs I, II, III, or VI resulted in inhibition of these activities. The RNA binding activity of HEL is cooperative and is based on conserved motifs I and II as well as on poorly conserved aa sequences. The N-terminal viral replicase MTR domain was proposed to have an AdoMet-dependent guanylyl-transferase activity which transfers the methyl group of AdoMet to GTP, leading to m7GTP formation. The m7GMP moiety of m7GTP is then transferred to the 50 end of the genomic RNA via the covalent [m7GMP-enzyme] intermediate. Analogous cap formation mechanisms have been found also in other members of the alphavirus-like superfamily of positive strand RNA viruses. Importantly, the first two domains

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Alphaflexiviruses (Alphaflexiviridae)

of potexvirus replicase (MTR and HEL) work in a concert to form the 50 cap-structure on the nascent viral genomic and sub-genomic RNAs. The potexvirus TGB1 product also contains an NTPase-helicase domain of SF1 with seven conserved motifs (I, Ia, II, III, IV, V, and VI). Motif I (“Walker A”) includes typical GKS/T tripeptide, and motif II (“Walker B”) is involved in binding of ATP and Mg2 þ . The TGB1 protein is necessary for the cell-to-cell movement and also functions as a suppressor of RNA silencing. It seems that these activities of the TGB1 are uncoupled. In recent years it was documented that potexvirus TGB1 also participates in formation of virus replication factories in infected cells. Thus, the PVX TGB1 is involved in reorganizing actin cytoskeleton and endoplasmic reticulum (ER) into the membrane-bound virus replication complexes (VRC). Moreover, TGB1 participates in positioning the VRC close to plasmodesmata thus helping to establish connection between replication and movement. The TGB2 and TGB3 proteins, containing blocks of hydrophobic residues, are associated with membranes derived from the ER. Potexvirus TGB2 protein has two predicted transmembrane domains that interact with ER membranes. It seems that TGB2 protein integrates into the ER membranes in a U-like structure, with the central part positioned in the ER lumen and the N- and C-termini located on the cytoplasmic side of the ER. Mutation analysis revealed that two conserved cysteine residues in the C-terminal region of potexvirus TGB2 protein are important for the cell-to-cell movement. Recently, it was found that PVX TGB2 may function as the molecular bridge allowing interaction between the VRC and TGB1, TGB3, and coat protein, thereby enhancing the efficiency of the virus spread in infected plant tissue. Potexvirus TGB3 protein also has a conserved C(x5)G(x6–9)C sequence positioned at the C-terminus of a predicted transmembrane domain. This C-terminal sequence is exposed to the cytoplasmic side of the ER and contains a sorting signal that is necessary for TGB3 oligomerization and for the targeting of cortical ER.

Replication and Propagation Cis-acting replication signals have been mostly studied using PVX and Bamboo mosaic virus (BaMV) as models. Replication of potexviruses is governed by the cis-signals at the 50 - and the 30 end of the genome (each representing a complex tertiary RNA structure). The synthesis of the negative strand and of the positive-strand genomic and sub-genomic RNAs requires long-distance interactions between the conserved sequences at the 30 ends of the positive and negative RNA strands, and the internal complementary octanucleotides. The consequence of the replication/transcription events is as follows: (1) using genomic RNA as template, the replicase recognizes the 30 -cis-signal base-paired with an internal octanucleotide and synthesizes the antigenomic (negative) RNA strand; (2) in the negative strand, the replicase interacts with the 30 -cis-signal base-paired with an internal octanucleotide and produces the progeny genomic RNA strands; (3) in the negative strand, contacts of the 30 -cis-signal with complementary octanucleotides localized, in 30 to 50 direction, prior to the genes for TGBp1, TGBp2, and CP, force the replicase to initiate internally and produce the respective sgRNAs.

Transmission, Host Range Most members of the family infecting higher plants are transmitted by mechanical contacts, seeds, or grafting. Citrus yellow vein clearing mandarivirus (CYVCV) is spread from infected to healthy citrus trees by the whitefly Dialeurodes citri (Ashmead). Alexiviruses are transmitted by eriophyd mites. PVX causes mild mosaic or a symptomless infection in solanaceous hosts, depending on the virus strain. In Gomphrena globosa L., a diagnostic host, the virus induces formation of ring necrotic lesions. Another potexvirus, PepMV causes tomato fruit discoloration (marbling, blotchy ripening and flaming), leaf distortion, chlorosis, yellow spots, interveinal leaf yellowing (Fig. 5) and, occasionally, leaf and stem necrosis. In addition, CYVCV infection in lemon and sour orange is characterized by strong yellow vein clearing, leaf distortion, ringspot lesions, and veinal necrosis. The natural host range of the family members vary from narrow (lolaviruses, mandariviruses, and the viruses infecting fungal hosts) to moderate (potexviruses).

Epidemiology and Control The genus Potexvirus includes species that are responsible for serious crop damage (PVX, BaMV, Pepino mosaic virus, and Citrus yellow vein clearing virus), whereas the economical impact of other family members is limited or not important. Phytosanitary control measures are used to diminish the virus spread in vegetatively propagated crops. Application of cross-protection with mild viral strains, which proved effective against damaging crop pathogens like PepMV, has been proposed. Transgenic plants of tobacco, tomato, and orchid, resistant to some potexviruses have been produced; they may have a potential for virus control.

Virus–Host Relationships Alphaflexiridae members are replicated in the cytoplasm of infected host cells. Infection is accompanied by formation of ER-derived vesicles that serve as replication sites (virus fabrics). Other cytopathic effects are the fibrous, beaded, banded, or irregular aggregates in the cytoplasm and, occasionally, in the nucleus.

Alphaflexiviruses (Alphaflexiviridae)

147

Fig. 5 Interveinal leaf yellowing symptoms after inoculation of RNA transcripts of PepMV infectious clone P-5-IYflc (variety Moneymaker). After Hasiów-Jaroszewska, B., Paeleman, A., Ortega-Parra, N., et al., 2013. Ratio of mutated versus wild-type coat protein sequences in Pepino mosaic virus determines the nature and severity of yellowing symptoms on tomato plants, Molecular Plant Pathology 14, 923–933.

On the molecular level, virus-host interactions involve contacts of the replicase and the movement proteins of the plantinfecting alphaflexiviruses with a variety of host factors that are required for efficient infection. After entering the host cell, the virus RNA genome is used as template for translation and replication. Several host factors involved in the movement of the potexvirus were identified. First, potexvirus TGB3 exhibits the affinity to highly curved subdomains of cortical ER, targets membrane bodies at the cell periphery and directs the TGB2 protein to these structures. Second, a serine/threonine kinase-like protein from N. benthamiana (NbSTKL) was demonstrated to be upregulated in BaMV infection and to be critical for the cell-to-cell movement. It is suggested that NbSTKL modifies a specific protein factor that regulates plasmadesmata (PD) gating for virus trafficking. In addition, another kinase, casein kinase 2a (CK2a), which interacts with BaMV CP in PD, can be involved in releasing viral RNA from the movement RNP after the passage of the latter through PD. Third, RabGTPase-activating protein (GAP or NbRabGAP1), is involved in the intercellular and systemic movement of the BaMV potexvirus. Rabs, a family of small GTPases, is involved in all aspects of intracellular vesicle budding, targeting, docking, and fusion. It was shown that NbRabGAP1 activates one of the RabGTPases to support the activity of the TGB proteins in transferring the viral movement complex through plasmodesmata. Involvement of NbRabGAP1 in the BaMV movement suggests that this movement is associated with the vesicle turnover pathway. Recently, several potential host factors affecting BaMV replication have also been identified. Among them, cytoplasmic 50 -30 exoribonuclease NbXRN4 promotes potexvirus replication, most probably, by removing uncapped genomic and subgenomic RNAs that are produced during replication/transcription as aberrant transcripts. At the replication stage, the 30 untranslated region (UTR) of the potexvirus genomic RNA plays an important role in capturing several host factors assisting the RNA synthesis. Glutathione transferase U4 (NbGSTU4), chloroplastic PGK (chlPGK), and heat shock protein 90 (Hsp90), were found to interact with the 30 UTR of BaMV. The protein chlPGK interacts with 30 UTR and seems to be vital for transmitting viral RNA and the replicase, to chloroplasts. The advantage of BaMV RNA targeting to the chloroplast can be explained by isolation of the VRC from the host protection systems, i.e., RNA silencing. The large viral RNA-protein complex including viral RNA, replicase, and possible host factors enters into chloroplasts, assisted by the Hsp90 acting as chaperone in protein complex folding and protein translocation across membranes. NbGSTU4, a protein involved in antioxidant protection, also stimulates BaMV replication. As soon as BaMV RNA and VRC components enter chloroplasts, they face the danger of high levels of reactive oxygen stress (ROS), since the chloroplast is one of the main producers of ROS. Movement of NbGSTU4 into chloroplasts along with BaMV RNA and the bound glutathione may play a role in eliminating the effects of ROS. Importantly, NbGSTU4 can interact with 30 UTR at physiological glutathione concentrations. Additionally, the S-adenosyl methionine synthetase and the respiratory explosion oxidase homolog were found to stimulate the BaMV replication. ER and myosins are employed in movement of potexviruses. Although the ER membrane allows the transport of membraneassociated molecules by diffusing within the lipid layer, the mobility of the ER membrane as well as the ER-directed transport of macromolecules is controlled by the ER-associated actin-myosin system. In accordance with this, the directed targeting of TGB proteins to PD and, consequently, the spread of infection also depends on actin and myosin activity. Antiviral plant protection is also controlled by complex interactions of host and viral factors. For example, antiviral reactions may be associated with nonsense-mediated degradation pathways of mRNA decay (NMD), which is a conservative mRNA quality control mechanism and selectively eliminates aberrant transcripts containing unusually long non-coding 30 terminal sections. By recognizing such viral RNAs and directing them to degradation, NMD neutralizes viral infection and functions as a viral restriction mechanism. A genetic screen in A. thaliana found that inactivating mutations in the UPF1 gene, which is a key NMD factor, increase the susceptibility of plants to PVX infection.

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Alphaflexiviruses (Alphaflexiviridae)

Alphaflexiviruses produce dsRNA as an intermediate in replication, making them targets for RNA silencing mechanism initiated by Dicer-like ribonucleases (DCL). After formation, siRNAs bind to AGOs, which are part of RISCs, to form a molecular platform for RNA silencing mechanism. To counteract RNA silencing, alphaflexiviruses express RNA silencing suppressors (VSRs) that inhibit various steps of RNA silencing. Thus, the potexvirus TGBp1 and mandarivirus CP act as VSRs. The TGBp1 of several potexviruses has been shown to have VSR activity inducing destabilization of AGO proteins. On the other hand, the PlAMV TGB1 protein inhibits secondary siRNA production by targeting the RDR6 pathway and additionally affects the AGO4 transfer from nucleus to cytoplasm. As alternative to RNA-targeted protective mechanisms, plants have developed a variety of resistance (R) proteins that can detect pathogen-associated molecular patterns (PAMPs) and induce protective reactions. Such detected effectors are called avirulence proteins (Avr). Most R-gene products have leucine-rich repeat domains (LRR) associated with the assumed nucleotide-binding site domain (NBS). Recent studies have identified molecular factors that affect the accumulation of NB-LRR and, consequently, the expression of resistance. Among them, the “susceptibility factor essential for potexvirus accumulation” (EXA1) protein containing the glycinetyrosine-phenylalanine (GYF) domain was identified as a negative regulator of NB-LRR protein accumulation and, as a result, involved in plant immunity. It is interesting that mutations affecting GYF domain protein EXA1 are also involved in resistance to potexviruses. Plantago asiatica mosaic virus (PlAMV), Alternanthera mosaic virus (AltMV), and PVX show significantly suppressed ability to establish infection in the absence of EXA1. Some plants contain a single EXA1 gene in their genomes. However, other plants, particularly N. benthamiana, soybean, and banana, encode multiple genes. It was found that EXA1 proteins in are important factors for successful potexvirus infection in a variety of plant species. Translation elongation factor eEF1a has been shown to regulate the RNA replication of potexviruses. As the eEF1a binding site in the 30 UTR of BaMV genome overlaps with the RdRp recognition site, eEF1a can play a role in template switch that blocks viral replication during translation. NADP þ - dependent isocitrate dehydrogenase, MAP kinase, phosphatase-like protein, and proliferating cell nuclear antigen (PCNA) also appear to suppress potexvirus replication. Recently, it was shown that phosphorylation of REMORIN protein changes its membrane dynamics. PVX TGBp1 is able to activate protein kinase(s), which in turn phosphorylate REMORIN1.3. Since phosphorylated REMORIN is associated with PD closure by induction of callose deposition, this protein causes the reduction of PD permeability and thereby restriction of viral cell-to-cell movement.

Diagnosis Diagnosis of viruses belonging to Alphaflexiviridae mostly relies on serological tests (enzyme-linked immunosorbent assay) and reverse transcriptase PCR methods, including the modifications of the latter like triplex PCR and droplet digital PCR.

Satellite RNAs Associated With Infections of Alphaflexiviridae The only satellite RNAs (satRNAs) so far known to accompany the infections of Alphaflexiviridae are those associated with BaMV potexvirus. The satRNAs are variable in sequence, but have the 50 - and 30 terminal hairpins resembling the cis-signals at the helper BaMV RNA termini. The vital functions provided by the helper BaMV to satRNAs are replication, encapsidation, and cell-to-cell movement. Co-replication of satRNAs has pronounced negative effect on the helper virus propagation. One of the BaMV satRNAs encodes a 20 kDa nonstructural protein which preferentially binds to the satRNA and promotes its long-distance transport in BaMV-infected N. benthamiana. The use of transgenic plants expressing satRNAs to control BaMV infection has been considered.

Further Reading Çağlayan, K., Roumi, V., Gaze, L.M., et al., 2019. Identification and characterization of a novel Robigovirus species from sweet cherry in Turkey. Pathogens 8, 57. Hasiów-Jaroszewska, B., Paeleman, A., Ortega-Parra, N., et al., 2013. Ratio of mutated versus wild-type coat protein sequences in Pepino mosaic virus determines the nature and severity of yellowing symptoms on tomato plants. Molecular Plant Pathology 14, 923–933. Lin, K.Y., Lin, N.S., 2017. Interfering satellite RNAs of bamboo mosaic virus. Frontiers in Microbiology 8, 787. Liu, Q., Yang, L., Xuan, Z., et al., 2020. Complete nucleotide sequence of Loquat virus A, a member of the family Betaflexiviridae with a novel genome organization. Archives of Virology 165, 223–226. Martelli, G.P., Adams, M.J., Kreuze, J.F., Dolja, V.V., 2007. Family Flexiviridae: A case study in virion and genome plasticity. Annual Review of Phytopathology 45, 73–100. Park, M.R., Seo, J.K., Kim, K.H., 2013. Viral and non-viral elements in potexvirus replication and movement and in antiviral responses. Advances in Virus Research 87, 75–112. Ur Rehman, A., Li, Z., Yang, Z., et al., 2019. The coat protein of Citrus yellow vein clearing virus interacts with viral movement proteins and serves as an RNA silencing suppressor. Viruses 11, 329. Verchot-Lubicz, J., Ye, C.M., Bamunusinghe, D., 2007. Molecular biology of potexviruses: Recent advances. Journal of General Virology 88, 1643–1655.

Alphasatellites (Alphasatellitidae) Rob W Briddon and Muhammad S Nawaz-ul-Rehman, University of Agriculture, Faisalabad, Pakistan r 2021 Published by Elsevier Ltd.

Nomenclature CR-M Common region major CR-SL Common region stem-loop mRep Master replication-associated protein ORF Open reading frame

Glossary Post-transcriptional gene silencing An epigenetic process for regulation of gene expression that results in the mRNA of a particular gene being destroyed. Post-transcriptional gene silencing is believed to protect the organism’s genome from, among other things, transposons and viruses. Rolling-circle replication A mechanism of DNA replication that synthesizes multiple copies of circular molecules of DNA. Rolling circle replication is initiated by an initiator protein that nicks one strand of a doublestranded, circular DNA molecule. The initiator protein remains bound to the 50 phosphate end of the nicked strand and the free 30 hydroxyl end is released to serve as a primer

PTGS Post-transcriptional gene silencing Rep Replication initiator protein ssDNA single-stranded deoxyribonucleic acid TGS Transcriptional gene silencing

for DNA synthesis using the unnicked strand as a template; replication proceeds around the circular DNA molecule, displacing the nicked strand as single-stranded DNA. Satellite like molecule A self replicating molecule which depends on helper virus for movement, encapsidation and transmission. Transcriptional gene silencing An epigenetic process for regulation of gene expression that results from DNA methylation and histone modifications. Transcriptional gene silencing is believed to protect the organism’s genome from, among other things, transposons and integrated viruses.

Introduction Viruses of the family Nanoviridae, some begomoviruses (genus Begomovirus, family Geminiviridae) and, in one case reported so far, a mastrevirus (genus Mastrevirus, family Geminiviridae) may be associated with additional circular ssDNA components that are known as alphasatellites. Recently coconut foliar decay, a severe disease of coconut palms in Vanuatu, has been shown to be associated with a multipartite ssDNA virus that is distinct from nanoviruses and geminiviruses and to be associated with multiple (possible) alphasatellites. The association with alphasatellites may thus extend to three families of plant-infecting ssDNA viruses. Nanoviruses (family Nanoviridae) are multipartite ssDNA viruses that infect plants and are transmitted by aphids. The genomes of nanoviruses are composed of 6–8 circular components, each B1100 nt in size, separately encapsidated in isometric particles. The component known as DNA-R encodes the master replication-associated protein (mRep) that is responsible for trans-replication of all bona fide virus components. mRep recognises the origin of replication (ori) which sits in the common region stem-loop (CR-SL), conserved between all bona fide virus components, and initiates rolling circle replication by nicking within the nonanucletide sequence TAGTATTAC that forms part of the loop of a predicted stem-loop structure. The bona fide nanovirus components also contain a second conserved sequence known as the common region major (CR-M). Geminiviruses (family Geminiviridae) are plant-infecting viruses with geminate (twinned quasi-icosahedral) particles that are transmitted by insect vectors. Begomoviruses (genus Begomovirus) have either monopartite or bipartite genomes and are transmitted by the whitefly Bemisia tabaci whereas mastreviruses (genus Mastrevirus) have monopartite genomes and are transmitted by specific leafhoppers. The first alphasatellite was identified in association with the nanovirus Subterranean clover stunt virus in 1995. Sequencing of the genomic components of the virus highlighted the presence of more than one component encoding a replication-associated protein, a feature that was shown for all subsequently characterized nanoviruses. Realisation that the additional Rep-encoding components are satellites came with the demonstration that only the mRep has the capacity to direct replication of the other genome components using infectious clones. The first geminivirus-associated alphasatellite was identified in 1999 in cotton affected by Cotton leaf curl disease (CLCuD). Cotton leaf curl Multan alphasatellite was identified fortuitously, by shotgun-cloning, in the search for additional components responsible for the symptoms of CLCuD in cotton. The symptoms were subsequently shown to be encoded by a component we now know as a betasatellite (family Tolecusatellitidae; genus Betasatellite).

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Fig. 1 Comparison of the structures nanovirus- and geminivirus-associated alphasatellites in comparison to the DNA-R genomic components of nanoviruses. The diagram shows the position of the replication-associated protein (Rep), the TATA box of the presumed Rep promoter, an adenine rich sequence (A-rich) in geminivirus-associated alphasatellites, as well as the common region major (CR-M) and common region stem-loop (CR-SL) of nanovirus DNA-R components that is conserved between all bona fide nanovirus genome components. All components have a predicted stem-loop structure (at position zero) with, in most cases, the nona-nucleotide sequence TAGTATTAC forming part of the loop. This likely forms the origin of virion-strand DNA replication that is nicked by Rep to initiate rolling-circle replication. Note that the DNA-R components of some nanoviruses contain an additional open reading frame (ORF) known as ORF 1.

Classification The additional circular ssDNA components associated with nanoviruses and geminiviruses are classified in the recently established family Alphasatellitidae. The family encompasses two subfamilies, Nanoalphasatellitinae and Geminialphasatellitinae, to accommodate the nanovirus and geminivirus alphasatellites, respectively. At present the subfamily Nanoalphasatellitinae encompasses seven genera and 19 species, whereas the subfamily Geminialphasatellitinae encompasses 4 genera with 43 assigned species. Additionally a divergent possible alphasatellite, associated with coconut foliar decay disease, has been assigned to a species but not a subfamily. Within this classification structure alphasatellites are assigned based on sequence and phylogenetic relatedness.

Structure of Alphasatellites In common with the DNA-R component of nanoviruses, the alphasatellites encode a Rep protein and have a predicted stem-loop structure with, in most cases, the nona-nucleotide sequence TAGTATTAC forming part of the loop (Fig. 1). Alphasatellites associated with nanoviruses have a size typical of nanovirus components (B1100 nt), but lack the conserved CR-SL and CR-M of their helper viruses and consequently are unable to trans-replicate the bona fide genome components of their helper viruses. Typically the geminivirus-associated alphasatellites comprise B1400 nt, being significantly larger than nanovirus components and nanovirus-associated alphasatellites. The size difference is due to the presence in geminivirus-associated alphasatellites of a sequence rich in adenine and a larger Rep-encoding gene (Fig. 1). The size difference is likely necessary for the movement and/or encapsidation of geminivirus alphasatellites by their helper viruses. Geminiviruses have a stringent size surveillance mechanism that operates during virus movement within the plant as well as size constraints for encapsidation. Being approximately half the geminivirus genome length (B1400 nt), geminialphasatellites may possibly be encapsidated in isometric (half geminate) virions. (Fig. 2).

Likely Origins of Alphasatellites Based on sequence relatedness it is likely that nanoalphasatellites evolved from the DNA-R components of nanoviruses that were captured by a second nanovirus during a co-infection. A similar “capture following co-infection” mechanism may have resulted in the geminialphasatellites, although the progenitor DNA-R then needed to adapt, by size increase, for efficient maintenance by a geminiviruses. Consistent with this hypothesis it is interesting to note that, having been uncoupled from the mechanism which controls the relative titers of bona fide nanovirus components, alphasatellites of both nanoviruses and geminiviruses in many cases vastly exceed the titers of their helper virus component(s) in plants. It is interesting to note that vast majority of geminivirus-associated alphasatellites from the Old World have been identified in association with monopartite begomoviruses infections that also include a betasatellite (family Tolecusatellitidae). In the New World, where only very few monopartite begomoviruses and no betasatellites have been identified, alphasatellites have only so far been reported to occur in association with bipartite begomoviruses. In contrast, nanoviruses have only been identified in the Old World, which may suggest that the association of alphasatellites with geminiviruses evolved in the Old World, although when this occurred is unclear.

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Fig. 2 (A) Maximum likelihood phylogenetic tree of representative geminivirus-associated alphasatellite sequences from each species. (B) Maximum likelihood phylogenetic trees of the nanovirus-associated alphasatellite sequences. Trees are inferred using IQ-TREE with TIM2 þ F þ I þ G4 chosen as the best fit model. Branches with less than 60% bootstrap support have been collapsed. The Maximum likelihood phylogenetic tree on the left is a guide for genera demarcation with the cyan line showing the rough 67% genus demarcation threshold. Both trees reproduced from Briddon, R.W., Martin, D.P., Roumagnac, P., et al., 2018. Alphasatellitidae: a new family with two subfamilies for the classification of geminivirus‑ and nanovirus‑associated alphasatellites. Archives of Virology 163, 2587–2600. doi:10.1007/s00705-018-3854-2.

Table 1

List of alphasatellites with subfamilies, genera, species, acronym and accession number. Type species in each genus is written in bold

Subfamily/Genus

Species

Acronym

Accession #

Ageratum yellow vein Singapore alphasatellite Cotton leaf curl Saudi Arabia alphasatellite

AYVSGA CLCuSAA

NC_003414 HG530543

Clecrusatellite

Cleome leaf crumple alphasatellite Croton yellow vein mosaic alphasatellite Euphorbia yellow mosaic alphasatellite Melon chlorotic mosaic alphasatellite Sida Cuba alphasatellite Tomato yellow spot alphasatellite Whitefly associated Guatemala alphasatellite 2 Whitefly associated Puerto Rico alphasatellite 1

ClLCrA CrYVMA EuYMA MeCMA SiCUA ToYSA WfaGUA2 WfaPRA1

NC_014646 NC_013801 NC_014630 NC_014379 NC_021708 NC_038993 NC_038973 NC_038975

Colecusatellite

Ageratum enation alphasatellite Ageratum yellow vein alphasatellite Ageratum yellow vein China alphasatellite Ageratum yellow vein India alphasatellite Bhendi yellow vein alphasatellite Cassava mosaic Madagascar alphasatellite Chilli leaf curl alphasatellite Cotton leaf curl Egypt alphasatellite Cotton leaf curl Gezira alphasatellite Cotton leaf curl Lucknow alphasatellite Cotton leaf curl Multan alphasatellite Gossypium darwinii symptomless alphasatellite Malvastrum yellow mosaic alphasatellite Malvastrum yellow mosaic Cameroon alphasatellite Pedilanthus leaf curl alphasatellite Sida leaf curl alphasatellite

AEA AYVA AYVCNA AYVINA BhYVA CMMGA ChLCuA CLCuEGA CLCuGeA CLCuLuA CLCuMuA GDarSLA MaYMA MaYMCMA PeLCuA SiLCuA

NC_019547 NC_033555 NC_023443 NC_019546 NC_029313 NC_018628 NC_039234 AJ512960 NC_013593 NC_015327 NC_018082 NC_039232 NC_039239 NC_014907 NC_033335 NC_039240 (Continued )

Geminialphasatellitinae Ageyesisatellite

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Table 1

Continued

Subfamily/Genus

Species

Acronym

Accession #

Sida yellow vein Vietnam alphasatellite Sunflower leaf curl Karnataka alphasatellite Synedrella leaf curl alphasatellite Tobacco curly shoot alphasatellite Tomato leaf curl Cameroon alphasatellite Tomato yellow leaf curl China alphasatellite Tomato yellow leaf curl Thailand alphasatellite Tomato yellow leaf curl Yunnan alphasatellite

SiYVVNA SLCuKaA SyLCuA TbCSA ToLCuCMA TYLCuCNA TYLCuTHA TYLCuYnA

NC_009563 NC_019499 NC_025244 NC_005057 NC_014747 NC_043449 AJ579359 NC_039229

Gosmusatellite

Gossypium mustelinum symptomless alphasatellite Hollyhock yellow vein alphasatellite Mesta yellow vein mosaic alphasatellite Okra enation leaf curl alphasatellite Okra yellow crinkle Cameroon alphasatellite Vernonia yellow vein Fujian alphasatellite

GMusSLA HoYVA MeYVMA OEnLCuA OkYCCMA VeYVFjA

NC_038954 NC_039084 NC_018573 NC_019944 NC_014906 NC_039233

Unassigned

Ageratum leaf curl Cameroon alphasatellite Dragonfly associated alphasatellite Parthenium leaf curl alphasatellite Tomato leaf curl Buea alphasatellite Whitefly associated Guatemala alphasatellite 1

ALCuCMA DfaA PLCuA ToLCuBuA WfaGUA1

NC_014744 NC_019498 NC_030226 NC_038903 NC_038974

Banana bunchy top alphasatellite 1 Banana bunchy top alphasatellite 2 Banana bunchy top alphasatellite 3 Cardamom bushy dwarf alphasatellite

BBTA2 BBTA1 BBTA3 CaBuDA

NC_038892 NC_038953 NC_038955 NC_022919

Clostunsatellite

Milk vetch dwarf alphasatellite 2 Pea necrotic yellow dwarf alphasatellite 2 Sophora yellow stunt alphasatellite 4 Sophora yellow stunt alphasatellite 5 Subterranean clover stunt alphasatellite 2

MVDA2 PNYDA2 SYSA4 SYSA5 SCSA2

NC_003639 NC_038959 NC_039000 NC_038996 NC_003818

Fabenesatellite Milvetsatellite Mivedwarsatellite

Faba bean necrotic yellows alphasatellite 2 Milk vetch dwarf alphasatellite 3 Faba bean necrotic stunt alphasatellite Milk vetch dwarf alphasatellite 1 Pea necrotic yellow dwarf alphasatellite 1 Sophora yellow stunt alphasatellite 2

FBNYA2 MVDA3 FBNSA MVDA1 PNYDA1 SYSA2

NC_003567 NC_003640 NC_023881 NC_003638 NC_038958 NC_038999

Sophoyesatellite Subclovsatellite

Sophora yellow stunt alphasatellite 3 Faba bean necrotic yellows alphasatellite 1 Sophora yellow stunt alphasatellite 1 Subterranean clover stunt alphasatellite 1

SYSA3 FBNYA1 SYSA1 SCSCA1

NC_038998 NC_024886 NC_038997 NC_003814

Coconut foliar decay alphasatellite

CFDA

NC_001465

Nanoalphasatellitinae Babusatellite

Unassigned

Life Cycle/Epidemiology Alphasatellites replicate autonomously of their helper viruses in the nuclei of permissive host cells by rolling circle replication. For all other functions alphasatellites depend upon factors provided by their helper viruses including movement within/between cells, and encapsidation for transmission between plants. However, there appears to be little, if any, specificity for helper virus of a particular alphasatellite. For example, the geminialphasatellite Ageratum yellow vein Singapore alphasatellite (AYVSGA; genus Ageyesisatellite), which was identified in association with the OW monopartite begomovirus Ageratum yellow vein virus, was shown to be maintained in plants by the NW bipartite begomovirus Tomato golden mosaic virus and the curtovirus (family Geminiviridae, genus Curtovirus) Beet curly top virus (BCTV). When maintained in plants by BCTV, AYVSGA co-insect transmitted by the leafhopper vector of BCTV, showing the vector of the alphasatellite to depend upon the vector specificity of the helper virus. The same work also showed AYVSGA not to be maintained in plants by the mastrevirus (family Geminiviridae, genus Mastrevirus) Chickpea chlorotic dwarf virus (CpCDV; earlier known as Bean yellow dwarf virus), possibly due to packaging constraints; the genome of CpCDV is significantly smaller than the genomes of the begomoviruses by which it is maintained.

Alphasatellites (Alphasatellitidae)

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Pathogenesis The precise impact of alphasatellites on nanovirus and geminivirus infections remains unclear. For some nanovirus infections the presence of nanoalphasatellites has been shown to reduce nanovirus infectivity. This suggests that there is competition between the nanoalphasatellite and the bona fide nanovirus genomic DNAs for factors required for replication, systemic movement and/or encapsidation. For geminivirus infections the presence of geminialphasatellites has been shown to result in either attenuated or exacerbated symptoms. For infections with attenuated symptoms the DNA titer of either the virus genome or the associated betasatellite were found to be reduced in comparison to plants not inoculated with the alphasatellite. The Rep proteins encoded by the geminialphasatellites Gossypium darwinii symptomless alphasatellite and Gossypium mustelinum symptomless alphasatellite suppress post-transcriptional gene silencing (PTGS), whereas those encoded by various other geminialphasatellites are suppressors of transcriptional gene silencing (TGS). This suggests that some, if not all, alphasatellites assist the helper virus in overcoming (suppressing) host resistance based on RNA interference (Table 1).

See also: Betasatellites and Deltasatelliles (Tolecusatellitidae). Geminiviruses (Geminiviridae). Nanoviruses (Nanoviridae)

Further Reading Abbas, Q., Amin, I., Mansoor, S., et al., 2019. The Rep proteins encoded by alphasatellites restore expression of a transcriptionally silenced green fluorescent protein transgene in Nicotiana benthamiana. Virus Disease 30 (1), 101–105. Boevink, P., Chu, P.W.G., Keese, P., 1995. Sequence of Subterranean clover stunt virus DNA: affinities with the geminiviruses. Virology 207, 354–361. Briddon, R.W., Martin, D.P., Roumagnac, P., et al., 2018. Alphasatellitidae: a new family with two subfamilies for the classification of geminivirus‑ and nanovirus‑associated alphasatellites. Archives of Virology 163, 2587–2600. doi:10.1007/s00705-018-3854-2. Gronenborn, B., Randles, J.W., Knierim, D., et al., 2018. Analysis of DNAs associated with coconut foliar decay disease implicates a unique single-stranded DNA virus representing a new taxon. Scientific Reports 8, 5698. Kumar, J., Kumar, J., Singh, S.P., Tuli, R., 2014. Association of satellites with a mastrevirus in natural infection: complexity of Wheat dwarf India virus disease. Journal of Virology 88, 7093–7104. Mansoor, S., Khan, S.H., Bashir, A., et al., 1999. Identification of a novel circular single-stranded DNA associated with cotton leaf curl disease in Pakistan. Virology 259, 190–199. Mar, T.B., Mendes, I.R., Lau, D., et al., 2017. Interaction between the new world begomovirus Euphorbia yellow mosaic virus and its associated alphasatellite: Effects on infection and transmission by the whitefly Bemisia tabaci. Journal of General Virology 98, 1552–1562. Nawaz-Ul-Rehman, M.S., Nahid, N., Mansoor, S., Briddon, R.W., Fauquet, C.M., 2010. Post-transcriptional gene silencing suppressor activity of two non-pathogenic alphasatellites associated with a begomovirus. Virology 405, 300–308. Saunders, K., Bedford, I.D., Stanley, J., 2002. Adaptation from whitefly to leafhopper transmission of an autonomously-replicating nanovirus-like DNA component associated with ageratum yellow vein disease. Journal of General Virology 83, 909–915. Sicard, A., Yvon, M., Timchenko, T., et al., 2013. Gene copy number is differentially regulated in a multipartite virus. Nature Communications 4. Timchenko, T., Katul, L., Aronson, M., et al., 2006. Infectivity of nanovirus DNAs: Induction of disease by cloned genome components of Faba bean necrotic yellows virus. Journal of General Virology 87, 1735–1743. Zhou, X., 2013. Advances in understanding begomovirus satellites. Annual Review of Phytopathology 51, 357–381.

Amalgaviruses (Amalgaviridae) Ioannis E Tzanetakis, University of Arkansas, Fayetteville, United States Sead Sabanadzovic, Mississippi State University, Starkville, MS, United States Rodrigo A Valverde, Louisiana State University Agricultural Center, Baton Rouge, United States r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

RdRp RNA-dependent RNA polymerase RT-LAMP Reverse transcription loop-mediated isothermal amplification RT-PCR Reverse transcription polymerase chain reaction

CP Coat protein dsRNA Double stranded RNA ORF Open reading frame

Glossary Bicistronic Nucleic acid that codes for two proteins. Nucleoprotein Protein protecting the genomic RNA of negative sense RNA viruses.

Ribosomal frameshift Translation strategy, quite widespread in viruses, that allows for the expression of two partially overlapping, non-in-frame ORFs in a single round of translation involving ribosome slipping by one base in either the 50 or 30 directions.

Classification The family Amalgaviridae, part of the realm Riboviria, was recognized by the International Committee on Taxonomy of Viruses (ICTV) in 2014. The family includes viruses infecting plants and fungi (yeasts) with a non-segmented, bicistronic dsRNA genome of 3.1–3.5 kbp and is currently composed of two genera: Amalgavirus and Zybavirus. The genus Amalgavirus contains 9 officially recognized species while the genus Zybavirus has only one species. The type species for genus Amalgavirus is Southern tomato virus, and the type species for the genus Zybavirus is Zygosaccharomyces bailii virus Z (Table 1). There are also another 27 possible members of the family for which sequences are available and phylogenetic studies indicate that at least 12 of them could be classified in the genus Amalgavirus while one (Antonospora locustae virus 1, AnLoV-1) could be classified either in the genus Zybavirus or represent a completely new genus in the family (Fig. 1).

Virion Structure Virus particles have never been observed for plant amalgaviruses under the electron microscope, but antibodies against the putative product of ORF1, with possible role of the viral CP, reacted with amorphous bodies in infected plants. Biochemical analysis points to protection of the genome by a protein. In silico analyses indicate that the putative CP of amalgaviruses is homologous to the nucleoprotein of animal and plant-infecting negative stranded RNA viruses suggesting the possibility of filamentous particles instead of the hypothesized spherical ones. The spherical particle hypothesis is based on Zygosaccharomyces bailii virus Z (ZbV-Z), where virus-like particles where purified from yeast. Notwithstanding, purification was performed using an isolate co-infected with a totivirus which is known to form spherical particles.

Genome The genome of amalgaviruses is comprised of a single double stranded RNA (dsRNA) molecule of B3.5 kbp (viruses in the genus Amalgavirus) or B3.1 kbp in size (Zygosaccharomyces bailii virus Z, sole member of the genus Zygovirus). All viruses in the family code for two proteins (Fig. 2). The 50 proximal protein is presumed to be the CP of the virus. As noted above, the protein is homologous to the nucleoprotein of animal and plant-infecting negative strand viruses. The second protein, encoded by ORF2, is predicted to be expressed via a programmed þ 1 ribosomal frameshift and contains RdRp motifs most similar to those found in members of the family Partitiviridae. Putative þ 1 ribosomal frameshifting motif, UUU_CGN, present in all but two plant amalgaviruses was identified recently by extensive in silico analyses. Phylogenetic analyses have shown that RdRp of amalgaviruses is evolutionarily closer to that of partitiviruses than to counterparts expressed by members of the family Totiviridae. Overall, the genome appears to be an amalgam of the monopartite family Totiviridae and the multipartite family Partitiviridae.

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Amalgaviruses (Amalgaviridae)

Table 1

155

List of members in the family Amalgaviridae. Type species are written in bold. Accessions of complete genome sequences are provided

Genus

Species

Virus

Acronym

Accession #

Amalgavirus

Allium cepa amalgavirus 1 Allium cepa amalgavirus 2 Blueberry latent virus Rhododendron virus A Southern tomato virus Spinach amalgavirus 1 Vicia cryptic virus M Zoostera marina amalgavirus 1 Zoostera marina amalgavirus 2

Allium cepa amalgavirus 1 Allium cepa amalgavirus 2 Blueberry latent virus Rhododendron virus A Southern tomato virus Spinach amalgavirus 1 Vicia cryptic virus M Zoostera marina virus 1 Zoostera marina virus 2

AcAV1 AcAV2 BlLV RhVA STV SpAV1 VCV-M ZmAV1 ZmAV2

NC_036580 NC_036581 NC_014593 NC_014481 NC_011591 NC_035070 NC_043095 NC_034614 NC_034615

Zybavirus

Zygosaccharomyces bailii zybavirus

Zygosaccharomyces bailii virus Z

ZbV-Z

NC_003874

Anthoxanthum odoratum amalgavirus 1 Anthoxanthum odoratum amalgavirus 2 Antonospora locustae virus 1 Camellia oleifera amalgavirus 1 Capsicum annuum amalgavirus Cistus incanus RNA virus 1 Cleome droserifolia amalgavirus Cucumis melo amalgavirus Erigeron breviscapus amalgavirus 1 Erigeron breviscapus amalgavirus 2 Festuca pratensis amalgavirus 1 Festuca pratensis amalgavirus 2 Festuca pratensis amalgavirus 3 Gevuina avellana amalgavirus 1 Lolium perenne amalgavirus 1 Medicago sativa amalgavirus 1 Neurachne minor latent virus 1 Phalaenopsis equestris amalgavirus 1 Pinus patula amalgavirus 1 Pterostylis amalgavirus Rubber dandelion latent virus 1 Rubber dandelion latent virus 2 Salicornia europaea amalgavirus Salvia hispanica RNA virus 1 Secale cereale virus

AoAV1 AoAV2 AlV1 CoAV1 CaAV-1 CiRV1 CdAV1 CmAV EbAV1 EbAV2 FpAV1 FpAV2 FpAV3 GaAV1 LpAV1 MsAV1 NmLV1 PeAV1 PpAV1 PdV RdLV1 RdLV2 SeAV1 ShRV1 ScV

NC_040432 BK010357 NC_035189 NC_040433 NC_040662 MG833407 NC_040777 MH479774 NC_040592 NC_040593 NC_040663 NC_040778 BK010350 BK010351 BK010352 NC_040591 MF325033 NC_040590 BK010353 NDa MF197380 MF197379 BK010404 NC_040690 BK010349

Unassigned viruses

a

ND, no complete genome sequence available.

Life Cycle Little is known about the life cycle of amalgaviruses. The plant-infecting members of the family are not mechanically, nor grafttransmissible. They are very efficiently transmitted through seed, reaching 100% transmission in the case of southern tomato virus (STV) and blueberry latent virus (BLV). ZbV-Z can be transmitted to other yeast species.

Epidemiology Given the lack of a identifiable movement protein it is hypothesized that the plant-infecting members of the family move within their hosts only during cell division. No symptoms have been observed in hosts infected with any member of the genus Amalgavirus although there are reports of STV having a mutualistic interaction with tomato or affecting the expression of tomato miRNAs. The effect of ZbV-Z on its hosts remains unknown.

Pathogenesis None known.

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Fig. 1 Phylogenetic tree of recognized (red font) and putative members (black) of the family Amalgaviridae. The tree was reconstructed on amino acid sequences of viral RdRPs initially aligned with MUSCLE (v3.8.31) and curated by Gblocks (v0.91b) in order to remove ambiguous regions. The phylogenetic tree was reconstructed with the maximum likelihood method implemented in the PhyML program (v3.1/3.0 aLRT), applying LG þ G substitution model, estimated to be best-fit for this dataset. Reliability for internal branch was assessed using the aLRT test (SH-Like). The tree was edited and visualized with FigTree (v1.44). Complete names of viruses used in the analyses are: Allium cepa amalgavirus 1 (AcAV1; BK010347), Allium cepa amalgavirus 2 (AcAV2; BK010348), Anthoxanthum odoratum amalgavirus 1 (AoAV1; BK010356), Antonospora locustae virus 1 (AnloV1; KX525322), blueberry latent virus (BlLV; HM029246), Camellia oleifera amalgavirus 1 (CoAV1; BK010409), Capsicum annuum amalgavirus 1 (CaAV1; BK010407), Cleome droserifolia amalgavirus 1 (ClDAV1; BK010408), Erigeron breviscapus amalgavirus 1 (EbAV1; BK010410), Erigeron breviscapus amalgavirus 2 (EbAV2; BK010411), Festuca pratensis amalgavirus 1 (FpAV1; BK010412), Festuca pratensis amalgavirus 2 (FpAV2; BK010413), Medicago sativa amalgavirus 1 (MsAV1; BK010406), Neurachne minor latent virus (NmLV1; MF325033) Phalaenopsis equestris amalgavirus 1 (PeAV1; BK010405), rhododendron virus A (RhVA; HQ128706) Salvia hispanica RNA virus 1 (ShRV1; MH988691), southern tomato virus (STV; EF442780), spinach amalgavirus 1 (SpAV1; KY695011), Vicia cryptic virus M (VCVM; EU371896), Zostera marina amalgavirus 1 (ZmAV1; KY783316), Zostera marina amalgavirus 2 (ZmAV2; KY783317), Zygosaccharomyces bailii virus Z (ZbV-Z; KU200450). Amino acid sequences of RdRPs of Nigrospora oryzae unassigned RNA virus 1 (NoURV1; KT258976) and Ustilaginoidea virens nonsegmented virus 1 (UvNV1; KJ605397) were used as outgroups. The two genera of the family, Amalgalvirus and Zybavirus, indicated.

Fig. 2 Genome depiction and putative protein products of Southern tomato virus (STV), the type member of the genus Amalgavirus. The two open reading frames are coded in the same strand and translate to the putative coat protein (red block) and its fusion with the RNA dependent RNA polymerase (RdRp; blue).

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157

Diagnosis There are many protocols developed for the type species of the genus Amalgavirus, southern tomato virus, which include RT-PCR, quantitative RT-PCR, molecular hybridization and RT-LAMP. The other viruses in the family are primarily detected by virus-specific RT-PCR.

Treatment Virus-elimination protocols such as thermo-/chemo-/cryotherapy have not been reported for any amalgavirus. Yeast can be cured from ZbV-Z using cycloheximide or elevated temperatures.

Prevention The best prevention approach for all plant viruses including amalgaviruses is to start with clean propagative materials. Independent of true seed or clonal propagation, mother plants should be tested free of the respective amalgavirus.

See also: Fungal Partitiviruses (Partitiviridae). Totiviruses (Totiviridae)

Further Reading Depierreux, D., Vong, M., Nibert, M.L., 2016. Nucleotide sequence of Zygosaccharomyces bailii virus Z: Evidence for þ 1 programmed ribosomal frameshifting and for assignment to family Amalgaviridae. Virus Research 217, 115–124. Elvira-González, L., Medina, V., Rubio, L., Galipienso, L., 2020. The persistent Southern tomato virus modifies miRNA expression without inducing symptoms and cell ultrastructural changes. European Journal of Plant Pathology 156, 615–622. Elvira-Gonzalez, L., Puchades, A.V., Carpino, C., et al., 2017. Fast detection of Southern tomato virus by one-step transcription loop-mediated isothermal amplification (RT-LAMP). Journal of Virological Methods 241, 11–14. Fukuhara, T., Tabara, M., Koiwa, H., Takahashi, H., 2020. Effect of asymptomatic infection with Southern tomato virus on tomato plants. Archives of Virology 165, 11–20. Isogai, M., Nakamura, T., Ishii, K., et al., 2011. Histochemical detection of Blueberry latent virus in highbush blueberry plant. Journal of General Plant Pathology 77, 304. Martin, R.R., Zhou, J., Tzanetakis, I.E., 2011. Blueberry latent virus: An amalgam of the Partitiviridae and Totiviridae. Virus Research 155, 175–180. Nibert, M.L., Pyle, J.D., Firth, A.E., 2016. A þ 1 ribosomal frameshifting motif prevalent among plant amalgaviruses. Virology 498, 201–208. Puchades, A.V., Carpino, C., Alfaro‐Fernandez, A., et al., 2017. Detection of Southern tomato virus by molecular hybridisation. Annals of Applied Biology 171, 172–178. Sabanadzovic, S., Valverde, R.A., Brown, J.K., Martin, R.R., Tzanetakis, I.E., 2009. Southern tomato virus: The link between the families Totiviridae and Partitiviridae. Virus Research 140, 130–137. Schmitt, M.J., Neuhausen, F., 1994. Killer toxin-secreting double-stranded RNA mycoviruses in the yeasts Hanseniaspora uvarum and Zygosaccharomyces bailii. Journal of Virology 68, 1765–1772.

Relevant Websites https://viralzone.expasy.org/4956?outline=all_by_species Amalgavirus ViralZone. https://talk.ictvonline.org/taxonomy/ Taxonomy ICTV.

Badnaviruses (Caulimoviridae) Andrew DW Geering, The University of Queensland, St. Lucia, QLD, Australia r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

NLS Nuclear localization signal nm Nanometre ORF Open reading frame pgRNA Pregenomic RNA ssRNA Single-stranded RNA tRNA Transfer RNA uORF Upstream ORF VAP Virion-associated protein

CP Capsid protein dsDNA Double-stranded DNA kbp Kilobase pairs kDa Kilodaltons MP Movement protein NC Nucleocapsid NID N-terminal intrinsically disordered

Glossary Discontinuity Gap in the DNA strand. Endogenous viral element Viral DNA integrated in the nuclear genome of the host organism and inherited from parents like any normal cellular gene. Minichromosome Viral DNA wrapped around histone proteins in the nucleus.

Reverse transcription Use of an RNA template to direct synthesis of a complementary strand of DNA. Virion The virus particle, consisting of a DNA genome surrounded by a protein shell and constituting the infective form of the virus. Viroplasms Virus factories.

Introduction The genus Badnavirus is a group of double-stranded (ds) DNA viruses with bacilliform particles in the family Caulimoviridae. They are referred to as ‘reverse-transcribing viruses’ as they produce a single-stranded RNA replicative intermediate, which acts as the template for reverse transcription back to dsDNA. Badnaviruses have been mainly identified in tropical regions and they have an extremely wide host range: a significant number of badnaviruses are pathogens of crop plants, such as banana, citrus, cacao, fig, grapevine, pepper, pineapple, raspberry, sugarcane, and sweet-potato, but many more are been found in ornamental plants such as bougainvillea, hibiscus, kalanchoe, lucky bamboo, pelargonium and wisteria. So far badnaviruses have been shown to be transmitted by two types of insect vector, aphids and mealybugs. A total of 68 species are currently recognized by the ICTV, and in addition there are six tentative species in the genus. There is also evidence in the form of endogenous viral elements (viral DNA that is integrated in the plant genome) that many more plant species have been exposed to badnaviruses throughout their evolution. The molecular species demarcation criteria of 80% identity of nucleotide sequences has been set 20 years ago, and now that so many more badnavirus complete genome sequences are available, this demarcation threshold should be revisited.

Classification The genus Badnavirus, typified by Commelina yellow mottle virus, currently contains 68 officially recognized species and six tentative virus species (Table 1), making it the largest of all genera in the family Caulimoviridae. All members of the family Caulimoviridae have the conserved core of replication and structural proteins found within the order Ortervirales, called the gag-pol, which are names originating from the retrovirus literature and are abbreviations of ‘group antigens’ and ‘polymerase’, respectively. In badnaviruses, the gag gene encodes the major capsid protein (CP) and the pol gene, the replication-associated enzymes, namely the aspartic protease and reverse transcriptase/ribonuclease H1 proteins. The caulimoviruses are sometimes referred to as ‘plant-infecting pararetroviruses’ but the term ‘pararetrovirus’ has no taxonomic standing and is purely a descriptive term used to describe reverse-transcribing viruses that encapsidate a double-stranded DNA (dsDNA) form of the viral genome rather than single-stranded RNA (ssRNA) in the case of retroviruses. Caulimoviruses are united with all members of the Ortervirales by alternating between double-stranded DNA and singlestranded RNA during the replication cycle through cycles of transcription and reverse transcription. The RNA intermediate in the replication cycle is termed the pregenomic (pg) RNA. The key characteristic that differentiates the genus Badnavirus from all other members of the family Caulimoviridae is genome organization. The criteria for demarcation of different badnavirus species is less than 80% nucleotide identity within the reverse transcriptase and ribonuclease H1 coding regions of ORF3, differences in host range and vector specificities. Evolutionary relationships within the genus are illustrated in Fig. 1.

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Ghana Ghana Indonesia (?), Ghana, Togo Cote d0 Ivoire, Togo Trinidad and Tobago USA

Theobroma cacao Theobroma cacao Theobroma cacao Theobroma cacao Theobroma cacao Canna indica

MF642725

MF642734

AJ781003

L14546

KX276641

KX066020

KU168312

CSSGNV

CSSGQV

CSSTAV

CSSTBV

CYVBV

CaYMaV

CaYMV

CiYMV

Citrus yellow mosaic virus

AF347695

Ghana

Theobroma cacao

CSSGMV MF642724

Austria, China, Italy, India, Kenya, Russia, The Netherlands, USA Citrus x aurantifolia, Citrus x jambhiri, Citrus x limonia, Citrus maxima, Citrus reticulata, India Citrus x sinensis

Alpinia purpurata, Canna indica, Canna hybrids, Piper betle

Cote d0 Ivoire, Ghana

Theobroma cacao

MF642720

CSSCEV

Puerto Rico, Trinidad and Tobago Cote d0 Ivoire

Theobroma cacao Theobroma cacao

KX276640 JN606110

CMMV CSSCDV

Sri Lanka

Theobroma cacao

MF642736

CBSLV

(Continued )

Yes

USA Brazil, China, Malaysia, Taiwan

Rubus hybrids Bougainvillea spectabilis

KJ413252 EU034539

BVF BCVBV

Endogenous?

Musa spp. Musa spp. Musa spp. Musa spp. Musa spp. Musa spp. Musa spp. Betula pubescens, Betula pendula

AY805074 AJ002234 HQ593107 HQ593108 HQ593109 HQ593110 AY750155 MG686419

BSMYV BSOLV BSUAV BSUIV BSULV BSUMV BSVNV BLRaV

Geographical distribution

Banana streak MY virus Banana streak OL virus Banana streak UA virus Banana streak UI virus Banana streak UL virus Banana streak UM virus Banana streak VN virus Birch leaf roll-associated virus Blackberry virus F Bougainvillea chlorotic vein banding virus Cacao bacilliform Sri Lanka virus Cacao mild mosaic virus Cacao swollen shoot CD virus Cacao swollen shoot CE virus Cacao swollen shoot Ghana M virus Cacao swollen shoot Ghana N virus Cacao swollen shoot Ghana Q virus Cacao swollen shoot Togo A virus Cacao swollen shoot Togo B virus Cacao yellow vein banding virus Canna yellow mottle associated virus Canna yellow mottle virus

Aglaonema modestum Musa spp. Musa spp.

Natural hosts

USA All banana-growing countries Yes Australia, China, Cuba, Guadeloupe, Indonesia, Yes Kenya, Uganda All banana-growing countries Yes All banana-growing countries Yes Kenya, Uganda Uganda Uganda Uganda China, Viet Nam Finland, Germany

MH384837 AY493509 HQ593112

Acronym Accession # of exemplar isolate

Aglaonema bacilliform virus ABV Banana streak GF virus BSGFV Banana streak IM virus BSIMV

Species

Table 1 List of species, tentative species and unassigned viruses in the genus Badnavirus, family Caulimoviridae. Their acronym, accession # of an exemplar, natural hosts, geographical distribution, and their capacity to be endogenous are also indicated

Badnaviruses (Caulimoviridae) 159

Acronym Accession # of exemplar isolate

MK048221

X52938

KX008573

KY827395

KY827394

KX008574

KX008577

DQ822073

KX430257

JF411989 JQ316114

MF781082 HG940503

JF301669 KX852476

AY180137 LN651258 GU121676

EU377672

KC808712

KM078034

MN850490 AF299074

CoVCV

CoYMV

DBALV

DBALV2

DBESV

DBRTV1

DBRTV2

DBSNV

DBTRV

FBV1 GVBaV

GBV 1 GRLDaV

GVCV JuMaV

KTSV MBV1 PBCOV

PBERV

PYMoV

RYNV

Codonopsis vein clearing virus Commelina yellow mottle virus Dioscorea bacilliform AL virus Dioscorea bacilliform AL virus 2 Dioscorea bacilliform ES virus Dioscorea bacilliform RT virus 1 Dioscorea bacilliform RT virus 2 Dioscorea bacilliform SN virus Dioscorea bacilliform TR virus Fig badnavirus 1 Gooseberry vein banding associated virus Grapevine badnavirus 1 Grapevine Roditis leaf discoloration-associated virus Grapevine vein clearing virus Jujube mosaic-associated virus Kalanchoe top-spotting virus Mulberry badnavirus 1 Pineapple bacilliform CO virus Pineapple bacilliform ER virus Piper yellow mottle virus

Rubus yellow net virus

SRV Schefflera ringspot virusa Spirea yellow leafspot virus SYLSV

Continued

Species

Table 1

Nigeria Nigeria, Samoa Benin French Guyana, Guadeloupe USA Bosnia and Herzegovina, Canada, Czech Republic, The Netherlands, United Kingdom, Croatia Croatia, Iran, Italy, Greece, Turkey

USA China USA Lebanon, Iran All pineapple-growing countries Australia China, India, Indonesia, Sri Lanka, Thailand, The Yes Philippines, Viet Nam

Dioscorea rotundata Dioscorea rotundata Dioscorea sansibarensis Dioscorea alata, Dioscorea trifida Ficus carica Riber nigrum, Ribes rubrum, Ribes uva-crispa

Vitis vinifera Ziziphus jujube Kalanchoë blossfeldiana Morus alba Ananas comosus var. comosus Ananas comosus var. erectifolius Piper argyrophyllum, Piper attenuatum, Piper barberi, Piper betle, Piper colubrinum, Piper galeatum, Piper longum, Piper ornatum, Piper nigrum, Piper sarmentosum, Piper trichostachyon Rubus idaeus Schefflera sp. Spirea spp.

Yes

Fiji, Nigeria

Dioscorea dumetorum, Dioscorea esculenta

Canada, Bosnia and Herzegovina, Finland, United Kingdom, USA USA USA

Papua New Guinea

Vitis vinifera Ficus carica

Brazil, Nigeria, Tonga, Vanuatu

Dioscorea alata, Dioscorea bulbifera, Dioscorea cayennensis, Dioscorea esculenta, Dioscorea rotundata, Dioscorea transversa, Dioscorea trifida Dioscorea alata

Yes

Guadeloupe

Commelina diffusa

Endogenous?

South Korea

Geographical distribution

Codonopsis lanceolata

Natural hosts

160 Badnaviruses (Caulimoviridae)

China, India, Morocco Cameroon, China, Peru, South Korea, South Africa, Spain, Tanzania China, Ethiopia, Kenya, Tanzania, Uganda, USA Australia, Fiji, French Polynesia, Kenya, New Caledonia, Papua New Guinea, Tanzania, Uganda, Vanuatu China South Korea USA USA Nigeria

Saccharum officinarum Ipomoea batatas

SCBMOV M89923

KX168422 KM229702 MN542417 EU853709 MF476845

DQ473478 MH396440

SPPV

TaBCHV TaBV

WBV1 YNMoV CLGV CLNV DBRTV3

DrMV EpMoaV

Taro bacilliform CH virus Taro bacilliform virus

Wisteria badnavirus 1 Yacon necrotic mottle virus Camelia lemon glow virus Cycad leaf necrosis virus Dioscorea bacilliform RT virus 3 Dracaena mottle virus Epiphyllum mottleassociated virus (syn. Pitaya badnavirus 1) Green Sichuan pepper vein clearing-associated virus Ivy ringspot-associated virus (syn. Schefflera ringspot virus)a Pagoda yellow mosaic associated virus Pelargonium vein banding virus Polyscias mosaic virus Tentative Species in the Genus South Africa, USA

China USA USA, Nigeria Geographical distribution

USA Japan Australia, China, Guadeloupe, Kenya, Mauritius China, France, Italy, USA China Ethiopia

Hedera helix, Schefflera sp. Styphnolobium japonicum Pelargonium x hortorum Polyscias fruticosa Natural hosts Agave tequilana Aucuba japonica Musa spp., Saccharum officinarum Castanea crenata x C. sativa, Castanea dentata, Castanea mollissima, Castanea sativa Dioscorea composita Ensete ventricosum

MN850490

KJ013302

GQ428155

IRSaV

PYMaV

PVBV

Endogenous?

Yes

Only a partial genome sequence is available for Schefflera ringspot virus (MH475920) and this sequence is 99.1 identical to the genome sequence of Ivy ringspot-associated virus (MN850490), suggesting the two species are synonymous.

a

PoMV MH475918 Acronym Accession # of exemplar isolate Agave badnavirus A ABVA MH898467 Aucuba ringspot virus AuRSV LC487411 Banana streak CA virus BSCAV HQ593111 Chestnut mosaic virus ChMV MT269853 Dioscorea bacilliform virus A DBVA MH898490 Enset leaf streak virus ELSV MF991909

China

Zanthoxylum schinifolium

Dracaena sanderiana China Discocactus spp., Epiphyllum spp. Hylocereus polyrhizus, Opuntia spp. Schlumbergera China, USA spp.

Wisteria sinensis Smallanthus sonchifolius Camellia japonica Zamia fischeri Dioscorea rotundata

GSPVCaV MK371353

KP710178 AF357836

FJ560943 Colocasia esculenta, Xanthosoma sp. Colocasia esculenta, Xanthosoma sp.

Australia, China, India

Saccharum officinarum

AJ277091

SCBIMV

Guadeloupe, China

Saccharum officinarum

FJ439817

SCBGDV

Guadeloupe, China

Saccharum officinarum

FJ824813

SCBGAV

Sugarcane bacilliform Guadeloupe A virus Sugarcane bacilliform Guadeloupe D virus Sugarcane bacilliform IM virus Sugarcane bacilliform MO virus Sweet potato pakakuy virus

Badnaviruses (Caulimoviridae) 161

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Badnaviruses (Caulimoviridae)

Fig. 1 Phylogram depicting relationships within the genus Badnavirus, based on a maximum likelihood analysis of a pol gene sequence alignment using the IQ-TREE web server. The pol gene sequence alignment of approved badnavirus species was downloaded from the ICTV Caulimoviridae Resources Page and amended with sequences of tentative badnavirus species that are available on GenBank. GenBank accession codes and virus acronyms (see Table 1) are provided at the tips of the branches; rice tungro bacilliform (RTBV) is the outgroup. Ultrafast bootstrap support values (%) from 1000 replicates are shown in the nodes of the branches (only values 470% are shown).

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Fig. 2 Virions of Banana streak IM virus (BSIMV), negatively stained with 1% ammonium molybdate pH 7. Micrograph courtesy of K.S. Crew.

Virion Structure Virions are bacilliform with parallel sides and rounded ends, and modal dimensions of 30 nm  130 nm (Fig. 1). However, virions that are two to three times this modal length are sometimes observed, and these contain non-covalently bonded concatemers of viral genomic DNA that are up to four monomeric units in length. Two types of virion are observed in mulberry plants infected by Mulberry badnavirus 1 (MBV1), one that has normal dimensions and a second that is thinner (c. 20 nm diameter) and has a looser capsid structure. These aberrant virions contain a shorter form of the MBV1 genome. The main structural protein of the virion is the capsid protein (CP), while a second protein, the virion-associated protein (VAP), is thought to decorate the surface of the virion by occupying the pores between the capsomers. At the C-terminus of the VAP, there is a proline-rich region that binds to double-stranded DNA in a sequence non-specific manner, and putatively interacts with the encapsidated DNA. Conversely, the N-terminus of the VAP is thought to be surface-exposed and contains a coiled-coil motif that could interact with other viral, host or insect proteins to mediate functions in the virus’ life cycle such as cell-to-cell movement and vector transmission (Fig. 2).

Genome The encapsidated form of the badnaviral genome is a non-covalently closed dsDNA molecule of 7.0–9.3 kbp. There are discontinuities in each strand of DNA, which arise as a consequence of synthesis of the DNA strands by reverse transcription. Synthesis of the negative strand of DNA is primed by a cytosolic initiator methionine tRNA (tRNAmet), and the discontinuity on this strand is adjacent to the 30 end of the homology to the tRNAmet. On the positive strand, the discontinuity is located within a polypurine-rich region within ORF3. By convention, position 1 of the genome is designated the first nucleotide of the tRNAmet binding motif (50 T1GGTATCAG…. 30 ) on the positive strand. The majority of badnavirus genomes have three conserved open reading frames (ORFs), which are in different translational reading frames (Fig. 3). For the most part, the start codons for these ORFs are AUG but for some species, non-conventional start codons are utilized, particularly at the beginning of ORF1 (e.g., CUG for BSMYV ORF1). Other ORFs have been described but these are not found in all species and evidence that these ORFs are translated is lacking. It is typical for the stop codons of ORFs 1 and 2 to overlap the start codons of the succeeding ORFs. ORF1 encodes a small protein of c. 21–25 kDa that has no assigned function but is recognizable through the presence of a conserved domain, pfam07028. ORF2 also encodes a small protein of c. 15 kDa, the VAP. ORF3 encodes a large polyprotein of c. 197–232 kDa, with conserved movement protein (MP), capsid protein (CP) and nucleocapsid, aspartic protease, reverse transcriptase and RNase H domains in that order. All domains in the polyprotein bar the MP have strong sequence conservation at primary and secondary structural levels with paralogous proteins of retroviruses and long terminal repeat retrotransposons, and are classified in the same protein families. Two variants of the typical badnavirus genome organization are known. MBV1 has a genome sequence with one long ORF, which appears to be the equivalent of a fusion of ORFs 1 and 3 in other badnaviruses based on the arrangement of conserved domains. The genome sequence of sweet potato pakakuy virus (SPPV), which contains four ORFs, represents yet another permutation of the typical badnavirus genome. ORF3 encodes the MP and CP, while ORF4 encodes the aspartic protease and reverse transcriptase/ribonuclease H1 proteins. Hence for SPPV, ORF3 in other badnaviruses is split in two. The intergenic region in the genome between the end of ORF3 and the beginning of ORF1 in the typical badnavirus genome is long and contains transcription regulatory motifs. Between the end of ORF3 and the tRNAmet motif, there is the equivalent of the cauliflower mosaic virus 35S promoter, identifiable by the presence of a TATA box. Due to prospects of using the badnavirus promoter for genetic engineering of plants, there has been considerable work done to identify the minimal promoter sequences and to identify cis-acting elements that control the specificity of tissue expression. Downstream of the promoter is a polyadenylation signal but only a small percentage of the RNA transcripts have a poly(A) tail.

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Fig. 3 Genome map for Commelina yellow mottle virus (ComYMV), type species of the genus Badnavirus. The black circle represents the genomic DNA, the purple circle is the pregenomic RNA and the gray curved boxes are the open reading frames. Marked on the genomic DNA are the positions of the TATA box (black diamond), polyadenylation signal (black arrow) and tRNAmet-binding site (black clover). The blue, green, red, orange and yellow curved boxes represent the movement protein, capside protein, aspartic protease, reverse transcriptase and RNaseHI domains on ORF 3, respectively.

Life Cycle The replication cycle of badnaviruses has not been investigated to any great extent, and most knowledge is based on extrapolation from model plant viruses such as cauliflower mosaic virus (CaMV), and to a lesser extent, rice tungro bacilliform virus (RTBV). Following entry of the badnavirus virion into the cell, the encapsidated viral DNA needs to be transported to the nucleus for replication to begin. A putative nuclear localization signal (NLS) is present in the basic, intrinsically disordered region of the NC domain of the CP. For RTBV, this NLS facilitates interaction between the CP and importin a but as the NLS is within the mature virion, is unlikely to play a role in docking of the virion to the nuclear pore but rather entry of the genomic DNA after either partial of complete virion decapsidation. There are two major CP isoforms that vary in length of the acidic N-terminus of the protein. It is not clear whether this variation in length is due to the presence of alternative aspartic protease cleavage sites or is a consequence of targeted degradation of the protein. With CaMV, the extended N-terminus inhibits nuclear targeting of the protein, ensuring that it is localized within the cytoplasm where virion assembly occurs. Once the genomic DNA enters the nucleus, it is assumed that replication follows the pathway of CaMV, whereby the discontinuities in the genomic DNA are repaired to form a covalently closed circle, and the DNA then associates with histone proteins and RNA polymerase II to form transcriptionally active complexes called minichromosomes. A single, terminally redundant, slightly greater than genome-length RNA transcript is produced called the pregenomic RNA (pgRNA), and there is no evidence of subgenomic or spliced RNA transcripts. Following the replication model proposed for CaMV, it is thought that the RNA polymerase II enzyme begins transcription, ignores the polyadenylation signal on the first pass, continues transcribing around the circle and drops off the DNA template when it reaches the polyadenylation signal for the second time. Hence, the region between the transcription start site and the polyadenylation signal is duplicated at the 50 and 30 ends of the pgRNA. It has been a mystery as to where in the cell, virion assembly occurs but recent studies suggest that badnaviruses produce viroplasms (virus factories) that are equivalent in function to those of CaMV but clearly different in composition because badnaviruses lack the major viroplasm matrix protein produced by caulimoviruses. These putative viroplasms have a characteristic structure comprising an undifferentiated core, an encircling lacuna where reverse transcription may occur, and an outside ring of differentiating virions. A gap is present in the outside ring, allowing the import of newly synthesized viral proteins from the surrounding ribosomes. The pgRNA, which also acts as the messenger RNA, is polycistronic and the downstream ORFs are likely translated by a leaky ribosome scanning mechanism, whereby the start codons of ORFs 1 and 2 are in sub-optimal initiation contexts and are bypassed by the ribosome on a fraction of occasions in favor of the next downstream start codon. Consistent with this translation mechanism, internal AUG codons are rare to absent in ORFs 1 and 2. The 50 leader sequence of the pgRNA is long and folds into a stable hairpin structure, which is immediately preceded by a short upstream ORF (uORF). It is proposed that translation initiates at the start of the uORF and the ribosome then shunts across the hairpin structure to a landing site near the beginning of ORF1, where translation recommences. The uORF could regulate sorting of the ribosomes between the three ORFs to maintain balanced production of the different proteins.

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The large polyprotein encoded by ORF3 must be post-translationally processed to release each individual component protein. Within the polyprotein is an aspartic protease, which belongs to the retrovirus aspartic protease family (pfam00077) and has a highly conserved active site (motif Asp-Ser/Thr-Gly). It is hypothesized that the aspartic protease autocleaves and then processes other proteins in the polyprotein. Based on studies with retroviruses, the active form of the protease is likely to be a ß homodimer. Substrate sites for the enzyme are not strictly conserved at the amino acid sequence level and cleavage sites must be empirically determined. No badnaviral aspartic protease has been biochemically characterized and there has only been one attempt to map a cleavage site, at the N-terminus of the BSMYV capsid protein. The final step in the badnavirus replication cycle, reverse transcription, is thought to follow the general model for CaMV. The (  ) strand of DNA is first synthesized using the pgRNA as the template, the virus-encoded reverse transcriptase as catalyst and a tRNAmet molecule to prime the reaction. When the reverse transcriptase reaches the 50 end of the pgRNA, the nascent end of the DNA melts away from the RNA and anneals to the identical stretch of sequence at the 30 end of the pgRNA. Reverse transcription then continues around the circle until the original tRNAmet-binding site is encountered. At some point, the pgRNA and tRNAmet molecule must be removed, presumably through the action of the virus-encoded RNaseH enzyme. Synthesis of the ( þ ) strand of DNA then begins, presumably using a host-encoded DNA polymerase and a primer that has yet to be characterized. Transmission electron microcopy studies suggest that badnaviruses are trafficked between cells as whole virions and this is enabled by the formation of a tubule that displaces the contents of the plasmadesmatum and increases its size exclusion limit. Badnaviruses produce an example of the 30K superfamily of viral MP, which is most closely related to those produced by nepoand idaeoviruses.

Epidemiology The epidemiology of many of the badnaviruses is poorly understood, and most knowledge derives from the study of a small number of diseases with the largest economic impacts, such as cacao swollen shoot and banana streak disease. The majority of badnavirus species are transmitted by mealybugs (family Pseudococcidae), which are most abundant in warm, moist climates or in greenhouses. Mealybugs inhabit protected sites on the plant such as the underside of plant leaves and stems, in leaf axiils and the underneath leaf sheaths, and on the roots, and hence their presence may be overlooked. Where known, the virus-vector specificity of mealybug transmission is weak, although there is very little data on this topic due to the difficulties of rearing mealybugs and doing controlled transmission experiments. At least fourteen species of mealybug are capable of transmitting the cacao swollen shoot complex of viruses, although three species are considered the most important vectors, namely Formicococcus njalensis, Planococcus citri and Ferrisia virgata. The most mobile growth stages of the mealybug are the first and second instars of nymph, and these are also known as ‘crawlers’. Crawlers move over most parts of the plant, whereas older nymphs and adults will retreat to more protected positions on the plant. Transmission of badnaviruses can occur when either the crawlers actively cross leaf bridges between plants or are passively transported by the wind or carried by attendant ants. Ants have a strong mutualistic relationship with mealybugs, as they feed on their honeydew secretions and in turn, provide protection from parasites and predators and improve hygiene by cleaning up waste products. Mealybugs can be indirectly controlled by reducing ant populations using insecticides. The second most important group of badnavirus vectors is aphids, and again these transmit the viruses in the semi-persistent manner. Only five species of badnavirus are known to be transmitted by aphids, namely chestnut mosaic virus (ChMV), gooseberry vein-banding associated virus (GVBaV), Rubus yellow net virus (RYNV), Spirea yellow leaf spot virus (SYLSV) and grapevine vein clearing virus (GVCV). All four viruses are closely related, and all originate from temperate regions of the Northern Hemisphere. Evolution of this novel virus-vector relationship may reflect the greater abundance of aphids in higher latitudes of Eurasia and North America relative to mealybugs. The only other type of insect that can vector badnaviruses is the black pepper lace bug (Diconocoris distanti), which together with the citrus mealybug (Planococcus citri), is reported to transmit Piper yellow mottle virus (PYMoV); but this finding does need independent verification. True seed transmission has been reported for several badnavirus species including PYMoV in black pepper, Kalanchoe top spotting virus (KTSV) in Kalanchoë blossfeldiana, SPPV in Ipomoea batatas and an unidentified species of taro bacilliform virus in Colocasia esculenta. A report that BSMYV is seed-transmitted in banana is ambiguous as there is the possibility that the infections in seedlings originated from activation of endogenous badnaviral elements. While seed transmission of many badnaviruses is theoretically possible, it is likely to be epidemiological insignificant as many of the host species are vegetatively propagated. Where known, most badnaviruses have narrow host ranges in nature and alternative hosts are unlikely to play an important role in the virus epidemics but there are important exceptions. It is well documented that the indigenous rainforest trees in West Africa are the origins of the badnaviruses associated with cacao swollen shoot disease. When the rainforest was cleared for cropping, some trees were left to provide shade for the cacao plants, and these acted as primary sources of inoculum for the epidemics. The most important alternative host trees for cacao swollen shoot in Ghana are Cola gigentia, Cola chlymydantha and Ceiba pentandra, all from the same Malvaceae family as Theobroma cacao. However, pockets of older, infected cacao trees in the same or neighboring plantations are now the most important sources of inoculum and disease control is dependent on operation of an effective roguing program.

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Other examples of cross-species transmission of badnaviruses include banana streak CA virus, which as the name suggests, was first described from banana but appears to be more common in sugarcane. These two crops grow on the same types of soil, and often on the same or neighboring farms, and it is thought that infections in banana originate from nearby sugarcane plants. Canna yellow mottle virus (CaYMV), a virus that is ubiquitous in ornamental canna plants in the tropics, is responsible for diseases in betel nut (Piper betel) in Pakistan and ornamental ginger (Alpinia purpurata) in Hawaii. Finally, wild members of the Vitaceae family such as Vitis rupestris and Ampelopsis cordata are important alternative hosts of GVCV, which causes grapevine decline in the American Midwest. The frequency of cross-species transmission of these viruses is unknown but does not need to be common for the infections to proliferate as all crops are vegetatively propagated.

Clinical Features Badnaviruses typically produce chlorotic streaks, mild mottles and sometimes ringspot patterns on the leaves of monocots, and mottle and mosaic patterns, vein clearing and ring spots on the leaves of dicots (Fig. 4). Plants affected by cacao swollen shoot disease have distinctive protuberances on the stems, caused by proliferation of the xylem, phloem and cortex cells. Plants infected by badnaviruses may be asymptomatic or symptoms sporadic in occurrence, as best documented with the various species of the banana streak disease complex (BSD). The factors triggering symptom expression in banana are poorly understood but there is a clear trend of symptom severity increasing with maturity of the plant, reaching a maximum at about the time of bunch initiation. For BSD, virus titer is also positively correlated with symptom severity but infection often leads to cell death, causing the chlorotic streaks to turn necrotic, and virus titer to decline again.

Fig. 4 Disease symptoms caused by badnaviruses: A, Musa sp. AAB group infected by banana streak MY virus (BSMYV) (photograph courtesy of A.D.W. Geering); B, Betula pubescens infected by birch leaf roll-associated virus (BLRaV) (photograph courtesy of A. Rumbou); C, Hedera helix infected by ivy ringspot-associated virus (IRSaV) (photograph courtesy of J.T. Burger); D, Vitis vinifera cv. Cabernet Franc infected by Grapevine vein clearing virus (GVCV) (photograph courtesy of W. Qiu); E, Citrus x sinensis infected by citrus yellow mosaic virus (CiYMV) (photograph courtesy of V.S. Baranwal); F, Aucuba japonica infected by Aucuba ringspot virus (AuRSV) (photograph courtesy of A. Uke).

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167

Pathogenesis There is scant information on how badnaviruses cause disease. The tissue tropism of badnaviruses is variable and dependent on the virus in question. Rubus yellow net virus infects all cell types of the raspberry leaf including the epidermis but is most common in the phloem companion cells. BSMYV most commonly occurs in the palisade and spongy mesophyll cells and rarely in the epidermal cells of the banana leaf. Cacao swollen shoot Togo B virus (CSSTBV) appears to be mainly vascular-limited, with most virions observed in the phloem companion cells and far less commonly in the xylem parenchyma cells. Cytopathological studies suggest that badnaviruses trigger cell apoptosis although further studies are required to confirm these observations. Older infected cells exhibit severe plasmolysis, accumulate vacuoles, the chloroplasts become deformed, starch granules aggregate and the cells eventually lose all contents. The putative viroplasms can reach relatively large diameters and are easily visible by light microscopy using stains such as periodic acid-Schiff-toluidine blue. One peculiar aspect of the biology of badnaviruses, particularly among plant-infecting viruses, is that new infections can arise following activation of viral sequences that are integrated in the nuclear genome of the host plant. These integrated sequences are commonly referred to as endogenous viral elements. Badnaviruses do not actively integrate into their host genome as do retroviruses but it is thought that the viral DNA is captured as a type of filler sequence during non-homologous or microhomologymediated end-joining repair of double-stranded DNA breaks in the chromosomal DNA. Integration of viral DNA is part of a broader phenomenon of horizontal gene transfer involving all types of DNA, such as that from the plastid genomes. The problem of endogenous badnaviral elements causing new infections was discovered in a quest to understand why new hybrid bananas generated by banana breeding programs around the world became infected with banana streak OL virus (BSOLV), despite the fact that the parent plants used to create the hybrids were healthy. A locus was discovered in one parent, cv. ‘Obino l0 Ewai’ (Musa AAB group), which contains the complete genome sequence of BSOLV but divided in two fragments separated by a large scrambled region of fragments of non-contiguous and sometimes inverted viral sequence. An activation model was proposed involving two homologous recombinations, the first to excise the scrambled region and the second to join either end of the locus to form a circular, transcriptionally active form of the genome. This locus has now been better characterized and shown to be diallelic, with only one allele being infective. Activation is not spontaneous but triggered by external factors such as tissue culture propagation and interspecific hybridization. It is hypothesized that in a normal state, the endogenous badnaviral elements are transcriptionally silenced but certain stresses cause the transient release of this silencing. The phenomenon of activation of endogenous badnaviral elements has now been extended to at least two more badnaviruses, banana streak GF virus (BSGFV) and banana streak IM virus (BSIMV) and as with BSOLV, activation involves recombinations to reconstitute complete viral genomes. All aforementioned replication-competent endogenous banana streak viruses are linked to the B genome of banana, which derives from Musa balbisiana. The A genome of banana (from Musa acuminata) also contains endogenous badnaviral elements but these are derived from different ancestral badnavirus species and all are regarded as being replication-defective due to the accumulation of mutations, rearrangements and deletions. Endogenous badnaviral elements have now been detected in an broad range of plant species, both monocots and dicots. While it is not particularly surprising that badnaviral DNA is captured in the host genome, a more perplexing question is why the DNA has become fixed in the genome and has persisted for hundreds of thousands, even millions of years. It is generally assumed that the endogenous badnaviral elements confer a selective advantage to the plant, such as imparting resistance against the cognate infectious virus through activation of RNAi defense pathways. However, it is likely that the endogenous badnaviral elements could also help shape the transcriptome by, for example, contributing novel promoter elements or being a source of small non-coding RNAs. Furthermore, it is conceivable that badnaviral proteins are generated that may have been coopted for some other function in normal plant metabolism.

Diagnosis Badnaviruses vary considerably in the difficulty of diagnosis, depending on the presence of endogenous badnaviral elements in the plant, as there is a need to distinguish these from encapsidated (exogenous) viral DNA. This problem is best exemplified by the various species of banana streak disease. When describing a new species of badnavirus, it is also essential to demonstrate that the DNA sequence derives from exogenous viral DNA. The nuclear genome of cultivated banana contains multiple copies of endogenous badnaviral DNA, representing a large range of ancestral virus species. However, dessert banana such as the Cavendish subgroup of cultivars have an AAA genome, which is devoid of endogenous forms of Banana streak OL virus (BSOLV), Banana streak GF virus (BSGFV), Banana streak IM virus (BSIMV) and Banana streak MY virus (BSMYV). Hence, conventional PCR using species-specific PCR primers and total plant DNA extracts can be used for diagnosis. However, for hybrid plantain or dessert banana cultivars with AAB or ABB genotypes, conventional PCR cannot be used because of the problem of endogenous viral DNA giving false positive diagnostic results. Two diagnostic technologies are routinely used to distinguish endogenous from exogenous viral DNA, these being immunocapture PCR and rolling circle amplification. The first step of immunocapture PCR is to trap virions on the sides of a PCR tube, and then to wash away other cellular components including plant nucleic acids. While this step does enrich for virions, plant DNA can bind directly to the tube wall and precautions need to be taken to prevent this from happening. Non-specific binding of DNA to the tube wall is a much slower process than the antibody-virion interaction, and limiting the immunocapture step to 3 h at room temperature helps prevent amplification of endogenous badnaviral elements. To alert the diagnostician of direct binding of

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plant DNA, duplex PCRs can be done using primers against repetitive DNA sequences in the plant such as microsatellites. Another option is to do a DNase I digestion step after the immunocapture step, as the capsid shell protects the encapsidated DNA from nuclease degradation. One of the major constraints to the wider adoption of immunocapture PCR for diagnosis is the lack of availability of antibodies against many of the viruses but use of antipeptide antibodies shows promise for rapidly resolving this problem. Some crop species such and banana, sugarcane and yam, are infected by a broad diversity of badnaviruses and universal badnavirus-group primers have been designed to cover most combinations of virus species. Rolling circle amplification (RCA) using Phi29 DNA polymerase is a very useful technique for characterizing badnaviral genomes and for the most part, the selected method has been to use a commercial RCA kit (TempliPhi™) that has been spiked with a mixture of degenerate and specific badnavirus primers to increase the sensitivity of detection. It is only the minichromosomal fraction of viral DNA that is amplified by RCA, as the discontinuities present in the encapsidated DNA cannot be bridged by the DNA polymerase. TempliPhi™ is a sequence independent amplification method, as DNA polymerization is primed with random hexamers, and this has sometimes created problems with interpreting results, as circular plant plastid DNA sequences may also be amplified. However, major improvements to the specificity of amplification by RCA have been made by formulating reaction mixtures from individual components, replacing the random hexamers with a panel of degenerate badnavirus primers, and optimizing amplification temperatures.

Prevention The two foundations of badnavirus prevention are the use of virus-indexed planting material and the operation of disease roguing programs. Virus indexing is particularly important for crops such as banana, which are propagated by tissue culture and a single meristem can give rise to 500 progeny plants. For chronically-infected, vegetatively propagated plants, somatic embryogenesis and meristem-tip culture can be used to eliminate virus, and the success of the technique is enhanced by combining with chemotherapy. The use of insecticides to control mealybug vectors is generally considered to be ineffective due to the waxy, water-repellant coatings on the insect and the cryptic places where the insects feed but there may be a place for systemic insecticides in a management program. A problem exists with hybrid (A x B genome) bananas and plantains as the practice of tissue culture in itself provides the trigger for activation of endogenous badnaviral elements and this is particularly a concern for the banana breeding programs as embryo rescue is integral to the breeding process. PCR diagnostic assays have now been developed to screen for replication-competent alleles of the endogenous banana streak viruses, and use of these diagnostic assays allows identification of cultivars that are particularly prone to becoming infected when propagated by tissue culture. The long term solution to the issue of endogenous badnaviral elements causing infection is to create plants that are devoid of replication-competent endogenous virus alleles by conventional breeding techniques or by inactivating these alleles using gene-editing techniques such as CRISPR/Cas9. Cacao swollen shoot is undoubtedly the most economically important disease caused by badnaviruses, and no plant disease resistance is available and therefore the primary control strategy has been roguing, resulting in the destruction of over 200 million trees in Ghana since 1946. Mild strain cross protection does not protect plants from superinfection by virulent strains but does slow the rate of spread by nearly half. Oil palm and citrus barriers can also be used to slow the reintroduction of cacao swollen shoot into newly planted, healthy crops and at the same time provide an alternative source of income for the farmers.

Further Reading Bhat, A.I., Hohn, T., Selvarajan, R., 2016. Badnaviruses: The current global scenario. Viruses 8, 177. Chabannes, M., Baurens, F.-C., Duroy, P.-O., et al., 2013. Three infectious viral species lying in wait in the banana genome. Journal of Virology 87, 8624–8637. Cheng, C.P., Tzafrir, I., Lockhart, B.E., Olszewski, N.E., 1998. Tubules containing virions are present in plant tissues infected with Commelina yellow mottle virus. Journal of General Virology 79, 925–929. Diop, S.I., Geering, A.D.W., Alfama-Depauw, F., et al., 2018. Tracheophyte genomes keep track of the deep evolution of the Caulimoviridae. Scientific Reports 8, 572. Krupovic, M., Blomberg, J., Coffin, J.M., et al., 2018. Ortervirales: New virus order unifying five families of reverse-transcribing viruses. Journal of Virology 92, e00515-18. Medberry, S.L., Lockhart, B.E., Olszewski, N.E., 1990. Properties of Commelina yellow mottle virus's complete DNA sequence, genomic discontinuities and transcript suggest that it is a pararetrovirus. Nucleic Acids Research 18, 5505–5513. Ngo, T.H., Webb, R., Crew, K.S., et al., 2020. Identification of putative viroplasms within banana cells infected by banana streak MY virus. Journal of General Virology. doi:10.1099/jgv.0.001498. Pooggin, M.M., Ryabova, L.A., 2018. Ribosome shunting, polycistronic translation, and evasion of antiviral defenses in plant pararetroviruses and beyond. Frontiers in Microbiology 9, 644. Sukal, A.C., Kidanemariam, D.B., Dale, J.L., Harding, R.M., James, A.P., 2019. Assessment and optimization of rolling circle amplification protocols for the detection and characterization of badnaviruses. Virology 529, 73–80. Teycheney, P.-Y., Geering, A.D.W., Dasgupta, I., et al., 2020. ICTV virus taxonomy profile: Caulimoviridae. Journal of General Virology 101, 1025–1026. Vo, J.N., Campbell, P.R., Mahfuz, N.N., et al., 2016. Characterization of the banana streak virus capsid protein and mapping of the immune-dominant continuous B-cell epitopes to the surface-exposed N terminus. Journal of General Virology 97, 3446–3457.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/reverse-transcribing-dna-and-rna-viruses/w/caulimoviridae Caulimoviridae. Reverse Transcribing DNA and RNA Viruses.

Banana Bunchy Top Virus (Nanoviridae) John E Thomas, The University of Queensland, Brisbane, QLD, Australia r 2021 Elsevier Ltd. All rights reserved. This is an update J.E. Thomas, Banana Bunchy Top Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00636-1.

Glossary Cell-cycle link protein A plant virus protein which most probably subverts the cell cycle control of the host, forcing cells into DNA synthesis or S phase favorable to viral replication. Circulative transmission Mode of transmission whereby the virus is internally borne in the insect vector but does not

replicate there. It is characterized by a latent period before which transmission cannot occur, and by retention of infectivity by the vector for a period of at least weeks. Nuclear shuttle protein A virus-encoded protein that transports the viral single-stranded DNA, as a complex with the viral movement protein, to and from its site of replication in the nucleus and into adjacent cells.

Introduction Banana bunchy top disease is the most economically important virus disease of banana and plantain (Musa spp.) worldwide, due to its devastating effect on crop yield, and the importance of banana and plantain as both a staple food and a major export commodity in much of the developing world. The causal agent is Banana bunchy top virus (BBTV), which has a multipartite circular ssDNA genome, encapsidated in small isometric virions, and is transmitted by the banana aphid, Pentalonia nigronervosa. Edible bananas are derived from wild seeded progenitors including predominantly subspecies of M. acuminata and M. balbisiana, which have a center of origin in South and South-East Asian-Australasian region. It is likely that BBTV and the aphid vector also originated within this area.

Taxonomy, Phylogeny, and Evolution BBTV is a member of the genus Babuvirus, in the family Nanoviridae. Recently, two new virus species have been identified which are clearly distinct members of the genus Babuvirus (Fig. 1). Abaca bunchy top virus, which also infects Musa in South-East Asia, and Cardamom bushy dwarf virus causing Foorkey disease of cardamom in India, both have homologous components for all six genome components of BBTV. Additional species in this family are classified in the genus Nanovirus, and include Faba bean necrotic yellows virus, Milk vetch dwarf virus Subterranean clover stunt virus, Black medic leaf roll virus, Faba bean necrotic stunt virus, Faba bean yellow leaf virus, Pea necrotic yellow dwarf virus and Pea yellow stunt virus. Members of this family are phylogenetically related and share a similar particle morphology and multi-component circular, ssDNA genome. However, members of the two genera are not serologically related. Although both genera possess five components vital to replication and spread in common, some additional components present in one genus do not have a homologous component in the other genus. However, some of these components encode proteins on unknown function, and these unrelated proteins may share similar functions.

Fig. 1 Phylogenetic tree based on the amino acid sequences of babuvirus Rep proteins, using the nanovirus Faba bean necrotic yellows virus as an outgroup. The South-East Asian and Pacific\Indian Ocean subgroups of BBTV, with representative isolates, are circled in blue and red, respectively.

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Fig. 2 Electron micrograph of BBTV virions, negatively contrasted with 1% ammonium molybdate.

BBTV isolates worldwide fall into two broad phylogenetic groups (Fig. 1), called the South Pacific (or Pacific/Indian Ocean) group (isolates from Australia, the Pacific islands the Indian sub-continent, Myanmar, Egypt and Sub-Saharan Africa) and the Asian (or South-East Asian) group (China, Philippines, Japan, Vietnam, Indonesia, Laos, Thailand and Taiwan). Sequence differences between the two groups are shared across all genome components. Within group sequence diversity is much greater for the Asian group suggesting a longer evolutionary period and possible evolutionary origin for the virus. The coat protein gene of BBTV is highly conserved, with a maximum difference of 3% at the amino acid level between isolates. No serological differences have been detected between any isolates using polyclonal or monoclonal antibodies. Recombination and reassortment has been detected between the two groups, supporting the temporal co-existence of the groups at one time. The most recent common ancestor of these BBTV sequences has been dated as occurring 1086 years ago (95% HPD 812–1399 years). The presence of the virus in many countries from the South Pacific group can be traced to introductions within the last century.

Virion Structure The viral nature of bunchy top disease was established by CJP Magee in Australia in the 1920s. However, it was not until 1990 that the virus particles were first isolated, in part due to lack of a suitable herbaceous experimental host, its low titer in infected plants and its restriction to the phloem in the fibrous vascular tissue of Musa. Typical of the family Nanoviridae, BBTV has icosahedral particles, 18–20 nm in diameter with T ¼ 1 symmetry (Fig. 2), a buoyant density of 1.29–1.30 g/cm3 in cesium sulfate, and a sedimentation coefficient of 46S.

Genome Organization and Functions of Gene Products BBTV has a multi-component genome comprising six transcriptionally active components (Fig. 3, Table 1), each ca. 1 kb in size. DNA-R encodes two proteins, one the master replication initiation protein (Rep), which can initiate and terminate replication of DNA-R and the other genome components, and a second internal ORF of unknown function. All other components are monocistronic. DNA-S encodes the coat protein, demonstrated by N-terminal sequencing and the production of specific antibodies against the cloned and expressed gene. It also contributes to RNA silencing. The DNA-C gene product contains an LXCXE motif and binds to a retinoblastoma protein, indicating that it is a cell cycle link protein which stimulates viral DNA replication. It has also been linked to suppression of RNA silencing. DNA-M has been shown to be both a movement protein, and the major pathogenicity determinant and suppressor of RNA silencing. DNA-N encodes the nuclear shuttle protein, while the function of the protein encoded by DNA-U3 is unknown. The untranslated regions of all six components share two areas of extensive homology, both concerned with the rolling circle method by which BBTV replicates. A stem-loop common region (CR-SL) of 69 nt includes a nonanucleotide sequence (TATTATTAC) which is shared between plant-infecting circular ssDNA nanovirids and geminivirids, and which is the site of the origin of viral replication. It also contains three short repeated sequences (iterons F1, F2 and R1) which are the presumed binding site for the Rep. The major common region (CR-M) varies between 66 and 92 nt in length amongst the various components. Virions contain a heterogeneous population of DNA primers, around 80 nt in length, which bind to the CR-M and prime complementary strand synthesis. The relative concentrations of the genome components can vary enormously with DNA-S generally being the least abundant in plants, and DNA-N the highest, at 60 to 41660 times the copy number of DNA-S. The intergenic regions of all six integral components of BBTV have been shown to have promoter activity. The highest activity in banana embryonic cells was shown by promoters from DNA-C and DNA-M, components thought to be intrinsic to the infection process. Studies on the DNA-N promoter demonstrated that expression in banana embryonic cells is limited to phloem tissue,

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Fig. 3 Diagram of the integral genome components of BBTV, showing transcribed ORFs (black arrows) and other main genome features. From Fauquet, C.M., Mayo, M.A., Maniloff, J., Desselberger, U., Ball, L.A., 2005. Virus Taxonomy. Eighth Report of the International Committee on Taxonomy of Viruses. San Diego, CA: Elsevier Academic Press.

Table 1

Size and function of the six integral components of BBTV

Genome component

Size (nt)

Size of encoded protein(s) (kDa)

Function

DNA-R DNA-S DNA-M DNA-C DNA-N DNA-U3

1111 1075 1043 1018 1089 1060

(i) 33.6 (ii) 5.2 20.1 13.7 19.0 17.4 10.4

(i) Replication initiation protein (ii) Unknown Capsid protein Movement protein Cell-cycle link Nuclear shuttle protein unknown

consistent with the circulative mode of transmission by the aphid vector and observations that histological effects were confined to the phloem and phloem parenchyma cells. Circulative transmission by aphid vectors usually involves specific feeding on the phloem tissue for virus acquisition. The CR-M and CR-SL are not essential for promoter activity. All essential elements of the promoter are located 30 of the stem-loop and within 239 bp of the translation start codon and within this region are an ASF-1 motif (TGACG), a hexamer motif (ACGTCA), rbcS-I-box (GATAAG), G-boxcore and the TATA box, all associated with promoter activity in other genomes. Some isolates of BBTV, especially those from the South-East Asian group (see Taxonomy and Phylogenetic Relationships), contain additional Rep proteins called alphasatellites, similar in size to DNA-R, but capable of self-replication only. These molecules have a CR-SL, though the nonanucleotide sequence TAGTATTAC forming part of the loop is not conserved with the six integral DNA components. Unlike DNA-R, the alphasatellites lack the internal ORF, their TATA boxes are 50 of the stem loop and they generally lack the CR-M. Interestingly, the amino acid sequences of BBTV alphasatellites- are more closely related phylogenetically to the alphasatellites associated with viruses in the related genus Nanovirus than they are to DNA-R of BBTV. The biological function of the alphasatellites is uncertain.

Host Range and Transmission The symptoms of bunchy top disease in banana are characteristic, especially in the Cavendish subgroup of cultivars, and easily distinguished from all other virus diseases of banana. Plants can become infected at any stage of growth, and the initial appearance of symptoms can depend on the manner of infection. Plants that are aphid-inoculated usually develop first symptoms in the second or later leaves to emerge after inoculation, and there display a few dark green streaks and dots on the lower part of the lamina and on the petiole. These symptoms become more general on subsequent leaves. These streaks form hooks as they enter the midrib and are best viewed from the underside of the leaf, with transmitted light (Fig. 4). However, these dark streaks can be rare or absent in some cultivars. Successive leaves become shorter and narrower, and have a brittle lamina with upturned, chlorotic, ragged margins. Leaves fail to emerge fully, giving the plant a bunched appearance (Fig. 5). Plants derived from infected planting material (suckers, bits) develop severe symptoms from the first leaf to emerge.

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DNA-R 1111 nt

DNA-S 1075 nt

DNA-M 1043 nt

DNA-C 1018 nt

DNA-N 1089 nt

DNA-U3 1060 nt

: Common stem-loop region

: TATA box

: Major common region

: Poly(A) signal

Fig. 4 Dark green dot-dash, hooking and vein clearing fleck symptoms on a Cavendish banana leaf.

Infected plants seldom produce a bunch, though if infected late in the current cropping cycle, a small, distorted bunch may result. With very late infections, the only symptoms to appear in the current season may be a few dark streaks on the tips of the flower bracts. No fruit is produced in subsequent years, and plants generally die within a couple of years. From Taiwan, symptomless strains of BBTV, and mild strains that produce only limited vein clearing and dark green streaks have been reported. Interestingly, DNA-N could not be detected in the mild strain infection. Also, some field plants of the Cavendish subgroup cultivar Veimama from Fiji have been observed to initially show severe symptoms, then to recover and display few if any symptoms. This phenomenon has also been observed in the greenhouse in Cavendish cv Williams. Confirmed hosts of BBTV are mostly confined to the family Musaceae. Known susceptible hosts include Musa species, cultivars in the Eumusa and Australimusa series of edible banana and Ensete ventricosum. Susceptible Musa species include M. balbisiana, M. acuminata ssp. banksii, Musa acuminata ssp. halabanensis, M acuminata ssp. longipetiolata, Musa acuminata ssp. malaccensis, M. acuminata ssp. zebrina, M. coccinea, M. jackeyi, and M. ornata. M. textilis, M. velutina. There are some valid reports of hosts outside the Musaceae, in species in the Zingiberales, including Alpinia purpurata, A. zerumbet, Canna indica, Colocasia esculenta and Hedychium coronarium and from the Alismatales, Colocasia esculenta. However, using other virus isolates, not all tests to infect these hosts have been successful. Although there are no confirmed reports of immunity to BBTV in Musa sp., there are many reports of differences in susceptibility and symptom expression between cultivars. The banana aphid (Pentalonia nigronervosa) has a worldwide distribution in tropical and sub-tropical regions where banana is grown, and in 1925 was shown to be a vector of BBTV. Using morphological and molecular methods, P. nigronervosa f sp. caladii was resurrected as a distinct species, P. caladii. Both species are vectors of this virus. The host ranges of both aphid species show considerable overlap, though P. nigronervosa is found predominantly on Musa sp., while P. caladii occurs mostly on related hosts in the Zingiberales. On banana, they commonly colonize the base of the pseudostem at soil level and for several centimeters below the soil surface, beneath the outer leaf sheaths and newly emerging suckers. Transmission by P. nigronervosa is of the persistent, circulative, non-propagative type, and the virus has been demonstrated to accumulates in the anterior midgut, hemolymph and principal salivary glands. Individuals from areas where BBTV is endemic and from where it is absent, both transmit the virus with equal efficiency. There is a minimum acquisition access period of 4 h, a minimum latent period of a few hours, and a minimum inoculation access period of 15 min. Aphids retain infectivity, after removal from a virus source, for at least 20 days and probably for life. Both nymphs and adults can acquire the virus, though the former a more efficient, and reported transmission rates for individual aphids are in the range 46%–67%. There is no evidence for transmission of BBTV to parthenogenetic offspring or for replication of the virus in the aphid. BBTV is also efficiently transmitted in vegetative planting material, both conventional corms, corm pieces (bits) and suckers, and through micropropagation. All meristems from an infected corm will eventually become infected.

Virus–Host Relationships BBTV is systemic within the banana plant, and following aphid inoculation, symptoms do not appear until at least two more new leaves have been produced (bananas produce single new leaves sequentially from a basal meristem). Histological examination suggests that BBTV is restricted to the phloem tissue, which shows hypertrophy and hyperplasia and a reduction in the

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BBTV Burundi

BBTV South-EastAsian subgroup

Cardamom bushy dwarf virus

Abaca bunchy top virus - Philippines Abaca bunchy top virus - Sarawak

BBTV Pacific/Indian Ocean Subgroup

0.05

Faba bean necrotic yellows virus

Fig. 5 Young BBTV-infected banana plants, showing stunting, and successively shorter narrow leaves with upturned, chlorotic margins. Leaves have failed to emerge fully, giving the plant a bunched appearance.

development of the fibrous schlerenchyma sheaths surrounding the vascular bundles. The cells surrounding the phloem contained abnormally large numbers of chloroplasts, resulting in the macroscopic dark green streak symptom. It has been demonstrated through, the use of RNA probes and PCR, that BBTV replicates briefly at the site of aphid inoculation, then moves down the pseudostem to the basal meristem, subsequently infecting the newly formed leaves, the corm and the roots. The virus apparently does not replicate in leaves formed prior to infection, consistent with the lack of symptoms on these leaves and an inability to recover the virus from them via the aphid vector. BBTV has been detected in most parts of the banana plant, including the leaf lamina and midrib, pseudostem, corm, meristems, roots, fruit stalk and fruit rind.

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Diagnosis The characteristic symptoms expressed by BBTV-infected plants allow symptomatology to be used to identify infected plants in field survey and control activities. However, BBTV is sometimes symptomless in tissue culture and the use of laboratory detection assays is desirable in the provision of clean planting material. Polyclonal and monoclonal antibodies to BBTV are used in ELISA to detect the virus in field and tissue culture plants and can detect the virus in single viruliferous aphids. Polymerase chain reaction (PCR) was shown to be about one thousand times more sensitive than ELISA or dot blots with DNA probes. Substances in banana sap inhibitory to PCR can be circumvented by simple extraction procedures or by immunocapture PCR. Nucleic acid-based loop mediated isothermal amplification (LAMP) and recombinase polymerase amplification (RPA) assays have also been described for the detection of BBTV.

Geographical Distribution Historical Records and Possible Origins The origin of BBTV is unclear. The first records were from Fiji in 1889, and though reports and photographs clearly indicate that it was present at least 10 years prior, soon after the establishment of an export industry based on Cavendish cultivars, evidence suggests that the disease did not originate there. Other early recorded are from Egypt (1901 – source unknown) and Australia and Sri Lanka both in 1913, and probably from planting material imported from Fiji. Considerably more genetic diversity, and also a greater incidence of alphasatellites is found in BBTV isolates from South-East Asia. The aphid vector and the wild progenitors of modern edible bananas also originated in the South and South-East AsianAustralasian region, and the Cavendish cultivars of international trade and associated with early outbreaks of bunchy top disease, are thought to have originated in South-southern China/northern Vietnam. These factors lead to speculation that BBTV also originated and evolved in this region. BBTV has a widespread, but scattered distribution in many of the banana-growing countries of the Asia-Pacific regions and Africa, but at present is not found in the Americas. In some countries the distribution is localized. For example, in Australia it is present only in southern Queensland and northern New South Wales, but not in the major production area of north Queensland. Banana bunchy top disease has been recorded in the following countries: Africa: Angola, Benin, Burundi, Cameroon, Central African Republic, Congo Republic (Congo-Brazzaville), Democratic Republic of Congo (formerly Zaire), Egypt, Gabon, Malawi, Mozambique, Nigeria, Rwanda, South Africa, Zambia. Asia: Bangladesh, Cambodia, China, Hong Kong, Indonesia, India, Iran, Japan (Ogasawara-gunto, formerly Bonin Island, Okinawa), Korea, Laos, Malaysia (Sarawak), Myanmar, Pakistan, Philippines, Sri Lanka, Taiwan, Thailand, Vietnam. Oceania: Australia, Fiji, Guam, Kiribati (formerly Gilbert Islands), Marianas Islands (Guam, Saipan, Tinian and Rota), New Caledonia, Samoa, Tahiti, Tonga, Tuvalu (formerly Ellice Islands), USA (American Samoa, Hawaii), Wallis Island.

Epidemiology and Control of BBTV Outbreaks of bunchy top disease can have a devastating effect on banana production, especially industries based on Cavendish cultivars. Production in Fiji fell by more than 80% from 1892 to 1895, primarily due to bunchy top disease. By 1925, around 10 years from the introduction of BBTV to Australia, the banana industry in northern New South Wales and southern Queensland had collapsed, with most plantations affected and production decreased by 90%–95%. Magee noted at the time “It would be difficult for anyone who has not visited these devastated areas to visualize the completeness of the destruction wrought in such a short time by a plant disease”. More recently, a severe outbreak of banana bunchy top disease occurred in Pakistan. From 1991–1992, production area fell by 55% and total production by 90%, as a direct result of the disease. The disease has also recently appeared in Hawaii, New Caledonia, Benin, Nigeria, Mozambique and Malawi, all with severe impact in the affected areas. The epidemiology of banana bunchy top disease is simplified by the occurrence of a single insect vector species and a limited host range for the virus, usually cultivated or feral edible bananas. Long distance spread is usually via infected planting material, and local spread via aphids and planting material. Analysis of actual outbreaks of bunchy top disease in commercial banana plantations in Australia showed that the average distance of secondary spread by aphids was only 15.5–17.2 m, with nearly two-thirds of new infections less than 20 m from the nearest source of infection and 99% less than 86 m. Isolation of new plantations has a marked effect on reducing the risk of infection. New plantations situated adjacent to affected plantations had an 88% chance of recording infections in the first year. This was reduced to 27% if the plantations were separated by 50–1000 m, and to 5% if separated by more than 1000 m. The disease latent period (i.e., period from inoculation of a plant until an aphid can transmit the virus from this plant to another) is equivalent to the time taken for 3.7 new leaves to emerge from the plant. The actual time varies seasonally depending on the rate of leaf production. Control strategies were devised by CJP Magee in the 1920s, and these measures still form the basis of the very successful control program in Australia today. The two major elements of the strategy are (1) exclusion of the disease from unaffected and lightly affected areas and (2) eradication of infected plants from both lightly and heavily-affected areas.

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These measures require the participation of all growers and are unlikely to succeed if left to the goodwill of growers alone and are thus enforced by legislation in Australia. The measures include: (1) (2) (3) (4) (5) (6)

Registration of all banana plantations. Establishment of quarantine zones. Restrictions on the movement and use of planting material. Regular inspections of all banana plantations for bunchy top. Prompt destruction of all infected plants. Ongoing education and extension programs for growers.

When adopted, these measures allowed the complete rehabilitation of the Australian banana industry. Sometimes total eradication of BBTV from a district has been achieved, but in most cases, incidence has been reduced to very low, manageable levels. Such successful control of bunchy top is rarely achieved in other countries, in most cases due to an inability to enforce an organized control program across whole districts. There are no confirmed reports of immunity to BBTV in Musa. However, it has frequently been observed that there are differences in susceptibility between cultivars to both field and experimental infection. Edible bananas have diploid, triploid or tetraploid genomes containing, predominantly, elements of the M. acuminata (A), or M. balbisiana (B) genomes. Cultivars in the Cavendish subgroup (AAA genome), which dominates the international export trade, and many other A genome cultivars are highly susceptible and show severe disease symptoms. Interestingly, however, Gros Michel (AAA) displays resistance to the disease under both experimental inoculation and field conditions. Compared with highly sensitive cultivars such as Cavendish, the cultivar is less susceptible to aphid inoculation, contains a lower level of virions in infected plants, and symptoms are less severe and develop more slowly. These factors may contribute to a reduced rate of aphid transmission and field spread in plantations of Gros Michel, and introduction of this cultivar may explain the partial recovery of the Fijian banana industry after devastation of the Cavendish-based industry in the early 1900s. Field observations and glasshouse inoculations suggest that some B genome-containing cultivars are less susceptible to infection and/or display more limited symptoms, but this needs to be further investigated. In glasshouse trials, the RNAi strategy for BBTV resistance based on DNA-R sequences has been shown to be effective in India and with DNA-M in Australia. In Malawi, initial field testing of transgenic bananas with RNAi-based BBTV resistance was less successful.

See also: Nanoviruses (Nanoviridae)

Further Reading Allen, R.N., 1987. Further studies on epidemiological factors influencing control of banana bunchy top disease, and evaluation of control measures by computer simulation. Australian Journal of Agricultural Research 38, 373–382. Amin, I., Ilyas, M., Qazi, J., et al., 2011. Identification of a major pathogenicity determinant and suppressors of RNA silencing encoded by a South Pacific isolate of banana bunchy top virus originating from Pakistan. Virus Genes 42, 272–281. Burns, T.M., Harding, R.M., Dale, J.L., 1995. The genome organization of banana bunchy top virus: Analysis of six ssDNA components. Journal of General Virology 76, 1471–1482. Dugdale, B., Becker, D.K., Beetham, P.R., Harding, R.M., Dale, J.L., 2000. Promoters derived from banana bunchy top virus DNA-1 to  5 direct vascular-associated expression in transgenic banana (Musa spp.). Plant Cell Reports 19, 810–814. Kumar, P.L., Hanna, R., Alabi, O.J., et al., 2011. Banana bunchy top virus in sub-Saharan Africa: Investigations on virus distribution and diversity. Virus Research 159, 171–182. Magee, C.J., 1953. Some aspects of the bunchy top disease of banana and other Musa spp. Journal and Proceedings of the Royal Society of New South Wales 87, 3–18. Magee, C.J.P., 1927. Investigation on the bunchy top disease of the banana. Bulletin of the Council for Scientific and Industrial Research. (30), 88. Niu, S., Wang, B., Guo, X., et al., 2009. Identification of two RNA silencing suppressors from banana bunchy top virus. Archives of Virology 154, 1775–1783. Stainton, D., Martin, D., Muhire, B., et al., 2015. The global distribution of banana bunchy top virus reveals little evidence for frequent recent, human-mediated long-distance dispersal events. Virus Evolution 1, vev009. Thomas, J.E., Dietzgen, R.G., 1991. Purification, characterization and serological detection of virus-like particles associated with banana bunchy top disease in Australia. Journal of General Virology 72, 217–224. Thomas, J., 2018. Diseases caused by viruses. In: Jones, D.R. (Ed.), Handbook of Diseases of Banana, Abacá and Enset. Wallingford: CABI, p. 632. Thomas, J.E., Smith, M.K., Kessling, A.F., Hamill, S.D., 1995. Inconsistent transmission of banana bunchy top virus in micropropagated bananas and its implication for germplasm screening. Australian Journal of Agricultural Research 46, 663–671. Vetten, H.J., Dale, J.L., Grigoras, I., et al., 2012. Nanoviridae. In: King, A.M.Q., Adams, M.J., Carstens, E.B., Lefkowitz, E.J. (Eds.), Virus Taxonomy: Ninth Report of the International Committee on Taxonomy of Viruses. Amsterdam: Elsevier, Academic Press, pp. 395–404. Watanabe, S., Borthakur, D., Bressan, A., 2016. Localization of banana bunchy top virus and cellular compartments in gut and salivary gland tissues of the aphid vector Pentalonia nigronervosa. Insect Science 23, 591–602. Watanabe, S., Greenwell, A.M., Bressan, A., 2013. Localization, concentration, and transmission efficiency of Banana bunchy top virus in four asexual lineages of Pentalonia aphids. Viruses 5, 758–775.

Barley Yellow Dwarf Viruses (Luteoviridae) Leslie L Domier, Agricultural Research Service, US Department of Agriculture, Urbana, IL, United States r 2021 Elsevier Ltd. All rights reserved. This is an update L.L. Domier, Barley Yellow Dwarf Viruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00637-3.

Glossary Hemocoel The primary body cavity of most arthropods that contains most of the major organs and through which the hemolymph circulates.

Hemolymph A circulatory fluid in the body cavities (hemocoels) and tissues of arthropods that is analogous to blood and/or lymph of vertebrates.

Introduction Barley yellow dwarf disease (BYD) is one of the most economically important virus diseases of cereals, and is found in almost every grain growing region in the world. Widespread BYD outbreaks in cereals were noted in the United States in 1907 and 1949. However, it was not until 1951 that a virus was proposed as the cause of the disease. The causal agents of BYD are obligately transmitted by aphids, which probably delayed the initial classification of BYD as a virus disease. Subsequently, BYD was shown to be caused by three species of Barley yellow dwarf virus (BYDV), a species of Cereal yellow dwarf virus (CYDV) and a species of Maize yellow dwarf virus (MYDV). Depending on the virulence of the virus strain, infection may contribute to winter kill in regions with harsh winters, induce plant stunting, inhibit root growth, reduce or prevent heading, or increase plant susceptibility to opportunistic pathogens and other stresses. Yield losses to wheat in the United States alone are estimated at 1%–3% annually, exceeding 30% in certain regions in epidemic years. The effects of BYD in barley and oats typically are more severe than in maize and wheat; sometimes resulting in complete crop losses. The existence of multiple strains and species of viruses that are transmitted in strain-specific manner made the viruses model systems to study interactions between viruses and aphid vectors in the circulative transmission of plant viruses. In addition, the compact genomes of the viruses have provided useful insights into the manipulation of host translation machinery by RNA viruses.

Taxonomy and Classification The viruses that cause BYD are members of the family Luteoviridae, and were first grouped because of their common biological properties. These properties included persistent transmission by aphid vectors, the induction of yellowing symptoms in grasses, and serological relatedness. Different viruses are transmitted more efficiently by different species of aphids, a fact that was originally used to distinguish the viruses. Around 1960, the viruses were separated into five ‘strains’ (now recognized as distinct species) based on their primary aphid vectors. BYD-causing viruses transmitted most efficiently by Sitobion (formerly Macrosiphum) avenae were assigned the acronym MAV, for Macrosiphum avenae virus. Similarly, viruses transmitted most efficiently by Rhopalosiphum maidis and Rhopalosiphum padi were assigned the acronyms RMV and RPV, respectively. Viruses transmitted most efficiently by Schizaphis graminum were assigned the acronym SGV. Finally, vector-nonspecific viruses, that is, viruses transmitted efficiently by both R. padi and S. avenae were assigned the acronym PAV. The abbreviations now are assigned to BYDV-MAV, BYDV-PAV, BYDV-SGV, CYDV-RPV and MYDV-RMV. Subsequently, additional species were described, including a severe BYDV (BYDV-PAS) and a severe CYDV (CYDV-RPS). Prevalent BYDcausing viruses from China also have been named, including a Luteovirus, BYDV-GAV, which is a strain of BYDV-MAV transmitted by S. graminum and S. avenae, and a Polerovirus named wheat yellow dwarf virus-GPV (WYDV-GPV) transmitted by R. padi and S. graminum. Recently, two new BYDV species, BYDV-kerII and BYDV-kerIII, were identified in native grasses on the Keruelen islands in the Antarctic that are thought to have been introduced along with R. padi during the second half of the 20th century. Based on genome organization and predicted amino acid sequence similarities, BYDV-MAV, BYDV-kerII, BYDV-kerIII, BYDVPAS, and BYDV-PAV have been assigned to the genus Luteovirus, and CYDV-RPS, CYDV-RPV, MYDV-RMV and WYDV-GPV to the genus Polerovirus. The RNA-dependent RNA polymerases (RdRps) encoded by open reading frames (ORFs) 1 and 2 of luteoviruses are phylogenetically related to those of members of the family Tombusviridae (Fig. 1). In contrast, the predicted amino acid sequences of the RdRps encoded by ORFs 1 and 2 of poleroviruses are phylogenetically related to those of viruses in the family Solemoviridae. The two polymerase types are distantly related in evolutionary terms. For this reason, BYDV-SGV has not been definitively assigned to a genus because its genome has not been completely sequenced. The divergent origin of the RdRp sequences suggest that the genomic RNAs of luteoviruses and poleroviruses were formed by recombination between RNAs expressing a common set of structural and movement proteins and RNAs expressing two different sets of replication proteins. Because of these differences, it has been suggested that luteoviruses should be placed in the family Tombusviridae and poleroviruses in the family Solemoviridae.

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Fig. 1 Phylogenetic relationships of the predicted amino acid sequences of RNA-dependent RNA polymerases (RdRps) encoded by ORF2s of barley yellow dwarf (BYD)-causing viruses and selected members of the families Solemoviridae and Tombusviridae. The RdRps of BYD-causing luteoviruses are more similar to those of members of the family Tombusviridae than to those of BYD-causing poleroviruses. Similarly, the RdRp amino acid sequences of BYD-causing poleroviruses and Potato leaf roll virus (type member of the genus Polerovirus) are more similar to those of members of the family Solemoviridae than to those of BYD-causing luteoviruses. The resulting consensus tree from 500 bootstrap replications is shown. The numbers next to each node indicate the percentage of bootstrap replicates in which that node was recovered.

Virion Properties and Composition All members of the family Luteoviridae have non-enveloped icosahedral particles with diameters of 25–28 nm (Fig. 2). Capsids are composed of major (22 kDa) and minor (65–72 kDa) coat proteins (CPs). The minor CP is formed by a carboxy-terminal extension to the major CP called the readthrough domain (RTD). According to X-ray diffraction and molecular mass analysis, virions consist of 180 protein subunits, arranged in T ¼ 3 icosahedra. Virus particles do not contain lipids or carbohydrates, and have sedimentation coefficients s20,w (in Svedberg units) that range from 115 to 118 S. Buoyant densities in CsCl are approximately 1.4 g cm3. Virions are moderately stable, insensitive to freezing, and are insensitive to treatment with chloroform or nonionic detergents but are disrupted by prolonged treatment with high concentrations of salts. Table 1.

Genome Organization and Expression Genomic RNAs of BYDV-causing viruses for which complete nucleotide sequences are available contain six to seven ORFs (Fig. 3). ORFs 1, 2, 3, 3a, 4 and 5 are shared among all the viruses. BYDVs lack ORF0. Genomic sequences of some BYDVs contain a small ORF, ORF6, downstream of ORF5. In poleroviruses, ORFs 0 and 1 and ORFs 1 and 2 overlap by more than 600 nt. In BYDVs, ORF1 overlaps ORF2 by less than 50 nt. In BYDV-causing virus genome sequences, ORF4 is contained within ORF3. An amber (UAG) termination codon separates ORFs 3 and 5. The genomes of BYD-causing viruses have relatively short 50 ends and intergenic non-coding regions. ORFs 2 and 3 are separated by about 200 nt. The lengths of non-coding sequences downstream of ORF5 are very different between BYDVs and BYD-causing viruses. BYDV-PAV contains over 860 nt downstream of ORF5 compared to just 170 nt for CYDV-RPV. The expression of BYDV-PAV RNA has been studied in detail and has revealed a complex set of RNA–RNA and RNA–protein interactions that are employed to express and replicate the virus genome. Less experimental data are available for BYDV-causing poleroviruses. However, expression and replication strategies and gene functions can be inferred from those of closely related poleroviruses, particularly Beet western yellows virus (BWYV) and Potato leaf roll virus (PLRV). ORFs 0, 1, and 2 are expressed directly from genomic RNAs. Downstream ORFs are expressed from subgenomic RNAs (sgRNAs) that are transcribed from internal initiation sites by virus-encoded RdRps from negative-strand RNAs and are 30 -coterminal with the genomic RNA. Since the initiation codon for ORF0 of poleroviruses is upstream of that of ORF1, translation of ORF1 is initiated by ‘leaky scanning’ in which ribosomes bypass the AUG initiation codon of ORF0 and continue to scan the genomic RNA until they reach the initiation codon of ORF1. The protein products of ORF2 are expressed only as a translational fusion with the product of ORF1. At a low frequency during the expression of ORF1, translation continues into ORF2 through a  1 frameshift that produces a large protein containing sequences encoded by both ORFs 1 and 2 in a single polypeptide. The frameshift is mediated by a

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Fig. 2 Scanning electron micrograph of barley yellow dwarf virus-PAV particles, magnified 200,000  . Virions are c. 25 nm in diameter, hexagonal in appearance, and have no envelope and encapsidate a c. 5.6 kb molecule of single-stranded, positive-sense RNA. CYDV-RPV virions also encapsidate a 322-nucleotide satellite RNA that accumulates to high levels in the presence of the helper virus.

Table 1

Viruses causing barley yellow dwarf in cereals

Genus

Species (alternative name)

Abbreviation

Accession numbera

Luteovirus

Barley yellow dwarf virus KerII Barley yellow dwarf virus KerIII Barley yellow dwarf virus-MAV Barley yellow dwarf virus-PAS Barley yellow dwarf virus-PAV (Barley yellow dwarf virus-RGV) (rice giallume) Cereal yellow dwarf virus-RPS Cereal yellow dwarf virus-RPV Maize yellow dwarf virus-RMV Wheat yellow dwarf virus-GPV Barley yellow dwarf virus-GAV Barley yellow dwarf virus-GPV Barley yellow dwarf virus-SGV

BYDV-KerII BYDV-KerIII BYDV-MAV BYDV-PAS BYDV-PAV

NC_021481.1 KC559092.1 NC_003680.1 NC_002160.2 NC_004750.1

CYDV-RPS CYDV-RPV MYDV-RMV WYDV-GPV BYDV-GAV BYDV-GPV BYDV-SGV

NC_002198.2 NC_004751.1 NC_021484.1 NC_012931.1

Polerovirus

Unassigned

a

Accession numbers beginning with NC_ represent reference genomic sequences.

“slippery heptanucleotide sequence” (in the form X XXY YYZ) and a downstream RNA secondary structure termed a pseudoknot that causes ribosomes to pause and then shift back one nucleotide before continuing translation in the new reading frame. In BYDV-PAV, frameshifting between ORFs 1 and 2 also is dependent upon the interaction of RNA sequences close to the site of frameshifting and a long-distance frameshift element (LDFE) located 4000 nt downstream in the 30 non-coding region of genomic RNAs. Mutations that disrupt the interactions between these two distal regions suppress frameshifting and abolish RNA replication. ORFs 3, 3a, and 4 are expressed from the 50 terminus of sgRNA1 by a leaky scanning mechanism much like that used to express ORF1 of BYD-causing poleroviruses. The 50 terminus of sgRNA1 is located about 200 nt upstream of ORF3, and the 30 terminus is co-terminal with the genome. ORF3a is initiated as a non-AUG codon. The product of ORF5 is expressed only as a translational fusion with the products of ORF3 by readthrough of the UAG termination codon at the end of ORF3. This produces a protein with the product of ORF3 at its amino terminus and the product of ORF5 at its carboxyl terminus. Readthrough is regulated by local and long-distance RNA-RNA interactions. BYDVs produce two additional sgRNAs. sgRNA2 contains ORF6, but sgRNA3 does not contained a predicted ORF. While genomic RNAs of BYD-causing poleroviruses contain 50 VPgs that interact with translation initiation factors, BYDV-PAV RNA contains only a 50 phosphate. Unmodified 50 termini usually are recognized poorly for translation initiation. To circumvent this problem, a short sequence located in the noncoding region just downstream of ORF5 in the BYDV-PAV genome, called the BYDV translation enhancer (BTE), promotes efficient cap-independent translation initiation by directly interacting with eukaryotic translation initiation factor 4E and sequences near the 50 termini of the genomic and sgRNA1.

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Fig. 3 Genome organizations of Barley yellow dwarf virus-PAV (BYDV-PAV) and Cereal yellow dwarf virus-RPV (CYDV-RPV). Individual open reading frames (ORFs) are shown as staggered open boxes. The predicted sizes of the protein products are indicated. The genome-linked protein (VPg) attached to the 50 terminus of CYDV RNA is indicated by a solid circle. Based on homology to other viruses ORF0 encodes a silencing suppressor and ORFs 1 and 2 encode replication-related proteins. ORFs 3 and 5 encode the major coat protein and readthrough domain, respectively. ORF3a, which is initiated at a non-AUG codon, encodes a protein required for long-distance movement. ORF4 encodes a protein required for virus cell-to-cell movement. The BYDV translation enhancer (BTE) facilitates translation initiation of BYDV-PAV genomic RNA and subgenomic RNA1 (sgRNA1). In both BYDV-PAV and CYDV-RPV, ORF2 is expressed as a translational fusion with the product of ORF1 via a  1 frameshift. In BYDV-PAV, frameshifting requires interaction between the 50 frameshift signals and the long-distance frameshift element (LDFE). Dashed lines indicate long-distance RNA–RNA interactions. GNRA tetraloops at the 30 terminus of BYDV-PAV genomic RNA are required for negative-strand RNA synthesis and virus replication.

Functions of BYDV and CYDV proteins have been ascribed based on homology to virus proteins with known functions and mutational characterization of protein coding regions. Similar to BWYV and PLRV, the proteins encoded by ORF0s of BYD-causing poleroviruses are inhibitors of local and systemic post-transcriptional gene silencing (PTGS). PTGS is an innate and highly adaptive antiviral defense found in all eukaryotes that is activated by double-stranded RNAs (dsRNAs), which are produced during virus replication. Furthermore, researchers noted a correlation between symptom severity and the strength of PTGS suppression of ORF0-encoded proteins of CYDV-RPV and CYDV-RPS. The genomes of BYDVs lack an ORF0, but the product of ORF4 functions as an inhibitor of systemic PTGS in those viruses. The ORF1-encoded proteins of BYD-causing poleroviruses contain the VPg and a chymotrypsin-like serine protease that is responsible for the proteolytic processing of ORF1-encoded polyproteins. The protease cleaves the ORF1 protein in trans to liberate the VPg, which is covalently attached to genomic RNAs. ORF2s, which are expressed as translational fusions with the product of ORF1, have coding capacities of 59–72 kDa and predicted amino acid sequences that are very similar to known RdRps and likely represent the catalytic portion of the viral replicase.

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Primary salivary gland

Mycetocytes

Accessory salivary gland

Midgut Hindgut Hemocoel

Foregut

Food canal Salivary duct

Fig. 4 Circulative transmission of BYD-causing viruses by vector aphids. While feeding from sieve tubes of an infected plant, an aphid (shown in cross section) acquires virions, which travel up the stylet, through the food canal, and into the foregut. Virions are actively transported across cells of the hindgut into the hemocoel and then passively diffuse through the hemolymph to the accessory salivary gland where they are again actively transported into the lumen of the gland. Once in the salivary gland lumen, the virions are expelled with the saliva into the vascular tissue of host plants. Viruses that are not transmitted by a particular species of aphid often accumulate in the hemocoel, but do not traverse the membranes of the accessory salivary gland.

ORF3a expresses highly conserved 4.8–5.3 kDa proteins that are essential for long-distance movement. ORF3 encodes the major 22 kDa CP. ORF5 has a coding capacity of 43–50 kDa, which is expressed only as a translational fusion with the product of ORF3 when translation reads through the termination codon at the end of ORF3 and continues through to the end of ORF5. The ORF5 portion of this readthrough protein has been implicated in aphid transmission and virus stability. Recombinant viruses that do not express ORF5 produce virions assembled from the major CP alone, which are not transmitted by aphid vectors and are less efficient in systemic infection of host plants than wild-type viruses. The amino-terminal portions of ORF5 proteins are highly conserved among BYD-causing viruses, while the carboxyl termini are much more variable. Proteins encoded by ORF4 are thought to facilitate intra- and intercellular virus movement because viruses with mutations in ORF4 are able to replicate in isolated plant protoplasts, but are deficient or delayed in systemic movement in whole plants. Some BYDV genomic sequences contain an ORF6 downstream of ORF5. The predicted sizes of the proteins expressed by ORF6 range from 4 to 7 kDa. The predicted amino acid sequences of the proteins encoded by ORF6 are poorly conserved among BYDV-PAV isolates. Repeated attempts to detect protein products of ORF6 have been unsuccessful. In addition, BYDV-PAV genomes into which mutations have been introduced that disrupt ORF6 translation are still able to replicate in protoplasts. Based on these observations, it has been concluded that ORF6 is not translated in vivo. However, sgRNA2, which contains ORF6, has been shown to inhibit translation of capped and polyadenylated mRNAs, and hence, may function as a negative riboregulator of host translation.

Host Range and Transmission BYD-causing viruses infect over 150 species of annual and perennial grasses in five of the six subfamilies of the Poaceae. The feeding habits of vector aphids have a major impact on the host ranges of virus species. Hence, the number of species naturally infected by the viruses is much lower than their experimental host ranges. As techniques for infecting plants with recombinant viruses have improved, the experimental host ranges of the viruses have been expanded to include plants on which aphid vectors would not normally feed. For example, BYDV-PAV and CYDV-RPV have been shown to infect Nicotiana species when inoculated using Agrobacterium tumefaciens harboring binary plasmids containing infectious copies of the viruses, which had not been described previously as experimental hosts for the viruses. Viruses that cause BYD are transmitted in a circulative strain-specific manner by at least 25 aphid species. Circulative transmission of the viruses is initiated when the piercing–sucking mouthparts of aphids acquire viruses from sieve tubes of infected plants during feeding. Aphids that do not probe into and feed from the vascular tissues of infected plants do not transmit the

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viruses. The virions travel up the stylet, through the food canal, and into the foregut (Fig. 4). After 12–16 h, virions then are actively transported across the cells of the hindgut into the hemocoel in a process that involves receptor-mediated endocytosis of the viruses and the formation of tubular vesicles that transport viruses through epithelial cells and into the hemocoel. Virions then passively diffuse through the hemolymph to the accessory salivary gland where virions must pass through the membranes of accessory salivary gland cells in a similar type of receptor-mediated transport process to reach the lumen of the gland. The accessory salivary gland produces a watery saliva, containing few or no enzymes, that is thought to prevent phloem proteins from clogging the food canal. Once in the salivary gland lumen, virions are expelled with the watery saliva into vascular tissues of host plants. Typically, hindgut membranes are much less selective than those of the accessory salivary glands. Consequently, viruses that are not transmitted by a particular species of aphid often are transported across gut membranes and accumulate in the hemocoel, but do not traverse the membranes of the accessory salivary gland. The specificity of aphid transmission and gut tropism has been linked to the RTD of the minor capsid protein. Even though large amounts of virions can accumulate in the hemocoel, there is no evidence for virus replication in their aphid vectors. Aphids may retain the ability to transmit virus for several weeks. Genetic and biochemical studies have been conducted to identify aphid determinants of strain-specific transmission of BYDV-MAV and BYDV-PAV. Protein–protein and protein–virus interaction experiments were used to isolate two proteins from heads of vector aphids that bind BYDV-MAV that were not detected in non-vector aphids. These two proteins are good candidates for the cell-surface receptors that are thought to be involved in strain-specific transport of viruses into accessory salivary gland lumens. In addition, endosymbiotic bacteria that reproduce in specialized cells called mycetocytes in abdomens of aphids express chaperonin-like proteins that bind BYDV particles and the amino-terminal region of recombinant BYDV-PAV RTD proteins. However, the role of these proteins in aphid transmission is unclear since they are found in both vector and non-vector aphid species. Interactions of virus particles with these proteins seem to be essential for persistence of the viruses in aphids. The proteins may protect virus particles from degradation by aphid immune systems.

Replication Like other viruses of the family Luteoviridae, BYD-causing viruses infect and replicate in sieve elements and companion cells of the phloem and occasionally are found in phloem parenchyma cells. The viruses induce characteristic ultrastructural changes in infected cells. BYDV-MAV, -PAV, and -SGV induce single-membrane-bound vesicles in the cytoplasm near plasmodesmata early in infection. Subsequently, filaments are observed in nuclei, and virus particles are first observed in the cytoplasm. In contrast, CYDV-RPV and MYDV-RMV induce double-membrane-bound vesicles in the cytoplasm that are continuous with the endoplasmic reticulum. Later, filaments and tubules form in the cytoplasm, and CYDV-RPV and MYDV-RMV particles are first observed in nuclei. The subcellular location of viral RNA replication has not been determined unequivocally. However, early in infection, negativestrand RNAs of BYDV-PAV are first detected in nuclei and later in the cytoplasm, which suggests that at least a portion of the BYDV-PAV replication occurs in the nucleus. A nuclear location for replication is supported by the observation that the movement protein encoded by ORF4, which also binds single-stranded RNA, localizes to the nuclear envelope and is associated with virus RNA in nuclei of infected cells. Synthesis of negative-strand RNA, which requires tetraloop structures at the 30 end of BYDV-PAV genomic RNAs, is detected in infected cells before the formation of virus particles. Because tetraloops have been implicated in RNA–protein interactions, these structures could be binding and/or recognition sites for BYDV replication proteins. BYDV-PAV sgRNAs are synthesized by internal initiation of RNA synthesis on negative-strand RNAs from three dissimilar subgenomic promoters. Late in infection, the BTE near the 50 terminus of BYDV-PAV sgRNA2 inhibits translation from genomic RNA, which may promote a switch from translation to replication and packaging of genomic RNAs. In addition to genomic RNAs, CYDV-RPV replicates a satellite RNA by a rolling-circle mechanism that generates multimeric satellite RNAs that self-cleave to unit length.

Virus–Host Relationships Visible symptoms induced by the viruses vary greatly depending on the genotype and developmental stage of the host and strain of the virus. The most common symptoms are stunting and chlorosis. While some infected plants display no obvious symptoms, most BYD-causing viruses induce characteristic symptoms that include stunting, leaves that become thickened, curled or serrated, and yellow, orange or red leaf discoloration, particularly of older leaves of infected plants. These symptoms result from phloem necrosis that spreads from inoculated sieve elements and causes symptoms by inhibiting translocation, slowing plant growth, and inducing the loss of chlorophyll. Symptoms may persist, may vary seasonally, or may disappear soon after infection. Temperature and light intensity often affect symptom severity and development. In addition, symptoms can vary greatly with different virus isolates or strains and with different host cultivars. Yield losses caused by BYD are difficult to estimate because the viruses are so pervasive, and symptoms often are overlooked or attributed to other agents. In Australia alone, losses in barley production have been valued at over 100 million US dollars annually. Plants infected with viruses causing BYD at early developmental stages suffer the most significant yield losses, which often are linearly correlated with the incidence of virus infection.

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Epidemiology BYD infections have been reported from temperate, subtropical, and tropical regions of the world. Even though the incidence of infections of individual viruses varies from year to year and can differ among annual and perennial hosts, BYDV-PAV usually is the most prevalent of the viruses causing BYD in small grains worldwide followed by CYDV-RPV or BYDV-MAV. The remaining BYD-causing viruses are typically much less prevalent. The viruses must be reintroduced into annual crops each year by their aphid vectors. Alate, that is, winged aphids may transmit viruses from local cultivated, volunteer, or weed hosts. Alternatively, alate aphids may be transported into crops from distant locations by wind currents. These vectors may bring the virus with them, or they may first have to acquire virus from locally infected hosts. In temperate regions of Europe and North America moderate and long-distance migration of viruliferous aphids is important to development of BYD epidemics. In Australasia, and other regions with Mediterranean climates, alate aphids usually transmit viruses from relatively close infected plants. Secondary spread of the viruses is often primarily by apterous, that is, wingless aphids. The relative importance of primary introduction of viruses by alate aphids and of secondary spread of viruses by apterous aphids in disease severity varies with the virus, aphid species, crop, and environmental conditions. Studies have shown both vector and non-vector aphids species preferred to feed on noninfected plants after having acquired BYDV virus particles, while nonviruliferous aphids preferred to feed on BYDV-infected plants. This modification of aphid behavior associated with virus acquisition could enhance the spread of virus within plant populations. In addition to crop plants, BYD-causing viruses native and invasive annual and perennial grasses. In some cases, prevalence of BYDV in native habitats was inversely correlated with distance from cultivated cereals. Grazing of herbivores on natural grass lands was also associated with increased incidence of BYDV infections. As observed on Keruelen islands in the Antarctic, the introduction of vector aphid species into new environments can increase incidence of virus infections that negatively impacts native plant communities.

Diagnosis Accurate diagnosis of infections has been important in understanding the transmission and epidemiology of the viruses and developing control strategies for BYD. Because BYD symptoms resemble those caused by other biotic and abiotic factors, visual diagnosis is unreliable and other methods have been developed. Initially, infectivity, or biological, assays were used to diagnose infections. In bioassays, aphids are allowed to feed on infected plants and then are transferred to indicator plants. These techniques have also been used to determine vector specificities of viruses causing BYD and to identify viruliferous vector aphids in epidemiological studies. These techniques are very sensitive but can require several weeks for symptoms to develop on indicator plants. The viruses causing BYD are strongly immunogenic, which has facilitated development of genus- and even strain-specific antibodies that have been used extensively in BYD diagnosis. Because the viruses causing BYD are present in infected tissues at very low levels, mice have been used to produce monoclonal antibodies against the viruses. Mice typically require much less viral antigen per immunization than rabbits, and hybridoma cell lines that produce monoclonal antibodies can be stored for extended periods and used for many years, which further reduces the amount of antigen needed to produce diagnostic antibodies. Techniques have also been developed to detect viral RNAs from infected plant tissues by reverse transcription polymerase chain reaction, which can be more sensitive and discriminatory than serological diagnostic techniques. Even so, serological tests are the most commonly used techniques for the detection of infections because of their simplicity, speed, and relatively low cost.

Control Planting of insecticide-treated seeds that protect emerging seedlings from aphid infestation has been shown to reduce losses caused by BYD in Africa, Australasia and North America. Foliar applications of insecticides on older plants typically have been less effective. Alternatively, planting of tolerant or resistant cereals has proved to be a much more cost-effective and sustainable management strategy for BYD. Breeding programs have successfully integrated genes conferring high levels of tolerance into barley and oat and to a lesser extent in wheat. Even though a limited number of single genes for BYD resistance/tolerance have been identified in cultivated barley and rice, in most instances, tolerance to BYD is conditioned by multiple genes in a quantitative fashion, which has made moving BYD tolerance into new plant lines challenging. Particularly in barley, molecular markers have begun to facilitate the process of breeding for BYD tolerance. Because of a lack of effective single-gene resistance in cultivated wheat, some researchers have moved BYD resistance genes from wheat grasses (Thinopyrum intermedium and Thinopyrum ponticum) into wheat, which have provided high levels of resistance. The lack of naturally occurring resistance in cereals to BYD has made transgene-mediated resistance very attractive. Transgenic barley and oat plants have been produced that express either intact or inverted-repeat regions of the BYDV-PAV genome that conferred high levels of resistance to BYDV-PAV and closely related viruses. However, the transgenic plant lines have not been released for use by growers possibly because of the expenses of obtaining regulatory approval and/or poor acceptance of genetically modified cereal crops by grower and consumer groups. In many small grain growing regions, viruliferous aphids arrive at similar times each spring and fall even though sizes of the aphid populations can vary significantly from year to year. In these areas, it is sometimes possible to plant crops so that young,

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highly susceptible plants are not in the field when the seasonal aphid migrations occur. However, crops planted later typically do not yield as well as those planted early in the growing season. Consequently, growers must weigh the probability of obtaining higher yields against possible yield losses caused by BYD. In some instances, biological control agents such as predatory insects and parasites have reduced aphid populations significantly.

See also: Luteoviruses (Luteoviridae). Solemoviruses (Solemoviridae). Tombusviruses (Tombusviridae)

Further Reading Crasta, O.R., Francki, M.G., Bucholtz, D.B., et al., 2000. Identification and characterization of wheat-wheatgrass translocation lines and localization of Barley yellow dwarf virus resistance. Genome 43, 698–706. D'Arcy, C.J., Domier, L.L., 2000. Barley Yellow Dwarf. United States: American Phytopathological Society, Available at: https://www.apsnet.org/edcenter/disandpath/viral/ pdlessons/Pages/BarleyYelDwarf.aspx (accessed 26.05.19). Falk, B.W., Tian, T., Yeh, H.H., 1999. Luteovirus-associated viruses and subviral RNAs. Current Topics in Microbiology and Immunology 239, 159–175. Fusaro, A.F., Barton, D.A., Nakasugi, K., et al., 2017. The Luteovirus P4 movement protein is a suppressor of systemic RNA silencing. Viruses 9, 16. Gray., S., Gildow, F.E., 2003. Luteovirus-aphid interactions. Annual Review of Phytopathology 41, 539–566. Gray, S., Cilia, M., Ghanim, M., 2014. Circulative, "non-propagative" virus transmission: An orchestra of virus-, insect-, and plant-derived instruments. In: Maramorosch, K., Murphy, F.A. (Eds.), Advances in Virus Research, vol. 89. San Diego: Elsevier Academic Press Inc., pp. 141–199. Hershman, D.E., Johnson, D.W., 2011. Barley Yellow Dwarf – Plant Pathology Fact Sheet. PPFS-AG-SG-03 Cooperative Extension Service. United States: University of Kentucky, p. 5. Available at: https://plantpathology.ca.uky.edu/files/ppfs-ag-sg-03.pdf (accessed 26.05.19). Hulo, C., de Castro, E., Masson, P., et al. 2019. Luteoviridae. ViralZone. ExPASy bioinformatics resource portal. Switzerland. Available at: https://viralzone.expasy.org/609? outline=all_by_species (accessed 26.05.19). Kendig, A.E., Borer, E.T., Mitchell, C.E., Power, A.G., Seabloom, E.W., 2017. Characteristics and drivers of plant virus community spatial patterns in US west coast grasslands. Oikos 126, 1281–1290. Koev, G., Mohan, B.R., Dineshkumar, S.P., et al., 1998. Extreme reduction of disease in oats transformed with the 50 half of the Barley yellow dwarf virus PAV genome. Phytopathology 88, 1013–1019. Miller, W.A., Jackson, J., Feng, Y., 2015. Cis- and trans-regulation of luteovirus gene expression by the 30 end of the viral genome. Virus Research 206, 37–45. Miller, W.A., Liu, S.J., Beckett, R., 2002. Barley yellow dwarf virus: Luteoviridae or Tombusviridae? Molecular Plant Pathology 3, 177–183. Miller, W.A., Rasochova, L., 1997. Barley yellow dwarf viruses. Annual Review of Phytopathology 35, 167–190. Miller, W.A., White, K.A., 2006. Long-distance RNA-RNA interactions in plant virus gene expression and replication. Annual Review of Phytopathology 44, 447–467. Ordon, F., Friedt, W., Scheurer, K., et al., 2004. Molecular markers in breeding for virus resistance in barley. Journal of Applied Genetics 45, 145–159. Smirnova, E., Firth, A.E., Miller, W.A., et al., 2015. Discovery of a small non-AUG-initiated ORF in Poleroviruses and Luteoviruses that is required for long-distance movement. PLOS Pathogens 11, e1004868. Xu, Y., Ju, H.J., DeBlasio, S., Carino, E.J., et al., 2018. A stem-loop structure in potato leafroll virus open reading frame 5 (ORF5) is essential for readthrough translation of the coat protein ORF stop codon 700 bases upstream. Journal of Virology 92, e01544-17. Zhu, S., Kolb, F.L., Kaeppler, H.F., 2003. Molecular mapping of genomic regions underlying barley yellow dwarf tolerance in cultivated oat (Avena sativa L.). Theoretical and Applied Genetics 106, 1300–1306.

Bean Common Mosaic Virus and Bean Common Mosaic Necrosis Virus (Potyviridae) Ramon Jordan, Agricultural Research Service, US Department of Agriculture, Beltsville, MD, United States John Hammond, Floral and Nursery Plants Research, Agricultural Research Service, US Department of Agriculture, Beltsville, MD, United States r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

MP Movement protein nm Nanometer(s) nt Nucleotide(s) ORF Open Reading Frame RdRp RNA-dependent RNA polymerase RT-PCR Reverse transcription-polymerase chain reaction ssRNA Single-stranded RNA UTR Un-translated region Vpg Genome-linked protein

aa Amino acid(s) CP Coat protein or capsid protein DAS-ELISA Double antibody sandwich ELISA DIA Dot immunoassay ELISA Enzyme-linked immunological assays HC-Pro Helper component-proteinase kb Kilobase kDa Kilo dalton McAbs Monoclonal antibodies

Glossary Pathogroup A series of isolates of the same strain or pathotype (i.e., groups of isolates or strains that can be differentiated by their biological response on common bean cultivars carrying different combinations of resistance genes). Pathotype An isolate or strain of a virus that is biologically distinct from other isolates of the same virus by virtue of differential host reactions.

Serogroup A series of isolates or strains of the same serotype. Serotype An isolate or group of isolates that are distinguished from biologically related isolates by reaction (or lack of reaction) with key serological reagents such as defined polyclonal antisera or monoclonal antibodies. Strain A genetic variant or subtype of a viral species.

History Bean common mosaic virus and Bean common mosaic necrosis virus are species within the genus Potyvirus, family Potyviridae, and cause some of the most economically important diseases of legume crops worldwide. Yield losses due to Bean common mosaic virus (BCMV) and Bean common mosaic necrosis virus (BCMNV) may vary between 6 to 98% depending on the cultivar and time of infection. Both viruses occur essentially wherever bean and cowpea (including Phaseolus, Vicia, Vigna), lupin (Lupinus), pea (Pisum), peanut (Arachis), and soybean (Glycine) are grown. BCMV was first reported from the US in 1917, and the associated disease initially known as bean mosaic. It was renamed bean common mosaic in 1934 to differentiate it from bean yellow mosaic, caused by Bean yellow mosaic virus (BYMV), another potyvirus. Several pathotypes or strains of BCMV were distinguished in the 1970s by differential reactions of a number of bean cultivars, and in the early 1980s strains were further divided by serology into serotype A and serotype B. Serogroup A isolates were also biologically differentiated by temperature-insensitive induction of necrosis in bean cultivars carrying the dominant I gene. Peptide profiling of the coat protein (CP) and sequence analysis of the CP gene and 30 untranslated region (UTR) demonstrated that the serotypes represented distinct viruses. Serotype A became known as bean common mosaic necrosis virus, while serotype B retains the name BCMV. ‘Bean necrosis mosaic virus’ was used for serogroup A isolates prior to acceptance of the species name bean common mosaic necrosis virus. BCMV and BCMNV also differ in particle length, CP size, and biological properties (see below). Complete genome sequences support the distinction between BCMV and BCMNV proposed using CP sequences and peptide profiles.

Taxonomy and Classification The current criteria for differentiation of potyvirus species are o76% nt identity and o82% aa identity for the complete single ORF sequence and resulting polyprotein. The thresholds for species demarcation using nucleotide identity values for the individual coding regions range from 58% for the P1 coding region to 74%–78% for other regions. For the CP, the optimal species demarcation criterion is 76%–77% nt and 80% aa identity. Different species also often have distinct polyprotein cleavage sites. Using these criteria, Bean common mosaic virus and Bean common mosaic necrosis virus are distinct potyviral species; BCMV also now includes strains previously described by other names, including Azuki bean mosaic virus, Blackeye cowpea mosaic virus,

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Dendrobium mosaic virus, Guar green sterile virus, Peanut chlorotic ring mottle virus, Peanut mild mottle virus, and Peanut stripe virus. At least 26 different strains of BCMV have been biologically authenticated as distinct on the basis of host range and host response, whereas only five biologically distinct strains of BCMNV have been recognized. Potyviruses can be separated into at least 11 subgroups, including two ‘supergroups’, of which the Bean common mosaic virus supergroup is the largest, and the Potato virus Y (PVY) supergroup the second. Although ‘supergroup’ and ‘subgroup’ have no formal taxonomic meaning, it is a useful concept to describe a subset of virus species that cluster together in a phylogenetic tree (Fig. 1) and which may also have common serological properties. BCMV and BCMNV, together with BSVA, CerMV, CABMV, CLLV, DsMV, EAPV, FVY, HarMV, IFBV, PMNV, PWV, SaLV, SaVY, SMV, TelMV, TrVY, WMV, WVMV, YBMV, ZaMMV, and ZYMV, form the BCMV supergroup (for full virus names, see Table 1). Some confusion exists in the literature over isolates described as Cowpea aphid-borne mosaic virus (CABMV); true CABMV isolates are distinct from BCMV, but some isolates initially described as CABMV were later shown to be synonymous with Blackeye cowpea mosaic virus (BlCMV), which is now recognized as a strain of BCMV.

Geographic Distribution Many believe BCMV probably originated in Latin America, the center of diversity of Phaseolus vulgaris; others suggest South or East Asia. Regardless, BCMV has now spread worldwide and can be found wherever beans are grown, and in association with other bean types, as well as soybean, pea, and cowpea. Blackeye cowpea mosaic isolates are also found worldwide. Peanut stripe isolates are found in peanuts in Asia and the United States, and also infect lupin, soybean, and sesame. In many areas of the world BCMV is the most important potyvirus affecting beans, although in some areas BCMNV or BYMV may predominate. Before major efforts were made to eradicate BCMV from US germplasm collections, more than 60% of germplasm accessions were found to be infected. BCMNV is thought to have evolved from BCMV in Central or Eastern Africa and is more important than BCMV in much of Central, Eastern, and Southern Africa, where BCMNV is common in a wide variety of wild and forage legumes as well as beans grown for human consumption. BCMNV has also been distributed throughout Africa, Europe, North and South America, largely through germplasm introductions, and has increased in importance in the United States partly as a consequence of deployment of resistance genes against BCMV (see below). The increase in diversity and severity of bean-infecting geminiviruses in many regions has displaced BCMV and BCMNV as the most important viruses infecting beans in some areas.

Host Range and Transmission BCMV naturally infects P. vulgaris (kidney bean), P. acutifolius (tepary bean), P. atropurpureus, P. coccineus (runner bean), Glycine max (soybean), Macroptilium lathyroides (horse gram), Pisum sativum (pea), Rhynchosia minima, Vicia faba (broad bean), Vigna mungo (black gram, urdbean), V. radiata (mung bean, green gram), V. angularis (azuki bean), and V. unguiculata (cowpea). Peanut stripe isolates naturally infect Arachis hypogea (peanut), Lablab purpureus (syn. Dolichos lablab; hyacinth bean), Indigofera amoena, G. max, Lupinus albus (lupin), Pueraria phaseoloides, Sesamum spp. (sesame), Stylosanthes capitata, and S. craba. Soybean isolates are adapted to soybean, and rarely infect peanuts. Dendrobium mosaic isolates infect Dendrobium orchids, while guar green sterile isolates infect guar (Cyamopsis tetragonoloba). In addition to its natural hosts, BCMV has a wide experimental host range, infecting about 100 species from 44 genera over nine families including Amaranthaceae, Chenopodiaceae, Fabaceae, Solanaceae, and Tetragoniaceae, although individual isolates may be much more restricted. BCMNV naturally infects P. vulgaris, P. lunatus, Centrosema pubescens, Crotalaria incana, Lablab purpureus, Senna bicapsularis, S. hirsuta, S. sophora, and V. vexillata, and has been detected using a BCMNV-specific monoclonal antibody in naturally infected plants of Albizia coriaria, Desmodium intortum, D. uncinatum, Rhynchosia resinosa, Tephrosia barbigera, and T. paniculata. Experimental hosts include several other leguminous species, such as Canavalia ensiformis, Crotalaria spinosa, Macroptilium lathyroides, Rhynchosia minima, and V. radiata. Both BCMV and BCMNV are transmitted by aphids in a non-persistent manner typical of viruses in the genus Potyvirus; the most important vector species in many countries are Acyrthosiphon pisum, Aphis fabae, and Myzus persicae. Aphis craccivora may be the most important vector in India. Minimum acquisition access and inoculation access periods are less than 1 min; no latent period is required between acquisition and inoculation. Aphid transmission is the cause of secondary transmission within the crop, but the primary means of introduction to the crop is through infected seed. Aphid transmission and infection rates tend to be higher when beans are grown under irrigation in dry regions. Both BCMV and BCMNV are also seed-transmitted. Seed transmission varies with isolates and host species, as well as the timing of infection, but rates of up to 93% of infected seed may result from diseased plants, with erratic distribution of infected seed within individual pods. Plants infected after flowering typically do not yield infected seed, probably because the virus is not found in the embryo and cotyledons, and the virus may not be able to spread into these locations if flowering occurs prior to infection of the mother plant. Infection of the seed, and perhaps of the mother plant, can occur through infected pollen. Infectivity of the virus is retained during prolonged storage of seed. Dendrobium mosaic virus (a strain of BCMV) is transmitted by aphids and by vegetative propagation of Dendrobium orchids.

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Fig. 1 Phylogenetic relationships of the Bean common mosaic virus subgroup. Phylogenetic relationship based on analysis of the polyprotein amino acid sequences of Bean common mosaic virus and Bean common mosaic necrosis virus with members of the BCMV subgroup, other legume-infecting potyviruses, and the genus Potyvirus type member, Potato virus Y (PVY). The tree was constructed by the neighbor-joining algorithm based on calculations from Clustal W pairwise amino acid sequence distances. The horizontal branch lengths are proportional to the genetic distance. The data set was subjected to 1000 bootstrap replicates. All nodes supported by 450% confidence values are shown. GenBank sequence sources are listed. Virus acronyms are as listed in Table 1; and, BYMV (Bean yellow mosaic virus), ClYVV (Clover yellow vein virus), and PSbMV (Pea seed-borne mosaic virus).

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Table 1

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The viruses included in the Bean common mosaic virus subgroup

Virus species recognized as members of the bean common mosaic virus subgroup

Acronym

Representative GenBank Accession Number

Bean common mosaic virusa Azuki bean mosaic virusb Bean common mosaic virus serotype B Blackeye cowpea mosaic virus Dendrobium mosaic virus Guar green sterile virus Peanut chlorotic ring mottle virus Peanut mild mottle virus Peanut stripe virus Bean common mosaic necrosis virus Bean common mosaic virus serotype A (Bean necrosis mosaic virus)c Blue squill virus A Calla lily latent virus Ceratobium mosaic virus Cowpea aphid-borne mosaic virus Sesame mosaic virus South African passiflora virus Dasheen mosaic virus (Vanilla mosaic virus)d East Asian passiflora virus Fritillary virus Y Hardenbergia mosaic virus Impatiens flower break virus Passionfruit woodiness virus Paris mosaic necrosis virus Saffron latent virus Sarcochilus virus Y Soybean mosaic virus Telosma mosaic virus Tricyrtis virus Ye Watermelon mosaic virus Watermelon mosaic virus 2 Vanilla necrosis virus Wisteria vein mosaic virus Yambean mosaic virus Zantedeschia mild mosaic virus Zucchini yellow mosaic virus

BCMV (AzMV) BCMV (BlCMV) (DeMV) (GGSV) (PCRMV) (PMMV) (PStV) BCMNV BCMNV (BNMV) BSVA CLLV CerMV CABMV (SeMV) (SAPV) DsMV (VanMV) EAPV FVY HarMV IFBV PWV PMNV SaLV SaVY SMV TelMV TrVY WMV (WMV-2) (VNV) WVMV YBMV ZaMMV ZYMV

AY112735 KP903372 KM023744 KC832501 U23564 AF045066 Y11772 Y11776 U05771 HQ229994 U19287 HQ229995 JQ807999 EF105297 AF022442 KM655833 U90326 S51666 AB219545 KX505964 AB246773 AM039800 HQ161081 KU981084 HQ122652 MF509898 KY562565 AF185957 S42280 DQ851493 AY864850 AY437609 D13913 L22907 AY656816 JN190431 AY626825 L31350

a

Names in bold italic font are species currently recognized by the ICTV. Names in regular font are no longer recognized as potyvirus species, but are indented to indicate their position as strains of the species listed above. Acronyms in parentheses are no longer valid as species, but refer to strains of the virus in bold font. c ‘Bean necrosis mosaic virus’ was a name used for strains of the A serotype prior to ICTV designation of these isolates as strains of BCMNV. d Vanilla mosaic virus was recognized in the ICTV Eighth Report as a tentative species in the genus Potyvirus; recent reports suggest that it is an isolate of DsMV. e Tricyrtis virus Y appears to meet the criteria for distinct virus species, but has not yet been recognized as such by the ICTV; the acronym is indicated only in bold font. b

Properties of the Virion and Genome Properties of particles BCMV and BCMNV are typical of viruses in the genus Potyvirus in that particle preparations contain a single sedimenting (154–158S) and buoyant density component (1.31–1.32 g cm3 in CsCl). Virions are non-enveloped flexuous filaments, 12–15 nm wide, and 847–886 nm (BCMV) or 810–818 nm (BCMNV) long. Virions are composed of one single-stranded RNA (ssRNA) of c. 9600 nt for BCMNV and c. 10,000 nt for BCMV (accounting for B5% of the particle weight) encapsulated in about 1700–2000 monomer units of one CP polypeptide species with a molecular mass of c. 30 kDa for BCMNV, and c. 33 kDa for BCMV (comprising B95% of the particle weight); a molecule of the genome-linked protein NIa-VPg is bound the 50 end of the RNA. In gel electrophoresis the CP of BCMNV migrates with apparent Mr 33 kDa, and that of BCMV of Mr 34.5–35 kDa. Virus preparations that have undergone limited proteolysis also contain lower-molecular-weight peptides of apparent Mr 29–34 kDa.

Serological relationships BCMV and BCMNV particles are strongly immunogenic and can be easily distinguished using polyclonal antisera and monoclonal antibodies (McAbs). Polyclonal antisera show only a very distant serological relationship between strains of these two viruses. BCMV- and

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BCMNV-specific antisera are available commercially for use in standard serological tests such as enzyme-linked immunosorbent assay (ELISA). Early work with BCMV and homologous antisera showed that BCMV was closely related serologically to AzMV, BlCMV, PStV, and several soybean infecting-isolates from Taiwan. As noted above, these viruses are now recognized as strains of BCMV. BCMV is distantly related to BYMV, Clover yellow vein virus (ClYVV), CABMV, PVY, Passion fruit woodiness virus (PWV), SMV, Tobacco etch virus (TEV), and WVMV. Several broad-spectrum McAbs have been identified that detect all of the strains of BCMV, including AzMV, BlCMV, DeMV, and PStV, as well as the BCMV subgroup member CABMV, but not with any strains of BCMNV. Other McAbs are reported which display various degrees of specificity for these viruses (e.g., can only detect CABMV or BCMV-PStV). While these McAbs can be used to discriminate among the BCMV isolates, biological diversity among these viruses may limit the use of many of these antibodies for diagnostic purposes. Most McAbs produced to BCMNV are specific only for strains of BCMNV (i.e., have no cross-reactivity with any strains of BCMV, nor other potyviruses). Another series of McAbs were raised against three strains of BCMV (AzMV, BlCMV, and PStV). Results of ELISA, immunosorbent electron microscopy, and western-blot analyzes indicated that these McAbs are specific for a series of epitopes located entirely on the virion surface. From western-blot analyzes of untreated and trypsin- as well as endopeptidase Lys-C-treated CPs, these virus-specific epitopes appear to be located exclusively on the amino-terminus of the CP. These McAbs were found to discriminate between strains of BCMV. These and other N-terminal targeted antibodies often enable clear distinction of strains in mixed infections, but a BCMV-PStV-specific McAb that reacts well with most BCMV-PStV blotch isolates failed to detect a necrotic isolate from Taiwan and several blotch isolates from Georgia, while a BCMNV-specific McAb misdiagnosed a naturally occurring genomic recombinant between BCMV and BCMNV.

Properties of the genome and replication The ssRNA genome, which has a VPg protein covalently linked to its 50 end, is about 10,000 nt for BCMV and 9600 nt for BCMNV. The organization of the BCMV and BCMNV genomes is similar to other potyviruses, consisting of short untranslatable sequences at the 50 and 30 ends, and a single, long open reading frame (ORF). The ORF is translated into a single polyprotein: c. 3222 aa for BCMV and c. 3071 aa for BCMNV. The polyprotein undergoes co- and post-translational proteolytic processing by three viralencoded proteinases to form ten individual gene products. Most of the polyprotein cleavage sites differ between BCMV and BCMNV, with the exception of the HC-Pro/P3 cleavage. The ten viral proteins (Fig. 2) include, in order from the 50 end of the genome, P1 proteinase; helper component-proteinase (HC-Pro); P3; a 6 kDa protein (6K1); cylindrical inclusion (CI); a second 6 kDa protein (6K2); the nuclear inclusion “a” (NIa)-VPg protein; NIa-proteinase; nuclear inclusion “b” (NIb); and the CP. An additional protein is translated by a frameshift within the P3 gene, resulting in production of a fusion protein, P3N-PIPO. Genomic RNA replicates via production of a full-length negative-sense RNA.

Pathogenicity, Pathology, and Resistance Genes Isolates of BCMV and BCMNV can be differentiated into at least nine pathotypes (0, and I through VIII) based on their reactions on a series of differential bean cultivars. In susceptible bean genotypes, BCMV and BCMNV induce highly similar symptoms including mosaic, dwarfing, chlorosis and leaf curling. The intensity and severity of the symptoms depend on various parameters including the virus strain, the bean cultivar, and the age and growing conditions (temperature) of the plant when infection occurs. The difference between the two viruses relies on the phenotypes generated on resistant cultivars. BCMV normally produces only mosaic symptoms in susceptible genotypes; BCMNV isolates can induce lethal systemic necrosis and plant death in bean cultivars carrying the incompletely dominant “I” gene. In most such cultivars the systemic necrosis develops at or above 201C; in hypersensitive genotypes the response is restricted to necrotic local lesions at normal temperatures, and systemic or “black root” symptoms only at temperatures above 301C. Hypersensitivity results in field resistance, and is genetically dominant; no seed transmission of BCMV occurs in genotypes with the dominant “I“ gene. Other resistance genes (bc-1, bc-12, bc-2, bc-22, bc-3, and bc-u) condition resistance to particular pathotypes in various combinations; the combination of the dominant I gene with the strain-specific recessive genes, bc-12, bc-22, or bc-u protects against systemic infection and seed transmission of BCMNV. In the absence of the recessive strain-specific resistance genes effective against necrosis-inducing isolates of BCMNV, the I gene alone is not sufficient to protect against systemic infection by BCMNV in many regions of Africa. The viruses carry various combinations of six pathogenicity determinants (P0, P1, P12, P2, P22, and Px), which to date have not been correlated with particular viral genes. Pathotype is not correlated with either CP or 30 UTR phylogeny, nor is there evidence for determinants controlling systemic movement in the 30 -terminal region. Evidence from

Fig. 2 Genome organization of BCMV and BCMNV. 50 , 50 UTR; P1, P1 proteinase; HC-Pro, helper component-proteinase; P3, P3 protein; P3N-PIPO; 6K1, a 6 kDa protein; CI, cylindrical inclusion; 6K2, a second 6 kDa protein; NIa, nuclear inclusion “a”, cleaved into VPg and Pro; VPg, genome-linked protein; Pro, proteinase; NIb, nuclear inclusion “b”; CP, coat protein; 30 ,30 -UTR. P3N-PIPO is expressed as a result of a frameshift within the P3 ORF.

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natural recombinants highlights the importance of the potyviral and the P1-HC-Pro precursor protein(s) in pathogenesis and the breakage of genetic resistance (see below). The incompletely dominant I gene was first identified in the cultivar “Corbett Refugee”, and the first recessive genes in “Robust” and “Great Northern 1” in the 1930s; strain-specificity of the recessive genes was identified in the 1950s, and the strain-unspecific gene bc-u shown in the 1970s to be necessary for expression of the strain-specific genes. Only one isolate has been identified so far that breaks resistance conferred by bc-3 (pathotype VIII); however, the gene originated in a single European breeding line, and it took many years to introgress bc-3 into genotypes adapted to the tropics. Cultivars carrying the dominant I gene may have increased susceptibility to Cowpea severe mosaic virus. There is genetic linkage between certain seed colors and BCMV susceptibility, including red-mottled seed coat; the dominant I gene is closely linked to darker purple-mottled seed color. The lighter seed color associated with susceptibility is preferred in many cultures and commercial uses of the beans; this linkage can be partially avoided by selecting resistant types with commercially acceptable intermediate seed colors. Introgression of bc-3 can be problematic, as lines carrying the gene can be symptomless; serological or infectivity assays may be necessary to differentiate resistant and susceptible lines. Introgression of I and bc-3 has become simpler with development of molecular markers associated with each gene, allowing marker-assisted selection without virus challenge and phenotypic evaluation. The I gene is closely linked to a multigene family of TIR-NBS-LRR resistance genes conferring resistance to several related potyviruses and a comovirus. In Latin America most black-seeded bean cultivars carry the I gene, and many additional bean lines bred at Centro International de Agricultura Tropical and associated national programs also carry resistance. Many land races grown by traditional farmers, and particularly beans with light-colored seed coats, are susceptible; infection rates of up to 100%, and yield losses of 35%–98% are reported. In North America many bean varieties have effective resistance to BCMV, although many pinto, navy, and red-seeded types have only strain-specific resistance. However, recent breeding has yielded pinto and navy bean cultivars carrying both I and bc-3. BCMNV was previously rare in the US, but combined resistance to BCMV and BCMNV is now available in some cultivars of most bean types. Many of the beans grown by small farmers in eastern and southern Africa are susceptible to both BCMV and BCMNV. The presence of necrosis-inducing strains of BCMNV, also found in wild legumes and species grown for fodder, means that the I gene cannot be deployed independent of recessive resistance genes, as infection by necrosis-inducing isolates could result in death of a significant proportion of plants in a field. In India infections of up to 100% are reported in bean, urdbean, mungbean, and cowpea, resulting in mottle, mosaic, blistering, stunting, and poor pod set. Many Asian bean types also lack resistance, due to linkage between susceptibility and desirable seed colors. Peanut stripe strains of BCMV causing symptoms described as blotch, blotch-CP-N, blotch stripe, chlorotic line pattern, chlorotic ring mottle, mild mottle, necrosis, and stripe have also been differentiated on the basis of disease reactions on specific host genotypes. Losses of 23%–38% to peanut mild mottle have been documented in China. Peptide profiles of five symptom variants from Thailand that were not serologically distinct confirmed close relationships. No resistance was identified among over 8000 cultivated peanut genotypes in the germplasm collection at the International Crops Research Institute for the Semi-Arid Tropics. Some wild Arachis species have been identified as immune or highly resistant, including accession PI475998 of A. cardenasii, which could not be infected. Resistance genes have been identified and incorporated into resistant cultivars of Pisum sativum, Lupinus angustifolius, Lablab purpureus, and Macroptilium lathyroides. Transgenic resistance has been demonstrated in peanut transformed with the CP gene of a peanut stripe isolate; a high level of resistance was observed in lines expressing either untranslatable or N-terminally truncated forms of the CP gene, suggesting that resistance resulted from RNA silencing (RNAi). While beans have been transformed for resistance to Bean golden mosaic virus (Geminivirus), and soybean for resistance to Soybean mosaic virus (Potyvirus), there are no reports of transgenic BCMV- or BCMNVresistant bean lines.

Recombination and Variability Although recombination between different isolates of BCMV had been reported for some years, no recombinants between BCMV and BCMNV were reported until recently. No recombinants were recovered from deliberate mixed infections of BCMV strain US-5 (pathogroup IV) and BCMNV strain NL-8 (pathogroup III) following many serial passages in either of two susceptible hosts. However, multiple recombinants were recovered within 28 days when the same strains were inoculated on opposite primary leaves of beans that were susceptible to one virus and resistant to the other; these recombinants fell into five different classes based on combinations of serotype (A, BCMNV; or B, BCMV) and pathogroup (IV, V, or VI). Subsequently, a naturally occurring strain of BCMNV (NL-3 K) with atypical responses on differential hosts was sequenced and shown to have a 50 -UTR and 50 region of the P1 gene almost identical to BCMV strain RU1, while the rest of the genome was almost identical to BCMNV strain NL-3 D. The P1 gene of the recombinant was larger than that of BCMNV, and similar to the BCMV P1. These results suggest that the P1 gene may play a significant role in pathogenicity and virulence, as previously suggested for BYMV. The observed phenotypes of recombinants indicate that serotype, pathogenicity, and symptom expression are independent traits with separate determinants in the genome. The occurrence of recombination in bean genotypes with only recessive, strainspecific resistance genes suggests that this may be the primary cause of emergence of new resistance-breaking strains. Much of the variability between isolates of either BCMV or BCMNV appears to be within the 50 portion of the genome, as has been shown both within and between other potyviruses. The P1 and P3 genes, and the N-terminal region of the CP gene are the most variable regions of the potyvirus genome, and may be involved in host–virus interactions. Differences in the 50 UTR and

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N-terminus of P1 of both BCMV and BCMNV have been associated with symptom differences in common bean (P. vulgaris), and for BCMV in asparagus bean (V. sesquipedalis). Variability in the P1 gene is extensive, with as little as 61% identity at the aa level between BCMV strains, with most of the differences in the N-terminal domain. Differences in the CP gene result in clustering of BCMV isolates according to host species (bean, cowpea, peanut, asparagus bean, guar, Dendrobium, etc.) and to some extent by geographic origin within host clusters; peanut stripe isolates from China, Indonesia, and Thailand form separate clusters.

Diagnosis Host range and symptomatology Potyviruses can be detected and identified by a variety of techniques that are based on biological, cytological, biochemical, antigenic, and structural properties. With respect to BCMV and BCMNV, host range and symptomatology are not very useful because there are many viruses that infect bean, and significant differences in symptoms may be reproduced depending on the virus pathotype host genotype, and environmental conditions. Morphology of cytoplasmic inclusions has been useful in the identification of many potyviruses since inclusions can be virus-specific. However, inclusion morphology is not reliable for detection and differentiation of BCMV and BCMNV. BCMNV induces pinwheel and scroll types of cytoplasmic inclusions (subdivision I) in infected tissue. Most strains of BCMV, including BCMV-AzMV and BCMV-BlCMV, also induce subdivision I type inclusions. However, PStV and Taiwan soybean isolates induce subdivision IV cylindrical inclusions.

Serological techniques BCMV- and BCMNV-specific polyclonal antisera and McAbs are useful for the detection and differentiation of these seed-borne viruses, and several methods have been used for detection in bean seed. When indirect ELISA and a dot immunoassay (DIA) with both McAbs and polyclonal antisera were evaluated, ELISA with McAbs proved the most sensitive method. However, BCMV antigen was found to be erratically distributed within the seed, with more than 50% of infected samples having detectable levels of BCMV antigen only in the seed coat. Tests on individual seeds proved unreliable and it is recommended that flour from a bulked sample of eight seeds be used when testing seed lots. Other assays include immunosorbent electron microscopy to detect BCMV in bean seed; or double antibody sandwich (DAS)-ELISA and bioassay on Chenopodium quinoa to test hydrated mature cowpea seed for presence of BCMV-BlCMV. In these tests infectious virus was detected in cotyledons and sometimes in the embryonic axis of infected seed. Viral antigen was found in or on the testae, but very little infectious virus was present. Others have used direct antigen-coated-ELISA to detect BCMV-BlCMV in cowpea seed.

Reverse transcription-polymerase chain reaction The sensitivity, capacity, and potential of the PCR technique has been applied to BCMV and BCMNV. Several sets of BCMV- and BCMNV-specific RT-PCR primers have been designed and utilized to sensitively and accurately detect, differentiate, and characterize the two viral species. In one study using virus-specific CP primers, the specific geographical distribution of these viruses in Mexico was determined. In addition, the alignment of nine nucleotide sequences from cloned amplicons from the RT-PCR reactions for each viral species confirmed the identities of the viruses and was helpful in assigning them tentatively to pathogroups. Other primer sets have been developed and utilized, for example, in the successful detection and differentiation of the necrotic and blotch isolates of BCMV-PStV, and in the differentiation of naturally occurring genomic BCMV/BCMNV recombinants. Considering the seed-borne nature of these viruses, RT-PCR has potential in quarantine programs for screening imported seed material and germplasm.

Prevention and Control It is possible to eradicate BCMV and BCMNV from a particular production region, or at least to significantly reduce primary inoculum, by eliminating susceptible genotypes and planting only resistant varieties. There are few important non-crop reservoirs of inoculum, except in eastern and southern Africa, and infected seed are responsible for primary infections in the crop. Rogueing of symptomatic plants is not recommended as a means of control, as it is highly likely that systemically infected plants without significant symptoms will remain to act as significant reservoirs for secondary infection by aphid transmission. Exclusion of the virus is highly effective in areas where the virus is not present, but this may preclude introduction of new germplasm or cultivars without effective resistance. One or both viruses are almost universally present in seed lots of varieties that do not carry the I gene, and the majority of Latin American land race types do not possess any of the strain-specific recessive resistance genes. Most cultivars derived through extensive breeding programs carry the dominant I gene for resistance to BCMV and BCMNV. Increasingly bc-3 is being combined with I to yield cultivars with effective resistance to both BCMV and BCMNV. Certified seed can be an effective means of control. A certified seed program in California with a limit of o0.5% BCMV has aided in the control of the disease, as resistant varieties are not yet available for all bean classes currently grown in California. Monitoring of BCMV for certification also aided in identification and containment of an outbreak of BCMNV introduced in seed of a navy bean cultivar. Vector exclusion may aid in control, but as aphids rarely colonize common beans, migratory aphids are responsible for most transmission in many areas. As BCMV and BCMNV can be acquired and transmitted in less than 1 min access to plants, insecticide

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treatment is unlikely to prevent, and may even encourage vector movement and transmission. Vector control through insecticide application may be effective in parts of Africa and Asia where aphids do colonize beans. Oil sprays are known to reduce transmission of stylet-borne viruses such as BCMV and BCMNV, but their lack of systemic action, and thus need for frequent reapplication to a growing crop, reduces the cost-effectiveness of their use.

See also: Recombination

Further Reading Adams, M.J., Antoniw, J.F., Fauquet, C.M., 2005. Molecular criteria for genus and species discrimination within the family Potyviridae. Archives of Virology 150, 459–479. Berger, P.H., Wyatt, S.D., Shiel, P.J., et al., 1997. Phylogenetic analysis of the Potyviridae with emphasis on legume-infecting potyviruses. Archives of Virology 142, 1979–1999. Chung, B.Y., Miller, W.A., Atkins, J.F., Firth, A.E., 2008. An overlapping essential gene in the Potyviridae. Proceedings of the National Academy of Sciences of the United States of America 105, 5897–5902. Dijkstra, J., Khan, J.A., 1992. A proposal for a Bean common mosaic subgroup of potyviruses. In: Barnett, O.W. (Ed.), Potyvirus Taxonomy. Archives of Virology, Suppl. 5 Springer, pp. 389–395. Drijfhout, E., 1978. Genetic interaction between Phaseolus vulgaris and Bean common mosaic virus with implications for strain identification and breeding for resistance. Agricultural Research Reports No. 872. Wageningen: Center for Agricultural Publishing and Documentation. Feng, X., Myers, J.R., Karasev, A.V., 2015. Bean common mosaic virus isolate exhibits a novel pathogenicity profile in common bean, overcoming the bc-3 resistance allele coding for the mutated eIF4E translation initiation factor. Phytopathology 105, 1487–1495. Gibbs, A.J., Ohshima, K., 2010. Potyviruses and the digital revolution. Annual Review of Phytopathology 48, 205–223. Higgins, C.M., Hall, R.M., Mitter, N., Cruickshank, A., Dietzgen, R.G., 2004. Peanut stripe potyvirus resistance in peanut (Arachis hypogea L.) plants carrying viral coat protein gene sequences. Transgenic Research 13, 59–67. Larsen, R.C., Miklas, P.N., Druffel, K.L., Wyatt, S.D., 2005. NL-3 K strain is a stable and naturally occurring interspecific recombinant derived from Bean common mosaic necrosis virus and Bean common mosaic virus. Phytopathology 95, 1037–1042. McKern, N.M., Ward, C.W., Shukla, D.D., 1992. Strains of bean common mosaic virus consist of at least two distinct potyviruses. In: Barnett, O.W. (Ed.), Potyvirus Taxonomy. Archives of Virology. Springer, pp. 407–414. (Suppl. 5). Meziadia, C., Blanche, S., Geffroy, V., Pflieger, S., 2017. Genetic resistance against viruses in Phaseolus vulgaris L.: State of the art and future prospects. Plant Science 265, 39–50. Milne, R.G. (Ed.), 1988. The Filamentous Plant Viruses. The Plant Viruses, vol. 4. New York: Plenum. Mink, G.I., Silbernagel, M.J., 1992. Serological and biological relationships among viruses in the Bean common mosaic virus subgroup. In: Barnett, O.W. (Ed.), Potyvirus Taxonomy. Archives of Virology. Springer, pp. 397–406. (Suppl. 5). Morales, F.J., 1998. Present status of controlling bean common mosaic virus. In: Hadidi, A., Kheterpal, R.K., Koganezawa, H. (Eds.), Plant Virus Disease Control. St. Paul: APS Press, pp. 524–533. Provvidenti, R., Hampton, R.O., 1992. Sources of resistance to viruses in the Potyviridae. In: Barnett, O.W. (Ed.), Potyvirus Taxonomy. Archives of Virology. Springer, pp. 189–211. (Suppl. 5). Sengooba, T.N., Spence, N.J., Walkey, D.G.A., Allen, D.J., Femi Lana, A., 1997. The occurrence of bean common mosaic necrosis virus in wild and forage legumes in Uganda. Plant Pathology 46, 95–103. Shukla, D.D., Ward, C.W., Brunt, A.A., 1994. The Potyviridae. Wallingford: CAB International. Vallejos, C.E., Astua-Monge, G., Jones, V., et al., 2006. Genetic and molecular characterization of the I locus of Phaseolus vulgaris. Genetics 172, 1229–1242. Vetten, H.J., Lesemann, D.-E., Maiss, E., 1992. Serotype A and B strains of bean common mosaic virus are two distinct potyviruses. In: Barnett, O.W. (Ed.), Potyvirus Taxonomy. Archives of Virology. Springer, pp. 415–431. (Suppl. 5). Worrall, E.A., Wamonje, F.O., Mukeshimana, G., et al., 2015. Bean common mosaic virus and bean common mosaic necrosis virus: Relationships, biology, and prospects for control. Advances in Virus Research 93, 1–46. Wylie, S.J., Adams, M., Chalam, C., et al., 2017. ICTV virus taxonomy profile: Potyviridae. Journal of General Virology 98 (3), 352–354. (accessed June 6, 2019). Available at: https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/w/potyviridae.

Bean Golden Mosaic Virus and Bean Golden Yellow Mosaic Virus (Geminiviridae) Francisco M Zerbini, Federal University of Viçosa, Viçosa, Brazil Simone G Ribeiro, Embrapa Genetic Resources and Biotechnology, Brasília, Brazil r 2021 Elsevier Ltd. All rights reserved. This is an update of F.J. Morales, Bean Golden Mosaic Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00695-6.

Glossary HTS High throughput sequencing technologies, refer to methods that are able of sequencing multiple molecules in parallel, allowing hundreds of millions of molecules to be sequenced at a time. Hybridization probe Labeled DNA or RNA molecule (radioactively or otherwise) that will hybridize with nucleic acids with complementary sequence and will detect their presence due to the label. PCR Polymerase chain reaction is a technique used to make multiple (millions to billions) copies of a DNA segment in vitro using a thermostable DNA polymerase. Resistance Relative ability of an organism to restrict the growth and attenuate the effects of a pathogen.

RFLP Restriction fragment length polymorphism is a method to examine the genetic variation of individuals by analyzing the pattern of the fragments generated by the digestion of a specific DNA segment with endonucleases. RNA silencing A conserved evolutionary process active in a wide variety of eukaryotic organisms that can lead to inhibition of transcription or translation of a target gene in a sequence-specific way. RNAi A process to control gene function by harnessing the expression of a dsRNA corresponding to the sequence of a target gene. The dsRNA triggers the RNA silencing mechanism resulting in the production of small RNAs that will induce the degradation of mRNA in a sequence-specific manner, hence blocking protein production.

Introduction The common bean (Phaseolus vulgaris L.) is one of the most widely cultivated legume crops in the world. Its center of origin is located in the Americas, with two major domesticated gene pools known as Andean (from the highlands of Bolivia, Ecuador and Peru) and Mesoamerican (from Mexico and Central America). Common bean is a source of carbohydrates, protein and fiber and is a major component of the human diet in Latin America, Africa and Southeast Asia. As of 2016, the world’s largest producers were Myanmar, India and Brazil. In the Americas the largest producers are Brazil, the United States and Mexico, but common bean is a major crop in every country in Central and South America. Bean golden mosaic was first reported in the mid-1960s in Brazil by A.S. Costa as a whitefly (Bemisia tabaci Genn.)-transmitted disease. Similar diseases were described in several Central American and Caribbean countries during the late 1960s and early 1970s, and were named either “golden mosaic” or “golden yellow mosaic”. In all countries from which it was reported, the disease was associated with an increase in the population of B. tabaci. The similar symptoms and association with the same vector led to the assumption that the same causal agent was responsible for the disease in all countries. Plants affected by the disease usually display striking yellow mosaic symptoms (Fig. 1), named “golden” mosaic to differentiate from the less severe common mosaic (which is usually associated with mosaic and blistering symptoms without yellowing) and yellow mosaic diseases. In reality, it may be difficult to distinguish these three diseases based solely on symptoms in the field. Additional symptoms of bean golden mosaic may include downward leaf curling, leaf distortion and stunting. A significant level of flower abortion may occur in warmer climates, contributing to yield losses. When first reported, bean golden mosaic was considered to be of “insignificant” economic importance. However, in less than ten years it became the main disease of common bean during the dry season (Jun–Sep) in south–central Brazil, mostly due to the exponential growth in the area planted with soybean, an excellent host of B. tabaci. Following the harvest of soybean (Feb–May), whitefly populations migrate massively to recently planted common bean fields, and the disease causes extensive losses. As soybean became a major crop in Brazil during the 1970s, bean golden mosaic reached alarming levels and the country’s bean production was greatly affected. A similar scenario occurred in Argentina. Bean golden mosaic remains a major disease in these countries and the bean industry underwent several changes as a result, including relocation to regions which are less favorable to the disease and the large-scale adoption of central pivot irrigation (which eventually led to the emergence of soil-borne diseases such as white mold). The disease also continues to be of great relevance in other Latin American countries, where the increase in whitefly populations is associated with poor management practices leading to the emergence of insecticide-resistant populations and climate change. Ultrathin sections of leaf tissue obtained from golden mosaic-affected bean plants in Brazil revealed the presence of isometric particles (20–25 nm) restricted to vascular tissues. Probably due to its phloem-restriction, the causal agent of bean golden mosaic

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Fig. 1 Bright yellow mosaic (“golden” mosaic) symptoms in common bean (Phaseolus vulgaris) caused by Bean golden mosaic virus (BGMV). Photograph: R. Ramos-Sobrinho.

Fig. 2 Characteristic geminate particles of BGYMV. Bar ¼ 20 nm.

in Brazil could not be transmitted by mechanical means or by seed. Conversely, the virus associated with the disease in Puerto Rico was readily sap-transmissible. This was an early indication that the similar diseases occurring in Brazil and in Central America/ Caribbean were caused by distinct, albeit related, viruses. The isolation of whitefly-transmitted viruses that induced golden mosaic symptoms in common bean was achieved in 1976 by G. Galvez and M. Castaño at CIAT, Colombia, using a bean golden yellow mosaic virus (BGYMV) isolate from El Salvador, and in 1977 by R. Goodman at the University of Illinois, USA, using a BGYMV isolate from Puerto Rico. In both cases the isolated viruses had quasi-isometric particles with a unique geminate morphology (Fig. 2). The isolate from Puerto Rico was used to characterize the viral nucleic acid. Strikingly, the viral nucleic acid was identified as a single-stranded (ss) DNA molecule, the first reported ssDNA virus in plants. In 1981, the Puerto Rican BGYMV isolate was shown to possess a two-component ssDNA genome, with each molecule (ca. 2600 nt) separately encapsidated in a geminate particle. In 1978, BGMV from Brazil and other BGMV/BGYMV-like viruses, as well as other similar ssDNA viruses transmitted by B. tabaci or leafhoppers (Homoptera: Cicadellidae) were included in the newly created Geminivirus group. BGMV was classified as a “subgroup II” geminivirus (whitefly-transmitted viruses with bipartite genomes), while the leafhopper-transmitted viruses with

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monopartite genomes (such as Maize streak virus) were classified as “subgroup I” geminiviruses. In 1995 the leafhoppertransmitted geminiviruses were further divided into two subgroups, and BGMV became a member of “subgroup III” geminiviruses. Studies performed by R. L. Gilbertson and D. Maxwell at the University of Wisconsin, USA in the early 1990s using BGMV isolates from Brazil and Central American/Caribbean countries provided definitive proof that two distinct geminiviruses were associated with bean golden mosaic disease in these regions. Broad-spectrum DNA probes were obtained which were capable of detecting all isolates tested (BGMV from Brazil, Guatemala and the Dominican Republic, plus an isolate of bean dwarf virus from Colombia) under low stringency hybridization conditions. A specific DNA probe was also prepared which detected only the isolate from Brazil, and another probe detected the isolates from Guatemala and the Dominican Republic (these two isolates could not be differentiated by nucleic acid hybridization even at high stringency hybridization conditions). These assays provided strong evidence that “BGMV” isolates from Brazil and Central America/Caribbean were actually distinct viruses. In the same year, a polymerase chain reaction (PCR)-based assay was used to obtain nucleotide sequences of a hypervariable region of the viral genome (the intergenic region between the MP gene and the common region of the DNA-B component). Nucleotide sequence identities between different BGMV isolates from the Dominican Republic (95% sequence identity amongst them), Guatemala (86%), BGYMV from Puerto Rico (75%) and BGMV from Brazil (46%) further suggested the existence of significant genetic variability, particularly between the Central American/Caribbean geminiviruses and the Brazilian BGMV isolate. In 1993, the complete nucleotide sequences of the Brazilian and Puerto Rican isolates were obtained. The percent nucleotide identities between these two viruses ranged between 71% and 82%, the latter value corresponding to the coat protein gene, which is the most conserved gene among whitefly-transmitted geminiviruses. These results clearly demonstrated that BGMV from Brazil was distinct from BGYMV from Puerto Rico. The former was classified as “type I” BGMV, and the latter as “type II” BGMV. In the same year, the isolates from Guatemala and the Dominican Republic were sequenced and shown to belong to type II BGMV, together with the isolate from Puerto Rico. In 2000 the Geminivirus group became the family Geminiviridae, including the genus Begomovirus (from bean golden mosaic). Type I BGMV isolates were classified as members of the species Bean golden mosaic virus, while type II BGMV isolates were classified as members of the species Bean golden yellow mosaic virus. The bean golden mosaic disease observed in northwestern Mexico was shown in 1992 to be caused by another distinct begomovirus originally isolated in 1981 from squash in southwestern United States, named squash leaf curl virus (SLCV). Some SLCV isolates sampled from diseased common beans in Mexico had already changed significantly and were thus classified as members of a distinct species named Bean calico mosaic virus. However, SLCV can still be found in northwestern Mexico causing bean calico mosaic, a severe disease that often progresses from the characteristic golden mosaic into severe foliar bleaching and plant death. In Nicaragua, bean golden mosaic was shown to be caused predominantly by BGYMV, but also by squash yellow mild mottle virus and Calopogonium golden mosaic virus. In 2010, a begomovirus detected in Macroptilium lathyroides and named Macroptilium yellow spot virus (MaYSV) was shown to infect common bean in the northeastern Brazilian state of Alagoas, inducing golden mosaic symptoms indistinguishable from those caused by BGMV and BGYMV. In a large-scale survey carried out in 2011–2012, MaYSV was the predominant begomovirus detected in common bean in that state, but was not detected in the midwestern states of Goiás and Minas Gerais. More recent surveys detected MaYSV in the states of Bahia and Pernambuco, also as the predominant begomovirus in common bean. The emergence of MaYSV adds an additional layer of complexity to the scenario of bean golden mosaic in Brazil.

Taxonomy, Phylogeny, and Evolution Currently, viruses known to cause bean golden mosaic disease are classified in three species of the genus Begomovirus (family Geminiviridae): Bean golden mosaic virus, Bean golden yellow mosaic virus, and Macroptilium yellow spot virus. Isolates of Bean golden mosaic virus (BGMV) are found in Brazil and Argentina, while isolates of Bean golden yellow mosaic virus (BGYMV) are found throughout Central America and the Caribbean. It is remarkable that after more than 50 years since bean golden mosaic has been described, BGMV and BGYMV remain geographically isolated. This is probably due to the presence of large geographical barriers and the lack of seed transmissibility of both viruses. Isolates of Macroptilium yellow spot virus (MaYSV) have only been found in the northeastern Brazilian states of Alagoas, Bahia and Pernambuco. Phylogenetically, begomoviruses are neatly divided into Old World and New World clades. Both BGMV and BGYMV are New World (NW) begomoviruses, but within the NW clade they are distantly related. Consistent with the geography-based (rather than host-based) grouping of begomoviruses, BGMV is phylogenetically close to other begomoviruses found in Brazil, such as tomato golden mosaic virus (TGMV) and Sida micrantha mosaic virus (SiMMV), while BGYMV is closer to other begomoviruses found in Central America and the Caribbean such as Macroptilium yellow mosaic virus (MaYMV). In 1999, samples of common bean (P. vulgaris), lima bean (P. lunatus) and the weed Leonurus sibiricus were collected in beangrowing areas of Brazil, and viral fragments encompassing part of the Rep gene, the common region and part of the CP gene were amplified, cloned and sequenced. The L. sibiricus samples were infected by a distinct virus eventually named tomato yellow spot virus, which also infects tomato and common bean but is not associated with bean golden mosaic. All sequences obtained from common bean samples were closely related to the original BGMV isolate, with o7% nucleotide sequence divergence. This was low degree of genetic variability (unlike what is found for other begomoviruses) was attributed to the lack of BGMV-resistant bean cultivars that could place selection pressure upon the viral population. A second study in 2014 confirmed the low genetic variability of BGMV populations. This study compared BGMV and MaYSV populations obtained from three hosts (common bean,

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lima bean and M. lathyroides), and reported a much higher degree of genetic variation in MaYSV isolates compared to BGMV. This was explained by numerous recombination events in MaYSV. Recombination was much less frequent in BGMV populations. Interestingly, the genetic variability of both BGMV and MaYSV was evenly distributed among isolates infecting the different hosts, indicating that genetic variability is an intrinsic viral property which is not affected by the host. Nevertheless, BGMV and MaYSV populations showed distinct distributions of genetic variation, with the BGMV population (but not the MaYSV population) being strongly structured by both host and geography, suggesting ongoing speciation. Purifying (negative) selection is the dominant force acting on BGMV populations. Positive selection is only rarely observed, and some positively selected sites in the coat protein gene could be associated with improving whitefly transmission. No evolutionary studies have been carried out with BGYMV populations.

Virion Structure, Genome Organization, Properties and Functions of Gene Products, Replication and Propagation Geminivirus particles are formed from two isometric, T ¼ 1 capsids fused to form a geminate particle (18  30 nm), from which the family derives its name. The geminate capsid is formed by assembly of 110 subunits of a single CP, arranged as 22 pentamers (i.e., each T ¼ 1 capsid is missing one pentamer). Virion structure has not been determined in detail for BGMV or BGYMV, but it is assumed to be similar to that determined for two other begomoviruses, African cassava mosaic virus (ACMV) and Ageratum yellow vein virus (AYVV). The CP would assume three distinct conformations, facilitating the formation of the interface between the two capsids. The solution structure of the AYVV capsid confirmed the long-standing assumption that each DNA component is encapsidated in a separate geminate particle. Complete DNA sequences of several BGMV and BGYMV isolates were determined in the late 1980s/early 1990s, and genomic organizations are essentially identical for both viruses. Thus, only the one for the Brazilian isolate of BGMV will be described in detail. The DNA-A (RefSeq accession number NC_004042, derived from M88686) and DNA-B (NC_004043, derived from M88687) are 2617 and 2580 nt long, respectively. The two components share no significant sequence identity, except for an approx. 180-nt long region known as the common region (CR), which is highly conserved (497% identity) between the two components. The CR sequence includes direct and inverted repeats that form a stem-loop structure, with the conserved 50 -TAATATTAC-30 nona-nucleotide in the loop. In addition, two direct repeats and one inverted repeat, known as iterons (high-affinity binding sites for the Rep protein), are present upstream from the stem-loop structure. The DNA-A has five open reading frames (ORFs): one in the viral sense (CP; previously known as AV1 or AR1) and four in the complementary sense [Rep (AC1 or AL1), TrAP (AC2 or AL2), REn (AC3 or AL3) and AC4] (Fig. 3). The complementary sense ORFs are located in different reading frames, with the 30 -end of Rep overlapping the 50 -end of TrAP, and TrAP overlapping the 50 and central regions of REn. The Rep ORF overlaps the AC4 ORF in its entirety. CP and REn overlap in opposite directions in an AT-rich region and share four nucleotides at their C-termini, which contain their translational stop codons. The DNA-B contains two non-overlapping ORFs: one in the viral sense, the nuclear shuttle protein (NSP, also BV1 or BR1), and one in the complementary sense, the cell-to-cell movement protein (MP, also BC1 or BL1). All ORFs except REn and MP have TATA boxes within 100 nt of the start codon. Polyadenylation signals are located within the AT-rich regions at or near the 30 -ends of the CP, REn and NSP ORFs. The Rep, TrAP and REn ORFs are assumed to share a single polyadenylation signal located inside the 30 -end of REn, analogous to what was shown for TGMV. The four complementary-sense genes in the DNA-A are involved in DNA replication (Rep and REn), suppression of host defenses (TrAP and AC4) and trans-activation of the viral-sense ORFs in both the DNA-A and DNA-B (TrAP). The CP, besides its structural role, is also necessary for whitefly transmission. Together, the two DNA-B-encoded proteins coordinate viral DNA movement from the nucleus to the cytoplasm (NSP) and cell-to-cell via plasmodesmata (MP).

Fig. 3 Genome and virion structure of Bean golden mosaic virus (BGMV). The circles represent the two genomic components (DNA-A and DNA-B). The CP and NSP open reading frames are located in the viral-sense strand. Rep, TrAP, REn, AC4, and MP are located in the complementary-sense strand. CR, common region. Each genomic component is encapsidated in a separate geminate particle, as indicated below the genome structures. Thus, a virion is comprised of two geminate particles, one containing the DNA-A and the other containing the DNA-B.

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Bipartite begomoviruses utilize both rolling-circle replication and recombination-dependent replication strategies. The common region contains the origin of rolling-circle replication, which is the conserved nona-nucleotide located in the stemloop structure. The replication-associated protein (Rep) functions as a specific DNA binding protein that acts as both nuclease and ligase in the initiation and termination of the rolling-circle replication process. In the infection process of begomoviruses, virus particles are inoculated into the plant by means of the insect vector, and the viral genome (ssDNA) disassociates spontaneously from the capsid. Viral DNA is then transported to the host cell nucleus, where the ssDNA is converted to a double-stranded dsDNA, referred to as the replicative form (RF). Indirect evidence, such as the need for local destabilization of dsDNA for initiation of rolling-circle replication in prokaryotes by strand-nicking enzymes, indicates that this is performed by host factors. The RF serves as a template for synthesis of new ssDNAs and viral mRNAs. The viral genome is replicated by the rolling-circle mechanism, similar to that used by bacteriophages phiX174 and M13, using the RF as a template. The origin of replication is the conserved nona-nucleotide (50 -TAATATTAC-30 ) located in the stem-loop structure in common region. Origin recognition is mediated by the high-affinity binding sites (iterons) located upstream of the stem-loop. After binding of Rep to viral DNA and stabilization of the complex formed by Rep, Ren and host factors, the Rep protein cleaves the nona-nucleotide (TAATATT//AC), initiating rolling-circle replication. Rep is the only viral protein which is essential for replication, with multiple roles. It functions as a DNA binding protein with sequence and structural requirements, and also as a nuclease and DNA ligase in the initiation and termination of the rolling-circle replication process (cleavage of the nona-nucleotide at the origin of replication; cleavage of the linear, multimeric DNA; ligation of the monomeric genome units). As shown for other begomoviruses (Abutilon mosaic virus, African cassava mosaic virus, tomato golden mosaic virus and tomato yellow leaf curl virus), replication of BGMV and BGYMV may involve a recombination-dependent mechanism. In this mechanism, fragments of viral DNA are recovered from incomplete synthesis or from nucleolytic attack to create recombinant viruses.

Transmission, Host Range, and Epidemiology BGMV and BGYMV have narrow host ranges. Host species are mostly restricted to the Fabaceae for both viruses. Among crop species, only Phaseolus spp. such as common bean (P. vulgaris) and lima bean (P. lunatus) are of significance. A.S. Costa and A.M. Almeida reported in 1979 that certain soybean (Glycine max) cultivars in Brazil are susceptible to BGMV. These susceptible cultivars are not considered important as virus sources for bean crops, but may probably serve as virus reservoirs between two consecutive bean plantings that coincide with the beginning and the end of the soybean growing season. The low degree of genetic variability of BGMV populations, which does not favor the emergence of new virus variants, may be one of the reasons why BGMV has never acquired economic relevance in soybean (in Brazil or in other countries). Non-cultivated hosts of BGMV and BGYMV include Macroptilium spp., Calopogonium and wild Phaseolus spp. Early reports of BGMV infection in malvaceous and solanaceous hosts were probably due to mis-identification of the virus, and have not been confirmed since the use of molecular tools for virus identification became routine. It is noteworthy that fabaceous plant species used as cover crops, such as Crotalaria spp. and velvet bean (Mucuna aterrina), are non-hosts. BGMV and BGYMV are not seedborne in any known host. BGMV from South America is not mechanically transmitted, while BGYMV from Central America and the Caribbean can be transmitted in this way with relative ease. Nevertheless, mechanical transmission is not relevant as far as virus spread in the field is concerned. The only significant form of natural transmission of both viruses in the insect vector, the whitefly B. tabaci. Populations of this insect have always been recognized as genetically complex. In the past, B. tabaci was divided into biotypes. These are now recognized as distinct species, morphologically indistinguishable but with reproductive isolation. Up until the late-1980s, only indigenous species of B. tabaci such as New World 1 (NW1, previously known as biotype A) and New World 2, were present in the Americas. During the 1990s, multiple introductions of an aggressive species, B. tabaci Middle East-Asia Minor 1 (MEAM1, previously known as biotype B), occurred in the continent. The dissemination of B. tabaci MEAM1 brought dramatic changes in the incidence of begomoviruses in solanaceous crops, which are not normally colonized by B. tabaci NW1 but are excellent hosts for MEAM1. Although not to the same extent, MEAM1 also impacted bean golden mosaic disease. Both BGMV and BGYMV are transmitted with similar efficiencies by indigenous species and by MEAM1, but MEAM1 has a shorter life cycle and increased reproductive efficiency compared to NW1/2. Thus, MEAM1 populations reach much higher levels than NW1/2 populations, and virus transmission occurs at even higher levels. A second invasive species, B. tabaci Mediterranean (MED, previously known as biotype Q) has recently been introduced to South America, and is now present in southern and southeastern Brazil. The consequence of the presence of this species in the dissemination of bean golden mosaic disease is unknown, but a recent report indicated a higher efficiency of BGMV transmission by MED compared to MEAM1. Bean golden mosaic, like other whitefly-transmitted diseases, is more severe under conditions of warm temperatures and low rainfall. In south-central Brazil, temperatures are warm during most of the year, and a pronounced dry season lasts from late June until late September. In addition, soybean crops are harvested from February until May, with massive migration of whitefly populations from soybean to other crops. As a result, June-Sep is the time of the year with the greater incidence and severity of bean golden mosaic in this region.

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Virus-Host Relationships BGMV was the object of pioneer studies on the cytopathology of geminiviruses. The 1978 work by K.S. Kim at the University of Arkansas and T. Shock and R. Goodman at the University of Illinois showed that these viruses induce major changes in the nuclear structure of infected cells, which resemble the cytopathology of animal cells infected with parvoviruses. The observed changes included the hypertrophy of the nucleolus, which eventually occupies most of the nuclear volume; the segregation of nucleolar components into granular and fibrillar regions composed of ribonucleoprotein; the presence of fibrillar rings in multiple numbers and sizes, which have been shown to contain DNA; and the appearance of viral particles either as loosely compacted aggregates or in hexagonally close-packed arrays. Virus particle aggregates were observed only in the nuclei of infected phloem cells.

Diagnosis The correct identification of the virus infecting a plant is crucial for the selection of effective management and control procedures of the disease in the field. Bean golden mosaic disease, which is characterized by yellow and golden mosaic, leaf distortion, flower abortion, pod deformities, and stunting of plants is caused mainly by BGYMV in North and Central America and the Caribbean and by BGMV in South America. However, similar symptoms may also be induced in beans by about other 20 begomoviruses, for example, MaYSV in northeastern Brazil, tomato yellow vein streak virus in northwestern Argentina, Calopogonium golden mosaic virus in Costa Rica and Nicaragua, and also viruses from other genera and families such as alfalfa mosaic virus (fam. Bromoviridae, gen. Alfamovirus), bean yellow mosaic virus (Potyviridae, Potyvirus) and cowpea chlorotic mottle virus (Betaflexiviridae, Carlavirus) which may be present in different areas and countries. Therefore, it is not possible to identify the virus solely by relying on the type of symptoms displayed by the plant. Different diagnostic methods may be used for begomovirus detection in bean plants. Even though antibody-based tests such as ELISA (enzyme-linked immunosorbent assay) and immunostrip assays are readily available for many plant viruses, specific antibodies and serological tests are lacking for bean-infecting begomovirus identification. Nucleic acid-based technologies are highly accurate and sensitive and are widely used for the detection of bean-infecting begomoviruses. BGYMV and BGMV were cloned and sequenced in the early 1990s, and the availability of genetic information of these and other begomoviruses allowed for the quick development of tests based on DNA molecular hybridization and polymerase chain reaction (PCR). Dot-blot and tissue blot are the most used assay formats in hybridization tests. Cloned viruses or virus genome fragments are used as probes in the hybridization tests, and the probe can be tailored to be a broad-spectrum probe and recognize several viruses or to be specific for a given virus, hybridizing only to that virus. Typically, a mixture of DNA-A components from different begomoviruses is used as a general probe under low stringency hybridization conditions, and the common region or a hypervariable fragment of the DNA-B of a particular bean-infecting begomovirus is used as a specific probe under high stringency hybridization conditions. Currently, PCR is the most popular technique for the detection of bean-infecting begomoviruses due to its speed, sensitivity and accuracy. Different PCR set-ups, depending on the primers used, will provide general begomovirus detection or specific virus diagnosis. Usually, as a first step in identifying if a begomovirus is infecting a bean plant, very robust and effective universal degenerate primers designed in 1993 by M.R. Rojas and D. Maxwell at the University of Wisconsin are still used. Then, the amplicon is sequenced, permitting the initial identification of the begomovirus. Subsequently, back-to-back primers can be designed to recover the complete virus genome by inverse PCR for sequencing and definitive taxonomic classification. An additional set of specific primers can be designed to amplify part of the virus genome for specific diagnosis of that particular begomovirus in the samples. More recently, alternative nucleic-acid based techniques that use isothermal amplification have been developed and are also employed for the identification of bean-infecting begomoviruses. RCA (rolling-circle amplification) uses random primers and a DNA polymerase with strand displacement properties, favoring ssDNA amplification into double-stranded products in a nonspecific manner. RCA products can be cloned and sequenced allowing the identification and study of unknown viruses from symptomatic or asymptomatic samples. RCA is also valuable in discovering multiple virus infections in plants. Digestion of RCA products with restriction endonucleases and analysis of the restriction fragment length polymorphisms (RFLP) allows for the identification of mixed infections. Other diagnostic tools using isothermal amplification include loop-mediated isothermal amplification (LAMP) and recombinase polymerase amplification (RPL) assays. Protocols for LAMP and RPL are currently being developed for begomovirus diagnostics, and only RPL has been applied so far for BGYMV detection. However, these methods have great potential since they are sensitive, can use crude extracts and may be implemented to be used in the field. High-throughput sequencing technologies (HTS), also known as next generation sequencing (NGS), have greatly advanced the detection and discovery of plant viruses. HTS coupled with appropriate bioinformatics is a very sensitive tool, having the potential to detect the whole virome of a sample, including known and unknown viruses. Hence, HTS has been considered an attractive alternative for diagnostic virology. For the identification of begomoviruses, diverse types of nucleic acid templates may be used as starting material in the sequencing process including RCA products, double-stranded RNA and small RNAs. The two latter ones enable the simultaneous detection of viruses with DNA and RNA genomes belonging to different genera and families, revealing complex multiple infections. Several HTS sequencing platforms are commercially available. However, the costs are still high for routine diagnosis.

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Management Bean golden mosaic disease is very challenging to control and manage. The disease occurs and is a serious problem in the tropical and subtropical regions of the Americas. These areas are also zones where the whitefly vector thrives, colonizing, besides common bean, other crops such as soybean, tomato, peppers, cucurbits, cotton, and numerous non-cultivated plants, which sometimes can act as virus reservoirs. Because these crops may be present in the field year-round, high populations of the whitefly are built and migrate to newly planted bean fields. Chemical control using insecticides is the most used approach to control the whitefly vector and hence the infection caused by BGMV and BGYMV. However, the use of insecticides is not always efficient. Several types of molecules with different mechanism of action exist including nicotinoids and insect growth regulators. The over-employment of a determined insecticide chemistry over time may lead to the selection of resistant whitefly populations shortening the useful life of insecticides and, as a consequence, limiting the usefulness of chemical control. Thus, the alternation of products with distinct active ingredients should be implemented to preclude or delay resistance buildup. Severe epidemics and high yield losses are mainly recorded in the dry season when whitefly populations are high and may transmit begomoviruses to beans in the early phase of the cycle when the plants are young. To try to avoid serious yield reduction, a common practice is to treat the seeds and spray the fields with insecticide five to six times starting ten days after emergence, and then every five days until the plants are 40 days old. This type of schedule aims to control both adults and nymph stages, and as the whitefly population decreases, their migration to newly planted fields also tends to reduce. This scheme should take into consideration the insecticides’ characteristics and are not appropriate for those that have longer residual activity. Due to the limitations to the use of insecticides regarding the development of resistance in the whiteflies and environmental concerns, one of the best and sustainable options is the use of resistant cultivars. Many institutions throughout the Americas have dedicated great efforts in research programs focusing on breeding beans for resistance to golden mosaic disease. Screening attempts to identify common bean germplasm with disease immunity were unsuccessful. Thousands of bean accessions were evaluated under BGYMV and BGMV infection, and not a single genotype with full immunity was found. Some black-seeded genotypes displayed partial resistance or tolerance to BGYMV, and crosses between these genotypes rendered several lines denominated DOR that were released and cultivated, for example, in Guatemala. Further screening led to the selection of additional golden mosaic-resistant bean types with different seed colors. Genetic studies identified a few genes associated with the resistance in the different genotypes (Bgp, bgm1 and bgm2). Pyramiding different genes into breeding lines allowed for the achievement of higher and more adequate levels of resistance and the introgression of these genes into resistant varieties. Cultivars with resistance to BGYMV were released in Honduras, Nicaragua, El Salvador, Costa Rica, Puerto Rico and Southern Florida with good acceptance by farmers. Breeding for resistance to BGMV initiated in the 1970s in Brazil. Surveys for resistant genotypes and breeding efforts rendered bean lines with low levels of resistance that was controlled by polygenic and quantitative factors, and though a few cultivars with enhanced resistance were released, the resistance was not satisfactory under moderate to high BGMV incidence levels. Because there were no good resistance sources to be exploited, bean improvement programs by crossing and selection of desired superior genotypes were unproductive in Brazil (the resistant cultivars planted in other Latin American countries are of different seed types which are poorly accepted by consumers in Brazil). Therefore, an alternative biotechnological approach was used to achieve BGMV resistance. The RNAi technology was explored to generate a transgenic bean line transformed with an intron-hairpin construction harboring a fragment of the BGMV Rep gene that expresses a Rep-derived double-stranded RNA and activates the RNA silencing mechanism in the plant. In this approach, the transgenic plant produces small interfering RNAs that induce the degradation of the BGMV-derived Rep mRNA, halting virus replication and disease progression. The transgenic line Embrapa 5.1 proved to be immune to BGMV, and after extensive risk assessment studies, this line was approved in 2011 by the Brazilian Biosafety Committee (CTNBio) for cultivation and human consumption in the country. The Embrapa 5.1 line was crossed with ‘Carioca’ type beans (the most widely consumed in Brazil), originating the cultivar BRS FC401 RMD which was tested in different environments, holding the resistance in the field since 2012. Even under high virus pressure, no symptoms or virus replication have been observed. BRS FC401 RMD is expected to be released for commercial production in 2020 and is recommended for cultivation in Central Brazil. It is important to point out that BRS FC401 RMD is most probably not resistant to other golden mosaic-inducing begomoviruses occurring in Brazil (MaYSV) or in other countries, and is not resistant to RNA viruses that infect beans such as bean rugose mosaic virus and cowpea chlorotic mottle virus. Hence, other measures still have to be employed to control those viruses. Cultural practices may play a role in controlling bean golden mosaic disease. These include the sanitation of the field after harvest to eliminate any infected plants that could act as virus sources for the next crop; the avoidance of successive and continuous cultivation of the same crop in the same area; and prevention of virus transmission by sowing the new crop when the vector population and begomovirus availability are low. One control measure that is contributing to the reduction of begomovirus-associated disease in beans is the adoption of a host-free period. The host-free period usually spans from four to eight weeks after harvesting, when beans are not allowed to be cultivated. This type of measure is employed on a regional basis. Indeed, a bean-free period of four weeks in some areas in Brazil contributed to the reduction of the use of insecticides to control whiteflies and succeeded in reducing the losses due to BGMV infection. Similar success was achieved with an eight to twelve weeks of a bean-free period in the Dominican Republic in decreasing the incidence of BGYMV.

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Further Reading Bird, J., Sanchez, J., Vakili, N.G., 1973. Golden yellow mosaic of beans (Phaseolus vulgaris) in Puerto Rico. Phytopathology 63, 1435. Costa, A.S., 1965. Three whitefly-transmitted diseases of beans in the state of São Paulo, Brazil. FAO Plant Protection Bulletin, 13. pp. 121–130. Faria, J.C., Aragão, F.J.L., Souza, T., et al., 2016. Golden mosaic of common beans in Brazil: Management with a transgenic approach. APS Features. Available at: www.apsnet. org/edcenter/apsnetfeatures/Pages/GoldenMosaic.aspx. Gilbertson, R.L., Hidayat, S.H., Martinez, R.T., et al., 1991. Differentiation of bean-infecting geminiviruses by nucleic acid hybridization probes and aspects of bean golden mosaic in Brazil. Plant Disease 75, 336–342. Kim, K.S., Shock, T.L., Goodman, R.M., 1978. Infection of Phaseolus vulgaris by bean golden mosaic virus: Ultrastructural aspects. Virology 89, 22–33. Morales, F.J., 2001. Conventional breeding for resistance to Bemisia tabaci-transmitted geminiviruses. Crop Protection 20, 825–834. Morales, F.J., Jones, P.G., 2004. The ecology and epidemiology of whitefly-transmitted viruses in Latin America. Virus Research 100, 57–65. Ramos-Sobrinho, R., Xavier, C.A.D., Pereira, H.M.B., et al., 2014. Contrasting genetic structure between two begomoviruses infecting the same leguminous hosts. Journal of General Virology 95, 2540–2552. Rojas, M.R., Gilbertson, R.L., Russel, D.R., Maxwell, D.P., 1993. Use of degenerate primers in the polymerase chain reaction to detect whitefly-transmitted geminiviruses. Plant Disease 77, 340–347. Rojas, M.R., Hagen, C., Lucas, W.J., Gilbertson, R.L., 2005. Exploiting chinks in the plant’s armor: Evolution and emergence of geminiviruses. Annual Review of Phytopathology 43, 361–394. Rojas, M.R., Macedo, M.A., Maliano, M.R., et al., 2018. World management of geminiviruses. Annual Review of Phytopathology 56, 637–677. Silva, S.J.C., Castillo-Urquiza, G.P., Hora-Junior, B.T., et al., 2012. Species diversity, phylogeny and genetic variability of begomovirus populations infecting leguminous weeds in northeastern Brazil. Plant Pathology 61, 457–467.

Beet Curly Top Virus (Geminiviridae) Robert L Gilbertson, Tomas A Melgarejo, and Maria R Rojas, University of California, Davis, CA, United States William M Wintermantel, Agricultural Research Service, US Department of Agriculture, Salinas, CA, United States John Stanley, John Innes Centre, Colney, United Kingdom r 2021 Elsevier Ltd. All rights reserved. This is an update of J. Stanley, Beet Curly Top Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00696-8.

Glossary Agroinoculation A method for inoculating plants with infectious clones of a virus inserted in between the T-DNA borders of a binary vector such that the T-DNA is inserted into the plant nucleus via a disarmed Agrobacterium tumefaciens strain. Curly top disease A viral disease of plants characterized by stunting, up or downward curling and distortion of leaves, vein swelling, enations and purpling; the disease can be caused by genetically diverse geminiviruses, including Beet curly top virus. Enation Virus-induced outgrowth from leaves (typically veins) or other plant tissues. Endoreduplication DNA replication in the absence of cell division. Etiology The cause of a disease. Hemolymph The insect circulatory system. Hyperplasia Unregulated cell division.

Hypertrophy Cell enlargement resulting from abnormal increase in cell size. Non-propagative transmission Virus transmission mode in which the virus does not replicate in the vector. Phloem-limited virus A virus that only infects cells and tissues of the phloem. RFLP Restriction fragment length polymorphism, a molecular tool for fingerprinting and identifying DNA viruses. Steckling A small plant composed mostly of the root of a biennial crop plant (e.g., carrot or sugar beet) that is usually exhumed, stored during the winter and replanted the next season for seed production. Transovarial transmission Transmission of a virus vertically, through the insect egg. Viruliferous The condition in which an insect vector is carrying a virus and can transmit the virus to an uninfected plant.

Introduction Beet curly top virus (BCTV) is the type species of the genus Curtovirus, family Geminiviridae. Like all members of this family, BCTV has twinned quasi-icosahedral virions that measure B18
30 nm and a circular single-stranded DNA genome. BCTV is a phloemlimited virus that is transmitted by the beet leafhopper (BLH; Circulifer tenellus (Baker) (Homoptera: Cicadellidae) (Fig. 1(A)), and causes curly top disease of several economically important crops including common bean, pepper, sugar beet and tomato (e.g., Fig. 1(B–F)) in the western USA, north-central Mexico and countries of the Mediterranean Basin and the Middle East (e.g., Iran and Turkey), where it can cause substantial economic losses. Disease outbreaks are driven by populations of the beet leafhopper vector and favorable environmental conditions. In 2013, the disease caused an estimated US$100 million loss to the California processing tomato industry. Research on various aspects of the virus has added important fundamental knowledge about geminiviruses and the complex epidemiology of plant viruses transmitted by insect vectors. BCTV and the beet leafhopper vector represent an early example of the introduction of an exotic virus-vector combination into a new geographic region, the New World (NW) through movement of plant materials (sugar beet roots and stecklings) by humans. The BCTV genome sequence provided early evidence of the important role of recombination in viral evolution. Studies of the etiology of curly top disease in different geographic regions revealed that this disease is caused by multiple genetically distinct geminiviruses. Curly top disease also serves as a success story for how the development and implementation of resistant varieties and other management strategies saved the sugar beet industry from economic collapse. Finally, the BCTV-BLH vector system may lead to the identification of the elusive receptor(s) involved in the persistent circulative transmission of plant viruses by their insect vectors.

History The first reports of curly top disease were from sugar beet fields in the western USA in 1888. Along with the establishment and rapid growth of the sugar beet industry in this region (e.g., states of California, Idaho and Utah), came widespread and damaging outbreaks of curly top disease of sugar beet and other crops such as common bean, squash and tomato. Outbreaks of curly top disease caused substantial yield losses to sugar beet production in the western USA from B1920–1940 and the disease became a limiting factor in sugar beet production during this time, causing abandonment of major production areas and the closing of sugar

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Fig. 1 (A) Beet leafhopper (Circulifer tenellus) feeding on a vein on the abaxial side of a sugar beet leaf infected with Beet curly top virus (BCTV); (B) Portion of a tomato field with plants infected with BCTV, showing severe stunting and yellowing and poor stand due to plants that have died from the disease; (C) BCTV-infected tomato plant showing typical symptoms of curly top including stunting, upward leaf curling and yellowing; (D) Close-up of foliar symptoms of curly top disease in tomato showing upward rolling, dull green to yellow coloration and swollen and purple veins; (E) BCTV-infected sugar beet plant showing foliar symptoms of curly top on the leaves, including yellowing, crinkling, and upward curling; (F) Abaxial side of leaves of a BCTV-infected sugar beet plant showing swollen veins and enations associated with curly top disease; and inset showing darkening of vascular tissue in the root of a sugar beet plant infected with BCTV and showing curly top symptoms.

refineries. Subsequently, curly top-like diseases were reported to occur in western Canada, north-central Mexico, Brazil, Argentina, the Caribbean Basin, Turkey, and Iran. Here, it is important to note that molecular characterization of the causal agent of curly top disease from different geographical regions has revealed that these disease symptoms can be caused by multiple genetically distinct geminiviruses, including some that are not in the genus Curtovirus. Thus, symptoms alone are insufficient for diagnosis, and molecular tests (PCR and sequencing) are needed to confirm the identity of the causal agent. An association between curly top disease and the beet leafhopper (Fig. 1(A)) was noted, and evidence that this insect was a vector of curly top disease was first presented in 1909. These phloem feeding insects acquired and transmitted the causal agent of curly top rapidly (minutes), although longer periods of acquisition were needed for high rates of transmission. The curly top agent also persisted within the leafhopper vector, but it did not appear to replicate in the insect. Although the leafhopper transmission properties supported a viral etiology for curly top disease, the low level of accumulation (viral titer) of the causal agent in plants made purification difficult. This was mostly due to the virus exclusively infecting cells of the phloem. It was not until the 1970s that dimeric or twinned virus-like particles, measuring B20–30 nm, were purified from curly top-infected plants. This suggested that the causal agent of curly top might be a geminivirus, and further evidence came from the development of curly top symptoms in plants exposed to beet leafhoppers that had fed on purified preparations of the twinned virus-like particles. These results indicated that curly top disease was caused by a geminivirus, which was named Beet curly top virus. The viral genomic DNA was eventually cloned from DNA isolated from leaves of sugar beet plants with curly top disease symptoms, and the DNA sequence was determined in 1986. Analysis of the BCTV sequence confirmed that BCTV was a geminivirus, and also revealed that the virus has a hybrid genome composed of complementary-sense genes having greatest identities with those of whitefly-transmitted geminiviruses (genus Begomovirus) and virion-sense genes having highest identities with those of leafhopper-transmitted geminiviruses (genus Mastrevirus) (Fig. 2). This clearly indicated that recombination played a major role in the emergence of BCTV. The cloned genomic DNA of BCTV induced curly top symptoms in sugar beet plants, when delivered by agroinoculation, thereby fulfilling Koch’s postulates for curly top disease. Progeny virions derived from the infectious BCTV clone were transmitted by beet leafhoppers and subsequent development of curly top symptoms in inoculated sugar beet plants confirmed leafhopper transmissibility of BCTV.

Classification and Nomenclature Early investigations of host range, symptoms, and virulence suggested that curly top disease in the western USA was caused by a complex of viral strains/species. To some extent, this hypothesis has been supported by molecular (RFLP and sequencing) studies showing genetic diversity among the viruses causing curly top disease. In the western USA three strains were characterized based on

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Fig. 2 The genome organization of Beet curly top virus (BCTV). The position and orientation of virion-sense (V) and complementary-sense (C) open reading frames (ORFs) are shown in relation to the intergenic region (IR) that contains the invariant nonanucleotide motif TAATATTAC and iterons (repeated sequences underlined with arrows) involved in replication-associated protein (Rep) recognition and the initiation of viral DNA replication. The recombinant nature of the BCTV genome is shown with the virion-sense genes having highest identities with those of leafhoppertransmitted geminiviruses (mastreviruses) shown in orange, and the complementary-sense genes having highest identities with those of whiteflytransmitted geminiviruses (begomoviruses) shown in blue. CP, capsid protein; and REn, replication enhancer protein. Table 1 History of the taxonomy and nomenclature for phenotypic and genotypic variants of Beet curly top virus (BCTV) and other curtoviruses before and after the advent of molecular tools Molecular eraa Before 1990

1990

1998

Species

Strains

Species

Strainsb

Speciesc

BCTV

Various

BCTV

BCTV-Cal/Logan BCTV-Worland

BCTV BMCTV

BCTV-CFH

BSCTV SpCTV PeCTV PeYDV

BCTV-HRCT

HrCTV

2014 Strains

Speciesc

Strainsb

BCTV

BCTV-Cal/Logan BCTV-Wor BCTV-Mld BCTV-Svr BCTV-SpCT BCTV-PeCT BCTV-PeYD BCTV-CO BCTV-LH71 BCTV-SvrPep BCTV-Kim1

HrCTV SpSCTV

a

Molecular tools employed: 1990, restriction fragment length polymorphism (RFLP); 1998, RFLP and DNA sequencing; and 2014, DNA sequencing and phylogenetic analysis. Strain abbreviations: BCTV-Wor, BCTV-Worland; BCTV-Mld, BCTV-Mild; BCTV-Svr, BCTV-Severe; BCTV-SpCT, BCTV-spinach curly top; BCTV-PeCT, BCTV-pepper curly top; BCTV-PeYD, BCTV-pepper yellow dwarf; BCTV-CO, BCTV-Colorado; BCTV-LH71, BCTV-leafhopper71; BCTV-SvrPep, BCTV-severe pepper; BCTV-Kim1, BCTV-Kimberly1; BCTV-HRCT, BCTV-Horseradish curly top. c Species abbreviations: BMCTV, Beet mild curly top virus; BSCTV, Beet severe curly top virus; SpCTV, Spinach curly top virus; PeCTV, Pepper curly top virus; PeYDV, Pepper yellow dwarf virus; HrCTV, Horseradish curly top virus; and SpSCTV, Spinach severe curly top virus. b

molecular and biological properties: BCTV-Cal/Logan, BCTV-Worland and BCTV-CFH, with the strain designation generally reflecting the geographic origin of these strains (Table 1). Another strain was recovered from a horseradish plant with brittle root disease in Illinois and was designated BCTV-HRCT. The first strain for which the complete sequence was determined was BCTV-Cal/Logan in 1986. As sequences of more BCTV strains and putative new curtoviruses became available, additional genetic diversity as well as differences in biological properties (e.g., host range, symptomatology in sugar beet plants, and transreplication) were revealed. The taxonomy of viruses causing curly top disease is now largely based on genome properties, e.g., genome-wide sequence identities, along with some consideration of biological properties of the viruses. At the same time, the properties used to classify geminiviruses were also being defined and used to establish genera within the family Geminiviridae. The key properties that were used to classify geminiviruses were: genome organization, DNA sequence relationships, type of insect vector and host range of the virus. The viruses causing curly top disease in the western USA were placed into the genus Curtovirus (name derived from curly top) based upon (1) having a monopartite genome, (2) a high degree of sequence identity among members of the genus and divergence from sequences of species of other genera, (3) being transmitted by the beet leafhopper and (4) infecting dicotyledonous plants. There were originally four geminivirus genera: Curtovirus, Begomovirus (whitefly-transmitted and dicot-infecting),

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Fig. 3 Phylogenetic network generated with the complete genomes of 104 isolates of Beet curly top virus (BCTV) and illustrating the grouping of the isolates into the eleven recognized strains of the virus and their relationships with one another. The other two species in the genus Curtovirus, Spinach severe curly top virus (SpSCTV) and Horseradish curly top virus (HrCTV), were used as out-groups. The P-value of the pairwise homoplasy index (PHI) statistical test of recombination is shown on the bottom of the network.

Mastrevirus (leafhopper-transmitted and mostly monocot-infecting) and Topocuvirus (treehopper-transmitted and dicot-infecting). However, with the advent of inexpensive DNA sequencing and rapid methods for the amplification and cloning of geminivirus genomes, new types of geminiviruses have been identified and characterized. Thus, the family is now subdivided into nine genera with five new genera recently established: Eragrovirus, Capulavirus, Grablovirus, Becurtovirus, and Turncurtovirus. Here, it is important to note that the genera Becurtovirus and Turncurtovirus contain viruses, some of which are transmitted by the leafhopper C. haematoceps, that cause curly top-like disease symptoms in sugar beet and other crops. Thus, curly top disease symptoms can be caused by multiple genetically divergent geminiviruses from three genera. Therefore, molecular methods are needed to conclusively identify the virus(es) associated with the disease in a given geographical region. The use of DNA sequences for classification and phylogenetic analysis of geminiviruses has provided important insight into viral evolution and the relationships among members of this large family of viruses. However, the relationships among some geminiviruses have been complicated by extensive recombination, especially those in the genera Begomovirus and Curtovirus. Thus, because recombination has played a major role in their evolution, it can be difficult to establish an exact lineage for some viruses, i.e., whether it represents the parent or progeny of a recombination event. Another important issue is where to establish the cut-off values for establishing species, strain and isolate relationships in different genera. With the large number of sequences now available, these values are typically established based upon genome-wide pairwise identities. The original species demarcation value for curtoviruses was 89% identity, i.e., sequences having identities >89% to a known species represent isolates or strains of that species. This was the same species demarcation value used for begomoviruses. Other criteria that were considered in establishing curtovirus species were (1) lack of transreplication of known species mediated by the Replication-associated protein (Rep), (2) serological properties, and (3) biological properties (e.g., symptoms in sugar beet). Based upon these criteria, three BCTV strains were elevated to species in 1998: Beet curly top virus (type species, BCTV-Cal/Logan strain), Beet mild curly top virus (BCTV-Worland strain [BCTV-Wor]) and Beet severe curly top virus (BCTV-CFH strain) (Table 1). In addition, BCTV-HrCT was elevated to species (Horseradish curly top virus, HrCTV), as were at least five other curtoviruses associated with curly top disease in various crops (formally or informally). However, in 2014, the taxonomy of curtoviruses was revisited and, based upon genome-wide pairwise identities of all available curtovirus sequences, a new species demarcation value of 77% identity or greater was established, with little or no consideration of other properties. Based upon these new criteria, the genus Curtovirus currently includes only three species: the type species BCTV, HrCTV and the more recently described Spinach severe curly top virus (SpSCTV) (Fig. 3 and Table 1). The species BCTV is comprised of 11 recognized strains, which share o94% identity and, in some cases have different serological and biological properties (Fig. 3 and Table 1). Like BCTV-Cal/Logan, most of the recognized strains of BCTV have recombinant genomes (e.g., Fig. 4). HrCTV and SpSCTV also have recombinant genomes and lack a C3 ORF. The latter two species appear to have limited distribution and minimal economic impact, perhaps reflecting reduced fitness due to these genomic properties. For example, HrCTV was isolated in 1990 from a horseradish plant with brittle root disease in Illinois, USA, but Koch’s postulates were not fulfilled for this disease nor has the virus emerged as a pathogen of economic importance. However, as the genomes of HrCTV and SpSCTV are composed mostly of curtovirus sequences and are well below the 77% species demarcation threshold, they clearly represent distinct curtovirus species.

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Fig. 4 Similarity plot analysis of complete genome sequences of three strains of Beet curly top virus (BCTV): BCTV-CO, BCTV-LH71 and BCTV-PeCT. Bootscan analysis was performed with a sliding window of 200 nucleotides and step size of 20 nucleotides with Neighbor-Joining phylogenetic trees with 100 bootstrap replicates inferred in each case. This analysis provides a schematic representation of the recombination profile of each of these strains (query) as revealed by comparisons with other BCTV sequences in the NCBI database. The horizontal arrows on the top indicate the position of the open reading frames (ORFs) encoded by the BCTV genome. LIR, left side of the intergenic region and RIR, right side of the intergenic region.

The Origin of BCTV and the Beet Leafhopper Vector: An Early Example of Global Transport of a Plant Pathogen and its Vector Curly top disease continues to have the greatest economic impact on sugar beet and processing tomato production in the western USA. Soon after the disease became a major problem for sugar beet production, there were questions regarding the origin of BCTV and the beet leafhopper vector, i.e., were they indigenous to the NW or was one or both introduced from another region? Multiple lines of evidence suggest that BCTV originated in the OW and was introduced, along with the beet leafhopper, into the NW via movement of sugar beet propagative materials (e.g., roots and stecklings) brought with settlers coming from the OW (e.g., Europe and the Mediterranean Basin). The evidence for the introduction of the beet leafhopper included (1) the presence of C. tenellus and its closest relative in the OW (Mediterranean Basin), (2) interbreeding between NW and OW beet leafhopper populations, and (3) transmission of the causal agent of curly top in the NW by beet leafhoppers from the OW. The origin of BCTV remained unclear as there were no reports of curly top disease in the OW. However, in 1955, curly top disease of sugar beet was reported from Turkey and, subsequently, from Iran. In 1998, an isolate of BCTV from sugar beet plants with curly top disease symptoms in Iran (BCTV-I) was reported to have a high level of nucleotide sequence identity with BCTV strains from the NW, including >98% identity with BCTV-CFH (now BCTV-Svr [US-SVR-Cfh]). Moreover, an infectious clone of BCTV-I induced symptoms in N. benthamiana and sugar beet that were indistinguishable from those induced by the infectious clone of BCTV-CFH. This suggested a common origin for BCTV in the NW and OW, though it did not resolve the origin of BCTV. Evidence for an OW origin of BCTV comes from (1) the high levels of resistance to curly top disease in wild beet species in the OW (where sugar beet originated), suggesting virus-host coevolution over a long period of time,

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and (2) a portion of the recombinant genome of BCTV was derived from an OW leafhopper-vectored mastrevirus, consistent with emergence and evolution of BCTV in the OW. Although more BCTV genetic diversity has been documented in the NW, this could simply reflect the greater amount of research conducted on BCTV in the NW, or multiple introductions of BCTV strains from the OW and subsequent evolution mediated by mutation and recombination, which are facilitated during major curly top disease outbreaks. Taken together, these results support the hypotheses that both BCTV and the beet leafhopper vector originated in the OW and were introduced into the NW by the activities of humans. Unfortunately, the desert environment of the western USA was ideal for establishment of populations of beet leafhoppers and the associated BCTV.

Geographic and Seasonal Distribution Curly top disease occurs widely throughout arid and semiarid regions of the western USA, where high populations of the beet leafhopper vector can occur. It is here that the epidemiology of the disease and the life cycle of the insect vector has been extensively investigated. At the end of the growing season (in the fall), adult female beet leafhoppers (some of which are viruliferous) migrate from agricultural fields to overwintering areas (e.g., foothills of mountain ranges located on the western side of the Central Valley of California), where they survive on perennial weeds. In late winter to early spring, these females lay eggs into stems and petioles of perennial and annual weeds. The resulting nymphs and adults of this new generation acquire BCTV as they feed in the phloem of infected weed reservoir hosts, most of which show few or no symptoms. Initially, the virus and vector survive on perennial weeds such as Russian thistle (Salsola sp.) and saltbush (Atriplex sp.), until winter rains lead to the emergence of annual weeds, such as filaree (Erodium sp.), peppergrass (Lepidium sp.) and plantain (Plantago sp.) on which the leafhoppers propagate. As these winter annuals become dry in mid- to late-spring, some combination of viruliferous and non-viruliferous adult beet leafhoppers migrate from the overwintering areas to agricultural areas (often valleys), where they transmit BCTV to crops including sugar beet (Beta vulgaris), tomato (Solanum lycopersicum), common bean (Phaseolus vulgaris), and pepper (Capsicum annuum), as well as to common agricultural weeds as they search for preferred hosts for reproduction. Depending on the geographic region, beet leafhoppers undergo 3–5 generations on weeds, such as wild mustards (Brassica sp.), London rocket (Sisymbrium irio), Shepherd's purse (Capsella bursa-pastoris) and goosefoot (Chenopodium sp.). As crops are harvested and summer annuals dry in the fall, some combination of viruliferous and non-viruliferous adult female leafhoppers migrate from agricultural areas back to their overwintering areas.

Changes in BCTV Strain Prevalence BCTV strain diversity was investigated in samples of sugar beets with curly top disease collected from commercial fields in Texas in 1994 with restriction fragment length polymorphism (RFLP) analysis. The predominant strains were BCTV-Wor (previously BMCTV) and BCTV-Svr [US-SVR-Cfh] (previously BSCTV), with no detection of BCTV-Cal/Logan (type species). BCTV-Wor and BCTV-Svr were also the predominant strains detected in a more comprehensive survey of sugar beets with curly top symptoms collected from California, Colorado, Idaho, New Mexico, Oregon, Texas, Washington, and Wyoming in 1995. In a survey of tomato plants with curly top symptoms, beet leafhoppers and weed species in the Central Valley of California conducted between 2002 and 2008, BCTV-Wor and BCTV-Svr were prevalent, although BCTV-Cal/Logan was detected in a small number of leafhopper samples. Another survey of BCTV strains associated with curly top of sugar beet in the western USA was conducted from 2006-2007 and, again, BCTV-Wor and BCTV-Svr were prevalent, although a small number of plants from Idaho and Oregon were infected with BCTV-Cal/Logan. In these surveys, it was not uncommon to find plants infected with genotypic variants of these strains or co-infections of more than one strain. Interestingly, a major shift in the BCTV population structure has been detected in recent years in sugar beet and processing tomato samples from the Pacific Northwest and the Central Valley of California, respectively. A survey of sugar beets with curly top symptoms from Idaho and Oregon was conducted between 2012 and 2015 in response to concerns about increased curly top disease severity. The results revealed that (1) BCTV-CO and BCTV-Wor had become the prevalent strains, (2) there was a re-emergence of BCTV-Cal/Logan (30% of isolates detected) and (3) there was a decline in the prevalence of BCTV-Svr. Although BCTV-CO and BCTV-Wor are closely related (Figs. 3 and 4), they were differentiated by whole genome sequencing. Following the 2013 curly top outbreak in processing tomatoes in the Central Valley of California (where there is currently little or no sugar beet production), surveys of tomato plants with curly top symptoms and symptomless weeds in the foothills conducted from 2014 to 2018 revealed that BCTV-CO and BCTV-LH71, a new recombinant strain initially identified from viruliferous leafhoppers (Fig. 4) were predominant, rather than BCTV-Wor and BCTVSvr. This was particularly true with samples of tomatoes with curly top symptoms, whereas more strain diversity was detected in the weeds. Interestingly, in crop and weed samples from the Imperial Valley of California, where sugar beets are still grown, more BCTV-Svr was detected. In addition, two other BCTV strains were detected in pepper and tomato samples collected in California in 2014–2018, BCTV-spinach curly top (BCTV-SpCT) and BCTV-pepper curly top (BCTV-PeCT) (Table 1). Together, these results revealed a dramatic shift in strain prevalence in two different crops in two geographical regions, and the emergence of BCTV-CO as a major strain in both regions. Although the basis for this population shift in BCTV strains remains unknown, it may reflect changes in cropping patterns or in the predominant weed reservoirs surrounding agricultural fields and selection of more virulent or fit strains. For example, BCTV-CO and BCTV-LH71 appear to be better adapted to tomato.

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Genome Organization and Gene Expression Like all geminiviruses, BCTV has a genome of circular single-stranded DNA (ssDNA) that is encapsidated in twinned (geminate) quasi-icosahedral particles measuring approximately 18
30 nm. The BCTV genome is composed of a single genomic DNA of B3000 nt that encodes seven genes distributed between the virion-sense (V) and complementary-sense (C) strands (also referred to as rightward [R] and leftward [L] strands in the literature), separated by an intergenic region (IR) of B300 nt, which contains the origin of replication (Fig. 2). A nonanucleotide motif (TAATATTAC), which is highly conserved among all geminiviruses, is located within the IR and lies in between inverted repeat sequences with the potential to form a stem-loop structure. Short repeat sequences, termed iterons, are upstream of the stem-loop (Fig. 2). The complementary-sense genes encode the Rep protein (ORF C1) that is required for viral DNA replication, the replication-enhancer REn (ORF C3), and two proteins that contribute to viral pathogenicity (ORFs C2 and C4). The virion-sense gene products include the capsid protein (CP, ORF V1) and proteins involved in the regulation of the relative levels of viral ssDNA and double-stranded DNA (dsDNA) (ORF V2) and virus movement (ORF V3) (Fig. 2).The complementary-sense gene products are required during the early stages of infection, whereas the virion-sense ORFs encode proteins that are required later in infection, as the virus shifts to virion formation and movement within the plant. The identification of BCTV virion- and complementary-sense transcripts is consistent with a bidirectional transcription strategy that, along with overlapping genes (in different frames), maximizes the number of genes encoded by a small genome (Fig. 2). The two most abundant complementary-sense transcripts are similar to their begomovirus counterparts, with the larger transcript mapping across all four ORFs and the smaller transcript across ORFs C2 and C3. Overall, the most abundant transcripts mapped across the virion-sense ORFs and downstream of two consensus eukaryotic promoter sequences. The most abundant transcript mapped to the ORFs V2 and V3, whereas a less abundant transcript mapped to ORF V1. Precisely how the viral proteins are translated from these transcripts is not yet understood, but the overlapping nature of both the virion-sense and complementarysense ORFs may provide temporal control during the infection cycle.

Replication The virion-sense ssDNA carried in the BCTV virion must gain access to the nucleus of phloem cells (e.g., companion and phloem parenchyma), where replication of viral DNA occurs. It is not known if virions or viral DNA released from disassembled virions enters the nucleus following introduction of virions by the beet leafhopper vector. Once in the nucleus, the virion-sense ssDNA is converted into dsDNA, mediated by host-derived short DNAs as primers and host DNA polymerases (note that geminiviruses do not encode a DNA polymerase). Fractionation of viral DNA forms by a combination of chromatography and two-dimensional gel electrophoresis has identified BCTV intermediates consistent with both rolling-circle and recombination-dependent replication strategies. During rolling-circle replication, Rep interacts with the iterons located in the IR and introduces a nick into the virionsense strand of the dsDNA within the nonanucleotide motif (TAATATT↓AC). By analogy with the replication strategy of begomoviruses, Rep then forms a covalent bond with the 50 terminus of the nicked virion-sense strand, whereas the 30 terminus must be extended by a host DNA polymerase. The full-length nascent strand is nicked and ligated by Rep to produce a genomic circular ssDNA that either re-enters the replication cycle (i.e., be converted to dsDNA early in infection) or is encapsidated to form virions (later in infection). The interaction between Rep and the origin of replication is defined by the amino acid sequence of the N-terminal region of Rep and the iteron sequences of the origin of replication. This interaction is highly specific and determines whether transreplication occurs between strains or species (i.e., the Rep of one strain/species mediates the replication of another strain/species). For example, the Rep of the BCTV-Cal/Logan strain functionally interacts with its own iteron, GGAGTATTGGAGT (iteron underlined) (Fig. 2), but not with those of BCTV-Wor [US-Mld-Wor], GGTGCTATGGGAG, and BCTV-Svr [US-SVR-Cfh], GGTGCTTTGGGTG. Conversely, the Rep proteins of BCTV-Wor and BCTV-Svr do not functionally interact with the iterons of BCTV-Cal/Logan, whereas the BCTV-Wor and BCTVSvr iterons are sufficiently similar to allow functional interaction with the Rep proteins of these strains and, thus, transreplication. Comparison of Rep N-terminal amino acid sequences has revealed residues conserved between BCTV-Wor and BCTV-Svr and that differ from those of the BCTV-Cal/Logan Rep. These residues, referred to as the iteron recognition domain (IRD), are involved in iteron recognition. Whereas replication incompatibility between cis- and trans-acting elements may be considered to maintain the integrity of a particular species, this constraint can be overcome by a recombination event in which the functional module comprising the 50 terminus of the Rep ORF and the origin of replication is exchanged between incompatible viruses. Indeed, in addition to mediating the emergence of the hybrid genome of the BCTV progenitor, recombination also has played a major role in the evolution and emergence of other BCTV strains, including BCTV-CO, BCTV-LH71, and BCTV-PeCT (Fig. 4). This is mediated by recombination-dependent replication, which typically occurs later in the infection cycle compared with rollingcircle replication, possibly due to higher DNA concentrations and the presence of ss- and dsDNA forms that mediate template switching and recombination. It remains to be established which, if any, viral proteins contribute to this process, although it is possible that Rep helicase activity could participate in both replication strategies. Additionally, the nick site for rolling-circle replication has been shown to be a recombinational hot spot. Thus, recombination-dependent replication likely explains the high frequency of recombinant geminiviruses. Subgenomic-sized defective viral DNAs of diverse size (ranging from 800 to 1800 nt) and complexity are rapidly produced from the cloned BCTV genomic DNA in sugar beet, tomato, and N. benthamiana. Deletions occur within all ORFs and only the IR

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sequence is conserved, consistent with its participation in viral DNA replication. BCTV-infected sugar beet plants grown in fields frequently contains such defective DNAs, indicating that they are also produced under natural conditions. They occur as both single- and double-stranded DNA forms but it is not known if they are encapsidated or have the ability to move systemically in the plant. However, in view of their rapid appearance, it is likely that they are produced de novo in every infected cell. Short repeat sequences of 2–6 nt at the deletion boundaries may specify the deletion endpoints and suggest that the defective DNAs are produced by recombination. These subgenomic-sized DNAs can interfere with infection, presumably by competing for a limited amount of Rep, and may have a biological role in modulating pathogenicity of the helper virus to encourage leafhopper feeding and virus transmission.

Virus Movement The CP is a multifunctional protein that is targeted to the nucleus, probably via one or more nuclear localization signals (NLSs) in the N-terminus, where virion formation and accumulation occurs. In transient expression experiments in protoplasts and bombarded common bean hypocotyl tissues, a BCTV-Wor [US-Mld-Wor4] (previously BMCTV) CP-GFP fusion protein was targeted to the nucleus and formed ring-like structures, reminiscent of fibrillary rings observed in nuclei of BCTV-infected cells in ultrastructural studies. Late in infection, virions and viral nucleic acids form large aggregates or inclusion bodies in nuclei of BCTVinfected phloem cells. The BCTV CP is essential for systemic infection and leafhopper transmission. BCTV-Wor [US-Mld-Wor4] CP alanine scanning mutants that retain the ability to form virions (mainly N-terminal mutations) were generally able to produce a systemic infection and were leafhopper-transmissible, whereas those unable to form virions (mainly C-terminal mutations) were not, despite being competent for replication. Thus, these results suggest that the C-terminal amino acids of the CP play an important role in virion assembly, and that long-distance movement of BCTV (in the plant) occurs in the form of virions, which is consistent with the observation of virus-like particles in phloem sap exudates of leaves of Amsinckia douglasiana plants with curly top symptoms. ORF V2 mutants accumulate high levels of dsDNA and greatly reduced levels of ssDNA compared with the wild-type virus, implicating the V2 protein in the regulation of the levels of these viral DNA forms. However, the V2 protein does not have a typical NLS, and in transient expression experiments in protoplasts and bombarded hypocotyl tissues, a V2-GFP fusion protein was localized to the endoplasmic reticulum (ER), including having a perinuclear distribution. If the function of the V2 protein is to ensure that ssDNA is available in sufficient amounts for encapsidation and systemic movement late in infection, this may involve V2-mediated nucleocytoplasmic transport of an infectious form of the virus, e.g., virions or some other nucleoprotein complex. The possibility of an infectious non-virion form of BCTV was suggested based on results with alanine scanning mutant CP 49–51, which infected and caused curly top symptoms in N. benthamiana plants, but did not form virions and was not leafhopper transmissible. ORF V3 mutants are competent for replication but produce only sporadic and symptomless infections with low levels of viral DNA in sugar beet and N. benthamiana plants, suggesting a role for the V3 protein in virus movement. In transient expression experiments in protoplasts and hypocotyl tissues, V3-GFP initially localized to the nuclear periphery and ER, similar to V2-GFP. However, V3-GFP subsequently was localized to motile ER-derived vesicle-like structures, which were observed around the nucleus, in the cytoplasm and at the cell periphery. It is tempting to speculate that this represents a transport mechanism, possibly delivering an infectious form of the virus to the cell periphery and plasmodesmata for limited cell-to-cell movement. Finally, it is important to consider how the virions are acquired by leafhoppers during the feeding process. Although virions seem to be the main form in which BCTV moves in the phloem, it is not known if this occurs as individual virion particles or aggregates (e.g., intact or degraded parts of inclusion bodies). It is also not clear how virions enter the phloem. One hypothesis involves the collapse and death of BCTV-infected phloem cells and the release of cell contents, including inclusion bodies, into the developing phloem sieve elements, possibly in the shoot apex. These virions would then be available for acquisition by beet leafhoppers or long-distance movement in the phloem in a source-to-sink manner to infect new protophloem and other phloem cells in the newly developing root and shoot apices. In this ‘wave of infection’ model, the virus is continually infecting and colonizing the nucleate phloem progenitor cells in newly developing shoots.

Host Range and Pathogenesis In contrast to most other geminiviruses that typically have a narrow host range, BCTV is reported to have an extremely wide host range, including over 300 species of dicotyledonous plants in 44 families, including members of the Chenopodiaceae, Compositae, Cruciferae, Leguminosae, and Solanaceae. Here, it is worth noting that the BCTV host range was established from early observations of diseased plants of undefined etiology, when it was difficult to precisely identify the causal virus(es) involved. Therefore, the reported host range of BCTV should be confirmed. However, using agroinoculation and leafhopper transmission of infectious clones and progeny virions, respectively, relatively wide host ranges (at least among crops) were demonstrated for BCTV-Wor, BCTV-Svr, BCTV-SpCT, BCTV-Mld, BCTV-CO and other strains. Furthermore, the curly top symptoms that developed in these plants were similar to those described in these crops in the

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field. BCTV-infected crop plants also had high concentrations (titers) of virus. In contrast, non-cultivated plants such as weeds typically show few or no symptoms when infected by BCTV, and such plants often have lower levels of virus. However, BCTV infects a wide variety of annual and perennial weed species and at rates as high as 20% (in the foothills of California), which serve as reservoirs of the virus, especially in leafhopper breeding areas. Moreover, even though the levels of virus are low in most infected weeds, the feeding characteristics of the beet leafhopper allows for concentration of the virus in viruliferous insects. The products of BCTV complementary-sense ORFs C2 and C4 have been associated with pathogenesis. ORF C2 mutants induce severe symptoms and accumulate to wild-type virus levels in sugar beet and N. benthamiana. However, plants infected with these mutants have a greater propensity for recovery from severe symptoms compared with those infected with wild-type virus, suggesting that the C2 protein suppresses a plant defense response. Consistent with this hypothesis, transgenic N. benthamiana and N. tabacum plants constitutively expressing the C2 protein exhibit enhanced susceptibility to BCTV infection, as revealed by a shorter latent period before symptom development. However, when infected with BCTV, these transgenic plants did not develop more severe symptoms or accumulate higher levels of viral DNA compared with non-transformed control plants. The transgenic plants were also more susceptible to the begomovirus Tomato golden mosaic virus (TGMV) and the unrelated RNA virus Tobacco mosaic virus, indicating that the C2 protein affects a nonspecific host defense response. Consistent with these observations, the C2 protein has been shown to suppress post-transcriptional gene silencing (PTGS) and transcriptional gene silencing (TGS) in N. benthamiana. In this respect, the BCTV C2 resembles the AC2 homolog in the begomovirus TGMV. However, the two proteins share only limited homology, and the BCTV C2 protein does not appear to have transcriptional activator activity for virion-sense gene expression as has been shown for TGMV AC2. Despite this divergence, both BCTV C2 and TGMV AC2 proteins bind to, and inactivate, adenosine kinase (ADK), an enzyme required for 50 -adenosine monophosphate (AMP) synthesis. ADK activity is also reduced in BCTV-infected plants and transgenic plants expressing C2 protein. Regulation of adenosine levels by ADK plays a key role in the control of intermediates required for methylation, which is the underlying mechanism of TGS and provides a possible link between C2 protein activity and suppression of TGS targeting the viral genome. Both BCTV C2 and TGMV AC2 proteins also bind to, and inhibit, sucrose non-fermenting 1 (SNF1)-related nucleoside kinase. Reduction of SNF1 protein kinase expression in N. benthamiana results in an enhanced susceptibility phenotype resembling that associated with the expression of C2 and AC2 proteins, whereas SNF1 overexpression produces plants with reduced symptom severity. This activity is not directly linked to silencing suppression, suggesting that C2 protein inhibition of SNF1 activity affects a different plant defense pathway. BCTV ORF C3 mutants produce severe symptoms and accumulate to wild-type levels in Nicotiana benthamiana plants, but they induce only mild symptoms and accumulate low levels of viral DNA in sugar beet plants. This suggests that the BCTV REn may enhance viral DNA replication in a host-dependent manner. BCTV typically induces upward curling of the leaves (Fig. 1(B–F)) and clearing, enations and swelling of veins on the abaxial side of leaves, particularly in leaves of sugar beet plants (Fig. 1(F)). Although the phloem-limited virus cannot access the apical meristem, it is possible that BCTV exploits undifferentiated cambium cells in the vascular bundles or nucleate protophloem cells in the root and shoot apices, as shown for the begomovirus Bean dwarf mosaic virus. BCTV infected tissues exhibit hyperplasia and hypertrophy within the phloem and adjacent parenchyma, and the abnormally dividing cells differentiate into sieve element-like cells. In older infected tissues, the affected phloem cells eventually become necrotic and collapse. Pathogenic effects occur in developing tissues only after maturation of sieve elements, consistent with the virus moving from source to sink cells with the flow of photoassimilates, either to newly developing shoots or roots, depending on where the leafhopper introduced the virus to the plant (there can be large amounts of BCTV in roots). Infected sugar beet leaves accumulate enhanced levels of sucrose, attributed to impaired transport resulting from disruption of the phloem. This is associated with a reduction in chlorophyll content, reduced activity of key photosynthetic enzymes and a concomitant reduction in the rate of photosynthesis and altered turgor pressure that causes an increase in mesophyll cell size. Geminiviruses are masters at manipulating the plant cell cycle to produce an environment suitable for their replication and spread. This is achieved by the interaction of Rep with the plant homolog of retinoblastoma-related tumor-suppressor protein (pRBR), which relieves the constraint imposed by pRBR binding to the E2F transcription factor and allows the expression of E2Fresponsive genes. BCTV infection induces cell division within vascular tissues, attributed to the action of the C4 protein. Sugar beet infected with ORF C4 mutants remain asymptomatic and N. benthamiana plants show an altered phenotype; neither host develops hyperplasia of the phloem or enations. Furthermore, transgenic N. benthamiana plants expressing C4 protein show abnormal plant development and tumorigenic growths. Consistent with this phenotype, C4 protein interacts with BIN2, a negative regulator of transcription factors in the brassinosteroid signaling pathway that controls cell division and tissue development. The reasons for the induction of cell division are not clear, particularly as the hyperplastic tissues often do not contain detectable levels of virus. However, it is likely that it does provide additional nucleate cells that can be infected, via cell division or a yet to be determined means of cell-to-cell movement, and produce virions that can be released into sieve elements for long-distance movement or acquisition by the beet leafhopper vector. The host range of BCTV includes Arabidopsis thaliana, which represents an important resource for the study of virus-host interactions. Most susceptible ecotypes become stunted and develop enations on infected leaves. The Sei-O ecotype is hypersusceptible to BCTV-Svr and develops callus-like structures containing high levels of the virus, suggesting a virus-induced hormonal imbalance. Symptom development correlates with enhanced expression of the cell cycle gene cdc2 and small auxin-upregulated RNA gene (saur). Hence, disruption of the phloem in infected tissues may affect auxin transport, causing localized increases that result in cell proliferation.

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Transmission of BCTV by the Beet Leafhopper Vector In nature, BCTV is transmitted, plant-to-plant, exclusively by the beet leafhopper (C. tenellus, Fig. 1(A)). BCTV is not transmitted by contact or mechanically (sap inoculation) and is not believed to be seed-transmitted, although seed transmission of BCTV in petunia has been reported. A search for additional insect vectors in the NW was unsuccessful, which indicated a high degree of specificity in the association of the curly top agent and the beet leafhopper. However, other species of Circulifer from the OW can vector BCTV, and C. opacipennis was reported to vector the curly top agent in the Mediterranean Basin. There is no evidence for strain specificity in transmission of BCTV by C. tenellus, and all strains tested to date have been leafhopper-transmissible. The mode of transmission is persistent circulative, and there is no evidence that BCTV replicates in the insect vector. The beet leafhopper is an efficient vector of BCTV and rapidly acquires the virus (in minutes) from the phloem of infected plants. Being a mobile, polyphagous phloem-feeding insect makes the beet leafhopper an ideal vector for BCTV. The leafhopper also prefers to feed from veins on the abaxial side of young- to medium-aged leaves, including the swollen veins of BCTV-infected plants (Fig. 1(F)), in which there are high concentrations of virions. It is not known if beet leafhoppers are preferentially attracted to plants with curly top symptoms, as has been reported for whiteflies (Bemisia tabaci species complex) and Tomato yellow leaf curl virus (TYLCV). The pathway BCTV follows in the beet leafhopper was investigated with a PCR-based approach. BCTV-Wor [US-Mld-Wor4] was detected in the digestive tract of the insect after an acquisition access period (AAP) of 1 h, in the hemolymph after 3 h, and in the salivary glands after 4 h, from where it is reintroduced into plants with saliva during feeding. This is consistent with earlier reports estimating the minimum time between virus acquisition and the leafhopper becoming viruliferous (the latent period) to be 4 h. The feeding time necessary for transmission of the virus can be as short as 1 min, though longer feeding times (e.g., inoculation access periods of 24–48 h) provide higher rates of infection. Depending on the length of the AAP, leafhoppers can remain viruliferous for life, although the amount of virus retained and the ability of leafhoppers to transmit the virus declines with time when viruliferous leafhoppers are maintained on plants that are not hosts for the virus. This indicates that the virus is not replicating in the leafhopper. There is also no evidence for transovarial transmission of the virus, a common feature of viruses that replicate in their insect vector. Several lines of evidence demonstrate that the CP is the determinant of leafhopper transmission specificity. First, BCTV-Wor [US-Mld-Wor4] CP mutants that failed to produce virions are not transmitted by C. tenellus. Second, a recombinant virus in which the CP coding sequence of the whitefly-transmitted begomovirus African cassava mosaic virus was replaced with that of BCTV produced virions that were transmissible by the beet leafhopper when injected into the hemocoel. Finally, the autonomously replicating nanovirus-like DNA-1 component, normally associated with the whitefly-transmitted begomovirus Ageratum yellow vein virus, can be maintained in sugar beet plants infected with BCTV, and is transmitted by the beet leafhopper. These results indicated that the B1.4 kb DNA-1 molecule is encapsidated by the BCTV CP, although the mechanism is not known, i.e., are two B1.4 kb ssDNAs or a single B2.8 kb dimeric ssDNA form (a product of rolling circle replication) encapsidated. The BCTV-beet leafhopper interaction was further investigated by immunofluorescence and immunolocalization, with antisera raised against E. coli-expressed BCTV-Wor [US-Mld-Wor4] CP. BCTV was detected in the mid- and hindgut of viruliferous beet leafhoppers, and distinct fluorescent punctate structures, far greater than the size of an individual virion, were observed on or in the gut walls. These may represent large aggregates of virions (possibly inclusion bodies) acquired from the phloem during feeding or some form of cooperative binding of virions during feeding to form aggregates following initial binding of a small number of virions to the gut wall. Immunolocalization studies revealed vesicle-like structures on the inside of the gut wall, possibly involved in endocytosis of BCTV virions. In the salivary glands, BCTV was mostly localized to the primary salivary gland. The BCTV-Wor alanine scanning mutant CP 25–28 formed virions, but was not leafhopper-transmitted. Dissection PCR and immunofluorescence and immunolocalization revealed that this mutant interacted with the gut in a similar fashion to wild-type virus and entered into the hemolymph, but was not detected in the salivary glands. Thus, this may reveal an amino acid motif of the CP that interacts with a receptor on the salivary gland surface. Together, these results suggest that surface features of the BCTV virion interact in a highly specific manner during circulative transmission through the beet leafhopper, possibly involving two receptor-mediated endocytosis events, i.e., during virus acquisition across the gut wall and movement into the salivary glands. An effort is ongoing to sequence the genome of the beet leafhopper with the goal of identifying the receptors involved in the transmission of BCTV and the microbes tomato big bud phytoplasma and citrus stubborn spiroplasma.

Experimental Systems for Infection of Plants by BCTV BCTV is poorly (at best) transmitted by mechanical inoculation, which reflects the phloem-limited life style of the virus. Infection can be achieved using a fine-gauge needle (pin-pricking) to introduce virions or viral DNA inoculum directly into the phloem, but the efficiency is low. Particle bombardment is an efficient method for infection of the laboratory host N. benthamiana with BCTV, but very inefficient for infection of other hosts. The reason for the difference in host susceptibility to particle bombardment inoculation is not known, and BCTV is phloem-limited in N. benthamiana. It is possible that phloem cells susceptible to BCTV infection are more accessible in the leaves of the young (two to four leaf stage) N. benthamiana plants inoculated by particle bombardment. In contrast, high rates of BCTV infection in numerous crops (common bean, cucurbits, pepper, sugar beet and tomato) can be achieved via agroinoculation, in which a multimeric infectious clone is inserted in between the T-DNA borders of a binary

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plasmid. This plasmid is transformed into a disarmed strain of Agrobacterium tumefaciens, and bacterial cells carrying the binary plasmid are inoculated into stems of young plants just beneath the shoot apex by the needle-puncture method. The A. tumefaciens cells bind to wounded plant cells, presumably including nucleate progenitor cells of the phloem, and deliver the T-DNA containing the multimeric BCTV DNA into the nucleus. In the nucleus, the genomic BCTV ssDNA is released from the integrated T-DNA (in transformed cells) by the expressed Rep, which recognizes the IR sequences. The viral infection then proceeds as if the leafhopper had delivered virions into the phloem. Agroinoculation has proven to be a rapid and efficient method of inoculation for BCTV and other geminiviruses, and can be used for screening tomatoes and other crops for resistance to BCTV. This eliminates the need to rely on field plots and natural inoculum, the occurrence of which is highly unpredictable. Agroinoculation has limitations, including difficulties in generating agroinoculation systems for some strains (e.g., BCTV-CO), and the recalcitrance of some plants to Agrobacterium-mediated transformation, e.g., pepper.

Management of Curly Top Disease Caused by BCTV Like many vector-borne diseases, management of curly top disease caused by BCTV has been challenging due to multiple factors. These include the sporadic nature of curly top outbreaks, difficulty in predicting ‘bad’ curly top years, the migratory nature of the vector and difficulty in managing leafhopper populations with insecticides. In years having severe outbreaks, plants are typically infected at an early stage of development, resulting in substantial yield losses. Resistance to curly top disease is not available in commercial varieties of a number of susceptible crops, including pepper and tomato. Therefore, differences in (1) crop production, (2) the biology of curly top disease in different crops, and (3) the availability of resistant varieties means that management approaches for different crops may vary. However, the most effective curly top disease management will be accomplished through an integrated pest management (IPM) approach. Below, some general strategies for curly top disease management and examples of IPM programs for sugar beet and tomato crops are presented.

Resistant Varieties The planting of resistant varieties is one of the most effective and environmentally friendly management strategies for any disease. Sources of resistance to BCTV were identified in germplasm of common bean, flax, squash and sugar beet. In the case of sugar beet, mass selection of plants from fields with high incidences of curly top disease led to the identification and release of the first curly top resistant sugar beet variety, US1, in 1933. This variety became widely grown in the western USA, despite some horticultural shortcomings. The identification and selection of resistant sugar beet varieties has been facilitated by the use of breeding plots, known as curly top nurseries, in which large numbers of leafhoppers viruliferous for the most prevalent and virulent strains of BCTV are released. As a result of the high selection pressure in these nurseries, combined with the development of commercial hybrids and molecular breeding technologies, curly top resistance in sugar beet has advanced tremendously despite being a complex quantitatively inherited trait. Modern curly top resistant varieties have greater uniformity, higher yield and improved resistance. However, plants of these resistant sugar beet varieties will develop curly top symptoms and may suffer yield losses when infected at an early stage of development (two to four leaf stage). Therefore, these resistant varieties need to be part of an IPM program that addresses the susceptibility of young plants. Curly top resistance was also found in a range of common bean varieties (e.g., Red Mexican type), and genetic studies identified a single dominant gene (Bct-1) associated with curly top resistance. Breeding efforts led to the release of the commercial curly topresistant varieties Great Northern UI 15 and Red Mexican UI 34. Subsequently, snap bean varieties with curly top resistance were developed and released. These varieties have provided acceptable yields under high curly top pressure. There is a need to develop curly top-resistant varieties for other market classes of common bean. In terms of tomato and pepper, curly top resistance was not found in commercial cultivars. High levels of resistance were found in accessions of some wild species of tomato, including Solanum chilense, S. habrochaites and S. peruvianum. Unfortunately, introgression of this resistance into commercial tomato varieties has been difficult. Agroinoculation screening of tomato breeding lines with different combinations of Ty genes, which confer resistance to the begomovirus TYLCV, with BCTV-Svr and BCTV-LH71 revealed gene combinations, e.g., Ty-1, Ty-2, Ty-3 and Ty-5, that provided high levels of curly top resistance. The tomato breeding Line 20 #12, carrying the Ty-1, Ty-2 and Ty-3 genes, has moderate resistance to curly top, which involves initial symptom development followed by recovery. This may indicate that TGS is targeting the viral genome for methylation.

Cultural Practices Because young plants are more susceptible to infection and suffer the greatest yield losses, some degree of protection can be achieved by adjusting planting dates to avoid exposure of young plants to flights of viruliferous leafhoppers. Typically these flights occur in mid- to late-spring. Thus, early or late planting dates may allow for avoidance of peak leafhopper flights when plants are young, but the difficulty in predicting leafhopper flights from year-to-year is a challenge and populations vary considerably depending on weather conditions and the composition of host plants in overwinter areas. Avoiding the establishment of susceptible crops near leafhopper breeding areas is another strategy that can reduce curly top incidence, though this is limited by land

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availability and options for planting crops that are not susceptible to curly top disease (e.g., garlic, melons, onions or small grains). Finally, over-seeding of tomatoes can reduce losses due to curly top disease by allowing for uninfected plants to compensate for yield losses due to infected plants, and to possibly make fields less attractive to migrating leafhoppers, which cue in on patches of brown and green.

Management of the Leafhopper Vector With Insecticides The most common management strategy is to reduce populations of the beet leafhopper vector in order to reduce the incidence and spread of BCTV. This involves spraying contact insecticides such as malathion or dimethoate and delivering systemic insecticides such as neonicotinoids via sprays, soil drenches or drip irrigation. More recently, technologies have been developed for coating seeds with systemic insecticides. Spraying of contact insecticides can be carried out on weeds in leafhopper breeding grounds or directly on crops such as beans, sugar beets and tomatoes. Ideally, sprays are applied based on monitoring for leafhopper populations with sticky cards or sweep nets. In California, it has been shown that high incidences of curly top in tomato (>10%) require high populations of viruliferous leafhoppers in the spring. This information can inform growers if sprays are necessary. The California Department of Food and Agriculture runs the Curly Top Virus Control Program, the only remaining program that sprays breeding grounds of beet leafhoppers. In this grower-funded program, breeding areas in the rangelands of the foothills on the western edge of the Central Valley are surveyed with sweep nets for beet leafhoppers and malathion is applied when thresholds are reached. The efficacy of this program in reducing incidence of curly top disease in crops remains unclear. Parasites and predators of the beet leafhopper are not uncommon, but it has proven difficult to assess their sustainability or impact on leafhopper populations under natural conditions.

Sanitation It is not possible to eradicate weed reservoirs in the vast areas that comprise the breeding areas for the beet leafhopper in the western USA. However, in the agricultural valleys where curly top disease is a problem, it is important to practice effective weed management in fields and in non-cultivated areas, as well as around canals and other bodies of water, to eliminate preferred hosts for reproduction of beet leafhoppers, such as Russian thistle (Salsola sp.). This can reduce beet leafhopper populations in areas near susceptible crops, as well as those that migrate to the foothills in the fall.

IPM for Curly Top Disease IPM packages for management of curly top in sugar beet (a preferred host of the beet leafhopper on which it feeds and reproduces) and tomato (a non-preferred host that the leafhopper tastes and then moves on) are different. For sugar beet the IPM package frequently includes: (1) planting cultivars with curly top resistance; (2) early planting dates (to allow plants to reach six to eight true leaves before viruliferous leafhoppers arrive in the fields); (3) management of beet leafhopper populations in fields with insecticides based on an IPM approach, e.g., seed treatment with systemic insecticides followed by mid-season application of a contact insecticide; and (4) sanitation of harvested fields and of weeds on a regional basis. Tomato is not a preferred host of the beet leafhopper, and there are no commercially available resistant varieties. Thus, the IPM package may involve (1) modifying planting dates or the location of fields to avoid leafhopper flights and proximity to leafhopper breeding grounds or flight paths; (2) drenching tomato transplants in the greenhouse and at transplanting with a systemic insecticide; (3) management of beet leafhopper populations in fields with insecticides based on an IPM approach; and (4) sanitation of harvested fields and regional weed management.

Further Reading Bennett, C.W., 1971. American Phytopathology Society Monograph No. 7: The Curly Top Disease of Sugar Beet and Other Plants. St. Paul, MN: American Phytopathology Society. Chen, L.-F., Brannigan, K., Clark, R., Gilbertson, R.L., 2011. Characterization of curtoviruses associated with curly top disease of tomato in California and monitoring for these viruses in beet leafhoppers. Plant Disease 94, 99–108. Chen, L.-F., Gilbertson, R.L., 2016. Transmission of curtoviruses (Beet curly top virus) by the beet leafhopper (Circulifer tenellus). In: Brown, J.K. (Ed.), Vector-Mediated Transmission of Plant Pathogens. St. Paul, MN: American Phytopathological Society, pp. 243–262. Creamer, R., Luque-Williams, M., Howo, M., 1996. Epidemiology and incidence of beet curly top geminivirus in naturally infected weed hosts. Plant Disease 80, 533–535. Esau, K., Hoefert, L.L., 1978. Hyperplastic phloem in sugarbeet leaves infected with the beet curly top virus. American Journal of Botany 65, 772–783. Hanley-Bowdoin, L., Settlage, S.B., Orozco, B.M., Nagar, S., Robertson, D., 1999. Geminiviruses: Models for plant DNA replication, transcription, and cell cycle regulation. Critical Reviews in Plant Sciences 18, 71–106. Rojas, M.R., Macedo, M.A., Maliano, M.R., et al., 2018. World management of geminiviruses. Annual Review of Phytopathology 56, 637–677. Soto, M.J., Chen, L.-F., Seo, Y.-S., Gilbertson, R.L., 2005. Identification of regions of the Beet mild curly top virus (family Geminiviridae) capsid protein involved in systemic infection, virion formation and leafhopper transmission. Virology 34, 257–270. Stanley, J., Markham, P.G., Callis, R.J., Pinner, M.S., 1986. The nucleotide sequence of an infectious clone of the geminivirus beet curly top virus. EMBO Journal 5, 1761–1767. Stenger, D.C., 1998. Replication specificity elements of the Worland strain of beet curly top virus are compatible with those of the CFH strain but not those of the Cal/Logan strain. Phytopathology 88, 1174–1178.

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Strausbaugh, C.A., Eujayl, I.A., Wintermantel, W.M., 2017. Beet curly top virus strains associated with sugar beet in Idaho, Oregon, and a Western U.S. Collection. Plant Disease 101, 1373–1382. Varsani, A., Martin, D.P., Navas-Castillo, J., et al., 2014. Revisiting the classification of curtoviruses based on genome-wide pairwise identity. Archives of Virology 159, 1873–1882.

Beet Necrotic Yellow Vein Virus (Benyviridae) Sebastian Liebe, Institute of Sugar Beet Research, Göttingen, Germany Annette Niehl and Renate Koenig, Julius Kühn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany Mark Varrelmann, Institute of Sugar Beet Research, Göttingen, Germany r 2021 Elsevier Ltd. All rights reserved.

Nomenclature þ ssRNA Positive-sense single-stranded RNA ARF Auxin response factors Aux/IAA Auxin/indole-3-acetic acid BdMV Burdock mottle virus BNYVV Beet necrotic yellow vein virus BSBMV Beet soil-borne mosaic virus CP Coat protein CP-RT Read-through minor coat protein ELISA Enzyme-linked immunosorbent assay ER Endoplasmic reticulum ncRNA Noncoding RNA

Glossary Benyvirus virus.

Siglum derived from Beet necrotic yellow vein

NES Nucleolar export signal NLS Nucleolar localization signal ORF Open reading frame RdRp RNA-dependent RNA polymerase RSNV Rice stripe necrosis virus SCF Skp1-Cullin-F-box siRNAs Small-interfering RNAs TGB Triple gene block TM Transmembrane domain VSR Viral suppressor of gene silencing Znf Zinc-finger domain

Rhizomania Root beardedness (root madness): Extensive proliferation of secondary rootlets at the expense of the main tap root.

Classification Beet necrotic yellow vein virus (BNYVV) is the type species of the genus Benyvirus. This genus is the only one in the new family Benyviridae which comprises viruses with multipartite genomes encapsidated in rigid rod-shaped particles. The closest known relative is the Beet soil-borne mosaic virus (BSBMV). More distant relationships exist with Burdock mottle virus (BdMV) and Rice stripe necrosis virus (RSNV). Morphologically similar viruses are found in the family Virgaviridae. Similarities and dissimilarities between the viruses in the two families have been summarized in the 10th Report of the International Committee on Taxonomy of Viruses (ICTV).

Virion Structure BNYVV virus particles are non-enveloped and rod shaped (Fig. 1). The particles have a diameter of about 20 nm and the length varies between 65 and 390 nm depending on the encapsidated RNA species. Deviations from the upper and the lower limits may be due to end-to-end aggregation or breakage of particles. The particles have a helical structure with a central canal. The pitch of the helix is estimated to be 2.6 nm. This helix has an axial repeat of four turns, involving 49 protein subunits of the 21 kDa major coat protein (CP). Each coat protein subunit occupies four nucleotides on the RNAs of BNYVV. Immunogold labeling revealed that the 75 kDa read-through minor coat protein (CP-RT) is predominantly localized close to one end of the virus particles. It had been believed that the CP-RT is necessary to initiate the encapsidation process, but recent investigations showed that it is dispensable for particle assembly and successful virus infection at least in the experimental host Nicotiana benthamiana.

Genome Organization and Functions of Gene Products The BNYVV genome consists of four to five positive-sense single-stranded RNAs ( þ ssRNAs) (Fig. 2). Each RNA is capped at the 50 end and possesses a 30 poly(A) tail. RNA1 (6.7 kb) contains one open reading frame (ORF) encoding a replicase protein of 237 kDa (P237), which is processed by a papain-like proteinase into one 150 kDa protein with methyltransferase, helicase and protease domains and one 66 kDa protein containing the RNA-dependent RNA polymerase. Localization studies revealed an association of the P237 protein as well as domains of the 150 kDa and 66 kDa replication-associated proteins with the endoplasmic reticulum (ER). The first ORF on RNA2 (4.6 kb) encodes the CP which is terminated by a leaky UAG stop codon allowing the translation of the readthrough protein CP-RT (P75). The C-terminal part of the CP-RT contains a KTER motif at position 553–556 which is essential for transmission by the vector P. betae. Several rounds of mechanical transmission can cause deletions within this region leading to a loss

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Fig. 1 Particles of Beet necrotic yellow vein virus in a purified preparation negatively stained with uranyl acetate. Courtesy D. E. Lesemannand J. Engelmann, BBA, Braunschweig, Germany.

Fig. 2 Organization of the BNYVV genome and expression of BNYVV genes. Self-cleavage of the replicase protein (P237) is indicated by a black arrow. At 50 end, the m7Gppp is indicated by a black circle and the poly(A) tail at the 3’ end by A(n). The suppressible leaky stop codon UAG is indicate by *. Abbreviations: methyltransferase (MTR); helicase (HEL); protease (PRO); RNA polymerase (POL); coat protein (CP); readthrough domain (RTD); triple gene block (TGB).

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of vector transmissibility. It was shown that a mitochondrial targeting sequence within the CP-RT targets modified CP-RT expressing GFP and reporter proteins to the cytoplasmic face of mitochondria. Furthermore, four trans-membrane domains (TM1-TM4) have been identified by bioinformatic analysis, but their function is not clarified yet. The TM helices are apparently tightly packaged with ridge/groove arrangements between the helices and strong electrostatic association. It is supposed that these helices act as anchors to the mitochondrial membrane in plant cells, but might also facilitate the movement of virus particles across the fungal membrane separating the plasmodium from the cytoplasm. The next three ORFs on RNA2 encode the triple gene block (TGB) proteins 1–3 that are translated from subgenomic RNAs and responsible for cell-to-cell movement. TGB1 (P42) is derived from subgenomic RNA2a whereas TGB2 (P13) and TGB3 (P15) are translated from the bicistronic subgenomic RNA2b. P42 possesses an N-terminal nucleic acid-binding activity and helicase motifs at the C-terminus. Electron microscopy revealed that P13 and P15 localize to callosecontaining cell wall thickenings which suggests plasmodesmal targeting by these two proteins. It is proposed that P13 and P15 provide a docking site for a P42-viral RNA complex which modifies plasmodesmatal size exclusion limit to improve transit of the viral RNA. The last ORF on RNA2 is translated from the subgenomic RNA2c and encodes the viral suppressor of gene silencing (VSR). It is a 14 kDa cysteine-rich protein (P14) which inhibits the production of small-interfering RNAs (siRNAs), especially secondary siRNAs by interfering with RDR6-dependent siRNA generation. A zinc-finger domain (Znf) containing an embedded nucleolar localization sequence (NLS) has been shown to be crucial for silencing-suppression activity and protein stability. The protein accumulates in the nucleolus and in the cytoplasm, but it’s suppression activity is probably independent of the subcellular localization. Long-distance movement is abolished in viral mutants carrying a defective VSR. The first two RNAs of BNYVV are sufficient for virus replication, systemic movement, and symptom development in the experimental host N. benthamiana, but characteristic disease symptoms in sugar beet are only induced when RNA3 (1.8 kb) is present. Thus, RNA3 carries the pathogenicity factor of BNYVV, encoded by the 25 kDa protein (P25). P25 possesses a nuclear localization signal (NLS) and nuclear export signal (NES) allowing the protein to shuttle between the nucleus and cytoplasma. The importance of this protein for viral pathogenicity is described in the section ‘Virus-Host Relationships’. Downstream of P25 is another ORF located encoding for a 6.8 kDa protein called N gene, but its expression could not be detected under natural infection conditions. However, the N gene is translationally activated by the deletion of upstream sequences leading to a truncated RNA. Expression of this RNA induces necrotic lesions in the local lesion host Tetragonia expansa. The last ORF on RNA3 encodes a 4.6 kDa protein of unknown function that is presumably expressed from a subgenomic RNA (RNA-3 sub). Its expression has never been confirmed supporting the presumption that the ORF is cryptic. Besides encoding proteins, the RNA3 additionally has a non-coding function that is crucial for systemic movement. A ‘core’ nucleotide sequence (at nt position 1033–1257) was identified that mediates systemic movement independent of P25 expression in Beta macrocarpa. Within this region, a 20 nt long sequence stretch (‘coremin’) is crucial for systemic movement. In a study to decipher the function of the ‘coremin’ nucleotide sequence, a non-coding RNA3 (ncRNA3) was identified, which is a cleavage product that accumulates only when the coremin sequence is present. Production of the ncRNA3 appears to depend on XRN1-type, i.e., plant XRN4 exo-ribonucleases. It has been shown that the ncRNA3 acts synergistically to P14 as a weak second VSR and promotes efficient systemic host infection. The 1.4 kb RNA4 encodes a single 31 kDa protein (P31), which is essential for virus transmission by the vector P. betae. Some isolates of BNYVV (see section ‘Epidemiology’) possess an RNA5 (1.3 kb), containing a single ORF which codes for a 26 kDa protein (P26). The presence of this protein induces necrotic lesions in the host Chenopodium quinoa when it is expressed either from RNA5 or from an RNA3-derived replicon. The nucleo-cytoplasmic protein displays a strong transcriptional activity. Interestingly, the 20 nt coremin sequence is also present on RNA5 outside the ORF of P26, indicating a non-coding function of RNA5 during virus movement similar to RNA3.

Transmission BNYVV is transmitted by the soil-borne ubiquitous plasmodiophorid Polymyxa betae, which infects sugar beet to complete its life cycle. P. betae. can survive in the soil as sporosorus (cytosori), which are resting spores with a very thick and strong cell wall providing persistence in soil for many years under unfavorable conditions. When a host plant is present and sufficient soil moisture available, resting spores germinate and primary zoospores are released. Motile zoospores move through free water in the soil and start encysting on root hairs or epidermal cells of roots. Zoospores penetrate host cells and inject their cytoplasma. A multinucleate plasmodium is formed within the host cell and differentiates either into a zoosporangium releasing secondary zoospores (sporangial phase) or into a sporosorus forming resting spores (sporogenic phase). The secondary zoospores penetrate new root cells and induce plasmodium development. It is suggested that P. betae. transmits BNYVV when zoospore cytoplasm is injected into the host cell. Likewise, the virus is acquired when a plasmodium develops within an infected host cell. Secondary zoospores released from this plasmodium will be viruliferous. Similary, sporosorus developed within infected cells will likely release viruliferous primary zoospores. Due to the vector biology, BNYVV can be disseminated by soil adhering to agricultural machines, by irrigation or even by wind. All viral proteins of BNYVV were detected in resting spores and zoospores, however, whether BNYVV is able to replicate within its vector remains to be demonstrated.

Epidemiology The host range of BNYVV is limited to certain members of the plant families Amaranthaceae, Caryophyllaceae, Chenopodiacea, and Tetragoniaceae. The major natural hosts are Beta vulgaris (sugar beet) and B. macrocarpa, which are infected and systemically invaded

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only when RNA1–4 are present. Spinach (Spinacia oleracea) was also identified as natural host, but is only of minor importance. The experimental host N. benthamiana is systemically infected in the absence of RNA3 and RNA4. Local lesion hosts for virus propagation are C. quinoa and T. expansa. Infection is restricted to small chlorotic or necrotic spots without systemic movement after mechanical inoculation. BNYVV was first described in 1950 in Italy and then spread rapidly into sugar beet growing areas wordwide. Restriction fragment length polymorphism, single-strand conformation polymorphism and phylogenetic analysis allowed the identification of four major virus types, namely the A-, B-, P-, and J-type. These different types are serologically indistinguishable, but their RNAs show sequence variability ranging between 1%–5%. The virus types are geographically differently distributed. The A-type is most prevalent with many reports in Europe, USA, and Asia. The B-type has been reported in Germany, France, and other central European countries. The P-type was first described in France in the area of Pithiviers (CenterVal de Loire) and was later discovered also in the UK, Kazakhstan, and a small area in Germany. A great diversity of BNYVV types can be found especially in the Pithiviers area. Within this region, the A-, B-, and P-type are present and occur frequently in mixed infections. Moreover, P-type RNA5 was found in plants infected only with A-or B-type RNAs 1–4 indicating the formation of reassortants. Although RNA5 increases viral aggressiveness, P-type isolates without RNA5 have been reported, indicating its dispensability for viral pathogenicity. The J-type was identified in Asia and has been distinguished from the P-type by sequence analysis. The evolutionary history of BNYVV has not yet been completely resolved, but several subpopulations are thought to have diverged from at least four original types of populations that developed in native hosts in East Asia long before sugar beet was cultivated. Afterwards, BNYVV populations from different origins were introduced into sugar beet cultivation areas and spread extensively. Sequence analysis indicates that the A-and P-type are more closely related than the A- and B-type. Therefore, it is suggested that the A-and B-type became separated before the P-type evolved from the A-type.

Disease Symptoms BNYVV causes rhizomania. The term signifies root madness (Rhizo: root; Mania: madness). The disease is characterized by excessive lateral root formation (‘root beardedness’) (Fig. 3). Infected sugar beet plants can display also to some extend dwarfism with a taproot which is reduced in size, has a wine-glass like shape and low sugar content. A brownish discoloration of the vascular

Fig. 3 Characteristic rhizomania disease symptoms on taproot (A) and leaf (B) from a BNYVV infected sugar beet plant compared to a healthy plant (C and D). The white bar indicates the scale (5 cm).

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system is often seen. Leaves may become pale and have an upright position. Although symptoms are mostly confined to the root system, infected plants can display vein yellowing and necrosis on leaves after systemic movement of BNYVV to upper plant parts. The name BNYVV refers to these symptoms. Yield losses associated with disease occurrence can range between 50%–60% and are mainly due to reduced root weight and sugar content.

Virus–host Relationships The pathogenicity of BNYVV is strongly linked to the activity of P25. The protein possesses a nucleo-cytoplasmic shuttling activity allowing the virus to manipulate many cellular processes. Upon BNYVV infection, plants display massive lateral root proliferation leading to the characteristic symptom of a “root beard” only in the presence of P25. Auxin as the major plant hormone controls an array of developmental processes including the development of lateral roots. Microscopic studies of sugar beet roots revealed that differentiated pericycle cells are converted into meristematic cells during BNYVV infection. Additionally, infected plants show elevated auxin concentration in the roots substantiating the hypothesis that BNYVV interacts with the auxin signaling pathway to induce lateral root proliferation. Transgenic Arabidopsis thaliana plants expressing P25 displayed also excessive lateral root formation and elevated auxin concentration, which strongly supports the role of P25 in viral pathogenicity. Recent investigations identified an auxin/indole-3-acetic acid (Aux/IAA) transcription factor in sugar beet that interacts with the pathogenicity factor P25 of BNYVV. Aux/IAA proteins are key regulators of the auxin signaling pathway in plants. They act as transcriptional repressors by suppressing the activity of auxin response factors (ARF). ARFs are transcription factors that regulate the expression of auxinmodulated genes responsible for lateral root formation. At low auxin concentration, the transcriptional activity of ARFs is suppressed by Aux/IAA proteins, whereas high auxin concentration promotes the degradation of Aux/IAA proteins, hence releasing ARFs. Aux/IAA proteins are nucleus-localized, whereas P25 shuttles between the nucleus and the cytoplasm. It has been shown that the nuclear localization of a specific Aux/IAA protein (AUX28) is disrupted by P25 leading to an accumulation of AUX28 within the cytoplasma. It is suggested that this interaction leads to the activation of auxin-modulated genes responsible for lateral root development. This hypothesis is further supported by the observation that the expression of genes downstream of Aux/IAA proteins involved in lateral root development are highly induced upon BNYVV infection. P25 is also responsible for the development of symptoms on leaves including vein yellowing and necrosis. Furthermore, P25 interacts with an F-box family protein that is part of the Skp1-Cullin-F-box (SCF) complex and mediates protein degradation. F-box proteins determine the selectivity of the SCF complex as they recruit target proteins. The exact function of the interaction of P25 with the F-box protein is yet unknown. Interestingly, a transcription activation domain could be mapped within the amino acid residues 103 and 160 of P25. This suggests that P25 can directly activate or suppress gene expression in sugar beet; however, target genes are still to be identified. Apart from its function in virus pathogenicity, P25 is also involved in the defense response of resistant cultivars (see section ‘Prevention’).

Diagnosis Although BNYVV infection induces very characteristic symptoms, the occurrence of leaf symptoms is rare in the field making visual disease diagnosis very difficult. Therefore, serological and molecular detection tools are required for BNYVV diagnosis. Enzymelinked immunosorbent assay (ELISA) is a common and reliable method to detect the virus within plant parts most likely to be infected (e.g., lateral roots), but BNYVV types are serologically undistinguishable. PCR techniques such reverse transcription-PCR and real-time PCR are also useful for detection using BNYVV specific primers.In addition, sequencing of (cloned) PCR products or next generation sequencing after RNA/dsRNA/virion extraction allows the differentiation of the different BNYVV types and identification of mutations associated with resistance breaking. Infestation of soil can be detected by means of bait plants which are tested for the presence of BNYVV with the aforementioned methods.

Prevention Methods for controlling the vector P. betae. by means of pesticides are currently not available. Although soil fumigation with pesticides can reduce the vector population in the soil, severe environmental damage and high economic costs prohibit an application. Resistance traits against the vector were found in B. patellaris and B. procumbens, but introduction of these traits into commercial sugar beet varieties failed. Therefore, growing of virus resistant cultivars is the only strategy to control rhizomania disease. In 1983, the Holly Sugar Company identified the first resistance gene effective against BNYVV (‘Holly’ resistance). Resistance is based on a single dominant gene (Rz1) located on chromosome III. Upon BNYVV infection, Rz1 confers partial resistance by reducing virus replication. Infected plants do not show disease symptoms or yield loss but the virus can be still detected in lateral roots in low quantities. Most commercial varieties today contain the Rz1 gene. Further analysis of wild-beet germplasm allowed the identification of a second resistance gene (Rz2) in B. vulgaris subsp. maritima WB42 originating from Denmark, which encodes a CC-NBS LRR (coiled-coil nucleotide binding site, leucine rich-repeat) resistance protein. The gene is also located on chromosome III, but Rz2 appears to be based on a different mechanism than Rz1. The Rz2 gene has also been

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introduced into commercial varieties in combination with Rz2. Further resistance genes (Rz3-Rz5) located on chromosome III could be identified, but have not been introduced into commercial varieties yet. With the introduction of the Rz1 resistance gene into commercial varieties, the virus population was under a high selection pressure. Thus, reports about the emergence of resistance breaking strains in Europe and the USA appeared soon after the introduction of Rz1. Such strains can replicate to a higher level and induce disease symptoms in Rz1 resistant plants, but the resistance breaking mutations are probably associated with a certain fitness loss in susceptible cultivars. Extensive sequence analysis of viral isolates revealed high sequence variability within a stretch of four amino acids (AS67–70) in the pathogenicity factor P25. Mutation analysis with infectious cDNA clones of RNA3 showed that a specific single amino acid substitution on position 67 already mediates Rz1 resistance breaking. However, the resistance breaking ability for most of the reported sequence variants at AS67–70 is unknown and needs to be determined. Moreover, besides AS67–70, further mutations in P25 are presumably involved in resistance breaking. Interestingly, resistance breaking has so far been reported only for the BNYVV A- and P-type but not for the B-type. The P-type displays a low variability of AS67–70, but is still able to overcome the Rz1 resistance. BNYVV P-type contains an additional RNA5 encoding the pathogenicity factor P26. It is believed that resistance breaking is mediated by RNA5 rather than by mutations in P25 at AS67–70. Hence, BNYVV has probably developed different strategies to overcome Rz1 resistance.

See also: Benyviruses (Benyviridae)

Further Reading Bornemann, K., Varrelmann, M., 2013. Effect of sugar beet genotype on the Beet necrotic yellow vein virus P25 pathogenicity factor and evidence for a fitness penalty in resistance‐breaking strains. Molecular Plant Pathology 14 (4), 356–364. Chiba, S., Kondo, H., Miyanishi, M., et al., 2011. The evolutionary history of Beet necrotic yellow vein virus deduced from genetic variation, geographical origin and spread, and the breaking of host resistance. Molecular Plant-Microbe Interactions 24 (2), 207–218. Flobinus, A., Chevigny, N., Charley, P., et al., 2018. Beet necrotic yellow vein virus non-coding RNA production depends on a 50 -30 XRN exoribonuclease activity. Viruses 10 (3), 137. Flobinus, A., Hleibieh, K., Klein, E., et al., 2016. A viral noncoding RNA complements a weakened viral RNA silencing suppressor and promotes efficient systemic host infection. Viruses 8 (10), 272. Galein, Y., Legrève, A., Bragard, C., 2018. Long term management of rhizomania disease – Insight into the changes of the Beet necrotic yellow vein virus RNA-3 observed under resistant and non-resistant sugar beet fields. Frontiers in Plant Science 9, 795. Gil, J.F., Liebe, S., Thiel, H., et al., 2018. Massive up‐regulation of LBD transcription factors and EXPANSINs highlights the regulatory programs of rhizomania disease. Molecular Plant Pathology 19 (10), 2333–2348. Koenig, R., Loss, S., Specht, J., et al., 2009. A single U/C nucleotide substitution changing alanine to valine in the Beet necrotic yellow vein virus p25 protein promotes increased virus accumulation in roots of mechanically inoculated, partially resistant sugar beet seedlings. Journal of General Virology 90 (Pt 3), 759–763. Laufer, M., Mohammad, H., Christ, D.S., et al., 2018. Fluorescent labelling of Beet necrotic yellow vein virus and Beet soil-borne mosaic virus for co-and superinfection experiments in Nicotiana benthamiana. Journal of General Virology 99 (9), 1321–1330. McGrann, G.R., Grimmer, M.K., Mutasa-Göttgens, E.S., Stevens, M., 2009. Progress towards the understanding and control of sugar beet rhizomania disease. Molecular Plant Pathology 10 (1), 129–141. Pakdel, A., Mounier, C., Klein, E., et al., 2015. On the interaction and localization of the Beet necrotic yellow vein virus replicase. Virus Research 196, 94–104. Peltier, C., Hleibieh, K., Thiel, H., et al., 2008. Molecular biology of the Beet necrotic yellow vein virus. Plant Viruses 2 (1), 14–24. Peltier, C., Klein, E., Hleibieh, K., et al., 2012. Beet necrotic yellow vein virus subgenomic RNA3 is a cleavage product leading to stable non-coding RNA required for longdistance movement. Journal of General Virology 93 (5), 1093–1102.

Benyviruses (Benyviridae) Annette Niehl, Julius Kühn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany Sebastian Liebe and Mark Varrelmann, Institute of Sugar Beet Research, Göttingen, Germany Renate Koenig, Julius Kühn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany r 2021 Published by Elsevier Ltd. This is an update of R. Koenig, Benyvirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00541-0.

Nomenclature aa Amino acid(s) CP Coat protein CP-RT Coat protein-readthrough protein ELISA Enzyme-linked immunosorbent assay ER Endoplasmic reticulum GFP Green fluorescent protein kDa Kilodalton ncRNA Non-coding RNA NGS Next generation sequencing NLS Nuclear localization signal nm Nanometer(s)

Glossary Benyvirus Siglum derived from beet necrotic yellow vein virus.

nt Nucleotide(s) ORF Open reading frame PCR Polymerase chain reaction PD Plasmodesmata RdRp RNA-dependent RNA polymerase RNA Ribonucleic acid siRNAs Small-interfering RNAs ssRNA Single-stranded ribonucleic acid TGB Triple gene block TM Trans-membrane domain VSR Viral suppressor of gene silencing

Rhizomania Root beardedness (root madness). Extensive proliferation of often necrotizing secondary rootlets at the expense of the main tap root.

History Beet necrotic yellow vein virus (BNYVV) is the type species of the genus Benyvirus. Originally, it was classified as a possible member of the tobamovirus group, because its rod-shaped particles resemble those of tobamoviruses. In 1991, the rod-shaped viruses with multipartite genomes transmitted by species of the plasmodiophorid Polymyxa were separated from the monopartite tobamoviruses to form a new group, named furovirus (for ‘fungus-transmitted rod shaped viruses’). Soil-borne wheat mosaic virus (SBWMV) became the type species of this new group; BNYVV and Rice stripe necrosis virus (RSNV) were listed as possible members. Molecular studies conducted in the following years revealed that the genome organization of the members belonging to this new virus group greatly differed from each other. Thus, eventually, four new genera were created, i.e., the genus Furovirus with SBWMV as type species, the genus Benyvirus with BNYVV as type species, the genus Pomovirus with Potato mop top virus as type species, and the genus Pecluvirus with Peanut clump virus as type species. The genus Benyvirus presently comprises four definitive species, i.e., BNYVV, Beet soil-borne mosaic virus (BSBMV), RSNV and Burdock mottle virus (BdMV), and two putative members: Magnifera indica latent virus (MILV) and wheat stripe mosaic virus (WhSMV). Moreover, a number of beny-like viruses were identified by high throughput sequencing in libraries prepared from several non-plant organisms: a study on invertebrate RNA virus diversity identified a benyvirus-like replication polyprotein gene, the source of this sequence was named Hubei beny-like virus 1. Similarly, sequences resembling those in benyviral replicase genes were found in sclerotium rolfsii and rhizoctonia solani in studies aimed at identifying mycoviruses.

Virion Structure, Particle Properties and Associations of Viral Particles With Cellular Structures Benyviruses are non-enveloped, rod-shaped viruses with multipartite (bi- to pentapartite) positive-single strand RNA (ssRNA) genomes. The RNAs are encapsidated with a coat protein (CP), and in the case of BNYVV also contain the CP-read-through (CP-RT) protein as minor CP in the capsid shell. The diameter of benyvirus particles is approximately 20 nm. The rods have a central canal and usually show several length maxima ranging from approximately 80 to 400 nm depending on the RNA species encapsidated (Fig. 1). Additional length maxima may be due to end-to-end aggregation or breakage of particles. The right-handed helix of BNYVV particles has a 2.6 nm pitch with an axial repeat of four turns involving 49 subunits of the 21 kDa major CP, which consists of 188 aa. Each CP subunit

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Fig. 1 Transmission electron micrographs of beet necrotic yellow vein virus (left) and beet soil-borne mosaic virus (right) particles negatively stained with uranyl acetate. Scale bars: 100 nm. BNYVV image (left): Courtesy of D.E. Lesemann and J. Engelmann, BBA, Braunschweig, Germany. BSBMV image: Modified from Kastirr, U., Richert-Pöggeler, K., 2018. Berichte aus dem Julius-Kühn Institut. Nr. 201, under the Creative Communs Attribution License (CC BY 4.0).

occupies four nucleotides on the BNYVV RNAs. The 75 kDa CP-RT protein has been detected by immunogold-labeling on one end of the particles in freshly extracted plant sap. Green fluorescent protein (GFP)-labeled particles of BNYVV localize to the cytoplasmic face of mitochondria early during infection, but later they relocate to semi-ordered clusters in the cytoplasm. In ultrathin sections, BNYVV particles are found scattered throughout the cytoplasm or occur in aggregates. Masses of particles with different densities arranged in parallel or angle-layer arrays may be formed.

Organization of the Genome and Properties of the Encoded Proteins The functions of the BNYVV proteins and RNAs have been studied in detail. Hence, most knowledge on the genome organization of benyviruses is based on BNYVV; however, aa sequence homologies indicate functional similarities between benyvirus proteins. The BNYVV genome consists of four to five positive-sense, ssRNAs, while the other benyviruses contain only two to four genomic RNAs (Fig. 2 and Table 1). Four genomic RNA segments have been identified for BSBMV, but only two for BdMV, RSNV, WhSMV and MILV. Each benyvirus RNA is capped at the 50 end and polyadenylated at the 30 end. On RNA1, all benyviruses possess one large ORF encoding the replicase. On RNA2, BNYVV, BSBMV, RSMV, BdMV, and WhSMV all contain six ORFs, of which the first two encode CP and CP-RT proteins, and the following three encode triple gene block (TGB)1, TGB2 and TGB3 movement proteins. The cysteine-rich proteins encoded by the 30 terminal ORF may encode the viral silencing suppressors (VSR), as a VSR function has been shown for BNYVV, BSBMV and BdMV cysteine-rich proteins. An RNA3 and RNA4 has only been identified for BNYVV and BSBMV, but some BNYVV isolates have an additional RNA5. RNA3 of BNYVV and BSBMV, RNA4 of BSBMV and RNA5 of BNYVV contain the ‘coremin’ sequence, which has been assigned RNA silencing suppressor activity and a function in virus systemic movement. RNA3-encoded BNYVV P25 and BSBMV P29 are the viral pathogenicity factors responsible for symptom development in their major host sugar beet; however, BSBMV RNA3-encoded P29 more resembles BNYVV RNA5encoded P26 in sequence and function, as it resembles BNYVV P25. The 31 kDa and 32 kDa P31 and P32, encoded on RNA4 of BNYVV and BSBMV, respectively, are involved in vector transmission and symptom development. Moreover, BNYVV P31 has been shown to enhance silencing suppressor activity in roots of Nicotiana benthamiana. The BNYVV large open reading frame (ORF) for the replicase protein encoded on RNA1 is cleaved autocatalytically by a papain-like proteinase. Virus replication may take place in association with the endoplasmic reticulum (ER), as the replicationassociated proteins localize to this organelle. In in vitro systems, translation of the replication protein may start either at the first AUG at position 154 or at a downstream AUG at position 496–498. The resulting proteins of 237 and 220 kDa, respectively, contain in their N-terminal part methyl-transferase motifs, in their central part helicase and papain-like protease motifs, and in their C-terminal part RNA dependent RNA polymerase (RdRp) motifs. BNYVV RNA2 contains six ORFs, i.e., the CP gene, which is terminated by a suppressible UAG stop codon, the CP-RT protein gene, a TGB coding for proteins of 42, 13, and 15 kDa, and a gene coding for a 14 kDa cysteine-rich protein. The CP is the major capsid protein important for efficient encapsidation of the virus. Within the 75 kDa CP-RT protein, specific domains appear to be important for virion assembly and vector transmission. A KTER motif at aa position 553–556 of the 75 kDa CP-RT protein is essential for efficient transmission of the virus by P. betae. Upon repeated mechanical inoculation of the virus, deletions within the C-terminal part of CP-RT occur and the area containing this motif may be lost, which results in abolished vector transmissibility of the virus. Apart from the KTER motif, trans-membrane domains have been predicted in the CP-RT protein, which presumably aid

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Fig. 2 Benyvirus genome structure. Benyviruses contain two to five genomic RNAs with a 50 guanosylmethionine cap (black circle) and a 30 poly A tail (An). Benyviruses share the genome structure of RNA1 and RNA2 and encode similar proteins on corresponding open reading frames (ORF). RNAs 3 and 4 encode proteins with partially overlapping functions. RNA5 is only present in BNYVV. RNA1: self-cleavage of the replicase polyprotein between protease (Pro) and RNA-dependent RNA polymerase (RDRP) domains is indicated by a black arrow. Met, methyltransferase domain; Hel, helicase domain. RNA2: Broken line between ORF1 and ORF2 indicates a read-through stop codon. CP, coat protein; CP-RT, CP-readthrough protein; TGB, triple gene block; VSR, viral silencing suppressor. Arrows indicate protein synthesis from coding sequences, broken arrows and question marks indicate that the proteins corresponding to the respective ORFs have not yet been detected.

Features

Functional RNAs

RNA3 Accession number Length ORFs Encoded proteins, Function

RNA2 Accession number Length ORFs Encoded proteins, functionb

13K TGB2, movement 15K TGB3, movement 14K CRP, silencing suppression

13K TGB2, movement 15K TGB3, movement 14K CRP, silencing suppression

P11?c N?c P4.6?c Coremin, function in systemic movement Coremin, function in systemic movement and silencing suppression and silencing suppression

EU410955 1720 bp 2 P29, pathogenicity factor

42K TGB1, movement

42K TGB1, movement

D84412 1774 bp 3 P25, pathogenicity factor

JF513083 4615 bp 6 75K CP-RT, virion assembly, vector transmission 21K CP, encapsidation

JF513082 6679 bp 1 238 kDa replicasea polyprotein

BSBMV

D84411 4609 bp 6 75K CP-RT, virion assembly, vector transmission 54K CP, encapsidation

150 kDa replicase 66 kDa RDRP

D84410 6746 bp 1 237 kDa replicasea polyprotein

BNYVV

Benyvirus RNAs and encoded proteins

RNA1 Accession number Length ORFs Encoded proteins, function

RNA

Table 1

12K TGB2 15K TGB3 17K CRP

12K TGB2 13K TGB3 13K CRP

38K TGB1

20K CP

20K CP 38K TGB1

AB818899 4315 bp 6 66K CP-RT

AB818898 7038 bp 1 249 kDa replicasea poly-protein

BdMV

EU099845 4657 bp 6 74K CP-RT

EU099844 6634 bp 1 236 kDa replicasea poly-protein

RSNV

14K TGB2 13K TGB3 3K

55K TGB1

19K CP

MH151801 4901 bp 6 81K CP-RT

MH151795 6583 bp 1 232 kDa replicasea poly-protein

WhSMV

39K helicase protein (TGB1?) 15K putative movement protein (TGB2?)

KU140663 2516 bp 3 21K CP

KU140662 6725 bp 1 234 kDa replicasea poly-protein

MILV

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Coremin, function in systemic movement and silencing suppression

D63936 1320 bp 1 P26, pathogenicity factor

D84413 1465 bp 1 P31, transmission factor, symptom determinant, VSR? P13?c Coremin, function in systemic movement and silencing suppression

FJ424610 1730 bp 2 P32, transmission factor

b

Experimental evidence for autocatalytic cleavage of the replicase protein only exists for BNYVV. Protein functions only experimentally verified for BNYVV and BSBMV. c Proteins have never been detected in natural infection conditions.

a

RNA5 Accession number Length ORFs Encoded proteins, Function Functional RNAs

Functional RNAs

RNA4 Accession number Length ORFs Encoded proteins, Function

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association with distinct vector and host membranes during the viral life cycle. Three sub-genomic RNAs are derived from BNYVV RNA2. The first TGB protein (P42) is translated from sub-genomic RNA2a, the second and third TGB proteins (P13 and P15) from the bicistronic sub-genomic RNA2b, and the 14 kDa cysteine-rich protein from subgenomic RNA2c (Fig. 2). The three TGB proteins are necessary for cell-to-cell movement of the virus. The N-terminal part of the 42 kDa TGB1 protein has nucleic acidbinding activity; its C-terminal part contains helicase motifs. GFP-tagged P42 is targeted by P15 and P13 to punctuate structures associated with plasmodesmata (PD). It has been speculated that P15 and P13 provide docking sites for a P42–viral RNA complex at PD. P42 may alter the PD size exclusion limit to allow virus movement between cells. The 30 proximal ORF encodes a cysteinerich 14 kDa protein acting as an RNA silencing suppressor. P14 inhibits secondary siRNA production by interfering with RDR6dependent siRNA synthesis. BNYVV RNA3 encodes the 25 kDa P25 pathogenicity factor, which determines symptom severity, likely through reprogramming of auxin signaling within the host plant. In the major host sugar beet, P25 induces massive lateral root proliferation and vein yellowing in systemically infected leaves. The appearance of bright yellow local lesions can be observed on Tetragonia expansa. Symptoms seem to depend on the ability of P25 to nucleocytoplasmic shuttling, as mutants unable to shuttle between the nucleus and the cytoplasm fail to induce yellow lesions on C. quinoa. The following ORF on RNA 3 encodes a 6.8 kDa N gene, which is not expressed to detectable levels under natural infection conditions. However, expression of the protein is activated by the deletion of upstream sequences, which positions it closer to the 50 end of RNA3. Expression of the N gene induces the formation of necrotic lesions in BNYVV infections and also when it is expressed from Cauliflower mosaic virus. The third ORF on BNYVV RNA3 encodes a protein of 4.6 kDa, which is presumably expressed from an abundantly present sub-genomic RNA; however, a function of this protein is not yet known. RNA3 also contains a functional non-coding RNA (ncRNA), which mediates systemic movement of the virus and functions complementary to P14 as VSR. The RNA3 20 nt ‘coremin’ sequence is responsible for accumulation of the noncoding RNA, as it stalls exoribonucleases of the yeast XRN1 or plant XRN4-type and inhibits RNA processing by the enzymes. The BNYVV RNA4-encoded 31 kDa P31 protein increases the transmission rate of the virus by P. betae and increases symptom severity in N. benthamiana and in some Beta species. Moreover, experimental evidence suggests that P31 is able to suppress RNA silencing in roots. BNYVV RNA5 only occurs in some BNYVV types (P-type and J-type, see section disease, host range and epidemiology below). It enhances symptom expression in sugar beet. Addition of P-type RNA5 to RNA1 and RNA2 results in the formation of necrotic local lesions in C. quinoa. P26, the only protein expressed by RNA5, exhibits a nucleo-cytoplasmic localization similar to RNA3-encoded P25 and has been proposed to act synergistically with P25 in symptom development. As RNA5 also contains the 20 nt ‘coremin’ motif it may, in analogy to RNA3, be involved in the systemic movement of the virus. Blast searches of reference aa sequences of the different benyviruses revealed that the highest sequence similarity between the different members of the benyvirus family is found at the level of the replicase; BNYVV and BSBMV replicase proteins share 85% nt sequence identity. BdMV, RSNV, MILV, and WhSMV replicase proteins share 51%, 44%, 39%, and 31% aa sequence identity, respectively, with the BNYVV replicase protein. Also, the TGB proteins are relatively well conserved with 75%, 82%, and 66% nt identity between TGB1, TGB2 and TGB3 proteins, respectively, of BNYVV and BSBMV; 48%, 49%, and 38% nt identity between TGB1, TGB2, and TGB3 proteins, respectively, of BNYVV and BdMV, and 38%, 44%, and 30% nt sequence identity between TGB1, TGB2, and TGB3 proteins, respectively, of BNYVV and RSNV. The TGB1 and TGB2 proteins of BNYVV and WhSMV share 29% and 45% nt sequence identity, while no significant similarity is found between the TGB3 proteins of the two viruses. The TGB1 and TGB2 proteins of MILV show 27% and 39% nt sequence identity, respectively, with those of BNYVV. At the level of the CP, BNYVV and BSBMV share 59% nt sequence identity, while the CP of BdMV, RSNV, MILV, and WhSMV have 39%, 35%, 27%, and 26% nt sequence identity with BNYVV CP, respectively. The cysteine-rich proteins share only little sequence identity; here BNYVV and BSBMV have 43% aa sequence identity, BNYVV and RSMV 30% and BNYVV and BdMV 22% sequence identity. Reassortment experiments showed that BNYVV and BSBMV RNA1 and RNA2 reassortants are viable and spread systemically in N. benthamiana. The small genomic RNAs 3 and 4 were also exchangeable and systemically infected B. macrocarpa and allowed vector transmission. Moreover, rhizomania-specific taproot symptoms in sugar beet could be induced by BSBMV in the presence of BNYVV RNA3 whereas BSBMV RNA3 in combination with BNYVV RNA1 and RNA2 failed to induce taproot symptoms. This underlines the close relationship between both viruses as well as the importance of RNA3 for viral pathogenicity.

Disease, Host Range, and Epidemiology BNYVV is the causal agent of rhizomania, one of the most damaging diseases of sugar beet. Natural hosts of the virus are B. vulgaris ssp. and B. macrocarpa. Infections of sugar beet with BNYVV are mainly confined to the root system. Susceptible sugar beet varieties show an extensive proliferation of nonfunctional, necrotizing secondary rootlets, a condition described by the names ‘root beardedness’, ‘root madness’, or rhizomania (Fig. 3). The taproots are stunted, their shape tends to be constricted, and their sugar content is low. A brownish discoloration of the vascular system is often seen. Leaves may become pale and have an upright position. Under dry conditions, wilting is often observed due to insufficient water uptake by the roots. The upper parts of the plants are invaded only rarely by the virus. In infected leaves, the veins turn yellow, and occasionally become necrotic, a condition after which the virus was named (Fig. 3). In susceptible varieties, rhizomania may cause yield losses of 50% and more. Like other diseases caused by soil-borne organisms, rhizomania often occurs in patches within infected fields. BNYVV can be mechanically

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Fig. 3 Benyvirus symptoms on sugar beet. Typical symptoms on roots and leaves of BNYVV (A) and BSBMV (B) infected sugar beet. Plants were mechanically inoculated with the virus (C) shows a root and a leaf of a healthy control plant.

transmitted to many species in the Chenopodiaceae and to some species in the Amaranthaceae and Tetragoniaceae as well as to N. benthamiana. B. macrocarpa, S. oleracea and N. benthamiana become infected systemically. BNYVV occurs in sugar beet-growing areas worldwide. Molecular analyses have revealed the existence of different BNYVV types (A type, B type, P type, and J-type) that cannot be distinguished serologically. The A type is most common and occurs in Europe, in the USA, and in East Asia. The B type is prevalent in Germany, France, and other central European countries. B-type-like BNYVV is also found in East Asian countries. The P type, which usually contains an RNA5 and causes especially severe symptoms, has only been found in restricted areas in central France (Pithiviers), the UK, Germany and in Kazakhstan. The J-type has been identified in Asia. The current BNYVV types seem to have evolved from at least four original populations that developed in native hosts in East Asia and spread from there. Naturally infected local hosts in different geographic regions may harbor various types of BNYVV. Once transmission from a native host to sugar beet has been successful, further spread in beet-growing areas by machinery, irrigation, or infested soil may be very rapid. The symptoms caused by BSBMV on its natural host B. vulgaris are more variable than those produced by BNYVV. Infected roots may remain symptomless (Fig. 3) or occasionally show rhizomania-like symptoms. Infections of the upper parts of the plants occur more frequently than with BNYVV. Leaves may develop vein banding or faint mottling. When plants are dually infected with BSBMV and BNYVV, foliar symptoms appear more frequently. In general, BSBMV causes much less damage to sugar beet than BNYVV. It has been reported that, when present in mixed infections, BNYVV suppresses BSBMV. In addition, at the cellular level, BNYVV and BSBMV remain spatially separated. BSBMV can be transmitted mechanically to C. quinoa, Chenopodium album, and T. tetragonioides, all of which become infected locally, and to Beta maritima, which becomes infected systemically. BSBMV is widely distributed in the USA, but has so far not been found in other parts of the world. Analyses of polymerase chain reaction (PCR) products indicated that this virus is genetically much more variable than BNYVV. This may explain the variability in the symptoms produced by this virus.

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RSNV was identified in 1983 in West Africa (Ivory Coast) as the causal agent of a long-known disease named ‘rice crinkling disease’. The percentage of infection varies greatly between varieties and in different years. RSNV also occurs in additional African countries, so in Liberia, Nigeria, and Sierra Leone. Recently, RSNV was identified in Burkina Faso (2014), Benin (2015), and Mali (2017). In South America, the virus was first noticed in Columbia, where it causes a severe disease of rice named ‘entorchamiento’ that is characterized by seedling death, foliar striping, and severe plant malformation and is also present in Ecuador, Brazil and Panama. Recently, (2018) RSNV was also reported in Argentina. The virus can be transmitted mechanically to C. quinoa, C. album, and Chenopodium amaranticolor where it produces local lesions, but not to rice. N. benthamiana does not become infected. BdMV has been isolated from leaves of naturally infected burdock (Arctium lappa L) plants in Japan in 1970. Leaves as well as roots of this plant are common vegetables in Japan. BdMV causes mild chlorosis or mottling symptoms on leaves of burdock plants, the only known natural host of the virus. The symptoms can become masked in older plants. BdMV can be transmitted mechanically to C. quinoa, C. murale, N. clevelandii, and N. rustica, which are infected systemically, and to B. vulgaris var. cicla and var. rapa, S. oleracea, Cucumis sativus, and T. expansa, which are infected only locally, some of them only with difficulty. So far, BdMV has only been reported to occur in Japan. WhSMV has only recently (2019) been isolated from wheat plants in Brazil showing chlorotic to necrotic stripes on the leaves, symptoms of soil-borne wheat mosaic disease typically associated with SBWMV and wheat spindle streak mosaic virus, and causes strong crop losses in Brazil.

Transmission BNYVV, BSBMV and presumably also RSNV and WhSMV are transmitted by species of the obligate biotrophic root parasite Polymyxa. While direct evidence for the transmission of BNYVV and BSBMV by Polymyxa betae exists through transmission studies, the evidence, that RSNV and WhSMV are transmitted by P. graminis remains correlative, based on the consistent occurrence of the pathogen in virus infected roots. Whether the other members of the benyvirus family are also transmitted by plasmodiophorids is not yet known. RSNV-harboring cystosori may be carried in soil adhering to the surface of rice seeds, but true seed transmission has not been observed. Under experimental conditions, benyviruses are readily transmitted mechanically among test plants. However, attempts to transmit BNYVV mechanically from sugar beet rootlets to test plants may not always be successful. The Polymyxa life cycle begins with the encystment of a zoospore on a host plant root epidermal or root hair cell, injection of the spore content into the host cytoplasm and the formation of a plasmodium inside the host cell. It is believed that virus transmission to the host and acquisition of virus from the host takes place during this plasmodial phase. The plasmodium then differentiates either into a zoosporangium or into resting spores; the mechanisms driving entry into a zoosprangial or sporogenic phase are not yet understood. BNYVV virus-like particles were found inside P. betae zoosporangia and proteins associated with viral replication and movement have been detected inside P. betae zoospores and resting spores, allowing the hypothesis that virus replication and movement may take place within Polymyxa. Whether Polymyxa may indeed be a host for the virus remains to be shown. Virus present in Polymyxa resting spores remains infective for many years. The spores may be distributed on agricultural equipment, by irrigation, or even by wind.

Diagnosis The presence of symptoms typical for BNYVV is a good first indicator for benyvirus infection. Root symptoms, i.e., rootlet proliferation and necrosis of vascular bundles are considered reliable indicators of infection. However, symptoms on the foliage of plants in the field can easily be confused with those resulting from other causes, e.g., infection with other viruses, nematodes, fungi, or abiotic stress such as nitrogen deficiency. Thus, benyvirus infections are diagnosed most reliably by serological or molecular biological techniques, bioassays with indicator plants, electron microscopy or a combination of these techniques. Benyviruses are moderately to strongly immunogenic. Polyclonal antibodies have been obtained for BNYVV, BSBMV, and RSNV. BNYVV and BSBMV are only very distantly serologically related – some antisera may detect only one of the two viruses. Particles of RSNV and BdMV do not react with antisera raised against BNYVV. Monoclonal antibodies against BNYVV have been used for diagnostic purposes and for determining the accessibility of aa on the virus particles. Single-chain antibody fragments have also been produced and used to detect BNYVV. Enzyme-linked immunosorbent assay (ELISA), immunoelectron microscopy, tissueprint immunoblotting, western blot, and immunocapture PCR are serological methods commonly used for the detection of benyviruses. Molecular biology techniques based on RT-PCR or sequencing are also routinely applied for the diagnosis of benyviruses. Especially the identification of the putative new benyvirus member species has been achieved by next generation sequencing (NGS) technology. An advantage of PCR-based methods alone or in combination with sequencing of PCR products as well as of NGS is that they allow the discrimination of different virus types or strains, which are serologically indistinguishable and, in the case of NGS, allow the identification of viruses without the requirement of a priori sequence information. Electron microscopy techniques, i.e., transmission electron microscopy, immunoelectron microscopy, or immunogold labeling are also possibilities to diagnose benyvirus infection. Bioassays relying on mechanical inoculation of indicator plants are also useful, though laborious. Infestation of soil may be detected by means of bait plants, which are planted into contaminated soil and tested for the presence of virus, usually by ELISA or with PCR-based methods.

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Control Polymyxa-transmitted plant viruses may survive in soil in the long-living resting spores of the vector for many years, probably even decades. The diseases caused by these viruses are therefore difficult to control. Chemical approaches, e.g., by soil treatment with methyl bromide, is neither efficient nor acceptable for economic and ecological reasons. Growing resistant or tolerant varieties currently represents the only practical and environmentally friendly means to lower the impact of these diseases on yield. BNYVVtolerant or partially resistant sugar beet varieties are available which enable good yields also on infested fields. Their resistance is based mainly on the Rz1 gene from a sugar beet line (‘Holly’ resistance) and the Rz2 gene from the B. maritima line WB42. Rz1 resistance-breaking BNYVV strains have been reported, but presumably lose their advantage in susceptible varieties. The resistance breaking properties have been associated with aa in the P25 pathogenicity factor as well as with the presence of RNA5. Additional resistance genes (Rz3 to Rz5) have been identified, but have not yet been introduced into commercial varieties. Among the different known loci conferring resistance to BNYVV, all located on Chromosome III in B. vulgaris, only Rz2 has been identified to encode a CC-NBS-LRR (coiled-coil, nt binding site, leucine rich-repeat) resistance protein. These proteins typically act to restrict virus infection during effector-triggered immunity. Genetically modified sugar beet expressing various portions of the BNYVV genome have been shown to be highly resistant to BNYVV. Resistance against the vector has been detected in B. patellaris and B. procumbens, but attempts to develop agronomically acceptable sugar beet cultivars resistant to P. betae have so far failed. The degree of resistance or tolerance to RSNV differs in different rice cultivars. A high degree of resistance is found in Oryza glaberrima. Studies with O. glaberrima x O. sativa introgression lines allowed the identification of an RSNV resistance locus on chromosome 11. This resistance locus has so far not been introduced into commercial varieties. Apart from growing resistant plants, prevention methods consisting of stringent hygiene and decontamination procedures for contaminated material and instruments may help to limit further spread of the pathogen.

Similarities and Dissimilarities With Other Taxa The morphology of benyviruses resembles that of other rod-shaped viruses, that is, of furoviruses, goraviruses, hordeiviruses, pecluviruses, pomoviruses, tobamoviruses and tobraviruses. Phylogenetic relationships between benyviruses and viruses from other genera and similarities with other taxa have been extensively reviewed in the 10th report of the International Committee on Taxonomy of Viruses. The benyvirus genomes are multipartite with two to five RNAs. Tobamoviruses have monopartite, goraviruses, furoviruses, pecluviruses, and tobraviruses bipartite, and hordeiviruses and pomoviruses tripartite genomes. The fact that the RNAs of benyviruses are polyadenlylated differentiates them from the RNAs of the other viruses listed above. Opposed to the other rod-shaped viruses, which encode their replication-associated proteins in two ORFs, benyviruses have a single large ORF on their RNA1 coding for a polypetide, which is cleaved post-translationally into two replication-associated proteins. The replicationassociated ORFs of hordeiviruses are located on two different RNAs; in the case of furoviruses, gora-, peclu-tobamo-, and tobraviruses, the ORFs coding for replication proteins are located on the same RNA, where ORF1 is terminated by a leaky stop codon and extends into an ORF2. The methyl-transferase, helicase, and RNA-dependent RNA polymerase motifs in the benyvirus replicase display higher similarity to e.g., Hepatitis virus E and Rubella virus than to other rod-shaped plant viruses, and database similarity searches of benyviral replicase sequences reveal similarities to sequences in plant-, fungal- and invertebrate-derived NGS sequencing libraries. Taken together, this supports a model in which benyviruses may have specialized from ancestors infecting a wide range of hosts. Like hordei-, gora-, peclu-, and pomoviruses, benyviruses have their movement function encoded on a TGB. Sequence comparisons reveal similarities within the first and second TGB-encoded proteins of hordei- and pomoviruses as well as to carla- and potexviruses. With respect to structural motifs in RNA, it is interesting to note that the ‘coremin’ motif present on BNYVV and BSBMV RNAs is also present on Cucumber mosaic virus RNAs. One of the functions assigned to this motif is stalling of host exonucleases, resulting in the production of ncRNAs with functions associated with virus movement and silencing suppression.

Further Reading Biancardi, E., Tamada, T. (Eds.), 2016. Rhizomania. Cham: Springer. Chiba, S., Hleibieh, K., Delbianco, A., et al., 2012. The benyvirus RNA silencing suppressor is essential for long-distance movement, requires both zinc-finger and NoLS basic residues but not a nucleolar localization for its silencing-suppression activity. Molecular Plant-Microbe Interactions 26, 168–181. Chiba, S., Kondo, H., Miyanishi, M., et al., 2011. The evolutionary history of Beet necrotic yellow vein virus deduced from genetic variation, geographical origin and spread, and the breaking of host resistance. Molecular Plant-Microbe Interactions 24, 207–218. Commandeur, U., Koenig, R., Manteuffel, R., et al., 1994. Location, size, and complexity of epitopes on the coat protein of Beet necrotic yellow vein virus studied by means of synthetic overlapping peptides. Virology 198, 282–287. D'Alonzo, M., Delbianco, A., Lanzoni, C., et al., 2012. Beet soil-borne mosaic virus RNA-4 encodes a 32 kDa protein involved in symptom expression and in virus transmission through Polymyxa betae. Virology 423, 187–194. Flobinus, A., Chevigny, N., Charley, P.A., et al., 2018. Beet necrotic yellow vein virus noncoding RNA production depends on a 5'-3' Xrn exoribonuclease activity. Viruses 10, 137.

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Flobinus, A., Hleibieh, K., Klein, E., et al., 2016. A viral non-coding RNA complements a weakened viral RNA silencing suppressor and promotes efficient systemic host infection. Viruses 8, 272. Hehn, A., Fritsch, C., Richards, K.E., Guilley, H., Jonard, G., 1997. Evidence for in vitro and in vivo autocatalytic processing of the primary translation product of Beet necrotic yellow vein virus RNA 1 by a papain-like proteinase. Archives of Virology 142, 1051–1058. Jupin, I., Guilley, H., Richards, K.E., Jonard, G., 1992. Two proteins encoded by Beet necrotic yellow vein virus RNA3 influence symptom phenotype on leaves. The EMBO Journal 11, 479–488. Laufer, M., Mohammad, H., Christ, D.S., et al., 2018. Fluorescent labelling of Beet necrotic yellow vein virus and Beet soil-borne mosaic virus for co- and superinfection experiments in Nicotiana benthamiana. Journal of General Virology 99, 1321–1330. Rahim, M.D., Andika, I.B., Han, C., Kondo, H., Tamada, T., 2007. RNA-4 encoded p31 of Beet necrotic yellow vein virus is involved in efficient vector transmission, symptom severity and silencing suppression in roots. Journal of General Virology 88, 1611–1619. Scholten, O.E., Jansen, R.C., Paul Keizer, L.C., De Bock, T.S.M., Lange, W., 1996. Major genes for resistance to Beet necrotic yellow vein virus (BNYVV) in Beta vulgaris. Euphytica 91, 331–339. Shi, M., Lin, X.-D., Tian, J.-H., et al., 2016. Redefining the invertebrate RNA virosphere. Nature 540, 539. Tamada, T., Schmitt, C., Saito, M., et al., 1996. High resolution analysis of the readthrough domain of Beet necrotic yellow vein virus readthrough protein: A KTER motif is important for efficient transmission of the virus by Polymyxa betae. Journal of General Virology 77, 1359–1367. Valente, J.B., Pereira, F.S., Stempkowski, L.A., et al., 2019. A novel putative member of the family Benyviridae is associated with Soilborne wheat mosaic disease in Brazil. Plant Pathology 68, 588–600.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/w/benyviridae/648/genus-benyvirus Genus: Benyvirus Benyviridae Positive-sense RNA Viruses.

Betaflexiviruses (Betaflexiviridae) Nobuyuki Yoshikawa, Iwate University, Morioka, Japan Hajime Yaegashi, Institute of Fruit Tree and Tea Science, NARO, Morioka, Japan r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) AlkB Alpha-ketoglutarate-dependent dioxygenase CP Coat protein or capsid protein dsRNA double-stranded ribonucleic acid Hel Helicase kb Kilobases; the size of a ssDNA or ssRNA molecule kbp Kilobase pairs; the size of a dsDNA or dsRNA molecule kDa Kilo Daltons; the size of a protein Met Methyl-transferase MP Movement protein

Glossary Alfavirus A genus in the family Togaviridae. The type species is Sindbis virus. Helical symmetry A form of capsid structure found in many RNA viruses in which the protein subunits which interact with the nucleic acid form a helix. Movement protein A virus-encoded protein which is essential for the cell-to-cell movement of the virus in plant tissues. RNA silencing suppressor Viral protein that suppresses the antiviral RNA silencing response by the host. Semi-persistent transmission The relationship between a plant virus and its arthropod vector which is intermediate between non-persistent transmission and persistent

NBP RNA-binding protein nm Nanometer(s) nt Nucleotide(s) ORF Open reading frame Pol RNA-dependent RNA polymerase P-Pro Papain-like protease Rep Replication initiator protein S20,w Corrected sedimentation coefficient; extrapolated in water at 201C and infinitely diluted sgRNA sub-genomic RNA ssRNA single-stranded ribonucleic acid TGB Triple gene block

transmission. It has the features of short acquisition feed and no latent period found in non-persistent manner, but the vector remains able to transmit the virus for periods of hours to days which is longer than the non-persistent manner. Sub-genomic RNA A segment of RNA generated from a genomic RNA via an international promoter that has the same 30 end as the genomic RNA, but has a deletion at the 50 end. The sub-genomic RNA makes it possible to efficiently translate the downstream open reading frame of the genomic RNA. The 30K superfamily The group of movement proteins related to the 30 kDa ('30K') movement protein of Tobacco mosaic virus.

Introduction Betaflexiviridae is a family of viruses in the order Tymovirales and contains viruses infecting plants and sharing a distinct lineage of alphavirus-like replication proteins. The viruses in this family have flexuous particles composed of a single stranded (plus-sense), polyadenylated RNA genome, and a single coat protein. In the ICTV 9th report (2009 release), six genera (Capillovirus, Carlavirus, Citrivivirus, Foveavirus, Trichovirus, and Vitivirus) were established in this family. Now in 2019, two subfamilies and six new genera have been added, and the family Betaflexiviridae now consists of two subfamilies and 12 genera including 107 species, and 27 tentative species in the family.

Taxonomy and Phylogeny The members of the family Betaflexiviridae are classified in two subfamilies Quinvirinae and Trivirinae and twelve genera: four genera; Carlavirus, Foveavirus, Robigovirus in the subfamily Quinvirinae, and eight genera; Capillovirus, Chordovirus, Citrivirus, Divavirus, Prunevirus, Tepovirus, Trichovirus, Vitivirus, and Wamavirus in Trivirinae in the subfamily Trivirinae (Table 1). Three species in the subfamily Quinvirinae; Banana mild mosaic virus, Banana virus X, and Sugarcane striate mosaic-associated virus are not assigned to any genera yet. The viruses in the subfamily Quinvirinae have the ‘triple gene block’ (TGB) as the proteins involved in cell-to-cell movement. On the other hand, the members in the subfamily Trivirinae have a single movement protein (MP) of the 30K superfamily (Table 2). The genus demarcation criteria in this family are less than 45% nt sequence identity and 40% aa identity in the replication-associated protein (Rep) and coat protein (CP), and the different species within the same genus should have less than 72% nt identity (or 80% aa identity) in the Rep and CP genes.

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Table 1

Subfamilies, genera, and species in the family Betaflexiviridae. Type species are written in bold

Subfamilies

Genera

Species

Acronyms

Accession#

Quinvirinae

Carlavirus

Carnation latent virus Aconitum latent virus American hop latent virus Atractylodes mottle virus Blueberry scorch virus Butterbur mosaic virus Cactus virus 2 Caper latent virus Chrysanthemum virus B Cole latent virus Coleus vein necrosis virus Cowpea mild mottle virus V Cucumber vein-clearing virus Daphne virus S Gaillardia latent virus Garlic common latent virus Helenium virus S Helleborus mosaic virus Helleborus net necrosis virus Hippeastrum latent virus Hop latent virus Hop mosaic virus Hydrangea chlorotic mottle virus Kalanchoe latent virus Ligustrum necrotic ringspot virus Ligustrum virus A Lily symptomless virus Melon yellowing-associated virus Mirabilis jalapa mottle virus Narcissus common latent virus Nerine latent virus Passiflora latent virus Pea streak virus Phlox virus B Phlox virus M Phlox virus S Poplar mosaic virus Potato latent virus Potato virus H Potato virus M Potato virus P Potato virus S Red clover vein mosaic virus Sambucus virus C Sambucus virus D Sambucus virus E Shallot latent virus Sint-Jan onion latent virus Strawberry pseudo mild yellow edge virus Sweet potato C6 virus Sweet potato chlorotic fleck virus Verbena latent virus Yam latent virus Alfalfa latent virus Chrysanthemum virus R Edelberry carlavirus Edelberry carlavirus A Jasmine virus C Narcissus latent virus Opuntia virus H Pepper virus A

CLV AcLV AHLV AtMV BlScV ButLV CV  2 CapLV CVB CoLV CVNV CPMMV CuVCV DVS GalLV GarCLV HVS HeMV HNNV HiLV HpLV HpMV HdLV KLV LNRSV LVA LSV MYaV MjMV NCLV NeLV PLV PeSV PhlVB PhlVM PhlVS PopMV PotLV PVH PVM PVP PVS RCVMV SVC SVD SVE SLV SJOLV SPMYV SPC6V SPCFV VeLV YLV ALV CVR EV EVA JVC NLV OV2 PVA

NC038865 NC002795 NC017859 NC038966 NC003499 NC013527 ND NC043080 NC009087 NC038322 NC009764 NC014730 NC043081 NC008020 NC023892 NC016440 NC038323 NC043082 NC012038 NC011540 NC00255 NC010538 NC012869 NC013006 NC010305 NC031089 NC005138 NC038324 NC016080 NC008266 NC028111 NC008292 NC027527 NC009991 NC04308 NC009383 NC005343 NC011525 NC018175 NC001361 NC009759 NC007289 NC012210 NC029087 NC029088 NC029089 NC003557 NC043084 ND NC018448 NC006550 NC043085 NC026248 NC026616 NC04070 NC029086 NC029085 NC030926 NC008552 LS999823 NC034376

Unassigned

Betaflexiviruses (Betaflexiviridae)

Table 1 Subfamilies

Continued Genera

Species

Acronyms

Accession#

Foveavirus

Apple stem pitting virus Apricot latent virus Asian prunus virus 1 Asian prunus virus 2 Grapevine rupestris stem pitting-associated virus Grapevine virus T Peach chlorotic mottle virus Rubus canadensis virus 1 Apple green crinkle virus Asian prunus virus 3

ASPV ApLV APV1 APV2 GRSPaV GVT PCMoV RuCV  1 AGCV APV3

NC003462 NC014821 NC025388 NC028868 NC001948 NC035203 NC009892 NC019025 NC018714 NC028975

Cherry necrotic rusty mottle virus African oil palm ringspot virus Cherry green ring mottle virus Cherry rusty mottle associated virus Cherry twisted leaf associated virus Banana mild mosaic virus Banana virus X Sugarcane striate mosaic-associated virus

CNRMV AOPRV CGRMV CRMaV CTLaV BanMMV BaVX SCSMaV

NC002468 NC012519 NC001946 NC020996 NC024449 NC002729 NC04308 ND

Apple stem grooving virus Cherry virus A Currant virus A Mume virus A Birch capillovirus Yacon virus A

ASGV CVA CuVA MuVA BCV YVA

NC001749 NC003689 NC029301 NC040568 MK402233 NC030657

Unassigned

Carrot Ch virus 1 Carrot Ch virus 2 Lettuce chordovirus 1

CChV  1 CChV  2 LeCV1

NC025469 NC025468 NC040627

Citrivirus Unassigned

Citrus leaf blotch virus Citrus leaf blotch virus 2

CLBV CLBV2

NC003877 MH144344

Divavirus

Diuris virus A Diuris virus B Hardenbergia virus A Occimum basilicum RNA virus 1 Apricot vein clearing associated virus Actinidia seed borne latent virus Caucasus prunus virus Potato virus T Prunus virus T Zostera virus T Apple chlorotic leaf spot virus Apricot pseudo-chlorotic leaf spot virus Cherry mottle leaf virus Grapevine Pinot gris virus Grapevine berry inner necrosis virus Peach mosaic virus Phlomis mottle virus Fig latent virus 1 Peach chlorotic leaf spot virus Peach virus M

DiVA DiVB HarMV OBRV1 AVCaV ASbLV CPV PVT PrVT ZVT ACLSV APCLSV CMLV GPGV GINV PcMV PhMV FLV  1 PCLSV PeVM

NC019029 NC019030 NC015395 NC035462 NC023295 NC040800 NC038325 NC011062 NC024686 MK514426 NC001409 NC006946 NC002500 NC015782 NC015220 NC011552 NC043412 FN377573 MH084695 MK012336

Grapevine virus A Actinidia virus A Actinidia virus B Arracacha virus V Blackberry virus A Grapevine virus B

GVA AcVA AcVB AVV BVA GVB

NC003604 NC043087 NC016404 NC03426 NC040630 NC003602 (Continued )

Unassigned

Robigovirus

Unassigned

Trivirinae

231

Capillovirus

Unassigned

Chordovirus

Unassigned Prunevirus

Tepovirus Unassigned Trichovirus

Unassigned

Vitivirus

232

Betaflexiviruses (Betaflexiviridae)

Table 1

Continued

Subfamilies

Genera

Unassigned

Wamavirus

Table 2 Subfamily

Species

Acronyms

Accession#

Grapevine virus D Grapevine virus E Grapevine virus F Grapevine virus G Grapevine virus H Grapevine virus I Grapevine virus J Heracleum latent virus Mint virus 2 Agave tequilana leaf virus Arracacha latent virus V Grapevine virus K Grapevine virus L Grapevine virus M

GVD GVE GVF GVG GVH GVI GVJ HLV MV  2 AtLV ALVV GVK GVL GVM

NC038326 NC011106 NC018458 NC040616 NC040545 NC037058 NC040564 NC039087 NC043088 NC034833 KY451036 NC035202 MH248020 MK492703

Watermelon virus A

WVA

NC034377

Distinguishing properties of sub-families and genera in the family Betaflexiviridae Genus

Virion length (nm)

Genome size (kb)

No. of ORFs

Unknown proteine

Proteins Repa

MP (s)b

CPc

RBPd

Quinvirinae

Carlavirus Foveavirus Robigovirus

610–700 800 800

5.8–9 8.4–9.3 8–8.5

6 5 5

215–225 230–250 209–232

TGB TGB TGB

32–36 28–44 29–30

11–16 – –

Trivirinae

Capillovirus Chordovirus Citrivivirus Dvavirus Prunevirus Trichovirus Tepovirus Vitivirus Wamavirus

640–700 640–760 960 640 960 640–890 640 725–785 –

6.5–7.5 8 8.7 6.5–7.5 8.7 7.5–8 6.5 7.6 8.4

2 3 3 2 4 3 or 4 3 5 4

210–245 213 227 237 193 215–220 206 190–200 212

30Ksf 30Ksf 30Ksf 30Ksf 30Ksf 30Ksf 30Ksf 30Ksf 30Ksf

25–27 24 41 20 25 21–24 25 18–22 27

– – – – 16 (15–16)f – 11–14

19 25

a

Replication-associated protein (kDa). Movement protein either of the 30K superfamily (30Ksf) or a triple gene block (TGB). c Coat protein (kDa). d RNA-binding protein (kDa). e The hypothetical protein of unknown function (KDa). f The presence of RBP is depending on the virus species. b

Phylogenetic analysis of the aa sequences of the Rep (POL) and CP of 106 members in the family Betaflexiviridae are shown in Fig. 1(a) and (b). A phylogenetic tree constructed using POL sequences showed that the two subfamilies are separated and members in most genera are grouped into distinct clusters (Fig. 1(a)). As an exception, ASGV, a type species of the genus Capillovirus, and another three species (Cherry virus A, Currant virus A, and Mume virus A) are separated into different clusters. In a phylogenetic tree using the CP sequence (Fig. 1(b)), members of the genus Foveavirus are grouped into two different clusters which were divided into two subfamilies. Phylogenetic trees based on aa sequences of TGB1 proteins for the members of the subfamily Quinvirinae and MPs of the members of the subfamily Trivirinae are shown Fig. 1(c) and (d), respectively. In the subfamily Quinvirinae -TGB1, members of the genus Carlavirus are divided into several clusters (Fig. 1(c)).

Members of the Family The number of virus species within each genus varies greatly depending on the genus, the largest genus Carlavirus contains 53 species and the smallest ones Citrivirus and Wamavirus consist of only one virus species (Table 1). There is another 27 betaflexiviruses that are tentative members for some genera, and for which full sequences are already available (Table 1).

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Fig. 1 Phylogenic relationships among virus species in the family Betaflexiviridae. Phylogenic tree was constructed by CLUSTAL W based on the amino acid sequences of the polymerase of viruses in the family Betaflexiviridae (1a), the coat protein of viruses in the family Betaflexiviridae (1b), the TGB1 protein of viruses in the subfamily Quinvirinae, and movement protein (MP) of viruses in the subfamily Trinivirinae. The POL amino acid sequences of 106 viruses classified in the family Betaflexiviridae (approved and tentative) were collected from Ref-seq database at July 2019. Virus species in each genus were selected from the virus reference sequence collection of the gene bank. The abbreviations of the viruses are listed in Table 1.

Virion Structure Virions are non-enveloped, flexuous filamentous particles, 610–960 nm long and 10–13 nm in diameter, depending on the genus and species (Table 2). They have helical symmetry with a pitch of about 3.4 nm (range 3.3–3.7 nm). Electron micrographs of members of the genera Potyvirus (Potyviridae), Potexvirus, (Alphaflexiviridae), Closterovirus (Closteroviridae), Carlavirus, Foveavirus, Capillovirus, Trichovirus, and Vitivirus (the last five are in the family Betaflexiviridae) suggests that the flexibility and surface structure of the virus particles are not significantly different between Zucchini yellow mosaic virus (Potyvirus), Potato virus X (Potexvirus), and three carlaviruses (Potato virus S, Southern potato latent virus, and Strawberry pseudo mild yellow edge virus). On the other hand, the particle flexibility and cross-banding structure of Apple stem pitting virus (ASPV) (Foveavirus), Apple stem grooving virus

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Betaflexiviruses (Betaflexiviridae)

Fig. 1 Continue.

(ASGV) (Capillovirus), Apple chlorotic leaf spot virus (ACLSV) (Trichovirus), and Grapevine virus A (Vitivirus) resembles to those of Citrus tristeza virus (Closterovirus) (Fig. 2). Virions of the members in the Betaflexiviridae sediment as a single species with a sedimentation coefficient (S20,w) of 92–176S, depending on the genus and species. Virions are composed of a linear positive-sense, single-stranded RNA (ssRNA) of 5.8–9.3 kb which is 5%–6% by weight of the virion and a single coat protein of Mr of 18–44 kDa (Table 2). The 30 terminus of the genomic RNA has a poly(A)-tail and the 50 terminus has a cap structure. The particles of ASPV (Foveavirus) readily form end-to-end aggregates with four prominent peaks appearing at 800 nm, 1600 nm, 2400 nm, and 3200 nm in length in purified preparation. ACLSV particles require the presence of divalent cations (for example, Mg2 þ ) to maintain the integrity of the quaternary structure. Potato virus T (Tepovirus) particles stained with uranyl formate showed cross-banding structure similar to ASGV (Capillovirus), whereas those stained with uranyl acetate showed ‘criss-cross’ and rope structure.

Genome Organization The number of open reading frames (ORFs) encoded by viral genomes is between two and six depending on the genus (Table 2 and Fig. 3). In all species, the ORF1-encoded product has homologies with polymerase proteins of the 'alphavirus-like' supergroup of RNA viruses. This protein (Rep) (190–250 kDa) contains the conserved domains for methyl-transferase (Met), helicase (Hel), and RNA-dependent RNA polymerase (Pol) activity (Fig. 3). Most members also have a DNA alkylation damage repair protein (AlkB) and/or a papain-like protease (P-pro) domains between the Met and Hel (Fig. 3). Smaller ORFs encode the proteins involved in cell-to-cell movement, either a 'triple gene block' (TGB) (Carlavirus, Foveavirus, and Robigovirus) or a single MP of the '30K' superfamily (Capillovirus, Chordovirus, Citrivirus, Divavirus, Prunevirus, Tepovirus, Trichovirus, and Vitivirus) (Table 2 and Fig. 3). These are usually located following (30 proximal) the polymerase but in the genomes of capilloviruses and divaviruses, ORF2 (MP) is nested within the ORF1 (Fig. 3). For vitiviruses, an extra ORF with unknown function is present between the polymerase and MP genes. Watermelon virus A (Wamavirus) also have an additional protein of unknown function between MP and CP (Fig. 3). The CP

Betaflexiviruses (Betaflexiviridae)

1

235

7

2

3

8

4

5

9

10

6

Fig. 2 Electron micrographs of (1) Zucchini yellow mosaic virus (Potyvirus), (2) Potato virus X (Potexvirus), (3) Potato virus S (Carlavirus), (4) Southern potato latent virus (Carlavirus), (5) Strawberry pseudo mild yellow edge virus (Carlavirus), (6) Citrus tristeza virus (Closterovirus), (7) Apple stem pitting virus (Foveavirus), (8) Apple chlorotic leaf spot virus (Trichovirus), (9) Grapevine virus A (Vitivirus), and (10) Apple stem grooving virus (Capillovirus). Bars represent 100 nm. (7) Courtesy of H. Koganezawa, (9) Courtesy of J. Imada.

gene always follows a single MP or TGB except a member of the genus Wamavirus. There is also another ORF at the 30 terminus in the genera Carlavirus, Prunevirus, and Vitivirus and in some species of Trichovirus. This latter ORF encodes a putative RNA-binding protein (NBP) with a zinc binding finger motif and the ability to bind nucleic acids (Fig. 3).

Properties and Functions of Gene Products All members in the family Betaflexiviridae encode functional proteins for replication (Rep), cell-to-cell movement (a single MP or TGB), and protection of the genome RNA (CP). A small RNA binding protein is also encoded in all, or some members, in the genera Carlavirus, Prunevirus, Vitivirus, and Trichvirus. As an additional function, an RNA silencing suppressor activity is found in some proteins. For PVM in the genus Carlavirus, both a TGP protein 1 and a cysteine-rich 11K protein (ORF6) act as viral suppressor for local and systemic silencing, respectively. An ORF5 protein of Grapevine virus A (GVA) in the genus Vitivirus also suppresses local and systemic silencing. On the other hand, MPs of Citrus leaf blotch virus in the genus Citrivivirus and ACLSV in the genus Trichovirus have activity of silencing suppressors in which the former suppresses both local and systemic silencing, and the latter inhibits only systemic silencing.

Replication and Propagation Members in the family Betaflexiviridae are alleged to replicate in the cytoplasm of infected cells. Virus particles are observed as aggregates in the cytoplasm but not frequently. Viral genomic RNA is infectious and serves as a viral messenger RNA. The ORF1 protein (Rep) is translated directly from the genomic RNA and probably the products are proteolytically processed by a P-Pro in Rep for maturation. The other ORFs located in the 30 terminus of the genome are expressed from sub-genomic mRNAs (sgRNAs). Northern blot hybridization analysis of virus-specific double-stranded (dsRNA) and ssRNAs in ASPV (Foveavirus)-infected tissues showed the presence of five distinct dsRNAs (ca. 9 kbp, 7.5 kbp, 6.5 kbp, 2.6 kbp, and 1.6 kbp) and three ssRNA (ca. 9 kb, 2.6 kb, and 1.6 kb) in infected tissues (Fig. 4). The slowest migrating ssRNA (9 kb) was equivalent to that of the ASPV genome,

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Betaflexiviruses (Betaflexiviridae)

Fig. 3 Gemone organization of representatives of virus species from the twelve genera in the family Betaflexiviridae. GaLV, Garlic latent virus; ASPV, Apple stem pitting virus; CNRMV, Cherry necrotic rusty mottle virus; ASGV, Apple stem grooving virus; CtChV1, Carrot Ch virus 1; CLBV, Citrus leaf blotch virus; DiVA, Diuris virus A; ApVCaV, Apricot vein clearing associated virus; PVT, Potato virus T; ACLSV: Apple chlorotic leaf spot virus; CMLV, Cherry mottle leaf virus; GVA, Grapevine virus A; WVA, Watermelon virus A. Met, methyltransferase; AlKB, a DNA alkylation damage repair protein; P-pro, papain-like protease; Hel, nucleotide triphosphate-binding helicase; Pol, RNA dependent RNA polymerase; CP, coat protein; MP, movement protein; TGB, triple gene block.

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(a)

(b) Fig. 4 (a) Northern blot analysis of double-stranded RNA (dsRNA) and single-stranded RNA (ssRNA) in leaves of Nicotiana occidentalis infected with Apple stem pitting virus (ASPV IF-38) using three different RNA probes specific to different sequences within the genomic RNA. Lane 1: a probe complementary to nt positions 6207 to 6683 (ORF1 region), lane 2; nt positions 6678 to 7447 (ORF2 region), and lane 3; nt positions 8717 to 9293 (CP and the 30 non-coding regions). gRNA, genomic RNA; sgRNA, sub-genomic RNA. (b) The predicted positions of ASPV-specific dsRNAs, genomic RNA (gRNA) for Rep, sub-genomic mRNA1 (sgRNA1) for TGB, and sgRNA2 for CP.

and other two ssRNAs (2.6 kb and 1.6 kb) are thought to be sgRNAs of TGB proteins (ORF2–4) and CP (ORF5), respectively. The dsRNA species with 7.5 kbp and 6.5 kbp, the function of which are unknown, are 50 co-terminal with genomic dsRNA. Similar patterns of dsRNAs and the 30 terminal sgRNAs are reported in infections with members of the genera Capillovirus, Carlavirus, and Trichovirus.

Transmission and Host Range Most of the viruses in the family Betaflexviridae are transmitted by mechanical inoculation, although the difficulty varies depending on the species. Transmission by grafting and dispersal through propagating materials is common in all species, especially in viruses that occur in woody fruit trees. Most members in the genus Carlavirus are transmitted by aphids in the non-persistent manner. Cucumber vein-clearing virus (Carlavirus) is transmitted by whiteflies. No vectors of any species have been reported for viruses in the genera Foveavirus, Robigovirus, Capillovirus, Divavirus, Citrivivirus, Chodovirus, Tepovirus, and Prunevirus. However, some trichiviruses, i.e., Cherry mottle leaf virus and Grapevine berry inner necrosis virus are known to be transmitted by the peach bud mite Eriophyes insidiosus and the grape mite Colomerus vitis, respectively. For vitiviruses, both GVA and Grapevine virus B are transmitted in nature by several species of the pseudococcid mealybug genera Planococcus and Pseudococcus in a semi-persistent manner, whereas Heracleum latent virus is transmitted from naturally infected hogweed plants by aphids in a semi-persistent manner, which depends on a helper virus present in naturally infected plants. ASGV (Capillovirus) has been known to be transmitted through seeds to progeny seedlings of lily (1.8%) and Chenopodium quinoa (2.5%–60%). PVT (Tepovirus) is readily transmitted through true potato seed and pollen and spreads to tubers produced by infected plants. With the exception of most viruses in the genus Carlavirus, many viruses in this family use woody plants (fruit trees) as hosts, for example, rosaceae fruit trees for foveavirus, robigoviruses, capilloviruses, pruneviruses, tepovirus, and trichoviruses, grapevines for foveaviruses, trichoviruses, and vitiviruses,

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and citrus plants for capillovirus and citrivivirus. Some viruses cause diseases in fruit trees, but many viruses are infecting fruit trees without showing obvious disease.

Geographical Distribution Geographical distribution of viruses in the family Betaflexiviridae ranges from wide to restricted according the genus and the species. For example, ACLSV, ASGV, and APSV occurs in all countries where apples are grown, in contrast that GINV occurs only in Japan.

See also: Quinviruses (Betaflexiviridae). Triviruses (Betaflexiviridae)

Further Reading Adams, M.J., Antoniw, J.F., Bar-Joseph, M., et al., 2016. Ratification vote on taxonomic proposals to the International Committee on Taxonomy of Viruses. Archives of Virology 161, 2921–2949. Adams, M.J., Candresse, T., Hammond, J., et al., 2012. Family Betaflexiviridae. In: King, A.M.Q., Adams, M.J., Carstens, E.B., et al. (Eds.), Virus Taxonomy-Ninth Report on the International Committee on Taxonomy of Viruses. Cambridge, MA: Elsevier Academic Press, pp. 920–941. Marais, A., Faure, C., Mustafayev, E., et al., 2015. Characterization of new isolates of Apricot vein clearing-associated virus and of a new Prunus-infecting virus: Evidence for recombination as a driving force in Betaflexiviridae evolution. Plos One. doi:10.1371/journal.Pone.0129469. Rubino, L., Russon, M., De Stradis, A., et al., 2012. Tepovirus, a novel genus in the family Betaflexiviridae. Archives of Virology 157, 1629–1633. Svanella-Dumas, L., Tsarmpopoulos, I., Marais, A., et al., 2018. Complete genome sequence of Lettuce chordovirus 1 isolated from cultivated lettuce in France. Archives of Virology 163, 2543–2545. Villamor, D.V., Druffel, K.L., Eastwell, K.C., 2013. Complete nucleotide sequence of a virus associated with Rusty mottle disease of sweet cherry (Prunus avium). Archives of Virology 158, 1805–1810. Vives, M.C., Galipienso, L., Navarro, L., et al., 2001. The nucleotide sequence and genome organization of Citrus leaf blotch virus: Candidate type species for a new virus genus. Virology 287, 225–233. Wylie, S.J., Li, H., Dixon, K.W., et al., 2013. Exotic and indigenous viruses infect wild populations and captive collections of temperate terrestrial orchards (Diuris species) in Australia. Virus Research 171, 22–32.

Betasatellites and Deltasatelliles (Tolecusatellitidae) Muhammad S Nawaz-ul-Rehman, University of Agriculture, Faisalabad, Pakistan Nazia Nahid, GC University, Faisalabad, Pakistan and University of Agriculture, Faisalabad, Pakistan Muhammad Hassan and Muhammad Mubin, University of Agriculture, Faisalabad, Pakistan © 2021 Published by Elsevier Ltd.

Nomenclature A Adenosine CP Coat protein or capsid protein GFP Green florescent protein JA Jasmonic acid MAPK Mitogen activated protein kinase NLS Nuclear localization signal

Glossary Betasatellite ssDNA satellite molecules about 1300 nt long and coding for a pathogenicity determinant protein, which depends on a geminivirus helper for replication, movement, encapsidation and transmission. Deltasatellite Small non-coding ssDNA satellites about 700 nt long, associated with helper begomoviruses. Iterons High-affinity binding sites located upstream of the stem-loop of geminiviruses.

nt Nucleotide(s) ORF Open reading frame PASC Pairwise sequence comparison RCR Rolling-circle replication RDR6 RNA-dependent RNA polymerase 6 SCR Satellite conserved region

Pathogenicity determinant protein Any protein coded by virus or the satellite which determines the virulence of the pathogen. Satellite conserved region A highly conserved region of B80 nucleotides, which is integral part of all the betasatellites. Satellite DNA Subviral agent consisting of ssDNA that becomes packaged in protein shells made from coat protein of the helper virus and whose replication, movement and transmission is dependent on that virus.

Introduction Betasatellites are small circular sub-viral components associated with geminiviruses and are mainly confined to the Old World. The word “beta” was used to differentiate with the DNA-B component of begomoviruses and the word “satellite” was added because they are effectively satellites. Their discovery started from the identification of a satellite molecule associated with Tomato leaf curl virus in Australia (now classified under the genus Deltasatellite of the family Tolecusatellitidae). Betasatellites have very conserved sequence structure or genome organization which includes an adenosine rich (A-rich) region, a satellite conserved region (SCR) and a small open reading frame coding for a protein called bC1. Initially, they were associated with monopartite begomoviruses, but nowadays they are routinely identified in association with bipartite begomoviruses as well. So far, betasatellites have been limited to the Old World, however a recent discovery suggests that a defective form of an African-origin betasatellite (Cotton leaf curl Gezira betasatellite – lacking bC1) has been found in the United States. The DNA-A component alone of begomoviruses, in many cases, is sufficient to induce the disease symptoms. Nevertheless, for an increasing number of begomoviruses, betasatellites are crucial to induce the typical disease symptoms. For example, Cotton leaf curl Gezira virus in Africa, Cotton leaf curl Multan virus in Pakistan and Tomato yellow leaf curl China virus in China require the presence of a betasatellite for disease development. Betasatellites are trans-replicated and encapsidated by their helper virus Rep protein and coat protein (CP) respectively. In the laboratory conditions Old World betasatellites can replace the functions of a DNA-B component. However, New World begomoviruses can also maintain the betasatellites without compromising the function of the DNA-B component. The first betasatellite was characterized for the Ageratum yellow vein disease in 2000. The presence of the betasatellite increased viral symptoms in ageratum plants infected with the geminivirus. Subsequently several betasatellites were characterized from Asia and Africa. For several diseases in tomato, tobacco, cotton, chillies, and okra, the helper viruses do not require the absolute presence of a betasatellite for the disease development. To date, more than 1880 sequences of betasatellites are present in GenBank. The betasatellites not only interfere with the normal plant development, but they act as a suppressor of transcriptional and post-transcriptional gene silencing.

Classification and Nomenclature Betasatellites and deltasatellites are satellite molecules, therefore dependent on a helper virus for their replication and/or other biological functions. Their classification is based on their genomic nucleic acid sequence. Since 2016, the species demarcation threshold for betaand deltasatellites has been set to 91% nucleotide sequence identity. This species demarcation threshold will be soon revised as it seems

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Table 1 List of species of the genera Betasatellite and Deltasatellite of the family Tolecusatellitidae. A list of betasatellite unassigned species is indicated. The acronym and isolate ID of the reference virus for each species, as well as the NCBI GeneBank accession number and the length of the satellite are also provided. Type species for each genus are written in bold Genus/Species

Acronym-[isolate ID]

Accession #

Length (nt)

Betasatellite Ageratum leaf curl Buea betasatellite Ageratum leaf curl Cameroon betasatellite Ageratum yellow leaf curl betasatellite Ageratum yellow vein betasatellitea Ageratum yellow vein India betasatellite Ageratum yellow vein Sri Lanka betasatellite Alternanthera yellow vein betasatellite Andrographis yellow vein leaf curl betasatellite Bhendi yellow vein mosaic betasatellite Cardiospermum yellow leaf curl betasatellite Chili leaf curl betasatellite Chili leaf curl Jaunpur betasatellite Chili leaf curl Sri Lanka betasatellite Cotton leaf curl Gezira betasatellite Cotton leaf curl Multan betasatellite Croton yellow vein mosaic betasatellite Eupatorium yellow vein betasatellite Eupatorium yellow vein mosaic betasatellite French bean leaf curl betasatellite Hedyotis yellow mosaic betasatellite Honeysuckle yellow vein betasatellite Honeysuckle yellow vein mosaic betasatellite Malvastrum leaf curl betasatellite Malvastrum leaf curl Guangdong betasatellite Mirabilis leaf curl betasatellite Momordica yellow mosaic betasatellite Mungbean yellow mosaic betasatellite Okra leaf curl Oman betasatellite Papaya leaf curl betasatellite Papaya leaf curl China betasatellite Papaya leaf curl India betasatellite Rhynchosia yellow mosaic betasatellite Rose leaf curl betasatellite Siegesbeckia yellow vein betasatellite Tobacco curly shoot betasatellite Tobacco leaf curl betasatellite Tobacco leaf curl Japan betasatellite Tobacco leaf curl Patna betasatellite Tomato leaf curl Bangalore betasatellite Tomato leaf curl Bangladesh betasatellite Tomato leaf curl betasatellite Tomato leaf curl China betasatellite Tomato leaf curl Gandhinagar betasatellite Tomato leaf curl Java virus betasatellite Tomato leaf curl Joydebpur betasatellite Tomato leaf curl Laguna betasatellite Tomato leaf curl Laos betasatellite Tomato leaf curl Malaysia betasatellite Tomato leaf curl Nepal betasatellite Tomato leaf curl Patna betasatellite Tomato leaf curl Philippine betasatellite Tomato leaf curl Sri Lanka betasatellite Tomato leaf curl Yemen betasatellite Tomato yellow leaf curl China betasatellite Tomato yellow leaf curl Rajasthan betasatellite Tomato yellow leaf curl Shandong betasatellite Tomato yellow leaf curl Thailand betasatellite Tomato yellow leaf curl Vietnam betasatellite

ALCBueB-[CM: LIO1: SatB33:09] ALCCMB-[CM: Man:AMBF:06] AYLCB-[PK: Fai4:00] AYVB-[SG:95] AYVINB-[IN: Mad:03] AYVLKB-[LK:Ag:03] AlYVB-[VN: Hue:05] AnYVLCB-[IN: Luc:10] BYVB-[IN: Mut:00] CaYLCB-[SL:04] ChLCB-[PK: MC:97] ChLCJauB-[IN: Jau:07] ChLCLKB-[LK: Mih:09] CLCGezB-[SD: Dat:06] CLCMulB-[PK: Mul:U89:97] CroYVMB-[PK: Pun:06] EpYVB-[JR: MNS2:00] EpYVMB-[JP: Suya:03] FBLCB-[IN: Kan:11] HeYMB-[VN: BinhDinh:13] HYVB-[UK: Nor1:99] HYVMB-[JP: Hy:04] MaLCB-[CN: Gx87:04] MaLCGuB-[CN: Gu:11] MiLCB-[IN: Him:13] MamYMB-[BJ:57:14:14] MYMB-[IN: Cowpea:12] OLCOMB-[OM: Barka:12] PaLCB-[IN: ND:03] PaLCCNB-[CN: Hn:14] PaLCINB-[IN: Pan:08] RhYMB-[IN: Pha:14] RoLCB-[PK: Fai:06] SiYVB-[CN: FZ02:12] TbCSB-[CN: Yn35:01] TbLCB-[PK: Lah:04] TbLCJPB-[JP: Miy:05] TbLCPatB-[IN: Pusa:09] ToLCBanB-[IN: Ban:03] ToLCBDB-[BD: Gaz:01] ToLCB-[PK: RYK:97] ToLCCNB-[CN: Gx14:02] ToLCGanB-[IN: pToGNbH14:12] ToLCJavB-[NP: R7: Pap:10] ToLCJoyB-[BD: Gaz:05] ToLCLagB-[PH:Lag2:06] ToLCLAB-[LA: Sav:01] ToLCMYB-[MY:13] ToLCNPB-[NP: Jhapa] ToLCPatB-[IN: Pat:07] ToLCPHB-[PH:Lag1:06] ToLCLKB-[LK] ToLCYEB-[YE: tob56:89] TYLCCNB-[CN: Yn45:01] ToLCRajB-[IN: Raj:03] ToYLCShB-[CN: SDSG:14] TYLCTHB-[CN: Yn72:02] TYLCVNB-[VN: Han:05]

NC014746 NC012557 NC005046 NC003403 NC043428 NC043429 NC009562 NC023876 NC014895 NC010297 NC005048 HM007103 NC038676 NC006935 NC009535 NC008579 NC004515 NC038677 NC018091 NC023015 NC005052 NC005953 NC007711 NC023896 NC038924 KT454829 NC018869 NC038678 NC004706 NC038679 HM143906 NC038681 NC024695 NC038682 NC004546 NC038891 NC009450 NC038683 NC038684 NC014594 NC004715 NC004544 NC023038 NC005497 NC010236 NC038685 NC038925 NC038686 NC038926 NC012493 NC009570 AJ542493 NC018864 NC019532 NC038687 NC038688 NC004903 NC009560

1259 1389 1351 1347 1353 1351 1344 1379 1373 1338 1387 1362 1371 1348 1346 1346 1356 1359 1379 1348 1344 1262 1354 1344 1367 1367 1360 1351 1372 1350 1333 1359 1349 1359 1354 1358 1350 1338 1377 1371 1424 1339 1365 1360 1370 1346 1348 1342 1350 1349 1349 1371 1352 1335 1371 1334 1337 1356

nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt nt

Betasatellites and Deltasatelliles (Tolecusatellitidae)

Table 1

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Continued

Genus/Species

Acronym-[isolate ID]

Accession #

Length (nt)

Tomato yellow leaf curl Yunnan betasatellite Vernonia yellow vein betasatellite Vernonia yellow vein Fujian betasatellite

ToYLCYnB-[CN: tob:] VYVB-[IN: Mad:09] VYVFuB-[CN:09]

NC038689 NC013423 NC015928

1337 nt 1364 nt 1377 nt

Unassigned species Datura leaf curl betasatellite Emilia yellow vein betasatellite Leucas zeylanica yellow vein betasatellite Lindernia anagallis yellow vein betasatellite Okra yellow leaf curl betasatellite Pea leaf distortion betasatellite Sida leaf curl betasatellite Tobacco leaf chlorosis betasatellite Tobacco leaf curl Yunnan betasatellite Tomato leaf curl Yunnan betasatellite Tomato yellow leaf curl Kanchanaburi betasatellite Zinnia leaf curl betasatellite

DaLCB-[IN: Lak:Raj:11] EmYVB-[V: Hoa:06] LzYVB-[LK:06] LaYVB-[VN: Han:06] OYLCB-[BF: Kaya:Sida30BA:14] PeaLDB-[NP:10] SiLCB-[CN: Hai:Hn57:16] TbLChB-[IN:10] TbLCYnB-[CN: Yn:12] ToLCYnB-[CN: Yn:Yn4980:15] ToLCKanB-[ID: Kan:Egg: BL1:19] ZLCB-[TH: Pat:Tz4:03]

JQ693149 NC012666 NC013424 NC009561 MK032309 NC033618 NC007639 NC018935 NC005030 NC040544 MK936043 NC005874

1042 1337 1363 1346 1356 1347 1365 1345 1349 1361 1323 1354

Deltasatellite Croton yellow vein deltasatellite Malvastrum leaf curl deltasatellite Sida golden yellow vein deltasatellite 1 Sida golden yellow vein deltasatellite 2 Sida golden yellow vein deltasatellite 3 Sweet potato leaf curl deltasatellite 1 Sweet potato leaf curl deltasatellite 2 Sweet potato leaf curl deltasatellite 3 Tomato leaf curl deltasatellite Tomato yellow leaf distortion deltasatellite 1 Tomato yellow leaf distortion deltasatellite 2

CrYVD-[IN: Mad:04] MalLCD-[PH: Mc1:12] SiGYVD:1-[CU:H1:09] SiGYVD:2-[CU:228H1:09] SiGYVD:3-[CU:412N1:10] SPLCD:1-[ES: SBG51:09] SPLCD:2-[VN: Mer:1764E13:09] SPLCD:3-[PR: PR3:2:09] ToLCD-[AU:96] ToYLDD:1-[CU: Sida:404N1:10] ToYLDD:2-[CU:603N1:11]

AJ968684 NC021929 NC043200 NC043201 NC043202 NC016536 NC025220 NC043203 NC002743 NC043204 NC043205

739 673 687 677 686 662 733 731 682 685 694

nt nt nt nt nt nt nt nt nt nt nt nt

nt nt nt nt nt nt nt nt nt nt nt

a

Unassigned betasatellite species.

that the distribution of pairwise comparison sequence (PASC) values indicate a clear difference between isolates, species and genera, pointing to a cut-off value closer to 73% for species demarcation. Currently the ICTV recognizes 61 species of betasatellites and 11 species of deltasatellites. In addition, 12 tentative species of betasatellites are considered for official recognition (Table 1, Fig. 1). Recently the family Tolecusatellitidae has been created to host betasatellites and deltasatellites. The word Tolecusatellitidae is coined from the first ever identified satellite in Australia (Tomato leaf curl virus-satellite). The nomenclature of betasatellites and deltasatellites species names is so far based on the name of the first helper geminivirus with which the satellite was identified, complemented, if necessary, with the geographical location of the first sample and a number if several species are identified on the same host with the same helper geminivirus. The family encompasses two genera, Betasatellite and Deltasatellite, and Ageratum yellow vein betasatellite and Tomato leaf curl deltasatellite are the type species for each genus respectively. Betasatellites and deltasatellites do not share any sequence similarity to each other nor with their helper viruses, except the nonanucleotide origin of replication (TAATATTAC).

Genome Organization All the known betasatellites have conserved sequence features with an A-rich region (B160–280 nt), a satellite conserved region (SCR, B80 nt) and a single open reading frame (ORF) coding for a protein called bC1 (B13.6 kDa) (Fig. 2). While deltasatellites lack the bC1 gene and are roughly half the size (B700 nt) of betasatellites (Fig. 2). The A-rich region is involved in the maintenance of betasatellite integrity, as its deletion can cause milder symptoms in the host plant. SCR is involved in interaction with the Rep protein of the helper virus, while bC1 is a multifunctional pathogenicity determinant protein, which interacts with several host factors to counter the host defense system by suppressing both transcriptional and post-transcriptional gene silencing. Deltasatellite do not code for any functional protein and are fully dependent on their helper virus.

Genetic Diversity and Center of Origin for Betasatellites Betasatellites are widely distributed in the Old World mainly with monopartite begomoviruses. Due to the lack of sequence similarity with their helper viruses or other related viruses, the origin of betasatellites is unknown. Betasatellites are present in more than

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Fig. 1 Phylogenetic trees for betasatellites and deltasatellites in the family Tolecusatellitidae. The acronyms are listed in Table 1. Betasatellites have B61 recognized species and 11 putative unassigned species (noted with *), while deltasatellites consist of 11 different recognized species. The reference sequences of each species were downloaded and aligned in MEGA software. The alignment was made through Clustal-W. Phylogenetic trees were constructed by neighbor joining method with 1000 bootstrap replications, and only scores above 50% were noted.

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Fig. 2 Genome organization (left to right) of betasatellites and deltasatellites. Betasatellites are of B1350 nt, with an A-rich region, a satellite conserved region and a small gene bC1. While, deltasatellites are of B680 nt and do not encode any gene. The genome organizations of Tomato leaf curl deltasatellites (ToLCD) from Australia, Sweet potato leaf curl deltasatellite-1 (SPLCD-1) and Tomato yellow leaf curl distortion deltasatellite (ToYLCDD-1) from Cuba are presented. The nucleotides region mentioned with magenta colors represent the satellite-conserved region; brown color represents the A-rich region, while green parts represent the secondary stem loop in the sequence.

Fig. 3 Worldwide distribution for 61 species and 11 putative species of betasatellites and 11 species of deltasatellites. Most of the betasatellites species are present in the Indian sub-continent, while a second center of diversity appears in China and Japan. Betasatellites are limited to the Old World, while deltasatellites are present in both the Old World and the New World. (mentioned with *).

20 countries of Asia, Africa, and Europe and none have been identified in the New World (Fig. 3). Potentially, there are B73 betasatellite species and they mostly belong to South, East, South East, and West Asia (67/73). Among these betasatellites, 35 species circulate in India, Pakistan, Bangladesh, Sri Lanka, and Nepal. While 19 betasatellite species are present in China and Japan. Another 11 distinct betasatellite species exist in Vietnam, Laos, Philippines, Singapore, Indonesia, and Malaysia. In addition, two different species infect okra and tomatoes in Oman and Yemen respectively. The whole African continent only hosts 5 different species of betasatellites that are phylogenetically close to the Oman and Yemen isolates (Fig. 1). While from Europe (United Kingdom) a single betasatellite species is known which was most probably imported from Japan (Fig. 1). A phylogenetic study carried with representatives of these 63 species reveals a very strong geographical distribution with 11 clusters almost 100% homogeneous for the regions mentioned above. Five clusters belong to the Indian subcontinent, 3 to the China-Japan region, 2 from south east Asia and 1 from Africa. Only 11 species are “misplaced” for having a perfect match. Therefore, considering the genetic diversity here described it can be concluded that the Indian sub-continent is possibly the center of origin of betasatellites (Figs. 1 and 3).

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Fig. 4 Impact of betasatellite on symptom severity in Nicotiana benthamiana plants. The Nicotiana benthamiana plants inoculated with Cotton leaf curl Multan virus DNA-A alone show very mild symptoms (center), while the addition of the Cotton leaf curl Multan betasatellite induces very severe symptoms (right). Non-inoculated control plants are on the left.

Unlike betasatellites, deltasatellites have been also identified in the New World (Fig. 3), however it is believed that they were introduced through infected root crops such as sweet potato. A contrario to betasatellites, It is unclear what is the competitive advantage for the helper geminivirus to carry a deltasatellite along. Recent studies suggest that deltasatellites may reduce the geminivirus impact and thereby extend the life of the infected plant, thus allowing a greater period for the geminivirus transmission and survival of the host.

Life Cycle, Epidemiology As mentioned earlier, the betasatellites are often found with monopartite begomoviruses. However, they are also found in combination with bipartite viruses such as Tomato leaf curl New Delhi virus, Tomato leaf curl Gujarat virus, and Sri Lankan cassava mosaic virus. In the laboratory conditions, betasatellites can also interact with the New World bipartite begomoviruses. Betasatellites are also known to interact with mastreviruses in wheat plants. These observations suggest that trans-replication of betasatellites is not very stringent in respect to the helper geminivirus. The experiments conducted on many economically important begomoviruses suggested that DNA-A alone could effectively produce the disease symptoms in Nicotiana benthamiana plants. However, in many other cases, like cotton leaf curl disease, Ageratum yellow vein virus and Eupatorium yellow vein virus, the betasatellites were necessary to induce the disease symptoms (Fig. 4). Further experiments suggested that the role of bC1 was important for disease induction. A mutated bC1 could not induce the proper disease symptoms in the inoculated plants, despite of its successful replication in the systemic leaves. The co-inoculation of betasatellite with bipartite begomoviruses like Tomato leaf curl New Delhi virus and Tomato leaf curl Gujarat virus resulted in increased viral titer and exacerbated symptoms. The presence of betasatellites in non-cultivated cotton species did not result in increased symptoms. Therefore it can be concluded that betasatellites result in severe disease symptoms in cultivated crops. Normally bipartite begomoviruses are accompanied by a DNA-B component, which provides the cell-to-cell movement functions for the virus. The experiments conducted on Sri Lankan cassava mosaic virus and Tomato leaf curl New Delhi virus suggested that betasatellites could make DNA-B dispensable to the helper virus. The exact mechanism of virus movement in the absence of DNA-B is unknown. There are however few indirect evidences, which suggest the possible role of bC1 encoded by betasatellite in viral movement. The green florescent protein (GFP) tagged to bC1 of TYLCCNB and CLCMulB were transiently expressed in tobacco cells. The bC1 accumulated inside the nucleus and the cell periphery, suggesting its possible role in transport of DNA from nucleus to plasmodesmata. However so far, all the experiments for DNA-B replacement were conducted in laboratory conditions, and there is no such example in nature, where a bipartite virus was converted into monopartite virus by capturing a betasatellite, while loosing its DNA-B component. However, there are recent examples, where a cognate betasatellite was replaced by a non-cognate one. For examples, Tomato leaf curl Karnataka virus in India adopted Cotton leaf curl Multan betasatellite replacing Tomato leaf curl Karnataka betasatellite. TYLCV-IR is a monopartite begomovirus in the Middle East, but in Oman, it captured Tomato leaf curl betasatellite. A similar example of re-assortment also exists in Mali, where Cotton leaf curl Gezira betasatellite was found in combination with the monopartite Tomato yellow leaf curl Mali virus.

Replication of Betasatellites The viruses belonging to family Geminiviridae multiply through rolling circle replication (RCR) mechanism. RCR starts by the binding of the Replication associated protein (Rep) at iteron sequences present at the upstream of the nonanucleotide origin of replication. Despite using the same Rep the trans-replication of betasatellites is very relaxed as compared to DNA-B component. A single betasatellite can be trans-replicated by several helper viruses. For example, Cotton leaf curl Multan betasatellite (CLCMulB) can be

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maintained in Nicotiana benthamiana plants by Ageratum yellow vein virus and Eupatorium yellow vein virus, but Cotton leaf curl Multan virus shows specificity to its cognate betasatellite. The SCR of betasatellites present in the upstream of origin of replication appears to be the analogous of the common region of DNA-A components. As a matter of fact all the defective betasatellites or deltasatellites contain intact SCR indicating its vital role in their replication. The experiments conducted on Tomato leaf curl virus associated betasatellites suggested that the stem loop structure and the satellite conserved region are very crucial sequences for their replication. The betasatellites contain a primary stem loop structure (common to all geminiviruses) and a secondary stem loop structure B380 nt upstream of origin of replication. The Rep binding affinity experiments suggested that the secondary stem loop structure is very crucial for betasatellites replication as its deletion can cause impaired replication. Despite of their relaxed transreplication betasatellites have certain unknown specificities with their helper begomoviruses. The Rep binding affinity experiments suggested that helper components have better adaptation with their cognate betasatellites compared to non-cognate ones.

Functional Role of bC1 in Disease Development The betasatellites strongly influence the pathogenicity of the helper begomovirus. The presence of betasatellites are resulting in very severe disease symptoms. The bC1 gene present in the complementary strand of betasatellite genomes encodes an important pathogenicity determinant protein for geminiviruses of the Old World. Most of the available information regarding bC1 is obtained from Tomato yellow leaf curl china betasatellite (TYLCCNB). The bC1 of TYLCCNB localizes in the nucleus and forms a multimeric complex due to two alpha helices present at the C-terminus of the protein. The self-interaction of bC1 is important to induce the disease symptoms. Indeed, a mutated version of bC1, that cannot make a multimeric complex, is unable to induce the typical disease symptoms. The nuclear localizing signal (NLS) of bC1 is also important in the disease induction. The deletion mutants for NLS in TYLCCNB resulted in its inability to induce typical disease symptoms. The protein bC1 is a multitasking protein, which suppresses the transcriptional and post-transcriptional gene silencing, the whole hormone based defense system, and the ubiquitin proteome degradation system. It also inhibits the activity of kinases like mitogen activated protein kinases (MAPK). The pleotropic interaction of bC1 with multiple pathways results in enhanced virulence and infection. The transgenic plants expressing the protein bC1 show severe disease-like symptoms. In the infected cells, bC1 interacts with the Asymmetric Leaves gene 1 (AS1), which can ultimately result in suppression of jasmonic acid (JA) production and its associated genes. The suppression of JA results in severe disease symptoms. The bC1 encoded by Radish leaf curl betasatellite (RaLCB) localizes in chloroplast and results in vein clearing symptoms. The symptoms were very severe in the presence of Tomato leaf curl New Delhi virus. The detailed biochemical analysis revealed that bC1 altered the starch accumulation, decreased the thylakoid contents and change the granum structure. bC1counters the gene silencing initiated by plants as a defense against betasatellite and geminiviruses. The interaction of bC1 with Calmodulin-like (CaM-endogenous suppressors of gene silencing) proteins results in higher accumulation of the helper virus by compromising the RDR6 based gene silencing. bC1 is also a target of the ubiquitination based protein degradation pathway. It has evolved a system to hijack the ubiquitination machinery. The interaction of bC1 with host proteins resulting in reduction of polyubiquitinated proteins. Autophagy is another evolutionary conserved pathway, which is targeted by bC1. Autophagy is actively involved in controlling several viruses. Studies have shown that, autophagy pathways targeted and degraded the bC1. The silencing of autophagy related genes; ATG5 and ATG7 resulted in increased susceptibility to begomoviruses and betasatellites.

Resistance Against Betasatellites So far, there is no example where geminiviruses/betasatellites have been controlled successfully at commercial level. However, limited success has been reported at the experimental levels. During the natural infection tobacco ring finger proteins (RFP- a functional E3 ubiquitin ligase) ubiquitinates and degrades the bC1 proteins. The impact of bC1 ubiquitination can be observed by attenuated symptoms. In the future, it is believed that exploiting such pathways to increase the plants immunity level against geminiviruses/ betasatellites complexes, is possible. Recent studies conducted on Tomato yellow leaf curl virus demonstrated that Ty-1 gene confers natural resistance against the virus. However, with the addition of a betasatellite the Ty-1 based resistance was compromised and viral titer was increased. Due to the global spread of betasatellites the coordinated efforts between different countries are required.

Deltasatellites Introduction Deltasatellites were the first satellites identified with an example from Australia in association with Tomato leaf curl virus (ToLCV). Initially it was named ToLCV-Sat, but now it has been renamed as Tomato leaf curl deltasatellite (ToLCD). The ToLCD (682 nt long) did not share any significant sequence identity with the helper virus except for the origin of replication. The sequence variability of ToLCD is unknown due to the lack of sequences available in the GenBank. The ToLCD can be maintained by very distinct geminiviruses including Tomato yellow leaf curl Sardinia virus, African cassava mosaic virus and Beet curly top virus. From the New World, deltasatellites were first reported in 2012 from the Caribbean islands. To date, deltasatellites are known from India, Vietnam, Philippines, Spain, Australia, Cuba, and Puerto Rico. There are 11 distinct species of deltasatellites, which are phylogenetically not linked to each other (Table 1, Fig. 1).

246

Betasatellites and Deltasatelliles (Tolecusatellitidae)

Genome Organization of Deltasatellites Deltasatellites do not encode any gene in their genomes. However, they share some common features. Most of the known deltasatellites contain a variable A-rich region, a nonanucleotide origin of replication (TAATATTAC) and a secondary stem loop structure. The deltasatellites from Australia and Spain, share common features of A-rich region and a satellite conserved region. However, the deltasatellites from the New World have very short satellite conserved region (Fig. 2).

Trans-Replication of Deltasatellites Deltasatellites are not self-replicating molecules therefore they require helper viruses for their replication and systemic movement. Although, ToLCD can be maintained by several heterologous Old World geminiviruses, but it does not contribute to any symptom severity or increase of the helper virus titer. The experiments conducted on New World deltasatellites suggested that they cannot be trans-replicated by Old World begomoviruses or curtoviruses, however they can be efficiently maintained by New World bipartite and monopartite begomoviruses. Deltasatellites significantly reduce their helper component’s accumulation. Deltasatellites do not encode any gene in their genome, therefore the biological significance and mode of action of deltasatellites is clearly not understood.

See also: Cotton Leaf Curl Disease (Geminiviridae). Emerging Geminiviruses (Geminiviridae). Geminiviruses (Geminiviridae). Nanoviruses (Nanoviridae)

Further Reading Amin, I., Hussain, K., Akbergenov, R., et al., 2011. Suppressors of RNA silencing encoded by the components of the cotton leaf curl begomovirus-betasatellite complex. Molecular Plant-Microbe Interactions 24, 973–983. Briddon, R.W., Brown, J.K., Moriones, E., et al., 2008. Recommendations for the classification and nomenclature of the DNA-beta satellites of begomoviruses. Archives of Virology 153, 763–781. Eini, O., 2017. A betasatellite-encoded protein regulates key components of gene silencing system in plants. Molecular Biology 51, 656–663. Fiallo‐Olivé, E., Tovar, R., Navas‐Castillo, J., 2016. Deciphering the biology of deltasatellites from the New World: Maintenance by New World begomoviruses and whitefly transmission. New Phytologist 212, 680–692. doi:10.1111/nph.14071. Gnanasekaran, P., Chakraborty, S., 2018. Biology of viral satellites and their role in pathogenesis. Current Opinion in Virology 33, 96–105. Gnanasekaran, P., Kishorekumar, R., Bhattacharyya, R.D., Kumar, V., Chakraborty, S., 2019. Multifaceted role of geminivirus associated betasatellite in pathogenesis. Molecular Plant Pathology 20 (7), 1019–1033. doi:10.1111/mpp.12800. Hassan, I., Orilio, A.F., Fiallo-Olive, E., Briddon, R.W., Navas-Castillo, J., 2016. Infectivity, effects on helper viruses and whitefly transmission of the deltasatellites associated with sweepoviruses (genus Begomovirus, family Geminiviridae). Scientific Reports 6, 30204. doi:10.1038/srep30204. Hohn, T., Vazquez, F., 2011. RNA silencing pathways of plants: Silencing and its suppression by plant DNA viruses. Biochimica et Biophysica Acta 1809, 588–600. Khan, A.J., Mansoor, S., Briddon, R.W., 2014. Oman: A case for a sink of begomoviruses of geographically diverse origins. Trends in Plant Science 19, 67–70. Kumar, J., Kumar, J., Singh, S.P., Tuli, R., 2014. Association of satellites with a mastrevirus in natural infection: Complexity of Wheat dwarf India virus disease. Journal of Virology 88, 7093–7104. Kumar, R.V., Singh, A.K., Singh, A.K., et al., 2015. Complexity of begomovirus and betasatellite populations associated with chilli leaf curl disease in India. Journal of General Virology 96, 3143–3158. Li, Y., Qin, L., Zhao, J., et al., 2017b. SlMAPK3 enhances tolerance to Tomato yellow leaf curl virus (TYLCV) by regulating salicylic acid and jasmonic acid signaling in tomato (Solanum lycopersicum). PLoS One 12, e0172466. Li, F., Zhao, N., Li, Z., et al., 2017a. A calmodulin-like protein suppresses RNA silencing and promotes geminivirus infection by degrading SGS3 via the autophagy path- way in Nicotiana benthamiana. PLoS Pathogens 13, e1006213. Lozano, G., Trenado, H.P., Fiallo-Olive,́ E., et al., 2016. Characterization of non-coding DNA satellites associated with Sweepoviruses (Begomovirus, Geminiviridae) – Definition of a distinct class of Begomovirus-associated satellites. Frontiers in Microbiology 7, 162. Mubin, M., Ijaz, S., Nahid, N., et al., 2020. Journey of begomovirus betasatellite molecules: From satellites to indispensable partners. Virus Genes 56, 16. Nawaz-ul-Rehman, M.S., Fauquet, C.M., 2009. Evolution of geminiruses and their satellites. FEBS Letters 583, 1825–1832. Nawaz-Ul-Rehman, M.S., Nahid, N., Mansoor, S., Briddon, R.W., Fauquet, C.M., 2010. Post-transcriptional gene silencing suppressor activity of two non-pathogenic alphasatellites associated with a begomovivirus. Virology 405, 300–308. Pumplin, N., Voinnet, O., 2013. RNA silencing suppression by plant pathogens: Defence, counter-defence and counter-counter-defence. Nature Reviews Microbiology 11, 745–760. Singh, A.K., Kushwaha, N., Chakraborty, S., 2016. Synergistic interaction among begomoviruses leads to the suppression of host defense-related gene expression and breakdown of resistance in chilli. Applied Microbiology and Biotechnology 100, 4035–4049. Verchot, J., 2016. Plant virus infection and the ubiquitin proteasome machinery: Arms race along the endoplasmic reticulum. Viruses 8, E314. Vinoth Kumar, R., Singh, D., Singh, A.K., Chakraborty, S., 2017. Molecular diversity, recombination and population structure of alphasatellites associated with begomovirus disease complexes. Infection, Genetics and Evolution 49, 39–47. Yang, X., Wang, Y., Guo, W., et al., 2011a. Characterization of small interfering RNAs derived from the geminivirus/betasatellite complex using deep sequencing. PLoS One 6, e16928. Yang, X., Xie, Y., Raja, P., et al., 2011b. Suppression of methylation-mediated transcriptional gene silencing by betaC1-SAHH protein interaction during geminivirus-betasatellite infection. PLoS Pathogens 7, e1002329. Zhang, T., Xu, X., Huang, C., et al., 2015. A novel DNA motif contributes to selective replication of a geminivirus-associated betasatellite by a helper virus-encoded replicationrelated protein. Journal of Virology 90, 2077–2089. Zhou, X., 2013. Advances in understanding begomovirus satellites. Annual Review of Phytopathology 51, 357–381. Zhou, X., Xie, Y., Tao, X., et al., 2003. Characterization of DNA beta associated with begomoviruses in China and evidence for co-evolution with their cognate viral DNA-A. Journal of General Virology 84, 237–247. Zubair, M., Zaidi, S.S., Shakir, S., Amin, I., Mansoor, S., 2017. An insight into Cotton leaf curl Multan betasatellite, the most important component of cotton leaf curl disease complex. Viruses 9, E280. doi:10.3390/v9100280.

Bluner-, Cile-, and Higreviruses (Kitaviridae) Diego F Quito-Avila, Department of Life Sciences, ESPOL Polytechnic University, Guayaquil, Ecuador Juliana Freitas-Astúa, Brazilian Agricultural Research Corporation (Embrapa) Cassava and Fruits, Cruz das Almas, Brazil Michael J Melzer, Department of Plant and Environmental Protection Sciences, University of Hawaii, Honolulu, HI, United States r 2021 Elsevier Ltd. All rights reserved.

Glossary Cytopathology

Symptomatology

The symptom complex of a disease.

The study of disease at the cellular level.

Introduction Kitaviridae is a family of positive-sense RNA viruses with linear, segmented genomes and bacilliform to spherical virions. All current members of the family infect plants, often causing chlorotic and/or necrotic lesions in their natural hosts. These infections are typically non-systemic, making them highly unusual among plant viruses. Despite being non-systemic, diseases caused by these viruses can be highly destructive and of tremendous economic importance, with citrus leprosis being the most prominent. Citrus leprosis virus C, a causal agent of citrus leprosis, represents the most well-studied member and functional model for the family. The name Kitaviridae is derived from the renowned Brazilian virologist and electron microscopist Dr. Elliot W. Kitajima, in recognition of his extensive contributions to our knowledge on the fundamental aspects of kitavirus structure, cytopathology, and biology. The family was established by the International Committee on Taxonomy of Viruses (ICTV) in 2018.

Taxonomy, Phylogeny, and Evolution The family Kitaviridae is currently composed of three genera, established by the ICTV, Cilevirus, Higrevirus, and Blunervirus. The genus Cilevirus derives its name from the type species Citrus leprosis virus C, the prevalent causal agent of citrus leprosis, one of the most aggressive and important citrus diseases in South and Central America. The genus Higrevirus is named after the type species Hibiscus green spot virus 2 (HGSV 2), a virus associated with leprosis-like lesions in some citrus species and chlorotic lesions on the leaves of hibiscus species. To date, the virus and its associated diseases have only been reported in the Hawaiian Islands. The genus Blunervirus was named after its type species Blueberry necrotic ring blotch virus, which was discovered in 2010 from symptomatic highbush blueberry (Vaccinum corymbosum) plants in Georgia, United States. At the species level, demarcation criteria have only been established for the genus Cilevirus and include the extent of serological relationship; o85% amino acid identity for the proteome, natural and experimental host range reactions, and vector species and transmission. Recently, a threshold of 75% amino acid identity for the polyprotein encoded by RNA1 has been proposed for species demarcation in the genus Blunervirus. Phylogenetic reconstructions using RNA-dependent RNA polymerase (RdRp) sequences, the standard for inferring relationships between RNA viruses, illustrate the family Kitaviridae as a monophyletic clade and sister taxon to the plant virus family Virgaviridae. The phylogenetic relationships that kitaviruses share with other viral taxa, however, are not this straightforward. For example, blueberry necrotic ring blotch virus (BNRBV) encodes two helicase (HEL) domains. Phylogenetic reconstructions using these two domains indicate that one (HEL-1) is most closely related to those of virgaviruses, and the other (HEL-2) most closely related to members of the plant virus family Bromoviridae. Furthermore, the methyltransferase (MTR) domain of BNRBV contains hallmark motifs that are found in bromoviruses. The predicted movement proteins (MP) of kitaviruses are also of different phylogenetic lineages, with blunerviruses and cileviruses possessing 3A MPs of the 30K superfamily, and the lone higrevirus possessing ‘triple gene block’ MPs. Taken together, the evolutionary history of kitaviruses is complex and highly indicative of recombination events between distinct viral taxa. High-throughput metagenomic sequencing of arthropod and other invertebrate specimens has provided evidence of several unclassified viruses that appear to be closely related (and even interspersed) within the kitavirus clade when using the RdRp as the phylogenetic signal. Furthermore, a predicted structural protein designated p22 (Blunervirus), p23 (Higrevirus) or p24 (Cilevirus) is present in all kitaviruses (but no other plant virus taxa), and also in some of these unclassified invertebrate viruses (Fig. 1). This, combined with the inability of recognized kitaviruses to infect their natural plant host systemically, has led to speculation that the evolutionary progenitor(s) of the kitavirus clade might be an arthropod virus.

Members of the Family The family Kitaviridae is sparsely populated, with only four recognized species among the three genera. Some tentative members have been partially described but are not currently recognized by the ICTV (Table 1).

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Fig. 1 Phylogenetic relationships across kitaviruses (BNRBV, HGSV-2 and CiLV-C) and unclassified invertebrate viruses, based on hypothetical protein p22–24. A Bayesian tree is shown using LG þ G þ I protein evolution model. NCBI (National Center for Biotechnology Information) accession numbers are provided on the right. Taxa where p22–24 have been annotated as host proteins are indicated by the asterisk (*).

Table 1

Species and tentative species in the family Kitaviridae

Virus Genus Blunervirus Definitive species Blueberry necrotic ring blotch virusa Tentative species Tea plant necrotic ring blotch virus Genus Cilevirus Definitive species Citrus leprosis virus Ca Citrus leprosis virus C2 Tentative species Ligustrum ringspot virus Passion fruit green spot virus Genus Higrevirus Definitive species Hibiscus green spot virus 2a

Genomic segments

Genome length (kb)

Natural host

4

14.15

Blueberry

4

15.08

Tea

2 2

13.73 13.71

Citrus, tropical spiderwort (Commelina benghalensis) Citrus, hibiscus

N/a N/a

N/a N/a

Privet Passion fruit

3

14.70

Citrus, hibiscus

a

Type species.

Virion Structure Virion morphology for members of the Cilevirus and Higrevirus genera has been determined by transmission electron microscopy (TEM). Infected cells contain congregations of bacilliform virions, which are typically detected in cytoplasmic vesicles associated with the endoplasmic reticulum. Virions range between 30 and 50 nm in width and 50–120 nm in length depending on the species (Fig. 2). Electron microscopy studies have not been done to investigate virion morphology of BNRBV (genus Blunervirus). However, TEM examination of ultrathin sections of symptomatic tissue infected with tea plant necrotic ring blotch virus, a proposed member of the genus, revealed cytoplasmic aggregates of spherical virus-like particles approximately 85 nm in diameter.

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Fig. 2 Transmission electron micrographs of ultrathin sections of soybean (an experimental host) leaves infected with Citrus leprosis virus C (CiLV-C). Asterisk (*) indicates electron-dense viroplasm, and arrows show host-derived vesicle structures containing virus-like particles. Photo courtesy of Dr. E.W. Kitajima.

Genome Organization and Gene Product Function Kitavirus genomes are multipartite and composed of linear, positive-sense RNA molecules (Fig. 3). The genomes of current cileviruses are bipartite and comprise a total length of 13.1 kb. RNA1 contains a large 50 -terminal open reading frame that encodes a replication-associated polyprotein containing MTR, HEL, and RdRp domains followed by an ORF encoding a 29 kDa protein predicted to be the capsid protein (CP). Homologs of this putative CP have not been found in other kitavirus genomes. RNA2 contains four ORFs, the first of which encodes p15, which appears to have a role in cellular remodeling for vesicle formation. The second ORF encodes p61, a predicted glycoprotein that also appears to remodel the host endoplasmic reticulum (ER). The third ORF encodes a 3A type MP, and the 30 -terminal ORF encodes p24, a predicted virion membrane protein. p24 is one of the few proteins encoded by all kitaviruses and is also present in some unclassified invertebrate viruses. The 50 -termini of the RNAs are capped and the 30 -termini are polyadenylated. The genome of the current higrevirus species is tripartite and comprises a total length of 14.7 kb. RNA1 is similar to that of cileviruses but lacks the putative 29 kDa CP. RNA2 possesses a 50 kDa protein of unknown function followed by a triple gene block-like suite of proteins of 39, 9, and 6 kDa which putatively serves as the MPs of the virus. However, recent findings have suggested that proteins p39 and p9, acting in a ‘binary movement block’, are sufficient for virus intercellular movement. RNA3 possesses three ORFs of 33, 29, and 23 kDa. The first two are of unknown function, and p23 is a homolog of the cilevirus p24 and blunervirus p22. Higrevirus RNAs are polyadenylated on their 30 -termini. Current blunervirus genomes are quadripartite and comprise a total length of 14.2 kb. RNA1 contains a single open reading frame (ORF), whose predicted protein contains MTR and HEL-1 domains. The single ORF predicted in RNA2 encodes the putative RdRp, with the HEL-2 domain at the N terminus. RNA3 comprises five short ORFs (ORF3a, 3b, 3c, 3d and 3e) putatively encoding proteins of unknown functions. One of these unknown proteins, p22, encoded by ORF3c, has homology to proteins of similar size in cileviruses (p24), higreviruses (p23) and unclassified invertebrate viruses. RNA4 has a single ORF coding for the putative MP, which possesses motifs conserved with the 3A family of the 30K superfamily of viral MPs (30Ksf). This protein is homologous to its counterpart in cileviruses. Non-coding regions (NCR) at 50 -ends vary in size, with an unusually long (4350 nt) NCR in RNA4. NCRs at 30 -ends of BNRBV contain conserved nucleotide stretches. Although the presence of poly (A) tails were not documented for the original BNRBV genome, adenosine stretches of varying size were detected on 30 -termini of BNRBV-RL (a red-lesion strain discovered in Florida) and the putative new blunervirus Tea plant necrotic ring blotch virus (TPNRBV); suggesting that polyadenylation is a common feature in kitaviruses.

Replication and Propagation The Cilevirus citrus leprosis virus C (CiLV-C) represents the most well-studied kitavirus and the main virus for which replication and propagation studies have been conducted. Some information on that regard has been obtained for Citrus leprosis virus C2 as well (CiLV-C2). CiLV-C RNA replication-associated proteins (MTR, HEL, and RdRp) are expressed from genomic RNA1 as a polyprotein. Other proteins involved with replication and propagation are expressed via 30 -terminal sub-genomic RNAs. Replication appears to occur in cytoplasmic vesicles formed through the modification of cortical endoplasmic reticulum (ER) by

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Bluner-, Cile-, and Higreviruses (Kitaviridae)

Fig. 3 Genomic organization of the type species for each genus in the family Kitaviridae. Boxes represent open reading frames (ORFs). ORFs are labeled with either the predicted molecular weight (kDa) of the protein product, or its function: MTR, methyltransferase; HEL, helicase; RdRp, RNA-dependent RNA polymerase; and MP, movement protein. Asterisk (*) indicates that the polyadenylated 30 -terminus has been reported for one isolate of the blueberry necrotic ring blotch virus, but not another. The scale at top represents approximate RNA length in kilobases (kb).

multiple virus-encoded proteins. Transfer of virus replication complexes and/or the viral ribonucleoproteins from the cytoplasmic vesicles to plasmodesmata may be facilitated by actin or ER trafficking. Viral MP then facilitate cell-to-cell movement through the plasmodesmata. Only non-systemic movement has been reported for any member of the family in plant hosts, although systemic movement has been shown for TPNRBV.

Transmission and Host Range Cileviruses are vectored by polyphagous mites in the genus Brevipalpus in a persistent manner. There is molecular evidence for replication in these mites, but this has not been confirmed by electron microscopy, despite extensive efforts. Preliminary transmission studies indicate higreviruses are also transmitted by brevipalpus mites. Cileviruses and higreviruses have been found to naturally infect Citrus and Hibiscus spp., and experimental host range experiments using viruliferous mites have demonstrated that CiLV-C is able to infect dozens of plant species, including several of economic importance such as common bean and soybean. Preliminary transmission experiments, both under field and greenhouse conditions, have demonstrated that the blunervirus BNRBV is transmitted by eriophyoid mites in the genus Calacarus. The specific mechanism by which the vector transmits the virus is not known. BNRBV has not been reported in any host other than highbush blueberry. TPNRBV, a putative blunervirus, was found to infect tea plants (Camellia sinensis), although no additional transmission or host range data is currently available.

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Diseases, Epidemiology, and Control Citrus leprosis disease has multiple causal agents, including CiLV-C and CiLV-C2, which are both members of the genus Cilevirus. Symptoms of infection include localized chlorotic and necrotic lesions on both leaves and fruit, which can lead to premature abscission. Citrus leprosis is extant only in South and Central American countries. Although historical reports of the disease in Florida, USA exist, it appears the pathogen at this location was an unrelated virus which causes nearly identical symptoms in citrus. In Brazil, citrus leprosis caused primarily by CiLV-C is often regarded as the most important virus disease of citrus. Management efforts to control the disease, which are primarily the application of acaricides to control vector populations, often reach $40–80 M USD per year. In Hawaii, the higrevirus Hibiscus green spot virus 2 (HGSV-2) is responsible for an isolated disease on some citrus varieties that resembles citrus leprosis. Unlike citrus leprosis, foliar lesions on citrus caused by HGSV-2 rarely become necrotic, and lesions on fruit are only visible on green, unripe fruit. HGSV-2 also causes lesions on hibiscus leaves which are most prominent during senescence. Severe infection may cause premature abscission of leaves which may be of concern to threatened and endangered hibiscus species native to Hawaii. The blunervirus BNRBV was discovered as the causal agent of an unusual disease observed in southern highbush blueberry (Vaccinium corymbosum interspecific hybrids) plantings in Georgia, USA. Infected plants display irregularly shaped rings on the adaxial and abaxial leaf surfaces, which coalesce to form brown-to-black necrotic blotches, followed by premature defoliation. Since its first report in 2006, the disorder has become commonplace in other southern states including Florida, Mississippi, North Carolina and South Carolina. In Florida, a variant of the virus was characterized and found to induce red concentric rings, which develop across leaves without turning into necrotic spots or falling out prematurely. This strain is referred to as BNRBV-red lesion (RL) and shares an overall 90% nucleotide identity with BNRBV. The impact of the disease caused by BNRBV has not been reported. None of the current kitaviruses can move systemically in plant hosts and are therefore not graft-transmissible. However, stem lesions or viruliferous mites present on propagative materials can serve as a reservoir for long-distant movement by humans. Unlike citrus and hibiscus, blueberry defoliates in winter, indicating a direct role of overwintering vectors and/or alternate (virus reservoir) hosts in the spread of BNRBV during subsequent seasons. This aspect of the virus life cycle has a direct implication in terms of disease management by controlling the vector.

Diagnosis Determining the localization of virions and viroplasms by TEM was initially used to distinguish the cytoplasmic citrus leprosis viruses (CiLV-C and CiLV-C2) from the unrelated nuclear virus which caused similar symptoms. Today, PCR-based molecular approaches are used for kitavirus detection. Serological detection approaches are also available for CiLV-C and CiLV-C2, which include enzyme-linked immunosorbent assays (ELISA) and cellular analysis and notification of antigen risks and yields (CANARY) platforms.

Concluding Remarks The kitaviruses are a small, but important group of emerging viruses that warrant study not only for control of the diseases they cause, but also to better understand the fundamentals of plant virology. The non-systemic nature of their infection opens intriguing opportunities for experimentation on virus movement and arthropod vector relationships. The putatively close evolutionary relationship with a growing number of unclassified invertebrate viruses suggests that in the future this taxon may not be restricted to plant viruses.

Further Reading Bastianel, M., Novelli, V.M., Kitajima, E.W., et al., 2010. Citrus leprosis: Centennial of an unusual mite-virus pathosystem. Plant Disease 94, 284–292. Cantu-Iris, M., Harmon, P.F., Londono, A., Polston, J.E., 2013. A variant of Blueberry necrotic ring blotch virus associated with red lesions in blueberry. Archives of Virology 158, 2197–2200. Hao, X., Zhang, W., Zhao, F., et al., 2018. Discovery of plant viruses from tea plant (Camellia sinensis (L.) O. Kuntze) by metagenomic sequencing. Frontiers in Microbiology 9, 2175. Lazareva, E.A., Lezzhov, A.A., Komarova, T.V., et al., 2017. A novel block of plant virus movement genes. Molecular Plant Pathology 18, 611–624. doi:10.1111/mpp.12418. Leastro, M.O., Kitajima, E.W., Silva, M.S., Resende, R.O., Freitas-Astúa, J., 2018. Dissecting the subcellular localization, intracellular trafficking, interactions, membrane association, and topology of Citrus leprosis virus C proteins. Frontiers in Plant Science 11 (9), 1299. Locali-Fabris, E.C., Freitas-Astúa, J., Souza, A.A., et al., 2006. Complete nucleotide sequence, genomic organization and phylogenetic analysis of Citrus leprosis virus cytoplasmic type. Journal of General Virology 87, 2721–2729. Quito-Avila, D.F., Brannen, P.M., Cline, W.O., Harmon, P.F., Martin, R.R., 2013. Genetic characterization of Blueberry necrotic ring blotch virus, a novel RNA virus with unique genetic features. Journal of General Virology 94, 1426–1434. Robinson, T.S., Scherm, H., Brannen, P.M., Allen, R., Deom, C.M., 2016. Blueberry necrotic ring blotch virus in southern highbush blueberry: insights into in planta and in-field movement. Plant Disease 100, 1575–1579. Roy, A., Hartung, J.S., Schneider, W.L., et al., 2015. Role bending: Complex relationships between viruses, hosts, and vectors related to citrus leprosis, and emerging disease. Phytopathology 105, 1013–1025.

Brome Mosaic Virus (Bromoviridae) Guijuan He, Virginia Tech, Blacksburg, VA, United States Zhenlu Zhang, Shandong Agricultural University, Tai’an, China Preethi Sathanantham, Virginia Tech, Blacksburg, VA, United States Arturo Diaz, La Sierra University, Riverside, CA, United States Xiaofeng Wang, Virginia Tech, Blacksburg, VA, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of X. Wang, P. Ahlquist, Brome Mosaic Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00560-4.

Nomenclature aa Amino acid(s) CAT Chloramphenicol acetyltransferase CP Coat protein or capsid protein CPR5 Constitutive expresser of Pathogenesis-Related genes 5 ER Endoplasmic reticulum ESCRT Components of the endosomal sorting complex required for transport GFP Green fluorescent protein kb Kilobases; the size of a ssDNA or ssRNA molecule

Glossary Host factor Host proteins, lipids, and metabolites subverted by the virus at multiple stages of the life cycle required for successful viral infection. Spherule Single-membrane vesicular invaginations of host membranes induced by positive-strand RNA viruses in which viral replication proteins, RNA templates, and specific host factors are enriched and viral genome replication takes place. Other names used to refer to RNA replication competent spherules are viral replication compartments,

kDa Kilodaltons; the size of a protein mRNA Messenger RNA NCR Non-coding region nm Nanometer(s) nt Nucleotide(s) ORF Open reading frame RdRp RNA-dependent RNA polymerase RNAi RNA interference tRNA Transfer RNA UTR Untranslated region VIGS Virus-induced gene silencing

viral replication complexes, or viral replication miniorganelles. Sub-genomic RNA A messenger RNA produced during viral replication to express downstream viral genes from a viral genomic RNA expressing multiple genes. Tripartite RNA genome The viral genome is composed of three pieces of RNAs, which are all required for a successful systemic infection, may differ in length and in viral proteins encoded but may share similar sequences and/or structures of untranslated regions as well as 50 and/or 30 ends.

Introduction Brome mosaic virus (BMV) is a positive-strand RNA virus that infects cereal plants, causing mosaic symptoms and stunting. BMV is the type member of the genus Bromovirus in the family Bromoviridae. BMV is also a representative member of the alphavirus-like super-family that include viruses infecting human, animal, and plants that share similar genome structures, gene expression strategies, and protein functions. BMV has been used as a model to study gene expression, RNA replication, virus-host interactions, recombination, and encapsidation by positive-strand RNA viruses. Major contributions have revealed insights and principles extending to many other viruses and to general cell biology. Among numerous advances, research in BMV defined the first ribosome binding sites in eukaryotic mRNAs, produced the first eukaryotic viral RNA-dependent RNA polymerase (RdRp) extract that was template specific, the first infectious transcripts from cloned cDNA, the first engineered RNA virus expression vector expressing foreign genes, the first definition of sub-genomic mRNA synthesis pathways and determinants, and the first demonstration of higher eukaryotic viral replication in yeast. BMV has contributed many insights into virion assembly, virus-host interactions, RNA recombination, and RNA replication, including parallels in the replication of positive-strand RNA, reverse-transcribing and dsRNA viruses.

Virion Structure and RNA Encapsidation BMV forms non-enveloped virions B28 nm in diameter. The outer capsid is composed of 180 copies of coat protein (CP) arranged with T ¼ 3 quasi-icosahedral symmetry. Cryo-electron microscope reconstructions and subsequently x-ray crystallography showed that the BMV capsid structure is extremely similar to that of the related bromovirus Cowpea chlorotic mottle virus (CCMV). Intriguingly, some features of these capsids are dissimilar from other known isometric RNA virus capsids, but similar to capsids of the DNA-

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Fig. 1 Organization of the Brome mosaic virus (BMV) RNA genome. BMV has three genomic RNAs, RNA1, RNA2, and RNA3, and a sub-genomic mRNA, RNA4, encoded by the 30 portion of RNA3 as shown. Each of these four BMV positive-strand RNAs bears a 50 m7G cap and 30 tRNA-like structure (cloverleaf). The recruitment element (RE) in each viral genomic RNA and the promoter of sub-genomic RNA4 (Pro sgRNA) in RNA3 are shown. Open boxes represent the genes for the viral proteins 1a and 2apol, which are required for RNA replication, and 3a and coat protein, which are required for systemic movement in host plants. Some specific functions and features of each viral protein are listed, including the amphipathic a-helices at the N-terminus of 1a. The N-terminus (bottom half) and C-terminus (top half) of 1a are labeled as N and C, respectively. BMV 1a interacts with 2apol via the C-terminus of 1a and the N-terminus of 2apol.

containing papovaviruses. These features include orientation of the core b-barrel nearly perpendicular to the capsid surface to form distinctively prominent hexameric and pentameric capsomeres, and linking of adjacent capsomere clusters by exchange of invading Cterminal arms. These shared features suggest that BMV CP and the CPs of polyoma- and papillomaviruses likely share a common ancestor. BMV virion RNA is arranged as an interior subshell inside the capsid, leaving a hollow virion center. The N-terminal 26 aa of the CP, which are highly basic and required for RNA packaging, interact with the RNA to neutralize its charge. In addition to the predominant 180-subunit capsid, BMV CP can also assemble in vivo a 120-subunit capsid, composed of 60 CP dimers, first discovered in yeast and subsequently confirmed in infected plants. This assembly polymorphism is controlled in vivo by the RNA packaged, with BMV RNA2 packaged in 180-subunit capsids, while a small chimeric mRNA containing the CP open reading frame (ORF) as its only BMV sequence is packaged in 120-subunit capsids. Structural features shared by 120- and 180-subunit capsids imply that a common pentamer of CP dimers is an important intermediate in BMV virion assembly. In vitro and in vivo encapsidation identified portions of the BMV 3a coding region whose deletion blocked RNA3 encapsidation and interfered with normal co-encapsidation of RNA3 and RNA4. Studies also implicated the 3 0 tRNA-like structure of BMV RNAs as a facilitator of encapsidation in cis or trans, possibly by nucleating CP interactions into productive assemblies such as pentamers of dimers, and showed that BMV RNA replication promotes selective encapsidation of viral RNAs, possibly by inducing coupled synthesis of viral RNA and CP in close proximity. In 1971, buoyant density gradients were used to separate BMV virions into three classes, all three have identical capsids but package different RNAs (Fig. 1). Heavy virions contain a single copy of RNA1 (3.2 kb), medium density virions contain one copy each of RNA3 (2.1 kb) and RNA4 (0.9 kb), and light virions contain a single copy of RNA2 (2.9 kb). The three classes of viral particles are indistinguishable by electron microscopy, however, distinct capsid-RNA contacts influence the stability of the virions, viral RNA release, and BMV infection. Mapping of the contacts made between CP and the viral RNAs showed that RNA1 contacts CP differently than RNA2 or RNA3/4. Virions containing RNA1 were more sensitive to protease digestion and treatment of virions with RNase A revealed that RNA1 was preferentially degraded, while RNA2 or RNA3/4 were more resistant to RNase A digestion. The more facile interaction between RNA1 and the capsid proteins could ensure earlier and higher expression of BMV 1a, which is responsible for the formation of the viral replication compartments (see below in the Section of “Properties and Functions of Viral Gene Products”). The interaction of the viral RNAs with capsid is also influenced by the phosphorylation state of the N-terminal arm of the capsid protein. Interestingly, virions produced from different plant hosts had different post-translational modifications that impacted the thermostability of virions as well as the capsid-RNA interactions.

Genome Organization Genome Structure, Gene Expression and Sequences Interest in characterizing the protein(s) encoded by each of the four BMV virion RNAs motivated early in vitro translation studies with purified BMV virion RNAs. RNAs 1, 2, and 4 were characterized as monocistronic while RNA3 is dicistronic. Although early

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infectivity studies showed that RNA3 encodes CP, tryptic analysis showed that the principle translation product of RNA3 was unrelated to CP, while RNA4 served as an excellent template for CP. Moreover, when mixtures of all BMV RNAs were added to wheat germ extracts, increasing amounts of RNA4 inhibited translation of genomic RNAs 1–3. These results showed that RNA4 is a sub-genomic CP mRNA derived from RNA3, and implied an elegant system of gene regulation by translational competition: early in infection when viral RNA concentrations are low, all viral proteins including nonstructural RNA replication factors are translated, while after the virion RNAs are sufficiently amplified, CP is preferentially translated to encapsidate these RNAs. By the early 1970s, RNA bacteriophage studies had provided valuable information on prokaryotic translation initiation sites, including the finding that the first AUG initiation codon usually was 100 or more nucleotides from the RNA 50 end. In 1975, the first eukaryotic translation initiation site was characterized by isolating two fragments of BMV RNA4 that were efficiently bound by wheat germ ribosomes. RNA sequencing (a challenging, chromatography-based process at the time) revealed that the two fragments were 50 terminal, overlapping, and encoded the first 4 and 14 aa of CP, respectively. The most distinguishing feature was 0 0 that the initiating AUG codon for CP began only 10 nucleotides from the m7G5 ppp5 Gp-capped RNA4 50 end, presaging the now well-known mechanistic linkage between most eukaryotic translation initiation and 50 mRNA caps. The B8.2 kb BMV genome sequence was completed in 1984. In good agreement with in vitro translation results, RNA1 and RNA2 encode single proteins 1a (109 kDa) and 2a (94 kDa), respectively, the 50 half of RNA3 encodes protein 3a (35 kDa), and the 30 half of RNA3 encodes for RNA4, the sub-genomic mRNA for CP (20 kDa) (Fig. 1). Later work discussed below showed that 3a is required for infection movement in plants. Comparisons with other emerging viral RNA and protein sequences quickly revealed that the BMV 1a and 2a proteins, already implicated in RNA replication by protoplast experiments, shared extensive amino acid sequence similarities with proteins encoded by a diverse set of plant and animal positive-strand RNA viruses. These similarities initially were recognized between BMV, Alfalfa mosaic virus, Tobacco mosaic virus and Sindbis virus, and subsequently were found to extend to many other viruses now grouped together as the alphavirus-like super-family. Similarities shared by these viruses include a polymerase domain in BMV 2a (hereafter 2apol) and RNA capping and RNA helicase-like domains in 1a (Fig. 1).

Noncoding Regions, Cis Signals, and Sub-Genomic Promoter The first region of BMV RNAs to attract attention as a potential regulatory element was the 30 end. Synergistic work by multiple groups showed that the 30 non-coding regions (NCR) of BMV RNAs are highly conserved, contain multifunctional domains that direct negative-strand RNA synthesis, contribute to RNA encapsidation, translation and stability, and possess multiple tRNA-like features and functions. Limited early sequence data showed that BMV RNAs 1–4 share a tRNA-like CCAOH 30 end. All four BMV RNAs are selectively amino-acylated in vitro with tyrosine. Further studies showed that tyrosylated BMV RNAs interacted with translation elongation factor 1a and that BMV RNAs were tyrosylated in vivo during infection of barley protoplasts. BMV RNA 30 ends were also found to interact with (ATP, CTP):tRNA nucleotidyl transferase, which can add 30 CCAOH ends to mature BMV RNAs or maintain incomplete BMV RNAs, thus acting as a primitive telomerase. Beginning in the 1970s, further sequencing, enzymatic structure probing, and three-dimensional computer modeling showed that the 30 B200 nt of all BMV RNAs were strongly conserved and folded into an extended, tRNA-like structure with at least two alternate forms that differed in pairing nucleotides near the 30 with local or distal partners. Similarly conserved, highly structured 30 regions with related alternate forms were also found in other members of the family Bromoviridae. In vitro and in vivo studies using RNA fragments and mutations showed that sequences within the 30 -terminal tRNA-like structure direct negative-strand RNA initiation for RNA replication. Recent results have also implicated the tRNA-like sequence in translation and encapsidation. Involvement in all of these functions led to early and continuing suggestions that the 30 region mediates co-regulation of the varied uses of BMV positive-strand RNAs to minimize conflicts between multiple essential processes. A second class of elements combining tRNA-like features, replication signals and possible interaction with translation are the BMV template recruitment element (Figs. 1 and 2). Deletion analysis revealed that, in addition to 30 and 50 sequences required for negative- and positive-strand RNA initiation, BMV RNA3 replication in vivo requires a segment of B150 nt in the 50 half of the intergenic non-coding region between the 3a and CP genes (Fig. 1). Subsequent studies showed that this region is required for a step prior to negative-strand RNA synthesis, and is necessary and sufficient for 1a to recognize and recruit an RNA to a membraneassociated, translationally inaccessible, nuclease-resistant state that appears to be the interior of the replication compartments, also known as spherules (see the Section “BMV 1a and Viral Replication Compartments”). Structure probing studies show that this intergenic RNA3 sequence folds into a long, bulged stem-loop, which presents at its apex the invariant sequence and structure of tRNA TCC stem-loops. In plant and yeast cells, the appropriate BMV residues in this consensus are modified to T and C, showing that, like the 30 end, this sequence interacts in vivo with tRNA-specific enzymes. Moreover, any mutations to this TCC stem-loop mimicry abolish 1a-mediated template recruitment. Similar stem-loops with apical TCC stem-loop regions are found at the extreme 50 ends of BMV RNA1 and RNA2, where they similarly direct 1a-mediated template recruitment (Fig. 1). A common approach to express downstream gene(s) in a viral genomic RNA encoding more than one gene is to produce sgRNA(s) efficiently. In vivo and in vitro analyses of the BMV sub-genomic mRNA promoter have complemented well to reveal a core promoter within the 20 nt immediately upstream of the RNA4 start site, which directs low level but accurate initiation of sub-genomic mRNA. In vivo, the activity of this core promoter is greatly enhanced by upstream sequences that include an oligo(A) tract of variable, B16–22 nt in length in the viral population as well as upstream, partially conserved repeats of core promoter

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Fig. 2 Host factors required for the formation of functional BMV spherules. Host genes are involved in every step of BMV genome replication, including viral protein synthesis, recruitment of the viral RNA from ribosomes to the interior of spherules, targeting 1a to the nuclear ER membrane, membrane lipid composition and 1a-membrane interactions, spherule formation, and activation of replication activity. Host genes and pathways whose involvement in viral replication has been well characterized are listed.

sequences. The important role of the oligo(A) tract suggests that while negative-strand RNA can serve as a sub-genomic mRNA template in vitro, the natural in vivo template might be a dsRNA within which the oligo(A) provides a melting site to facilitate internal initiation. As one important aspect of cis signals in BMV replication, minimal core promoters were defined and dissected for the synthesis of negative-strand, positive-strand, and sub-genomic RNA, using a variety of approaches. Among other results, mutational studies imply that the BMV RdRp-promoter interaction has the characteristics of an induced fit, wherein the RdRp has some tolerance to adjust its binding to a range of promoter variants as long as some key sequence features remain. This model potentially reconciles the specificity of BMV RNA synthesis with the ability of the RdRp to synthesize different forms of viral RNA from separate promoters with distinct primary sequences and secondary structures. DNA or DNA/RNA hybrid templates containing the key BMV promoter sequences can be recognized in vitro by BMV RdRp extracts and copied into RNA. Although the efficiency of copying DNA templates is B15-fold lower than for BMV RNA templates, these results have significant potential implications for virus evolution.

Replication and Propagation In Vitro and In Vivo Replication Studies Positive-strand RNA viruses, like BMV, replicate their genomes in a completely RNA-dependent manner, producing a negativestrand RNA replication intermediate for each genomic RNA. Studies on BMV RNA replication were greatly advanced by the development and subsequent use of such tools as in vitro RdRp extracts and cultured plant protoplasts. In 1979, a virus-specific in vitro RdRp extract was developed from BMV-infected barley leaves that synthesized full-length negative-strand RNAs using added BMV virion RNAs as templates. This was the first eukaryotic in vitro RdRp preparation exhibiting a high level of template specificity, with other viral RNAs having less than 20% of the template activity of BMV RNAs. As noted in part below, this and similar BMV in vitro systems have been utilized to make many advances regarding promoters for positive- and negative-strand RNA synthesis, initiation mechanisms, and other issues. In parallel to in vitro systems, cultured plant protoplasts provided a valuable substrate for in vivo replication studies due to their ability to be infected or transfected with nearly 100% efficiency with virions, virion RNAs, or in vitro transcripts from cloned viral cDNAs. For BMV, barley protoplast systems have allowed examining all aspects of BMV RNA replication, sub-genomic RNA synthesis, progeny RNA encapsidation, and the like. The highly synchronized infections obtained also allow detailed kinetic studies. In the early 1990s, it was shown that BMV also would direct RNA replication, sub-genomic RNA synthesis, selective viral RNA encapsidation, RNA recombination and the like in the well-studied yeast, Saccharomyces cerevisiae. This host provides some of the same advantages as plant protoplasts together with rapid growth, a particularly small genome, well-characterized gene functions and cell biology, and powerful classical and molecular genetic tools including genome-wide arrays of isogenic yeast strains with each gene systematically modified by deletion, GFP-tagging, etc. Yeast cells expressing BMV RNA replication proteins 1a and 2apol support the

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replication of BMV genomic RNAs introduced by transfection or DNA-dependent transcription from plasmids or chromosomally integrated expression cassettes, duplicating all major features of replication in plant cells. To facilitate yeast genetic studies, BMV RNA replicons can express selectable or screenable markers and are transmitted to yeast daughter cells during cell division with 85%–90% efficiency, rivaling the transmission of yeast DNA plasmids. Some key findings identified and characterized in the BMV-yeast system have been confirmed in plants, including the enrichment of phosphatidylcholine at the viral replication sites in BMV-infected barley cells and an inhibition of BMV replication in BMV-infected Nicotiana benthamiana by overexpressing host gene PAH1 (Phosphatidic Acid Hydrolase 1) or a dominant-negative mutant of host SNF7 (Sucrose Non Fermenting 7)(see below in the Section “Host Factors in RNA Replication”). However, it should be noted that there are some differences in the replication features between plant and yeast cells as further discussed in the Section “BMV 1a and Viral Replication Compartments”.

Sub-Genomic mRNA Synthesis Early observations that the nature of CP in BMV infections is dictated by RNA3 rather than RNA4, and that RNA4 was regenerated when omitted from BMV inoculum, were partially explained when sequencing revealed that RNA4 was encoded by the 30 portion of RNA3. Nevertheless, whether RNA4 was produced from RNA3 by cleavage or any of several possible RNA synthesis pathways remained uncertain. In 1984, it was shown that a BMV RdRp extract produced sub-genomic RNA4 in vitro when supplied with negative-strand RNA3 templates, and that the product RNA4 could be labeled by g-32P-GTP incorporation, demonstrating de novo initiation. This first elucidation of a pathway for sub-genomic mRNA synthesis appears to provide a meaningful precedent for similar sub-genomic mRNA synthesis by many positive-strand RNA viruses, and an important foundation for understanding the diversity of alternate mechanisms that has begun to emerge with the demonstration of distinctly different sub-genomic mRNA synthesis pathways used by coronaviruses, nodaviruses and some other positive-strand RNA viruses.

RNA Recombination In 1986, BMV was used to provide an early demonstration of RNA recombination in a plant virus. Subsequent work demonstrated many forms and features of inter- and intra-molecular, homologous and non-homologous RNA recombination in BMV. Mutations in BMV RNA replication proteins 1a and 2apol could alter the frequencies and distributions of crossover sites, implying that at least a significant portion of such recombination was a byproduct of RNA replication, as by template switching. These and other results show that RNA recombination is a major force for repairing BMV genomes damaged by the high mutation rates of viral RNA replication and other events, thereby contributing to BMV survival and adaptability. Subsequently, it was further demonstrated that the recruitment of parental RNAs to a shared replication compartment is a limiting step in intermolecular RNA recombination. These results support a model in which BMV genomic RNAs are individually recruited into replication compartments, and intermolecular recombinants arise only in replication compartments receiving multiple viral RNAs.

Properties and Functions of Viral Gene Products BMV RNA3 encodes two proteins, 3a and CP, that are dispensable for RNA replication but are required for systemic spread (Fig. 1). Disruption of the 3a gene blocks cell-to-cell movement, limiting infection to individual, directly inoculated cells. The 3a protein shares multiple properties with cell-to-cell movement proteins of other viruses, including cooperative binding to single-stranded RNA, localization to the plasmodesmatal connections, and induction of virion-containing tubules from the surface of BMVinfected protoplasts. Disruption of the CP gene stops virus spread to uninoculated leaves. Whether local cell-to-cell spread occurs in the absence of CP depends on several factors including the 3a allele and the host plant. Protoplast studies showed that BMV RNA replication and sub-genomic RNA4 synthesis require 1a and 2apol but not 3a or CP. The conserved central domain of 2apol shows similarity to RdRps encoded by picornaviruses and many other RNA viruses. The N-terminal 1a domain is related to alphavirus protein nsp1 and contains m7G methyltransferase and m7GMP covalent binding activities required for capping viral RNA in vivo. The C-terminal 1a domain has sequence similarity to super-family I NTPase/helicases and NTPase activity that is required for RNA template recruitment and RNA synthesis. The N- and C-proximal domains are separated by a short proline-rich sequence that may serve as a flexible spacer. As shown in several systems, the C-terminus of BMV 1a interacts with the N-terminus of 2apol and this interaction is required for the recruitment of 2apol into the viral replication compartments.

BMV 1a and Viral Replication Compartments In yeast cells replicating BMV RNAs, the outer perinuclear endoplasmic reticulum (ER) membrane is invaginated towards the lumen to form numerous 50–80 nm spherular invaginations, which are referred to as spherules. Similar membrane invaginations are seen in plant cells infected by viruses from Bormoviridae, Tombusviridae, and Tymoviridae families, and in animal cells infected by viruses in and beyond the alphavirus-like super-family. However, in BMV-infected N. benthamiana cells, there are three types of polymorphic vesicles that are associated with cytoplasmic ER membranes but are absent from the perinuclear ER region. Type 1 vesicles are B66 nm in diameter with some spherule-like structures that remain connected to the ER through neck-like structures and as such, type 1 vesicles resemble those formed in yeast cells. Type 2 vesicles are B359 nm in diameter and likely are induced

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by CP. Type 3 vesicles are most probably formed by fusing type 1 and type 2 vesicles. Some possible reasons for the differences in BMV-induced membrane rearrangements between yeast and N. benthamiana are likely due to the organization and lipid composition of the ER membranes in yeast and N. benthamiana or possible variation in relative protein expression levels and replication efficiency in each system. Using immunogold-labeling and electron microscopy in yeast, replication proteins 1a, 2apol, and nascent BMV RNAs were found to be localized to the spherule interiors, which remain connected to the cytoplasm by a narrow neck that likely allows for the import of ribonucleotides required for replication and the export of offspring RNAs (Fig. 2). Immunogold and biochemical studies show that each spherule contains one or a few hundred 1a proteins and B25-fold fewer 2apol proteins along with specific host proteins (Fig. 2 and see the Section on “Host Factors in RNA Replication” below). The structure, assembly, and operation of these spherular replication complexes have functional and perhaps evolutionary links to the replicative cores of retrovirus and dsRNA virus virions, which sequester viral RNA replication templates and their polymerases in a protein shell. BMV 1a serves as the primary organizer to form active replication compartments: it invaginates the outer ER membranes into the ER lumen to form spherules, recruits 2apol via 1a-2apol interactions, recruits RNA templates into the interior of preformed spherules by recognizing the cis-element RE present only in viral genomic RNAs, and also interacts with and recruits specific host factors to the site of viral replication (Fig. 2, see below in the Section of “Host Factors in RNA Replication”). Spherular invaginations are formed when 1a is expressed in yeast cells in the absence of any other viral proteins or RNA templates. Localization of 1a to the perinuclear ER membrane requires an N-terminal amphipathic a-helix, termed Helix A (amino acids 392–407). NMR analysis shows that three leucine residues are deeply immersed into SDS micelles that mimicked a lipid bilayer. Substitutions of the three leucine residues with alanine prevent 1a from associating with the perinuclear ER membrane, and block spherule formation. In addition, Helix A plays critical roles in regulating the size and frequency of spherules as several individual substitutions in Helix A lead to similar viral replication defects including the formation of spherules that are more numerous in number but smaller in size. A second amphipathic a-helix, Helix B (amino acids 415–433), plays a crucial role in 1a’s function as a viroporin to permeabilize ER membranes to release oxidizing potential, which requires host thiol oxidase Ero1p (ER Oxidation 1), from the ER lumen to promote the formation of disulfide-bound 1a multimers, 1a’s RNA capping activity, and active genome replication (See Ero1p in the Section “Host Genes Required for Viral Replication Activity”).

Host Factors in RNA Replication As for many other viruses, the small size of BMV genome relative to the complexity of BMV replication suggests that many, if not most, steps in BMV RNA replication involve contributions from host factors. Indeed, genetic and systematic genome-wide screens have identified more than 100 genes that, when deleted, inhibited or enhanced BMV RNA replication by at least 3-fold. The mechanisms by which BMV usurps host factors to complete the various steps of its life cycle are illustrated in Fig. 2.

Host Genes That Regulate Lipid Metabolism and Membrane Composition Since positive-strand RNA viruses replicate on intracellular membranes, cellular lipid synthesis and appropriate lipid composition are crucial for their replication. BMV replication increases total fatty acids per yeast cells by 33%. Mutation or deletion of the OLE1 (OLEic acid requiring 1) gene, which encodes for the D9 fatty acid desaturase that converts saturated fatty acids to unsaturated fatty acids (UFAs), revealed that BMV RNA replication requires UFA levels five times higher than those required for normal yeast growth. Moreover, the gene products of DOA4 (Degradation Of Alpha 4), SPT23 (SuPpressor of Ty 23), and MGA2 (Multicopy suppressor of GAm 2), which are involved in regulating the expression of lipid metabolism genes, including OLE1, are also critical for proper BMV RNA replication. As major structural constituents of membranes, phospholipids are particularly important for the formation and activation of viral spherules. Phosphatidic acid is a precursor for both phospholipids and storage lipids. phosphatidic acid can be used to make phospholipids for membrane synthesis during active cell growth, or it can be converted to the storage lipid precursor diacylglycerol by Pah1p when cell growth reaches the stationary stage. Deleting PAH1 led to a 2-fold increase in total phospholipids levels, resulting in twice as many spherules and a 3-fold increase in viral replication. Further studies showed that BMV replication leads to a 25% increase in the levels of phosphatidylcholine, which is predominantly enriched at the sites of BMV replication in both yeast and barley cells. At least in yeast, BMV 1a interacts with and recruits Cho2p (Choline requiring 2), an enzyme involved in the conversion of phosphatidylethanolamine into phosphatidylcholine, to the viral replication sites to promote phosphatidylcholine synthesis, and facilitates BMV replication. In addition, deleting ACB1 (Acyl-CoA-Binding 1), another gene that regulates general lipid transport and synthesis, reduced BMV replication by up to 30-fold and it led to the formation of smaller but more abundant spherules compared to those in wild-type cells.

Host Genes That Induce/Stabilize Spherules Host factors are also essential to form and/or maintain BMV-induced spherules. The reticulons are a family of proteins that stabilize highly curved tubules in the peripheral ER. Through an interaction with BMV 1a, the reticulons are relocalized from the

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peripheral ER into the interior of the spherules. The reticulons were shown to stabilize the membrane curvature in the neck of the spherules and to also regulate the size and number of spherules. Components of the endosomal sorting complex required for transport (ESCRT) are also essential for spherule formation. One group of ESCRT genes regulate BMV RNA replication and the frequency of spherule formation while a second group of genes regulate RNA replication independent of spherule formation. In particular, deleting SNF7 significantly reduced viral replication and abolished detectable spherule formation. The current model suggests that BMV 1a initiates the invagination of the perinuclear ER, followed by recruitment of Snf7p and other ESCRT components to constrict the wide membrane rim, allowing the formation of the spherule body and neck. The reticulons in turn stabilize the curvature in the neck and regulate the size of the spherules.

Host Genes That Regulate Translation of Viral Proteins and 1a Targeting Until recently the mechanism by which BMV 1a is targeted to the perinuclear ER was unclear. It was shown that Erv14p (ER-vesicle protein of 14 kDa), a cargo receptor in COPII (COat Protein complex II) coated vesicles, not only interacts with 1a but helps localize it to the perinuclear ER. Deletion of ERV14 disrupts the proper distribution of 1a and significantly inhibits BMV RNA replication. Host ribosomes recognize the genomes of positive-strand RNA viruses to initiate translation of the viral proteins. Once sufficient viral replication proteins accumulate, viral genomic RNAs are recruited out of the cellular translational machinery and into spherules to serve as templates for viral replication. For BMV, control of these two functions is tightly regulated by the host complex consisting of Pat1p (Protein associated with topoisomerase 1) and 7 members of the Lsm (Like SM) family: Lsm1p to Lsm7p. Lsm1–7-Pat1 complex binds to the 50 and 30 untranslated regions (UTR) of BMV RNAs to circularize the RNAs to facilitate translation. When BMV 1a is present, 1a binds to the viral RNA and interacts with the Lsm1–7-Pat1 complex, likely disrupting Lsm1–7-Pat1 mediated 30 -50 circularization, thus repressing translation to allow recruitment of the viral RNA to the spherules. Moreover, classical yeast genetic approaches identified DED1 (Defines Essential Domain 1), an RNA helicase required for translation of all yeast mRNAs, to regulate translation of 2apol relative to other replication factors.

Host Genes Required for Viral Replication Activity Host factors are also essential to activate the viral replication proteins after spherules have been assembled. Ydj1p (Yeast dnaJ 1), a J-protein co-chaperone of Hsp70 and Hsp90, is required to activate the BMV RNA replication complexes. In yeast with a mutation in the YDJ1 gene, negative-strand RNA synthesis is inhibited even though 1a, 2apol, and RNA3 were recruited to ER membranes. The results suggest that the Ydj1p chaperone likely modulates 2aPol folding or assembly into the complex. BMV RNA replication also depends on the ER luminal thiol oxidase Ero1p. 1a can act as a viroporin by permeabilizing ER membranes to release the Ero1p-generated oxidizing potential from the ER lumen to activate 1a’s capping function via a new state of 1a multimerization involving oxidized linkages and potential conformational changes within the protein. 1a mutants lacking the viroporin function were able to recruit and stabilize positive-strand RNA3 templates but did not support significant RNA replication.

Transmission and Host Range BMV transmission in plants is not well characterized. BMV can be transmitted in the field by human activities involved in agricultural production, such as machinery trampling. In laboratory settings, plants can be easily infected by mechanical inoculation using sap prepared from BMV-infected tissues, purified virions or virion RNAs, or in vitro transcripts from cloned viral cDNAs. Vectors transmitting BMV in the field are unknown. Spotted cucumber beetles and nematodes in the genus Xiphinema have been reported to be able to transmit BMV in the laboratory. BMV replicates and encapsidates its RNAs in directly inoculated cells from a wide variety of plants, but has a fairly restricted host range for systemic infection of whole plants beyond monocotyledonous plants. The effective host range for BMV infection thus appears to be determined at the level of initiating or sustaining infection spread from the sites of primary infection. Exchanging genomic RNAs, individual genes and gene segments among BMV strains and between BMV and other viruses shows that adaptation for infection spread in particular host plants depends not only on 3a and CP but also on features of RNA1 and RNA2. Host adaptation of 3a generally exerts the predominant effects on infection spread, and only a few amino acid changes in 3a are required to extend BMV host range from monocotyledonous to dicotyledonous plants. However, changes modulating the efficiency of systemic spread also map to 1a and 2apol. Such changes may alter systemic spread through host-specific effects on RNA replication, as by influencing the ability of the virus to replicate and spread faster than host defense responses. Alternatively, 1a and 2apol may possess additional functions, as for some C-terminal 2apol sequences that are dispensable for RNA replication but required for efficient systemic infection. BMV primarily infects monocotyledonous cereal crops such as barley, maize, rice, wheat, and sorghum, as well as model plants Brachypodium distachyon and Setaria viridis. While N. benthamiana is a dicotyledonous host for BMV, Arabidopsis thaliana is generally considered as a non-host even though it allows BMV to infect in rare conditions. Screening numerous accessions and mutants of Arabidopsis, CPR5 (Constitutive expresser of Pathogenesis-Related genes 5) was identified as a key restriction factor for BMV systemic infection in Arabidopsis. Although Arabidopsis cpr5 mutant plants show an enhanced resistance to multiple bacterial and oomycete

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pathogens and several positive-strand RNA viruses compared to wt plants, they all support efficient systemic BMV infection. BMV RNA replication increased at the single-cell level by 6-fold in protoplasts prepared from cpr5 mutant plants compared to wt. The allowance for BMV replication in various cpr5 mutants is independent of host defense-related salicylic acid, jasmonic acid, and ethylene. Arabidopsis CPR5 is a component of nuclear pore complexes and serves as a selective barrier to restrict nuclear access of signaling cargos. However, the mechanisms whereby Arabidopsis cpr5 mutants allow BMV to replicate efficiently in single cells and promote systematic infection are unclear. The role of CPR5 might be direct, or it could indirectly regulate specific downstream effectors that might be responsible for the non-host resistance in Arabidopsis and possibly other dicotyledonous plants.

BMV Application in Biotechnology BMV was used to produce the first infectious transcripts from cloned RNA virus cDNA in 1984. Specially designed BMV cDNA clones were transcribed to produce capped in vitro transcripts of genomic RNAs 1–3, each with the natural viral RNA 50 end and only a few extra nucleotides at the 30 end. Mixtures of all three BMV RNA transcripts, but not their parent cDNA clones, were infectious to barley plants, a natural BMV host. This ability to engineer the expression of infectious transcripts provided a means to manipulate the viral RNA genome at the cDNA level using recombinant DNA technology, which has subsequently proved applicable to many other RNA viruses. In one of the first applications of these new reverse genetics approaches, it was successfully demonstrated that foreign genes could be inserted into the viral genome while retaining the ability to replicate and express genes. Using a transcribable BMV RNA3 cDNA clone, the CP gene was replaced with the bacterial reporter gene chloramphenicol acetyltransferase (CAT). When in vitro transcribed and inoculated onto barley protoplasts with RNA1 and 2 transcripts, this RNA3 derivative was replicated and produced CAT activity at higher levels than previously achieved by DNA-based transformation. This first demonstration that RNA viruses can be engineered at the cDNA level showed that the viral RNA genome functions in a sufficiently flexible and modular fashion to tolerate even large changes such as whole gene replacements without substantial optimization, which has significant implications for virus evolution, biotechnology research and applications such as additional gene expression vectors. RNA interference (RNAi) or gene silencing has been widely employed as a powerful tool to reveal gene functions in plants by degrading gene-specific transcripts. Virus-induced gene silencing (VIGS) knocks down gene expression of targeted cellular genes during virus infection, without a need to make transgenic plants. A BMV strain (fescue strain) isolated from tall fescue has been engineered as a VIGS vector by inserting a fragment of the target genes into the 30 UTR of RNA3. These BMV vectors have been successfully used in barley, maize, rice, and sorghum to understand gene function otherwise hard to achieve.

Future Perspectives Through a variety of intrinsic features and the work of many investigators, research on BMV not only has advanced understanding of bromoviruses, but also has contributed significantly to general virology and molecular biology. Some of the challenges for the future include improved definition and analysis of distinct substeps in viral RNA synthesis including initiation, elongation, termination and capping; better characterization at molecular, cellular and tissue levels of the pathways and mechanisms involved in infection spread and the interplay of virus-directed processes and host defenses; and improved understanding of the linkages between different infection processes including regulated gene expression, RNA replication, encapsidation and spread. Since the mechanism of action has been characterized for several host genes involved in BMV replication, the manipulation of host genes via knockout, knockdown, targeted substitutions, or overexpression could potentially be used as a way to achieve viral control in crop plants.

See also: Alfalfa Mosaic Virus (Bromoviridae). Bromoviruses (Bromoviridae). Cucumber Mosaic Virus (Bromoviridae). Ilarviruses (Bromoviridae)

Further Reading Ahlquist, P., 2006. Paralles among positive-strand RNA viruses, reverse-transcribing viruses and double-stranded RNA viruses. Nature Review Microbiology 4, 371–382. Chaturvedi, S., Rao, A.L.N., 2018. Molecular and biological factors regulating the genome packaging in single-stranded positive-sense tripartite RNA plant viruses. Current Opinion in Virology 33, 113–119. Cheng, K.C., Sivakumaran, K., 2000. Brome mosaic virus, good for an RN virologist’s basic needs. Molecular Plant Pathology 1, 91–97. Diaz, A., Wang, X., 2014. Bromovirus-induced remodeling of host membrane during viral RNA replication. Current Opinion in Virology 9, 104–110. Noueiry, A.O., Ahlquist, P., 2003. Brome mosaic virus RNA replication: Revealing the role of the host in RNA virus replication. Annual Review Phytopathology 41, 77–98. Rao, A.L., Cheng, K.C., 2015. The brome mosaic virus 30 untranslated sequence regulates RNA replication, recombination, and viral assembly. Virus Research 206, 46–52. Sztuba-Solinska, J., Bujarski, J.J., 2008. Insights into the single-cell reproduction cycle of members of the family Bromoviridae: Lessons from the use of protoplast systems. Journal of Virology 82, 10330–10340.

Bromoviruses (Bromoviridae) Jozef J Bujarski, Northern Illinois University, DeKalb, IL, United States and Polish Academy of Sciences, Poznan, Poland r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein ELISA Enzyme-linked immuno-sorbent assay ER Endoplasmic reticulum kb Kilobases; the size of a ssDNA or ssRNA molecule kDa Kilodaltons; the size of a protein MP Movement protein Mr Relative molecular mass NGS Next generation sequencing nm Nanometer(s) ORF Open reading frame(s)

Glossary Cross Protection It describes a phenomenon in that an initial infection of a host plant with a mild strain of a virus induces resistance in that plant to the infection of another, closely related virus potentially protecting the plant from disease caused by a more virulent isolate. Genome Activation Ilarviral genomic RNAs alone are unable to establish infection in plants, unless the coat protein is present. This function of the coat protein is termed genome activation and is specific for ilarviruses and closely related alfamoviruses. The event is triggered by binding of the coat protein RNA binding domain to the 30 terminus of genomic RNA. RNA Silencing A fundamental, evolutionarily conserved and sequence-specific mechanism that is triggered by double-stranded RNA and regulates gene expression in

PCR Polymerase chain reaction RBD RNA binding domain RdRp RNA-dependent RNA polymerase RT-PCR Reverse transcription polymerase chain reaction sgp Sub-genomic promoter sgRNA Sub-genomic RNA ssRNA Single-stranded ribonucleic acid TLS Transfer RNA-like structures UTR Untranslated region VIGS Virus-induced gene silencing VRC Virus replication complexes

eukaryotes. It is a primary antiviral defense mechanism in plants and other living organisms. RNA Silencing Suppressor A countermeasure to RNA silencing, often a protein encoded by a virus which interrupts a single, or multiple steps in the RNA silencing pathway such as binding to small interfering RNA and thereby preventing their incorporation into the RNA induced silencing complex. Sub-genomic RNA A segment of RNA generated from a genomic RNA via an internal promoter that has the same 30 end as the genomic RNA, but has a deletion at the 50 end. The sub-genomic RNA makes it possible to efficiently translate the downstream open reading frame of the genomic RNA. Tripartite Genome A viral genome consisting of three genomic fragments, which are encapsidated into three separate virions.

Introduction The family Bromoviridae contains important genera of plant viruses, with host ranges varying from narrow to very wide, and infecting herbaceous plants, shrubs and even trees. Several of them are responsible for major epidemics in fodder crops such as tomato, cucurbits, bananas, or alfalfa. The members of the family Bromoviridae have spherical or bacilliform particles with a trisegmented, positive-sense, single-stranded RNA (ssRNA) genome, packaged in separate virions. Bromovirids can be transmitted mechanically, via the pollen, seeds or by insect vectors. As shown in Tables 1 and 2, the Bromoviridae family includes six genera: Alfamovirus (one member, type species: Alfalfa mosaic virus, AMV), Anulavirus (two members, type species: Pelargonium zonate spot virus, PZSV), Bromovirus (six members, type species: Brome mosaic virus, BMV), Cucumovirus (four members, type species: Cucumber mosaic virus, CMV), Ilarvirus (22 members, type species: Tobacco streak virus, TSV), and Oleavirus (one member, type species: Olive latent virus 2, OLV-2). Genus demarcation criteria are based on natural host range, method of transmission, detailed morphology and properties of particles, organization of RNA genome, replication schemes and producing defective RNAs and satellite RNAs. The two prototype genera, Bromovirus and Cucumovirus, are the genera mostly related, with the latter being agriculturally important. Both bromo- and cucumoviruses share such properties like the molecular and genetic features of their tripartite RNA genome, the number of encoded proteins and similar virion structure. The computer-assisted comparisons of aa sequences reveal significant similarity among their RNA replication proteins, much beyond the presence of characteristic GDD motif for 2a or for helicase/transferase domains in 1a. More broadly, the replication proteins share aa sequence similarity within the alphavirus-like super-family of positive-strand RNA viruses, which includes numerous plant and important animal/human viruses. The type members of different genera, such as CMV, BMV and

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Table 1

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Main characteristics of the RNA genome in six genera of the family Bromoviridaea

Genus

Acronym

RNA1

RNA2

RNA3

30 UTR

sgRNA/diRNA

Alfamovirus Alfalfa mosaic virus

(AMV)

3644

2593

2037

Complex

–/–

Anulavirus Pelargonium zonate spot virus

(PZSV)

3383

2435

2639

Complex

–/–

Bromovirus Brome mosaic virus

(BMV)

3234

2865

2117

tRNA-like

þ /–

Cucumovirus Cucumber mosaic virus

(CMV)

3357

3050

2216

tRNA-like

þ/þ

llarvirus Tobacco streak virus

(TSV)

3491

2926

2205

Complex

–/–

Oleavirus Olive latent virus 2

(OLV-2)

3126

2734

2438

Complex

þ /–

a

Partial sequence.

AMV, have and continue to constitute excellent models for molecular research on viral gene expression, RNA replication, virion assembly, RNA recombination, epidemiology or the role of cellular genes in basic virology.

Phylogeny and Biodiversity of the Family Bromoviridae Although RNA1, 2 and 3 overall keep a great similitude in their sequences, clearly rearrangements of RNAs has been done for members of the Bromoviridae. Both RNA recombination but also segment reassortment played a major role as the sources of variation in shaping the bromovirids member groups, being important contributors to the evolutionary history of the family, especially for the genera Bromovirus, Cucumovirus and Ilarvirus (Figs. 1 and 2). However, doubts have been shed on the biological significance of the official taxonomy of the family Bromoviridae. To better understand the taxonomy, attempts have been made to reconcile the incongruences observed in the viruses’ evolutionary radiation caused by recombination and reassortment. These two processes could create new genetic variability and then these primary variants would undergo further selection for functional genomes of individual viruses. Consequently, the variants generated by reassortment and recombination events must have been initially viable which represents the first selective filter while further directional selection fine tunes the newly created RNAs. RNA segment reassortment was probably common at the origin of the bromoviruses and cucumoviruses as well as at the origin of Alfalfa mosaic virus, American plum line pattern virus and Citrus leaf rugose virus. Furthermore, recombination analyzes done for each of the three genomic RNAs revealed very common crossovers within the members of the genera Bromovirus, Cucumovirus and Ilarvirus, but also mixed recombination involving species from different genera. It seems that bromoviruses and cucumoviruses did split from a common ancestor forming distinct clades due to crossover events in RNA3, whereas protein 2b promoted the selection of a CMV-TAV RNA1/2-RNA3 recombinant. In the 50 untranslated regions (UTR) of CMV RNA3 the sequence rearrangements have likely been the precursors of the radiation of three cucumovirus subgroups. The results illustrated in Fig. 2 confirm a clear separation between the genera Bromovirus and Cucumovirus, while the ilarviruses constitute their own cluster; two other genera (Anulavirus and Oleavirus) are more unique within the family. Although these results suggest that AMV should be included in the ilarviruses, its unequivocal assignment has yet to be resolved, especially because AMV differs from other ilarviruses with the mode of transmission, by aphids versus by pollen and thrips. Bromovirids are members of a larger alpha-like supergroup based upon both 1a and 2a proteins whereas 3a proteins cluster the bromovirids together with other viral groups into a separate pool of movement-associated proteins. In general, the constructed phylogenetic network not only reflects the initial genetic exchanges but also confirms the taxonomic status of the different genera within the family Bromoviridae, notwithstanding the phylogenetic disturbances caused by genetic exchange.

Virion Properties and Structure Virions of bromovirids are non-enveloped, being either spherical or pseudo-spherical, with T ¼ 3 icosahedral symmetries, and a diameter of 26–35 nm (genera Anulavirus, Bromovirus, Cucumovirus and Ilarvirus), whereas genera Alfamovirus and some ilarviruses have bacilliform virions, of diameters 18–26 nm and lengths of 30–85 nm. In Oleavirus virions have different shape. In genus Bromovirus three types of icosahedral particles are composed of 180 molecules of a 20 kDa CP, and encapsidate different RNA components: RNA1 (Mr c. 1.1  106), RNA2 (Mr c. 1.0  106) and RNA3 plus sgRNA4 (Mr c. 0.75  106 and 0.3  106). In addition to viral RNAs, BMV virions have been recently reported to package small amounts of host RNAs, with their

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Table 2

List of genera and species in the family Bromoviridae. Type species are written in bold

Genus

Species

Acronym

GenBank accession no. RNA 1 (P1)

RNA 2 (P2)

RNA 3 (MP and CP)

Alfamovirus Anulavirus

Alfalfa mosaic virus Amazon lily mild mottle virus Pelargonium zonate spot virus

(AMV) (ALMMoV) (PZSV)

NC_001495 NC_018402 NC_003649

NC_002024 NC_018403 NC_003650

NC_002024 NC_018404 NC_003651

Tentative species

Cassava Ivorian bacilliform virus

(CsIBV)

NC_025482

NC_025483

NC_025484

Bromovirus

Broad bean mottle virus Brome mosaic virus Cassia yellow blotch virus Cowpea chlorotic mottle virus Melandrium yellow fleck virus Spring beauty latent virus

(BBMV) (BMV) (CsYBV) (CCMV) (MeYFV) (SBLV)

NC_004006 NC_002026 NC_006999 NC_003543 NC_013266 NC_004120

NC_004007 NC_002027 NC_007000 NC_003541 NC_013267 NC_004121

NC_004008 NC_002028 NC_007001 NC_003542 NC_013268 NC_004122

Cucumovirus

Cucumber mosaic virus Gayfeather mild mottle virus Peanut stunt virus Tomato aspermy virus

(CMV) (GMMoV) (PSV) (TAV)

NC_002034 NC_012134 NC_002038 NC_003837

NC_002035 NC_012135 NC_002039 NC_003838

NC_001440 NC_012136 NC_002040 NC_003836

(ALV) (BChRSV) (PaMoV) (PrRSV) (StNSV) (TSV)

NC_022127 NC_011553 NC_005848 NC_027928 NC_008706 NC_003844

NC_022128 NC_011554 NC_005849 NC_027929 NC_008707 NC_003842

NC_022129 NC_011555 NC_005854 NC_027930 NC_008708 NC_003845

(AsV2) (CiLRV) (CVV) (EMoV) (LRMoV) (SpLV) (ToNSV) (TAMV)

NC_011808 NC_003548 NC_009537 NC_003569 EU919668a NC_003808 NC_039075 NC_003833

NC_011809 NC_003547 NC_009538 NC_003568 NC_038777 NC_003809 NC_039074 NC_003834

NC_011807 NC_003546 NC_009536 NC_003570 NC_038776 NC_003810 NC_039076 NC_003835

(ApMV) (BlSV) (LiLChV) (PNRSV)

NC_003464 NC_022250 NC_025477 NC_004362

NC_003465 NC_022251 NC_025478 NC_004363

NC_003480 NC_022252 NC_025481 NC_004364

(FCLV) (PDV)

NC_006566 NC_008039

NC_006567 NC_008037

NC_006568 NC_008038

(APLPV) (ANMV) (CGV1) (HuJLV) (TPLPV)

NC_003451 NC_040469 NC_040393 NC_006064 NC_040435

NC_003452 NC_040471 NC_040392 NC_006065 NC_040436

NC_003453 NC_040470 NC_040394 NC_006066 NC_040437

(OLV)

NC_003673

NC_003674

NC_003671

Ilarvirus Subgroup 1 Ageratum latent virus Blackberry chlorotic ringspot virus Parietaria mottle virus Privet ringspot virus Strawberry necrotic shock virus Tobacco streak virus Subgroup 2 Asparagus virus 2 Citrus leaf rugose virus Citrus variegation virus Elm mottle virus Lilac ring mottle virus Spinach latent virus Tomato necrotic streak virus Tulare apple mosaic virus Subgroup 3 Apple mosaic virus Blueberry shock virus Lilac leaf chlorosis virus Prunus necrotic ringspot virus Subgroup 4 Fragaria chiloensis latent virus Prune dwarf virus Unassigned species American plum line pattern virus Apple necrotic mosaic virus Cape gooseberry virus 1 Humulus japonicus latent virus Tea plant line pattern virus Oleavirus

Olive latent virus 2

a

Partial sequence.

functions yet to be determined. The members of the genus Cucumovirus, in addition to three genomic RNAs, encapsidate two subgenomic RNAs (sgRNAs) and a satellite RNA. The crystal structures of both BMV and CCMV have been resolved showing very similar organization (Fig. 3), with the CP subunits folded into a beta-barrel core organized within the protruding both pentameric and hexameric capsomers. The interactions among hydrophobic aa residues stabilize the capsomers, and the hexameric subunits are further stabilized via interactions between N-terminal portions, where six short beta-strands form a tubule called beta-hexamer. Mutational analysis demonstrated that beta-hexamer was not required for virion formation but rather modulated virus spread in planta. In addition, the capsomers

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Fig. 1 Phylogenetic trees obtained for the three genomic segments RNA1, RNA2 and RNA3, by concatenating coding and non-coding regions and fitting a heterogeneous nucleotide-substitution model using CODEML. See Table 2 for virus names and abbreviations. The three trees have been aligned with a color-coded system for each genus to show similitudes and differences. The trees are reproduced from Codoner, F.M., Elena, S.F., 2008. The promiscuous evolutionary history of the family Bromoviridae. Journal of General Virology 89, 1739–1747 with permission (modified by C. Fauquet).

Fig. 2 Evolution of the family Bromoviridae. Unrooted phylogenetic network illustrating the evolutionary history of the family Bromoviridae. The different viral species in the family are linked to each other via multiple paths of interconnecting network rather than as single tree, which suggests the effects of recombination and reassortment events. Figure reproduced from Codoner, F.M., Elena, S.F., 2008. The promiscuous evolutionary history of the family Bromoviridae. Journal of General Virology 89, 1739–1747 with permission (modified by C. Fauquet).

are hold together by interactions through C-terminal portions that extend radially from the capsid. The C-termini are anchored between the beta-barrel core and the N-proximal loop, and this interaction might be responsible for initiation of assembly of CCMV capsids. The structure of BMV is organized similarly to that of CCMV such that both capsids undergo well-studied reversible structural transitions where shifting pH from 5.0 to 7.0 causes capsid expansion. However, some CP mutations can further stabilize the capsids. Capsids are also stabilized by metals at multiple biding sites that coordinate the amino acids from adjacent CP subunits. The packaged RNAs interact with the basic N-terminal aa of the CP in the torus-shaped sub-shell inside the BMV capsid so to neutralize the phosphate groups. Other sites of RNA interaction localize to the internally-proximal basic aa of the CP subunits. The RNA encapsidation signals have been mapped on BMV RNAs, especially to both the 30 UTR and a central sequence in RNA3. The co-packaging of sgRNA4 is contingent upon both RNA replication and translation of CP. The detailed knowledge of CCMV capsids provided opportunities in nanotechnology, e.g., for the reversible pH-dependent gating, useful during the regulation of size-constrained biomimetic mineralization. The interior surface of CCMV capsids can be engineered of as differentially functionalized CP subunits, to act e.g., as a ferritin surrogate that spatially constrains the formation

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Fig. 3 Montage of transmission electron microscopic photographs and computer rendering of molecular particle structures of bromovirids. A & B. Surface capsid structure of the Brome mosaic virus (BMV) (A) and Cowpea chlorotic mottle virus (CCMV) (B). The hexameric and the pentameric structural elements are visible. Both pictures are from the virion picture collection at the web-site of the Institute for Molecular Virology at the University of Wisconsin-Madison. C–F. Negative-contrast electron micrograph of particles of (C) Brome mosaic virus (Bromovirus), (D) Prunus necrotic ringspot virus (Ilarvirus), (E) Alfalfa mosaic virus (Alfamovirus), and (F) Olive latent virus 2 (Oleavirus). Photographs A-C IMV-Michigan State University, 1994 Thorben Lundsgaard, C.J. Woolston and Ed Rybicki, and Photograph D courtesy of A. Paredes, NCTR/ ORA, Arkansas USA. Photographs E and F, courtesy of A. De Stradis, IPSP-CNR, Bari, Italy. Bars ¼ 50 nm.

of iron oxide nanoparticles. Also, the electrostatically driven adsorption on Si and amine-functionalized Si as well as the fabrication of multilayer CCMV films have been reported. The electrostatically patchy protein cages of CCMV can be used to direct the assembly of super lattices for RNA encapsulation.

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Fig. 4 Genome organizations representatives of the six genera of the family Bromoviridae: (A) genus Alfamovirus, Bromovirus, Ilarvirus subgroups 3 and 4 and Oleavirus. (B) genus Anulavirus. (C) genus Cucumovirus and Ilarvirus subgroups 1 and 2. The 30 termini form either tRNA-like (B) or complex structures (A, C) shown as black square boxes. Figure from Bujarski, J., Gallitelli, D., García-Arenal, F., et al., 2019. ICTV Virus Taxonomy Profile: Bromoviridae. Journal of General Virology 100, 1206–1207.

Genome Organization and Expression The total length of the Bromoviridae RNA genome is approximately 8 kb, with three RNA segments capped at the 50 terminus. Whereas the highly conserved within members, the not polyadenylated 30 termini fold into strong secondary structures. These structures are either aminoacylable tRNA-like (genera Bromovirus and Cucumovirus) or forming other not aminoacylated arrangements (genera Alfamovirus, Anulavirus, Ilarvirus and Oleavirus) (Table 1). RNAs 1 and 2 are monocistronic and they code, respectively, for viral replicase proteins 1a and 2a (Fig. 4). Protein 1a has two distinct domains of guanylyl transferase and helicase, and it is involved in anchoring the replicase complex to the endoplasmic reticulum membrane, and induces the formation of membranous vesicular mini-organelles called spherules where RNA replication occurs. In AMV the replicase proteins interact with the tonoplast. Protein 2a is the actual RNA-dependent RNA polymerase enzyme that interacts with protein 1a and synthesizes the vRNAs. Mutations and deletions in 1a/2a ORFs helped to identify the regions active in BMV RNA replication as well as regions responsible for interaction with the cellular membrane or for interactions between 1a/2a polypeptides. An active BMV replicase preparation has been extracted. The anulavirus encodes the smallest RdRp (2a) protein within the family. For cucumoviruses and in some ilarviruses RNA2 is dicistronic encoding a protein 2b, that is part of the C-terminal region of the protein 2a. Protein 2b was found to act as suppressor of RNA interference, being involved in the inhibition of viral gene silencing but also in systemic movement and affecting the symptoms.

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RNA3 encodes the movement (MP) and coat (CP) proteins, the latter being translated from the subgenomic (sg)RNA4. BMV CP is a multi-functional protein. In addition to its structural/encapsidation role, it also coordinates the viral infection processes including (1) participation in the formation of replication factories, (2) repression of RNA replication but also translation, and (3) stimulation of BMV RNA accumulation at lower CP levels. Moreover, the BMV CP participates in RNA recombination events; an analogous function to that of BMV CP was assigned to nucleocapsid proteins in retroviruses and coronaviruses. The multiple functions of CP are exercised by effective binding to several distinct sites in RNA3, including the 30 non-coding hairpin, two central regions, and possibly at the 50 end. The contact BMV CP amino acids have been mapped at distinct parts of the CP monomers. Since BMV replicase complex also binds to most of these sites it has been postulated that the CP is involved in the regulation of BMV RNA replication such that the repression of RNA accumulation and translation occurs at higher levels of BMV CP, while stimulation of BMV RNA accumulation at the lower CP levels. While CP is usually translated from the encapsidated 30 sgRNA4, Olive latent virus 2 encapsidates a sgRNA with no apparent messenger activity whereas CP is translated from another non-encapsidated sgRNA4. The purified genomic RNAs are directly infectious but for some bromovirids the presence of CP is necessary (e.g., for AMV or ilarviruses). The 32 kDa MP of bromovirids has a conserved RNA binding domain that binds to vRNA and assists with viral transport. There are two groups among the members of the family Bromoviridae regarding the cell-to-cell transport; those that do not require participation of virions and those that transport whole virions. In the first group only the MP-RNA entity is transported (non-virion transport) in a form of either the complex of MP and vRNA, or a triple complex of vRNA, MP and CP. For the first group, the prototype example is TMV whereas members of the genus Bromovirus have a similar mechanism, which has been demonstrated for CCMV. The MP of these viruses belongs to the 30K superfamily of MPs that appear to interact with cellular microtubules. In the second subtype, where the transported complex is vRNA-MP-CP, the typical example is CMV, a comovirus. In this case, the CMV MP exhibits the binding affinity to the actine microfilaments. The second group transports whole virions intercellularly through plasmodesmata inside the microtubules (virion transport). BMV and AMV are the members of two genera that are transported this way. Also for ilarviruses such as PDV or PNRSV, their MPs likely assist during translocation of the entire viral particles alongside the tubular structures. It appears that MPs of these viruses interact with virion CP subunits via a 44C-terminal key aa domain. For long distance transport, most viruses require the CP which suggests that they are transported in the form of viral particles (virions). This has been demonstrated by showing that bromovirids require unmodified CP and the wild type C-terminus of MP for long distance spread, such as AMV, BMV and CMV. Very likely the CP-MP interactions enhance the systemic transport, independently of the mechanism of short distance (cell-to-cell) transport.

Genomic RNA Replication and Recombination Replication of bromovirus RNAs are the best studied among the members of the family Bromoviridae. Only viral proteins 1a (helicase and methyl-transferase domains) and 2a (RdRp), but not the proteins 3a or CP, are required for RNA synthesis (Fig. 4), first demonstrated for BMV, both in plant and in yeast cells. The cytoplasmic RNA replicase complex localizes to the endoplasmic reticulum membranes called spherules. The extracted bromoviral RdRp preparations have allowed for mapping in vitro on three genomic BMV RNAs the promoter of (–) strand synthesis within the 30 UTR. The promoters of ( þ ) strand synthesis have also been mapped to the 50 proximal non-coding region in BMV RNAs. Likewise, the sub-genomic promoter (sgp) has been localized as a 100 nt subset of the 250 nt intercistronic region in (–) strand in BMV RNA3, being responsible for synthesis (transcription) of sgRNA4. In addition, in the plus strand the 250 nt intercistronic sequence supports other functions including the efficient RNA3 replication, the maintenance of a proper ratio of ( þ ) to (–) strands of RNA3, stabilization of RNA3 via interaction with protein 1a, synthesis (transcription) of sgRNA4, and the assembly of the active RNA replication complex. It also serves as an efficient RNA recombination hot spot. Some bromoviruses as well as togaviruses carry the internal poly(A) tract, as part of the intercistronic region of the RNA3 segment. Besides viral RNA sequences and viral proteins, a variety of essential host genes affecting BMV RNA replication have been identified, by using yeast knockout libraries, as apparently the yeast cells can support a complete replication cycle of this virus Fig. 5. Both homologous and non-homologous RNA-RNA crossovers has been demonstrated between bromoviral RNAs during infection. Homologous recombination has also been shown during co-infection between two strains of BMV, with some distinct hot spots localized within both the coding and non-coding regions. The role in recombination of proteins 1a and 2a as well as of the CP have been demonstrated in BMV, suggesting the template switching as a likely mechanism for RNA recombination. For the cucumoviruses, the control of recombination frequency resides mainly in the 2a gene. Members of the genera Bromovirus and Cucumovirus are capable of producing the defective (d)RNAs and defective-interfering (di)RNAs during infection (Table 1). In particular, strains of Broad bean mottle virus (BBMV) do accumulate RNA2-derived deletion variants after serial passages through broad bean. In BMV, both replicating and non-replicating truncated RNA2-derived artificial diRNAs have been shown to interfere with BMV RNAs in protoplasts. For CMV, several types of RNA3-derived diRNAs have been described.

Biology The family Bromoviridae is one of the most important families of plant RNA viruses, with some members widely distributed in the world. In its entirety, the family has a wide host range (more than 10,000 species) and some members are causing agronomically

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Fig. 5 Severe outbreaks of Cucumber mosaic virus (CMV; Cucumovirus) containing the necrogenic satRNA (A) and Pelargonium zonate spot virus (PZSV; Anulavirus) (B) in crops of canning tomato in southern Italy. Insets show disease symptoms on fruits. Photographs courtesy of ICTV.

important diseases. However, the host range of members of individual genera ranges from significantly narrow (genera Bromovirus, Oleavirus) to extremely broad (genus Cucumovirus). CMV can infect one of the largest number of plant species among plant viruses. Some of these viruses cause major disease epidemics in vegetables, fodder and fruit crops, e.g., in tomato, cucurbits, bananas, or alfalfa, and in fruit trees (ilarviruses). Different virus species are transmitted mechanically, via pollen/thrips, through seeds or by insect vectors like aphids or beetles. It has been speculated that lack of inefficient vectors evolved some bromovirids toward producing larger concentration of viral particles. For CMV, although the virus is prone to recombination, the recombinants are rare during infection, suggesting the presence of strong selection bottlenecks. No direct correlation between virus yield and symptom severity have been observed, but rather the symptoms seem to be associated with changes in specific regions in the RNA genome, as it has been mapped by using the natural strains of BMV, BBMV or CCMV.

Further Reading Bujarski, J., Gallitelli, D., García-Arenal, F., et al., 2019. ICTV Virus Taxonomy Profile: Bromoviridae. Journal of General Virology 100, 1206–1207. Chaturvedi, S., Rao, A., 2018. Molecular and biological factors regulating the genome packaging in single-strand positive-sense tripartite RNA plant viruses. Current Opinion in Virology 33, 113–119. Codoner, F.M., Elena, S.F., 2008. The promiscuous evolutionary history of the family Bromoviridae. Journal of General Virology 89, 1739–1747. Diaz, A., Wang, X., 2014. Bromovirus-induced remodeling of host membranes during viral RNA replication. Current Opinion in Virology 9, 104–110. Gancarz, B.L., Hao, L., He, Q., Newton, M.A., Ahlquist, P., 2011. Systematic identification of novel, essential host genes affecting bromovirus RNA replication. PLoS One 6 (8), e23988. Kostiainen, M.A., Hiekkataipale, P., Laiho, A., et al., 2013. Electrostatic assembly of binary nanoparticle superlattices using protein cages. Nature Nanotechnology 8, 52–56. Pallas, V., Aparicio, F., Herranz, M.C., Sanchez-Navarro, J.A., Scott, S.W., 2013. The molecular biology of ilarviruses. Advances in Virus Research 87, 139–181. Schoelz, J.E., Harries, P.A., Nelson, R.S., 2011. Intracellular transport of plant Viruses: Finding the door out of the cell. Molecular Plant 4 (5), 813–831. Sztuba-Solin´ska, J., Urbanowicz, A., Figlerowicz, M., Bujarski, J.J., 2011. RNA-RNA recombination in plant virus replication and evolution. The Annual Review of Phytopathology 49, 415–443. Takeshita, M., Matsuo, Y., Suzuki, M., et al., 2009. Impact of a defective RNA 3 from cucumber mosaic virus on helper virus infection dynamics. Virology 389, 59–65. Weber, P.H., Bujarski, J.J., 2015. Multiple functions of capsid proteins in ( þ ) stranded RNA viruses during plant-virus interactions. Virus Research 196, 140–149.

Bymoviruses (Potyviridae) Annette Niehl, Julius Kühn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany Frank Rabenstein, Julius Kühn Institute, Quedlinburg, Germany r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

NIa-Pro Nuclear inclusion protein a-proteinase NIb Nuclear inclusion protein b nm Nanometer nt Nucleotide(s) ORF Open reading frame PCR Polymerase chain reaction RNA Ribonucleic acid siRNAs Small-interfering RNAs UTR Untranslated region VPg Viral protein genome-linked

bp Base pair CI Cylindrical inclusion protein CP Coat protein dsRNA Double-stranded RNA eIF4E Eucaryotic translation initiation factor 4E ELISA Enzyme-linked immunosorbent assay ER Endoplasmic reticulum GFP Green fluorescent protein kDa Kilodalton MB Membranous body

Glossary Bymovirus

From barley yellow mosaic virus.

Classification The bymoviruses are single stranded positive-sense RNA viruses with a bipartite genome in the family Potyviridae.

Member Species The Bymovirus genus currently comprises six member species, Barley mild mosaic virus (BaMMV), Barley yellow mosaic virus (BaYMV), Oat mosaic virus (OMV), Rice necrosis mosaic virus (RNMV), Wheat spindle streak mosaic virus (WSSMV), and Wheat yellow mosaic virus (WYMV). The type member of the genus is BaYMV. The currently unclassified Soybean leaf rugose mosaic virus seems to be a related virus species, since the C-terminal region of the NIb gene exhibits about 60% amino acid sequence identity with barley mild mosaic virus and other bymoviruses.

Virion Structure, Serological- and Cytological Properties Bymoviruses are flexuous filamentous particles of 250–300 nm and 500–600 nm in length and approximately 13 nm in width (Fig. 1(A)). The bipartite, positive-sense RNA genome is encapsidated by a single coat protein (CP). BaYMV is serologically related to WSSMV; WSSMV and WYMV appear to be serologically indistinguishable as neither polyclonal antisera or monoclonal antibodies are able to differentiate between them. OMV and BaMMV are serologically distinct from the other members of the genus. Bymoviruses induce pinwheel-like inclusions and endoplasmic reticulum (ER)-derived inclusions (membranous body, MB) with crystalline lattice-like or tubule dominated morphology (tube-MB) and lamellar structures (lamella-MB) in infected host cells (Fig. 1(B) and (C)). Electron tomography analysis and immuno-gold labeling revealed that both MB types contain viral proteins and components derived from ER membranes. The irregularly arranged lamellar-MB was composed of “rough” ER like structures, whereas regularly arranged tubule-MB was composed of “smooth” ER structures.

Nucleic Acid Properties and Differentiation of Bymoviruses The bymovirus positive-sense genomic RNAs are approximately 7.5–8.0 kb (RNA1) and 3.5–4.0 kb (RNA2) in length. The RNAs contain the virus-encoded genome-linked viral protein VPg covalently bound to their 50 end and a poly-A tail. The WYMV poly-A tail in naturally infected wheat is variable in length and can even be absent. Recent experiments using in-vitro translation and replication showed that polyadenylation of the viral 30 ends impacted translation and negative strand RNA synthesis, indicating that polyadenylation may represent a means for the virus to fine tune translation or replication or may even represent a switch

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Fig. 1 Electron microscopy of bymoviruses and infection-derived cellular structures. (A) Transmission electron microscopy of a virus purification from BaMMV infected plants reveals flexuous particles of approximately 250–300 and 500–600 nm in length. (B) and (C) Ultra thin sections of BaYMV (A) and BaMMV (B) infected cells showing ER-derived membranous bodies (MB) of tubular (B) or lamellar morphology (C). V virus particles, PW pinwheel inclusion bodies, scale bars 500 nm.

Fig. 2 Phylograms depicting relationships between the different bymovirus member species. Trees were made with alignments of polyprotein amino acid sequences in CLC main workbench 8.1.3 using the UPGMA algorithm, Jukes-Cantor distance measure and 100 bootstraps, indicated as numbers next to the branches.

between translation and replication of viral proteins. Interestingly, positive effects of cap-independent translation by the 50 UTRs (untranslated region) of WYMV RNA1 and RNA2 was counteracted by the presence of a poly-A tail at the 30 UTR of RNA1 and further increased by the presence of a poly-A tail at the 30 UTR for RNA2. Given that bymovirus resistance is at least in part based on VPg-eIF4 interaction (see below, paragraph on virus control), efficient cap-independent translation by bymoviruses due to a variable polyadenylation status may provide a possibility for the virus for host adaptation. Sequence similarity comparisons of polyprotein sequences from bymovirus reference numbers published in the 10th ICTV report on the family Potyviridae (last revised in October 2018, see “Relevant Websites section”) and the genebank reference sequence assembly data for WSSMV (RefSeq assembly: GCF_004117675.1) revealed relationships between polyproteins derved from RNA1 and RNA2 (Fig. 2). For RNA1 polyproteins, WYMV, WSSMV, BaYMV and OMV are more closely related to each other than to RNMV and BaMMV. For RNA2 polyproteins, RNMV and BaMMV form a subgroup, while WYMV, WSSMV, BaYMV and OMV form a different subgroup.

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Fig. 3 Genome organization of bymoviruses. Black lines represent viral RNAs with VPg protein bound to the 50 end and variable poly A tail at the 30 end. Square boxes along the RNA represent encoded proteins and their relative localization.

Organization of the Genome and Properties of the Encoded Proteins Bymovirus RNAs encode for polyproteins, which are proteolytically cleaved into functional proteins. RNA1 contains the typical potyvirus organization of genes except for P1 and HcPro proteins (Fig. 3). Bymovirus RNA2 encodes a P1 protein with amino acid sequence similarity to HcPro, which is restricted to the C-terminal cysteine protease domain of the protein, and a bymovirusspecific P2 protein. As the genome organization for RNA1 is shared among members of the Potyviridae family, it is likely that the proteins also have common functions. The large polyprotein derived from RNA1 is proteolytically cleaved into eight mature proteins, P3, 7K, CI (cylindrical inclusion) protein, 14K, VPg (viral protein genome-linked), nuclear inclusion protein a-proteinase (NIa-Pro), nuclear inclusion protein b (NIb), and CP (coat protein). The NIa-Pro is responsible for cleavage. Potyviral CI is thought to be involved in virus movement, VPg is a multifunctional protein involved in translation, replication and possibly movement, NIb is involved in replication, and CP is important for viral encapsidation. Apart from inferring viral protein functions from studies of other potyviruses, the functions of some of the proteins encoded by bymovirus RNAs have been studied in detail for WYMV. The small PIPO open reading frame (ORF) overlapping with the P3 ORF has been predicted in the WYMV genome. Bymovirus RNA2 encodes a polyprotein that is cleaved into the P1 and P2 proteins. The P1 protease cleaves at a site with the consensus sequence (Y,F, G)xG↓(A, N, S). P2 is believed to be important for virus transmission by its vector, Polymyxa graminis. VPg proteins are covalently attached to viral genomic RNA and involved in efficient translation of viral RNA. The WYMV VPg protein interacts with the other viral proteins P1 and CP, contains nuclear export and import signals and has been shown to localize to nucleus and cytoplasm during infection, indicating functions of the protein in both cellular compartments. Translocation to the cytosol depends on interaction of VPg with CP, however, the mechanism, by which CP may increase cytosolic accumulation of VPg remains unresolved. The viral VPg interacts with translation initiation factors of the eIF4E family to initiate translation and replication of viral proteins. Recently, WYMV CP has been shown to interact with TaHSP23.6, a wheat small heat shock protein and expression of a range of heat shock proteins was induced upon WYMV infection. Bymovirus infection causes strong modifications of the ER membrane (membranous body (MB)). The inclusions derived from ER membranes produced during WYMV infection have been shown to contain multiple viral proteins. The integral membrane protein P2 interacts with itself, P1, P3, VPg, NIa-Pro, and CP, rearranges the ER and forms large aggregate structures, thus likely induces the formation of the inclusions by interacting with the ER and recruiting other viral proteins including P1, P3, NIa-Pro, and VPg, but not 7K, CI and CP into these inclusions. Moreover, P2 has been described to recruit host proteins into P2-derived inclusions. P2 interacts with the small GTPase Sar1 important for COPII-vesicle mediated ER-to-Golgi transport in P2-derived inclusions. The formation of the inclusions appears to depend on a functional secretory pathway. As the inclusions are associated with RNA synthesis, they likely represent sites of viral replication. Moreover, P2 appears to be important for transmission of the virus by its vector, P. graminis. Upon repeated mechanical inoculation, deletions within the 30 region of the P2 cistron occur for BaYMV, BaMMV and OMV. Two transmembrane domains have been predicted in bymovirus P2, as well as in Furovirus, Pomovirus and Benyvirus CP-Read-through protein (CP-RT). The second of these domains appears to be crucial for vector transmission and is lost in non-transmissible deletion mutants. For C-terminally truncated P2 forms replication still occurs in host cells, indicating that the C-terminal region of P2 is dispensable for virus replication. For BaYMV, the entire P2 protein appears to be dispensable for virus replication, but appears to be important for efficient systemic virus accumulation and symptom development. The peripherally ER membrane-associated WYMV P1 protein interacts with itself and with the other WYMV-encoded proteins, except 14K, and the WYMV integral membrane protein P3 localizes to the ER and Golgi and interacts with itself, P1, P2 and CP. P1 contains cysteine protease activity and believed to cleave the RNA2-derived polyprotein into the mature P1 and P2 proteins. In protoplasts, BaYMV P1 expression increases accumulation of CP, suggesting that P1 enhances RNA1 replication. BaYMV P1 also appears to be essential for systemic infection. Consistent with these functions and in analogy to potyviral P1/HCPro, bymoviral P1 may be involved in the suppression of RNA silencing; however, up to now, there is no experimental evidence for a role of bymoviral P1 as viral silencing suppressor. Nevertheless, that bymovirus double-stranded (ds)RNA replication intermediates are targeted by RNA silencing has been shown. RNA silencing differs between leaves and roots of WYMV infected plants in the way that more small interfering RNAs (siRNAs) – but also more virus- accumulate in roots versus leaves of the plants and that siRNAs in leaf but not root tissues demonstrate a bias towards 50 A or U, and may hence be incorporated into different Argonaute complexes. Moreover, Dicer-like gene expression seems to be differently regulated upon WYMV infection in leaves and in roots.

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WYMV 7K was shown to localize to ER membranes when fused to GFP. The WYMV CI protein has been shown in reassortant experiments to determine the pathogenicity of virus infection. Three amino acids in the N-terminal region of CI involved in the change of pathogenicity were identified. As replication in protoplasts remained unchanged between different WYMV pathotypes, it was assumed that the reason for different pathogenicity between WYMV pathotypes is related to CI function in virus movement. Nucleotide sequences of several German WSSMV isolates used for virus resistance trials differ in their 14K protein region from the sequences of isolates found in France, Denmark, and the USA by a nona-nucleotide insertion. At present, it is not clear whether the amino acid changes in these virus isolates lead to biological effects and differences in resistance behavior.

Virus Transmission and Movement The bymoviruses are transmitted by the soil-borne plasmodiophorid P. graminis. Despite challenging for some species of the genus, bymoviruses can also be transmitted mechanically. Transmission takes place after the virus-carrying zoospores of the vector penetrate host root epidermal or root hair cells; however, the detailed mechanism by which the virus crosses the zoosporangial plasmalemma to enter the host cell cytoplasm has not been illuminated. No evidence exists to demonstrate that the virus is able to replicate in its vector. WSSMV CP was detected inside P. graminis resting spores. Virus inside resting spores remains infectious over many years. Once transmitted to the host plant, the virus moves systemically. Whether the virus moves cell to cell and systemically as virion or as viral ribonucleoprotein complex in its host plants has not been determined.

Symptoms, Epidemiology and Host Range Typical symptoms of bymovirus infection constitute of chlorotic mosaic of dark green areas on the younger leaves and chlorotic streaks on leaves (Fig. 4), growth depression, reduced vigor, fewer tillers and reduced grain yield. In the field, winter crops usually develop symptoms in late winter and early spring. Symptomatic plants are usually found in patches in the fields. Barley yellow mosaic disease caused by BaMMV and BaYMV was first reported in Japan in 1940. Since, it has also been detected in China, South Korea, and Europe including Belgium, Bulgaria, France, Germany, Greece, Hungary, Italy, the Netherlands, Poland, Russia, Spain, the UK, and Ukraine. In Japan, at least seven strains of BaYMV and two BaMMV strains are known, while in Europe currently two BaYMV and three BaMMV strains have been described. In China, at least six strains of BaYMV are probably present. Crop loss due to the barley-infecting bymoviruses can reach up to 50% in infected areas. Bymovirus-caused wheat yellow mosaic disease can be due to WSSMV and WYMV infection. WSSMV was first identified 1960 in Canada and ten years later in the USA, where it quickly spread throughout the country. In Europe, WSSMV is present in several countries, among them France, Italy, Germany, and Belgium. WSSMV has also been reported to occur in Africa. Wheat losses due to WSSMV have been reported to range between 20 and 25%. WYMV was first reported in Japan and also occurs in India and China. RNMV was first detected in Japan, and then in India in the late 1970th and causes yellow flecks and streaks on rice leaves. OMV was first reported in 1944 in the USA, where it is widely distributed. It is also present in Europe (England, France, Ireland, and Italy). Yield losses caused by oat mosaic disease of more than 50%, even to complete losses, have been reported. With respect to yield losses, it should be noted here that bymovirus infection often occurs as co-infection with furoviruses and the contributions of each virus species to the disease phenotype and associated crop losses have not been dissected. Thus, it remains unclear whether other virus factors contribute to the losses. Apart from other biotic parameters, also abiotic parameters such as temperature, humidity, soil conditions etc. contribute to the yield loss. Under natural conditions, BaYMV and BaMMV infect only barley, WSSMV and WYMV wheat, OMV oat and RNMV rice. Interestingly, for RNMV mechanically inoculated onto Ludwigia, Corchorus, and Hibiscus species, growth promotion phenotypes have been described.

Fig. 4 Disease symptoms caused by bymoviruses. (A) Barley field showing patches of plants with barley yellow mosaic disease symptoms. BaYMV (B) and BaMMV (C) leaf mosaic symptoms on winter barley leaves.

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Diagnosis Depending on the cultivated variety and the environmental conditions, the severity of disease symptoms caused by bymovirus infection can vary greatly and normally, virus symptoms disappear with increasing temperatures. The frequent occurrence of mixed infections with other viruses makes a precise diagnosis based on the symptoms very difficult or impossible. Thus, immunoassays such as double-antibody sandwich (DAS)- and triple antibody sandwich (TAS)-ELISA are still most commonly used for practical diagnosis. Commercial test kits are available for all bymoviruses except for RNMV. The detection of virus by electron microscopy can be made specific through the use of immuno-gold labeled antibodies. For the detection and differentiation of the serologically closely related WSSMV and WYMV, a multiplex RT-PCR method was developed. Quantification and improvement of detection sensitivity for WYMV could be achieved by real-time quantitative RT-PCR. In addition, a sensitive reverse transcription loop-mediated isothermal amplification (RT-LAMP) method was developed for detection of WYMV and BaYMV. Reliable detection of BaYMV and BaMMV by real-time PCR has also been reported. Furthermore, a RT-PCR protocol to specifically detect OMV has been developed. The application of High-Throughput Sequencing (HTS), also known as next-generation sequencing (NGS), has proven very successful also for virus detection of bymoviruses in field samples. In barley, the rym4 gene confers resistance to BaMMV and the common BaYMV pathotype, BaYMV-1. However, a new virus pathotype designated BaYMV-2, which overcomes the rym4-controlled resistance, appeared in several European countries. In order to discriminate between BaYMV-1 and BaYMV-2 by molecular methods a dCAPS (derived Cleaved Amplified Polymorphic Sequences) tool was developed and successfully applied to track the spread of pathotype BaYM-2 in France. It has been recently shown that deep small (s)RNA sequencing is a universal approach for virome reconstruction and RNAi characterization. This approach was used on dried barley leaves from field surveys. Illumina sequencing of sRNAs from plant samples identified besides BaYMV an endornavirus and, in one plant, a novel strain of Japanese soil-borne wheat mosaic virus (JSBWMV).

Control Apart from preventive measures by decontamination of machinery and man, the planting of resistant crop varieties presently represents the only efficient means of control for bymovirus infection. Cultural practices remain ineffective due to the robustness of resting spores and soil treatments are ecologically unacceptable. While cereal varieties with resistance to P. graminis have not been identified, several crop varieties with resistance to one or more bymoviruses are available. Against barley yellow mosaic disease caused by BaMMV and BaYMV, 19 rym resistance genes are known that are attributed to at least nine loci in the barley genome. 15 of these resistance genes are inherited in a recessive manner and are effective against one or more pathotypes or strains of the same virus or against more than one virus. Some of the resistance genes have been mapped and identified. One encodes for the eukaryotic translation initiation factor 4E (eIF4E). rym4, rym5, rym6, rymHor4224, and rymHor3298 resistance based on eIF4E are allelic. Alleles confer resistance against different strains of BaYMV and/or BaMMV. The viral VPg interacts with eIF4 and is presumed to be important for virus translation, replication and movement. The second resistance gene encodes a protein disulfide isomerase like 5-1 (PDIL5-1). To our knowledge, nothing is known about the mechanisms underlying protein disulfide isomerase–based resistance to BaMMV and BaYMV. The monogenic resistances are relatively frequently broken and lead to the emergence of new virus pathotypes. Thus, apart from the identification of novel resistance genes and introduction of these genes into breeding programs, the currently most promising way to achieve durable resistance is by pyramiding resistance genes. In the case of WYMV, there is evidence that resistance is, similar to the barley infecting bymoviruses, eIF4E-mediated. Replication studies of WYMV in barley and wheat protoplasts indicated that expression of wheat eIF4E enabled the replication of WYMV in barley, which is otherwise restricted to wheat. Moreover, approaches in transgenic wheat overexpressing the viral replicase gene in antisense orientation resulted in durable resistance against WYMV. Against WSSMV, the resistant trait in commercial winter wheat varieties encodes a single dominant resistance gene on the long arm of chromosome 2D or is controlled in an additive dominant manner by two loci. WSSMV resistance probably limits virus spread from the roots to the shoots of the plants, as leaves of resistant varieties are readily mechanically infected and lead to systemic distribution of virus. OMV tolerance, in which virus accumulates to high titers but symptoms are mild or absent, has been identified in different Avena species and is inherited as a quantitative polygenic trait. Most of the Japanese paddy rice varieties can become infected when grown in soil infested with RNMV. To our knowledge, no resistance genes for RNMV have been identified so far. However, a study using transgenic plants overexpressing a HAP (heme activator protein) transcription factor revealed that the plants displayed a resistance phenotype towards RNMV infection by accumulating lower virus titer and the absence of visible disease symptoms. Moreover, infection induced expression of the transcription factor.

Further Reading Adams, M.J., 2002. Bymovirus. In: Tidona, C.A., Darai, G., Büchen-Osmond, C. (Eds.), The Springer Index of Viruses. Berlin, Heidelberg: Springer. Geng, G., Yu, C., Li, X., Yuan, X., 2019. Variable 30 polyadenylation of Wheat yellow mosaic virus and its novel effects on translation and replication. Virology Journal 16, 23. Ghosh, S.K., 1982. Growth promotion in plants by rice necrosis mosaic virus. Planta 155, 193–198. Hooper, G.R., Wiese, M.V., 1972. Cytoplasmic inclusions in wheat affected by wheat spindle streak mosaic. Virology 47, 664–672.

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Kühne, T., 2009. Soil-borne viruses affecting cereals – Known for long but still a threat. Virus Research 141, 174–183. Lapierre, H.D., Hariri, D., 2008. Cereal viruses: Wheat and barley. In: Mahy, B.W.J., van Regenmortel, M.H.V. (Eds.), Encyclopedia of Virology, third ed. Amsterdam: Elsevier, pp. 490–497. Li, L., Andika, I.B., Xu, Y., et al., 2017. Differential characteristics of viral siRNAs between leaves and roots of wheat plants naturally infected with wheat yellow mosaic virus, a soil-borne virus. Frontiers in Microbiology 8, 1802. 1802. Li, H., Kondo, H., Kühne, T., Shirako, Y., 2016. Barley yellow mosaic virus VPg is the determinant protein for breaking eIF4E-mediated recessive resistance in barley plants. Frontiers in Plant Science 7, 1449. 1449. Ordon, F., Perovic, D., 2013. Virus resistance in barley. In: Varshney, R.K., Tuberosa, R. (Eds.), Translational Genomics for Crop Breeding. Wiley. doi:10.1002/9781118728475.ch5. Rolland, M., Villemot, J., Marais, A., et al., 2017. Classical and next generation sequencing approaches unravel Bymovirus diversity in barley crops in France. PLoS One 12, e0188495. Stein, N., Perovic, D., Kumlehn, J., et al., 2005. The eukaryotic translation initiation factor 4E confers multiallelic recessive Bymovirus resistance in Hordeum vulgare (L.). The Plant Journal 42, 912–922. Sun, L., Andika, I.B., Shen, J., Yang, D., Chen, J., 2014. The P2 of Wheat yellow mosaic virus rearranges the endoplasmic reticulum and recruits other viral proteins into replication-associated inclusion bodies. Molecular Plant Pathology 15, 466–478. Sun, L., Jing, B., Andika, I.B., et al., 2013. Nucleo-cytoplasmic shuttling of VPg encoded by Wheat yellow mosaic virus requires association with the coat protein. Journal of General Virology 94, 2790–2802. Timpe, U., Kühne, T., 1995. In vitro transcripts of a full-length cDNA of a naturally deleted RNA2 of barley mild mosaic virus (BaMMV) replicate in BaMMV-infected plants. Journal of General Virology 76, 2619–2623. You, Y., Shirako, Y., 2010. Bymovirus reverse genetics: Requirements for RNA2-encoded proteins in systemic infection. Molecular Plant Pathology 11, 383–394. You, Y., Shirako, Y., 2013. Evaluation of host resistance to Barley yellow mosaic virus infection at the cellular and whole-plant levels. Plant Pathology 62, 226–232.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/w/potyviridae/568/genus-bymovirus Bymovirus. http://www.dpvweb.net/dpv/showdpv.php?dpvno=374 Barley yellow mosaic virus.

Cacao Swollen Shoot Virus (Caulimoviridae) Emmanuelle Muller, The French Agricultural Research Center for International Development, Joint Research Units–Biology and Genetics of Plant-Pathogen Interactions, Montpellier, France and Biology and Genetics of Plant-Pathogen Interactions, University of Montpellier, The French Agricultural Research Center for International Development, French National Institute for Agricultural Research, Montpellier SupAgro, Montpellier, France r 2021 Elsevier Ltd. All rights reserved.

Glossary Badnavirid Virus belonging to the genus Badnavirus, family Caulimoviridae. Badnavirus Virus genus of plant double strand DNA virus encapsidated in bacilliform particles. Pararetrovirus Reverse transcribing viruses that replicate through an RNA intermediate.

Retrotransposons Are genetic elements that can amplify themselves in a genome and are ubiquitous components of the DNA of many eukaryotic organisms. RTase/RNaseH Enzymes responsible for the replication of the pararetrovirus group.

Introduction Cacao swollen shoot disease (CSSD) is caused by Cacao swollen shoot virus (CSSV), member of the genus Badnavirus, family Caulimoviridae. CSSD can be regarded as the major viral disease on cacao and has been recognized as one of the most important diseases in West Africa limiting cacao production. The disease appeared only after the introduction of cacao in West Africa and is probably the result of a host jump from native reservoir plants. It was highlighted for the first time in Ghana in 1922 but was described and named in 1936, then in Nigeria in 1944, in Ivory Coast in 1946, in Togo in 1949, and in Sierra Leone in 1963. CSSV is naturally transmitted to cacao (Theobroma cacao) in a semi-persistent manner by several mealybug species. Extensive research on this emerging disease has been conducted since the 1940s in West African Cacao Research Institute (WACRI- now CRIG for Cacao Research Institute of Ghana) and then in other institutes, in virology, epidemiology, and genetic improvement of cacao tolerance to CSSD. Nevertheless yet, the disease has not been controlled and continues to spread into new areas. Other viral diseases have been also described on cacao in Trinidad and Tobago and in Sri Lanka but associated with slightly different symptomatologies and have not caused epidemics similar to cacao swollen shoot disease. Complete viral genomes have been obtained recently from those diseases and correspond to clearly different badnavirid species: Cacao yellow vein-banding virus (CYVBV) and Cacao mild mosaic virus (CMMV) infecting cacao in Trinidad, Cacao bacilliform Sri-Lanka virus (CBSLV) infecting cacao in Sri Lanka.

Taxonomy The genus Badnavirus, one of the eight genera of the family Caulimoviridae, are double stranded DNA (dsDNA) viruses with bacilliform particles that emerge mainly on tropical plants. The most commonly encountered on crops in addition to CSSV on cocoa are Banana streak viruses - BSV - on banana, Dioscorea bacilliform viruses -DBV- on yam and Citrus yellow mosaic virus -CiYMV- on citrus. The viruses member of this genus have an extremely wide host range: some viruses are found in many ornamental plants such as Yucca, Dracaena, Kalanchoe, Hibiscus, Pelargonium, Bougainvillea, Cycas, Canna, Jasminum, Wisteria and Commelina as well as in a significant number of cultivated plants, such as pineapple, fig, sweet potato, grapevine, black pepper, jujube, currant, raspberry, ginger, mulberry, and blackberry-ripe in addition to the major crops mentioned above (Banana, cocoa, and citrus). New badnavirids are regularly identified on new plant species such as Sophora from Japan, Yacon, birch, or Aralia Ming (Polyscias fruticosa L.) are found hosting badnaviruses. A total of 57 species are currently recognized by ICTV (Table 1) and many more are unassigned species in the genus (450). The description of some of these new viruses is certainly linked to the improvement of virus detection and testing techniques, rather than viruses a particular health problem for plant. It should be noted that the species demarcation of 80% identity of sequences has been set more than 15 years ago, and now that so many badnavirid sequences are available, this demarcation threshold should be revisited. It is remarkable that BSV, DBV, CSSV and even RTBV (Tungrovirus, Caulimoviridae) seem to have many species for a single disease in each case, or would it be possible that in the case of badnaviruses, or even caulimoviruses, that the species threshold might be much lower than 80%?

Host Range and Symptomatology Experimental host range is limited to species of the large botanical family of Malvaceae but the principal host of the virus is Theobroma cacao. Symptoms on cacao trees are mostly seen in leaves, but stem and root swellings, as well as pod deformation also occur. In some

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Cacao Swollen Shoot Virus (Caulimoviridae) Table 1 List of members of the genus Badnavirus (Caulimoviridae). The list of assigned and unassigned species in the genus is indicated. The accession numbers for the reference complete genome sequences are indicated in the last column Species in the genus Badnavirus

Abbreviation-Isolate

Accession #

Aglaonema bacilliform virus Banana streak GF virus Banana streak IM virus Banana streak MY virus Banana streak OL virus Banana streak UA virus Banana streak UL virus Banana streak UI virus Banana streak UM virus Banana streak VN virus Birch leaf roll associated virus Blackberry virus F Bougainvillea spectabilis chlorotic vein-banding virus Cacao bacilliform Sri Lanka virus Cacao mild mosaic virus Cacao swollen shoot CD virus Cacao swollen shoot CE virus Cacao swollen shoot Ghana M virus Cacao swollen shoot Ghana N virus Cacao swollen shoot Ghana Q virus Cacao swollen shoot Togo A virus Cacao swollen shoot Togo B virus Cacao yellow vein-banding virus Canna yellow mottle associated virus Canna yellow mottle virus-AP01 Citrus yellow mosaic virus Commelina yellow mottle virus Dioscorea bacilliform AL virus Dioscorea bacilliform AL virus 2 Dioscorea bacilliform ES virus Dioscorea bacilliform RT virus 1 Dioscorea bacilliform RT virus 2 Dioscorea bacilliform SN virus Dioscorea bacilliform TR virus Fig badnavirus 1 Gooseberry vein banding virus BC isolate Grapevine roditis leaf discoloration associated virus Grapevine vein clearing virus Jujube Mosaic associated virus Kalanchoe top-spotting virus Mulberry badnavirus 1 Pagoda yellow mosaic associated virus Pineapple bacilliform CO virus Pineapple bacilliform ER virus Piper yellow mottle virus Rubus yellow net virus Schefflera ringspot virus Spiraea yellow leafspot virus Sugarcane bacilliform Guadeloupe A virus Sugarcane bacilliform Guadeloupe D virus Sugarcane bacilliform IM virus Sugarcane bacilliform MO virus Sweet potato pakakuy B virus Taro bacilliform CH virus Taro bacilliform virus Wisteria badnavirus 1 Yacon necrotic mottle virus

ABV BSGFV BSImV BSMyV BSOLV BSUAV BSULV BSUIV BSUMV BSVNV BLRaV BVF BCVBV CBSLV CMMV CSSCDV CSSCEV CSSGMV CSSGNV CSSGQV CSSTAV CSSTBV CYVBV CaYMaV CaYMV CiYMV ComYMV DBALV DBALV2 DBESV DBRTV1 DBRTV2 DBSNV DBTRV FBV 1 GVBV-BC GRLDaV GVCV JuMaV KTSV MBV1 PYMaV PBCoV PBErV PYMoV RYNV SRV SYLSV SCBGAV SCBGDV SCBIMV SCBMOV SPPV TaBCHV1 TaBV WBV1 YaNMoV

MH384837 AY493509 HQ659760 AY805074 AJ002234 HQ593107 HQ593109 HQ593108 HQ593110 AY750155 MG686419 KJ413252 EU034539 MF642736 KX276641 JN606110 MF642719 MF642724 MF642725 MF642726 AJ781003 L14546 KX276640 KX0660020 MF074075 AF347695 X52938 X94576 KY827395 KY827394 KX008574 KX008577 DQ822073 KX430257 JF411989 HQ852250 KT965859 JF301669 KX852476 AY180137 LN651258 KJ013302 GU121676 KC808712 KF241951

FJ824813 FJ439817 AJ277091 M89923 FJ560943 KP710177 AF357836 KX168422 KM229702 (Continued )

275

276

Cacao Swollen Shoot Virus (Caulimoviridae)

Table 1

Continued

Species in the genus Badnavirus

Abbreviation-Isolate

Accession #

Unassigned species Banana streak CA virus Banana streak UJ virus Banana streak UK virus Banana streak PE virus Cacao swollen shoot Ghana T virus Cycad leaf necrosis virus Dracaena mottle virus 1 Hibiscus bacilliform virus Pelargonium vein banding virus

BSCAV BSUJV BSUKV BSPEV CSSGTV CLNV DrMV HiBV PVBV

HQ593111

MN187554 MN179342 EU853709 DQ473478 KF875586 GQ428155

Fig. 1 (A) Symptoms of red-vein banding observed on young flush leaves of cocoa tree in Togo, Kloto area. (B) Symptoms of swellings on chupons of cocoa tree, in Togo, Litimé area. (C) Symptoms of swollen shoot observed on Theobroma cacao plantlets four months after agroinoculation with the Togolese Agou 1 isolate. From left to right: a plantlet inoculated with the wild strain Agrobacterium tumefaciens LBA4404 and two plantlets inoculated with the recombinant A. tumefaciens bacteria LBA4404 (pAL4404, pBCPX2) containing CSSV insert.

varieties of cacao, particularly Amelonado cacao, reddening of primary veins and veinlets in flush leaves is characteristic (Fig. 1(A)). This red vein banding later disappears. There can be various symptoms on mature leaves, depending on the cacao variety and virus strain. These symptoms can include: yellow clearing along main veins; tiny pin-point flecks to larger spots; diffused flecking; blotches or streaks. Chlorotic vein flecking or banding is common and may extend along larger veins to give angular flecks. Stem swellings may develop at the nodes, internodes, or shoot tips. These may be on the chupons, fans, or branches (Fig. 1(B)). Many strains of CSSV also induce root swellings. Stem swellings result from the abnormal proliferation of xylem, phloem, and cortical cells. Infected trees may suffer from partial defoliation initially due to the incompletely systemic nature of the infection. Ultimately, in highly susceptible varieties, severe defoliation and dieback occurs. Smaller, rounded to almost spherical pods may be found on trees infected with severe strains. Occasional green mottling of these pods is seen and their surface may be smoother than the surface of healthy pods. Various isolates were described in Ghana as in Togo with a variability and a gradation in the type of symptoms observed. Severe strains of this virus can kill susceptible cacao trees within 2–3 years. They affect Amelonado cacao, widely considered to give the bestquality cacao, more seriously than Upper Amazon cacao and hybrids which have been selected for resistance to the virus. A few avirulent strains occur in limited, widely scattered outbreaks, usually inducing stem swellings only, and having little effect, if any, on growth or yield. As for other perennial crops, we observe a latency period between the time of infection and the time of symptom expression, which complicates control measures. Moreover, there are periods of remission during which symptoms are not visible. The natural host range of the virus includes Ceiba pentandra, Cola chlamydantha, Cola gigantea var. glabrescens, Sterculia tragacantha, and Adansonia digitata with associated symptoms of transient leaf chlorosis or conspicuous leaf chlorosis. Viral sequences belonging to different species have been detected in C. pentandra as well as in Commelina sp.

Transmission Fourteen species of mealybugs (Pseudococcidae spp.), including Planococcoides njalensis, Planococcus citri, Planococcus kenyae, Phenacoccus hargreavesi, Planococcus sp. Celtis, Pseudococcus concavocerrari, Ferrisia virgata, Pseudococcus longispinus, Delococcus tafoensis, and Paraputo anomalus, have been reported to transmit CSSV. Only the nymphs of the first, second, and third larval stages and the adult females are able to transmit the virus. The virus does not multiply in the vector and is not transmitted to its progeny. CSSV is transmitted neither by seed nor by pollen. CSSV can infect cacao at any stage of plant growth. The virus is transmitted experimentally to susceptible species by grafting, particle bombardment, by agro-inoculation using transformed

Cacao Swollen Shoot Virus (Caulimoviridae)

277

Fig. 2 Electron microscopic picture of virus particles of Cacao swollen shoot virus (CSSV), genus Badnavirus, family Caulimoviridae, 28
130 nm. The bar represents 200 nm.

Agrobacterium tumefaciens and with difficulty by mechanical inoculation. Seedlings usually produce acute red vein banding within 20–30 days and, 8–16 weeks later, swellings on shoots and tap roots (Fig. 1(C)). The indigenous species are more difficult to infect than cacao, and mealybugs do not become infective as readily when feeding on them as when feeding on infected cacao.

Virion Structure It was not until 1940 that through experimentation, the causative organism of the CSSD was identified to be a virus named CSSV. CSSV possesses small non-enveloped bacilliform particles and a double-stranded DNA genome (Fig. 2). The bacilliform particles measure 130 nm
28 nm in size and have been shown by dot–blot hybridization to occur in the cytoplasm of phloem companion and xylem parenchyma cells.

Genome Organization Molecular characterization of CSSV had a boost in 1990, thanks to the improvement of the techniques of purification. The first complete genome of a CSSV isolate (Agou1 from Togo) was determined in 1993. Five putative open reading frames (ORFs) are located for this isolate on the plus strand of the 7.16 kbp CSSV genome (Fig. 3). ORF1 encodes a 16.7 kDa protein whose function is not yet determined. The ORF2 product is a 14.4 kDa nucleic acid-binding protein. ORF3 codes for a polyprotein of 211 kDa which contains, from its amino- to carboxyl-terminus, consensus sequences for a cell-to-cell movement protein, an RNA binding domain of the coat protein, an aspartyl proteinase, a reverse transcriptase (RTase), and an RNase H. The last two ORFs X (13 kDa) and Y (14 kDa) overlap ORF3 and encode proteins of unknown functions. More than 30 complete viral genomes have since been obtained by different technologies such as Sanger sequencing of PCR amplified full genome and Illumina sequencing. Their size varies from 6.8 kbp to 7.4 kbp. ORFY has been identified in all other complete genomes sequenced. ORFs 1, 2 and 3 are characteristic of all badnavirids unlike additional ORFs which are present only for some badnaviruses (CSSV, Citrus yellow mosaic virus –CiYMV-, Taro bacilliform virus –TaBV).

Phylogeny Badnaviruses are highly variable at both the genomic and serological level, a feature which complicates the development of both molecular and antibody-based diagnostic tests. Moreover, CSSV isolates were for a long time classified according to the variability of the symptoms expressed on T. cacao, but it is not known if there is a correspondence between this variability of symptoms and the intrinsic molecular variability of the viruses. A first study analyzed the molecular variability of the area of the ORF3 coding for N-terminal coat protein (CP) for 1A-like isolates from Ghana. Successive studies have analyzed molecular variability of CSSV isolates from Togo, Ghana and Côte d0 Ivoire in the first part of ORF3 coding for the cell to cell movement protein. These analyses showed a high variability of CSSV isolates and permitted to define several groups (A to P) of sequences (Fig. 4). Groups Q, R and T were never detected with these primers. Recently the analysis of all complete viral genomes of isolates from those countries made it possible to describe more exhaustively the variability of this virus and to show that the disease is caused by a complex of many viral

278

Cacao Swollen Shoot Virus (Caulimoviridae)

Fig. 3 Organization of the circular genome of the CSSV species. Gray arrows indicate the deduced ORFs 1–3, X and Y capable of encoding proteins larger than 9 kDa. In the ORF3, we indicate the position of the movement protein (MP); coat protein (CP); the RTase/RNaseH region, with the domains for Aspartyl Proteinase (AP); Reverse Transcriptase (RT); and Ribonuclease H (RNase H).

species. Results on phylogenetic relationships between the first Ghanaian and Togolese full sequences showed that these relationships were more influenced by their geographical origin than on whether the sequences originated from mild or severe isolates. Table 2 shows the nt sequence identity of pairwise combinations of representative CSSV isolates of each molecular group in the RT/RNase H region of ORF3 (500 bp). Considering the 20% divergence threshold in the RTase/RNaseH region indicated by ICTV in a region of 1500 nt for the determination of new badnavirid species, we could define 8 species. Seven species are now recognized by ICTV: Cacao swollen shoot Togo B virus (CSSTBV, first described as CSSV, first isolate sequenced, Agou1), Cacao swollen shoot Togo A virus (CSSTAV), Cacao swollen shoot CD virus (CSSCDV), Cacao swollen shoot CE virus (CSSCEV), Cacao swollen shoot Ghana M virus (CSSGMV), Cacao swollen shoot Ghana N virus (CSSGNV), Cacao swollen shoot Ghana Q virus (CSSGQV). Another full genome has been obtained recently and potentially corresponding to the eighth species Cacao swollen shoot Ghana T virus (CSSGTV). Recently a phylogenetic tree was built from all the available complete sequences of badnaviruses (Fig. 5, Table 3). It shows that the group of viral species associated with the cacao swollen shoot disease is polyphyletic since the MBV1, HiBV and CiYMV species are positioned at the junction between the two phyla of CSSV species TA, TB (groups B and C), CD, CE (groups E, F, G, H, J, K, L, P) GM and GN on the one hand and GQ (groups Q and groups Q and R) and GT on the other hand. As has been observed for most viruses studied, that badnavirus populations contain recombinant viruses. This was illustrated for the first time for CSSV with the sequencing of isolate Wobe12 (type member of species CSSTAV). Wobe12 possesses an ORF1 close to ORF1 (96% nt identity) of isolate Agou1 (group C of species CSSTBV) but shares only 75% nt identity with the complete sequence of Agou1. Recombinations allow the emergence of new viral isolates and increases the variability of viral populations beyond what is observed by studying a single region of the genome. The sites of recombination in the genome and their frequency should be extensively investigated in the future to have a better understanding of virus evolution in epidemics.

Epidemiology: Geographical Distribution of Viral Molecular Diversity Epidemiological characteristics of the disease derive directly from its natural transmission properties. The mealybug vectors (nymph of both sexes and adult females) spread the disease radially over short distances around the periphery of outbreaks by crawling through the canopy from infected trees to adjacent healthy trees. New outbreaks were shown to be occasioned by “jump spread” over greater distances by wind borne viruliferous mealybugs and mainly by the very active small first instar nymphs. A systematic molecular characterization of viral isolates make it possible to study the molecular epidemiology of CSSV. Viral molecular diversity studied in the first region of the ORF3 for isolates from the three countries: Togo, Côte d0 Ivoire and Ghana

Cacao Swollen Shoot Virus (Caulimoviridae)

279

Fig. 4 Maximum likelihood phylogenetic tree of CSSV sequences based on alignment of the 50 end of open reading frame 3 (ORF3). Numbers on the branches represent the SH-aLRT (approximate likelihood ratio test) branch supports over 0.7. The names of the CSSV groups A, B, C, D, E, F, G, H, J, K, L, M, N and P are indicated. as well as the names of the corresponding species. The Citrus yellow mosaic virus sequence – CiYMV (AF347695) – is used as the outgroup. The names of sequences include the abbreviation of the country: CI for Côte d0 Ivoire, G or Gha for Ghana, To for Togo, a sampling number along with a region code or a number corresponding to the clone number in brackets, and the year of sampling 1993–2016 coded as 93–16. When sequences belonging to different groups are obtained for the same isolate, the letter corresponding to the group is affixed to the name of the isolate.

280

Cacao Swollen Shoot Virus (Caulimoviridae)

Table 2 Nucleotide sequence identity of pairwise combinations of representative CSSV isolates of each molecular group in the RT/RNase H region of ORF3 (500 bp) Species name CSSTAV CSSTBV CSSCDV CSSCEV

CSSGMV CSSGNV CSSGQV CSSGTV

A B C D E F G J K L P M N Q R T

CSSV group isolate name

A

B

C

D

E

F

G

J

K

L

P

M

N

Q

R

T

ToWOB12–02 GhaNew Juaben-00 ToAgou1–93 CI152–09 CI632–10 CI617–10 Gha62–15 GWR198–13 GWR3–14 GCR329–14 Gha59–15 Gha57–15 Gha63–15 Gha1–00 Gha60–16 Gha2–00

100

73 100

76 89 100

75 72 71 100

72 73 73 69 100

72 75 74 69 83 100

73 73 75 70 83 80 100

70 70 70 68 82 84 78 100

72 76 76 69 86 85 82 80 100

71 74 74 69 82 85 83 80 83 100

72 72 73 69 85 81 82 81 83 82 100

70 74 74 69 72 71 72 71 71 72 71 100

69 70 71 67 71 71 72 68 72 73 70 76 100

60 59 60 62 59 60 58 59 58 58 58 60 57 100

58 57 59 61 56 60 59 58 58 58 56 57 55 76 100

56 59 61 60 60 59 58 56 59 60 56 57 56 60 62 100

has provided a picture of the geographical distribution of this diversity in these countries (Fig. 6). In most cases, sequences from the same farm/plot always belonged to the same group, but there were some instances where isolates belonging to two different groups were identified on the same plot/farm. The geographical distribution of the viral molecular diversity shows that some group (species TA or CD, or group C of species TB) have remained well localized in a single country or at the border between two countries, while other species (group B of species TB) have a more ubiquitous distribution in the different regions of the three countries. The ubiquitous nature of the B group may mean it is readily transmitted by mealybugs, probably because of higher concentrations in the cacao phloem than the other groups. This group from species TB tends to induce more easily the typical red vein banding symptom seen in young or flush leaves of infected cacao plants, since most of the samples collected in the Eastern Region containing only this group, whether young or mature, had the conspicuous red vein banding or vein clearing symptom. When CSSV groups identified in Ghana are compared to those found in Côte d0 Ivoire, only species TB and CE are common to both countries. In addition to these two species, Côte d0 Ivoire has a species (CD) which have not been detected in Ghana up to now and Ghana also has species TA, GM, GN, GQ, group C (species TB) and groups G, J, K, L (species CE) which have not been identified or detected in Côte d0 Ivoire yet. This distribution may reflect differences in cacao crop planting dates in the regions of these countries. These observations make it difficult to predict a single origin for CSSV in West Africa amongst the cacao growing neighboring countries and we can therefore make the assumption that the different putative CSSV species emerged from different sources at different times. In fact, the high variability observed within CSSV populations compared to its very short evolutionary history on cacao trees further suggests the existence of several parallel emergences, more likely by host shift from different native hosts to cacao trees in the various countries of West Africa. Studies on CSSV more exhaustively in adventitious plants or insect vectors would make it possible to better understand the geographical distribution and the epidemiologic factors that lead to the rapid expansion of the disease in some plots. However, the exact role played by these wild hosts in the epidemiology of CSSV is today difficult to investigate, because the prevalence of CSSD is greater in cacao trees than in these alternative host plants. The diversity of CSSV observed in the CRIG Museum (a collection of viral isolates collected in Ghana since the 1940s) compared to cacao farms in Ghana is interesting from a historical point of view, because this range can be seen as a picture of the diversity of the CSSV population at the time the Museum was established. Indeed, it should be noted that this viral collection has not been regularly updated. The species CSSGQV, CSSGTV currently only detected in the CRIG Museum may have been present previously in cacao farms but were absent or only present at a frequency that avoided detection in the farms. Conversely, groups of the species CE (J, K and L) and group C only detected in cacao farms of the species TB appear to correspond to recently emerged groups in the cacao farms. Group C has been detected in Togo since 1993 but could have emerged either in Togo or in Ghana, since the establishment of the CRIG Museum. Species CE, is underrepresented in the CRIG Museum but present in a high proportion in field samples (29% of the total isolates characterized in Ghana) and appears to correspond to a species that has recently spread to cacao plots especially in the Western and Brong Ahafo regions of Ghana. The complexity of the molecular diversity of viral species found in symptomatic cacao leaves means that there is a need to study the range of aggressiveness of the different species or groups. In future the symptomatology/aggressiveness associated with single strain infections arising from representative species should be explored as should the impact on hosts from mixed infections of these genomes.

Cacao Swollen Shoot Virus (Caulimoviridae)

281

Fig. 5 Maximum likelihood phylogenetic tree generated by PhyML based on complete nucleotide sequences of badnaviruses. The blue sequences correspond to the viral sequences infecting the cocoa tree and associated with the cacao swollen shoot disease. Numbers on the branches represent the SH-aLRT (approximate likelihood ratio test) branch supports over 0.7. The scale bar represents the substitution number per base. The GenBank accession numbers of Badnavirus sequences used are listed in Table 1 and the CSSV sequences are listed in Table 3. Rice tungro bacilliform virus (RTBV-Phi – X57924) was used as outgroup.

Diagnosis Serological Diagnostics The virus is not strongly immunogenic; however, several antisera have been raised and shown to react with CSSV. Enzyme-linked immunosorbent assay (ELISA) has been used to detect CSSV but high background values were obtained and difficulties were found in detecting CSSV in plants suspected of having only a low virus titer. Immunosorbent electron microscopy has been used for detection and comparison of some isolates of CSSV in Ghana. Both of these techniques cannot detect latent infection.

282

Cacao Swollen Shoot Virus (Caulimoviridae)

Table 3

List of CSSV isolates with complete genome sequences and their NCBI accession number

Cacao species in the genus Badnavirus

Abbreviation-Isolate

Accession #

Cacao swollen shoot CD virus Cacao swollen shoot CD virus – CI152-09 Cacao swollen shoot CD virus – CIDivo-15

CSSCDV-CI152-09 CSSCDV-CIDivo-15

JN606110 MF642718

Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao

swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen

shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot

CE CE CE CE CE CE CE CE CE CE

CSSCEV-GI632-10 CSSCEV-GWR198E-13 CSSCEV-GWR198J-13 CSSCEV-GWR3-14 CSSCEV-GCR329-14 CSSCEV-GH64 CSSCEV-GH67 CSSCEV-CI275 CSSCEV-CI286

MF642719 MF642720, MF642721 MF642722 MF642723 KX592572 KX592571 KX592573 KX592584

Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao

swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen

shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot

Ghana Ghana Ghana Ghana Ghana Ghana Ghana Ghana Ghana Ghana Ghana

CSSGMV-Gha57-15 CSSGMV-NIG1 CSSGMV-NIG10 CSSGMV-NIG12 CSSGMV-NIG13 CSSGMV-NIG16 CSSGMV-NIG18 CSSGMV-NIG5 CSSGMV-NIG7 CSSGMV-NIG9

MF642724 MH785297 MH785300 MH785301 MH785302 MH785303 MH029282 MH029281 MH785298 MH785299

Cacao swollen shoot Ghana N virus Cacao swollen shoot Ghana N virus - Gha63-15

CSSGNV-Gha63–15

MF642725

Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao

CSSGQV-Gha2-15 CSSGQV-Gha34Q-15 CSSGQV-Gha64-99 CSSGQV-Gha40-15 CSSGQV-Gha29-15 CSSGQV-Gha34R-15 CSSGQV-Gha37-15 CSSGQV-Gha3915 CSSGQV-Gha53-15 CSSGQV-Gha54-15

MF642726 MF642727 MF642729 MF642728 MF642730 MF642731 MF642732 MF642733 MF642734 MF642735

Cacao swollen shoot Togo A virus Cacao swollen shoot Togo A virus – Wobe12-02 Cacao swollen shoot Togo A virus – Gha25-15

CSSTAV-Wob12-02 CSSTAV-Gha25-15

AJ781003 MF642716

Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao Cacao

CSSTBV-Agou1-93 CSSTBV-NB2-00 CSSTBV-N1A-00 CSSTBV-Peki-00 CSSTBV-New Juaben-00 CSSTBV-CI303-09 CSSTBV-CI563-10 CSSTBV-CI569-10 CSSTBV-CI311 CSSTBV-CI134 CSSTBV-CI135 CSSTBV-CI214 CSSTBV-CI301 CSSTBV-CI44 CSSTBV-CIS2 CSSTBV-CIS3 CSSTBV-CIT5

L14546 AJ534983 AJ609020 AJ609019 AJ608931 MN179343 MN179344 MF642717 KX592580 KX592578 KX592581 KX592577 KX592579 KX592574 KX592583 KX592576 KX592582

swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen

swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen swollen

shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot

shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot shoot

virus virus virus virus virus virus virus virus virus virus

Ghana Ghana Ghana Ghana Ghana Ghana Ghana Ghana Ghana Ghana Ghana

Togo Togo Togo Togo Togo Togo Togo Togo Togo Togo Togo Togo Togo Togo Togo Togo Togo Togo

M M M M M M M M M M M

Q Q Q Q Q Q Q Q Q Q Q

B B B B B B B B B B B B B B B B B B

– – – – – – – – –

GI632-10 GWR198E-13 GWR198J-13 GWR3-14 GCR329-14 GH64 GH67 CI275 CI286

virus virus virus virus virus virus virus virus virus virus virus

virus virus virus virus virus virus virus virus virus virus virus

virus virus virus virus virus virus virus virus virus virus virus virus virus virus virus virus virus virus

– – – – – – – – – – – – – – – – –

– – – – – – – – – –

– – – – – – – – – –

Gha57-15 NIG1 NIG10 NIG12 NIG13 NIG16 NIG18 NIG5 NIG7 NIG9

Gha2-15 Gha34Q-15 Gha64-99 Gha40-15 Gha29-15 Gha34R-15 Gha37-15 Gha3915 Gha53-15 Gha54-15

Agou1-93 Nyongbo2-00 N1A-00 Peki-00 New Juaben-00 CI303-09 CI563-10 CI569-10 C1311 CI134 CI135 CI215 CI301 CI44 CIS2 CIS3 CIT5

Cacao Swollen Shoot Virus (Caulimoviridae)

Table 3

283

Continued

Cacao species in the genus Badnavirus

Abbreviation-Isolate

Accession #

Cacao swollen shoot Togo B virus – GH75

CSSTBV-Gh75

KX592575

Unassigned species Cacao swollen shoot Ghana T virus Cacao swollen shoot Ghana T virus – Gha4-15

CSSGTV-Gha4-15

MN179342

Geographical Distribution of Virus Species Causing CSSD

CRIG Museum Gontougo Marahoué Haut Sassandra Iffou

IndéniéJuablin

Brong Ahafo

Volta

Limé

Guémon Moronou Agneby Gôh Tiassa Mé

Nawa Gbôkle

Lôh Djiboua

Kloto

Ashan Eastern

Central Grands Sud Comoé Western Ponts

100 km

TA

TB

CD

CE

GM

GN

GQ

GT

Fig. 6 Geographical distribution of the species complex associated with cacao swollen shoot disease in the different outbreaks in West Africa, (Côte d0 Ivoire, Ghana and Togo).

The viro-bacterial agglutination (VBA) test has been found to be a useful test for detecting CSSV in leaf tissue from infected trees. Using this assay, CSSV can be detected in trees showing symptoms as well as in infected, but symptomless, trees. The immunocapture polymerase chain reaction (IC-PCR) technique was adapted to the only detection of CSSV-1A isolates from Ghana (CSSTBV species). There is considerable strain variation among the many recognized CSSV isolates, some of which react only weakly with certain antisera. Using monoclonal antibodies, four serotypes of CSSV have been distinguished by ELISA analysis of 31 samples of the virus from different geographical locations in Ghana. The efficiency of serological diagnosis depends on the use of polyvalent antiserum able to detect all serotypes. Since the 2000s and the first results on viral molecular variability, no work has been published on the development of serological methods for CSSV but these methods should not be excluded for the implementation of a versatile and sensitive diagnosis.

PCR Diagnosis For a reliable PCR diagnosis, it is necessary to design primers from conserved regions of the genome. Until 2000, only few badnaviruses were completely sequenced, and only the end of the ORF3, which contains the conserved motifs specific of RTase and RNaseH made it possible to obtain primers useful for diagnosis. After alignment of the first six full-length sequences of CSSV, polyvalent primers pairs were designed in two different parts of the genome, in the movement protein (MP, first part of ORF3) and in the RTase/RNaseH region (third part of ORF3) (Fig. 3)and were then improved to be able to detect new species that were subsequently identified. However, the presence in cacao genomes of retrotransposons containing also the motifs specific of RTase and RNaseH makes it difficult to develop specific CSSV primers in this area.

284

Cacao Swollen Shoot Virus (Caulimoviridae)

PCR-based diagnosis is able to detect CSSV not only in symptomless leaves of symptomatic plants, but also in symptomless plants as early as one week post-inoculation. As the sensitivity of diagnosis is better when young symptomatic host plant leaves are tested, it is recommended that validation of the diagnostic test is done using this type of material in the first instance, especially when testing isolates from new areas. Given the heterogeneous distribution of the virus in the plant, especially in adult trees, this PCR diagnosis should be used primarily to check for infection of plants with suspicious symptoms, or of grafts or rootstocks.

Control of the Disease Attempts at CSSV control in Ghana have required substantial financial and manpower inputs. The “cutting-out” policies in place in Ghana since the early 1940s which attempted to control cacao swollen shoot disease resulted in the removal of over 190 million trees up to 1988. Over ten million infected trees which still required ‘cutting out’ were identified in the field by 1990. Control of the swollen shoot disease based only on ‘cutting-out’ campaigns has not been successful due to several factors including political and socioeconomic problems. The better strategy for dealing with the disease is to develop a combination of control measures in an integrated approach. Moreover, this approach should implicate cacao farmers as much as possible. Mild strain protection is a possibility which is being investigated. Mild strains which appear to confer some protection against the severe strains from species CSSTBV are available and are being tested in the field in Ghana. However, only a closely related mild strain is able to protect against another strain, as has been demonstrated for the Citrus tristeza virus. Now that there are so many different CSSV species, it would seem that it would be impossible to protect the cacao trees against them all. The control of insect vectors such as mealybugs is difficult to implement due to the excessive cost of the molecules to be used and their toxicity which results to residues in the cacao beans produced by treated trees. The most effective replantation method is to clear whole areas and replant in large compact block because of the relatively small proportion of trees in the vulnerable peripheral areas. The possibility of isolating new cacao plantings from infected cacao by an isolation distance of 10 m or planting barriers with CSSV-immune crops should be considered. Barrier crops such as banana, plantain, oil palm, coffee, cola, and citrus can form a barrier to the movement of vectors. The ‘cutting out’ of the adventitious plants of the genus Commelina recognized as alternative hosts should also be done. The use of resistant cacao is advocated for replantation as many of the new hybrids available in West Africa do have some partial resistance to CSSV. Replanting with resistant cacao trees, however, requires the installation of a protocol of effective screening for resistance. Severe isolates representative of the different molecular groups should be used as well as a suitable screening method. However, a standardized inoculation method is not yet available because particle bombardment is difficult to develop on a large scale and agro-inoculation needs biosafety confinement. Mealybug transmission tests can nevertheless be set up by the national research institutes of the different producing countries, taking all precautions in terms of confinement (insect proof greenhouses). Intermediate quarantine facilities are at present hampered by the lack of suitable indexing methods for the virus particularly if the germplasm is to be moved as stem cuttings. New polyvalent PCR diagnosis could be easily tested in intermediate quarantine facilities and compared with the grafting procedure used for indexation on Amelonado cacao seedlings.

See also: Caulimoviruses (Caulimoviridae)

Further Reading Abrokwah, F., Dzahini-Obiatey, H., Galyuon, I., Osae-Awuku, F., Muller, E., 2016. Geographical distribution of Cacao swollen shoot virus molecular variability in Ghana. Plant Disease 100 (10), 2011–2017. Brunt, A.A., 1970. Cacao Swollen Shoot Virus. CMI/AAB Descriptions of Plant Viruses N110. Wellesbourne: Association of Applied Biologists. CABI – Crop Protection Compendium, Crop protection Compendium, 2002. Cacao Swollen Shoot Virus. Wallingford: CABI Publishing. Castel, C., Amefia, Y.K., Djiekpor, E.K., Partiot, M., Segbor, A., 1980. Le swollen shoot du cacaoyer au Togo. Les différentes formes de viroses et leurs conséquences économiques. Café, Cacao, Thé 24 (2), 131–146. Fauquet, C., Mayo, M., Maniloff, J., Desselberger, U., Ball, L. (Eds.), 2005. Virus Taxonomy: VIIIth Report of the International Committee on Taxonomy of Viruses. Elsevier Academic Press. Hagen, L.S., Jacquemond, M., Lepingle, A., Lot, H., Tepfer, M., 1993. Nucleotide sequence and genomic organization of Cacao swollen shoot virus. Virology 196 (2), 619–628. Hagen, L.S., Lot, H., Godon, C., Tepfer, M., Jacquemond, M., 1994. Infection of Theobroma cacao using cloned DNA of Cacao swollen shoot virus and particle bombardment. Molecular Plant Pathology 84, 1239–1243. Jacquot, E., Hagen, L.S., Michler, P., et al., 1999. In situ localization of Cacao swollen shoot virus in Theobroma cacao. Archives of Virology 144, 259–271. Lot, H., Djiekpor, E., Jacquemond, M., 1991. Characterization of the genome of Cacao swollen shoot virus. Journal of General Virology 72, 1735–1739. Muller, E., Sackey, S., 2005. Molecular variability analysis of five new complete Cacao swollen shoot virus genomic sequences. Archives of Virology 150, 53–66. Muller, E., 2016. Cacao swollen shoot virus (CSSV): History, biology, and genome. In: Bailey, Bryan A., Meinhardt, Lyndel W. (Eds.), Cacao Diseases. A History of Old Enemies and New Encounters. Cham: Springer International Publishing, pp. 337–358. Muller, E., Ravel, S., Agret, C., et al., 2018. Next generation sequencing elucidates cacao badnavirus diversity and reveals the existence of more than ten viral species. Virus Research 244, 235–251.

Carmo-Like Viruses (Tombusviridae) Miryam Pérez-Cañamás and Carmen Hernández, Institute for Plant Molecular and Cell Biology (Spanish National Research Council–Polytechnic University of Valencia), Valencia, Spain r 2021 Elsevier Ltd. All rights reserved. This is an updte of F. Qu, T.J. Morris, Carmovirus. In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00365-4.

Nomenclature aa Amino acid(s) AGO Argonaute CP Coat protein diRNAs Defective interfering RNAs dsRNA Double-stranded RNA eIF Eukaryotic translation initiation factor gRNA Genomic RNA ICTV International Committee for Taxonomy of Viruses kb Kilobase kDa Kilodalton MP Movement protein mRNA Messenger RNA

Glossary Defective interfering RNAs RNAs derived from the RNA genome of a virus in which a critical portion of such genome has been lost usually due to defective replication. These RNAs are associated to and depend on the parent virus for infection as it provides the lost factor. Leaky-scanning Phenomenon in which a weak initiation codon triplet on mRNA is sometimes skipped by ribosome in translation initiation. The 40S ribosomal subunit continues scanning to further initiation codon. The weak initiation codon can be an AUG in a weak Kozak consensus context or a non-AUG codon. Readthrough Consists of the “translation” of a stop codon, such that the synthesis of a larger proteins is allowed. It is a common phenomenon in viruses, and it has a regulatory function. RNA silencing A mechanism of transcriptional and post-transcriptional gene regulation operating in many

NCR Non-coding region nm Nanometer nt Nucleotide(s) ORF Open reading frame Poly(A) Polyadenylated RdRp RNA-dependent RNA polymerase satRNA Satellite sgRNA Sub-genomic RNA spp Species (plural) ssRNA Single-stranded RNA tRNA Transfer RNA VRC Viral replicase complex

eukaryotic organisms. It is mediated by 20–25 nt small RNAs produced from various RNA sources and functions in regulating RNA stability and translation as well as in chromatin modification underlying numerous developmental processes. It also plays a significant role in microbe–host interaction. Satellite RNA A subviral RNA that depends on the co-infection of a host cell with a helper virus for its replication. Silencing suppressor Virally encoded proteins from diverse families that interfere with host production or effectiveness of small RNAs involved in RNA silencing. Sub-genomic RNA Viral RNA that derives from and is usually 30 co-terminal to the viral genomic RNA and that is produced during infection to serve as mRNA for translation of internal and 30 -proximal genes. Tombusvirid Member of family Tombusviridae.

Introduction In 1988, a series of viruses forming polyhedral virions which enclosed monopartite single-stranded (ss) RNA genomes of B4 kb and that were represented by Carnation mottle virus (CarMV) were described. Since then, more than twenty-five related viruses sharing significant sequence homology and overall gene arrangement have been identified and are currently known under the generic name of carmo-like viruses. Investigations in the last years have significantly improved our knowledge on the biological and molecular properties of carmo-like viruses which, moreover, has helped to refine their classification. In this article, summary and updated information on this group of infectious agents is provided.

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Taxonomy, Classification, and Evolutionary Relationships The International Committee for Taxonomy of Viruses (ICTV) currently recognizes sixteen genera within the family Tombusviridae which, in turn, are clustered into three subfamilies: Calvusvirinae, Procedovirinae, and Regressovirinae. Four out of the fourteen genera included in Procedovirinae comprise carmo-like viruses: Alphacarmovirus, Betacarmovirus, Gammacarmovirus, and Pelarspovirus. These genera show differential branching in phylogenetic trees based on their encoded RNA-dependent RNA polymerases (RdRps) (Fig. 1). The generic name “carmo” is derived from the first member of the group to be described, CarMV, currently being the type species of the genus Alphacarmovirus. Members of each of the four genera share recognizable, yet variable, sequence similarity with members of other genera of carmo-like viruses as well as with other genera of the family Tombusviridae. As other components of the

Fig. 1 Phylogenetic tree of the RdRps of tombusvirids. The alignment of was made using MUSCLE while tree was generated with the Maximum Likelihood (ML) algorithm in MEGA7 (https://www.megasoftware.net/). The analysis involved 64 amino acid sequences (those corresponding to the RdRps of all recognized tombusvirids). Condensed triangles mark monophyletic lineages. Tombusviridae subfamily names are on the right, and members of the four carmo-like virus genera are colored in red (Alphacarmovirus), green (Pelarspovirus), blue (Betacarmovirus), and purple (Gammacarmovirus). Virus names and acronyms are as follows: Angelonia flower break virus (AnFBV), Calibrachoa mottle virus (CbMV), Cardamine chlorotic fleck virus (CCFV), Carnation mottle virus (CarMV), Clematis chlorotic mottle virus (ClCMV), Cowpea mottle virus (CPMoV), Cucumber Bulgarian latent virus (CBLV), Cymbidium ringspot virus (CyRSV), Elderberry latent virus (ELV), Furcraea necrotic streak virus (FNSV), Galinsoga mosaic virus (GaMV), Hibiscus chlorotic ringspot virus (HCRSV), Honeysuckle ringspot virus (HnRSV), Japanese iris necrotic ring virus (JINRV), Maize chlorotic mottle virus (MCMV), Maize necrotic streak virus (MNeSV), Melon necrotic spot virus (MNSV), Nootka lupine vein clearing virus (NLVCV), Oat chlorotic stunt virus (OCSV), Pea stem necrosis virus (PSNV), Pelargonium chlorotic ring pattern virus (PCRPV), Pelargonium flower break virus (PFBV), Pelargonium line pattern virus (PLPV), Pelargonium ringspot virus (PelRSV), Rosa rugosa leaf distortion virus (RrLDV), Saguaro cactus virus (SCV), Soybean yellow mottle mosaic virus (SYMMV), Trailing lespedeza virus 1 (TLV1), and Turnip crinkle virus (TCV). Hepatitis C virus (HCV) RdRp was used as outgroup.

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family, carmo-like viruses have icosahedral virions of about 30 nm in diameter, with T¼3 symmetry, that is composed of 180 coat protein (CP) subunits of about 37–42 kDa and a ssRNA genome ranging in size from 3.9 to 4.3 kb. In all cases, the genome consists of a unique positive ( þ ) sense ssRNA molecule that contains, at least, five definitive open reading frames (ORFs). Genome organizations of Alphacarmovirus, Betacarmovirus, and Gammacarmovirus are very similar and can be exemplified by those of their type species, corresponding to CarMV, Turnip crinkle virus (TCV), and Melon necrotic spot virus (MNSV), respectively (Fig. 2). TCV is probably the best studied virus among carmo-like viruses. It was the first carmo-like virus for which infectious transcripts were produced from a cDNA clone of the genome and detailed information about genome functions and virion configuration has been obtained, in the latter case including crystal structure determination. TCV genome harbors five ORFs encoding proteins of about 28, 88, 8, 9, and 38 kDa from the 50 - to the 30 end, respectively (Fig. 2). Leaving aside some specific features that will be mentioned in the next sections, the genome organization of pelarspoviruses resemble those of other carmo-like viruses and can be illustrated by the one from its type species, Pelargonium line pattern virus (PLPV). PLPV genome contains five genes encoding proteins of about 27, 87, 7, 9.7, and 37 kDa from the 50 - to the 30 end, respectively (Fig. 2). Some carmo-like viruses, such as Hibiscus chlorotic ringspot virus (HCRSV, genus Betacarmovirus), have additional ORFs whose functions are not entirely clear. To date, the nucleotide sequences of the 21 definitive carmo-like viruses have been determined (Table 1). These sequenced viruses share similar morphological and physicochemical properties with five additional viruses listed in Table 1, that are recognized as unassigned species of the family Tombusviridae by the ICTV. Moreover, genome sequences of four of these viruses have not yet been determined. Most carmo-like viruses are sufficiently distant from each other that they do not cross-react in standard RNA hybridization or serological tests. Carmo-like viruses have properties in common with viruses belonging to other genera of the family Tombusviridae. Their particle structure and CP sequences are closely related to tombus-, aureus-, diantho-, macana-, gallanti-, and avenaviruses. Their RdRp genes share a remarkable level of homology with other tombusvirids and particularly with those belonging to subfamily Procedovirinae: Machlomovirus, Panicovirus, Alphanecrovirus, Betanecrovirus, Aureusvirus, Gallantivirus, Macanavirus, Avenavirus, Zeavirus, and Tombusvirus. RdRp genes of carmo-like viruses and of other tombusvirids also share similarity with more distantly related viruses, such as luteoviruses. These latter viruses currently belong to family Luteoviridae though their reassignment to the family Tombusviriadae is expected in the near future. In a broader context, phylogenetic comparisons of viral RNA polymerase

Fig. 2 Genome organizations of species of genera Alphacarmovirus, Betacarmovirus, Gammacarmovirus, and Pelarspovirus exemplified, respectively, by those of their type species, Carnation mottle virus (CarMV), Turnip crinkle virus (TCV) and Melon necrotic spot virus (MNSV), and Pelargonium line pattern virus (PLPV). The colored boxes represent open reading frames with the sizes of the encoded proteins, in kilodaltons, indicated within the boxes. Blue, yellow and pink boxes have been used, respectively, for genes encoding proteins involved in replication, movement and encapsidation/suppression of RNA silencing, as indicated in the inset at the right. The two proteins required for replication are translated from the genomic RNA (gRNA) with the viral RNA-dependent-RNA polymerase (86–89 kDa in the Figure) being translated by readthrough (RT) of an amber codon at the end of the 50 -proximal gene. Thin lines under genomes indicate sub-genomic RNAs (sgRNAs) produced by members of each genera to translate internal and 30 proximal genes. Dashed line of PLPV MP2 marks a noncanonical start codon. The bottom panel at the right corresponds to Northern blot results for detection of viral RNAs in TCV- (left) and PLPV- (right) infected plants to illustrate the typical pattern of accumulation of genomic and sub-genomic RNAs in genera Alphacarmovirus, Betacarmovirus, and Gammacarmovirus, in the first case, and genus Pelarspovirus, in the second.

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Table 1

List of species in each genus composing the carmo-like viruses and unassigned species in the family Tombusviridae

Genus/Species

Virus acronym

Genbank accession

Alphacarmovirus Angelonia flower break virus Calibrachoa mottle virus Carnation mottle virus Honeysuckle ringspot virus Nootka lupine vein clearing virus Pelargonium flower break virus Saguaro cactus virus

AnFBV CbMV CarMV HnRSV NLVCV PFBV SCV

NC_007733.2 NC_021926.1 NC_001265.2 NC_014967.1 NC_009017.1 NC_005286.1 NC_001780.1

Betacarmovirus Cardamine chlorotic fleck virus Hibiscus chlorotic ringspot virus Japanese iris necrotic ring virus Turnip crinkle virus

CCFV HCRSV JINRV TCV

NC_001600.1 NC_003608.1 NC_002187.1 NC_003821.3

Gammacarmovirus Cowpea mottle virus Melon necrotic spot virus Pea stem necrosis virus Soybean yellow mottle mosaic virus

CPMoV MNSV PSNV SYMMV

NC_003535.1 NC_001504.1 NC_004995.1 NC_011643.1

Pelarspovirus Clematis chlorotic mottle virus Elderberry latent virus Pelargonium chlorotic ring pattern virus Pelargonium line pattern virus Pelargonium ringspot virus Rosa rugosa leaf distortion virus

ClCMV ELV PCRPV PLPV PelRSV RrLDV

NC_033777.1 NC_026239.1 NC_005985.1 NC_007017.2 NC_026240.1 NC_020415.1

Species not assigned to a genus Ahlum waterborne virus Bean mild mosaic virus Cucumber soil-borne virus Trailing lespedeza virus 1 Weddel waterborne virus

AWBM BMMV CSBV TLV1 WWBV

Unsequenced Unsequenced Unsequenced NC_015227.2 Unsequenced

genes have identified the tombusvirids as a representative virus cluster for one of three RNA virus super-groups with relatedness to animal viruses of the family Flaviviridae and small RNA phages (Leviviridae).

Distribution, Host Range, Transmission, and Economic Significance Carmo-like viruses are distributed worldwide and, with some exceptions, are reported to cause mild or asymptomatic infections in relatively restricted natural host ranges. Despite their usually moderate phenotypic effects, many of these viruses, although not all, accumulate to high concentrations in infected tissues. Notwithstanding their narrow natural host ranges, carmo-like viruses have, in general, broad experimental host ranges that frequently include herbaceous plant species amenable to experimental manipulation. A number of carmo-like viruses have been identified in ornamental hosts. CarMV (genus Alphacarmovirus) is one of the most noteworthy, because its high incidence and wide geographical spread in cultivated carnations. PLPV (genus Pelarspovirus) and Pelargonium flower break virus (PFBV, genus Alphacarmovirus) are other examples of highly widespread viruses, in this latter case in Pelargonium spp., causing diseases either alone or in association with other viruses. Information on the prevalence of other carmo-like viruses affecting ornamentals, such as Angelonia flower break virus and Calibrachoa mottle virus (CMoV) (both in the genus Alphacarmovirus), HCRSV and Japanese iris necrotic ring virus (JINRV) (both in the genus Betacarmovirus), Pelargonium chlorotic ring pattern virus, Pelargonium ringspot virus, and Rosa rugosa leaf distortion virus (RrLDV) (the three of them in the genus Pelarspovirus), is more scarce. However, such prevalence is likely to be highly influenced by the distribution of the corresponding infected nursery stocks as most ornamental plants are vegetatively propagated. Illustrating the harmful effects that infection by these viruses may cause, the severe disease symptoms induced by HCRSV on ornamental hibiscus varieties has led to a massive removal of this type of plants from many recreational areas in Singapore. Interestingly, HCRSV also infects kenaf (Hibiscus cannabinus L.), an annual crop of major interest to the wood-pulp industry. Legume species can also be naturally infected by several carmo-like viruses, such as Nootka lupine vein clearing virus (NLVCV, genus Alphacarmovirus), or Cowpea mottle virus (CPMoV), Pea stem necrosis virus (PSNV), and Soybean yellow mottle mosaic virus (SYMMV),

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all three belonging to genus Betacarmovirus. The geographical distribution of each of these viruses does not seem very broad so far, with CPMoV being potentially the most important since it may seriously limit the cultivation of cowpeas that are important for food supply throughout Sub-Saharan Africa and particularly in Nigeria. On the opposite side, MNSV is considered to be an endemic virus in greenhouses and open-field systems for the production of cucurbits, significantly reducing yields and causing vast economical damage worldwide. Two carmo-like viruses, TCV and Cardamine chlorotic fleck virus (CCFV) (genus Betacarmovirus), naturally infect Brassicaceae species. TCV is neither common, nor widespread in nature, whereas the known distribution of CCFV is limited to the Mount Kosiusko alpine region of Australia. Both are able to experimentally infect Arabidopsis thaliana, the most relevant model plant for basic plant research, in which they accumulate to extremely high concentrations, often approaching a level equivalent to 0.5% of the fresh weight of the plant tissue. Honeysuckle ringspot virus (HnRSV) and Saguaro cactus virus (SCV) (genus Alphacarmovirus), are presumably of little agricultural concern, and have been isolated from very particular hosts and areas, honeysuckle and San Luis Obispo (California) in the first case, and giant saguaro cactus from Sonoran Desert (Arizona), in the second. The infrequent isolation of these genetically similar viruses in remote locations around the world has prompted the speculation that ancestor carmo-like viruses may have been introduced into their natural hosts well before the last Ice Age and have since co-evolved in isolation in their diverse host plants. Spread of carmo-like viruses may occur through infected soil, irrigation water, vegetative propagation, grafting and/or mechanical inoculation. In addition, transmission by biological vectors has been reported in some instances. Thus, TCV and CPMoV can be transmitted by beetles, PFBV by thrips and MNSV and PSNV by the root-inhabiting fungus Olpidium bornovanus. Seed transmission of carmo-like viruses is not well documented but it could also be an important factor for the distribution of, at least, MNSV and CPMoV.

Virion Structure and Assembly The molecular structure of five carmo-like viruses (TCV, CarMV, CPMoV, HCRSV, and MNSV) has now been studied in detail, with TCV being the first studied by high-resolution X-ray crystallography. The detailed information about CP structure and inter-subunit interactions established that TCV and Tomato bushy stunt virus (TBSV, genus Tombusvirus) show remarkable structural conservation. In this regard, the common structural features shared by other members of the family Tombusviridae have been primarily deduced from alignment of the amino acid sequence of their CPs with those of TBSV and TCV. TCV consists of a T¼3 icosahedral capsid of 180 subunits of the 38 kDa CP. The individual CP subunit folds into three distinct domains typical of CP subunit of tombusvirids. The relatively basic N-terminal R domain extends into the interior of the virus particle and presumably interacts with viral RNA. The R domain is connected by an arm to the S domain which constitutes the virion shell. The S domain is attached through a hinge to the P domain which projects outward from the virion surface. The resulting surface protrusions give the virions a bumpy appearance in electron micrographs. TCV CP subunits are believed to form dimers in solution and during assembly. TCV is the only carmovirus for which detailed in vitro assembly studies have been performed. The virion has been shown to dissociate at elevated pH and ionic strength to produce a stable RNA–CP complex (rp-complex) and free CP subunits. Reassembly under physiological conditions in solution could be demonstrated using the isolated rp-complex and the soluble CP subunits. This rp-complex, consisting of six CP subunits tightly attached to viral RNA, could be generated in vitro and was shown to be important in selective assembly of TCV RNA. A model for assembly was proposed in which three sets of dimeric CP interact with a unique site on the viral RNA to form an initiation complex to which additional subunit dimers could rapidly bind. Preliminary characterization of the ‘origin of assembly’ for this virus identified two possible sites based on the identification of RNA fragments in the rp-complex protected from RNase digestion by CP. Further in vivo studies narrowed the assembly origin site to a bulged hairpin-loop of 28 nt within a 180 nt region at the 30 end of the CP gene.

Genome Structure and Protein Functions As mentioned above, complete nucleotide sequences of all virus species currently included in the genera Alphacarmovirus (7), Betacarmovirus (4), Gammacarmovirus (4), and Pelarspovirus (6) have been determined (Table 1). Comparative studies of the deduced ORFs revealed that all of these viruses encode a similar set of genes that are closely related and are present in the same gene order as illustrated in Fig. 2. Such set of genes is preceded by a 50 non-coding region (NCR), whose length ranges between 5 nt (PLPV) to 134 nt (PSNV), and is followed by a 30 NCR that varies from 224 nt (SCV) to 405 nt (NLVCV) in length. Distinct viral species do not share extensive sequence homology within these regions and none of them possesses neither a poly(A) tail nor a tRNA-like structure, in their 30 end. In addition, the 50 end of the genomes is not capped. The genome organization of the carmolike viruses is quite compact and most of the identified ORFs are overlapping each other. As shown for one or more members of each genus, the product of the most 50 proximal ORF (25–29 kDa) and its readthrough product (83–89 kDa, the RdRp) are essential for replication of the virus genome. In addition, all carmo-like viruses characteristically present two small ORFs in the middle of the genome that encode two proteins (6–13 KDa) involved in cell-to-cell movement (movement proteins or MPs). Both MPs have been proposed to function in a coordinate fashion, one recruiting the viral RNA and the other exploiting the plant

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endomembrane system to promote intra- and inter-cellular movement of carmo-like viruses. The 30 proximal gene encodes the viral CP which varies from 37 to 42 kDa for the different viruses. Interestingly, as proven for several carmo-like viruses, including CarMV, MNSV, PFBV, PLPV, and TCV, this protein also acts as suppressor of RNA silencing, a potent antiviral mechanism in plants that is conserved in nearly all eukaryotic organisms. This function is performed by other viral products in other genera of the family Tombusviridae, underlining the uniqueness of the structural subunit of carmo-like viruses. Although the genome organizations of all sequenced carmo-like viruses are very similar, those of pelarspoviruses are distinguished by some specific features such as the presence of a non-AUG start codon, instead of the canonical AUG, initiating the MP2 ORF and the lack of an AUG codon in any frame between the AUG initiation codon of MP1 and the CP ORFs. In addition, some individual carmo-like viruses display specific characteristics. For example, the two small central ORFs in MNSV (p7a and p7b) are connected by an in-frame amber codon that could result in the production of a 14 kDa fusion protein of the two ORFs by a readthrough mechanism. Similarly, a potential double readthrough could lead to translation of a C-terminal extended RdRp in the case of CarMV, HnRSV, PFBV, and JINRSV. On their side, genomes of CPMoV, HCRSV, SYMMV, and RrLDV contain a sixth ORF nested within the 30 proximal CP gene. Three-four additional small ORFs have also been predicted in RrLDV genome. Nevertheless, the biological functionality of most of these extra ORFs remains to be ascertained. Indeed, unequivocal proof of activity has only been provided for the novel ORFs that are nested within the RdRp and the CP gene, respectively, of HCRSV.

Replication and Gene Expression Some carmo-like viruses, such as TCV, replicate to very high concentrations in protoplast infections, with the genomic (g) RNA accumulating to levels approaching that of the ribosomal RNAs. This facilitates identification and characterization of both cis-acting RNA elements and viral products involved in viral multiplication. For expression of genes located in internal and 30 -proximal positions of the gRNA, carmo-like viruses transcribe either one (genera Alphacarmovirus, Betacarmovirus, and Gammacarmovirus) or two (genus Pelarspovirus) 30 -coterminal sub-genomic RNAs (sgRNAs). In the first case, the larger sgRNA (B1.7 kb) functions as the messenger (m) RNA for the two MP genes utilizing a leaky- scanning mechanism whereas the smaller sgRNA (B1.5 kb) is the mRNA for the CP gene. When only one sgRNA (B1.7 kb) is produced, it serves as mRNA for translation of the two MP genes as well as the CP gene. Such translation is accomplished through leaky-scanning processes that are favored by the suboptimal translational context of the initiating AUG of the MP1 gene, the non-canonical start codon of MP2 gene (Fig. 2), and the lack of an AUG codon between the MP1 AUG and CP AUG. Both models for gene expression are supported by in vitro translation experiments performed with, at least, TCV, SCV, and PLPV. The number of sgRNAs (one or two) together with the monophyletic lineage of the encoded RdRps (Fig. 1) are the main criteria that define the distinct genera of carmo-like viruses. As other viruses with RNA genome, carmo-like viruses are thought to replicate through a negative (–) strand intermediate because virus-specific double-stranded (ds) RNAs corresponding in size to the gRNA characteristically accumulate in infected plant tissue. As shown for TCV, the protein encoded by the 50 proximal ORF and its readthrough product are the only virus-encoded components of the polymerase complex. When expressed from two separate mRNAs, both viral products complemented in trans to enable the genome replication. Besides interactions between viral products, the assembly of viral replicase complexes (VRCs) involves intricate interactions between viral and host factors, including cellular membranes that serve as platforms for the formation of VRCs. Associations of the VCRs of PFBV and MNSV with mitochondrial membranes have been described. The VRCs also catalyze synthesis of sgRNAs most likely through a premature termination mechanism, as suggested from results with TCV and PLPV. This mechanism involves generation of a sg (–) strand on a genomic ( þ ) template because of the presence of a structural motif in the latter that causes the RdRp to pause before it reaches the template 50 end. The sg (–) molecules serve as templates for generation of sg ( þ ) RNAs. A membrane-containing extract prepared from evacuolated protoplasts of uninfected Arabidopsis plants has been shown to faithfully produce both genomic and sub-genomic RNAs from a full-length TCV RNA template. Such an extract may be useful for future characterization of viral as well as host elements required for virus replication and transcription. Numerous studies have demonstrated that, besides the virus-encoded proteins, a plethora of structural elements in the viral RNA play critical roles in the genome replication of carmo-like viruses. These elements are located throughout the entire viral genome and occur in both the ( þ ) and (–) strands. Their roles range from promoters, enhancers or repressors for RNA replication and transcription, enhancers for translation, signals for readthrough translation and specificity determinants for virus assembly. The current knowledge of these RNA structures suggests that different secondary, or even tertiary, structural motifs, some of them mutually exclusive, are formed at the different stages of virus multiplication, and the highly coordinated nature of their formation ensures optimal utilization of the compact viral genome.

Satellites, Defective-Interfering RNAs TCV is the only carmo-like virus in which replication of associated small subviral RNAs in infected plants has been characterized, and the situation for this virus is curiously complex. TCV infections are associated with defective interfering (DI) RNAs derived totally from the parent genome (e.g., RNA G of 342–346 nt), satellite (sat) RNAs of non-viral origin (e.g., RNAs D, 194 nt and F, 230 nt), and chimeric RNAs (e.g., RNA C of 356 nt) with a 50 region derived from sat RNA D and a 30 region derived from the 30 end of the TCV genome. All three types of small RNAs depend on the helper virus for their replication and encapsidation within

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the infected plant. The different satellites and DI RNAs have been shown to affect viral infections in different ways. Both RNAs C and G intensify viral symptoms while interfering with the replication of the helper virus, while RNAs D and F seem to produce no detectable effects on either expression of symptoms or helper virus replication.

Virus–Host Interaction Considerable progress has been made in the last two decades in our understanding of the molecular mechanisms of virus–plant interactions with significant contributions coming from studies utilizing TCV and its host plant Arabidopsis. Most notably, one ecotype of Arabidopsis thaliana has been determined to be resistant to TCV infection and the resistance gene (HRT) and its encoded protein have been characterized. Additional host factors that contribute to the resistance response have also been elucidated including the identification of a novel transcription factor (TIP) whose interaction with TCV CP is critical to prevent basal resistance. The determination that TCV CP is targeted by HRT resistance protein suggests that TCV CP plays a key role in combating antiviral defense mechanisms of the plant host, in addition to being required for virus assembly. Indeed, as mentioned above, distinct studies have established that TCV CP is a strong suppressor of RNA silencing, a host defense mechanism that targets invading RNA. TCV CP seems to develop this suppressor function through different mechanisms, one of which consists in interacting with Argonaute (AGO) proteins, the key effectors of RNA silencing. In addition, it has been reported that TCV CP requires RAV2, an ethylene inducible factor, for suppression of RNA silencing. The role of RNA silencing in the fight against TCV has been also evidenced by the high accumulation of suppressor-deficient TCV mutants in A. thaliana plants lacking either AGO1 or one or more of the four Dice-like enzymes (dsRNA-specific endoribonucleases that play an essential role in triggering RNA silencing). Besides host factors involved in the resistance response against TCV, a host factor that most likely assists the virus during its life cycle has been identified. This factor corresponds to eukaryotic translation initiation factor (eIF) 4G of Arabidopsis that has been found to augment TCV accumulation, presumably by promoting more efficient translation of viral genes. In case of MNSV, genetic and biochemical evidence showing that translation is eIF4E-dependent in melon has been obtained. Similarly to that found for the CP of TCV, the CP of PLPV, which also acts as RNA silencing suppressor, interacts with AGO proteins. Interestingly, this PLPV viral product has been reported to interact as well with distinct importins alpha that are crucial nuclear-transport receptors. Such interaction seems to favor the accumulation of PLPV although the precise underlying molecular mechanism remains to be known. Other relevant aspect of virus-host interactions refers to symptom elicitation. This question has been particularly tackled with TCV and its satellite RNA C. It has been shown that TCV CP is important in satellite RNA C interactions in the host plant. Normally, the presence of RNA C results in symptom intensification in TCV infections. However, when the TCV CP ORF is either deleted or replaced by the CCFV CP ORF, RNA C attenuates symptoms caused by the helper virus suggesting that CP either downregulates the replication of RNA C or enhances its own competitiveness. Finally, the replicase gene of TCV has also been implicated in the symptom modification by satellite RNA C by two independent groups. The 30 end of the TCV genome, a sequence common in TCV RNA, RNA C, and DI RNA G, was also suggested to be a symptom determinant. Recent results also suggest that viral pathogenesis could be related with virus-induced alterations in the levels of long noncoding RNAs, which are involved in diverse and substantial biological processes, and/or of microRNAs, small RNAs that act as developmental regulators. In this context, floral structure disorders typically observed in TCV-infected A. thaliana plants have been correlated with upregulation of a long noncoding RNAs which, in turn, seems to lead to downregulation of a floral structurerelated APETALA2 gene. However, changes in levels of regulatory RNAs induced by viral infections do not always correlate with phenotypic effects as shown with PLPV-infected Nicotiana benthamiana plants which, despite being asymptomatic, exhibited modifications in microRNA contents. These observations underline the complexity and the multiplicity of factors that may be involved in viral pathogenesis.

Further Reading Blanco-Pérez, M., Hernández, C., 2016. Evidence supporting a premature termination mechanism for sub-genomic RNA transcription in Pelargonium line pattern virus: Identification of a critical long-range RNA-RNA interaction and functional variants through mutagenesis. Journal of General Virology 97, 1469–1480. Castaño, A., Ruiz, L., Hernández, C., 2009. Insights into the translational regulation of biologically active open reading frames of Pelargonium line pattern virus. Virology 386, 417–426. Csorba, T., Kontra, L., Burgyán, J., 2015. Viral silencing suppressors: Tools forged to fine-tune host-pathogen coexistence. Virology 479–480, 85–103. Hacker, D.L., Petty, I.T.D., Wei, N., Morris, T.J., 1992. Turnip crinkle virus genes required for RNA replication and virus movement. Virology 186, 1–8. Kachroo, P., Yoshioka, K., Shah, J., Dooner, H.K., Klessig, D.F., 2000. Resistance to Turnip crinkle virus in Arabidopsis is regulated by two host genes and is salicylic acid dependent but NPR1, ethylene, and jasmonate independent. Plant Cell 12, 677–690. Komoda, K., Naito, S., Ishikawa, M., 2004. Replication of plant RNA virus genomes in a cell-free extract of evacuolated plant protoplasts. Proceedings of the National Academy of Sciences of the United States of America 101, 1863–1867. Lommel, S.A., Martelli, G.P., Rubino, L., Russo, M., 2005. Tombusviridae. In: Fauquet, C.M., Mayo, M.A., Maniloff, J., Desselberger, U., Ball, L.A. (Eds.), Virus Taxonomy: Eighth Report of the International Committee on Taxonomy of Viruses. San Diego, CA: Elsevier Academic Press, pp. 907–936.

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Koenig, R., 1988. Carnation mottle virus and viruses with similar properties. In: Morris, T.J., Carrington, J.C. (Eds.), The Plant Viruses: Polyhedral Virions With Monopartite RNA Genomes 3. New York: Plenum, pp. 73–112. Nagy, P.D., Pogany, J., Simon, A.E., 1999. RNA elements required for RNA recombination function as replication enhancers in vitro and in vivo in a plus-strand RNA virus. The EMBO Journal 18, 5653–5665. Navarro, J.A., Pallás, V., 2017. An update on the intracellular and intercellular trafficking of carmoviruses. Frontiers in Plant Science 8, 1801. Pérez-Cañamás, M., Blanco-Pérez, M., Forment, J., Hernández, C., 2017. Nicotiana benthamiana plants asymptomatically infected by Pelargonium line pattern virus show unusually high accumulation of viral small RNAs that is neither associated with DCL induction nor RDR6 activity. Virology 501, 136–146. Pérez-Cañamás, M., Hernández, C., 2015. Key importance of small RNA binding for the activity of a glycine-tryptophan (GW) motif-containing viral suppressor of RNA silencing. Journal of Biological Chemistry 290, 3106–3120. Ren, T., Qu, F., Morris, T.J., 2000. HRT gene function requires interaction between a NAC protein and viral capsid protein to confer resistance to Turnip crinkle virus. The Plant Cell 12, 1917–1925. Russo, M., Burgyan, J., Martelli, G.P., 1994. Molecular biology of Tombusviridae. Advances in Virus Research 44, 381–428. Truniger, V., Miras, M., Aranda, M.A., 2017. Structural and functional diversity of plant virus 30 -cap-independent translation enhancers (30 -CITEs). Frontiers in Plant Science 8, 2047.

Relevant Websites http://www.rcsb.org/structure/3ZX8 3ZX8: Cryo-EM reconstruction of native and expanded Turnip Crinkle virus RCSB PDB. https://www.ncbi.nlm.nih.gov/genomes/GenomesGroup.cgi?taxid=2560077 Complete genomes: Procedovirinae NCBI. http://www.els.net/WileyCDA/ElsArticle/refId-a0022338.html eLS. http://www.els.net/WileyCDA/ElsArticle/refId-a0000756.html eLS. https://viralzone.expasy.org/53?outline=all_by_species Viralzone.

Cassava Brown Streak Viruses (Potyviridae) Basavaprabhu L Patil, ICAR–Indian Institute of Horticultural Research, Bengaluru, India r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) amiRNAs Artificial microRNAs Cas9 CRISPR-associated nuclease 9 CBSD Cassava brown streak disease CMGs Cassava mosaic geminiviruses CRISPR Clustered regularly interspaced short palindromic repeats eIF4E Eukaryotic translation initiation factor 4E HAM1h Homologue of HAM1

Glossary Artificial microRNAs They are similar to micro RNAs, that are single-stranded, 7 21 nt long, and designed by replacing the mature miRNA sequences of duplex within precursor-miRNAs. CRISPR-associated nuclease 9 A genome editing technology. Eukaryotic translation initiation factor 4E An eukaryotic translation initiation factor involved in directing ribosomes to the cap structure of mRNAs. Gene silencing A method to regulate gene expression to prevent the expression of a targeted gene. Genome editing A method to edit the gene employing different editing technologies. HAM1-like sequence A gene that protects against the mutagenic effects through detoxification of abnormal pyrimidine as well as purine nucleotides. Helper component proteinase A multifunctional protein encoded by viruses of the genus Potyvirus and other viruses of the family Potyviridae, involved in aphid transmission, polyprotein processing, and suppression of host antiviral RNA silencing. microRNA A type of small non-coding RNA of 721 nt in size involved in gene regulation and gene silencing. Next generation sequencing A ultra-high throughput massively parallel sequencing technology.

HCPro Helper component proteinase kDa Kilo dalton NGS Next generation sequencing Nia Nuclear inclusion protein a Nib Nuclear inclusion protein b nt Nucleotide(s) NTP Nucleoside triphosphate RNAi RNA-interference siRNAs Small interfering RNAs VPg Viral protein-genome linked

Nuclear inclusion protein a It is one of the proteases encoded by potyviruses. Nuclear inclusion protein b It is a RNA-dependent RNA polymerase encoded by potyviruses. Phytosanitation It is one of the plant disease management strategy that involves removal and destruction of diseased plant parts to prevent further spread of the pathogen. RNA silencing Sequence-specific regulation of gene expression triggered by double-stranded RNA, also termed RNA-interference (RNAi). Silencing suppression Gene silencing suppression mechanism by certain specialized proteins encoded by plant viruses. Small interfering RNAs A class of double-stranded non-coding RNA molecules, 20–25 base pairs in length, operating within the RNA interference pathway. Transgenic plants Transgenic or genetically modified plants have genes that are derived from another species inserted into their genome through recombinant DNA technology. Viral protein-genome linked A potyviral multifunction protein covalently linked to 50 end of viral RNA, which interacts with the eukaryotic translation initiation factor eIF4E, a cap-binding protein, which is imminent for virus infection.

Introduction There are at least 20 different viruses that infect the tropical plant cassava (Manihot esculenta Crantz), of which the cassava infecting begomoviruses and the brown streak associated ipomoviruses are the economically most important viruses resulting in monetary loss exceeding US$ 1 billion per year. Here in this article we intend to describe and discuss about the ipomoviruses associated with the cassava brown streak disease (CBSD) in Africa. Although the first report of CBSD was in 1930s from the coastal region of Tanzania, it did not receive much attention since it was restricted to the coastal lowlands of east Africa and appeared sporadically. However, post 2004, CBSD has been spreading to several parts of Africa putting the food security of millions of Africans at stake.

Aetiology, Host Range and Transmission CBSD is known to affect cassava alone, while it has also shown its presence in the wild relative of cassava Manihot glaziovii. CBSD does not have any known alternate hosts and it is not known to infect weed species. However, under artificial inoculation

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Fig. 1 Symptoms of CBSD on Root and Leaves of cassava. (a). Brown streaks on the young stem, (b) Different types of feathery chlorosis on leaves, (c) Constrictions on the root surface, (d) Necrosis in the roots. Adapted with minor modification from Patil, B.L., Legg, J.P., Kanju, E., Fauquet, C.M., 2015. Cassava brown streak disease: A threat to food security in Africa. Journal of General Virology 96, 956–968.

conditions, it can infect some of the herbaceous hosts and the model host Nicotiana benthamiana, widely used in virology studies. Two diverse and distinct ipomovirus species Ugandan cassava brown streak virus (UCBSV) and Cassava brown streak virus (CBSV) are known to cause CBSD, either as independent infection or sometimes as mixed infections. In this article both UCBSV and CBSV isolates are sometimes collectively referred as CBSVs. The most common symptoms on the foliage is feathery chlorosis along the veins of the oldest leaves, occasionally with circular patches of chlorosis in between the primary veins (Fig. 1(b)), while the stem shows characteristic brown necrotic streaks and stem die-back in severe cases (Fig. 1(a)). Young leaves do not show symptoms. However, CBSVs cause major damage in the tuberous roots, the edible part of cassava. CBSD affected tuberous roots show brown corky necrosis in the starchy tissue, resulting in reduction of starch and cyanide content, occasionally the tuberous roots develop radial constrictions (Fig. 1(c)). However, the symptoms can vary depending on the virus isolate involved, cassava genotype and its age, and the prevailing environmental conditions. In one of the studies involving mechanical inoculation of diverse CBSV/UCBSV isolates showed production of variable symptoms in both the natural host cassava and the model host N. benthamiana. Comparison of symptom severity of CBSV and UCBSV indicated CBSV to be more virulent, causing necrosis in N. benthamiana, while sap inoculation of UCBSV induced mosaics and rugosity. Similar comparative studies in cassava by grafting of infected

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scions showed CBSV to be more virulent than UCBSV, and the CBSV infected stem cuttings significantly reduced the cassava sprouting, as a result of higher virus accumulation in case of CBSV infection compared to infection by UCBSV. Agro-infiltration of infectious clones of cassava mosaic geminiviruses (CMGs) followed by sap inoculation of UCBSV indicated a synergistic interaction between the geminivirus and ipomoviruses in the case of N. benthamiana, however there are no reports of synergism in field situation. Initially, Bemisia afer (Priesner-Hosny) was presumed to be the vector for CBSD transmission, since they were predominantly present on lower leaves of cassava, where the CBSD symptoms are prominent. Later, experiments showed that the whitefly (Bemicia tabaci (Gennadius)) was the vector transmitting CBSVs. It was capable of transmitting this ipomovirus in a semi-persistent manner unlike persistent transmission of the geminiviruses. The transmission studies done in the lab conditions showed that the CBSVs were acquired in 5–10 min and retained by the whiteflies for less than 48 h. A maximum transmission of 60% could be achieved by using 20–25 viruliferous whiteflies per plant and the virus spread in the field was maximum up to a distance of 17 meters per cropping season. This study clearly indicates that the whitefly vectors could transmit CBSVs semi-persistently and only for a short distance, while the long-distance transmission of CBSD was mostly through transportation and use of infected planting material. Effective virus diagnostics is of utmost importance for accurate disease forecasting and management of any disease outbreaks. Early and accurate detection of CBSVs in infected cassava is a great challenge. There are several molecular methods for diagnostics of CBSD, of which ELISA and its modified versions are common. Using ELISA based techniques it is difficult to differentiate between CBSV and UCBSV. However, PCR based techniques such as RT-PCR, multiplex RT-PCR, Real Time PCR and loopmediated isothermal amplification are developed to detect and differentiate CBSVs and UCBSVs. Sequences of several unknown viruses including CBSVs have been unraveled by next generation sequencing (NGS) strategies.

Current Status of CBSD in Africa The first report of CBSD was from cassava cultivated in the Usambara mountains of Tanzania. Subsequent surveys showed that CBSD was prevalent in almost the entire coastal lowlands of East Africa, stretching from north-eastern border of Kenya, to the Tanzanian border with Mozambique, and lower altitudes of Malawi. CBSD incidence was not recorded in inlands above an altitude of 1000 m above sea level (a.s.l) and hence for a long time CBSD was considered as a low altitude disease. However, subsequent studies in 2004 have shown that CBSD was more widespread and also present at altitudes beyond 1000 m a.s.l., such as Uganda, western Kenya, north-western Tanzania, Burundi and the Democratic Republic of Congo. A countrywide systematic survey for incidence of CBSD in Tanzania was done in 1994 and the maximum incidence was recorded from southern lowland coastal districts of Mtwara (36%) and Masasi (25%), while CBSD was completely absent in higher altitudes of north-western Tanzania. Contrastingly, at the same time CBSD symptoms were reported in cassava cultivated at altitudes 1200 m above sea level, in a location near Entebbe in central/southern Uganda and also from Tabora in north-western Tanzania. Except for these reports of occurrence of CBSD at higher altitudes the notion that CBSD is a low altitude disease remained unchanged until the time a major report on occurrence of CBSD was made from central/southern Uganda in 2004. An unpublished estimate by NARO (Uganda) put the overall incidence of CBSD at 16% in 2008 and 29% in 2009, clearly depicting significant spread of CBSD in East Africa. Similar such reports on spread of CBSD were made in western Kenya and Lake Victoria zone of Tanzania. Surveys for CBSD were done in 19 districts of Tanzania and revealed a drastic increase in CBSD incidence in north-western regions of Tanzania. In recent years, additional reports have been made from Rwanda, Burundi and the eastern Democratic Republic of Congo (DRC). In early 2000, symptoms similar to CBSD have also been reported from cassava cultivated in Bas Congo Province in western DRC, Mulanje Province in central Angola and parts of Madagascar. However, none of these reports has been confirmed for the presence of CBSV by using appropriate diagnostic tools. All of these surveys were primarily based on visual assessments of leaf symptoms, and they may be underestimates of the true level of CBSV infection, since under the unfavourable weather conditions the CBSD leaf symptoms may not be expressed.

Genome Organization and Gene Functions The viruses involved in CBSD belong to the family Potyviridae and the genus Ipomovirus. The family Potyviridae is one of the largest families of plant viruses, consisting of six genera, classified based on their genome organization, sequence similarity and insect vectors involved in their transmission. Infected tissues with ipomoviruses show characteristic pinwheel-like structures or cylindrical inclusions within the phloem tissue, typical of potyviruses. Similar to other viruses of Potyviridae family, both the ipomoviruses CBSV and UCBSV have a positive-sense single stranded RNA genome with an average length of 9 Kb. All the viruses of the Potyviridae family except for the members of the genus Bymovirus have monopartite genome, which is translated into a single polyprotein which is later proteolytically autocleaved into B10 mature proteins by three distinct proteases expressed by the virus itself. Since the first report of CBSD in 1935, the first complete genome sequence of an ipomovirus involved in CBSD was published in 2009. This isolate from Tanzania sampled in 2007 was an Ugandan cassava brown streak virus isolate abbreviated as UCBSV[TZ:MLB3:07] with a sequence length of 9069 nt, which was shorter than other ipomoviruses. This UCBSV genome has 134 nt and 226 nt untranslated regions (UTRs) on both of its ends referred as 50 UTR and 30 UTR respectively (Fig. 2(b)). These UTRs

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Fig. 2 (a) Depiction of endemic (red-striped) and epidemic zones (blue-striped) of CBSD and the distribution of isolates belonging to CBSV (pink circles) and UCBSV (blue circles) in East Africa, (b) A representative picture showing the genome organization of CBSV/UCBSV encoding for 10 proteins (P1, first serine proteinase; P3, third protein; 6K1 and 6K2, two 6 kDa proteins; CI, cylindrical inclusion protein; VPg, viral protein genome-linked; NIa-Pro, main viral proteinase; NIb, replicase; Ham1, Maf/HAM1 pyrophosphatase homologue; CP, coat protein) and 30 and 50 UTRs, the poly(A) tail and PIPO translated by þ 2 frame shift within the P3. Two amino acids at the proteolytic cleavage sites of the polyprotein are shown below the polyprotein, and the estimated molecular mass (in kDa) for each protein is given in the box, (c, d) Phylogenetic analysis of selected CP sequences and the complete genome sequences of CBSV/UCBSV isolates. GenBank accession numbers are indicated and the scale bar represents the pairwise distance expressed as percentage dissimilarity. Adapted from Patil, B.L., Legg, J.P., Kanju, E., Fauquet, C.M., 2015. Cassava brown streak disease: A threat to food security in Africa. Journal of General Virology 96, 956–968.

encompass the regulatory sequences involved in viral gene/protein expression. Later a second distinct ipomovirus species named as Cassava brown streak virus (CBSV), with a sequence length of B9008 nt, with similar genome organization as UCBSV, was also found to be involved in CBSD. Unlike other members of the Potyviridae family, both the above ipomoviruses infecting cassava encode for a single P1 serine proteinase and are devoid of helper component cysteine proteinase (HCPro) (Fig. 2(b)). Such a high level of diversity in the P1 gene is characteristic of viruses of the Potyviridae family. The P1 of CBSVs are most similar those of another ipomovirus species called Sweet potato mild mottle virus (SPMMV). However, in contrast to CBSVs, the SPMMVs possess the HCPro, indicating it to be an evolutionary link between ipomoviruses and potyviruses. In contrast to both CBSVs and SPMMVs, another two ipomovirus species, namely Cucumber vein yellowing virus (CVYV) and Squash vein yellowing virus (SqVYV) possess two P1 serine proteinases (P1a and P1b). Silencing suppression studies done by individually expressing the CBSV genes has demonstrated P1 to be a silencing suppressor for CBSVs. The P1 of CBSVs encompass the basic LxKA and Zn-finger motifs, and the LxKA motif of P1 is diverse within different ipomoviruses, with exchanges of lysine (K) and arginine (R) at positions 2 and 3, and in the case of CBSVs, the P1 protein contains LRRA. Whereas in case of SPMMV although the P1 protein has the silencing suppression activity, the

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HCPro is also known to extend the durability of the silencing suppression. Whereas in the case of ipomoviruses lacking HCPro, the P1b protein complements for the silencing suppression activity. The other eight ORFs namely, P3 (the third protein), 6K1 and 6K2 which code for the two 6 kDa proteins, the cylindrical inclusion protein (CI), viral protein genome-linked (VPg), the main viral proteinase (NIa-Pro), the replicase (NIb) and the coat protein (CP) of CBSVs are more similar to the other ipomovirus sequences such as CVYV (Cucumber vein yellowing virus), SqVYV (Squash vein yellowing virus) and SPMMV. A recently reported potyviral protein P3N-PIPO expressing by 2 nucleotide frameshift within the P3 ORF is also found in CBSVs. Both CI and P3N-PIPO complex coordinate the intercellular movement of potyviruses by formation of plasmodesmata-associated structures. In addition to the above described proteins, there are nine ORFs in CBSVs which are characteristic of the Potyviridae family. In addition, they contain an unusual ORF called as HAM1, encoding a polypeptide of 226 aa, flanked by proteolytic cleavage sites, and it is placed in between the replicase (NIb) and the CP. This additional ORF had homology with the Maf/HAM1 superfamily of proteins reported in several prokaryotes and eukaryotes, which are nucleoside triphosphate (NTP) pyrophosphatases. These proteins are known to reduce mutations by intercepting non-canonical forms of NTPs and thus prevent their incorporation into nucleic acids, which otherwise may result in unfavourable mutations. Euphorbia ringspot virus belonging to the Potyvirus genus is the only other viral species reported to contain an HAM1-like sequence. The presence of such an anti-mutator gene may have a beneficial role under oxidative stress conditions, during which mutations occur rampantly. Such a scenario may exist in the plants belonging to the Euphorbiaceae family, particularly the older leaves with higher levels of senescence where CBSVs are shown to accumulate. Blackberry virus Y, another species of the Potyviridae family is also known to contain such unusual heterologous sequences encompassing the AlkB domains within their P1 proteinase, which may also counter the occurrence of deleterious mutation. The presence of such AlkB domains are also known to be present in some of the viruses of the families Flexiviridae and Closteroviridae. However, a more recent investigation about the properties of Ham1 protein of CBSVs by in vitro and in vivo overexpression has indicated Ham1 to be a necrosis determinant. In vitro enzyme assays showed that the Ham1 proteins of both CBSV and UCBSV had ITPase activities. However, deep sequencing experiments did not show any evidence of Ham1 mutagenic protection during infection in N. benthamiana. Deletion and mutation studies with an infectious clone for an isolate of CBSV from Tanzania showed that knock out of Ham1 in CBSV failed to produce necrotic symptoms in the model host N. benthamiana. Thus it was proposed that the Ham1 of CBSVs with highly conserved ITPase motifs may have a highly selectable function during infections in cassava and may represent a euphorbia host adaptation that could be targeted in antiviral strategies.

Evolution, Diversity and Distribution of Viruses Involved in CBSD Phylogenetic analysis of the CBSV/UCBSV CP sequences from East Africa resulted in the formation of two distinct clusters, with all the CBSV CP sequences clustering together and the UCBSV CP sequences forming another distinct cluster (Fig. 2(c)). To date, 27 complete genome sequences of CBSVs are available in the NCBI database, and of these sequences, 11 are UCBSV isolates and 16 are isolates of CBSV. These sequences are isolated from 6 East African countries, 5 from Uganda, 14 from Tanzania, 3 from Mozambique, 3 from Comoros Islands, and one each from Kenya and Malawi (Fig. 2(a)). All the sequences are mostly of 9 Kb length, with minor variation in their sizes, the smallest one reported is 8748 nt long and the longest one is 9097 nt. Phylogenetic analysis of the full length sequences of CBSV/UCBSV also result in the formation of two main clusters corresponding to CBSV and UCBSV isolates and these major clades are further subdivided into minor clades. The cluster with UCBSV isolates has more subclusters than the CBSV species, though the genomes of CBSV isolates are genetically more diverse than those of UCBSV and the evolutionary rate of CBSV is considered to be much higher than UCBSV. Recent studies indicate that the accelerated rate of evolution is mainly contributed by NIa, 6K2, NIb and P1 sequences. P1 was the highest contributor to diversity of CBSVs and phylogenetic analysis of N-terminal sequences of P1 gene alone with advanced bioinformatics tools revealed the presence of a third distinct clade tentatively called as CBSV-Tanzania (CBSV-TZ). Sequence comparison of the complete genomes of CBSV and UCBSV isolates revealed an identity range of 69.0%–70.3% at the nucleotide level and 73.6%–74.4% at the polyprotein amino acid sequence levels. Thus, in view of the sequence divergence between these two groups, two distinct virus species are recognized, CBSV and UCBSV (Fig. 2(d)). While the nucleotide sequence identities among the UCBSV isolates ranged from 87% to 99% and among the CBSV isolates it was 79%–95%. Recombination analysis using RDP4 for all the CBSV and UCBSV sequences indicated 11 recombination events within CBSV isolates and 3 in UCBSV. There was at least one recombination in each of the ten genes, with maximum recombinations (5) detected in CP followed by CI. However, till date no recombinations have been detected between the CBSV and UCBSV isolates, which is consistent with the previous reports on absence of recombination between diverse genomes of RNA viruses due to reduced fitness in the recombinants. The size of the gene sequences that encoded for CI, VPg and CP proteins was variable among isolates of CBSV and UCBSV. The amino acid sequence identity for HAM1 sequences between CBSV and UCBSV was 55%, which was lowest among all gene sequences of the viruses. Such a low identity may indicate that both the Ham1 sequences may have been acquired from two different hosts at two different time points or they may have evolved rapidly than the other genes as evidenced by adaptive selection pressure on HAM1 of both CBSV and UCBSV. Sequence analysis revealed that there are 33 highly conserved amino acid

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residues across HAM1 homologues from different organisms, indicating a strong negative selection on HAM1 and CP genes. The CP of CBSV was 9 aa longer than the CP of UCBSV and the size of P1 protein was also highly variable, with only B60% aa identity between isolates of the two species. These size differences in the CP sequences may have an implication on their differential transmission by the whitefly vector. Although the transmission of CBSVs by whiteflies is well established under experimental conditions the differential transmission efficiency by whiteflies between the two viruses has not been significant. A recent report on detailed sequence analysis for several CBSV isolates revealed the presence of the DAG motif in their CP genes, however such a motif was not recorded in the UCBSV CP sequences. DAG sequences are present in the HCPro encoded by all the potyviruses which is an important determinant for aphid transmission. Hence it needs to be experimentally verified if aphids are capable of transmitting the CBSVs or if it is most probably an evolutionary mark in the CP of the ipomoviruses. The diversity in the P1 sequences may result in variation in their ability to suppress RNA silencing and which may have a correlation with the virulence of CBSV and UCBSV isolates. Although in the fields there are no apparent symptom differences among the isolates of CBSV and UCBSV, under controlled conditions distinct phenotypes are produced by CBSV and UCBSV isolates. Mixed infections are found in regions where both CBSV and UCBSV are found. However, there is no evidence of synergistic interaction between the isolates of CBSV and UCBSV, unlike the interaction between different species of cassava infecting geminiviruses. A RT-PCR based study in 2014, in eight major districts of Rwanda on distribution of CBSVs revealed that single infections of cassava by UCBSV were most common (74.2%) and were associated with wide range of symptom phenotypes. However, single infections by CBSV were predominant (15.3%) in plants showing CBSD symptoms on stem alone, while mixed infections of CBSV and UCBSV were in 10.5% of the total number of plants infected and were predominantly present in combinations of leaf þ stem þ root phenotypes. Further, phylogenetic analysis of a partial sequence of CP of all the isolates in this study indicated that in Rwanda there is only one type of CBSV while diverse types of UCBSVs are involved in CBSD.

Management and Control of CBSD Phytosanitary practices are most important for virus free cultivation of cassava, because of its high vulnerability to virus transmission. In this regard the group led by Dr. James Legg based in the CGIAR institute IITA (Tanzania) is playing a major role in containing CBSD spread. CBSD is transmitted by the whitefly vectors for a shorter distance since the mode of transmission of these ipomoviruses is semi-persistent in nature. While long distance transmission of CBSD can only happen through long distance movement of vegetative propagules. Some of the important steps for control of CBSD are production of clean or disease free planting material, virus indexing of parent material and the pre-basic germplasm propagation, and regular roguing of diseased plants. Further, cooperation among group of farmers at a community level is important for a collective action and implementation of phytosanitary measures. Currently, such large scale initiatives are being implemented in Eastern and Southern Africa, and efforts are made to contain the regional spread of CBSD. Further, national and regional quarantine measures are enforced to prevent spread of CBSD during exchange of cassava germplasm.

Sources of CBSD Resistance and Their Introgression Crop improvement through resistance breeding has been the most successful approach for control of crop diseases. Although, breeding for virus resistance in cassava was initiated in 1935 at Amani, Tanzania; it has been a major challenge since cassava is cross pollinated, with high level of heterozygosity. In this breeding programme, 46106/27 was the most CBSD resistant cassava variety developed by backcrossing M. glaziovii with M. esculenta. This is one of the most successful products of the research programme at Amani, presently cultivated by the farmers and this CBSD resistance still persists in the farmers’ fields in Kenya, which locally is called as Kaleso. Hundreds of SNP markers have been developed for CBSD tolerance and a linkage map has also been developed which is used for detection of CBSD tolerant quantitative trait loci (QTLs). Mapping populations developed by crossing CBSD-tolerant cultivar, Namikonga and a susceptible cultivar, Albert are being used for identification of QTLs associated with CBSD resistance. Now that cassava genome sequence is published, it could be used for identifying genes that control CBSD resistance and for development of new markers associated with CBSD resistance. Next generation sequencing technologies are also being employed for identification of genes associated with CBSD tolerance or the genes that respond to CBSD infection. Gene expression studies, using cassava varieties with differential resistance to CBSD revealed that at least 700 genes were overexpressing in the CBSD resistant variety Kaleso when compared to the susceptible variety Albert. Although none of these differentially expressed genes resembled a known disease resistance gene, many of them encoded enzymes or factors involved in hormone signalling or secondary metabolites pathways, that may be associated with disease resistance. However, none of the cassava genotypes/hybrids identified are immune to CBSD, but majority of them are tolerant since they show clear foliar symptoms, but necrosis in the roots is absent or delayed. In one of the recent studies, 238 cassava germplasm accessions from South America were screened for CBSD resistance. There were 7 South American accessions that were completely free of CBSD symptoms and were negative for RT-PCR detection. The breeding lines DSC167 and DSC188 showed broad spectrum resistance to diverse isolates of CBSVs. This clearly reflects that

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Fig. 3 (a) Screening of RNAi-based transgenic N. benthamiana for resistance against CBSVs (b) Confined field trials for RNAi-based transgenic cassava lines in Uganda, (c, d) Brown necrotic lesions seen on the stems of CBSD-infected control cassava plants, compared with non-infected transgenic cassava, (e, f) Cross section of cassava roots showing CBSD symptom distribution in non-transgenic vs transgenic cassava plants, (g) accumulation of small interfering (si)-RNA in transgenic cassava as revealed by Northern hybridization technique. Adapted from Patil, B.L., Legg, J.P., Kanju, E., Fauquet, C.M., 2015. Cassava brown streak disease: A threat to food security in Africa. Journal of General Virology 96, 956–968.

the germplasm collections are potential reservoirs of several disease resistance genes and their conservation is critically important for resistance breeding programmes. A project led by the International Institute of Tropical Agriculture (IITA) aims to ensure that the farmers in Africa have access to diverse disease-free improved cassava varieties with resistance to both CBSD and CMD, and with consumer preferred traits. Studies have revealed that both additive and non-additive genetic effects are required for expression of CBSD resistance trait, and of several crosses studied, the cassava variety Kaleso (Namikonga) had the best general combining ability for CBSD resistance.

Engineering CBSD Resistance Although there are sources of resistance against CBSD, it could be impossible to introgress CBSD resistance while retaining the original agronomic/quality traits of cassava. Furthermore, the resistance reported against CBSD is more of tolerance to virus infection, rather than immunity against the CBSVs. Hence, genetic engineering is an important option for controlling plant viruses and CBSVs in particular. There are several strategies used for control of viruses that infect plants, which will not be elaborated here, since it is an extensively reviewed topic. The first successful control of CBSV was demonstrated using RNA-interference (RNAi) strategy (Fig. 3), result confirmed later by other groups. Similarly, CBSD resistance has been accomplished by over-expression of artificial micro RNAs that targeted the CBSVs, however this work was restricted to the model host N. benthamiana. An interaction between the viral genome-linked protein (VPg) and host eukaryotic translation initiation factor 4E (eIF4E) isoforms is pre-requisite for a potyvirus to establish infection in a host cell.

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Recently, the CRISPR/Cas9-mediated genome editing was employed to alter the sequence of cassava eukaryotic translation initiation factor 4E (eIF4E) isoforms, namely ncbp-1 and ncbp-2. When the ncbp-1/ncbp-2 cassava mutants were challenged with CBSV, there was a delay in symptom development, as well as reduced severity and incidence of storage root necrosis, which corresponded with reduced virus titer in storage roots. However, despite all these very positive results showing the possible control of CBSD through biotechnology, none of the putative commercial products have been distributed to farmers so far, due to the lack of a biotechnology official framework in East African countries to permit it’s use in safe conditions.

Cross Reading With Other Chapters Cassava Mosaic Viruses (Geminiviridae); Potyviruses (Potyviridae).

See also: Cassava Mosaic Viruses (Geminiviridae). Potyviruses (Potyviridae)

Further Reading Anjanappa, R.B., Mehta, D., Okoniewski, M.J., et al., 2018. Molecular insights into Cassava brown streak virus susceptibility and resistance by profiling of the early host response. Molecular Plant Pathology 19, 476–489. Beyene, G., Chauhan, R.D., Ilyas, M., et al., 2017. A virus-derived stacked RNAi Construct confers robust resistance to Cassava brown streak disease. Frontiers in Plant Science 7, 2052. Gomez, M.A., Lin, Z.D., Moll, T., et al., 2019. Simultaneous CRISPR/Cas9-mediated editing of cassava eIF4E isoforms nCBP-1 and nCBP-2 reduces cassava brown streak disease symptom severity and incidence. Plant Biotechnology Journal 17 (2), 421–434. Legg, J.P., Lava Kumar, P., Makeshkumar, T., et al., 2015. Cassava virus diseases: biology, epidemiology, and management. Advances in Virus Research 91, 85–142. Legg, J., Ndalahwa, M., Yabeja, J., et al., 2017. Community phytosanitation to manage cassava brown streak disease. Virus Research 241, 236–253. Mbanzibwa, D.R., Tian, Y., Mukasa, S.B., Valkonen, J.P., 2009. Cassava brown streak virus (Potyviridae) encodes a putative Maf/HAM1 pyrophosphatase implicated in reduction of mutations and a P1 proteinase that suppresses RNA silencing but contains no HC-Pro. Journal of Virology 83, 6934–6940. Mbanzibwa, D.R., Tian, Y.P., Tugume, A.K., et al., 2011. Evolution of cassava brown streak disease-associated viruses. Journal of General Virology 92, 974–987. Patil, B.L., Legg, J.P., Kanju, E., Fauquet, C.M., 2015. Cassava brown streak disease: A threat to food security in Africa. Journal of General Virology 96, 956–968. Patil, B.L., Ogwok, E., Wagaba, H., et al., 2011. RNAi-mediated resistance to diverse isolates belonging to two virus species involved in Cassava brown streak disease. Molecular Plant Pathology 12, 31–41. Rwegasira, G.M., Momanyi, G., Rey, M.E., Kahwa, G., Legg, J.P., 2011. Widespread occurrence and diversity of Cassava brown streak virus (Potyviridae: Ipomovirus) in Tanzania. Phytopathology 101, 1159–1167. Taylor, N.J., Halsey, M., Gaitán-Solís, E., et al., 2012. The VIRCA project: Virus resistant cassava for Africa. GM Crops & Food: Biotechnology in Agriculture and the Food Chain 3, 93–103. Tomlinson, K.R., Bailey, A.M., Alicai, T., Seal, S., Foster, G.D., 2018. Cassava brown streak disease: historical timeline, current knowledge and future prospects. Molecular Plant Pathology 19, 1282–1294. Winter, S., Koerbler, M., Stein, B., et al., 2010. Analysis of cassava brown streak viruses reveals the presence of distinct virus species causing cassava brown streak disease in East Africa. Journal of General Virology 91, 1365–1372. Yadav, J.S., Ogwok, E., Wagaba, H., et al., 2011. RNAi-mediated resistance to Cassava brown streak Uganda virus in transgenic cassava. Molecular Plant Pathology 12, 677–687.

Relevant Websites https://www.cabi.org/isc/datasheet/17107 Cassava brown streak viruses. https://www.plantwise.org/knowledgebank/datasheet/17107 Cassava Brown Streak Disease - Plantwise Knowledge Bank. https://mel.cgiar.org/projects/424 Consultative Group for International Agricultural Research. https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/w/potyviridae/569/genus-ipomovirus Genus: Ipomovirus.

Cassava Mosaic Viruses (Geminiviridae) James Legg, International Institute of Tropical Agriculture, Dar es Salaam, Tanzania Stephan Winter, Leibniz Institute – DSMZ – German Collection of Microorganisms and Cell Cultures GmbH, Braunschweig, Germany r 2021 Elsevier Ltd. All rights reserved. This is an update of J.P. Legg, African Cassava Mosaic Disease, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00693-2.

Nomenclature CMG Cassava mosaic geminivirus CP Coat protein or capsid protein CR Common region dsDNA Double-stranded deoxyribonucleic acid ELISA Enzyme-linked immuno-sorbent assay IR Intergenic region kb Kilobase kbp Kilobase pair kDa Kilo Dalton

Glossary Nonanucleotide A codon containing nine nucleotides. Pandemic An outbreak of a disease over several countries or a large part of the world.

MP Movement protein NSP Nuclear shuttle protein PCR Polymerase chain reaction PTGS Post-transcriptional gene silencing RCA Rolling-circle amplification Ren Replication enhancer protein siRNA Small interfering RNA ssDNA Single-stranded deoxyribonucleic acid TrAP Transcriptional activator protein

Pseudo-recombinant A viral infection involving complementary genome components from different virus species.

Taxonomy Cassava mosaic viruses belong to the genus Begomovirus in the family Geminiviridae which comprises viruses with single-stranded circular DNA genomes that are transmitted by Bemisia tabaci (Gennadius) whiteflies. The cassava-infecting begomoviruses have two DNA components (DNA-A and DNA-B) of which the DNA-A genome sequences are used for taxonomic assignment of virus species. The species demarcation threshold is at 91% sequence identity, thus virus genomes having less than 91% of nucleotides on DNA-A that are identical to those of their closest neighbor are assigned to a unique species. Eleven cassava mosaic geminiviruses (CMGs) are currently recognized as distinct virus species by the ICTV (See Relevant Websites Section). These are: African cassava mosaic virus (ACMV), East African cassava mosaic virus (EACMV), South African cassava mosaic virus (SACMV), Indian cassava mosaic virus (ICMV), Sri Lankan cassava mosaic virus (SLCMV), East African cassava mosaic Malawi virus (EACMMV), East African cassava mosaic Cameroon virus (EACMCV), East African cassava mosaic Zanzibar virus (EACMZV), East African cassava mosaic Kenya virus (EACMKV), Cassava mosaic Madagascar virus (CMMGV), and African cassava mosaic Burkina Faso virus (ACMBFV) (Table 1; Fig. 1). Several of these originated from recombination events that generated genomic diversities beyond species boundaries. The names given reflect the relatedness to their putative parental virus.

Intraspecific Diversity New, recombinant cassava viruses described provide evidence for on-going diversification of CMG genomes. Interspecific recombination of DNA-A and DNA-B genome components of cassava viruses also include sequences from other begomoviruses. For example, genome portions of the monopartite Tomato leaf curl virus are present in ACMBFV. Similarly, the genome components of CMMGV include, besides elements from other CMGs, sequences from non-related monopartite begomoviruses infecting tomato in the Indian Ocean islands. Additionally, there is ample evidence for molecular diversity of CMG genomes from complete DNA-A and DNA-B genome sequences of virus isolates available in GenBank. Nevertheless, in spite of contrasting molecular characteristics, there is little evidence for unique biological features of each of the virus species in terms of their symptom phenotypes, natural host range, transmission parameters or the association of particular virus isolates with B. tabaci species that would provide additional information for the discrimination of virus strains or the identification of a particular virus species. The 11 acknowledged CMG species and the numerous isolates that are present in Africa and Asia are merely characterized by distinctive sequences of their DNA-A genome components.

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Table 1

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Cassava mosaic geminivirus species

Virus

Reference

Genome

Occurrence

African cassava mosaic virus

Bock and Woods (1983)

NC_001467 NC_001468

Africa

East African cassava mosaic virus

Swanson and Harrison (1994)

NC
004674 NC
004676

East Africa

Indian cassava mosaic virus

Hong et al. (1993)

NC_001932 NC_001933

India

Sri Lankan cassava mosaic virus

Saunders et al. (2002)

NC_003861 NC_003862

India, Sri Lanka, Vietnam, Cambodia, China

South African cassava mosaic virus

Berrie et al. (1998)

NC_003803 NC_003804

Southern Africa

East African cassava mosaic Cameroon virus

Fondong et al. (1998)

NC_004625 NC_004630

East and West Africa

East African cassava mosaic Malawi virus

Zhou et al. (1998)

NC_022645 NC_022644

East and southern Africa

East African cassava mosaic Zanzibar virus

Maruthi et al. (2004)

NC_004655 NC_004656

East Africa

East African cassava mosaic Kenya virus

Bull et al. (2006)

NC_011583 NC_001584

East Africa

Cassava mosaic Madagascar virus

Harimalala et al. (2012)

NC_017004 NC_017005

Madagascar

African cassava mosaic Burkina Faso virus

Tiendrebeogo et al. (2012)

HE616777 HE616778

West Africa

Fig. 1 Estimated phylogeny of DNA-A genome sequences of cassava mosaic geminivirus species within the genus Begomovirus, family Geminiviridae. The evolutionary history was inferred using the Maximum Likelihood method with the percentage of trees in which the associated taxa clustered together shown next to the branches. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The evolutionary analyses were conducted in MEGA X.

History Cassava (Manihot esculenta Crantz) is a root crop that is grown widely throughout the tropics, primarily for its value as a starchy staple food. From its origins in Latin America, cultivated cassava was introduced to Africa in the sixteenth century, and subsequently spread through much of Africa south of the Sahara. The first cultivation of cassava in Asia occurred in south India in the

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19th century, before it spread more widely throughout South-East Asia and eventually as far as Southern China. Although initially used as a food crop, most cassava in Asia is now cultivated for industrial starch production. None of the viruses infecting cassava in Latin America is known to have been carried to either Africa or Asia, but over time, the crop became infected by indigenous viruses. The first report of a virus-like disease in African cassava was made in 1894 from what is now northeastern Tanzania. The original German descriptor, ‘Krauselkrankheit’, made reference to the characteristic mosaic symptoms elicited in affected plants. It was not until just over a decade later, however, that the first firm indication was given that the disease had a viral etiology. In spite of these early reports, there seems to have been little concern about the impact of cassava mosaic disease (CMD) until the 1920s. Between 1929 and 1937, however, numerous reports were made of the spread and damaging effect on cassava crops of CMD from diverse locations throughout the continent, from the island of Madagascar off the southeastern shores of the African mainland, to Sierra Leone in West Africa. These developments provided the stimulus for the earliest concerted efforts to develop approaches to controlling the viruses that caused this damaging crop disease. Although substantial progress was made in the development of cassava varieties that were resistant to cassava mosaic-causing viruses in the 1930s and 1940s, the viruses themselves remained poorly understood, and it was not until 1983 that the first definitive study confirming the viral etiology of CMD was published. Geminate virions were shown to encapsidate a bipartite genome of singlestranded circular DNA, leading to the designation of these viruses as geminiviruses in the genus Begomovirus. In Asia, symptoms of cassava mosaic disease were first observed from southern India in the 1950s, and subsequently from Sri Lanka in the 1980s. Since 2015, the disease has spread more widely, affecting Cambodia and Vietnam in South-East Asia.

Virion Structure The two single-stranded circular DNA molecules (DNA-A and DNA-B) of CMGs are encapsidated within a virion having a unique architecture composed of two incomplete icosahedral particles (hemicapsids) fused to form a twinned, geminate particle from a single type of capsid protein. For each icosahedral half, 55 CP chains assemble into 11 capsomers. The near-atomic resolution of the virion structure of ACMV has been resolved by cryo-electron microscopy to 4.2 Å revealing that alternating conformations of the N-terminus of the CP confer stability of the twinned particle and most probably mediate the interaction with the encapsidated DNA (Fig. 2). This was further confirmed by similar studies with Ageratum yellow vein virus, and resolution at an even higher level

Fig. 2 Structure of African cassava mosaic virus determined by cryoelectron microscopy. Courtesy of Katharina Hipp.

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(3.3 Å ) showed that three distinct conformations of CP form the capsid and arrange the hemicapsid interface. The proposed model for geminivirus capsid assembly and DNA binding suggests that genomic DNA is crucial for the formation of the capsids and for directing the assembly of the capsomers and the virus particle. The high resolution model of the ACMV virion structure has also shown the position of amino acid side chains implicated in B. tabaci transmission of begomoviruses. The most critical, Q130 and N131, are located on the particle surface and are accessible to interact with vector insect receptors while other residues may influence particle stability and thus have indirect effects on virus transmission.

Genome The eight genes on the DNA-A and DNA-B genome components are located in sense and complementary sense orientations on the circular covalently-closed single-stranded DNA molecules (Fig. 3). A conserved intergenic region (IR) of ca. 200 bp is shared between both genome components. This unique IR identifies DNA-B to be transreplicated by its cognate DNA-A only. Embedded in the IR is a stem loop comprising the TAATATTAC sequence that is highly conserved among almost all geminiviruses and which represents the origin of replication. While DNA-A can infect and replicate autonomously in a host plant it requires DNA-B for movement and systemic infection of the host. CMG genes are multifunctional and proteins are responsible for encapsidation and for directing the host machinery to provide cellular functions favorable for replication. In virion sense orientation on DNA-A, the structural protein gene CP (AV1) codes for the capsid protein, which is involved in whitefly transmission and transport, and which interacts with host proteins and binds to ss and double-stranded (ds)DNA. A functional CP, however, is not required for infectivity and movement. Thus, DNA-A with a defective or missing CP is still infectious and capable of systemic movement within its host, however its capacity for vector transmission is compromised. Upstream of CP, the AV2 gene also provides movement function. On the complementary-sense strand, the replication initiation protein Rep (AC1) initiates specific steps in DNA replication; TrAP (transcriptional activator protein)(AC2) and REn (replication enhancer protein)(AC3) further downstream of Rep, function as suppressers of post-transcriptional gene silencing (PTGS) and support efficient DNA replication respectively. AC4 is nested within Rep and involved in symptom expression and possibly in counter-defense. Genes on DNA-B enable virus movement within the cell and nucleus (nuclear shuttle protein, NSP)(BV1) as well as sustaining long distance spread and movement within the plant (movement protein, MP)(BC1).

Replication Geminiviruses replicate in the nuclei of infected cells and depend on host DNA and RNA polymerases for their replication and transcription. Geminiviruses predominantly replicate in the phloem companion and parenchyma cells and some geminiviruses can also invade mesophyll cells but are never found in meristems. Thus, geminiviruses replicate in differentiated cells. Using host proteins the ssDNA genome is replicated by rolling circle- and recombination-mediated mechanisms in which double-stranded DNA intermediates are formed that are chromatinized with host histone proteins to form viral minichromosomes inside the nucleus. Rep recruits host factors, binds specifically to iteron sequences within IR and creates a nick in the conserved nonanucleotide TAATATT↓AC to initiate DNA replication. Rep also binds to a plant homolog of retinoblastoma protein to catalyze a series of intracellular reactions that move the cell from G to S phase to mobilize host factors essential for DNA replication. Double-stranded DNA (dsDNA) produced in the replication process then assembles under the control of TrAP and REn into

Fig. 3 Genome organization of bipartite cassava mosaic viruses. The proteins encoded by the components are indicated as follows: DNA-A component on the sense orientation encodes for the following proteins: CP or AV1; the coat protein, and the AV2 protein; and on the antisense orientation: Rep or AC1; replication associated protein, TrAP or AC2; the transcriptional activator protein, REn or AC3; the replication enhancer protein, and the AC4 protein. The DNA-B component encodes for the following proteins: NSP or BV1; the nuclear shuttle protein on the sense orientation and MP or BC1; the movement protein on the antisense orientation. The conserved region (CR) is the common region between the DNA-A and DNA-B components. For each component the conserved hairpin structure, containing the nonanucleotide sequence TAATATTAC within the loop structure, is shown at position zero.

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transcriptionally active mini-chromosomes. The nuclear export is mediated by NSP, from nucleus to the cytoplasm, and movement out of the cell through plasmodesmata and long distance transport and systemic movement are facilitated by MP.

Transmission CMGs, in common with all other members of the genus Begomovirus, are transmitted by the whitefly vector B. tabaci. Transmission can also be achieved through grafting, and relatively inefficiently through mechanical inoculation of indicator plants, but there is no seed-borne transmission. Cassava is normally propagated through the use of vegetative cuttings, and this is perhaps the most frequent source of infection in new crops under field conditions. Bemisia tabaci adults feed, mate, and reproduce preferentially on the upper newly-emerged leaves of cassava plants, and almost all transmission occurs here. Transmission is persistent, and ACMV has been shown to be retained by B. tabaci adults for up to 9 days. Transtadial, but not transovarial transmission, has been demonstrated, although the larval instars are unimportant in the epidemiology of CMGs in view of their sessile nature. Minimum periods for each of the stages of transmission for ACMV by B. tabaci adults are: acquisition (3 h), latent period (3 h), and inoculation (10 min). Inoculated plants begin to show symptoms of infection after 3–5 weeks. Varying levels of transmission efficiency have been reported, ranging from 0.3% to 10%, depending primarily on the nature of the CMG infection in the plants from which virus is acquired. There is evidence for a limited degree of co-evolutionary adaptation between the whitefly vector and CMGs, as Indian whiteflies are significantly better at vectoring Indian than African CMGs and vice versa. However, within Africa, there is currently no indication that whiteflies from a particular location are more efficient at vectoring locally-occurring CMGs than they are at vectoring CMGs from another part of the continent. There is, however, an important balance between the pattern of transmission and the nature of the virus infection. More severe infections, caused either by more virulent virus species/strains or by mixed virus infections, lead to a greater frequency of whitefly-borne infection but diminished propagation through cuttings by farmers. Conversely, more moderate or mild infections, caused by less virulent virus species/strains occurring in single infections, lead to less frequent whitefly-borne infection and increased propagation through cuttings by farmers. This balance dictates the epidemiological characteristics of CMG infection in any given location, area, or region.

Pathogenesis and Symptoms Following initial infection of a previously uninfected cassava plant by a CMG, viral DNA moves through the phloem to the newly developing phloem and parenchyma cells immediately behind the meristem where rapid multiplication of virus particles takes place. Plants have developed defense responses to virus multiplication through the process of PTGS, but in response, CMG species/ strains have developed various effective mechanisms to overcome this, and the degree of this effectiveness seems to be the main factor in determining the severity of disease resulting from infection. Both AC2 and AC4 have been shown to act as suppressors of PTGS. Significantly, in mixed ACMV þ EACMV-like virus infections, the responses of the two viruses to host plant PTGS are complementary, leading to a synergistic interaction between the two viruses, greatly increased titers of both, and a concomitant increase in severity of the disease symptoms expressed in the plant. Molecular studies of these interactions have shown that the abundance of short interfering RNA (siRNA) molecules associated with the host plant PTGS response increases over time in pure ACMV infections, yet remains low over similar time periods for ACMV þ EACMV-UG co-infections. Synergism has been described for ACMV þ EACMCV mixtures in Cameroon, but its significance is greatest in the widely occurring ACMV þ EACMV-UG mixed infections that cause severe CMD in many parts of East and Central Africa. As CMG virions replicate in infected cells, chloroplasts become distorted and reduced in number leading to the appearance of a patchy mosaic of chlorotic leaf portions interspersed with normal green leaf tissue. The chlorotic mosaic patches have discrete, non-diffuse borders, but these may vary greatly in size, ranging from small portions of individual leaflets to the entire leaf (Fig. 4). In moderate to severe infections, leaves are distorted and reduced in size and the growth of the plant is stunted. The severity of symptoms is affected by several factors, including the virulence of the CMG strain, the occurrence of mixed virus infection, temperature, host susceptibility and the stage of growth of the plant. Varieties display a full range of responses to CMG infection, ranging from near immunity to extreme sensitivity resulting in plant death. The most frequent cause of severe symptoms is where ACMV and EACMV (or related viruses) occur in mixed infection. This is a consequence of the synergistic interaction between these viruses which leads to increased virus titer of both. There are currently no anecdotal or published records of synergistic interactions between ICMV and SLCMV in South Asia. Plants infected through the cutting typically express symptoms in the first-formed leaves immediately on sprouting. Plants that sprouted without virus infection, but which were subsequently infected by the whitefly vector express symptoms only in leaves above the point of infection, although there is a latent period of 3–5 weeks between inoculation and the appearance of symptoms. These contrasting patterns of symptom expression make it possible to distinguish plants infected through the cutting from those infected during crop growth by the whitefly vector. Separating these two infection types has become an important facet of field research into the epidemiology of CMGs. Symptoms of CMD caused by ICMV and/or SLCMV in Asia vary in severity, depending on the factors described above, but the characteristics of the symptoms are the same as those of African CMD, as is the distinction between cutting-borne and whitefly-borne infection.

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Fig. 4 Cassava mosaic disease (CMD) symptoms: (A) & (B) – Sri Lankan cassava mosaic virus in Vietnam; (C) – African cassava mosaic virus in Gabon; (D) – mixed infection of African cassava mosaic virus and East African cassava mosaic virus-Uganda in Burundi.

There are currently no reported cases of cassava varieties with immunity to infection by CMGs. However, there are many varieties that are highly resistant, and when these are infected, symptoms expressed are often confined to small portions of a few leaves. A common feature of resistant varieties is symptom recovery, as lower symptomatic leaves drop naturally and new foliage is asymptomatic. This phenomenon is linked to reversion, which is the absence of virus in a proportion of cuttings taken from an infected parent plant. This is thought to be a consequence of the incomplete systemicity of CMGs in resistant varieties.

Epidemiology Diversity and Distribution Distribution of the African CMGs The earliest studies of CMG diversity and distribution used serological techniques to characterize variability, and then utilized that variability to develop diagnostic tests based on the use of monoclonal antibodies in the enzyme-linked immunosorbent assay (ELISA). Using these methods, two principal groups of CMGs were recognized that were subsequently confirmed as distinct species following the sequencing of DNA-A molecules. These were ACMV and EACMV. The earliest distribution maps of the African CMGs showed EACMV to occur in the coastal east African areas of Kenya, Tanzania, and Madagascar, as well as Malawi and Zimbabwe. ACMV, by contrast, occurred throughout the remainder of the cassava-growing areas of Africa, from South Africa and Mozambique in the southeast, to Senegal in the northwest. Significantly, at this time there was no reported zone of co-occurrence of the two virus species. With the increased use of polymerase chain reaction (PCR)-based diagnostics from 1990s onward, it became possible to identify differences not solely associated with the coat protein. This led to two important developments in the understanding of CMGs in Africa. First, several new species were identified, most of which were more closely related to EACMV than to ACMV. Second, it was shown that virus mixtures belonging to different species occurred frequently. A notable consequence of this finding was the concomitant evidence for the more widespread distribution of the EACMV-like viruses than had hitherto been recognized. New virus identifications included: SACMV (1998), EACMMV (1998), EACMCV (2000), EACMZV (2004), EACMKV (2006), ACMBFV (2012), and CMMGV (2012). Significantly, all but one (ACMBFV) of the CMGs occur in different parts of East and Southern Africa, while ACMV predominates in West Africa (Fig. 5). EACMCV, the only EACMV-like virus occurring in West Africa, is less frequent and often occurs in mixed infections together with ACMV. ACMV is absent from coastal areas of Kenya and

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Fig. 5 Distribution of cassava mosaic geminiviruses in Africa: ACMV (African cassava mosaic virus); ACMBFV (African cassava mosaic Burkina Faso virus); EACMV (East African cassava mosaic virus); EACMV-UG (East African cassava mosaic virus-Uganda); EACMCV (East African cassava mosaic Cameroon virus); EACMZV (East African cassava mosaic Zanzibar virus); EACMMV (East African cassava mosaic Malawi virus); EACMKV (East African cassava mosaic Kenya virus); SACMV (South African cassava mosaic virus); CMMGV (Cassava mosaic Madagascar virus).

Tanzania. Since there has been very little CMG characterization in many of the cassava-growing countries of Africa, it seems likely that further variability within this group of viruses still remains to be revealed. An assessment of the data currently available has led to the conclusion that East Africa is the center of diversity for the EACMV-like CMGs and is probably the home for yet-to-beidentified wild hosts of the proto-CMGs that were first introduced to cassava by B. tabaci sometime between the earliest introductions of cassava to this part of Africa in the eighteenth century, and the first report of CMD in 1894.

Distribution of CMGs in Asia ICMV and SLCMV are considered to be native to South Asia and have been recorded from cassava-producing areas of India and Sri Lanka for several decades. Early serology-based work identified ICMV from southern India. However, following the characterization of SLCMV as the causal agent of CMD in Sri Lanka in 2002, subsequent surveys in India showed that SLCMV was more widely distributed there than ICMV. The CMGs in India have been identified from Kerala, Karnataka, Tamil Nadu, Andhra Pradesh, Madhya Pradesh, and West Bengal states, a zone covering all of the major cassava-producing parts of the country. Although it has been suggested that SLCMV is a more aggressive virus than ICMV, and has been expanding its geographical coverage, the absence of systematic surveys conducted over several years means that the evidence for this remains circumstantial. CMD was reported for the first time from Southeast Asia (SE Asia) from a single location in Ratanakiri Province in Northeastern Cambodia in 2015. The causal virus was SLCMV, and it was assumed that this new outbreak had arisen through the introduction of infected planting material from South Asia. Since that time there has been rapid spread of CMD and associated SLCMV through most of the major cassava-producing areas of Cambodia, as well as southern Vietnam. An isolated report of the disease from Ha Tinh Province in northern Vietnam presents a risk for spread to neighboring Laos, and there are also several reports of CMD in Thailand that have yet to be officially confirmed. The most recent confirmed reports of the occurrence of CMD in Asia come from the southern Chinese regions of Hainan and Fujian, in both cases coupled with positive diagnoses of SLCMV. Rapid spread of SLCMV through SE Asia is thought to be driven by extensive cross-border and intra-regional commercial trade in cassava stems, although there is also clear evidence of local spread through whitefly transmission. The progress of the SE Asian pandemic has also been exacerbated by the extreme susceptibility to CMG infection of virtually all of the cassava varieties currently being cultivated. Given the rapid spread of CMD through this region from 2015 to 2019, it seems likely that new occurrences will be reported from

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Laos, Thailand and Myanmar in the near future, and there is also a greatly increased threat to other major-cassava producing countries in the region – Philippines and Indonesia.

Recombination and the CMD pandemic The CMGs represent a very dynamic group of viruses, and evidence has been presented for the occurrence of both pseudorecombinants, in which the DNA-A of one virus species co-replicates with the DNA-B of another species, as well as true recombinants, in which portions of the DNA-A or -B of one species have been spliced into the DNA-A or -B of another. In fact, all of the EACMV-like viruses other than EACMV show evidence for recombination events either with known or as yet unknown begomoviruses. One of the most important developments in the study of CMGs in Africa was the recognition that an unusually damaging strain, referred to as EACMV-UG, had arisen through a recombination event between EACMV and ACMV in which a 340 nt portion of ACMV AV1, had replaced the equivalent portion of the DNA-A of EACMV. The consequence of this was that ELISA-based diagnostic tests erroneously identified this strain as ACMV. This is one of the reasons why PCR-based diagnostic methods are now used almost exclusively for CMG monitoring work. EACMV-UG was associated with the rapid epidemic-like spread of CMD in East and Central Africa through the latter part of the twentieth and early 21st century. Countries affected by the severe CMD pandemic included: Angola, Burundi, Cameroon, Central African Republic, Democratic Republic of Congo (DRC), Gabon, Kenya, Republic of Congo, Rwanda, South Sudan, Tanzania, and Uganda. Localized identifications of EACMV-UG have also been made from Burkina Faso, Chad, Equatorial Guinea, South Africa, Swaziland, and Zimbabwe. The dynamic character of CMG diversity and distribution cannot be overemphasized, and the propensity that this group of viruses shows to produce both naturally occurring pseudo-recombinants, as well as novel true recombinants, ensures that the patterns of diversity and distribution will continue to evolve. It is significant to note, however, that most of the CMG variability occurs amongst the EACMV-like viruses, which show a high propensity for recombination. This contrasts strongly with ACMV, for which all 311 publicly-available accessions have 494% homology and can be considered to be a single strain of the same species.

Field-Level Epidemiology Epidemiological studies of the CMGs can be broadly categorized into two groups: those that describe patterns of infection at the field level and those that relate to area or region-wide spread. In the case of the former, external sources of infection have been shown to be most important for determining the rate and quantity of infection in initially CMD-free plantings in the normal field environment. Gradients of infection occur in which new infections are most frequent on the windward sides of cassava crops and these gradients are matched by similar patterns of vector distribution. Multiple regression relationships have been used to describe the association between measures of inoculum pressure and final CMD incidence in test plots. Gompertz curves model patterns of infection increase in plots of CMG-susceptible cassava cultivars, and incidence increases rapidly to 100% over the first 3–6 months of growth. For resistant varieties, however, under similar conditions of inoculum pressure, rates of infection increase are much lower and final incidences typically range from 0% to 50%. Mathematical models have been generated that predict patterns of CMD spread under varied conditions of virus infection and vector abundance. The great majority of epidemiological research has focused on CMD in Africa, although several studies in India have confirmed that patterns of cutting-borne and vector-borne spread of CMGs are largely the same as those observed in Africa.

Regional Epidemiology Following the earliest ‘first colonization’ descriptions of African CMD epidemics in the 1920s and 1930s, a few reports were made of rapid area-wide spread of severe CMD at other times during the twentieth century, some of the most notable of which were epidemics in Cape Verde and south-eastern Nigeria in the 1990s. Of much greater importance, however, has been the African CMD pandemic that was first reported from the northern-central part of Uganda in the late 1980s. CMD associated with the epidemic was unusually severe and was rapidly spread by super-abundant populations of B. tabaci. During the 1990s it became apparent that the zone affected by this severe CMD was expanding southwards at a rate of 20–30 km per year, and in 1997 molecular studies revealed that the severe disease phenotype was associated with the occurrence and spread of an ‘invasive’ recombinant CMG, EACMV-UG, commonly in mixed infection with the locally occurring, but now synergized ACMV. Regular monitoring surveys conducted throughout the East and Central African region through the 1990s and early part of the 21st century have used PCRbased diagnostics to map the spread of the EAMCV-UG associated with this ‘pandemic’ of severe CMD. This work has given rise to first reports of EACMV-UG and resulting severe CMD in ten additional countries in East and Central Africa: Kenya (1996), South Sudan (1997), Tanzania (1999), DRC (1999), Republic of Congo (1999), Rwanda (2001), Burundi (2003), Gabon (2003), Angola (2008), and Cameroon (2010). Significantly, the pandemic ‘front’ advanced through north-western Tanzania from the Uganda border a distance of c. 400 km between 1999 and 2004. Although the severe CMD pandemic threatened to spread further through southern Africa and westwards to Nigeria, the period post-2010 saw a decline in new reports of spread. Evidence from Uganda in 2019 indicates that EACMV-UG has now largely been replaced by ACMV, although the reasons for this pattern of change are currently unclear. An important contributing factor to this amelioration in the CMD pandemic in Africa is likely to be the increasingly widespread adoption by farmers of CMD-resistant varieties.

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Diagnosis CMD symptoms are very characteristic, although they cannot be used to discriminate between the CMG virus species causing the infection. Additionally, symptoms are almost always expressed in infected plants at some stage during their period of growth, which means that symptoms are widely used as an indication of infection during surveillance activities and by regulatory authorities during certification inspections. The characteristic mosaic pattern of CMD symptoms has also made it possible to use machine learning and artificial intelligence to develop a smartphone app – Nuru AI – which allows untrained users to recognize CMD with a high degree of accuracy. Laboratory-based methods are required, however, to increase the accuracy of detection as well as to identify the various CMG species. ELISA was the first of these approaches to be widely applied to CMG detection. Monoclonal antibodies allowed a distinction to be made between ACMV, EACMV, and ICMV, at a time when only these three species were recognized. The emergence of the severe CMD epidemic in Uganda, however, highlighted the weakness with this serology-based method, as EACMV-UG was misdiagnosed as ACMV by ELISA, since EACMV-UG was a recombinant virus with the coat protein of ACMV. From the late 1990s onwards PCR was used almost exclusively for the detection and identification of CMG species. PCR presents an easy and robust detection system for CMGs and specific primer sets and a diverse array of PCR protocols are available to achieve reliable detection of these viruses in cassava. To verify specificity, sequencing of PCR amplicons confirms virus identity. However, because of recombination at various positions across DNA-A, amplification of genome fragments may not fully resolve the diversity of DNA-A and the association of sequence to a particular virus species. Hence amplification followed by sequencing of the entire DNA-A genome component is required for reliable virus identification. This is achieved by using inverse, abutting PCR primers located in a conserved region of DNA-A (CP) to amplify the entire circular DNA-A molecule. A more general approach is rolling-circle amplification (RCA) using phi29 (F29) DNA polymerase. Plant DNA including the circular virus DNA is amplified in an isothermal process using random oligonucleotide primers. Phi29 polymerase is highly processive and in this unique RCA process, small circular DNA molecules are replicated nearly infinitely. By enzymatic restriction of the RCA products using rarely-cutting restriction enzymes, CMG DNA fragments or complete virus genome components can be resolved and the entire begomovirus composition of a plant sample can be analyzed without prior knowledge of virus sequences. With this approach it is possible to detect unexpected and unknown begomoviruses, mixed infections and satellites. Sample preparation and DNA extraction are identical to other methods and commercial kits are available. RCA generates genome DNA sequences as concatemers. Virus identification can then be performed after restriction digestion, cloning and sequence analysis.

Economic Importance CMD symptoms of leaf chlorosis, reduction of leaf size and plant stunting lead to a reduction in the quantity of photosynthetic assimilates channeled into the tuberous roots, and through this, a reduction in yield. The degree of yield loss varies greatly, depending primarily on the susceptibility of the cassava cultivar, the virulence of the CMG species/strain, and the stage of growth at which infection occurred (being most severe for cutting-infected plants). Typically, individual plants sustain yield losses ranging from 20% to 100%. The only study of the effects of specific viruses on yield showed average yield losses to be 42% for ACMV alone, 12% for EACMV-UG mild, 68% for EACMV-UG severe, and 82% for mixed ACMV þ EACMV-UG. The relatively moderate losses attributable to ACMV infection, coupled with moderate to low incidences, have been the reasons for the apparent lack of concern through large parts of cassava-growing Africa about the impact of CMGs and CMD. This outlook changed markedly, however, following the emergence and regional spread of the CMD pandemic. Extensive surveys of CMD incidence, CMD severity, and the occurrence of the causal viruses have made it possible to make continent-wide yield loss estimates for CMD. The last of these (2006) provided estimates for losses in pandemic-recovered (16% loss), pandemic-affected (47%), and as yet unaffected (18%) countries in sub-Saharan Africa, leading to an overall loss figure of 34 million tons per year, which was equivalent to roughly a third of total African production of fresh cassava roots. Following the region-wide spread of the pandemic, major control programs were implemented, and anecdotal evidence suggests that this has resulted in reductions in CMD incidence and reduced losses. This declining impact of the disease in Africa is likely to be the reason why there have been no comprehensive assessments of losses since 2006. For CMD in India, losses of up to 84% have been recorded at field level, although these are said to vary, as in Africa, depending on the varietal response, type of infection and virus species/species mixture causing the infection. There are currently no published records of overall losses due to CMD in South Asia or for the more recently affected countries of Southeast Asia.

Control The most widely practiced approaches to controlling the CMGs that cause CMD include the deployment of host plant resistance and the use of cultural methods, particularly phytosanitation. More recently, considerable attention has been directed toward the use of genetic engineering techniques to produce transgenic virus-resistant cassava plants.

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Host Plant Resistance The potential value of introgressing virus resistance genes from wild relatives of cassava was recognized from the earliest days of CMD research in the 1920s/1930s, and inter-specific crosses combining cultivated cassava with Ceara rubber (Manihot glaziovii) were developed independently through breeding programs in modern-day Tanzania and Madagascar. F1 progeny were triple backcrossed with cultivated cassava to produce plants that combined acceptable food quality with significantly enhanced resistance to CMGs. Germplasm developed in this way formed the basis for the later continental breeding program run from the Nigeria-based International Institute of Tropical Agriculture (IITA), which from its establishment in 1967, to the present day, has developed thousands of CMD-resistant cassava clones. Many of these have been sent to cassava-producing countries in Africa for use either specifically for CMD management programs, or more generally for cassava development. Germplasm derived from the initial interspecific crosses uses the name prefix ‘TMS’ for ‘Tropical Manihot species’. This resistance source is multigenic and has provided high levels of resistance which have been very durable when used against all CMGs and CMG combinations. Four distinct mechanisms of resistance are recognized: resistance to infection, resistance to virus multiplication, resistance to virus movement (leading to incomplete systemicity), and resistance of normal plant function to the effects of virus infection. During the 1990s, resistant landraces from West Africa, given the name prefix ‘TME’ for ‘Tropical Manihot esculenta’, were incorporated into the breeding program. These have been shown to possess alternative sources of resistance, one of the most important of which has been characterized through genetic analyses as a single dominant gene designated CMD2. Molecular marker approaches have been used subsequently to combine the multigenic M. glaziovii-derived resistance with CMD2. In recent years, improvements in sequencing technologies have enabled researchers to undertake genome-wide association studies. These have been used to identify major QTLs associated with CMD resistance and they have provided new tools to improve the efficiency of the breeding process. Hundreds of CMG-resistant varieties have been released and disseminated throughout the cassava-growing areas of subSaharan Africa. However, their adoption by farming communities has been most widespread in CMD pandemic-affected countries, where they have provided the only effective means of restoring cassava production to pre-pandemic levels. In South Asia, sources of resistance have been drawn from the breeding work undertaken in Africa. Evaluations of seedling populations obtained from introduced germplasm have led to the development of varieties such as CMR1, CMR129, CR21–10, and CR43–11 which combine high starch content with strong resistance to ICMV and SLCMV. Some of this germplasm, together with other CMD-resistant varieties from IITA in Nigeria are currently being introduced to Southeast Asia as part of regional efforts to control the CMD outbreak there.

Cultural Methods A range of cultural methods has been proposed for the control of CMGs. The methods most widely recommended have been the removal of infected plants (roguing) or the selection of disease-free planting material for the establishment of a new crop (selection). Crop isolation, adjusting crop disposition in relation to the prevailing wind, varying planting date, varietal mixtures, and intercropping cassava with other ‘putative’ protective crops have all been suggested at various times as potentially useful control options for CMGs. No convincing experimental evidence has yet been presented to confirm the value of any of these methods, however, and current field practice is restricted to selection and occasional roguing. Roguing is considered to be of value within the framework of institutional programs for the multiplication of CMD-resistant germplasm, in view of the requirement for the production of high-quality planting material. Experiments conducted in ‘post-epidemic’ areas of East Africa, first affected by the CMD pandemic five or more years previously, have provided clear evidence for the value of selection of CMD-free stems when choosing planting material. Local cultivars treated in this way provided equivalent yields to those of CMD-resistant varieties after two cropping cycles. A key drawback to the wider adoption of this approach, however, is the variability in effectiveness of the approach in relation both to the virus inoculum pressure of the location, as well as the relative susceptibility of the cultivar. Systems for the phytosanitary control of CMGs have been greatly strengthened in several parts of Africa in recent years. Most notably, certification guidelines for cassava have been developed and appended to Seed Acts in countries such as Nigeria, Rwanda and Tanzania. These guidelines include maximum tolerance levels for CMD incidence. This has facilitated the certification of early generation cassava ‘seed’ ( ¼ planting material) and promoted the production of near virus-free planting material. The approach is helping to ensure that newly developed virus-resistant varieties are multiplied and disseminated in a healthy condition and it is reducing overall incidences of CMD.

Virus Resistance by Engineering Engineering virus resistance by expression of virus sequences in transgenic plants is a strategy that is also used to achieve resistance against gemininiviruses. The most common approach is to engineer RNAi (RNA interference) constructs as inverted-repeats of virus sequence which are spaced by an intron. Splicing of the intron upon transcription generates a “hairpin” dsRNA that is targeted by the plants RNA silencing machinery. RNAi triggered antiviral defense using constructs comprising Rep sequences is effective against Tomato yellow leaf curl virus (TYLCV) and against Bean golden mosaic virus (BGMV). The stability of this resistance was verified in field trials showing virus resistance under field conditions for several seasons and the feasibility of the

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approach to provide protection in both crops. In Brazil, transgenic common beans resistant to BGMV were released in 2011 as a commercial variety. Thus RNAi strategies may protect plants against DNA viruses as shown also for other crops including cotton as well as cassava, by boosting the antiviral defense. The effectiveness of the RNAi protection depends on the choice of the viral sequence and the abundance of small interfering RNAs generated from the hairpin RNA transgene. In addition, temperature and the intrinsic interaction of virus and plant host influence the effectiveness of silencing and the level of resistance achieved. RNAi strategies have been used to induce gene silencing against cassava infecting geminiviruses. In all approaches, symptoms were attenuated and virus DNA accumulation was drastically reduced. Broad spectrum resistance against several CMGs e.g., ACMV, EACMCV, and SLCMV can be achieved when there is sufficient sequence homology between the siRNAs and transcripts of the viruses. Several studies have shown the effectiveness of CRISPR-Cas9 to engineer resistance against the geminiviruses Tomato yellow leaf curl virus (TYLCV) and Beet curly top virus (BCTV). A single-guide RNA (sgRNA) carrying the TAATATTAC nonanucleotide origin of replication of geminiviruses was used to induce a broad spectrum virus resistance. In cassava, CRISPR-Cas9 was used to engineer resistance to ACMV targeting AC2 and AC3 with a single sgRNA. However, CRISPR-Cas9 did not show any difference to the wild-type susceptible plants because of the emergence of a cleavage mutant of ACMV in transgenic plants. The selection of cleavage resistant mutants, as shown for ACMV, or new virus mutants that evade the sequence-specific action of transgene derived RNAi in TYLCV transgenic plants or ACMV transgenic cassava demonstrate the challenges of any of the engineering strategies employing targeting sequences of a replicating virus. However, this can be addressed by using multiplexed CRISPR/Cas9 with multiple sgRNAs to target virus complexes, as shown for the control of Cotton leaf curl virus (CLCuV). With more information on functional networks and genes involved in vital virus functions, CRISPR-Cas9, RNAi or similar approaches will be increasingly used for the editing or interfering with expression of host genes to achieve virus resistance in cassava.

Biosafety considerations Cassava plants transformed for resistance to CMGs have been tested under field conditions in several African countries (Kenya, Nigeria, Uganda) through confined field trial arrangements, with promising results confirming the effectiveness of the strategy. However, uncertainty in most African countries about the consequences of the widespread cultivation of GMOs has slowed the adoption of biosafety guidelines and meant that at the time of writing (2019) there are no transgenic cassava plants that have been evaluated under ‘real-world’ conditions in farmers’ fields. In January 2019, however, the National Biosafety Management Agency of Nigeria made its first approval of a GM crop – podborer-resistant cowpea. Since Nigeria is the world’s largest producer of cassava, and since its cassava crop is widely affected by CMD, this development offers significant promise for the future release of GM CMD-resistant cassava varieties. Furthermore, this is a move that could provide a signal for other countries similarly affected by cassava viruses to follow suit. Given the huge impact that CMD has on the production of one of the world’s most important staple food crops, the widespread adoption of science-based strategies to control may be essential if the food security of hundreds of millions of cassava-dependent households in Africa and elsewhere is to be assured.

See also: Bean Golden Mosaic Virus and Bean Golden Yellow Mosaic Virus (Geminiviridae). Beet Curly Top Virus (Geminiviridae). Cassava Brown Streak Viruses (Potyviridae). Cotton Leaf Curl Disease (Geminiviridae). Emerging Geminiviruses (Geminiviridae). Geminiviruses (Geminiviridae). Maize Streak Virus (Geminiviridae). Plant Resistance to Geminiviruses. Tomato Leaf Curl New Delhi Virus (Geminiviridae). Tomato Yellow Leaf Curl Viruses (Geminiviridae)

Further Reading Bock, K.R., Woods, R.D., 1983. The etiology of African cassava mosaic disease. Plant Disease 67, 994–995. Dubern, J., 1994. Transmission of African cassava mosaic geminivirus by the whitefly (Bemisia tabaci). Tropical Science 34, 82–91. Dutt, N., Briddon, R.W., Dasgupta, I., 2005. Identification of a second begomovirus, Sri Lankan cassava mosaic virus, causing cassava mosaic disease in India. Archives of Virology 150 (10), 2101–2108. Fauquet, C., Fargette, D., 1990. African cassava mosaic virus: Etiology, epidemiology and control. Plant Disease 74, 404–411. Fondong, V.N., 2017. The search for resistance to cassava mosaic geminiviruses: How much we have accomplished, and what lies ahead. Frontiers in Plant Science 8, 408. doi:10.3389/fpls.2017.00408. Hanley-Bowdoin, L., Bejarano, E.R., Robertson, D., Mansoor, S., 2013. Geminiviruses: Masters at redirecting and reprogramming plant processes. Nature Reviews Microbiology 11, 777–788. Harrison, B.D., Zhou, X., Otim-Nape, G.W., Liu, Y., Robinson, D.J., 1997. Role of a novel type of double infection in the geminivirus-induced epidemic of severe cassava mosaic in Uganda. Annals of Applied Biology 131, 437–448. Hipp, K., Grimm, C., Jeske, H., Bottcher, B., 2017. Near-atomic resolution structure of a plant geminivirus determined by electron cryomicroscopy. Structure 25, 1303–1309. Legg, J.P., Lava Kumar, P., Makeshkumar, T., et al., 2015. Cassava virus diseases: Biology, epidemiology and management. Advances in Virus Research 91, 85–142. Legg, J.P., Owor, B., Sseruwagi, P., Ndunguru, J., 2006. Cassava mosaic virus disease in East and Central Africa: Epidemiology and management of a regional pandemic. Advances in Virus Research 67, 355–418. Mehta, D., Sturchler, A., Anjanappa, R.B., Zaidi, S.S., Hirsch-Hoffmann, M., Gruissem, W., Vanderschuren, H., 2019. Linking CRISPR-Cas9 interference in cassava to the evolution of editing-resistant geminiviruses. Genome Biology 20, 80. Otim-Nape, G.W., Bua, A., Thresh, J.M., et al., 1997. Cassava Mosaic Virus Disease in Uganda: The Current Pandemic and Approaches to Control. Chatham, UK: Natural Resources Institute, p. 65. Swanson, M.M., Harrison, B.D., 1994. Properties, relationships and distribution of cassava mosaic geminiviruses. Tropical Science 34, 15–25.

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Thresh, J.M., Otim-Nape, G.W., Thankappan, M., Muniyappa, V., 1998. The mosaic diseases of cassava in Africa and India caused by whitefly-borne geminiviruses. Review of Plant Pathology 77, 935–945. Wang, H.L., Cui, X.Y., Wang, X.W., et al., 2016. First report of Sri Lankan cassava mosaic virus infecting cassava in Cambodia. Plant Disease 100, 1029.

Relevant Websites https://gd.eppo.int/taxon/ACMV00 African cassava mosaic virus (ACMV00)[Overview] EPPO Global Database. https://www.cabi.org/isc/datasheet/2747 Cassava mosaic disease (African cassava mosaic) CABI. https://www.youtube.com/watch?v=rY8xyO44Z18 Cassava Mosaic Disease (CMD) in Southeast Asia. https://ciat.cgiar.org/event/cmd-sea/ Cassava Mosaic Disease (CMD) in Southeast Asia CIAT. https://www.accessagriculture.org/cassava-mosaic-virus Cassava mosaic virus Access Agriculture. https://www.sciencedirect.com/topics/agricultural-and-biological-sciences/cassava-mosaic-virus Cassava Mosaic Virus ScienceDirect.com. https://www.dsmz.de DSMZ (homepage). https://talk.ictvonline.org/files/master-species-lists/m/msl/8266 ICTV Master Species List 2018b.v2 International Committee on Taxonomy of Viruses (ICTV). https://talk.ictvonline.org/ International Committee on Taxonomy of Viruses (ICTV). https://www.plantvillage.psu.edu PlantVillage Penn State. https://www.iita.org The International Institute of Tropical Agriculture (IITA).

Caulimoviruses (Caulimoviridae) James E Schoelz, University of Missouri, Columbia, MO, United States Mustafa Adhab, University of Baghdad, Baghdad, Iraq r 2021 Elsevier Ltd. All rights reserved. This is an update of T. Hohn, Caulimoviruses: Molecular Biology, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00367-8.

Nomenclature

kDa Kilo dalton MP Movement protein nt Nucleotide(s) ORF Open Reading Frame TAV Translational transactivator

aa Amino acid(s) CP Coat protein or capsid protein dsDNA Double stranded DNA IBs Inclusion bodies kb Kilobase

Glossary Agro-inoculation An inoculation technique in which Agrobacterium tumefaciens is used to deliver a full-length infectious clone of a virus into a plant cell. Plasmodesmata A narrow channel of cytoplasm that functions as a bridge between two plant cells to facilitate movement of macromolecules. Reverse transcriptase An enzyme that utilizes an RNA template for synthesis of DNA.

Ribosome shunt A translational mechanism in which ribosomes enter the RNA at the 50 end and scan for a short distance before being translocated to a downstream point. Semi-persistent transmission Vector acquires the virus in minutes to hours, and can transmit to other plants for hours after the initial feeding. Transgene A gene introduced into an organism through any one of a number of genetic engineering techniques.

Introduction The members of the family Caulimoviridae are plant viruses that replicate by reverse transcription of an RNA intermediate and whose virions contain circular, double stranded DNA. They belong to the order Ortervirales, a virus order that encompasses five families of reverse transcribing viruses. Although caulimoviruses replicate by reverse transcription, integration into host chromosomes is not required for completion of their replication cycle. The circular DNA encapsidated in virions is not covalently closed, as it contains at least one discontinuity in each DNA strand, and these discontinuities occur as a consequence of the replication strategy of the virus. There are eight genera in this family and they can be divided into two groups based on virion morphology; the members of the genera Caulimovirus, Petuvirus, Cavemovirus, Rosadnavirus, Solendovirus, and Soymovirus contain viruses that form icosahedral particles that are largely found within amorphous inclusion bodies in the cell (Fig. 1(A) and (B)). In contrast, the members of the genera Badnavirus and Tungrovirus form bacilliform particles that are not associated with inclusion bodies (Fig. 1(C)). Their virions are found in the cytoplasm either individually or clustered in palisade-like arrays. Cauliflower mosaic virus (CaMV), the type species of the genus Caulimovirus, was the first of the plant viruses to be shown to contain double-stranded DNA (dsDNA) in its icosahedral virion in 1968. This discovery led to an extended investigation into its replication strategy throughout the 700 s and early 800 s, culminating in 1983 when it was shown that CaMV replicated through reverse transcription of an RNA intermediate. Perhaps because the genome of CaMV is composed of dsDNA, it was also the first of the plant viruses to be completely sequenced and cloned into bacterial plasmids in an infectious form. In the early 1980s, CaMV was thought to have some promise as a vector for foreign genes in plants. However, the effort to convert CaMV into a vector was scaled back when it was shown that the virus genome could tolerate only small insertions of up to a few hundred base pairs of DNA. A few small genes, such as dihydrofolate reductase and interferon, were eventually expressed in plants via a CaMV vector, but other viruses have been shown to be much more versatile as vectors for foreign genes. Although the caulimoviruses have had only limited utility as plant virus vectors, they continue to have a great impact on plant biotechnology. The 35S promoter of CaMV is capable of directing a high level of transcription in most types of plant tissues. This promoter was used to drive expression of one of the first transgenes introduced into transgenic plants, and it is still widely used for expression of transgenes for both research and commercial applications. The promoter regions of several other caulimoviruses have also been evaluated for expression of transgenes in both monocots and dicots, and they can be good alternatives to the CaMV 35S promoter.

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Fig. 1 Inclusion bodies and virions of members of the family Caulimoviridae. (A). Amorphous inclusion body of CaMV (I) adjacent to a chloroplast (Chl). Individual virions can be seen in the vacuolated regions of the inclusion body. (B). Icosahedral virions of CaMV visualized within an inclusion body. (C). Purified bacilliform virions of CoYMV. CoYMV photo courtesy of Ben Lockhart (University of Minnesota).

For many years, only plant viruses that had icosahedral virions of 50 nm in diameter were thought to have genomes composed of dsDNA. However in 1990, it was shown that the bacilliform virions of Commelina yellow mottle virus (CoYMV) contained circular, dsDNA, and Commelina yellow mottle virus became the type species for the genus Badnavirus. Nucleic acid isolated from the bacilliform virions was resistant to RNase and degraded by DNase. Furthermore, DNA treated with S1 nuclease revealed that the CoYMV genome contained at least two single-stranded discontinuities, a hallmark of the plant viruses that replicate by reverse transcription. In addition to CoYMV, DNA purified from Banana streak virus (BSV), Kalanchoe top-spotting virus (KTSV) and Canna yellow mottle virus, providing evidence that they should also be placed in the new genus Badnavirus. Currently, there are 57 species in the Badnavirus genus (Table 1), many of which cause economically important diseases in the tropics. Soon after the discovery that CoYMV was a DNA virus, the bacilliform component of rice tungro disease was also shown to contain circular, dsDNA. Rice tungro is the most important virus disease of rice in South and Southeast Asia, with annual losses approaching $1.5 billion dollars. The disease is caused by a complex of an RNA virus, called Rice tungro spherical virus (RTSV) coupled with the bacilliform dsDNA virus called Rice tungro bacilliform virus (RTBV). The genus Tungrovirus is distinguished from the genus Badnavirus because RTBV has one more open reading frame (ORF) than CoYMV and overall, the RTBV genome has only 20%–25% nt sequence identity with members of the genus Badnavirus. There is only one species in the genus Tungrovirus, although several isolates of the species Rice tungro bacilliform virus have been collected and sequenced.

Taxonomy, Phylogeny, and Evolution The family Caulimoviridae consists of eight genera (Table 1), and they can be conveniently divided into two groups based on virion morphology. Viruses with icosahedral virions (Fig. 1(B)) include the genera Caulimovirus, Petuvirus, Soymovirus, Rosadnavirus, Solendovirus, and Cavemovirus. Viruses with bacilliform virions (Fig. 1(C)) include the genera Badnavirus and Tungrovirus. Genera can be further distinguished because of differences in genome organization (Fig. 2(A)) as well as phylogenetic analyses of proteins such as the reverse transcriptase, as illustrated in Fig. 2(B). All members of the family Caulimoviridae infect only plants. There are no animal or insect viruses in this family. Complete and partial nt sequences of members of the family Caulimoviridae are listed in Table 1. Only a single accession number is given for CaMV and RTBV, although multiple strains of each have been sequenced.

Virion Structure The viruses in the genera Caulimovirus, Cavemovirus, Soymovirus and Petuvirus form non-enveloped, isometric particles that vary in size from 43 to 50 nm. The virion is composed of 420 subunits with a T ¼ 7 structure. The viruses in the genera Badnavirus and Tungrovirus form non-enveloped, bacilliform particles that are 30 nm in width, but can vary in length from 60 to 900 nm. The length most commonly observed is 130 nm. Their structure is based on an icosahedron, in which the ends are formed from pentamers and the tubular section is made up of hexamers. The capsid also is loosely decorated with the 15 kDa virus-associated protein, which is encoded by ORF-III. The virions typically accumulate in amorphous inclusion bodies, which are composed primarily of a 62 kDa multifunctional protein encoded by ORF-VI. The virions of the family Caulimoviridae contain a single, circular dsDNA that is 7.2–9.3 kbp in length.

Caulimoviruses (Caulimoviridae)

Table 1

315

Virus genera and virus species members in the family Caulimoviridae

Genus

Virus

Acronym

Host (Family)

Distribution

Accession #

Caulimovirus

Cauliflower mosaic virusa Angelica bushy stunt virus Atractylodes mild mottle virus Carnation etched ring virus Dahlia mosaic virus Figwort mosaic virus Horseradish latent virus Lamium leaf distortion Mirabilis mosaic virus Rudbeckia flower distortion virus Soybean Putnam virus Strawberry vein banding virus Thistle mottle virus

CaMV AnBSV AMMV CERV DMV FMV HRLV LLDV MiMV RuFDV SPuV SVBV ThMoV

Brassica sp. (Crucifereae) Angelica (Apiaceae) Atractylodes (Compositae) Carnation (Caryophyllaceae) Dahlia (Compositae) Figwort (Scrophulariaceae) Horseradish (Crucifereae) Lamium (Lamiaceae) Mirabilis (Nyctaginaceae) Rudbeckia (Asteraceae) Soybean (Leguminosae) Strawberry (Rosaceae) Thistle (Compositae)

Worldwide South Korea South Korea Worldwide Worldwide U.S.A. Denmark U.S.A. U.S.A. U.S.A. U.S.A. Worldwide Europe

V00140 KU508800 KR080327 X04658 KC967481 X06166 JX429923 EU554423 AF454635 NC_011920 JQ926983 X97304

Petuvirus

Petunia vein clearing virus

PVCV

Petunia (Solanaceae)

Worldwide

U95208

Soymovirus

Soybean chlorotic mottle virus Blueberry red ringspot virus Cestrum yellow leaf curling virus Peanut chlorotic streak virus

SbCMV BRRSV CmYLCV PCSV

Soybean (Leguminosae) Blueberry (Ericaceae) Cestrum sp. (Solanaceae) Peanut (Leguminosae)

Japan U.S.A. Italy India

X15828 AF404509 AF364175 U13988

Cavemovirus

Cassava vein mosaic virus Sweet potato collusive virus

CsVMV SPCV

Cassava (Euphorbiaceae) Sweet potato (Convolvulaceae)

Brazil Worldwide

U59751

Rosadnavirus

Rose yellow vein virus

RYVV

Rose (Rosaceae)

U.S.A.

JX028536

Solendovirus

Tobacco vein clearing virus Sweet potato vein clearing virus

TVCV SPVCV

Nicotiana sp. (Solanaceae) Sweet potato (Convolvulaceae)

Worldwide Africa, Americas

AF190123 NC_015228

Badnavirus

Commelina yellow mottle virus Aglaonema bacilliform virus Banana streak GF virus Banana streak IM virus Banana streak Mysore virus Banana streak OL virus Banana streak UA virus Banana streak UI virus Banana streak UL virus Banana streak UM virus Banana streak VN virus Birch leafroll-associated virus Blackberry virus F Bougainvillea chlorotic vein banding virus Cacao bacilliform Sri Lanka virus Cacao mild mosaic virus Cacao swollen shoot CD virus Cacao swollen shoot CE virus Cacao swollen shoot Ghana M virus Cacao swollen shoot Ghana N virus Cacao swollen shoot Ghana Q virus Cacao swollen shoot Togo A virus Cacao swollen shoot Togo B virus Cacao yellow vein banding virus Canna yellow mottle associated virus Canna yellow mottle virus Citrus yellow mosaic virus Dioscorea bacilliform AL virus Dioscorea bacilliform AL virus 2 Dioscorea bacilliform ES virus Dioscorea bacilliform RT virus 1 Dioscorea bacilliform RT virus 2 Dioscorea bacilliform SN virus Dioscorea bacilliform TR virus

ComYMV ABV BSGFV BSIMV BSMyV BSOLV BSUAV BSUIV BSULV BSUMV BSVNV BLRaV BVF BsCVBV CBSLV CaMMV CSSCDV CSSCEV CSSGMV CSSGNV CSSGQV CSSTAV CSSTBV CYVBV CaYMAV-1 CaYMV CiYMV DBV DBALV 2 DBESV DBRTV1 DBRTV2 DBSNV DBTRV

Commelina (Commelinaceae) Aglaonema sp. (Araceae) Banana (Musaceae) Banana (Musaceae) Banana (Musaceae) Banana (Musaceae) Banana (Musaceae) Banana (Musaceae) Banana (Musaceae) Banana (Musaceae) Banana (Musaceae) Birch (Betulaceae) Blackberry (Rubus) Bougainvillea (Nyctaginaceae) Cacao (Malvaceae) Cacao (Malvaceae) Cacao (Malvaceae) Cacao (Malvaceae) Cacao (Malvaceae) Cacao (Malvaceae) Cacao (Malvaceae) Cacao (Malvaceae) Cacao (Malvaceae) Cacao (Malvaceae) Canna (Cannaceae) Canna (Cannaceae) Citrus sp. (Rutaceae) Yam (Dioscoreaceae) Yam (Dioscoreaceae) Yam (Dioscoreaceae) Yam (Dioscoreaceae) Yam (Dioscoreaceae) Yam (Dioscoreaceae) Yam (Dioscoreaceae)

Caribbean Southeast Asia Worldwide Worldwide Worldwide Worldwide Worldwide Worldwide Worldwide Worldwide Worldwide Europe U.S.A. Worldwide Sri Lanka Trinidad Africa Africa Africa Africa Africa Africa Africa Trinidad Worldwide Worldwide. India Africa Africa Africa Africa Africa Africa Worldwide

X52938 AY493509 KJ013508 AY805074 AJ002234 NC_015502 HQ593108 HQ593109 HQ93110 NC_007003 MG686420 KJ413252 NC_011592 MF642736 KX276640 JN606110 MF642719 MF642724 MF642725 MF642728 AJ781003 L14546 KX276641 NC_030462 NC_038380 NC_003382 X94576 KY827395 KY827394 KX008574 KX008577 NC_009010 KX430257 (Continued )

316

Caulimoviruses (Caulimoviridae)

Table 1

Continued

Genus

Virus

Acronym

Tungrovirus

Fig badnavirus 1 FBV1 Gooseberry vein banding associated virus GVBAV Grapevine roditis leaf discoloration associated virus GRLDaV Grapevine vein clearing virus GVCV Jujube mosaic-associated virus JuMAV Kalanchoe top-spotting virus KTSV Mulberry badnavirus 1 MBV1 Pagoda yellow mosaic associated virus PYMAV Pineapple bacilliform CO virus PBCOV Pineapple bacilliform ER virus PBERV Piper yellow mottle virus PYMoV Rubus yellow net virus RYNV Schefflera ringspot virus SRV Spiraea yellow leaf spot virus SYLSV Sugarcane bacilliform Guadeloupe A virus SCBGAV Sugarcane bacilliform Guadeloupe D virus SCBGDV Sugarcane bacilliform IM virus SCBIMV Sugarcane bacilliform MO virus SCBMOV Sweet potato pakakuy virus SPPV Taro bacilliform CH virus TaBCHV-1 Taro bacilliform virus TaBV Wisteria badnavirus 1 WBV1 Yacon necrotic mottle virus YV1 Rice tungro bacilliform virus RTBV

Host (Family)

Distribution

Accession #

Fig (Moraceae) Ribes sp. (Grossulariaceae) Grapes (Vitaceae) Grapes (Vitaceae) Chinese Jujube (Rhamnaceae) Kalanchoe (Crassulaceae) Mulberry (Moraceae) Pagoda tree (Fabaceae) Pineapple (Bromeliaceae) Pineapple (Bromeliaceae) Pepper (Piperaceae) Raspberry (Rosaceae) Schefflera (Araliaceae) Spiraea (Rosaceae) Sugarcane (Poaceae) Sugarcane (Poaceae) Sugarcane (Poaceae) Sugarcane (Poaceae) Sweet potato (Convolvulaceae) Taro (Araceae) Taro (Araceae) Wisteria (Fabaceae) Yacon (Asteraceae) Rice (Poaceae)

Worldwide Worldwide Greece U.S.A. China Worldwide Lebanon China Worldwide Worldwide Brazil, India, Asia Eurasia, U.S.A. Worldwide Worldwide West Indies West Indies U.S.A., Morocco U.S.A., Morocco Worldwide China South Pacific China Korea East Asia, China

JF411989 AF298883 HG940503 NC_015784 NC_035472 AY180137 NC_026020 NC_024301 GQ398110 EU377672 AF468454 AF299074 FJ824814 FJ439817 AJ277091 M89923 NC_015655 KP10178 AF357836 KX168422 KM229702 X57924

a

The type species of each genus is highlighted in bold font.

Genome Organization Several genome features are common to all members of the family Caulimoviridae, in addition to their circular dsDNA genomes. For example, the dsDNA is not covalently closed; it has at least two discontinuities, but may have up to four. The most significant difference among the caulimoviruses concerns the arrangement and number of ORFs (Fig. 2). All of the ORFs are found on only one of the DNA strands and all of the viruses have at least one strong promoter that drives the expression of a terminally redundant mRNA. In addition, all of the viruses have a reverse transcriptase, and in most cases, it is located downstream from the coat protein (CP). Several active sites can be identified within the reverse transcriptase protein, including a protease, the core reverse transcriptase, and RNaseH activity. The reverse transcriptase of PVCV is distinguished from all other members of the family Caulimoviridae because it has the core features of an integrase function, in addition to other functions. The genome organization of CaMV is the best characterized of the family and it is discussed in detail here. The CaMV genome is approximately 8000 bp in size and it contains three single-stranded discontinuities. One discontinuity occurs in the negative sense DNA strand and by convention, this is the origin of the DNA sequence. Two other discontinuities occur in the positive sense strand, one at nt position 1600 and a second at approximately nt position 4000. The virus genome consists of seven ORFs (Fig. 2(A)). ORF1 encodes a protein necessary for cell-to-cell movement (P1). ORFs 2 and 3 (proteins P2 and P3) are both required for aphid transmission. P3 also has an additional role in cell-to-cell movement. ORF4 encodes the CP, and ORF5 encodes a reverse transcriptase that also has protease and RNaseH domains. With the exception of Petunia vein clearing virus (PVCV), none of the reverse transcriptases of the caulimoviruses have evidence for an integrase function. The ORF6 product (P6) was originally described as the major inclusion body protein, but it also has a function as a translational transactivator (TAV). It physically interacts with host ribosomes to reprogram them for reinitiation of translation of the polycistronic 35S RNA. Furthermore, P6 is an important symptom and host range determinant. No protein product has been found for ORF7. Its nt sequence appears to play a regulatory role in aligning ribosomes for translation of other CaMV gene products. Two transcripts are produced from the CaMV genomic DNA. The 19S RNA serves as the mRNA for P6, whereas the terminally redundant 35S RNA serves as a template for reverse transcription and as a polycistronic mRNA for ORFs 1–5. One feature common to many of the caulimoviruses is the ribosomal shunt mechanism of translation. The ribosomal shunt has undoubtedly evolved over time to compensate for the complexity and length of the leader sequence of the genomic RNA. In the case of CaMV, this leader sequence is approximately 600 bp in length and it contains up to nine short ORFs that vary in size from 9 to 102 nt. The complexity of the CaMV 35S RNA leader sequence is bypassed through the formation of a large stem-loop structure, which allows ribosomes to bypass most of the leader and to initiate translation at ORF7. CaMV utilizes several other strategies for expression of the 35S RNA, including splicing and reprogramming of host ribosomes by the TAV for re-initiation of translation.

Caulimoviruses (Caulimoviridae)

317

Fig. 2 Genome structure and phylogeny of members of the family Caulimoviridae. (A). Genome structure of representatives viruses of the eight genera that comprise the family Caulimoviridae. Open reading frames (ORFs) are indicated by boxes, and are identified according to the information in the GeneBank nucleotide accession. The ORFs common to all genera include a movement protein (yellow), a capsid protein (green), and a protease/reverse transcriptase (purple). ORFs unique to a virus or whose function remains unidentified are in gray. The slashes on either end of the diagram (/) indicate where the genome is circular. The first base of the nucleotide sequence deposited in GenBank is located on the left, and the size of the genome is located on the right. Virus genera including Petuvirus, Cavemovirus, Tungrovirus, and Badnavirus produce polyproteins, which are presented as a single ORF. (B). Evolutionary relationships of the members of the family Caulimoviridae, based on the amino acid sequences of the reverse transcriptases. The evolutionary history was inferred using the Neighbor-Joining method. The optimal tree with the sum of branch length ¼ 3.32101427 is shown. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (500 replicates) are shown next to the branches. The evolutionary distances were computed using the p-distance method and are in the units of the number of amino acids differences per site. All ambiguous positions were removed for each sequence pair (pairwise deletion option). There was a total of 2020 positions in the final dataset. Evolutionary analyses were conducted in MEGA X. The outgroup for comparison is hepatitis B virus strain A (HBV-A).

Properties and Functions of CaMV Gene Products Movement Protein (MP) CaMV ORF-I encodes a 37 kDa protein dedicated to cell-to-cell movement. The CaMV movement protein (MP) dramatically reconfigures the host plasmodesma by removal of the desmotubule, leading to the formation of a tubule structure though which the virions move to adjacent cells. The MP is delivered to the plasma membrane through a vesicular transport pathway where it

318

Caulimoviruses (Caulimoviridae)

makes its way to the plasmodesma. Excess MP is thought to be recycled back to the central vacuole. By contrast, the CaMV virion is thought to be delivered separately to the tubules by inclusion bodies formed from the P6 protein. Other plant viruses whose MPs are capable of forming tubules include the tospoviruses, comoviruses, nepoviruses and bromoviruses.

Aphid Transmission Factor (P2) The product of ORF-II (or P2) is an 18 kDa protein that is required for transmission by aphids. P2 exists in cells in the form of inclusion bodies. When an aphid probes a leaf, this triggers the release of CaMV virions from the P6 inclusion bodies, the dissolution of the P2 inclusion bodies and the distribution of the P2 protein on host microtubules, where they come into contact with the virions. The N-terminus of the P2 protein is capable of binding to the aphid stylet, whereas the C-terminus of P2 binds to the CaMV P3 protein, which is located on the surface of virions. Consequently, CaMV virions are acquired by aphids through an association with the P2 and P3 proteins. Inactivation of the P2 protein abolishes aphid transmission but the mutant virus remains fully infectious through mechanical inoculation.

Virion-Associated Protein (P3) The product of ORF-III, a 15 kDa virion-associated protein (P3) has key roles in both cell-to-cell movement and aphid transmission. P3 is found on the surface of virions and functions as a bridge between the virion and the MP and P2 proteins. The N-terminus of P3 binds to either to MP or to P2 to facilitate cell-to-cell movement or aphid transmission, respectively. Although P3 is located on the surface of virions, it is not uncommon for it to be released during purification of CaMV virions.

Capsid Protein (P4) ORF-IV encodes the main capsid component (P4) has a molecular weight of 56 kDa and is processed into sizes of 37 and 44 kDa. The p37 protein is included within p44 and antibodies raised against virions react against both proteins. The p44 protein is phosphorylated by host casein kinase II, whereas p37 is not phosphorylated. Both proteins contain a Zn-finger motif, which is implicated in RNA binding. It is not clear which of p37 and p44 is the mature capsid protein and if one or both are required for full infection.

Protease/Reverse Transcriptase (P5) The CaMV P5 protein is a polyprotein which is cleaved into a 15 kDa aspartic protease on the N-terminus and a 60 kDa reverse transcriptase/RNase H. With the exception of Petunia vein clearing virus, the reverse transcriptases of the caulimoviruses do not have an integrase function and do not integrate into host chromosomes as part of their replication cycle. Nonetheless, caulimovirus nt sequences are frequently found in the genomes of plants due to illegitimate recombination events.

Transactivator/Viroplasmin (P6) The P6 protein of CaMV has a size of 62 kDa and is a target for plant defenses and it plays a key role in multiple steps of the viral replication cycle. P6 is the matrix protein for the formation of the electron dense inclusion bodies (IBs) (Fig. 1(A)), which are the site for translation of the polycistronic 35S RNA, reverse transcription of the 35S RNA into DNA, and encapsidation of the viral DNA into virions. Nearly all of the CaMV virions accumulate and are retained within the P6 IBs, and this has led to its designation as a “virion factory”. The role of P6 in translation is especially noteworthy, because the P6 protein interacts with and modifies host ribosomes to facilitate translation of the 35S RNA, essentially reprogramming eukaryotic ribosomes to deal with a polycistronic message. This function, designated the translational transactivator (TAV), is unique in nature. New functions have recently been discovered for the P6 protein, including (1) movement of P6 inclusion bodies on microfilaments for delivery of virions to modified plasmodesmata for transit to adjacent cells, (2) modification of host defenses associated with SA and JA pathways, and (3) functioning as a gene silencing suppressor. Furthermore, P6 is an important pathogenic determinant. In susceptible hosts, P6 has been shown to be a prominent symptom determinant, responsible for the chlorosis symptom in turnips and a systemic necrosis symptom in Nicotiana clevelandii. P6 has also been shown to be a target for host resistance proteins present in Nicotiana species and Arabidopsis; recognition of the P6 protein by a host resistance protein triggers a cascade of plant defense responses that results in limitation of spread of CaMV. To date, P6 has been shown to physically interact with 14 host or viral proteins to carry out such diverse roles as translational transactivation, intracellular movement and silencing suppression.

Replication and Propagation Caulimoviruses replicate by reverse transcription of an RNA intermediate, but integration of the viral DNA into host chromosomes is not required to complete the replication process. Fig. 3 illustrates the replication of an icosahedral caulimovirus, but the same steps

Caulimoviruses (Caulimoviridae)

319

Fig. 3 Replication strategy of the members of the family Caulimoviridae.

are applicable for the bacilliform viruses. After virions enter a plant cell, the viral DNA becomes un-encapsidated and is transported into the nucleus. The viral DNA contains two to four single-stranded discontinuities, which form as a consequence of the reverse transcription process. Once in the nucleus, the single-stranded discontinuities are covalently closed and the DNA is associated with histones to form a minichromosome. The host RNA polymerase II is responsible for synthesizing an RNA that is terminally redundant; the sequence on the 50 end is reiterated on the 30 end. In the case of CaMV, the terminal redundancy is 180 nt in size, whereas the terminal redundancy in the CoYMV RNA is 120 nt. The terminally redundant RNA is transported out of the nucleus into the cytoplasm where it can either serve as a template for translation of viral proteins or as a template for reverse transcription. Reverse transcription is thought to occur in nucleocapsid-like particles. First strand DNA synthesis is primed by a methionine (Met) tRNA that binds to a complementary sequence near the 50 end of the terminally redundant RNA. In the case of CaMV, the Met tRNA binds to a sequence approximately 600 nt from the 50 end of its 35S RNA. The virally encoded reverse transcriptase synthesizes DNA up to the 50 end of the terminally redundant RNA and the RNase H activity of the reverse transcriptase degrades the 50 end of the RNA. The terminal redundancies present in the genomic RNA provide a mechanism for a template switch, as the reverse transcriptase is able to switch from the 50 end of the genomic RNA to the same sequences on the 30 end of the genomic RNA and continue to synthesize the first strand of DNA. The RNase H activity of the reverse transcriptase degrades the viral RNA template and small RNA fragments serve as the primers for second strand DNA synthesis. The RNA fragments bind to guanosine-rich tracts present in the first DNA strand, and these priming sites determine the positions of the discontinuities in the second DNA strand. A second template switch is required to bridge the gap from the 50 to the 30 end of the first DNA strand. The completion of the second DNA strand results in the formation of a dsDNA molecule that contains the characteristic single-stranded discontinuities. Although caulimoviruses do not integrate into the host as part of their replication strategy, the sequences of several viruses have been detected in the genomes of their hosts. Furthermore, the integrated copies of several of these viruses, the Banana streak species, Tobacco vein clearing virus (TVCV) and PVCV, can be activated to yield episomal infections. All three viruses can form virions once episomal infections are initiated, but they differ in their capacity to be transmitted to other plants. Banana streak virus can be transmitted by mealybugs, whereas PVCV is transmitted only by grafting. TVCV and PVCV are transmitted vertically, through seed, but this probably involves only the integrated copies of the virus rather than the episomal forms.

320

Caulimoviruses (Caulimoviridae)

The sites of integration are complex, as the viral sequences have undergone rearrangements, and multiple copies are arranged in tandem. Banana streak virus (BSV) is integrated at two loci in Musa chromosomes, TVCV is integrated at multiple loci in Nicotiana edwardsonii chromosomes, and PVCV is integrated into four loci in Petunia hybrida chromosomes. The mechanism that results in episomal infections remains to be elucidated, but there are some features common to all three viruses. First, the infections arise in interspecific hybrids. Consequently, each parent plant must contribute some factor to activate the integrated virus. Second, the infections arise after the hybrid has been exposed to some sort of stress. PVCV is activated when plants are exposed to water or nutrient stress, or the plants are wounded. TVCV infections arise in N. edwardsonii in the winter months in the greenhouse and are thought to be related to changes in light quality or duration. Integrated Banana streak virus sequences are activated in otherwise healthy Musa species when they are subjected to tissue culture. In each case, the virus is likely released through a series of recombination events or through reverse transcription of an RNA template. Portions of other DNA viruses have been detected in plant chromosomes, but they have not been associated with episomal infections. This has led to speculation that plant DNA viruses might be capable of recombination in every infected plant, likely through a mechanism involving non-homologous recombination. However, since episomal forms of the caulimoviruses and badnaviruses have not been found in tissues fated for seed formation, the integrated forms would not appear in the next generation of plants. Consequently, the banana streak viruses, TVCV, and PVCV must have gained access to germline cells at some point such that their integrants would be passed through seed. This is likely to have been a relatively rare event.

Transmission, Host Range Most members of the genus Caulimovirus are transmitted by as many as 27 species of aphids in a semi-persistent manner. The virions are acquired rapidly by the aphid and can be transmitted immediately upon acquisition. Virions can be maintained by the aphid for as little as 5 h up to three days, but are not retained after the aphid molts and are not passed on to aphid progeny. The protein products encoded by ORF-II (P2) and ORF-III (P3) are both required for aphid transmission. P3 forms a tetramer that binds to both the virion and to the C-terminus of P2. P2 is responsible for the binding of this complex to the aphid, as the N-terminus of P2 binds to a site in the aphid foregut. No insect vectors have been identified for species in the genera Petuvirus, Rosadnavirus, Solendovirus, Soymovirus or Cavemovirus. None of the virions of the icosahedral viruses are transmitted through seed or pollen. Some viruses such as PVCV are transmitted from one generation to the next integrated into host chromosomes; at some point during the growth of the plant the integrated form of the virus is released from the chromosome and becomes capable of episomal replication. Most caulimoviruses are transmitted through mechanical inoculation, but there are a few exceptions, as PVCV, TVCV, Blueberry red ringspot virus, and Strawberry vein banding virus cannot be mechanically inoculated. In addition, most of these viruses are transmitted by grafting. In particular, Dahlia mosaic virus and Blueberry red ringspot virus infections in the field may be initiated through vegetative propagation or through grafting of infected plant material. Badnaviruses are transmitted primarily by mealybugs, but a few are transmitted by aphids. Badnaviruses transmitted by mealybugs include the BSV species, the Sugarcane bacilliform viruses, Cacao swollen shoot virus (CSSV), Dioscorea bacilliform virus, Kalanchoe top spotting virus (KTSV), Piper yellow mottle virus (PYMoV), Taro bacilliform virus (TBV), and Schefflera ringspot virus. These viruses are transmitted in a semi-persistent manner and can be retained after molts, but do not multiply in the mealybug and are not transmitted to progeny. The badnaviruses shown to be transmitted by aphids include Gooseberry vein banding associated virus, Rubus yellow net virus (RYNV), Grapevine vein clearing virus, and Spiraea yellow leaf spot virus. Other modes of transmission of the badnaviruses vary with the species. For example, CoYMV and KTSV can be mechanically inoculated, CSSV and PYMoV are mechanically inoculated with some difficulty, and RYNV and TBV have not been shown to be mechanically inoculated. The preferred method for inoculation of infectious badnavirus clones is agro-inoculation. Some of the badnaviruses are transmitted through seed, including the BSV, KTSV, and Mimosa bacilliform virus. KTSV is very efficiently transmitted by seed, with transmission rates from 60% to 90%, and is also transmitted in pollen. RTBV is dependent on the RNA virus, RTSV, for its transmission. RTSV is vectored by a leafhopper, but causes only very mild symptoms. RTBV is only transmitted by the leafhopper in the presence of RTSV, but is responsible for the severe symptoms associated with the rice tungro disease. RTBV is not mechanically transmitted or carried in seed or pollen. The host range of most caulimoviruses is fairly narrow, as in nature they generally infect plants within a single family (Table 1). Their experimental host range may extend to members of one or two other families, but in many instances, they may only infect a single genus of plants. There are a few exceptions. For example, the Sugarcane bacilliform viruses have a broader host range than most badnaviruses, as they can infect Sorghum, Rottboellia, Panicum, rice and banana. A limited host range can also be associated with similar limitations in the geographic distribution of the virus. For example, Soybean chlorotic mottle virus has only been recovered from a few samples in Japan. Perhaps the virus with the smallest geographic distribution is Stilbocarpa mosaic bacilliform virus, which has only been found on a single, small island in the subantarctic, midway between Tasmania and Antarctica. Other viruses, such as CaMV, Carnation etched ring virus and Dahlia mosaic virus are found worldwide, wherever their hosts are grown. Furthermore, the distribution of viruses that originate from integrated copies in their host’s genomes, PVCV, TVCV, and the BSV species, are also closely aligned with the locations of their hosts. Interestingly, the icosahedral viruses of the caulimoviruses tend to infect hosts in temperate climates, whereas the bacilliform viruses of the badnavirus group are more likely to infect hosts in tropical or subtropical climates.

Caulimoviruses (Caulimoviridae)

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Virus-Host Relationships Caulimoviruses induce a variety of systemic symptoms in their hosts, from chlorosis, streaking, and mosaics, to necrosis. The bestcharacterized pathogenicity determinant is the P6 protein of CaMV, as it has been shown to play a key role in the formation of chlorotic symptoms in turnips. This virulence function was first associated with P6 through gene-swapping experiments between CaMV isolates. It was confirmed when P6 was transformed into several species of plants, and in most cases, they exhibited viruslike symptoms. The P6 protein is also responsible for triggering systemic cell death in Nicotiana clevelandii, as well as a non-necrotic resistance response in N. glutinosa, and a hypersensitive resistance response in N. edwardsonii. One feature that distinguishes the icosahedral viruses from the bacilliform viruses is that the former have the capacity to aggregate into amorphous inclusion bodies (Fig. 1(A)), whereas the latter do not form inclusions. The inclusion bodies formed by the icosahedral viruses are not bound by a membrane, can range in size from 5 to 20 mm, and occur in virtually all types of plant cells. The inclusions can be visualized with a light microscope in strips of epidermal tissue that has been stained with phloxine B. Close examination of CaMV inclusion bodies by electron microscopy reveals that there are actually two types. One type contains many vacuoles and consists of an electron-dense, granular matrix that is composed primarily of the P6 protein (Fig. 1(A)). A second, electron translucent type is made up of the P2 protein, a protein required for aphid transmission. Both types of inclusions are thought to have a role in the biology of the virus. The vacuolated inclusion bodies may be considered pathogen organelles, as they are thought to serve as the sites for replication of the viral nucleic acid, as well as translation of the 35S RNA and assembly of the virions. The electron translucent inclusions are considered to have a role in aphid transmission. A second feature characteristic of the caulimoviruses is that the plasmodesmata of infected cells are enlarged enough to accommodate the virions, as electron micrographs have revealed the presence of CaMV and CoYMV virions in the enlarged plasmodesmata. For both CaMV and CoYMV, the alteration in size is mediated by their proteins required for cell-to-cell movement. In infected protoplasts, the CaMV P1 protein has been shown to induce the formation of tubular structures that extend away from the protoplast surface. It is hypothesized that virions are assembled in the cell and then are escorted to the enlarged plasmodesmata by the cell-to-cell movement proteins.

Further Reading Bak, A., Gargani, D., Macia, J.-L., et al., 2013. Virus factories of Cauliflower mosaic virus are virion reservoirs that engage actively in vector transmission. Journal of Virology 87, 12207–12215. Bhat, A.I., Hohn, T., Selvarajan, R., 2016. Badnaviruses: The current global scenario. Viruses 8, 177. doi:10.3390/v8060177. Geering, A.D.W., Maumus, F., Copetti, D., et al., 2014. Endogenous florendoviruses are major components of plant genomes and hallmarks of virus evolution. Nature Communications 5, 5269. doi:10.1038/ncomms6269. Hull, R., 1996. Molecular biology of rice tungro viruses. Annual Review of Phytopathology 34, 275–297. Krupovic, M., Blomberg, J., Coffin, J.M., et al., 2018. Ortervirales: New virus order unifying five families of reverse-transcribing viruses. Journal of Virology 92, e00515–e00518. Leisner, S.M., Schoelz, J.E., 2018. Joining the crowd: Integrating plant virus proteins into the larger world of effectors. Annual Review of Phytopathology 56, 89–110. Lockhart, B.E.L., 1990. Evidence for a double-stranded circular DNA genome in a second group of plant viruses. Phytopathology 80, 127–131. Mollov, D., Lockhart, B., Ziesak, D.C., Olszewski, N., 2013. Complete nucleotide sequence of Rose yellow vein virus, a member of the family caulimoviridae having a novel genome organization. Archives of Virology 158, 877–880. Pooggin, M.M., Ryabova, L.A., 2018. Ribosome shunting, polycistronic translation and evasion of antiviral defenses in plant pararetroviruses and beyond. Frontiers in Microbiology 9, 459. doi:10.3389/fmicb.2018.00459. Richert-Pöggeler, K.R., Shepherd, R.J., 1997. Petunia vein-clearing virus: A plant pararetrovirus with the core sequences for an integrase function. Virology 236, 137–146. Schoelz, J.E., Leisner, S., 2017. Setting up shop: The formation and function of the viral factories of Cauliflower mosaic virus. Frontiers in Plant Science 8, 1832. doi:10.3389/fpls.2017.01832. Shepherd, R.J., Wakeman, R.J., Romanko, R.R., 1968. DNA in Cauliflower mosaic virus. Virology 36, 150–152. Staginnus, C., Richert-Poggeler, K.R., 2006. Endogenous pararetroviruses: Two-faced travelers n the plant genome. Trends in Plant Science 11, 485–491. Uzest, M., Gargani, D., Drucker, M., et al., 2007. A protein key to plant virus transmission at the tip of the insect vector stylet. Proceedings of the National Academy of Sciences of the United States of America 104, 17959–17964.

Cheraviruses, Sadwaviruses and Torradoviruses (Secoviridae) Toru Iwanami, Tokyo University of Agriculture, Tokyo, Japan Rene AA van der Vlugt, Wageningen University and Research Center, Wageningen, The Netherlands r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

Pol RNA-dependent RNA polymerase PRO Protease Pro-Co Protease cofactor; Pro-glu Glutamic protease RT-PCR Reverse transcription polymerase chain reaction Vpg Viral protein genome-linked

CP Coat protein or capsid protein ELISA Enzyme-linked immunosorbent assay MP Movement protein nm Nanometer(s) NTB Nucleotide-binding protein PCR Polymerase chain reaction

Glossary Enzyme-linked immunosorbent assay A commonly used detection method of any protein that uses specific antibodies against the protein. All viruses have proteins on their particle surface, and are applicable for ELISA, if specific antibodies are prepared and mixed with an enzyme that accelerate colorization in the presence of the viruses. Reverse transcription polymerase chain reaction A laboratory technique that consists of reverse transcription of

an RNA molecule into a DNA molecule and subsequent amplification of the DNA molecule using polymerase chain reaction (PCR). Specific amplification of the DNA molecule is ensured by using a set of specific short DNA molecules (designated “primers”) that are complementary to the target DNA.

Classification The genera Cheravirus, Sadwavirus, and Torradovirus are taxa in the subfamily Comovirinae within the family Secoviridae. Each genus is distinguished from the other genera by it’s specific genomic organization, biological properties and phylogenetic relations. The member species of the genus Cheravirus are Apple latent spherical virus (ASLV), Arracacha virus B (AVB), Cherry rasp leaf virus (CRLV), Currant latent virus (CLV), and Stocky prune virus (StPV). The member species of the genus Sadwavirus is Satsuma dwarf virus (SDV). The member species of the genus Torradovirus are Carrot torradovirus 1 (CaTV1), Lettuce necrotic leaf curl virus (LNLCV), Motherwort yellow mottle virus (MYMoV), Squash chlorotic leafspot virus (SCLSV), Tomato marchitez virus (ToMarV) and Tomato torrado virus (ToTV) (Table 1 and Fig. 1).

Virion Structure Cheraviruses, sadwaviruses, and torradoviruses have the same virion structure as other viruses in the family Secoviridae. Virions are non-enveloped, and 25–30 nm in diameter. Cheraviruses and torradoviruses have three capsid proteins. Sadwaviruses have two capsid proteins. All cheraviruses, sadwavirus, and torradoviruses have two genomic RNAs that are encapsidated in separate virions. Many virus preparations also contain empty particles.

Genome Organization The genome organization of cheraviruses, sadwaviruses, and torradoviruses consists of two single-stranded ( þ )-sense RNA molecules and their genomic organization is similar to that of the other viruses in the subfamily Comovirinae. The distinguishing features in the genome of cheraviruses are three capsid protein genes, and a comparatively small RNA2 that apparently encodes a single polyprotein. The unique features in the genome of sadwaviruses are the extensive amino acid sequence similarity in the N-terminal regions of the polyproteins encoded by RNA1 and RNA2. A unique feature of the torradovirus genome is the presence of a small open reading frame (ORF1) that is located upstream of, and partially overlaps with the large ORF2 in RNA2. The protein potentially encoded by RNA2-ORF1 shows no apparent homologies with other known proteins (Fig. 2).

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Table 1 List of all ICTV classified members of the genera Cheravirus, Sadwavirus and Torradovirus in the family Secoviridae, their taxonomic order, acronyms and GenBank sequence information. # Seq - number of sequences for each virus registered in NCBI Taxonomy Browser data from which a complete genome has been assembled to give a RefSeq assembly number (GCF_number). The last column to the right provides explanation if no RefSeq assembly number is available in the form of accession numbers for incomplete viral sequences. Type species are written in bold Subfamily

Genus

Subgenusa

Species

Acronym

# Seq

RefSeq assembly

Unassigned

Cheravirus

–

Sadwavirusc

Satsumavirus Stramovirus

Apple latent spherical virus Arracacha virus B Cherry rasp leaf virus Currant latent virus Stocky prune virus Satsuma dwarf virus Strawberry mottle virus Black raspberry necrosis virus Chocolate lily virus A Dioscorea mosaic associated virus Carrot torradovirus 1 Lettuce necrotic leaf curl virus Motherwort yellow mottle virus Squash chlorotic leaf spot virus Tomato marchitez virus Tomato torrado virus

ALSV AVB CRLV CuLV StPV SDV SMoV BRNV CLVA DMaV CaTV1 LNLCV MYMoV SCLSV ToMarV ToTV

4 14 10 6 4 33 83 26 4 4 10 4 4 35 25 148

GCF_000861545 GCF_000907115 GCF_000859565 GCF_001503195 – GCF_000860985 GCF_000850445 GCF_000867325 GCF_000895075 GCF_001876835 GCF_000927615 GCF_002219685 GCF_002220005 GCF_002219665 GCF_000879575 GCF_000872965

Choliavirus Torradovirus

–

GenBank accession

DQ143874, DQ143875

a

subgenera organization of the Sadwavirus genus has yet to be ratified by the ICTV.

Fig. 1 Maximum-likelihood phylogenetic analysis of the cheraviruses, sadwaviruses, and torradoviruses using the amino acid sequence between protease domain and RNA polymerase domain in the RNA1 polyprotein. The percentage of trees in which the associated viruses clustered together in the bootstrap test (100 replicates) is shown next to the branches. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. The amino acid sequence between protease domain and RNA polymerase domain in the RNA1 polyprotein of Enterovirus C (EVC) was used as an outgroup. For acronyms of the other viruses, see Table 1.

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Fig. 2 Genomic organization of viruses in the genera Cheravirus, Sadwavirus, and Torradovirus. Schematic representation and the size of the genomes not reflecting their actual sizes. The figure is re-drawn from the figures published previously (Petrzik, et al., 2016, Sanfacon et al., 2009, van der Vlugt et al., 2015) with modifications made by the authors. The same color box indicates the same domains in the polyprotein. For SMoV, the genomic organization of Canadian isolate SMoV-NsPer3 is shown here. The German isolate SMoV-1134 lacks the C-terminal protein domain of the RNA2 polyprotein. Abbreviations: CP, coat protein; CP-L and CP-S, large and small components of coat protein; MP, movement protein; NTB, nucleotide-binding protein (putative helicase); Pol, RNA dependent RNA polymerase; Pro, 3C-like protease; Pro-Co, protease cofactor; Pro-glu, glutamic protease; Vp-35, Vp-26 and Vp-24, viral protein (¼coat protein) of about 35 kDa, 26 kDa, and 24 kDa; VPg, viral genome-linked protein. X1 domain was first reported in nepovirus and the function has not been characterized. X2 was also first reported for nepoviruses and it has sequence homologies with the Pro-Co of comoviruses. VPgs of viruses in the genera Cheravirus, Sadwavirus, and Torradovirus are putative.

Life Cycle and Epidemiology Host Range Natural hosts of cheraviruses include pome fruits, stone fruits, raspberry and currant as well as potato. The experimental host range of cheraviruses is broad or narrow, depending on the virus and strain. For example, CRLV, a cheravirus infects commercially cultivated cherry, apple, and potato, as well as wild herbaceous including dandelion, balsam root, and plantain. SDV, the type species of the genus Sadwavirus infects mainly citrus and occasionally Daphniphyllum teijsmannii and Viburnum odoratissimum var. awabuki. Torradoviruses were described to infect tomato or other crops like lettuce, carrot, squash and cassava, depending on virus species and strains. Little is known about other hosts but Tomato torrado virus (ToTV), the type species, infects over 20 weed species belonging to the family Amarancathaceae, Chenopodiaceae, Solanaceae etc., and the weeds apparently are ToTV sources for transmission to tomato (Figs. 3 and 4).

Geographic Distribution Cheraviruses have been described from different countries worldwide; Apple latent spherical virus (Japan), Arracacha virus B (Peru), Cherry rasp leaf virus (the United States), Currant latent virus (the Czech Republic), and Stocky prune virus (France). Satsuma dwarf virus (Sadwavirus) occurs mostly in Japan but has been found also in China, Korea and Turkey. Tomato-infecting torradoviruses occur in some countries in Europa, in Australia, as well as some parts of North, Central and South America. LNLCV was found in the Netherlands and a CaTV1 torrado-like virus was found in the United Kingdom, France and Germany. MYMoV was detected only in Korea, while SLCV was detected only in Sudan.

Transmission Among cheraviruses, CRLV is transmitted by a nematode (Xiphinema americanum) in the field and is seed-transmitted. Apple latent spherical virus, another cheravirus, is transmitted through seeds obtained from infected trees, and rarely through pollen. No vector

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Fig. 3 Foliar symptoms of satsuma dwarf induced on satsuma mandarin. Satsuma mandarin showing small leaves with shortend internodes.

Fig. 4 Foliar symptoms induced on tomato with infection of ToTV. Necrotic spots surrounded by a light green or yellow area on young leaflets of ToTV-infected tomato. Photo courtesy by Martin Verbeek, Wageningen University and Research; copyright, Wageningen University and Research.

transmission was demonstrated for any sadwavirus, although field observation strongly indicated a soil-borne nature. Seed transmission of sadwavirus in citrus has not been tested, presumably because the main host of sadwavirus is a seedless satsuma mandarine, however it has been demonstrated in an experimental leguminous host. Tomato-infecting torradoviruses are the first spherical viruses reported to be transmitted by whiteflies (Trialeurodes vaporariorum, Bemisia tabaci, and T. abutilonea). No vector is known yet for carrot- and lettuce-infecting torradoviruses but these viruses are spreading in the open field, suggesting vectorinvolvement. Seed transmission of torradoviruses is very rare, if it occurs. Seed transmission of torradoviruses is still uncertain.

Clinical Features and Pathogenesis Cheraviruses usually induce mild or no symptoms. Sadwavirus causes various foliar symptoms including smaller leaves, boat-shaped leaves with shortend internodes that lead to serious reduction in fruit quality and yield in citrus. Sadwavirus symptomlessly infects some wild trees including D. teijsmannii and V. odoratissimum var. awabuki near citrus fields. Some torradoviruses induce severe necrosis (in Spanish “torrado” meaning burned or roasted)” on tomato leaves and fruits. The overall growth of the infected tomato plant is reduced and serious economic damage is inflicted. Torradoviruses infect other vegetables including carrot and lettuce, as well as herbs and wild plants.

Diagnosis Cheraviruses, sadwaviruses, and torradoviruses are readily detected by RT-PCR. Generic RT-PCT that detect a wide range of viruses are available for cheravirus, sadwaviruses and tomato-infecting torradoviruses. Detection by ELISA using specific antibodies is possible for some cheraviruses, sadwaviruses and tomato-infecting torradovirus viruses.

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Treatment No treatment is available.

Prevention Cheraviruses and sadwaviruses mainly infect fruit trees. In this case, movement of contaminated budwoods plays an important role in the long-distance dissemination. For example, occurrence of SDV in China, Korea and Turkey is obviously related to un-checked budwoods from Japan. Severe restriction on such practice is the first step of prevention. Some whitefly-transmitted torradoviruses affect tomato in greenhouses. In this case, damage would be alleviated by preventing whiteflies by suitable screens or sticky traps. Some level of genetic resistance appears to be available against ToTV. Possible sources of resistance for the other torradoviruses are not known.

References Petrzik, K., Koloniuk, I., Prǐ bylova´, J., Špak, J., 2016. Complete genome sequence of currant latent virus (genus Cheravirus, family Secoviridae). Archives of Virology 161, 491–493. Sanfaçon, H., Wellink, J., Le Gall, O., et al., 2009. Secoviridae: A proposed family of plant viruses within the order Picornavirales that combines the families Sequiviridae and Comoviridae, the unassigned genera Cheravirus and Sadwavirus, and the proposed genus Torradovirus. Archives of Virology 154, 899–907. van der Vlugt, R.A.A., Verbeek, M., Dullemans, A.M., et al., 2015. Torradoviruses. Annual Review of Phytopathology 53, 485–512.

Further Reading Adams, I.P., Glover, R., Souza-Richards, R., et al., 2013. Complete genome sequence of Arracacha virus B: A novel cheravirus. Archives of Virology 158, 909–913. Candresse, T., Svanella-Dumas, L., Le Gall, O., 2006. Characterization and partial genome sequence of Stocky prune virus, a new member of the genus Cheravirus. Archives of Virology 151, 1179–1188. Iwanami, T., 2010. Properties and control of Satsuma dwarf virus. Japan Agricultural Research Quarterly 44, 1–6. James, D., Cieslinska, M., Pallás, V., et al., 2017. Viruses, viroids, phytoplasmas and disorders of cherry. In: Quero-Garcia, J., lezzoni, A., Pulawska, J., Lang, G. (Eds.), Cherries, Botany, Production and Uses. Oxfordshire: CAB International, pp. 386–419. Kenten, R.H., Jones, R.A.C., 1979. Arracacha virus B, a second isometric virus infecting arracacha (Arracacia xanthorrhiza, Umbelliferae) in the Peruvian Andes. Annals of Appllied Biology 93, 31–36. Koganezawa, H., Ito, T., 2011. Apple latent spherical virus. In: Hadidi, A., Barba, M., Candresse, T., Jelkmann, W. (Eds.), Virus and Virus-Like Disease of Pome and Stone Fruits. St. Paul, MN: American Phytopathological Society, pp. 23–24. Petrzik, K., Koloniuk, I., Prǐ bylova´, J., Špak, J., 2016. Complete genome sequence of currant latent virus (genus Cheravirus, family Secoviridae). Archives of Virology 161, 491–493. Rubio, M., Martínez-Gómez, P., Marais, A., et al., 2017. Recent advances and prospects in Prunus virology. Annals of Appllied Biology 171, 125–138.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/picornavirales/w/secoviridae/583/genus-cheravirus Cheravirus. https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/picornavirales/w/secoviridae/584/genus-sadwavirus Sadwavirus. https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/picornavirales/w/secoviridae/585/genus-torradovirus Torradovirus.

Citrus Tristeza Virus (Closteroviridae) Moshe Bar-Joseph, Agricultural Research Organization, Volcani Center, Bet Dagan, Israel Scott J Harper, Washington State University, Prosser, WA, United States William O Dawson, Citrus Research and Education Center, Lake Alfred, FL, United States and University of Florida, Lake Alfred, FL, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of M. Bar-Joseph, W.O. Dawson, Citrus Tristeza Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00639-7.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein dRNAs Defective RNAs ELISA Enzyme-linked immunological assays ER Endoplasmic reticulum kb Kilobase kDa Kilodalton LAMP Loop mediated amplification LMT Low molecular weight tristeza MP Movement protein nt Nucleotide(s)

Glossary Cross-protection Prevention of the symptomatic phase of a disease by prior inoculation with a mild or non-symptomatic isolate of the same virus. Defective RNA Subviral RNA molecules lacking parts of the genome while maintaining the signals enabling their synthesis by the viral replication system. Genome The complete genetic information encoded in the RNA of the virus, including both translated and non-translated sequences.

ORF Open Reading Frame PDR Pathogen derived resistance RdRp RNA-dependent RNA polymerase sgRNA Sub-genomic RNA SL Stem-and-loop structures SP Stem pitting SY Seedling-yellow reaction UTR Untranslated region VLPs Virus-like particles VPg Viral protein genome-linked VRC Virus replication complexes

Sub-genomic RNA Shorter than full length genomic RNA molecules produced during the replication process to allow translation of open reading frames. Transgenic pathogen-derived resistance Plants genetically transformed to harbor viral or other pathogen derived sequences expected to confer resistance against related virus isolates.

Introduction The virions of Citrus tristeza virus (CTV) are long flexuous particles (2,000
12 nm) with an unusual architecture that results from two different coat proteins, the major coat protein (CP) and minor coat protein (CPm) encapsidation approximately 97% and 3%, respectively, of the helical structures. Two additional viral proteins, p61 and p64 (HSP 70 homolog) are required for efficient assembly. The 19.3 kb single stranded RNA genome is the longest among the plant viruses. Genomic analysis showed a division into twelve open reading frames (ORFs), the organization of which is conserved among all isolates. The ORF1a encodes a 349 kDa polyprotein containing two papain-like protease domains plus methyltransferase-like and helicase-like domains. Translation of the polyprotein is thought occasionally to continue through the polymerase-like domain (ORF1b) by a þ 1 frameshift. Ten 30 ORFs are expressed by 30 co-terminal sub-genomic messenger RNAs. Infected cells contain a myriad of other sub-genomic RNAs. In addition, most CTV isolates contain one or more defective RNAs of different sizes and genomic rearrangements. Among the CTV translation products, at least 3 function as RNA silencing repressors, which are serving to protect the extremely large size CTV genome and correlated with unusual difficulty in conferring effective pathogen derived resistance to transgenic citrus plants. This article is essentially reporting all the basic elements of the knowledge about CTV including CTV population genomic studies, as well as the use of bioinformatic tools, and the development of the CTV gene cloning vector, that allows elucidating the functional genomics of both CTV and its citrus hosts. Although CTV is still considered to be among the most important plant viruses, the attention of the world’s citrus growers to the significance of CTV research has apparently diminished. We may attribute this decline in interest, at least in part, to the efficacy of control measures introduced in modern citriculture which are based on (1) CTV-tolerant or resistant rootstocks; and (2) effective selection of mild CTV strains that are providing commercially acceptable cross-protection against severe stem-pitting CTV isolates. However, despite

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the general decline in CTV research, the major reason that supports recent CTV research is related to the catastrophic damage experienced during the last 15 years by citrus growers in Brazil and Florida as a result of infection with “Candidatus Liberibacter asiaticus” bacteria. Finding effective means for controlling this bacterium, the causal agent of huanglongbing/greening (HLB) disease, is currently the focus of intense research efforts. Nevertheless, CTV-related problems have not been completely eliminated, and the danger of spread of severe CTV isolates, once the brown citrus aphid reaches the Mediterranean and Californian citrus, remains a continuous threat.

History Over the last 90 years Citrus tristeza virus (CTV) has killed or rendered unproductive millions of trees throughout most of the world’s citrus growing areas and hence it is rightfully considered as the most important virus of the world’s largest fruit crop, citrus, hence the name “tristeza” which means “sadness” in Spanish and Portuguese. However, as with many other disease agents, the actual damage of CTV infections and the timings varied considerably at different periods and geographical regions. The origins of CTV infections remain unknown; the virus however existed for centuries in Asia as an unidentified disease agent, but growers in these areas adapted citrus varieties and rootstock combinations with resistance or tolerance to CTV infections. Out of Asia, citrus production moved to the Mediterranean region, primarily through the introduction of fruit and propagation of seed which does not transmit CTV to the resulting plantings; hence, the new cultivation areas started from seed sources remained free of the virus for centuries. However, the improvement in maritime transportation, allowing long distance transport of rooted citrus plants – led to the outbreak of the deadly citrus root rot disease caused by Phytophthora sp., which was managed by adapting the more tolerant sour orange rootstock. The considerable horticultural advantages offered by this rootstock coincided with the large-scale expansion of plantings throughout the Mediterranean an American countries, and as a result citrus production in many areas was almost entirely based on that single rootstock. This decision had grave subsequent effects when CTV pandemics swept throughout the world, causing “quick decline” (death) of trees on this rootstock. Although the CTV problems were first noticed in South Africa and Australia where, by the end of the 19th century, the growers have found that sour orange was an unsuitable rootstock, it was only in the 1930s that the extent of this deadly disease problem manifested itself first in Argentina and shortly later in Brazil and California, with the death of millions of trees. The virus-like nature of the tristeza-related diseases was demonstrated experimentally by graft and aphid transmission of the disease agent in 1946; however, description of CTV properties began with the seminal findings of thread-like particles associated with tristeza-infected trees. These unusually long and thin particles presented a challenging problem for isolation (purification), which was an essential step towards virus characterization. Development of effective purification methods and biophysical characterization of CTV and similar viruses led to their assignment to the Closterovirus group of elongated viruses. The association of infectivity with the thin-particle-enriched preparations was demonstrated early on; however, the unequivocal completion of Koch’s postulates was only completed in 2001 with the mechanical infection of citrus plants with RNA transcripts of an infectious CTV cDNA clone amplified through serial passaging in Nicotiana benthamiana protoplasts. Development of CTV purification methods paved the way to antibody preparations and to improved CTV diagnosis by ELISA which demonstrated considerable serological diversity among CTV isolates. Later, RNA extracts from CTV-enriched particle preparations were used for molecular cloning of cDNA molecules, which when used as probes confirmed considerable genomic variation among CTV isolates. Nucleic acid probes also demonstrated that plants infected with CTV and other closteroviruses contained many defective RNAs (dRNAs) in addition to the normal genomic and sub-genomic RNAs. Infected plants also contain unusually large amounts of dsRNAs corresponding to the genomic and sub-genomic RNAs. Because of the ease of purifying these abundant dsRNAs, they often have been the template of choice for producing cDNAs. The advent of cDNA cloning of CTV led to the sequencing of its genome and to description of the defected (d)RNAs.

Taxonomy and Classification CTV belongs to genus Closterovirus, family Closteroviridae. The Closteroviridae family contains more than 30 plant viruses with flexuous, filamentous virions and with either mono- or bipartite single-stranded (one tripartite) positive sense RNA genomes. A recent revision of taxonomy of the Closteroviridae based on vector transmission and on phylogenetic relationships using three proteins highly conserved among members of this family (a helicase, an RNA dependent RNA polymerase, and a homolog of the HSP70 heat-shock proteins) led to defining of three genera: Closterovirus including CTV and other aphidborne viruses with monopartite genomes, Ampelovirus comprising viruses with monopartite genomes transmitted by mealybugs, and Crinivirus that includes whitefly-borne viruses with bipartite or tripartite genomes. Closteroviruses are all phloemlimited viruses. Among the hallmarks of the group is the presence of a conserved five-gene–module that includes the four proteins involved in assembly of virions, the major (CP) and minor (CPm) coat proteins, p61, and p64, a homolog of the HSP70 chaperon. In addition, infected phloem-associated cells have clusters of vesicles considered specific to closterovirus cytopathology (Fig. 1).

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250 nm

Fig. 1 An electron micrograph of a thin section from a Mexican lime (Citrus aurantifolia) young leaf vein showing typical symptoms of vein clearing following infection with the Floridian T36 isolate of Citrus tristeza virus. Note the clusters of vesicles typical of infections by closteroviruses. The bar represents 250 nm.

Geographic Distribution Citrus is a common fruit crop in areas with sufficient rainfall or irrigation from the equator to about 41 degrees of latitude north and south. CTV is now endemic in most of the citrus growing areas, with only a few places in the Mediterranean basin and Western USA remaining free of CTV infections. The recent spread of the brown citrus aphid (Toxoptera citricida) to parts of Portugal and Spain are threatening most of the remaining CTV-free areas of the Mediterranean basin. One important aspect of CTV geographic distributions is that some area that have endemic isolates of CTV still do not have the most damaging stem-pitting isolates. With the spread of the vector comes the threat of extremely severe CTV isolates which so far have not spread to North America and Mediterranean countries.

Economic Costs of CTV Control strategies varied at different geographic regions and periods, depending on the extent to which CTV was spreading within the newly infested areas, the sensitivity of the specific local varieties, and the economical availability of alternative crops to replace diseased groves. Most citrus growing countries that managed to remain free of CTV did so by enforcing sanitation practices to prevent virus spread. These included certification schemes aimed to ensure that citrus nurseries propagate and distribute to growers only CTV-free planting material. Other areas had far more costly and ambitious programs of eradication. These efforts were mostly only temporarily successful, mainly because of lack of long-term grower and governmental commitment to such costly operation. In areas were CTV was endemic, alternative strategies to enable continued commercial citrus production in the presence of CTV were developed. Indeed, long before the viral nature of the tristeza disease was realized, Japanese citrus growers were grafting CTV-tolerant Satsuma mandarins on the cold-tolerant and CTV-resistant trifoliate orange (Poncirus trifoliata) rootstock, thus enabling them to produce quality fruits despite endemic CTV infections. Similarly, change of rootstocks from the “quick declining” sour orange rootstocks to rough lemon rootstocks, and to less extent mandarin rootstocks, allowed the South African growers to produce good crops of oranges despite the presence of the most efficient aphid vector and infections with severe CTV strains. Similarly, half a century later the Brazilian citrus industry that was completely decimated during the 1940s, when all trees were grafted on sour orange rootstocks, was saved by adopting CTV-tolerant rootstocks and mild strain cross-protection. Historically the costs of CTV epidemics were estimated to be of the order of tens and even hundreds of millions of dollars. These estimates, however, varied depending on actual market value of the lost production capacity, alternative uses of the land, and the time needed for the CTV-tolerant replants to enter production. There were also indirect costs resulting in poorer performance and/or sensitivity of some CTV-tolerant rootstocks used to replace the widely adapted sour orange rootstocks to other citrus diseases such as citrus blight and citrus sudden death.

Host Range and Cytopathology CTV infects all species, cultivars and hybrids of Citrus sp., some citrus relatives such as Aeglopsis chevalieri, Afraegle paniculata, Fortunella sp., Pamburus missionis and some intergeneric hybrids. Species of Passiflora are the only non-Rutaceous hosts, infected naturally. The CTV decline isolates in trees on sour orange rootstocks (Fig. 2) are associated with the death of the phloem near the

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Fig. 2 Symptoms of CTV on different citrus species: (A). decline of sweet orange on sour orange rootstock, (B & C) seedling yellows on Duncan grapefruit.

bud union, resulting in a girdling effect that may cause the overgrowth of the scion at the bud union, loss of feeder roots and thus drought sensitivity, stunting, yellowing of leaves, reduced fruit size, poor growth, dieback, wilting, and death. However, other virulent and damaging CTV strains cause stem pitting (SP), which results in pits in the wood under depressed areas of bark and are often associated with severe stunting and considerably reduced fruit production. The seedling-yellow reaction (SY) which includes severe stunting and yellowing on seedlings of sour orange, lemon and grapefruit is primarily a disease of experimentally inoculated plants but might also be encountered in the field. CTV infection is closely restricted to the phloem tissues of infected plants that first show chromatic cells filled with either aggregates of virus particles and/or necrotized cytoplasm. Interestingly, the death of trees on the sensitive sour orange rootstock is not associated with increased amounts of chromatic cells, thus suggesting a different patho-physiological condition lead to the chromatic type of cell death, which could also be observed in tissues of sweet orange trees which are tolerant to CTV when growing on their own roots. Nicotiana benthamiana has become an important experimental host for CTV since it was discovered that plants become systemically infected by Agrobacterium tumefaciens containing recombinant DNA plasmids of CTV (agro-inoculation).

The CTV Virions The CTV virions are long flexuous particles, 2000 nm long
10–12 nm in width (Fig. 3). The virions have helical symmetry with a pitch of 3–4 nm, about 8–9 capsids per helix turn, and a central hole of 3–4 nm. Unlike the virions of other elongated plant viruses that consist of cylindrical nucleocapisds made of a single CP, the CTV virions consist of bipolar helices with a long body and short tail. Immuno-electron microscopy showed antibodies to CPm attached to only one end of the virions, with the major part (497%) of the virion is encapsidated by CP. Interestingly, the “tail” corresponds to the 50 end region of the viral genome and the tails of other closteroviruses have been associated with small amounts of p61 and HLP60h.

The CTV Genome Fig. 4 shows a schematic presentation of the 19.3 kb CTV RNA. Generally, the CTV genome is divided almost equally into two parts, the 50 part consisting of ORF1a and 1b harboring the viral replication machinery and the 30 end half harboring 10 ORFs

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Fig. 3 Particles of CTV, with a diameter of 10–12 nm, negatively stained with Uranyl acetate. Note their flexible thread like shape, considerably limiting the possibility of their isolation from infected citrus tissues. The bar represents 50 nm.

Fig. 4 A schematic presentation of the citrus tristeza virus (CTV) genomic (g) and subgenomic (sg) RNAs. The replication-associated ORFs are highlighted in blue, the five-gene structural and assembly block common to all closteroviruses in yellow, suppressors of silencing in red, and three genes on undetermined function in green. Lines shown on the left and right side below the genomic map indicate 50 large molecular single stranded (ss) 50 sgRNA (LaMT) and two low molecular ssRNAs (LMT1 and LMT2) and a nested set of 30 co-terminal sgRNAs, respectively. The left insert shows Northern blot hybridization of dsRNA enriched extracts from an Alemow plant infected with the VT strain of Citrus tristeza virus (CTV-VT) using Riboprobes specific to the 50 end of the viral RNA. Note the presence of the large replicative form (RF) molecules (upper band), LaMT and two abundant LMT molecules. The right insert shows hybridization of dsRNA with a 30 UTR probe. Note the hybridization bands of the different 30 sgRNAs indicated by arrows.

encoding a range of structural components and other gene products involved in virion assembly and host and vector interactions. CTV replicons containing only ORFs 1a and 1b plus the 50 and 30 non-translated ends (Fig. 4) fulfill all the requirements for efficient replication in protoplasts. Interestingly, while the sequences of the 30 half of all CTV isolates that have been sequenced are 97% and 89% identical, the 50 half sequences often differ considerably more. For instance, the isolates T36 and VT show only 60%–70% identities for their 50 halves.

The Untranslated Regions (UTRs) A remarkable feature of CTV isolates are the close identities (97%) of their 30 NTR primary sequences compared to the considerable divergence of their 50 UTR sequences. Computer assisted calculations, however, suggest that not only the 30 UTRs of different isolates fold into similar predicted structures, but surprisingly the dissimilar sequences of the 50 NTRs also are predicted to fold into similar secondary structures. The 30 replication signal of several CTV isolates was mapped to 230 nt within the UTR and predicted to fold into a secondary structure composed of 10 stem-and-loop (SL) structures. Three of these SL regions and a terminal 30 triplet, CCA, are essential for replication. Replacement of T36 30 UTR with 30 UTRs from other strains allowed

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replication, albeit with slightly less efficiency, suggesting on the significance of the primary sequence of this part. The plus-strand sequence of the 50 UTRs from different strains are predicted to form similar secondary structures consisting of two stem-loops (SL1 and SL2) separated by a short spacer region, that were essential for replication and initiation of assembly by CPm.

Gene Functions In infected cells, the 12 ORFs of CTV (Fig. 4) are expressed through a variety of mechanisms including; proteolytic processing of the polyprotein, translational frame shifting, and production of 10 30 co-terminal sub-genomic RNAs. The first two mechanisms are used to express proteins encoded by the 50 half of the genome while the third mechanism is used to express ORFs 2–11. The ORF1a encodes a 349 kDa polyprotein containing two papain-like protease domains plus methyltransferase-like and helicase-like domains. Translation of the polyprotein could also continue through the polymerase-like domain (ORF1b) by a þ 1 frameshift. These proteins along with the signals at the 50 and 30 ends of the genome are the minimal requirements for replication of the RNA. The function of p33 (ORF2) is required for superinfection exclusion and is required for infection of a subset of the viral host range. The next five gene products from the 30 genes include the unique signature block characteristic of closteroviruses, which consist of the small, 6 kDa hydrophobic protein (ORF3), 65 kDa cellular heat-shock protein homolog (HSP70h, ORF4), 61 kDa protein (ORF5), and the tandem pair of p27 (CPm, ORF6) followed by p25 (CP, ORF7). The four proteins with the exception of p6 are required for efficient virion assembly. The small hydrophobic p6 is a single-span trans-membrane protein not required for virus replication or assembly, but it is required for infection of all plants of the host range. CP is also a suppressor of RNA silencing. The function of p18 (ORF8) and p13 (ORF9) are required for infection of certain hosts. Protein p20 (ORF10) accumulates in amorphous inclusion bodies of CTV-infected cells and has been shown to be a suppressor of RNA silencing. p23 (ORF11) is a multifunctional protein with no homolog in other closteroviruses, that: (1) binds cooperatively both single-stranded and dsRNA molecules in a non-sequence-specific manner (2) contains a zinc finger domain that regulates the synthesis of the plus- and minus-strand molecules and controls the accumulation of plus-strand RNA during replication (3) is an inducer of CTV-like symptoms in transgenic C. aurantifolia plants and (4) is a potent suppressor of intracellular RNA silencing in Nicotiana tabacum and N. benthamiana and (5) controls the level of negative-stranded of genomic an sub-genomic RNAs. For additional p33 functions, see also Virus-Virus, interactions.

The CTV Sub-Genomic RNAs The replication of CTV involves the production of large number of less than full-length RNAs. These include ten 30 co-terminal sg mRNAs, and ten negative-stranded sgRNAs corresponding to the ten 30 sg mRNAs, plus ten 50 co-terminal sgRNAs that apparently are produced by termination just 50 of each of the 10 ORFs (Fig. 4). The amounts of different sg mRNAs vary with the highest levels for sg mRNAs p23 and p20, located at the distal 30 end. Infected cells also contain abundant amounts of two other small 50 co-terminal positive-stranded sgRNAs of B600 and 800 nt designated as low molecular weight tristeza (LMT).

Defective RNAs Most CTV contain defective RNAs (dRNAs), which consist of the two genomic termini, with extensive internal deletions, resulting in different sizes of 50 and 30 sequences. CTV dRNAs accumulate abundantly in infections with wild type virus. The dRNAs contain different sizes of 50 and 30 sequences. The biological roles of CTV dRNAs remain obscure as no clear cut associations have been established with any of the dRNA.

Virus-Virus Interactions The CTV isolates were categorized as strains based on biological properties. Major genetic differences between CTV isolates were reported using a CTV specific cDNA probe of the VT strain. Cross-protection is a practice in which a mild population of CTV is purposely pre-inoculated into trees to prevent infection and disease caused by endemic isolates. Cross-protection has been widely used to allow commercial citrus production in regions with severe endemic isolates of CTV. It works well when an effective protecting isolate is found, but this process is empirical and difficult. Most mild isolates fail to protect and finding the correct combination has consumed whole careers. The successful protecting isolate must affect a virus-virus interaction such that the protecting isolate interferes with establishment of infection and induction of disease symptoms. A component of this process is superinfection exclusion. It has been shown that previously established infections of CTV can completely exclude superinfection by another isolate. However, this is limited to isolates from the same strain of CTV. Challenges with isolates of different strains of CTV are not inhibited. This superinfection exclusion is dependent on the p33 gene and/or the 50 protease. For example, as the fulllength virus can completely exclude an isolate from the same strain, CTV with the p33 gene deleted does not. Thus, one component of finding or creating a protecting virus against a pathogenic endemic isolate is to identify the sequence group (strain) of the cause of disease, challenging isolates and choose or create a mild isolate from the same strain.

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Fig. 5 A diagram of class 1 CTV defective (d) RNAs with three different sizes and class 2 CTV-dRNAs that show 30 moieties of the size and structure of the full-length sgRNA of ORF11. Inserts A and B show Northern hybridization of class 1-dsRNAs from three different CTV-VT sub isolates, hybridized with Riboprobes specific for the 50 and 30 ends of the CTV gRNA, respectively. Note the intense bands resulting of the hybridization with both probes of the abundantly present dRNAs. Lower part is a diagrammatic presentation of direct repeats at the junction sites of some class 1 dRNAs. The bottom part shows a schematic presentation of a class 2 dRNA, with a 30 terminus corresponding to the ORF11 sgRNA of CTV-VT.

However, since most endemic isolates are populations made up of components from several strains, in practice it is not that simple because so far effective protecting isolates also are population. Thus, this requires an understanding of the rules of populations. For example, a typical population might consist of 50% strain A, 20% strain B, 29% strain C, and 1% strain D. CTV has at least seven characterized genotypes, defined by nucleotide sequence divergence of B10%–20% from each other. Fig. 5 shows a Phylogenetic tree of 60 CTV isolates belonging to different CTV strains (circles). These CTV genotypes differ in their infectivity in different citrus species, tropism and ability to move systemically, vector transmissibility and to some extent, pathogenicity. It should be noted, however, that there is variability in biological properties even between isolates or variants of the same genotype (Fig. 6). As a virus with multiple, distinct genotypes, and a long-lived host system, it is unsurprising CTV isolates from field trees almost always consist of populations of multiple genotypes in mixture. The existence of multiple genotypes within an individual plant, and co-infection of the same cells also explains the frequency of recombination observed for CTV isolates. The CTV genotypes within individual plants both compete and interact with one another, reaching equilibrium based on their relative fitness vis-à-vis one another, and the host. A major rule of equilibrium of mixtures of genotypes in a population is determined by the host, which could explain why cross-protection often breaks down upon utilization of new citrus varieties. Also, individual CTV genotypes or variants are also capable of interacting via complementation to increase fitness and overcome tropism limitations as well as increasing the efficacy of aphid transmission.

Transmission Although the virus is phloem-limited, mechanical inoculation can be done experimentally at relatively low efficiency by slashing the trunks of small citrus trees with blades containing sap extracts. In commercial groves, the virus is spread naturally by aphid vectors and by vegetative propagation in infected budwood. Long distance spread of the virus, particularly from country to country, had been through the long-distance transfer of plant propagation materials. Several aphid species including Aphis gossypii, A. spiraecola, A. craccivora, and T. citricida transmit CTV semi-persistently. The rate of transmissibility varies considerably between different virus isolates and different aphid species. Transmission efficacy of T. citricida is approximately 6 to 25 times more efficient than A. gossypii, and different CTV isolates or genotypes can differ dramatically, for example 40%–60% for the type isolate of strain T68 to less than 1% for type isolate or strain T36. However, it

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Fig. 6 (A) Phylogenetic tree and evolutionary analysis of 60 full-length citrus tristeza virus (CTV) genome sequences using the maximumparsimony method. The CTV isolates belonging to different CTV strains are indicated by circles, with permission from the author Yokomi, R.K., Selvaraj, V., Maheshwari, Y., et al., 2017. Identification and characterization of citrus tristeza virus isolates breaking resistance in trifoliate orange in California. Phytopathology 107, 901–908. (B) A phylogeny of displaying the major Citrus tristeza virus (CTV) genotypic groupings, generated using the maximum-parsimony method from complete genomic sequences.

should be noted that other isolates of the T36 strain are much more highly transmitted and the type isolate was originally isolated from a tree infected by aphid transmission. Apparently, more than 50 years of graft transmission of the type isolate of T36 resulted in loss of aphid transmission. Transmission efficacy can also be affected by the interaction of CTV genotypes in mixture, where the presence of highly transmissible variants can increase the transmission rate of poorly transmissible variants through complementation. For the semi-persistent mode of transmission, the virion has been found to bind to the cibarium and foregut of the aphid vector via interaction of the p27, p61 and p65 proteins. Additionally, the p33 protein has also found to be involved in enhanced transmission.

CTV Control Measures Strategies to control CTV have varied at different locations and periods and included (1) quarantine and budwood certification programs to prevent the introduction of CTV, (2) costly and ambitious eradication programs to contain situations of virus spread, (3) the use of CTV-tolerant rootstocks and mild (or protective) strain cross-protection, often also named preimmunization, (4) breeding for resistance, and (5) attempts to obtain resistance by genetic engineering. Mild strain cross-protection has been widely applied for millions of citrus trees in Australia, Brazil, and South Africa to protect against stem pitting of sweet orange and grapefruit trees. However, mild strain cross-protection has not provided field protection against CTV isolates causing quick decline of trees on the sensitive sour orange rootstock. Useful resistance to CTV has been found in a citrus relative, Poncirus trifoliata. This resistance has been mapped and shown to be controlled primarily by a single genome region and sequenced. However, a specific gene has not been identified by transformation into susceptible citrus resulting in a resistance phenotype.

CTV Diagnosis The outcome of CTV infections varies considerably depending the virulence of the prevailing virus isolates, and sensitivity of infected citrus varieties and rootstock combinations. Hence, the need for diagnosis is of utmost importance. Biological indexing of the disease by grafting sensitive citrus indicators has been the definitive assay, although it is costly and time consuming. It has largely been replaced with more rapid immunoassays (ELISA) that have been widely practiced for almost 30 years for a variety of CTV sanitation programs. Development of recombinant antigens considerably advanced diagnostic possibilities and production of monoclonal antibodies has allowed for more precise differentiation of isolates. PCR and combinations of immuno-capture PCR allow more sensitive CTV diagnosis. Recently, high-

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throughput sequencing has proven an effective tool for the rapid identification of CTV isolates and populations. However, despite the development of better detection tools, none are effective for predicting the biological properties of new CTV isolates.

Attempts to Apply Transgenic Resistance to CTV Conventional breeding for CTV resistance is a long, difficult, and inconvenient process mainly because most citrus varieties are hybrids of unknown parentage. Hence the considerable interest in applying pathogen derived resistance (PDR) to render both citrus rootstocks and varieties resistant to stem pitting isolates and decline causing CTV isolates. Yet, literally hundreds of independent transformations with a range of different configurations of CTV sequences has resulted in failure to obtain durable protection against CTV. These failures are especially frustrating since the control of severe stem pitting isolates is needed in several major citrus producing. Several reports on transgenic resistance to CTV infection including effective protection of plants expressing the three CTV silencing suppressor genes need still validation under field conditions. The lack of RNA silencing against CTV in citrus might be due to more effective suppression by the combination of the three suppressors of this virus.

Epilogue As indicated in the preface, in much of the world’s citrus-growing regions, CTV is now considered a manageable disease. Nevertheless, research is still needed to address several important questions such as the spread of severe CTV isolates and the brown citrus aphid into regions previously free of them, and a re-emergence of CTV as serious economic plant pathogen of citrus. We still do not know the origin of CTV; did it originate from one of the many wild relatives of domesticated citrus, or from another plant family and subsequently adapted to citrus? CTV and its host interactions especially in Pumello, one of the three ancestors of citrus also remains to be elucidated, especially as both the highly CTV sensitive sour orange and the tolerant sweet oranges share both Pumello and mandarins as parents. Similarly questions of CTV genome variation are worth attention noting the considerable variation between the 50 -terminal regions of the genomes of T36 and VT strains raises the possibility of a long-time separation from the original source, does the parental strain still survive and, if so, could the related genome re-emerge as a new pathogen based on its genome and biology? Finally, it is interesting to note that recent molecular studies of phloem-limited bacteria, such as the greening agent, indicated a drastic reduction in their genome size, compared to the majority of non-phloem-limited bacteria; whereas, CTV, with the largest genome of the family Closteroviridae, behaves just the opposite. The question thus remains why and how CTV adopted such a different survival strategy within the phloem vessels of its host plants.

See also: Closteroviruses (Closteroviridae). Plant Viral Diseases: Economic Implications

Further Reading Albiach-Marti, M.R., 2013. Chapter 1 – The complex genetics of citrus tristeza virus. In: Romanowski, V. (Ed.), Current Issues in Molecular Virology – Viral Genetics and Biotechnological Applications. InTech, pp. 1–25. Atta, S., Zhou, C., Zhou, Y., et al., 2012. Distribution and research advances of citrus tristeza virus. Journal of Integrative Agriculture 11, 346–358. Bar-Joseph, M., Marcus, R., Lee, R.F., 1989. The continuous challenge of citrus tristeza virus control. Annual Review of Phytopathology 27, 291–316. Catara, A.F., Bar-Joseph, M., Licciardello, G., 2019. Citrus Tristeza Virus: Methods and Protocols. Springer. (ISBN 978-1-4939–9558-5). Chen, A.Y.S., Watanabe, S., Yokomi, R., Ng, J.C.K., 2018. Nucleotide heterogeneity at the terminal ends of the genomes of two California citrus tristeza virus strains and their complete genome sequence analysis. Virology Journal 15, 141. Dawson, W.O., Bar-Joseph, M., Garnsey, S.M., Moreno, P., 2015. Citrus tristeza virus: Making an ally from an enemy. Annual Review of Phytopathology 53, 137–155. Flores, R., Moreno, P., Falk, B., Martelli, G.P., Dawson, W.O., 2013. E-book on Closteroviridae. Frontiers in Microbiology 4, 411. doi:10.3389/fmicb.2013.00411. (PMID: 24409172). Folimonova, S.Y., 2013. Developing an understanding of cross-protection by citrus tristeza virus. Frontiers in Microbiology 4, 76. Harper, S.J., 2013. Citrus tristeza virus: Evolution of complex and varied genotypic groups. Frontiers in Microbiology 4, 93. Karasev, A.V., Hilf, M.E. (Eds.), 2010. Citrus Tristeza Virus Complex and Tristeza Diseases. APS Press. (SBN: 978-0-89054-378-8). Moreno, P., Ambrós, S., Albiach-Martí, M.R., Guerri, J., Peña, L., 2008. Citrus tristeza virus: A pathogen that changed the course of the citrus industry. Molecular Plant Pathology 9 (2), 251–268. Satyanarayana, T., Bar-Joseph, M., Mawassi, M., et al., 2001. Amplification of Citrus tristeza virus from a cDNA clone and infection of Citrus trees. Virology 280, 87–96. Wallace, J.M., 1978. Virus and virus like diseases. In: Reuther, W., Calavan, E.C., Carman, G.E. (Eds.), The Citrus Industry, vol. 4. Berkeley, CA: Div. Agric. Sci. Univ. Calif., pp. 67–184. Yokomi, R.K., Selvaraj, V., Maheshwari, Y., et al., 2018. Molecular and biological characterization of a novel mild strain of citrus tristeza virus in California. Archives of Virology 163, 1795–1804.

Closteroviruses (Closteroviridae) Marc Fuchs, Cornell University, Geneva, NY, United States r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa amino acid(s) AlkB alpha-ketoglutarate-dependent hybroxylase domain CP coat protein or capsid protein CPm minor coat protein ELISA enzyme-linked immunological assays ER endoplasmic reticulum HC-Pro helper component-protease HEL helicase HSP70h heat shock protein 70 homolog

Glossary Ampelovirus One of the four genera in the family Closteroviridae. The name ampelo derives from the Greek ampelos, “grapevine”, the natural host of grapevine leafrollassociated virus 3, the type species of the genus Ampelovirus. Closterovirus Another genus in the family Closteroviridae. The name clostero derives from the Greek kloster, “spindle, thread” due to the size and appearance of very long, flexuous, and helically constructed filamentous virions. Crinivirus It is another genus in the family Closteroviridae. The name crini derives from the Latin crinis, “hair”, for the appearance of the very long thread-like particles.

kb kilobase kDa kilo dalton LAMP loop mediated amplification L-Pro leader papain-like protease MET methyltransferase MP movement protein nt nucleotide(s) ORF open reading frame RdRp RNA-dependent RNA polymerase UTR un-translated region

RNA-dependent RNA polymerase An enzyme that catalyses the transcription of RNA from an RNA template. tRNA-like structure An RNA sequence with a similar tertiary structure to transfer RNA. Velarivirus It is the most recently identified genus in the family Closteroviridae. The name velari derives from the Latin velari, “cryptic or veiled”, because Grapevine leafroll-associated virus 7, the type species of this genus, does not cause any apparent disease symptoms on its natural host.

Introduction Plant viruses of the family Closteroviridae belong to the higher phylum Kritrinoviricota in the realm Riboviria. Their common features include the morphology and size of virions, the genome structure and expression, the presence of canonical conserved proteins, characteristic cytopathic structures, tissue tropism, transmission by specific hemipteran insects in a semi-persistent manner, and extreme difficulties at mechanical transmission to herbaceous hosts (Table 1). Virions have a diameter of about 12 nm and lengths ranging from 650 nm for species with a fragmented genome to over 2000 nm for species with a monopartite genome (Table 2). Virions are helically constructed filaments with a pitch of the primary helix in the range of 3.4–3.8 nm, containing about 10 protein subunits per turn of the helix and showing a central hole of 3–4 nm (Fig. 1). Two types of coat protein (CP), the major CP and a CP analog or minor CP (CPm), are the most abundant protein components involved in the formation of most virions with CPm encapsidating the 600–700 nt 50 -terminal of the viral RNA and coating one extremity (75–100 nm) of the particles, thus forming a distinct structure referred to as rattlesnake or bipolar. The virus-encoded heat shock protein 70 homolog (HSP70h) of cell chaperones and the B60 kDa proteins are also integral virion components. The B60 kDa protein is required for incorporation of both HSP70h and CPm to virions, which also incorporate a 20 kDa protein that may form the tip segment of the virion head. Species demarcation criteria used for all genera in the family Closteroviridae are particle size, size of the CP, genome structure and organization, vector species and specificity, cytopathological features, host range, as well as aa sequence of the RNA-dependent RNA polymerase (RdRp), CP, and HSP70h differing by more than 25%.

Classification Plant viruses in the family Closteroviridae are divided into four genera: Closterovirus (monopartite genome), Ampelovirus (monopartite genome), Velarivirus (monopartite genome) and Crinivirus (bipartite or tripartite genome) (Table 2). Collectively these viruses are referred to as closteroviruses, the generic name for viruses in the family that was initially coined for this group of plant viruses by

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Genome structure

Genome organization

Common features among members of the family Closteroviridae Genome expression

A very large positive sense, Long, helically Two conserved gene modules encoding proteins Proteolytic single-stranded RNA (12,000 to associated with replication with a close processing, þ 1 constructed nearly 19,000 nt) with an filamentous particles phylogenetic relationship (ORF1a and 1b) and a ribosomal organization distinct from those quintuple gene module encoding proteins frameshift, and the (650–2200 nm in of other plant viruses production of sublength and 12 nm in associated with replication, virion formation, genomic RNAs diameter) cell-to-cell movement and suppression of RNA silencing; and a unique set of genes coding a heat shock protein 70 homolog, and for a duplicated, diverged copy of the coat protein (minor coat protein, CPm)

Virions

Table 1 Tissue tropism

Cytoplasmic Phloem aggregates of restriction with virions intermingled occasional with single or location in the clustered mesophyll and membranous epidermis vesicles

Cytopathic structures

Aphids, Extremely whiteflies, difficult for a mealybugs extremely and soft limited scale number of insects virus species

Transmission Mechanical inoculation

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Table 2 Genus

Distinguishing properties of the four genera in the family Closteroviridae Virion length (nm) RNA species (#)

Genome size (kb)

ORF (No.)

Replicase (kDa)

HSP70h (kDa)

CP (kDa)

CPm (kDa)

Vector

Closterovirus 1350–2000 Ampelovirus 1400–2200

1 1

14.5–19.3 13.0–18.5

8–12 7–12

349–367 245–293

65–67 57–59

22–25 28–36

24–27 50–56

Crinivirus

2 or 3

7.8–9.1/7.9–8.5 8.0–5.3–3.9 16.2–16.9

9–13

267–280

62–65

28–29

53–80

Aphids Mealybugs, soft scales Whiteflies

9

260–270

62–69

34–46

69–76

None known

Velarivirus

650–850 and 700–900 1500–1700

1

Fig. 1 Electron micrographs of virions of Beet yellows virus (BYV) from the genus Closterovirus that are negatively stained and decorated with an antiserum specific to the (a) coat protein (CP) -note that the CP minor (CPm) tail is not decorated-, and (b) CPm at the 75 nm tail of a virus particle. (c) Close-up of the 75 nm tail of four selected BYV particles decorated with an antiserum specific to CPm. Scale bars represent 300 nm. From Agranowsky, A.A., Lesemann, D.E., 2000. Beet yellows virus. Descriptions of Plant Viruses. Association of Applied Biologists. Available at: http://www.dpvweb.net/dpv/showdpv.php?dpvno=377, with permission.

the International Committee on Taxonomy of Viruses in 1976. In addition to classified viruses in the family, a few species are unassigned members in the family Closteroviridae: Actnidia virus 1, Alligatorweed stunting virus, Blueberry virus A, Megakepasma mosaic virus, Mint vein banding-associated virus, Olive leaf yellowing-associated virus, and Persimmon virus B. The genus Closterovirus comprises 14 species with particles ranging from 1350 to 2000 nm in length, and a monopartite, positive-sense, single-stranded RNA genome, 14.5–19.3 kb in size, in which the CPm is uniquely located upstream of the CP. The natural transmission of closteroviruses is by aphids (Table 2). Beet yellows virus (BYV) is the type species of the genus Closterovirus. The genus Ampelovirus comprises 13 species with particles ranging from 1400 to 2000 nm in length, and a monopartite positive-sense, single-stranded RNA genome, 13.0–18.5 kb in size. A wide variation in genome size and organization is typical for species in this genus. The natural transmission of ampeloviruses is by pseudococcid mealybugs and soft scale insects (Table 2). The type species of the genus Ampelovirus is Grapevine leafroll-associated virus 3 (GLRaV-3). The genus Crinivirus comprises 13 species transmitted by whiteflies. Virions usually have two modal lengths (650–850 and 700–900 nm) and a bipartite positive-sense, single stranded RNA genome with a size ranging from 7801 to 9127 nt (RNA-1) and from 7903 to 8530 nt (RNA-2) (Table 2). The type species of the genus Crinivirus is Lettuce infectious yellows virus (LIYV). Noteworthy, Potato yellow vein virus (PYVV) has a tripartite positive-sense, single-stranded RNA genome of 8035 nt (RNA-1), 5339 nt (RNA-2) and 3892 nt (RNA-3) in size (Table 2). The genus Velarivirus comprises seven species with particles ranging from 1500 to 1700 nm in length, a monopartite, positivesense, single-stranded RNA genome, 16–16.9 kb in size, and no known hemipteran vectors (Table 2). The type species of the genus Velarivirus is Grapevine leafroll-associated virus 7 (GLRaV-7).

Genome Organization The 50 end of the genome of viruses in the family Closteroviridae is likely capped, and the 30 end is not polyadenylated and does not possess a tRNA-like structure. The genome organization is conserved among species in the family although the number and relative

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position of the different open reading frames (ORFs) differ between genera and/or individual viral species. However, the dual gene module ORF1a–ORF1b at the 5’ end of the genomic RNA invariably encodes the replication-associated proteins with conserved domains for a papain-like cysteine protease (L-Pro), a methyl-transferase (MET), a helicase (HEL), and an RNA-dependent RNA polymerase (RdRp). Downstream ORFs form a conserved five-gene module with few modifications among most members of the family. This quintuple gene module codes for, in the 50 -30 direction, a 6K small hydrophobic protein, the HSP70h, a B60 kDa protein, a CP, and a CPm. The HSP70h and the B60 kDa proteins are integral virion components present in all the sequenced members of the family. The functions postulated for HSP70h are mediation of cell-to-cell movement through plasmodesmata, involvement in the assembly of multisubunit complexes for genome replication and/or sub-genomic RNAs synthesis, as well as assembly of virus particles. The B60 kDa protein is required for incorporation of both HSP70h and CPm to virion heads. In general, CP and CPm show a significant degree of sequence conservation for each species in the family and the duplicate copies probably retain the general spatial folding and some crucial properties of the CPs. Noteworthy, a group of ampeloviruses [e.g., Grapevine leafroll-associated virus 4 (GLRaV-4) and related strains, Pineapple mealybug wilt-associated virus 1 (PMWaV-1) and Pineapple mealybug wilt-associated virus 3 (PMWaV-3), Air potato ampelovirus 1 (AiPoV-1), and Plum bark necrosis stem pitting-associated virus (PBNSPaV)] with the smallest genome in the family does not appear to possess a CPm; and grapevine leafroll-associated virus 1 (GLRaV-1) carries two copies of CPm. The genome expression strategy of viruses in the family Closteroviridae is based on: (1) proteolytic processing of the polyprotein encoded by ORF1a; (2) þ 1 ribosomal frameshift for the expression of the RdRp domain encoded by ORF1b; and (3) expression of the downstream ORFs via the formation of a nested set of 30 co-terminal sub-genomic RNAs (sgRNAs). Analysis of double stranded (ds) RNAs in infected plant tissue reveals very complex and variable patterns among species of the family, reflecting the different numbers and sizes of ORFs present in individual genomes and, in some cases, the existence of defective RNAs. The largest dsRNA is usually the replicative form of the entire genome and sgRNAs generate a range of smaller dsRNAs. For species in the genus Closterovirus (Table 3), the order of the CP and CPm ORFs is reversed compared to that of species in the genera Ampelovirus, Crinivirus, and Velarivirus. BYV contains eight ORFs flanked by 50 and 30 untranslated (UTRs) of 107 and 181 nt, respectively, including an extra ORF (ORF2) encoding a 30 kDa polypeptide with no similarity to any other protein in databases (Fig. 2). Citrus tristeza virus (CTV) has 12 ORFs and UTRs of 107 nt at the 50 end and 275 nt at the 30 end. It differs from the BYV genome in having two L-Pro domains in ORF1a [so does Grapevine leafroll-associated virus 2 (GLRaV-2)], and two extra 30 proximal ORFs (ORF9 and ORF11). The genome organization of Beet yellows stunt virus (BYSV) is intermediate between that of BYV and CTV with 10 ORFs and a 30 UTR 241 nt in size, suggesting that these three viruses might represent three distinct stages in the evolution of closteroviruses. Non-structural proteins common to all members of the genus are: (1) a large polypeptide (over 300 kDa) containing the conserved domains of L-Pro, MET, and HEL; (2) a B50 kDa protein with all sequence motifs of viral RdRp; (3) a 6 kDa hydrophobic protein with membrane-binding properties; (4) the homolog of the cellular HSP70; and (5) a 55–64 kDa product, referred to as the B60 kDa protein. Some of the structural and non-structural proteins function as suppressors of the RNA silencing plant defense mechanisms. For instance, CP, p20 and p23 proteins of CTV have suppressor activity, much the same as the homologs of p21 of BYSV, BYV, and GLRaV-2. Silencing suppressors contribute to the accumulation of virus particles and are important determinants of pathogenesis. The CTV p23 is a unique protein in the family with a nucleolar localization. For species in the genus Ampelovirus (Table 3), Grapevine leafroll-associated virus 3 (GLRaV-3) has the largest genome of the family with 18,498 nt and 12 ORFs coding for the replication related proteins (ORFs 1a and 1b), two small hydrophobic proteins (6 kDa), the HSP70h, the B60 kDa protein, CP, CPm, and five additional proteins of 21, 20, 20, 4, and 7 kDa in size, respectively (Fig. 3). The 50 and 30 UTRs are 737 and 277 nt in size, respectively. GLRaV-3 is a sub-group I ampelovirus. Subgroup II species in the genus have only seven ORFs and lack the CPm. For example, PMWaV-1 has a genome 13,071 nt in size, beginning with a 535 nt UTR at the 50 end, followed by the ORFs expressing the replication related proteins, a 6 kDa hydrophobic protein, the HSP70h, the B60 kDa protein, the CP, and a 24 kDa protein, respectively. A UTR 132 nt in size terminates the genome at the 30 end. Structural and non-structural proteins are similar in type and function to those reported for the genus Closterovirus although ORF1a harbors, in addition to a L-Pro, an AlkB domain for RNA demethylation. For species in the genus Crinivirus (Table 3), the genome is divided between two linear, positive sense, single-stranded RNAs (Fig. 4) although PYVV is an exception with a tripartite genome. All RNA molecules are needed for infectivity and are separately encapsidated. RNA1 of LIYV contains three ORFs, i.e., the ORF1a–ORF1b complex coding for the replication-related proteins, plus a 30 -most ORF coding a 34 kDa protein with no similarity to any protein in databases. This ORF is similar in size and location to ORF2 of CTV and BYSV although the respective expression products are not related. RNA1 has 50 and 30 UTRs of 97 and 219 nt, respectively. RNA2 has seven ORFs flanked by a 50 UTR of 326 nt and a 30 UTR of 187 nt. RNA2 contains the quintuple gene module, which, however, differs from that of members of the genera Closterovirus, Ampelovirus, and Velarivirus by the insertion of an extra gene (ORF4, p9-10) upstream of the CP gene. The ultimate ORF at the 30 end of RNA2 of LIYV codes p26; this protein induces plasmalemma deposits, is localized in plasmodesmata, and is required for LIYV systemic plant infection. For PYVV, RNA1 (8035 nt in size) is composed of three ORFs, i.e., the ORF1a-ORF1b complex and a 7 kDa hydrophobic protein containing a potential transmembrane helix; RNA2 (5339 nt in size) comprises five predicted ORFs that encode the HSP70h, a 7 kDa protein similar to a comparable protein of cucurbit yellow stunting disorder virus (CYSDV), the B60 kDa protein, a 9.8 kDa product with no significant similarity to any other sequence in database, and the 28.2 kDa putative CP; and RNA-3 (3892 nt) has three potential ORFs coding for a protein 4 kDa in size with no counterpart with other proteins in the family and no significant sequence homology in databases, the 77.5 kDa CPm, and a 26.4 kDa protein present in other members of the genus. Sweet potato chlorotic stunt virus (SPCSV) and Tomato chlorosis virus (ToCV) have a particularly large CPm (75–80 kDa) compared to LIYV (53 kDa). Both genomic

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Table 3 List of virus species in the four genera Closterovirus, Ampelovirus, Crinivirus and Velarivirus, in the family Closteroviridae. Their name, abbreviation and accession numbers of their genome sequence are indicated Genus name

Species name

Acronym

NCBI accession #

Closterovirus

Arracacha virus 1 Beet yellow stunt virus Beet yellows virus Blackcurrant closterovirus 1 Carnation necrotic fleck virus Carrot yellow leaf virus Citrus tristeza virus Grapevine leafroll-associated virus 2 Mint virus 1 Pistachio ampelovirus A Raspberry leaf mottle virus Rehmannia virus 1 Rose leaf rosette-associated virus Strawberry chlorotic fleck-associated virus Tobacco virus 1

AV-1 BYSV BYV BCCV1 CVFV CYLV CTV GLRaV-2 MV-1 PAVA RLMoV ReV-1 RLRaV SCFaV TV1

MG919988 U51931 AF056575 MH267701 GU234166a FJ869862 AF260651 AY881628 AY792620 MF198462 DQ357218 MH033657 KJ748003 DQ860839 KT203917

Ampelovirus

Subgroup I Blackberry vein banding-associated virus Fig leaf mottle-associated virus 2 Grapevine leafroll-associated virus 1 Grapevine leafroll-associated virus 3 Grapevine leafroll-associated virus 13 Little cherry virus 2 Pineapple mealybug wilt-associated virus 2 Pistachio ampelovirus A Subgroup II Air potato ampelovirus 1 Grapevine leafroll-associated virus 4 Pineapple mealybug wilt-associated virus 1 Pineapple mealybug wilt-associated virus 3 Plum bark necrosis stem pitting-associated virus

BVBaV FLMaV-2 GLRaV-1 GLRaV-3 GLRaV-13 LChV-2 PMWaV-2 PAVA

KC904540 FJ473383a JQ023131 AF037268 LC052212 AF531505 AF283103 MF198462

AiPoV-1 GLRaV-4 PMWaV-1 PMWaV-3 PBNSPaV

MH206615 FJ467503 AF414119 DQ399259 EF546442

Crinivirus

Bean yellow disorder virus Beet pseudoyellows virus Cucurbit yellow stunting disorder virus Cucumber yellows virus Diodia vein chlorosis virus Blackberry yellow vein-associated virus Lettuce chlorosis virus Lettuce infectious yellows virus Potato yellow vein virus Strawberry pallidosis-associated virus Sweet potato chlorotic stunt virus Tetterwort vein chlorosis virus Tomato chlorosis virus Tomato infectious chlorosis virus

BYDV BPYV CYSDV CYV DVCV BYVaV LCV LIYV PYVV SPaV SPCSV TwVSV ToCV TICV

EU191904, EU191905 AY330918, AY330919 AY242077, AY242078 AB085612, AB085613 GQ225585, GQ376201 AY776334, AY776335 FJ380118, FJ380119 U15440, U15441 AJ557128, AJ557129, AJ508757 AY488137, AY488138 AJ428554, AJ428555 KR002686, KR002687 AY903447, AY903448 FJ815440, FJ814441

Velarivirus

Areca palm velarivirus 1 Cordyline virus 1 Cordyline virus 2 Cordyline virus 3 Cordyline virus 4 Grapevine leafroll-associated virus 7 Little cherry virus 1

ArPV1 CoV-1 CoV-2 CoV-3 CoV-4 GLRaV-7 LChV-1

KR349464 HM588723 JQ599282a JQ599283a JQ599284a HE588185 Y10237

a

Partial sequence.

RNAs of ToCV encode RNA silencing suppressors, e.g., the p22 protein in RNA1, and CP and CPm in RNA2. The p25 protein of CYSDV, a viral RNAse III, and the p22 gene present in a few isolates of SPCSV also have RNA silencing suppressor activity. For the virus species in the genus Velarivirus (Table 3), Little cherry virus 1 (LChV-1) has the largest genome of the genus with 16,934 nt and nine ORFs coding for the replication associated proteins (ORF1a with a L-Pro, a MET and a HEL; and ORF1b with a

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Fig. 2 Genome organization and strategy of replication of species in the genus Closterovirus with the relative position of the open reading frames, their expression products, and 30 nested sub-genomic RNAs (sgRNAs). Information on genomic organization and sgRNAs is provided for Beet yellows virus (BYV), the type species of the genus Closterovirus: UTR, untranslated region; L-Pro, leader papain-like cysteine proteinase; MET, methyltransferase; CR-1, central region 1; CR-2, central region 2; HEL, helicase; RdRp, RNA-dependent RNA polymerase; HSP70h, heat shock protein 70 homolog; B60 kDa protein; CPm, minor coat protein; CP, coat protein. The five boxes under cell-to-cell movement represent the quintuple gene module conserved among most closteroviruses. Information on proteolytic processing of the polyprotein precursors is provided for Citrus tristeza virus (CTV).

RdRp), a small hydrophobic protein (4–8 kDa) with a transmembrane domain, the HSP70h, the B60 kDa protein, CP, CPm, and two additional proteins 25 and 27 kDa in size (Fig. 5) (ICTV, see “Relevant Websites Section”). The genome of GLRaV-7 is 16,496 nt in size and has nine ORFs encoding structural and non-structural proteins similar in type and function to those of LChV-1. Phylogenetic relationships based on HSP70h aa sequences reveal the grouping of species into four distinct genera with a few species unassigned to the family (Fig. 6). Similar trees are obtained when analyzing phylogenetic relationships based on CP or RdRp aa sequences. The genic diversity of species in the family Closteroviridae is primarily influenced by (1) strong negative selection for some viruses, (2) co-infection by genetically similar sequence variants ibn distant geographic regions, likely as a result of the extensive exchange of infected propagative planting material, (3) recombination between divergent sequence variants, (4) preferential accumulation of certain virus strains in relation to mixed infections, and (5) positive selection of sequence variants due to host change or transmission by insect vectors. From an evolutionary point of view, viruses in the family Closteroviridae represent a monophyletic virus lineage that might have evolved from a smaller filamentous virus when higher plants have differentiated. This progenitor virus is hypothesized to be composed of three genes encoding replication-associated proteins, a protein (p6) with affinity for cell membranes, and a single CP, and then to have acquired the HSP70h and a B60 kDa protein derived from a fusion of two domains, a N-terminal domain of unknown evolutionary provenance, and a duplicated CP domain. Under the pressure of additional modular evolutionary events, i.e., duplication of the CP gene, acquisition of diverse suppressors of RNA silencing and of additional genes acquired via horizontal gene transfer (e.g., papain-like cysteine proteinase and AlkB domains), this family ancestor gave rise to the progenitors of the four extant genera of the family. Species in the genus Crinivirus differentiated further by splitting their genome in two or three components. Based on their extreme genomic stability, a few species in the family Closteroviridae have been engineered as gene expression and RNA interference vectors. For instance, BYV, LIYV, CTV, GLRaV-2, and GLRaV-3 have been engineered as vectors by elegantly inserting a heterologous sgRNA promoter into an optimal genomic location to drive the expression of the green fluorescence protein reporter, or silence host genes such as the phytoene desaturase via inverted-repeat sequences. The most notable success of a closterovirus-derived vector has been achieved with GLRaV-2.

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Fig. 3 Genome organization of Grapevine leafroll-associated virus 3 (GLRaV-3), the type species of the genus Ampelovirus, showing the relative position of the open reading frames and their expression products: UTR, untranslated region; L-Pro, leader papain-like protease; MET, methyltransferase; AlkB, alpha-ketoglutarate-dependent hybroxylase domain; HEL, helicase; RdRp, RNA-dependent RNA polymerase; HSP70h, heat shock protein 70 homolog; B60 kDa protein; CP, coat protein; CPm, minor coat protein. The predicted sub-genomic RNAs (sgRNAs) are indicated below the genome map. Putative sgRNAs are indicated by dashed lines.

Infectious Cycle Positive-sense RNA viruses, such as members of the family Closteroviridae, express their replication-associated proteins in infected plant cells following translation of the genomic RNAs. The replication complex then initiates the synthesis of complementary RNA strand while remodeling host membranes, thus creating cytoplasmic compartments that are referred to as virus factories. Such compartmentalization secures RNA replication from the action of nucleases and pattern recognition receptors that trigger host innate immunity responses while providing opportunities for exchange with the cytoplasm for the export of, for instance, newly synthesized genomic and sub-genomic RNAs. The 30 UTR of the genomic RNA of species in the family Closteroviridae, particularly CTV, is highly conserved and contains several hairpin structures and a putative pseudoknot that is essential for replication, while the 50 UTR is variable but adopts a conserve secondary structure that is important for replication and proper encapsidation of the viral RNA. Replication occurs in the cytoplasm, possibly in association with endoplasmic reticulum-derived membranous vesicles and vesiculated mitochondria. Virus encoded polyproteins 1a and 1b direct membrane remodeling and formation of multi-vesicular replication factories. Polyprotein 1a of BYV contains a variable central region (CR) between the MTR and HEL domains of which a stretch (aa 1368–1432) induces the formation of 1-mm mobile globules originating from endoplasmic reticulum membranes. Part of the CR is conserved in all members of the genus Closterovirus and contains a predicted amphipathic helix (aa 1368–1385). This region may be involved in the biogenesis of closterovirus replication factories. Ultrastructural modifications arise by membrane proliferation, degeneration and vesiculation of mitochondria, and formation of inclusion bodies. These are made up of aggregates of virions or membranous vesicles, or a combination of the two. Virions accumulate in conspicuous cross-banded fibrous masses or, more typically, in more or less loose bundles intermingled with single or clustered membranous vesicles. The vesicles contain a fibrillar network and derive either from the endoplasmic reticulum, or from peripheral vesiculation of mitochondria.

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Fig. 4 Genome organization of Lettuce infectious yellows virus (LIYV), the type species of the genus Crinivirus, showing the relative position of the open reading frames and their expression products: UTR, untranslated region; L-Pro, papain-like protease; MET, methyltranferase; HEL, helicase; RdRp, RNA-dependent RNA polymerase; HSP70h, heat shock protein 70 homolog; B60 kDa protein; CP, coat protein; CPm, minor coat protein.

Fig. 5 Genome organization of Grapevine leafroll-associated virus 7 (GLRaV-7), the type species of the genus Velarivirus, showing the relative position of the open reading frames and their expression products: UTR, untranslated region; L-Pro, papain-like protease; MET, methyltranferase; HEL, helicase; RdRp, RNA-dependent RNA polymerase; HSP70h, heat shock protein 70 homolog; B60 kDa protein; CP, coat protein; CPm, minor coat protein.

For GLRaV-3, putative 30 co-terminal sgRNAs are differentially expressed with four of them accumulating at high levels (CP, p20A, p20B, p21), two at intermediate levels (p4 and p7), and others at low levels (CPm, p55, HSP70h, and p5). This suggests temporal and quantitative regulation of sgRNAs transcription during plant infection. Each of these sgRNAs serves as a monocistronic messenger for translation of the corresponding 50 -proximal ORF. The 50 transcriptional start sites were determined for several GLRaV-3 sgRNAs. Additionally, the leader sequence of sgRNAs is variable in size and does not share conserved motifs. For LIYV, the replication of both genomic RNAs is asynchronous as RNA1 genomic and sgRNAs accumulate before significant accumulation of RNA2 can be detected, suggesting that RNA1 likely replicates in cis while RNA2 replicates in trans. Knockout mutation studies showed that the single-stranded RNA-binding protein p34 that is encoded by RNA1 is a trans enhancer of RNA2 replication.

Epidemiology Transmission of viruses in the family Closteroviridae by natural vectors is semi-persistent regardless of the type of vector, i.e., aphids (genus Closterovirus), whiteflies (genus Crinivirus), and pseudococcid mealybugs and soft scale insects (genus Ampelovirus). No insect vector is known for viruses in the genus Velarivirus. Additionally, some species in the family, particularly velariviruses, can be transmitted through dodder (Cuscuta spp.). Transmission through seeds has not been documented. Only a few species of the genus Closterovirus (CTV, GLRaV-2, BYV) are transmissible by mechanical inoculation to herbaceous hosts, though with difficulty, but none of the viruses in the genera Ampelovirus, Crinivirus and Velarivirus are mechanically transmissible. Viruses in the family Closteroviridae that infect vegetatively propagated hosts are transmitted by grafting and, thus, disseminated over long distances with propagating material that is not adequately screened for their presence. The natural and experimental host ranges are usually restricted, except for a few members of the genus Crinivirus. Similarly, the geographical

344

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Fig. 6 Maximum likelihood phylogenetic tree showing the relationships among recognized members of the family Closteroviridae based on an alignment of the complete amino acid sequence of the heat shock protein 70 homolog using MUSCLE. The maximum likelihood tree was inferred using RAxML in the T-REX web server. Distances are proportional to branch lengths and the bar represents the genetic distance. The heat shock protein 70 from Arabidopsis thaliana (AEE75218) was used as outgroup. The GenBank accession number used for each virus is as follows: Actnidia virus 1 (AcV-1, KX857665), Air potato ampelovirus 1 (AiPiV-1, MH206615), arracacha virus 1 (AV-1, MG919988), Blackcurrant closterovirus 1 (BCCV1, MH267701), Bean yellow disorder virus (BYDV, EU191904), Beet pseudoyellows virus (BPYV, AY330918), Beet yellow stunt virus (BYSV, U51931), Beet yellows virus (BYV, AF056575), Blackberry vein banding-associated virus (BVBaV, KC904540), Blueberry virus A (BVA, AB733585), Carnation necrotic fleck virus (CVFV, GU234166), Carrot yellow leaf virus (CYLV, FJ869862), Citrus tristeza virus (CTV, U16304), Cordyline virus 1 (CoV-1, HM588723), Cordyline virus 2 (CoV-2, JQ599282), Cordyline virus 3 (CoV-3, JQ599283), Cordyline virus 4 (CoV-4, JQ599284), Cucurbit yellow stunting disorder virus (CYSDV, AY242077), Diodia vein chlorosis virus (DVCV, CQ376201), Fig leaf mottle-associated virus 2 (FLMaV-2, FJ473383), Fig mild mottle-associated virus (FMMaV, FJ611959), Grapevine leafroll-associated virus 1 (GLRaV-1, JQ023131), Grapevine leafroll-associated virus 2 (GLRaV-2, JX513891), Grapevine leafroll-associated virus 3 (GLRaV-3, EU259806), Grapevine leafroll-associated virus 4 (GLRaV-4, FJ467503), Grapevine leafroll-associated virus 7 (GLRaV-7, HE588185), Grapevine leafroll-associated virus 13 (GLRaV-13, LC052212), Lettuce chlorosis virus (LCV, FJ380118), Lettuce infectious yellows virus (LIYV, U15440), Little cherry virus 1 (LChV-1, EU715989), Little cherry virus 2 (LChV-2, AF531505), Mint vein banding-associated virus (MVBaV, KJ572575), Mint virus 1 (MV-1, AY792620), Palm velarivirus 1 (ArPV1, KR349464), Persimmon virus B (PeBV, AB923924), Pineapple mealybug wilt-associated 1 (PMWaV-1, AF414119), Pineapple mealybug wilt-associated 2 (PMWaV-2, AF283103), Pineapple mealybug wilt-associated 3 (PMWaV-3, DQ399259), Pistachio ampelovirus A (PAVA, MF198462), Plum bark necrosis stem pitting-associated virus (PBNSPaV, EF546442), Raspberry leaf mottle virus (RLMoV, DQ357218), Rehmannia virus 1 (ReV-1, MH033657), Rose leaf rosette-associated virus (RLRaV, KJ7488003), Strawberry chlorotic fleck-associated virus (SCFaV, DQ860839), Potato yellow vein virus (PYVV, AJ557128), Strawberry pallidosis-associated virus (SPaV, AY488138), Sweet potato chlorotic stunt virus (SPCSV, AJ428554), Tetterwort vein chlorosis virus (TwVSV, KR002687), Tobacco virus 1 (TV1, KT203917), Tomato chlorosis virus (ToCV, AY903447), and Tomato infectious chlorosis virus (TICV, FJ815440).

distribution of viruses varies from restricted [Grapevine leafroll-associated virus 13 (GLRaV-13), Mint virus 1 (MV-1), Tobacco virus 1 (TV1)] to widespread (CTV, GLRaV-2, GLRaV-3, BYV). The range of vectors for species in the genus Closterovirus varies from rather wide to restricted. For example, BYV is transmitted by at least 23 aphid species (Myzus persicae and Aphis fabae being the main natural vectors), CTV by seven species (Toxoptera citricida and Aphis gossypii being the most efficient vectors), MV-1 by a single aphid species while GLRaV-2 does not have any known vectors. Recent studies of specific interactions between CTV and its aphid vector, Toxopetra citricida, indicate the cibarium of the

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aphid foregut as the virus retention site, and the critical role of a protein-carbohydrate complex for transmission with three viral proteins, i.e., the CPm, p61, and p65, binding to sugar moieties on the surface of the foregut. For species in the genus Ampelovirus, the range of vectors varies from large to narrow. For instance, GLRaV-1 is transmitted by species of several genera of pseudococcid mealybugs (Heliococcus, Phenacoccus, Pseudococcus) and soft scale insects (Pulvinaria, Neopulvinaria and Parthenolecanium). Similarly, GLRaV-3 is transmitted by a wide range of pseudococcid mealybugs (Planococcus, Pseudococcus, Heliococcus, Phenacoccus) and soft scale insects (Pulvinaria, Neopulvinaria, Parthenolecnium, Coccus, Saissetia, Parasaissetia, Ceroplastes). To the contrary, vectors of PMWaV-1, PMWaV-2 and PMWaV-3 are two species of the genus Dysmicoccus, and LChV-2 is transmitted by Phenacoccus aceris. Species in the genus Crinivirus are transmitted by whiteflies of the genera Trialeurodes and Bemisia. The specificity of transmission has been used for species differentiation. For example, viruses of group 1 [PYVV, Blackberry yellow vein-associated virus (BYVaV), Beet pseudoyellows virus (BPYV) and Strawberry pallidosis-associated virus (SpaV)] are transmitted by T. vaporariorum, viruses of group 2 [ToCV, SPCSV, CYSDV, and Bean yellow disorder virus (BYDV)] by B. tabaci, whereas one of the viruses of group 3 is transmitted by B. tabaci (LIYV) and the other by T. vaporariorum (TICV). These three groups are sustained by comparative phylogenetic analyses of RdRp aa sequences.

Disease Symptoms Viruses in the family Closteroviridae infect fruit, vegetable, tuberous, and berry crops, as well as herbs, flowering plants, shrubs, and weeds. They cause foliar discolorations (chlorosis, discoloration, or reddening) and rolling (Fig. 7), stem pitting and grooving on the woody cylinder of woody hosts, stunting, or symptomless infections. Disease symptoms also include small and late ripening fruits.

Pathogenesis Plant infection by species in the family Closteroviridae is systemic but virions are usually found in the phloem (sieve tubes, companion cells and parenchyma), and only occasionally in the mesophyll and epidermis. Virus-host interactions have been

Fig. 7 Disease symptoms caused by selected viruses of the family Closteroviridae on their natural host. (A) Downward rolling and reddening of the leaf blade of Vitis vinifera cv. Pinot noir infected with Grapevine leafroll-associated virus 3 from the genus Ampelovirus, (B) Aerial view of declining and dead citrus trees grafted onto the sour orange rootstock in a grove infected with Citrus tristeza virus from the genus Closterovirus. Infected trees show necrosis at the graft union with reduced carbohydrate movement into the roots, and leaves fall off (courtesy of W.O. Dawson, University of Florida, USA), (C) Interveinal discoloration of melon leaves infected by Cucurbit yellow stunting disorder virus from the genus Crinivirus (courtesy of W. Wintermantel, USDA-ARS, Salinas, California, USA), (D) Discoloration of tomato leaves infected by Tomato chlorosis virus from the genus Crinivirus (courtesy of W. Wintermantel, USDA-ARS, Salinas, California, USA), and (E) Tomato infectious chlorosis virus from the genus Crinivirus (courtesy of W. Wintermantel, USDA-ARS, Salinas, California, USA). Note the similarities of symptomatology caused by ToCV and TICV, and by magnesium deficiency.

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Closteroviruses (Closteroviridae)

extensively studied for CTV. Protein p23 is a multifunctional RNA-binding protein with a putative zinc finger domain that accumulates in the nucleolus and plasmodesmata. It regulates the asymmetric balance of plus and minus RNA strands during replication, induces disease symptoms similar to those caused by CTV infection in certain hosts, and enhances systemic infection, and virus accumulation. Recently, it was shown that CTV uses the cytosolic glyceraldehyde 3-phosphate dehydrogenase via interaction with p23 to facilitate infection. In addition, p23 like p25 and p20 is a suppressor of RNA silencing. Proteins p33, p18, and p13 are dispensable for systemic infection of certain hosts, and involved in stem pitting disease symptoms.

Diagnosis The visual identification of disease symptoms incited by viruses in the family Closteroviridae can be challenging due to confounding factors such as asymptomatic infections, virus co-infections, or similarities with nutritional deficiencies. Several techniques have been developed and applied to diagnose closteroviruses, including biological assays, serological assays, nucleic acid-based methods, and large-scale sequencing. Biological indexing consists of chip budding a candidate plant to a disease susceptible, indicator plant. The indicator plant with the newly grafted material is then observed for the development of disease symptoms. This approach is labor intensive, time consuming, costly, and often not very robust. Biological assays are progressively replaced by more powerful diagnostic assay that identify viruses rather than diseases. Serological diagnostic techniques such as enzyme-linked immunosorbent assays have been developed for simple, cost-effective and high-throughput testing. Although virion proteins are often moderately antigenic, serological reagents are available for many viruses in the family Closteroviridae, and most species within genera are serologically unrelated or distantly related to one another, for instance, GLRaV-1 and GLRaV-3, or GLRaV-1 and GLRaV-13, but no intergeneric serological relationship has been detected. Nucleic acid-based diagnostic methods are increasingly used for the detection of closteroviruses. Such assays include reverse transcription (RT) polymerase chain reaction (PCR) and variations thereof (immunocapture-RT-PCR, spot RT-PCR, RT-qPCR), loop-mediated amplification of nucleic acid, and oligonucleotide microarrays. These assays are often more robust than serological assays. More recently, large scale sequencing is increasingly used for discovery of new viruses, including closteroviruses, and for an unbiased diagnosis of a virome in a plant sample without any prior knowledge of the pathogens present. This technology should facilitate the safe and rapid exchange of propagative plant material.

Treatment Management of diseases caused by viruses in the family Closteroviridae is challenging, due mainly to the absence of curative practices. Strategies to mitigate the impact of viruses rely almost exclusively on preventive approaches.

Prevention Screening nursery stocks for viruses is critical for the production of clean planting material and the establishment of clean production fields. Planting material derived from virus-tested nursery stocks minimizes virus incidence. Technological advances that enable the efficient, rapid identification and removal of virus-infected stocks facilitate disease management tactics. Additionally, reducing disease inoculum via, for instance, rogueing, is critical to limit disease spread, especially in perennial crops.

Further Reading Al Rwahnih, M., Dolja, W.W., Daubert, S., Koonin, E.V., Rowhani, A., 2012. Genomic and biological analysis of Grapevine leafroll-associated virus 7 reveals a possible new genus within the family Closteroviridae. Virus Research 163, 302–309. Agranovsky, A.A., 2016. Closteroviruses: Molecular biology, evolution and interactions with cells. In: Gaur, R., Petrov, N., Patil, B., Stoyanova, M. (Eds.), Plant Viruses: Evolution and Management. Singapore: Springer. Dawson, W.O., Bar-Joseph, M., Garnsey, S.M., Moreno, P., 2015. Citrus tristeza virus: Making an ally from an enemy. The Annual Review of Phytopathology 53, 137–155. Dolja, V.V., Koonin, E.V., 2013. The closterovirus-derived gene expression and RNA interference vectors as tools for research and plant biotechnology. Frontiers in Microbiology 4. doi:10.3389/fmicb.2013.00083. Flores, R., Ruiz-Ruiz, S., Soler, N., et al., 2013. Citrus tristeza virus p23: A unique protein mediting hey virus-hoist interactions. Frontiers in Microbiology 4. doi:10.3389/ fmicb.2013.00098. Gushchin, V.A., Karlin, D.G., Makhotenko, A.V., et al., 2017. A conserved region in the closterovirus 1a polyprotein derives extensive remodeling of endoplasmic reticulum membranes and induces motile globules in Nicotiana benthamiana cells. Virology 502, 106–113. Jarugula, S., Gowda, S., Dawson, W.O., Naidu, R.A., 2018. Development of infectious cDNA clones of Grapevine leafroll-associated virus 3 and analyses of the 5’ nontranslated region for replication and virion formation. Virology 523, 89–99. Killi, N., Harper, S.J., Alfaress, S., El Mohtar, C., Dawson, W.O., 2016. Minor coat and heat shock proteins are involved in the binding of Citrus tristeza virus to the foregut of its aphid vector, Toxoptera citricida. Applied and Environmental Microbiology 82, 6294–6302. Kiss, Z., Medina, V., Falk, B.W., 2013. Crinivirus replication and host interactions. Frontiers in Microbiology 4. doi:10.3389/fmicb.2013.00099. Kurth, E.G., Peremyslov, V.V., Prokhnevsky, A.I., et al., 2012. Virus-derived gene expression and RNA interference vector for grapevine. Journal of Virology 86, 6002–6009.

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Maree, H.J., Almeida, R.P.P., Bester, R., et al., 2013. Grapevine leafroll-associated virus 3. Frontiers in Microbiology 4. doi:10.3389/fmicb.2013.00082. Qiao, W., Medina, V., Kuo, Y.-W., Falk, B.W., 2018. A distinct, non-virion plant virus movement protein encoded by a crinivirus essential for systemic infection. mBio 9 (6), e02230. Rubio, L., Guerri, J., Moreno, P., 2013. Genetic variability and evolutionary dynamics of viruses of the family Closteroviridae. Frontiers in Microbiology 4. doi:10.3389/ fmicb.2013.00151. Ruiz-Ruiz, S., Navarro, B., Peña, L., et al., 2019. Citrus tristeza virus: Host RNA silencing and virus counteraction. In: Catara, A.F., Bar-Joseph, M., Licciardello, G. (Eds.), Citrus Tristeza Virus: Methods and Protocols. Methods in Molecular Biology 2015. New York: Humana, pp. 195–207. Tzanetakis, I.E., Martin, R.R., Wintermental, W.M., 2013. Epidemiology of criniviruses: An emerging problem in world agriculture. Frontiers in Microbiology 4. doi:10.3389/ fmicb.2013.00119.

Relevant Websites http://www.dpvweb.net/dpv/showdpv.php?dpvno=377 Beet yellows virus Show DPV and Refs in Frame. https://talk.ictvonline.org/ictv-reports/ictv_9th_report/positive-sense-rna-viruses-2011/w/posrna_viruses/255/closteroviridae Closteroviridae Positive Sense RNA Viruses. https://www.ncbi.nlm.nih.gov/genomes/GenomesGroup.cgi?taxid=69973 Complete genomes: Closteroviridae NCBI.

Comoviruses and Fabaviruses (Secoviridae) George P Lomonossoff, John Innes Centre, Norwich, United Kingdom r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

nt Nucleotide(s) ORF Open reading frame RdRp RNA-dependent RNA polymerase UTR Untranslated region VLPs Virus-like particles VPg Viral protein genome-linked

aa Amino acid(s) Co-Pro Protease cofactor CP Coat protein or capsid protein kb Kilobase kDa Kilo Dalton MP Movement protein

Glossary Viral protein genome-linked A protein covalently attached to the 50 end of positive strand viral RNA and acting as a primer during RNA synthesis. Virus re-assortment The mixing of the genome segments of a virus into new combinations.

Virus replication complexes Intracellular complexes formed by association with intracellular membranes that are sites for stages in the replication cycle of a virus. b-barrel Beta-sheet composed of tandem repeats forming a closed structure.

Introduction The genera Comovirus and Fabavirus are non-enveloped plant viruses that are classified within the the family Secoviridae. They typically cause mottle or mosaic symptoms on susceptible hosts and are characterized by having a bipartite RNA genome with the individual components separately encapsidated in isometric particles (Fig. 1). Though having similar genetic and particle structures, the two genera are distinguished on the basis of their insect vectors.

Taxonomy and Classification The genus Comovirus is named after the type species, Cowpea mosaic virus (CPMV). The genus Fabavirus derives its name from the host species (Vicia faba) from which the first member of the genus, Broad bean wilt virus (BBWV), was originally isolated as the causative agent of vascular wilt symptoms. At present, 15 and 7 distinct virus species are recognized by the International Committee for the Taxonomy of Viruses (ICTV) as being members of the genus Comovirus and Fabavirus, respectively (Table 1). Together with the genus Nepovirus, they are classified in the subfamily Comovirinae within the family Secoviridae. Comoviruses and fabaviruses are insect-transmitted by beetles and aphids, respectively, while nepoviruses are nematode-transmitted. The relationship between members of the two genera and their relationship to other members of the family Secoviridae is shown in (Fig. 2).

Fig. 1 Fundamental features of comoviruses and fabaviruses. (A) Symptoms of the fabavirus, BBWV-1, on Vicia faba. (B) The three-component nature of comoviruses and fabaviruses. The photograph shows fractionation of CPMV on a cesium chloride gradient. The positions of the three nucleoprotein components and their RNA contents are indicated. Photo in (A) courtesy of Dr. I. Ferriol, CRAG, Barcelona, Spain

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Table 1 List of species members of the genera Comovirus and Fabavirus in the family Secoviridae. The genera type species are written in bold. The information regarding their molecular biology is indicated for each member Genus/Species

Acronym Genome sequence Accession #

Infectious clones Particle structure Biotechnology use

Comovirus Andean potato mottle virus

APMoV

Complete

Bean pod mottle virus

BPMV

Complete

Bean rugose mosaic virus

BRMV

Complete

Broad bean stain virus

BBSV

Partial

Broad bean true mosaic virus BBTMV

Complete

Cowpea mosaic virus

CPMV

Complete

Cowpea severe mosaic virus

CPSMV

Complete

Glycine mosaic virus Pea green mottle virus Pea mild mosaic virus Quail pea mosaic virus Radish mosaic virus

GMV PGMV PMiMV QPMV RaMV

Complete

Red Clover mottle virus

RCMV

Complete

Squash mosaic virus

SqMV

Complete

Ullucus virus C

UVC

RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA

1: 2: 1: 2: 1: 2: 1: 2: 1: 2: 1: 2: 1: 2:

MN176101 MN176102 NC003496/U70866 NC003495/M62738 NC028139/KP404602 NC028146/KP404603 NC043386/KJ746622 NC043385/FJ028650 NC022004/GU810903 NC022006/GU810904 NC003549/X00206 NC003550/X00729 NC003545/M83830 NC003544/M83309

RNA RNA RNA RNA RNA RNA

1: 2: 1: 2: 1: 2:

NC010709/AB295643 YES NC010710/AB295644 NC003741/X64886 NC003738/M14913 NC003799/AB054688 NC003800/AB054689

RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA

1: 2: 1: 2: 1: 2: 1: 2: 1: 2: 1: 2: 1: 2:

NC005289/AB084450 YES NC005290/AB084451 NC003003/AF225953 YES NC003004/AF225954 NC038760/EU881936 NC038759/EU881937 AB084452 AB084453 NC039073/KX241482 NC039072/KX241485 NC023016/KC595304 NC023017/KC595305 NC039077/KX269865 NC039078/KX269871

YES

YES

YES

YES YES

YES

YES

YES

YES

YES

Fabavirus Broad bean wilt virus 1

BBWV-1 Complete

Broad bean wilt virus 2

BBWV-2 Complete

Cucurbit mild mosaic virus

CuMMV

Complete

Gentian mosaic virus

GeMV

Complete

Grapevine fabavirus

GFabV

Nearly complete

Lamium mild mosaic virus

LMMV

Complete

Prunus virus F

PrVF

Complete

In several instances, individual comoviruses and fabaviruses have been shown to exist in several different strains. For example, three distinct strains of Red Clover mottle virus (RCMV), termed -S, -N and -O, have been described. In the case of Bean pod mottle virus (BPMV) at least two distinct strains have been reported and these can reassort. Detailed molecular characterization of BPMV reassortants has revealed the presence of strains that are diploid for RNA1 and haploid for RNA2. The fabaviruses, BBWV-1 and BBWV-2 were originally classified as strains of the same species but are now considered to be distinct species.

Physical Properties of Viral Particles The yields of comovirus particles from infected tissue are typically higher than those of fabaviruses, particles of which have a tendency to aggregate on purification. The thermal inactivation point of viral particles in plant sap is in the range 60–751C and the particles have a longevity in sap of between 2 and 112 days at room temperature. Comovirus and fabavirus preparations consist of non-enveloped isometric particles, approximately 30 nm in diameter, which can be separated into three components designated top (T), middle (M) and bottom (B) by centrifugation on sucrose density

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Fig. 2 Phylogenetic relationships within the family Secoviridae. The comoviruses and fabaviruses form two distinct branches within the subfamily Comovirinae, which includes the nepoviruses. Alignment figure courtesy of Prof. J. Thompson.

gradients (Fig. 1(B)). The exact sedimentation coefficients of the components vary slightly with the virus being analyzed but are typically in the range 50-60S for T, 90-100S for M and 110-120S for B components. The three components have identical protein compositions, containing 60 copies each of a large (L) and small (S) coat protein (CP). The sizes of the two proteins vary depending on the virus but lie in the ranges 37–49 kDa for the L protein and 18–26 kDa for the S protein. The difference between the three centrifugal components is in their RNA contents. Top components are devoid of RNA, while middle and bottom components each contain single molecules of RNA of approximately 3–4 kb and 6 kb, respectively. The two RNA molecules were originally termed middle (M) and bottom (B) component RNA after the nucleoprotein component from which they were isolated but are now termed RNA2 and RNA1, respectively (Fig. 3(A)). Because of their differing RNA contents, the three components of comoviruses and fabaviruses also differ in density and can hence be separated by isopycnic centrifugation. Generally, T, M and B components have densities of approximately 1.29, 1.40 and 1.41 g ml1 respectively. However, the pattern obtained is often more complex than that seen with sucrose gradients and depends on the precise conditions used and virus being examined. For example, while BPMV gives the expected three components when centrifuged on cesium chloride gradients under a wide range of conditions, the bottom component of CPMV can be resolved into two forms of differing density under alkaline conditions. Preparations of comoviruses are often not only centrifugally heterogeneous but can also be separated in two forms, fast and slow, electrophoretically. Both electrophoretic forms contain all three centrifugal components. The proportion of the two electrophoretic forms in a given viral preparation varies both with the time after infection at which the virus was isolated and the age of the preparation itself. Conversion of one form to the other is caused by loss of amino acids from the C terminus of the S protein. In the case of CPMV, this segment has been shown to be critical for virus assembly, and structural studies on Broad bean stain virus (BBSV) suggest that comoviruses share a common assembly mechanism. Though not studied extensively, it is likely that fabavirus particles also undergo a similar cleavage of the S protein. In a number of instances, comovirus particles have been shown to contain polyamines, particularly spermidine. Such polyamines are believed to have a role in the neutralization of the negative charges of the RNA within the particles. It is the exchange of such polyamines for cesium ions that leads to the two forms of bottom component of CPMV seen under alkaline conditions.

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Fig. 3 Molecular details of comoviruses and fabaviruses. (a) Organization of the two single-stranded RNA molecules that comprise the genome. Both RNAs have a VPg (red circle) linked to their 50 ends and both are polyadenylated. RNA1 contains a single ORF that is processed by the 24K protease (Pro) to give the viral proteins required for RNA replication. The functions of the individual processed products are indicated. RNA2 encodes the structural CPs, Large (L) and Small (S) as well as the MP required for the movement of CPMV virions from cell-to-cell. The solid line indicates translation from the upstream initiation codon that results in an N-terminal extension to the MP, while translation from the downstream initiation codon indicated by the dotted line results in the MP. (b) Arrangement of the CP subunits of comoviruses and fabaviruses. The asymmetric unit consists of one single-domain S subunit (blue) and one two-domain L subunit (green and red), 60 copies of which are arranged with icosahedral symmetry to form a pseudo T ¼ 3 (P ¼ 3) capsid. (c) Ribbon diagram of the asymmetric unit, showing b-barrel structure of the L (green and red) and S (blue) subunits.

Viral Structure X-ray crystallographic and cryo-EM studies on CPMV, BPMV, RCMV and BBSV have provided a detailed picture of the arrangement of the two viral CPs in the three-dimensional structure of comoviruses. Though, at the time of writing, there is no detailed structure of a fabavirus available, it is highly probably that their overall architecture is very similar. Comovirus virions are icosahedrally symmetrical, with 12 axes of fivefold and 20 axes of threefold symmetry and resemble a classic T ¼ 3 particle (Fig. 3(B)). The two CPs taken together consist of three distinct b-barrel domains, two being derived from the L and one from the S protein. Thus, in common with the T¼ 3 viruses, each CPMV particle is made up of 180 b-barrel structures. The S protein, with its single domain, is found at the fivefold symmetry axes and therefore occupies a position analogous to that of the A type subunits in T ¼ 3 particles (Fig. 3(c)). The N- and C-terminal domains of the L protein occur at the threefold axes and occupy the positions equivalent to those of the C and B type subunits of a T ¼ 3 particle respectively. Comovirus (and, by inference, fabavirus) capsids are also structurally homologous to those of the picornaviruses, with the N- and C-terminal domains of the L protein being equivalent to viral protein VP2 and viral protein VP3, respectively, and the S protein being equivalent to viral protein VP1. In the case of BPMV, ordered segments of RNA2 could be seen in the crystallographic structure of M components. RNA has also been detected in the cryo-EM structures of the M and B components of CPMV and in an unfractionated mixture of BBSV.

Genome Structure The genomes of both comoviruses and fabaviruses consist of the two RNAs (RNA1 and RNA2) encapsidated in bottom and middle components. Both RNA molecules are positive-sense and both are required for an infection of whole plants. In the case of CPMV, RNA1 is capable of independent replication in individual plant cells but this results in the establishment of gene silencing in the absence of RNA2, which encodes a suppressor of gene silencing. Both genomic RNAs have a small basic protein (VPg) covalently linked to their 50 termini and are polyadenylated. The VPg is linked to the viral RNA via the b-hydroxyl group of its N-terminal serine residue. The VPg is not required for the viral RNAs to be infectious. The complete nt sequences of both RNAs of most comoviruses and fabaviruses have been determined (Table 1); in addition, several partial sequences are also available. The two genomic RNAs of a given virus have no sequence homology apart from at the 50 and 30 termini. Full-length infectious cDNA clones of several viruses have been constructed (Table 1), allowing the genomes of these viruses to be manipulated.

Expression of the Viral Genome Both genomic RNAs contain a single long open reading frame (ORF) which occupies over 80% of the length of the RNA. A combination of in vitro translation and protoplast studies has unraveled the basic mechanism of expression of the genomic RNAs

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of CPMV, the type species of the genus Comovirus. More limited studies on other members of the group suggest that the mode of gene expression deduced for CPMV is applicable to all members of the genera. Both RNAs of the comoviruses and fabaviruses are expressed through the synthesis and subsequent cleavage of large precursor polyproteins. On RNA1, initiation of translation occurs at the first AUG encountered on the sequence and results in the synthesis of a protein of approximately 200 kDa. This initial product undergoes rapid co-translational autopoteolysis to give proteins with apparent sizes of 32 and 170 kDa (the 32 and 170K proteins). The 170K protein undergoes further cleavages to give the range of virus-specific proteins, the functions of which are indicated in Fig. 2(A). The proteolysis of the polyprotein is catalysed by the region marked Pro in Fig. 2(A). Initiation of translation of RNA2 occurs at two different positions on the RNA and results in the synthesis of two carboxy coterminal proteins. This double initiation phenomenon, which occurs as a result of “leaky scanning” of two in-frame AUG codons, is found with the RNA2 molecules of all comoviruses and fabaviruses. The distance between the two initiation codons varies significantly between different viruses, as does the sequence of the encoded aa. Both RNA2-encoded primary translation products are cleaved by the RNA1-encoded Pro to give the movement protein (MP) and a version with an N-terminal extension as well as the two viral CPs, L and S. Processing of the RNA2-encoded polyproteins at the site between MP and L CP also requires the presence of the protease cofactor (ProC) as well as Pro.

Functions of the Viral Proteins Functions have been ascribed to most of the regions of the polyproteins encoded by both RNA1 and RNA2 based on extensive investigations with CPMV. Far less information is available for other comoviruses and fabaviruses, with what little that is available indicating that the analyzes conducted with CPMV are generally applicable. Therefore, the reader is referred to the article on CPMV for a detailed description of the functions of the viral proteins.

Relationships With Other Viruses Together with the genus Nepovirus, the genera Comovirus and Fabavirus are grouped within the subfamily Comovirinae, in the family Secoviridae. Consideration of genome structure and organization, translational strategy and aa homologies between the virus-encoded proteins has led to the family Secoviridae being grouped with the family Picornaviridae, within the order Picornavirales. Members of this order are all non-enveloped icosahedral positive-strand RNA viruses with 30 -polydenylated genomic RNAs which have the VPg covalently linked to their 50 ends. All members of the order have a similar mode of gene expression which involves the synthesis of large precursor polyproteins and their subsequent cleavage by a virus-encoded proteinase. The members of the order, all contain the similar gene order, membrane-bound protein-VPg-proteinase-polymerase, a feature shared with members of the filamentous family Potyviridae.

Use in Biotechnology CPMV has been extensively used as a vector for the expression of foreign peptides and proteins in plants. The only other comovirus that has been deployed in biotechnology is BPMV. Several BPMV RNA2-based vectors have been developed which permit the insertion of foreign sequences for either protein expression or gene silencing in soybean. At present there are no reports of the use of a fabavirus in biotechnology.

Geographic Distribution Individual comoviruses tend to have a somewhat restricted distribution. Clearly, the apparent distribution of a virus is governed, at least in part, by whether or not infection is diagnosed and reported. The following list, therefore, contains only those geographical areas where the presence of the virus has, to date, been confirmed. CPMV: Nigeria, Kenya, Tanzania, Japan, Surinam and Cuba; Andean potato mottle virus (APMV): Central and South America; BPMV: USA; Bean rugose mosaic virus (BRMV): Central America; BBSV: Europe and Africa; Broad bean true mosaic virus (BBTMV): Europe and North-west Africa; Cowpea severe mosaic virus (CPSMV): North, Central and South America; Glycine mosaic virus-GMV: Australia; Pea mild mosaic virus (PMiMV): New Zealand; Quail pea mosaic virus (QPMV): North and Central America; Radish mosaic virus (RaMV): USA, Japan and Europe; RCMV: Europe; Squash mosaic virus (SqMV): North Africa, Israel, China, Japan, Australia and North, Central and South America; Ullucus virus C (UCV): Peru and Bolivia. Fabaviruses occur throughout the world and a wide variety of different isolates have been reported from different geographical regions have been reported.

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Host Range The host ranges of individual comoviruses tend to be rather narrow. Of the 15 members of the genus, ten (CPMV, BPMV, BRMV, BBSV, BBTMV, CPSMV, GMV, Pea green mottle virus-PGMV, PMiMV, QPMV and RCMV), in nature, infect predominantly, if not exclusively, legumes. The other four members of the group, APMV, RaMV, SqMV and UCV, infect predominantly members of the Solanaceae, Cruciferae, Cucurbitaceae and Basellaceae respectively. By contrast, the experimental host range of Fabaviruses is wide: 328 species of 186 genera in 44 families have been listed as infectable by BBWV. Lamium mild mosaic virus (LMMV) has been less extensively tested but differs from BBWV in not infecting, for example, cowpea (Vigna unguiculata) or tomato (Lycopersicum officinale cv. Marmande).

Transmission In nature, comoviruses are usually transmitted by leaf-feeding beetles, especially by members of the family Chrysomelidae. However, CPMV has been shown also to be transmitted by thrips and grasshoppers. The beetle vectors can acquire the virus by feeding for as little as one minute and can retain and transmit the virus for a period of days or weeks. Experimentally, all comoviruses are mechanically transmissible. In the case of BBSV, BBTV, CPMV, CPSMV and SqMV, transmission through seed has been reported. Fabaviruses are transmitted by aphids in a non-persistent manner. The list of known vector aphids extends to some 20 species including Acyrthosiphon pisum, Aphis craccivora, A. nasturtii and Macrosiphum euphorbiae. However, there are unpredictable and unexplained differences between fabavirus isolates in the efficiencies of their transmission by different aphid species. There is only one report showing that BBWV is seed-transmitted. The transmission of fabaviruses by aphids is not known to require the presence in source plants of another 'helper' virus.

Epidemiology The primary sources for infection of crop plants with comoviruses are transmission through seed and infection from wild hosts act as reservoirs of the viruses. It is also possible that virus which has overwintered in the beetle vector can act as a primary source of infection. Within-field spread is by the beetle vectors. Fabaviruses can be readily spread by their aphid vectors.

Symptomatology and Cytopathology Generally, each virus has certain hosts in which the virus can spread systematically, causing mosaic or mottling symptoms, and other hosts in which the infection is confined to local lesions. Stunting of the host plant is sometimes observed, as is occasional systematic wilting and seed discoloration. Infection of plant cells with comoviruses results in a number of characteristic cytological changes. These include the appearance of viral particles, either individually or as crystalline arrays, in the cytoplasm, a proliferation of cell membranes and vesicles in the cytoplasm, the appearance of amorphous inclusion bodies near or surrounding the nucleus and a variety of modifications to plasmodesmata. Virions of all the fabaviruses occur in the cell cytoplasm of foliar mesophyll and epidermis. In some isolates, virions, virus-like shells and probably also capsid protein assemble into a variety of ordered arrays (tubules, rings, square patterns) that can be recognized by electron microscopy. Amorphous X bodies (spindle-shaped fibrous or crystalline inclusions visible using light microscopy) may have some diagnostic value.

Economic Importance At present, BPMV, CPMV, CPSMV and RCMV are considered to be significant pathogens of legumes. Infection of soybeans with BPMV alone can cause yields to be reduced by 10%–17%. However, this figure can rise to 60% in plants doubly infected with BPMV and the potyvirus, Soybean mosaic virus. In Nigeria, infection of cowpeas with CPMV causes a great reduction in leaf area, flower production and yield. Infection of cowpeas with CPSMV has been shown to cause a 50% reduction in plant fresh weight and in the number and weight of pods. RCMV is of economic importance in forage production as it sometimes heavily invests clover. SqMV is a significant pathogen of cucurbits, problems associated with it being exacerbated by its high rate of seed transmission. Fabaviruses kill lettuce, bean and spinach plants and cause necrotic streaks in foliage of some pea cultivars. Symptoms in other crops are usually transient and have limited commercial significance – except in the People's Republic of China where complex intercropping patterns favor damaging epiphytotics also involving celery (Apium graveolensis) and pakchoi (Brassica chinensis).

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Further Reading Cooper, J.I., Lomonossoff, G.P., 2000. Fabaviruses. In: Maloy, O.C., Murray, T.D. (Eds.), Encyclopedia of Plant Pathology. New York, NY: John Wiley and Sons, Inc., pp. 445–446. Ferriol, I., Ferrer, R.M., Luis-Arteaga, M., et al., 2014. Genetic variability and evolution of Broad bean wilt virus 1: Role of recombination, selection and gene flow. Archives of Virology 159, 779–784. Gergerich, R.C., Scott, H.A., 1996. Comoviruses: Transmission, epidemiology, and control. In: Harrison, B.D., Murant, A.F. (Eds.), The Plant Viruses 5: Polyhedral virions and bipartite RNA genomes. New York: Plenum Press, p. 77. Ghabrial, S.A., 2018. My life and virus research journey. Annual Review of Virology 5, 1–32. Goldbach, R.W., Wellink, J., 1996. Comoviruses: Molecular biology and replication. In: Harrison, B.D., Murant, A.F. (Eds.), The Plant Viruses 5: Polyhedral virions and bipartite RNA genomes. New York: Plenum Press, p. 35. Kobayashi, Y.O., Kobayashi, A., Nakano, M., et al., 2003. Analysis of genetic relations between broad bean wilt virus 1 and broad bean wilt virus 2. Journal of General Plant Pathology 69, 320–326. Lecorre, F., Lai-Kee-Him, J., Blanc, S., et al., 2019. The cryo-electron microscopy structure of Broad bean stain virus suggests a common capsid assembly mechanism among comoviruses. Virology 530, 75–84. Lin, T., Johnson, J.E., 2003. Structure of picorna-like plant viruses: Implications and applications. Advances in Virus Research 62, 167–239. Lisa, V., Boccardo, G., 1996. Fabaviruses: Broad bean wilt and allied viruses. In: Harrison, B.D., Murant, A.F. (Eds.), The Plant Viruses 5: Polyhedral virions and bipartite RNA genomes 1996. New York: Plenum Press, pp. 229–250. Lomonossoff, G.P., Ghabrial, S.A., 2000. Comoviruses. In: Maloy, O.C., Murray, T.D. (Eds.), Encyclopedia of Plant Pathology. New York, NY: John Wiley and Sons, Inc, pp. 239–242. Pflieger, S., Blanchet, S., Meziadi, C., et al., 2014. The "one-step" bean pod mottle virus (BPMV)-derived vector is a functional genomics tool for efficient overexpression of heterologous protein, virus-induced gene silencing and genetic mapping of BPMV R-gene in common bean (Phaseolus vulgaris L.). BMC Plant Biology 14, 232–248. Zhang, C., Ghabrial, S.A., 2006. Development of bean pod mottle virus-based vectors for stable protein expression and sequence-specific virus-induced gene silencing in soybean. Virology 344, 401–411.

Relevant Websites https://viralzone.expasy.org/728?outline=all_by_species Comovirinae. https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/picornavirales/w/secoviridae/589/genus Comovirus. http://viperdb.scripps.edu/genuslist.php?genus=Comovirus Virus Genus Index VIPERdb Scripps Research.

Cotton Leaf Curl Disease (Geminiviridae) Nasim Ahmed, Imran Amin, and Shahid Mansoor, National Institute for Biotechnology and Genetic Engineering, Faisalabad, Pakistan r 2021 Elsevier Ltd. All rights reserved. This is an update of S. Mansoor, I. Amin, R.W. Briddon, Cotton Leaf Curl Disease, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00703-2.

Nomenclature aa Amino acid(s) amiRNA Artificial microRNA CP Coat protein or capsid protein DAS Days after sowing kb Kilobase kDa Kilo Dalton mRep Master replication-associated protein nm Nanometer(s) nt Nucleotide(s)

Glossary Epidemic A wide spread of an infectious disease in the population at a specific time. Host plant resistance The natural host plant species have the ability to control the parasite population. This is the most desired approach to control disease. Koch’s postulates The criteria first time developed in 1890 by Robert Koch that establishes the relationship between a disease and microbe. Following conditions must be fulfilled; the pathogen should be present in all symptomatic hosts, and after its isolation from the host it should be able to grow in pure culture, and when

NW New world OW Old world PDR Pathogen derived resistance PTGS Post-transcriptional gene silencing Rep Replication initiator protein SCR Satellite conserved region ssDNA Single-stranded deoxyribonucleic acid TGS Transcriptional gene silencing TrAP Transcriptional activator protein

reintroduced into a healthy host it must cause the disease. At the end, the pathogen must be isolated from this infected host. Pathogen-derived resistance The type of resistance to specific pathogens that is created by engineering host to express genes or sequences derived from the pathogen, it interferes with the infection caused by this pathogen or its most related pathogen which has the same sequence. Resistance breaking strain A strain of pathogen which breaks the resistance of host against a specific pathogen.

Introduction Cotton leaf curl disease (CLCuD) is the most destructive disease of cotton (Gossypium spp. L.) and also for several other plants in the family Malvaceae. The disease is widespread in Africa (mainly Sudan, Egypt, Malawi, Nigeria and South Africa) and southern Asia where it is endemic across Pakistan and northwestern India. Recently, it is also been reported from southern China in noncotton plants. Typical symptoms exhibited by CLCuD affected plants include vein thickening, upward or downward leaf curling, and on the main vein (underside the leaves) enations are formed that evolve into cup-shaped leaf-like structures (Fig. 1). CLCuD affected cotton plants are conspicuously greener than non-infected plants which is attributed to a higher density of chloroplasts in infected tissues. Symptoms of CLCuD vary between different varieties of cotton and also depend on the age of the infected plants. Plants infected at the early stage show severe symptoms like tightly rolled leaves, highly stunted growth and subsequently they produce no harvestable lint whereas plants which are infected at the late stage show just mild symptoms and their yield is not significantly reduced. CLCuD is caused by begomoviruses (genus Begomovirus, family Geminiviridae) in association with betasatellites (Tolecusatellitidae) and alphasatellites (Alphasatellitidae), all being transmitted by insect vector whiteflies (Bemisia tabaci). The disease complex is distinct between the two continents. Since prehistoric times, cotton is being grown in almost all tropical and subtropical areas of the world and it is increasingly adapted to temperate climates. China, India, USA, Pakistan and Brazil are the major cotton producing countries in the world. The genus Gossypium consists of approximately 52 species, five of which are tetraploid, and all others are diploid. Only four species are cultivated which produce spinnable fiber for the textile industry. These include two allotetraploid species, G. barbadense (Pima/ Egyptian cotton) and G. hirsutum and two diploid species, G. arboreum and G. herbaceum. G. hirsutum (also known as upland cotton) accounts for more than 90% of the cotton production in the world. Diploid cotton species are native to Asia and Africa and they were largely grown prior to G. hirsutum species that was introduced from the New world (NW). The Old world (OW) native species have natural resistance to CLCuD.

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Fig. 1 Infected cotton (Gossypium hirsutum) plants showing typical symptoms of CLCuD. Symptoms include vein darkening, vein swelling, upward curling of the leaves and enations on the undersides of leaves (left). A severely infected plant showing a reduced leaf size, downward curling of leaf, leaf crumpling and severely stunted growth (right).

History of CLCuD CLCuD in Africa CLCuD was reported for the first time in 1912 from Nigeria, where it was sporadic and remained a minor problem. Its second outbreak occurred in the same country in 1924. Later in the year 1924, CLCuD was also reported from Sudan and in 1926 from Tanzania. In North-East Africa, (mostly Egypt and Sudan) species of cotton mostly cultivated is G. barbadense. CLCuD is endemic in these areas, although it is only sporadically a problem. An epidemic was reported in 1927–28 in Sudan which stimulated interest in this disease, and it was shown that whiteflies were responsible for its transmission. At that time extensive research established that the disease is also transmissible through graft and a virus-like agent was suspected as the causative agent although the actual causing agent was not identified. In those days the disease was controlled by imposing a period in which cotton was not grown and later they cultivated virus resistant G. barbadense varieties. CLCuD sporadically occurs in Africa but with no major economic losses.

CLCuD in Southern Asia Prior to the 1980s, CLCuD was found sporadically in the Indian subcontinent. During the early-1990s, in Pakistan, the cotton production started to suffer heavy losses due to an epidemic of CLCuD. The disease was first reported near the city of Multan and soon it spread to almost all major cotton growing areas of Pakistan and also into the western part of India. Most probably this epidemic of CLCuD was attributed to the large-scale cultivation of two cotton varieties viz. CIM70 and S-12 which were highyielding, but unfortunately highly susceptible to this local disease. G. arboreum, the native species of cotton is naturally immune to CLCuD, but this species does not produce cotton lint of high quality demanded by the processing industry. However, still farmers grow this species on a small scale. At the end of the 1990s, the cotton production reached again pre-epidemic levels, through the development and gradual adoption of tolerant and resistant cotton varieties. Unfortunately, this resistance did not last a long time, as it was broken in 2001, by the emergence of a resistance breaking combination of a virus strain of begomovirus and a betasatellite. This resulted in the 2nd epidemic of CLCuD in this region. Currently, there is no commercial cotton variety resistant to this disease complex. In 2015, begomoviruses associated with the first epidemic reappeared in cultivated cotton fields and the resistance breaking strain disappeared after 2015.

Etiology of CLCuD In both CLCuD affected regions [Africa (Sudan and Egypt) and Indian subcontinent] the etiology of the disease has been determined. In both regions, it was found that the disease is caused by monopartite begomoviruses (genus Begomovirus, family; Geminiviridae) associated with single-stranded DNA satellite molecules viz., betasatellites (Tolecusatellitidae) and alphasatellites (Alphasatellitidae). Betasatellites encode a single protein bC1 which is a pathogenicity determinant, whereas alphasatellites play no apparent role in the etiology of CLCuD. However, it has been found in all cases of CLCuD in the field.

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Elucidation of the Etiology of CLCuD In Africa, since 2002 CLCuD is caused by a single species of begomovirus viz. Cotton leaf curl Gezira virus (CLCGezV) in association with Cotton leaf curl Gezira betasatellite (CLCGezB) and Cotton leaf curl Gezira alphasatellite (CLCGezA). Although this disease complex remained permanently associated with CLCuD in Africa, recently and for the first time, in west and central Africa, a bipartite begomovirus, Cotton yellow mosaic virus (CYMV) was also reported in a non-cultivated cotton species i.e. Gossypium raimondii. CLCuD caused huge losses to cotton industries in both Pakistan and India in the early 1990s. This initiated a deep investigation into the etiology of the CLCuD. Initially, several species of begomoviruses were isolated and sequenced from cotton in both countries but their infectivity was not tested in cotton. So, the Koch’s postulate was not fulfilled at that time. Later, efforts were made to introduce one of these begomoviruses i.e., CLCMulV into cotton which showed atypical and mild symptoms. These results indicated that either another component was required for the disease or CLCMulV was not responsible for the CLCuD. Begomoviruses are typically divided into two groups i.e., monopartite and bipartite based on their genomic components. The bipartite begomovirus has two component DNA-A and DNA-B approximately equal size (2.7 kb) whereas monopartite begomoviruses have single genomic components equivalent to DNA-A of bipartite begomoviruses. Old world monopartite begomoviruses are often also associated with DNA satellites referred to as betasatellites and alphasatellites. During the first epidemic, it was troubling that CLCMulV alone was unable to infect cotton to develop the typical symptoms of CLCuD and the same situation was noticed with several other begomoviruses. When CLCuD-affected plants were further analyzed then a novel additional, subviral component was identified. Similar components had been identified in association with several monopartite begomoviruses. These components had been named as DNA-1 (now known as alphasatellite). However, in the case of CLCuD, it was found that alphasatellites did not play an essential role in the disease. Nevertheless, this finding promoted the reinvestigation of subviral DNA molecules, which might be associated with monopartite begomoviruses. In the mean-time, with a different study done with the monopartite Ageratum yellow vein virus (AYVV), another sub-genomic DNA component, named DNA-b (betasatellite), was identified. Shortly after this discovery a similar component DNA-b was also identified associated with CLCMulV. Inoculation of CLCMulV with the associated betasatellite induced all typical symptoms of CLCuD in cotton, hereby establishing the etiology of the disease and Koch’s postulate was fulfilled. Betasatellites encode a single protein referred to as bC1 on its complementary strand which acts as a pathogenicity determinant.

Components of the CLCuD Complex Begomoviruses Associated With CLCuD CLCuD in Africa A single OW monopartite begomovirus viz. Cotton leaf curl Gezira virus (CLCGezV), has been found in association with CLCuD in Africa. However, recently Cotton yellow mosaic virus (CYMV); a bipartite OW begomovirus has been isolated from non-cultivated cotton (Gossypium raimondii) for the first time in west and central Africa. CLCGezV is more closely related to begomoviruses originating from the African Mediterranean region infecting malvaceous plants, and distantly related to begomoviruses associated with the Asian CLCuD. Begomoviruses are geographically related, supporting the distinct evolutionary origin of begomoviruses causing CLCuD in Asia and Africa. In 2005, CLCGezV-EG (Egyptian strain) was isolated from cotton in Sindh, Pakistan, in a mixed infection, but its cognate betasatellite viz., CLCGezB was not found, instead it was trans-replicating two asian betasatellites; CLCMulB and Chili leaf curl betasatellite (ChLCB). Interestingly, until now, CLCGezV has not been reported from Punjab in Pakistan and from India.

CLCuD in the Indian subcontinent The first epidemic of CLCuD in the Indian subcontinent was associated with several distinct begomovirus species which are CLCMulV, Cotton leaf curl Alabad virus (CLCAlaV), Cotton leaf curl Kokhran virus (CLCKokV), Tomato leaf curl Bangalore virus (ToLCBanV) and Papaya leaf curl virus (PaLCV). They were associated with the disease either in single or multiple co-infections (Fig. 2). Some of these viruses induced CLCuD in cotton plants when they were co-inoculated with CLCMulB whereas, only ToLCBanV was able to cause tomato leaf curl disease in the absence of a betasatellite. However recently, it was shown to transreplicate multiple betasatellites which resulted in the increased pathogenicity in tomato and a model plant. CLCBanV was reported from G. barbadense in southern India but it was not affecting the north of India where CLCuD was epidemic. In addition, experiments to demonstrate that it could infect cotton and cause CLCuD were not successful. In the late 1990s scientists had developed some cotton resistant varieties which helped to restore the production to preepidemic levels. But, unfortunately in the year 2001, a resistance breaking begomovirus appeared. It was a recombinant of CLCMulV and CLCKokV and referred to as CLCKokV-Burewala strain (CLCKokV-Bur). It was first identified from Punjab in Pakistan then also spread to north-western India and it became the second epidemic of CLCuD in the region. This strain had a truncated TrAP (AC2) protein which was speculated to cause the resistance breaking. However, since that time, the existence of CLCKokV-Bur with an intact C2 gene has been shown. Therefore, the reason for this strain to break the resistance and cause the second epidemic has not been elucidated.

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In 2014, two bipartite begomoviruses have also been identified from cultivated cotton plants in Pakistan. The most important one is Tomato leaf curl New Delhi virus (ToLCNDV), which was frequently identified from cotton in both Sindh and Punjab provinces of Pakistan, whereas the other was Tomato leaf curl virus (ToLCV) reported from the same country. Moreover, the identification of Chickpea chlorotic dwarf virus (CpCDV), a Mastrevirus (Geminiviridae), and Okra enation leaf curl virus (OELCV), a typical begomovirus from infected cotton, has further complicated the situation. It has been shown experimentally that ToLCNDV enhances the accumulation of CLCMulB-Bur by synergistically interacting with CLCKokV-Bur in the infected plants. Moreover,

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CLCMulB has been shown to be trans-replicated by the DNA-A component of ToLCNDV, in the absence of its DNA-B component, and causing typical symptoms of CLCuD in cotton. Interestingly, in 2015, multiple begomoviruses which were associated with the first epidemic of CLCuD in 1990s were found in cultivated cotton in Vehari, Punjab, in Pakistan and in the same year CLCMulV was exclusively identified from Punjab, in India. In both studies CLCKokV-Bur was not identified in cultivated cotton. Identification of multiple begomoviruses with a single species of betasatellite in cotton while causing a single disease, suggests that these begomoviruses, also identified from many alternative hosts, most probably moved to cotton, where they could interact with CLCMulB to cause a unique cotton disease. Moreover, recently it was reported that non-cultivated cotton species from Multan, Punjab, in Pakistan (the region considered as hotspot for begomoviruses and satellites associated with CLCuD) were continuously infected by multiple begomoviruses, associated with multiple DNA satellites but with the unique identical betasatellite, CLCMulB. These plants are therefore reservoirs for a collection of begomoviruses and satellites. The presence of multiple begomoviruses makes it easy to overcome resistance against individual viruses and facilitates adaptation to other hosts. In addition, in the Indian subcontinent there are many other monopartite begomoviruses that are not found in cotton, possibly suggesting that these few species of begomoviruses recently adapted to cotton. Moreover, some begomoviruses that are not reported in the region, such as ToLCV, originating from Australia, and Ageratum yellow vein virus originating from southeast Asia, as well as Tomato yellow leaf curl virus (TYLCV), were shown capable experimentally to trans-replicate the CLCMulB (the unique betasatellite associated with CLCuD). The capability of CLCMulB to interact with a multitude of begomoviruses provides a huge flexibility for the cotton disease to occur and makes it a real threat to cotton production worldwide. It has been shown experimentally that CLCMulV is efficiently transmitted by the whitefly species Asia II 1 as compared to Middle East Asia minor 1 (MEAM1), and Asia II 1 is the dominant species in major cotton growing areas of Pakistan and India. Moreover, the Asia II 1 whitefly genome has been sequenced, and compared to that of MEAM1, revealing variations in genes involved in insecticide resistance and virus transmission. These findings may explain the rapid spread of CLCuD but also its current limitation to the Indian subcontinent. If Asia II 1 was to become dominant in other parts of the world, most probably it may allow the rapid transfer of CLCuD associated begomoviruses and satellites to become a serious global threat. In addition, the Faisalabad strain of CLCMulV was recently isolated from Hibiscus rosasinensis and from okra in the southern China, but it did not become epidemic in China, most probably due to the local whitefly species. With all this information, a third epidemic of CLCuD could be expected to occur soon, possibly with distinct begomoviruses, but with the same associated DNA satellite components (Fig. 3). A recent Pakistan report survey about whiteflies and begomoviruses from all major cotton growing areas (both Sindh and Punjab in Pakistan), indicates that the dominant whiteflies are of the species Asia II 1 and they all carry CLCMulV-Raj, a new strain of CLCMulV. The most recent isolates of CLCMulV from India and Pakistan indicate a novel clade in the phylogenetic tree of begomoviruses associated with CLCuD and this may be associated with a potential third epidemic of CLCuD. Similarly, the most recent strain of betasatellite viz., CLCMulBVeh has been identified from whiteflies as well as cotton, so it may also be associated with a possible third epidemic of the disease.

CLCuD-Associated Alphasatellite Components Alphasatellites (Alphasatellitidae) are the satellite components associated with disease complexes of monopartite begomovirus (Fig. 3). They are circular, single-stranded (ss)DNA molecules have about half the size (B1380 nt) of their helper viruses. With a single gene on virion-sense orientation which encodes a protein required for their replication (rolling-circle replication) initiation. The product of this gene is called replication associated protein (Rep) of the plant-infecting ssDNA viruses. In the cells of host plants, alphasatellites are capable of autonomous replication so for their replication they do not depend on their helper viruses. However, for movement within plants and transmission between plants, these molecules require the helper geminiviruses. Presumably, they are trans-encapsidated in the coat protein of the helper geminiviruses. Although alphasatellites are not apparently required to induce the disease symptoms in host plants it was however recently reported that Rep of alphasatellites may have a role in gene silencing mechanism in plants. Fig. 2 Neighbor-joining phylogenetic tree of begomoviruses was reconstructed using bootstrap method with 1000 replicates (percentage bootstrap value is shown on the right corner of each branch when 450%). The size of the triangles is roughly proportionate to the number of isolates in the cluster. The color code is for identifying viruses originally described on cotton in the Indian subcontinent (yellow), or for newly identified viruses on cotton but from other origin in the Indian subcontinent (green), light brown is for African viruses, light blue is for ACMV identified in Pakistan but originally described in Africa, pink is from cotton viruses from America and orange is for a mastrevirus identified in cotton in Pakistan. The triangles with a double line are those for the bipartite viruses versus 1 line for the monopartite viruses. Abbreviations used for different species and strains are as follows: CChSV (Cotton chlorotic spot virus), ChiLCV (Chili leaf curl virus), CLCAlaV-Ala (Cotton leaf curl Alabad virus-Alabad), CLCAlaV-Har (Cotton leaf curl Alabad virus-Haryana), CLCAlaV-Kar (Cotton leaf curl Alabad virus-Karnal), CLCAlaV-Lob (Cotton leaf curl Alabad virus-Lobatum), CLCAlaV-Mul (Cotton leaf curl Alabad virus-Multan), CLCBanV (Cotton leaf curl Bangalore virus), CLCBarV (Cotton leaf curl Barasat virus), CLCGezV-(Cotton leaf curl Gezira virus), CLCKokV-Bur (Cotton leaf curl Kokhran virus-Burewala), CLCKokV-Kok (Cotton leaf curl Kokhran virus-Kokhran), CLCKokV-Sha (Cotton leaf curl Kokhran virus-Shadadpur), CLCMirV (Cotton leaf curl Mirzapur virus), CLCMulV-Dar (Cotton leaf curl Multan virus-Darvinii), CLCMulV-Fai (Cotton leaf curl Multan virus-Faisalabad), CLCMulV-His (Cotton leaf curl Multan virus-Hisar), CLCMulV-PK(Cotton leaf curl Multan virus-Pakistan), CLCMulV-Raj (Cotton leaf curl Multan virus-Rajasthan), CLCrV (Cotton leaf crumple virus), CpCDV (Chickpea chlorotic dwarf virus), CYMV-DNA-A (Cotton yellow mosaic virus-DNA-A genome), OELCV (Okra enation leaf curl virus), PaLCV (Papaya leaf curl virus), SLCV (Squash leaf curl virus), ToLCBanV (Tomato leaf curl Bangalore virus), ToLCGujV-DNA-A (Tomato leaf curl Gujarat virus-DNA-A genome), ToLCNDV-DNA-A (Tomato leaf curl New Delhi virus-DNA-A genome).

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Fig. 3 Genomic organization of begomoviruses and associated alphasatellites and betasatellites. A; genome organization of a monopartite begomovirus and associated satellites in the first epidemic of CLCuD in Asia, B; genome organization of a monopartite begomovirus and associated satellites found in the second epidemic of CLCuD in Asia, C; genome organization of a monopartite begomovirus and associated satellites expected in a third epidemic (left), and genome organization of a bipartite begomovirus expected in a third epidemic (right), D; genome organization of a monopartite begomovirus and associated satellites found in Africa and E; genome organization of a bipartite begomovirus found in Africa. Abbreviations; CR; conserved region, Rep; Replication-associated protein, TrAP; Transcription associated protein, Ren; Replication Enhancer protein, CP; Coat Protein, MP; Movement protein, NSP; Nuclear Shuttle protein, bC1; Beta C1; SCR; Satellite Conserved Region. A-rich; Adenine rich region.

Surprisingly, the alphasatellite are more closely related to nanovirus genomic components that also encode a Rep protein. Nanoviruses (family Nanoviridae) are a group of plant-infecting, ssDNA viruses that are transmitted by aphids. Similarly, the master Rep (mRep) component which encodes the Rep required for replication of all nanovirid components. These satellite-like molecules and mRep components are however quite distinct from each other based on their genomic sequence and genome organization. The position of the promoter of these components is different from that of the mRep. It is believed that alphasatellites associated with begomoviruses have evolved directly from the satellite-like components of nanoviruses. The major difference between alphasatellites and satellite-like components of nanovirus is their genomic size. The average size of alphasatellites is B1380 bp, which is almost half the size of helper begomoviruses, whereas the genome of nanovirid components is typically of 1000–1100 bp. Most of this increased size is occupied by an adenine (A) rich region of about 200 bp. The function of this A-rich region it still not clear; it was observed that alphasatellites with deleted A-rich region were capable to replicate and move in plants in the presence of a helper begomovirus. There is an understated selection mechanism to maintain

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the size of alphasatellites, most probably due to efficient encapsidation by the coat protein of their helper virus. Since the first identification of Cotton leaf curl Multan alphasatellite (CLCMulA) associated with the cotton begomovirus, multiple alphasatellites have been reported from CLCuD infected cotton in the indian subcontinent. They include Cotton leaf curl Multan alphasatellite (CLCMulA), Gossypium darwinii symptomless alphasatellite (GDarSLA), Ageratum conyzoides symptomless alphasatellite (AConSLA), Cotton leaf curl Burewala alphasatellite (CLCBurA), Okra enation leaf curl alphasatellite (OELCA), Ageratum yellow vein India alphasatellite (AYVINA) and Tomato leaf curl alphasatellite (ToLCA). Interestingly, in North-East Africa, the only alphasatellite component reported with the CLCuD complex is Cotton leaf curl Gezira alphasatellite (CLCGezA).

CLCuD-Associated Betasatellite Betasatellites are ssDNA satellites (genus Betasatellite, family Tolecusatellitidae) which are associated with many monopartite begomoviruses. Presumably, betasatellites are trans-encapsidated by the coat protein of helper viruses to allow movement in plants, as well as transmission by whiteflies. Betasatellites also depend on the helper geminivirus for their replication. In infectivity assays betasatellites were found responsible for the high titer of the helper virus, to levels found in their original host in the field. This showed that betasatellites are involved in either the replication of their helper virus, their systemic movement in the host, and/or suppressing the gene silencing host defense mechanism against viruses. The size of the betasatellites is about half (1350 bp) of the size of their helper begomovirus. Interestingly, they share no sequence homology with their helper virus except the stem–loop structure which contains the conserved nonanucleotide (TAATATTAC). This stem-loop structure is present in a satellite conserved region (SCR) which is about 100 nt and highly conserved among betasatellites. Betasatellites encode a single gene, the product of which is known as bC1 protein (B118 aa) which has a critical role in pathogenicity and symptoms determination as it alters several cellular processes of the host like ubiquitination, autophagy and suppression of gene silencing (post-transcriptional gene silencing (PTGS) or transcriptional gene silencing (TGS)). After the first discovery of the Ageratum yellow vein betasatellite (AYVB), they have been found from a diverse range of hosts. More than 1800 complete betasatellite genomes were sequenced and their comparison shows that they are highly diverse. Surprisingly, only a single species of betasatellite, Cotton leaf curl Multan betasatellite (CLCMulB) is associated with CLCuD. Both experimentally and in the field, this species was found to be trans-replicated by several distinct begomoviruses. When bC1 of CLCMulB is expressed in tobacco plants, either stably transformed or transient expression, it induces virus-like symptoms, which demonstrates that in the CLCuD complex, the bC1 is the most important pathogenicity and symptom determinant. The second epidemic of CLCuD, which was caused by a recombinant begomovirus (from two different species), was also associated with a recombinant CLCMulB, in which CLCMulBMul (Multan strain) recombined with Tomato leaf curl betasatellite (ToLCB). In Sindh province of Pakistan, another novel strain of CLCMulB viz. CLCMulBSha (Shadadpur strain) was also found to be associated with CLCuD whereas in Punjab only CLCMulBBur was found. Most recently, a novel strain CLCMulBVeh (Vehari strain) was identified from Vehari, Punjab, in Pakistan and from India. Identification of different strains of CLCMulB indicates the co-evolution of CLCMulB with helper begomoviruses. In Africa, Cotton leaf curl Gezira betasatellite (CLCGezB) is the only betasatellite found so far in association with CLCuD in Sudan and it is quite distinct from CLCMulB in the indian subcontinent.

Control of CLCuD Strategies to Control the CLCuD Complex The CLCuD is a serious threat to cotton cultivation in the countries where whiteflies are a major pest. There are mainly two types of strategies i.e. a short term strategy, involving management practices and a long term strategy involving the development of resistant cotton varieties either by conventional or non-conventional methods.

Short Term Strategies After the spread of CLCuD on a massive scale during the first epidemy, many short-term strategies were adopted to minimize the losses by reducing the whitefly population in the field. For instance, treatment of seeds to avoid early establishment of vector populations, spraying insecticide on cotton crop to control whitefly populations, eradication of weeds (alternative hosts of viruses), providing balanced dose of fertilizers and biological agents…. Application of selective insecticide is most important in controlling the whitefly population. Recently, the whole genome sequence of Asia II 1 showed genetic variance in insecticide resistance related genes as compared to MEAM1. Most probably, this may be the impact of misuse of insecticides to control whitefly population. Moreover, it is advisable to spray insecticide on cotton crop 70–90 DAS (days after sowing) to control the whitefly population as seed treatment protects from whitefly infestation up to 75 DAS in some reports. After this period, cotton plants are grown enough that they can escape the disease, and losses are reduced. As CLCuD associated viruses are not seed-borne they depend upon the alternative hosts to survive the off season. Alternative hosts are many other plants like okra, tomato, tobacco, ageratum, hibiscus and many weeds. Avoiding the cultivation of alternative host in off season and proper eradications of weeds may reduce the disease incidence.

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Further damage due to this disease can be reduced by applying a balanced dose of nutrients especially using potassium to boost the resistance against the disease whereas excessive use of nitrogen reduces the resistance. So, it is advised to use the balanced ratio of potassium and nitrogen which helps to reduce the disease severity.

Long Term Strategies Genetics of host plant resistance to CLCuD For durable resistance strategy to control the disease, it is most important to breed virus resistant varieties with enough genetic diversity. Before developing disease resistant varieties of cotton, the key components is designing a breeding plan for the genetic basis of disease resistance and its inheritance. Unfortunately, very little is known about the molecular basis of the disease resistance in cotton. G. arboreum, native species of Asia is naturally immune to the Asian CLCuD complex, but it is grown on a small scale because it produces a poor quality fiber. The transcriptomics study of G. arboreum under virus stress revealed multiple resistance mechanisms against CLCuD. The exotic species G. hirsutum produces good quality and quantity of fiber and is commercially cultivated on large scale. Unfortunately, this species showed limited resistance/tolerance to CLCuD and only two varieties, LRA5166 and CP15/2, have been used to develop resistance in the late 1990s. Losses due to CLCuD were successfully alleviated by cultivating these varieties and cotton production was regained to the level prior to the epidemic. Unfortunately, this resistance was broken by the evolution of novel strain of begomovirus viz., CLCKokV-Bur. Currently, all commercial varieties are highly susceptible to CLCuD. Recently, transcriptomic studies in G. hirsutum under whitefly transmitted virus stress provided some insights into disease susceptibility in cotton and predicted some important genes to be involved in the compatible interaction of CLCuD associated disease complex and cotton. These findings will have to be validated to pinpoint the genes that make G. hirsutum susceptible and G. arboretum resistant. Although some efforts were made in introgression virus resistance from G. arboreum into G. hirsutum, they were not successful in developing a resistant variety suitable for agriculture. it was also tried to transfer the desired traits from G. hirsutum to G. arboreum (after doubling its chromosomes) but unfortunately, it also remained unsuccessful in the development of commercial varieties. Finally, about 5000 accessions of G. hirsutum from USA and some introgression lines were screened in Pakistan against CLCuD. This work identified the “Mac-07” accession, and 95 other accessions, that remained asymptomatic under field conditions, hoping that a good source of resistance could be used in the near future.

Engineered Resistance to the CLCuD Complex Because of the difficulty to identify and transfer useful genes of resistance to CLCuD from G. hirsutum germplasm, multiples strategies have been deployed using genetic engineering approach to develop cotton plants resistant to this viral disease and its insect vector whitefly. Generally, these strategies are of two types; firstly the “pathogen derived resistance” in which full length viral genes or different small conserved portion of the virus genome are used and secondly the ‘non-pathogen derived resistance’ in which genes from distantly related genetic source are used.

Pathogen Derived Resistance (PDR) Introducing the genomic part (gene or its part) of virus that is conserved among different viruses into the host plant is the most desirable strategy for broad spectrum resistance to control disease. Following strategies have been adopted for the development of resistance in which genomic information of pathogen is used.

Antisense RNA technology In this technology antisense RNA molecule suppresses the complementary target mRNA. When antisense RNA against Rep gene of the virus was introduced in cotton, it suppressed the replication of the virus, similarly in another study when the coat protein gene (AV1) was targeted by antisense RNA in transgenic cotton, it arrested the movement, replication and encapsidation of the target virus. Transgenic cotton was also developed using antisense AV2 (350vbp) gene and transgenic plants were resistant to CLCuD. Similarly, two truncated forms of the Rep gene were introduced in G. hirsutum, which inhibited the replication of both viral and satellite genomes. Recently antisense ßC1 gene was also introduced in cotton and transgenic plants were symptomless under virus stress. However, none of these transgenic cotton lines were successful under field conditions.

RNAi technology The principle of the RNAi technology utilizes the transcriptional gene silencing (TGS) and post-transcriptional gene silencing (PTGS) and has been deployed to develop resistance against different viruses, such as the African cassava mosaic virus (ACMV), the Mung bean yellow mosaic virus (MYMV) and many others. A 21 nt of V2 of CLCKokV-Bur strain was used to develop the artificial microRNA (amiRNA) and construct was transformed in Nicotiana benthamiana plants and transgenic plants showed resistance to CLCKokV-Bur. Recently, RNAi based construct against the V2 gene of CLCKoKV-BuR was transformed in two cotton species viz. MNH-786 and VH-289 and transgenic cotton plants, when challenged to whitefly under control conditions, showed a low titer of

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the virus for transgenic plants as compared to non-transgenic. It was concluded that these transgenic plants could be used in the fields to control the CLCuD but have so far not proven successful in the field.

Non-Pathogen Derived Resistance In this strategy, genes from host or non-host are used to develop the resistance against the disease. For instance, genes which confer DNA binding proteins, antiviral antibodies, coat binding proteins etc. have been introduced in plants to develop resistance to CLCuD. Recently, Tma12 a protein identified from fern to confer resistance to whitefly, was transformed in cotton, and one of the transgenic lines showed increased resistance (499%) to whitefly. Moreover, this protein was non-toxic in rats which shows that this is safe to use in other crops.

Genome Editing Strategy: CRISPR/Cas9 Approach to Control CLCuD CRISPR/Cas9 system has been exploited in many organisms for genome editing by transforming the Cas9 protein. One of the desirable properties of this system is that it can targets multiple sites, which can be utilized to control multiple viruses in plants. Resistance to different geminiviruses has already been achieved through this system. For example, genome of Bean yellow dwarf virus (BeYDV) was cut down when CRISPR–Cas9 system was applied in bean against this virus and resulted in reduced disease symptoms. Similarly, this system was used against TYLCV and Beet curly top virus (BCTV) and resulted in the reduced disease symptoms. In cotton this system is in progress to target CLCKokV-Bur and associated betasatellite genes and conserved regions. However, its success is yet to be demonstrated. Recently, it was reported that expressing Cas9 in cassava was unable to protect it from geminiviruses infections where viruses multiply faster than edited by Cas9 and this also helps in rapid evolution of viruses. This makes an alarming sign for future results of CRISPR/Cas9 use in cotton against CLCuD associated disease complex (as multiple sgRNAs are being used). In this scenario it is most important to explore the actual molecular mechanisms involved in compatible interactions of virus-host vector and identify some susceptible genes in cotton which can be knocked out by CRISPR/ Cas9 system, or resistance genes for introgression in susceptible commercial varieties.

See also: Alphasatellites (Alphasatellitidae). Bean Golden Mosaic Virus and Bean Golden Yellow Mosaic Virus (Geminiviridae). Beet Curly Top Virus (Geminiviridae). Betasatellites and Deltasatelliles (Tolecusatellitidae). Cassava Mosaic Viruses (Geminiviridae). Emerging Geminiviruses (Geminiviridae). Geminiviruses (Geminiviridae). Maize Streak Virus (Geminiviridae). Tomato Leaf Curl New Delhi Virus (Geminiviridae). Tomato Yellow Leaf Curl Viruses (Geminiviridae)

Further Reading Amin, I., Mansoor, S., Amrao, L., et al., 2006. Mobilisation into cotton and spread of a recombinant cotton leaf curl disease satellite. Archives of Virology 151, 2055–2065. Asad, S., Haris, W.A.A., Bashir, A., et al., 2003. Transgenic tobacco expressing geminiviral RNAs are resistant to the serious viral pathogen causing cotton leaf curl disease. Archives of Virology 148, 2341–2352. Briddon, R.W., 2003. Cotton leaf curl disease, a multicomponent begomovirus complex. Molecular Plant Pathology 4, 427–434. Briddon, R.W., Markham, P.G., 2000. Cotton leaf curl virus disease. Virus Research 71, 151–159. Briddon, R.W., Bull, S.E., Amin, I., et al., 2004. Diversity of DNA-1: A satellite-like molecule associated with monopartite begomovirus-DNA b complexes. Virology 324, 462–474. Datta, S., Budhauliya, R., Das, B., et al., 2017. Rebound of cotton leaf curl Multan virus and its exclusive detection in cotton leaf curl disease outbreak, Punjab (India), 2015. Scientific Reports 7 (1), 17361. Fauquet, C.M., Stanley, J., 2005. Revising the way we conceive and name viruses below the species level: A review of geminivirus taxonomy calls for new standardized isolate descriptors. Archives of Virology 150, 2151–2179. Leke, W.N., Khatabi, B., Mignouna, D.B., Brown, J.K., Fondong, V.N., 2016. Complete genome sequence of a new bipartite begomovirus infecting cotton in the Republic of Benin in West Africa. Archives of virology 161 (8), 2329–2333. Sattar, M.N., Iqbal, Z., Tahir, M.N., Ullah, S., 2017. The prediction of a new CLCuD epidemic in the old world. Frontiers in microbiology 8, 631. Sattar, M.N., Kvarnheden, A., Saeed, M., Briddon, R.W., 2013. Cotton leaf curl disease – An emerging threat to cotton production worldwide. Journal of General Virology 94 (4), 695–710. Sohrab, S.S., Kamal, M.A., Ilah, A., et al., 2016. Development of cotton leaf curl virus resistant transgenic cotton using antisense ßC1 gene. Saudi Journal of Biological Sciences 23 (3), 358–362. Zaidi, S.S.E.A., Shafiq, M., Amin, I., et al., 2016. Frequent occurrence of tomato leaf curl New Delhi virus in cotton leaf curl disease affected cotton in Pakistan. PLoS One 11 (5), e0155520. Zubair, M., Zaidi, S.S.E.A., Shakir, S., et al., 2017. Multiple begomoviruses found associated with cotton leaf curl disease in Pakistan in early 1990 are back in cultivated cotton. Scientific Reports 7 (1), 680. Zubair, M., Zaidi, S., Shakir, S., Amin, I., Mansoor, S., 2017. An insight into cotton leaf curl Multan betasatellite, the most important component of cotton leaf curl disease complex. Viruses 9 (10), 280.

Cowpea Mosaic Virus (Secoviridae) George P Lomonossoff, John Innes Centre, Norwich, United Kingdom r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

MP Movement protein nt Nucleotide(s) ORF Open reading frame RdRp RNA-dependent RNA polymerase UTR Untranslated region VLPs Virus-like particles VPg Viral protein genome-linked

aa Amino acid(s) Co-Pro Protease cofactor CP Coat protein or capsid protein ER Endoplasmic reticulum kb Kilobase kDa Kilo Dalton

Glossary Affimers Small proteins that mimic antibodies. Nanobiotechnology Refers to the intersection of nanotechnology and biology. Given that the subject is one that has only emerged very recently, bionanotechnology and nanobiotechnology serve as blanket terms for various related technologies. Nanotechnology Also known as “nanotech” is manipulation of matter on an atomic, molecular, and supramolecular scale.

Viral protein genome-linked A protein covalently attached to the 50 end of positive strand viral RNA and acting as a primer during RNA synthesis. Virus replication complexes Intracellular complexes formed by association with intracellular membranes that are sites for stages in the replication cycle of a virus. b-barrel Beta-sheet composed of tandem repeats forming a closed structure.

Introduction Since its first isolation in 1959, Cowpea mosaic virus (CPMV) has served as a model icosahedral plant virus due to the high yield, ease of isolation and stability of its particles. As a consequence, it has been at the forefront of both genetic and structural studies of plant viruses and has been widely deployed in bionanotechnology. Thus, CPMV can be considered alongside viruses such as Tobacco mosaic virus (TMV) and Brome mosaic virus (BMV) as one of the “classic” plant viruses.

Taxonomy and Classification CPMV is the type species of the genus Comovirus (which includes 14 additional members). The genus Comovirus is classified, together with the genera Fabavirus and Nepovirus, within the sub-family Comovirinae of the family Secoviridae. Within the subfamily the greatest affinity is between the genera Comovirus and Fabavirus, both of which are insect-, as opposed to nematode-, transmitted. On a wider scale, consideration of genome structure and organisation, translational strategy and aa homologies between the virusencoded proteins has led to the family Secoviridae being one of the 6 families within the order Picornavirales. Historically, the family Secoviridae had been grouped within the same “superfamily” as the family Potyviridae but the two families are now separately classified.

Biology CPMV was first isolated from infected cowpeas (Vigna unguiculata) in Nigeria in 1959. Subsequently it has been found to occur in Nigeria, Kenya, Tanzania, Japan, Surinam and Cuba. It causes typical mosaic symptoms on infected plants (Fig. 1). While its natural host is cowpea, it can infect other legumes and Nicotiana benthamiana has proven to be an extremely valuable experimental host. In nature, transmission is usually by leaf-feeding beetles, especially by members of the Chrysomelidae. However, CPMV has been shown also to be transmitted by thrips and grasshoppers. The beetle vectors can acquire the virus by feeding for as little as one minute and can retain and transmit the virus for a period of days or weeks. The virus does not, however, multiply in the insect vector. Experimentally, CPMV is mechanically transmissible. In Nigeria, infection of cowpeas with CPMV causes a great reduction in leaf area, flower production and yield. Infected plant cells show a number of characteristic cytological changes. These include the appearance of viral particles, a proliferation of cell membranes and vesicles in the cytoplasm, and a variety of modifications to plasmodesmata.

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Fig. 1 CPMV symptoms and particles (A) Mosaic symptoms on the trifoliate leaves of a CPMV-infected Vigna unguiculata plant (right) compared to an uninfected plant (left). (B) Transmission electron micrographs of purified particles of CPMV at two different magnifications. The particles were negatively stained with 2% (w/v) uranyl acetate.

Physical Properties of Viral Particles CPMV particles are icosahedral, non-enveloped with an average diameter of 28 nm (Fig. 1). Up to 2 g of particles can be obtained per kg of fresh infected cowpea tissue and they can be readily purified by polyethylene glycol precipitation and differential centrifugation. The particles are very stable with a thermal inactivation point in plant sap of 65–751C and a longevity in sap of 3–5 days at room temperature. Once purified, the particles can be stored for prolonged periods at 41C. The ease with which virus particles can be propagated, purified and stored has undoubtedly contributed to the early popularity of CPMV as an object of study. Preparations of CPMV particles can be separated into three components on density gradients that are designated top (T), middle (M) and bottom (B), with sedimentation coefficients of 58S, 95S and 115S, respectively. The three components have identical protein compositions, containing 60 copies each of a large (L) and small (S) coat protein (CP), with sizes 42 kDa and 24 kDa, respectively, as calculated from the nucleotide sequence. The discovery, in 1971, that CPMV particles contained equimolar amounts of two different polypeptides suggested that the capsids had an architecture more similar to the animal picornaviruses than to other plant viruses which has been previously examined. This provided an early clue as to the common origins of plant and animal viruses. The difference in the sedimentation behaviour of the three centrifugal components of CPMV lies in their RNA contents. Top components are devoid of RNA, while middle and bottom components each contain single molecules of positive-strand RNA of 3.5 and 6.0 kb respectively. The two RNA molecules were originally termed middle (M) and bottom (B) component RNA after the component from which they were isolated. However, more recently they have been referred to as RNA2 and RNA1, respectively. The three-component nature of CPMV preparations is summarized in Fig. 2. The determination of the component structure of the virus, and particularly the relationship between this and infectivity, was an important step in establishing the principle that plant viruses frequently have divided genomes, the individual components of which are separately encapsidated. CPMV preparations are not only centrifugally heterogeneous but can also be separated in two forms, fast and slow, electrophoretically. Both electrophoretic forms contain all three centrifugal components. The proportion of the two electrophoretic forms in a given virus preparation varies both with the time after infection at which the virus was isolated and the age of the preparation itself. Conversion of one form to the other is caused by loss, through proteolysis, of 24 aa from the C terminus of the S protein.

Viral Structure X-ray crystallographic and cryo-electron microscopy (cryo-EM) studies on CPMV, using either a natural mixture or individual components, have provided a detailed picture of the arrangement of the two viral CPs in the three-dimensional structure of the particle. Overall, the virions have icosahedral symmetry, with 12 axes of fivefold and 20 axes of threefold symmetry and resemble a classic T¼ 3 particle. However, since there are two different types of CP subunit (L and S), their arrangement is usually referred two as pseudo T¼3 or P¼3. The two CPs taken together consist of three distinct b-barrel domains, two being derived from the L and one from the S protein. Thus, in common with the T¼ 3 viruses, each CPMV particle is made up of 180 b-barrel structures. The S protein, with its single domain, is found at the fivefold symmetry axes and therefore occupies a position analogous to that of the A type subunits in T¼3 particles. The N- and C-terminal domains of the L protein occur at the threefold axes and occupy the positions equivalent to those of the C and B type subunits of a T¼ 3 particle respectively (Fig. 2). This detailed analysis confirmed the earlier suggestion that CPMV particles are structurally homologous to those of picornaviruses, with the N- and C-terminal domains of the L protein being equivalent to viral protein VP2 and viral protein VP3, respectively, and the S protein being equivalent to viral protein VP1; there is no equivalent of VP4. While no RNA could be seen in the crystallographic structures of either the M or B components of CPMV, cryo-EM analysis revealed that the encapsidated RNA forms concentric shells, with the strongest density being found directly beneath the twofold symmetry axes of the capsid. In common with other members of the order Picornavirales, the specific incorporation of the genomic RNAs into particles appears to be based on their ability to be replicated.

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Fig. 2 Molecular structure of CPMV (a) Organisation of the two single-stranded RNA molecules that comprise the genome. Both RNAs have a VPg (red circle) linked to their 50 ends and both are polyadenylated. Excluding the poly(A) tail, RNA1 is 5889 nt in length and contains a single ORF that is processed by the 24K protease (Pro) to give the viral proteins required for RNA replication. RNA2 is 3481 nt in length and encodes the structural CPs, Large (L) and Small (S), as well as the protein required for the movement of CPMV virions from cell-to-cell. (b) Gradient separation of a natural mixture permits separation into three components. Empty CPMV particles sediment at the top (CPMV-T), RNA2-containing particles sediment in the middle (CPMV-M) and RNA1-containing particles sediment at the bottom of the gradient (CPMV-B). (c) An asymmetric unit of synthetic CPMV empty virus-like particles (eVLP) as revealed by cryo-electron microscopy. Both domains of the CP-L subunit are shown in green, while single domain of the CP-S is shown in blue. The C-terminal extension of the CP-S, visible in eVLPs, but not in particles from infectious virus particles, is coloured pink. (d) The icosahedral organisation of CPMV. Each of the 60 asymmetric units comprises one copy of the L subunit and the S subunit (coloured as in c). A view down the two-fold axis is shown. Reproduced from Hesketh, E.L., Meshcheriakova, Y., Dent, K.C., et al. 2015. Mechanisms of assembly and genome packaging in an RNA virus revealed by high-resolution cryo-EM. Nature Communications 6, 10113.

As well as the natural components, the structure of synthetic empty virus-like particles (eVLPs), that lack any detectable RNA, has been determined by both crystallography and cryo-EM. These particles were shown to be essentially identical to the natural components; however, it was possible to visualize some of the 24 aa that are normally cleaved from the C-terminus of the S protein. These structural studies, together with mutagenesis of the 24 aa segment, has enabled a model for the assembly of CPMV particles to be generated in which the 24 aa promote pentamer formation followed by the formation of RNA binding sites at the two-fold symmetry axes (Fig. 3). The presence of the C-terminal portion of the 24 aa segment on the particle surface is also important in permitting efficient systemic movement of the virus.

Genome Structure Both M and B (but not T) components of a virus preparation are essential for infection of whole plants. As CPMV is a positivestrand RNA virus, a mixture of the genomic RNAs within the particles can also be used to initiate an infection. However, RNA1 is capable of independent replication in individual plant cells but this leads to the establishment of gene silencing, rather than a productive infection, in the absence of RNA2. Both genomic RNAs have a small basic protein (VPg) covalently linked to their 50 termini and both are polyadenylated at their 30 ends. The elucidation of the overall structure of the RNA segments once more underscored the similarity between CPMV and picornaviruses. However, unlike picornaviruses, the VPg is linked to the viral RNA via the b-hydroxyl group of its N-terminal serine residue rather than via a tyrosine. The VPg is not required for the viral RNAs to be infectious. The complete nucleotide sequences of both genomic RNAs were reported in 1983, making CPMV one of the first RNA plant viruses to be completely sequenced. The length of the RNAs are 5889 and 3481 nt, for RNA1 and RNA2, respectively, excluding the poly(A) tails. The two genomic RNAs have no sequence homology apart from at the 50 and 30 termini. Full-length infectious cDNA clones of both RNAs of CPMV have been constructed, allowing the genome to be manipulated.

Expression of the Viral Genome Both genomic RNAs contain a single long open reading frame (ORF) which occupies over 80% of the length of the RNA. A combination of in vitro translation and protoplast studies has unravelled the basic mechanism of gene expression of the virus. Both RNAs are expressed through the synthesis and subsequent cleavage of large precursor polyproteins (Fig. 2). On RNA1, initiation of

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Fig. 3 A model for CPMV Assembly. The L (green) and S (blue) subunits associate to form a CP penton. The C-terminal extension of the S protein (pink circle) appears to act as ‘molecular glue’, stabilizing the formation of the penton due to extensive interactions with the neighbouring S subunit. In the absence of RNA (RNA-free assembly shown in the top half of the diagram), C-terminal cleavage is slow, and eVLPs are produced. Once an extensive protein: protein interaction network is formed, the eVLP structure is stable and no longer depends on the presence of the C-terminal extension. In the presence of genomic RNA, assembly follows a different path (bottom half of the diagram). Initially, pentons are formed in the same way as in the eVLP, using the C-terminal extension for stability. Two pentons interact to form a twofold axis, and thus the RNA-binding site appears to exist at that position. The model shows genomic RNA (represented as an orange rectangle) initially binding to the two pentons at this twofold axis, the first step in genome encapsidation, followed by the stepwise addition of pentons, to form RNA-containing capsids. Reproduced from Hesketh, E.L., Meshcheriakova, Y., Dent, K.C., et al. 2015. Mechanisms of assembly and genome packaging in an RNA virus revealed by high-resolution cryo-EM. Nature Communications 6, 10113.

translation occurs at the first AUG encountered on the sequence (at position 207) and results in the synthesis of a protein of approximately 200 kDa (the 200K protein). This initial product undergoes rapid co-translational autoproteolysis to give proteins with apparent sizes of 32 kDa (Protease co-factor; ProC) and 170 kDa (170K) proteins. The 170K protein undergoes further cleavages to give the range of virus-specific proteins shown in Fig. 2, specifically the 58K Helicase, VPg, the 24K protease and the 87K polymerase. Initiation of translation of RNA2 occurs at two different positions on the RNA and results in the synthesis of two carboxy coterminal proteins, the 105K and 95K proteins. This double initiation phenomenon occurs through “leaky scanning”. Synthesis of the 105K protein is initiated from an AUG at position 161, while initiation from an in-frame AUG at position 512 directs the synthesis of the 95K protein; RNA-2 also has an additional AUG (position 115) upstream of both these initiation sites but in a different reading frame. Both RNA2-encoded primary translation products are cleaved by the RNA1-encoded 24K protease to give either the 58K or the 48K protein (depending on whether it is the 105K or 95 K protein being processed) and the precursor two viral CPs, VP60, which is then subsequently processed to give the mature L and S CPs. Processing of the RNA2-encoded polyproteins, at least at the site between the 48K and L CP, has been shown to require the presence of the 32K ProC protein as well as the 24K protease.

Functions of the Viral Proteins Functions have been ascribed to most of the regions of the polyproteins encoded by both RNA1 and RNA2 of CPMV. In most cases, however, it is not certain at what stage(s) in the cleavage pathway they manifest their activity. In the case of RNA1, the 32K ProC protein, which is rapidly cleaved from the N-terminus of the 200K primary translation product, is a cofactor which modulates the activity of the virus-encoded 24K protease. As described above, the presence of ProC is required for the cleavage of the RNA2-encoded 105K and 95K proteins but is not essential for the cleavage of the RNA1-encoded 170K protein. It does, however, seem to play a role in determining the rate at which cleavage of the 170K protein occurs. When mutant RNA1 molecules carrying deletions in the region encoding ProC are translated in vitro, the rate of processing of the 170K protein is greatly increased, indicating that the 32K protein acts as an inhibitor of processing. This inhibition may be achieved through the interaction of the 32K with the 58K Helicase domain of the 170K protein. The mechanism by which the 32K protein enables the 24K proteinase to cleave in trans is unclear. It could be due to a direct association between the 32K protein and the 24K proteinase (or an intermediate containing the proteinase domain) or it could be an indirect result of the 32K protein modulating the cleavage of the 170K protein, thereby allowing a 24K-containing intermediate with in trans processing activity to accumulate. The RNA1-encoded 58K Helicase protein is associated with cell membranes and contains a nucleotide-binding motif. A protein containing the aa sequence of the Helicase protein linked to VPg, is involved in rearrangements in the endoplasmic reticulum of CPMV-infected cells and acts in concert with ProC. The 24K protease carries out all the cleavages on both the RNA1- and RNA2encoded polyproteins. Its proteolytic activity has been shown to be expressed in processing intermediates that contain its sequence. Indeed, it is not known whether the free form of the protein has any biological significance. Although the proteinase contains a

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Fig. 4 Diagrammatic representation of the T-DNA region of the transient expression vector, pEAQ-HT showing the polylinker in detail. This polylinker allows the expression of the target protein with or without His-tags. The black boxes represent the T-DNA borders, the green arrows represent promoter sequences and the red arrows represent terminator sequences. The locations of the CPMV RNA2-derived 50 and 30 UTRs, between which the target sequence is inserted, is indicated. Reproduced from Sainsbury, F., Thuenemann, E.C., Lomonossoff, G.P., 2009. pEAQ: Versatile expression vectors for easy and quick transient expression of heterologous proteins in plants. Plant Biotechnology Journal 7, 682–693.

cysteine at its active site, it is structurally related to serine proteases, such as trypsin, rather than cellular thiol proteases, such as papain, a feature similar to the 3C proteinases of members of the family Picornaviridae. All CPMV cleavage sites have glutamine (Q) residue at the  1 position. The 87K Polymerase contains all the sequences associated with it being the virus-encoded RNA-dependent RNA polymerase (RdRp), including the GDD sequence motif found in all such enzymes. It also has aa sequence homology to the 3D polymerases encoded by the picornaviruses. Consistent with this identification, mutation of the GDD motif to GAD or AAA reduces or abolishes replication, respectively. When replication complexes capable of elongating nascent RNA chains were isolated from CPMV-infected cowpea plants they were found to contain the 110K protein (Fig. 3), consisting of the sequence of 87K polymerase protein linked to the 24K proteinase, rather than the free 87K protein, and this 110K protein is often referred to as the viral “replicase”. In the case of RNA2, the 48K protein, derived from processing of the 95K protein, is involved in potentiating the spread of the virus from cell to cell. This protein is found in tubular structures that are formed in the plasmodesmata of infected cells. Tubules extending into the culture medium can also be seen in protoplasts either infected with CPMV or transiently expressing the 48K protein. Virus particles can be seen within these tubules when protoplasts are infected with CPMV but not when only 48K protein is expressed. At present, no definite role has been assigned to the 58K protein which is produced by processing of the 105K protein. Elimination of AUG161, preventing production of the 105K protein, results in a substantial reduction of RNA2 replication. The mature viral CPs, L and S, are produced by cleavage of VP60 by the 24K protease. As well as protecting the genomic RNAs, capsid formation is essential for the virus to be able to spread from cell to cell through modified plasmodesmata and long-distance movement also requires capsid formation. An additional function in suppressing gene silencing is also provided by the C-terminal region of the S protein.

Replication CPMV replicates to high level in infected cells. Replication is believed to involve the initial transcription of the incoming positivesense RNA into minus-strands followed by initiation and synthesis of new plus-strands from the recently formed minus-strands. It has been shown that the 50 ends of both the plus- and minus-strands are covalently linked to the VPg, suggesting that this protein has an essential role in the initiation of RNA synthesis. There also appears to be a tight linkage between the translation of the viral RNAs and their replication and between replication and the encapsidation of the viral RNAs. Replication of the viral RNAs has been shown to occur in the membranous cytopathological structures which are formed in the cytoplasm of cells during infection through the action of the RNA1-encoded 32K and 60K proteins. Both CPMV-specific double-stranded replicative form (RF) RNA and an enzyme activity capable of completing nascent RNA strands can be isolated

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from such structures. Purified preparations of the enzyme activity contain the RNA1-encoded 110K replicase and two hostencoded proteins of 68 and 57 kDa. However, at present no enzymatic activity capable of initiating RNA synthesis in vitro has been described.

Use in Biotechnology CPMV particles have been extensively used in bio and nanotechology. In the first instance, antigenic peptides (epitopes) were genetically fused to exposed loops on the surface of the viral capsids using sites within both the L and S CPs. The resulting chimeric virus particles could be propagated in plants and the modified virions purified, provided the inserted peptide was not too long or excessively positively-charged. When injected into experimental animals, the modified particles could elicit the production of antibodies against the inserted epitope and in several instances were shown to confer protective immunity against the pathogen from which the epitope was derived. In addition to genetic modification, it is possible to chemically attached compounds to the surface of the particles via addressable aa such as lysine, glutamic and aspartic acid. Chemical attachment has the advantage that it allows moieties other than aa to be displayed on the surface of particles. It has also proved possible to combine both genetic and chemical methods to enable the mineralisation of virus particles. The development of a system for the production of eVLPs has also enabled CPMV particles to be loaded with foreign materials, such as drugs and heavy metals. The eVLPs have also proven useful for the selection of affimers which can subsequently be used to diagnose CPMV infections. CPMV RNA2 has been extensively used as a gene vector for the expression of heterologous proteins in plants via transient expression. Initially, such applications relied on the ability of the modified RNA to be replicated by RNA1 to achieve high level expression. However, the deployment of suppressors of gene silencing and the removal on inhibitory AUG codons, coupled with efficient agro-infection technology, has led to the development of a series of RNA2-based expression vectors (the pEAQ series) that do not rely on replication. In this system, the gene to be expressed is positioned between a modified version of the RNA2 50 UTR, lacking the AUG codons upstream of AUG512, and the RNA2 30 UTR to create a synthetic mRNA cassette (Fig. 4). Transcription of the cassette is driven by the Cauliflower mosaic virus 35S promoter and terminated by the nos terminator. The whole transcription unit is placed between the left and right T-DNA borders within a binary plasmid together with a transcription unit for the P19 suppressor of gene silencing. Agro-infiltration is used to deliver the plasmids to plant tissue and expression is usually detected within one week. The pEAQ expression system has been used for the expression of a wide variety of different proteins, including those capable of forming VLPs and enzymes for bio-transformations. The elimination of the need for replication means that it is possible to co-express multiple proteins within the same cell, thus allowing the manipulation of biosynthetic pathways.

See also: Cheraviruses, Sadwaviruses and Torradoviruses (Secoviridae). Nepoviruses (Secoviridae). Secoviruses (Secoviridae). Sequiviruses and Waikaviruses (Secoviridae)

Further Reading Czapar, A.E., Steinmetz, N.F., 2017. Plant viruses and bacteriophages for drug delivery in medicine and biotechnology. Current Opinion in Chemical Biology 38, 108–116. Gergerich, R.C., Scott, H.A., 1996. Comoviruses: Transmission, epidemiology, and control. In: Harrison, B.D., Murant, A.F. (Eds.), The Plant Viruses 5: Polyhedral Virions and Bipartite RNA Genomes. New York: Plenum Press, p. 77. Goldbach, R.W., Wellink, J., 1996. Comoviruses: Molecular biology and replication. In: Harrison, B.D., Murant, A.F. (Eds.), The Plant Viruses 5: Polyhedral Virions and Bipartite RNA Genomes. New York: Plenum Press, p. 35. Hesketh, E.L., Meshcheriakova, Y., Dent, K.C., et al., 2015. Mechanisms of assembly and genome packaging in an RNA virus revealed by high-resolution cryo-EM. Nature Communications 6, 10113. Hesketh, E.L., Meshcheriakova, Y., Thompson, R.F., Lomonossoff, G.P., Ranson, N.A., 2017. The structures of a naturally empty Cowpea mosaic virus particle and its genomecontaining counterpart by cryo-electron microscopy. Science Reports 7, 539. Kruse, I., Peyret, H., Saxena, P., Lomonossoff, G.P., 2019. Encapsidation of Viral RNA in Picornavirales: Studies on Cowpea mosaic virus demonstrate dependence on viral replication. Journal of Virology 93.(pii: e01520-18). Lin, T., Johnson, J.E., 2003. Structure of picorna-like plant viruses: Implications and applications. Advances in Virus Research 62, 167–239. Lomonossoff, G.P., Evans, D.J., 2011. Applications of plant viruses in bio-nanotechnology. In: Palmer, K., Gleba, Y. (Eds.), Current Topics of Microbiology and Immunology vol. 375, pp. 61–87. Marsian, J., Lomonossoff, G.P., 2016. Molecular pharming – VLPs made in plants. Current Opinion in Biotechnology 37, 201–206. Peyret, H., Lomonossoff, G.P., 2015. When plant virology met Agrobacterium: The rise of the deconstructed clones. Plant Biotechnology Journal 13, 1121–1135. Sainsbury, F., Cañizares, M.C., Lomonossoff, G.P., 2010. Cowpea mosaic virus: The plant virus-based biotechnology workhorse. Annual Review of Phytopathology 48, 437–455. Sainsbury, F., Thuenemann, E.C., Lomonossoff, G.P., 2009. pEAQ: Versatile expression vectors for easy and quick transient expression of heterologous proteins in plants. Plant Biotechnology Journal 7, 682–693. Steinmetz, N.F., 2019. Biological and evolutionary concepts for nanoscale engineering: Viruses as natural nanoparticles have great potential for a wide range of nanoscale products. EMBO Reports 20 (8), (e48806).

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Relevant Websites http://www.dpvweb.net/dpv/showdpv.php?dpvno=378 Cowpea mosaic virus. https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/picornavirales/w/secoviridae/589/genus-comovirus Genus: Comovirus. http://viperdb.scripps.edu/genuslist.php?genus=Comovirus Virus Particle ExploreR.

Cucumber Mosaic Virus (Bromoviridae) Judith Hirsch and Benoît Moury, Plant Pathology Unit, INRAE – French National Research Institute for Agriculture, Food and Environment, Montfavet, France r 2021 Elsevier Ltd. All rights reserved. This is an update of F. García-Arenal, P. Palukaitis, Cucumber Mosaic Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00640-3.

Nomenclature aa Amino acid(s) AGO Argonaute proteins CP Coat protein or capsid protein CRISPR/Cas9 Clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 ELISA Enzyme-linked immunosorbent assays ER Endoplasmic reticulum kb Kilobase kDa Kilo dalton miRNA MicroRNA

Glossary Cross-protection Inhibition of systemic virus accumulation and disease by prior inoculation of plants with a mild or symptomless strain of the same virus. Filiformism Narrowing of the leaf blade, often leading to symptoms referred to as shoestring. Non-circulative A mode of virus transmission in which the virus does not need to circulate through the vector digestive system, hemocoel or salivary glands. The virus is carried by attachment to the mouthparts or foregut of the vector. Non-persistent A mode of virus transmission characterized by short acquisition and inoculation times (from seconds to minutes).

MP Movement protein nt Nucleotide(s) ORF Open reading frame RISC RNA-induced silencing complex RdRp RNA-dependent RNA polymerase RT-PCR Reverse transcription-polymerase chain reaction satRNA Satellite RNA sgRNA Sub-genomic RNA siRNA Small interfering RNA TILLING Targeting induced local lesions in genomes tRNA transfer RNA UTR Untranslated region

Pseudo-recombination Reassortment of the genomic segments of two or more strains of a virus to generate novel combinations of the full complement of genomic segments. Reassortant A virus resulting from reassortment (or pseudo-recombination), i.e., exchange of entire genome segments between two or more parental strains. Satellite RNA A subviral RNA dependent on a helper virus for both replication and encapsidation. Tonoplast The membrane surrounding the central vacuole of plant cells. Virulence The impact of a parasite on its host’s health or fitness. Yellowing Light green symptoms induced by virus infection affecting chlorophyll accumulation.

Introduction Cucumber mosaic virus (CMV) is the type species of the genus Cucumovirus in the family Bromoviridae, a group of single-stranded positive-sense RNA viruses. It was first described in 1916 as a disease of cucurbits in the USA. CMV has a worldwide distribution and a host range of over 1000 plant species. CMV is one of the most important agronomic pathogens on vegetable and ornamental plants as well as some important fruit crops. The virus is transmitted horizontally by about 80 aphid species in a non-persistent and noncirculative manner. In some plants, CMV is also transmitted vertically through the seed with variable, but usually low, efficiencies. CMV has a segmented genome and is multipartite, meaning that the three main genomic RNAs are encapsidated in separate particles (virions). CMV virions are isometric particles of 29 nm in diameter (T¼ 3 symmetry). The CMV genome encodes five proteins. The 1a and 2a proteins are involved primarily in virus replication occurring on tonoplast membranes, while the 2b, 3a, and 3b proteins are involved in virus cell-to-cell or systemic movement. The 2b protein is one of the best studied viral suppressors of plant defenses based on RNA silencing. Over 100 years of research on this agronomically important pathogen have yielded much knowledge on diverse aspects of virus biology, from epidemiology, ecology and evolution to the molecular mechanisms involved in virus-host interactions. Different isolates of CMV induce a variety of disease responses, which are attenuated by a satellite RNA (satRNA) associated with some isolates of CMV, although some satRNAs can enhance pathogenicity. Control of CMV is principally based on sanitary selection, prophylactic measures and use of resistant plant cultivars.

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Fig. 1 Schematic topology of phylogenetic trees representing CMV genetic diversity from full-genome sequences. Whereas numerous isolates from subgroups IA and IB and group II have been entirely sequenced (indicated by gray triangles), only one or two isolates of group III or outliers (isolates Rom from rosemary, 209 from soybean and BX and PHZ from Pinellia ternata) have been sequenced. Only branches supported by bootstrap values 470% are depicted (branch lengths are not at scale). The tree was rooted with other members of the genus Cucumovirus.

Taxonomy and Phylogeny CMV isolates differ from members of the other three species in the genus Cucumovirus, Tomato aspermy virus (TAV), Peanut stunt virus (PSV) and Gay feather mild mottle virus (GMMV), having only 50%–67% nt sequence identity, depending on the RNA and isolates being compared. Isolates of CMV are heterogeneous in symptoms, host range, transmission, serology, physicochemical properties, and nucleotide sequence of the genomic RNAs. CMV isolates can be classified into two major genetic groups, I and II. The nt identity between isolates belonging to these groups ranges from 69% to 77%, depending on the pair of isolates and the RNA segment compared, dissimilarity being highest for RNA2. Within a group, nt identity is above 88% for group I and above 96% for group II. Isolates of groups I and II can also be distinguished using monoclonal antibodies. RNA2 sequences of group I CMV isolates correspond to two monophyletic subgroups named IA and IB (Fig. 1). Subgroup IA, but not subgroup IB, is also monophyletic for RNA3, while RNA1 shows no clear division into subgroups IA and IB. This indicates that the different genomic segments have followed different evolutionary histories. Crossprotection occurs between isolates from all groups or subgroups. A few CMV isolates do not belong to groups I or II (Fig. 1).

Geographic Distribution CMV isolates have a worldwide distribution, having been reported from both temperate and tropical regions. Most reported isolates belong to group I. Group II isolates are found more frequently in cooler areas or seasons of temperate regions. This has been associated with lower temperature optima for in planta virus accumulation shown for the few isolates characterized for this property. Most isolates in subgroup IB have been reported from East Asia, which is presumed to be the origin of this subgroup. Subgroup IB isolates have also been reported from other areas, for example, the Mediterranean region, California, Brazil, Africa and Australia. Those in the Mediterranean Basin could have been introduced recently from East Asia.

Host Range Considering all isolates collectively, the host range of CMV includes 1071 plant species belonging to 100 different families, monocot or dicot. Of these plants, more than 500 species were found infected naturally, whereas the others were only found infected in laboratory conditions. Four families include more than 100 host species each: the Solanaceae, Asteraceae, Fabaceae and Brassicaceae. CMV infects most of the major horticultural crops as well as many weed species acting as reservoirs for the virus. However, considerable differences in host range breadth exist between individual CMV isolates. Most isolates have a broad host range while a few isolates have a very narrow host range, like certain isolates from soybean, lily or rosemary. These ‘soybean’ and ‘rosemary’ isolates, as well as isolates from Pinellia ternata, are phylogenetically distant from groups I and II (Fig. 1). Recent evidence has expanded the host range of CMV outside of the plant kingdom: a phytopathogenic fungus, Rhizoctonia solani, was described as a natural alternative host for CMV. This non-plant host may have an important impact on epidemiology of the disease, since the fungus could acquire CMV from an infected plant and transmit it to a healthy plant under laboratory conditions.

Symptoms and Diagnosis Symptoms induced by CMV include leaf mosaic or mottling, vein-clearing, filiformism (the shoestring appearance of the leaf blade, a typical CMV symptom on tomato leaves), stunting, chlorosis and necrosis (including necrotic “oak-leaf” symptoms on older leaves) as well as decreased fruit or seed yield (Fig. 2). Pepper and cucurbit fruits, among others, can display symptoms such as mottling, mosaic, ring spotting or deformations. Symptom type and severity can vary greatly depending on the host species, the particular combination of plant and virus genotypes, the phenological stage at the time of infection, environmental conditions, as well as the presence of other

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Fig. 2 Symptoms associated with CMV infection. (A) CMV-induced symptoms in a pepper field. (B) Mosaic symptoms on tobacco leaves infected with CMV I17F isolate (belonging to the IA subgroup). (C) Pepper plants mechanically inoculated with CMV I17F isolate displaying stunting and mosaic symptoms two weeks after inoculation (left infected plant, right mock-inoculated plant). (D) Necrotic oak-leaf pattern on naturally infected pepper leaf. Photographs Judith Hirsch, INRAE Avignon.

biotic factors such as satRNAs and other viruses, which can modulate the disease symptoms induced by CMV. CMV infection can also be symptomless. This is the case in many weed species which can serve as asymptomatic CMV reservoirs. Host species which are systemically infected by most CMV isolates and commonly used as indicator plants include squash (Cucurbita pepo), cucumber (Cucumis sativus), tobacco (Nicotiana tabacum), as well as N. benthamiana, N. clevelandii, and N. glutinosa. Chenopodium amaranticolor, C. quinoa and cowpea (Vigna unguiculata) are local lesion hosts. Serological methods like enzyme-linked immunosorbent assay (ELISA) have been successfully used for the large scale detection and diagnosis of CMV. Although molecular methods like reverse transcription-polymerase chain reaction (RT-PCR) are more sensitive, since CMV usually accumulates to high levels in infected plants, it is readily detected using ELISA, which makes this serological assay the method of choice for detecting CMV in plant samples. Another widely used serological diagnostic method is the lateral flow test (immune-strip tests based on this method are commercially available). Although lateral flow tests are relatively expensive, they constitute a rapid means of screening crops for the presence of CMV, particularly adapted for diagnostics in the field or greenhouse.

Cytopathology Cytopathology associated with infection by CMV includes viral inclusions within the cytoplasm and vacuoles, or forming membrane-bound clusters in sieve elements. In some cases, angular inclusions corresponding to virus crystals can be seen in vacuoles by staining and light microscopy. CMV infection usually also leads to proliferation of cytoplasmic membranes, which originate from the plasma membrane, the endoplasmic reticulum (ER), or the tonoplast. Effects on the nucleus or nucleolus (e.g., vacuolation), usually due to virion accumulation, have been observed with some isolates of CMV. Similarly, some isolates have caused effects on mitochondria or chloroplasts. Yellowing isolates, in particular, lead to smaller and rounded chloroplasts that have fewer grana and starch granules. These various effects appear to be host specific. How virus–plant interactions lead to cytopathic effects remains unknown; however, as the virus expands from the initial site of infection, rings or zones of responses occur, in which the expression patterns of numerous plant genes are altered in spatial- and temporal-specific manners.

Virion Structure Transmission electron microscopy observations allow visualization of the uniform isometric (roughly spherical) particles of CMV. The atomic structure of CMV has been determined to a 3.2 Å resolution using X-ray crystallography. CMV virions are icosahedral particles 29 nm in diameter. They are composed of 180 subunits of a single capsid protein (CP) of approximately 24 kDa, and contain approximately 18% RNA. RNA-protein interactions stabilize the virions and particle assembly requires a nucleic acid as a nucleating factor (no empty particles are formed). Virions are disrupted by sodium dodecyl sulfate and concentrated chloride salts, but can be

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Fig. 3 Cryo-transmission electron microscopy reconstruction of CMV particles. Icosahedral CMV virions are composed of 12 pentamers and 20 hexamers. Reproduced from Wikoff, W.R., Tsai, C.J., Wang, G., Baker, T.S., Johnson, J.E., 1997. The structure of cucumber mosaic virus: Cryo-electron microscopy, X-ray crystallography, and sequence analysis. Virology 232 (1), 91–97.

Fig. 4 Schematic representation of the genome organization of CMV. The CMV genome is composed of 3 genomic RNAs, RNAs 1, 2, and 3 encoding 5 proteins. Two sub-genomic RNAs, RNAs 4A and 4 are transcribed from the minus RNA2 and RNA3 strands, respectively. The open reading frames are depicted as colored bars. The approximate length of the RNAs are indicated. nt, nucleotides; sgRNA, sub-genomic RNA; MP, movement protein; CP, capsid protein.

reassembled in vitro by removing salts or sodium dodecyl sulfate. Each CP subunit adopts a canonical b-barrel structure, and all subunits assume one of three different configurations (A, B, or C). The A subunits cluster to form pentameric capsomers (or pentamers) and the B and C subunits form hexameric capsomers (or hexamers). The capsid is composed of 12 pentamers and 20 hexamers arranged with T¼ 3 symmetry (Fig. 3). The N termini of the B and C subunits form a bundle of six alpha helices. This hexameric bundle constitutes a hydrophobic pore lined with leucine residues that extends down to the RNA-capsid interface. The first 30 residues of the CP (upstream of the alpha helix) are rich in positively charged aa, including 7 conserved arginine residues, that are proposed to form the interface with the RNA core. RNAs that can be encapsidated in CMV virions include genomic RNAs 1, 2, and 3, sub-genomic RNAs (sgRNAs) 4 and 4A, RNA5, transfer RNAs (tRNAs), as well as satellite and defective RNAs (see below).

Genome Organization The genome of CMV consists of five genes distributed over three, single-stranded, positive-sense, capped genomic RNAs designated RNAs 1, 2, and 3 in descending order of size (Fig. 4). CMV is a multipartite virus, meaning that the three genomic RNAs are encapsidated in separate virus particles. RNA1 (B3.3 kb) encodes the 111 kDa 1a protein. RNA2 (B3.0 kb) encodes the 98 kDa 2a protein, as well as the 13–15 kDa 2b protein, which is translated from a B0.7 kb sub-genomic RNA designated RNA 4A that is co-terminal with the 30 end of RNA2. The Open Reading Frame (ORF) expressing the 2b protein overlaps with the ORF encoding the 2a protein, but in a þ 1 reading frame. RNA3 (B2.2 kb) encodes the 30 kDa 3a protein, as well as the 25 kDa 3b protein, which is expressed from a B1 kb sub-genomic RNA designated RNA4 that is co-terminal with the 30 end of RNA3. RNA4 is co-encapsidated with RNA3. The 30 untranslated regions (UTRs) of all three genomic and both sub-genomic RNAs are highly conserved, forming a tRNA-like structure as well as several pseudoknots. The 50 UTR regions of RNA1 and RNA2 are more conserved in sequence with each other than with those of RNA3. CMV also produces an RNA5 of unknown function, which is co-terminal with the 30 UTR regions of RNA1 and RNA2. RNA4A and RNA5 are only encapsidated by group II isolates. CMV

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particles also encapsidate a low level of tRNAs, which have been reported in the literature as CMV RNA6. Although rarely reported, some isolates of CMV can also encapsidate defective RNAs derived from CMV RNA3. CMV can also support satRNAs varying in size from 333 to 405 nt. These satRNAs do not encode any proteins. They are dependent upon CMV as the helper virus for both their replication and encapsidation, but have sequence similarity to the CMV RNAs limited to no more than 6–8 contiguous nts. The origin of satRNAs remains enigmatic. The de novo emergence of satRNAs after serial passages of CMV devoid of satRNA in tobacco plants suggests that they may have arisen from the host genome. Failure to detect long satRNA homologs in host plant genomes has led to an interesting hypothesis: that host genome‐encoded small RNAs, possibly induced by viral infection, could be assembled by the viral replicase to form satRNAs. More than 180 satellite variants have been found associated with over 65 isolates of CMV from both of the CMV groups. These satRNAs usually reduce the accumulation of the helper viruses and on most hosts also reduce the virulence of CMV. However, certain satRNAs can enhance the disease induced by CMV in some hosts. In the case of tomato plants infected by CMV and certain satellites, this has led to systemic necrosis observed in the fields of several Mediterranean countries.

Properties and Functions of Gene Products The CMV 1a and 2a proteins contribute to virus replication but also promote virus movement in several host species. The 2b protein is an RNA-silencing suppressor that interferes with the salicylic acid (SA) and jasmonic acid (JA) plant defense pathways and also influences virus movement in some hosts. The predominant mechanism by which the 2b protein inhibits antiviral silencing is by sequestering virus-derived small RNAs. Additionally, the 2b protein binds to Argonaute (AGO) proteins, the catalytic component of the RNA-induced silencing complex (RISC), and inhibits their RNA cleavage activity. The contribution of the 2b-AGO interaction to antiviral silencing is minor, likely through the inhibition of signal amplification mediated by host RNA-dependent RNA polymerases (RdRps). Conversely, the 2b-AGO interaction plays an important role in the induction of developmental symptoms, through the disruption of microRNA- (miRNA) mediated post-transcriptional gene silencing. The 3a protein is the major movement protein (MP) of the virus and is essential for both cell-to-cell and long-distance (systemic) movement. The 3b protein is the sole viral CP and is also required for cell-to-cell and systemic movement, although the ability to form virions is not a requirement for intercellular trafficking of the virus. All of the CMV-encoded proteins are RNA-binding proteins. Viral RNAs from different groups and subgroups can be exchanged to form novel viruses, allowing mapping of some phenotypes to specific RNAs. Higher-resolution mapping requires the use of biologically active cDNA clones for generating chimeras and site-directed mutants. By these approaches, a large number of genetic determinants of pathogenicity or transmission have been mapped (Table 1). All CMV RNAs and genes can be involved in infection and symptomatology, depending on the plant species or genotype. The biochemical mechanisms responsible for the variety of symptoms induced by CMV have rarely been determined. In the case of an isolate of CMV that induces chlorotic symptoms in tobacco, interaction of the virus CP with the ferredoxin I protein precursor interferes with its import into the chloroplast and is therefore responsible for the symptoms. Moreover, particular satRNAs can also contribute to CMV symptomatology, sometimes causing an attenuation of symptoms, a property that has been used commercially in tomato production. Attenuation of disease appears to result from the production of satellite-derived small interfering RNAs (siRNAs) that can either directly target the conserved 30 UTR region of the CMV genomic RNAs and trigger their degradation through the host RNA silencing machinery or interfere with the RNA silencing suppressor activity of the 2b protein by binding to it.

The CMV Infection Cycle Replication CMV replication takes place on the vacuole membrane (the tonoplast). Replication involves the 1a and 2a proteins of the virus and a number of host proteins. The purified CMV replicase contains the 1a and 2a proteins, as well as a 50 kDa host protein of unknown function. The 1a protein of CMV contains an N-terminal methyltransferase domain involved in capping of the RNAs, as well as a putative C-terminal helicase domain, presumed to be required for the unwinding of the viral RNAs during replication. The 2a protein possesses a GDD (Glycine-Aspartic acid-Aspartic acid) motif found in most RdRps. The N-terminal region of the 2a protein interacts with the C-terminal region of the 1a protein in vivo and in vitro. CMV replication is initiated by the binding of the tonoplast-associated replicase to the tRNA-like structure and various pseudoknots present in the 30 UTR region of the positive-sense CMV RNAs. Minus-sense RNAs are then synthesized from each of the genomic RNAs and act as templates for synthesis of new plus-stranded genomic RNAs. The minus-sense RNA2 and RNA3 also serve as templates for the synthesis of the two plus-sense subgenomic RNAs (4 and 4A), through recognition of the sub-genomic promoters present on the minus-sense RNAs. The subgenomic RNAs are not themselves replicated, but defective and satellite RNAs are replicated by the CMV replicase. Differences in the relative levels of accumulation of the various CMV genomic and satellite RNAs have been observed in different host species, which is probably due to host-specific differences in template copying. Like most plant viruses, CMV encodes a very small number of proteins and relies on plant host factors to fulfill the different steps of its infection cycle. CMV replication is intricately dependent on various host proteins. Several host factors which support or

Nicotiana tabacum Nicotiana spp. Nicotiana glutinosa

Symptoms Chlorosis, mosaic, or necrosis Green mosaic, yellow chlorosis Top necrosis

Genome region

N. tabacum N. tabacum

Legumes

C. pepo

Aphid transmission Transmission efficiency Transmission efficiency

Seed transmission Transmission efficiency

Others Satellite replication

Abbreviations: CP, coat protein (i.e., 3b protein); HR, hypersensitive reaction; MP, movement protein (i.e., 3a protein); UTR, untranslated region.

C. pepo, Phaseolus vulgaris, Pisum sativum Cucumis melo Vicia faba Zea mays Tetragonia expansa, Momordica charantia, Physalis floridana Lactuca saligna, Raphanus sativus Arabidopsis thaliana (RCY1 gene) Vigna unguiculata (Cry gene) Phaseolus vulgaris (RT4–4 gene) N. tabacum Nicotiana spp. (9 species) C. melo (cmv1 gene) Capsicum annuum (Cmr1 gene)

Infection Infection at 271C but not at 371C Infection Infection Infection Infection HR triggering HR triggering HR triggering HR triggering HR triggering Resistance breakdown Resistance breakdown

Host range and interaction with plant resistance Infection Lilium spp.

nucleotides 148, 149, 153, 158, 170 (chlorosis)

267 of 2a and 168 of 3a

193

129, 111, 124 111, 124

Mutation (codon)

RNA 1

RNA 1

CP gene 2b gene

25, 129, 162,168, 214

RNA 1 (50 UTR and/or 1a 485–826 of 1a gene) RNA 2 RNA 1 RNA 3 CP gene 129, 162 CP gene 129 RNAs 2 and 3 CP gene 2a gene 631, 641 2a gene 1a gene 461 CP gene 36 MP gene 64–68, 81, 171, 195 1a gene 865, 896, 957, 980

CP gene CP gene 2b gene alone or 2b and 2a genes Necrotic etch N. tabacum RNA 3 (50 UTR and/or MP gene) Stunting N. glutinosa CP gene Severe symptoms Cucurbita pepo RNA 1 Stunting, filimorfism N. tabacum RNAs 1 and/or 2 Fast infection and severe symptoms C. pepo RNAs 1, 2, and 3 Severe symptoms Datura stramonium, Gomphrena globosa, Nicotiana clevelandii, Nicotiana x edwardsonii, N. glutinosa, RNA 2 or RNAs 2 and 3 N. tabacum, Solanum melongena White-yellow leaves, systemic necrosis, N. tabacum, Solanum lycopersicum Satellite RNAs fruit necrosis

Plant

Molecular determinants of pathogenicity and transmission identified in CMV

Phenotype

Table 1

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restrict virus multiplication have been identified (Table 2). Although the exact roles of these proteins in virus replication are not always clear, they are proposed to act through regulation, assembly and/or anchoring of the CMV replication complex to the tonoplast. One such protein is glyceraldehyde 3-phosphate dehydrogenase (GAPDH) which is critical for CMV replication and appears to be involved in assembly of the viral replication complex by promoting the interaction between the 1a and 2a proteins. Comparatively, very few plant factors involved in cell-to-cell or systemic propagation of the virus have been identified (see below and Table 2) although CMV presumably co-opts a number of host factors for these steps of its infection cycle. Post-translational modifications of viral replication proteins play an important role in the regulation of CMV replication. For example, phosphorylation of the 2a protein prevents its association with the 1a protein and thus regulates assembly of the replicase complex. This mechanism probably plays an important role in the regulation of plus-strand and minus-strand synthesis and the asymmetry of CMV replication. Indeed, the kinetics of accumulation of positive and negative strands are distinct and accumulation of CMV positive strand RNAs is approximately 100-fold that of negative strand RNAs. Free 2a protein is present in the cytoplasm, while the 1a protein is strictly membrane-associated. The 2a protein alone is sufficient to synthesize positive strands from negative strand templates, while both the 1a and 2a proteins are required for synthesis of negative strands. Therefore the switch from negative to positive strand synthesis may be controlled by 2a phosphorylation, resulting in dissociation of 1a-2a complexes. Positive-sense viral RNAs serve as templates for replication and translation of viral proteins and are encapsidated to form virions. The balance between replication, translation, encapsidation and probably also movement of these viral RNAs determines the efficiency of virus multiplication.

Cell-to-Cell and Long-Distance Movement CMV moves cell to cell via plasmodesmata until it reaches the vasculature, when the virus moves systemically via the phloem. The viral RNAs move as a nucleoprotein complex between cells involving the 3a MP. Although the virus does not appear to migrate as virus particles, the CP is also necessary for cell-to-cell movement. This movement does not involve interactions with microtubules. Conversely, the actin cytoskeleton appears to be involved in CMV cell-to-cell movement (Table 2). The MP has been shown to interact with and sever actin filaments, an activity associated with increase of the plasmodesmal size exclusion limit. CMV appears to move between epidermal cells as well as from epidermal cells down to mesophyll cells toward vascular cells. The virus replicates in all of these cell types, but not in the sieve elements of the vasculature. Long-distance movement includes three main steps: viral loading into the phloem tissues, movement within sieve elements, and unloading from these elements into cells of the sink tissues. Virion assembly may take place inside sieve elements from RNAs and capsid protein moving from neighboring vascular cells. Virion assembly is necessary in some, but not all, species for systemic infection. A 48 kDa phloem protein (PP1) from cucumber phloem exudates interacts with CMV particles in vitro and increases virus particle stability. How the virus moves from the vasculature back to mesophyll and epidermal cells is unknown. The 2b protein is also involved in local and systemic movement of the virus. The 2b protein has indirect effects on virus movement through its RNA silencing suppressor activity, as well as direct effects on cell-to-cell movement, independently of this suppressor activity, as has been revealed in an alanine scanning mutagenesis study that identified residues in the 2b protein that are required for local movement but not for silencing suppression. The elucidation of resistance mechanisms targeted against systemic spread of CMV is revealing some of the probably manifold resistance barriers preventing long-distance movement of the virus. For example, a Piezo-like mechanosensitive cation channel which suppresses systemic spread of CMV has been identified in the model plant Arabidopsis (see also in the “Control” section the molecular mechanisms involved in resistance conferred by the cmv1resistance gene in melon). Conversely, other host proteins like Tcoi1 (tobacco CMV 1a-interacting protein 1) promote systemic spread of the virus (Table 2). Plant proteins that are essential to the virus infection cycle can serve as targets to screen the plant natural genetic diversity in order to identify resistance alleles or to engineer new resistances, using technologies such as CRISPR/Cas9 (Clustered Regularly Interspaced Short Palindromic Repeats/CRISPR associated protein 9) genome editing or TILLING (Targeting Induced Local Lesions in Genomes).

Transmission Seed transmission of CMV has been reported in 41 plant species, mostly in the families Fabaceae (26 species) and Cucurbitaceae (seven species), with efficiencies varying from less than 1% up to almost 100%. The virus may be present in the embryo, endosperm, and seminal integuments, as well as in pollen. RNA1, possibly via the encoded protein 1a, affects the efficiency of seed transmission. Horizontal transmission of CMV is mediated by aphids (family Aphididae) in a non-persistent and noncirculative manner. This mode of virus transmission (also called stylet-borne) is characterized by short acquisition and inoculation times (from seconds to minutes). The stylets of the aphid become contaminated with the virus during probing of infected tissue, through a series of short intracellular stylet punctures of epidermal or neighboring cells. The virions are retained at sites at the tip of the aphid stylet, and released by salivation during probing of non-infected plants. Over 80 species of aphids have been reported to transmit CMV, Aphis gossypii and Myzus persicae being two efficient and most studied vectors. Transmission efficiency depends on several factors, particularly the specific combination of virus isolate and aphid species, and the

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Table 2 Plant proteins having a positive or negative effect on one or more steps of the virus infection cycle or that interact with CMV proteins and play a putative role in assembly, anchoring or regulation of the CMV replicase complex

Cucumber Mosaic Virus (Bromoviridae)

Table 2

379

Continued

a

Are not included in this table plant factors that are part of the anviral silencing machinery or involved in its regulaon, which are not specific to CMV. Text in blue font: no funconal data available concerning the role of the host protein in the CMV infecon cycle. Abbreviaons: APUM5, arabidopsis pumilio 5; BRP1, bromodomain-containing protein 1; CIPK12, CBLinteracng protein kinase 12; Co-IP, co-immunoprecipitaon; CRISPR/Cas9, clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9; CRT3, calreculin-3 precursor; CsAO4, cucumis savus ascorbate oxidase 4; eIF, eukaryoc translaon iniaon factor; EMS, ethyl methane-sulfonate; FDH, formate dehydrogenase; GAPDH, glyceraldehyde 3-phosphate dehydrogenase; LC-MS/MS, liquid chromatographytandem mass spectrometry; MP, movement protein; PP1, phloem protein 1; RNAi, RNA interference; Tcoi, tobacco CMV 1a-interacng protein; TIP, tonoplast intrinsic protein; TRV, tobacco rale virus; Tsip1, Tobacco stress induced 1(Tsi1) interacng protein; UTR, untranslated region; VIGS, virus-induced gene silencing; Y2H yeast two-hybrid. accumulation of particles in the source leaf. Differences in transmissibility of various isolates by different aphid species are determined primarily by the virus coat protein and aa determinants for transmission have been mapped (Table 1). The aa positions that determine transmission are either exposed on the outer surface of capsids (e.g., aa position 129) or lay in the inner surface (e.g., position 162). Hence, the effect could be through direct interaction with receptors in the aphid mouth parts or by affecting particle stability. On the vector side, there is accumulating evidence for the role of cuticular proteins of the CPR (cuticular proteins with the Rebers and Riddiford Consensus) family in the transmission of plant viruses. Putative CMV receptors belonging to this family are present in the acrostyle, a small structure inside the fused food/salivary common canal at the tip of aphid stylets. One of these proteins has been shown to act as a probable caulimovirus receptor. However, there is no direct evidence of binding of CMV in the common canal, and validating the role of these cuticular proteins in CMV transmission will require further biochemical and functional studies. The 2b protein of CMV also seems to have an indirect effect on virus transmission by manipulating the behavior of aphids. By altering the jasmonate signaling pathway and increasing the plant reactive oxygen species content, the 2b protein simultaneously enhances attractiveness of host plants for aphid vectors and increases probing and dispersal of the aphids, thus favoring spread of the virus. Transmission efficiency also depends on the host plant, as shown by the resistance to transmission of melon genotypes carrying the Vat gene, which does not confer resistance to the virus directly, but impairs CMV transmission by Aphis gossypii.

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Variation and Evolution CMV populations share evolution patterns of many plant RNA viruses, including high mutation rates and high intensities of selection and genetic drift. Moreover, phylogenetic analysis showed that recombination and particularly reassortment have largely contributed to the diversification of CMV populations, as seen in other viruses with segmented genomes. Using infectious cDNA clones and experimental evolution, CMV mutation and recombination frequencies were estimated. Mutation frequency varied from 0.6  10–4 to 1.8  10–3 mutation per nt in a 12–15 days passage in plants. Significant differences in mutation frequency were observed between plant species and between CMV isolates. The recombination frequency was also highly variable between isolates. Indeed, depending on the parental isolates, from 1.8% to 28.2% of RNA3 recombinants were observed 15 days after inoculation with parental CMV clones. The 2a gene is a major determinant of recombination frequency differences. Recombination also seems to be facilitated by stem–loop structures in the RNA. The 30 UTR of CMV genomic RNAs is also a recombination hotspot. Recombinants in this region may have an increased fitness in some hosts, as shown for isolates infecting Alstroemeria spp. However, analyses of field populations of CMV show that recombinants are not frequent, suggesting that selection operates against most of them. A second mechanism of genetic exchange is reassortment of genomic segments also called pseudo-recombination. Reassortants exchanging any genomic segment have been obtained between different CMV isolates and multiply efficiently under experimental conditions. Natural CMV reassortants also have been described but are generally rare and seem to be selected against, as is the case for recombinants, which suggests co-adaptation of the different viral genes that, when disrupted, results in a decreased fitness. However, reassortants may reach higher frequencies as shown in pepper (Capsicum spp.) crops in Tunisia and South Korea, where they comprised 90% and 20%, respectively, of the CMV populations. Selection by the host plant and possibly by resistant cultivars may be responsible for this over-representation of reassortants. Genetic diversity of CMV has been shown to be countered by genetic drift associated to population bottlenecks during plant colonization and horizontal transmission by aphids. For the latter, from one to two CMV infectious units are inoculated by a single aphid. Selection is also an evolutionary force that drastically restricts the diversity within virus populations. Sequence analysis have shown different evolutionary constraints for the different viral proteins. Protein 1a and the MP are the most constrained, whereas the 2b protein is the least constrained. The evolution constraint acting on the CP differs greatly between CMV groups and subgroups. Positively-selected aa positions have been detected in all CMV proteins and may be responsible for CMV adaptation to host plants or to the vector. Analyses of the population structure of CMV in Spain and California indicate a metapopulation structure, with local extinctions and recolonizations, which suggests that population bottlenecks occur, probably associated with unfavorable seasons for the host plants and/or the aphid vectors. Interestingly, this is not the case for the population structure of the satRNA. Analyses in Italy and Spain during epidemics of CMV and the associated satRNAs have shown that the satRNAs have an undifferentiated population. The different population structures for CMV and its satRNAs indicate that the satRNAs expanded as a molecular parasite on the CMV population, rather than satellite expansion was linked to a particular CMV isolate.

Epidemiology and Control CMV infects a wide range of food crops, ornamentals and wild plant species. Economic losses in crops are highest in field-grown vegetables and ornamentals, pasture legumes and banana. In recent times, CMV has caused severe epidemics in many crops, including necrosis of tomato in Italy, Spain, and Japan; mosaic and heart rot of banana worldwide; mosaic of melons in California and Spain; mosaic of pepper in Australia, California, Turkey and Tunisia; and mosaic of lupins and other legumes in Australia and the USA. As for many plant viral diseases for which there are no curative methods, control of CMV begins with sanitary selection, planting or sowing virus-free material and prophylactic measures. In crops in which seed transmission is effective (e.g., pasture or fodder legumes), the primary inoculum for epidemics may be the seedlings from infected seeds and even a low frequency of seed infection may have dramatic consequences because of accelerated secondary spread of the virus. In most vegetable and ornamental crops, seed transmission does not occur or is negligible, and the primary inoculum must come from outer sources as other crops or weeds, which should be near the crop as aphid transmission is non-persistent. In the absence of crops during unfavorable seasons, infected perennial weeds or crops, and infected seeds from weeds act as inoculum reservoirs. Seed transmission has been shown to be important in several weed species from different regions. The dynamics of virus infection may differ largely in weeds and crops within a region, indicating that the relevant reservoirs and inoculum sources for crops need not be the most frequently infected weeds. In banana, secondary spread of infection within the crop is ineffective for most isolates, and alternative hosts are both primary and secondary inoculum sources for epidemics. Since many species of weeds act as reservoirs for the virus and many of these are asymptomatic hosts, it is important to remove them from the borders of fields. This also applies to removal of infected crop plants during the growing season. A number of agricultural practices can be implemented to reduce CMV epidemics. For example, avoiding overlap of the growing cycles of CMV-susceptible crops, adapting sowing or planting dates in order to create a lag time between peaks of aphid flights and the susceptibility period of crop plants and using cultivar or crop mixing to benefit from inoculum dilution and barrier effects can slow down CMV epidemics. The use of mineral oils or reflective mulches can also affect aphid behavior and contribute to CMV control.

Cucumber Mosaic Virus (Bromoviridae)

Table 3

381

Characteristics of CMV resistance genes or loci

Plant

Gene or allele name

Nature of encoded protein

Resistance mechanism

Arabidopsis thaliana Capsicum annuum C. annuum C. annuum

RCY1 Cmr1 cmr2 Several QTLs

NLR – – –

Capsicum frutescens Cucumis melo C. melo Cucumis sativus

At least 2 recessive genes cmv1 Vat Several QTLs or 1 recessive gene Cmv-2

– VPS41 NLR –

HR Inhibition of systemic movement Inhibition of cell-to-cell movement Inhibition of infection, virus multiplication and systemic movement Inhibition of cell-to-cell and systemic movements Inhibition of systemic movement of group II strains Resistance to Aphis gossypii-mediated inoculation Symptom attenuation

–

High-level resistance

Three QTLs Nam-1 Several QTLs RT4–4 Cmr polygenic Single major gene Cry Single dominant gene

–

Inhibition of systemic movement HR Inhibition of infection, symptom attenuation HR Symptom attenuation Tolerance to satRNA-induced lethal necrosis Inhibition of systemic movement HR HR

Cucurbita moschata, Cucurbita pepo Glycine max Lupinus luteus Nicotiana tabacum Phaseolus vulgaris Solanum chilense Solanum habrochaites Solanum tuberosum Vigna unguiculata Zea mays

– NLR – – – – –

Abbreviations: HR, hypersensitive reaction; NLR, nucleotide-binding and leucine-rich repeat-containing protein; QTL, quantitative trait locus; satRNA, satellite RNA; VPS, vacuolar protein sorting; –, unknown data.

Control of CMV can also be achieved by planting of resistant crop varieties. Genetic resources for CMV resistance are available in many crop species and their genetic determinism has been unraveled in a few of them (Table 3). However, there is generally a lack of high-level resistance and of major-effect resistance genes against CMV. A large number of resistance mechanisms are polygenic, which hampers their exploitation by breeders, and partial. They frequently slow down the systemic movement of the virus and reduce symptoms. Some of them are ontogenic, i.e., they are only expressed when plants achieve a given developmental stage. Their spectrum of action is also frequently incomplete and covers a few CMV isolates or isolate groups. A novel mechanism has been discovered for the resistance conferred by the cmv1 gene in Cucumis melo. cmv1 inhibits CMV systemic infection by restricting the virus to the bundle sheath cells and impeding its loading into the phloem. cmv1 was cloned and shown to encode a novel class of resistance factors, the Vacuolar Protein Sorting 41 (VPS41) which is required for post-Golgi vesicle trafficking towards the vacuole. CMV might use VPS41 for its transport towards the phloem and a single aa mutation in the protein abolishes or reduces this transport and converts the VPS41 gene into a recessive resistance gene. However, the resistance is efficient only against group II CMV isolates, which reduces its agronomical interest. In pepper (Capsicum spp.), several resistance sources and mechanisms have been described and exploited to breed resistant cultivars. In France, cultivars carrying a polygenic resistance were bred using a recurrent selection program involving several parental lines with different partial resistance mechanisms (inhibition of inoculation efficiency, decrease of virus multiplication and inhibition of systemic movement). Whereas all resistance sources were partial, the composite resistance resulting from the combination of these mechanisms is almost total. The resistance was efficient and durable in the field but is threatened by adapted CMV isolates, belonging to the IB subgroup, that have been introduced in the Mediterranean basin. In South Korea, the majoreffect dominant resistance gene Cmr1 was used in breeding programs and deployed in the field. It inhibits the systemic movement of CMV. However, as in the previous case, it is impaired by subgroup IB isolates. The cmr2 recessive resistance gene seems to possess a broader spectrum of action and may be a valuable resource for resistance breeding in pepper. In addition to resistance, mechanisms of plant tolerance against CMV have also been studied. Tolerance is defined as the ability of the host to limit the damage caused by a given virus load, as opposed to resistance, which is the capacity of the host to reduce virus load. Tolerance to CMV has been studied in detail in the model plant Arabidopsis thaliana but has also been observed in several agronomically important plants, including bean, cucumber, pepper and tomato. The effect of infection by different CMV isolates on biomass and viable seed production of different Arabidopsis accessions was evaluated. Tolerance to different CMV isolates was higher in accessions with longer life cycles and was associated with resource reallocation from growth to reproduction. This tolerance response is CMV-specific as it was not observed in response to infection by other more virulent viruses. Whether similar tolerance mechanisms exist in cultivated plants remains to be determined. As it exerts a weaker selection pressure on virus populations, tolerance is expected to be more durable than resistance. Thus, whatever the mechanisms involved, breeding CMV tolerant crops is a potential source of sustainable protection against CMV. Other sources of resistance include the use of transgenic plants expressing either protein-mediated or RNA silencing-mediated resistance. Although most of these approaches have led to resistance to only members of one of the two major groups, the use of

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pyramiding of viral segments from different groups offers the promise of obtaining a broad-spectrum resistance to CMV together with other viruses infecting the same crop species. Transgenic expression of satRNAs has also been used to confer resistance to CMV. This has met with success, but has raised concerns about favoring the emergence of a virulent pathogen, as did the use of mild isolates of CMV for cross-protection against severe isolates. The use of insecticides to control the aphid vectors of the virus has met with only limited success, since the virus is transmitted in a non-persistent manner and thus the aphids can transmit the virus before being killed by the insecticide. Rather, insecticides are used to reduce aphid population size and thus reduce the spread of infections. Similar results were obtained through the biological control of aphid populations by the use of predators.

See also: Brome Mosaic Virus (Bromoviridae). Ilarviruses (Bromoviridae). Satellite Nucleic Acids and Viruses. Viral Suppressors of Gene Silencing

Further Reading Ben Tamarzizt, H., Montarry, J., Girardot, G., et al., 2013. Cucumber mosaic virus populations in Tunisian pepper crops are mainly composed of virus reassortants with resistance-breaking properties. Plant Pathology 62, 1415–1428. Deshoux, M., Monsion, B., Uzest, M., 2018. Insect cuticular proteins and their role in transmission of phytoviruses. Current Opinion in Virology 33, 137–143. Giner, A., Pascual, L., Bourgeois, M., et al., 2017. A mutation in the melon vacuolar protein sorting 41 prevents systemic infection of Cucumber mosaic virus. Scientific Reports 7, 10471. Jacquemond, M., 2012. Cucumber mosaic virus. Advances in Virus Research 84, 439–504. Lewsey, M.G., Murphy, A.M., Maclean, D., et al., 2010. Disruption of two defensive signaling pathways by a viral RNA silencing suppressor. Molecular Plant-Microbe Interactions 23 (7), 835–845. Pagán, I., Alonso-Blanco, C., García-Arenal, F., 2007. The relationship of within-host multiplication and virulence in a plant-virus system. PLoS One 2 (8), e786. Palukaitis, P., García-Arenal, F., 2019. Cucumber Mosaic Virus. The American Phytopathological Society. Seo, J.-K., Kwon, S.-J., Choi, H.-S., Kim, K.-H., 2009. Evidence for alternate states of Cucumber mosaic virus replicase assembly in positive- and negative-strand RNA synthesis. Virology 383, 248–260. Wang, M., Smith, N.A., 2016. Satellite RNA pathogens of plants: impacts and origins – An RNA silencing perspective. Wiley Interdiscip. Rev. RNA 7, 5–16. Wu, D., Qi, T., Li, W., et al., 2017. Viral effector protein manipulates host hormone signaling to attract insect vectors. Cell Research 27, 402–415.

Dianthovirus (Tombusviridae) Kiwamu Hyodo, Okayama University, Kurashiki, Japan Masanori Kaido, Kyoto University, Kyoto, Japan r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

nt Nucleotide(s) RdRp RNA-dependent RNA polymerase sgRNA Sub-genomic RNA SL Stem loop ssRNA Single-stranded RNA UTR Untranslated region VRC Viral replicase complex

aa Amino acid(s) CP Coat protein gRNA Genomic RNA kb Kilobase kDa Kilo dalton MP Movement protein

Glossary Dicer Eukaryotic ribonuclease that cuts long dsRNAs and structured ssRNAs into B20–25 bp fragments with 2-nt 50 extensions. Frameshift A change of the reading frame, which results in a different translation and a different protein product.

Plasmodesmata (Singular: plasmodesma), channels traversing the plant cell walls, allowing for the cell-to-cell transport of substances. Plasmodesmata are internally coated by the endoplasmic reticulum. Silencing suppressor (Suppressor of silencing): virally encoded proteins from diverse families that interfere with host production or effectiveness of small RNAs.

Introduction The members of the family Tombusviridae have uncapped, single-stranded, positive-sense RNA [( þ )ssRNA] genomes. The family Tombusviridae can be subdivided into three subfamilies: Calvusvirinae, Procedovirinae, and Regressovirinae. The subfamily Regressovirinae contains the genus Dianthovirus, the members of which have icosahedral virions and a genome consisted of two ( þ )ssRNA molecules. Although two genomic RNAs share viral and pro-viral host factors during infection, they exploit different strategies to regulate their translation, transcription, replication, packaging, and cell-to-cell movement.

Taxonomy and Classification Dianthoviruses belong to the family Tombusviridae and the subfamily Regressovirinae. Of the 16 genera in the family, only the dianthoviruses have bipartite genomes. The genus Dianthovirus is composed of three viruses: Carnation ringspot virus (CRSV), Red clover necrotic mosaic virus (RCNMV), and Sweet clover necrotic mosaic virus (SCNMV) (Table 1). Furcraea necrotic streak virus, which was provisionally classified as a member of the genus Dianthovirus, is now classified as a member of the genus Macanavirus in the family Tombusviridae. The amino acid (aa) sequence of dianthovirus coat protein (CP) has high homology with those of the tombusviruses and aureusviruses that belong to the family Tombusviridae. Interestingly, the aa sequence of dianthovirus RNA-dependent RNA polymerase (RdRP) has higher homology with that of the luteoviruses that belong to the family Luteoviridae, than with those of the viruses in the family Tombusviridae.

Genome Organization The genome of dianthoviruses consists of two single-stranded positive-sense RNA {( þ )RNA} molecules, RNA1 (3.9 kb) and RNA2 (1.4 kb) (Fig. 1). RNA1 encodes two replication-associated proteins: an auxiliary 27 kDa protein (p27), an N-terminally overlapping 88 kDa RdRP (p88pol); and the 37 kDa CP, which is the building block of the mature virion. A non-coding viral RNA, called SR1f, accumulates during dianthovirus infection. SR1f is generated from the 30 -untranslated region (UTR) of RNA1 through the action of a host 50 -30 exoribonuclease. The 30 UTR of dianthovirus RNA1 contains a sequence that inhibits 50 -30 exoribonucleolytic decay in cis. SR1f may suppress translation from RCNMV RNA1 and thus could play a regulatory role in infection by dianthoviruses. RNA2 encodes a 35 kDa movement protein (MP), which plays an essential role in viral cell-to-cell movement and systemic spread in plants.

Virions, Transmission, and Epidemiology Similarly, to the viruses in other genera, members of the genus Dianthovirus have icosahedral virions of approximately 35 nm in diameter with T ¼ 3 symmetry that consist of 180 CP subunits of approximately 36–37 kDa. RCNMV virions consist of two

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Table 1

Dianthovirus (Tombusviridae) List of members of the genus Dianthovirus, the length of their genomes, and their accession numbers

Name

Abbreviation

Isolate

Genome segment length (nt)

Accession no.

Carnation ringspot virus

CRSV

a

RNA1: RNA2: RNA1: RNA2: RNA1: RNA2: RNA1: RNA2: RNA1: RNA2: RNA1: RNA2: RNA1: RNA2: RNA2:

NC_003530 NC_003531 LC461179 LC461180 LC461178 LC461177 LC460998 LC461176 NC_003756 NC_003775 AB034916 AB034917 NC_003806 NC_003807 S46027

1.30 PV-21 PV-0097 Red clover necrotic mosaic virus

RCNMV

Australiaa Canada

Sweet clover necrotic mosaic virus

SCNMV

59a 38

3840 1403 3843 1404 3872 1405 3851 1406 3890 1448 3890 1456 3876 1449 1446

a

Example isolate.

Fig. 1 A schematic representation of the dianthovirus genome structure. The genome of dianthovirus consists of RNA1 and RNA2, which lack a 50 -cap structure and a 30 -poly(A) tail. RNA1 encodes an auxiliary replication protein p27 and an RNA-dependent RNA polymerase (RdRp) p88pol which is translated via a –1 ribosomal frameshift mechanism. RNA1 also encodes a coat protein (CP), which is translated from sub-genomic RNA (CPsgRNA). SR1f, a small noncoding RNA, is generated from the 30 -untranslated region (UTR) of RNA1 by a host 50 -30 exoribonuclease. RNA2 encodes a movement protein (MP) that is required for viral cell-to-cell movement.

distinct types: one contains one copy of each genomic RNA and the other contains four copies of RNA2. The origin of assembly sequence (OAS) is located in the MP coding region of RNA2. The stem-loop (SL) structure formed from the OAS is also known as the transactivator (TA), and is a multifunctional region that is required for the transcription of sub-genomic RNA and the replication of RNA2 (see the sections below that describe gene expression and RNA replication). RNA1 does not have an independent packaging signal and is co-packaged with RNA2 through an interaction with the SL structure in RNA2. The N-terminal 50 aa of RCNMV CP are rich in lysine residues and are essential for virion formation. RCNMV virions are stable in conditions in which divalent cations such as calcium and magnesium ions are abundant. In contrast, the virions are unstable when divalent cations are depleted. Low-divalent-cation environments such as those found within the cytoplasm of a cell might be favorable for uncoating the virion and exposing the genomic RNAs for gene expression. The unique response of RCNMV virions to changes in divalent cation concentrations has been exploited for the development of plant viral nanoparticles for controlled delivery of the anticancer drug doxorubicin. In addition to medical applications, a study has shown that RCNMV nanoparticles improve the performance of abamectin, a potent nematicide that targets a broad spectrum of plant parasitic nematodes, by enhancing the poor mobility of abamectin in soil. Dianthoviruses multiply to high concentrations in infected plants and the virions are stable in vitro. The release of dianthoviruses from infected roots into surrounding environments such as soil and drainage water has been reported. However, it has not been determined whether soil-inhabiting organisms transmit dianthoviruses in the field. Olpidium spp., nematodes, and bees have been proposed as vectors of RCNMV, CRSV, and SCNMV, respectively.

Gene Expression After entry into host cells, the genomic RNA of ( þ )RNA viruses recruits host ribosomes to act as a template for translation. Like all viruses in the family Tombusviridae, the genomic RNAs of dianthoviruses lack both a 50 -cap structure and a 30 -poly(A) tail.

Dianthovirus (Tombusviridae)

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Thus, to produce their encoded proteins efficiently, dianthoviruses use cellular translational machinery in a non-canonical, cap-independent mechanism. RCNMV uses different modes for cap-independent translation of its bipartite genomic RNAs. The protein p27 is directly translated from RNA1 in a cap-independent manner. Two cis-acting RNA elements play essential roles in the translation of RNA1: a 30 -cap-independent translation enhancer termed 30 TE-DR1 and an adenine-rich sequence (ARS) in the 30 UTR. 30 TE-DR1 is essential for recruiting the eukaryotic initiation factors eIF4F/eIFiso4F and ribosomes to RNA1. ARS, located 60 nucleotides upstream from 30 TE-DR1, recruits a host poly(A)-binding protein (PABP) to RNA1 through a direct RNA–protein interaction. The ARS–PABP interaction mediates the recruitment of eIF4F/eIFiso4F to 30 TE-DR1. Although 30 TE-DR1 can bind to both eIF4F and eIFiso4F, RNA1 preferentially utilizes eIF4F for its translation. Because the cap-independent translation of RNA1 is scanning dependent, ribosomes should be recruited to the 5-end of the viral RNA. However, the mechanism by which the 30 UTR of RNA1 coordinates the recruitment of ribosomes to the 50 end remains elusive. The protein p88pol is translated via –1 programmed ribosomal frameshifting (–1PRF) from RNA1. Three cis-acting RNA elements required for –1PRF have been identified: a shifty heptanucleotide sequence GGAUUUU where the reading frame shifts, a highly structured bulged SL immediately downstream from the slippage site, and a small stable SL structure located between ARS and 30 TE-DR1. Intramolecular base-pairing between the bulge of the second element and the loop sequence of the third element mediates efficient –1PRF of RCNMV RNA1. This long-distance RNA–RNA interaction is likely to be conserved in all dianthoviruses. CP is translated from sub-genomic RNA (CPsgRNA). CPsgRNA is likely to be generated via a premature termination mechanism, where the viral replicase terminates internally during negative-strand RNA {(  )RNA} synthesis. The 30 -truncated (  )RNA is then used as a template for CPsgRNA synthesis. CPsgRNA transcription requires intermolecular base-pairing between RNA2 (TA element) and RNA1 (TA-binding site located immediately upstream of the transcription start site), which is likely to be important for premature termination of the viral replicase complex (VRC) during (  )RNA synthesis. CPsgRNA is identical to the 30 -terminal 1.5 kb of RNA1 and therefore possesses the same 30 TE-DR1, which is also essential for translation from CPsgRNA. MP is translated from RNA2. Unlike RNA1, RNA2 lacks cis-acting RNA elements such as 30 TE-DR1 and ARS. Instead, cap-independent translation of RNA2 seems to be coupled with RNA2 replication. That is, only the RNA2 that is generated de novo through replication seems to act as a template for translation. Interestingly, RNA2 preferentially utilizes eIFiso4F rather than eIF4F. The cis-acting RNA elements that recruit eIFiso4F and/or ribosomes to RNA2 have not yet been identified. A translation-silencer element has been identified in the basal stem structure of the 50 UTR (50 BS) of RNA2, and translation-enhancer activity is found in the 50 UTR other than 50 BS. Therefore, the 50 UTR may coordinate cap-independent and replication-coupled translation of RNA2. The molecular mechanisms underlying replication-coupled translation of RNA2 remain elusive.

RNA Replication Replication of RCNMV takes place at the cytoplasmic surface of the host endoplasmic reticulum (ER) membrane. At the early phase of replication, RCNMV replicates at punctate structures on cortical ER. Later, such punctate structures disappear, and instead, a large aggregated ER structure emerges at the perinuclear region. p27 plays a key role in the biogenesis of the viral replication compartments. An amphipathic a-helix of p27 is essential for targeting p27 to ER membranes and the assembly of the VRC (termed the 480 kDa replicase complex). The p27–p27 and the p27–p88pol interactions are prerequisite for the assembly of the 480 kDa replicase complex. The C-terminal half of p27 is responsible for the p27–p27 and the p27–p88pol interactions, whereas the nonoverlapping region of p88pol is responsible for the p27–p88pol interaction. After translation in the cytoplasm, the genomic RNA of ( þ )RNA viruses must be recruited to the viral replication compartment and used as a template for (  )RNA synthesis. To distinguish viral genomic RNAs from host-derived RNAs, viral replication proteins must have mechanisms to specifically recognize viral RNAs as replication templates. Cis-acting RNA elements that are important for replication have been identified in the 30 UTR of both genome segments. In RNA1, two SLs, termed SLDE and SLF, and the intervening sequence are likely to act as the core promoter for (  )RNA synthesis. The corresponding cis-acting RNA elements are also present in RNA2. Interestingly, RNA2 has additional RNA elements that are also crucial for (  )RNA synthesis: 50 BS in the 50 UTR, a TA element in the coding region of MP, and a Y-shaped RNA element (YRE) in the 30 UTR. Because RNA2 utilizes a replication protein supplied from RNA1, these additional cis-acting RNA elements probably contribute to recruiting viral replication proteins. Indeed, YRE is an essential cis-acting RNA element that is recognized by the p27 replication protein. The p27–YRE interaction is required for the recruitment of RNA2 to the sites of viral replication. In contrast, RNA1 has no RNA element that interacts with p27 and/or p88pol supplied in trans, and p88pol is only able to act in cis on RNA1 during replication. This cis-preferential requirement for p88pol in the replication of RNA1 may be explained by the finding that p88pol appears to bind to the 30 UTR of RNA1 in a translation-coupled manner. The translation-coupled binding of p88pol may enable rigorous template selection of RNA1 without special cis-acting elements such as YRE.

Host Factors in Dianthovirus RNA Replication Accumulating evidence has suggested that interactions between viral proteins and host factors are important determinants of successful viral infection. Recent proteomics approaches followed by functional studies have suggested that RCNMV hijacks a number of host proteins for its robust replication.

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Heat shock protein 70 (Hsp70) and Hsp90 are abundant and highly conserved molecular chaperones that play central roles in protein homeostasis, including the folding of newly synthesized proteins and the assembly of macromolecular protein complexes. Like many other ( þ )RNA viruses, RCNMV requires these molecular chaperones for replication. p27 binds to Hsp70 and Hsp90 in vitro and in vivo. Hsp70 regulates the assembly of the 480 kDa viral replicase complex by preventing its aggregation, while Hsp90 promotes the p27–YRE interaction. Thus, RCNMV utilizes Hsp70 and Hsp90 to regulate different steps in RNA replication before starting (  )RNA synthesis. One of the hallmark features of eukaryotic ( þ )RNA virus replication is the establishment of viral replication compartments, which are generated de novo during virus infection via massive rearrangement of host membranes and lipid metabolism. A host small GTPase ADP ribosylation factor 1 (Arf1) is a central regulator of the biogenesis of the COPI vesicle that mediates Golgi-to-ER retrograde vesicle trafficking. Arf1 is critical for RCNMV replication. p27 directly interacts with and recruits Arf1 from the Golgi to the viral replication compartment; in fact, RCNMV seems to hijack Arf1 function to establish the viral replication compartment. Another important host factor for RCNMV replication is phospholipase D (PLD). PLD mediates the biogenesis of phosphatidic acid (PA) through hydrolysis of phosphatidylethanolamine or phosphatidylcholine. The interaction of p27 with PA was demonstrated by a protein–lipid overlay assay, and RCNMV was shown to induce an approximately threefold accumulation of PA in infected plant leaves. p88pol interacts with and recruits PLDa and PLDb of Nicotiana benthamiana to the viral replication compartment during RCNMV replication. PLD-derived PA is required for RCNMV replication, and exogenously added PA was shown to stimulate RNA replication in protoplasts and (  )RNA synthesis in plant-derived cell-free extracts. Therefore, RCNMV is likely to induce enrichment of PA at the viral replication compartment via PLD to establish a suitable environment for viral RNA replication. It is known that many plant viruses induce production of reactive oxygen species (ROS) during infection. ROS are often associated with host defense against biotrophic pathogens such as fungi and bacteria. However, the role of ROS in plant–virus interactions has been obscure. RCNMV strongly induces intracellular ROS production in infected cells. Respiratory burst oxidase homolog (RBOH) is one of the major host-plant enzymes responsible for the production of ROS in response to various types of stresses. Interestingly, RCNMV replication protein p27 interacts with the RBOHB of N. benthamiana and recruits it to the viral replication compartment. In addition, p27 is sufficient for induction of ROS production in an RBOH-dependent manner and gene silencing or pharmacological inhibition of RBOH activity compromises viral replication. These findings suggest that RCNMV hijacks a host ROS-generating enzyme to boost viral replication. RCNMV appears to activate RBOH to mediate virus-induced intracellular ROS production. A subgroup II calcium-dependent protein kinase (CDPKiso2) of N. benthamiana phosphorylates the N-terminal region of RBOHB and activates its enzyme activity. CDPKiso2 is required for p27-triggered ROS production and viral RNA replication. Although p27 can form a complex with CDPKiso2, this interaction seems to be indirect. Instead, a host scaffold protein, receptor for activated C kinase 1 (RACK1), bridges the interaction of p27 with CDPKiso2. Indeed, RACK1 is required for the p27–CDPKiso2 interaction, p27-mediated ROS production, and viral RNA replication. Together, these findings suggest that RCNMV co-opts RACK1 to integrate CDPKiso2 into the viral replication compartment to promote RBOH activation and ROS production.

Cell-to-Cell and Systemic Movement A 35 kDa MP encoded by RNA2 is essential for the cell-to-cell and systemic movement of dianthoviruses. The amino acid sequences of the MPs of dianthoviruses suggest that they belong to the 30K superfamily. RCNMV MP has the ability to bind single-stranded nucleic acids cooperatively. However, unlike the MP of tobacco mosaic virus (TMV), which also belongs to the 30K superfamily, RCNMV MP does not unfold single-stranded RNA molecules in vitro. Despite having such different characteristics, the MPs of TMV and RCNMV are functionally homologous because RCNMV MP can support the systemic infection of a movement-deficient mutant of TMV in N. benthamiana. Although it has not been determined whether dianthoviruses move cell-to-cell as virions or as RNA–MP complexes, the former is unlikely because the CP is dispensable for the cell-to-cell movement of dianthoviruses in several host plants. RCNMV MP localizes at plasmodesmata (PD) and increases their size exclusion limit. In addition, MP also localizes to small punctate structures at the cortical ER that contain the VRCs during the early phase of viral infection. Colocalization of MP to the cortical VRCs is associated with the replication of RNA1, but not that of RNA2, and is required for the cell-to-cell movement of RCNMV. Glyceraldehyde 3-phosphate dehydrogenase A is involved in this colocalization event and is required for the efficient cell-to-cell movement of RCNMV in N. benthamiana. Although a CP-deficient mutant of RCNMV is able to systemically infect N. benthamiana at lower temperatures such as 171C, virion formation is required but not sufficient for the systemic movement of dianthoviruses.

RNA Silencing Suppression The importance of RNA silencing mediated by small RNAs in the defense of various organisms, including plants, against viruses has been well established. To counteract antiviral RNA silencing, most plant viruses encode at least one viral silencing suppressor protein (VSR). RCNMV has at least two strategies to counteract host RNA silencing. Suppression of RNA silencing by RCNMV

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closely correlates with viral (  )RNA synthesis and accumulation of the 480 kDa replicase complex, implying that RCNMV interferes with host RNA silencing in an RNA replication-associated manner. RCNMV replication also inhibits microRNA biogenesis that is catalyzed by the host RNaseIII-like enzyme dicer-like 1 (DCL1). Higher viral accumulation was observed during RCNMV infection in a dcl1 mutant Arabidopsis. However, the role of DCL1 or its homologs in suppression of RNA silencing during RCNMV replication remains unknown. In addition to replication-associated RNA-silencing suppression, it has been reported that RCNMV MP has VSR activity. The functional domains required for VSR activity in MP differ from those required for cell-to-cell movement.

Further Reading Cao, J., Guenther, R.H., Sit, T.L., et al., 2015. Development of abamectin loaded plant virus nanoparticles for efficacious plant parasitic nematode control. ACS Applied Materials & Interfaces 7, 9546–9553. Cao, J., Guenther, R.H., Sit, T.L., et al., 2014. Loading and release mechanism of red clover necrotic mosaic virus derived plant viral nanoparticles for drug delivery of doxorubicin. Small 10, 5126–5136. Heinlein, M., 2015. Plant virus replication and movement. Virology 479–480, 657–671. Hiruki, C., 1987. The dianthoviruses: A distinct group of isometric plant viruses with bipartite genome. Advances in Virus Research 33, 257–300. Hyodo, K., Hashimoto, K., Kuchitsu, K., Suzuki, N., Okuno, T., 2017. Harnessing host ROS-generating machinery for the robust genome replication of a plant RNA virus. Proceedings of the National Academy of Sciences of the United States of America 114, E1282–E1290. Hyodo, K., Nagai, H., Okuno, T., 2017. Dual function of a cis-acting RNA element that acts as a replication enhancer and a translation repressor in a plant positive-stranded RNA virus. Virology 512, 74–82. Hyodo, K., Suzuki, N., Okuno, T., 2019. Hijacking a host scaffold protein, RACK1, for replication of a plant RNA virus. New Phytologist 221, 935–945. Hyodo, K., Taniguchi, T., Manabe, Y., et al., 2015. Phosphatidic acid produced by phospholipase D promotes RNA replication of a plant RNA virus. PLoS Pathogens 11, e1004909. Kaido, M., Abe, K., Mine, A., et al., 2014. GAPDH – A recruits a plant virus movement protein to cortical virus replication complexes to facilitate viral cell-to-cell movement. PLoS Pathogens 10, e1004505. Kaido, M., Nagano, H., Omote, K., et al., 2019. 50 – Terminal stem-loop of carnation ringspot virus RNA1 is required for the efficient amplification of viral RNAs. Virus Research 265, 138–142. Okuno, T., Hiruki, C., 2013. Molecular biology and epidemiology of dianthoviruses. Advances in Virus Research 87, 37–74. Steckelberg, A.L., Akiyama, B.M., Costantino, D.A., et al., 2018. A folded viral noncoding RNA blocks host cell exoribonucleases through a conformationally dynamic RNA structure. Proceedings of the National Academy of Sciences of the United States of America 115, 6404–6409. Tajima, Y., Iwakawa, H.O., Hyodo, K., et al., 2017. Requirement for eukaryotic translation initiation factors in cap-independent translation differs between bipartite genomic RNAs of red clover necrotic mosaic virus. Virology 509, 152–158. Wang, A., 2015. Dissecting the molecular network of virus-plant interactions: The complex roles of host factors. Annual Review of Phytopathology 53, 45–66.

Endornaviruses (Endornaviridae) Toshiyuki Fukuhara, Tokyo University of Agriculture and Technology, Fuchu, Japan r 2021 Elsevier Ltd. All rights reserved. This is an update of T. Fukuhara, H. Moriyama, Endornavirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00580-X.

Glossary Cytoplasmic male sterility Male sterility caused by a cytoplasmic factor. Endophyte A bacterium or fungus that lives within a plant for at least part of its life cycle without causing any apparent disease. Horizontal transmission The transmission of infections (e.g., viruses) between members of the same species. in silico An expression meaning “performed on computer” in reference to biological experiments. Isogenic line Genetically identical pure-breeding group of individuals. Mycorrhiza A symbiotic association between a green plant and a fungus. Mycovirus A virus that infects fungal species. Oomycete A fungus-like eukaryotic microorganism that is a filamentous, microscopic, and absorptive organism.

Open reading frame The part of a reading frame that has the ability to be translated into a protein. Phytopathogen An organism that is pathogenic to plants. Polyprotein A protein (produced by viruses) that is cleaved to produce a number of functional proteins. Positive-sense, single-stranded RNA virus A virus that uses positive sense, single-stranded RNA as its genome. RNA silencing The mechanism of transcriptional and post-transcriptional gene silencing mediated by 21–24 nt small RNAs. RNA-dependent RNA polymerase An enzyme that synthesizes RNA from an RNA template. Vertical transmission The transmission of parasites (e.g., virus) or symbionts from parents to children.

Introduction Viruses are thought to affect the phenotype of their hosts, and plants infected with viruses usually exhibit visible disease symptoms. Thus, infection of crop plants and vegetables with viruses severely damages their yield and quality. Therefore, commercially available crops, vegetables, and fruits are believed to be virus-free, but they are often asymptomatically infected with persistent viruses. As these plants are infected with persistent viruses and exhibit no symptoms, they are sold commercially. Double-stranded RNAs (dsRNAs) have frequently been detected from various healthy plants. These dsRNAs are not transcribed from the host genome DNAs, and replicate (propagate) independent of their host genome DNAs as bacterial plasmids do. These dsRNA replicons have common properties that differ from those of conventional RNA viruses: (1) Most of these dsRNAs have no obvious effect on the phenotype of their host plants; (2) they are distributed throughout all tissues of the host plants at a stable low concentration; (3) they are transmitted to the next generation only through the gametes; (4) their horizontal transfer to other plants has never been proven. The size of these dsRNAs varies from 1.5 to 20 kbp. Smaller dsRNAs (about 2.0 kbp) are often found with virus-like particles, and some of these dsRNAs have been classified as viruses in the family Partitiviridae. Partitiviruses have two unrelated linear dsRNA segments, each about 2.0 kbp in size, which encode a viral RNA-dependent RNA polymerase (replicase, RdRp) and a capsid (coat) protein (CP). These partitiviruses are typical persistent viruses that asymptomatically infect crop plants and vegetables. Larger dsRNAs (more than 10 kbp) are probably not associated with distinct virus-like particles, because no virus-like particles have been isolated in preparations obtained with various purification procedures. Thus, these large dsRNAs were previously referred to as RNA plasmids, enigmatic dsRNAs or endogenous dsRNAs. However, now these large dsRNAs are also recognized as persistent viruses, and named endornaviruses in the genus Alphaendornavirus of the family Endornaviridae. Acute virus infection severely damages the yield and quality of crop plants and vegetables, so phytopathologic studies aimed at protecting crop plants and vegetables from virus infections have been conducted, and many acute plant viruses have been extensively studied. In contrast, because persistent viruses asymptomatically infect host plants, studies on persistent plant viruses have been more limited than those on acute plant viruses. A typical persistent RNA viruses, endornaviruses, are the focus of this article.

Plant Endornaviruses Linear dsRNAs of approximately 14 kbp in length were often found in rice (Oryza sativa), barley (Hordeum vulgare), melon (Cucumis melo), and bell pepper (Capsicum annuum). These dsRNAs were not transcribed from their host DNA genomes. Furthermore they propagate independent of their host genomes but have several common properties that are different from those of conventional viruses. Complete nucleotide sequences of these dsRNAs isolated from broad bean (Vicia faba), cultivated rice (O. sativa) and wild rice

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Fig. 1 Genome organizations of Oryza sativa endornavirus (OsEV), the type species of Alphaendornavirus, and Sclerotinia sclerotiorum endornavirus 1 (SsEV1), the type species of Betaendornavirus. The site-specific nick is only present in the coding strand of alphaendornaviruses.

(O. rufipogon) were reported about two decades ago. These dsRNAs encoded a single open reading frame (ORF) of approximately 4600 amino acid (aa) residues, in which the conserved viral RdRp and RNA helicase (Hel) domains were found (Fig. 1). Phylogenetic analyses of these two conserved domains indicated that these dsRNAs share a common ancestor with the alpha-like supergroup of positive-sense single-stranded RNA (ssRNA) viruses including many plant ssRNA viruses, such as Tobacco mosaic virus (TMV) and Cucumber mosaic virus (CMV). Consequently, the International Committee on Taxonomy of Viruses (ICTV) classified these dsRNAs as members of a new virus genus Endornavirus within a new virus family Endornaviridae, and named the three dsRNAs as Vicia faba endornavirus (VfEV), Oryza sativa endornavirus (OsEV) and Oryza rufipogon endornavirus (OrEV). Deep-sequencing (RNA-seq) analyses have revealed asymptomatic infections of various viruses in crop plants and vegetables as well as wild plants. Some of these virus sequences have been recognized as members of the family Endornaviridae. To date, 14 endornaviruses have been found in plants, including barley, melon, bell and hot peppers (C. annuum), bottle gourd (Lagenaria siceraria), common bean (Phaseolus vulgaris), avocado (Persea americana) and others (Table 1). The eighth and ninth reports of the ICTV classified the family Endornaviridae in the dsRNA virus group. However, an alternative opinion based on phylogenetic analyses claims that the family Endornaviridae should be classified in the positive-sense ssRNA virus group, and large dsRNAs of endornaviruses should be considered as their replication intermediates. Consequently, the tenth report of the ICTV, published recently, classifies the family Endornaviridae in the positive-sense ssRNA virus group.

Fungal Endornaviruses A wide variety of viruses have been found in many fungi, including phytopathogenic and endophytic fungi and mushrooms. Fungal viruses are termed mycoviruses, and most mycoviruses contain dsRNAs as their genome. dsRNA mycoviruses are primarily classified into four families, Partitiviridae, Totiviridae, Chrysoviridae, and Endornaviridae, but novel mycoviruses recently discovered should be classified into a new genus and/or family. In general, mycoviruses persistently infect the host fungus and are transmitted vertically via host cell division, and infections are usually symptomless. The violet root rot fungus, Helicobasidium mompa, occurs on various plants, and contains various sized dsRNAs. A large dsRNA (L1 dsRNA) in the V670 strain of H. mompa has been identified as a hypovirulence factor. The nucleotide sequence of this dsRNA encodes a single long ORF containing conserved RdRp and Hel domains exhibiting significant similarity to those encoded by plant endornaviruses described above, indicating that this dsRNA is a member of the family Endornaviridae. The virus was subsequently named Helicobasidium mompa endornavirus 1 (HmEV1). An increasing number of recent reports have described nucleotide sequences that likely belong to viruses of the family Endornaviridae, especially from plant pathogenic fungi. Currently, the family Endornaviridae consists of two genera, Alphaendornavirus and Betaendornavirus, which are classified based on genome size and the presence of unique protein domains. Seven fungal endornaviruses, which have shorter genomes (o12.6 kbp) than alphaendornaviruses and lack the UDP (uridine diphosphate) glucosyltransferase (UGT) domain present in the majority of alphaendornaviruses, have been classified as members of the genus Betaendornavirus (Fig. 1). To date, the genus Alphaendornavirus includes 20 viruses that infect plants, fungi, and oomycetes, whereas the genus Betaendornavirus includes seven viruses that infect ascomycete fungi (Table 1, Fig. 2). Eight novel endornavirus-like sequences were recently identified from isolates of mycorrhizal (Ceratobasidium) fungi isolated from pelotons within root cortical cells of wild indigenous orchid species in Western Australia. The partial and complete sequences of viral RdRps shared low (9%–30%) identities with one another and with those of endornavirus RdRp described previously. Four putative viruses had longer genomes (420 kbp) than those of endornaviruses, and three had two ORFs within their genomes.

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List of species in the family Endornaviridae

Host type

Host species

Species name

Abbreviation

Malabar spinach Bell pepper Melon Barley Hot pepper Bottle gourde Wild rice (W1714) Cultivated rice Avocado Common bean Common bean Common bean Broad bean (447) Winged bean Mate

Basella alba endornavirus 1 Bell pepper endornavirus Cucumis melo endornavirus Hordeum vulgare endornavirus Hot pepper endornavirus Lagenaria siceraria endornavirus Oryza rufipogon endornavirus Oryza sativa endornavirusa Persea americana endornavirus 1 Phaseolus vulgaris endornavirus 1 Phaseolus vulgaris endornavirus 2 Phaseolus vulgaris endornavirus 3 Vicia faba endornavirus Winged bean endornavirus Yerba mate endornavirus

BaEV1 BPEV CmEV HvEV HPEV LsEV OrEV OsEV PaEV1 PvEV1 PvEV2 PvEV3 VfEV WBEV1 YmEV

Fungus

Powdery mildew fungus Endophyte White root rot fungus Rhizoctonia

Erysiphe cichoracearum endornavirus Grapevine endophyte endornavirus Helicobasidium mompa endornavirus 1 Rhizoctonia cerealis endornavirus 1

EcEV GEEV HmEV1 RcEV1

Protista

Phytophthora

Phytophthora endornavirus 1

PEV1

Alternaria Gray mold fungus Gremmeniella Soil-borne fungus Sclerotinia blight fungus White mold fungus Truffle

Alternaria brassicola endornavirus 1 Botrytis cinerea endornavirus 1 Gremmeniella abietina endornavirus 1 Rosellinia necatrix endornavirus 1 Sclerotinia minor endornavirus 1 Sclerotinia sclerotiorum endornavirus 1a Tuber aestivum endornavirus

AbEV1 BcEV1 GaEV1 RnEV1 SmEV1 SsEV1 TaEV

Alphaendornavirus Plant

Betaendornavirus Fungus

a

The type species.

These two features of novel endorna-like viruses are unlike endornaviruses currently described, and they may challenge current taxonomic criteria for membership of the family Endornaviridae.

Endornaviruses From Other Eukaryotic Kingdoms Plant pathogens of the genus Phytophthora have many of the same biological properties as fungi. However, on the basis of sequence similarities, plant pathogens of the genus Phytophthora, an oomycete, have been classified together with diatoms and brown algae into a protist group known as the Stramenopiles. Sequencing and phylogenetic analyses of a large dsRNA from a Phytophthora isolate from Douglas fir (Pseudotsuga menziesii) revealed that it encodes a single long ORF similar to those encoded by known plant and fungal endornaviruses. This was recognized as a member of the genus Alphaendornavirus and named Phytophthora endornavirus 1 (PEV1). Using an in silico bioinformatics approach to obtain full or partial cDNA sequences of genes and comparing them against known viral sequences in the NCBI (National Center for Biotechnology Information) Expressed Sequence Tag database, 119 novel virus-like sequences related to members of the families Partitiviridae, Totiviridae, Chrysoviridae, and Endornaviridae, were discovered. Among them one endornavirus-like sequence that exhibited significant similarity to the fungal endornavirus HmEV1 was identified from the sea louse (Caligus rogercresseyi). As the sea louse is a member of the phylum Arthropoda, this finding suggests that endornaviruses are also found in the animal kingdom.

Genome Organization A recent ICTV report claimed, based on phylogenetic analyses, that endornaviruses are classified in the positive-sense ssRNA virus group, and that large dsRNAs should be recognized as their replication intermediates. However, ssRNA genomes of endornaviruses have never been isolated. Consequently, all cDNA cloning experiments were performed by using purified dsRNAs of

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Fig. 2 Phylogenetic tree of selected endornaviruses, as modified from a tree constructed by Valverde et al. using the aa sequences of RdRp domains of endornaviruses. Abbreviations of virus names are indicated in Table 1. Reproduced from Valverde, R.A., Khalifa, M.E., Okada, R., et al., 2019. ICTV virus taxonomy profile: Endornaviridae. Journal of General Virology 100 (8), 1204–1205.

endornaviruses as templates for cDNA synthesis. In this case, most endornavirus sequences were obtained from nucleotide sequences of cDNA clones. Recently, nucleotide sequences of endornaviruses have been obtained more easily from RNA-seq data of endornavirus-infected organisms. The endornavirus genomes sequenced to data range from 9639 to 17,635 bp in length, and electron microscopic observations revealed that the dsRNAs of endornaviruses are linear (Fig. 3). They commonly encode a single ORF of 3148 to 5821 aa residues, and this long ORF is one of the unique molecular features of endornaviruses (Fig. 1). Conserved Hel and RdRp domains located in the central and C-terminal regions, respectively, of the long ORF are common features of all endornaviruses described to date (Fig. 1). Other conserved domains found in some, but not all, endornaviruses are viral methyltransferase (MTR), UGT, capsular polysaccharide synthetase (CPS), and cysteine-rich region (CRR) (Fig. 1). Four plant endornaviruses [OsEV, OrEV, VfEV and Bell pepper endornavirus (BPEV)], one fungal endornavirus (HmEV1), and one oomycete endornavirus (PEV1) belonging to the genus Alphaendornavirus were shown to contain a site-specific nick in the 50 region of the coding strand, which divides not only the coding strand but also the single long ORF (Fig. 1), but the nick has never been found in betaendornaviruses. The biological implications of this nick in the coding strand and the mechanism by which it is generated are unknown. However, because the divided coding strand can no longer be used as a template for non-coding strand synthesis (replication) or mRNA for translation of the protein, the nick must affect at least two important steps in the life cycle of endornaviruses. Therefore, endornavirus dsRNAs containing the nick may be remains (dead virus bodies) because they have a defect in translation and replication, even though these unique dsRNA molecules predominantly accumulate in the hosts infected by some alphaendornaviruses described above. The continuous ssRNA of the coding strand may play a vital role in the virus life cycle, but

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Fig. 3 Electron micrograph of dsRNA molecules of Oryza sativa endornavirus (OsEV). Bar represents 1 mm. Figure reproduced with permission from Fukuhara, T., Moriyama, H., Pak, J.K., Hyakutake, T., Nitta, T., 1993. Enigmatic double-stranded RNA in Japonica rice. Plant Molecular Biology 21, 1121–1130.

there are no reports describing the isolation of ssRNA molecules of endornaviruses. These two molecular features – the single long ORF and the site-specific nick – have not been reported in other known RNA viruses.

Gene Products All endornaviruses reported to date encode a single long ORF of approximately 3100 to 5800 aa residues, in which several conserved protein domains are found. This suggests that this long ORF encodes a polyprotein that is cleaved to produce a number of functional proteins. Indeed, potyviruses, which have a single long ORF of approximately 3000 aa residues, encode a single polyprotein consisting of eight functional proteins (see Potato Virus Y, Potyvirus). Therefore, endornaviruses likely encode a polyprotein containing more than 10 functional proteins. Indeed, several conserved protein domains have been found in all endornaviruses (Fig. 1), as discussed below. However, these putative proteins have never been experimentally detected by a specific antibody using a technique such as Western blot. Conserved motifs encoding RdRp and Hel are found in the same regions of long ORFs encoded by most endornaviruses (Fig. 1), and these two proteins are structurally similar to those encoded by ssRNA viruses such as CMV and TMV. Thus, these putative RdRp and Hel probably function in the replication of the genome RNA, as their homologues encoded by ssRNA viruses are essential for the replication of their genomes. MTR (a capping enzyme for viral genomic RNAs) also probably functions in the replication of endornaviruses, similar to their homologues encoded by ssRNA viruses, but the conserved MTR motif is not found in some endornaviruses. The enzyme UGT is commonly found in eukaryotes. This enzyme catalyzes the addition of the glycosyl group from a UDP-sugar to a small hydrophobic molecule. UGTs have been identified in dsDNA viruses, including baculoviruses and nucleopolyhedroviruses from insects. In insect cells, the viral ecdysteroid UDP-glycosyltransferase inactivates the molting hormones (ecdysteroids) of the host insect by sugar conjugation. Although a UGT gene has never been found in the genomes of the alpha-like supergroup of ssRNA viruses, it was found in strains of two hypoviruses (fungal ssRNA viruses), Cryphonectria hypovirus 3 and 4 (CHV3 and CHV4). A conserved motif significantly similar to that of UGT was found in some alphaendornaviruses (Fig. 1). A conserved motif similar to that of CPS is also found in some endornaviruses (Fig. 1). However, the functions of both putative UGT and CPS proteins in endornaviruses remain unknown. The ORF of endornaviruses is very large (Fig. 1), so it likely encodes several other proteins in addition to those already discussed. The ORF must encode a polyprotein, so it must also encode one or more proteinases to cleave the putative polyprotein into functional units such as RdRp, Hel, MTR, UGT and CPS. A similar situation is well known in potyviruses, which encode a polyprotein of approximately 3000 aa residues containing two proteinases (see Potato Virus Y, Potyvirus). The conserved CRR located in the N-terminal half of endornavirus ORFs may function as a proteinase (Fig. 1), because many proteinases (also known as cysteine proteinases) containing a conserved cysteine residue in their catalytic domain have been reported so far. CP and movement proteins (MP) are essential for conventional plant viruses such as TMV and CMV. CP is essential for viral horizontal transmission from one host plant to another, and MP is essential for the movement of viruses from one cell to surrounding (neighboring) cells and systemic infection in host plants. Systemic propagation (infection) within the host plant mediated by MP and horizontal transmission (infection) from one host to other hosts mediated by CP constitute a fundamental propagation strategy for acute plant viruses. In contrast, endornaviruses have a symbiotic relationship with the host plant. Therefore, both CP and MP are probably not necessary for persistent propagation of endornaviruses.

Replication (RdRp Activity) A limited number of reports have demonstrated the functions (or enzymatic activities) of proteins encoded by endornaviruses. The RdRp activity of endornaviruses was initially reported from a study of VfEV in the “44700 line of broad bean (V. faba).

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Purified cytoplasmic membranous vesicles containing VfEV had RdRp activity, which required Mg2 þ ions and the four nucleotide triphosphates, and was unaffected by inhibitors of cellular DNA-dependent RNA polymerases such as alphaamanitin and actinomycin D. RdRp activity was also detected in the crude microsomal fraction of OsEV-harboring rice cells. Like the VfEV RdRp activity, this RdRp activity was also highest in the presence of all four nucleotide triphosphates and Mg2 þ ions, and was resistant to inhibitors of DNA-dependent RNA polymerases. Treatment of the purified microsomal fraction with proteinase K plus deoxycholate suggested that the RdRp enzyme complex is located in vesicles. So far, RdRp activity has been found only in two endornavirus-harboring plants mentioned above. Therefore, the available data concerning endornavirus replication are limited. There is a need to identify and characterize the endornavirus- and/or host-encoded proteins that are involved in endornavirus replication.

Vertical Transmission Although horizontal transmission of endornaviruses has never been reported in plants, these viruses are frequently found in a variety of plants. For example, OsEV and BPEV are commonly detected in many cultivars of japonica rice and bell pepper, respectively. To understand why OsEV is widely distributed in japonica rice cultivars, crossing experiments with OsEV-harboring and OsEV-free plants were carried out to examine the efficiency of vertical transmission of OsEV. OsEV-harboring and OsEV-free plants were found to coexist in the japonica rice cultivar Nipponbare, and these two isogenic lines could not be distinguished on the basis of their appearance. The results of crossing experiments indicated that the efficiency of OsEV transmission via pollen was more than 98%, and that of transmission via ovules was 100%. Consequently endornaviruses are efficiently transmitted to progeny plants via both pollen and ovules, though differential centrifugation and sucrose density-gradient centrifugation experiments revealed that OsEV localizes in the cytoplasm of host cells. The high efficiency of OsEV transmission via both pollen and ovules is likely responsible for the wide distribution of OsEV in many rice cultivars. Analyses of the F2 progeny plants of cv. Nipponbare indicated that the absence of OsEV was not associated with any particular gene(s) in the OsEV-free plants. The observed inheritance of OsEV was different from that of other cytoplasmic genetic elements (e.g., chloroplasts and mitochondria), which are usually inherited only via egg cells. OsEV has been found in many cultivars of japonica rice but not in any cultivars of indica rice (a subspecies of O. sativa). Japonica and indica are distinguishable on the basis of their phenotype (e.g., grain shape) and genotype (e.g., the distribution of transposons in their genomes). To determine the reason why OsEV is not found in indica rice cultivars, reciprocal crosses between an OsEV-harboring japonica variety (cv. Nipponbare) and an OsEV-free indica variety (cv. IR26 or Kasalath) were performed. When cvs. IR26 and Nipponbare were used, efficient transmission of OsEV via ovule (93%) and pollen (89%) was observed. However, when cvs. Kasalath and Nipponbare were used, OsEV transmission efficiency to F1 progeny was 68% via ovule and 20% via pollen, and transmission to F2 progeny plants also followed a complicated, non-Mendelian inheritance pattern. These results suggest that OsEV is unstable in indica rice plants, which may lack one or more genes that are involved in the maintenance (replication) of OsEV. Similar crossing experiments with endornavirus-harboring bell-pepper plants (cv. Yolo Wonder) and endornavirus-free plants (cv. Jalapeño M or Hungarian Wax) demonstrated that BPEV is also inherited maternally and paternally to the next generation. Thus, the propagation of OsEV and BPEV (and probably other plant endornaviruses as well) may depend on seed-mediated transmission, and endornaviruses may survive in cooperation with their host plants. No horizontal spread of plant endornaviruses has been observed in the field, and no potential vectors have been identified.

Horizontal Transmission In general, fungal endornaviruses as well as other mycoviruses persistently infect the host fungus, and are transmitted vertically via host cell division. Recently, however, it has been reported that natural infections of a phytopathogenic soil-borne fungus, Rosellinia necatrix, with a novel endornavirus, Rosellinia necatrix endornavirus 1 (RnEV1), can occur under experimental soil conditions. Furthermore, it has also been demonstrated that Sclerotinia minor endornavirus 1 (SmEV1), which is found in a phytopathogenic fungus Sclerotinia minor, was readily transmitted horizontally via hyphal contact to isolates of different vegetative compatibility groups of S. minor, and transmitted vertically as well. Therefore, some fungal endornaviruses are able to transmit horizontally as well as vertically.

Regulation of Copy Number All plant endornaviruses reported so far are likely to be found in all tissues of host plants at a stable low concentration, suggesting that their propagation strategy is probably different from that of acute ssRNA viruses. However, studies on the copy number of endornaviruses are limited. OsEV in rice plants is the only report that studied the regulation of copy number of endornaviruses. A comparison of the relative amount of OsEV with genomic DNA in host rice plants of cv. Nipponbare showed that OsEV is maintained at approximately 100 copies per cell. No significant difference in OsEV concentration was found in seedlings, roots,

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and mature leaves, indicating that a mechanism to regulate copy number may exist in host rice cells. In contrast, the copy number of OsEV in cultured rice cells was found to be about 10-fold higher than in the cells of seedlings. Although no changes in the copy number of OsEV was found in suspension cultures over the course of three years, copy number returned to the original low level in rice plants regenerated from cultured cells. Furthermore, the copy number of OsEV in pollen grains was found to be 50-fold higher than in seedlings. The steady replication of plant endornaviruses before every host cell division (both mitosis and meiosis) and their efficient transmission to the next generation via ovules and pollen are essential for endornavirus propagation because horizontal transmission likely does not occur. The unregulated increase of endornavirus replication could cause disease or lead to the death of host plants, as occurs with acute virus infections. Conversely, a decrease in copy number could cause the virus to disappear from the host plant (germ cells). The mating of host plants provides an opportunity for endornavirus propagation, so an increase in copy number only in pollen grains is a reasonable strategy to ensure virus transmission, despite their cytoplasmic localization. Unlike almost all other viruses, endornaviruses are not able to transmit horizontally, but they are capable of vertical transmission.

Pathogenesis Endornavirus-harboring crop plants and vegetables are commercially available, indicating that these plants are healthy and common cultivars. Current data also indicate that most endornaviruses found in plants, with the exception of VfEV, cause asymptomatic and persistent infections in the host plant. For instance, BPEV and OsEV have been frequently found in commercially available bell pepper fruits and rice grains, respectively. Consequently we usually consume these fruits and grains that contain endornaviruses. In the model cultivar of cultivated rice, Nipponbare, molecular biological analyses identified OsEV-harboring and OsEV-free plants. However, these two isogenic lines did not exhibit distinguishable phenotypes, and even rice breeders and farmers were unable to distinguish them. If the OsEV-harboring line had a lower harvest yield than the OsEV-free line, breeders and/or farmers would recognize this phenotype and discard the lower-yielding line. In commercial cultivars of bell pepper grown in the United States, an endornavirus was found at a prevalence rate of 100%. The overall appearance of the two bell pepper near-isogenic lines, BPEV-harboring and -free, was similar. However, the BPEV-free line had a significantly higher percentage of seed germination than the BPEV-harboring line. Plant height, number of fruits, and total fruit weight in plants of the BPEV-free line were higher than in plants of the BPEV-harboring line, but these differences were not statistically significant. Other characteristics between the two lines, such as, stem diameter, percentage of dry weight, fruit volume, chlorophyll, carotenoid, and anthocyanin content, were similar. These results suggest that BPEV appears to have a weak parasitic relationship with the host. Recently, variations in physiological traits have been reported in eight lines of common bean (P. vulgaris) cv. Black Turtle Soup, four of which were double-infected with Phaseolus vulgaris endornavirus 1 (PvEV1) and 2 (PvEV2), and four of which were endornavirus-free. Plants from all eight lines were morphologically similar and did not show statistically significant differences in plant height, wet weight, number of days to flowering and pod formation, pods per plant, pod thickness, seed size, number of seeds per pod, and anthocyanin content. However, the endornavirus-infected lines had faster seed germination, longer radicles, lower chlorophyll content, higher carotene content, longer pods, and higher weight of 100 seeds, all of which were statistically significant. Over four decades ago, a cytoplasmic male sterility (CMS) trait in the “447” line of broad bean (V. faba) and cytoplasmic spherical bodies (CSBs) found in the cytoplasm of 447 plants were reported. These CSBs contained a large dsRNA of approximately 17.6 kbp in length, and a correlation between the CMS trait and the presence of a CSB with a large dsRNA in the “447” strain was reported. This dsRNA was the genome of VfEV, which is the only plant endornavirus that affects the visible phenotype of the host. In general, mycoviruses asymptomatically infect host fungi. However, HmEV1, which was first reported as a fungal endornavirus, was identified as the hypovirulence factor from the strain V670 of the violet root rot fungus, Helicobasidium mompa. Recently, the endornavirus, SmEV1, was isolated from the hypovirulent strain LC22 of S. minor, and hypovirulence and associated traits of strain LC22 and SmEV1 were readily co-transmitted horizontally via hyphal contact to isolates of different vegetative compatibility groups of S. minor. Therefore, SmEV1 is also a hypovirulence-associated mycovirus with a wide spectrum of transmissibility and has the potential for biological control (virocontrol) of diseases caused by S. minor. Therefore, most endornaviruses asymptomatically infect their hosts. However, in combinations between specific endornaviruses and their hosts, endornaviruses are pathogenic to hosts and their infection affects the host phenotype.

Virus-Host Relationship (Host RNA Silencing Against Endornaviruses) RNA silencing (RNA interference) is the process of sequence-specific post-transcriptional gene silencing triggered by dsRNAs. Most eukaryotes, such as fungi, insects, and plants, sense exogenous long dsRNAs, such as viruses, because replication intermediates of ssRNA viruses are long dsRNAs. They then activate the RNA silencing pathway for defense against virus infection. Extensive studies on the interactions between acute ssRNA viruses and host plants such as Arabidopsis thaliana revealed that RNA silencing functions as an innate defense system against virus infections in plants. In the RNA silencing pathway, a Dicer or Dicer-like (DCL) endoribonuclease cleaves long viral dsRNAs into viral small interfering RNAs (vsiRNAs), which then associate with an Argonaute

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(AGO) protein. The vsiRNA-loaded AGO proteins then slice the viral genomic ssRNAs. However, many acute ssRNA viruses encode one or several viral suppressors of RNA silencing (VSR), which inhibits the host RNA silencing system, thus enabling the virus to overcome the host defense system and propagate systemically in the host plant. Plant endornaviruses are likely maintained within the host plant for hundreds of generations, suggesting that the host plant regulates virus propagation and that some host factors control virus replication so that it is coordinated with host cell division. If host plants did not have such factors (proteins), those harboring endornaviruses would exhibit disease symptoms presumably due to unregulated virus propagation, or some of the plant’s somatic or germ cells would lose endornaviruses. Therefore, host plants likely express factors that control endornaviruses as symbiont-like parasites. Candidate host factors for regulating endornaviruses could be proteins that constitute the RNA silencing machinery. Indeed endornavirus-derived vsiRNAs have been detected in host plants infected with OsEV, BPEV, PvEV1 and PvEV2, indicating that the host RNA silencing machinery recognizes endornaviral dsRNAs. The copy number and inheritance of OsEV were examined in transgenic rice plants with a knock-down (KD) construct of genes for the RNA silencing machinery, OsDCLs or host RNA-dependent RNA polymerases. A low vertical transmission rate and the disappearance of OsEV during somatic cell divisions were observed in some OsDCL2-KD plants, suggesting that KD of a Dicer gene negatively affects the maintenance of OsEV in host plants. The host RNA silencing system may be necessary for persistent infection by endornaviruses because it provides stringent regulation of low copy number and efficient vertical transmission to the next generation. RNA silencing functions as a defense against acute virus infection, but it may also function as a host factor that maintains persistent infections of endornaviruses by keeping their copy number low.

Further Reading Boccardo, G., Lisa, V., Luisini, E., Milne, R.G., 1987. Cryptic plant viruses. Advances in Virus Research 32, 171–214. Brown, G.G., Finnegan, P.M., 1989. RNA plasmids. International Review of Cytology 117, 1–56. Dodds, J.A., Morris, T.J., Jordan, R.L., 1984. Plant viral double-stranded RNA. Annual Review of Phytopathology 22, 151–168. Fukuhara, T., 1999. Double-stranded RNA in rice. Journal of Plant Research 112, 131–138. Fukuhara, T., 2019. Endornaviruses: Persistent dsRNA viruses with symbiotic properties in diverse eukaryotes. Virus Genes 55, 165–173. Fukuhara, T., Gibbs, M.J., 2012. Family Endornaviridae. In: King, A.M.Q., Adams, M.J., Carstens, E.B., Lefkowitz, E.J. (Eds.), Virus Taxonomy: Ninth Report of the International Committee on Taxonomy of Viruses. San Diego: Elsevier, pp. 519–521. Roossinck, M.J., 2018. Evolutionary and ecological links between plant and fungal viruses. New Phytologist 221, 86–92. Roossinck, M.J., Sabanadzovic, S., Okada, R., Valverde, R.A., 2011. The remarkable evolutionary history of endornaviruses. Journal of General Virology 92, 2674–2678. Tavantzis, S.M., 2001. dsRNA Genetic Elements: Concepts and Applications in Agriculture, Forestry, and Medicine. Boca Raton: CRC Press. Valverde, R.A., Khalifa, M.E., Okada, R., et al., 2019. ICTV virus taxonomy profile: Endornaviridae. Journal of General Virology 100 (8), 1204–1205.

Fimoviruses (Fimoviridae) Toufic Elbeaino and Michele Digiaro, International Center for Advanced Mediterranean Agronomic Studies (CIHEAM), Mediterranean Agronomic Institute of Bari, Valenzano, Italy r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) AAP Acquisition access period CP Coat protein DMB Double-membraned body ELISA Enzyme-linked immunosorbent assay ER Endoplasmic reticulum GP Glycoprotein precursor HPD High Plains disease kb Kilobase kDa Kilo Dalton

MP Movement protein NGS Next generation sequencing nt Nucleotide(s) ORF Open reading frame PCR Polymerase chain reaction RdRP RNA-dependent RNA polymerase RNP RNA ribonucleoprotein RRD Rose rosette disease RT-PCR Reverse transcription polymerase chain reaction ssRNA Single-stranded RNA UTR Untranslated terminal regions

Glossary Double-membraned bodies

Double-membraned proteinaceous structures enveloping the virus particles.

Introduction Although they were associated with typical symptoms on their hosts, i.e., chlorotic ringspot and mosaic (Fig. 1), the nature and identity of fimoviruses remained unknown for a long time. This was mainly due to the difficulty of isolating the fimoviruses by mechanical transmission onto herbaceous hosts and to the abnormal conformation of virus particles observed under the electron microscope, since their proteinaceous structures, known as double-membraned bodies (DMBs), was different from all other plant viruses known at the time. Today, the family Fimoviridae, order Bunyavirales, includes only one genus (Emaravirus), which comprises nine species: European mountain ash ringspot-associated emaravirus (as a type species), Actinidia chlorotic ringspot-associated emaravirus, Fig mosaic emaravirus, Pigeonpea sterility mosaic emaravirus 1, Pigeonpea sterility mosaic emaravirus 2, Redbud yellow ringspot-associated emaravirus, Raspberry leaf blotch emaravirus, Rose rosette emaravirus and High Plains wheat mosaic emaravirus. It also includes several tentative emaravirus species: Actinidia emaravirus 2 (AcEV-2), Blackberry leaf mottle-associated virus (BLMaV), Camellia japonica associated emaravirus 1 (CjEV-1), Camellia japonica associated emaravirus 2 (CjEV-2), Jujube yellow mottle-associated virus (JYMaV), Lilac chlorotic ringspot-associated virus (LiCRaV), Palo verde broom virus (PVBV), Pistacia virus B (PiVB), Ti ringspot-associated virus (TiRSaV), Perilla mosaic virus (PerMV) and Woolly burdock yellow vein virus (WBYVV). All these viruses are characterized by a multipartite negative-sense ssRNA genome composed of four to eight segments (nine, in the case CjEV-1) contained within spherical virions. Most of these viruses are vectored in nature by eriophyid mites, which are known to have a highly specific relationship with their plant hosts, thus explaining the rather restricted host range that characterize them.

Classification Fimoviruses are distantly related to orthotospoviruses and orthobunyaviruses of the families Tospoviridae and Peribunyaviridae, respectively. They share with them the following common features: (1) a multipartite, negative-sense, ssRNA genome, which in the case of fimoviruses consists of four to eight segments; (2) a high sequence identity with orthologous proteins of members of the order Bunyavirales at equivalent genome positions in the first three RNAs (corresponding to L, M and S RNA segments); (3) the presence of five conserved motifs (A-E) in the amino acid sequence of their RNA-dependent RNA Polymerase (RdRP), similar to those in the L segment-encoded protein of members of the order Bunyavirales; (4) the virus particles are typically enveloped in membranes; (5) the presence of conserved nucleotide sequences at both the 50 - and 30 -termini of all genomic RNAs segments of fimoviruses, but not identical to those of other members of the order Bunyavirales.

Virion Structure Fimoviruses are approximately spherical and enveloped virions with a diameter of 60–200 nm (Fig. 2). DMBs have an envelope consisting of a unit membrane approximately 12 nm thick, which is apparently acquired from the endoplasmic reticulum, containing proteinaceous

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Fig. 1 Host symptoms caused by emaraviruses: FMV-infected plants showing chlorotic ringspots on leaves (a) and fruits (c), together with mosaic-like symptoms, i.e., vein clearing and leaf mottling (b). FMV-infected cyclamen plants showing flower discoloration (d), mosaic (e) and leaf deformation (f). PPSMV-1-infected plants showing ringspot (g), chlorosis and mosaic (h), yellowing and sterility of flowers (i). EMARaV-infected plants showing ringspots (j), mottling (k) and mosaic (l) symptoms.

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Fig. 2 Groups of double-membraned bodies (DMB) in the cytoplasm of cells from a fig seedling infected with FMV. Arrows point to the aggregates of convoluted electron-dense filamentous elements (DMBs). Bar ¼ 200 nm.

material and fine fibrils. In ultrathin sections they are frequently found in association with amorphous electron-dense material. DMBs are present in all cell types except those of the vascular bundles and are rarely observed in bundle sheath parenchyma cells.

Genome The virus genome comprises from four to eight segments of negative sense ssRNA (Table 1). Genomic RNAs are not capped or polyadenylated and all contain complementary sequences (13–20 nt, depending on the RNA segment) at their 50 - and 30 -termini conserved in all viral genomic RNA components. The lengths of 50 and 30 untranslated terminal regions (UTR) of each viral segment are very variable in the different species, ranging from 40 to 50 nt up to 600–700 nt in length. All emaraviruses have a similar genomic organization, and each RNA is composed of a single ORF in negative polarity (Fig. 3). (1) RNA-1 is 6981-7217 nt long and contains ORF1 that encodes a RdRP of a predicted molecular mass of 265–272 kDa (P1). In a delimited area of this ORF there are the five motifs also conserved in all bunyavirids, which correspond to the core polymerase module of the RdRP active site. (2) RNA-2 is 2054-2399 nt long and contains ORF2, encoding a glycoprotein precursor (GP) with a predicted molecular mass of 73–83 kDa (P2). P2 shares several conserved motifs, including one predicted cleavage site that may generate two larger glycoproteins, Gn (20–23 kDa) and Gc (52–53 kDa). A third smaller glycoprotein (Gs), consisting of 23 aa (2.6 kDa), has been predicted in the case of Actinidia chlorotic ringspot-associated virus (AcCRaV). (3) RNA-3 is 1220-1678 nt long and contains ORF3, encoding a nucleocapsid protein (NP) of 32–35 kDa (P3). In the case of High Plains wheat mosaic virus (HPWMoV), two RNA-3 consensus sequences encoding the NP were found, with 12.5% sequence divergence. This protein contains stretches of positively charged aa (BindN specificity, 80%) that are probably involved in RNA binding. (4) RNA-4 is 1342-1675 nt long (1154 nt in CjEV-2) and contains ORF4, encoding a putative movement protein (MP) of 38–43.6 kDa (P4), having structural elements resembling the consensus secondary structure of the 30K superfamily of plant virus MPs. In the case of European mountain ash ringspot-associated virus (EMARaV), the MP is encoded by the RNA-5 (# LR536382) and not by RNA-4 (# NC_013108). More uncertainty surrounds the exact number and functionality of the other genomic RNAs that complete the emaravirus genome. Unlike the first four RNAs examined so far, the sequence homologies and the lengths of orthologs RNA-5 to RNA-8 in the different emaraviruses are much less significant. A ninth RNA (RNA-9) has been to date detected in the only CjEV-1. An exception is the high sequence identity (93%) shown in the RNA-6 of Pigeonpea sterility mosaic viruses 1 and 2 (PPSMV-1 and PPSMV-2), thus suggesting that the RNA-6 encoded protein is performing a conserved function in virus biology for both PPSMV‐1 and 2. Instead of one ORF in all genomic RNAs of emaraviruses, unexpectedly, only RNA-6 of RRV has shown the presence of two different ORFs (7.4 kDa and 27.1 kDa). Studies on the genetic variability within the population of different emaraviruses [Fig mosaic virus (FMV), Raspberry leaf blotch virus (RLBV), HPWMoV, Redbud yellow ringspot-associated virus (RYRSaV), PPSMV-1 and PPSMV-2] have shown significant differences in nucleotide sequences among isolates of different origins, which in the case of RNA-3 (NP) never exceeded 15%. This variability was generally lower than that found in RNA-1 and RNA-2 of FMV, whereas RNA-4 (MP) was more conserved than RNA-3 in the case of PPSMV-1 and PPSMV-2.

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Table 1 List of members and tentative members of the Genus Emaravirus in the family Fimoviridae. Name, Acronym and Accession # of each genomic RNA is provided Species names virus names

Acronym Accession # RNA-1

Actinidia chlorotic ringspotassociated emaravirus Actinidia chlorotic ringspotassociated virus European mountain ash ringspotassociated emaravirus European mountain ash ringspotassociated virus Fig mosaic emaravirus Fig mosaic virus

AcCRaV

RNA-2

RNA-3

RNA-4

RNA-5

RNA-6

RNA-7

RNA-8

NC038769 NC038770 NC038772 NC038771 NC038773

EMARaV NC013105 NC013106 NC013108 LR536382a NC013107a LR536383

FMV

NC029562 NC029565 NC029563 NC029564 NC029566 NC029568

High Plains wheat mosaic emaravirus High Plains wheat mosaic virus HPWMoV NC029570 NC029549 NC029550 NC029551 NC029552 NC029553 NC029554 NC029555 Pigeonpea sterility mosaic emaravirus 1 Pigeonpea sterility mosaic virus 1

PPSMV-1 NC029575 NC029556 NC029557 NC029574 NC029569 KX363891

Pigeonpea sterility mosaic emaravirus 2 Pigeonpea sterility mosaic virus 2

PPSMV-2 NC030660 NC030662 NC030661 NC030663 NC030658 NC030659

Raspberry leaf blotch emaravirus Raspberry leaf blotch virus

RLBV

NC029567 NC029558 NC029559 NC029560 NC029561 NC029571 NC029572 NC029573

RYRSaV

NC038852 NC038856 NC038854 NC038853 NC038855

RRV

NC015298 NC015299 NC015300 NC015301 NC034979 NC034980 NC034981

AcEV-2 BLMaV

MK602171 MK602172 MK602173 MK602174 MK602175 MK602176 KY056657 KY056658 KY056659 KY056660 KY056661

CjEV-1

MN385573 MN385574 MN385575 MN385576 MN557024 MN557025 MN557026 MN557027

CjEV-2

MN385577 MN385578 MN385579 MN385580

JYMaV LiCRaV

MK305894 MK305895 MK305896 MK305897 MK305898 MK305899

BPVBV PerMV PiVB TiRSaV WBYVV

MF766024 MF766029 MF766034 MF766039 LC430222d MH727572 MH727573 MH727574 MH727575 MH727576 MH727578 MH727579 MH223635 MH223636 MH223637 MH223638 MH223639 JQ354894d

Redbud yellow ringspot-associated emaravirus Redbud yellow ringspot-associated virus Rose rosette emaravirus Rose rosette virus Tentative Members Actinidia emaravirus 2 Blackberry leaf mottle-associated virus Camellia japonica associated emaravirus 1b Camellia japonica associated emaravirus 2 Jujube yellow mottle-associated virus Lilac chlorotic ringspot-associated virusc Palo verde broom virus Perilla mosaic virus Pistacia virus B Ti ringspot-associated virus Woolly burdock yellow vein virus (Arctium tomentosum virus) a

The former RNA-4 of EMARaV reported in the GenBank is here reported as RNA-5 and vice versa. An additional RNA segment (RNA 9, Accession #. MN557028) has been identified. c No sequences are available in GenBank. d Only the sequence of a partial RNA segment is available in GenBank. b

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Fig. 3 Schematic representation of the organization of the six RNA segments constituting the genome of European mountain ash ringspotassociated virus (EMARaV); the type species of the genus Emaravirus. Expression products of each RNA (p1 to p6) are represented as dark gray boxes. The function and estimated molecular weight of each protein are reported below boxes. Arrows represent the virion-complementary sense RNA, from which the proteins are translated. Figure not drawn to scale.

In addition, interspecies recombination events were detected among PPSMV-1 and PPSMV-2 segments, two species commonly found in mixed infections in pigeonpea plants. Similar RNA segment reassortments were reported for FMV and BLMaV. The phylogenetic trees inferred from the sequences of structural proteins (P1 to P4) of emaraviruses and putative emaraviruses in general displayed two clades (Fig. 4). The first group includes AcCRaV, AcEV-2, BLMaV, EMARaV, FMV, PiVB, PPSMV-1, PPSMV-2, RRV and RYRSaV, whereas the second group comprises HPWMoV, RLBV, PVBV, CjEV-1, CjEV-2, TiRSaV and JYMaV, among which the genomes of RLBV, HPWMoV and CjEV-1 have been reported to contain eight genomic RNA segments.

Life Cycle (Replication) Like other negative sense ssRNA multipartite viruses, some fimoviruses are known to use “cap snatching” to initiate transcription and facilitate translation of their mRNAs, since, unlike the polymerases from non-segmented negative strand RNA viruses, they do not possess a capping activity. Cap-snatching involves binding of host capped mRNAs to the ribonucleoproteins (RNPs), cleavage of these RNAs close to the 50 cap by a viral endonuclease activity and use of the short-capped fragments as primers for viral mRNA transcription. As for other bunyavirids, the replication of fimoviruses probably occurs in the cytoplasm.

Epidemiology Transmissibility by eriophyid mites is one of the most significant features shared by the different emaraviruses. The virus-eriophyid relationship is highly specific; it follows that only rarely the same viral species is transmitted by different eriophyid species and vice versa,

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Fig. 4 Phylogenetic relationships among emaraviruses. Neighbor joining (NJ) phylogenetic trees generated on the deduced amino acid sequences of the putative RNA-dependent RNA polymerase (RdRP, RNA-1), glycoprotein precursor (GP, RNA-2), nucleocapsid protein (NP, RNA-3) and movement protein (MP, RNA-4). Orthologs of representative members in the genera Emaravirus, Orthotospovirus and Orthobunyavirus are included. Members and tentative members of the emaraviruses reported in the phylogenetic trees are the following: AcCRaV (Actinidia chlorotic ringspot-associated virus), AcEV-2 (Actinidia emaravirus 2), BLMaV (Blackberry leaf mottle-associated virus), CjEV-1 (Camellia japonica associated virus 1), CjEV-2 (Camellia japonica associated virus 2), EMARaV (European mountain ash ringspotassociated virus), FMV (Fig mosaic virus), HPWMoV (High Plains wheat mosaic virus), JYMaV (Jujube yellow mottle-associated virus), PiVB (Pistacia virus B), PPSMV-1 (Pigeonpea sterility mosaic virus 1), PPSMV-2 (Pigeonpea sterility mosaic virus 2), PVBV (Palo verde broom virus), RLBV (Raspberry leaf blotch virus), RRV (Rose rosette virus), RYRSaV (Redbud yellow ringspot-associated virus), TiRSaV (Ti ringspot-associated virus). RNAs accession numbers of emaraviruses are reported in Table 1. TSWV (Tomato spotted wilt virus, AY070212; AY870390; KM365066) and BUNV (Bunyamwera orthobunyavirus, MH091919) were used to root the phylogenetic trees. The bar represents the number of amino acid replacements per site.

i.e., PPSMV-1 and PPSMV-2. This also explains the narrow range of host plants generally infected by each emaravirus. To date, the ascertained virus-vector transmission ratios are those between EMARaV with Phytoptus pyri, FMV with Aceria ficus, PPSMV-1 and PPSMV-2 with Aceria cajani, RLBV with Phyllocoptes gracilis, RRV with Phyllocoptes fructiphilus, HPWMoV with Aceria tosichella, and BPVBV with Aculus cercidis. In other cases (BLMaV and RYRSaV) transmission by eriophyid mites is highly suspected, but the exact species responsible has not yet been identified. Spontaneous host plants in marginal and uncultivated areas contribute to preserving populations of eriophyids and viruses when the sensitive and elective host plant is temporarily unavailable. It has also been observed that eriophyid (A. cajani, A. tosichella and P. fructiphilus) populations are larger on infected plants than on healthy ones.

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Fimoviruses (Fimoviridae)

Eriophyid mites possess a short stylet, with which they can penetrate only epidermal and, at most, the underlying mesophyll cells. Therefore, mites can acquire the disease agent only if it is present in these plant cells. Transmission efficiency increases as the number of individual mites present on the host plant increases. In experimental trials, the efficiency of PPSMV-1 transmission by a single A. cajani mite was up to 53% but rose to 100% when more than 5 mites were used per plant. Moreover, A. cajani and A. tosichella acquired PPSMV-1 and HPWMoV, respectively, after a minimum acquisition access period (AAP) of 15 min (although 50% transmission of HPWMoV was achieved only with an AAP of 16 h) and inoculated virus after a minimum inoculation access period of 90 min. Starvation before or following virus acquisition can reduce these periods. Mites retain viruses for up to 13 h in A. cajani, 4 days in A. tosichella, and 10 days in A. ficus. No latent period was observed with PPSMV-1 in A. cajani, whereas a 6–7 h latent period was reported for FMV in A. ficus. These data suggest a semipersistent fimoviruses transmission mode. Trans-stadial transmission of the virus was ascertained for A. ficus and FMV, A. cajani and PPSMV, and A. tosichella and HPWMoV. The latter virus is transmitted by all stages of A. tosichella and is retained through the molt. However, adults cannot acquire the virus and therefore transmit only if they acquired the virus during their immature stages. The mechanical transmission of fimoviruses onto herbaceous hosts is rare, and has been shown only for a few species, i.e., AcCRaV onto N. benthamiana, PPSMV-1 and PPSMV-2 onto N. benthamiana and N. clevelandii, and TiRaV onto N. benthamiana, N. tabacum and some cucurbits. The leaf stapling technique was effectively used to transmit PPSMV-1 and PPSMV-2 from infected plants onto Phaseolus vulgaris, while mite transmission was used to transmit FMV to Catharanthus roseus.

Symptomatology and Economic Impact Given the novel establishment of this virus family and the recent identification of many of its members, mainly thanks to the use of the innovative NGS technique, it is difficult to determine the actual economic impact of these viruses on crops. For many of them, their effective distribution, apart from the countries where they were first identified, remains unclear, except for FMV, which is ubiquitous. It seems certain, however, that the range of hosts naturally infected by each emaravirus species is rather restricted, due to the high specificity of the relationship between virus/vector/plant. FMV is the main, if not the exclusive agent of Fig mosaic (FMD), a worldwide fig tree disease first described in 1933. FMV causes many disorders in fig trees: poor fruit quality, reduced productivity and stunted growth (Fig. 1). Given the widespread and great incidence of this disease, the economic impact of FMV should be considered as particularly relevant. Another natural host of FMV outside the genus Ficus is the species Cyclamen persicum, while members of family Moraceae, Cudrania tricurpidata and Morus indica can be artificially infected. HPWMoV is the agent of High Plains disease (HPD), an economically important disease of wheat and maize, first found in the Great Plains region of the United States in 1993. Since then, HPD has been reported in different regions of the USA, Australia, and New Zealand. Symptoms on sweet corn consist of stunting, yellowing (chlorosis) and mosaic foliar patterns along veins, which progress into streaking patterns and necrosis. Infected wheat develops severe mosaic symptoms. PPSMV-1 and PPSMV-2 are the agents of sterility mosaic (SMD), which is the most damaging disease of pigeonpea (Cajanus cajan) in the Indian subcontinent (India, Bangladesh, Nepal, Thailand, Myanmar, Sri Lanka and China). SMD is manifested by a wide array of symptoms on pigeonpea (Fig. 1). These include various mosaic patterns on leaves, small misshapen leaves, shoots proliferation, stunting and complete or partial cessation of flower production (sterility), but the disease is not lethal to the host. Plant infection by PPSMVs at an early stage can result in a 95%–100% loss of yield, while losses from late infection depend on the level of infection and range from 26% to 97%. In the 1970s it was estimated that this disease caused annual grain losses worth over $300 million. Rose rosette disease (RRD), induced by RRV, is one of the most destructive diseases of cultivated roses in North America, regardless of cultivar. Symptoms were described in the United States as early as 1941. Spread of the disease in the USA was linked to the introduction from Japan of the multiflora rose, an exotic plant used as a rootstock for ornamental roses. Symptoms include excessive lateral shoot growth, excessive thorniness, witches' broom, leaf proliferation and malformation, mosaic, red pigmentation, and eventually plant death. EMARaV mainly infects European mountain ash (Sorbus aucuparia L.), a widely distributed indigenous tree in many parts of Europe and the Middle East, widely present in forests, but also frequently cultivated in gardens, public green areas and along avenues. The virus mostly causes chlorotic ringspots and mottling on leaves (Fig. 1), leading to the slow dieback of affected trees within a few years. Its presence has been reported in Austria, the Czech Republic, Sweden, Finland, Germany, Russia, Norway, Poland and the United Kingdom. Other host plants of EMARaV apart from S. aucuparia have been identified, such as various Rosaceae species and hybrids. The impact of the other known emaravirus species appears less important; their diffusion seems to be relatively contained at present, their etiological meaning is still uncertain, and they attack ornamental or cultivated plants with less commercial value. AcCRaV infects kiwifruits trees (Actinidia spp.), causing leaf ringspots, vein yellowing and chlorotic spots. BLMaV has been detected in many cultivated and wild blackberries affected by yellow vein, the major constraint for blackberry production in the southern United States. RYRSaV has been detected in eastern redbud trees (Cercis canadensis) grown in Arkansas, which present yellow ringspot symptoms on the leaves. Grafting can artificially infect Glicine max, Phaseolus vulgaris, Vigna unguiculata and Robinia pseudoacacia plants.

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RLBV is associated with severe raspberry leaf blotch disorder in wild and cultivated Rubus spp. plants in many European countries. It is a long-known disease characterized by large yellow blotches or rings on leaves, twisting of the leaves, distortion of leaf margins, leaf malformation, necrosis, and reduced vigor of the severely affected plants. TiRSaV is a tentative emaravirus responsible for an emerging disease of Cordyline fruticosa in Hawaii and produces small chlorotic ringspots on leaves that often coalesce to form larger lesions. PiVB is a tentative emaravirus infecting symptomless Pistacia plants in Turkey. AcEV-2 is a tentative emaravirus affecting kiwifruit tree showing leaf mottle and chlorosis symptoms in China. CjEV-1 and CjEV-2 are two tentative emaraviruses found in Italy in infected Camellia plants associated to symptoms of chlorotic and necrotic ringspots, deformations and yellowing on leaves, and deformations and color breaking of petals. JYMaV is a tentative emaravirus found in China associated to Jujube yellow mosaic, a disease characterized by mottling, yellowing and distortion of leaves, malformation and discoloration of fruits, with necrotic areas mainly around the calix. LiCRaV is a tentative emaravirus found in China on ornamental shrub common lilac (Syringa vulgaris L.), associated to leaf chlorotic ringspots and mottling symptoms. PVBV is a tentative emaravirus infecting the ornamental plant Parkinsonia florida in Arizona, causing witches’ broom symptoms. WBYVV is a tentative emaravirus attacking Arctium spp. (Asteraceae) in Finland.

Diagnosis In recent years there has been a sharp rise in the number of fimoviruses due to the impulse given using molecular techniques. Of the many such techniques available, Next-generation sequencing (NGS) technology is a powerful tool for the detection of unknown disease-associated viruses and has made a great contribution to the discovery of these viruses. Fimoviruses are easily discernable in the field because of the outstanding symptoms, i.e., typical chlorotic ringspot and mosaic, that they induce in infected plants. However, when are present together with mixed viral infections, the specific recognition of their symptoms may be disorientating. Their mechanical transmission to herbaceous hosts from infected sap, which is a simple way for recovering viruses, often is unsuccessful. This is due on the one hand to the non-transmissible nature of many of these viruses and on the other hand to the presence of plant inhibitors impeding the success of the inoculation. In the absence of serological means for the detection of most fimoviruses, except for FMV and PPSMV-1, DNA-based molecular assays with all their various techniques remain the most suitable methods for detecting these viruses. The high number of RNA segments composing the genome and the presence of conserved motifs in many of their genes has led to the development of specific and degenerate primers, which have proved effective in RT-PCR to detect fimoviruses at the species and genus levels, respectively. Furthermore, TaqMan Realtime PCR and LAMP assays have also been developed for FMV and BLMaV, providing a greater sensitivity than conventional RT-PCR assay in the detection of both viruses. At the serological level, immunological detection is based on two different polyclonal antisera raised against the recombinant RNP of FMV and on purified virus particles of HPWMoV and PPSMV-1.

Treatment and Prevention Given the impossibility of combating virus diseases in the field, control of fimoviruses is essentially preventive. It mostly consists of avoiding the introduction of infected plants that can act as a reservoir for the spread of viruses to healthy plants via vectors. Thus, rapid detection of infected plants and their immediate and complete removal, and the elimination of alternative host species, where these exist, (e.g., multiflora roses for RRV, grass-type weeds and volunteer grain crops for HPWMoV) or of infected residues from previous crops (e.g., for RRV and PPSMVs) can be very useful in reducing the amount of inoculum. Good spacing between plants so that leaves do not touch can prevent eriophyid mites from moving within a crop. Containment of the infection at very low levels in the field also serves to delay the onset of the disease and significantly attenuates the extent of symptoms, especially in herbaceous crops (e.g., in the case of PPSMVs, RRV and HPWMoV). Delayed planting of winter wheat reduces the chance for Aceria tosichella to transmit the virus and prevents an overwintering reservoir for HPWMoV. Early season planting of corn has been shown to reduce the effect of HPWMoV infections by allowing the plants to mature before the eriophyid vector becomes active. The use of genetic resistance sources is one of the most promising approaches in preventing or lessening the impact of fimoviruses. Several varieties of HPWMoV-resistant maize have been identified, although there are no known resistant varieties of wheat, barley, oats or rye. Genetic resistance lines have also been identified regarding pigeonpea by evaluating the behavior of numerous varieties preserved in germplasm collections when infected with PPSMVs. Control of the eriophyid vectors, mainly involving pesticides, is the other pillar in control of these viruses. Pesticide use alone entails a high level of risk because the eriophyid mite may develop resistance. Therefore, the most effective way to manage fimovirus diseases is a multi-tactic approach that includes both cultural and chemical practices.

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Further Reading Buzkan, N., Chiumenti, M., Massart, S., et al., 2019. A new emaravirus discovered in Pistacia from Turkey. Virus Research 263, 159–163. Di Bello, P.L., Laney, A.G., Druciarek, T., et al., 2016. A novel emaravirus is associated with redbud yellow ringspot disease. Virus Research 222, 41–47. Elbeaino, T., Digiaro, M., Alabdullah, A., et al., 2009. A multipartite single-stranded negative-sense RNA virus is the putative agent of fig mosaic disease. Journal of General Virology 90, 1281–1288. Elbeaino, T., Digiaro, M., Mielke-Ehret, N., Muehlbach, H.P., Martelli, G.P., 2018. ICTV Report Consortium. (2018) ICTV Virus Taxonomy Profile: Fimoviridae. Journal of General Virology 99 (11), 1478–1479. Elbeaino, T., Digiaro, M., Uppala, M., Sudini, H., 2014. Deep sequencing of pigeonpea sterility mosaic virus discloses five RNA segments related to emaraviruses. Virus Research 188, 27–31. Elbeaino, T., Digiaro, M., Uppala, M., Sudini, H., 2015. Deep sequencing of dsRNAs recovered from mosaic-diseased pigeonpea reveals the presence of a novel emaravirus: Pigeonpea sterility mosaic virus 2. Archives of Virology 160, 2019–2029. Laney, A.G., Keller, K.E., Martin, R.R., Tzanetakis, I.E., 2011. A discovery 70 years in the making: Characterization of the Rose rosette virus. Journal of General Virology 92, 1727–1732. McGavin, W.J., Mitchell, C., Cock, P.J.A., Kathryn, K.M., MacFarlane, S.A., 2012. Raspberry leaf blotch virus, a putative new member of the genus Emaravirus, encodes a novel genomic RNA. Journal of General Virology 93, 430–437. Mielke, N., Muehlbach, H.P., 2007. A novel multipartite negative-strand RNA virus is associated with the ringspot disease of European mountain ash (Sorbus aucuparia L.). Journal of General Virology 88, 1337–1346. Olmedo-Velarde, A., Park, A.C., Sugano, J., et al., 2019. Characterization of Ti ringspot-associated virus, a novel emaravirus associated with an emerging ringspot disease of Cordyline fruticosa (L.). Plant Disease 103 (9), 2345–2352. doi:10.1094/PDIS-09-18-1513-RE. Peracchio, C., Forgia, M., Chiapello, M., et al., 2019. A complex virome that includes two distinct emaraviruses is associated to virus-like symptoms in Camellia japonica. bioRxiv. doi:10.1101/822254. Tatineni, S., McMechan, A.J., Wosula, E.N., et al., 2014. An eriophyid mite-transmitted plant virus contains eight genomic RNA segments with unusual heterogeneity in the nucleocapsid protein. Journal of General Virology 88, 11834–11845. Wang, Y., Zhai, L., Wen, S., et al., 2020. Molecular characterization of a novel emaravrius infecting Actinidia spp. in China. Virus Research 275. doi:10.1016/j.virusres.2019.197736. Yang, C., Zhang, S., Han, T., et al., 2019. Identification and characterization of a novel emaravirus associated with jujube (Ziziphus jujuba Mill.) yellow mottle disease. Frontiers in Microbiology 10 (1417), 1–12. doi:10.3389/fmicb.2019.01417. Zheng, Y., Navarro, B., Wang, G., et al., 2017. Actinidia chlorotic ringspot-associated virus: A novel emaravirus infecting kiwifruit plants. Molecular Plant Pathology 18, 569–581.

Furoviruses (Virgaviridae) Annette Niehl and Renate Koenig, Julius Kühn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany r 2021 Elsevier Ltd. All rights reserved. This is an update of R. Koenig, Furovirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00408-8.

Nomenclature bp Base pair CP Coat protein CP-RT Coat protein-readthrough protein ELISA Enzyme-linked immunosorbent assay ER Endoplasmic reticulum GFP Green fluorescent protein HSP Heat shock protein kDa Kilodalton MP Movement protein NGS Next generation sequencing nm Nanometer

nt Nucleotide ORF Open reading frame PCR Polymerase chain reaction PD Plasmodesmata PME Pectin methyl-esterase RdRp RNA-dependent RNA polymerase RNA Ribonucleic acid siRNAs Small-interfering RNAs ssRNA Single-stranded ribonucleic acid vRNP Viral ribonucleoprotein VSR Viral suppressor of gene silencing

Glossary Furovirus

Siglum from fungus-borne, rod-shaped virus.

Classification Furoviruses are positive sense, single stranded RNA (ssRNA) viruses in the family Virgaviridae. Furoviruses have a bipartite genome and each RNA is encapsidated by a single SBWMV of coat protein (CP). Soil-borne wheat mosaic virus (SBWMV) is the SBWMV species of the genus Furovirus. The genus Furovirus presently comprises six species; Soil-borne wheat mosaic virus, Japanese Soil-borne wheat mosaic virus (JSBWMV), Soil-borne cereal mosaic virus (SBCMV), Chinese wheat mosaic virus (CWMV), Oat golden stripe virus (OGSV), and Sorghum chlorotic spot virus (SrCSV). In 2011, a barley-infecting furovirus named soil-borne barley mosaic virus (SBBMV) was identified. Because of high nt sequence identity to JSBWMV (approximately 94% on RNA2) and the fact that barley is infected as primary host species, we suggest that this virus is classified as strain of the JSBWMV. (Table 1).

Virion Structure Furoviruses have bipartite genomes which are encapsidated in non-enveloped rod-shaped particles with a helical symmetry and a diameter of approximately 20 nm. The maximum length of the particles ranges between ca. 140–160 nm and 260–300 nm. Due to internal deletions in the CP-readthrough protein genes on RNA2, particles shorter than 140 nm may arise in naturally infected plants and in laboratory isolates at later infection stages or after passage of the virus from plant to plant, and may outcompete the 140–160 nm particles. The furovirus CP has a molecular mass of approximately 20 kDa. Virions occur scattered, or in aggregates or inclusion bodies in the cytoplasm and vacuole of infected cells. Inclusion bodies may be crystalline or consist of clusters of virus particles in association with microtubules. In plants doubly infected by Sorghum chlorotic spot virus (SrCSV) and potyviruses, SrCSV virions have been shown to bind to cylindrical inclusions induced by potyviruses present in the same cell.

Nucleic Acid Properties and Differentiation of Furoviruses The nt sequences of the RNAs of all six furovirus member species have been determined. The RNAs are 50 capped and terminate in a tRNA-like structure. For SBWMV, it has been shown that the tRNA-like structure can be amino acetylated with valine in vitro. The furoviral tRNA-like structure is preceded in the 30 untranslated region by an upstream hairpin and an upstream pseudoknot domain with two to seven possible pseudoknots. The genome organization is rather similar for all furoviruses although there are considerable differences in the nt sequences (Fig. 1). The percentages of sequence identities between SBWMV and CWMV, SBCMV and JSBWMV,

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Table 1

Furoviruses (Virgaviridae) List of species in the genus Furovirus, family Virgaviridae. The SBWMV species is written in bold

Species name

Acronym

# segments

Length (nt)

Accession #

Chinese wheat mosaic virus

CWMV

2

(7147 nt) (3569 nt)

NC_002359 NC_002356

Japanese soil-borne wheat mosaic virus

JSBWMV

2

(3574 nt) (7226 nt)

NC_038851 NC_038850

Oat golden stripe virus

OGSV

2

(3232 nt) (7111 nt)

NC_002357 NC_002358

Soil-borne cereal mosaic virus

SBCMV

2

(7025 nt) (3683 nt)

NC_002351 NC_002330

Soil-borne wheat mosaic virus

SBWMV

2

(7099 nt) (3593 nt)

NC_002041 NC_002042

Sorghum chlorotic spot virus

SCSV

2

(3418 nt) (6878 nt)

NC_004015 NC_004014

Fig. 1 Phylogenetic trees generated by the DNAman software for furoviral RNA1 (A) and RNA2 (B). The Genbank accession numbers are given in parentheses.

respectively, ranges from approximately 70%–77% for RNA1. OGSV and SrCSV are more distantly related to SBWMV. For RNA2, SBWMV, JSBWMV, and SBCMV are most closely related. The percentage of sequence identities between the different members of the furoviruses for RNA2 ranges between 67% and 83%. Sequence dissimilarities justify that CWMV and SBCMV may be considered as separate species and that JSBWMV is considered a separate species rather than a strain of SBWMV (Fig. 1). However, the fact that reassortment experiments between RNA1 of JSBWMV and RNA2 of the SBWMV strain of SBWMV yielded an infectious progeny and the observation that the biological properties of the wheat-infecting furoviruses are very similar may be taken as evidence that SBWMV - and JSBWMV as well as CWMV and SBCMV may also all be regarded as distantly related strains of the same species. Moreover, differences at the nt level do not necessarily translate into differences at the aa level as exemplified with the CP sequences of the New York and Nebraska strains of SBWMV, which are 100% identical at the aa level, despite the fact that their coding regions differ in 68 nt positions.

Organization of the Genome and Properties of the Encoded Proteins Furoviral RNA1 codes for two replication-associated proteins of approximately 150 kDa and 209 kDa and for a 37 kDa movement protein (MP)(Fig. 2). The shorter of the two replication-associated proteins contains methyl-transferase and helicase motifs. The longer one results from the read-through of a stop codon and contains the RNA-dependent RNA polymerase (RdRp) in addition to the methyl-transferase and helicase domains. Over-expression experiments using fluorescent protein tagged CWMV replicase showed that it localized to membrane-associated inclusions leading to the hypothesis that virus replication might occur on these host membranes. However, the identity of the host membranes was not further investigated. The CWMV replicase was also shown to interact with heat shock proteins (HSP) of the HSP70 family. The furovirus 37 kDa MP belongs to the ‘30K’ superfamily of viral MPs. Experimental evidence for the function of the CWMV 37 kDa protein as MP came from studies showing that the green fluorescent

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Fig. 2 Genome organization of SBWMV. The RNAs are capped and possess a t-RNA structure at the 30 end. RNA1 contains the genetic information for two replicase proteins, the 150K protein containing methyltransferase (Mtr) and helicase (Hel) domains and the 209K protein containing in addition the RNA-dependent RNA polymerase (RdRP) domain. A 37K MP is encoded by ORF2. RNA2 encodes for a 19K major CP, a 25K minor CP initiated at an upstream CUG start codon, a 84K CP-RT protein and a 19K cysteine-rich silencing suppressor.

protein (GFP)-labeled 37 kDa CWMV MP moves from cell to cell when bombarded into wheat leaf cells and can trans-complement the intercellular spread of potato virus X. Moreover, CWMV MP exhibits plasmodesmata (PD) labeling and forms endoplasmic reticulum (ER)-derived vesicular and aggregate structures. Export from the ER appears to be required for intracellular movement and depends on the secretory pathway. Moreover, the CWMV MP interacts with host pectin methyl-esterase (PME), a host cell-wall modifying enzyme, which has been shown to interact with a variety of MPs of the ‘30K’ superfamily. In the case of tobacco mosaic virus, it has been shown that interaction between MP and PME is important for virus movement. Furoviral RNA2 contains the genetic information for the approximately 20 kDa major coat protein (CP), an approximately 24 kDa minor CP initiated at an upstream CUG start codon, an approximately 84 kDa CP-readthrough (CP-RT) protein and an approximately 19 kDa cysteine-rich protein. Neither the CP with N-terminal extension nor the CP-RT are required for SBWMV and CWMV virion formation and systemic infection. The 24 kDa N-terminally extended CP may be incorporated into virus particles, as it has been detected in purified CWMV virions. For SBCMV and CWMV, the 24 kDa extended CP, but not CP itself, has been shown to interact with the CWMV 19 kDa RNA silencing suppressor. In the case of SBWMV, both CP forms were able to interact with CWMV P19. As silencing activity was not significantly affected by the interaction, a function of the interaction between forms of CP and P19 in silencing suppression cannot be deduced. Despite dispensable for virion assembly and systemic infection, CP-RT can be associated with virions, as CWMV CP-RT was detected on the surface of virus particles by immunogold labeling. The CP-RT protein of furoviruses appears to be involved in vector transmission, similar to the CP-RT proteins of beny- and pomoviruses. Prolonged cultivation of field-infected plants, virus propagation by repeated mechanical inoculations or growth at elevated temperatures result in spontaneous deletions within the CP-RT domain. These mutants can no longer be transmitted by the vector. Two highly conserved trans-membrane domains within CP-RT proteins are thought to be involved in vector transmission by facilitating the transport of the virion or infectious viral ribonucleoprotein complex across the plasmodiophorid membrane into the host cell cytoplasm. The CP-RT protein also appears to affect symptom development as spontaneous mutations occurring in SBWMV CP-RT or engineered mutations affecting expression of the CP-RT protein in infectious CWMV clones increased symptom severity. Interestingly, mutations affecting expression of the N-terminally extended minor CP slowed symptom development for CWMV and SBWMV, thus indicating that the different CP-variants and the ratio of the different CP-forms produced may be involved in fine-tuning the infection cycle. The cysteine-rich 19 kDa protein acts as a suppressor of RNA silencing (VSR) and enhances symptom severity. For the CWMV protein it has been shown that it localizes to the ER and is capable to self-interact. Self-interaction, but not ER-localization correlates with VSR activity of the protein. Residues important for protein stability, silencing suppression activity, symptom development, as well as targeting to the ER have been identified. The exact mode of action of the furovirus VSR is as yet unknown; however, for the CWMV CRP, it appears not to be related with local siRNA production and accumulation. Instead, CWMV CRP protein appears to promote viral cell-to-cell spread, possibly by interfering with the spread of RNA silencing signals ahead of

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infection, as movement of a heterologous virus, whose movement depends on the presence of a functional silencing suppressor to be provided in trans, is complemented in the presence of over-expressed CWMV CRP. The localization of the CWMV CRP protein to the ER may be important for symptom development.

Virus Transmission and Movement Furoviruses are soil-borne. SBWMV has been demonstrated to be transmitted by the zoospores of Polymyxa graminis, a ubiquitous plamodiophorid protozoan. P. graminis is probably also the vector for the other viruses in the genus. Polymyxa-transmitted viruses are believed to be taken up by the plasmodia of the vector in infected root cells. The multinucleate plasmodia, which are separated from the host cytoplasm by a distinct membrane, may either develop into zoosporangia from which secondary viruliferous zoospores are released. Alternatively, the plasmodia may develop into cytosori, in which resting spores mature. These resting spores are released into the soil from decaying plant material and may survive in the soil for many years even under extreme conditions. Spores are distributed by agricultural equipment, by irrigation or even by wind. Upon germination, they release primary zoospores transmitting the virus. Zoospores inject their contents into the cytoplasm of root cells and initiate infection by the formation of plasmodia. Zoospores treated with antisera to SBWMV or resting spores treated with 0.1 N NaOH or HCl retained their ability to transmit the virus, indicating that the infectious viral material is present inside the spores. Immunolabeling and in situ RNA hybridization studies have revealed the presence of SBWMV MP and RNA but not of SBWMV CP in the resting spores of P. graminis, indicating that SBWMV may be transmitted in form of viral ribonucleoprotein (vRNP) complexes. Moreover, the presence of viral RNAs and MP may indicate that the virus is able to replicate inside the vector. Transmission in form of a vRNP complex containing viral MP and RNA may have the advantage, that the RNA can be directly transported from the primarily infected cell into adjacent cells for the initiation of virus replication and thus, competition between virus and vector for reprogramming of host cell metabolism for replication may be avoided. The ‘30K’ superfamily of MPs, to which the furovirus MPs belong, are known to mediate cell-to-cell and vascular transport of viruses by binding viral nucleic acids and carrying them through plasmodesmata and through the vasculature. However, with respect to systemic movement, SBWMV may use the xylem in order to move from infected roots to the leaves as suggested from light and electron microscopy experiments. The virus may enter primary xylem elements before cell death occurs. Moreover, the virus may move laterally between adjacent xylem vessels.

Symptoms, Epidemiology and Host Range SBWMV is known since the early 1920s. It causes chlorotic leaf streaking, stunting, and severe losses of yield in winter wheat. SBWMV is widely distributed in the USA and Canada, mainly in the east and has been reported to occur in Europe in Great Britain, Germany, and Poland. It has also been described to occur in Brazil, Africa (Zambia), and Japan. Two types of SBWMV are described, the original “Nebraska-like” SBWMV and a “New York-like” SBWMV. Symptoms often occur in patches in the fields. SBWMV naturally infects wheat, barley, rye, and triticale. An additional strain of SBWMV naturally infecting barley and wheat, termed Japanese soil-borne wheat mosaic virus, has been identified in Japan (SBWMV-JP), France (SBWMV-FR), and Germany (SBBMV-DE). Diseases on wheat with symptoms similar to those described for SBWMV are caused by CWMV in China and SBCMV in Europe. Apart from wheat, SBCMV naturally also infects rye and triticale. SBCMV occurs in a number of European countries including Denmark, Great Britain, Belgium, Poland, Ukraine, France, Italy, Germany, and Turkey. SBCMV occurs in different SBWMV with different aggressivity on wheat. In Italy, France, England, Belgium, and Denmark, wheat becomes heavily infected, while SBCMV in Germany and Poland usually develops weaker symptoms in wheat. OGSV causes yellow striping on leaves and has been detected in oats in Great Britain, France, and the USA. SBWMV, SBCMV, CWMV, and OGSV may be transmitted mechanically to some species of the Chenopodium and Nicotiana genus. SrCSV was isolated in 1986 Kansas/USA. It is mechanically transmissible to maize where it produces a bright yellow mosaic and elongated ringspot symptoms several weeks after inoculation. Local infections are produced on mechanically inoculated Chenopodium quinoa, C. amaranticolor, and Nicotiana clevelandii. Attempts to transmit the virus back to Sorghum bicolor or to winter wheat either mechanically or by growing plants in soil, in which infected sorghum was growing, were unsuccessful. Plants naturally infected by furoviruses are often also infected by bymoviruses, that is, Wheat spindle streak mosaic virus in Europe and North America, Wheat yellow mosaic virus in East Asia or Oat mosaic virus in Europe and the USA. The symptoms caused by these bymoviruses are very similar to those caused by the furoviruses, and both, the furoviruses and the bymoviruses are transmitted by P. graminis.

Diagnosis Visible symptoms caused by furoviruses may vary depending on environmental conditions, such as moisture and temperature as well as dependent on the plant cultivar. Nutrient deficiencies, winter injury or other viruses (especially bymoviruses), may produce symptoms that are easily confused with those caused by furoviruses. Thus, serological or molecular biological techniques are the methods of choice for reliable virus detection. Serological tests such as the enzyme-linked immunosorbent assay (ELISA), tissue

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print immunoassay, and immunoelectron microscopy as well as polymerase chain reaction (PCR) techniques are established for the specific detection and discrimination of the viruses. Real-time reverse transcriptase-PCR (RT-PCR) based on intercalation dyes and TaqMan probes have been developed for SBCMV and allow the quantification of virus titer within the tested plants. In the case of Next generation sequencing (NGS) it is possible to identify a virus in a sample without a priori sequence information.

Control Furoviruses inside the long-living resting spores of P. graminis may survive for decades. Growing resistant or tolerant varieties currently represents the only practical and environmentally friendly means to lower the impact of these diseases on yield. Furovirus resistance appears to be quantitative and more than one gene appears to control the resistance phenotype. The only resistance genes known to date to be effective against furoviruses in wheat are the Sbm1 gene located on chromosome 5DL and the sbm2 gene located on chromosome 2BS. Both genes confer the so called “translocation resistance”, in which roots still become infected but the virus is inhibited to efficiently infect the shoots of the plants. Although disease symptoms are reduced in the resistant plants, virus ingress into the soil remains unaffected, thus necessitating the development of additional resistance strategies to control these viruses. Cereal genotypes with resistance to P. graminis have so far not been identified.

Similarities and Dissimilarities with Other Taxa The morphology of furoviruses resembles that of other rod-shaped viruses, i.e., of benyviruses, pecluviruses, pomoviruses, hordeiviruses, tobraviruses, and tobamoviruses. The CPs of all these viruses have a number of conserved residues. Furoviruses, pecluviruses and tobraviruses have bipartite genomes. Furoviruses differ from pecluviruses by having their movement function encoded on a single ORF rather than a triple gene block. They also differ from pecluviruses as well as from tobraviruses by having a CP-RT gene. The gene for their cysteine-rich protein is located on RNA2, whereas with pecluviruses and tobraviruses it is located on RNA1. The cysteine-rich proteins of furoviruses, pecluviruses, and tobraviruses act as suppressors of post-transcriptional gene silencing and are phylogenetically interrelated. Furoviruses encode two N-terminally overlapping replication-associated proteins (Fig. 2), which are related to those of pomoviruses, pecluviruses, tobraviruses, and tobamoviruses. The 37 kDa MP of the furoviruses belongs to the ‘30K’ superfamily of MPs and relates the furoviruses to the dianthoviruses and the tobamoviruses, whereas the valine-accepting tRNA-like structure on the 30 ends of furovirus RNAs relates them to the tymoviruses. Furoviruses like tobamoviruses have an upper pseudoknot domain in the 30 untranslated regions of their genomic RNAs.

Further Reading Adams, M.J., Adkins, S., Bragard, C., et al., 2017. ICTV virus taxonomy profile: Virgaviridae. Journal of General Virology. 1999–2000. An, H., Melcher, U., Doss, P., et al., 2003. Evidence that the 37 kDa protein of Soil-borne wheat mosaic virus is a virus movement protein. Journal of General Virology 84, 3153–3163. Andika, I.B., Zheng, S., Tan, Z., et al., 2013. Endoplasmic reticulum export and vesicle formation of the movement protein of Chinese wheat mosaic virus are regulated by two transmembrane domains and depend on the secretory pathway. Virology 435, 493–503. Bass, C., Hendley, R., Adams, M.J., Hammond-Kosack, K.E., Kanyuka, K., 2006. The Sbm1 locus conferring resistance to Soil-borne cereal mosaic virus maps to a gene-rich region on 5DL in wheat. Genome 49, 1140–1148. Campbell, R.N., 1996. Fungal transmission of plant viruses. Annual Review of Phytopathology 34, 87–108. Dreher, T.W., 2009. Role of tRNA-like structures in controlling plant virus replication. Virus Research 139, 217–229. Driskel, B.A., Doss, P., Littlefield, L.J., Walker, N.R., Verchot-Lubicz, J., 2004. Soilborne wheat mosaic virus movement protein and RNA and wheat spindle streak mosaic virus coat protein accumulate inside resting spores of their vector, Polymyxa graminis. Molecular Plant-Microbe Interactions 17, 739–748. Goodwin, J.B., Dreher, T.W., 1998. Transfer RNA mimicry in a new group of positive-strand RNA plant viruses, the furoviruses: Differential aminoacylation between the RNA components of one genome. Virology 246, 170–178. Hao, Y., Wang, Y., Chen, Z., et al., 2012. A conserved locus conditioning Soil-borne wheat mosaic virus resistance on the long arm of chromosome 5D in common wheat. Molecular Breeding 30, 1453–1464. Koenig, R., Bergstrom, G.C., Gray, S.M., Loss, S., 2002. A New York isolate of soil-borne wheat mosaic virus differs considerably from the Nebraska type strain in the nucleotide sequences of various coding regions but not in the deduced amino acid sequences. Archives of Virology 147, 617–625. Kühne, T., 2009. Soil-borne viruses affecting cereals – Known for long but still a threat. Virus Research 141, 174–183. Miyanishi, M., Roh, S.H., Yamamiya, A., Ohsato, S., Shirako, Y., 2002. Reassortment between genetically distinct Japanese and US strains of Soil-borne wheat mosaic virus: RNA1 from a Japanese strain and RNA2 from a US strain make a pseudorecombinant virus. Archives of Virology 147, 1141–1153. Shirako, Y., Suzuki, N., French, R.C., 2000. Similarity and divergence among viruses in the genus Furovirus. Virology 270, 201–207. Sun, L., Andika, I.B., Kondo, H., Chen, J., 2013. Identification of the amino acid residues and domains in the cysteine-rich protein of Chinese wheat mosaic virus that are important for RNA silencing suppression and subcellular localization. Molecular Plant Pathology 14, 265–278. Te, J., Melcher, U., Howard, A., Verchot-Lubicz, J., 2005. Soilborne wheat mosaic virus (SBWMV) 19K protein belongs to a class of cysteine rich proteins that suppress RNA silencing. Virology Journal 2, 18. [18]. Verchot, J., Driskel, B.A., Zhu, Y., Hunger, R.M., Littlefield, L.J., 2001. Evidence that soilborne wheat mosaic virus moves long distance through the xylem in wheat. Protoplasma 218, 57–66. Yamamiya, A., Shirako, Y., 2000. Construction of full-length cDNA clones to soil-borne wheat mosaic virus RNA1 and RNA2, from which infectious RNAs are transcribed in vitro: Virion formation and systemic infection without expression of the N-terminal and C-terminal extensions to the capsid protein. Virology 277, 66–75.

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Yang, J., Zhang, F., Xie, L., et al., 2016. Functional identification of two minor capsid proteins from Chinese wheat mosaic virus using its infectious full-length cDNA clones. Journal of General Virology 97, 2441–2450. Yang, J., Zheng, S.-L., Zhang, H.-M., et al., 2014. Analysis of small RNAs derived from Chinese wheat mosaic virus. Archives of Virology 159, 3077–3082. Ziegler, A., Klingebeil, K., Papke, V., Kastirr, U., 2014. Quantification of Wheat spindle streak mosaic virus and Soil borne cereal mosaic virus in resistance testing of field samples of triticale using real-time RT-PCR. Journal of Plant Diseases and Protection 121, 149–155.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/w/virgaviridae/667/genus-furovirus Genus: Furovirus Virgaviridae Positive-sense RNA Viruses.

Geminiviruses (Geminiviridae) Jesús Navas-Castillo and Elvira Fiallo-Olivé, Institute for Mediterranean and Subtropical Horticulture “La Mayora”–Spanish National Research Council–University of Malaga, Algarrobo-Costa, Málaga, Spain r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein CR Common region dsDNA Double-stranded deoxyribonucleic acid EM Electron microscopy HR Hypersensitive response IPM Integrated pest management IR Intergenic region kb Kilobase kDa Kilo dalton MP Movement protein

Glossary CRISPR-Cas The acronym for Clustered Regularly Interspaced Short Palindromic Repeat-CRISPR-associated protein, an adaptive immune system used by bacteria and archaea against viruses and mobile genetic elements, which has been adapted for genome editing in eukaryotes. Emerging disease A disease of infectious origin that has appeared in a population for the first time, or that has rapidly increased in incidence or geographic range. Geminate virion A type of twinned virions unique to geminiviruses consisting of two joined, incomplete T¼1 icosahedra. Nucleic acid satellites Subviral agents depending on helper viruses for their replication and/or transmission;

NSP Nuclear shuttle protein nt Nucleotide(s) ORF Open reading frame PCR Polymerase chain reaction PTGS Post-transcriptional gene silencing RCA Rolling circle amplification RDR Recombination-dependent replication RdRp RNA-dependent RNA polymerase ssDNA Single-stranded DNA TGS Transcriptional gene silencing TrAP Transcriptional activator protein

those associated to geminiviruses (alphasatellites, betasatellites and deltasatellites) are circular single stranded DNA molecules. Pseudorecombination A particular type of genetic exchange which involves swapping of complete genome segments that make up the segmented viral genomes, as bipartite geminivirus genomes. Recombination A process in which exchange of genome segments occurs between DNA (or RNA) molecules during replication. Rolling circle amplification A technique that allows amplification of circular DNA molecules (as geminivirus genomes) with j29 DNA polymerase and random primers under isothermal conditions.

Introduction Geminiviruses (family Geminiviridae) are a large group of plant viruses that possess small circular single-stranded (ss) DNA genomes encapsidated in unique twinned (geminate) virions. Many geminiviruses cause economically important diseases to vegetable and fiber crops worldwide, although major losses are caused in tropical and subtropical regions. Many others are found infecting weeds and other wild plant species which, in turn, are potential reservoirs for geminivirus emergence. A geminivirus is thought to have been responsible for an unseasonal change in the appearance of eupatorium plants – autumnal yellowing in summer – described in a poem by the Empress Koken in Japan in 752 CE. Thus, geminiviruses are probably the cause of the first record of a viral disease in plants. With nine recognized genera and a number of species approaching 500, the family Geminiviridae is revealed as the plant virus family with the greatest evolutionary success. Their members infect both dicot and monocot species and use as vectors a wide range of insects including whiteflies, aphids, leafhoppers, and treehoppers. The present entry focuses on current knowledge about viruses in the family Geminiviridae, dealing with taxonomy, phylogeny, evolution, genome structure and expression, diagnosis, pathogenesis, epidemiology and control.

Taxonomy, Phylogeny, and Evolution Genus demarcation criteria in the family Geminiviridae include host range (monocots or dicots), type of vector (aphids, leafhoppers, treehoppers, whiteflies), genome organization (monopartite or bipartite) and phylogenetic relationships. Currently, geminiviruses are classified in nine genera: Becurtovirus, Begomovirus, Capulavirus, Curtovirus, Eragrovirus, Grablovirus, Mastrevirus, Topocuvirus and Turncurtovirus, briefly described below (Table 1).

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Table 1 Classification of members of the family Geminiviridae into genera, with indication of type member, number of virus species, number of DNA genome components, transmission vector and host type Genus

Type species

Acronym

Numbera

Genome

Vector

Host type

Becurtovirus Begomovirus Capulavirus Curtovirus Eragrovirus Grablovirus Mastrevirus Topocuvirus Turncurtovirus

Beet curly top Iran virus Bean golden yellow mosaic virus Euphorbia caput-medusae latent virus Beet curly top virus Eragrostis curvula streak virus Grapevine red blotch virus Maize streak virus Tomato pseudo-curly top virus Turnip curly top virus

BCTIV BGYMV EcmLV BCTV ECSV GRBV MSV TPCTV TCTV

3 409 4 3 1 3 40 1 2

Monopartite Monopartite/bipartite Monopartite Monopartite Monopartite Monopartite Monopartite Monopartite Monopartite

Leafhoppers Whiteflies Aphids Leafhoppers Unknown Treehoppers Leafhoppers Treehoppers Leafhoppers

Dicots Dicots Dicots Dicots Monocots Dicots Monocots/Dicots Dicots Dicots

a

Number of members according to the International Commitee on Taxonomy of Viruses (ICTV) as per March 2019.

Fig. 1 Symptoms caused by the begomoviruses Tomato yellow leaf curl virus in tomato (A), the complex Tomato yellow leaf distortion virus and Tomato yellow leaf distortion deltasatellite 1 in the wild plant Sidastrum micranthum (B) and African cassava mosaic virus in cassava (C), and the mastrevirus Chickpea chlorotic dwarf virus in chickpea (D). (D) Reproduced from Kanakala, S., Kuria, P., 2019. Chickpea chlorotic dwarf virus: An emerging monopartite dicot infecting mastrevirus. Viruses 11, 5, CC BY 4.0 licence.

Becurtovirus This genus includes three recognized species, Beet curly top Iran virus, Exomis microphylla latent virus and Spinach curly top Arizona virus. Members of all species are monopartite viruses transmitted by leafhoppers to dicot plants.

Begomovirus This genus, with 409 accepted species, is the largest in the entire virosphere. Its members infect dicot plants and are transmitted by whiteflies of the Bemisia tabaci cryptic species complex. The genome can be either bipartite or monopartite. Begomoviruses usually induce severe symptoms in their hosts, including yellow/golden mosaics, leaf curl/deformation, yellow veins and stunting (Fig. 1(A)–(C)). Devastating viruses include members of the species African cassava mosaic virus, Bean golden yellow mosaic virus, Cotton leaf curl Multan virus, and Tomato yellow leaf curl virus. Three classes of circular ssDNA satellites have been described associated with begomoviruses: alphasatellites, betasatellites, and deltasatellites.

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Capulavirus This genus includes four species, members of which have monopartite genomes. Members of the species Alfalfa leaf curl virus are transmitted by the aphid Aphis craccivora.

Curtovirus This genus includes three recognized species, including Beet curly top virus, whose members are economically important pathogens in North America and Iran and have the widest host range of any geminivirus (over 300 species reported in over 40 dicot plant families). Members are transmitted by leafhoppers.

Eragrovirus This genus includes a single species, Eragrostis curvula streak virus, members of which have monopartite genomes. All known isolates have been found infecting the monocot plant weeping lovegrass (Eragrostis curvula) in South Africa. The vector of the only known eragrovirus remains unknown.

Grablovirus This genus includes three species, members of which have monopartite genomes and infect Vitis and Prunus spp. Members of Grapevine red blotch virus are transmitted by the three-cornered alfalfa treehopper, Spissistilus festinus.

Mastrevirus This genus include 40 species, members of which have monopartite genomes, infect mostly monocots and some dicots, and are transmitted by several species of leafhoppers. It is worth mention Maize streak virus and Chickpea chlorotic dwarf virus, members of which cause severe diseases of maize in Africa and chickpea (Fig. 1(D)) and other crops in Africa and Asia, respectively. Recently, members of Wheat dwarf India virus have been shown to have associated alphasatellites and betasatellites.

Topocuvirus Members of the single species in this genus, Tomato pseudo-curly top virus, have monopartite genomes, infect dicots and are transmitted by the treehopper Micrutalis malleifera.

Turncurtovirus This genus include two species, members of which have monopartite genomes and have been isolated from symptomatic turnip (Brassica rapa) or radish (Raphanus sativus) plants from Iran, being transmitted by the leafhopper Circulifer haematoceps. Also, two unassigned species in the family Geminiviridae have been recognized, Citrus chlorotic dwarf associated virus and Mulberry mosaic dwarf associated virus. Phylogenetic analysis of complete genome sequences (DNA-A sequences for bipartite genomes) from isolates of representative species shows that geminiviruses group in clusters corresponding to the nine existing genera (Fig. 2). In addition, they are grouped to some extent according to geographic distribution, at least within the begomoviruses, probably reflecting their evolutionary divergence as a consequence of isolation. Begomoviruses can be classified into four lineages: Old World (OW) begomoviruses, New World (NW) begomoviruses, sweepoviruses and legumoviruses. OW and NW begomoviruses are originated from the Old World (Africa, Asia, and Europe) and the New World (The Americas), respectively. OW begomoviruses can have monopartite or bipartite genomes whereas most of the NW begomoviruses have bipartite genomes. A few exceptions of monopartite begomoviruses natives to the NW exist, including tomato leaf deformation virus, reported from Peru and Ecuador. Legumoviruses are restricted to legumes from Asia and Africa and most of them have bipartite genomes. Sweepoviruses infect sweet potato (Ipomoea batatas) and other members of the family Convolvulaceae and have monopartite genomes. Sweepoviruses seem to have diverged prior to the separation between the Old World and the New World begomovirus branches, seeming to represent one of the earliest points of divergence among this genus. Genetic diversity and evolution of geminiviruses are driven by mutation, recombination and pseudorecombination (reassortment of genome components) which contribute significantly to the appearance of new viral variants, increasing their potential of adaptation to different hosts and environmental conditions. Recombination, a process in which exchange of genome segments occurs between DNA strands during replication, is common among members of the family Geminiviridae, notably among members of the genus Begomovirus, contributing greatly to genetic diversification of viral populations and speciation. Replication of geminiviruses, in addition to a rolling-circle replication (RCR) mechanism, also involves a recombination-dependent replication (RDR) mechanism. RDR provides a tool by which damaged or incomplete geminivirus DNA could be recovered for productive infection by homologous recombination and converted into full-size genomic DNA. The existence of this mechanism might explain in part the extent at which recombination is shaping geminivirus populations. Recombination in geminiviruses has been shown to occur either at the strain, species, genus and family levels. Many studies have reported that an important number of begomovirus epidemics are directly linked to recombinant viruses, for example, cassava mosaic disease in Africa, tomato yellow leaf curl disease in the Mediterranean basin and cotton leaf curl disease in the Indo–Pakistan sub-continent.

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Fig. 2 Neighbor-joining phylogenetic tree of the full genome sequences (or DNA-A sequences for bipartite geminiviruses) of representative isolates of all genera (Becurtovirus, Begomovirus, Capulavirus, Curtovirus, Eragrovirus, Grablovirus, Mastrevirus, Topocuvirus, and Turncurtovirus) and unassigned species in the family Geminiviridae. OW (Old World), Legumo (legumoviruses), NW (New World) and Sweepo (sweepoviruses) are the four phylogenetic groups in the genus Begomovirus. Bootstrap values (1000 replicates) are shown. Viruses used to generate the phylogenetic tree are as follows: ACMV, African cassava mosaic virus; ALCV, Alfalfa leaf curl virus; BCTIV, Beet curly top Iran virus; BCTV, Beet curly top virus; BGYMV, Bean golden yellow mosaic virus; CCDaV, Citrus chlorotic dwarf associated virus; CLCuKoV, Cotton leaf curl Kokhran virus; CpCDV, Chickpea chlorotic dwarf virus; DesMoV, Desmodium mottle virus; EcmLV, Euphorbia caput-medusae latent virus; ECSV, Eragrostis curvula streak virus; GRBV, Grapevine red blotch virus; HrCTV, Horseradish curly top virus; MMDaV, Mulberry mosaic dwarf associated virus; MSV, Maize streak virus; PlLV, Plantago lanceolata latent virus; SCTAV, Spinach curly top Arizona virus; SPLCV, Sweet potato leaf curl virus; TCTV, Turnip curly top virus; TPCTV, Tomato pseudo-curly top virus; TYLCV, Tomato yellow leaf curl virus; WDV, Wheat dwarf virus. The bar below the tree indicates nucleotide substitutions per site.

Fig. 3 (A) Electron micrograph of Maize streak virus virions negatively stained with uranyl acetate. (B) Complete atomic model for all 110 subunits in the capsid of Ageratum yellow vein virus obtained by cryo-electron microscopy at 3.3 Å resolution, with a polyhedral cage showing the symmetry of the particle. Reproduced with permission from (A) Zhang, W., Olson, N.H., Baker, T.S., et al., 2001. Structure of the maize streak virus geminate particle. Virology 279, 471–477. (B) Hesketh, E.L., Saunders, K., Fisher, C., et al., 2018. The 3.3 Å structure of a plant geminivirus using cryo-EM. Nature Communications 9, 2369, CC BY 4.0 licence.

Virion Structure Geminiviruses have evolved a unique capsid structure. Virions are typically twinned or “geminate” particles of 22  30 nm which give the virus family its name. They consist of two joined, incomplete T¼ 1 icosahedra (hemicapsids) (Fig. 3). Cryo-electron microscopy (EM) has shown that virions of the begomovirus African cassava mosaic virus, and the mastrevirus maize streak virus containing 110 coat protein (CP) subunits organized as 22 pentameric capsomers. Cryo-EM structure at 3.3 Å resolution has further shown that the

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capsid of the begomovirus ageratum yellow vein virus is built from three distinct conformations of a single CP, and that these conformational differences facilitate the formation of the interface between hemicapsids. Each twinned virion contains a single copy of circular ssDNA. Thus, for viruses with bipartite genomes, two virions containing different genomic components are required for infection. Defective genome components and ssDNA satellites associated with begomoviruses have also been found to be encapsidated.

Genome and Viral Proteins Geminivirus genomes contain coding regions in both virion-sense and complementary-sense strands diverging from an intergenic region (IR) that includes the origin for RCR. Transcription is bi-directional, with independently controlled transcripts initiating within the IR. Multiple overlapping transcripts are used by geminiviruses for gene expression and transcript splicing is also used by members of the genus Mastrevirus and probably also by becurtoviruses, capulaviruses, and grabloviruses. The best-characterized genomes of geminiviruses are those of members of genera Begomovirus, Curtovirus, and Mastrevirus. Fig. 4 shows the genome organization of representative members of the nine genera in the family Geminiviridae. The genome of begomoviruses, the largest and more studied group in the family, is described more in detail below. The genomes of bipartite begomoviruses consist of DNA-A and DNA-B components, each of 2.5–2.7 kb. Both genome components share an approximately 200 nt segment (common region, CR) within the IR that includes the origin of replication. The genomes of monopartite begomoviruses resemble the bipartite DNA-A component. DNA-A and monopartite genomes have six open reading frames (ORFs), two in the virion sense (AV1/V1 and AV2/V2) and four in the complementary sense (AC1/C1 to AC4/C4). DNA-B has two ORFs, the virion-sense BV1 and complementary-sense BC1. DNA-A and monopartite genomes encode a coat protein (CP, ORF AV1/V1), a protein putatively involved in viral movement (ORF AV2/V2, absent in bipartite New World begomoviruses), a replication-associated protein (Rep, ORF AC1/C1), a transcriptional activator (TrAP, ORF AC2/C2), a replication enhancer (REn, ORF AC3/C3), and C4 protein (ORF AC4/C4). DNA-B encodes a nuclear shuttling protein (NSP, ORF BV1) and a movement protein (MP, ORF BC1). The CP is the only structural protein of geminivirus particles and, in addition to form the viral capsid, has been associated with other functions as insect transmission, shuttling of viral DNA into and out of the nucleous and cell-to-cell and systemic spread of the virus in for monopartite viruses. ORF AV2/V2 starts upstream ORF AV1/V1 and encodes for a protein whose exact function is uncertain, although in the case of at least some monopartite begomoviruses it is a movement protein. Rep is a multi-functional protein essential for viral DNA replication by initiating (endonuclease activity) and terminating (ligase activity) RCR. For that, Rep binds to dsDNA during origin recognition and introduces a nick in a highly conserved nonanucleotide (TAATATT↓AC) contained within a stem-loop structure that is part of the origin of replication. TrAP protein activates the transcription of the genes coding for CP and MP proteins. REn protein, although not essential for virus preplication, enhances viral DNA accumulation. ORF AC4/C4 is contained entirely within ORF AC1/C1, but in a different frame, and C4 proteins are the least conserved of all geminivirus proteins, having diverse functions including virus movement and are involved in symptom development. In addition to their specific functions, C4, TrAP and V2 proteins have been shown to suppress transcriptional (TGS) and post-transcriptional gene silencing (PTGS), whereas Rep protein supresses TGS. In the DNA-B, BV1 encodes the NSP required for trafficking viral ssDNA between the nucleus and the cytoplasm in the form of a viral DNA–NSP complex. For cell-to-cell and long-distance movement, the NSP–viral DNA complex is trapped by the MP (encoded by BC1) in the cytoplasm and redirected to adjacent cells, where NSP directs the viral genome to the nucleus to initiate replication again.

DNA Satellites Associated With Geminiviruses Three types of circular single-stranded DNA satellites are associated with begomoviruses, alphasatellites, betasatellites, and deltasatellites. Alphasatellites and betasatellites are about half size begomovirus genome components whereas deltasatellites are about half size alphasatellites and betasatellites. Alphasatellites are mainly associated with monopartite Old World begomoviruses but also with some bipartite New World begomoviruses. They encode a Rep protein needed for their replication with similarity with the Rep encoded by nanoviruses (family Nanoviridae). Betasatellites are associated with many monopartite Old World begomoviruses and are essential for induction of typical disease symptoms. The bC1 protein encoded by the betasatellites has important roles in symptom induction, suppression of transcriptional and post-transcriptional gene silencing, and can affect jasmonic acid responsive genes. Only in one case alphasatellites and betasatellites have been found associated with a mastrevirus, Wheat dwarf India virus. Deltasatellites have been found associated with bipartite Old World begomoviruses, monopartite Old World begomoviruses and sweepoviruses. Deltasatellites do not encode for any protein and, although in some cases reduce the accumulation of the helper begomovirus, rarely modify the symptoms caused by the begomovirus. All these satellites depend on their helper begomoviruses for replication (except alphasatellites), cell-to-cell and systemic spread throughout the plant, encapsidation, and transmission to new host plants by insect vectors.

Diagnosis There are a number of leaf symptoms typically associated with geminivirus – mainly begomovirus – infection, including yellow and golden mosaics, leaf curl and deformation and vein yellowing. Also, monocots infected with mastreviruses use to show typical

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Fig. 4 Genome organization of representative members of the nine genera in the family Geminiviridae. ORFs are color-coded according to the function of their protein products (rep, replication-associated protein; ren, replication enhancer protein; trap, transcriptional activator protein; cp, capsid protein; mp, movement protein; nsp, nuclear shuttle protein). Old World bipartite begomoviruses contains ORF AV2, not shown in the figure. LIR, long intergenic region; SIR, short intergenic region; CR, common region. The hairpin which includes the origin of replication is indicated in the LIR. Reproduced with permission from Varsani, A., Roumagnac, P., Fuchs, M., et al., 2017. Capulavirus and Grablovirus: Two new genera in the family Geminiviridae. Archives of Virology 162, 1819–1831.

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leaf streaks and striates. However, these symptoms, although suggestive of geminivirus infection, may be caused by other viruses, and rarely they can be used to identify the specific virus involved. Thus, diagnosis based on symptoms alone is not sufficient and specific diagnostic tests must be used to confirm geminivirus infection. For some geminiviruses, such as the begomoviruses African cassava mosaic virus and Tomato yellow leaf curl virus or the curtovirus Beet curly top virus, antibodies against the coat protein have been generated to be used in serological tests. However, although in some cases antisera have been commercialized, they are not commonly used for diagnosis because the presence of common epitopes in the coat protein hinder them to be used for specific virus identification. DNA-based methodologies have allowed the development of rapid and specific tests for detection and characterization of geminiviruses. From dot- and squash-blot hybridization using cloned geminivirus DNAs as probes to polymerase chain reaction (PCR) technologies, a wide range of methods have revolutionized the diagnosis of geminiviruses. PCR can be considered currently the method of choice for geminivirus detection due to sensitivity, specificity and rapidity. Also, PCR can be adapted to detection of a specific virus or a group of them. Sequencing of PCR products can be used later for precise identification of the virus. Loop-mediated isothermal amplification has been also used to develop tests for a number of geminiviruses. Rolling circle amplification (RCA), which amplifies circular DNA molecules in a nonspecific manner using random primers and j29 DNA polymerase, coupled with restriction enzyme digestion, has resulted recently in a valuable and powerful tool for cloning of uncharacterized geminiviruses. More recently, next-generation sequencing (Roche 454, Illumina, Ion Torren, etc.) has become increasingly accessible, allowing characterization of mixed infections and readily discovery of novel geminiviruses. Nanoporebased platforms, including portable versions, are also being used in metagenomic studies applied to the discovery of geminivirus diversity.

Life Cycle Geminiviruses, exemplified by begomoviruses, initiate infection when an insect vector (whitefly) carrying the virus feeds on the sap transported through the phloem of a healthy leaf and transmits virions to phloem-associated cells (Fig. 5). In the plant cell, viral ssDNA is released from virions and, since geminiviruses do not encode a DNA polymerase, it is copied by host DNA polymerases to generate double-stranded (ds) DNA. The dsDNA is transcribed by host RNA polymerase II, allowing the production of Rep. This protein initiates viral replication, which occurs by a combination of RCR and RDR mechanisms. Nascent circular ssDNA can be converted to dsDNA to re‑enter the replication cycle. Later in infection, Rep represses its own transcription, leading to activation of transcriptional activator protein (TrAP) expression, which in turn activates coat protein (CP) and nuclear shuttle protein (NSP) expression. Circular ssDNA can then be encapsidated by CP into virions, which are available for insect vector acquisition. The infection is propagated inside the plant by the movement of viral DNA out of the nucleus into the next cell or the phloem through the action of two viral movement proteins, nuclear shuttle protein (NSP) and movement protein (MP). An important phase of the begomovirus life cycle is the transmission between plants by the insect vector, the whiteflies of the B. tabaci complex. For that, begomoviruses need first to cross the midgut wall to reach the hemolymph and then accumulate in the primary salivary glands, from where they are secreted in the saliva to infect new plants. The transport across midgut wall and

Fig. 5 The geminivirus life cycle, exemplified by begomoviruses. See main text for details.

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through primary salivary glands are key steps determining differential transmission in different whitefly-begomovirus combinations. Although whether circulative begomovirus transmission is non-propagative (the virus does not replicate in the whitefly) or propagative (the virus replicates in the whitefly) has been a topic of debate, there is accumulating evidence that at least Tomato yellow leaf curl virus (TYLCV) is able to replicate within B. tabaci. Certain begomoviruses have been also shown to be transmitted transovarially to the descendence. From the virus side, the CP has been found to be the sole determinant of the persistent transmission of begomoviruses by B. tabaci. Inside the whiteflies, the transmission also involves certain endosymbiotic bacteria. For instance, it has been shown that transmission of TYLCV depends on chaperonin GroEL homologs produced by endosymbiotic bacteria, which have been proposed to protect the virus from degradation during passage through the hemolymph.

Pathogenicity Although the information about how geminiviruses cause diseases in their host plants is very limited, there are a number of cases in which the mechanisms involved in the production of symptoms has been studied. Determinants of pathogenicity include C4, TrAP, V2, and NSP proteins encoded by different begomoviruses and curtoviruses. C4 encoded by the curtovirus Beet curly top virus, for example, is the major determinant of the characteristic vein swelling phenotype observed in infected plants caused by abnormal cell division. In this case, C4 induces the expresion of RKP, a RING finger E3 ligase, which may be involved in cell cycle regulation. Another well studied case is that of NSP encoded by the begomovirus Tomato leaf curl New Delhi virus (ToLCNDV) which is an avirulence determinant inducing a hypersensitive response (HR) in tomato (Solanum lycopersicum) and tobacco (Nicotiana tabacum). ToLCNDV TrAP is also a pathogenicity factor which may counter NSP-induced HR in infected cells. The involvement of NSP in virus pathogenicity is supported by its interactions with membrane receptor kinases implicated in a wide range of signal transduction pathways. On the other hand, TrAP compromises the ability of COP9 signalosome (a protein complex that functions in the ubiquitin–proteasome pathway) to bind to Cullin-1 (CUL1), an essential component of the SCF (SKP1, CUL1/CDC53, F box proteins) ubiquitin E3 ligase complex; TrAP–COP9 interaction alters the cellular processes regulated by SCF complexes, including jasmonate signaling, thereby regulating host response to infection.

Control Many factors affect the occurrence and prevalence of geminivirus diseases and the losses they cause to crops worldwide. As for plant viruses in general and particularly for those transmitted by an insect vector, control of diseases caused by geminiviruses would benefit from the application of integrated pest management (IPM) strategies, which take in account all those factors. Some of the general strategies that can be used with more or less success to manage geminivirus diseases and epidemics are considered below. One of the most desirable situation is the availability of resistant or tolerant cultivars to a given geminivirus disease, as this can provide complete protection. These materials are obtained by conventional breeding methods by introgressing pertinent genes in commercial cultivars usually from wild relative species. Crops for which there are materials with effective resistance to specific begomovirus diseases include cassava (African cassava disease), cotton (cotton leaf curl disease) and tomato (tomato yellow leaf curl disease). When resistant cultivars are not available, simple control measures can be taken in order to initiate crops with virus-free planting materials, thus reducing the amount of primary inoculum. In the case of vegetable propagated from seeds, whenever possible, transplants should be produced in greenhouses instead of in open fields to protect them from insect vectors. An extreme case of this approach is to protect the crop during the entire season, as it is done when high-value vegetables are grown in greenhouses. For crops propagated from cuttings, as cassava or sweet potato, mother plants should be selected based on absence of symptoms and molecular tests to discard infected plants. Another action to be taken in annual crops is to establish them at the moment when the source of inoculum and the populations of the insect vector are the lowest possible, thus delaying infection and providing protection to young, highly susceptible, plants. In some cases, mandatory host-free periods can be established in a given area. Such host-free periods have been implemented for example in Costa Rica and Brazil to protect tomato crops from begomovirus infections. Just as vector insects play a key role in the life cycle of geminiviruses, their control is also essential when controlling the diseases that these viruses cause. A first measure, mostly used by farmers in many circumstances, is the use of insecticides to reduce vector populations. Options include systemic insecticides such as the neoicotinoids, contact insecticides, and insect growth regulators. An important limitation to the repeated application of the same active ingredient in a given area is that insect populations with high resistance to various insecticides can readly emerge, as it has been largely documented for certain species of the B. tabaci complex. Even though an insect population is susceptible to insecticide treatment, the effect may not be rapid enough to avoid virus transmission, thus limiting the practical effectiveness of this strategy for the control of geminivirus diseases. An alternative to chemical insect vector control is biological control using natural enemies, mainly parasitoids and predators. For management of whiteflies and begomovirus diseases in protected crops, numerous commercial preparations are available including the parasitoids Encarsia formosa and Eretmocerus eremicus and the predators Nesidiocoris tenuis, Macrolophus pygmaeus and Amblyseius swirskii. Two biotechnological approaches have also been evaluated to generate geminivirus-resistant crops, the production of transgenic plants and the most recent strategy of genome editing using the CRISPR-Cas system. Although many examples of transgenic

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plants resistant to geminiviruses have been evaluated and proved to be effective, mostly using gene silencing by RNA interference, these plants have not been commercialized due to controversial public acceptance. Although not yet materialized, commercialization of transgenic bean plants resistant to the begomovirus Bean golden moaic virus have been approved in Brazil. The CRISPR-Cas methodology has been evaluated against geminiviruses by editing the viral genome or host genes encoding factors required for the virus life cycle.

Further Reading Brown, J.K., Zerbini, F.M., Navas-Castillo, J., et al., 2015. Revision of Begomovirus taxonomy based on pairwise sequence comparisons. Archives of Virology 160, 1593–1619. Fiallo-Olivé, E., Pan, L.L., Liu, S.S., Navas-Castillo, J., 2019. Transmission of begomoviruses and other whitefly-borne viruses: Dependence on the vector species. Phytopathology 110, 10–17. Fondong, V.N., 2013. Geminivirus protein structure and function. Molecular Plant Pathology 14, 635–649. García-Arenal, F., Zerbini, F.M., 2019. Life on the edge: Geminiviruses at the interface between crops and wild plant hosts. Annual Review of Virology 6, 411–433. Hanley-Bowdoin, L., Bejarano, E.R., Robertson, D., Mansoor, S., 2013. Geminiviruses: Masters at redirecting and reprogramming plant processes. Nature Reviews Microbiology 11, 777–788. Hesketh, E.L., Saunders, K., Fisher, C., et al., 2018. The 3.3 Å structure of a plant geminivirus using cryo-EM. Nature Communications 9, 2369. Lefeuvre, P., Moriones, E., 2015. Recombination as a motor of host switches and virus emergence: Geminiviruses as case studies. Current Opinion in Virology 10, 14–19. Lozano, G., Trenado, H.P., Fiallo-Olivé., E., et al., 2016. Characterization of non-coding DNA satellites associated with sweepoviruses (genus Begomovirus, Geminiviridae) – Definition of a distinct class of begomovirus-associated satellites. Frontiers in Microbiology 7, 162. Navas-Castillo, J., Fiallo-Olivé, E., Sánchez-Campos, S., 2011. Emerging virus diseases transmitted by whiteflies. Annual Review of Phytopathology 49, 219–248. Rojas, M.R., Macedo, M.A., Maliano, M.R., et al., 2018. World management of geminiviruses. Annual Review of Phytopathology 56, 637–677. Saunders, K., Bedford, I.D., Yahara, T., Stanley, J., 2003. The earliest recorded plant virus disease. Nature 422, 831. Varsani, A., Navas-Castillo, J., Moriones, E., et al., 2014. Establishment of three new genera in the family Geminiviridae: Becurtovirus, Eragrovirus and Turncurtovirus. Archives of Virology 159, 2193–2203. Zerbini, F.M., Briddon, R.W., Idris, A., et al., 2017. ICTV virus taxonomy profile: Geminiviridae. Journal of General Virology 98, 131–133. Zhou, X., 2013. Advances in understanding Begomovirus satellites. Annual Review of Phytopathology 51, 357–381.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/ssdna-viruses/w/geminiviridae ICTV Report Geminiviridae. https://talk.ictvonline.org/taxonomy/ International Committee on Taxonomy of Viruses Taxonomy.

Hordeiviruses (Virgaviridae) Zhihao Jiang, Meng Yang, Yongliang Zhang, Andrew O Jackson, and Dawei Li, China Agricultural University, Beijing, China r 2021 Elsevier Ltd. All rights reserved. This is an update of J.N. Bragg, H.-S. Lim, A.O. Jackson, Hordeivirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00426-X.

Glossary Autophagy A conserved eukaryotic mechanism that removes damaged or dysfunctional components. CRISPR/Cas9 A genome editing tool that enables directly edit the genome by removing, adding or altering sections of the DNA sequence. Cryo-EM (Cryo-electron microscopy) An electron microscopic technique that involves freezing and applies on protein structure analysis. PTM (Posttranslational modification) A biochemical modification that occurs to amino acid(s) on a protein after the protein has been translated.

ROS (Reactive oxygen species) Chemically reactive oxygen radicals as well as non-radical derivatives of oxygen. VIGS (Virus-induced gene silencing) A reverse genetics tools for gene function analysis that uses viral vectors carrying a target gene fragment to produce dsRNA which initiates the silencing of the target gene. VSR (Viral suppressor of RNA silencing) Viral proteins which can counteract the host silencing-based antiviral process.

Introduction The genus Hordeivirus has been assigned to the family Virgaviridae. Hordeiviruses represent a unique genus of serologically related viruses with rigid rods that are composed of 96% protein and 4% RNA. Although members of the genus Hordeivirus have similar particle structure, they collectively infect both monocots and dicots and exhibit considerable biological diversity. Barley stripe mosaic virus (BSMV), the type member of the genus, can be traced back to B750 years ago at a site near the Nile River in modernday Egypt, and the virus has subsequently spread around the world via global movement of seeds. BSMV has been known to cause serious worldwide disease problems in cultivated barley for more than 85 years, and during this time has also been isolated infrequently from wheat and wild oats. Because of the yield losses caused in barley, a number of strains of the virus have been isolated and their biological properties evaluated. The survival of BSMV in nature depends solely on direct plant-to-plant contact and seed transmission; consequently, virus spread can be prevented by planting virus-free seed. The advent of sensitive virus detection methods has permitted seed to be screened prior to planting, thus virus-free seed production has resulted in eradication of the virus in most areas of North America, Europe, and Asia.

Taxonomy and Characteristic In addition to Barley stripe mosaic virus, three additional species, Poa semilatent virus (PSLV), Lychnis ringspot virus (LRSV), and Anthoxanthum latent blanching virus (ALBV) have been included in the genus Hordeivirus in the Eighth Report of the International Committee on Taxonomy of Viruses (ICTV). This assignment is based on serological relatedness of the coat proteins, genome organization, and sequence relatedness of BSMV, LRSV, and PSLV. The coat proteins of BSMV, LRSV, and PSLV are similar in size, but have different mobilities on polyacrylamide gels and are distantly related serologically. Serology and analyses of coat protein sequences indicate that BSMV and PSLV are more closely related to each other than to LRSV. Serological studies also show that ALBV is closely related to BSMV, and may be a strain of BSMV. However, ALBV has not been investigated by hybridization or sequence analysis to determine whether it is a strain of BSMV or is a distinct virus. PSLV has been recovered on two occasions from native grasses in widely separated locations within Canada, and from one location in Hungary, throughout North America and Europe, so the virus may be relatively abundant amongst members of the Poaceae throughout North America and Europe. LRSV was first isolated from Lychnis divaricata seeds imported into the USA, and an additional mild strain of LRSV has been isolated in Hungary. Infectious cDNA clones of LRSV and PSLV have been constructed and used to compare some biological and biochemical properties with BSMV. ALBV has been described only in Great Britain and little is known about its properties. As is the case with BSMV, LRSV is highly seed transmitted, but seed transmission has not been reported for PSLV or ALBV. All of the viruses reach high concentrations in infected plants and are easily transmitted from plant to plant by mechanical contact. In addition to their limited natural host ranges, numerous experimental hordeivirus hosts have been identified by mechanical inoculation and agro-infiltration of recombinant DNA clones. BSMV has been shown to infect several monocots and a few dicots, and recently, Brachypodium distachyon, wild emmer (Triticum turgidum var. dicoccoides), Dasypyrum villosum, ginger (Zingiber officinale and Z. zerumbet) have been shown to support BSMV replication. Local lesions on Chenopodium species have been used for several

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genetic studies of BSMV disease phenotype, and systemic Nicotiana benthamiana infections have provided a basis for analyses of virus life cycle and virus-host interactions in dicot hosts. PSLV infects several monocots while LRSV has been shown to infect several experimental dicot hosts, including several Nicotiana species and members of the family Caryophyllaceae. Reverse genetic systems for BSMV, PSLV, and LRSV have recently resulted in considerable information about biological properties of the viruses and the molecular biology of their infection processes.

Virus Particle Structure The CP interacts with the genomic RNAs to form short rod-shaped particles and always undergo end-to-end aggregation during extraction in vitro. The length of BSMV virions range from about 110–150 nm, similar with LRSV but 20% shorter than PSLV. A cryo-EM structure of BSMV virion suggests that BSMV has an average of 23.2 subunits per helix turn and has two types of virions. A wide virion form has an additional CP per turn compared with the narrow form, and has specific contact differences between the CP subunits. The diameters of the wide and narrow virions are 22.4 and 21.6 nm, respectively, and the corresponding helical parameters are 106 and 111 subunits/period. In addition, the inter-subunit of the narrow virion has two more salt bridges than the wide virion. In agreement with their serological relatedness, the BSMV and PSLV coat protein sequences are more similar to each other than to the CP of LRSV. Although the CP shares several conserved motifs with other rod-shaped viruses, the hordeivirus CPs are closely related to those of members of the fungal-transmitted virus species Peanut clump virus (PCV) and Indian peanut clump virus (ICPV), but have limited similarity to Soil-borne wheat mosaic virus (SBWMV).

Genome Structure and Expression Hordeiviruses are composed of three genomic (g) RNAs that have been designated a, b, and g based on hybridization of the viral RNAs. The sizes of the BSMV a, b, and g gRNAs vary between strains, with the ND18 strain sizes being 3.8, 3.2, and 2.8 kb, respectively. RNAs a and b are similar in size among strains of BSMV, but the sizes and complexity of the g RNAs vary considerably. The PSLV and LRSV a, b, and g gRNAs are similar in size (3.9, 3.6, and 3.2 kb, and 3.8, 3.0, and 2.7 kb, respectively) to those of BSMV. Each of the hordeivirus RNAs has a 7-methylguanosine cap at the 50 terminus, and amongst the gRNAs within a species, the 50 untranslated regions (UTRs) that are necessary for replication have very little sequence similarity. Obvious sequence similarity is not evident among the 50 UTRs of the a and g RNAs of BSMV, PSLV, and LRSV, but the RNAb components of the three viruses have considerable sequence similarity. The hordeiviruses gRNAs also contain a conserved 30 terminal non-translated sequence that is required for replication. The 30 termini are variable in length between the different viruses, ranging from 184 nt in LRSV to 290 B 330 nt in PSLV. The sequences form tRNA-like structures that in BSMV and PSLV are capable of binding tyrosine in vitro. Sequence comparisons indicate that the structures are very similar except that the 30 UTRs upstream of the conserved tRNA regions of the PSLV and BSMV RNAs have a large stem loop element and an array of possible pseudoknot stem loop regions that make the 30 UTR longer than that of LRSV. An internal poly(A) sequence of varying length is located directly after the stop codon of the 30 proximal gene of each RNA, and resides immediately upstream of the conserved 30 tRNA termini in the BSMV and LRSV genomes. Intriguingly, the poly(A) sequence is absent in the PSLV genome. Hordeivirus gRNAs (Fig. 1) encode seven proteins. The replicase protein subunits are encoded by the a and g gRNAs. RNAa encodes a single protein (aa) that contains methyltransferase and helicase domains that are highly conserved between Sindbislike viruses. The bicistronic gRNAs encodes the polymerase (ga) subunit, which contains the conserved GDD motif found in RNA-dependent RNA polymerase (RdRp) proteins of RNA viruses, plus a 30 encoded cysteine-rich protein (gb). RNAb encodes the coat protein in the first open reading frame (ORF), and is followed by a series of overlapping ORFs termed the ‘triple gene block’ (TGB) that are also present in allexi-, beny-, carla-, fovea-, peclu-, pomo-, and potexviruses. The first ORF of the TGB encodes the TGB1 protein (formerly designated bb). The remaining two TGB ORFs encode the small hydrophobic proteins, TGB2 and TGB3 (formerly called bd and bc, respectively). The aa, ba (coat protein), and ga proteins encoded by gRNAa, gRNAb, and gRNAg, respectively, are translated directly from the gRNAs (Fig. 1). Expression of the TGB proteins is mediated by transcription of two subgenomic (sg) RNAs, designated sgRNAb1 and sgRNAb2. TGB1 is translated from sgRNAb1, and the other overlapping proteins, TGB2 and TGB3, are expressed from sgRNAb2. Despite sequence predictions, in vitro translation assays, and barley protoplasts showing that the TGB20 protein is translated from sgRNAb2, the TGB20 inactivated mutant has no detectable effects on BSMV infection in N. benthamiana. The gb protein is translated from sgRNAg. Each of the encoded proteins is expressed at different levels and times during the life cycle, and the three sgRNAs have similar variances in their abundance during replication. High-level constitutive expression of sgRNAg occurs throughout the replication cycle, but expression of sgRNAb1 and sgRNAb2 is temporal. In addition, sgRNAb1 is considerably more abundant than sgRNAb2. The three sgRNA promoters have little obvious sequence relatedness; However, the individual BSMV, LRSV, and PSLV sgRNA promoters share a number of blocks of conserved sequence. Pairwise combinations indicate that the BSMV and PSLV promoters have the highest conservation, whereas the LRSV promoters have undergone more divergence. These results and the protein comparisons described below support existing evidence for a common origin of the hordeiviruses and buttress biological evidence suggesting that BSMV and PSLV are more closely related to each other than to LRSV. Additional comparisons of

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Fig. 1 The tripartite genome of BSMV and sgRNAs used for expression of the genes encoded by the b and g RNAs. Each genomic (g) RNA and sub-genomic (sg) RNA contains a 50 cap structure (m7G) and an internal polyadenylate sequence (An) preceding a 238 nt tRNA-like region that terminates with a UAA sequence. The three gRNAs are designated a, b, and g and contain open reading frames (ORFs) represented by rectangular blocks. RNAa consists of a single ORF (aa) that is translated directly from the gRNA. The aa protein forms the ‘helicase subunit’ of the RNA-dependent RNA polymerase (RdRp) and contains characteristic chloroplast transit peptides (cTP), methyltransferase (MT) and helicase (Hel) motif signatures. RNAb encodes the 50 proximal coat protein (CP) and three overlapping triple gene block (TGB) proteins (TGB1, TGB2, and TGB3), that are each required for cell-to-cell movement. The CP ORF is translated directly from gRNAb and the TGB proteins are expressed from two subgenomic (sg) RNAs, sgRNAb1 and sgRNAb2, that are illustrated below gRNAb. TGB1 contains two positively charged regions, NoLS and NLS, toward the N-terminus of the protein and a Hel domain, whereas TGB2 and TGB3 are small hydrophobic transmembrane proteins. The bicistronic RNAg encodes the 50 ga GDD-containing polymerase protein subunit, and the 30 proximal gb cysteine-rich pathogenesis protein. The ga protein is translated directly from gRNAg and the gb protein is expressed from sgRNA gb.

the TGB promoter sequences of several other viruses possessing TGB movement proteins have revealed no strong correlations in sequence or secondary structure between these putative promoter sequences and those of BSMV. Replication of BSMV gRNAs requires the 50 and 30 terminal regions, but additional internal cis-elements are required for replication of each of the RNAs. RNAa replication is cis-preferential, with replication appearing to be coupled to translation of a functional aa protein. The 117 nt intergenic region separating the ba (coat protein) ORF and the TGB1 ORF has a cis-acting function required for replication. In contrast, RNAg replication is not dependent upon the 42 nt intergenic region separating ga and gb, but essential cis-acting elements are present in the first 500 nt of the ga gene.

Function and Relatedness of the Hordeivirus Proteins Replicase Proteins (aa and ca) BSMV gRNAs expressing aa and ga are able to replicate in protoplasts. The aa and ga proteins are essential subunits of the RdRp and are members of the tobamo-lineage of supergroup III RdRps. A histidine-tagged ga protein recovered from infected barley fractionates with the aa protein and the recovered complex exhibits BSMV RNA-specific polymerase activity. Therefore, the aa and ga proteins constitute the helicase and polymerase subunits of the RdRp complex. The aa protein contains amino-terminal methyltransferase and carboxy-terminal NTPase/helicase domains. The methyltransferase domain likely functions in capping of the viral RNAs and the DEAD box helicase domain consists of at least six amino acid motifs that are conserved among the RdRps of different groups of RNA viruses. Software predictions show that aa has a chloroplast-targeted transit peptides (cTP) at the amino-terminus, and both subcellular localization assays and immuno-gold labeling experiments demonstrate that BSMV aa is localized on the chloroplast surface. A subcellular localization assay has also shown that LRSV aa has a chloroplast localization signal and targets to the chloroplasts when transient expressed in N. benthamiana. The ga RdRp hordeivirus subunits contain a GDD polymerase motif toward their C-termini that is characteristic of the supergroup III polymerases, and this domain falls into the ‘tobamo-lineage’ among the plant viruses. Phylogenetic comparisons of hordeivirus polymerase subunits reveal more than 75% conservation within the genus and lower levels of conservation extend into members of the genera Pecluvirus, Furovirus, Tobravirus (465%), and Tobamovirus (430%). Sequence comparisons of the NTPase, helicase, and GDD motifs have revealed significant conservation among the polymerase subunits of Sindbis-like viruses (supergroup III). Among the plant

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viruses within this group, the conserved domains of BSMV and PSLV are more closely related than those of LRSV, and these sequences also share close relatedness with viruses in the genera Furovirus and Pomovirus. Lower levels of sequence relatedness are observed when comparisons are extended to the genera Tobravirus and Tobamovirus and members of the family Bromoviridae.

Coat Protein The coat protein (CP) is translated directly from gRNAb and is the most abundant of the viral proteins in infected plants. The CPs of all hordeiviruses are approximately 22 kDa in size, but exhibit different electrophoretic mobilities. Studies of BSMV show that the CP is not essential for infectivity of two systemic hosts (barley and N. benthamiana) or Chenopodium species that form local lesions. An LRSV CP deletion mutant is also dispensable for systemic infection of N. benthamiana. Moreover, infections elicited by CP-deficient BSMV mutants appear to be more aggressive, and the phenotype is more severe and protracted in barley and N. benthamiana than derivatives expressing the CP. Plants infected with CP deletion mutants have serious mosaic and deformity symptoms in systemically infected leaves and appear to lack the recovery phase of infection that is usually observed in barley and N. benthamiana inoculated with BSMV. Molecular analysis also reveal that CP-deficient mutants have higher levels of viral RNA accumulation in barley and N. benthamiana than wild type BSMV, which suggests that the CP may affect replication through a feedback mechanism during the late stages of the virus life cycle when RNA replication switches to virion assembly.

TGB Movement Proteins The hordeiviruses encode Class I TGB proteins that differ in several features from the Class II TGB proteins of Potato virus X (PVX) and other viruses of the family Alphaflexiviridae. Coordinated actions of each of the three TGB proteins are required for cell-to-cell and systemic movement. The proteins are expressed transiently and simultaneously during the early stages of infection and decline in abundance at later stages of infection.

TGB1 The TGB1 protein, encoded by the 50 proximal TGB gene, is expressed from sgRNAb1 and accumulates to high levels early in infection. Among the hordeivirus species, the TGB1 proteins range from 50 to 63 kDa in size. These proteins can be distinguished from the B25 kDa Class II TGB proteins of the Alphaflexiviruses by having substantially larger N-terminal domains preceding the helicase domain. The N-terminal half of the hordeivirus TGB1 proteins contains two positively charged regions rich in lysine and arginine residues. The N-terminal region also contains a nuclear localization signal (NLS) and a localization signal (NoLS), both of which are required for nuclear-cytoplasmic trafficking of TGB1 and viral cell-to-cell movement. The C-terminal half of the TGB1 protein contains an NTPase/helicase domain with seven conserved motifs (I, IA, II, III, IV, V, and VI) that are characteristic of superfamily I helicases of alpha-like viruses. The TGB1 NTPase/helicase domain is similar to the helicase domain present in the aa protein and is also essential for BSMV movement. The PSLV TGB1 protein has RNA helicase activity in vitro that is dependent on Mg2 þ and ATP and can unwind RNA duplexes in both 50 - to  30 and 30 - to  50 directions. The BSMV and PSLV TGB1 proteins also bind ATP and dATP, and exhibit ATPase activity that maps to motifs I, IA, and II in the helicase domain. The BSMV TGB1 protein has RNA-binding activity at high ionic strength and exhibits a high binding affinity for both double-stranded (ds) RNAs and single-stranded (ss) RNAs. RNA sequence specificity has not been detected in binding studies, and TGB1 has little detectable affinity for DNA. The TGB1 proteins of both BSMV and PSLV contain multiple RNA-binding regions, and in PSLV, the N-terminal half of TGB1 containing the positively charged regions is able to bind RNA under high-salt conditions, whereas the C-terminal half containing the helicase motif binds RNA only under low-salt conditions. A BSMV TGB1/RNA ribonucleoprotein (RNP) complex has been purified from infected barley, demonstrating that TGB1 also binds to RNA in vivo. In addition, the TGB1 protein also participates in both homologous interactions and TGB3 binding. When GFP-TGB1 is transiently expressed in the absence of TGB2 or TGB3, the TGB1 protein is localized in cytoplasmic membranes. However, when GFP-TGB1 was expressed in cis during BSMV infection, fluorescence was evident on opposite sides of the cell wall, suggesting that TGB1 localizes to the plasmodesmata (PD). Due to its RNA-binding activities and presence in an RNP complex recovered from BSMV-infected plants, the TGB1 protein appears to function in formation of movement complexes that are transported to plasmodesmata and to adjacent cells during interactions with TGB2 and TGB3. Additionally, TGB1 interacts directly with a host factor, nucleolar protein fibrillarin (Fib2), and is a key component of a BSMV ribonucleoprotein (RNP) movement complex that functions in cell-to-cell movement.

TGB2 protein Among the hordeivirus TGB proteins, the TGB2 protein contains the highest amount of sequence similarity within the hordeiviruses, and the degree of sequence similarity extends to the class I TGB2 proteins encoded by PVX and other members of the family Alphaflexiviridae. The TGB2 sequence contains two hydrophobic stretches and a conserved hydrophilic region that separates the hydrophobic residues. The hydrophobic regions are predicted to integrate into membranes with a U-shaped topology that directs the termini of the proteins to the cytoplasmic side of the membrane, and the strong conserved central portion of the protein is exposed to the ER lumen.

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When expressed alone, the GFP-TGB2 protein is associated with the cortical ER where actin and TGB3 interactions result in dramatically increased filament thickening at the infection front of N. benthamiana leaves and motile vesicles resembling Golgi stacks that co-localize to plasmodesmata. However, in BSMV RNAa and RNAg infections, ectopically expressed GFP-TGB2 proteins associated with chloroplasts, which suggests that TGB2 may have unknown roles in chloroplast infection processes.

TGB3 protein The TGB3 protein is encoded by the 30 proximal ORF of sgRNAb2 and is translated by leaky scanning of ribosomes past the TGB2 AUG, which is in a poor context for translation initiation. This mechanism normally produces about a 100:10:1 ratio of TGB1:TGB2:TGB3 during in vitro translation. The TGB3 proteins are the least conserved of the TGB proteins, and the class 1 and 2 TGB3 proteins are thought to have a polyphyletic origin, in contrast to the TGB2 proteins that are thought to have originated from a common ancestor. The N-termini of the approximately 17 kDa TGB3 proteins from each of the hordeivirus species contains a stretch of conserved amino acids that includes one histidine and three cysteine residues. The TGB3 protein has two hydrophobic domains separated by a hydrophilic region that contains a conserved tetrapeptide (QDLN) sequence. The transmembrane regions are predicted to direct the TGB3 termini toward the ER lumen. This topology is quite distinct from that of the class II TGB3 proteins of the alphaflexiviruses, which have only a single membrane-spanning domain that results in localization of the C-terminus on the cytoplasmic side of the ER. When expressed alone, TGB3 localizes to paired peripheral bodies at the cell wall (CW) that appear to lie on opposite sides of the PD channels connecting adjacent cells. The targeting of PSLV TGB3 to the CW requires the central hydrophilic and the C-terminal trans-membrane regions, whereas the BSMV TGB3 protein requires the five C-terminal residues for PD localization. Furthermore, protein interaction studies demonstrate that TGB3 binds both TGB1 and TGB2, and that TGB3 directs targeting of co-expressed TGB1 and TGB2 to paired peripheral bodies at the CW.

Pathogenesis Protein (cb) The 30 proximal ORFs of the hordeivirus gRNAs encode cysteine-rich gb proteins that range in size from 16 to 20 kDa. The hordeivirus gb proteins share some structural similarities, but have little direct amino acid sequence similarity with the cysteinerich proteins of tobra- and carlaviruses. Studies of the BSMV ND18 strain demonstrate that gb is not strictly required for infectivity in plants, but that it affects pathogenicity and virus accumulation. Extensive mutational analyses of gb have revealed a number of distinct phenotypes that can ameliorate or exacerbate disease symptoms in barley, Chenopodium species, and N. benthamiana. The majority of the cysteine residues in gb are concentrated in the N-terminal half of the protein. These residues are clustered in two zinc-finger-like motifs, designated C1 and C2. A basic motif (BM) rich in lysine and arginine residues is located between the cysteine-rich regions. The BSMV gb protein has been expressed and purified from Escherichia coli and shown to bind ssRNAs and zinc in vitro. The basic motif mediates RNA-binding activity and is involved in replication processes, and the C1, BM, and C2 regions each show independent zinc-binding activity. A coiled-coil domain near the C-terminus of the gb protein mediates self-interactions that affect pathogenicity. The hordeivirus gb protein also has viral suppressor of RNA silencing (VSR) activities that interfere with host RNA silencing mechanisms, and recombinant BSMV mutants that lack gb protein functions have reduced virulence. Cis- or trans- rescue with other VSR proteins, such as the Tomato bushy stunt virus (TBSV) P19 and Potyviral HC-Pro proteins, provide RNA silencing activities that partially restore BSMV virulence. The coiled-coil region of BSMV gb is required for RNA silencing suppression, suggesting that homologous interactions of gb are required for RNA silencing suppressor activity. In addition, individual site-specific mutations within the C1, BM, and C2 region of gb also affect VSR activity of the protein, which suggests that RNA and metal binding are critical virulence components. Fluorescence of the BSMV gb-GFP fusion protein is dispersed throughout the cytoplasm, and subcellular fractionation of infected barley tissue and N. benthamiana leaf cells confirms that the protein is located in both the soluble and membrane fractions. In the presence of RNAa and RNAg, a majority of the gb proteins are recruited to the chloroplasts by interactions with the aa replication subunit protein. These interactions directly affect the BSMV replication cycle by binding progeny ssRNA, to facilitate enhanced dsRNA unwinding and BSMV replication. In addition, circumstantial evidence suggests that gb may be involved in multiple steps of BSMV infection.

Cytopathology and Replication Microscopy studies of BSMV-infected barley indicate that most of the visible changes in leaf appearance can be attributed to the disruption of organelles, most notably in chloroplasts of expanding barley and N. benthamiana leaves. Microscopy studies in N. benthamiana reveal cytoplasmic invaginations (CI) of the chloroplast outer membrane that contain B50 nm diameter spherule structures within the invaginations that are characteristic of virus replication vesicles (VRCs) (Fig. 2). Several lines of evidence have confirmed that the BSMV aa replication protein has a major role in initiating chloroplast membrane remodeling to form the VCRs. At early stages of virion assembly, large numbers of virions accumulate in association with the CI’s, suggesting that CI’s may

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Fig. 2 Ultrastructural changes and 3D model of chloroplasts membrane rearrangement during BSMV infection of N. benthamiana. (A) Invaginations present around the chloroplast envelope (white arrowheads). The insert area illustrates a spherule magnification occasionally observed inside the invagination. Scale bar, 250 nm. (B) Immunogold labeling of dsRNA in BSMV-infected N. benthamiana cells. Insets I and II show magnifications of the corresponding boxed regions. Scale bar, 200 nm. (C) CP-specific antiserum showed specific labeling of virion-like particles (black arrows) in the cytoplasm and CIs. Scale bar, 200 nm. (D) Tomogram slices of altered chloroplast membranes from leaves of BSMV-infected N. benthamiana. The yellow arrowheads indicate the same spherules in different slices. Scale bar, 100 nm. (E–G) 3D model of remodeled chloroplast membranes induced by BSMV. Translucent white, inner chloroplast membrane; Light blue, outer chloroplast membrane; Yellow, spherules derived from the outer membrane. Scale bars, 100 nm. (H–I) A 3D surface rendering of the chloroplast tomogram. Green, chloroplast; Yellow, CI. To emphasize the CI (yellow) inside the chloroplast, the appearance of chloroplast shown in H was changed to translucent (I). White arrowheads indicate chloroplast envelope invaginations similar to those in Fig. 2G; black arrow point to the CI opening. The image inset in H is an enlarged view of the CI. Scale bars, 500 nm. This figure is adapted with permission from Jin, X., Jiang, Z., Zhang, K., et al., 2018. Three-dimensional analysis of chloroplast structures associated with virus infection. Plant Physiology 176, 282–294. (© 2018 by the American Society of Plant Biologists).

provide sites for virion assembly, but at later stages of infection, so many virions are present in the cytoplasm and associated vacuoles that it is difficult to determine specific sites of accumulation. A model for hordeivirus replication posits that virions are disassembled by host factors in the plant cell, followed by translation of RdRp subunits from RNAa and RNAg (Fig. 3). The available evidence indicates that the aa subunit replication protein localizes to chloroplasts via an N-terminal chloroplast localization signal and induces chloroplasts outer membrane invaginations. The aa protein is then posited to recruit gRNAs, the ga subunit, gb, and relevant host factors to the chloroplasts to facilitate replication. Current evidence suggests that the gb protein is recruited to the VCRs by interactions with the methyltransferase (MT) hinge-like domain of the aa protein. During replication, the gb protein enhances the helicase activity of the aa protein via its N-terminal ssRNA binding activity to facilitate efficient RNA unwinding and replication. Subsequently, progeny vRNAs in the VRCs interact with TGB1 proteins to assemble RNP movement complexes. These complexes participate in movement by interacting with TGB2

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Fig. 3 Model of the hordeivirus infection cycle. Hordeivirus transmission from plant to plant occurs via mechanical or pollen transmission, and infections may be subsequently maintained by seed transmission through the embryo to germinating seedlings. Entry during mechanical abrasion is followed quickly by uncoating of gRNAs in the cytoplasm and translation of gRNAs a, b, and g. Then, the translated aa and ga proteins interact with gRNAs to form RNA-Dependent-RNA polymerase complexes that localize at chloroplast outer membranes where membrane invaginations are induced to form nascent VRCs. The RdRp then recruits relevant host factors (HF) to the VRCs to initiate robust gRNA replication. Production of negative- and positive- sense gRNAs results in formation of dsRNA replicative intermediate species. The RdRp also initiates transcription of sgRNAs from internal promoters on negative-strand RNAb and RNAg to generate sgRNAs that function in translation of proteins encoded by the sgRNAs. The gb protein, translated by sgRNAg, functions as a helicase enhancer to facilitate RNA replication and subvert host autophagy processes. During the course of these events, phosphorylated gb proteins suppress RNA silencing pathways and host cell death responses. The TGB1 protein has a nuclear phase during movement and relies on the importin a/b protein for some aspects of nuclear-cytoplasmic export and trafficking. The exported TGB1 protein interacts with the fibrillarin protein and binds to positive-sense gRNAs to create ribonucleoprotein (RNP) complexes. The TGB2 protein functions in ER/actin membrane thickening at the infection front, and binds to TGB3 proteins that interact with CK2 kinase phosphorylated TGB1-RNP complexes during transport to the plasmodesmata (PD). The TGB3 C-terminal amino acids have a central role in PD recognition and transit of RNP gRNAs complexes to adjacent cells, and the gb protein may also be involved in some aspects of movement. The TGB2 and 3 proteins also may be recycled by the endocytic pathway to facilitate the movement of additional RNP complexes. As the later stages of infection progress, increasing amounts of CP are translated from gRNAb and encapsidate positive-sense gRNAs at chloroplast encapsidation sites to form progeny virions that accumulate in the cytoplasm.

and TGB3 to elicit actin remodeling that produces dramatically thickened filaments that associate with the endoplasmic reticulum. TGB2 and 3 actin interactions then facilitate transit of RNP complexes to peripheral vesicles and through the PD to adjacent cells. Molecular events involved in virion assembly are currently obscure, but it is likely that the CP has a major role in interacting with nascent gRNAs at CIs during the late stages of replication for virion assembly.

Pathogenesis Natural mutants in both the non-coding and coding regions of BSMV RNAs have been shown to have strain-specific effects on pathogenesis. For example, the ND18 strain is unable to infect oats, whereas the CV42 strain is pathogenic in this host. ND18 virulence in oats can be engineered by altering a single amino acid in the aa protein, so this phenotype may result from specific associations of aa with a host protein. Some site-specific amino acids of BSMV are potential targets of host resistance genes. Most of the known resistance genes in barley and oats are recessive, but analysis of the inheritance of resistance in B. distachyon indicates that a single dominant gene, designated Barley stripe mosaic virus resistance 1 (Bsr1), recognizes an arginine residue at position 390 (R390) and a threonine residue at position 392 (T392) of BSMVND18 TGB1 and elicits the BD3–1 Bsr1 hypersensitive response, whereas the BSMVNW containing lysines at both positions does not. Natural variation amongst BSMV strains has also facilitated identification of factors affecting seed transmission. Primary determinants influencing the efficiency of seed transmission reside in RNAg sequences in the 50 UTR and in the gb gene. In a fourth case of strain-specific virulence, ND18 is able to systemically infect N. benthamiana, whereas the Type strain is able to replicate and move in inoculated leaves, but is unable to invade systemically. Infections with the ND18 and the Type strains also result in

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different local lesion phenotypes on C. amaranticolor. These aberrant phenotypes appear to be caused by the presence of a short ORF within an amino-terminal 372-nucleotide duplication in the 50 UTR of the Type strain that reduces ga translation. However, the ga replicase/host interactions contributing to these distinct movement properties have not been defined molecularly, and differences in the amino acid composition of the gb proteins may also affect the disease phenotypes. Advances in understanding pathogenesis have also been obtained by reverse genetic mutations. The results show that gb is not strictly required for infectivity in plants, but that it has important pathogenicity effects. Individual site-specific mutations within the cysteine-rich region of gb can also result in a variety of phenotypes ranging from striking white regions to necrotic streaks on systemically infected leaves of barley and N. benthamiana. Moreover, plants infected with site-specific gb mutants have secondary effects that culminate in greatly reduced abundance of viral RNA accumulation. Although the CP is not required for infectivity in barley, N. benthamiana, or Chenopodium species, infections elicited by CP-deficient BSMV mutants appear to be more aggressive, because recovery is protracted and the symptoms are more severe than in the presence of the CP. These results suggest that in addition to being essential for formation of virions, the coat protein may have a critical role in the kinetics of replication late in infection. Overall, these analyses suggest that complex interactions may affect virulence and disease phenotype, and that the levels of replication may have important effects on movement, host range, and seed transmission. Molecular genetic studies also suggest substantial specificity amongst genes encoded by BSMV, LRSV and PSLV RNAs, because heterologous strand reassortments of BSMV, LRSV, and PSLV gRNAs, fail to establish systemic infections in plants. However, substitution of the PSLV TGB for the BSMV TGB in RNAb results in systemic infections of N. benthamiana, C. amaranticolor, and wheat (Triticum durum), whereas similar LRSV TGB substitutions failed to establish systemic infections of any of these plants. Interestingly, BSMV infects all 4 hosts tested, whereas PSLV infects only wheat and LRSV infects N. benthamiana and C. amaranticolor, but not wheat. These results suggest that factors other than the TGB sequences per se contribute to host range, but this appears not to be a consequence of gb because our recent studies show that heterogeneous substitutions of PSLV and LRSV gb for BSMV gb do not interfere with the ability of BSMV to infect N. benthamiana.

Hordeivirus-Host Interactions cb Interference With Host Defenses RNA silencing pathways Several lines of evidence indicate that a major function of gb is to serve as an inhibitor of host defense mechanisms that target RNA replication. A VSR deficient gb mutant compromises BSMV replication, but the gb function can be partially complemented by expression of tombusvirus p19 and potyvirus HC-Pro VSRs in cis or in trans. Similarly, expression of the gb protein is able to suppress silencing in heterologous virus-induced silencing systems. An agrobacterium-mediated transient silencing assay also provides strong support for a role of gb in suppression of RNA silencing. In this assay, gb interferes with silencing of GFP fluorescence induced by expression of dsRNAs. These results thus provide a persuasive argument that a major function of gb is to counter host innate RNA interference defenses, as well as to assist in helicase unwinding during replication.

Autophagy responses Autophagy is a conserved defense pathway against abiotic and biotic stresses including virus infection. However, plant virus counterdefense mechanisms that interfere with autophagy to promote viral infection have not been explored extensively. We recently found that BSMV infection suppresses autophagy and that expression of the gb protein is sufficient to inhibit autophagy. Molecular analyses indicate that gb interferes with interactions of the autophagy-related protein 7 (ATG7) with ATG8 in a competitive manner and the conversion of ATG8 to ATG8-PE. These findings reveal that the gb protein subverts autophagy-mediated antiviral defenses by disrupting ATG7-ATG8 interactions to promote BSMV infection, and that ATG7 is a newly discovered target of pathogen effectors in the plant defense and virus counter-defense arms race.

ROS bursts Reactive oxygen species (ROS) such as hydrogen peroxide (H2O2) or superoxide anions (O2-) are amongst the earliest cellular responses against pathogen infection. BSMV infection reduces both H2O2 and O2- elevation by B10 fold in inoculated leaves compared with the empty vector. The gb protein has a major role in reduction of ROS bursts by interacting directly with glycolate oxidase (GOX), which is localized in peroxisomes and is a key constitutive enzyme in the photorespiration pathway. These results demonstrate that gb interacts with GOX to suppress peroxisomal ROS production and facilitate virus infection, and contributes to an understanding of roles played by ROS and plant ROS-producing enzymes during viral infection.

Post-Translational Modifications of BSMV Proteins Post-translational modifications are ubiquitous in eukaryotic cells, and are defined by addition of a covalent group or an enzymatic modification of proteins after translation. Viruses have evolved many strategies to hijack host post-translational modification pathways to maximize their pathogenicity. During BSMV infection, protein kinase CK2 from barley and N. benthamiana phosphates the TGB1 protein mainly at Thr-401, and Thr-395 kinase docking sites. The TGB1T395A/T401A mutation compromises BSMV cell-to-cell

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movement, and the virus fails to develop systemic infections in barley and N. benthamiana. The phosphorylated TGB1 protein interacts more strongly than unphosphorylated TGB1 with the TGB3 protein, and this interaction promotes BSMV cell-to-cell movement. Post-translational modifications also affect gb functions during BSMV infection. A recent study also indicates that gb Ser96 is phosphorylated in vitro and in vivo by a protein kinase A-like kinase. Non-phosphorylatable gb substitution mutants (BSMV gb S96A and S96R) reduced BSMV accumulation and induced necrosis in barley, wheat, and N. benthamiana. Phosphorylation of the gb protein at Ser-96 also enhances VSR activity by promoting 21-bp dsRNA binding activity. These results in toto suggest that phosphorylation may affect several aspects of pathogenesis and infection, including movement, cell death, and RNA silencing responses.

Hordeivirus Applications to Cereal Genomic Analyses Virus-induced gene silencing (VIGS) has provided useful approaches to improve our understanding of the functional genomics of monocots and dicots, and several virus vectors have been developed for this purpose. BSMV has been developed into an effective VIGS vector by modifying sequences downstream of the gb gene to enable expression of untranslatable foreign inserts. This strategy has been successfully applied to N. benthamiana, wheat, barley, and B. distachyon by targeting phytoene desaturase (PDS), magnesium chelatase subunit H (ChlH), and plastid transketolase (TK) genes for VIGS. Other BSMV VIGS applications have been developed to downregulate host genes known to be involved in disease resistance, and these have enabled identification of genes required for powdery mildew resistance in barley, and rust resistance in wheat. In addition, high levels of foreign proteins and chimeric proteins such as GFP proteins fused to the C-terminus of the gb protein have been expressed from BSMV vectors. Heterologous gb proteins less than 110 amino acids have been expressed during systemic infections, and the fusion proteins can be efficiently and constitutively expressed during infections of N. benthamiana and some monocots. These studies indicate that BSMV VIGS provides a powerful and robust system for gene silencing and gene overexpression from BSMV may have numerous applications for genetic analyses of a large number of cereal crops.

Future Perspectives The broad host range of hordeiviruses and the information accumulated about virus genes involved in replication, movement, and responses to host resistance provides a rich resource for obtaining insight into the mechanics of infection. Considerable efforts have recently focused on BSMV replication and interactions between hordeivirus-encoded proteins and host factors. However, very little is known about the roles of host interactions that facilitate hordeiviruses replication cycles, switch from replication to RNP movement complexes that move across PD and into vascular tissues, or how host resistance pathways interfere with hordeivirus infection. An understanding of events leading to the establishment of replication factories and initiation and regulation of replication cycles requires additional information about the interplay between chloroplasts and cytoplasmic elements during the early stages of infection. An understanding of BSMV-chloroplast interactions may also reveal mechanisms whereby recessive resistance genes employed in agriculture affect BSMV replication. Additional biochemical and cell biological analyses of the TGB-encoded proteins and their interaction proteins could also hone our understanding of the cell-to-cell and vascular movement events that permit systemic hordeivirus infections, and how hordeiviruses invade vegetative tissue, seed, and pollen. Such investigations may also provide insights into global aspects of the subcellular transport network, and functions of organelles and PD functions in plant development. New genome-editing techniques like CRISPR/Cas9 have developed rapidly over the past decade, and the use of hordeiviruses for novel applications of these techniques will accelerate future genomics methods. Hence, we envision that more sophisticated analyses of host genes, cellular interactions, and the ability to modulate their expression will accelerate the pace of future hordeivirus research findings.

Further Reading Adams, M.J., Adkins, S., Bragard, C., et al., 2017. ICTV virus taxonomy profile: Virgaviridae. Journal of General Virology 98, 1999–2000. Bragg, J.N., Lim, H.S., Jackson, A.O., 2008. Hordeivirus. In: Mahy, B.W.J., Regenmortel, M.H.V.V. (Eds.), Encyclopedia of Virology. Oxford: Academic Press, pp. 459–467. Clare, D.K., Pechnikova, E.V., Skurat, E.V., et al., 2015. Novel inter-subunit contacts in Barley stripe mosaic virus revealed by cryo-electron microscopy. Structure 23, 1815–1826. Cui, Y., Lee, M.Y., Huo, N., et al., 2012. Fine mapping of the Bsr1 Barley stripe mosaic virus resistance gene in the model grass Brachypodium distachyon. PLOS One 7, e38333. Holzberg, S., Brosio, P., Gross, C., Pogue, G.P., 2002. Barley stripe mosaic virus-induced gene silencing in a monocot plant. The Plant Journal 30, 315–327. Jackson, A.O., Lim, H.-S., Bragg, J., Ganesan, U., Lee, M.Y., 2009. Hordeivirus replication, movement, and pathogenesis. Annual Review of Phytopathology 47, 385–422. Jiang, Z., Li, Z., Yue, N., et al., 2018. Construction of infectious clones of lychnis ringspot virus and evaluation of its relationship with barley stripe mosaic virus by reassortment of genomic RNA segments. Virus Research 243, 106–109. Jin, X., Jiang, Z., Zhang, K., et al., 2018. Three-dimensional analysis of chloroplast structures associated with virus infection. Plant Physiology 176, 282–294. Lee, W-S., Hammond-Kosack, K.E., Kanyuka, K., 2012b. Barley stripe mosaic virus-mediated tools for investigating gene function in cereal plants and their pathogens: Virus-induced gene silencing, host-mediated gene silencing, and virus-mediated overexpression of heterologous protein. Plant Physiology 160, 582–590. Lee, M.Y., Yan, L., Gorter, F.A., et al., 2012a. Brachypodium distachyon line Bd3-1 resistance is elicited by the Barley stripe mosaic virus triple gene block 1 movement protein. Journal of General Virology 93, 2729–2739. Lim, H-S., Lee, M.Y., Moon, J.S., et al., 2013. Actin cytoskeleton and golgi involvement in Barley stripe mosaic virus movement and cell wall localization of triple gene block proteins. The Plant Pathology Journal 29, 17–30.

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Yang, M., Zhang, Y., Xie, X., et al., 2018a. Barley stripe mosaic virus gb protein subverts autophagy to promote viral infection by disrupting the ATG7-ATG8 interaction. Plant Cell 30, 1582–1595. Yuan, C., Li, C., Yan, L., et al., 2011. A high throughput Barley stripe mosaic virus vector for virus induced gene silencing in monocots and dicots. PLOS One 6, e26468. Zhang, X., Dong, K., Xu, K., et al., 2018. Barley stripe mosaic virus infection requires PKA‐mediated phosphorylation of gb for suppression of both RNA silencing and the host cell death response. New Phytologist 218, 1570–1585. Zhang, K., Zhang, Y., Yang, M., et al., 2017. The Barley stripe mosaic virus gb protein promotes chloroplast-targeted replication by enhancing unwinding of RNA duplexes. PLOS Pathogens 13, e1006319.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/w/virgaviridae/669/genus-hordeivirus Genus: Hordeivirus.

Idaeoviruses (Mayoviridae) Robert R Martin and Karen E Keller, Horticultural Crops Research Unit, Agricultural Research Service, US Department of Agriculture, Corvallis, OR, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of A.T. Jones, H. Barker, Idaeovirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00539-2.

Nomenclature

MTR Methyl-transferase nt Nucleotide(s) ORF Open reading frame PCR Polymerase Chain Reaction PVP Polyvinyl pyrrolidone RdRP RNA-dependent RNA polymerase RISC RNA-induced silencing complex satRNA Satellite RNA UTR Un-translated region

aa Amino acid(s) CP Coat protein ELISA Enzyme-Linked Immuno Sorbent Assay GFP Green fluorescent protein HEL Helicase kb Kilobase kDa Kilo dalton MP Movement protein

Glossary Certification Program Is a comprehensive process established and authorized by a state or other governmental entity for the production of plants free of regulated pests and diseases. The regulations for each program define the program participation, plant production, plant identification and labeling, and quality assurance requirements. ELISA An acronym for Enzyme-Linked Immuno Sorbent Assay, which is a serological test in which antibodies are used to detect plant viruses. Foundation Block A foundation block is a group of plants that have been tested for viruses or other diseases and are maintained in isolation under conditions that prevent (re) infection. These blocks are considered Generation 1 (G1) stock within the meaning of the ‘Generation level’ concept recommended by the NAPPO Guidelines. Heat therapy Is a method used to eliminate viruses in which plants are grown at 371C–401C for 4–6 weeks. This is used in combination with meristem tip culture for efficient elimination of most plant viruses. Horizontal Transmission Transmission of a pathogen to individuals within the same generation. For pollen-borne plant viruses, this is virus movement from the pollen grain

to maternal tissue during the pollination process resulting in the infection of the pollinated plant. Meristem tip-culture The process of removing pieces of plant 0.5 mm or less in length, consisting of a meristem and a few leaf primordia that is grown in vitro with the goal of generating a plant free of plant viruses. Pollen-borne Carried on or in the pollen, plant viruses can be carried on the exterior of the pollen or internally in the pollen grain. Polymerase Chain Reaction Is a laboratory detection technique for plant pathogens that amplifies a segment of DNA or RNA from the target organism (for example, a virus) many times by using short segments of DNA (primer) that are complimentary to the target nucleic acid. For RNA targets, reverse transcription is required to convert the RNA to DNA that is then used in PCR. Tissue Culture The cultivation of plants (cells, tissues, or organs) under aseptic conditions in a synthetic medium in vitro. Vertical Transmission Transmission of a pathogen from one generation to the next, for plant viruses this is the result of seed transmission.

Introduction Until recently, Raspberry bushy dwarf virus (RBDV) was the sole recognized member of the genus Idaeovirus. More recently, Privet leaf blotch associated virus (PrLBaV) has been listed as a member of the genus. A novel virus from blackcurrant, Blackcurrant leaf chlorosis associated virus (BCLCaV), or Blackcurrant idaeovirus (BCIV) has been described by two groups (the acronym BCLCaV will be used here), which is phylogenetically most similar to PrLBaV and RBDV. A third virus, Japanese holly fern mottle virus has similarities in the RNA1 to that of RBDV, but its RNA2 is quite distinct from the other three viruses. A partially sequenced virus from citrus showed significant similarities to the coat protein of RBDV, but complete sequence information is lacking. The initial isolation of RBDV was from plants that exhibited stunting and proliferation, thus the name Raspberry bushy dwarf virus. RBDV was initially isolated from red raspberry by mechanical transmission from symptomatic plants to herbaceous hosts. It was shown later that these plants were coinfected with Black raspberry necrosis virus and that RBDV in single infections did not cause the ‘bushy dwarf’ symptoms. However, in single infections, RBDV does induce

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crumbly fruit in many cultivars of raspberry and in some cases leaf chlorosis. The virus genus name is from the species of the host (Rubus idaeus). RBDV is serologically indistinguishable from Loganberry degeneration virus and synonymous with Raspberry yellows virus. It is also probably the same as Raspberry line pattern virus reported from Poland. Antisera has not been prepared to the other idaeoviruses. Studies on the molecular biology of RBDV have shown it to possess a novel combination of properties and was assigned as the sole known member in the genus Idaeovirus. Though RBDV and the other idaeoviruses have a bipartite genome, they are most similar to the ilarviruses that have tripartite genomes in the family Bromoviridae.

Geographical Distribution RBDV probably occurs wherever red raspberry (R. idaeus, R. strigosus), black raspberry (R. occidentalis), or blackberry (R. fruticosus), are grown. The virus has been reported from Australasia, China, Eastern and Western Europe, North and South America, South Africa, and the former Soviet Union. Three strains of RBDV have been characterized based on serological or biological properties. The common or Scottish (S) isolates have been identified in most countries where raspberries are grown. The black raspberry (B) isolate from a North American black raspberry, R. occidentalis are serologically distinct from the S isolates. However, more data are needed to determine whether B isolates are restricted to R. occidentalis, or representative of isolates from North America, and whether S isolates can occur in R. occidentalis in nature. B isolates have only been found in North America in native Rubus. A third strain (RB) for resistance breaking, has limited distribution in Europe and was identified in North America for the first time in 2016. These isolates are indistinguishable from S isolates, except that they are able to infect cultivars that are resistant to the S isolates. A fourth strain has been partially characterized from R. multibracteatus collected in China, which has 23 aa substitutions in the coat protein (CP) when compared to the B isolate and is the most divergent isolate of RBDV reported to date. It is also distinguishable from D, S, and RB isolates using monoclonal antibodies. A fifth strain has been reported from grapevines in Slovenia and Hungary, which forms a separate cluster from the isolates from Rubus species, and can also be distinguished using monoclonal antibodies. BCLCaV and BCIV were identified using high throughput sequencing from dsRNA purified from symptomatic plants. The distribution and population structure of the virus have not been studied. The virus isolates were from two different blackcurrant cultivars, ‘Belaruskaya Slodkaya’ held in the Ribes germplasm collection at the USDA-ARS National Clonal Germplasm Repository in Corvallis, Oregon, and ‘Baldwin’ from British Columbia, Canada. In both cases the plants exhibited a general chlorosis symptom. PrLBaV was identified using high throughput sequencing of small RNAs purified from privet (Ligustrum spp.) plants exhibiting yellow mosaic symptoms. Infectious chlorosis of privet, variegation of privet, mosaic of ligustrum are considered synonyms of yellow mosaic of privet. In parallel experiments with symptomless privet, the virus was not detected. Additionally, none of the other viruses reported to infect privet were detected in these experiments. There was no mention if there were any other viruses detected in the study that had not been previously reported in privet. The disease has been reported to be widespread in Europe based on symptoms, though it is not known if the multiple reports refer to the same disease or if there are multiple causal agents involved.

Natural and Experimental Transmission In red raspberry and black raspberry, RBDV is readily transmitted to progeny seedlings via either the pollen or the ovule of infected plants and transmission is greatest when both parents are infected. If raspberry plants were deflowered they did not become infected when adjacent flowering plants of the same cultivar did become infected. Therefore, the flower is believed the route for natural transmission of the virus. Healthy plants can be infected after the first flowering season when planted close to virus sources. The rate of virus spread horizontally in the field varies greatly by cultivar. Horizontal transmission occurs as pollen tubes penetrate the stigma and virus is able to move into the stigmatic tissue, and from there move systemically to infect the plant. This transmission has been documented using pollen from RBDV infected Rubus to infect healthy Rubus plants. It was also shown that transmission to the maternal tissue requires pollen tube growth into the stigma, but does not require fertilization. Pollen from RBDV infected raspberry placed on stigmas of Torenia fournieri resulted in horizontal transmission of RBDV to the maternal tissue if the pollen tube penetrated the stigma. If the pollen was inactivated and not able to grow into the stigma transmission did not occur. Thus, horizontal transmission via infected pollen can occur between sexually incompatible species. RBDV is transmitted easily from Rubus by grafting to susceptible Rubus species and cultivars. However, graft-inoculated plants may need to go through a dormant season before the virus becomes fully systemic. RBDV is also transmissible from infected Rubus to herbaceous test plants by mechanical inoculation of sap extracts in aqueous solutions of either 2% nicotine or in 0.05 M phosphate buffer containing 1% polyvinyl pyrrolidone (PVP) (mol. wt. 6000–40,000 Da). Mechanical transmission from RBDV-infected herbaceous plants to susceptible red raspberry cultivars is possible, but with great difficulty. BCLCaV is graft transmissible to other blackcurrants and PrLBaV is graft transmissible to symptomless privet plants. In both cases, limited work on graft transmission has been reported, so symptoms in different cultivars or species is unknown. BCLCaV was transmitted mechanically to Nicotiana benthamiana and N. occidentalis. There are no reports of transmission of PrLBaV to herbaceous hosts.

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Fig. 1 Top. Leaves of RBDV infected ‘Caroline’ red raspberry (A) and ‘Marion’ blackberry infected with RBDV, Blackberry calico virus, and Black raspberry necrosis virus (B). ‘Marion’ infected with the latter two viruses shows a fine line pattern and do not have fruit symptoms. Center: Symptoms of crumbly fruit of ‘Meeker’ red raspberry: (C). fruit from healthy plant (D). fruits from RBDV-infected plant. Bottom: Fruit of ‘Marion’ blackberry infected with Blackberry calico virus and Black raspberry necrosis virus; (E). plants without RBDV, (F). plants with RBDV infection. Photographs courtesy of R.R. Martin.

Disease Symptoms and Effects Naturally Infected Plants In nature, RBDV occurs in wild and cultivated Rubus species and cultivars. In single infections RBDV is often symptomless in raspberry and blackberry plants depending on cultivar. However, in sensitive raspberry cultivars and under certain environmental conditions, it can induce chlorosis. This disease is characterized by an initial yellowing of the main or minor veins of the lower leaves that can progress to affect the entire leaf (Figs. 1(A) and (B)). Some raspberry cultivars may also develop chlorotic/yellow rings or line patterns in leaves. RBDV infection also causes ‘crumbly fruit’ in some raspberry cultivars (Fig. 1). This condition arises from the abortion of some drupelets and the uneven development of others and frequently results in an abnormally shaped fruit, which, on picking, may disintegrate into individual drupelets or drupelet clusters. The same disease syndrome can be induced by other unrelated causes, such as poor pollination conditions, infection with other viruses, or genetic aberrations. Chlorosis and crumbly fruit is also observed in blackberry, but usually when RBDV is present in combination with other viruses (Fig. 1(C)–(F)). Additional symptoms have been attributed to RB isolates. In field studies an association was reported between infection with RBDV-RB and premature defoliation of fruiting canes, decreased vigor, leaf curling, necrosis, and death of laterals, and increased winter kill in some red raspberry cultivars that are immune to S isolates. In commercial fields, RBDV rarely occurs in single infections. The virus name comes from symptoms in a plant that was co-infected with RBDV and Black raspberry necrosis virus. In single infections RBDV does not cause bushiness or dwarfing symptoms. In mixed infections with Raspberry leaf mottle virus the titer of RBDV in ‘Meeker’ red raspberry was incre ased approximately 400-fold over the titer of RBDV in single infections. In mixed infections of RBDV with Raspberry leaf mottle virus or Raspberry latent virus, raspberry plants growth was significantly reduced during the establishment year compared to plants infected singly with RBDV. The mixed infections also resulted in smaller fruit, more severe crumbliness of the fruit and lower yields, than in plants singly infected with RBDV in the first two fruiting years. RBDV was detected in cultivars of white-fruited and red-fruited grapevines (Vitis vinifera) in Slovenia and Hungary. RBDV infected ‘Laški Rizling’ exhibited line pattern and mild leaf yellowing. In most cultivars RBDV infections were symptomless. RBDV is reported to be widespread in grapevines in Slovenia. Of 390 seedlings grown from seed collected from RBDV infected cv. Laška Rizling, all tested negative for RBDV, suggesting a very low levels or no seed transmission in grapevines. It would be curious to examine horizontal transmission in grapevines using pollen from infected plants. RBDV isolates from grapevine spread systemically in Chenopodium murale, which has not been observed with isolates from Rubus spp.

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Fig. 2 Electron micrograph of a purified preparation of RBDV Particles stained with uranyl acetate. Scale¼100 nm. Photographs courtesy of R.R. Martin.

PrLBaV was sequenced from plants exhibiting cream colored blotches, some ringspots, leaves with coalesced blotches leading to almost completely cream colored leaves and in some cases necrotic spots within the cream colored blotches. There was not any testing for the PrLBaV in a large number of symptomatic and asymptomatic plants to document an association between the symptoms and the presence of the virus. BCIV was sequenced from blackcurrant ‘Belaruskaya Slodkaya’ and BCLCaV was sequenced from blackcurrant ‘Baldwin’ both exhibiting a general chlorosis of the leaves.

Experimentally Infected Plants RBDV has been transmitted by graft inoculation from Rubus to perennial plants in other genera. It infected Fragaria vesca and Prunus mahaleb seedlings without any symptoms, and induced pronounced chlorotic vein-banding and line patterns in Cydonia oblonga cv. C7–1. RBDV has a moderately wide host range in herbaceous test plants infecting over 50 plant species in 12 families of dicots. The idaeovirus from blackcurrants has been transmitted mechanically to Nicotiana occidentalis and N. benthamiana. There is no information on mechanical transmission of PrLBaV to herbaceous hosts.

Virion Structure Chenopodium quinoa is the preferred propagation host for RBDV. Purified virus particles are about 33 nm in diameter, and appear quasiisometric because they tend to collapse and deform on the electron microscope grid (Fig. 2). They appear more spherical in shape when fixed with formaldehyde. Particle preparations have A260/A280 of 1.62, suggesting that RBDV particles contain about 24% RNA. Purified preparations of RBDV particles typically yield a single major protein estimated by polyacrylamide gel electrophoresis to have a molecular weight of ca. 30 kDa. The particles contain three species of single-stranded RNA (ssRNA) of approximately 5.5 kb (RNA1), 2.2 kb (RNA2), and 1.0 kb (RNA3). RNA3 is a sub-genomic of RNA2 (Fig. 3). The particles are unstable and readily disrupt in the presence of 0.01% sodium dodecyl sulfate, indicating that they are stabilized by protein–RNA interactions.

Genome The complete nucleotide sequences have been determined for the two genomic RNAs (RNA1 and RNA2) of RBDV isolates S and RB, BCLCaV, and PrLBaV. The genome organization of idaeoviruses is shown in Fig. 3, at this time the 12 kDa putative ORF has only been predicted in RBDV. RNA1 of RBDV (R15 strain), PrLBaV and BCLCaV are 5,449, 5377, and 5349 nt long, respectively. RNA1 codes for a multi-functional protein of B19 kDa, which contains the methyl-transferase (MTR), helicase (HEL), and RNAdependent RNA polymerase (RdRp) domains. RNA2 of RBDV (R15 strain), PrLBaV, and BCLCaV are 2,231, 2348, and 2280 nt long, respectively. RNA 2 codes for two proteins, a putative movement protein (MP) and a coat protein (CP) with a molecular masses of about B39 kDa and B30 kDa, respectively. The CP of RBDV is expressed via a sub-genomic RNA (RNA3, 946 nt), and this is likely the case for PrLBaV and BCLCaV. This predicted size corresponds to that of protein obtained from purified virus particles and that of the main polypeptide made by in vitro translation of RNA3. Fig. 4 shows phylogenetic relationship among the idaeoviruses and two closely related viruses. RNA1 also contains a small ORF in a different translation frame that overlaps with the 190 kDa protein, which if expressed could code for a 12 kDa protein. The position of this ORF corresponds to that of the 2b gene in the tripartite genome of Cucumber

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Fig. 3 Schematic showing the genome organization of RBDV. RNA molecules are represented by solid lines, the yellow blocks show the open reading frames (ORFs). The relative positions of the functional domains of the polyprotein encoded by RNA1 are MET (methyl-transferase), HEL (helicase) and RdRp (RNA-dependent RNA polymerase), RNA2 codes for the MP (movement protein), and RNA 3, a sub-genomic of RNA 2 codes for the CP (coat protein). PrLBaV and BCLCaV have similar organizations but lack the 12K putative open reading frame.

Fig. 4 Phylogenetic relationship of the idaeoviruses Raspberry bushy dwarf virus (RBDV), Privet leaf blotch associated virus (PrLBaV) and Blackcurrant leaf chlorosis associated virus (BCLCaV), the Japanese holly fern mottle virus (JHFMoV) is also included in the polymerase protein since its RNA1 resembles those of idaeoviruses, and Blackberry chlorotic ringspot virus (BCRV), since idaeoviruses have nearest affinities to ilarviruses. Phylogenetic analysis was performed using the coat proteins (a), putative movement proteins (b) and polymerase proteins (c), the BCRV was not included in the polymerase phylogeny due to the multiple functional domains on two RNAs. The trees were generated by the maximum-likelihood PhyML 3.0, using the default parameters. Branch lengths are proportional to genetic distances between sequences and the scale represents substitutions per amino acid site. RBDV, red raspberry isolate (NC_003739, NC-003740); RBDV-Vitis, grapevine isolate (EU796086); RBDV-R.multi, Rubus multibracteatus isolate (DQ120126); BCLCaV-Can, Canadian isolate (KX838923, KX838924); BCLCaV-OR, NCGR isolate (KY399998, KY399999); PrLBaV (NC_031341, NC_031342); JHFMoV, (FJ907327); and BCRV (NC_011554).

mosaic virus (CMV) that encodes a 10 kDa polypeptide and overlaps the C-terminal 69 codons of ORF 2a that encodes the RNA polymerase protein. The CMV 2b protein is important in long distance systemic movement and the suppression of post- transcriptional gene silencing, but it is not necessary for infectivity. However, experiments failed to demonstrate the activity of the 12 kDa RBDV protein in suppressing post-translational gene silencing. There is a report of RNA1 of RBDV replicating and moving systemic in the absence of RNA2, which suggests the expression of ORF 1a if it serves a long distance movement function similar to that of ORF 2a in CMV. There are similarities among the 50 - or 30 terminal non-coding sequences of each of the genomic RNA species of RBDV, PrLBaV and BCLCaV. The 50 termini of both RNAs of the three viruses start with the same four nt 50 -AUAU-30 . The 30 terminal regions for RNA1 and RNA2 of each virus share 5-AACCCC-30 , with BCLCaV having an extra C residue. Additionally, the 30 termini of RNA1 and RNA2 of each of the three viruses form four similar stem-loop structures. BCLCaV lacks the putative 12 kDa ORF on RNA1, An isolate of RBDV from Rubus glaucus in Ecuador and the BCLCaV from Canada each had a concatenated form of RNA2 that contained an inverted CP gene. The RBDV isolate was cloned and sequenced from a dsRNA template using standard DOP-PCR and Sanger sequencing combined with RACE to obtain the terminal sequences. The dsRNA pattern on an agarose gel showed an unexpected band of about 3.2 kbp. The BCLCaV was sequenced using high throughput sequencing from a dsRNA template. The presence of this unusual RNA in two idaeoviruses suggest that there may be some secondary structure that leads to a rare formation of overhanging sequence that can be transcribed to yield this inverted repeat. In the case of RBDV it was found in a single plant,

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Table 1 List of members of the genus Idaeovirus (top table). Percent amino acid identity between coat proteins and movement proteins sequences of Raspberry bushy dwarf virus (RBDV), Privet leaf blotch associated virus (PrLBaV), and Blackcurrant leaf chlorosis associated virus (BCLCaV) (bottom table) Species names

Acronym

RNA1 accession #

RNA2 accession #

Raspberry bushy dwarf virus Privet leaf blotch associated virus Blackcurrant leaf chlorosis associated virus Tentative members Japanese holly fern mottle virus

RBDV PrLBaV BCLCaV

NC_003739 NC_031341 NC_034408

NC_003740 NC_031342 NC_034390

JHFMoV

NC_013133

NC_013134

Virus

Coat Protein

Movement Protein

RBDV PrLBaV BCLCaV

RBDV

PrLBaV

BCLCaV

RBDV

PrLBaV

BCLCaV

100

26 100

23 61 100

100

26 100

21 61 100

suggesting it may be maintained in vegetatively propagated material, but not encapsidated and therefore not pollen transmitted. It is not known if this is a common occurrence with BCLCaV since there are only two sources that have been sequenced.

Viral Proteins The Coat Protein The CPs of the idaeoviruses show no striking similarities with the CPs of other virus genera. An alternative to this sequence-based alignment program is to use secondary structure predictions to produce an alignment. This type of prediction with the RBDV CP predicted a long N-terminal sequence containing two a-helices and followed by a region containing eight b-sheets. If correct, this arrangement would resemble that of the CPs of many viruses that have isometric particles. The N-terminal parts of the CPs of RBDV and of viruses in the family Bromoviridae are relatively rich in basic aa, which are thought to be internal in virus particles and to be involved RNA encapsidation. The CPs of the ilarviruses and Alfalfa mosaic virus (AMV) are involved in ‘genome activation’ and required for virus replication. This phenomenon has also been reported for RBDV, where combining infectious clones of RNA1 and RNA2 gave very low level of infectivity but the addition of RNA3 greatly increased virus replication. With the ilarviruses and AMV addition of CP without adding the sub-genomic for the CP also was very effective at enhancing replication. RNA-binding domains have been shown to play a role in genome activation. In recent analyses using BindN several putative RNA-binding domains were identified in the CP of PrBLaV and similar domains were predicted in the CP of RBDV, though their involvement in genome activation has not been documented (Table 1).

The Movement Protein RNA2 of RBDV corresponds in size and gene content to RNA3 of viruses in the family Bromoviridae. The ORF for the 39 kDa protein therefore corresponds in position to the ORF for the putative MP of members of the Bromoviridae. A limited sequence identity was detected in multiple alignment tests between the 39 kDa protein and the putative or actual MPs of several viruses. In further alignment tests of this sort, Mushegian and Koonin have detected a match between part of the 39 kDa protein sequence and a motif conserved in the MPs of some other viruses, such as Red clover necrotic mosaic virus (Tombusviridae family, Dianthovirus genus) and Soil-borne wheat mosaic virus (Virgaviridae family, Furovirus genus). These proteins were proposed to belong to a ‘30 kDa superfamily’ of MPs. With PrLBaV the localization of the putative MP (38 kDa) in planta was studied by fusing green fluorescent protein (GFP) to its C-terminus. This fused protein was agro-infiltrated into leaves of N. benthamiana. When examined using confocal laser microscopy the GFP was accumulated at the plasmodesmata, which has been observed for other MPs. This provides strong evidence that the 38–39 kDa proteins coded by RNA2 are indeed MPs of the idaeoviruses and involved in virus trafficking.

The Replicase Protein The sequence of the 190 kDa protein has similarities with replicase proteins of viruses in the Alphavirus superfamily. PFAM analysis identified three conserved domains in the predicted protein of the three idaeoviruses (RBDV, PrLBaV, and BCLCaV) coded by ORF1. The viral methyltransferases (MTR) belong to PF01660, which if found throughout the Alphavirus superfamily. The viral helicases (HEL) belong to PF01443 and the RdRp belong to PF00978. There was homology between parts of the idaeovirus replicase proteins and parts of the 183 kDa protein of Tobacco mosaic virus, but they showed greater similarity with the 90 kDa

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protein (RdRp) of AMV (encoded by RNA2), and 125 kDa protein (HEL and MET), which are encoded by RNA1 of AMV. RBDV RNA1 therefore corresponds in size and gene functions to RNA1 plus RNA2 of the tripartite viruses in the family Bromoviridae.

Diagnosis and Detection In Rubus, RBDV may induce yellowing, line patterns, or crumbly fruit, but such symptoms are unreliable for diagnosis because they may be due to infection with other disease agents, poor pollination, environmental conditions or poor nutrient management. Also, with many cultivars of raspberry (red and black) and blackberry RBDV is often asymptomatic. Detection and diagnosis therefore depend on bioassays, serological tests and/or reverse transcription PCR assays. RBDV can be detected reliably by ELISA. Many isolates of RBDV are readily detected by mechanical transmission to C. quinoa test plants, but this is most effective early in the season and is not as reliable as serological or RT-PCR assays. As the distribution of RBDV in individual Rubus plants can be erratic, leaves/leaflets for mechanical transmission or ELISA should be taken and pooled from at least three different nodes. For mechanical transmission newly emerging leaves should be used. For ELISA and RT-PCR, the youngest but fully expanded leaves re the optimal tissue for testing.

Serological Testing Polyclonal antisera raised to S isolates have a moderate titer of ca. 1/512 in gel double diffusion tests. In agarose gel doublediffusion serological tests spur formation was used to document serological differences between S isolates from B isolates. Monoclonal antibodies to RBDV were able to distinguish three different epitopes of a Canadian isolate of RBDV. Each of the three monoclonal antibodies reacted with all isolates of RBDV from raspberry and blackberry. One of the monoclonal antibodies did not react with the isolates from grape or from Rubus multibracteatus. These monoclonal antibodies reacted in double antibody sandwich ELISA (DAS-ELISA) with all tested isolates from red raspberry, including B, S, and RB isolates. They are usually used in combination with a polyclonal antibody for trapping the virus onto the plate, the monoclonal antibodies as the second antibody after the virus has been trapped, and then followed with a goat anti-mouse conjugated antibody. This combination of antibodies are available commercially from several companies and can be used to detect RBDV with minimal equipment. There are not antibodies available for PrLBaV or BCLCaV, so testing for these two viruses is limited to biological assays or RT-PCR.

Reverse Transcriptase – PCR (RT-PCR) There are more than 75 partial or full sequences of RBDV in GenBank, which is useful for designing detection primers that can capture the diversity of the virus. Detection primers have been designed for end-point RT-PCR and for quantitative RT-PCR. For most applications end-point PCR is used. Also, primers have been designed specific for RNA1 and for RNA2. There has been a report of RBDV RNA1 replicating and spreading systemic in raspberry plants. In this case, the ELISA tests and the end-point RT-PCR based on RNA2 sequences were negative, while the RT-PCR based on RNA1 was positive. It is not known if this is a common occurrence or was a rare event, but care should be taken to include some testing with primers specific for RNA1.

Therapy Unlike many other viruses infecting Rubus, RBDV is not readily eradicated from Rubus plants by meristem tip-culture alone. However, heat treatment for several weeks at 361C followed by meristem tip-culture can be used to eliminate RBDV from infected plants. Even with this combination, virus elimination of RBDV can be challenging. Cryotherapy has been used successfully to eliminate RBDV from raspberry, but again it was not very efficient. Chemotherapy combined with thermal therapy and meristem tip culture has shown some success in eliminating RBDV. RBDV appears to move very readily cell-to-cell and therefore can move into the meristematic dome more efficiently than most viruses.

Prevention and Control The most important component of virus control in vegetatively propagated crops is to start with virus-tested planting stocks. This requires a clean plant production system. Such a system includes: (1) A facility to produce clean plants, hold the top-tier or G1 plants generally under protected culture (screenhouses to minimize vectors) or in long-term tissue culture at 41C (Fig. 5), and that can retest G1 plants on a regular schedule to ensure they remain clean; (2) A certification scheme that performs audit testing and ensures best management practices are followed during the multiple stages of increase (G2 and G3 blocks) required between the G1 (Foundation Block) and the certified plants sold to growers, (3) Increase blocks that are monitored by the certifying agency and tested for viruses and other targeted pathogens as outlined in the certification standard; (4) A system to track the certified plants under the certification scheme so that disease issues can be traced-back or traced-forward as needed.

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Fig. 5 Plants of ‘Columbia Star’ blackberry in gas permeable sealed bags. These plants can be held at 41C for up to three years without any maintenance or risk of reinfection. Photographs courtesy of R.R. Martin.

Because RBDV is transmitted in association with infected pollen, the only methods of controlling it in crops are to grow resistant cultivars, or to plant healthy susceptible cultivars in isolation from possible sources of infection. If the virus is present in native Rubus spp., isolation from infected sources can be challenging in many growing regions. All black raspberry, blackberry, and blackberry/raspberry hybrids tested were susceptible to the S strains of RBDV. Many red raspberry cultivars are resistant, possibly immune, to S isolates (the most widespread isolates worldwide). This resistance is conferred by the presence of a single dominant gene, Bu. All Bu-containing cultivars are susceptible by grafting with RB isolates, though the cultivars, ‘Heritage’, ‘Haida’, ‘Rannaya Sladkaya’ were difficult to infect and did no become infected when exposed to the RB isolates under field conditions. There are cultivars that show ‘field resistance’ and are slower to become infected in the field to either the S or RB isolates. In some cases, cultivars that are susceptible to grafting infection with the S isolates remained RBDV-free in the field up to 20 years, even under conditions where adjacent plantings were nearly 100% infected after 5 years. The mechanism for this has not been studied, but if high fruit quality can be combined with slow to become infected, such cultivars could have a place in production systems. Breeding for field resistance to the RB strain may be a more realistic goal since resistance to this strain has not been identified. RBDV susceptibility has resulted in some cultivars with excellent fruit quality to be abandoned by growers. Studies on this form of resistance indicated that its mode of inheritance was complex but seemed to depend on the presence of gene Bu, together with a second resistance component whose inheritance was probably multigenic. In field studies, some raspberry cultivars, including some that do not contain gene Bu, are more resistant than others to natural infection with RB isolates. ‘Meeker’ red raspberry has been developed with resistance to RBDV that makes use of RNAi. The plants were free of RBDV when challenged in the field for six years under extreme disease pressure when all 202 control ‘Meeker’ plants in the same test plots were infected after three years in the field. The resistance was stable and effective against graft inoculations that were repeated three times and the transgenic raspberry produced fruit of quality indistinguishable from wild-type ‘Meeker’. Though the resistance is effective, the plants are on hold and applications for approval for human consumption to the Food and Drug Administration in the U.S. have not been made, pending a change in public perception on the use of this technology. Additionally, there has been 20 years of breeding since these transgenic lines were developed, and the standard for fruit quality and yield has surpassed that of ‘Meeker’.

Relationships With Other Viruses Idaeoviruses resemble viruses in the genus Ilarvirus, family Bromoviridae in having quasi-spherical particles that are easily distorted in preparations viewed in an electron microscope. Also, they are transmitted vertically and horizontally in association with pollen as are the ilarviruses. RNA1 of idaeoviruses contains the HEL, MET, and RdRp functions, while in the ilarviruses, RNA1 contains the HEL function and RNA2 contains the MET and RdRp functional domains. RNA2 of idaeoviruses resemble RNA3 of ilarviruses in the arrangement and sizes of its encoded gene products, and the generation of a 30 terminal sub-genomic RNA (sgRNA). RBDV is serologically unrelated to all recognized ilarviruses tested. Also, RBDV differs from llarviruses in the sedimentation behavior of

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its particles, in the number and sizes of its RNA molecules, and in having a bipartite genome. Taken together, these properties distinguish idaeoviruses from all other well-characterized viruses and support their status as a separate genus. The genus Idaeovirus has not been assigned to a family, but has many molecular properties in common with members in the family Bromoviridae, the major difference being that the multi-functional replicase protein of idaeoviruses is coded one RNA whereas in the family Bromoviridae these functional domains are coded by two RNAs. The idaeoviruses have bipartite genomes whereas the six genera in the Bromoviridae have tripartite genomes.

Further Reading Derrick, K.S., Beretta, M.J., Barthe, G.A., 2006. Detection of an idaeovirus in citrus with implication as to the cause of citrus blight. Proceedings of the Florida State Horticultural Society 119, 69–72. ICTV Virus Taxonomy, 2018. Release. Available at: https://talk.ictvonline.org/taxonomy/ (accessed 08.07.2019). Isogai, M., Yoshida, T., Nakanowatari, C., Yoshikawa, N., 2014. Penetration of pollen tubes with accumulated Raspberry bushy dwarf virus into stigmas is involved in initial infection of maternal tissue and horizontal transmission. Virology 452–453, 247–253. James, D., Phelan, J., 2017. Complete genome sequence and analysis of Blackcurrant leaf chlorosis associated virus, a new member of the genus Idaeovirus. Archives of Virology 162, 1705–1709. Jones, A.T., Mayo, M.A., 1998. Raspberry bushy dwarf virus. Descriptions of Plant Viruses 360, 6. Available at: https://www.dpvweb.net/dpv/showdpv.php?dpvno=360 (accessed July 2019). MacFarlane, S.A., McGavin, W.J., 2009. Genome activation by Raspberry bushy dwarf virus coat protein. Journal of General Virology 90, 747–753. MacFarlane, S.A., 2011. Genus Idaeovirus. In: King, A.M.Q., Adams, M.J., Carstens, E.B., Lefkowitz, E.J. (Eds.), Virus Taxonomy: Ninth Report of the International Committee on Taxonomy of Viruses. London: Elsevier Academic Press, pp. 1173–1175. Martin, R.R., MacFarlane, S., Sabanadzovic, S., et al., 2013. Viruses and virus diseases of Rubus. Plant Disease 97, 169–182. Navarro, B., Loconsole, G., Giampetruzzi, A., et al., 2017. Identification and characterization of Privet leaf blotch-associated virus, a novel idaeovirus. Molecular Plant Pathology 18, 925–936. Pleško, I.M., Marn, M.V., Širca, S., Urek, G., 2009. Biological, serological and molecular characterization of Raspberry bushy dwarf virus from grapevine and its detection in the nematode Longidorus juvenilis. European Journal of Plant Pathology 123, 261–268. Quito-Avila, D.F., Ibarra, M.A., Alvarez, R., Peralta, E.L., Martin, R.R., 2014. A Raspberry bushy dwarf virus isolate from Ecuadorean Rubus glaucus contains an additional RNA that is a rearrangement for RNA2. Archives of Virology 159, 2519–2521. Thekke-Veetil, T., Ho, T., Postman, J.D., Tzanetakis, I.E., 2017. Characterization and detection of a novel idaeovirus infecting blackcurrant. European Journal of Plant Pathology 149, 751–757. Valverde, R.A., Sabanadzovic, S., 2009. A novel plant virus with unique properties infecting Japanese holly fern. Journal of General Virology 90, 2542–2549.

Ilarviruses (Bromoviridae) Aaron Simkovich, Agriculture and Agri-Food Canada, London, ON, Canada and The University of Western Ontario, London, ON, Canada Susanne E Kohalmi, The University of Western Ontario, London, ON, Canada Aiming Wang, Agriculture and Agri-Food Canada, London, ON, Canada r 2021 Elsevier Ltd. All rights reserved. This is an update of K.C. Eastwell, Ilarvirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00642-7.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein ELISA Enzyme-linked immuno-sorbent assay ER Endoplasmic reticulum kb Kilobases; the size of a ssDNA or ssRNA molecule kDa Kilodaltons; the size of a protein MP Movement protein NGS Next generation sequencing nm Nanometer(s)

Glossary Bicistronic A viral RNA that encodes two genes which are often separated by a non-coding intergenic region. Cross protection It describes a phenomenon in that an initial infection of a host plant with a mild strain of a virus induces resistance in that plant to the infection of another, closely related virus potentially protecting the plant from disease caused by a more virulent isolate. Genome activation Ilarviral genomic RNAs alone are unable to establish infection in plants, unless the coat protein is present. This function of the coat protein is termed genome activation and is specific for ilarviruses and closely related alfamoviruses. The event is triggered by binding of the coat protein RNA binding domain to the 30 terminus of genomic RNA. RNA silencing A fundamental, evolutionarily conserved and sequence-specific mechanism that is triggered by double-stranded RNA and regulates gene expression in

ORF Open reading frame(s) PCR Polymerase chain reaction RBD RNA binding domain RdRp RNA-dependent RNA polymerase RT-PCR Reverse transcription polymerase chain reaction sgRNA Sub-genomic RNA TLS Transfer RNA-like structures UTR Untranslated region VIGS Virus-induced gene silencing VRC Virus replication complexes

eukaryotes. It is a primary antiviral defense mechanism in plants and other living organisms. RNA silencing suppressor A countermeasure to RNA silencing, often a protein encoded by a virus which interrupts a single, or multiple steps in the RNA silencing pathway such as binding to small interfering RNA and thereby preventing their incorporation into the RNA induced silencing complex. Sub-genomic RNA A segment of RNA generated from a genomic RNA via an international promoter that has the same 30 end as the genomic RNA, but has a deletion at the 50 end. The sub-genomic RNA makes it possible to efficiently translate the downstream open reading frame of the genomic RNA. Tripartite genome A viral genome consisting of three genomic fragments, which are encapsidated into three separate virions.

Introduction Ilarviruses are a group of isometric and labile viruses that are distributed worldwide and infect many agriculturally relevant herbaceous and woody hosts including fruit trees, vegetables, and ornamentals. For long time, the economic importance of ilarviruses has been underestimated as they are often considered as latent pathogens, particularly when they infect fruit trees. Since 1990s, a large number of ilarviruses have been molecularly identified, and this group of viruses has received more and more attention. In this article, we briefly discuss the classification, virion structure, genomic organization, and infection cycle of ilarviruses, summarize the current knowledge of their replication mechanisms and transmission modes, highlight the control strategies, viral symptoms and pathogenesis of the viruses, and finally introduce diagnosis of ilarviruses.

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Table 1

A list of recognized species in the genus Ilarvirus, family Bromoviridae

Subgroup Subgroup 1

Subgroup 2

Subgroup 3

Subgroup 4 Unassigned

Species

Acronym

GenBank accessions

Ageratum latent virus Blackberry chlorotic ringspot virus Parietaria mottle virus Privet ringspot virus Strawberry necrotic shock virus Tobacco streak virus

ALV BCRSV PMoV PrRSV SNSV TSV

JX463341 DQ091194 AY496069 KT290040 AY743591 U75538

Asparagus virus 2 Citrus leaf rugose virus Citrus variegation virus Elm mottle virus Lilac ring mottle virus Spinach latent virus Tomato necrotic streak virus Tulare apple mosaic virus

AV-2 CiLRV CVV EMoV LiRMoV SpLV TomNSV TAMV

EU919667 U17726 EF584665 U34050 EU919669 U93193 KT779205 AF226161

Apple mosaic virus Blueberry shock virus Lilac leaf chlorosis virus Prunus necrotic ringspot virus

ApMV BlShV LLCV PNRSV

AF174585 KF031038 FN669168 AF278535

Fragaria chiloensis latent virus Prune dwarf virus

FCiLV PDV

AY707771 AF277662

American plum line pattern virus Humulus japonicus latent virus

APLPV HJLV

AF235165 AY500237

Classification The Ilarvirus genus was first recognized during the third meeting of the International Committee on the Taxonomy of Viruses in 1975. The genus, along with five other genera including Alfamovirus, Anulavirus, Bromovirus, Cucumovirus and Oleavirus, belongs to the Bromoviridae family. Among the six genera, Ilarvirus is particularly closely related to Alfamovirus. The name “Ilarvirus” is in fact a siglum derived from isometric labile ringspot viruses, describing several characteristics of ilarviruses: the shape of virions, the fragility or labile nature of viral particles and lastly, the frequently observed ring spotting symptom on infected hosts. Currently, the genus consists of 22 recognized members (Table 1). Previously ilarviruses were sub-grouped mainly based on serological properties with antibodies against the coat protein (CP). Accumulated sequence data support the serological relationships for most ilarviruses. However, extensive phylogenetic analyses also identify some species that had been inappropriately classified possibly due to conserved amino acid (aa) sequences and secondary structures in their CPs. Currently, the 22 recognized ilarviruses have been phylogenetically classified into four sub-groups in addition two unassigned viruses (Table 1, Fig. 1).

Virion Structure Virions of ilarviruses are most commonly found as quasi-isometric shapes (Fig. 2(B)). The capsid of the virus is comprised of 180 coat protein (CP) units arranged in a T ¼ 3 symmetry, ranging in diameter from 26 to 35 nm. Additionally, some members, such as Prune dwarf virus (PDV) and Prunus necrotic ringspot virus (PNRSV) also produce bacilliform particles with a diameter of 18–26 nm and 30–85 nm in length. As the virion contains the encapsulated genomic RNA, the size of virions is often related to the length and amount of RNA contained. The regions of the CP in some members is implicated in the formation and degree of lability of the virion.

Genome Members of the Ilarvirus genus have a tripartite genome consisting of three separate positive-sense single-stranded RNA molecules, i.e., RNA1, RNA2 and RNA3, all of which bear a 50 -7-methyl-G cap structure but lack a poly A tail. The 30 ends of genomic RNAs code for hairpin loop structures potentially forming intricate pseudoknots. The genomic organization of Tobacco streak virus (TSV), the type member of the Ilarvirus genus is presented in Fig. 2(A). Among three ilarviral genomic RNAs, RNA1 is the largest

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Fig. 1 Phylogram showing nucleotide sequence relationships of ilarvirus RNA dependent RNA polymerase (ORF2a) with Barley stripe mosaic virus (BSMV) included as an outgroup (Accession NC_003478). Subgrouping (shown at right) is based on phylogenetic data, host range and serology. *Species unassigned to subgroups. Numbers at branch nodes shows percentage of bootstrap replicates in which that branch occurs. Abbreviations and sequence accession numbers are listed in Table 1.

Fig. 2 (A) The genome structure of Tobacco streak virus, the type member of the Ilarvirus genus. Each single stranded positive sense RNA genomic fragment is shown and encoded proteins are shown as boxes with approximate nt and aa lengths, respectively. The complete genome of TSV isolate WC has a total length of 8622 nt. The P1 protein of TSV has 1094 aa with a molecular mass of 123.4 kDa. Encoded from ORF2a, the P2 protein has a sequence length of 800 aa and a mass of 91.5 kDa. The 2b protein has a sequence length of 205 aa, and a mass of 22.4 kDa. The movement protein (MP) of TSV has a length of 289 aa and a mass of 31.6 kDa. The coat protein (CP) encoded by ORF3b located in sgRNA4 is 237 aa in length and has a mass of 26.2 kDa. Each RNA fragment has a 50 cap shown as C and 30 tRNA-like structure shown as *. The large arrow denotes the transcription of sub-genomic RNA4. (B) A transmission electron micrograph showing virions of Prunus necrotic ringspot virus (PNRSV). Photograph courtesy of the Rothamsted Experimental Station, UK. The bar represents 100 nm.

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genomic component, and contains a single open reading frame (ORF1), which encodes the P1 protein. This protein contains both methyltransferase (near the N-terminus) and helicase (near the C-terminus) domains. P1 may also contain a transmembrane domain at the C-terminus. It has been suggested that it is the P1 protein that recruits and anchors the viral RNA to the assembly site to support virus replication. Consistent with this assumption, subcellular localization studies suggest that P1 is associated with membranes of the vacuole, the endoplasmic reticulum (ER) and chloroplasts. Therefore, P1 is considered as an essential component of the viral replication complex (VRC). In subgroups 1 and 2, RNA2 is bicistronic with ORF2a and ORF2b. ORF2a encodes the P2 protein, which is the putative RNA dependent RNA polymerase (RdRp), whereas ORF2b codes for the 2b protein. P2 contains several conserved domains of RdRp such as the RNA binding domain and the GDD motif. Therefore, P2 is believed to be part of the VRC and directly responsible for biosynthesis of viral RNA. The 2b protein was originally proposed to be involved in systemic infection and was later proven to be a suppressor of post-transcriptional gene silencing to counteract against RNA silencing, a primary host antiviral defense mechanism. Members of subgroups 3 and 4 do not possess ORF2b. The third fragment, RNA3 is bicistronic and contains two ORFs, ORF3a and ORF3b. The former encodes a movement protein (MP), and the latter is expressed via a sub-genomic fourth RNA fragment (sgRNA4) for the translation of the CP. The ilarviral MP is a member of the 30K MP superfamily. Members of this superfamily are known to locate to the plasmodesmata (PD) and alter size exclusion limits of these cellular channels to assist virus cell-to-cell movement. The CP is a multifunctional protein and plays essential roles in different stages of the virus infection cycle such as virion formation and replication. A zinc finger motif has been identified in CPs of many plant viruses including some ilarviruses such as TSV and is presumed to be associated with nucleotide binding and virion formation. However, the fact that some ilarviruses such as PDV lack this motif supports the notion that this motif is not an absolute requirement for virion formation.

Infection Cycle The infection cycle of ilarviruses may be artificially divided into several stages. As plant viruses usually have a small genome with limited coding capacity, they must hijack host pathways and factors to support their infection. Upon entry into host cells, ilarviruses undergo uncoating to remove the CP shell to expose the viral genome to the host translation machinery for translation on the rough ER. Subsequently, the P1 protein may target the membranes of preferred organelles such as the ER, vacuole and/or tonoplast and induces the formation of spherules or small vesicular structures where VRCs are assembled. The VRC catalyzes biosynthesis of the minus (  ) strand RNA using the viral genomic RNA as template and further multiplicate the ( þ ) strand RNA using the newly synthesized (  ) RNA as template. The ( þ ) and (  ) RNA strands are biosynthesized in an asymmetrical manner. During AMV replication, the ratio of ( þ ) RNA versus (  ) RNA is approximately 100:1. The progeny ( þ ) RNAs are encapsidated by CPs to form virions. The newly synthesized RNA or virions may interact with MP, and the ribonucleoprotein complexes are transported intracellularly to the PD where they move from the primarily infected cells to neighboring cells. MP is definitely required for this process. For some ilarviruses, virion is not necessary for virus cell-to-cell spread, but the presence of CP is required for all ilarviruses to move intracellularly. For long-distance movement from the local infection to remote sites of the plant, the virus is transported from the mesophyll via bundle sheath cells, phloem parenchyma, and companion cells into phloem sieve elements and then following the source-to-sink flow of photo-assimilates and unloaded to sink tissues. Ilarviruses are likely transported for the long distance in the form of virions.

Replication Most members in the Bromoviridae family possess transfer RNA-like structures (TLS) in the 30 untranslated region (UTR) of their genomic RNAs, and these TLS regions interact with the cap at the 50 end to circularize the viral RNA and promote the translation process. Different from these viruses, alfalfa mosaic virus (AMV) and ilarviruses lack TLS in their genomes. In this case, the CP is required to establish infection possibly by activation of viral protein translation and regulating transcription of new ( þ )/(  ) viral RNA, a phenomenon termed genome activation. In lieu of TLS, this requirement of CP for virus replication is now a distinguishing feature of both Alfamovirus and Ilarvirus genera. Viral replication mechanism has been studied more in depth for alfamoviruses than ilarviruses and due to the common feature of the CP requirement for genome activation has led to analogous conclusions for replication strategies. The process of genome activation is based on interactions between the viral CP and secondary structures found within the 30 UTR of viral RNAs. A highly conserved RNA binding domain (RBD) containing crucial arginine (R) residue(s) is present in the N-terminal CP sequences of AMV and some ilarviruses, and the RNA-binding consensus motif in this RBD has the aa sequence Q/K/R-P/N-T-X-R-S-R/Q-Q/N/S-W/F/Y-A. Additional sequence analyses have identified a second putative RNA binding consensus sequence V(T/S)(R/N)RQ(S/R)RNA(A/R)RAAX(Y/F)R which is also conserved in at least six other ilarviruses. Some ilarviruses such as PNRSV do not have any of the two consensus RNA-binding sequences. However, the CP sequence in these viruses is R-rich. In the CP of PNRSV, there is a stretch of about 20 aa with five R residues that has the capacity to bind to the 30 UTR of RNA3. In addition, the N-terminal region of the CPs of many ilarviruses including PNRSV possess a zinc finger motif that is believed to increase RNA binding affinity. The process of genome activation is not species-specific, and some ilarviruses are able to use CPs from other ilarviruses to activate their genomes. This ability to reciprocally activate infectivity of the different viruses by their CPs can extend to the intergenus level. It

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has also been shown that CP of AMV (an alfamovirus) and that of some ilarviruses can be interchanged to induce genome activation. In the absence of a translated CP, the presence of sgRNA4 is sufficient to result in an active genome as the CP is translated from this sub-genomic RNA. In the absence of sgRNA4 or CP, the transcription of viral RNA is preferred, however the addition of either one of these two elements leads to a shift towards translation. It is suspected that the binding of CP inhibits transcription, and that the RNA secondary structures enable binding by the CP. Complex secondary structures have been identified among sequenced ilarviruses, as the 30 UTRs of genomic RNAs possess conserved single stranded motifs of (A/U)(U/A/G)GC, which flank the regions potentially forming hairpin loop structures acting as binding sites for the CP. It has been suggested that the CP-RNA complex formed in ilarviruses are functionally analogous to the complex formed by poly A tract and the poly A binding protein in eukaryotes which is known to enhance translation of proteins. This has been further supported by the finding that the AMV CP does in fact bind to eukaryotic translation initiator complexes eIF4F and eIFiso4F. Two models have been proposed to explain the role of the CP in the early stages of the ilarviral infection cycle. The conformational switching model proposes that the 30 UTR of AMV and ilarviruses can fold into two separate structures. The first is a pseudoknot which is functionally equivalent to the TLS and promotes the binding of viral RdRp and synthesis of negative strand RNA. The second structure is a linearized series of hairpin loops. This model proposes that early binding of CP causes a conformational change, or switch, unwinding the pseudoknot into to a series of hairpin loops which cannot be bound by the RdRp. This model is supported by several lines of evidence from studies on AMV. The pseudoknot in the 30 UTR is essential for AMV replication and disruption of this structure inhibits viral replication. CP binding to the 30 UTR is inhibited when magnesium is added to stabilize the pseudoknot structures. The CP can bind to mutated 30 UTR sequences which are unable to form pseudoknots even after addition of magnesium. Consistent evidence has also obtained from studies with ilarviruses. For example, the CP of PNRSV is unable to bind to the 30 UTR of genomic RNAs in the presence of magnesium. Overall, this model explains the role of CP in the regulation of RNA synthesis and protein translation. The second model suggests that CP binding to the 30 UTR forms a complex critical for viral replication. This model contradicts the first model by predicting that the structure of the 30 UTR undergoes a different conformational change and CP binding to the 30 UTR creates a more compact structure. Structural analyses show that the unbound 30 UTR of AMV is flexible, supporting this model. When a short peptide identical in sequence to the AMV CP RNA binding domain is bound, the resulting CP-RNA complex is more rigid and compact due to base pairing between the AUGC repeats that flank hairpin loops. Modeling of the CP binding along the entire length of the AMV 30 UTR, shows that the predicted molecule has a compact, rod shaped structure with hairpins protruding from the center. In vitro studies have also shown that in absence of CP, the binding of the RdRp protein to labeled RNA is weak, and addition of CP significantly enhances the binding ability of RdRp to the labeled RNA. Therefore, CP binding to the 30 UTR promotes RNA transcription.

Transmission Ilarviruses have several modes of transmission. The majority of ilarviruses have a relatively limited natural host range, with TSV as an exception which can infect a number of plant species naturally. All ilarviruses can infect a broad range of experimental herbaceous hosts via mechanical transmission. Ilarviruses naturally infecting herbaceous plants are easily transmitted mechanically via accidental damage during farming practices or by insects. Many ilarviruses naturally infect woody hosts such as stone fruit trees. However, the probability of mechanical transmission in the woody hosts is low. Perhaps the most common mode of ilarvirus transmission in these hosts is by vegetative propagation, which is a common horticultural practice. For example, grafting of scions to rootstocks is used for clonal propagation of woody fruit trees. In some cases, it is also used for herbaceous crop production (i.e., tomato, cucumber and watermelon) to take advantage of desirable traits of rootstocks such as disease resistance. In grafting, the use of infected material as either a root stock or scion facilitates the spread of viruses. Another method of vegetative propagation involves the direct planting of cuttings (i.e., alfalfa and hops), or division of crowns (i.e., rhubarb and asparagus). The propagation of new explants from an infected mother plant provides another simple yet common vector for virus spread. The finding that some ilarviruses are spread via seed poses a further challenge as rootstocks are commonly grown from seedlings. The spread of viruses by seed presents a challenge in the production of seed grown crops, and planting of uncertified seed stock has the potential to spread members of this genus at the beginning of a new growing season and even to new agricultural production regions. Aside from transmission by vegetative propagation and seeds, ilarviruses may also transmit via plant-to-plant, i.e., hops, pollen grains and insects. For example, pollinating honeybees have been shown to transmit both PNRSV and PDV. The spread of TSV by thrips is well documented, and several species of thrips have been identified as vectors of ilarviruses. Feeding by thrips that carry virus-contaminated pollen, or feed on leaves covered by virus-contaminated pollen causes mechanical damage to the host plant, providing a point of entry for the virus.

Control Treatment of any infections by plant viruses including ilarviruses is difficult. Therefore, methods to prevent viral infection should be implemented. Cross protection, also known as superinfection exclusion, describes the process in which the exposure to a mild

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virus isolate induces the resistance against a more virulent, closely related isolate. The virus isolate selected for inoculation should be endemic to the growing region, ideally latent and showing minimal impact on the host. Cross protection does not offer broad resistance to unrelated viruses, and in fact, for some viruses, cross protection is only strain specific. A very successful example for the control of plant viruses using cross protection is to protect citrus from the devastating diseases caused by virulent isolate of Citrus tristeza virus, which belongs to the genus Closterovirus in the family Closteroviridae. Cross protection has also been reported to control infections by ilarviruses. Cherry rugose mosaic disease is a serious disease of sweet cherry and the causal agent is a virulent strain of PNRSV. Less virulent isolates of PNRSV have been identified for the control of the disease in sweet cherry. One possible mechanism of cross protection is RNA silencing. It has been suggested that virus infection is involved in the formation of doublestranded RNAs which trigger virus induced gene or RNA silencing (VIGS). VIGS explains why infection by an attenuated version of a virus can induce resistance to the virus isolates sharing similar sequences with the attenuated isolate. VIGS can also be used to silence host genes by insertion of a short fragment of a target gene into the attenuated virus. Recently, using a modified PNRSV, we have successfully silenced a gene in plum that is required for plum pox virus infection. The resulting plum plants show resistance to Plum pox virus. Crop management to minimize risk of infection, and methods of early detection are the effective strategy to combat infections by ilarviruses and other plant viruses. Certification programs are important measures which should be implemented for introduction of viral pathogens. Many countries have programs to certify growing materials (seed, rootstocks, scion cuttings) as being free of viruses after testing. Using certified virus free materials is important when starting both perennial and annual crops. In particular for fruit trees which are propagated over decades in densely populated orchards such as members of the Prunus genus, the use of virus free material is a worthwhile long-term investment. Hygiene practices including cleaning of pruning shears and other implements which contact plants should be routinely performed. When virus infections are found, common practices place an emphasis on sanitation and eradication including the removal and destruction of infected plants and plant material, removal of perennial weeds which may act as alternative hosts near annual growing fields. In response to the spread of viral diseases caused by transport of infected plant materials, some countries have imposed legislative methods to fight plant disease by enforcing strict regulations to restrict the movement of plant material. Control of vectors such as thrips or other insect vectors is another effective approach to minimize spread of insect- and/or pollentransmitted viruses. It is worth mentioning that the introduction of resistance using modern genetic engineering technologies has been demonstrated to be highly effective in the control of plant viruses. This type of resistance operates through RNA silencing by generation of transgenic plants that express a double-stranded RNA fragment sharing the sequence of the target virus. However, due to public concerns and regulatory barriers, this technology has not been widely used for the control of ilarviruses.

Symptoms and Pathogenesis Symptoms induced by ilarviruses vary for different virus-plant combinations. As the name suggests, ringspot is a common symptom found on young leaves of ilarvirus-infected plants (Fig. 3(A)). Ringspots often become necrotic and the necrotic tissue dies leaving a shot-hole symptom pattern. Infection by TSV induces various symptoms in tobacco leaves including vein clearing, streaking, ringspots, stunting, leaf deformation and systemic necrosis. Cucumber plants infected with PDV initially show small chlorotic lesions, then a more severe mottling of leaves and stunted growth (Fig. 3(B)). Ilarvirus-infected plants often undergo an initial shock phase with the development of disease symptoms such as necrotic ringspots, followed by symptom attenuation and eventual recovery on new leaves. On woody plants, this disease process may take a few years. For example, infection of blueberry and cranberry by Blueberry shock virus, which is pollen-transmitted with the aid of honeybees, induces clear symptoms on the entire plant or some branches in the first 1–2 years. Leaves and flowers of infected plants appear blighted, and gradually drop off. After 1–3 years, the infected plants become asymptomatic. Cranberry bushes infected with TSV also show symptom recovery. However, different from the typical recovery phenomenon in which the abatement of symptoms is associated with antiviral RNA silencing and decreased viral accumulation (termed viral clearance), symptom recovery following infection by some ilarviruses may be caused by different mechanisms. The molecular determinant for the recovery phenomenon in TSV has been identified. A single nucleotide substitution (A - G) in the intergenic region of RNA3, upstream of the transcription start site of sgRNA4 is responsible for the failure of a TSV isolate to initiate recovery in the host. Plants infected with this recovery deficient isolate develop disease symptoms which never subside. Interestingly, the viral titer in distal plant tissues is lower than that in the corresponding tissues of plants infected with the TSV wild type virus that can induce symptom recovery. These data support the assumption that symptom recovery in plants infected by at least some ilarviruses is not necessary to be related to viral titers. It is possible that ilarvirusinduced RNA silencing downregulates host resistance genes, which may contribute to an infection with abated symptoms where a higher viral titer is maintained. Determinants of pathogenicity have been identified in some ilarviruses such as Asparagus virus 2 (AV-2) and PNRSV. The 2b protein encoded by AV-2 ORF2b acts as a suppressor of systemic RNA silencing to indirectly function as a determinant by counteracting the host plants defense mechanism. A more direct example of a disease determinant came from a study on PNRSV. In this study, full-length infectious cDNA clones derived from a pathogenically aggressive isolate (Pch12) and a mild isolate (Chr3) of PNRSV (Fig. 3(C)) were used to identify determinants of pathogenicity. By reassortment of genomic segments, swapping

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Fig. 3 Ilarvirus-infected herbaceous experimental and natural host plants. (A) Healthy, asymptomatic leaf from sweet cherry (Prunus avium cv. Vogue) compared to leaf from an individual infected with PDV showing leaf deformation and yellowing. (B) Experimental herbaceous host cucumber (Cucumis sativus cv. Wisconsin) showing viral symptoms after being mechanically inoculated with homogenate prepared from PDV infected foliar tissue shown in A. Newly emerging and fully expanded symptomatic first true leaf and symptomatic second true leaf are magnified to show mottling and vein clearing symptoms at 5, 7, and 10 days post inoculation (DPI). (C) Symptoms induced by two infectious cDNA clones of PNRSV on its natural host cherry (Prunus avium cv. Bing) at 18 dpi. The two PNRSV isolates differ in symptom severity Chr3 induces the development of milder symptoms (necrotic spots on young leaves) whereas Pch12 causes severe top wilting and more severe necrosis. (D) Identification of pathogenicity determinants of PNRSV. Herbaceous host cucumber plants (Cucumis sativus cv. Straight Eight) were inoculated with two isolates of PNRSV via agrobacterium mediated infiltration of infectious cDNA clones. The mild isolate Chr3 shows minimal disease however the more aggressive isolate, Pch12 causes stronger disease symptoms in cucumber as well. The chimeric virus formed from the Chr3 isolate of PNRSV by the substitution of the 30 region of RNA1 from Pch12, and the substitution of asparagine to lysine at position 279 in RNA2 shows severe symptom development leading to the identification of pathogenicity determinants in PNRSV. Figs. 3(C and D) reproduced with permission from the American Phytopathological Society.

of partial genomic segments, and site-directed mutagenesis, it was shown that the 30 terminal nt sequence in RNA1 and aa K at residue 279 in RNA2-encoded P2 are the severe virulence determinants and are required for severe virulence and high levels of viral accumulation (Fig. 3(D)). PNRSV RNA1 and RNA2 seem to codetermine viral pathogenicity to adapt to alternating natural Prunus hosts, likely through mediating viral accumulation.

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Diagnosis Diagnosis of an infection by ilarviruses is temporally sensitive as initial shock symptoms subside, making the distinction of healthy and infected plants difficult. Symptoms are easily seen at the beginning of growing seasons before peak seasonal temperatures are reached; young, newly emergent leaves usually display the clearest symptoms. Common diagnosis techniques for other plants viruses are applicable to ilarviruses. Since ilarviruses usually have a wide experimental host range, indexing on herbaceous and woody indicators is an option for detection. The main advantages of this method are the relatively low costs and minimal reagents required. It should be noted that chemical additives such 2-mercaptoethanol, ethylenediaminetetraacetic acid, cysteine hydrochloride, sodium diethyl-dithiocarbamate and polyvinyl-pyrrolidone are routinely added to inoculum preparations. These additives protect the virion from degradation while infected host tissue is homogenized which results in the release of endogenous RNA degrading enzymes, oxidizing agents, and tannins, all compounds that are known to degrade virions or inactivate them. There are several disadvantages to indexing including the long duration of experiments, the labor intensive nature of these studies and the low specificity. Generation of antibodies specific to some ilarviruses CPs has led to the commercial availability of serological diagnostic kits based on enzyme-linked immunosorbent assay (ELISA), which is fast, accurate, highly sensitive and specific, compared to other diagnostic methods. Disadvantages of serological diagnostic kits are that they are often highly specific, and are not sensitive to detect low titer of viruses. Nucleic acid-based detection techniques such as reverse transcription polymerase chain reaction (RT-PCR) serve as more powerful diagnostic tools. Multiplex PCR can be used to detect multiple viruses simultaneously with greater sensitivity and specificity compared to serological techniques. Indeed, multiplex PCR has also been shown to allow the detection of several ilarviruses simultaneously. Another powerful molecular technique is next generation sequencing (NGS). This method requires the greatest amount of time for preparation and is the most expensive option. However, NGS is a broad detection tool and additionally provides the user with incredibly extensive sequence data. Such sequencing data from infected hosts have been used to identify causal agents of plant disease including novel viruses, and even to obtain full-length genome sequences of some viruses.

Further Reading Bujarski, J.J., Gallitelli, D., García-Arenal, F., et al., 2019. ICTV virus taxonomy profile: Bromoviridae. Journal of General Virology 100 (8), 1206–1207. doi:10.1099/jgv.0.001282. Cui, H., Hong, N., Wang, G., Wang, A., 2013. Genomic segments RNA1 and RNA2 of Prunus necrotic ringspot virus codetermine viral pathogenicity to adapt to alternating natural Prunus hosts. Molecular Plant-Microbe Interactions 26, 515–527. Cui, H., Wang, A., 2017. An efficient viral vector for functional genomic studies of prunus fruit trees and its induced resistance to plum pox virus via silencing of a host factor gene. Plant Biotechnology Journal 15 (3), 344–356. Kozieł, E., Bujarski, J., Otulak, K., 2017. Molecular biology of Prune dwarf virus – A lesser known member of the family Bromoviridae but a vital component in the dynamic virus–host cell interaction network. International Journal of Molecular Sciences 18 (12), 2733. Moreno, A., Fereres, A., 2012. Virus diseases in lettuce in the Mediterranean basin. II Alfalfa mosaic virus (Bromoviridae, Alfamovirus). Advances in Virus Research 8, 250–253. Pallas, V., Aparicio, F., Herranz, M.C., et al., 2012. Ilarviruses of Prunus spp.: A continued concern for fruit trees. Phytopathology 102, 1108–1120. Pallas, V., Aparicio, F., Herranz, M.C., Sanchez-Navarro, J.A., Scott, S.W., 2013. The molecular biology of ilarviruses. Advances in Virus Research 87, 139–181. Rubio, M., Martínez-Gómez, P., Marais, A., et al., 2017. Recent advances and prospects in Prunus virology. Annals of Applied Biolology 171, 125–138. Wang, A., 2015. Dissecting the molecular network of virus-plant interactions: The complex roles of host factors. Annual Review of Phytopathology 53, 45–66. Wang, A., 2018. Virus and host plant interactions. In: eLS. Chichester: John Wiley & Sons Ltd. Available at: https://onlinelibrary.wiley.com/doi/10.1002/9780470015902. a0000758.pub3.

Luteoviruses (Luteoviridae) Leslie L Domier, Agricultural Research Service, US Department of Agriculture, Urbana, IL, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of L.L. Domier, C.J. D’Arcy, Luteoviruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00438-6.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein IRES Internal ribosomal entry site kb Kilobases; the size of a ssDNA or ssRNA molecule kDa Kilodaltons; the size of a protein nm Nanometer(s) nt Nucleotide(s) ORF Open reading frame Poly(A) Polyadenylated

Glossary Hemocoel The primary body cavity of most arthropods that contains most of the major organs and through which the hemolymph circulates. Hemolymph A circulatory fluid in the body cavities (hemocoels) and tissues of arthropods that is analogous to blood and/or lymph of vertebrates.

PTGS Post-transcriptional gene silencing RdRp RNA-dependent RNA polymerase RTD Readthrough domain RT-PCR Reverse transcription polymerase chain reaction S20,w Corrected sedimentation coefficient; extrapolated to that in water at 201C and infinitely diluted sgRNA Sub-genomic RNA ssRNA Single-stranded ribonucleic acid Vpg Genome-linked protein

Pseudoknot A folded RNA that contains two or more stem-and-loop structures where the loop of one stem-andloop structure interacts with the stem of a second stem-andloop structure.

Introduction Viruses of the family Luteoviridae (luteovirids) cause economically important diseases in many monocotyledonous and dicotyledonous crop plants, including cereal grasses, cucurbits, legumes, lettuce, potatoes, sugar beets, and woody perennials. Yield reductions as high as 30% have been reported in epidemic years, although in some cases crops can be totally destroyed. Diseases caused by the viruses were recorded decades and even centuries before they were associated with the causal viruses. In many cases, the stunted, deformed, and discolored plants that result from luteovirid infections were thought to be the result of abiotic factors, such as mineral imbalances or stressful environmental conditions, or other biotic agents. This, along with their inabilities to be transmitted mechanically, delayed the initial association of the symptoms with plant viruses. For example, curling of potato leaves was first described in Lancashire, UK, in the 1760s, but was not recognized as a specific disease of potato until 1905 and to be caused by an aphid-transmitted virus until the 1920s. The causal agent, Potato leaf roll virus (PLRV), was not purified until the 1960s. Similarly, widespread disease outbreaks in cereals, probably caused by Barley yellow dwarf virus (BYDV), were noted in the United States in 1907 and 1949. In 1951, a virus was proposed as the causal agent. Other diseases caused by luteovirids, like yellowing of Suakwa vegetable sponge, which is caused by Suakwa aphid-borne yellows virus (SABYV), were not described until recently.

Taxonomy and Classification Members of the family Luteoviridae were first grouped because of their common biological properties. These properties included persistent transmission by aphid vectors and the induction of yellowing symptoms in many infected host plants. ‘Luteo’ comes from the Latin luteus, which translates as yellowish. All luteovirids have small (c. 25 nm diameter) icosahedral particles, composed of one major and one minor protein component and a single molecule of positive-sense single-stranded RNA of approximately 5600 nt in length. The family Luteoviridae is divided into three genera – Luteovirus, Polerovirus (derived from potato leaf roll), and Enamovirus (derived from pea enation mosaic) – based on the arrangements, sizes, and phylogenetic relationships of the predicted aa sequences of the open reading frames (ORFs). In some plant virus families, a single gene can be used to infer taxonomic and

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Luteoviruses (Luteoviridae)

Fig. 1 Phylogenetic relationships of the predicted amino acid sequences of the (a) RNA-dependent RNA polymerase (ORF2) and (b) major capsid protein (ORF3). When predicted amino acid sequences from ORF2 are used to group virus species the genera form three distinct groups. Using predicted amino acid sequences from ORF3, species of the genera Luteovirus and Polerovirus are intermingled in the tree. The resulting consensus trees from 500 bootstrap replications are shown. The numbers above each node indicate the percentage of bootstrap replicates in which that node was recovered. For virus abbreviations and GenBank accession numbers, see Table 1.

phylogenetic relationships. Within the family Luteoviridae, however, different taxonomic relationships can be predicted depending on whether sequences of the replicase (ORF2) or coat protein (CP; ORF3) genes are analyzed (Fig. 1). ORFs 1 and 2 of the luteoviruses are most closely related to the polymerase genes of viruses of the family Tombusviridae, while ORFs 1 and 2 of the poleroviruses and enamoviruses are related to those of the family Solemoviridae. These polymerase types are distantly related in evolutionary terms. Consequently, it has been suggested that the family Luteoviridae be abolished and the Luteovirus genus be moved to the family Tombusviridae and the Enamovirus and Polerovirus genera be moved to the family Solemoviridae. It is hypothesized that luteovirid genomic RNAs arose by recombination between ancestral genomes containing the CP genes characteristic of the family Luteoviridae and genomes containing either of the two polymerase types. For taxonomic purposes, the polymerase type has been the primary determinant in assigning a virus to a genus. For this reason, viruses for which only CP sequences have been determined have not been assigned to a genus. The current members of the family are listed in Table 1. The genus Luteovirus contains nine species, and the Polerovirus genus has 19 species. The genus Enamovirus contains four species. In addition, with recent advances in virus detection by next-generation sequencing, over 60 partial or complete genome sequences have been deposited in GeneBank that potentially represent new virus species in the family.

Virion Properties The sedimentation coefficients S20,w (in Svedberg units) for luteoviruses and poleroviruses range from 106 S to 127 S. Buoyant densities in CsCl are approximately 1.40 g cm
3. The particles formed as result of the mixed infections by PEMV-1

Luteoviruses (Luteoviridae)

Table 1

449

Virus members in the family Luteoviridae

Genus

Species

Abbreviation

Accession numbera

Luteovirus

Barley yellow dwarf virus KerII Barley yellow dwarf virus KerIII Barley yellow dwarf virus – MAV Barley yellow dwarf virus – PAS Barley yellow dwarf virus – PAV Bean leafroll virus Nectarine stem pitting-associated virus Rose spring dwarf-associated virus Soybean dwarf virus Beet chlorosis virus Beet mild yellowing virus Beet western yellows virus Carrot red leaf virus Cereal yellow dwarf virus – RPS Cereal yellow dwarf virus – RPV Chickpea chlorotic stunt virus Cotton leafroll dwarf virus Cucurbit aphid-borne yellows virus Maize yellow dwarf virus RMV Maize yellow mosaic virus Melon aphid-borne yellows virus Pepo aphid-borne yellows virus Pepper vein yellows virus Pepper vein yellows virus 5 Potato leafroll virus Suakwa aphid-borne yellows virus Sugarcane yellow leaf virus Tobacco vein distorting virus Turnip yellows virus Alfalfa enamovirus 1 Citrus vein enation virus Grapevine enamovirus 1 Pea enation mosaic virus 1 Barley yellow dwarf virus – GPV Barley yellow dwarf virus – SGV Chickpea stunt disease associated virus Groundnut rosette assistor virus Indonesian soybean dwarf virus Sweet potato leaf speckling virus Tobacco necrotic dwarf virus

BYDV-KerII BYDV-KerIII BYDV-MAV BYDV-PAS BYDV-PAV BLRV NSPaV RSDaV SbDV BChV BMYV BWYV CRLV CYDV-RPS CYDV-RPV CpCSV CLRV CABYV MYDV-RMV MYMV MABYV PABYV PVYV PVYV-5 PLRV SABYV ScYLV TVDV TuYV AEV-1 CVEV GVEV-1 PEMV-1 BYDV-GPV BYDV-SGV CpSDaV GRAV ISDV SPLSV TNDV

NC_021481.1 KC559092.1 NC_003680.1 NC_002160.2 NC_004750.1 NC_003369.1 NC_027211.1 NC_010806.1 NC_003056.1 NC_002766.1 NC_003491.1 NC_004756.1 NC_006265.1 NC_002198.2 NC_004751.1 NC_008249.1 NC_014545.1 NC_003688.1 NC_021484.1 KU248489.1 NC_010809.1 NC_030225.1 NC_015050.1 NC_036803.1 NC_001747.1 NC_018571.2 NC_000874.1 NC_010732.1 NC_003743.1 NC_029993.1 NC_021564.1 NC_034836.1 NC_003629.1 NC_039035.1 AY541039.1 Y11530.1 NC_038509.1

Polerovirus

Enamovirus

Unassigned

NC_038510.1

a

Accession numbers beginning with NC_ represent reference genomic sequences.

and PEMV-2 sediment as two components. The S20,w are 107–122 S for B components (PEMV-1) and 91–106 S for T components (PEMV-2, an umbravirus). Virions are moderately stable and are insensitive to treatment with chloroform or nonionic detergents but are disrupted by prolonged treatment with high concentrations of salts. Luteovirus and polerovirus particles are insensitive to freezing.

Virion Structure and Composition All members of the Luteoviridae have nonenveloped icosahedral particles with diameters of 25–28 nm (Fig. 2). Capsids are composed of major (21–23 kDa) and minor (54–76 kDa) CPs, which contain a C-terminal extension to the major CP called the readthrough domain (RTD). According to X-ray diffraction and molecular mass analysis, virions consist of 180 protein subunits, arranged in T ¼ 3 icosahedra. Virus particles do not contain lipids or carbohydrates. Using an X-ray crystallography-derived structure of virions of the Rice yellow mottle sobemovirus, which shares a CP aa sequence similarity of 33% with PLRV, it was possible to predict the virion structure of PLRV and other luteovirids. Virions contain a single molecule of single-stranded positive-sense RNA of 5300–5900 nt. The RNAs do not have a 30 terminal poly(A) tract. A small protein (VPg) is covalently linked to the 50 end of polerovirus and enamovirus genomic RNAs. Cereal yellow dwarf virus - RPV (CYDV-RPV) also encapsidates a 322 nt satellite RNA that accumulates to high levels in the presence of the

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Luteoviruses (Luteoviridae)

Fig. 2 Transmission electron micrograph of Soybean dwarf virus particles magnified 240,000  . Virions (stained with uranyl acetate) are c. 25 nm in diameter, hexagonal in appearance, and have no envelope.

helper virus. Complete genome sequences have been determined for more than 30 species of the family Luteoviridae (Table 1). For several viruses, genome sequences have been determined from multiple isolates.

Genome Organization and Expression Genomic RNAs of luteovirids contain five to seven conserved ORFs (Fig. 3). ORFs 1, 2, 3, and 5 are shared among all members of the Luteoviridae. Luteoviruses lack ORF0. Enamoviruses lack ORF4. Luteo- and polerovirus genomes contain a small ORF (ORF3a) upstream of ORF3. Luteovirus genomes contain a small ORF (ORF6) downstream of ORF5. The PLRV genome contains ORFs 6 and 7, within ORF5 and ORF8 within ORF1. In the enamo- and poleroviruses ORF0 overlaps ORF1 by more than 600 nt, which also overlaps ORF2 by more than 600 nt. In the luteoviruses, ORF1 overlaps ORF2 by less than 50 nt. In most luteo- and polerovirus genome sequences, ORF4 is contained completely within ORF3. A single, in-frame amber (UAG) termination codon separates ORF5 from ORF3. Luteovirids have relatively short 50 and intergenic non-coding sequences. The first ORF is preceded by 21 nt in CABYV RNA and 142 nt in Soybean dwarf virus (SbDV) RNA. ORFs 2 and 3 are separated by 112–200 nt of non-coding RNA. There is considerable variation in the length of sequence downstream of ORF5, which ranges from 167 nt for CYDV-RPV to 650 nt for SbDV. Luteovirids employ a wide range of strategies to express their compact genomes. ORFs 0, 1, 2, and 8 are expressed directly from genomic RNAs. Downstream ORFs are expressed from sub-genomic RNAs (sgRNAs) that are transcribed from internal initiation sites by virus-encoded RNA-dependent RNA polymerases (RdRps) from negative-strand RNAs and are 30 co-terminal with the genomic RNA. Since the initiation codon for ORF0 of polero- and enamoviruses is upstream of that of ORF1, translation of ORF1 is initiated by ‘leaky scanning’ in which ribosomes bypass the AUG of ORF0 and continue to scan the genomic RNA until they reach the ORF1 AUG. The protein products of ORF2 are expressed as a translational fusion with the product of ORF1. At a low but significant frequency during the expression of ORF1, translation continues into ORF2 through a –1 frameshift that produces a large protein containing sequences encoded by both ORFs 1 and 2 in a single polypeptide. The frameshift is mediated by a “slippery hepta-nucleotide sequence” (in the form X XXY YYZ) and a downstream RNA secondary structure termed a pseudoknot that causes ribosomes to pause and then shift back one nt before continuing translation in the new reading frame. ORF8, which has only been identified in PLRV, resides entirely within ORF1 in a different reading frame and encodes a 5 kDa replication-associated protein. To express ORF8, sequences within the ORF fold into a structure called an internal ribosome entry site (IRES), which recruits ribosomes to initiate translation about 1600 nt downstream of the 50 terminus of PLRV RNA. ORFs 3a, 3, 4, and 5 are expressed through a leaky scanning mechanism from the 50 terminus of sgRNA1, which is located about 200 nt upstream of ORF3 at the end of ORF2 and extends to the 30 terminus of the genome. Translation of ORF3a is initiated at a non-AUG codon. ORF4 of most luteo- and poleroviruses is contained within ORF3. In all luteovirids, ORF5 is expressed only as a translational fusion with the products of ORF3 by readthrough of the UAG stop codon at the end of ORF3, which produces a protein with the product of ORF3 at its N-terminus and the product of ORF5 at its C-terminus. Readthrough is regulated by local and long-distance RNA-RNA interactions and in the case of luteoviruses and some poleroviruses requires the presence of CCXXXX repeats (where X is any base) downstream of the ORF3 stop codon. Luteo- and poleroviruses produce second smaller sgRNA capable of expressing ORFs 6 and 7. Third sgRNAs, which do not appear to encode proteins, are produced at very high levels in luteoviruses, but at only low levels in PLRV.

Luteoviruses (Luteoviridae)

451

Fig. 3 Maps of the virus genomes of genera in the family Luteoviridae. Individual ORFs are shown with open boxes. The ORFs are staggered vertically to show the different reading frames occupied by each ORF. The yellow boxes indicate protein products with the predicted sizes listed to the right of each. The polyproteins encoded by ORF1 of enamo- and poleroviruses contain the protease and the genome-linked protein (VPg). The predicted amino acid sequences of proteins encoded by ORF2 are similar to RNA-dependent RNA polymerases. ORF3, which encodes the major coat protein, is separated from ORF5 by an amber termination codon. ORF4, when present, is most often contained within ORF3 and encodes a protein required for virus cell-to-cell movement. Luteoand poleroviruses contain an ORF3a for which translation is initiated at a non-AUG codon. The 30 non-coding regions of luteoviruses contain translation enhancer elements (BTE). In PLRV, ORF7 is in-frame with the C-terminus of ORF5, and translation of ORF8 is mediated by an internal initiation ribosome entry site (IRES). Luteo- and poleroviruses produce three sub-genomic RNAs (sgRNAs), but enamoviruses produce a single sgRNA.

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Luteoviruses (Luteoviridae)

While enamo- and polerovirus RNAs contain 50 VPgs that interact with translation initiation factors, luteovirus RNAs contain only a 50 phosphate. Unmodified 50 termini are recognized poorly for translation initiation. To circumvent this problem, the BYDV-PAV genome contains a short sequence (BYDV translation element; BTE) located in the 30 non-coding region downstream of ORF5 that interacts with sequences near the 50 termini of genomic RNA and sgRNA1 to promote cap-independent translation initiation. Post-transcriptional gene silencing (PTGS) is an innate and highly adaptive antiviral defense found in all eukaryotes that is activated by double-stranded RNAs (dsRNAs), which are produced during virus replication. Research into the functions of the proteins encoded by luteovirids has shown that the 28–34 kDa proteins encoded by ORF0 are strong suppressors of local and systemic PTGS for polero- and enamoviruses. Luteovirus genomes lack ORF0, but the product of ORF4 in luteoviruses functions to suppress systemic PTGS. The ORF1-encoded proteins of enamo- and poleroviruses contain the VPg and a chymotrypsin-like serine protease that is responsible for the proteolytic processing of ORF1-encoded polyproteins. The protease cleaves the ORF1 protein internally to liberate the VPg, which is covalently attached to genomic RNA. The protein expressed by ORF8 of PLRV is required for virus replication. Luteovirid ORF2s have a coding capacity of 59–67 kDa for proteins that are very similar to known RdRps and hence likely represent the catalytic portion of the viral replicase. For luteo- and poleroviruses ORF3a produces highly conserved 4.8–5.3 kDa proteins that are essential for long-distance movement. ORF3 encodes the major CP of the luteovirids, which ranges in size from 21 to 23 kDa. ORF5 has a coding capacity of 29–56 kDa. However, ORF5 is expressed only as a translational fusion with the product of ORF3 when, about 10% of the time, translation does not stop at the end of ORF3 and continues through to the end of ORF5. The ORF5 portion of this readthrough protein has been implicated in aphid transmission and virus stability. Experiments with PLRV and BYDV-PAV have shown that the N-terminal region of the ORF5 readthrough protein determines the ability of virus particles to bind to proteins produced by endosymbiotic bacteria of aphid vectors. Interactions of virus particles with these proteins seem to be essential for persistence of the viruses in aphids. Nucleotide sequence changes within ORF5 of PEMV-1 abolish aphid transmissibility. The N-terminal portions of ORF5 proteins are highly conserved among luteovirids while the C-termini are much more variable. The luteo- and polerovirus genomes possess an ORF4 that is contained within ORF3 and encodes proteins of 17–21 kDa. Viruses that contain mutations in ORF4 are able to replicate in isolated plant protoplasts, but are deficient or delayed in systemic movement in whole plants. Hence, the product of ORF4 seems to be required for movement of the virus within infected plants. This hypothesis is supported by the observation that enamoviruses lack ORF4. While luteo- and poleroviruses are limited to phloem and associated tissues, the enamovirus PEMV-1 is able to move systemically through other plant tissues in the presence of PEMV-2, which under natural conditions invariably coexists with PEMV-1. Some luteo- and polerovirus genomes contain small ORFs within and/or downstream of ORF5. In luteoviruses, no protein products have been detected from these ORFs in infected cells. BYDV-PAV genomes that do not express ORF6 are still able to replicate in protoplasts. The predicted sizes of the proteins expressed by ORFs 6 and 7 of PLRV are 7.1 and 14 kDa, respectively. Based on mutational studies, it has been proposed that these genome regions may regulate transcription late in infection.

Evolutionary Relationships As mentioned above, viruses in the family Luteoviridae have replication-related proteins that are similar to those in other plant virus families and genera. The luteovirus replication proteins encoded by ORFs 1 and 2 resemble those of members of the family Tombusviridae. In contrast, the serine protease-VPg-polymerase complex of polero- and enamoviruses resemble those of viruses in the family Solemoviridae. The structural proteins of some members of the Sobemovirus genus in the family Solemoviridae also are similar to the major CP of luteovirids.

Host Range and Transmission Several luteovirids have natural host ranges largely restricted to one plant family. For example, BYDV and CYDV infect many grasses, BLRV infects mainly legumes, and CtRLV infects mainly plants in the family Apiaceae. Other luteovirids infect plants in several or many different families. For example, Beet western yellows virus (BWYV) infects more than 150 species of plants in more than 20 families. As techniques for infecting plants with recombinant viruses have improved, the experimental host ranges of luteovirids have been expanded to include plants on which aphid vectors would not normally feed. For example, BYDV, CYDV, PLRV, and SbDV have been shown to infect Nicotiana species when inoculated biolistically with viral RNA or using Agrobacterium tumefaciens harboring binary plasmids containing infectious copies of the virus genomes even though those species had not been described previously as experimental hosts for the viruses. These results suggest that feeding preferences of vector aphids play important roles in defining luteovirid host ranges. Luteovirids are transmitted in a circulative manner with varying efficiencies by at least 25 aphid species. With the exception of the enamovirus PEMV-1, members of the family Luteoviridae are transmitted from infected plants to healthy plants in nature only by the feeding activities of specific species of aphids. There is no evidence for replication of the viruses within aphid vectors. Myzus

Luteoviruses (Luteoviridae)

Accessory salivary gland

453

Primary salivary gland

Midgut

Foregut

Hemocoel

Hindgut Food canal Salivary duct

Fig. 4 Circulative transmission of viruses of the family Luteoviridae by vector aphids. While feeding from sieve tubes of an infected plant, an aphid (shown in cross section) acquires virus particles, which travel up the stylet, through the food canal, and into the foregut. The virions are actively transported across cells of the posterior midgut and/or hindgut into the hemocoel in a process that involves receptor-mediated endocytosis. Virions then passively migrate through the hemolymph to the accessory salivary gland where they are again transported by a receptor-mediated process to reach the lumen of the gland. Once in the salivary gland lumen, the virions are expelled with the saliva into the vascular tissue of host plants. Aphids can retain the ability to transmit virus for several weeks. Hindgut membranes usually are much less selective than those of the accessory salivary glands, which is why viruses that are not transmitted by a particular species of aphid often accumulate in the hemocoel, but do not traverse the membranes of the accessory salivary gland.

persicae is the most common aphid vector of luteovirids that infect dicots. Several different species of aphids transmit luteovirids that infect monocotyledenous plants (BYDV and CYDV) in a species-specific manner. Circulative transmission of the viruses is initiated when aphids acquire viruses from sieve tubes of infected plants during feeding. The viruses travel up the stylet, through the food canal, and into the foregut (Fig. 4). The viruses then are actively transported across the cells of the alimentary tract into the hemocoel in a process that involves receptor-mediated endocytosis of the viruses and the formation of tubular vesicles that transport the viruses through epithelial cells and into the hemocoel. Luteovirids are acquired at different sites within the gut of vector aphids. PLRV and BWYV are acquired in the posterior midgut. BYDV, CYDV, and SbDV are acquired in the hindgut. CABYV is taken up at both sites. Viruses then passively migrate through the hemolymph to the accessory salivary gland where the viruses must pass through the membranes of the accessory salivary gland cells in a similar type of receptor-mediated transport process to reach the lumen of the gland. Once in the salivary gland lumen, viruses are expelled with saliva into vascular tissues of host plants. Since large amounts of virus can accumulate in the hemocoel of aphids, they may retain the ability to transmit virus for several weeks. Typically, hindgut membranes are much less selective than those of the accessory salivary glands. Consequently, viruses that are not transmitted by a particular species of aphid often are transported across gut membranes and accumulate in the hemocoel, but do not traverse the membranes of the accessory salivary gland. The RTD of the minor capsid protein plays a major role in aphid transmission of luteovirids. The RTD interacts with symbionin produced by endosymbiotic aphid-borne bacteria, which may protect virions from degradation by the aphid immune system. The specificity of aphid transmission and gut tropism has been linked to the RTD in multiple luteovirids. Unlike other luteovirids, PEMV-1 can be transmitted by rubbing sap taken from an infected plant onto a healthy plant, in addition to being transmitted by aphids. This difference in transmissibility is dependent on its multiplication in cells co-infected with PEMV-2, but aphid transmissibility can be lost after several mechanical passages.

Replication Luteovirids infect and replicate in sieve elements and companion cells of the phloem and occasionally are found in phloem parenchyma cells. PEMV-1 can move systemically into other tissues in the presence of PEMV-2. Virus infections commonly result in cytopathological changes in cells that include formation of vesicles containing filaments and inclusions that contain viral RNA and virions. The subcellular location of viral RNA replication has not been determined unequivocally. However, early in infection, negative-strand RNAs of BYDV-PAV are first detected in the nucleus and later in the cytoplasm, which suggests that at least a

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Luteoviruses (Luteoviridae)

portion of luteovirus replication occurs in the nucleus. Synthesis of negative-strand RNA, which requires tetraloop structures at the 30 end of BYDV-PAV genomic RNAs, is detected in infected cells before the formation of virus particles. Late in infection, BYDVPAV sgRNA2 inhibits translation from genomic RNA, which may promote a switch from translation to replication and packaging of genomic RNAs.

Virus–Host Relationships While some infected plants display no obvious symptoms, most luteovirids induce characteristic symptoms that include stunting, leaves that become thickened, curled or brittle, and yellow, orange, or red leaf discoloration, particularly of older leaves of infected plants. These symptoms result from phloem necrosis that spreads from inoculated sieve elements and causes symptoms by inhibiting translocation, slowing plant growth, and inducing the loss of chlorophyll. Symptoms may persist, may vary seasonally, or may disappear soon after infection. Temperature and light intensity often affect symptom severity and development. In addition, symptoms can vary greatly with different virus isolates or strains and with different host cultivars. Yield losses caused by luteovirids are difficult to estimate because the symptoms often are overlooked or attributed to other agents. US Department of Agriculture specialists estimated that yield losses from BWYV, BYDV, and PLRV infections were over $65 million during the period 1951–60. Plants infected at early stages of development by luteovirids suffer the most significant yield losses, which often are linearly correlated with the incidence of virus infection.

Epidemiology Luteovirid infections have been reported from temperate, subtropical, and tropical regions of the world. Some of the viruses are found worldwide, such as BWYV, BYDV, and PLRV. Others have more restricted distribution, such as tobacco necrotic dwarf virus, which has been reported only from Japan, and groundnut rosette assistor virus, which has been reported only in African countries south of the Sahara. Luteovirids infect annual and perennial, cultivated and wild plants. In annual crops, the viruses must be reintroduced each year by their aphid vectors, sometimes from native perennial hosts. Some viruses are disseminated in infected planting material. For example, infected potato tubers are the principal source of inoculum for new epidemics of PLRV. Consequently, programs to produce clean stock are operated around the world to control these viruses. Alate, that is, winged, aphid vectors may transmit viruses from local cultivated, volunteer, or weed hosts. Alternatively, alate aphids may be transported into crops from distant locations by wind currents. These vectors may bring the virus with them, or they may first have to acquire virus from locally infected hosts. The agronomic impact of luteovirid diseases depends both on meteorological events that favor movement and reproduction of vector aphids and susceptibility of the crop at the time of aphid arrival. Only aphid species that feed on a particular crop plant can transmit virus. Aphids that merely probe briefly to determine a plant’s suitability will not transmit the viruses. Secondary spread of the viruses is often primarily by apterous, that is, wingless, aphids. The relative importance of primary introduction of virus by alate aphids and of secondary spread of virus by apterous aphids in disease severity varies with the virus, aphid species, crop, and environmental conditions. Some members of the family Luteoviridae occur in complexes with other members of the family or with other plant viruses. For example, BYDV and CYDV often are found co-infecting cereals; BWYV and SbDV are often found together in legumes; and PLRV is often found co-infecting potatoes with Potato virus Y and/or Potato virus X. Some other plant viruses depend on luteovirids for their aphid transmission, such as the Groundnut rosette virus, Carrot mottle virus, and Bean yellow vein banding virus (all umbraviruses), which depend on Groundnut rosette assistor virus, Carrot red leaf virus, and PEMV-1, respectively.

Diagnosis An integral part of controlling luteovirid diseases is accurate diagnosis of infection. Because symptoms caused by luteovirids often resemble those induced by other biotic and abiotic factors, visual diagnosis is unreliable and other methods have been developed. Initially, infectivity, or biological, assays were used to diagnose infections. These techniques also have been used to identify species of vector aphids and vector preferences. In bioassays, aphids are allowed to feed on infected plants and then are transferred to indicator plants. These techniques are very sensitive but can require several weeks for symptoms to develop on indicator plants. The strong immunogenicity of luteovirids has facilitated development of very specific and highly sensitive serological tests that can discriminate different luteovirids and sometimes even strains of a single virus species. Poly- and monoclonal antibodies for virus detection are produced by immunizing rabbits and/or mice with virus particles purified from infected plants. Techniques also have been developed to detect viral RNAs from infected plant tissues by reverse transcription-polymerase chain reaction (RT-PCR), which can be more sensitive and discriminatory than serological diagnostic techniques. High-throughput sequencing of plant transcriptomes has become a powerful tool for identification of new luteovirids and characterization of virus complexes infecting plants. Even so, serological tests are the most commonly used techniques for the detection of infections because of their simplicity and speed.

Luteoviruses (Luteoviridae)

455

Control Because methods are not available to cure luteovirid infections after diagnosis, emphasis has been placed on reducing losses through the use of tolerant or resistant plant cultivars and/or on reducing the spread of viruses by controlling aphid populations. Many luteovirids are transmitted by migrating populations of aphids that occur at similar times each year. For those virus–aphid combinations, it is sometimes possible to plant crops so that young, highly susceptible plants are not in the field when the seasonal aphid migrations occur. Insecticides have been used in a prophylactic manner to reduce crop losses. While insecticide treatments do not prevent initial infections, they can greatly limit secondary spread of aphids and therefore of viruses. In some instances biological control agents such as predatory insects and parasites have reduced aphid populations significantly. Genes for resistance or tolerance to infection by luteovirids have been identified in most agronomically important plant species infected by the viruses. For BYDV, PLRV, and SbDV, transgenic plants that express portions of the virus genomes have been produced through DNA-mediated transformation. In some cases, the expression of these virus genes in transgenic plants confers higher levels of virus resistance than naturally occurring plant resistance genes.

Further Reading Brault, V., Perigon, S., Reinbold, C., et al., 2005. The polerovirus minor capsid protein determines vector specificity and intestinal tropism in the aphid. Journal of Virology 79, 9685–9693. Fusaro, A.F., Barton, D.A., Nakasugi, K., et al., 2017. The Luteovirus P4 movement protein is a suppressor of systemic RNA silencing. Viruses 9, 16. Gray, S., Cilia, M., Ghanim, M., 2014. Circulative, “nonpropagative” virus transmission: An orchestra of virus-, insect-, and plant-derived instruments. In: Maramorosch, K., Murphy, F.A. (Eds.), Advances in Virus Research, vol. 89. San Diego: Elsevier Academic Press Inc., pp. 141–199. Hipper, C., Monsion, B., Bortolamiol-Becet, D., Ziegler-Graff, V., Brault, V., 2014. Formation of virions is strictly required for Turnip yellows virus long-distance movement in plants. Journal of General Virology 95, 496–505. Hogenhout, S.A., van der Wilk, F., Verbeek, M., Goldbach, R.W., van den Heuvel, J.F., 2000. Identifying the determinants in the equatorial domain of Buchnera GroEL implicated in binding potato leafroll virus. Journal of Virology 74, 4541–4548. Hulo, C., de Castro, E., Masson, P., et al. 2019. Luteoviridae. ViralZone. ExPASy Bioinformatics Resource Portal. Switzerland. Available at: https://viralzone.expasy.org/45 (accessed 26.05.2019). Lee, L., Palukaitis, P., Gray, S.M., 2002. Host-dependent requirement for the Potato leafroll virus 17-kDa protein in virus movement. Molecular Plant–Microbe Interactions 10, 1086–1094. Miller, W.A., Jackson, J., Feng, Y., 2015. Cis- and trans-regulation of luteovirus gene expression by the 30 end of the viral genome. Virus Research 206, 37–45. Miller, W.A., Liu, S.J., Beckett, R., 2002. Barley yellow dwarf virus: Luteoviridae or Tombusviridae? Molecular Plant Pathology 3, 177–183. Moonan, F., Molina, J., Mirkov, T.E., 2000. Sugarcane yellow leaf virus: An emerging virus that has evolved by recombination between luteoviral and poleroviral ancestors. Virology 269, 156–171. NCBI, 2019. Complete Genomes: Luteoviridae. Bethesda, MD: National Center for Biotechnology Information, Available at: https://www.ncbi.nlm.nih.gov/genomes/ GenomesGroup.cgi?taxid=119163 (accessed 26.05.2019) Pfeffer, S., Dunoyer, P., Heim, F., et al., 2002. P0 of Beet western yellows virus is a suppressor of posttranscriptional gene silencing. Journal of Virology 76, 6815–6824. Robert, Y., Woodford, J.A., Ducray-Bourdin, D.G., 2000. Some epidemiological approaches to the control of aphid-borne virus diseases in seed potato crops in Northern Europe. Virus Research 71, 33–47. Siddell, S.G., Walker, P.J., Lefkowitz, E.J., et al., 2019. Virus Taxonomy: 2018b Release. International Committee on Taxonomy of Viruses. Washington, DC, July 2018. Available at: https://talk.ictvonline.org/taxonomy (accessed 26.05.2019). Smirnova, E., Firth, A.E., Miller, W.A., et al., 2015. Discovery of a small non-AUG-initiated ORF in poleroviruses and luteoviruses that is required for long-distance movement. PLOS Pathogens 11, e1004868. Taliansky, M., Mayo, M.A., Barker, H., 2003. Potato leafroll virus: A classic pathogen shows some new tricks. Molecular Plant Pathology 4, 81–89. Terradot, L., Souchet, M., Tran, V., Giblot Ducray-Bourdin, D., 2001. Analysis of a three-dimensional structure of Potato leafroll virus coat protein obtained by homology modeling. Virology 286, 72–82. Thomas, P.E., Lawson, E.C., Zalewski, J.C., Reed, G.L., Kaniewski, W.K., 2002. Extreme resistance to Potato leafroll virus in potato cv. Russet Burbank mediated by the viral replicase gene. Virus Research 71, 49–62. Xu, Y., Ju, H.J., DeBlasio, S., Carino, E.J., et al., 2018. A stem-loop structure in Potato leafroll virus open reading frame 5 (ORF5) is essential for readthrough translation of the coat protein ORF stop codon 700 bases upstream. Journal of Virology 92. (e01544-17).

Machlomovirus and Panicoviruses (Tombusviridae) Kay Scheets, Oklahoma State University, Stillwater, OK, United States r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

Poly(A) Polyadenylated RAP Replicase-associated protein RdRp RNA-dependent RNA polymerase satRNA satellite RNA SDS-PAGE Sodium dodecyl sulfate-polyacrylamide gel electrophoresis UTR Untranslated region vRNA virion RNA xrRNA exoribonuclease-resistant RNA

aa Amino acid(s) CITE Cap-independent translation enhancer CP Coat protein or capsid protein kb Kilobase kDa Kilo Dalton LDRI Long distance RNA–RNA interaction MP Movement protein nt Nucleotide(s) ORF Open reading frame

Glossary 30 CITE An RNA tertiary structure located in the 30 UTR which binds to translation initation factor(s) and to a structure nearer the 50 end of the RNA which allows assembly with a ribosome and initiation factors for translation. Cryo-electron microscopy Technique for rapidly cooling dilute aqueous biomolecules to determine structures at the molecular level. Movement proteins Viral proteins that assist in the movement of vRNA or virions through plasmodesmata, the tube-like structures connecting one plant cell to another. Phloem Vascular tissue that transports small molecules throughout the plant. Plasmodesmata Microscopic membrane channels through plant cell walls that allow small molecules to readily move between plant cells. Plasmodesmata are internally coated by the endoplasmic reticulum. Poaceae A family of monocotyledenous plants commonly called the grass family. Potyvirid A virus in the family Potyviridae which contains the genera Potyvirus and Tritimovirus.

Protoplast A plant cell (either from an undifferentiated plant cell culture or from a plant tissue such as a leaf) that has been treated in an isotonic solution with a mixture of enzymes that degrades the cell wall, leaving membranebound structures similar to single animal cells. Satellite RNA Non-coding ssRNAs dependent on a helper virus for replication. Satellite virus ssRNA encoding only a CP that requires a helper virus to replicate. Silencing suppressor - (suppressor of silencing) Virally encoded proteins from diverse virus families that interfere with host production or effectiveness of siRNAs. Small interfering RNAs (or silencing RNAs), 20–25 bp RNAs with 2 nt 3' overhanging ends that target invading viral RNAs or cellular RNAs for degradation in eukaryotic cells. Synergism/synergistic disease When disease symptoms of a double infection are more severe than the sum of each single infection. Tombusvirid Member of the family Tombusviridae.

Classification Realm Riboviria, family Tombusviridae, subfamily Procedovirinae, genera Machlomovirus, and Panicovirus. The genus Machlomovirus contains only one species, Maize chlorotic mottle virus, while the genus Panicovirus contains three species: Cocksfoot mild mosaic virus, Panicum mosaic virus (the type species) and Thin paspalum asymptomatic virus (Table 1).

Virion Structure Like the majority of plant viruses, Maize chlorotic mottle virus (MCMV) and Panicum mosaic virus (PMV) are composed of only the viral genome within a protein shell made of multiple copies of a single coat protein (CP). Similar smooth virion structures have been determined for MCMV (cryo-electron microscopy) and PMV (X-ray diffraction). Both viruses are B30 nm in diameter, exhibit icosahedral symmetry, and are composed of 180 subunits of a single 25 or 26 kDa CP that encapsidates one copy of their single ssRNA genomes (Fig. 1).

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List of members of the genera Machlomovirus and Panicovirus in the family Tombusviridae. Type species are in bold

Subfamily/Genus Procedovirinae Machlomovirus Panicovirus

Species

Acronym

Accession #

Length (nt)

# sgRNAsa

Maize chlorotic mottle virus Cocksfoot mild mosaic virus Panicum mosaic virus Thin paspalum asymptomatic virus

MCMV CMMV PMV TPAV

NC003627 NC011108 NC002598 NC021705

4437 4198 4326 4195

1 1 1 1

a

protein-coding sgRNAs only.

Fig. 1 Purified maize chlorotic mottle virus negatively stained with uranyl acetate. The bar represents 100 nm.

Genomes The genome organizations of the 16 genera in the family Tombusviridae fit one of four patterns. MCMV and panicoviruses have a carmovirus-like genome organization in which the cell-to-cell MP ORFs immediately follow the RdRp ORF, and the CP ORF is located at the 30 end of the genome. Like other members of the family Tombusviridae, the vRNA for both viruses is uncapped and does not contain a poly(A) tail. MCMV (4437 nt), PMV (4326 nt) and the other two recognized panicoviruses have larger genomes than other tombusvirids that form smooth virions (lacking a protruding domain [24.5–29.9 kDa CPs]), and are longer than some aureusviruses, which encode CPs with protruding domains (35.1–42.2 kDa). The size reflects the presence of larger genes for the pre-readthrough region (RAP) of their replicases, which encode carboxyl regions with high homology to other family members while the amino-terminal extensions are unique to each genus. MCMV encodes two overprinted genes that, 30 years after the first vRNA submission to GenBank, are still completely unique. And unlike all other tombusvirids, the RAP ORF start codon is preceded by the start codon of the unique 32 kDa ORF. P32 and p50, due to the p32 ORF overlap of the p50 ORF, are highly charged. P32 has a pI B4 while p50 has a pI B10 which causes an anomalous (opposite) migration pattern of the two proteins in SDS-PAGE. Initially, it was predicted that the second unique MCMV ORF encoding an extension of the upstream MP (p7a) was expressed via stop codon readthrough to make a 31 kDa protein. But the p7a ORF stop codon is not in a context that is likely to support readthrough, so it is more likely that the second unique gene is expressed from another non-canonical start codon to make a 23 kDa protein. PMV encodes an unrelated unique gene overprinted within its CP ORF that encodes a 15 kDa protein which is only found in viruses in the genus Panicovirus (Fig. 2).

Hosts and Distribution MCMV was initially found in maize (Zea mays), also known as corn, in Peru in 1973. MCMV next spread long-distance via contaminated seeds to Kansas and Nebraska in midwestern USA (1976). Testing of native grasses and common cereal crops

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Fig. 2 Genome organizations of MCMV and PMV. Thick black lines mark the gRNAs while the thinner short lines are sgRNAs. ORFs encoding proteins with the same function are the same color while unique ORFs have unique colors (identified below the genomes) along with symbols for readthough stop codons and noncanonical start codons. ORFs are located relative to the RdRp reading frame (  1 above and þ 1 below).

indicated that MCMV-K and MCMV-P systemically infected 19 and 15 species, respectively, including wheat and barley. Since MCMV seed transmission rate varies widely for different maize cultivars, and maize production has increased worldwide, the virus has spread to most continents. Over the years MCMV has appeared in Mexico, additional South American countries, Hawaii, Thailand, Mainland China, and Taiwan, seven countries in Eastern Africa, and most recently was detected in maize and johnsongrass in Spain. MCMV has infected additional Poaceae crops in Africa (sorghum, sugarcane, and finger millet) and was found in johnsongrass, a common agricultural weed which grows nearly worldwide. China has reported MCMV in maize, sugarcane, sorghum, and Job's tears (Coix lacryma-jobi). PMV infects switchgrass (Panicum virgatum) which grows in most of the USA and Canada except the farthest Western & Northern States/Provinces. PMV mostly occurs in 12 states: Nebraska-to-Texas, and Eastern States below parallel 361300 N except Tennessee. PMV also infects the warm-season lawn grasses St. Augustine grass (Stenotaphrum secundatum) and centipedegrass (Eremochloa ophiuroides), and the model organism purple false brome (Brachypodium distachyon). For the two other recognized panicoviruses, CMMV infects a cool season perennial grass (Dactylis glomerata) native to northern Europe, temperate Asia, and northern Africa that is grown in hay meadows and pastures, while TPAV was found in a native tallgrass prairie in Oklahoma, USA. Bermuda grass latent virus (BGLV), found in South-Eastern USA, is likely another asymptomatic panicovirus that infects Bermuda grass (Cynodon dactylon) which is a warm-season grass used for sports fields and lawns.

Life Cycle For both MCMV and PMV, upon entering a plant cell the virion disassembles, exposing the vRNA, which, based on work with many tombusvirids, folds into complex secondary and tertiary structures that regulate the various stages of virus replication and gene expression via LDRIs, including translation, replication, and sgRNA synthesis. Both viruses replicate in the cytoplasm, and the vRNA (and later the sgRNA) engages with ribosomes via unique genus-specific 3 0 CITEs: the PMV-like 30 CITE (PTE) and a PTE variant for MCMV in their 30 UTRs. Base-pairing between a small stem loop in the 50 UTRs of both vRNA and sgRNA with a sequence in the 30 CITE allows the circularization function for each RNA that is typically performed for eukaryotic mRNAs by translation factors that bind poly(A) tails and m7 G-caps. RAP and lower quantities of RdRp are synthesized from vRNA, and RAP and RdRp are the only viral proteins required for replication in protoplasts for both viruses. Replication, including (  )strand synthesis, most likely occurs in membrane-bound vesicles on mitochondrial or peroxisomal membranes as has been shown for some tombusviruses. During a viral infection (  )RNA accumulates to levels that allow isolation of dsRNA from leaf tissue. For MCMV, p32 is also synthesized from vRNA at low levels from the 5 0 overprinted ORF. MCMV p32 is not required for replication in protoplasts or plants, but infectious transcripts lacking p32 expression accumulate virus at B1/3 of the level of wild type in protoplasts, and produce delayed, mild infections in maize plants, suggesting this gene might encode a unique suppressor of silencing. A single large sgRNA is synthesized for MCMV and PMV, which contains all the ORFs necessary for cell-to-cell and long distance movement. For both species, proteins expressed from the two small ORFs (MP1 and MP2) are required for cell-to-cell movement, while MP1, MP2, and CP are required for efficient long-distance movement. The unique ORFs in PMV and MCMV overlapping their CP ORFs encode proteins required for long distance viral movement. Fortuitously, when MCMV gene functions were analyzed, the mutations introduced to “stop” the ability to read through the MCMV p7a stop codon also altered the noncanonical start codon for p23, so the function of the downstream ORF was accurately identified. MCMV infections also produce a small 337 nt 3 0 coterminal RNA, sgRNA2. This is likely an exoribonuclease-resistant RNA (xrRNA) produced by a cellular exoribonuclease, which enhances the viral infection. xrRNAs have been identified in viruses from five other tombusvirid genera.

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Epidemiology PMV and other panicoviruses are transmitted via mechanical inoculation, usually lawnmowers (St. Augustine grass and centipedegrass) or other grass-harvesting/cutting tools such hay swathers and balers for switchgrass. MCMV is also readily transmitted via mechanical inoculation, but it has also been shown to be transmitted via corn rootworm beetles and thrips. It is likely that transmission via these insects occurs through the masceration of the tissue (beetles) or rasping/sucking mouthparts of the thrips after the insects have fed on infected plants, which would be mechanical inoculation via insects. MCMV also exhibits transmission through the soil since immediately replanting maize in fields that contained MCMV-infected maize results in a high rate of infection of the new crop. Some chytrid fungi in the genus Olpidium can transmit tombusviruses via binding of assembled virions to fungal zoospores, so a fungus may be the vector for MCMV transmission via soil. Growing a nonhost crop such as soybeans between maize crops stops soil-transmission.

Pathogenesis Both MCMV and PMV are part of highly destructive synergistic diseases that are produced by different co-infections. SAD is caused by coinfection of PMV and Satellite panicum mosaic virus (SPMV) or the closely related Satellite St. Augustine decline virus (SSADV) in St. Augustine grass. PMV supports the replication of SPMV (824 nt) and two subviral RNAs, satS (375 nt) and satC (444 nt). SPMV encodes its own CP (17 kDa), and SPMV forms icosahedral virions that exclusively encapsidate its own RNA or PMV satRNA. Encapsidated SPMV accumulates in the cytosol, and SPMV CP also accumulates in the cell wall, nuclear membranes, and nucleolus. The satRNA satS replicates in pearl millet protoplasts or plants when RNA from PMV or PMV þ SPMV are present, and satS decreases PMV and SPMV accumulation, thereby decreasing disease symptoms. While single infections of MCMV produce relatively mild symptoms in maize that vary with the cultivar, double infections with the maize-infecting potyviruses [(Maize dwarf mosaic virus (MDMV), Johnson grass mosaic virus (JGMV), Sugarcane mosaic virus (SCMV)], or the tritimovirus Wheat streak mosaic virus (WSMV) causes a severe synergistic infection called CLN or MLN. Potyviruses are transmitted by aphids while WSMV is transmitted by the wheat curl mite. In most synergistic diseases involving potyvirids, the potyvirus accumulates to about the same level in single or double infections, while the other virus (MCMV in CLN) can increase dramatically, and this occurs for MLN caused by potyviruses. This is probably due to the highly effective potyvirid silencing suppressor, HC-Pro. In MCMV þ WSMV infections of maize, the level of WSMV also increased in doubly-infected plants, and the increase was temperature-dependent. The recent MLN outbreak in Africa involved SCMV and JGMV, while SCMV is associated with CLN in China.

Diagnosis Antibody-based detection kits are commercially available for both MCMV and PMV, and both viruses are readily detected from leaf tissue via RT-PCR using primers within the sgRNA region.

Prevention Scrupulous cleaning of lawn equipment and farming equipment (balers, tillers, combines) decreases the chance of transmitting the viruses to additional fields. There are no treatments for living infected plants. Crop rotation of maize with any dicot crop stops soil transmission of MCMV.

Further Reading Chkuaseli, T., White, K.A., 2018. Intra-genomic long-distance RNA–RNA interactions in plus-strand RNA plant viruses. Frontiers in Microbiology 9, 529. Kraft, J.J., Peterson, M.S., Cho, S.K., et al., 2019. The 30 untranslated region of a plant viral RNA directs efficient cap-independent translation in plant and mammalian systems. Pathogens 8, 8010028. Miller, W.A., Koev, G., 2000. Synthesis of sub-genomic RNAs by positive-strand RNA viruses. Virology 273, 1–8. Miller, W.A., Shen, R., Staplin, W., Kanodia, P., 2016. Non-coding RNAs of plant viruses and viroids: sponges of host translation and RNA interference machinery. Molecular Plant-Microbe Interactions 29, 156–164. Miras, M., Miller, W.A., Truniger, V., Aranda, M.A., 2017. Non-canonical translation in plant RNA viruses. Frontiers in Plant Science 8, 494. Mwando, N.L., Tamiru, A., Nyasani, J.O., et al., 2018. Maize chlorotic mottle virus induces changes in host plant volatiles that attract vector thrips species. Journal of Chemical Ecology 44, 681–689. Nagy, P.D., 2016. Tombusvirus-host interactions: Co-opted evolutionarily conserved host factors take center court. Annual Review of Virology 3, 491–515. Palukaitis, P., 2016. Satellite RNAs and satellite viruses. Molecular Plant-Microbe Interactions 29, 181–186. Pyle, J.D., Scholthof, K.B.G., 2017. Biology and pathogenesis of satellite viruses. In: Hadidi, A., Flores, R., Randles, J., Palukaitis, P. (Eds.), Viroids and Satellites. Academic Press. Scheets, K., 1998. Maize chlorotic mottle machlomovirus and Wheat streak mosaic rymovirus concentrations increase in the synergistic disease corn lethal necrosis. Virology 242, 28–38.

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Scheets, K., 2000. Maize chlorotic mottle machlomovirus expresses its coat protein from a 1.47-kb sub-genomic RNA and makes a 0.34-kb sub-genomic RNA. Virology 267, 90–101. Scheets, K., 2016. Analysis of gene functions in maize chlorotic mottle virus. Virus Research 222, 71–79. Steckelberg, A.L., Vicens, Q., Kieft, J.S., 2018. Exoribonuclease-resistant RNAs exist within both coding and noncoding subgenomic RNAs. mBio 9, e02461-18. Wang, Z., Parisien, M., Scheets, K., Miller, W.A., 2011. The cap-binding translation initiation factor, eIF4E, binds a pseudoknot in a viral cap-independent translation element. Structure 19, 868–880.

Maize Streak Virus (Geminiviridae) Darren P Martin, University of Cape Town, Cape Town, South Africa Aderito L Monjane, Norwegian Veterinary Institute, Oslo, Finland r 2021 Elsevier Ltd. All rights reserved. This is an update of D.P. Martin, D.N. Shepherd, E.P. Rybick, Maize Streak Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00707-X.

Abbreviations

Rep

C-ori

complementary strand (i.e., the half of the DNA duplex that is not packaged into virus particles) origin of replication. CP coat protein. The only protein component of the virus particle also believed to be involved in nuclear trafficking and cell-to-cell movement of viral DNA. LIR long or large intergenic region containing the origin of virion strand replication and gene promoters. MP movement protein. A small (c. 10 kDa) protein believed to be involved in intercellular virus movement via plasmodesmata.

Glossary Agro-infection Technique used for the infection of host plants with cloned virus genomes involving transfer of virus DNA into the nuclei of host cells by the bacterium Agrobacterium tumefaciens. Agro-inoculation See agro-infection. Bicistronic Contains two protein-coding regions within a single mRNA transcript. Capsomer A subunit of the mature virus particle containing an ordered series of polymerized coat protein molecules.

replication-associated protein involved in the initiation of virion strand replication. RepA a truncated version of Rep with a unique C-terminal domain believed to be involved in regulation of host and/or virus gene expression. SIR short or small intergenic region containing gene polyadenylation signals and the origin of complementary strand replication. V-ori virion strand (i.e., the half of the DNA duplex that is packaged into virus particles) origin of replication.

Cicadulina spp. A group of leafhopper species involved in transmission of MSV. Oviposition Egg-laying. To oviposit means to lay eggs. Plastochron The time interval between successive leaf primordia, or the attainment of a certain stage of leaf development. Viruliferous A state in which an MSV vector species is carrying and is capable of transmitting the virus.

Introduction Maize streak virus (MSV) is the causal agent of maize streak disease (MSD), one of the most serious viral diseases of maize in Africa. It is a major contributor to the continent’s food security problems and is endemic throughout Africa south of the Sahara. It is also found on the Indian Ocean islands of Madagascar, Mauritius, and La Réunion. There is no obvious barrier to spread of the virus outside of this region; hence it should be considered as a serious potential problem for other as yet unaffected maize-growing areas.

History and Taxonomy “The disorder of the mealie plant, locally described as “Mealie Blight”, “Mealie Yellows”, or “Striped Leaf Disease”, belongs to a group of plant troubles arising from obscure causes …” was how MSD was first described by Claude Fuller in 1901 in Natal, South Africa. Fuller mistakenly attributed the disease to a soil disorder, but in retrospect it is quite clear that the “mealie (a local word for maize or corn) variegation” he described and drew in minute detail can be attributed to MSV. The first milestone in MSD research was reached in 1924, when Storey determined that a virus transmitted by leafhopper species of the genus Cicadulina (Fig. 1) was the causal agent of MSD. Storey named the virus “maize streak virus”, and was also the first to determine both the genetic basis of MSV transmission by Cicadulina mbila, and the heritability of MSD resistance in maize. When MSV particles were first purified in 1974 they were found to have a novel twinned quasi-icosahedral (geminate) shape (Fig. 2), from which the name “geminivirus” was derived. This was followed by the unexpected discovery in 1977 that geminivirus particles contain circular single-stranded DNA (ssDNA); a genome type never before observed in plant viruses. These novel characteristics led to the proposal of a new virus group – the geminiviruses – comprising MSV and other viruses with geminate particles and ssDNA genomes. Maize streak virus is now recognized as the type species of the genus Mastrevirus, in the family Geminiviridae.

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Fig. 1 The leafhopper vector of MSV, Cicadulina mbila Naudé. Photograph courtesy of Dr. Benjamin Odhiambo, Kenyan Agricultural Research Institute (KARI).

Fig. 2 MSD symptoms on a maize leaf: note characteristic veinal streaks. Photograph courtesy of Dr. Frederik Kloppers, PANNAR (Pty) Ltd., Greytown, KwaZulu-Natal, South Africa.

Host Range and Symptoms While most notorious for the yield losses it causes when infecting maize, MSV also infects over 80 other grass species including the economically important crops wheat, barley, and rye. In susceptible maize and grass genotypes, the virus first causes symptoms between 3 and 7 days after inoculation. These first appear as almost circular pale spots of 0.5–2 mm diameter in the lowest exposed portions of the youngest leaves. Later, fully emerged symptomatic leaves show veinal streaks from a few millimeters long to the entire length of the leaf and between 0.5 and 3 mm wide. These streaks often fuse laterally and symptomatic leaves may become 495% chlorotic (Fig. 2). Plants are worst affected when infected within a few days of coleoptile emergence; symptoms only develop above the site of inoculation on newly emerging leaves. Susceptible varieties may display severe stunting as well as very severe streaking. Afflicted plants will frequently produce deformed cobs or may fail to produce cobs altogether and per-field yield losses can reach 100% when susceptible maize genotypes are infected early. Of the eleven major MSV strains so far identified (designated MSV-A through MSV-K; (Fig. 3)), only MSV-A produces economically significant infections in maize. The “grass-adapted” MSV strains (MSV-B, -C, -D, -E, -F, -G, -H, -I, -J and -K) differ from MSV-A types by 5%–25% in nucleotide sequence, and produce substantially milder symptoms in maize than do MSV-A viruses. In many cases these grass-adapted strains are incapable of producing symptomatic infections in MSV-resistant maize genotypes.

Diversity and Evolution MSV is closely related to the other distinct ‘African streak’ mastreviruses, Panicum streak virus, Sugarcane streak virus, Sugarcane streak Egypt virus, and Sugarcane streak Reunion virus, with which it shares B65% genome sequence identity. It is, however, most similar to, although not necessarily more closely related to, an isolate of Digitaria streak virus from the Pacific island of Vanuatu, with which it shares B67% genome sequence identity (Fig. 3).

Maize Streak Virus (Geminiviridae)

463

MSV-A

MSV-H 0.05

MSV-B

MSV-I MSV-E

DSV MSV-G MSV-F

MSV-J

MSV-D

MSV-K MSV-C

Fig. 3 Phylogenetic relationships between the full genome sequences of different MSV strains. The tree is constructed using the maximumlikelihood method (HKY model transition and transversion weight determined from the data and 100 bootstrap replicates) and numbers associated with branches indicate degrees of bootstrap support for those branches. Branches with less than 70% support have been collapsed and the genome sequence of a Digitaria streak virus (DSV) from Vanuatu is included as an outgroup. Only viruses in the MSV-A group have been isolated from maize. All viruses in the other groups have been isolated from wheat, barley, or wild grass species.

The full genome nucleotide sequences of MSV-A isolates display relatively low degrees of diversity, with any two MSV-A isolates obtained from anywhere in Africa invariably having genome sequences that are more than 97% identical. MSV isolates from La Réunion share B95% identity with mainland isolates. Although Maize was first introduced at multiple points into Africa and its neighboring islands in the 16th century, it is apparent that the most recent common ancestor of all presently sampled MSV-A genomes only came into existence in the mid 1800s somewhere in SouthEastern Africa. It is also likely that this ancestral virus was the recombinant offspring of grass adapted-adapted parental viruses: one parent an MSV-B variant, and the other parent belonging to a presently unsampled MSV strain that is most closely related to MSV-F and MSV-G.

Transmission In nature, MSV and other African streak viruses are neither seed nor contact transmissible and rely instead on transmission by cicadellid leafhoppers in the genus Cicadulina (including, among others, C. mbila, C. storeyi, C. bipunctella zeae, C. latens, and C. parazeae). Of these, C. mbila is considered the most important MSV vector as it is the most widely distributed. Also, a greater proportion of C. mbila individuals are capable of transmitting the virus than is found in other Cicadulina populations. The virus may be acquired by leafhoppers at any developmental stage in less than 1 h of feeding with a minimum acquisition time of 15 s. A latent period within the vector, during which the virus cannot be transmitted, lasts between 12 and 30 h at 301C. Once this latent period is over (signaled by the appearance of virus within the leafhopper’s body fluids) the virus can be transmitted within 5 min of feeding. Once MSV has been acquired by a leafhopper, the insect will transmit the virus for the rest of its life.

Particle Structure MSV particles (Fig. 4) consist of two incomplete icosohedra with a T ¼ 1 surface lattice, comprising 22 pentameric capsomers each containing five coat protein (CP) molecules. Particle dimensions are 38 nm  22 nm with 110 CP molecules in each virion packaging a B2690 nt covalently closed mostly single-stranded circular DNA genome. The packaged DNA has annealed to it a complementary B80 nt sequence believed to act as a primer for complementary strand synthesis following infection and uncoating.

Genome Organization As with all other mastreviruses discovered to date, the MSV genome encodes four proteins: a movement protein (MP), a coat protein (CP), a replication-associated protein (Rep), and a regulatory protein (RepA). Whereas CP and MP are expressed off alternatively spliced virion sense transcripts, Rep and RepA are expressed of alternatively spliced complementary sense transcripts. MSV genomes also contain two intergenic regions: these are a short or small one (SIR), and a long or large one (LIR), which are the complementary sense strand and virion sense strand origins of replication, respectively (Fig. 5).

The Long Intergenic Region Besides containing divergent RNA polymerase II-type promoters and other transcriptional regulatory features necessary for the expression of the complementary and virion sense genes, the LIR also contains sequence elements that are essential for replication. The

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Fig. 4 Electron micrograph of MSV purified from infected maize, showing particles of 18 nm  30 nm stained with uranyl acetate. Scale ¼ 50 nm. Photograph courtesy of Kassie Kasdorf; copyright EP Rybicki.

A A

TA T

T

T A C

Nick site

V1 start

C1 start

−101 TATA

−57 −62 TATA

Repeated GC box

V sense transcript TATA box

T tracts

(a)

LIR

RepA

MP

C1 C sense genes

V sense genes

Rep CP

C2

SIR Rep stop codon

(b)

Primer-binding site

C sense

V sense

CP stop codon

Polyadenylation signals

Fig. 5 A schematic representation of the MSV LIR (a) and SIR (b), shown in context with the MSV genome. In (a) the main features of the MSV LIR are shown. These include a stem-loop structure with the loop’s nonanucleotide sequence conserved among all geminiviruses and other rollingcircle systems. The site at which Rep introduces an endonucleolytic nick to initiate virion strand replication is shown. Iterated sequences (iterons) are shown in the V sense, with blue arrows indicating their location in the LIR. Iterons are potentially specific Rep-recognition sequences via which Rep may bind to the LIR. 50 of the stem–loop is a repeated GC-box, which binds host transcription factors. A series of T tracts 30 of the stem–loop may be involved in DNA bending of this region of the LIR. TATA boxes 50 and 30 of the stem–loop are potential C sense and V sense transcription initiation sites, respectively. In (b), the main features of the SIR include polyadenylation signals for V and C sense transcripts, and a primer-binding site on the plus strand. An B80 bp DNA primer-like molecule, encapsidated with the viral genome and annealed to this site, is thought to be involved in initiating complementary strand replication. Both the MSV LIR and SIR are essential for viral replication.

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most striking of these is an inverted repeat sequence that is capable of forming a stable hairpin loop structure. All geminiviruses sequenced to date have the highly conserved nonanucleotide sequence TAATATT↓AC within the loop sequence of similar hairpin structures: this sequence contains the virion sense strand origin of replication (V-ori;↓). A sequence 6–12 nt long occurring in all known mastreviruses between the TATA box that directs rep/repA transcription and the repA initiation codon is directly repeated in the stem near the V-ori hairpin, and is probably involved in Rep and/or RepA binding during replication. The hairpin and two GC boxes on the 50 side of the stem also forms part of an upstream activator sequence (UAS) required for efficient CP expression. The GC boxes bind nuclear factors to the UAS and are known as the rightward promoter element (rpe1).

The Short Intergenic Region The MSV SIR occurs between the termination codons of the CP and rep genes (Fig. 5) and contains the polyadenylation and termination signals of the virion and complementary sense transcripts. The SIR also contains the origin of complementary strand synthesis (C-ori). A small 80-nt-long primer-like molecule is bound to the SIR of encapsidated MSV DNA and, at the onset of an infection, probably enables synthesis of double-stranded DNA (dsDNA) replicative forms (RFs) of the genome from newly uncoated virion strand DNA.

The Complementary Sense Genes (Rep and RepA) Rep is the only MSV gene product that is absolutely required for virus replication. In mastreviruses Rep is encoded within two open reading frames (ORFs) referred to as C1 and C2 (Fig. 5). Beginning at the same transcription initiation site, two C sense transcripts (1.5 and 1.2 kbp in size) are produced during MSV infections. Splicing of the larger transcript removes an intron, which permits expression of full-length Rep from the two ORFs. It is very probable, although as yet unproven in vivo, that RepA is translated from both the unspliced 1.5 kbp transcript and the 1.2 kbp transcript. If expressed in infected cells, MSV RepA would have the same N-terminal 214-amino-acid sequence as Rep, but would have a different C-terminus. RepA is likely a multifunctional protein that modifies the nuclear environment to favor viral replication. The N-terminal portions of Rep and RepA contain three conserved amino acid sequence motifs commonly found in replication-associated proteins of many extremely diverse rolling-circle replicons. Other significant landmarks include a plant retinoblastoma-related protein (pRBR) interaction motif (via which RepA binds to host pRBR molecules to manipulate the cell cycle), and oligomerization domains (via which Rep and RepA bind to other Rep/RepA molecules to form homo- and heterooligomers). It is also likely that the approximately 100 N-terminal amino acids of both Rep and RepA are involved in binding these proteins to the viral LIR during replication. The C-terminal portion of Rep contains a dNTP-binding domain with motifs similar to those found in proteins with kinase and helicase activities. The dNTP-binding domain also sits within a region with similarity to the DNA-binding domains of the myb-related class of plant transcription factors: this domain may be functional in the induction of virus and/or host gene transcription. The C-terminal portion of RepA, which is different from that of Rep, contains another potential transactivation domain also possibly involved in the regulation of virus and/or host gene expression. A second domain within the C-terminal portion of RepA possibly interacts with host proteins involved in developmental regulation.

The Virion Sense Genes (MP and CP) Transcription of the MSV virion (V) sense genes is directed by two TATA boxes within the LIR 26 and 214 nucleotides 50 of the MP start codon. Each TATA box directs the production of different-sized transcripts, both of which terminate at the same place. Splicing of an intron within the MP portion of V sense transcripts appears to be an important determinant of relative MP and CP expression levels. Whereas CP is expressed from both long and short, spliced and unspliced V sense transcripts, MP is most likely only expressed from unspliced long transcripts. The MP is post-translationally modified and contains a hydrophobic domain that may either facilitate its interaction with host cell membranes or be involved in homo- or hetero-oligomerization with the CP. The N-terminal B100 amino acids of the CP contain both nuclear localization signals and a sequence-nonspecific dsDNA- and ssDNA-binding domain. The CP and MP interact with one another and it is possible that this interaction is involved in trafficking of naked and/or packaged virus DNA from nuclei through nuclear pores, to the cell periphery, through plasmodesmata, and into the nuclei of neighboring cells.

Molecular Biology of MSV Replication As with other geminiviruses, MSV replicates by both a rolling-circle mechanism (rolling-circle replication, or RCR; (Fig. 6)) and a recombination-dependent mechanism (recombination dependent replication or RDR). As with other rolling-circle replicons, MSV

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Induction of host cell’s DNA replication machinery Gene expression

2

MP expression 4

Earlier

RepA expression? CP expression

Later Later

5

Movement into the nucleus

Rep expression Earlier Later Oligomer formation

Completion of (−)strand synthesis

ssDNA:CP complex formation?

Replicativeform DNA Initiation of RCR and (+)-strand synthesis

Earlier 1 Later

Later

Leafhopper transmission Movement as particles?

Negative regulation of replication

3 Rolling-circle replication

Earlier

Encapsidation

ssDNA:CP:MP complex formation?

Negative regulation of C sense transcription and induction of V sense transcription

Movement of dsDNA?

Movement and encapsidation

Movement out of the nucleus

Earlier

Initiation of (−)strand synthesis

Termination of (+)-strand synthesis and release of old (+)-strand

Transport to the nucleus of new cell

Movement as ssDNA:CP:MP?

Fig. 6 Summary of the MSV infection process. Early during an infection following the synthesis of a dsDNA replicative form (RF; 1) RepA is most likely expressed and induces a cellular state in which viral DNA replication can occur (2). Rep is also expressed early and RCR begins (3). At a later point in the infection process, following genome amplification and possibly Rep and/or RepA induction of the V sense promoter, MP and CP are expressed (4), and movement and encapsidation occur (5). Represented here is movement of un-encapsidated ssDNA, but it should be noted that it is possible that dsDNA and/or encapsidated ssDNA may also be moved either cell to cell or systemically within the phloem of plants. Whereas the involvement of MSV CP and MP in movement has been demonstrated, the mechanics of the process are obscure. While the probable timing of events is indicated, it is unlikely, for example, that absolutely no MP and CP expression occurs during the earlier stages of the infection process. ssDNA is represented by blue lines, dsDNA by bold black lines, and RNA by orange lines.

replication is discontinuous with virion strand replication being initiated from the hairpin structure in the LIR and complementary strand synthesis being initiated from a short 80 nt primer-like molecule synthesized on the SIR of newly replicated virion strands.

Particle Assembly and Movement Besides being the primary location of replication, the nucleus is also the site of virus particle assembly. CP molecules in the nucleus nonspecifically bind virion strands released during RCR (there is no known encapsidation signal in mastrevirus genomes), arresting the synthesis of new RF DNAs. Viral ssDNA molecules are packaged into particles that aggregate to form large paracrystalline nuclear inclusions. Crystalline arrays of MSV particles have also been detected outside nuclei within physiologically active phloem companion cells, and inside the vacuoles of dead and dying cells within chlorotic lesions. These lesions are caused by an as yet unexplained degeneration of chloroplasts in infected cells. The mechanistic details of MSV cell-to-cell movement are still obscure, but it seems to involve an interaction between the CP, MP, and viral DNA. Besides requiring the coordinated interactions of viral gene products and DNA, the successful movement of MSV genomes from infected to uninfected cells is strongly dependent on the extent of plasmodesmatal connections between neighboring cells. Also, in maize it appears as though certain cell types are more sensitive to MSV infection than others. For example, in maize leaves the virus infects all photosynthetic cell types (e.g., mesophyll and bundle sheath cells) but despite abundant plasmodesmatal connections between photosynthetic, epidermal, and parenchyma cells, MSV is only rarely detectable in the latter two cell types. It is unknown whether systemic movement of geminiviruses within plants simply relies on normal cell-to-cell movement to deliver genomic DNA into the phloem, or whether viral DNA is specifically packaged for long-distance transport. It is possible that

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cell-to-cell movement might involve un-encapsidated ss- or dsDNA but that long-distance movement in the phloem might require encapsidation. Long-distance movement of MSV within infected plants occurs via phloem elements and it is believed that MSV is incapable of invading the root apical, shoot apical, and reproductive meristems due to the absence of developed vasculatures in these tissues. Thus, the virus is not found in tissues that develop into gametes and is therefore not seed-borne. It also does not appear to travel within plants from sites of infection into older uninfected tissues. Within the shoot apex where most productive MSV replication occurs, MSV first enters developing leaves at approximately plastochron five. While the virus is restricted to the developing leaf vasculature before plastochron 12, it is likely that the development of metaphloem elements at approximately plastochron 12 provides an opportunity for the virus to escape the vasculature into the photosynthetic cells of the leaf. Metaphloem develops with the abundant plasmodesmatal connections required for efficient loading of photoassimilates once the leaf emerges from the whorl. Before emergence, however, the developing photosynthetic tissues are still net importers of photoassimilates and the virus most likely moves into these cells through their plasmodesmatal connections with the metaphloem. On the leaves, the pattern of chlorotic streak-like lesions that characterizes MSV infections is directly correlated with the pattern of virus accumulation within the leaves and the virus can only be acquired by leafhoppers from these lesions. The degree of chlorosis that occurs within lesions can differ between MSV isolates and is related to the severity of chloroplast malformation that occurs in infected photosynthetic cells.

Control of MSD Although effective control of MSD in cultivated crops is possible with the use of carbamate insecticides, and it is possible to avoid leafhopper infestations by varying planting dates, the fact that small farmers cannot generally use these options means that the development and use of MSV-resistant crop genotypes is probably the best way to minimize the impact of MSD on African agriculture. MSV resistance is associated with up to five separate alleles conferring a mixture of both dominant and recessive traits, none of which are sufficient by themselves to prevent MSV infections. Despite great successes achieved in the development of maize genotypes that tolerate MSV infection without significant yield loss, there has been only limited success in the field. For example, severe infections of so-called MSV-tolerant genotypes can occur when they are grown under environmental conditions different from those in which the plants were selected, meaning that distinct geographical growing areas may each require genotypes with MSV resistance that is tailored to those areas. The problem facing breeders is that natural genetic resistance to MSD is not usually associated with desirable agronomic traits such as good yield. It can therefore be difficult to transfer resistance traits without also transferring undesirable characteristics. Moreover, the number of alleles involved means that successful breeding takes years for each release. Even in the absence of any predictive modeling of sporadic MSD outbreaks, most farmers would still prefer to gamble on the use of higher-yielding MSV-sensitive genotypes. Efforts are also currently underway to introduce MSV resistance traits into commercial maize genotypes by genetic engineering. This could have the advantage of enabling the direct transfer of single-gene resistance, without linkage to undesirable characteristics, to many different breeding lines suited to different environmental conditions. However, up till now this strategy has been limited by negative public perception of genetically modified organisms, and the expensive and time-consuming risk assessment necessary to ensure a safe feed and food product.

MSD Epidemiology There are loose correlations between MSD incidence and both environmental conditions and agricultural practices. Environmental influences on MSD epidemiology are mostly driven by a strong correlation between rainfall and leafhopper population densities. For example, drought conditions followed by irregular rains at the beginning of growing seasons tend to be associated with severe MSD outbreaks. Also, maize planted later in the growing season tends to get more severely infected than that planted at the beginning of the season, probably due to steady increases in leafhopper numbers and inoculum sources over the course of the season. As is the case with most insect-borne virus diseases, however, the incidence of MSD is erratic. Whereas MSD can devastate maize production in some years, in others it has only a negligible effect. The reason for this is that, apart from MSD epidemiology being strongly dependent on environmental variables, it is also the product of complex interactions between the various MSV leafhopper vector and host species, and an as yet unknown number of virus strains.

Leafhopper Vectors Serious MSD outbreaks are absolutely governed by leafhopper acquisition and movement of severe MSV isolates from infected plants (wild grasses or crop plants) to sensitive, uninfected crop plants. The distance that MSV spreads from a source of inoculum

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is determined by the movement behavior of leafhoppers. Distinct long- and short-distance flight morphs have been detected among certain Cicadulina populations. It is believed that the long-flight morphs are a migratory form and, as such, these may play an important part in the rapid long-distance spread of virulent MSV variants. Migratory movement is more common in certain Cicadulina species than in others and it is probably influenced by environmental conditions. The dynamics of primary infection following leafhopper invasion of a susceptible maize crop are influenced by leafhopper population densities, the proportions of viruliferous individuals in populations, and virus titers within these individuals. Disease spread within individual maize fields is apparently linear when only a few viruliferous leafhoppers are involved in transmission, but becomes exponential once the number of insects exceeds one individual per three plants.

Plant Hosts Although attempts to understand the dynamics of MSD epidemics have focused primarily on vector population dynamics and behavior, an important component of MSD epidemiology is the population density, turnover, and demographics of the over 80 grass species that are both MSV and vector hosts. Because Cicadulina species favor certain annual grass hosts for mating and oviposition, the species composition of grass populations that vary seasonally in any particular area will directly influence leafhopper population densities and feeding behaviors in that area. The species composition and age distribution of grasses (including cultivated crops) in an area may also affect the amount of MSV inoculum available for transmission in that area. While MSV infects at least 80 of the 138 grass species that leafhoppers feed on, both the susceptibility of these grasses to MSV infection and the severity of symptoms that occur following their infection may be strongly influenced by a number of factors. While sensitivity to infection can vary substantially from species to species, it can also vary within a species with genotype and plant age at the time of inoculation: for example, plants from many species, including maize, generally become more resistant to MSV infection with age, thereby reducing the inoculum available for transmission to other plants.

The Virus While efforts are underway to promote the widespread cultivation of MSV-resistant maize in Africa, surprisingly little is known about the MSV populations that will confront these new genotypes. Although to date eleven major MSV strain groupings have been discovered, it is unknown whether any other than the maize-adapted MSV-A strain play an important role in the epidemiology of MSD. MSV-B, -C, -D, and -E isolates only produce very mild symptoms in MSV-sensitive maize genotypes and are therefore unlikely to pose any significant direct threat to maize production. Mixed MSV-A and -B infections have, however, been detected in nature and there is also strong evidence of recombination occurring between these strains. It is therefore possible that MSV-B, and possibly other MSV strains, may indirectly influence MSD epidemiology through recombination with MSV-A-type viruses. Recombination has been linked with the emergence of a number of geminivirus diseases and it may also have contributed to the original emergence of MSV-A. It is plausible therefore that it could also eventually contribute to the evolution of MSV-A variants (or even variants of the other MSV strains) with elevated virulence in resistant maize genotypes.

Future Threat MSV is rightly regarded as a significant potential threat to maize production outside of Africa: while the vectors do not occur outside of Africa, there is no obvious reason that they would not survive in other equatorial or temperate regions of the world such as the southern USA, South America and Eurasia. It is a distinct possibility that they could inadvertently spread, or even be deliberately taken, to these regions. If, following the establishment of a MSV vector species outside of Africa, MSV was then introduced, the virus would inevitably begin spreading within native grasses, cultivated maize and the other cereals that it is known to infect. As none of the maize varieties grown outside of Africa has the slightest resistance to MSV, the probability of severe economic consequences would be very high.

See also: Emerging Geminiviruses (Geminiviridae). Geminiviruses (Geminiviridae). Plant Resistance to Geminiviruses

Further Reading Bosque-Pérez, N.A., 2000. Eight decades of maize streak virus research. Virus Research 71, 107–121. Boulton, M.I., 2002. Functions and interactions of mastrevirus gene products. Physiological and Molecular Plant Pathology 60, 243–255. Damsteegt, V., 1983. Maize streak virus. Part I: Host range and vulnerability of maize germ plasm. Plant Disease 67, 734–737. Efron, Y., Kim, S.K., Fajemisin, J.M., et al., 1989. Breeding for resistance to maize streak virus – A multidisciplinary team-approach. Plant Breeding 103, 1–36. Fuller, C., 1901. First Report of the Government Entomologist, 1899–1900. P. Davis. http://www.mcb.uct.ac.za//msv/fuller.htm. (accessed February 2008).

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Harrison, B.D., Barker, I., Bock, K., et al., 1977. Plant viruses with circular single-stranded DNA. Nature 270, 760. Martin, D.P., Willment, J., Billharz, R., et al., 2001. Sequence diversity and virulence in zea mays of maize streak virus isolates. Virology 288, 247. McLean, A.P., 1947. Some forms of streak virus occurring in maize, sugarcane and wild grasses. Science Bulletin of Department of Agriculture for Union of South Africa 265, 1–39. Palmer, K.E., Rybicki, E.P., 1998. The molecular biology of mastreviruses. Advances in Virus Research 50, 183–234. Storey, H.H., 1925. The transmission of streak disease of maize by the leafhopper. Balclutha mbila naudé. Annals of Applied Biology 12, 422–443. Varsani, A., Shepherd, D.N., Dent, K., et al., 2009. A highly divergent South African geminivirus species illuminates the ancient evolutionary history of this family. Virology Journal 25 (6), 36. doi:10.1186/1743-422X-6-36. (PubMed PMID: 19321000; PubMed Central PMCID: PMC2666655). Zhang, W., Olson, N.H., Baker, T.S., et al., 2001. Structure of the maize streak virus germinate particle. Virology 279, 471–477.

Relevant Websites http://www.mcb.uct.ac.za Molecular & Cell Biology.

Nanoviruses (Nanoviridae) Bruno Gronenborn, Institute for Integrative Biology of the Cell, CNRS, University of Paris-Sud, CEA, Gif sur Yvette, France H Josef Vetten, Julius Kühn Institute, Braunschweig, Germany r 2021 Elsevier Ltd. All rights reserved. This is an update of H.J. Vetten, Nanoviruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00644-0.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein CR-M Common region major CR-SL Common region stem-loop DAS-ELISA Double antibody sandwich form of ELISA ELISA Enzyme-linked immuno-sorbent assay EM Electron microscopy kb Kilobases; the size of a ssDNA or ssRNA molecule kDa Kilodaltons; the size of a protein MAb Monoclonal antibody

Glossary Alphasatellite Alphasatellites are single-stranded DNA molecules of 1.0–1.3 kb, which behave like virus satellites in many respects. Unlike the true non-coding satellites, alphasatellites code for a protein that specifically catalyzes only their own replication initiation. They are encapsidated in particles formed by the capsid protein of the respective helper virus. Clink The protein named Clink (for “cell cycle link”) interacts with the cell cycle regulator protein retinoblastoma and thereby modulates cell cycle regulation in favor of DNA synthesis (S-phase). Clink enhances nanovirus replication. Enzyme-linked immunosorbent assay The enzymelinked immunosorbent assay (ELISA) is a sensitive and versatile serological test with two common variants, the double antibody sandwich (DAS)-ELISA and the triple antibody sandwich (TAS)-ELISA. A double antibody sandwich (DAS)-ELISA consists of four steps (separated by individual washings): (1) the immunoglobulins (IgG) isolated from a virus-specific antiserum are used for coating of microtiter plates; (2) IgG-coated plates are incubated with extracts of samples to capture antigen, if present; (3) incubation with enzyme (usually alkaline phosphatase)labeled IgG [see (1)]; (4) incubation with substrate of the respective enzyme followed by measuring its activity, for instance by colorimetry. The triple antibody sandwich (TAS)-ELISA differs from DAS-ELISA in step (3), which is

MP Movement protein nm Nanometer(s) NSP Nuclear shuttle protein nt Nucleotide(s) PCR Polymerase chain reaction pRB Protein retinoblastoma RCA Rolling-circle amplification RCR Rolling-circle replication Rep Replication initiator protein ssDNA Single-stranded deoxyribonucleic acid TAS-ELISA Triple antibody sandwich-ELISA

replaced by two steps consisting of (i) incubation with a specific (often monoclonal) antibody from a different animal species than the animal used to produce the coating IgG in step (1), and (ii) incubation with enzyme-labeled IgG raised against IgG of that same different animal species. Replication initiator protein A replication initiator protein (Rep protein or Rep, also referred to as RC-Rep) initiates rolling-circle replication of DNA by cleaving double-stranded DNA at a defined sequence (replication origin). The Rep protein remains bound to the 50 phosphate of the cleaved strand while a polymerase uses the liberated 30 hydroxyl of the cleaved strand as primer for DNA synthesis. After one round of DNA synthesis another subunit of Rep cleaves the newly synthesized DNA at the replication origin sequence. The thus generated 30 hydroxyl attacks the phosphotyrosyl ester bond of the bound Rep subunit, and by a nucleotidyltransfer a covalently closed circular single-stranded DNA molecule is created. Rolling-circle replication Rolling-circle replication (RCR) is a process of unidirectional nucleic acid amplification, by which multiple copies of circular DNA or RNA molecules such as plasmids, bacteriophage- or eukaryote virus genomes are reproduced. Satellites Satellites are subviral nucleic acids that depend for replication, systemic movement, encapsidation, and/or vector transmission on a helper virus, with which they coinfect a host; they do not code for a protein.

Introduction In 1988 and 1991 a novel type of single-stranded DNA (ssDNA) virus causing yellowing and dwarfing diseases of legumes or banana was described. The viruses were persistently transmitted by aphids and not transmissible by sap. Unusually small, icosahedral particles measuring only 18–20 nm in diameter were isolated from infected plants. Because of the small size of the virions, the small size of individual genome components of only about 1 kb, and the dwarfing symptoms they caused, these

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viruses were referred to as nanoviruses. They differ from geminiviruses, the other group of ssDNA plant viruses, in particle morphology, genome size, number and size of DNA components, genomic organization, mode of transcription, and vector species. Common with other ssDNA viruses including geminiviruses is the mode of nanovirus replication. Unlike bipartite geminiviruses, in which the two genomic DNAs are individually encapsidated, the nanovirus genome is multipartite and consists of six or eight ssDNAs each of which is individually encapsidated in a separate particle. Here we provide a brief synopsis of our current knowledge of these viruses.

Classification and Taxonomy To date, eleven established and three recently identified members of the family Nanoviridae have been described. On the basis of differences in biology (host range, aphid vectors) and both genome size and organization, the family members (nanovirids) are subdivided into the genera Nanovirus and Babuvirus. The respective type species of these two genera are Subterranean clover stunt virus and Banana bunchy top virus. Three babuvirus and eight nanovirus species have been approved by the International Committee on Taxonomy of Viruses (ICTV). Information on the geographic distribution, important aphid vectors and some natural hosts of the nanovirids as well as the amount and type of genome sequence data available for them is given in Table 1. Crops in mainly tropical and subtropical countries of the Old World harbor babuviruses, such as Abaca bunchy top virus (ABTV) in the Philippines, Cardamom bushy dwarf virus (CBDV) in India and Banana bunchy top virus (BBTV). The latter occurs widely on Pacific Islands (including Hawaii), in Australia and Indochina but has an erratic geographic distribution in South Asia and Africa. While Subterranean clover stunt virus (SCSV) occurs only on legumes in Australia, Milk vetch dwarf virus (MDV) has been reported Table 1

Assigned and tentative members of the genera Babuvirus and Nanovirus of the family Nanoviridae Geographic distribution

Aphid vector

Natural host

Genome components

Complete sequences

Genus Babuvirus Banana bunchy top virus (type species) Abaca bunchy top virus Cardamom bushy dwarf virus

Africa, Australia, Pacific Islands, southern Asia, Borneo, Philippines India

Pentalonia nigronervosa

Musa spp.

6

4200

no data Micromyzus kalimpongensis

Musa spp. Elettaria cardamomum, Amomum subulatum

6 6

2 4160

Genus Nanovirus Subterranean clover stunt virus (type species)

Australia

8

2

8

3

8

8

8

8

8 8

1 8

8

8

8 8 7b

1 1 1

8c

2

Germany, Austria, Hungary, Denmark, Serbia, Netherlands Austria France Germany

Aphis craccivora, Aphis Trifolium subterraneum and gossypii, Acyrthosiphon other legumes pisum Aphis craccivora, Medicago lupulina, Pisum Acyrthosiphon pisum sativum Aphis craccivora, Aphis Vicia faba and other legumes fabae, Acyrthosiphon pisum Aphis craccivora, Vicia faba, Lens culinaris, Acyrthosiphon pisum Phaseolus vulgaris and other legumes No data Vicia faba Aphis craccivora, Vicia faba and many legumes Acyrthosiphon pisum, and non-legumes i.e., Megoura viciae, Myzus Nicotiana tabacum, Carica persicae papaya, Catharanthus roseus Aphis craccivora, Aphis Pisum sativum, Vicia faba and fabae, Acyrthosiphon other legumes pisum, Megoura viciae No data Pisum sativum No data Vicia cracca No data Petroselinum crispum

Iran

No data

Black medic leaf roll virus Austria, Sweden, Azerbaijan Faba bean necrotic yellows Near East, North Africa, virus Ethiopia, Spain Faba bean necrotic stunt virus

Ethiopia, Morocco, Azerbaijan, Iran

Faba bean yellow leaf virus Ethiopia Milk vetch dwarf virus China, Japan, Bangladesh

Pea necrotic yellow dwarf virus Pea yellow stunt virus Cow vetch latent virusa Parsley severe stuntassociated virusa Sophora yellow stuntassociated virusa a

Sophora alopecuroides et al.

Recently described viruses not yet assigned to the genus Nanovirus by ICTV. No DNA-U4 found; four different types of DNA-R found but M-Rep not experimentally identified. c Two different DNA-C types found; four different DNA-R types found but M-Rep not experimentally identified. b

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Fig. 1 Faba bean necrotic yellows virus particles. Negative contrast electron micrograph of virions of faba bean necrotic yellows virus. The scale bar corresponds to 50 nm. Courtesy of D.E. Lesemann and L. Katul.

from several legumes and non-legumes in Japan, China, and recently also in Bangladesh, suggesting that it has a wide distribution in several other countries of South-east Asia. Whereas Faba bean yellow leaf virus (FBYLV) was reported only from Ethiopia, Faba bean necrotic yellows virus (FBNYV) appears to have a much wider geographic distribution in West Asia, the Middle East, North- and East Africa and Europe (Spain). Faba bean necrotic stunt virus (FBNSV) has been found in Ethiopia, Morocco, Azerbaijan and Iran. Also from Iran, Sophora yellow stunt associated virus (SYSaV), a nanovirus infecting Sophora alopecuroides, a leguminous fodder crop, was reported. During recent years a number of nanoviruses have been described from Central- and Northern Europe. This situation is exemplified by Pea necrotic yellow dwarf virus (PNYDV) from Germany, Denmark, the Netherlands, Austria, Hungary and Serbia, Black medic leaf roll virus (BMLV) from Austria, Sweden and Azerbaijan, Pea yellow stunt virus (PYSV) from Austria, and Cow vetch latent virus (CvLV) from France. The most recently identified nanovirid is Parsley severe stunt associated virus (PSSaV) from Germany, the only nanovirus potentially restricted to a non-leguminous host. No nanovirids have as yet been found in the Americas. All available evidence suggests that nanovirids are quite variable in their biological and molecular properties.

Particle Properties Virions of the nanovirids are not enveloped, 17–20 nm in diameter, and presumably of an icosahedral T ¼ 1 symmetry structure containing 60 subunits. Capsomeres may be evident, producing an angular or hexagonal outline (Fig. 1). Virions are stable in Cs2SO4 but may not be stable in CsCl. The buoyant density of virions is about 1.24–1.30 g/cm3 in Cs2SO4, and 1.34 g/cm3 in CsCl. They sediment as a single particle component in sucrose rate-zonal and isopycnic Cs2SO4 density gradients. Virions have a single capsid protein (CP) of about 19 kDa. No other proteins have been found associated with virions. Each ssDNA component is encapsidated in a separate particle. Virions are strong immunogens. Most nanovirid species are serologically distinct from one another.

Genome Organization Based on DNA sequence analyses of a range of geographical isolates there is evidence that six and eight DNAs form the genome of a babu- and nanovirus, respectively (Table 1). Both encapsidated genomic nanovirid DNAs and nanovirid-associated DNAs (Alphasatellitidae) are covalently closed ssDNA circles of about 1 kb in size. A characteristic feature of all nanovirid and nanoviridassociated DNAs are two short (8–13 nt) inverted repeat sequences flanking a stretch of 11–13 nt containing a conserved nonanucleotide motif: TATTATTAC in babuviruses and TAGTATTAC in nanoviruses (exception: PSSaV, which has TATTATTAC). These sequences have the potential to form a stem-loop structure in ssDNA or a cruciform extrusion in dsDNA. Comparable DNAsequence or DNA-structure motifs, also referred to as stem-loop or hairpin-loop, are common in all ssDNA viruses that multiply their genome by rolling-circle replication (see below). Sequences 50 and 30 of the inverted repeats are also conserved among the individual genome components, and their respective extensions in both directions are quite variable depending on the virus. This region has been designated common region stem-loop (CR-SL). In extreme cases sequences of up to 160 nt (PSSaV) or 250 nt (MDV), located 30 of the inverted repeat are conserved among individual genomic DNAs.

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Fig. 2 Genome organization of nanovirids and associated alphasatellites. Individual genomic DNAs are represented as circles. A knob, often also referred to as hairpin, indicates the position of the inverted repeat sequences flanking the conserved nonanucleotide at the origin of viral strand replication. Component designations are displayed inside each circle. Arrows represent coding sequences of the respective encoded proteins. Protein names are given below DNA component names. Constituents of the babuvirus genome are framed in red and those of the nanovirus genome in green. DNAs of the top row have homologs in viruses of both genera; DNAs of the bottom row are unique for either nano- or babuviruses as indicated by the respective frame. Alphasatellites differing in numbers and types are frequently found associated with members of each genus of the Nanoviridae, hence included in the black frame. Graphics courtesy of I. Grigoras.

Apart from the CR-SL both babu- and nanoviruses possess a second conserved region that is common to all genomic DNAs. In babuviruses it is located 50 of the inverted repeats at distances that vary from about 50 nt to about 350 nt. The region is referred to as common region major (CR-M) and has a length of around 60 nt in ABTV, 65–92 nt in BBTV and 75–80 nt in CBDV. There is no such rather uniform CR-M in nanoviruses. Instead, short stretches of conserved sequences from 17 to 34 nt constitute a so-called common region II (CR-II), which is located about 70 nt 50 of the CR-SL. In some cases (e.g., FBYLV) there is no CR-II, and in others (e.g., MDV and PNYDV) rather long sequence stretches 50 of the inverted repeats are conserved between all genomic DNAs, except DNA-R. The genomic DNAs of nanoviruses are about 100 nt shorter than those of babuviruses and the coding region of DNA-R terminates closely 50 of the CR-SL. This restricts the possibility for longer common conserved sequences due to constraints dictated by aa conservation. Hence only short CR-II sequences are common to all genomic nanovirus DNAs whereas longer CR-II stretches may occur between nanovirus genomic DNAs other than DNA-R. The genomic information of each nanovirid is distributed over six or eight different molecules of the aforementioned ssDNA type (Table 1). Upon infection, the circular ssDNAs are converted into circular double-stranded DNAs (mini-chromosomes) that serve as templates for gene expression. Each of these DNA molecules contains a single open reading frame of positive (virion-sense) polarity, preceded by a promoter sequence with a TATA box and followed by a polyadenylation signal (Fig. 2). The fact that a typical set of respectively six or eight distinct DNAs has been consistently identified from babu- and nanoviruses suggests that the babuvirus genome consists of six DNAs and the nanovirus genome of eight DNAs. DNA-R, -S, -C, -M and -N have been identified from all nanovirids (Fig. 2, Table 1). DNA-U1, -U2 and -U4 homologs are typical for all nanoviruses but are absent from babuvirus genomes. In contrast, DNA-U3 is restricted to babuviruses and is absent from nanovirus genomes. Transcript analyses of BBTV demonstrated a single virion-sense transcript per genome component. A minor transcript was found internal of the DNA-R ORF. For nanoviruses only transcripts of FBNYV and SCSV DNA-R and FBNYV DNA-C, along with a FBNYV-associated alphasatellite (see below) transcript have been mapped in detail. They are 30 -polyadenylated and their 50 ends map shortly after the TATA sequences preceding the respective ORFs. Consistent with these transcript analyses is the notion that each nanovirid DNA encodes a single gene defined by the respective ORFs and their flanking transcription initiation and termination signals. Interestingly the DNA-R transcripts of the two nanoviruses studied, FBNYV and SCSV, are terminally redundant by 54–161 nt (FBNYV) and 49 nt (SCSV). Considering the positions of TATAA box- and polyadenylation sequences of DNA-R molecules of all nanoviruses, such a terminal redundancy is predicted to be a unique and distinctive feature of nanovirus DNA-R components in general. Its biological significance remains, however, unknown. Although eight DNAs (DNA-R, -S, -C, -M, -N, -U1, -U2, and -U4) had invariably been found associated with nanovirus infections, the question of ‘how many DNAs constitute a nanovirus genome’ had remained open for some time until experimental infection using individually cloned FBNSV DNAs became possible. In this way, the type of disease symptoms induced as well as the efficiency of vector transmission of the thus reconstituted virus could be analyzed. Due to this considerable technical accomplishment it was demonstrated that without DNA-R, DNA-S and DNA-M no infection was possible. DNA-C, DNA-U1, and DNA-U2 were required for

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efficient disease development and vector transmission but not absolutely essential. Interestingly, infections devoid of DNA-N resulted in a disease phenotype indistinguishable from infections with wild-type virus. However, neither cowpea- nor pea aphids, the most efficient nanovirus vectors, were able to transmit virions produced in DNA-N-less infections. Hence DNA-N encodes the transmission helper factor postulated earlier. Finally, DNA-U4 proved to be non-essential for infection and sustained virus transmission under experimental conditions. Although these results were obtained with FBNSV in faba bean, it is very likely that they are equally valid for other nanoviruses. Experimental infection using cloned genome DNAs has been successfully demonstrated also for BMLRV, FBNYV and PNYDV. To date, no comparable experimental infection with DNAs of any babuvirus has been achieved. There are indications that the genome organization of nanoviruses can sometimes be even more complex. In infections by each of the recently discovered SYSaV and PSSaV four DNA-R-like components were identified. However, which one of them encodes a Rep protein capable of acting as a master-Rep protein, i.e., catalyzing replication initiation of the other genomic DNAs, has not been determined. These findings illustrate well that the current picture of nanovirus genome organization may still be in flux.

Properties and Functions of Nanovirid Proteins In addition to the capsid protein (CP) of about 19 kDa encoded by DNA-S, at least 5–7 non-structural proteins are encoded by the other 5 or 7 genomic DNAs of nanovirids (Table 1, Fig. 2). Besides acting as the structural protein required for virion formation, the protein encoded by DNA-S of BBTV has also been identified as a silencing suppressor. One of the best-studied nanovirid proteins is the ca. 33 kDa replication initiator protein (Rep) encoded by their DNA-R. It belongs to the vast protein family of rolling-circle replication (RCR) initiator proteins of ssDNA viruses, bacteriophages, RCR plasmids and RCRtransposable elements (helitrons). Nanovirids express a unique Rep protein that is essential for replication initiation of all genome segments, six in case of babuviruses and eight in case of nanoviruses. This Rep protein recognizes specific sequence motifs in the so-called origin region, DNA stretches flanking the inverted repeat sequences that are shared by all genome components. By contrast to the Rep proteins encoded by DNA-R, those encoded by the nanovirid-associated alphasatellites are not able to trigger replication initiation of any nanovirid genome component. This led to the name ‘master replication initiator protein’ (M-Rep) for the DNA-R-encoded protein. Several salient features characteristic of RCR-initiator proteins have been demonstrated also for a nanovirus (FBNYV) M-Rep protein, e.g., origin DNA cleavage and nucleotidyltransferase activity and ATPase activity. Also, the three-dimensional structure of the DNA-binding and endonuclease domain of FBNYV M-Rep has been determined, revealing a striking similarity to those of geminiviruses, circoviruses, parvoviruses, polyomaviruses and papillomaviruses. By analogy to geminivirus Rep proteins, nanovirus M-Rep proteins are probably also the smallest helicases of the superfamily 3 helicases (S3H), experimental proof however pending. Replication initiator and capsid proteins are, in addition to the so-called ‘stem-loop’ motif in the ssDNA, common elements shared by the very large group of so-called CRESS (circular Rep-encoding single-stranded DNA) viruses. The third protein essential for nanovirid infection, apart from CP and M-Rep, is the 13–14 kDa protein encoded by DNA-M. Experimental evidence obtained for the protein encoded by DNA-M of BBTV indicates that it is involved in cell-to-cell movement. A stretch of 25–30 hydrophobic residues at its N-terminus is conserved in all its nanovirid homologs, consistent with a function as movement protein (MP). As shown for the BBTV CP, the MP of BBTV is also able to act as a suppressor of RNA silencing by reversing systemically established gene silencing. The protein encoded by DNA-C is 19–20 kDa in size and has an N-terminal F-box sequence and the aa motif LxCxE characteristic for proteins binding to the cell-cycle regulator protein retinoblastoma (pRB). Moreover, the DNA-C encoded protein has been shown to interact with pRB. Therefore, the name “Clink” for ‘cell cycle link’ protein was coined for it. Based upon evidence obtained for the activity of the BBTV DNA-N encoded 17–18 kDa protein it was proposed to act as a nuclear shuttle protein (NSP). In addition to that it has been shown that the NSP homologs of FBNSV and PNYDV are mandatory for virus transmission by vector aphids. Another biological property of a nanovirus NSP is its interaction with cellular proteins involved in stress granule formation, as shown for PNYDV. It is speculated that this way NSP may inhibit stress granule formation, a protective cellular response to virus infection. Little is known about the biochemical properties or biological functions of the proteins encoded by DNA-U1, -U2, -U3, and -U4, where “U” stands for “unknown function”. As mentioned above, FBNSV DNA-U1 and DNA-U2 influence disease symptoms and the total amount of virions produced. It is plausible to assume that this is due to the action of the respective encoded proteins U1 (17–18 kDa) and U2 (14–15 kDa). The experimental disruption of a nuclear localization signal in the U1 protein altered its subcellular localization, yet no specific function was thus revealed for the U1 protein. The U3 protein deduced from an ORF on DNA-U3 of BBTV is ca. 9 kDa in size. There are no data about its potential function. A transcript spanning the BBTV DNA-U3 ORF has been mapped. A homologous ORF is absent in the DNAs termed U3 of ABTV and CBDV. It remains, therefore, questionable whether these DNAs can be considered being true homologs of BBTV DNA-U3. The deduced U4 protein encoded by DNA-U4 of the nanoviruses has a size of ca. 13 kDa. The fact that DNA-U4 homologs were detected in all nanovirus infections except those by PSSaV underpins the importance of the respective DNA-U4 product in natural infections. This notion appears valid despite the observation that DNA-U4 was not required for development of disease symptoms in experimental infections of faba bean seedlings with FBNSV. Moreover, FBNSV lacking DNA-U4 was serially transmitted to faba bean by aphids for 42 years. To date, only in planta protein-protein interaction data of bi-molecular fluorescence complementation assays employing PNYDV U4 protein are available and revealed auto-interaction and interaction with the PNYDV movement protein (MP).

Nanoviruses (Nanoviridae)

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Nanovirid Replication The knowledge about geminivirus replication can largely be transposed also to nanovirid replication. Upon inoculation by aphid vectors virions enter host cells, presumably phloem parenchyma cells. As shown for BBTV and geminiviruses the ssDNAs are converted with the help of virion-encapsidated primers into double-stranded transcriptionally active mini-chromosomes. Most probably, this step occurs in the nucleus, but whether virions or only ssDNAs enter the nucleus is not known. Once a sufficient amount of M-Rep protein is available, a subunit of M-Rep cleaves the phosphodiester backbone of the origin DNA between nt T7 and A8 of the nonanucleotide TAG/TTATT-AC, becomes covalently linked to the 50 phosphate of A8 via its active site tyrosine (Y79 in case of FBNYV M-Rep), and a host DNA polymerase utilizes the 30 -OH of T7 as primer for DNA synthesis. After one round of replication, a second subunit of M-Rep cleaves the newly synthesized nonanucleotide sequence between T7 and A8, becomes linked to the 50 phosphate of the newly synthesized A, and the 30 -OH of T7 attacks the phosphotyrosyl ester bond between the first M-Rep subunit and the DNA, which leads to a nucleotidyltransfer of the M-Rep-linked 50 phosphate to the newly available 30 -OH. As a result, a phosphodiester bond is formed resulting in a liberated circular single-stranded DNA molecule, and a free tyrosine of the first M-Rep subunit, a step that requires no energy. These steps are repeated as long as rolling-circle replication continues. This so-called ‘nicking and joining’ activity by M-Rep proteins of DNA with the conserved nonanucleotide sequence has been experimentally demonstrated in vitro for BBTV and FBNYV as well as for one Rep of a FBNYV-associated alphasatellite. Replication of nanovirus DNAs is enhanced by the action of Clink, the nanovirid-encoded cell cycle modulator protein that is assumed to reprogram the cell cycle of non-proliferating plant cells towards S-phase. One interesting and quite unique characteristic of nanovirus replication has been discovered using FBNSV as a model: multiplication of each individual genomic DNA to a different and component-specific maximum level, the so-called genome formula. This genome formula differs between the two hosts faba bean (V. faba) and barrel medic (Medicago truncatula) and is also different in its vector, the pea aphid A. pisum.

Taxonomy and Phylogeny Nanovirids share a set of five homologous DNA components, referred to as DNA-R, -S, -C, -M and –N (Fig. 2). Three other DNAs (DNA-U1, -U2 and -U4) encoding proteins of as yet unknown functions have been identified from nanoviruses and one further DNA (DNA-U3) only from babuviruses. Viruses of the individual species typically have narrow host ranges. Under natural and experimental conditions all nanoviruses only infect dicotyledonous plants. Most nanoviruses have largely overlapping host ranges and predominantly infect a range of leguminous species (Fabaceae) and a few other dicotyledonous species (Table 1), whereas babuviruses have been reported only from a few monocotyledonous species, such as the Musaceae or Zingiberaceae. Since these plants are colonized by a restricted number of (often monophagous) aphid species, the nanoviruses studied for their biological properties are transmitted by the cowpea aphid Aphis craccivora, the pea aphid Acyrthosiphon pisum and some other aphid species predominantly feeding on legumes. The vector of PSSaV, a nanovirus identified from parsley, a non-leguminous host, is unknown. The currently known babuviruses are vectored by the banana aphid Pentalonia nigronervosa and Micromyzus kalimpongensis. The other major differences between babu- and nanoviruses are that (1) nanovirus DNA components range in size from 962 to 1045 nt and are thus slightly smaller (by ca. 100 nt) than those of babuviruses (1013–1116 nt), (2) babuviruses are serologically unrelated to members of the genus Nanovirus, and (3) nanoviruses share low levels of aa sequence identities ranging from 14% to 58% in individual genes with babuviruses. Moreover, nanovirus DNA-R transcripts are terminally redundant unlike those of the babuviruses. Phylogenetic analyses of the nt sequences of nano- and babuviruses demonstrated (Fig. 3) that the five babuvirus sequences representing three species form a cluster clearly distinct from the nanoviruses. The latter consist of a group of eight related species from Asia, Europe and North Africa and three more distantly related species each from Australia (SCSV), Iran (SYSaV) and Europe (PSSaV). Comparison of the deduced aa sequences of the five proteins encoded by DNA-R, -S, -C, -M, and –N by representative isolates of the nanovirid species listed in Table 1 shows that the master Rep protein (M-Rep) is the most conserved protein. It has an overall aa identity of 80%–97% between members of the same genus and 54%–58% between the two genera. The second most conserved protein is the NSP with 55%–91% intra-genus aa identity and 41%–50% inter-genus aa identity. Capsid protein, MP and Clink protein follow with maximum intra-genus aa identities of 88%, 81% and 76%, respectively. The sequence conservation of these proteins between babu- and nanoviruses is lower than that of M-Rep and NSP with maximum sequence identities of 33% (MP), 27% (CP) and 25% (Clink), respectively. If one considers intra-genus variability of the respective proteins there is no significant difference in variability among homologous nanovirus proteins and among homologous babuvirus proteins. The variability of nanovirus proteins appears slightly higher, but this is probably due to the greater number of nanovirus genomes sequenced, including one from parsley, a non-leguminous species. With respect to the currently used criteria for species discrimination in the family, the reader is referred to the latest ICTV description of the family Nanoviridae. Based on similarities in particle morphology and size and based on the identification of an alphasatellite-like DNA (1.3 kb), Coconut foliar decay virus (CFDV), the causal agent of a severe disease of coconut palms in Vanuatu, is listed as an unassigned species of the family Nanoviridae. However, following rolling-circle amplification and deep sequencing analyses of historic and fresh samples twelve distinct circular DNAs have recently been identified. Sequence analysis suggested that nine of the DNAs represent alphasatellites (Fig. 4) and that the CFDV genome is possibly tripartite, consisting of an alphasatellite-like DNA (1.3 kb), a 1.3 kb DNA encoding a capsid protein related to that of grabloviruses (Geminiviridae), and a unique DNA-gamma (641 nt) that

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Fig. 3 Maximum likelihood phylogenetic tree of nanovirid genomes. Individual DNAs of homologous genome components were concatenated in the order DNA-R, -S, -C, -M, -N and aligned using Muscle. The tree was constructed in MEGA7 using the GTR þ G þ I nt substitution model and is mid-point rooted. The scale bar corresponds to 0.1 substitutions per site. Percent bootstrap support of 5000 non-parametric replicates is shown. Branches below 50% support are collapsed.

may encode a movement protein. Thus, CFDV differs clearly from established members of the family Nanoviridae not only in genomic organization, DNA sequence as well as in the type of insect vector (planthopper versus aphids), implicating that CFDV possibly represents a member of a new family of plant viruses.

Nanovirid Evolution Virus genomes that consist of single-stranded nucleic acids are known to undergo rapid sequence evolution. This is also true for nanovirids as has been shown for FBNSV. It has a rate of 1.8  10–3 substitutions per nt per year, the highest substitution rate among ssDNA viruses and is only outrivaled by those of influenza A viruses and human immunodeficiency viruses. Any potential disadvantage caused by such an elevated nt substitution rate is probably compensated for by the fact that every type of genome component acts individually as a quasi-species-like swarm of molecules. Also, reassortment of individual genomic DNAs or groups thereof and recombination between them would alleviate the genetic load imposed by such a high nt substitution rate. Indeed, both recombination and reassortment of genome component has been observed in nanovirids, for instance BBTV and a range of nanoviruses. In fact, given the extent of common sequences shared by several genomic DNAs of the hitherto identified nanoviruses, recombination between them is probably frequent during replication. Interestingly, each type of nanovirus genomic DNA does not need to be present simultaneously in an individual cell of a host plant to trigger a productive infection, as has been shown recently for FBNSV. This finding led to the assumption that nanovirus infection, exemplified by FBNSV, does not necessarily need to operate at the level of individual cells but rather at the level of an organism, here a plant host. Such a behavior, coined “multicellular way of life”, is unique for multipartite viruses and represents a novel concept in virology. A multicellular way of life would alleviate the problems caused by the very high multiplicity of infection required for a virus with an octopartite genome if it were productively operating only at the level of a single cell. This observation also fits with the proposition that the de novo origin of multipartite ssDNA viruses may postdate the emergence of multicellular eukaryotes.

Nanovirid-Associated Alphasatellites In addition to the genomic DNAs of nanovirids, a large number of additional DNAs encoding Rep proteins have been found associated with almost all nanovirid infections. These Rep-encoding DNAs are quite diverse and phylogenetically distinct from any nanovirid DNA-R, however related to Rep-encoding DNAs associated with begomoviruses and CFDV (Fig. 4). Collectively, these DNAs are now referred to as alphasatellites (Alphasatellitidae). Using conventional approaches of sequence analysis, nanovirusassociated alphasatellites were often detected prior to the essential nanovirid DNA-R components. This is explained by the fact that these alphasatellites are much more abundant than any genomic DNA of the nanovirus they associate with, as was revealed by

Nanoviruses (Nanoviridae)

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Fig. 4 Maximum likelihood phylogenetic tree of nanovirid DNA-R components and nanovirid-associated alphasatellites. DNAs were aligned using Muscle and a mid-point rooted tree was constructed in MEGA7 using the GTR þ G þ I nt substitution model. The scale bar corresponds to 0.1 substitutions per site. Percent bootstrap support of 1000 non-parametric replicates is shown. Branches below 80% support are collapsed.

deep sequencing analyses. Deep sequencing also permitted the detection of up to seven or fourteen distinct alphasatellites in single plants infected with PSSaV or SYSaV, respectively. Begomovirus- and CFDV-associated alphasatellites are larger (ca. 1300 nt) than nanovirid-associated alphasatellites (ca. 1000 nt) due to the inclusion of an A-rich sequence within their non-coding region. Nanovirus-associated alphasatellites are individually encapsidated in nanovirid-like particles. Proof of their encapsidation stems from the first sequenced FBNYV-associated Rep-encoding DNA and was recently also provided by sequencing DNA extracted from CFDV virions. In contrast to the activity of the M-Rep proteins encoded by DNA-R, alphasatellite-encoded Rep proteins are only capable of initiating replication of their cognate DNA, as shown for FBNYV- and BBTV-associated alphasatellites. Despite the fact that almost all nanovirid infections are accompanied by varying numbers of alphasatellites, little is known about their influence on the pathogenicity or epidemiology of the respective helper viruses. In this context it is noteworthy that two begomovirusassociated alphasatellites were shown to act as silencing suppressors. Recently alphasatellites were classified in the newly established family Alphasatellitidae, subdivided in the subfamilies Geminialphasatellitinae (five genera) and Nanoalphasatellitinae (seven genera).

Transmission and Host Range Tissue Tropism and Means of Transmission SCSV and FBNYV have been shown to replicate in inoculated protoplasts. All assigned viruses are restricted to the phloem tissue of their host plants and are not transmitted mechanically and through seeds. Apart from graft transmission, vector transmission had been the only means of experimentally infecting plants with nanovirids for a long time, until infectivity of purified FBNYV virions by biolistic bombardment was demonstrated. Meanwhile, vector transmission of virus derived from experimental infections of various hosts by cloned nanovirus genome components is possible for BMLRV, FBNSV, FBNYV, and PNYDV, and has been used to identify the virus-encoded helper factor of aphid transmission.

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Transmission by Aphids Under natural conditions, all nanovirids are transmitted by certain aphid species, in which they can persist for many days or weeks without replicating in their vectors. At least 10 aphid species belonging to the tribes Macrosiphini and Aphidini were reported as vectors of nanovirids. Whereas only one aphid species, the banana aphid (Pentalonia nigronervosa) and Micromyzus kalimpongensis has been reported as respective vector of BBTV and CBDV, several aphid species transmit FBNYV, MDV, and SCSV. The cowpea aphid (Aphis craccivora) appears to be the major natural vector of these viruses as it is the most abundant aphid species on legume crops in the afflicted areas and was among the most efficient vectors under experimental conditions. Other aphid vectors of FBNYV are the black bean aphid (Aphis fabae) and the pea aphid (Acyrthosiphon pisum). SCSV has been reported to be vectored also by the cotton aphid (Aphis gossypii), the green peach aphid (Myzus persicae) and the potato aphid (Macrosiphum euphorbiae), and PNYDV by the cowpea- and pea aphid and the vetch aphid (Megoura viciae). The vector aphids of ABTV, FBYLV, PYSV, SYSaV, and PSSaV have not been determined (Table 1). Transmission studies showed that aphids are able to transmit BBTV, FBNYV and SCSV following short acquisition and inoculation access feeding periods of about 30 min each. Although viruliferous aphids often retain transmission ability for life, nanovirids do not multiply in their insect vectors. Together with the observation that longer acquisition and inoculation access feeding periods resulted in higher transmission rates, the long persistence of nanovirids in their insect vectors indicates that they are transmitted in a circulative persistent manner similar to that of luteoviruses. In contrast to observations for luteoviruses, however, immunofluorescence localization of BBTV in the banana aphid Pentalonia nigronervosa indicates that BBTV antigens occur in cells of the anterior midgut and the principal salivary glands. This translocation resembles that of the geminiviruses, the other family of plant ssDNA viruses. For FBNYV it has been demonstrated that purified virions alone are not transmissible by its aphid vector, regardless of whether they are acquired from artificial diets using membrane feeding techniques or directly microinjected into the aphid’s hemocoel. However, faba bean seedlings biolistically inoculated with intact virions or viral DNA developed symptoms typical of FBNYV infections and were efficient sources for FBNYV transmission by aphids. These observations together with results from complementation experiments suggest that FBNYV (and other nanovirids) require a virus-encoded helper factor for its vector transmission that is absent from purified virion preparations. Using the individually cloned DNAs of FBNSV and PNYDV, it has recently been demonstrated that the proteins encoded by DNA-N of FBNSV and PNYDV are mandatory for virus transmission by vector aphids. This sets the nanovirids clearly apart from other viruses transmitted in a circulative manner, such as geminiviruses and luteoviruses. The uniqueness of nanovirids is also substantiated by the observation that the so-called genome formula of FBNSV in its vector, the pea aphid A. pisum, differs from those in two host plants, faba bean (V. faba) and barrel-medic (Medicago truncatula).

Host Range Nanoviruses often have largely overlapping host ranges by infecting over 50 legume species and only a few non-legume species under experimental and natural conditions. FBNYV, MDV and SCSV naturally infect a range of leguminous species, whereas BBTV and ABTV have been reported only from Musa species and closely related species within the Musaceae, such as abaca (M. textilis Née) and ensete (Ensete ventricosum Cheesem). There are no confirmed non-Musa hosts of BBTV. Symptoms of BBTV include plant stunting, foliar yellowing, and most characteristic dark green streaks on the pseudostem, petioles, and leaves. CBDV has been reported from black cardamom (Amomum subulatum) and green cardamom (Elettaria cardamomum), two closely related species of the family Zingiberaceae. All economically important natural hosts of FBNYV, MDV, and SCSV are legumes, in which these viruses generally cause plant stunting and a range of foliar symptoms, such as leaf deformations and chlorosis or reddening. Although FBNYV infects 450 legume species and only few non-legume species (Stellaria media, Amaranthus and Malva spp.) under experimental and natural conditions, major legume crops naturally infected by FBNYV are faba bean, lentil (Lens culinaris), chickpea (Cicer arietinum), pea (Pisum sativum), French bean (Phaseolus vulgaris) and cowpea (Vigna unguiculata). Likewise, SCSV experimentally infects numerous legume species, but its economically important natural hosts only include subterranean clover, common bean, faba bean, pea, and medics. In addition to Sophora alopecuroides, SYSaV was found in Iran in a few other legumes, such as milk vetch (Astragalus sp.), chickpea, licorice (Glycyrrhiza glabra), and lentil but also in a non-leguminous plant, wild rue (Peganum harmala; Nitrariaceae). MDV is known to cause yellowing and dwarfing in Chinese milk vetch (Astragalus sinicus L.), a common green manure crop in Japan, as well as in cowpea, faba bean, pea, and soybean. The host range of MDV appears to be larger than that of most other nanoviruses as a number of non-leguminous plants have been reported as experimental hosts of MDV, and natural infections by MDV have recently been recorded from tobacco (Nicotiana tabacum; Solanaceae) in China and papaya (Carica papaya; Caricaceae) and periwinkle (Catharanthus roseus; Apocynaceae) in Bangladesh. This and the recent identification of a novel type of nanovirus from parsley (Petroselinum crispum; Apiaceae) in Germany may indicate that there are further nanovirids which naturally infect several other non-leguminous plants.

Epidemiology and Control Based on symptomatology and transmission characteristics, luteoviruses were earlier suspected as the causative agents of several diseases now known to have a nanovirid etiology. Unlike luteoviruses, however, nanovirids generally cause more severe

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symptoms. In many nanovirid-host combinations, early infections lead to very severe effects and even premature plant death. The disease caused by BBTV is considered the most serious viral disease of banana worldwide. SCSV and FBNYV are also of considerable economic importance as they have caused serious diseases of subterranean clover in Australia and of faba bean in Egypt, respectively, leading to repeated crop failures. The same applies to PNYDV, which has occurred in certain years (e.g., 2016) at an epidemic scale in central European countries where it caused considerable yield losses in legume crops such as green pea and faba bean. Since PNYDV seems to have a wide geographic distribution in Central Europe, it poses a potential threat to legume production in Europe. This threat might be increased by future changes in climatic conditions that favor survival of aphid vector populations during milder winters in Central Europe and thus facilitate the spread of PNYDV. The economic importance and control of other nanovirids such as ABTV, FBYLV, PYSV, SYSaV and CvLV cannot be commented on as they were identified from single samples and the biological properties of these viruses have not been studied so far. Disease control is based on the results from ecological and epidemiological studies. Since nanoviruses and legume luteoviruses are transmitted by almost the same aphid species, are phloem-limited in their host plants, cause nearly identical symptoms and are not seed-transmitted, they appear to have similar ecologies. Because of this similarity and because luteoviruses have been better studied than nanoviruses, information generated in relation to the control of luteoviruses are useful in the development of control strategies for nanoviruses. Control measures can be subdivided into three major categories: (1) those aimed directly at the virus, (2) those aimed directly at the vector, and (3) integrated approaches, which combine all possible components in a way to complement each other and be applied as one control package. Since virus control is a complex topic, only two examples of virus control, one for a babuvirus (CBDV) in a perennial crop and the other for a nanovirus (FBNYV) in an annual crop are briefly presented. The control of BBTV is dealt with in a separate article (see BBTV (Nanoviridae)). CBDV causes foorkey disease of large cardamom by inducing a remarkable reduction in size of leaves of the infected plants, stimulating proliferation of a large number of stunted shoots arising from the rhizome, transforming the inflorescence into leafy vegetative parts and suppressing fruit formation. CBDV-infected plants remain unproductive and gradually degenerate. The primary spread of disease from one area to another is through infected rhizomes and further spread within the plantation is by aphids. For the management of foorkey disease a regular rogueing of diseased plants by uprooting and immediately destroying them is recommended. As preventive measures, cardamom growers should use CBDV-free planting material preferably seedlings, avoid suckers as planting material from diseased areas and avoid keeping nurseries in the vicinity of infected plantations. In countries of West Asia and North Africa, FBNYV can occur at a high incidence and cause severe yield losses in cool-season legumes, such as chickpea, faba bean, lentil and pea. A significant decrease in disease incidence can be expected if sources of infection are eliminated from within or near crops. In practice, this is not easy and it is rather impossible to eliminate all weed hosts of FBNYV. Prior to advocating such an approach for FBNYV control, more information on the relative importance of wild legumes and weeds as sources of FBNYV infection is needed. Severe epidemics of FBNYV can be prevented by proper timing of sowing dates, avoiding peak aphid population and by spraying the fields with appropriate insecticides once or twice during the period when viruliferous aphids introduce FBNYV to the crop. Another measure to prevent further spread and to decrease disease incidence is to eliminate or reduce primary infection foci as sources of infection from within fields. Since sources of resistance to FBNYV have been identified in wild legume species a classical breeding approach might be used for incorporating resistance into established cultivars of some of the aforementioned cool-season legumes.

Diagnosis Nanovirids can be detected and identified by a range of methods, such as electron microscopy (EM), serology, polymerase chain reaction (PCR), isothermal rolling-circle amplification (RCA) and various sequencing techniques. Because of the small size of the nanovirid particles, which can be easily confused with cell constituents, EM (i.e., use of leaf extracts for adsorption preparates) is not the method of choice for nanovirid detection. The virions can only be visualized following particle purification and by immunosorbent EM using nanovirid-specific antisera for coating of EM grids. Most nanovirid species are serologically distinct from one another. BBTV and the two other babuviruses are serologically unrelated to members of the genus Nanovirus. However, antisera and some monoclonal antibodies (MAbs) to BBTV cross-react strongly with ABTV and CBDV. Therefore, BBTV antibodies have been used for the detection of ABTV and CBDV, the two other babuviruses. Of the 10 MAbs raised to BBTV, only two reacted with ABTV. This is consistent with a CP aa sequence difference of about 20% between ABTV and BBTV. On the contrary, antisera to FBNYV and SCSV cross-reacted weakly with SCSV and FBNYV, respectively, in Western blots and immunoelectron microscopy but not at all in double-antibody sandwich enzyme-linked immunosorbent assays (DAS-ELISA), indicating that the serological relationship between these two nanoviruses is remote. However, MDV antigen reacts strongly not only with an antiserum to FBNYV in DAS-ELISA but also with the majority of MAbs to FBNYV in TAS-ELISA, indicating a close serological relationship between FBNYV and MDV (CP aa sequence identity of about 83%). Some of the FBNYV MAbs also allowed detection and differentiation of PNYDV, PYSV, and BMLRV, indicating that these nanoviruses share certain epitopes with FBNYV. However, species-specific MAbs are indispensable for specific detection of all nanovirids by serological means. Under routine conditions and for the analysis of large numbers of samples, nanovirids are typically tested in TAS-ELISA using a cross-reactive antiserum (or a mixture of antisera) for coating of plates and a mixture of broad-spectrum MAbs for antigen detection.

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Samples that give positive reactions can then be used for aphid transmission experiments, PCR analysis and various sequencing techniques. After the initial screening of samples using TAS-ELISA, PCR or immuno-capture PCR using nanovirus component-specific primers have been used for the molecular characterization of nanovirids. Also, isothermal multiple displacement amplification (MDA) or RCA using Phi29 polymerase has become a widely used technique to enrich nanovirus DNAs in DNA preparations from tissue samples or even ancient particle preparations. For instance, the multiple DNAs associated with coconut foliar decay disease were uncovered this way from virion samples stored for about 30 years. At present, RCA- or PCR-amplified DNA is routinely used for deep sequencing analyses and has yielded considerable insights into the diversity of known nanovirids and has also led to the discovery of PSSaV, a novel and quite different nanovirus.

Concluding Remarks Apart from graft transmission and biolistic bombardment with purified virions, vector transmission had been the only means of experimentally infecting plants with nanovirids for a long time, until infectivity of cloned nanovirus DNA was demonstrated for FBNSV and fully infectious and aphid transmissible virus could be reconstituted this way. Due to this technical achievement functional analyses of individual nanovirus genes have become possible. Meanwhile infection using cloned genome DNAs has been successfully demonstrated also for BMLRV, FBNYV and PNYDV and allowed the application of reverse genetics to studying also nanoviruses, which was not possible before. As a consequence, the experimental independence of vector transmission has substantially advanced nanovirus research during the last years and yielded new and unexpected findings about nanovirus biology. In particular, the recent discovery of a ‘multicellular lifestyle’ of these multipartite ssDNA plant viruses represents a remarkable feat that would have been impossible without reverse genetics.

Further Reading Aronson, M.N., Meyer, A.D., Györgyey, J., et al., 2000. Clink, a nanovirus encoded protein binds both pRB and SKP1. Journal of Virology 74, 2967–2972. Chu, P.W.G., Helms, K., 1988. Novel virus-like particles containing circular single-stranded DNA associated with subterranean clover stunt disease. Virology 167, 38–49. Franz, A.W., van der Wilk, F., Verbeek, M., Dullemans, A.M., van den Heuvel, J.F., 1999. Faba bean necrotic yellows virus (genus Nanovirus) requires a helper factor for its aphid transmission. Virology 262, 210–219. Grigoras, I., Timchenko, T., Katul, L., et al., 2009. Reconstitution of authentic nanovirus from multiple cloned DNAs. Journal of Virology 83, 10778–10787. Grigoras, I., Vetten, H.J., Commandeur, U., et al., 2018. Nanovirus DNA-N encodes a protein mandatory for aphid transmission. Virology 522, 281–291. Gronenborn, B., Randles, J.W., Knierim, D., et al., 2018. Analysis of DNAs associated with coconut foliar decay disease implicates a unique single-stranded DNA virus representing a new taxon. Scientific Reports 8, 5698. Koonin, E.V., Dolja, V.V., Krupovic, M., 2015. Origins and evolution of viruses of eukaryotes: The ultimate modularity. Virology 479–480, 2–25. Sicard, A., Yvon, M., Timchenko, T., et al., 2013. Gene copy number is differentially regulated in a multipartite virus. Nature Communications 4, 2248. Sicard, A., Michalakis, Y., Gutierrez, S., Blanc, S., 2016. The strange lifestyle of multipartite viruses. PLoS Pathogens 12, e1005819. Sicard, A., Pirolles, E., Gallet, R., et al., 2019. A multicellular way of life for a multipartite virus. eLife 8, 43599. Timchenko, T., de Kouchkovsky, F., Katul, L., et al., 1999. A single rep protein initiates replication of multiple genome components of faba bean necrotic yellows virus, a single-stranded DNA virus of plants. Journal of Virology 73, 10173–10182. Timchenko, T., Katul, L., Sano, Y., et al., 2000. The master rep concept in nanovirus replication: identification of missing genome components and potential for natural genetic reassortment. Virology 274, 189–195. Vetten, H.J., Dale, J.L., Grigoras, I., et al., 2012. Family Nanoviridae. In: King, A.M.Q., Adams, M.J., Carstens, E.C., Lefkowitz, E.J. (Eds.), Virus Taxonomy: Ninth Report of the International Committee on Taxonomy of Viruses. London: Elsevier/Academic Press, pp. 395–404. Vega-Rocha, S., Gronenborn, B., Gronenborn, A.M., Campos-Olivas, R., 2007. Solution structure of the endonuclease domain from the master replication initiator protein of the nanovirus faba bean necrotic yellows virus and comparison with the corresponding geminivirus and circovirus structures. Biochemistry 46, 6201–6212. Wanitchakorn, R., Hafner, G.J., Harding, R.M., Dale, J.L., 2000. Functional analysis of proteins encoded by banana bunchy top virus DNA-4 to -6. Journal of General Virology 81, 299–306.

Necro-Like Viruses (Tombusviridae) Luisa Rubino, Institute for Sustainable Plant Protection, National Research Council, Bari, Italy Giovanni P Martelli†, University of Bari Aldo Moro, Bari, Italy r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) BTE Barley yellow dwarf virus-like translational enhancer CITE Cap-independent translational enhancer CP Coat protein CsCl Cesium chloride kb Kilobase kDa Kilo dalton

Glossary Procedovirinae A subfamily in the family Tombusviridae, which RNA-dependent RNA polymerase is translated after the read through of a stop codon (from the Latin “procedo”, go ahead). Read-through Read-through consists of the “translation” of a stop codon, such that the synthesis of a larger protein is

MP Movement protein nt Nucleotide(s) ORF Open reading frame RdRp RNA-dependent RNA polymerase S20,w Sedimentation coefficient satRNA Satellite RNA TED Translational enhancer domain UTR Untranslated region

allowed. It is a common phenomenon in viruses, and it has a regulatory function. Riboviria A higher taxonomic rank including all viruses possessing a RNA genome. Tombusviridae A family of single-stranded positive sense RNA viruses within realm Riboviria.

Taxonomy and Classification As the representative of a monotypic group, Tobacco necrosis virus A (TNV-A) was among the 16 groups of plant viruses described in 1971, and became the type species of the genus Necrovirus when it was established in 1995. Since 2012, the genus Necrovirus does not exist anymore as such, but phylogenetic analyses of the RNA-dependent RNA polymerase (RdRp) and movement proteins 1 and 2 (MP1 and MP2) supported the division of the former genus Necrovirus into two genera, Alphanecrovirus and Betanecrovirus. Due to the expression of their RNA-dependent RNA polymerase after translational readthrough of a stop codon, both genera belong to the subfamily Procedovirinae in the family Tombusviridae (realm Riboviria). The genus Alphanecrovirus comprises four definitive member species, i.e., the type species Tobacco necrosis virus A (TNV-A), Olive latent virus 1 (OLV-1), Olive mild mosaic virus (OMMV), and Potato necrosis virus (PoNV). The genus Betanecrovirus, besides the type species Tobacco necrosis virus D (TNV-D), includes two additional virus species: Beet black scorch virus (BBSV), and Leek white stripe virus (LWSV). Chenopodium necrosis virus (ChNV), formerly a member of the genus Necrovirus and for which sequence data are not available, is now an unassigned species in the subfamily Procedovirinae.

Virion Properties, Structure and Composition Virus particles are very stable, resist temperatures in excess of 901C, and chloroform, ether and non-ionic detergents. TNV-A virions sediment as a single component with a coefficient (S20,w) of 118S and have buoyant density of 1.399 g ml
1 at equilibrium in CsCl. Icosahedral virions are approximately 28 nm in diameter, have angular profile and a capsid made up of 60 copies of a trimer consisting of three chemically identical but independent protein subunits (A, B, and C) stabilized by Ca2 þ ions, arranged in a T ¼ 3 lattice. Subunit size ranges from 24 to 27 kDa (BBSV and LWSV, respectively) to 29–30 kDa (other viral species). The capsid has a smooth appearance as protein subunits have an internal R domain and a shell (S), but lack the protruding domain proper of members of the majority of the other genera in the family Tombusviridae (Fig. 1). Necro-like virions encapsidate one molecule of single-stranded, positive-sense RNA, neither capped at the 50 end, nor polyadenylated at the 30 -terminus, approximately 3.6–3.7 kb in size, constituting c. 19% of the particle weight.

Genome Organization and Expression The monopartite genome contains four open reading frames (ORFs) which, in the order from the 50 to the 30 terminus, code for replication-associated proteins (ORF1 and ORF1-RT), movement proteins (ORF2 and ORF3), and the coat protein (CP) (ORF4). Some species possess a fifth ORF either located in the 30 terminal region (TNV-A) or in the middle of the genome (TNV-DH and †

Deceased.

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Fig. 1 Negatively stained TNV-D virions. Bar, 100 nm.

Fig. 2 Genome organization and expression of tobacco necrosis virus A. G-RNA, genomic RNA; sg1 RNA, sub-genomic RNA 1; sg2 RNA, sub-genomic RNA 2; RT, readthrough; CP, coat protein. Molecular weights of the ORF-encoded proteins (p) are indicated.

BBSV), partially overlapping the C-terminus of ORF1-RT and the N-terminal region of ORF2. The genome is very compact, having noncoding regions of limited size (Fig. 2). To compensate for the lack of a 50 cap or a 30 poly(A), RNA cis-elements are required for cap-independent translation of the virus genome. A cap-independent translational enhancer (CITE) has been identified in the 30 -end untranslated region (UTR) of all necro-like viruses, belonging to the Barley yellow dwarf virus (BYDV) like-element (BTE) class of CITEs. The BTE has a helical cloverleaf-like structure and contains a highly conserved nucleotide stretch able to form a stem-loop structure, and binds the eukaryotic translation initiation factor 4F (eIF4F) complex. Genomic RNA acts as messenger for the translation of a protein of 22–24 kDa from ORF1. By translational readthrough of the UAG termination codon of ORF1, a protein of 82–83 kDa is synthesized (Fig. 2), which contains, in the readthrough portion, the conserved GDD motif of RNA-dependent RNA polymerases (RdRp). This protein, together with the expression product of ORF1, is indispensable for virus replication. RNA elements have been identified in TNV-D genome, which are able to modulate the readthrough efficiency. Genes downstream ORF1-RT are expressed via the synthesis of two sub-genomic RNAs, 1.4–1.6 nt and 1.1–1.3 nt in size (Fig. 2). The larger sub-genomic RNA is responsible for the expression of ORFs 2 and 3, whereas the smaller sub-genomic RNA

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constitutes the translational template for the expression of CP. ORFs 2 and 3 are two small, centrally located ORFs which, depending on the viral species, encode proteins of 7–8 and 6–7 kDa, respectively. Both of these proteins are involved in cell-to-cell transport but are dispensable for viral replication. LWSV ORF2 differs in size from that of other sequenced necro-like viruses, for it codes for a protein of 11 kDa. The additional central ORFs of TNV-DH and BBSV code for a 7 kDa and a 5 kDa protein, respectively, which, like the products of ORFs 2 and 3, are necessary for cell-to-cell movement. A possible transmembrane motif has been predicted in OLV-1, OMMV and PoNV ORF3, which may have a membrane anchoring function. ORF4 is the CP gene encoding a 24–30 kDa protein, the building block of the capsid. Necro-like virus CP was shown to be a multifunctional protein: besides encapsidation, it is required also for symptom modulation and efficient systemic virus spreading in the host and in the environment, since it has been reported to interact with the zoospores of the fungus Olpidium brassicae. Moreover, TNV-D CP was shown to modulate virus replication, since its deletion enhances virus RNA accumulation, whereas BBSV CP is able to interact with viral and host factors during virus replication. Silencing suppression ability was shown for OMMV CP, which acts coordinately with the ORF4 p6 product. No experimental evidence is available for the expression of TNV-A ORF5.

Intraspecific Relationships Necro-like viruses are serologically distinguishable from one another. Some are unrelated (e.g., LWSV with TNV-A and TNV-D), whereas others share antigenic determinants that result in weak relationships. Serological relationships are independent from whether viruses belong to genera Alphanecrovirus or Betanecrovirus. This is the case of OLV-1 (Alphanecrovirus) which is distantly related (serological differentiation index from 6 to 9) with both TNV-A (Alphanecrovirus) and TNV-D (Betanecrovirus), and of OMMV and PoNV (Alphanecrovirus) which are serologically related with TNV-D (Betanecrovirus). The level of molecular similarity varies very much with the viral species. The RdRp sequence identity is high within the members of the genera Alphanecrovirus or Betanecrovirus, respectively, but low between alphanecroviruses and betanecroviruses. Thus, TNV-A, OLV-1, OMMV and PoNV share 86%–92% sequence identity with one another in the RdRp at the amino acid level. Likewise, TNV-D, BBSV and LWSV share 57%–69% sequence identity with one another, but sequence identity ranges from only 32%–36% between members of the two genera. MP1 aa sequences show high identities among each other alphanecrovirus (66%–90%) and betanecrovirus (39%–49%), respectively, but only 6%–23% identity if alphanecroviruses are compared with betanecroviruses. Similarly, alphanecrovirus MP2 amino acid sequences share 95%–100% identity, but they show only 10%–26% identity with betanecrovirus MP2 sequences. Members of the genus Betanecrovirus share 39%–47% sequence identity with one another at the MP2 amino acid level. The CP sequences of both alpha- and betanecroviruses are related and share 26%–86% sequence identity. OMMV (genus Alphanecrovirus), a putative recombinant between TNV-D and OLV-1, shows the highest CP sequence identity (86.2%) with that of TNV-D (genus Betanecrovirus), which may account for the cross reactivity with this virus, and 91.2% identity at the amino acid level with the RdRp of OLV-1.

Satellites Satellites are subviral agents depending on the presence of a helper virus for replication, movement and transmission. Satellite nucleic acids are encapsidated by the helper virus encoded coat protein, whereas satellite viruses are able to encode for their own capsid protein. Three satellite viruses are associated with necro-like viruses: Satellite tobacco necrosis virus (STNV)-1, STNV-2 and STNV-C. They are all belong to the genus Albetovirus (from alphanecrovirus, betanecrovirus, tobacco) (realm Riboviria) and are classified as representative isolates of the species Tobacco albetovirus 1, Tobacco albetovirus 2 and Tobacco albetovirus 3, respectively. STNV-1 and -2 are associated with the replication of TNV-A, whereas STNV-C is associated with TNV-D. The association of the satellite virus with its helper virus is specific, since no heterologous helpers have been reported to support STNV replication. STNV-1, -2 and -C are transmitted in association with TNV-A or -D by the fungus Olpidium brassicae, using the same mechanism as the helper virus. The three STNV have isometric particles c. 17 nm in diameter, sedimenting as a single component with a coefficient of 50S, for which the crystal structure has been determined. The capsid is constructed with 60 identical subunits c. 21–22 kDa in size, arranged in a T ¼ 1 lattice, and contains a single-stranded, positive-sense RNA molecule 1221–1245 nt in size. STNV RNA accounts for c. 20% of the particle weight, and comprises a single ORF encoding the CP. Similarly to the helper virus genome, STNV RNA is uncapped and lacks a 30 poly(A) tail. The 50 and 30 untranslated regions share limited identity (47% and 36% for STNV-C) with the corresponding regions in the cognate virus genome, but are interchangeable between satellite and helper viruses, thus suggesting that they share also the same replication mechanism. Short stem-loop structures are responsible for the specific packaging of the satellite virus RNA in its own CP. The first 30 -translational enhancer domain (TED) ever characterized is present downstream the CP encoding sequence in the 30 -untranslated region of STNV RNA, which is able to function as a 50 cap and to interact with the eIF4F complex. Long-distance interactions between the 30 -TED and a 38 nt stretch in the 50 UTR are important in enhancing and regulating the translation of satellite virus RNA. The satellite virus genome contains signals for the correct packaging and specific assembly of the virus particles. The CP sequences of the three satellite viruses share 49%–62% sequence identity, but are different from any other viral protein, with the

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exception of a limited sequence homology with the CP of Satellite maize white line mosaic virus (SMWLMV). The TNV satellite viruses are serologically distinguishable. STNV-1 interferes to some extent with helper virus infections, ameliorating the TNV-induced symptomatology, probably because of competition for factors essential for shared replication mechanisms, for its presence in TNV inocula reduces slightly the virus concentration and the size but not the number of local lesions. Whereas satellite viruses are not associated with any other member of the genus, BBSV supports the replication of a small, linear, single-stranded, noncoding satellite RNA 615 nt in size, which is encapsidated in the virions in monomeric or, more rarely, dimeric form. Monomers are thought to be produced from multimeric templates. BBSV satRNA enhances the aggressiveness of the helper virus for more lesions are produced on infected hosts when mixed inocula are used. Moreover, the effect of BBSV satRNA on the helper virus pathogenicity is further enhanced at low temperatures probably because of some satRNA interference with the expression of the host plant antiviral silencing genes.

Transmission and Host Range All necro-like viruses are readily transmitted by mechanical inoculation to experimental herbaceous hosts, which usually react with necrotic local lesions, not followed by systemic infection. Under natural conditions, infection is often restricted to the roots. All necro-like viruses are transmitted through the soil, either by the chitrid fungus Olpidium brassicae (TNV-A, TNV-D, BBSV), or without the apparent intervention of a vector (OLV-1). Particles of species transmitted by O. brassicae are acquired by the vector from the soil where they are released from roots of infected plants through sloughing off epidermal cell layers and/or following decay of plant debris. Virions are bound tightly to the plasmalemma and the axoneme (flagellum) of the fungal zoospores and are transported inside the zoospore cytoplasm when the flagellum is retracted prior to encystment that precedes penetration in the host. The same mechanism is exploited by STNV to gain entrance into both vector and host. Since necro-like viruses and their satellites (STNV and BBSV satRNA) multiply in the plant cells but not in the fungal plasmodium, zoospores released from infected roots are virus-free. Transmission can therefore occur only if they come again in contact with and adsorb virus particles. Seed transmission of necro-like viruses has not been reported, except for OLV-1, which was detected in the integuments and internal tissues of 82% olive seeds, and transmitted to 35% of the seedlings. Pollen transmission has not been reported for necro-like viruses so far. The natural host range and geographical distribution of necro-like viruses varies with the species. Thus, TNV-A and TNV-D are ubiquitous and infect a wide range of cultivated and wild plants. OLV-1 was recorded from olive in several Mediterranean countries, citrus in Turkey, tomato in Poland and tulip in Japan; OMMV from olive in Portugal, spinach in Greece and tulip in the Netherlands; PoNV, the most recently characterized alphanecrovirus, was found on potato in UK; BBSV was recorded from beet in China, LWSV from leek in France.

Cytopathology Necro-like viruses replicate very actively in their hosts, which translates into the production of a large number of virus particles in infected cells of all tissue types, including vessels. Virions are either scattered in the cytoplasm, gathered in bleb-like evaginations of the tonoplast into the vacuole, or arranged in crystalline arrays of various sizes. STNV particles can also give rise to crystals that can be found in the same cells along with those of the helper virus. Cells infected by TNV and OLV-1 also contain two types of inclusions, that is, clumps of electron-dense amorphous material resembling accumulations of excess CP and fibrous bundles made up of thin filaments with a helical structure. In OLV-1 infected cells, these bundles were identified as accumulations of the 8 kDa movement protein expressed by ORF2. The same protein and the 6 kDa movement protein coded for by ORF3 were detected by immunogold labeling near plasmodesmata. Cytoplasmic clusters of membranous vesicles with fibrillar material, derived from the endoplasmic reticulum or lining the tonoplast, were observed in cells infected by OLV-1 and LWSV.

Further Reading Bringloe, D.H., Pleij, C.W., Coutts, R.H., 1999. Mutational analysis of cis-elements in the 30 - and 50 -untranslated regions of satellite tobacco necrosis virus strain C RNA. Virology 264, 76–84. Huang, Y.W., Hu, C.C., Hsu, Y.H., Lin, N.A., 2017. Replication of satellites. In: Hadidi, A., Flores, R., Randles, J.W., Palukaitis, P. (Eds.), Viroids and Satellites. London: Elsevier Academic Press, pp. 577–586. Kotta-Loizou, I., Peyret, H., Saunders, K., Coutts, R.H., Lomonossoff, G.P., 2019. Investigating the biological relevance of in vitro-identified putative packaging signals at the 50 terminus of satellite tobacco necrosis virus 1 genomic RNA. Journal of Virology 93, e02106–e02118. doi:10.1128/JVI.02106-18. Miras, M., Miller, W.A., Truniger, V., Aranda, M.A., 2017. Non-canonical translation in plant RNA viruses. Frontiers in Plant Science 8, 494. doi:10.3389/fpls.2017.00494. Monger, W., Jeffries, C., 2018. A new virus, classifiable in the family Tombusviridae, found infecting Solanum tuberosum in the UK. Archives of Virology 163, 1585–1594. Newburn, L.R., White, K.A., 2017. Atypical RNA elements modulate translational readthrough in tobacco necrosis virus D. Journal of Virology 91, e02443. doi:10.1128/ JVI.02443-16. Pu, H., Li, J., Li, D., Han, C., Yu, J., 2013. Identification of an internal RNA element essential for replication and translational enhancement of tobacco necrosis virus AC. PLoS One 8 (2), e57938. doi:10.1371/journal.pone.0057938.

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Pyle, J.D., Scholthof, K.-B.G., 2017. Biology and pathogenesis of satellite viruses. In: Hadidi, A., Flores, R., Randles, J.W., Palukaitis, P. (Eds.), Viroids and Satellites. London: Elsevier Academic Press, pp. 627–636. Rubino, L., 2017. Biology of satellites. In: Hadidi, A., Flores, R., Randles, J.W., Palukaitis, P. (Eds.), Viroids and Satellites. London: Elsevier Academic Press, pp. 567–575. Simon, A.E., Miller, W.A., 2013. 30 Cap-independent translation enhancers of plant viruses. Annual Review of Microbiology 67, 21–42. doi:10.1146/annurev-micro-092412-155609. Truniger, V., Miras, M., Aranda, M.A., 2017. Structural and functional diversity of plant virus 30 -Cap independent translation enhancers (30 -CITEs). Frontiers in Plant Science 8 (2047), doi:10.3389/fpls.2017.02047. Varanda, C.M., Materatski, P., Campos, M.D., et al., 2018. Olive mild mosaic virus coat protein and p6 are suppressors of RNA silencing, and their silencing confers resistance against OMMV. Viruses 10 (416), doi:10.3390/v10080416. Vilar, M., Saurì, A., Monné, M., et al., 2002. The insertion and topology of a plant viral movement protein in the endoplasmic reticulum membrane. Journal of Biochemical Chemistry 277, 23447–23452. Wang, X., Cao, X., Zhang, R., et al., 2018. Hsc70–2 is required for beet black scorch virus infection through interaction with replication and capsid proteins. Scientific Reports 8 (4526). Xu, J., Liu, D., Zhang, Y., et al., 2016. Improved pathogenicity of beet black scorch virus variant by low temperature and co-infection with its satellite RNA. Frontiers in Microbiology 7 (1771), doi:10.3389/fmicb.2016.01771.

Relevant Websites https://talk.ictvonline.org/taxonomy/ Taxonomy.

Nepoviruses (Secoviridae) Hélène Sanfaçon, Agriculture and Agri-Food Canada, Summerland, BC, Canada Crown Copyright r 2021 Published by Elsevier Ltd. All rights reserved.

Glossary Lectin Protein that has the ability to bind carbohydrates (sugars). Odonstyle Anterior section of the stylet of a longidorid nematode. The odonstyle consists of a needle-like mouth spear used to penetrate plant root cells. Odontophore Posterior section of the stylet of a longidorid nematode. The odontophore is located at the

base of the nematode mouth. Stylet protractor muscles are attached to the ondotophore. Triradiate lumen Posterior region of the lumen (or food channel) of the esophagus of a longidorid nematode. The triradiate lumen is located in a muscular bulb at the base of the esophagus. The radial muscles attached to the triradiate lumen are used for food ingestion.

Introduction and Historical Perspective Nepoviruses were among the first 16 groups of viruses recognized by the International Committee on Taxonomy of Viruses (ICTV). The name stands for nematode-transmitted viruses with polyhedral particles. From its origin as the “nepovirus group”, it became the genus Nepovirus within the family Comoviridae, which also included the genera Fabavirus and Comovirus. Taxonomic updates later led to the dissolution of the family Comoviridae and the creation of the family Secoviridae, to which the genus Nepovirus is assigned as part of the sub-family Comovirinae. Although nematode transmission was one of the original defining characteristics of the genus Nepovirus, the primary criteria for inclusion of viruses in the genus are now the structure of the RNA genome and of the particles and phylogenetic relationships. As a result, not all viruses belonging to the genus Nepovirus are nematode transmitted (Table 1), and some nematode-transmitted polyhedral viruses that were originally classified as nepoviruses, have been reclassified (e.g., Strawberry latent ringspot virus, currently an unassigned species within the family Secoviridae). As of 2019, there are 40 recognized nepovirus species (Table 1). Nepoviruses cause a variety of diseases of economic importance, notably on grapevine, fruit trees, small fruits and other horticultural crops.

Taxonomy, Phylogeny, and Evolution The genus Nepovirus belongs to the sub-family Comovirinae (also including the genera Comovirus and Fabavirus), family Secoviridae (encompassing the genera Cheravirus, Sadwavirus, Torradovirus, Sequivirus and Waikavirus, in addition to the sub-family Comovirinae), order Picornavirales (consisting of six families including Secoviridae and Picornaviridae), realm Ribovaria. Nepoviruses share several characteristics in common with other members of the order Picornavirales: the small polyhedral viral particles, the positive-strand RNA genomes that are polyadenylated at their 30 end and linked to a small genome-linked protein (VPg) at their 50 end, the production of mature viral proteins by proteolytic cleavage of large polyproteins, and the conserved signature sequences and order of viral replication protein domains within the polyproteins including the nucleotide triphosphate-binding protein (a putative helicase abbreviated either as NTB or as Hel in the literature), the VPg, a chymotrypsinlike protease (Pro), and the RNA-dependent RNA polymerase (Pol). Nepoviruses have a bipartite genome with each RNA encoding one large polyprotein, a property they share with several other members of the family Secoviridae (referred to as secovirids from hereon). Similar to other secovirids with a bipartite genome, the RNA1 polyprotein includes the domains for the replication proteins and the RNA2 polyprotein contains the domains for the movement protein (MP) and coat protein (CP). Nepoviruses encode a single large coat protein of 53–60 kDa, a characteristic they only share with a few other secovirids (e.g., Strawberry mottle virus). In contrast, other secovirids have either two (Comovirus, Fabavirus, Sadwavirus) or three (Cheravirus, Torradovirus, Sequivirus, Waikavirus) coat proteins. In phylogenetic trees based on the Pro-Pol amino acid sequence, which is defined by the sequence between the conserved catalytic cysteine (or serine, see below) of the Pro domain and the conserved GDD motif of the Pol domain, nepoviruses branch together as a sub-branch within a larger branch corresponding to the subfamily Comovirinae (Fig. 1). Nepoviruses are genetically very diverse, more so than most other genera in the family. Phylogenetic trees based on different protein domains are not always congruent (Fig. 1). This is likely due to reassortment between the two viral RNAs and/or recombination events that have contributed to the evolution of nepoviruses. Nepoviruses have been divided into three subgroups (A, B and C) based on the length and packaging of RNA2, and serological properties (Table 1). These subdivisions may need to be further refined in the future. Indeed, while subgroup B nepoviruses confidently branch together in the phylogenetic trees based on the Pro-Pol or CP amino acid sequences, subgroup C nepoviruses only cluster together in the CP phylogenetic tree and subgroup A nepoviruses do not consistently group together (Fig. 1).

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Table 1

Recognized nepovirus species

Name

Abbreviation

Subgroup A Aeonium ringspot virus Arabis mosaic virus

AeRSV ArMV

Arracacha virus A Artichoke Aegean ringspot virus Cassava American latent virus Grapevine deformation virus Grapevine fanleaf virus Melon mild mottle virus Mulberry mosaic leaf roll associated virus Olive latent ringspot virus Potato black ringspot virus Raspberry ringspot virus

AVA AARSV CsALV GDefV GFLV MMMoV MMLRaV OLRSV PBRSV RpRSV

Tobacco ringspot virus

TRSV

Subgroup B Artichoke Italian latent virus Beet ringspot virus Cocoa necrosis virus

AILV BRSV CNV

Crimson clover latent virus Cycas necrotic stunt virus Grapevine Anatolian ringspot virus Grapevine chrome mosaic virus Mulberry ringspot virus Myrobalan latent ringspot virus Potato virus B Tomato black ring virus

CCLV CNSV GARSV GCMV MRSV MLRSV PVB TBRV

Subgroup C Artichoke yellow ringspot virus

AYRSV

Blackcurrant reversion virus

BRV

Blueberry latent spherical virus Blueberry leaf mottle virus

BLSV BLMoV

Cassava green mottle virus Cherry leaf roll virus Chicory yellow mottle virus Grapevine Bulgarian latent virus Grapevine Tunisian ringspot virus Hibiscus latent ringspot virus Lucerne Australian latent virus Peach rosette mosaic virus Potato virus U Soybean latent spherical virus Tomato ringspot virus

CsGMV CLRV ChYMV GBLV GTRSV HLRSV LALV PRMV PVU SLSV ToRSV

a

Vectora

Xiphinema diversicaudatum

X. index

Longidorus elongatus; L. macrosoma Paralongidorus maximus X. americanum L. apulus; L. fasciatus L.elongatus

RNA1b

RNA2b

NC_038762 NC_006057

NC_038761 NC_006056

NC_017939 NC_003615 NC_038765 NC_038767 NC_022798 NC_005266

NC_017938 NC_003623 NC_038766 NC_038768 NC_038863 NC_022799 NC_005267

NC_005097

NC_005096

LT608395 NC_003693 EU741694 (partial)

LT608396 NC_003694

NC_003791 NC_018383 NC_003622

NC_003792 NC_018384 NC_003621

KX656670 NC_004439

þ KX656671 NC_004440

NC_003890

NC_038862 (partial) NC_003509

NC_003502

NC_003872

L. martini L. attenuatus

Cecidophyopsis ribis (mite)

X. americanum X. americanum; X. bricolensis X. californicum; X. rivesi

NC_038764

NC_038763 U20621 (partial)

NC_015414

NC_015415

NC_015492

NC_015493

NC_034214

NC_034215

NC_032270 NC_003840

NC_032271 NC_003839

Satellite RNAsb Type B Type D

NC_003523

NC_001546

NC_003203

NC_003889

NC_003778 þ

Only cases for which a specific association with a nematode vector has been verified experimentally are listed. GenBank accession numbers are listed. A ( þ ) symbol indicates that a type B satellite is known to be associated with the virus but has not been sequenced.

b

487

NC_003971

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Fig. 1 Phylogenetic analyses of nepoviruses and other members of the family Secoviridae. The amino acid sequence of the CP or Pro-Pol domains from recognized species within the family Secoviridae were aligned using Clustal W. Phylogenetic analyses were conducted using the maximum likelihood method as implemented in MEGA 6. Enterovirus C (EVC, genus Enterovirus, family Picornaviridae) was used as an outgroup. The results of 1000 bootstraps are shown for each node (only values above 50% are shown). A scale for the genetic distance is shown below the trees. Nepovirus acronyms are as listed in Table 1, the letter in parenthesis after each acronym represents the subgroup. Branches corresponding to genera other than nepoviruses were collapsed. The Sadwavirus branch includes Satsuma dwarf virus (the only recognized species of the genus) and four related species that are unassigned within the family Secoviridae (Strawberry mottle virus, Black raspberry necrosis virus, Chocolate lily virus A and Dioscorea mosaic associated virus). SLRSV: Strawberry latent ringspot virus, an unassigned species in the family Secoviridae. StPV: Stocky prune virus.

Fig. 2 Electron micrograph depicting purified nepovirus particles and cytopathological structures typical of nepovirus-infected cells. (A) Purified ToRSV particles in negative staining. Note the empty particle (T-particle) which is penetrated by the negative stain (arrow). (B) Proliferation of membrane vesicles observed in the vicinity of the nucleus (Nc) in ToRSV-infected cells. (C) Tubular structures containing virus-like particles accumulating near the cell wall (CW) in PRMV-infected cells. (D) Tubular structure traversing the cell wall in ArMV-infected cells. Bars represent 25 nm in panel A and 200 nm in panels B–D.

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Virion Structure Nepoviruses have isometric particles of 26–30 nm in diameter with sharp hexagonal outlines (Fig. 2(A)). Equilibrium centrifugation in CsCl of purified virus particles typically reveals the presence of three types of particles. T-particles sediment at 50S and do not contain an RNA molecule. In electron microscopy, these empty particles are penetrated by a negative stain. B-particles sediment at 115–134S and contain a single molecule of RNA1. In the case of sub-group A nepoviruses, B-particles can also contain two molecules of RNA2. M-particles sediment at 86–128S and contain a single molecule of RNA2. M- and B-particles of sub-group C nepoviruses are often difficult to separate due to the larger size of RNA2. Nepovirus particles contain 60 molecules of the single CP. The crystal structure of virus particles was solved for TRSV, GFLV and ArMV (see Table 1 for acronyms). The structure of BRV particles was also inferred by cryo-electron microscopy. The pseudo T ¼3 structure is similar to that of comoviruses and picornaviruses. The CP contains three functional domains. Each functional domain corresponds to one of the three smaller CPs found in picornaviruses and consists of eight b-strands that form a b-barrel. Comoviruses also share a similar structure with one large CP containing two b-barrels and one small CP with a single b-barrel. The N-terminal region of the CP is buried inside the particle while the C-terminal region of the CP is exposed to the outside of the particle. In many nepoviruses, a small fragment of the C-terminus of the CP is cleaved off from the virions at late stages of infection, although the biological function of this cleavage is not known. Several loops connecting the b-strands are exposed to the surface of the particle and interact with the viral movement protein to support cell-to-cell movement or with vector receptors to enable transmission. The mechanism of encapsidation is not well-characterized, but the presence of empty particles with structures similar to full particles suggests that presence of the viral RNAs is not strictly required to initiate capsid assembly.

Genome Organization and Function of Viral Proteins The genomic organization of a representative virus from each subgroup is shown in Fig. 3. The C-terminal protein domains of the polyproteins are conserved amongst nepoviruses, while the N-terminal regions of the polyproteins are more variable. The C-terminal portion of the RNA1 polyprotein contains the NTB, VPg, Pro and Pol domains, which are implicated in viral RNA replication and regulation of polyprotein processing. Identification of cleavage sites upstream of NTB defined two protein domains in the N-terminal region of the ToRSV and ArMV RNA1 polyproteins, termed X1 and X2. Putative cleavage sites at corresponding positions were found in the RNA1 polyproteins of other nepoviruses. Thus, the presence of two protein domains upstream of NTB is likely a common feature of nepovirus RNA1 polyproteins, which is not shared by comoviruses or fabaviruses. The X2 protein domain is highly hydrophobic, shares conserved sequence motifs with the 32 kDa protein of comoviruses and has been suggested to play a role in the assembly of replication complexes. The X1 domain shows a high degree of sequence diversity amongst nepoviruses and its function is unknown. Because RNA1 encodes all the essential replication proteins, it is able to replicate in single plant cells without the assistance of RNA2. The RNA2 polyprotein includes the domains for the CP and MP at its C-terminus and one or two additional protein domains at its N-terminus: the single 2a domain for subgroup A and B nepoviruses and the two X3 and X4 domains for ToRSV, a subgroup C nepovirus. The GFLV 2a protein plays a role in the replication of RNA2. The function of X3 and X4 is not known. Both RNA1 and RNA2 are required for cell-to-cell and systemic movement of the virus in the plant. Infectious cDNA clones have been produced for subgroup A (GFLV, ArMV, RpRSV, TRSV) and subgroup B (TBRV) nepoviruses, allowing reverse genetic experiments for these viruses.

Fig. 3 Genomic organization of representative nepoviruses of subgroup A (ArMV), B (BRSV) and C (ToRSV). Each RNA is represented with the covalently attached VPg (pink circle) and the polyA tail (An). The coding regions are represented by the boxes. Cleavage sites confirmed by in vitro processing experiments or by the detection of viral proteins in infected plants are indicated by the continuous vertical lines. Putative cleavage sites that have not been confirmed experimentally are shown with dashed lines. Dipeptide sequence of confirmed or putative (in parenthesis) cleavage sites are indicated above each line. Thick bars below each RNA indicate regions with high degree of sequence identity between RNA1 and 2 (black for regions with near 100% sequence identity and gray for regions with 75%–83% sequence identity).

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Each RNA includes untranslated regions (UTR) at its 50 and 30 ends. The 50 UTR is 70–300 nucleotides (nts) long while the 30 UTR varies in length from 200 to 400 nts for subgroup A and B nepoviruses to 1300–1600 nts for subgroup C nepoviruses. RNA1 and 2 often share regions of complete or partial sequence identity in the UTRs (Fig. 3). For example, the 5’ UTRs of ToRSV RNA1 and 2 share a region of 100% sequence identity which extends into the coding region. On the other hand, the 50 UTRs of the BRV RNAs share only 78% sequence identity and do not extend beyond the UTRs. The 30 UTRs of RNA1 and 2 are often nearly identical, especially those of subgroup C nepoviruses. Conservation of sequence identity in the UTRs has been attributed to recombination between the two RNAs occurring during replication of the viral RNAs and/or to selection pressure.

Production of Viral Proteins: Translation and Polyprotein Processing The viral RNAs are polyadenylated but are not capped, requiring a cap-independent translation mechanism. Studies on BRV RNAs suggested that long-distance interaction between the 50 and 30 UTRs (possibly directed by sequences in the loops of putative stemloop structures) is required for efficient translation of the viral RNAs. In addition, the polyA tail enhances translation efficiency. Further work will be necessary to elucidate the exact mechanism of BRV RNAs translation, in particular the requirement for plant translation factors, and to determine whether other nepoviruses utilize similar or distinct mechanisms to translate their RNAs. Nepoviruses encode a single protease which is responsible for processing the two polyproteins. Nepovirus Pros are related to the 3C-Pro of picornaviruses. The catalytic triad normally consists of a histidine, glutamic acid and cysteine, although the Pros of a few subgroup C nepoviruses have a serine instead of the cysteine as the third component of the triad. Proteases also contain a substrate-binding pocket that determines their cleavage site specificity. Similar to picornaviruses, a conserved histidine is found in the substrate-binding pocket of all subgroup C nepovirus Pros characterized so far and of one subgroup A nepovirus Pro (MMLRaV). The cleavage sites recognized by these Pros normally include a glutamine, asparagine or aspartate at the -1 position. This is similar to the conserved glutamine or glutamate found at the -1 position of picornavirus cleavage sites. However, there are reported exceptions, including a characterized cleavage site recognized by the BLSV Pro with a cysteine at the -1 position and several putative cleavage sites of subgroup C nepoviruses with a histidine at the -1 position. The Pros of subgroup A and B nepoviruses contain a leucine instead of a histidine in their substrate-binding pocket and recognize a variety of cleavage sites that have lysine, cysteine, arginine or glycine at the -1 position. Thus, nepovirus proteases have very diverse specificities, often making the prediction of cleavage sites difficult. The RNA1 polyprotein is cleaved predominantly intra-molecularly (cis-cleavage). The RNA2 polyprotein is cleaved in trans by the Pro. The processing cascade results in the release of mature proteins and stable processing intermediates containing two or more protein domains. These intermediate polyproteins accumulate in infected plant cells and may have different activities from the mature proteins. In ToRSV-infected cells, several polyprotein intermediates containing the NTB domain are detected in addition to the mature NTB protein. In BRSV-infected cells, an intermediate containing the Pro and Pol domains accumulates rather than the mature Pro and Pol proteins. This suggests preferential recognition of some cleavage sites by the viral Pro. Slow release of mature proteins by processing of stable intermediates at suboptimal cleavage sites provides a regulatory mechanism to control the accumulation of specific proteins during the replication cycle. In fact, the activity of the Pro itself is regulated by its release from larger polyprotein precursors. Indeed, the mature Pro of GFLV and ToRSV processes the cleavage sites of the RNA2 polyprotein more efficiently than the VPg-Pro precursor.

Viral RNA Replication Infection of plant cells by nepoviruses results in membrane proliferation and the formation of membrane vesicles (Fig. 2(B)). The use of cerulenin, an inhibitor of de novo phospholipid synthesis, has confirmed that membrane proliferation is required for the replication of GFLV RNAs. Brefeldin A also inhibits GFLV replication suggesting a requirement for intact vesicle trafficking between the endoplasmic reticulum (ER) and the Golgi apparatus. The replication complexes of two nepoviruses (GFLV and ToRSV) were shown to co-localize with ER-derived membranes in infected cells. Double-strand RNA replication intermediates, viral replication proteins and replication activity are associated with the membrane-bound complexes. GFLV VPg antibodies were used to isolate membrane vesicles that had a rosette-like structure similar to that associated with picornavirus replication complexes. As shown for ToRSV, the mature NTB protein and the intermediate NTB-VPg and X2-NTB-VPg polyproteins co-fractionate with the replication complex, suggesting that they act as membrane anchors for the replication complex. The NTB protein contains two membrane-binding domains: a C-terminal transmembrane domain and a putative N-terminal amphipathic helix. The ToRSV X2 protein also contains several transmembrane helices that traverse the ER membrane. Other replication proteins (Pro, Pol or intermediate polyproteins containing these domains) are soluble when expressed individually but are found in association with the membrane-bound replication complex in infected cells. Thus, they are probably brought to the replication complex either as part of a larger polyprotein that includes the NTB domain or through protein-protein interaction with the viral membrane anchors. By analogy with picornaviruses, the VPg protein may act as a primer for viral replication, although this has not been demonstrated experimentally for nepoviruses. The GFLV RNA2-encoded 2a protein is also

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associated with the replication complex. The 2a protein is required for the replication of RNA2 but not RNA1 and probably interacts with RNA1-encoded replication proteins.

Cell-to-Cell and Systemic Movement in the Plant Nepovirus-infected cells are characterized by the presence of tubular structures containing virus-like particles in or near the cell wall (Fig. 2(C) and (D)). These tubules are similar to the ones found in comovirus-infected cells and have been suggested to direct the cell-to-cell movement of intact virus particles. The viral movement protein is a structural component of the tubular structure. Expression of the GFLV MP alone is sufficient to induce the formation of empty tubular structures in intact plant cells or in protoplasts. In the latter case, tubular extensions are found projecting from the surface of the protoplasts, a phenomenon also induced by comovirus MP. The GFLV MP is an integral membrane protein. The secretory pathway and the cytoskeleton are involved in the intracellular targeting of the GFLV MP from its site of synthesis (probably in association with the ER-bound replication complex) to specific foci in the cell wall where it assembles into tubules. GFLV cell-to-cell movement also requires myosin-motor plant proteins, which may assist in transporting other plant proteins to the plasmodesmata to initiate the formation of the MP tubules. By analogy with comoviruses, it is likely that an interaction between MP and CP is necessary to enable nepovirus cell-to-cell movement. A surface-exposed region of the CP and the C-terminal region of the MP are probably involved in this interaction. Long-distance movement of TRSV occurs through the phloem resulting in the invasion of most tissues of the plant including meristematic tissues. The virus probably reaches the phloem through cell-to-cell movement from inoculated cells to phloem sieve tubes.

Host Range, Symptom Determinants and Interaction With Plant Defense Responses Most nepoviruses have a wide host range that includes woody and herbaceous hosts. In nature, many hosts remain symptomless while others display symptoms such as necrotic or chlorotic rings which can appear as concentric rings (ringspots) or lines. Other symptoms can include leaf flecking and mottling, vein necrosis, plant stunting and in some cases death. Common experimental hosts used for virus propagation are Chenopodium quinoa (in which most nepoviruses induce obvious symptoms), C. amaranticolor, C. murale, Cucumis sativus, Nicotiana clevelandii, N. benthamiana, Petunia hybrida and Phaseolus vulgaris. The intensity of symptoms produced by nepovirus infection depends on the specific virus-host combination and on environmental conditions. In many herbaceous hosts and in particular in Nicotiana species, symptoms develop on the inoculated leaves and on the first upper systemic leaves. Later in infection, new leaves remain free of symptoms although the virus is present. This phenomenon is termed symptom recovery. Recovered leaves of nepovirus-infected plants often show reduced concentration of viral RNAs and this has been attributed at least in part to the induction of the plant antiviral RNA silencing. Recovered leaves are also resistant to reinfection in a sequence-dependent manner. In the Nicotiana benthamiana-ToRSV interaction, the establishment of symptom recovery depends on the specific isolate and also on the growth temperature (Fig. 4). The onset of symptom recovery from a severe ToRSV isolate is not accompanied with a significant reduction of viral RNA levels. However, translation of the viral RNAs is suppressed limiting the accumulation of viral proteins and encapsidation. It has been hypothesized that the low nepovirus titers typically found in late stages of infection contribute to their ability to escape plant surveillance systems and invade meristematic tissues. When they occur, necrotic symptoms share characteristics of a hypersensitive response (HR) normally induced by activation of plant dominant resistance gene. Although the virus can be restricted to local necrotic lesions developing on inoculated leaves in some hosts, the HR-like response often fails to contain the virus and systemic infection is observed, which may be followed by recovery or not. The plant genetic determinants of this HR-like response have been characterized in the Arabidopsis thaliana-TRSV interaction. Most A. thaliana ecotypes are tolerant to TRSV and do not display visible symptoms. However, the lethal systemic necrosis induced by TRSV in some ecotypes was linked to a dominant mutation in the TTR1 resistance gene. Viral symptom determinants have also been identified in a few interactions. In the Nicotiana occidentalis-GFLV interaction, construction of chimeric clones between two isolates identified the 2a protein as the elicitor for the necrotic HR-like response. Using a similar approach, a single amino acid in the GFLV Pol was found to determine vein clearing symptoms in N. benthamiana.

Transmission Many nepoviruses are transmitted by soil dagger nematodes belonging to the genera Xiphinema, Longidorus or Paralongidorus in the order Dorylaimida, family Londigoridae. The nematodes feed ectoparasitically on the roots using long mouth stylets. There is no evidence that nepoviruses replicate in the nematodes. The acquired viruses remain transmissible for varying periods of time (9 weeks for Longidorus species and up to 4 years for Xiphinema species), suggesting different modes of retention and release of the virus. Because of the restricted movement of the nematodes through the soil, the spread of nematode-transmitted nepoviruses through an infected field is slow and often occurs in patches. The interaction between nepoviruses and nematodes is

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Fig. 4 Symptom recovery in ToRSV-infected Nicotiana benthamiana plants. Plants inoculated with a severe (Rasp1) or mild (GYV) isolate of ToRSV were grown at either 271C or 211C. GYV-infected plants recover from infection at both temperature, while Rasp1-infected plants only recover from infection at 271C. Detached inoculated leaves (I), upper symptomatic leaves (S) or recovered leaves (R) are shown at the indicated days post-infection (dpi). The lower panel compares plants that were mock-inoculated to plants infected with the two isolates at 20 dpi after growth at 271C (left) or 211C (right). Modified from Ghoshal and Sanfacon, 2014. Temperature-dependent symptom recovery in Nicotiana benthamiana plants infected with tomato ringspot virus is associated with reduced translation of viral RNA2 and requires ARGONAUTE 1. Virology 456–457, 188–197. Paudel et al., 2018. Expression and antiviral function of ARGONAUTE 2 in Nicotiana benthamiana plants infected with two isolates of tomato ringspot virus with varying degrees of virulence. Virology 524, 127–139. Both under Crown Copyright.

usually specific with only one or two species of nematodes transmitting a given nepovirus (Table 1). In the case of GFLV, the viral determinant for the specificity of nematode transmission has been mapped to the CP, in particular a positively charged pocket exposed to the surface of virus particles. It is not known whether specific nematode receptors recognize the viral CP. Earlier studies suggested that carbohydrates may be involved in the retention of ArMV particles in its vector. However, further experiments will be required to determine if the viral coat protein has lectin properties. It has been suggested that pH changes may be involved in the release of viral particles from their site of retention within the nematode. Nepoviruses transmitted by Longidorus species are usually associated with the odontostyle, while viruses transmitted by Xiphinema species are found associated with the cuticle lining the lumen of the odontophore and the esophagus. Interestingly, while TRSV and ToRSV are both transmitted by X. americanum, immunofluorescence labeling of these viruses in the nematode vector revealed different sites of retention. TRSV is retained predominantly in the lining of the lumen of the stylet extension and the anterior esophagus, while ToRSV is localized only in the triridiate lumen. Although the predominant vector for transmission of TRSV is a nematode, possible aerial vectors have been suggested including Thrips tabaci and Epitrix hirtipennis (flea beetle). The importance of these vectors in natural epidemics of TRSV-induced diseases needs to be confirmed. A recent study also reported detection of TRSV double-strand RNA replication intermediates in bees, suggesting viral replication. However, TRSV is not transmitted by bees and the biological relevance of this observation, including its possible association with bee decline, is not clear. Other nematode-transmitted nepoviruses do not have known aerial vectors. BRV is transmitted by the eriophyid gall mite of black currant (Cecidophyopsis ribis) and possibly other Cecidophyopsis species but not by nematodes. Plant-to-plant transmission can occur rapidly (in only 4 h). Virus particles have not been found inside mites suggesting that the transmission may be non-persistent or semi-persistent. Two surface-exposed amino acid triplets are conserved between the CP of BRV and other mite-transmitted viruses from unrelated genera, suggesting that they may play a role in the interaction between the virus and its vector. However, this remains to be determined experimentally. Seed transmission has been reported for most but not all nepoviruses. For example, BRV is apparently not seed transmitted. Infection can occur through the ovule or the pollen, although nepovirus-infected pollen may not compete effectively with healthy pollen. An exception to this is BLMoV and CLRV, which are efficiently transmitted by pollen. Honeybees likely contribute to the pollen transmission of BLMoV by increasing the movement of infected pollen. Many nepoviruses are readily transmitted through grafting and mechanical inoculation. The propagation of infected seed and plant stocks plays an important role in the long-

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distance movement of nepoviruses. Because many nepoviruses have a wide host range including common weeds, dormant weed seeds may provide a reservoir for the virus in the field.

Population Structures: Quasi-Species, Satellite RNAs and Defective-Interfering RNAs Examination of nepovirus isolates recovered from different hosts or geographic locations has revealed a degree of sequence diversity of 2%–20% at the nucleotide level. Isolates can also differ in the length of their RNAs with the RNA2 polyprotein N-terminal domains being the most variable, as shown for the GFLV and ArMV 2a protein and the ToRSV X4 protein. Evidence for recombination and reassortment between viral RNAs is well-documented resulting in the emergence of new viral strains or species, although interspecies recombination has so far only been reported among closely related nepovirus species. Mixed infections between viral isolates and/or closely related species frequently occur in the field. In agreement with the quasi-species model, detailed analyses of GFLV population structure highlighted the presence of a few predominant variants along with other minor variants. An ArMV field isolate was shown to contain two species of RNA2, each encoding a distinct polyprotein. It is possible that co-existing variants act synergistically to facilitate infection or determine symptoms but this has not been tested experimentally. Analysis of a number of CLRV isolates revealed that the degree of diversity is defined primarily by the host rather than the geographic location. This is an unusual situation among plant viruses and may reflect the fact that this virus is pollen-transmitted. Two classes of satellite RNAs (satRNAs) have been found in association with some nepoviruses (Table 1). SatRNAs depend on the helper virus for their replication and are encapsidated in virus particles. Type B satRNAs are 1100–1500 nts in length. They are linked to a VPg molecule at their 50 end, polyadenylated at their 30 end and encode a non-structural protein which is essential for their replication. The exact function of the encoded protein is not known but it has been suggested that it interacts with the viral replication complex. The 50 -ends of type B satRNAs often have short sequence motifs that are identical or nearly identical to the 50 ends of the viral RNAs and are likely recognized by the viral replication complex. One or several copies of type B satRNAs are packaged in the viral particles, either alone or together with one molecule of RNA2. As a general rule, type B satRNAs are replicated specifically by the virus with which they are associated although there are exceptions (e.g., the replication of GFLV satRNA is supported by ArMV). Type B satRNAs are usually found in low concentration and do not affect significantly the replication of the helper virus or the symptomatology. Type D satRNAs are less than 500 nts long. They are not linked to a VPg molecule or polyadenylated and do not encode a protein. Type D satRNAs are encapsidated as monomeric or multimeric linear molecules. A circular form of the molecule is present in infected cells and serves as the polymerase template for replication through a rolling circle mechanism. The multimeric linear forms produced during replication are cleaved in an autocatalytic reaction which allows the release of linear monomers. These monomers are circularized to form new templates of positive or negative polarity. Type D satRNAs have been shown to either attenuate (TRSV) or intensify (ArMV) symptoms associated with the helper virus. TBRV is the only nepovirus reported to be associated with defective-interfering RNAs (diRNAs). These 400–500 nts RNAs are derived exclusively from RNA1 and are probably produced during viral RNA replication. Presence of the diRNAs was associated with attenuated symptoms and reduced virus accumulation in some hosts. However, further work will be necessary to confirm the biological significance of the diRNAs and to determine whether they are also observed in other nepovirus-plant pathosystems.

Diseases, Economic Considerations and Control Nepoviruses cause a wide range of diseases on a variety of crops including: grapevine, small fruits such as strawberry, raspberry, blueberry, black currant and red currant, fruit trees such as peach, apricot, almond, cherry, plum, walnut and apple, and other horticultural crops including but not limited to hop, soybean, potato, beet and tobacco. Many nepoviruses also induce diseases in ornamental species. Most nepoviruses are restricted geographically by the natural distribution of their nematode vector. An exception to this is GFLV which has been disseminated worldwide along with its vector. GFLV is the most significant nepovirus at the economic level and can reduce yield in grapevine by as much as 80%. Other nepoviruses can cause significant diseases where they occur. Nepoviruses are mainly controlled through the removal of infected plants and replanting with resistant cultivars (when available) or with virus-free certified plant material. Weed control can reduce potential reservoirs for further nepovirus infection. In the case of nematode-transmitted nepoviruses, soil can be treated with broad-spectrum fumigant nematicides. However, because nematode populations can occur at considerable depths in the soil (one meter or more), fumigation rarely completely eliminates plant parasitic nematodes from a site and population densities typically return to pre-fumigation levels within a few years in the presence of a suitable host. Further, many fumigant nematicides are now banned due to their toxicity. Few natural sources of resistant cultivars are available for nepoviruses. Transgenic expression of small regions of the viral genome was shown to induce the plant antiviral RNA silencing and provide virus resistance in herbaceous hosts (several nepoviruses) and in grapevine (GFLV). Broad spectrum resistance against a range of GFLV isolates was also obtained in N. benthamiana and in grapevine upon transgenic expression of camelid-derived heavy-chain only GFLV-specific antibodies. The characterization of plant factors essential for nepovirus infection combined with emerging gene editing approaches will likely provide new opportunities for generating non-transgenic nepovirus-resistant plants in the future.

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Concluding Remarks Although the infection cycle of nepoviruses is relatively well-understood, at least in herbaceous hosts, there are many outstanding questions. Many plant factors facilitating the translation, replication and movement of nepoviruses remain to be identified and/or characterized. Such plant factors may prove useful new targets for antiviral strategies, based on gene editing techniques. It will also be critical to evaluate the impact of nepoviruses and their complex populations (including the presence of multiple variants) in the field and to gain further insights into the emergence of virulent strains or new species. Additional research is needed to better understand the complex interaction between nepoviruses and various plant defense responses leading to the range of symptoms, including tolerance, symptom recovery and lethal systemic necrosis. Finally, evaluating the contribution of environmental factors on virus titers, disease severity and seed or vector transmission will also likely be a focus of future studies.

Further Reading Amari, K., Lerich, A., Schmitt-Keichinger, C., Dolja, V.V., Ritzenthaler, C., 2011. Tubule-guided cell-to-cell movement of a plant virus requires class XI myosin motors. PLoS Pathogens 7, e1002327. Fuchs, M., Schmitt-Keichinger, C., Sanfacon, H., 2017. A renaissance in nepovirus research provides new insights into their molecular interface with hosts and vectors. Advance in Virus Research 97, 61–105. Ghoshal, B., Sanfacon, H., 2014. Temperature-dependent symptom recovery in Nicotiana benthamiana plants infected with tomato ringspot virus is associated with reduced translation of viral RNA2 and requires ARGONAUTE 1. Virology 456–457, 188–197. Hemmer, C., Djennane, S., Ackerer, L., et al., 2018. Nanobody-mediated resistance to Grapevine fanleaf virus in plants. Plant Biotechnology Journal 16, 660–671. Martin, I.R., Vigne, E., Berthold, F., et al., 2018. The 50 distal amino acids of the 2A(HP) homing protein of Grapevine fanleaf virus elicit a hypersensitive reaction on Nicotiana occidentalis. Molecular Plant Pathology 19, 731–743. Nam, M., Koh, S., Kim, S.U., et al., 2011. Arabidopsis TTR1 causes LRR-dependent lethal systemic necrosis, rather than systemic acquired resistance, to Tobacco ringspot virus. Molecules and Cells 32, 421–429. Osterbaan, L.J., Choi, J., Kenney, J., et al., 2019. The identity of a single residue of the RNA-dependent RNA polymerase of grapevine fanleaf virus modulates vein clearing symptoms in Nicotiana benthamiana. Molecular Plant Microbe Interactions 32 (7), 790–801. Paudel, D.B., Ghoshal, B., Jossey, S., et al., 2018. Expression and antiviral function of ARGONAUTE 2 in Nicotiana benthamiana plants infected with two isolates of tomato ringspot virus with varying degrees of virulence. Virology 524, 127–139. Sanfacon, H., 2013. Investigating the role of viral integral membrane proteins in promoting the assembly of nepovirus and comovirus replication factories. Frontiers in Plant Science 3, 313. Santovito, E., Mascia, T., Siddiqui, S.A., et al., 2014. Infection cycle of Artichoke italian latent virus in tobacco plants: Meristem invasion and recovery from disease symptoms. PLoS One 9, e99446. Schellenberger, P., Sauter, C., Lorber, B., et al., 2011. Structural insights into viral determinants of nematode mediated Grapevine fanleaf virus transmission. PLoS Pathogens 7, e1002034. Thompson, J.R., Dasgupta, I., Fuchs, M., et al., 2017. ICTV virus taxonomy profile: Secoviridae. Journal of General Virology 98, 529–531.

Ophioviruses (Aspiviridae) Anna M Vaira, Institute for Sustainable Plant Protection, National Research Council of Italy, Torino, Italy John Hammond, Floral and Nursery Plants Research, Agricultural Research Service, US Department of Agriculture, Beltsville, MD, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of A.M. Vaira, R.G. Milne, Ophiovirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00645-2.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein ELISA Enzyme-linked immunosorbent assay EVEs Endogenous viral elements GFP Green fluorescent protein kb Kilobase kDa Kilodalton LAMP Loop mediated isothermal amplification MP Movement protein NES Nuclear export signal NGS Next generation sequencing NLS Nuclear localization signal

Glossary Bunyavirales Order of viruses comprising species that have a segmented linear negative-sense RNA genome and a lipid envelope. The order includes 12 families: Arenaviridae, Cruliviridae, Fimoviridae, Hantaviridae, Leishbuviridae, Mypoviridae, Nairoviridae, Peribunyaviridae, Phasmaviridae, Phenuiviridae, Tospoviridae, and Wupedeviridae.

NP Nucleoprotein nt Nucleotide(s) ORF Open reading frame PD Plasmodesmata RdRp RNA-dependent RNA polymerase RT-PCR Reverse transcriptase-polymerase chain reaction SEL Size exclusion limit TEM Transmission electron microscopy UTR Un-translated region vcRNA Viral complementary RNA VIGS Virus-induced gene silencing VSR Viral suppressor of RNA silencing

Mononegavirales Order of viruses comprising species that have a linear, negative-sense RNA genome and a lipid envelope. The order includes 11 families: Artoviridae, Bornaviridae, Filoviridae, Lispiviridae, Mymonaviridae, Nyamiviridae, Paramyxoviridae, Pneumoviridae, Rhabdoviridae, Sunviridae, and Xinmoviridae. Serpentovirales Order of viruses comprising 1 family, Aspiviridae, and 1 genus, Ophiovirus.

Introduction The genus Ophiovirus is the only genus listed in the family Aspiviridae (formerly Ophioviridae), order Serpentovirales. The genus comprises plant-infecting virus species. In some cases ophioviruses are known to be the cause of well-known and “classical” major plant diseases, but in other cases the presence of the virus has not been linked to any specific symptom, due to almost invariable occurrence in mixed infections with other viruses. Ophioviruses occur in monocots and dicots, in vegetables, ornamentals, trees and shrubs, in the New and Old World, suggesting a well-adapted group of viruses. Where identified, the vectors of ophioviruses have proved to belong to Olpidium spp., soil-inhabiting chytrid fungi. Ophioviruses have been slow to emerge mainly because the virions are not easy to see in the electron microscope; indeed, the first ophiovirus particle was observed only in 1988 and its morphology understood in 1994. The genus name derives from the Greek word ophis, meaning snake, in reference to the serpentine appearance of the virus particles and the family name derives from the Latin word aspis, with the same meaning.

Taxonomy, Phylogeny and Evolution There are currently seven species in the genus Ophiovirus (Table 1).The accepted species in the genus are: Citrus psorosis ophiovirus (the type species)(Citrus psorosis virus, CPsV), Mirafiori lettuce big-vein ophiovirus (Mirafiori lettuce big-vein virus, MLBVV), Lettuce ring necrosis ophiovirus (Lettuce ring necrosis virus, LRNV), Ranunculus white mottle ophiovirus (Ranunculus white mottle virus, RWMV), Tulip mild mottle mosaic ophiovirus (Tulip mild mottle mosaic virus, TMMMV), Freesia sneak ophiovirus (Freesia sneak virus, FreSV) and Blueberry mosaic associated ophiovirus (Blueberry mosaic associated virus, BlMaV). To date, the genomes of four species, MLBVV, CPsV, LRNV and BlMaV, are fully sequenced. For species demarcation within the genus, different criteria have been considered: the different coat protein (CP) sizes; absence of, or distant serological relationship between the CPs; differences in natural host range; and different number, organization, and/or size of genome segments. From available data, there is 95%–100%

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Table 1

Virus members in the Genus Ophiovirus with relevant NCBI database accession numbers

Species (alternative name)

Acronym

Accession numbersa

Blueberry mosaic associated ophiovirus Citrus psorosis ophiovirus (Citrus ring-spot virus) Freesia sneak ophiovirus (Freesia ophiovirus) Lettuce ring necrosis ophiovirus Mirafiori lettuce big-vein ophiovirus (Mirafiori lettuce virus) Ranunculus white mottle ophiovirus Tulip mild mottle mosaic ophiovirus

BlMaV CPsV

NC036635, NC024476, NC036634 NC006314, NC006315, NC006316

FreSV

MN365016, DQ885455

LRNV MLBVV

NC006051, NC006052, NC006053, NC006054 AF525933, AF525934, AF525935, AF525936

RWMV TMMMV

NC043389, AY542957 NC043390

a

Complete genome sequences are reported when available.

Fig. 1 Unrooted phylogenetic tree of selected ophiovirus isolates from different geographical origin, based on their NP amino acid sequences. The evolutionary history was inferred using the Neighbor-Joining method. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) are shown next to the branches. The evolutionary distances were computed using the Poisson correction. Evolutionary analyzes were conducted in MEGA X. The TMMMV, RWMV and BlMaV AJR35847 nucleoprotein sequences analyzed are partial. Rectangles grouping isolates represent established viral species.

identity among complete CP aa sequences of isolates belonging to the same species detected in different world areas, in particular for MLBVV, CPsV, BlMaV and FreSV. The percentage of identity falls to 28%–59% for interspecies alignments, with the exception of the identity between MLBVV and TMMMV CP sequences, which is slightly above 80%, confirming the closeness of these two viruses. They are currently considered as two different species at least because of different host ranges; more information on TMMMV sequences is needed prior to any revision of their taxonomic position. The percentage of identity/similarity between CP aa sequences may also be proposed as a further tool for species demarcation as several full sequences are now available. A phylogenetic tree based on alignment of representative CP aa sequences is shown in Fig. 1. CPsV and BlMaV, presently the only ophioviruses infecting perennial hosts with asexual means of propagation, are more correlated. These differences, if confirmed and reinforced, could lead to considering placement of these species in a different genus within the family Aspiviridae. RT-PCR amplification, using a pair of degenerate primers (see below), is currently a good tool for detecting and identifying official and potential new species within the genus, providing also indications of the taxonomic position of newly diagnosed isolates. The analysis of RdRp aa sequences shows the Aspiviridae family as a monophyletic group among negative-stranded RNA virus families of orders Bunyavirales and Mononegavirales, newly assigned to the order Serpentovirales. Recently, due to the wide use

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Fig. 2 Negative contrast electron micrograph (uranyl acetate) of partially purified virion preparation from field lettuce showing big-vein symptoms. Note large and small ophiovirus particles. The bar represents 100 nm. Courtesy of RG Milne, IPSP-CNR, Torino, Italy.

of NGS technologies on complex samples, reports regarding aspivirus-like isolates in fungi note similarities of their RdRp protein sequences to those of LRNV and MLBVV, but further characterization is needed to assess their putative taxonomic position in the family Aspiviridae. Partial ophiovirus-like sequences have also been reported from NGS studies of Plantago lanceolata. Interestingly, a recent report describes the presence of endogenous viral elements (EVEs) putatively derived from an ophiovirus in the genome of the eelgrass Zostera marina. EVEs are sequences in eukaryotic genomes that are derived from the ancestral integration of viral sequences into germline cells; in this case the ancestral sequence is represented by the ophiovirus CP ORF; this finding, if confirmed, would extend the Aspiviridae host range to seagrasses and expand ophiovirus evolutionary history.

Virion Structure The virions are naked filamentous nucleocapsids about 3 nm in diameter forming circularized structures of different lengths, presumably corresponding to the different lengths of the genomic segments. The virions appear to form internally coiled circles that in some cases can collapse into pseudolinear duplex structures (Fig. 2) and can be difficult to visualize in TEM assays due to a low contrast with the background. There is no evidence of a lipoprotein envelope and the mechanism underlining the apparent circularization of ophiovirus particles remains incompletely explained. Ophiovirus particles can be purified from infected plant tissue using differential centrifugation, clarifying the suspension using PEG6000 and sodium chloride, and density gradient ultracentrifugation. Particles are unstable in cesium chloride but the particle structure remains intact in cesium sulfate gradients (buoyant density is 1.22 g/cm3 for RWMV and MLBVV). Particle structure survives limited treatment with organic solvents and nonionic or zwitterionic detergents.

Genome Organization and Replication The ophiovirus genome is ssRNA, 11.3–12.5 kb in size, divided into three or four segments (RNAs 1–4) of mainly negative sense. Positive- and negative-sense RNAs are individually encapsidated in ophiovirus particles, and the more abundant negative sense RNA is referred to as viral RNA (vRNA), while the complementary RNA will be described as viral complementary (vcRNA). The 30 termini of the vRNAs show sequences of 6–13 nt conserved within the RNAs of each species: A7GUAUC for CPsV, A4–6UAAUC for MLBVV, A7GUAUCA and A3UA3GUAUCA for LRNV; A2UAUC for BlMaV; A4UGUAUC for FreSV (preliminary results); these may be involved in the recognition of the RNAs by the RdRp. The 50 termini of BlMaV RNAs were shown to fold into conserved stem-loop structures. These structures may be involved in packaging of genomic RNAs or in the long-distance interactions for transcription and replication. The pseudo-circularized appearance of ophiovirus particles suggests the presence of panhandle structures at genomic RNA ends, but no conclusive results were inferred for the four fully sequenced species. In CPsV, the first 12 nt of 50 end of vcRNAs were found almost identical in the three RNAs, but unexpectedly no identity among the three RNAs is found at their 30 ends, and no self-complementary panhandle structures between the 30 and 50 ends of each RNA could be hypothesized. In the case of MLBVV and LRNV, both 50 and 30 ends are highly or partially conserved among the four

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Table 2

Ophioviruses (Aspiviridae)

Details of genomic organization of fully sequenced viral segments for viruses of the genus Ophiovirus

Virus

RNA1

RNA2

RNA3

RNA4

BlMaV

7963 nt 2 ORFs in vcRNA ORF1: 23 kDa ORF2: 272 kDa 8186 nt 2 ORFs in vcRNA ORF1: 24 kDa ORF2: 280 kDa 7651 nt 2 ORFs in vcRNA ORF1: 22 kDa ORF2: 261 kDa 7794 nt 2 ORFs in vcRNA ORF1: 25 kDa ORF2: 263 kDa

1934 nt 1 ORF in vcRNA ORF1: 58 kDa

1570 nt 1 ORF in vcRNA ORF1: 50 kDa

–

1645 nt 1 ORF in vcRNA ORF1: 54 kDa

1447 nt 1 ORF in vcRNA ORF1: 49 kDa

–

1830 nt 1 ORF in vcRNA ORF1: 50 kDa

1527 nt 1 ORF in vcRNA ORF1: 48 kDa

1417 nt 1 ORF in vcRNA ORF1: 37 kDa

1788 nt 1 ORF in vRNA ORF1: 10 kDa (?) 1 ORF in vcRNA ORF1: 55 kDa 1722 nt 1 ORF in vRNA ORF1: 54 kDa

1515 nt 1 ORF in vcRNA ORF1: 49 kDa

1402 nt 2 ORFs in vcRNA ORF1: 37 kDa ORF2: 11 kDa

nd 1 ORF in vcRNA ORF1: 48.8 kDa

?

CPsV

LRNV

MLBVV

FreSV

nda

a

not fully determined; - not detected.

Fig. 3 Genome organization of ophioviruses. The four genomic RNAs of MLBVV, the first ophiovirus to be full sequenced, are represented to scale. An RNA4 is not described in all species, see text and Table 2 for details; v-RNA and vc-RNA indicate the viral and the viral complementary RNAs, respectively. Boxes indicate ORFs and the function of the encoded protein, if known, is reported.

viral RNAs but are not predicted to form panhandle structures; one hypothesis is that their RNAs ends fold into structures resembling the “corkscrew” conformation of RNA termini of members of the family Orthomyxoviridae, but this structure was not found for CPsV. Regarding BlMaV no complementarity was found between the 50 and 30 ends of each genomic RNA, as found for CPsV; in conclusion, alternative explanations for the pseudo-circular structure of ophiovirus particles are required. The genome organization of ophioviruses is summarized in Table 2 and Fig. 3. All RNA segments have 50 and 30 untranslated regions. RNA1, the largest genome segment, is negative sense and contains two ORFs separated by an AU-rich intergenic region. The presence of two ORFs, the longest encoding the RdRp, with the same polarity in the largest RNA is a distinct genomic feature of Aspiviridae when compared to all other segmented negative-strand viruses. RNA2 contains one ORF in the negative strand in all four fully sequenced viruses. An additional minor ORF, in the virus-sense strand, is present for MLBVV, but to date there is no evidence for expression of the predicted 10 kDa protein. RNA3 contains one ORF in the vcRNA in all viruses, which has been identified as coding for the nucleocapsid protein. Up to now, an RNA4 has been found only in MLBVV and LRNV. Both encode a 37 kDa protein in the vc strand, with only MLBVV having a smaller (10.6K) ORF which lacks an initiation codon and is potentially expressed by a þ 1 translational frameshift near the 30 end of the 37K ORF. A slippery sequence, GGGAAAU, can be recognized immediately in front of the UGA stop codon of the 37 kDa ORF. This unique feature of ORF positioning has been shown for two different isolates of MLBVV, and therefore seems not to be caused by cloning artifacts. RNAs 1, 2, and 3 of the other partially sequenced species are similar in size to those described. No RNA4 has been found in CPsV, RWMV, BlMaV and TMMMV, in spite of multiple NGS reactions, or PCRs using primers designed from the NGS data and conserved regions of the MLBVV/LRNV 37K proteins, which have been performed at least for BlMaV; its presence in FreSV is under study. Analysis for the presence of sub-genomic RNAs has not extensively been pursued for all species, even though several studies have used minus-strand and plus-strand probes in Northern blot hybridization using RNA extracted from infected tissues as well as virus particles, to obtain information regarding the polarity of encapsidated viral RNA. No subgenomic RNAs were ever validated for any of RNA1, 2, or 3 of CPsV. Reassortment between genome segments from different isolates may occur in plants infected with more than one isolate, as it has been shown for BlMaV.

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The presence of one or more NLS reported for the RdRp protein (see below) suggests that replication may occur in the nucleus; however, gold immunolabeling and electron microscopy of RWMV-infected tissues showed accumulation of CP in the cytoplasm.

Properties and Functions of Gene Products Two proteins are encoded in the viral complementary (vc) strand of RNA1, separated by an intergenic region of about 100 nt; the first of these is a protein of approximately 22–25 kDa. For CPsV and MLBVV this protein has been shown to function as viral suppressor of RNA silencing (VSR), and it is likely that the equivalent proteins of other ophioviruses have the same function; consistent with this function, a CPsV 24K:GFP fusion protein was found to accumulate in the nucleus and probably the nucleolus despite having an identified NES. Nucleolar import of CPsV 24K may be aided by an N-terminal proximal “GW” motif, while the BlMaV 23K protein has an identified bipartite NLS not seen in the other ophioviruses for which the small protein from RNA1 has been characterized. Indeed, the BlMaV 23K has low identity to the equivalent proteins of CPsV, MLBVV, and LRNV. The CPsV 24K and MLBVV 25K proteins have both been demonstrated to bind long (114 bp) but not short (21 bp) dsRNAs, a property shared with their 54–55K MPs which have also been identified as VSRs (see below), and which also affect miRNA maturation. The second, and major protein encoded by RNA1 is the viral RdRp, which is of 261–280 kDa. The RdRp of each virus has the signature motifs present in RdRp of viruses of the order Mononegavirales, having a core polymerase module of about 240 aa residues which includes five conserved motifs; pre-motif A, plus motifs A to D. Of these, motif C is absolutely conserved between the five viruses for which the core polymerase sequence is available (CPsV, MLBVV, LRNV, RWMV, and BlMaV), and includes the presumed RdRp active site “SDD” triad found in all negative strand RNA viruses with segmented genomes, including ophioviruses. In motif D, a Gly residue is conserved between MLBVV, LRNV, and RWMV, but is substituted by an Asp residue in CPsV and BlMaV. The RdRps of CPsV, MLBVV, RWMV, and BlMaV all have identified bipartite NLSs, with CPsV having dual NLSs, and the other viruses having only one, though no NLS has been reported for the RdRp of LRNV. In most viruses with a negative sense RNA genome, the RdRp is also incorporated in the virions, as in the absence of translation from the incoming genomic RNA, the RdRp is required to initiate infection. However, a variable proportion of positive (vc) strand RNA of ophioviruses is also encapsidated, which may overcome the necessity for other viruses with negative sense genomes to incorporate the RdRp into virions. All of the characterized ophioviruses produce a protein of 50–58 KDa translated from the RNA2 vc strand, initially identified as a movement protein belonging to the “30K superfamily” of MPs, and later shown to also include VSR activity. The MPs of CPsV and MLBVV interact with PD (presumably increasing its SEL, a characteristic of many different MPs), and with cellular microtubules, and also spread into adjacent cells. CPsV and MLBVV MPs interact with their homologous nucleocapsid protein (NP); interaction of MP with NP/coat protein is a common feature of MPs of other viral genera and families. The predicted structures of ophiovirus MPs align with each other, and have a similar organization comprising a core domain, common to all MPs of the “30K superfamily”, including one a-helix, seven b-strands, and the conserved Aspartic acid (D) residue associated with increased SEL of PD and full MP function. The C-terminal domain has conserved secondary structure potentially involved in RNA binding and interactions with host proteins, and a conserved DTG tripeptide not observed in any other “30K superfamily” MPs. The 54K MP of CPsV is autocatalytically cleaved by an aspartic protease to generate an N-terminal 34K protein including the “30K core domain”, which forms tubules at PD to allow virus movement between cells, and a 20K product. The 20K includes the protease activity within the MP conserved C-terminal domain, with the absolutely conserved DTG tripeptide near the N-terminus of this domain; the cleavage site occurs about 30 residues upstream. The D residue is critical for cleavage; however, a high proportion of the CPsV 54K protein remains unprocessed even in native infections. Cleavage affects MP subcellular localization, and is required for efficient nuclear localization (presumably of the 20K cleavage product). Uncleaved 54K accumulated to a greater extent at chloroplasts, and although also observed at PD, no tubular structures were observed; whereas 34K expressed alone resulted in tubule production at PD. Chloroplast localization is presumably due to a predicted 21 aa residue N-terminal chloroplast transit peptide. The 34K cleavage product complements a movement-deficient TMV more efficiently than the full-length 54K, confirming movement function is due to the 34K. Both the 34K and 20K cleavage products form homodimers, consistent with 34K forming tubules, and the presumed dimeric catalytic triad for aspartic protease activity of the 20K product. Most viruses with tubule-forming MPs have isometric virions; it has been suggested that interaction between CPsV MP and NP may allow guidance of virions (or a ribonucleoprotein complex of viral RNA with both MP and NP) through tubules linking adjacent cells through PD. To date, only protease activity has been ascribed to the 20K cleavage product. Both (54K) MP and the RNA1 24K protein each have VSR activity, which is enhanced when the two proteins are co-expressed; each presumably functions at a different step of the RNAi pathway. As a 20K:GFP fusion protein traffics to the nucleus, the 20K cleavage product may have the VSR activity originally assigned to the 54K product. It has been proposed that the CPsV 54K protein should be renamed “MP-PRO”, with the 30K cleavage product identified as “MP”, and the 20K cleavage product as “PRO”. This suggested nomenclature may be further modified if the VSR activity first associated with the uncleaved 54K protein is attributed to a specific cleavage product. A minor protein of about 10 kDa is apparently encoded in the viral (v) strand of MLBVV RNA2 and this would represent a unique case among the known ophiovirus genome structures, but its significance is still unclear. RNA3 of all characterized ophioviruses encodes a single protein of 43–50 kDa, which is the viral nucleoprotein, alternatively described as the “coat protein” (CP) forming at least the bulk of the filamentous virions together with the genomic RNAs. The NP protein sizes of different ophioviruses ranges between 43 and 50 kDa (see Table 2), with FreSV NP of 48.4 kDa, and the sizes of RWMV (B43 kDa) and TMMMV (B47 kDa) NPs estimated by western blotting rather than from partial NP sequence data.

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Ophiovirus NPs are relatively poor antigens. The antisera or, in one case, monoclonal antibodies, have been produced using purified virus preparations or recombinant protein. Western blots show that the NPs of RWMV, TMMMV, MLBVV, and LRNV are slightly to moderately related; CPsV NP appears to be unrelated to the others; no clear information is available for FreSV or BlMaV. Genetic variability of the NP gene has been extensively studied in CPsV and MLBVV. Variability of the NP gene of CPsV was assessed serologically, and by sequence analysis of two regions located in the 30 and 50 halves of the gene. CPsV serological variability yielded 14 reaction patterns, without correlation between serogroups and specific aa sequences, field location, or citrus cultivar. Sequence analysis showed limited nucleotide diversity in the NP gene within the population, with diversity slightly higher in the 50 region. The ratio between non-synonymous and synonymous substitutions (dN/dS) for the two regions indicated a negative selective pressure for aa changes, with greater selection pressure at the 30 end. When the entire CP sequences were considered, two clusters were identified, one comprising 19 isolates from Italy and an isolate from Spain and a second one containing only the Florida CPsV isolate. Phylogenetic analysis of MLBVV NP genes has revealed two distinct clades, A and B, further divided into subclades A1 and A2, B1 and B2; however, this grouping was not correlated with symptom severity on lettuce or the geographic origin of the isolates, except that all Australian isolates fell into subclades A1 and A2; whereas European isolates were found in all four subclades, only European isolates fell into B1 and B2, including all reported isolates from Sonchus spp. in subclade B2. Whether these two clades or subclades show different characteristics with regard to virulence in indicator plants or serological relationships remains to be determined; as with CPsV, a low ratio of dN/dS was estimated for all MLBVV isolates and between the two subgroups, implying negative selection pressure for aa changes. A similar study of BlMaV MP and NP sequences of isolates from British Columbia (Canada), Slovenia, and several regions within the United States found a low ratio of dN/dS for both genes. Under natural conditions, genetic stability of ophiovirus NPs seems to be the rule rather than the exception, perhaps due to the necessity both for structural integrity of the virions, and interaction with host or vector proteins. The NP encapsidates both negative (v) and positive (vc) strands of the genomic RNA. Each NP includes a domain of almost 180 aa residues that is conserved among the NPs of viruses with negative strand ssRNA genomes, and is rich in a-helices. The NP of MLBVV and partial NP sequence of TMMMV share about 80% identity, showing the closest relationship between ophiovirus species; although distant serological relationships between MLBVV and TMMMV, and MLBVV and RWMV indicate the presence of some conserved epitopes between these viruses, no serological relationship was detected between these viruses and CPsV. CPsV and BlMaV NPs share only 38% identity, despite their current status as the only ophioviruses infecting woody hosts. A putative leucine zipper motif of six (CPsV) or four turns (MLBVV, TMMMV, FreSV, LRNV, and RWMV) followed by an SRCK tetrapeptide has been identified in the NPs of the six ophioviruses for which all or most of the NP sequence is available; for the complete NP of BlMaV, a five turn leucine zipper motif followed by a TRCK tetrapeptide at the same spacing can be identified. Leucine zippers are associated with protein oligomerization and genome encapsidation, consistent with NP self-interaction and role of the NP in virion formation. Whether the SRCK/TRCK tetrapeptide may be involved in a specific interaction with zoospores of the various Olpidium spp. known to be vectors of most ophiovirus species remains to be determined. An RNA4 has been identified only for MLBVV and LRNV; the 37 kDa proteins of MLBVV and LRNV share just over 40% identity, with a higher proportion of conserved residues in the N-terminal 60 aa residues than any other portion of the protein, suggesting that this is a functional domain; both the MLBVV and LRNV 37K proteins are predicted to have three trans-membrane domains, with the N-terminus ‘outside’, and the C-terminus “inside”. Expression of the putative 10.6K frame-shift ORF present only in RNA4 of MLBVV has not been demonstrated. To date no function has been assigned to either of the ORFs encoded by RNA4.

Pathogenicity and Geographic Distribution Psorosis is a graft-transmissible disease of citrus characterized primarily by bark scaling of the trunk and main branches of various Citrus species (Fig. 4). These symptoms were first reported from Florida and California, USA in the 1890s, and in the great majority of cases psorosis symptoms are associated with infection by CPsV. However, in a small proportion of trees with “psorosis-like” barkscaling CpSV is not detected, and the idea of a non-CPsV psorosis-like disease of unknown etiology is emerging. In the past, different kinds of symptoms were described: “psorosis A” (PsA), characterized by causing bark-scaling in trunk and limbs of infected field trees, and staining of the wood; “psorosis B” (PsB), causing rampant scaling of thin branches in field trees and chlorotic blotches in old leaves with gummy pustules in the underside; “ringspot”, characterized by presence of chlorotic blotches and rings in the old leaves of inoculated seedlings but apparently no specific symptoms on infected field trees or in other cases, chlorotic flecks and ringspots on leaves, and trunk and fruit symptoms. All these appear to be caused by CPsV. CPsV has been documented in many citrus production areas, including the USA, Argentina, Uruguay, Spain, Italy, Turkey, Lebanon, Egypt, Morocco, Tunisia, China, and Australia. The disease caused by CPsV has been brought under control in most advanced citrus-growing countries through implementation of rigorous indexing and quarantine. In Argentina it remains a severe problem, and it is still reported in the Mediterranean areas, while in citrus-growing parts of Asia the disease may well be widespread, although rigorous testing for the presence or absence of CPsV has generally been lacking. Among citrus types, sweet orange, grapefruit, and mandarin are most severely affected, whereas sour orange and lemon usually show foliar symptoms without obvious bark lesions. Infection with PsA isolates offers some cross-protection against the more severe symptoms induced by PsB isolates (also sometimes referred to as “Citrus ringspot” isolates). Disease severity is also affected by temperature, with cool temperatures increasing the probability of a shock reaction in the first growth flush following inoculation, whereas warmer temperatures may result in masking of foliar symptoms. Increased temperatures appear to allow greater RNA silencing by the host, resulting in lower viral accumulation.

Ophioviruses (Aspiviridae)

501

Fig. 4 Bark scaling in citrus, typical of severe psorosis.

Lettuce big-vein disease was first reported in the United States in 1934 and occurs in all major lettuce producing areas in the world. The disease becomes more serious during cooler periods of the year. The main symptoms are vein-banding in the leaves, due to zones parallel to the vein cleared of chlorophyll (Fig. 5); there is associated leaf distortion, delayed head formation, and decreased head size. The causal agent of the disease, long known to be soil-transmitted, was first identified as “Lettuce big-vein virus” (LBVV) (genus Varicosavirus); more recently, re-evaluation of the etiology was necessary as a second less easily detected virus, MLBVV, was found in lettuces with big-vein symptoms. Following experimental inoculation of the two viruses together and separately, MLBVV was shown to be the etiological agent of big-vein, while LBVV, renamed as Lettuce big-vein associated virus (LBVaV), apparently plays no part in the disease, although it is almost always present, and is, like MLBVV, transmitted by Olpidium. Studies on symptom development in the field have confirmed that both viruses very commonly occur together in lettuce crops. MLBVV likely occurs worldwide; it is reported in California (USA), Brazil, Chile, France, Germany, Italy, Spain, the Netherlands, England, Denmark, Turkey, South Africa, Saudi Arabia, Japan, Australia, and New Zealand. To date, natural infection has been reported only in cultivars of Lactuca sativa, endive (Cichorium endivia), and two species of sowthistle (Sonchus asper and S. oleraceus). LRNV is closely associated with lettuce ring necrosis disease, first described in the Netherlands and in Belgium as “kring necrosis” and also in France as “maladie des taches orangées”, in the 1980s. It is an increasingly important disease of butterhead lettuce crops in Europe. Definitive proof that LRNV is the cause of ring necrosis is still awaited. In southern France the disease is observed primarily in winter lettuces (September–January) under plastic or glass, when the crop is maturing, the day length is short, and both light intensity and temperature are low. Symptoms depend on the lettuce type and environmental conditions, and mainly consist of necrotic rings and ring-like patterns on leaves, which may render the product unmarketable. LRNV is often found together with MLBVV (and, of course, LBVaV) with which it shares host and vector. The virus has also been reported in California.

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Ophioviruses (Aspiviridae)

Fig. 5 Leaf of butterhead lettuce showing big-vein symptoms. Close-up showing vein-banding.

Fig. 6 Shoot of blueberry infected with BlMaV showing part of the range of symptoms often observed, which include chlorotic fleck and mottling, strong chlorotic mosaic, and combinations of yellow mosaic with areas of pink or orange coloration.

BlMaV was initially described from an infected plant from Arkansas, USA, showing symptoms of mild to brilliant foliar mottle and mosaic (Fig. 6) observed since the 1950s in blueberry production areas across North America, Argentina, Chile, Europe, New Zealand, and South Africa. The disease is associated with late ripening, reduced yield, and poor quality fruit. Some leaves may turn partly or completely yellowish green, yellow, orange, or pink. BlMaV has been specifically identified by molecular methods from several states in the USA, from British Columbia (Canada), Slovenia, Serbia, Turkey, and Japan, with the Slovenian isolates being most closely related to isolates from New Jersey, USA. Most of the Slovenian isolates were propagated from plants originally imported from New Jersey. Necrotic disorders of freesia (Freesia refracta hyb., family Iridaceae), known as “leaf necrosis” and “severe leaf necrosis”, were first described in freesia crops in the Netherlands before 1970, and a similar disease, named “freesia streak”, was reported in England and Germany in the same years. The ‘severe leaf necrosis’ appeared to be caused by mixed infection with the potyvirus Freesia mosaic virus and a virus with varicosavirus morphology and the same mode of transmission, that has been tentatively named Freesia leaf necrosis virus. More recently in northwest Italy, a necrotic disease of freesia spread and caused considerable economic losses (Fig. 7), even though it was present since 1989. Both in Italy and in the Netherlands, the disease has been linked to the presence of an ophiovirus, named Freesia sneak virus (FreSV). Typical symptoms are chlorotic inter-veinal spots and streaks at the leaf tips that expand downwards and turn necrotic, and may vary according to cultivar and climate. FreSV has since also been identified in freesia in England, Germany, the Netherlands, Bulgaria, New Zealand, Korea, Japan, and the USA, and from Lachenalia hybrids (family Asparagaceae) from South Africa. RWMV was initially reported in two species, ranunculus (Ranunculus asiaticus hyb.) and anemone (Anemone coronaria) in northwest Italy since the 1990s. The pathogenic impact is uncertain as it was almost always found in mixed infection with other viruses commonly infecting the two species. The symptom description “white mottle” present in its name derives from the bright white mottling symptoms consistently observed on Nicotiana benthamiana leaves mechanically infected by RWMV. In some cases the bright mottle has also been observed in naturally infected ranunculus, always in mixed infection. Indexing for the presence of RWMV in ranunculus crops done in 1996 showed an incidence of 2.5% among symptomatic plants; furthermore, the very few plants apparently infected only with RWMV did not show any distinctive symptoms. Infection of ranunculus seedlings by mechanical inoculation results in limited necrosis and

Ophioviruses (Aspiviridae)

503

Fig. 7 Symptoms of freesia necrotic disease on freesia leaves. Courtesy of D. Salvi, Studio Ferrari Salvi, Sanremo (IM) Italy.

Fig. 8 TMMMV symptoms on tulip (healthy tulip on the left). Courtesy of T. Morikawa, Toyama Agric. Res. Center, Tonami, Toyama, Japan.

deformation of stems and leaves; however, it is one of the most prevalent viruses in ranunculus crops in Italy, possibly as a result of establishment of the Olpidium vector in production fields. Infection of anemone was also typically in mixed infections, leaving the pathogenic potential of RWMV unclear. A report published by Horticulture Australia Ltd notes RWMV infecting pepper showing vein yellowing disease in Australia; the disease showing incidence up to 30% is widespread on the North Adelaide Plain but does not appear to cause significant yield loss or reduction in fruit quality. The symptoms are easily seen in young plants and fading as plants mature. More recently RWMV was detected in Pittosporum tobira in Italy, in mixed infection with Eggplant mottled dwarf rhabdovirus. Tulip mild mottle mosaic disease, caused by TMMMV, is one of the most serious diseases in some bulb-producing areas of Japan (which produces bulbs on a large scale for the Southeast Asian market) and has been reported since 1979. Symptoms on tulips include color-attenuating mottle on flower buds and color-increasing streaks on petals (Fig. 8); mild chlorotic mottle and mosaic may appear along the leaf veins. TMMMV infection is apparently restricted to Tulipa species and cultivars, and has not been reported in other geographical areas.

Diagnosis A pair of degenerate primers was originally designed from a conserved region of RNA1 (in the RdRp gene, located outside the sequence coding the polymerase domain). These primers, used in RT-PCR experiments, amplify a 136 bp DNA fragment from all recognized ophiovirus species and not from representative plant-infecting species of Rhabdoviridae, Tenuivirus, Orthotospovirus and Varicosavirus. These genus-specific primers, by a simple RT-PCR assay, may lead to the detection of new ophioviruses, with the establishment, at least provisionally, of their taxonomic position, through phylogenetic analysis of the predicted 45 aa translation of the amplified fragment. When an ophiovirus infection is suspected, RT-PCR and qRT-PCR with specific primers are the most commonly used and sensitive method for specific diagnosis, both in plants and in the natural vector, even if ELISA using polyclonal antisera is still used for mass diagnosis. An RT-LAMP protocol was also developed for MLBVV; compared with DAS-ELISA and RT-PCR, RT-LAMP and IC-RT-LAMP (Immuno-Capture of particles) seem to have higher sensitivity (100-fold) but

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Ophioviruses (Aspiviridae)

similar specificity, with the advantage of a shorter assay time and no need for RNA extraction in IC-RT-LAMP. Recently NGS technology gained high importance in virus diagnosis, and the “small RNA-Seq” technique has been used to characterize the causal viruses of lettuce big vein disease, confirming the crucial involvement of MLBVV. NGS technology has a huge potential and may be easily used to find new hosts for ophiovirus species (as happened for the RWMV infection of Pittosporum tobira) and help to find new ones (as happened for the disclosure of blueberry infection by BlMaV, the most recently recognized official ophiovirus).

Transmission, Experimental Host-Range and Control Most ophiovirus infections in nature are recognized as soil-borne. The natural vector of most ophiovirus species has been identified as Olpidium spp., ubiquitous obligate root-infecting fungal parasites (Chytridiomycota, chytrids) with cosmopolitan distribution. At least two species, O. brassicae and O. virulentus seem to be involved in ophiovirus transmission. Studies on MLBVV and O. virulentus suggest that the virus is carried internally (in vivo) in zoospores and resting sporangia. Virus-free zoospores acquire the virus during the vegetative part of their life cycle in the roots of virus-infected host plants. Zoospores released from infected sporangia carry the virus internally in their protoplasts and transmit it to healthy roots. In vivo acquisition is also supported given that resting spores can remain viruliferous in soil for decades in the absence of a growing host plant. The mechanism by which MLBVV and likely other ophioviruses are acquired and released into plant hosts by Olpidium is unknown; the possibility of ophioviruses multiplying in the fungal vector has not been reported to date. Olpidium spp. transmission has been proved for TMMMV, MLBVV (O. virulentus), LRNV, FreSV (O. brassicae) and BlMaV (O. virulentus). No information on natural vectors is available for RWMV but soil transmission is likely. CPsV is commonly transmitted by vegetative propagation and no natural vectors have been so far identified; in some cases natural tree-to-tree spread of psorosis in limited citrus areas has been reported, but the spatial patterns would suggest a hypothetic aerial vector instead of a soil-borne one. However, CPsV has been detected in the roots of infected grapefruit trees, and in zoospores of an unidentified Olpidium-like fungus released from sporangia in infected roots, but not from healthy controls. The thick cell-walled nature of Olpidium spp resting spores and their persistence in the soil for greater than 20 years makes ophiovirustransmitted disease control most challenging, especially after the banning (Montreal Protocol) of use of chemicals for soil sterilization due to deleterious effects on the environment. Environmentally more sensitive control methods can be applied in order to reduce the spread and severity of the diseases. For nurseries, for example, disinfection of soil and potting mix, surface sterilization of seed, and the use of filtered/UV-treated water and nutrients, and clean production areas may all be important in the management of MLBVV and LRNV in lettuce. Crop covers, which affect the favorable environmental conditions for the viruses by lowering soil temperature and raising air temperature, were also shown to reduce the disease symptoms in lettuce. Moreover, the use of resistant or tolerant crops may be a good choice. In Japan, the use of resistant tulip cultivars is the most important component of managing TMMMV disease, as it can be highly effective and has no deleterious effect on the environment; resistance assays have allowed researchers to identify highly resistant tulip lines and use them for breeding new resistant cultivars. In the case of CPsV, control of the sanitary status of mother plants for producing propagating material is essential. Shoot-tip grafting in vitro associated with thermotherapy or somatic embryogenesis from stigma and style cultures have been successfully used to eliminate CPsV from plant propagating material. Preliminary results of a shoot apical meristem culture procedure performed on RWMV-infected ranunculus was also successfully used for ophiovirus eradication: the use of seeds and in vitro virus eradication can also be applied to currently vegetatively propagated ornamentals like freesia, ranunculus and anemone, to control ophiovirus infection. Genetic engineering of citrus and lettuce in recent years has shown promising results for obtaining ophiovirus resistance; several pineapple sweet orange plant lines transformed with a hairpin construct derived from the CPsV nucleoprotein gene showed varying responses from tolerance to full resistance traits toward both psorosis variants, a strategy that seems promising for a biotech product aimed at eradicating psorosis. Stable, marker-free and MLBVV-resistant lettuce lines were also engineered, in which MLBVV nucleoprotein gene is controlled by plant regulators; the new lines can be used as resistant cultivars or parental source for breeding. All of the characterized ophioviruses are mechanically transmissible to a limited range of test plants, including Chenopodiaceae and Solanaceae, inducing local lesions; in some cases systemic mottle or apical leaf deformation and necrosis can occur. Systemically infected test plants can, in many instances, be advantageous for virus characterization, because of the enhanced virus multiplication and the possibility to obtain a high amount of purified virus. Chilling of source tissue, buffers, and equipment, together with additives such as antioxidants, enhance mechanical transmission of most, if not all ophioviruses. Nicotiana benthamiana, N. clevelandii, N. occidentalis, and Chenopodium quinoa are among the bioassay plants most often reported to be susceptible (Table 3). However, mechanical transmission is often difficult, and serial transfer between test plants is not always successful; there are also differences in ability or efficiency of mechanical transmission between isolates of CPsV. Transmission by the natural vectors, Olpidium spp. (O. brassicae, O. virulentus, or O. bornovanus) has rarely been used for host range tests, while dodder (Cuscuta spp.) has been demonstrated to transmit CPsV between citrus types. CPsV is efficiently transmitted to susceptible citrus or citrus-related species by grafting, infecting Microcitrus inodora and Fortunella hindsii symptomlessly but with high ELISA values; other citrus relatives (Citrus depresa, Carrizo citrange, Eremocitrus glauca, Microcitrus virgata, and Clausena excavata) produced a severe shock reaction in the first flush of leaves, but did not flush again. Symptom intensity of some susceptible hosts (typically chlorotic flecking in young leaves) is correlated with virus accumulation as measured by ELISA, but mild symptoms were observed in Cleopatra mandarin, sour orange (C. aurantium L.), and citrumelo (C. paradisi  Poncirus trifoliata), which were negative by ELISA. Mechanical inoculation to test plants is also achieved. MLBVV is reported to be more readily mechanically-transmissible than LBVaV, with which it is commonly found in mixed infection in lettuce; serial transfers from an initial mixed infection have resulted in recovery of only MLBVV after the third transfer to either lettuce or N. benthamiana. Continued mechanical transfers may result in loss of transmission by O. brassicae, and loss of

Ophioviruses (Aspiviridae)

Table 3

505

Symptomatology on test plants commonly used for ophiovirus biological assays

Test plant

CPsV

MLBVV

LRNV

RWMV

TMMMV

FreSV

C. quinoa C. amaranticolor Gomphrena globosa Datura metel N. benthamiana N. rustica N. occidentalis N. clevelandii N. tabacum N. hesperis N. megalosiphon

CLLa C/NLL NLLc, SSd CLL, SS – SS – – – – –

LLb – – – SSIe – LL, SS SS LL SS –

C/NLL C/NLL – – SSI – C/NLL, SS SSI – C/NLL, SS –

LL LL – – CLL, SS – – CLL, SS LL – CLL, SS

LL LL – – LL – – – LL – –

CLL CLL – – – – NLL, SS – – NLL, SS –

a

Chlorotic local lesions. Local lesions. c Necrotic local lesions. d Systemic symptoms. e Symptomless systemic infection. - no information available. b

ability to cause big vein symptoms in lettuce. Mechanical transmission of LRNV to test plants has been reported, with the most evident systemic symptoms in N. occidentalis and N. hesperis. BlMaV is the most recently identified ophiovirus, and to date there are no reports of mechanical transmission, and no host range data are available, beyond blueberry infection. Transmission assays using blueberry seedlings and BlMaV-infected Olpidium virulentus were positive. RWMV is able to infect several test plants; the inoculum source can maintain infectivity only if stored in liquid nitrogen. Local lesions are observed also in C. capitatum, C. murale, C. album, and local infection is also detected in Spinacia oleracea and Vigna unguiculata. Ranunculus seedlings (Ranunculus sardous and Ranunculus hybrid) challenged by mechanical inoculation with RWMV became infected, showing local and systemic necrosis. FreSV can be transmitted to test plants; in some cases serial passage on N. occidentalis may yield evident chlorotic mosaic. Mechanical inoculation of purified FreSV to healthy freesia seedlings was never successful, however, freesia seedlings challenged by FreSV-infected O. brassicae resting spores became positive for FreSV infection. Transmission of TMMMV to common test plants and to Tetragonia expansa, Sonchus oleracea, and Beta vulgaris resulted in only local infection.

Cytopathology Thin sections of Nicotiana clevelandii leaves mechanically infected by RWMV have been examined by electron microscopy, but no distinctive inclusions were observed and no virus particles were seen. In classical thin sections, viral nucleic acids stain well but protein coats show low contrast, so it is not surprising that very thin nucleoprotein threads in random orientations should escape detection. After gold immune labeling with RWMV-specific polyclonal antibodies, sections of the cytoplasm of parenchyma cells were seen in the EM to be clearly labeled, but nuclei, chloroplasts, mitochondria, and microbodies were unlabeled. Confocal observation for subcellular localization of CPsV and MLBVV coat proteins linked to the fluorescent GFP molecule show cytoplasmic localization. Ultrastructural changes have been reported in leaves of grapefruit infected with CPsV, but apparently free of Citrus tristeza virus (CTV) and Citrus exocortis viroid (CEVd), resulting in decreased thickness of both the phloem and the xylem, including reduced vessel diameter. Fewer oil glands, of smaller diameter than normal were observed, together with many abnormal chloroplasts with disorganized grana stacks and thylakoid membrane. Crystal idioblasts (cells containing calcium oxalate crystals) were found to be lacking in the palisade layer below the upper epidermis, in contrast to leaves of healthy plants, while inclusion bodies and tubular multi-membrane bodies were also observed. Although negative for CTV and CEVd, these plants may have been infected with other viruses, and some of the effects reported may result from reduced auxin levels rather than a direct presence of viral components.

Concluding Remarks Ophioviruses are probably more widespread than currently known and they apparently have a very long evolutionary history. Psorosis of Citrus spp. and big-vein disease of lettuce, together with the necrotic disease of freesia and the blueberry syndrome, are relevant in agricultural production. Crop protection against ophiovirus infection is prevention centered; for this reason, there is a

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need for resistance genes to be identified, or for introduction of resistance through biotechnological methods, together with application of good agricultural practices.

Further Reading Andika, I.B., Kondo, H., Sun, L., 2016. Interplays between soil-borne plant viruses and RNA silencing-mediated antiviral defense in roots. Frontiers in Microbiology 7, 1458. doi:10.3389/fmicb.2016.01458. García, M.L., Dal Bó, E., da Graça, J.V., et al., 2017. ICTV virus taxonomy profile: Ophioviridae. Journal of General Virology 98, 1161–1162. Garcia, M.L., Dal, Bo, E., Grau, O., Milne, R.G., 1994. The closely related citrus ringspot and citrus psorosis viruses have particles of novel filamentous morphology. Journal of General Virology 75, 3585–3590. Garcia, M.L., Goodin, M., Sasaya, T., Haenni, A.-L., 2011. Negative-strand RNA viruses: The plant-infecting counterparts. Virus Research 162, 184–202. Robles-Luna, G., Peña, E.J., Borniego, M.B., Heinlein, M., García, M.L., 2018. Citrus psorosis virus movement protein contains an aspartic protease required for autocleavage and the formation of tubule-like structures at plasmodesmata. Journal of Virology 92, e00355 doi:10.1128/JVI.00355-18. Robles-Luna, G., Reyesa, C.A., Peña, E.J., et al., 2017. Identification and characterization of two RNA silencing suppressors encoded by ophioviruses. Virus Research 235, 96–105. Thekke-Veetil, T., Ho, T., Keller, K.E., Martin, R.R., Tzanetakis, I.E., 2014. A new ophiovirus is associated with blueberry mosaic disease. Virus Research 189, 92–96. Vaira, A.M., Accotto, G.P., Costantini, A., Milne, R.G., 2003. The partial sequence of RNA 1 of the ophiovirus Ranunculus white mottle virus indicates its relationship to rhabdoviruses and provides candidate primers for an ophio-specific RT-PCR test. Archives of Virology 148, 1037–1050.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/negative-sense-rna-viruses/w/aspiviridae Aspiviridae.

Orthotospoviruses (Tospoviridae) Renato O Resende, University of Brasilia, Brasilia, Brazil Hanu R Pappu, Washington State University, Pullman, WA, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of M. Tsompana, J.W. Moyer, Tospovirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00517-3.

Nomenclature aa Amino acid(s) AGO Argonaute 1 CAPS Cleaved amplified polymorphic sequence CP Coat protein or capsid protein diRNA defective interfering RNA ELISA Enzyme-linked immunological assays ER Endoplasmic reticulum HC-Pro Helper component-protease IGR Intergenic region kb Kilobase kDa Kilo Dalton LAMP Loop mediated amplification MP Movement protein

Glossary Ambisense genome Viral RNA genome with open reading frames in both the viral- and viral complementary (vc) sense on the same genome segment. Envelope Membrane-like structure that packages genome segments. Intergenic region It is the untranslated, A-U rich region found between the two open reading frames on the S and M RNA segments. Negative sense genome Viral RNA genome that codes for

NBS-LRR Nucleotide-Binding site - Leucine-Rich Repeat NGS Next-generation sequencing nt Nucleotide(s) ORF Open reading frame RdRp RNA-dependent RNA polymerase RISC RNA-induced silencing complex RNP Ribonucleoprotein satRNA satellite RNA UTR Un-translated region VIGS Virus-induced gene silencing VLPs Virus-like particles VPg Viral protein genome-linked VRC Virus replication complexes vRNA virion RNA

proteins in the vc sense. Transcription of vc mRNA is required for translation of viral proteins. Nucleocapsid Viral RNA encapsidated by nucleoproteins. Protoplast Plant cell lacking its cell wall. Ribonucleoprotein complex Consisting of the viral RNA genome segment, nucleoprotein, and a small number of polymerase molecules. Virion Quasispherical structure containing the viral genome and bounded by a membrane-like envelope.

History Diseases now known to be caused by Tomato spotted wilt virus (TSWV) were first reported in 1915 and were shown to be of viral etiology by 1930. This taxon of plant viruses was categorized as a monotypic virus group consisting of a single virus (TSWV) until the report of impatiens necrotic spot virus (INSV) in 1991. Thus, most of the characteristics which define the genus Orthotospovirus were obtained through investigation of TSWV even after the discovery of additional viruses in the genus (Table 1). Biological investigations beginning in the 1940s revealed a virus that had an unusually large host range and occurred in nature as a complex mixture of phenotypic isolates. However, it was one of the least stable viruses and most difficult plant viruses to mechanically transmit. Although the enveloped virions were observed in the 1960s, molecular characterization and elucidation of the genome organization were not completed until the early 1990s. The virus was shown to be vectored by thrips in the 1930s and later transmitted in a persistent manner. Thrips were demonstrated to be a host for replication of the virus and that replication was required for transmission in the early 1990s. Later, it was recognized that limited, localized replication may occur in thrips that does not result in the thrips becoming viruliferous. Advances in gene function and cellular biology have been limited due to the absence of a robust in vitro plant or thrips cell culture system, and lack of an efficient reverse genetics system. However, limited progress has been made utilizing gene expression systems and classical viral genetics.

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Table 1

Orthotospoviruses (Tospoviridae) Viruses members and tentative members of the genus Orthotospovirus (family Tospoviridae)

Virus species names

Acronym

Reported thrips vectors

GenBank accession #

Alstroemeria necrotic streak orthotospovirus Alstroemeria yellow streak orthotospovirus Bean necrotic mosaic orthotospovirus Calla lily chlorotic spot orthotospovirus Capsicum chlorosis orthotospovirus

ANSV AYSV BeNMV CCSV CaCV

Frankliniella occidentalis Thrips tabaci Unknown Thrips palmi Ceratothripoides claratris Thrips palmi Frankliniella schultzei

L: L: L: L: L:

Chrysanthemum stem necrosis orthotospovirus

CSNV

Frankliniella occidentalis Frankliniella intonosa Frankliniella schultzei

L: KM114546; M: KM114547; S: KM114548

Groundnut bud necrosis orthotospovirus

GBNV

Frankliniella schultzei Scirtothrips dorsalis Thrips palmi Scirtothrips dorsalis Frankliniella occidentalis Frankliniella schultzei Frankliniella gemina Scirtothrips dorsalis Unknown

L: AF025538; M: U42555; S: U27809

Frankliniella occidentalis Frankliniella fusca Frankliniella intonosa Frankliniella fusca Thrips tabaci Unknown Thrips palmi Unknown

L: X93218; M: M74904; S: X66972

Groundnut chlorotic fan-spot orthotospovirus GCFSV Groundnut ringspot orthotospovirus GRSV

MG696851; M: MG696852; S: MG696853 MF469033; M: MF469034; S: MF469035 JF417980; M: JN587269; S: JN587268 FJ822962; M: FJ822961; S: AY867502 DQ256124; M: DQ256125; S: DQ256123

L: KP146140; M: KP146141; S: AF080526 L: KT972590; M:KT972592; S: KT972594

Groundnut yellow spot orthotospovirus Hippeastrum chlorotic ringspot orthotospovirus Impatiens necrotic spot orthotospovirus

GYSV HCRV

Iris yellow spot orthotospovirus

IYSV

Melon severe mosaic orthotospovirus Melon yellow spot orthotospovirus Mulberry vein banding-associated orthotospovirus Pepper chlorotic spot orthotospovirus Polygonum ringspot orthotospovirus Soybean vein necrosis orthotospovirus

MeSMV MYSV MVBaV PCSV PolRSV SVNV

Unknown Dictyothrips betae Neohydatothrips variabilis Frankliniella fusca Frankliniella tritici

L: KX247379; M: KX247378; S: KX247377 L: KJ541746; M: KJ541745; S: KJ541744 L: HQ728385; M:HQ728386; S: HQ728387

Tomato chlorotic spot orthotospovirus

TCSV

L: MH742959; M: MH742960; S: MH742961

Tomato spotted wilt orthotospovirus

TSWV

Tomato yellow ring orthotospovirus Tomato zonate spot orthotospovirus Watermelon bud necrosis orthotospovirus Watermelon silver mottle orthotospovirus Zucchini lethal chlorosis orthotospovirus Tentative species in the genus Lisianthus necrotic ringspot virusa Pepper necrotic spot virusa Tomato necrotic spot virusa

TYRV TZSV WBNV WSMoV ZLCV

Frankliniella occidentalis Frankliniella schultzei Frankliniella intonosa Frankliniella occidentalis Frankliniella fusca Frankliniella schultzei Frankliniella intonosa Frankliniella bispinosa Frankliniella cephalica Frankliniella gemina Thrips setosus Thrips tabaci Thrips tabaci Frankliniella occidentalis Thrips palmi Thrips palmi Frankliniella zucchini

LNRV PNSV TNSV

Unknown Unknown Unknown

INSV

S: HQ402596 L: HG763861; M: JX833565; S: JX833564

L: FJ623474; M: AF214014; S: AF001387 L: KX698424; M: KX698423; S: KX698422 L: AB061774; M: AB061773; S: AB038343 L: KM819698; M: KM819699; S: KM819701

L: D10066; M: S48091; S: D00645

L: L: L: L: L:

JN560178; M: JN560177; S: AY686718 EF552435; M: EF552434; S: EF552433 GU735408; M: GU584185; S: GU584184 AF133128; M: U75379; S: U78734 KU641378; M: KU641379; S: KU641380

S: AB852525 S: HE584762

Tentative orthotospoviruses that have been described and proposed to the International Committee on Taxonomy of Viruses (ICTV) and waiting for the final approval as definitive species. Note: Updated table from Rotenberg, D., Jacobson, A.L., Schneweis, D.J., Whitfield, A.E., 2015. Thrips transmission of tospoviruses. Current Opinion in Virology 15, 80–89. doi:10.1016/j.coviro.2015.08.003. a

Orthotospoviruses (Tospoviridae)

509

Taxonomy and Classification In addition to the genus Tenuivirus (family Phenuiviridae), the genus Orthotospovirus (family Tospoviridae) emcompasses the plantinfecting viruses of the order Bunyavirales. All these plant viruses share many molecular characteristics with animal-infecting viruses classified in this order. The orthotospoviruses, for example, have an enveloped virion containing the viral genome which is distributed among three RNA segments that replicate in a manner consistent with that of other vertebrate-infecting bunyaviruses such as hantaviruses, phleboviruses, orthobunyaviruses, etc. All three RNA segments have highly conserved, complementary termini resulting in a pan-handle structure and genes with similar functions. Differently, the small (S) and medium (M) segments of orthotospoviruses encode two genes in opposite or ambisense polarity. Classification of an orthotospovirus population as a distinct viral species is based upon the similarity of sequence between the nucleocapsid genes of the respective viruses (Fig. 1). This is in contrast to the system used to differentiate animal viruses in other genera which traditionally relied on serological neutralization of infectivity or other biological properties (hemagluttination)

Fig. 1 Phylogeny of the complete S-RNA genome segment sequences of orthotospoviruses belonging to recognized and proposed species within the genus Orthotospovirus in the family Tospoviridae. The midpoint-rooted tree was deduced in MEGA X v. 10.0.5 after alignment in Muscle, using the neighbor-joining method based on the maximum composite likelihood substitution model with uniform rates and 1000 bootstrap replications. The scale bar indicates the number of substitutions per site. Bootstrap support for branches is shown at the junctions of branches where it was 450%. Accession numbers are given in Table 1.

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Orthotospoviruses (Tospoviridae)

mediated by the glycoproteins. Orthotospovirus isolates with greater than 90% nt similarity in the nucleocapsid gene are classified as isolates of the same viral species.

Geographic Distribution TSWV, the type species member of the orthotospoviruses is found worldwide in association with its thrips vector. The wide hostrange of TSWV and its thrips vectors is consistent with the geographic distribution. Other orthotospoviruses have more well-defined distribution (Fig. 1). For example, Groundnut bud necrosis virus (GBNV), Watermelon bud necrosis virus (WBNV), and Watermelon silver mottle virus (WSMoV), that are transmitted by Thrips palmi, a thrips species found only in the subtropics and only known to occur in South-East Asia. Another anomaly is INSV. While INSV is reported to occur around the world, it is almost entirely limited to greenhouse-grown floral crops.

Host-Range and Virus Propagation TSWV has one of the most diverse host-ranges of any plant-infecting virus. This virus infects over 900 plant species belonging to 90 botanical families, both monocots and dicots. In addition, TSWV is reported to be transmitted by at least nine thrips species. Important economic plants susceptible to TSWV include tomato, potato, tobacco, peanut, pepper, lettuce, papaya, and chrysanthemum. Other orthotospoviruses (e.g., IYSV) have much narrower host-ranges and thus the broad host-range of TSWV is not characteristic of the genus. These viruses can be transmitted mechanically or by their thrips vectors. Recently, it has been shown that Soybean vein necrosis virus (SVNV) can be seed-transmitted at a rate of 6% in soybean plants, but this phenomenon has not been observed for any other orthotospovirus. RNA preparations are not infectious. There are no robust cell culture systems for orthotospoviruses. However, plant protoplasts and insect cells have been successfully inoculated.

Virion Properties Orthotospovirus virions are quasispherical, enveloped particles with 80 – 120 nm in diameter (Fig. 2). Two viral coded glycoproteins, GN and GC, are embedded in the viral envelope and form 5–10 nm surface projections. Ribonucleoprotein (RNP) particles consisting of the viral RNA encapsidated by nucleoproteins (ribonucleocapsid), and a small number of polymerase molecules are contained within the envelope. Ribonucleocapsids are pseudocircular due to noncovalent bonding of the complementary RNA termini. Intact

Fig. 2 Orthotospovirus quasispherical virion particles. Left. Electron micrograph of negatively stained particles of Tomato spotted wilt virus (TSWV). The bar represents 100 nm. Courtesy of J. van Lent. Right. The S, M, and L RNA genomic segments are encapsidated by nucleoproteins, which are in association with L protein (polymerase) molecules, and form pan-handle structures due to the complementarity of their 50 - and 30 -ends. The glycoproteins GN and GC are embedded within the viral envelope.

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virions as well as carefully prepared RNPs retrieved from sucrose or CsSO4 gradients are infectious. There are several reports that TSWV and INSV isolates, while infectious, are defective for virion formation.

Genome Properties Orthotospoviruses have a single-stranded, tripartite RNA genome with segments designated as L, M, and S in order of decreasing size (Fig. 3). The termini of each of the RNA segments consist of an eight nt sequence (50 -AGAGCAAU-30 ) that is strictly conserved among all orthotospoviruses. The remaining untranslated region at the termini also has a high degree of complementarity. Base pairing at the termini between the inverted complementary sequences supports a pan-handle structure that most likely serves as a promoter for replication. For TSWV, the L RNA has 8.9 nt and codes for the L or RNA-dependent RNA polymerase (RdRp) protein in the viral complementary (vc) sense (Fig. 3). The M and S RNAs are in ambisense orientation. The M RNA has 4.8 nt and codes in the viral sense for the non-structural protein NSm and for the GN/GC precursor glycoprotein in the vc sense. The S RNA presents 2.9 nt and codes in the viral sense for the nonstructural protein NSs and the nucleocapsid (N) protein in the vc sense (Fig. 3). Intergenic regions (IGRs) of TSWV M and S RNA have variable lengths, are A–U rich, and are the most hypervariable regions of the genome. The 50 - and 30 -ends of the IGRs are conserved, separated by variable sequences, deletions, and insertions. In addition, highly conserved sequences are embedded within the S RNA IGR. A 33 nt duplication occurring in the S RNA IGR of some isolates has been correlated with loss of competitiveness in mixed infections of isolates with and without the duplication. A 31 nt conserved sequence, with significantly higher GC-content compared to the remaining S RNA IGR, has also been found in some TSWV isolates. The IGRs of the M and S segments have high inclination for base pairing thought to be involved in initiation and termination of transcription.

Fig. 3 Orthotospovirus ambisense genome organization (inset in figure) and expression strategy. Positive ( þ ) and negative (  ) sense ORFs are dark and light shaded tubes, respectively. Proteins from the S and M segments are translated from subgenomic mRNAs which are capped with 10–20 nt of host-derived RNA at the 50 -end.

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There is speculation that the termination of transcription is dictated by a conserved nt sequence (CAACUUUGG) in the center of the S and M RNA IGR or that it is due to a secondary structure highly stabilized in the 31 nt region referred to above. Full length molecules of the M and S RNAs are found in infected tissue and purified virions in both the viral and vc sense (approximate ratio of 10:1), consistent with ambisense segments from other viruses. Defective interfering RNAs (DIs) associated with attenuated symptom expression and increased replication rate are also frequently observed. DIs from the L ORF in TSWV infected tissue are the result of and associated with attenuated infectivity. Deletions, frameshift, and nonsense mutations in the GN/GC ORF have been shown to interfere with thrips transmissibility and virion assembly. Frameshift and nonsense mutations with unknown effect have also been identified in the N ORF. The formation of DIs is favored by repeated mechanical passage in certain plant hosts, high inoculum concentration, and low temperatures. Available evidence supports the hypothesis that secondary structure rather than the RNA sequence itself is the primary determinant of the site of deletion. There is also a high frequency of DIs that maintain the original reading frame resulting in translation of truncated proteins whose existence was confirmed in nucleocapsid preparations.

Protein Properties The 330–336 kDa L protein encoded by the L RNA has been identified as a RdRp through sequence homology with other members of the order Bunyavirales and identification of sequence motifs characteristic of polymerases. RdRp activity has been associated with detergent-disrupted TSWV virion preparations. The 34 kDa NSm protein encoded by the M RNA has been shown to induce tubule structures in plant protoplasts and insect (Spodoptera and Trichoplusia) cells. Induction of tubules in plants, ability to change the size exclusion limit of plasmodesmata, an early expression profile and complementation of cell-to-cell and systemic movement in a movement-defective Tobacco mosaic virus vector is evidence that NSm is the TSWV movement protein and that it supports long-distance movement of viral RNAs. In thrips, NSm does not aggregate into tubules, indicating that this protein might not have cell-to-cell movement function in the vector's life cycle. It is also known that NSm specifically interacts with the N protein, the At-4/1 intra- and intercellular trafficking plant protein and binds single-stranded RNA in a sequence-nonspecific manner. An NSm homolog is absent in the vertebrateinfecting viruses of the order Bunyavirales. The 127 kDa GN/GC precursor glycoprotein also coded by the M RNA contains a signal sequence that allows its translation on the endoplasmic reticulum. Proteolytic cleavage of the polyprotein does not require other viral proteins. The molecular mass of GN and GC is 78 kDa and 54 kDa, respectively, for TSWV. Evidence for the involvement of the glycoproteins in thrips transmission is provided by: (1) their interaction with proteins of the thrips vector, (2) their association with the insect midgut during acquisition, (3) the loss of thrips transmissibility of envelope-deficient mutants, (4) the presence of a glycoprotein sequence motif that is characteristic for cellular attachment domains, and (5) the observation that only reassortants with the M RNA of a thrips-transmissible isolate rescue thrips transmissibility. Specifically, GN is involved in virus binding and/or entry in thrips midgut cells, whereas GC is a possible fusion protein playing a significant role in pH-dependent virus entry. It is also believed that these proteins are implicated in virion assembly. The NSs protein encoded on the S RNA is 52 kDa and accumulates to high levels as loose aggregates or paracrystalline arrays of filaments. NSs has RNA silencing suppressor activity, affects symptom expression in TSWV-infected plants, and is not present in the mature virus particle. The N protein, also encoded by the S RNA, ranges in size from 28.1 to 30.9 kDa depending on the virus. This protein encapsidates the viral RNA segments, is highly abundant, and is the predominant protein detected in serological assays. A ‘head-to-tail’ interaction of the nucleoprotein N terminus (aa 1–39) with the C terminus (aa 233–248) results in multimerization.

Replication Replication of viral RNA and assembly of virions occurs in the cytoplasm of both plant and insect cells. Orthotospovirus replication, however, has mainly been described based on plant infection. Upon entry into the plant cell, the virus loses its membrane and releases infectious nucleocapsids into the cytoplasm. In thrips cells, infection by orthotospoviruses is accommodated by binding of the viral surface glycoproteins to a host cell receptor(s) (possibly a 50 kDa and/or a 94 kDa protein). This is followed by release of infectious nucleocapsids into the cytoplasm, through fusion between the viral and thrips membranes possibly initiated by low pH. Depending on the concentration of N protein, the viral RNA is either transcribed or replicated. At low N concentrations, the polymerase transcribes mRNAs that are translated into the virus proteins. Translation of proteins from the S and M ambisense RNAs occurs from subgenomic mRNAs (Fig. 3). The S and M subgenomic mRNAs are capped at the 50 terminus with 10–20 nt of non-viral origin indicating that orthotospoviruses utilize a cap-snatching mechanism to regulate transcription. Leader sequences of Alfalfa mosaic virus have also been detected as caps of TSWV mRNA in mixed infections of the two viruses. The TSWV transcriptase has a reported preference for caps with multiple base complementarity with the viral template. Upon increase of N protein concentration, the polymerase switches its mode to replication with the viral RNA serving as the template. Replicated viral RNAs form RNPs that can presumably associate with the NSm protein for movement through plasmodesmata to adjacent plant cells through tubular structures. Alternatively, RNPs form new virions by associating with the glycoproteins and budding through the Golgi membranes. Virions are initially double-membraned, but soon coalesce and form groups of virions with a single membrane surrounded by another membrane.

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Pathogenicity and Cytopathology Orthotospoviruses are noted for the severity of the diseases they cause in plants. Symptoms are highly variable, depending on the virus, the virus isolate, the host plant, time of the year, and environment, and are thus of little diagnostic value. Chlorosis, necrosis, ring or line patterns, mottling, silvering, and stunting often appear on inoculated and systemically infected leaves. Systemic invasion of plants is frequently nonuniform. Stems and petioles may exhibit necrotic lesions. Observed symptoms often mimic disease and injury caused by other biotic and abiotic stresses. Infection of younger plants results in severe stunting and high mortality rates. TSWV has been shown to affect more severely Datura, Nicotiana, and Physalis plants under a specific temperature regime (daytime, 29 7 21C, nighttime, 24 7 31C). The effect of orthotospovirus infection on thrips has been controversial, due to the confounded effects of plant host, virus, and environment on the insect vector and the genetic variability of thrips and virus populations. TSWV infection of F. occidentalis provided evidence that thrips exhibit an immune response to the virus. Recent work with TSWV-infected F. fusca reared on infected foliage indicated a direct effect of the virus on thrips resulting in reduced fitness. The same study showed that the plant infection status and the TSWV isolate have also an effect on the insect, explaining the variable results obtained from independent studies of virus pathogenicity on the insect vector. Orthotospoviruses induce characteristic cytopathic structures that are host and virus-isolate dependent. In addition to virions, inclusions of viroplasms consisting of the NSs or N protein may be abundant in the cytoplasm. NSs may aggregate in loose bundles (e.g., TSWV) or in highly ordered paracrystalline arrays (e.g., INSV). Excess N protein occurs in granular electron dense masses. NSs and nucleocapsid protein inclusions have been observed in infected plant and insect cells. NSm protein induces tubule structures in plant protoplasts and insect cells.

Transmission and Epidemiology Orthotospoviruses are transmitted from plant to plant by at least sixteen thrips species in the genera Ceratothripoides, Dictyothrips, Frankliniella, Neohydatothrips, Scirtothrips, and Thrips. Among the more common vectors are Frankliniella occidentalis, F. fusca, F. schultzei, F. intonsa, F. bispinosa, Thrips palmi, T. setosus, and T. tabaci. Thrips feed on the cytoplasm of plant cells. The contents of infected cells are ingested and the virus is transported along the lumen of the digestive tract to the midgut, the primary binding and entry site into the insect cells. The brush border of the midgut lumen is the first membrane barrier that the virus encounters. The virus replicates in the midgut and crosses the basement membrane into the visceral muscle cells. The virus subsequently enters the primary salivary glands. It has been hypothesized that the virus moves from the midgut to the salivary glands through infection of ligament-like structures, or when there is direct contact between membranes of the visceral muscles and the primary salivary glands during the larval stages of development. A less plausible hypothesis is that the virus infects the salivary glands after entry and circulation in the hemocoel. Viral inoculum is introduced into plants in the insect saliva coincident with feeding on the plant by adult thrips. The process of successful acquisition occurs only by larvae and acquisition rates decrease as larvae develop, affecting adult vector competency. Vector competency is also determined by the thrips’ feeding preference on a particular host, the uniformity of distribution of virus in plant cells, the rate of virus replication in the midgut, and the extent of virus migration from the midgut to the visceral muscle cells and the salivary glands. In some instances the virus can be acquired by adult thrips and infects midgut cells, but is unable to spread further possibly due to the formation of an age-dependent midgut barrier (e.g., basal lamina). Research has shown the existence of thrips transmitters with detectable levels of virus, nontransmitters with detectable virus, and nontransmitters with no detectable virus, supporting multiple sites for vector specificity between orthotospoviruses and thrips. Evidence for replication of the virus in the insect vector is based on the accumulation of NSs and the visualization of viral inclusions in midgut epithelial cells, muscle cells, and the salivary glands. Although the virus is maintained transtadially throughout the life of the insect, there is no evidence for transovarial transmission. Thus, each generation of thrips must acquire the virus during the larval stages. The primary dispersal of orthotospoviruses is by adult thrips and dissemination of infected somatic tissue in vegetatively propagated crops. These viruses are thought to move long distances in thrips carried by wind currents. They may also survive in commercial agricultural systems in weeds that serve as a bridge between crops. Infected summer weeds (e.g., in NC I. purpurea, I. hederacea, M. verticillata, A. palmeri, C. obtusifolia, R. scabra, Ambrosia artemisiifolia L., Polygonum pensylvanicum L., and Chenopodium album L.) are the principal source for spread of TSWV to winter annual weeds, from which the virus is spread to susceptible crops in spring. Secondary spread within a crop can only occur in crops that concomittantly support virus infection and reproduction of the vector as only the larval stage can acquire the virus for transmission. Transmission through plant seed has only been demonstrated for SVNV but at a low rate. The emergence of these viruses as serious pathogens in crops has been attributed to the increased prevalence of F. occidentalis as an agricultural pest on a worldwide basis.

Genetics and Evolution The knowledge base for genetics and evolution of orthotospoviruses has been derived almost exclusively from TSWV. TSWV has a characteristic ability to adapt to new or resistant hosts and to lose phenotypic characters following repeated passages in experimental

514

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hosts, especially Nicotiana benthamiana. The virus occurs in plants as a heterogeneous mutant population with one or two predominant haplotypes and 9–21 rare haplotypes. Recent research shows that natural TSWV variants evolve in nature through recombination, random genetic drift, and mutation. Intergenomic recombination is important for the genesis and evolution of ancestral TSWV lineages. Genetic drift during thrips transmission and mutation concurrent with virus population growth, shape the genetic architecture of the most recently evolved lineages. The existence of single viral strains as mutant populations and recombination in ancestral viral lineages arm TSWV with a unique genetic reservoir for causing disease and spreading in epidemic proportions in nature. Additional research at the species level supports a distinct TSWV geographical structure and the occurrence of species-wide population expansions. TSWV is also known to use reassortment of genome segments to adapt to resistant hosts under specific laboratory conditions. The determinants of adaptation to resistance in tomato and pepper have been mapped to the M and S RNAs, respectively. Tomato chlorotic spot virus (TCSV) and Groundnut ringspot virus (GRSV) share the same M segment due to host adaptation reasons. Phylogenetic analyses suggest that TCSV putatively incorporated this genomic segment from GRSV upon a reassortment event since the parental genotype of TCSV remains unknown. Little is known about the thrips–orthotospovirus coevolution and the genetic diversity of the thrips vector itself. The altered status of Thrips tabaci as a TSWV vector is one of the very few likely examples of coevolution between orthotospoviruses and their insect vector.

Detection and Diagnosis Orthotospoviruses have certain unique biological properties that are useful for diagnosis. These viruses can be mechanically transmitted by gently rubbing inoculum on plants dusted with carborundum. Nicotiana glutinosa L., Chenopodium quinoa Wild., and garden petunia give characteristic lesions that progress as spots or concentric zones, and sometimes as lethal necrosis. Orthotospoviruses can also be identified by electron microscopy of leaf-dip preparations on thin sections of infected plants. Additional techniques for identification are based on the enzyme-linked immunosorbent assay (ELISA) using polyclonal and monoclonal antibodies, and detection of viral-specific nucleic acids using ribo- and cDNA-probes. The reverse transcription-polymerase chain reaction (RT-PCR) is the most powerful and commonly used technique for detecting small amounts of orthotospovirus RNA. Real time RT-PCR has been successfully used to detect and quantify TSWV in leaf soak and total RNA extracts from infected plants and thrips. RT-PCR with degenerate primers can detect five distinct orthotospovirus species. Tissue selection and sampling strategy are critical factors in TSWV diagnosis and detection regardless of the technique. Because, TSWV titer varies throughout the plant and does not spread uniformly throughout plants that are ‘systemic’ hosts, sampling strategies should be validated in each situation. Nowadays, with the availability of high-throughtput sequencing techniques, also referred to as next-generation sequencing (NGS), viruses have been identified and had their genomes completely sequenced following a fast and unbiased manner. Thus, reports of new viruses include the genetic characterization of entire genomes, which facilitates evolutionary and biological studies.

Prevention and Control Orthotospoviruses cause significant economic losses annually, due to suppressed growth, yield, and reduced quality. These viruses can be partially managed in well-defined cropping systems such as glasshouses by obtaining uninfected plant propagules, implementing a preventative thrips control program in high risk areas, together with constant monitoring of production areas for thrips and infected plants. However, these strategies are costly and require intensive management. Control in field crops is problematic due to the array of external sources of inoculum. Vector control is generally ineffective against the introduction of virus from external sources, due to thrips’ high fecundity, ability to develop insecticide resistance, and to infest many TSWVsusceptible crops. Some measure of control can be achieved using thrips-proof mesh tunnels in the field and reflective mulches. Cultural practices such as utilization of virus-tested planting stock, careful selection of planting dates, removal of cull piles and weeds, rotation with nonsusceptible crops, prevention of planting TSWV-susceptible crops adjacent to each other, reduced in-field cultivation to avoid movement of thrips from infected sources, can reduce the spread of orthotospoviruses. In peanuts, higher plant density, planting from early until late May and application of selected insecticides have reduced the incidence of TSWV. In flue-cured tobacco, early-season treatment with activators of plant defenses and insecticides have also significantly reduced TSWV incidence. Deployment of resistant cultivars has provided benefits in few crops infected by orthotospoviruses. Single-gene resistance is available for orthotospovirus in a limited number of tomato (Sw-5b) and pepper (Tsw) cultivars. Naturally occurring, resistancebreaking isolates of TSWV have been recovered from pepper and tomato cultivars containing their respective resistance genes. Nevertheless, a correct management of Sw-5b-resistant cultivars has shown that this resistance can be durable. The main strategy is to decrease the selection pressure on virus populations by alternating resistant and susceptible cultivars or by planting susceptible tomatoes alongside production fields. The Sw-5b protein is a Nucleotide-Binding site and Leucine-Rich Repeat (NB-LRR) receptor with nucleocytoplasmic distribution that confers a dominant, high-spectrum resistance against isolates of Alstroemeria necrotic streak virus (ANSV), Chrysanthemum stem necrosis virus (CSNV), GRSV, INSV, TCSV, and TSWV. The Sw-5b protein activates resistance by direct recognition of orthotospovirus NSm proteins. Co-dominant markers have been developed for screening Sw-5bcontaining tomato cultivars. Co-dominant cleaved amplified polymorphic sequence (CAPS) marker has also been developed for TSWV marker-assisted selection in pepper. ‘Field’ resistance has been reported for some peanut varieties. Progress has been made in

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understanding the genetic basis of the ability of TSWV to overcome single gene resistance by mapping determinants to specific segments of the TSWV genome and characterizing the selection process. Pathogen-derived resistance utilizing the N and NSm genes has been effective in some greenhouse and field tests; however, isolates have been obtained that overcome nucleocapsid mediated resistance. Best suppression of TSWV epidemics has been achieved with the integrated use of resistant cultivars, chemical, and cultural practices.

Future Perspectives The last few years have been characterized by exciting developments in our understanding of orthotospovirus molecular biology, diversity, evolution, and virus–host relationships. Within this context, several new viruses and thrips vectors have been reported. Advances have also been made in understanding orthotospovirus resistance mechanisms in plants, especially concercing Sw-5 resistance genes, which have brought some insights in the management of orthotospovirus in the field. Unfortunatelly, efficient reverse genetic systems of orthotospoviruses still remain to be developed as well as stable insect cell cultures for in vitro orthotospovirus propagation. The avaiablity of editing DNA tools as CRISPR Cas9 may allow relevant changes in how to control orthotospovirus infection from dismissing key elements in plant hosts necessary for virus infection.

Further Reading Abudurexiti, A., et al., 2019. Taxonomy of the order Bunyavirales: Update 2019. Archives of Virology 164. doi:10.1007/s00705-019-04253-6. de Oliveira, A.S., Boiteux, L.S., Kormelink, R., Resende, R.O., 2019. The Sw-5 gene cluster: Tomato breeding and research toward orthotospovirus disease control. Frontiers in Plant Science 19. doi:10.3389/fpls.2018.01055. Oliver, J.E., Whitfield, A.E., 2016. The genus Tospovirus: Emerging bunyaviruses that threaten food security. Annual Review of Virology, 3, 101–124. doi:10.1146/annurevvirology-100114-055036. Pappu, H.R., Jones, R.A.C., Jain, R.K., 2009. Global status of tospovirus epidemics in diverse cropping systems: Successes achieved and challenges ahead. Virus Research 141 (2), 219–236. doi:10.1016/j.virusres.2009.01.009. Rotenberg, D., Jacobson, A.L., Schneweis, D.J., Whitfield, A.E., 2015. Thrips transmission of tospoviruses. Current Opinion in Virology 15, 80–89. doi:10.1016/j.coviro.2015.08.003. Turina, M., Kormelink, R., Resende, R.O., 2016. Resistance to tospoviruses in vegetable crops: Epidemiological and molecular aspects. Annual Review of Phytopathology 54, 347–371. doi:10.1146/annurev-phyto-080615-095843. Zhu, M., van Grinsven, I.L., Kormelink, R., Tao, X., 2019. Paving the way to tospovirus infection: Multilined Interplays with plant innate immunity. Annual Review of Phytopathology 57. doi:10.1146/annurev-phyto-082718-100309.

Ourmiaviruses (Botourmiaviridae) Gian Paolo Accotto, Institute for Sustainable Plant Protection, National Research Council of Italy, Torino, Italy Cristina Rosa, Pennsylvania State University, University Park, PA, United States r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein HVT Horizontal virus transfer. kb Kilobase

Glossary Monocistronic polypeptide.

RNA that encodes for only one

kDa Kilo dalton MP Movement protein nt Nucleotide(s) ORF Open Reading Frame RdRp RNA-dependent RNA polymerase

Reassortment viruses.

Exchange of genomic segments between

Introduction The only three virus species of this genus were described a long time ago. Since then, only a handful of articles have been published, mainly concerning Ourmia melon virus (OuMV), the type species. The other two members of the genus are Cassava virus C (CsVC), reported only once in a mixed infection in cassava from Malawi, and Epirus cherry virus (EpCV), found in a single cherry tree. This tree has since died. OuMV has been detected on several occasions in melons in north-western Iran. In general, OuMV has been found in mixed infections with other cucurbit viruses, so the extent of economic losses is unknown. Ourmiaviruses are potentially damaging, as they have been experimentally transmitted to a rather wide range of dicot species, including important crops in which they produce symptoms of mosaic, mottle, and necrosis.

Taxonomy, Phylogeny and Evolution The genus is not assigned to a specific family. The members have the same structural features but their coat proteins (CPs) share less than 70% aa identity. Phylogenetic analysis of the ourmiavirus RdRp places them in a clade originated from ourmia-like viruses of invertebrates, infecting members of the subphylum Chelicerata and Crustacea, and of the superphylum Lophotrochozoa. This clade was shown to be sister to the one containing ourmia-like mycoviruses. Such a phylogeny suggests that ourmia-like viruses infecting fungi, crustacea, and plants could belong to distinct genera in the same virus family. These genera are distinct from the Narnavirus genus. Since most of the ourmia-like viruses have been found in invertebrates, it has been hypothesized that the plant ourmiaviruses evolved by horizontal virus transfer (HVT) from herbivorous invertebrates. In the new host the acquisition of a movement protein (MP) from a plant virus was necessary. Actually, analyzes of the ourmiavirus MP indicates similarities with the MPs of viruses in the family Tombusviridae. The CP of ourmiaviruses shows distant affinities with the CPs of sobemo-, tombusluteo- and nodaviruses. As of 2017, three species in the genus Ourmiavirus were recognized by ICTV: Cassava virus C (CsVC), Epirus Cherry Virus (EpCV), and Ourmia melon virus (OuMV).

Virion Structure The virions are non-enveloped and made of subunits of a single CP (Fig. 1). They have cylindrical bodies of discrete lengths, 18 nm in diameter and capped by conical (probably hemi-icosahedral) ends. The bodies of the particles are composed by a series of disks, commonly two or three for particles with a length of 30–37 nm, more rarely four or six disks for particles with a length of 45.5–62 nm. Particles with five disks and with a hypothetical length of 54 nm have never been observed. The buoyant density in CsCl of the particles of all sizes is 1.375 g cm–3.

Genome Organization Viruses in this genus have a tripartite single stranded positive sense RNA genome. Each genomic segment is monocistronic. The lengths of the genomic segments are approximately 2800 nt, 1100 nt and 970 nt (Fig. 2).

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Fig. 1 Ourmiavirus virion morphology: (A, B, C) Negative contrast electron micrographs (uranyl acetate) of purified particles of Ourmia melon virus. The bar represents 100 nm. (D, E) Features of the two commonest particle types, enhanced by photographic superimposition. (from ICTV 9th report, page 1178, Fig. 2, with permission from Elsevier).

Fig. 2 Diagram summarizing genome organization of members of the genus Ourmiavirus. Green bars: genomic RNAs. Yellow bars: ORFs.

Properties and Functions of Gene Products Each genomic segment contains a single ORF. RNA1 encodes for a putative RdRp of ca. 860 aa and 97 kDa, RNA2 for a putative MP of ca. 290 aa and 32 kDa, and RNA3 for a putative CP of ca. 200 aa and 23 kDa (Fig. 2). The functions of the RdRp, MP and CP have been experimentally demonstrated for OuMV, the better characterized virus in the genus. OuMV RdRp is necessary for virus replication and contains the polymerase motif GDD. OuMV MP belongs to the “30K” Superfamily of movement proteins. It is a symptoms determinant, is localized at plasmodesmata of infected plant cells and forms tubular structures inside the cytoplasm and between adjacent cells. The OuMV CP is required for assembly of the virions and systemic movement. It is localized in the nucleolus when expressed as fusion with GFP and without the expression of RNA1 and RNA2. The KR-rich region at the N-terminus of the CP has been shown to induce tissue tropism, to be a host determinant and to interfere with the plant silencing pathway. Virion assembly requires an active replication and RNA3, but not RNA2.

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Replication and Propagation OuMV replication occurs in the cytoplasm of infected plant cells. OuMV MP is required for cell-to-cell movement. While effective systemic infection requires the expression of the OuMV CP, RNA1 and 2 can move up to some vascular tissue exit sites without CP expression.

Transmission, Host Range Ourmiaviruses are plant infecting viruses. OuMV was isolated from melon, CsVc from cassava and EpCV from cherry. Extensive surveys have not been conducted to reveal other natural hosts of these viruses in the geographic areas where the 3 species were originally isolated, aside from one survey in Iran on cucurbits. Experimentally, OuMV and other ourmiaviruses have been mechanically transmitted to dicots in ca. 15 plant families, including Solanaceae, Brassicaceae and Cucurbitaceae. Transmission studies found that these viruses could not be transmitted by several aphid species, the whiteflies Trialeurodes vaporariorum and Bemisia tabaci, and the mite Tetranychus urticae. Transmission experiments via soil or water were also unsuccessful. Seedtransmission rates in Nicotiana benthamiana and N. megalosiphon are 1%–2%.

Epidemiology and Control Not much is known about the epidemiology of these viruses. A survey conducted on melon and squash in 16 regions in Guilan Province (Iran) in 2005 and 2006 revealed that OuMV was present in 4 regions. Based on this survey, OuMV was the most prevalent virus detected in melon (Cucumis melo L.) and was frequently found in squash (Cucurbita sp.). Based on the limited information available, standard agronomic practices aimed at reducing the economic damage caused by ourmiaviruses could include screening for virus presence in seeds and other plant propagative material, exclusion and eradication in areas where the viruses are not present, and rouging of symptomatic plants.

Virus–Host Relationships Plants systemically infected with ourmiaviruses show an array of symptoms, such as vein chlorosis, rasp-leaf, necrotic spots and rings, stunting, yellowing and necrosis.

Diagnosis Virions and tubular structures of OuMV have been used to produce species specific polyclonal antibodies for ELISA and western blotting, but commercial diagnostic kits are not available. PCR primers used in published experiments can be adopted for diagnosis via molecular techniques.

Concluding Remarks Due to the little effort dedicated so far to this genus, in spite of its very special characteristics, many questions remain unsolved. Evidence from metagenomics suggests that the origin of this genus could have been genomic reassortment of RNA viruses infecting hosts outside the plant kingdom, accompanied by HVT. More studies are necessary to elucidate the missing information about this interesting genus.

Further Reading Accotto, G.P., Boccardo, G., Riccioni, L., Barba, M., 1997. Comparison of some molecular properties of Ourmia melon and Epirus cherry viruses, two representatives of a proposed new virus group. Journal of Plant Pathology 78, 87–91. Accotto, G.P., Milne, R.G., 2008. Ourmiavirus. In: Mahy, B.W.J., Van Regenmortel, M.H.V. (Eds.), Encyclopedia of Virology. London, UK: Elsevier, pp. 500–501. Aiton, M.M., Lennon, A.M., Roberts, I.M., Harrison, B.D., 1988. Two new cassava viruses from Africa. In: Proceedings of the 5th International Congress of Plant Pathology. p. 43. Kyoto, Japan. Avgelis, A., Barba, M., Rumbos, I., 1989. Epirus cherry virus, an unusual virus isolated from cherry with rasp-leaf symptoms in Greece. Journal of Phytopathology 126, 51–58. Crivelli, G., Ciuffo, M., Genre, A., Masenga, V., Turina, M., 2011. Reverse genetic analysis of Ourmiaviruses reveals the nucleolar localization of the coat protein in Nicotiana benthamiana and unusual requirements for virion formation. Journal of Virology 85, 5091–5104. Dolja, V.V., Koonin, E.V., 2018. Metagenomics reshapes the concepts of RNA virus evolution by revealing extensive horizontal virus transfer. Virus Research 244, 36–52. Gholamalizadeh, R., Vahdat, A., Hossein-Nia, S.V., Elahinia, A., Bananej, K., 2008. Occurrence of Ourmia melon virus in the Guilan province of Northern Iran. Plant Disease 92 (7), 1135.

Ourmiaviruses (Botourmiaviridae)

Lisa, V., Milne, R.G., Accotto, G.P., et al., 1988. Ourmia melon virus, A virus from Iran with novel properties. Annals of Applied Biology 112, 291–302. Margaria, P., Anderson, C.T., Turina, M., Rosa, C., 2016. Identification of Ourmiavirus 30K movement protein amino acid residues involved in symptomatology, viral movement, subcellular localization and tubule formation. Molecular Plant Pathology 17, 1063–1079. Rastgou, M., Habibi, M.K., Izadpanah, K., et al., 2009. Molecular characterization of the plant virus genus Ourmiavirus and evidence of inter-kingdom reassortment of viral genome segments as its possible route of origin. Journal of General Virology 90, 2525–2535. Rossi, M., Genre, A., Turina, M., 2014. Genetic dissection of a putative nucleolar localization signal in the coat protein of Ourmia melon virus. Archives of Virology 159, 1187–1192. Rossi, M., Vallino, M., Abba, S., et al., 2015. The importance of the KR-rich region of the coat protein of Ourmia melon virus for host specificity, tissue tropism, and interference with antiviral defense. Molecular Plant-Microbe Interactions 28, 30–41. Shi, M., Lin, X.D., Tian, J.H., et al., 2016. Redefining the invertebrate RNA virosphere. Nature 540, 539–543. Turina, M., Hillman, B.I., Izadpanah, K., et al., 2017. ICTV virus taxonomy profile: Ourmiavirus. Journal of General Virology 98, 129–130.

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Papaya Ringspot Virus (Potyviridae) Cécile Desbiez and Hervé Lecoq, Plant Pathology Unit, INRAE – French National Research Institute for Agriculture, Food and Environment, Montfavet, France r 2021 Elsevier Ltd. All rights reserved. This is an update of D. Gonsalves, J.Y. Suzuki, S. Tripathi, S.A. Ferreira, Papaya Ringspot Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00731-7.

Nomenclature

HC-Pro

aa Amino acid CP Capsid protein DAS-ELISA Double antibody sandwich enzyme-linked immunosorbent assay

Glossary Gene silencing Plant defense mechanism based on the detection and fragmentation of double-stranded RNAs, including the replicative forms of RNA viruses. Mild-strain cross-protection A plant systemically infected by a mild virus strain will not develop additional symptoms when inoculated by a severe strain of the same virus. Molecular clock The molecular clock hypothesis considers that mutations between homologous nucleotide or amino-

Helper component-protease protein of potyviruses kDa kilodalton (1 Da ¼ 1 g/mol) LAMP Loop-mediated isothermal amplification nt Nucleotide

acid sequences accumulate in a clock-like fashion, i.e., at a constant rate with time. Molecular changes along the branches of a phylogenetic tree can thus be used to estimate the time of divergence from a common ancestor. Pathogen-derived resistance Engineered resistance to pathogens through the synthesis in transgenic plants of pathogen proteins or nucleic acids.

History and Taxonomy The term Papaya ringspot virus (PRSV) was first used in the 1940s to describe a viral disease of papaya, initially observed in Hawaii. The name was used primarily to describe the ringspots that appeared on fruits from infected plants. Early investigations showed that the virus was transmitted by several species of aphids in a non-persistent manner. That is, the aphid vector could acquire the virus in a short period of time while feeding on infected plants and likewise transmit the virus in a span of few seconds to less than a minute during subsequent feeding. In the same decade researchers from India and other places like Puerto Rico reported the occurrence of an aphid-transmitted disease of papaya; based on the symptoms on the leaves, it was identified as Papaya mosaic virus. Work in the 1980s showed that the aphid-transmitted papaya mosaic virus and PRSV were the same virus, and the name of PRSV was adopted. PRSV is a member of the family Potyviridae, a large and arguably the most economically important group of plant viruses. Today, the term Papaya mosaic virus (PapMV) is reserved for a virus that is not aphid transmitted, belonging to the genus Potexvirus in the family Alphaflexiviridae, and causes the papaya mosaic disease which is seldom observed and not important commercially, even if mixed infections between PRSV and PapMV can increase symptoms and virus accumulation. PRSV should not be confused with another potyvirus, Papaya leaf distortion mosaic virus (PLDMV), which occurs in Okinawa and other parts of Asia, such as Taiwan. This virus causes very similar symptoms as PRSV on papaya and cucurbits but is serologically unrelated and its sequence shares only 55%–59% similarity to that of PRSV. In the 1930s a mosaic disease was described in cucurbits in the USA and named Watermelon mosaic virus (WMV). Later on, biological and serological studies showed that the disease might be caused by two distinct viral entities called WMV-1 and WMV-2. More recently, serological and molecular characterization showed that PRSV and WMV-1 are virtually identical. Based on their close relationship, a single name was adopted to unify both viruses into one species. The name PRSV was chosen due to its being named before WMV-1. To clarify host range, ‘P’ (PRSV-P) or ‘P type’ is used to designate virus infecting papaya and cucurbits, while ‘W’ (PRSV-W) or ‘W type’ refers to virus infecting cucurbits only.

Classification and Relationship With Other Potyviruses: The “PRSV Cluster” PRSV was classified as a member of the genus Potyvirus in the family Potyviridae based on particle morphology, aphid transmissibility, serological relationships, ability to induce pinwheel cytoplasmic inclusions in host cells, genome organization and nucleotide (nt) sequences. Several other potyviruses infect papaya or cucurbits. The main other potyvirus infecting papaya is PLDMV, that shares no particular relationship with PRSV. More than 10 potyviruses infect cucurbits. Some of them are not closely

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related to PRSV based on serological and molecular data: Zucchini yellow mosaic virus (ZYMV), Watermelon mosaic virus (WMV), Melon vein banding mosaic virus (MVBMV), Telfairia mosaic virus (TeMV), Cucurbita vein-banding virus (CVBV), Turnip mosaic virus (TuMV), Clover yellow vein virus (ClYVV) and Bean yellow mosaic virus (BYMV). However, in the 1990s, several viral isolates, mostly infecting cucurbits, were found to share biological and serological relationships with PRSV, forming what was called a “PRSV cluster” of related members that could not always be unambiguously distinguished. When sequences became available, the different criteria of the International Committee for the Taxonomy of Viruses (ICTV) could be applied, mainly: (1) less than 76% nt identity (80% aa identity) in the coat protein (CP)-coding region or in the whole genome; (2) difference in polyprotein cleavage site; (3) host range and key host reactions; (4) different inclusion body morphology; (5) serological relatedness. Partial or complete sequences have revealed the presence of at least seven species besides PRSV in the “PRSV cluster”: Moroccan watermelon mosaic virus (MWMV), Algerian watermelon mosaic virus (AWMV), Zucchini yellow fleck virus (ZYFV), Zucchini tigre mosaic virus (ZTMV), Zucchini shoestring virus (ZSTV), Sudan watermelon mosaic virus (SuWMV) and Wild melon vein banding virus (WMVBV). Partial sequences suggest that more species are present in Asia, although they remain to be fully characterized (Fig. 1). Most members of the PRSV cluster were found in a limited geographic area and/or presented a low prevalence in the countries where they were present. AWMV, MWMV, SuWMV, WMVBV, ZYFV and ZSTV were found only in Africa and/or the Mediterranean Basin, indicating that this region is most probably their center of origin. ZTMV was found in Europe, Asia, America and Africa but at a rather low prevalence, although it has probably been frequently mis-diagnosed as PRSV because of the frequent cross-reactions observed in serological tests.

Host Range, Symptomatology, and Geographic Distribution The systemic host range of PRSV is confined to plants in the families Caricaceae and Cucurbitaceae, with the primary economically important host being papaya and a range of cucurbits such as squash, cucumber, watermelon, and melons. PRSV-P isolates cause generally severe symptoms in papaya and mild symptoms in cucurbits, while PRSV-W do not infect papaya and cause severe symptoms in cucurbits. In addition, some PRSV isolates cause local lesions on plants of the family Chenopodiaceae such as Chenopodium quinoa and C. amaranticolor. In papaya, PRSV infection is characterized by mosaic and chlorosis symptoms on leaves, water-soaked streaks on the petiole, and deformation of leaves that can result in shoestring-like symptoms that resemble mite damage (Fig. 2). The virus can cause deformation and ringspot symptoms on the fruit, hence the name PRSV. PRSV-P is present in all papaya-growing areas. PRSV-W infects cucurbits worldwide, but is more prevalent in tropical, subtropical and Mediterranean conditions than in temperate ones. This may be related to the difficulty for the virus to overwinter in areas where cucurbits are not present all year round. On cucurbits, PRSV-W causes severe mosaic and chlorosis on leaves, which are also often deformed with shoestring-type symptoms. The fruits are often deformed and bumpy. Some PRSV-W isolates induce systemic necrotic spots and top necrosis in some melon cultivars.

Transmission and Epidemiology PRSV is transmitted in a non-persistent manner by more than 24 aphid species among which Aphis gossypii, Myzus persicae and A. craccivora are the most efficient. PRSV is acquired and transmitted during very short probes, what makes its spread within a field generally rapid. However PRSV persists only few hours on the stylets: therefore long-distance spread by aphids is limited. Virus sources are generally neighboring infected crops, but for PRSV-W several wild cucurbits (Melothria pendula, Momordica sp.) were shown efficient virus sources in tropical and sub-tropical regions. There is no report of PRSV-P or -W seed transmission. Longdistance spread may occur through the movement of infected material (seedlings, fruits). Indeed, aphids were shown to be able to acquire and transmit efficiently PRSV-W from commercial infected melon fruits.

General Properties and Genome of PRSV Like other potyviruses, PRSV has flexuous filamentous particles about 760–800 nm  12 nm encapsidating a single-stranded RNA of about 10,326 nt in length. The genome is translated as a single polyprotein that is further cleaved autocatalytically in 10 functional proteins (P1, HC-Pro, P3, 6K1, CI, 6K2, NIa-VPg, NIa-Pro, NIb and CP), with a smaller 11th protein P3N-PIPO expressed through translational frameshift. Virus particles consist of 94.5% protein and 5.5% nucleic acid by weight. PRSV has a single coat protein (CP) of about 36 kDa. The density of the virion in purified preparations is 1.32 g cm3 in cesium chloride. A rather interesting feature of the PRSV is that sequence analysis predicts two potential cleavage sites at the N-terminus of the CP. One of the sites (VFHQ/SKNF) predicts a CP of 33 kDa and a polymerase (nuclear inclusion b protein, NIb) of 537 aa about 20 aa larger than those of other potyviruses. The second predicted cleavage site (VYHE/SRGTD) generates a CP of 36 kDa and a NIb of 517 aa. There is no firm evidence to suggest that only one cleavage site is used. If both sites are used in polyprotein processing, one would expect heterogeneous products. This may explain why the analysis of purified CP preparations that are stored frequently shows the major B36 kDa form and CPs that are 2–5 kDa smaller.

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PRSV-W_Japan PRSV-P_Taiwan PRSV-W_Korea PRSV-W_Myanmar Asia PRSV-W_China 95 PRSV-P_Philippines PRSV-W_Thailand PRSV-P_Brazil PRSV-W_Venezuela 100 PRSV-P_Hawaii PRSV 76 PRSV-W_France America, PRSV-W_Australia Europe, PRSV-P_Mexico Australia 100 PRSV-P_Florida PRSV-P_India PRSV-W_India Indian «PRSV cluster» PRSV-W_Myanmar Peninsula PRSV-P_India PRSV-P_India 100 98 PRSV-P_India New? Unknown_Myanmar-Mp10 Unknown_Myanmar_Msg New? ZTMV_Myanmar ZTMV_Pakistan 100 ZTMV 77 ZTMV_Guadeloupe WMVBV_Sudan ZYFV-Italy AWMV-Algeria ZSTV-South Africa SuWMV_Sudan MWMV_Tunisia PLDMV WLMV Outgroups WMV ZYMV 86 79

99

88 98

88 87

99

0.05

Fig. 1 Taxonomy, based on coat protein nucleotide sequences, of potyviruses in the “PRSV cluster” including PRSV, six acknowledged species and two putative new species. PRSV-P isolates of PRSV infect papaya whereas PRSV-W isolates infect cucurbits. Branch lengths indicate the molecular divergence of viruses (the scale bar represents 0.05 mutations per residue). Figures at some nodes represent bootstrap values (in %), indicating the robustness of each node. Only values above 75% are indicated.

Sequence Diversity and Evolution; Molecular Clock Analyzes Knowledge of the sequence diversity among isolates of a virus has great implications in developing an effective virus disease management program and in understanding the origin and biology of the virus. Numerous PRSV-P and -W sequences from virus isolated from different parts of the world have been reported. Nucleotide and aa sequences among PRSV isolates differ by as much as 21% and 13% respectively. Heterogeneity in CP length ranging from 837 to 870 nt has been noted, resulting in CPs of between 279 and 290 aa. The first 50 or so aa of the N-terminal region of the PRSV CP gene are highly variable and all differences in CP length were confined to this region. The N-terminal part of PRSV CP is characterized by the presence of several –up to 8-Glutamic acid (E) and Lysine (K) repeats, and the majority of the size differences were due to differences in the number of EK repeats. Phylogenetic analyzes based on HC-Pro and CP nt sequences indicated the presence of two main clades of PRSV isolates, one from Asia and one from Africa, Australia and America. Isolates from India presented a basal location in the phylogenetic tree, suggesting that the Indian Peninsula could correspond to the center of origin of the virus. Two major radiations appear to have arisen from different branches of the Indian population. The earliest included all the isolates in east and south east Asia (China, Indonesia, Malaysia, Philippines, Taiwan, Thailand and Vietnam). The other includes all the isolates from the Americas and Europe with small separate sub-clusters of isolates from Hawaii, Taiwan, and Australia. A “molecular clock” approach, relying on the hypothesis that mutations accumulate in viral populations at an almost constant yearly rate, was applied to PRSV. Based on the molecular divergence between the Indian basal group and the “American” lineage, PRSV introduction to America presumably happened about 300 years

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Fig. 2 Symptoms of PRSV-P on papaya (left) and of PRSV-W on zucchini (right, top) and on melon (right, bottom).

ago, maybe through Europe. Isolates from the “Asian” cluster have been observed recently in Europe and the Caribbean, showing that population exchanges are currently taking place, probably through the worldwide trade of plant materials. There was no relationship between the biotype (W or P) and the viral phylogeny, suggesting that jumps between W and P happened several times during the evolution of the virus. Evidences from various sources suggest that PRSV is primarily a pathogen of cucurbits, and that PRSV-P originated from PRSV-W. This suggestion is supported by the diversity in cucurbit-infecting potyviruses that are phylogenetically related to PRSV. In Australia, the outbreak of PRSV-P about 25 years ago most probably arose from the population of PRSV-W already present in the country.

Molecular Determinants for Biological Properties The easy jump between PRSV-W and PRSV-P pathotypes suggested a simple genetic determinism related to a few mutations. Studies utilizing infectious transcripts from recombinant viruses followed by bioassays of the transcripts have demonstrated that the region of PRSV genome encoding the NIa-Protease was critical for papaya infection. In particular, two aa at positions 2309 (K - D) and 2487 (I - V) of PRSV are significantly different between papaya-infecting type P and non-papaya-infecting type W. Further point mutational studies in these sites indicated that the K-4 D mutation was responsible for conferring the ability to infect papaya. As noted earlier, PRSV causes local lesions on C. amaranticolor and C. quinoa. The severe strain PRSV HA strain from Hawaii causes local lesions on C. quinoa but a mild nitrous acid mutant of it, PRSV HA 5–1, does not. Recombinant infectious viruses were generated by exchanging genome parts between PRSV HA and PRSV HA 5–1. The study revealed that the pathogenicity-related region is present between nt positions 950 and 3261 of the PRSV HA genome and mutations in the P1 and HC-Pro resulted in the attenuation of PRSV HA symptoms and the loss of ability to produce local lesions on C. quinoa. The HC-Pro coding region of PRSV is thus the major determinant factor for local lesion formation. Contrary to other potyviruses where the N-terminal part of the HC-Pro is dispensable for infectivity, this region was shown to be required in PRSV-W for systemic infection of zucchini. Virus–host interaction studies based on recombinant analyzes between severe and mild strains of PRSV indicated that the HC-Pro gene plays an important role in viral pathogenicity and virulence and acts as a suppressor of the gene-silencing defense mechanism in the papaya host plant. In addition, the comparative reaction of recombinant PRSV with chimeric CP gene sequences showed that heterologous sequences and their position in the CP gene influences their pathogenicity on PRSV-resistant transgenic papaya. As with other potyviruses the HC-Pro/virions interaction is essential for aphid transmission of PRSV.

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Diagnostics As for many other potyviruses, serological methods (DAS-ELISA) are widely used for PRSV diagnostics. Polyclonal antibodies against purified virus particles have been obtained, and commercial DAS-ELISA kits are available. However serological crossreactivity between viruses in the “PRSV cluster” can be observed, potentially resulting in misdiagnosis. Since many PRSV CP or complete sequences are now available, specific primers have been designed for the molecular diagnostic of PRSV using reverse transcription and polymerase chain reaction (RT-PCR), alone or in multiplex with other papaya- or cucurbit-infecting viruses. More recently, isothermal amplification techniques (LAMP) have been developed for PRSV.

Virus Control Cultural Practices: Prophylactic Measures Prophylactic measures are not specific to one virus. They generally tend to prevent or limit the contact of viruliferous aphids with susceptible crops. Insecticide treatments are usually inefficient for non-persistently transmitted viruses. The control or management of PRSV on papaya has been approached through practices such as quarantine, eradication, avoidance by planting crops in areas isolated from the virus, and continual rogueing of infected plants. In Taiwan a successful but costly method used was the cultivation of papaya under insect proof nets. For cucurbits, the same precautions can be used, particularly through careful weeding near plantings and avoiding overlapping crops in the same areas to reduce virus and aphid sources. Besides, plastic mulches are frequently used, as they limit weeds but they also have a repelling effect on aphids and contribute to delay virus spread, at least at the beginning of the crop before the plants grow and cover most of the mulch surface. Different covers (plastics, unwoven) can also be used to prevent winged aphids from reaching low-growing cucurbit plants, but they must be removed to allow pollination.

Cross-Protection Cross-protection can be defined as the use of a mild strain of virus to infect plants that are subsequently protected against economic damage caused by a severe strain of the same virus. This practice has been used successfully for many years to minimize damage by Citrus tristeza virus in Brazil, for example. In the early 1980s, a mild strain of PRSV (described above as PRSV HA 5–1) was developed through nitrous acid treatment of a severe strain, PRSV HA isolated from Oahu Island in Hawaii. This mild strain was tested on Oahu Island and showed good protection against damage by severe strains but produced symptoms that were very obvious on certain cultivars, especially in the winter months. This prominent symptom induction on certain cultivars and the logistics of mild strain buildup and inoculation of plants, among others, were factors that caused it not to be consistently used on the island of Oahu. Interestingly, the mild strain was used extensively for several years in Taiwan, but it did not afford sufficient protection against the severe strains from Taiwan and thus its use was abandoned after several years.

Breeding for Resistance The use of resistant cultivars, when they are available, is the easiest and cheapest way for farmers to control plant viruses. However resistance must be present in the germplasm of the species of interest, and introducing the resistance in commercial cultivars is a time-consuming process. It is important, before starting the breeding process, to check if the resistance is not broken by preexisting virus trains, or if the virus can easily evolve to overcome it. Research to develop PRSV-resistant papaya varieties started in the 1970s. Since resistance to PRSV has not been identified in Carica papaya, researchers have used tolerant germplasm in an attempt to develop papaya cultivars with acceptable PRSV tolerance and horticultural characteristics. However, tolerance to PRSV is apparently governed by a family of genes that is inherited quantitatively, which makes it technically difficult to develop cultivars of acceptable horticultural quality. Furthermore, the tolerant lines do become infected with PRSV, although fruit production continues still at a lower level. Indeed, in Thailand, the Philippines, and Taiwan, a number of tolerant lines have been developed and are used. However, tolerance could not be introduced in all types of papayas and infected tolerant plant may act as virus sources to contaminate susceptible crops. Resistance to PRSV has been described in the germplasm of different cucurbits, including melon, cucumber, squash and watermelon. The resistance was characterized in most cases as monogenic, either dominant or recessive. Interestingly in melon, a dominant resistant gene was found efficient for PRSV-W as well as for several other virus species of the “PRSV cluster”. Resistance is now present in several commercial cultivars of melon, cucumber and squash.

Pathogen-Derived Resistance for Controlling PRSV: The Hawaii Case The control or management of PRSV in Hawaii has been approached through cultural practices, cross-protection with mild virus strains, tolerance, and resistance using the “pathogen-derived resistance” technology. Ultimately, the most successful has been the “pathogen-derived resistance” approach.

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PRSV was first detected in the 1940s on Oahu Island where Hawaii’s papaya industry was located at that time. Efforts to control the virus on Oahu largely consisted of state officials and farmers continually monitoring for infected plants and rogueing them, especially in areas where the virus was not prevalent. However, by the late 1950s, PRSV was causing extensive damage, which caused the papaya industry to relocate to Puna on the island of Hawaii. The relocation of the industry to Puna was timely and effective because Puna had an abundance of land that was suitable to grow Kapoho, a cultivar of excellent quality that adapted to the volcanic soil base there, allowing excellent drainage, had high rainfall and yet lots of sunshine, and the land there could be bought or leased at reasonable prices. By the 1970s, the Kapoho papaya grown in Puna accounted for 95% of the state’s papaya production, making papaya the second most important fruit crop behind pineapple. Despite strict quarantine on movement of papaya seedlings between islands, PRSV was discovered in Puna in May 1992 (Fig. 3(a)) and the Hawaiian papaya industry would be forever changed. By 1995, a third of the papaya growing area was completely infected and much of the rest of Puna had widespread infection (Fig. 3(b)). By 1998, the production of papaya in Puna had dropped to 27 million pounds of papaya from 52 million pounds in 1992 when PRSV was discovered in Puna. In retrospect, the efforts of quarantine, monitoring and rogueing of infected plants in Hilo, and suppression efforts of PRSV in Puna all played key roles in helping Hawaii’s papaya industry, because it gave researchers time to develop control measures for PRSV.

Transgenic Resistance In the late-1980s, an exciting development on Tobacco mosaic virus (TMV) provided a rationale that resistance to plant viruses could be developed by expressing the viral CP gene in a transgenic plant. This approach was called CP-mediated protection, and, at about the same time, a report introduced the concept of “parasite-derived resistance”. The report on transgenic resistance to TMV set off a flurry of work in many laboratories to determine if this approach could be used for developing resistance to other plant viruses. Likewise, work was initiated in 1985 to use this approach for developing PRSV-resistant transgenic papaya for Hawaii. Key requirements for successful development and commercialization of transgenic virus-resistant plants are the isolation and engineering of the gene of interest, vectors for mobilization into and expression of the gene in the host, transformation and subsequent regeneration of the host cells into plants, effective and timely screening of transformants, testing of transformants, and the ability to deregulate and commercialize the product. The CP gene of the mild strain of PRSV was chosen as the “resistance” gene because it had been recently cloned and it was of the PRSV P type. The commercial cultivars Kapoho, Sunrise, and Sunset were chosen for transformation. In 1991, tests of the R0 lines identified a transgenic Sunset that expressed the CP gene of PRSV HA 5–1, and showed resistance to PRSV from Hawaii. A field trial of R0 plants was started in April 1992 on as the island of Oahu, and a month later PRSV was discovered at Puna in May 1992, as discussed above. The Oahu field trial showed that R0 plants of line 55–1 were resistant, and line 55–1 was further developed to obtain the cultivar “SunUp” which is line 55–1 that has the CP gene in a homozygous state, and “Rainbow” which is an F1 hybrid of SunUp and the nontransgenic “Kapoho”. SunUp is red-fleshed and Rainbow is yellow-fleshed. In 1995, SunUp and Rainbow were tested in a subsequent field trial in Puna and showed excellent resistance (Fig. 3(c)). Due to its yellow flesh and good shipping qualities, Rainbow was especially preferred by the growers. Line 55–1 was deregulated by the US government and commercialized in May 1998. The deregulation also applied to plants that were derived from line 55–1. The timely commercialization of the transgenic papaya in 1998 was crucial since PRSV had decreased papaya production in Puna by 50% that year compared to 1992 production levels. The transgenic Rainbow papaya was quickly adopted by growers and recovery of papaya production in Hawaii was underway (Fig. 3(d)). The transgenic papaya is sold throughout Hawaii (Fig. 3(e)) and the mainland USA, and to Canada where it was deregulated in 2003. Japan, the main importer of Hawaii-grown papaya, approved transgenic papayas for import in 2011. However, several challenges remain: coexistence, exportation, deregulation, and the adoption of transgenic papaya in other countries that suffer from PRSV. The use of transgenic papayas in Hawaii has now been going on for more than 20 years and the resistance is still highly durable. One limitation of larger-scale deployment of the transgenic papayas lies in the fact that the efficiency of resistance is related to the percentage of identity between the transgene and the challenging PRSV isolate: the transgenic papayas from Hawaii were found susceptible to PRSV isolates from Hainan. The success of transgenic papaya in Hawaii has encouraged other papaya-cultivating states and countries to develop transgenic papayas resistant to their local PRSV strains. Transgenic varieties have thus been developed in America (Florida, Brazil, Venezuela, Jamaica), Asia (China, Indonesia, Malaysia, the Philippines, Taiwan) and Australia. Depending on the construct and the country, the plants were more or less resistant, ranging from complete and durable resistances to easily broken ones. Resistance-breaking could be related to the emergence of new strains that presented a reduced sequence identity with the transgene. In Taiwan, a transgenic papaya initially conferred resistance to PRSV, but the plants became infected with the emerging PLDMV. Transgenic papayas containing partial CP genes of PRSV and PLDMV conferred resistance to both viruses, but the resistance was later broken by a recombinant “super-strain” of PRSV. This shows that viral diversity and rapid evolution remain challenging for a durable control. Pathogen-derived resistance appears as the most effective way to control papaya ringspot disease, but its use should be associated with a good knowledge of the local viral diversity and the use of cultural practices for an integrated management of PRSV. Although the transgenic approach was not as successful to control PRSV-W in cucurbits, it is used in the USA to control ZYMV, WMV and Cucumber mosaic virus in squash. Interestingly a commercial squash cultivar possessing transgenic resistance to these three viruses and conventional resistance to PRSV-W shows a good protection against these four viruses.

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Fig. 3 (a) Healthy Puna papaya in 1992. (b) Severely infected papaya orchards in Puna in 1994. (c) Field trial of transgenic papaya in Puna in 1994: PRSV-resistant transgenic papaya (center) surrounded by PRSV-infected non-transgenic papaya (d) Commercial planting of transgenic papaya one year after releasing seeds of PRSV-resistant transgenic papaya. (e) Transgenic papaya commonly sold in supermarkets. (f) Risk of growing non-transgenic papaya still exists (photograph from 2005). Foreground is PRSV-infected non-transgenic papaya that are cut, and background shows healthy PRSV-resistant transgenic papaya.

Summary Remarks PRSV has been thoroughly characterized and is a typical member of the family Potyviridae, arguably the largest and economically most important plant virus group. The complete genome sequence has been elucidated and infectious transcripts have provided a means to determine the genetic determinants of some important biological functions such as host range and virulence. Furthermore, pathogen-derived resistance has been used to control PRSV-P in Hawaii through the use of virus-resistant transgenic papaya. The transgenic virus-resistant papaya provides a potential means to test the global acceptance of genetically modified (GM) organisms while presenting a plausible approach to control a disease affecting papaya worldwide. In order to avoid resistance-breaking, GM papayas presenting resistance to local strains should be included in an integrated pest management of PRSV.

Cross Check With Other Chapters Potyviruses (Potyviridae); Plum Pox Virus (Potyviridae); Potato Virus Y (Potyviridae); Watermelon Mosaic Virus and Zucchini Yellow Mosaic Virus (Potyviridae); Plant Resistance to Viruses: Engineered Resistance; Plant virus disease: Economic aspects, Vector transmission of plant viruses.

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See also: Plant Resistance to Viruses: Engineered Resistance. Plant Viral Diseases: Economic Implications. Plum Pox Virus (Potyviridae). Potato Virus Y (Potyviridae). Potyviruses (Potyviridae). Vector Transmission of Plant Viruses. Watermelon Mosaic Virus and Zucchini Yellow Mosaic Virus (Potyviridae)

Further Reading Chen, K.C., Chiang, C.H., Raja, J.A., et al., 2008. A single amino acid of NIaPro of papaya ringspot virus determines host specificity for infection of papaya. Molecular Plant Microbe Interactions 21, 1046–1057. Desbiez, C., Wipf-Scheibel, C., Millot, P., et al., 2017. New species in the papaya ringspot virus cluster: Insights into the evolution of the PRSV lineage. Virus Research 241, 88–94. Fuchs, M., Gonsalves, D., 2007. Safety of virus-resistant transgenic plants two decades after their introductions: Lessons from realistic field risk assessments studies. Annual Review of Phytopathology 45, 173–202. Gibbs, A.J., Ohshima, K., Phillips, M.J., Gibbs, M.J., 2008. The prehistory of potyviruses: Their initial radiation was during the dawn of agriculture. PLoS One 3, 1–11. Gonsalves, D., 1998. Control of papaya ringspot virus in papaya: A case study. Annual Review of Phytopathology 36, 415–437. Gonsalves, D.A., Vegas, A., Prasartsee, V., et al., 2006. Developing papaya to control papaya ringspot virus by transgenic resistance, intergeneric hybridization, and tolerance breeding. Plant Breeding Reviews 26, 35–78. Hamim, I., Borth, W.B., Marquez, J., et al., 2018. Transgene-mediated resistance to Papaya ringspot virus: Challenges and solutions. Phytoparasitica 46, 1–18. Olarte-Castillo, X.A., Fermin, G., Tabima, J., et al., 2011. Phylogeography and molecular epidemiology of papaya ringspot virus. Virus Research 159, 132–140. Quiot-Douine, L., Lecoq, H., Quiot, J.-B., Pitrat, M., Labonne, G., 1990. Serological and biological variability of virus isolates related to strains of papaya ringspot virus. Phytopathology 80, 256–263. Tripathi, S., Suzuki, J.Y., Ferreira, S.A., Gonsalves, D., 2008. Papaya ringspot virus-P: Characteristics, pathogenicity, sequence variability and control. Molecular Plant Pathology 9, 269–280. Yeh, S.D., Jan, F.J., Chiang, C.H., et al., 1992. Complete nucleotide sequence and genetic organization of papaya ringspot virus RNA. Journal of General Virology 73, 2531–2541.

Pecluviruses (Virgaviridae) Hema Masarapu and Pothur Sreenivasulu, Sri Venkateswara University, Tirupati, India Philippe Delfosse, University of Luxembourg, Esch-sur-Alzette, Luxembourg Claude Bragard and Anne Legreve, University of Louvain, Louvain-la-Neuve, Belgium DVR Reddy, International Crops Research Institute for the Semi-Arid Tropics, Hyderabad, India r 2021 Elsevier Ltd. All rights reserved. This is an update of D.V.R. Reddy, C. Bragard, P. Sreenivasulu, P. Delfosse, Pecluvirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00542-2.

Nomenclature

NCR Noncoding regions nt Nucleotide(s) OAS Origin of assembly ORF Open reading frame pAbs Polyclonal antibodies PCR Polymerase chain reaction RdRp RNA-dependent RNA polymerase RISC RNA-induced silencing complex RT-qPCR Reverse transcription quantitative PCR satRNA satellite RNA UTR Untranslated region VIGS Virus-induced gene silencing VLPs Virus-like particles VPg Viral protein genome-linked VRC Virus replication complex vRNA virion RNA

aa Amino acid(s) AGO Argonaute 1 CITE Cap-independent translation enhancer Co-Pro Protease-cofactor CP Coat protein or capsid protein CRP Cysteine-rich protein ELISA Enzyme-linked immunosorbent assay ER Endoplasmic reticulum HC-Pro Helper component-proteinase IRES Internal ribosome entry site kb Kilobase kDa Kilodalton LAMP Loop-mediated isothermal amplification mAbs monoclonal antibodies MP Movement protein

Glossary Forma specialis An informal rank in a classification, allowed by the International Code of Nomenclature for Algae, Fungi, and Plants, that is applied to a parasite (most frequently a fungus), which is characterized from a physiological standpoint (e.g., by the ability to infect a specific host), but scarcely or not at all from a morphological standpoint. Fortuitous hosts of obligate biotrophic Plasmodiophorida Host on which some primary infections evolve into resting spores without going through the zoosporangial stage. Hetero-encapsidation Partial or full coating of the genome of one virus with the coat protein of a differing

virus. Also termed trans-capsidation or heterologous encapsidation. Leaky scanning Mechanism by which the ribosomes fail to initiate translation at the first AUG start codon and scan downstream for the next AUG codon. Obligate endoparasite A parasitic organism that cannot complete its life cycle without exploiting a suitable living host, and that lives within that host. Post-transcriptional gene silencing Mechanism for sequence-specific RNA degradation in plants. Triple gene block A specialized evolutionarily conserved gene module involved in the cell-to-cell and long-distance movement of plant viruses.

History Peanut (groundnut, Arachis hypogaea) clump disease, characterized by severe stunting and clumping, was first described from India in 1927 and subsequently from West Africa in 1931. The causal agent in West Africa as well as in India was identified as a virus, Peanut clump virus (PCV) and Indian peanut clump virus (IPCV), respectively. Annual losses to the peanut crop due to PCV and IPCV were estimated to exceed US$38 million in the late 1990s. Pecluviruses also cause diseases on several dicotyledonous and monocotyledonous crops.

Taxonomy and Classification The genus Pecluvirus has only two species, isolates from West Africa were grouped under the species Peanut clump virus (Peanut clump virus; PCV) and those from the Indian subcontinent were grouped under the species Indian peanut clump virus (Indian peanut clump

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virus; IPCV). The two virus species were initially assigned to the genus Furovirus mainly based on virion morphology and vector transmission by Polymyxa graminis (Rhizaria, Phytomyxea, Plasmodiophorida). Subsequently, based on molecular features, the ICTV in 1997 assigned PCV and IPCV to the newly established genus Pecluvirus (siglem from Peanut clump virus). They differ in their geographical distribution, host range, antigenic properties and genomic sequences. Later, the genera Tobamovirus, Tobravirus, Hordeivirus, Furovirus, Pecluvirus, Pomovirus, and Goravirus were assigned to the newly established family Virgaviridae (from the Latin Virga, meaning rod).

Geographic and Seasonal Distribution PCV occurs in Burkina Faso, Benin, Chad, Congo, Côte d0 Ivoire (Ivory Coast), Niger, Mali, Gabon, Senegal, and Sudan, and IPCV in the Indian states of Punjab, Andhra Pradesh, Telangana, Tamil Nadu, Rajasthan, Haryana, and Gujarat, and in Pakistan (Sind and Punjab provinces). IPCV and PCV infections are noticed mainly in the peanut crops cultivated during the rainy season (July–November in India and April–October in West Africa). Nevertheless, post-rainy season crops can also be infected.

Host Range and Symptoms PCV infects peanut, cowpea, sugarcane, maize, sorghum, pearl millet, and finger millet, whereas IPCV infects peanut, cowpea (Vigna unguiculata), pigeonpea (Cajanus cajan), chili (Capsicum annuum), wheat (Triticum aestivum), barley (Hordeum vulgare), maize (Zea mays), sorghum (Sorghum bicolor), pearl millet (Pennisetum glaucum), foxtail millet (Setaria italica), and finger millet (Eleusine coracana). Cynodon dactylon, Cyperus rotundus, Oldenlandia aspera, and Vigna subterranea (bambara groundnut) that can play a significant role in both virus survival and dissemination.

Fig. 1 Symptoms induced by Peanut clump virus (PCV), in West Africa. (a) field-infected peanut plants occurring in patches of various sizes, intercropped with pearl millet in Maradi, Niger Republique, (b) peanut leaflets showing initial symptoms of chlorosis and mild mosaic, (c) chlorosis along the veins resembling oak leaf, (d) PCV-infected sugarcane plants exhibiting red leaf mottle in Burkina Faso, (e) IPCV and PCV vector, Polymyxa graminis, in transversal section of sorghum roots, stained with lactophenol blue showing zoosporangia, (f) sporosori, bars represent 25 mm. Photographs courtesy of P. Delfosse.

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Fig. 2 Symptoms induced by Indian peanut clump virus (IPCV) on various hosts. (a) on peanut plants showing severe stunting, occurring in patches in the field, and an apparently uninfected plant, (b) young peanut leaflets with initial symptoms of chlorosis and mosaic symptoms, (c) early infected pigeonpea plants showing severe stunting with an apparently uninfected plant, (d) inset showing chlorotic symptoms on young pigeonpea leaflets (left) and by its side leaflets from healthy plant, (e) infected wheat plants occurring in patches, (f) early infected wheat plants showing dark green leaves and chlorotic streaks, (g) late infected wheat plants showing panicle formation. Seed derived from them can transmit IPCV up to 1%, (h) early infected barley plants showing stunting and severe yellowing. Photographs courtesy of P. Delfosse.

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Diseased peanut plants are conspicuous under field conditions because of their dark green appearance, stunting, and occurrence in patches (Fig. 1(a) and Fig. 2(a)). The disease incidence is high in fields where peanut crops are grown in rotation with wheat, barley, and pearl millet. Early-infected peanut plants often did not yield pods and late-infected crops showed decreased crop yields up to 60%. In peanut, the symptoms incited by PCV are essentially similar to those induced by IPCV. In West Africa, peanut clump disease symptoms are often confused with green rosette because of the presence of stunted dark green plants occurring in patches in the fields. The peanut clump disease occurs in the same areas year after year with marginal increase in the periphery of the patches in the successive peanut crops. Initial symptoms on young leaflets of peanut are mottling, mosaic and chlorotic veins or the 'oak leaf symptom' (Fig. 1(b) and (c), Fig. 2(b)). When such leaflets mature, they turn dark green with or without faint mottling. Late infected plants may not show conspicuous symptoms and clumping is restricted to a few branches in some plants. PCV causes the red leaf mottle disease in sugarcane (Saccharum officinarum) (Fig. 1(d)). Pigeonpea plants infected with Hyderabad isolate of IPCV (IPCV-H) showing stunting are presented in Fig. 2(c). Fig. 2(d) shows chlorosis on infected pigeonpea leaflets, left, and uninfected leaflets, right. Wheat plants, up to three weeks old, infected by IPCV-H show rosette symptoms similar to those caused by the Soil-borne wheat mosaic virus (SBWMV) and grain yield losses up to 58% are recorded. Wheat cultivar RR-21 infected with IPCV-H showing stunting is presented in Fig. 2(e). Early infected wheat plants show chlorotic streaks (Fig. 2(f)) and will produce panicles (Fig. 2(g)). Seed from these plants can transmit IPCV up to 1%. Barley plants infected with IPCV-H are stunted and bushy with chlorotic or necrotic leaves (Fig. 2(h)) and the majority of these plants die. The plants that reached maturity produced poorly developed spikes. IPCV-H also infected pearl millet, finger millet, foxtail or Italian millet, and sorghum plants. In maize IPCV-H caused aerial biomass losses up to 33% and grain loss up to 36%. Various isolates of PCV and IPCV are readily detected in the cells of roots, stems, and leaves of systemically infected hosts. In wheat cells, PCV particles are found in the cytoplasm, near the nucleus or along the plasmalemma, and arranged in angledlayer aggregates. In experimental host range studies by sap inoculation, PCV and IPCV infected a wide range of both dicotyledonous and monocotyledonous plants. For IPCV, Nicotiana clevelandii x N. glutinosa hybrid and Phaseolus vulgaris (cv. Top crop) are suitable for virus propagation and as assay/diagnostic hosts, respectively. Chenopodium amaranticolor and C. quinoa are local lesions hosts. IPCV isolates collected from clump diseased peanut crops from different locations in India differ slightly in their host ranges. Canavalia ensiformis and N. clevelandii x N. glutinosa hybrid are found to be useful for differentiating the isolates of IPCV. In the case of PCV, C. amaranticolor, N. benthamiana, N. glutinosa, P. vulgaris, and Triticum aestivum are of diagnostic value. N. benthamiana and P. vulgaris are suitable for virus propagation. The symptoms induced in C. amaranticolor by various PCV isolates collected from Burkina Faso, Senegal, and Niger are shown to differ markedly.

Transmission Seed and Sap Transmission Pecluviruses can be transmitted by sap inoculation and through peanut seed. The transmission rate through the peanut seed depends on the cultivar, the mode of infection (up to 24% for plants infected through soil and up to 50% for plants infected through seed) and the age at which the plants are infected. IPCV is transmissible up to rates of o2% through the seeds of pearl millet, finger millet, foxtail millet, wheat, and maize. Pearl millet CV GGP-16 did not transmit IPCV through seed.

Soil-Borne Vector Transmission Both PCV and IPCV are naturally vectored by soil-borne protists, P. graminis (Fig. 1(e) and (f)). Two formae speciales of this species are associated with the transmission of PCV and IPCV: P. graminis. f. sp. subtropicalis and P. graminis f. sp. tropicalis. These obligate root endoparasites complete their entire life cycle, including sporangial and sporogenic (resting spores) phases (Fig. 1(e) and (f); Fig. 3 right panel), only in monocotyledonous hosts in the family Poaceae (P. graminis tropicalis on sorghum, pearl millet, and maize but rarely on wheat and barley and P. graminis subtropicalis on sorghum, pearl millet, wheat, and barley). Nonetheless, they can infect and transmit viruses to dicotyledonous hosts, including peanut, which are considered to be fortuitous hosts of the vector. The presence of P. graminis in infested soils can be identified by growing bait plants such as pearl millet, sorghum, or wheat on infested soils or by PCR. In vitro transmission studies under controlled conditions is complicated because of the obligate parasitic nature of P. graminis and consequent difficulties in maintaining the cultures of both the virus and the vector in the same hosts. Nevertheless, the acquisition and the transmission of PCV by the vector is demonstrated using certain hosts such as sugarcane (CV CP-89–2377) and Sorghum arundinaceum supporting abundant zoospore production and virus infection. Presence of P. graminis in roots can also be quantified by reverse transcription quantitative PCR (RT-qPCR). The P39 viral protein, encoded by RNA2 and expressed by leaky scanning of P23, is involved in the transmission by P. graminis based on its similarity with P75 protein encoded by RNA2 of Beet necrotic yellow vein virus (BNYVV).

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Fig. 3 Electron micrographs of the Indian Peanut clump virus (IPCV; Pecluvirus, Virgaviridae) and its vector Polymyxa graminis. (left) Virus particles with two predominant lengths of 249 and 184 nm are decorated with homologous antiserum for the Hyderabad isolate, bar represents 100 nm. (right) Transversal section of a sorghum root heavily infested by P. graminis showing sporosori, bar represents 25 mm. Photographs courtesy of P. Delfosse and Larry Littlefield.

Virion Morphology The virions of IPCV (Fig. 3 left panel) and PCV are non-enveloped rigid straight rod shaped, 24 nm (IPCV) and 21 nm (PCV) in diameter with two predominant lengths of c. 250 and c. 180 nm. They have helical symmetry with a pitch of 2.6 nm and contain a single coat protein (CP) of 23 kDa.

Physical Properties Virions sediment as two major components with S20w of 183S and 224S. Buoyant density in CsCl is 1.32 g cm3 and isoelectric point is pH 6.45. Thermal inactivation of virus infectivity occurs at 641C. Virions are stable in frozen leaves.

Molecular Properties Genome Structure and Expression Strategies The genome of pecluviruses is bipartite, with two positive-sense, linear single-stranded RNAs (ssRNAs) (4% in weight). RNA1 is c. 5900 nt long and RNA2 is c. 4500 nt long. The PCV RNAs show little sequence identity with those of IPCV RNAs except for the 30 terminal 273 nt, which are conserved among the two RNAs. Both the RNAs have methylated cap structure (to be confirmed) at their 50 ends. The 30 ends of the RNAs have tRNA-like structure and are not polyadenylated. Both RNAs are flanked by untranslated regions (50 - or 30 UTR) or noncoding regions (50 - or 30 NCR). The genome organization and expression of the PCV is shown in the Fig. 4.

Coding Sequences RNA1 contains two open reading frames (ORFs) that encode two proteins involved in viral RNA replication (P131 and P191, Fig. 4; Table 1). The proximal ORF encodes a 131 kDa polypeptide and P191 is a C-terminally extended form of P131 produced by translational readthrough of the UGA termination codon. The P15 ORF is downstream of the P191 ORF and separated from it by a noncoding region of about 60 nt. The polypeptides of 131 and 191 kDa contain NTP-binding methyl transferase, helicase and RNA-dependent RNA polymerase (RdRp) domains and are involved in the putative replication complex (Table 1). The 15 kDa polypeptide, a cysteine-rich protein (CRP) is translated from a sub-genomic RNA (Fig. 4). It is a suppressor of posttranscriptional gene silencing (PTGS). The P15 possesses four C-terminal proximal heptad repeats that can generate a coiled-coil interaction and is targeted to peroxisomes via a C-terminal Ser-Lys-Leu (SKL) motif. Such a motif is conserved among pecluviruses from both Africa and India. It has been demonstrated that a coiled-coil motif is necessary for the anti-PTGS activity of P15, but the peroxisomal localization signal is not, although it is required for efficient intercellular movement of the virus. P15 resembles CRPs of Barley stripe mosaic virus (BSMV), Poa semilatent virus (PSLV), and SBWMV. These proteins have been suggested to act as regulatory factor during virus replication as well as for long-distance movement and contribute to the virulence factor. RNA1 can replicate in the absence of RNA2 in protoplasts of tobacco BY-2 cells. However, both RNAs are required for infection and systemic invasion of plants. Experiments using enhanced 50 green fluorescent protein and 50 -bromouridine 50 -triphosphate labels have suggested that PCV replication complexes co-localize with endoplasmic reticulum (ER) bodies accumulating around

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Fig. 4 Genome organization of PCV (genus Pecluvirus, family Virgaviridae). The tRNA structure motifs at the 30 termini of the RNAs are represented by a dark square. ORFs are indicated by rectangles and a suppressible termination codon (UGA) by an arrow. Mtr, methyl transferase; Hel, helicase; RdRp, RNA-dependent RNA polymerase; CRP, cysteine-rich protein; CP, coat protein; TGB, triple gene block. Modified from Adams, M.J., Adkins, S., Bragard, C., et al., 2017. ICTV virus taxonomy profile: Virgaviridae. Journal of General Virology 98, 1999–2000.

Table 1

Pecluvirus open reading frames (ORFs), polypeptides and their functions

Genomic RNA

ORF

Mr of polypeptide (kDa)

Function

RNA1

P131 P191 P15

131 191 15

Methyltransferase, helicase, replicase

P23 P39 P51 P14 P17

23 39 51 14 17

RNA2

Suppressor of PTGS Coat Protein Putative vector transmission factor Virus movement (triple gene block)

the nucleus during infection. P15 does not act directly at sites of viral replication but intervenes indirectly to control viral accumulation levels. RNA2 contains five ORFs (Fig. 4). The 50 proximal ORF (23 kDa) encodes the CP, and the adjacent ORF (ORF2) encodes a 39 kDa polyprotein which is expressed by leaky scanning mechanism in vitro and thought to be involved in the transmission of PCV by its vector. ORF2 starts 1 nt upstream of the first residue of UGA stop codon of the CP cistron. Further, downstream and separated by 135 nt intergenic region, is a triple gene block (TGB) that codes for proteins of 51, 14, and 17 kDa that are presumed to be involved in the cell to cell movement.

50 and 30 Noncoding Sequences The 50 and 30 NCRs of RNA1 of pecluviruses are about 130 and 300 nt in length, respectively (Fig. 4), whereas the RNA2 has more diverse NCR of 390–500 nt at 50 end and c. 300 nt at 30 end. Within the 30 NCR of RNA2, approximately 100 terminal nt are conserved among pecluvirus RNAs sequenced so far. The NCRs differ in size among isolates from the different serotypes. The 30 NCR of pecluviruses, as in the case of furoviruses, forms a t-RNA-like structure (TLS) that aids in the replication of both RNAs.

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Fig. 5 (a) Phylogenetic tree constructed using coat protein (CP-P23) amino acid sequences from eight PCV/IPCV isolates. (b) Phylogenetic tree constructed using 18 sequences of the 30 UTR of RNA1 and RNA2 of PCV/IPCV isolates. The evolutionary distances were computed using Maximum Composite Likelihood method by Tamura, K., Dudley, J., Nei, M. Kumar, S., 2007. MEGA4: Molecular Evolutionary Genetics Analysis (MEGA) software version 4.0. Molecular Biology and Evolution 24, 1596-1599. GenBank Accession numbers are given in parentheses.

Sequence Comparisons and Phylogeny Only one complete RNA1 genome sequence of each of PCV (X78602) and IPCV (X99149) are available, while five and three complete RNA2 sequences of PCV (L07269, AF447401, AF447400, AF447399, AF447398), and IPCV (AF447397, AF239729, AF447396), respectively are available in the database. PCV and IPCV exhibited considerable differences in homology depending on the part of the genome sequence compared. The complete nt sequence of RNA1 of PCV is 78% similar to that of IPCV. The RNA1-encoded polypeptides (P191, P131, and P15) of PCV and IPCV share identities ranging from 75% (P15) to 95% (read-through portion of P191) and show significant similarities with furoviruses (e.g., 56% identity with polymerase of SBWMV). Comparison of the P15 gene shows a close relationship between IPCV isolates and a relatively high diversity among PCV isolates. The full-length sequence comparisons between RNA2 of PCV and IPCV isolates revealed a high degree of variability in size (between 58%–79%) and the proteins encoded by RNA2 are 39%–89% identical between species. Among the five ORFs of RNA2, ORF4 encoding P14 of TGB is highly conserved (90%–98%), whereas ORF2 encoded P39 is less conserved (25%–60%). Phylogenetic comparisons, based on complete RNA2 sequences, showed that the eight isolates could be grouped into three distinct clusters with no geographical distinction between PCV and IPCV isolates. Phylogenetic analysis of individual ORFs revealed an overall similarity with that obtained from complete RNA2 sequences, but the relative positions of individual isolates varied within each cluster. These studies indicate that there is substantial divergence among the RNA20 s of pecluviruses and suggest that different polypeptides have evolved differently, possibly due to different selection pressures. In the most members of the family Virgaviridae, CP is conserved among the isolates, but in the case of PCV and IPCV isolates, coat proteins are highly diverse (aa sequence identities between 37% and 89%) and have c. 30% similarity with the CP of BSMV (genus Hordeivirus). A conserved motif ‘F-E-X6-W0 is present near the CP C-terminus of all three IPCV serotypes and PCV, as in the CPs of other rod-shaped viruses [Tobacco mosaic virus (TMV) and Tobacco rattle virus (TRV)]. Phylogenetic analysis of CP of eight PCV and IPCV isolates (PCV-B, PCV-M, PCV-N, PCV-Ni, PCV-S, IPCV-D, IPCV-H, IPCV-L) revealed that the PCV isolates grouped into two major clusters and IPCV into one cluster (Fig. 5(a)). The TGB proteins resemble those of Potato mop top virus (PMTV, genus Pomovirus). Comparison of 30 end nt sequences revealed that they are highly conserved (88% for RNA1 and 85% for RNA2), thus offering a potential target for broad-spectrum detection by RT-PCR. Conservation of 30 UTR is verified by the alignment of eighteen 30 UTR (124 nt long) sequences of RNA1 and RNA2 (collected from NCBI database) of PCV/IPCV isolates from different geographical regions and a phylogenetic tree has been constructed using the neighbor-joining method (Fig. 5(b)). The isolates were grouped into one Indian and two African clusters. Indian cluster appeared to be intermediate between the African clusters. Though there is high percentage of identity between 30 UTR of RNA1 and RNA2 of PCV and IPCV isolates from different geographical regions, there is no evidence of reassortment between isolates. Putative recombination events and their frequencies in the complete genome of 168 accessions extracted from the International databases representing several members of six genera Viz. Furovirus, Pecluvirus, Comovirus, Tobravirus, Hardeivirus, and Tobamovirus indicate that exchange of heredity material occurred between non-homologous sequences like those of IPCV and PCV.

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Assembly of Virus Particles Sequences required for assembly of PCV virions have been established by testing the ability of encapsidation of deletion mutants of RNA1 and RNA2 in vivo. A putative origin of assembly (OAS) is mapped in the 50 proximal part of the P15 gene of RNA1. Two sequence positions, 50 proximal CP gene and the other in the p14 gene near the RNA2 30 terminus that can drive encapsidation of RNA2, have been identified. No obvious sequence similarities between different assembly initiation sequences are noted. The possible localization of an OAS in the CP gene of IPCV is realized as a result of the formation of virus-like particles (VLPs) in Escherichia coli and N. benthamiana, confirming the results observed for PCV. The monomer VLP size corresponded to the length of the encapsidated CP gene transcript RNA. Using immunocapture RT-PCR (IC-RT-PCR), such VLPs have been demonstrated to contain RNA encoding IPCV-H CP. Transgenic N. benthamiana expressing IPCV-H CP gene inoculated with the serologically distinct IPCV-L serotype accumulated virus particles that contained both types of CP, indicating that hetero-encapsidation occurs in transgenic plants.

Detection and Diagnosis Pecluvirus diseased peanut plants occur in patches, mostly restricted to sandy/sandy-loam soils. The diseased peanut plants are stunted and dark green in color with or without chlorotic symptoms on leaves. Pecluvirus affected cereal and millet plants and grass weeds are difficult to recognize because they show mild or no visible symptoms. Diseases caused by pecluviruses reappear nearly in the same location year after year.

Immunoassays Several immunological tests utilizing polyclonal (pAbs) and monoclonal antibodies (mAbs) have been used for the detection and identification of pecluvirus isolates. This approach revealed a diversity of serotypes and serogroups in Africa and India. The pAbs of PCV did not react with IPCV, BSMV, TMV, BNYVV, SBWMV, and PMTV. Wide serological diversity exists among PCV and IPCV isolates. Thus, serological tests have limitations to detect more than one isolate from a single antibody source. Antisera to different isolates of IPCV facilitated the grouping of isolates into three serotypes, IPCV-H (Hyderabad), IPCV-D (Durgapura), and IPCV-L (Ludhiana). All IPCV isolates are serologically distinct from PCV isolates and vice-versa. Utilizing a panel of mAbs raised against a PCV isolate and four different ELISA formats, several PCV isolates are grouped into five serotypes. Surprisingly, one of the mAbs reacted with IPCV-D in triple antibody sandwich ELISA. ISEM and alkaline phosphate or penicillinase-based ELISA formats have been applied for detection of both PCV and IPCV in various investigations.

Molecular Assays It is essential to employ diagnostic tests that are highly sensitive and broadly specific for the detection of pecluviruses in disease surveys, to eliminate virus-contaminated sources in quarantine, and to devise strategies for disease control. A broad-spectrum RT-PCR protocol, based on the conserved sequences of the 30 end region of genomic and sub-genomic RNAs of several virus isolates, has been developed to detect all the currently known IPCV and PCV isolates in various plant samples originated from different countries. Primers are also designed for quantitation of pecluvirus isolates using a TaqMan-based real time RT-PCR (RT-qPCR). RT-PCR is compared with penicillinase-based DAS-ELISA for the detection of pecluvirus isolates in crop and weed plants. The virus is detected in 70% of suspected plants by RT-PCR as compared to 24% detected by DAS-ELISA. The RT-PCR enabled the detection of virus in the plants that did not exhibit overt symptoms. The detection and quantification of PCV and IPCV isolates (IPCV-L, IPCV-H, IPCV-D, PCV-B, PCV-M, PCV-N, PCV-S) is performed by RT-qPCR. These PCR-based techniques are very useful for detecting PCV and IPCV especially in non-symptomatic hosts. RT-qPCR is recognized as an indispensable tool for tracing resistant cultivars and for the maintenance of pecluvirus-free germplasm. Immunocapture RT-PCR has also been used to demonstrate the presence of RNA in PCV VLPs.

Vector Separation of P. graminis resting spores from sorghum root material facilitated pAbs production. In DAC-ELISA, they detected one sporosorus per well of the ELISA plate. In spiked root samples, the procedure detected one sporosorus per mg of dried sorghum roots. The majority of P. graminis isolates from Europe, North America, and India reacted strongly with pAbs. However, P. graminis isolates from Rajasthan state (North India), from Pakistan, and an isolate from Senegal (West Africa) reacted weakly with pAbs. The DAC-ELISA procedure is applied to detect various stages in the life cycle of P. graminis and to detect infection that occurs under natural and controlled environments. Resting spores are detected in root sections by using fluorescein 5-isothiocyanate - labeled pAbs. Probes specific for nuclear ribosomal DNA (rDNA) detected the presence of P. graminis f. sp. subtropicalis, tropicalis, and temperata in roots by RT-PCR. P. graminis f. sp. tropicalis is quantified in plants inoculated with viruliferous zoospores using a specific RT-qPCR. The ribotypes of P. graminis f. sp temperata and P. graminis f. sp. tepida are identified by restriction fragment length polymorphism analysis of PCR amplified rDNA, nested and multiplex PCR.

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Fig. 6 Disease development cycle of Pecluviruses showing ways of acquisition, multiplication, and transmission by Polymyxa graminis through a variety of host plants. Modified from Delfosse P., 2000. Epidemiology and Management of the Indian peanut clump virus. Louvain-la-Neuve, Belgium: Université catholique de Louvain, PhD Thesis. Dieryck, B., Otto, G., Doucet, D., et al., 2009. Seed, soil and vegetative transmission contribute to the spread of pecluviruses in West Africa and the Indian sub-continent. Virus Research 141, 184–189. Seed transmission data from Reddy, A.S., Hobbs, H.A., Delfosse, P., Murthy, A.K., Reddy, D.V.R., 1998. Seed transmission of Indian peanut clump virus (IPCV) in peanut and millets. Plant Disease 82, 343–346.

Epidemiology and Disease Cycle The study of epidemiology and formulation of cost-effective disease control practices depend on the precise and sensitive detection of PCV and IPCV and their vector in plants, seeds, and soil. Peanut clump disease is largely confined to sandy soils and loams. Detection is required to identify infested fields, to select virus-free seed lots and vegetative propagules, to identify alternate hosts, and to assess the resistance of peanut, sugarcane and cereal and millet crops breeding lines.

Disease Development Cycle The presence of viruliferous P. graminis vector, adequate soil moisture to facilitate the mobility of the vector zoospores from plantto-plant in the sandy and sandy-loam fields, the optimum temperatures conducive to vector transmission and planting with seeds of virus-infected cereal crops contribute to primary spread. Presence of cereal hosts, ambient temperatures near 301C and adequate moisture are vital for secondary spread. The relative contributions of favorable hosts for both the pecluvirus and its vector in the disease cycle are summarized (Fig. 6). Dicotyledonous hosts limit the multiplication of the plasmodiophorid vector and hence considered them as fortuitous hosts that may not contribute to the perpetuation of the virus inoculum. Indeed, virus-infected peanut roots could not transmit or establish the disease. Monocotyledonous plants such as maize, sorghum, and pearl millet are ‘preferred’ hosts of the vector and contribute to the build-up of vector population potential in the soil. Seed of millets, maize and wheat, rhizomatous grasses (e.g., C. dactylon) and sugarcane vegetative propagules are likely to contribute to the disease establishment in new areas by supporting

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the multiplication of the virus and vector. Soils from disease infested areas containing roots/rhizomes harbor resting spores of the vector. Dispersal of such infested soils by wind and displacement during tillering and plowing can contribute to the establishment of the disease in disease-free areas. During the rainy season, prolonged dry spells followed by supplementary irrigation in peanut fields may also contribute to the dispersal of resting spores and disease spread.

Disease Control Research during the past four decades has led to the accumulation of considerable information on IPCV and PCV. It is now apparent that pecluviruses are not the viruses of peanut but should be regarded as viruses of cereals that opportunistically infect peanut. The difficulty in controlling such viruses is that they are both seed (in peanut, finger millet, foxtail millet, pearl millet, maize, wheat) and soil transmitted (through thick walled resting spores of the vector embedded in roots/rhizomes) and can also be propagated through sugarcane stem cuttings or the rhizomes of grass weeds. Continuous monocropping with fortuitous hosts such as peanut, bambara groundnut and pigeon pea will contribute to a reduction in plasmodiophorid population in soil. Seeds from infected cereal hosts should be avoided for planting. Thus, understanding precisely the sources of virus and vector and transmission mechanisms for each type of host plant in virus endemic agro-ecosystems using sensitive and specific diagnostic tools may led to formulation of effective disease control practices. Initial experiments with IPCV in India showed that application of soil biocides, e.g., DD (a mixture of 1,3-dichloropropene and 1,2-dichloropropane), fumigant nematicides that also have fungicidal action and soil solarization are effective on a limited scale in decreasing the disease incidence in peanut fields. Clump affected peanut plants always occur in patches and easy to avoid. Additionally, under field conditions early infected, which is the case, hardly produce any seed.

Cultural Practices The following crop cultural practices are suggested for reducing disease incidence:

• • • •

Early planting of peanut, before the onset of monsoons under judicious irrigation. Planting with trap crops such as pearl millet at a high density, plowing of two-week-old seedlings into the soil followed by planting with peanut seed. Avoid crop rotation and intercropping with cereal and millet crops. Adopt continuous cropping with dicotyledonous crops (peanut, bambara groundnut, cowpea, pigeon pea) for at least three successive growing seasons.

Host Plant Genetic Resistance No resistance to IPCV is found in over 9000 cultivated Arachis germplasm. Unfortunately attempts to incorporate resistance to IPCV by genetic engineering, deploying CP gene, have not produced successful results. However, cultivation with cereal hosts that did not show any seed transmission may help in limiting the disease establishment at hitherto disease-free locations.

Future Perspectives At present the geographical distribution of pecluviruses is restricted to the Indian subcontinent and West Africa. Locations where peanut crops are grown in rotation or as mixed crops with cereal crops on sandy soils can be regarded as targets for establishment of clump disease. Hence measures must be taken to avoid planting with seeds from cereal crops resourced from clump infested fields. They also should not be used towards germplasm exchange because they pose a quarantine risk of introduction of pecluviruses into new geographical regions. Mapping of locations with peanut crops grown in sandy soils in rotation with cereal crops and peanut crops intercropped with cereals, along with the temperature range required for infection, should help in determining the potential for occurrence of pecluviruses in new locations. RNA interference (RNAi)-mediated resistance has been found to be highly effective in controlling RNA virus infections, and such approach needs to be exploited to produce pecluvirus-resistant crops. The suspected role of P39 in vector transmission of the pecluvirus needs to be confirmed by mutational analysis in order to deploy this ORF in transgenic research. The presence of subgenomic RNAs encoding P14 and P17 in infected tissues needs to be verified. The cultural control measures tested for IPCV are worth exploiting in West Africa to minimize the impact of PCV. Development of duplex or multiplex RT-PCR is necessary to distinguish peanut clump and groundnut green rosette diseases that visually look alike. Broadly specific nonradioactive probes must be made available to researchers in Africa and India.

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Further Reading Adams, M.J., Antoniw, J.F., Kreuze, J., 2009. Virgaviridae: A new family of rod-shaped plant viruses. Archives of Virology 154, 1967–1972. Adams, M.J., Adkins, S., Bragard, C., et al., 2017. ICTV virus taxonomy profile: Virgaviridae. Journal of General Virology 98, 1999–2000. Boulila, M., 2012. Molecular evolution and genetic distinctness among rod-shaped plant RNA viruses of the family Virgaviridae. In: Chiheb, B., Khalil., B. (Eds.), Bioinformatics Research: New Developments. New York: Nova Science Publishers, pp. 107–130. CABI/EPPO, 2006. Peanut clump virus. Distribution Maps of Plant Diseases. Wallingford: CAB International, (No. 988). Delfosse, P., Reddy, A.S., Legrève, A., et al., 1999. Indian peanut clump virus (IPCV) infection on wheat and barley: Symptoms, yield loss and transmission through seed. Plant Pathology 48, 273–282. Delfosse, P., 2000. Epidemiology and Management of the Indian peanut clump virus. Louvain-la-Neuve: Université catholique de Louvain, (PhD Thesis). Delfosse, P., Reddy, A.S., Legreve, A., et al., 2000. Serological methods for detection of Polymyxa graminis, an obligate root parasite and vector of plant viruses. Phytopathology 90, 537–545. Dieryck, B., Otto, G., Doucet, D., et al., 2009. Seed, soil and vegetative transmission contribute to the spread of pecluviruses in West Africa and the Indian sub-continent. Virus Research 141, 184–189. Dieryck, B., Delfosse, P., Reddy, A.S., Bragard, C., 2010. Targeting highly conserved 30 -untranslated region of pecluviruses for sensitive broad-spectrum detection and quantitation by RT-PCR and assessment of phylogenetic relationships. Journal of Virological Methods 169, 385–390. Dieryck, B., Weyns, J., Doucet, D., Bragard, C., Legreve, A., 2011. Acquisition and transmission of Peanut clump virus by Polymyxa graminis on cereal species. Phytopathology 101, 1149–1158. Pandey, V., Mandal, B., Jain, R.K., Roy, A., 2017. Indian peanut clump virus, a fungal transmitted Pecluvirus infecting both monocotyledonous and dicotyledonous plants in India. In: Mandal, B., Rao, G.P., Baranwal, V.K., Jain, R.K. (Eds.), A Century of Plant Virology in India. Singapore: Springer Nature Singapore Pvt. Ltd, pp. 351–359. Reddy, A.S., Hobbs, H.A., Delfosse, P., Murthy, A.K., Reddy, D.V.R., 1998. Seed transmission of Indian peanut clump virus (IPCV) in peanut and millets. Plant Disease 82, 343–346. Tamada, T., Kondo, H., 2013. Biological and genetic diversity of plasmodiophorid-transmitted viruses and their vectors. Journal of General Plant Pathology 79, 307–320. Vaïanopoulos, C., Bragard, C., Moreau, V., Maraite, H., Legrève, A., 2007. Identification and quantification of Polymyxa graminis f. sp. temperata and P. graminis f. sp. tepida on barley and wheat. Plant Disease 91, 857–864.

Pepino Mosaic Virus (Alphaflexiviridae) Rene AA van der Vlugt, Wageningen University and Research Center, Wageningen, The Netherlands CCMM Stijger, Wageningen University and Research Center, Bleiswijk, The Netherlands r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein DAS-ELISA Double antibody Sandwich - Enzyme-linkedImmunosorbent-Assay ELISA Enzyme-linked immunological assays EM Electron microscope HEL NTPase/helicase domain IEM Immunosorbent electron microscopy IR Intergenic region

Glossary Double antibody sandwich - Enzyme-Linked-ImmunoSorbent-Assay An assay in which by the use of a virusspecific antibody the presence of a virus in plants or plant parts can be established. Immunosorbent electron microscopy Identification of a virus by visualization in the electron microscope of the reaction of the virus coat protein with a specific antiserum.

kDa Kilo Dalton MET Methyl-transferase domain nm Nanometer nt Nucleotide(s) ORF Open reading frame RdRp RNA-dependent RNA polymerase RISC RNA-induced silencing complex RT-PCR Reverse transcriptase polymerase chain reaction RT Reverse transcriptase TGB Triple gene block

Reverse transcriptase polymerase chain reaction A PCR reaction whereby first a cDNA copy of the viral RNA has to be generated by Reverse Transcriptase (RT) before PCRamplification is possible. Virus symptomatology Typical local or systemic symptoms on natural host plants or experimental test plants that can be used to characterize and differentiate plant viruses and plant virus strains.

Introduction Pepino mosaic virus (PepMV) was first found in 1974 in field samples of pepino plants (Solanum muricatum) located near some potato fields, collected in the Canete valley in coastal Peru. These plants showed some yellow mosaic in young leaves. Electron microscope (EM) investigations showed the presence of typical filamentous potexvirus particles of approximately 500 nm, which did not react with antisera against Potato virus X (PVX). Serologically it appeared most closely related to Narcissus mosaic virus (NaMV) but the host ranges from both viruses differ considerably. It was concluded that Pepino mosaic virus was a new and distinct potexvirus species. Since its first description in 1980 the virus was never reported and not considered to be of any agricultural significance until it was described in 1999 from commercial protected tomato crops (S. lycopersicum) in the UK and The Netherlands. Since then the virus has spread very rapidly through commercial tomato crops and is now reported worldwide (See Relevant Websites Section). It is particularly present in Europe and The America’s. In Africa and Asia its presence appears to be restricted and it has not been reported from Australia or New Zealand.

Taxonomy and Classification Pepino mosaic virus is a species member of the genus Potexvirus within the family Alphaflexiviridae.

Virion Properties PepMV has typical filamentous potexvirus particles with a normal length of 510 nm. Particles are comprised of a single capsid protein (CP) of approximately 26 kDa. Ultrathin section of infected leaf material may contain inclusions consisting of arrays of filamentous virus-like particles. The virus particles are less clearly cross-banded as those of PVX. As a member of the genus Potexvirus, PepMV is fairly stable. In endpoint dilution studies, sap from infected N. glutinosa was always infectious at dilutions of 10
4, occasionally at 10
5 but never at 10
6. Sap lost most of its infectivity after 10 min at 651C and was no longer infectious at 701C. Sap stored at 201C still shows some infectivity after 3 months while leaves of N. glutinosa desiccated over silica gel were still infectious after 6 months. Under practical conditions in greenhouses and fields the virus may

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Fig. 1 Schematic organization of the RNA genome of Pepino mosaic virus with its five open reading frames (ORFs). M7G ¼ 50 cap, AAAAAn ¼ poly-A tail.

easily survive for several weeks in plant debris and on surfaces (e.g., tools, clothes, containers) that have come in contact with virus-infected leaves or fruits.

Genome Organization and Expression The genome of PepMV resembles that of typical potexviruses. For all strains its positive single-stranded RNA is around 6410 nt long, capped at the 50 -end, polyadenylated at the 30 -end and contains 50 - and 30 -untranslated regions. It encodes five putative partly overlapping open reading frames (ORFs 1–5, see Fig. 1). The following data are based on the RNA sequence of the original EU-tomato isolate PD99901066 (FJ940223) as found in 1999 in The Netherlands. Positions for regions and motifs in other isolates and other strains may vary slightly. The 50 -untranslated region (50 -UTR) of the virus is 86 nt longs and like all other potexviruses it starts with the 50 -GAAAA pentanucleotide. ORF1 (nt 86–4406) encodes a 164 kDa protein of 1439 aa. It contains a putative methyl-transferase domain (MET; aa 59–224) specific for the supergroup of ‘Sindbis-like’ viruses, a NTPase/helicase domain (HEL; aa 708–934) with the NTP-binding motifs GCGGSGKS and VVIFDD and a RNA dependent RNA polymerase (RdRp) domain (aa 1217–1374) characterized by the SGEGPTFDANT-X22-GDD motif. The stop codon of the first ORF is followed by a short intergenic region (IR1) of 25 nt and a set of three partially overlapping ORFs typically known as the triple gene block (TGB). ORF2 (nt 4432–5136) encodes the first Triple Gene Block protein (TGB1), a 234 aa protein of 26 kDa. This protein contains a typical NTPase/helicase motif (aa 26–233) characterized by 7 conserved motifs, two of which may be involved in NTP binding. TGBp1 belongs to the superfamily I of RNA helicases. ORF3 (nt 5117–5488) overlaps 19 nt with the 30 -end of ORF2 and extends 148 nt past the start codon of ORF4. It encodes the TGB2 protein, a small 14 kDa protein which contains a potexvirus specific consensus motif: PxxGDxxHxL/FPxGGxYxDGTKxxxY. ORF4 (nt 5340–5594) encodes the third TGB protein (TGB3), a 85 aa protein of 9 kDa. This protein is the most variable among potexviruses. It contains a CxV/IxxxG consensus motif among potexvirus TGB3 proteins. Substitution of Lysine (K) for glutamic acid (E) at position 67 in TGB3 was described as being involved in increased virus accumulation and the induction of systemic necrosis both in tomato and test plants. The TGB proteins are essential for virus movement, suppression of RNA silencing and the formation of the virus factory. The second intergenic region (IR2) of 38 nt (nt 5595–5632) precedes ORF5 (nt 5633–6346) which encodes the 238 aa coat protein (CP) of 25 kDa. This CP contains the amphipathic core sequence KFAAFDFFDGVT. A similar sequence is also found in the CP of other potexviruses and might be responsible for binding of virus RNA to the CP through hydrophobic interactions. The PepMV CP is also required for cell-to-cell and long-distance movement and is involved in the suppression of RNA silencing. The 64 nt long 30 -untranslated region (30 -UTR: nt 6347–6410) precedes the poly-A tail and contains the hexameter 50 -ACUUAA sequence which is also present in the 30 -UTR of all potexviruses sequenced so far. This motif is proposed to be a cis-acting element involved in the positive and negative viral RNA synthesis. The 50 -AAUAAA polyadenylation signal terminates the RNA genome whereby the AAA portion form the first A residues of the poly(A) tail.

Host Range and Symptomatology The host range of PepMV is relatively broad infecting plants from different families including both cultivated and wild hosts. PepMV has not been reported to infect plants from the Cucurbitaceae or Fabaceae. Most plant species belong to the Solanaceae of which many species are infected systemically. Its main hosts appear to be Solanum spp. The originally described pepino strain was found to infect wild and commercial tuber-bearing S. tuberosum sp. (potato), mostly with a symptomless systemic infection or with mild mosaic symptoms but in two local Peruvian S. tuberosum cultivars and S. stoloniferum PI 230557 it caused a severe systemic necrosis. The virus was shown to be transmitted through the tubers. Its best-known natural hosts are pepino (S. muricatum) and cultivated tomato (S. lycopersicum) but surveys showed infection with PepMV of several related wild Solanum spp. These include S. peruvianum, S. parviflorum, S. chilense, S. chmielewskii, and S. pimpinellifolium. Infections in these wild species were generally symptomless. PepMV was also reported to infect other solanaceous crop and test plants; Solanum melongena (eggplant), S. nigrum, S. tuberosum (potato), Datura stramonium, D. metel, D. inoxia, Nicotiana bethaminana, N. occidentalis, N. glutinosa, and N. tabacum cv Xanthi. Sweet

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pepper (Capsicum annuum) does not seem a suitable host since most PepMV strains were not able to induce a systemic infection, with the exception of the US1 strain which could induced systemic necrotic lesions. Several weed hosts from various plant families (e.g., Convolvulaceae, Brassicaceae, Boraginaceae, Asteraceae, Plantaginaceae and Polygonaceae were found to be hosts for PepMV. The original virus description reports weed hosts in Peru including Datura stramonium, Nicandra physaloides and Physalis peruvianum. Studies on sampled material from mainland Spain and the Canary Islands show a large number of weeds species latently infected (i.e., showing no symptoms)). The initially described PepMV strain caused a distinct yellow mosaic on young leaves of pepino and most infected plant also showed dark green enations on the lower surface of some leaves. In Solanum spp. it caused a symptom less systemic infection as shown by back-inoculation on a sensitive indicator plants like N. glutinosa. Only D. metel, D. stramonium, and a number of Nicotiana sp. showed distinct symptoms upon systemic infection. In contrast to the original Pepino isolate the EU-tomato virus isolate found in 1999 in commercial tomato crops in the UK and The Netherlands did cause distinct although generally mild symptoms in commercial tomato crops. In general, PepMV symptoms depend on the particular virus isolate in combination with the tomato cultivar, the age of the plant when first infected and environmental conditions. Symptoms may include small yellow leaf spots, yellow-green mosaic, mottle on the older leaves and/or slight curling in the top leaves (nettle-head symptom) or a greyish appearance of the top of the plant. Fruit symptoms may range from discoloration in the form of blotchy ripening or flaming, fruit marbling, mild sometimes concentric yellow/orange mottling and uneven ripening to netting and cracking (‘open fruits’) and shape distortion. Basically, severity of symptoms in tomato may vary with the particular virus isolate involved. Both mild and severe symptoms inducing PepMV isolates have been described and there appears to be no correlation between symptom severity in tomato and the PepMV virus strain. Mild symptoms may vary from small yellow spots on leaves to a slight reduction in fruit production. Certain isolates can cause much more pronounced plant symptoms: more severe leaf chlorosis, bright yellow mosaic, leaf distortions (‘bubbling’), green striation on the stem and sepals up to more or less pronounced leaf and stem necrosis. Because of fruit symptoms, fruit quality can be affected which may result in economic damage. Symptom expression is reported to depend on environmental conditions with low light conditions and low temperatures favouring more pronounced symptoms. Inoculation studies of three PepMV strains (EU, CH2, and US1) clearly showed a number of differences between these strains on tomato and other solanaceous crops and test plants. Tomato is susceptible to all three strains, while sweet pepper (C. annuum) is generally not a systemically susceptible host for the three strains. Eggplant is susceptible to all three strains however local symptoms are seldom seen and systemic leaf symptoms may depend on locality and virus isolate. In potato systemic infections may develop in a sensitive cultivar but local and systemic symptoms are seldom seen. The virus however can infect the tubers and can thus be transmitted. In test plants N. occidentalis 37B proved a useful susceptible host which is able to differentiate between strains based on typical local and systemic symptoms.

Virus Isolates and Strains The first PepMV isolate was isolated in 1974 from pepino crops in Peru and only in 1999 an isolate was described on commercial tomato. Comparison of this tomato isolate with the original Peruvian pepino isolate (#BBA1137 in the DSMZ plant virus collection in Germany) showed some differences in a number of characteristics. These were most pronounced in the reactions of both isolates on tomato (S. lycopersicum) in which the pepino isolate is symptomless and the particular isolate of the tomato strain caused mild but distinct symptoms in the form of small sharp yellow flecks on leaves. Also on a number of test plants the two isolates can be distinguished, most clearly on N. glutinosa and D. stramonium. Since 1999 a large number of different PepMV isolates have been described. These were either isolated from commercial tomato crops or tomato seed lots in different countries or from wild Solanum sp. Most show the typical characteristics of the tomato strain but some deviant isolates were observed. A number of isolates show more severe symptoms, (including leaf and stem necrosis and sever fruit symptoms) in tomato or wild Solanum sp. There is considerable biological variation between isolates and severity of symptoms is not related to the virus strain and in most strains both mild and severe isolates have been described. Currently available biological data (i.e., host and indicator plant symptoms), serological relationships and multiple nucleotide sequence alignments (Fig. 2) suggest at least five clusters of relationship between the PepMV isolates which are now considered as separate genotypes or strains: (1) (2) (3) (4) (5)

The The The The The

Peruvian pepino strain (LP), European tomato strain (EU), Chile-2 strain (CH2), US1 strain (US1), PES strain, identified from wild Solanum species in Peru.

A significant number of full genome sequences are now available for PepMV with most sequences derived EU and CH2 isolates. Only a limited number of sequences from the LP, US1, and PES strains are published. Sequence identity between isolates of the same strain is high (i.e., above 98%) and several studies have shown that single strain populations of PepMV shown very low genetic variation over time. Direct full genome nucleotide sequence comparisons between strains showed that LP and EU strains

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Fig. 2 Phylogenetic Maximum Likelihood tree of full-length RNA sequences of selected Pepino mosaic virus isolates of the five main strains. LP ¼ Peruvian strain, EU ¼ European tomato strain, CH2 ¼ Chile-2 strain, US1 ¼ US1 strain, PES ¼ wild Solanum strain.

are most closely related with around 95% sequence identity. The US1 and PES strains share around 86% identity to each other and around 81% identity to the LP and EU strain. The CH2 strain is the most deviant as it generally shares around 78% sequence identity with the four other strains Fig. 3.

Virus Transmission and Epidemiology PepMV clearly originates from South America. Surveys in Peru showed that the virus was widespread, even in isolated wild Solanum sp, and phylogenetic analyses suggested that the virus most likely originated from a wild Solanum sp. Interestingly most infected Solanum sp. showed no distinct symptoms. Several studies have confirmed that the original Peruvian pepino isolate (LP) is characterized by the absence of, or only very mild symptoms on commercial tomato crops. The EU-strain isolate first found in 1999 in Europe also induces only mild symptoms in commercial tomato crops. This likely contributed to the initial rapid spread of the virus within and outside Europe. Initial studies between 2000 and 2005 employing comparisons of viral gene sequences revealed that the tomato strain was prevalent in all countries reporting the virus. All tomato isolates found were all highly similar and could not be grouped according to geographic origin, or symptoms on naturally or experimentally infected plants. These data indicated a fairly recent introduction from limited sources and expansion of the tomato strain into commercial tomato crops, as the most likely cause for the virus epidemic.

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Fig. 3 Symptoms of Pepino mosaic virus on tomato. A. Severe symptom on tomato plant. B. Detail of mild leaf symptom on tomato leaf. C. Severe symptom on tomato fruits. Source: Courtesy of R. van der Vlugt.

The first known occurrence of PepMV in North America can be traced back to 2000. Three strains were identified from pooled symptomatic leaf material collected in Arizona; US1, US2, and US3. US1 and US2 clearly differed from the EU and LP strains while US3 was highly homologous to the EU strain. Later studies revealed that US1 is indeed a distinct strain while US2 was a recombinant sequence between US1 and another strain now known as Chile-2 (CH2). US1 and CH2 were later confirmed as distinct strains on tomato seed originating from Chile. In 2005 the CH2 strain was first found in commercial tomato crops in Europe and soon after that it started to spread rapidly, apparently because it seems to have a biological advantage over the EU strain in tomato. Detailed population studies from Spain on material collected between 1998 and 2004 confirmed the prevalence and homogeneity of the tomato strain isolates as well as their presence since 1998. However, there were clear indications for several independent introductions of this virus strain, both on the Canary Islands and the Spanish Main land. Interestingly in addition to the tomato strain, also the original LP strain and the CH2 strain were found, but always in mixed infections with the tomato strain. Both strains were shown to be present in Spain since 2000. Since 2005 the CH2 strain spread rapidly and is now the most prevalent strain in most commercial tomato crops where the virus is found however, it did not replace the EU tomato strain. Isolates for both strains can occur in mixed infections the same plant. The US1 strain is found only occasionally. PepMV is a mechanically transmitted virus. The most important transmission routes are through contaminated tools, clothes, and surfaces. The virus is relatively stable at room temperature and can survive and stay infectious for several weeks in plant debris and on contaminated surfaces. Fruits from infected plants can contain high concentrations of virus and do not necessarily show symptoms. Long distance transmission of the virus in this way is relatively likely. Implementation of strict hygiene protocols during the growing season and thorough cleaning of greenhouses at the end of the growing season have shown to effectively control the introduction and spread of the virus. PepMV is also transmitted by tomato seed at low levels whereby the virus is likely located on the seed coat. The virus was also shown to be efficiently transmitted between tomato plants in closed recirculating hydroponic systems as well as by bumblebees. The latter two transmission routes are likely to contribute to the spread of the virus in commercial closed greenhouse production systems whereby bumblebees are used for pollination. The possible involvement of Olpidium virulentus in the transmission of PepMV has been reported as well as low levels of transmission from tomato to tomato under experimental conditions by the greenhouse whitefly (Trialeurodes vaporariorum) and the aphids Myzus persicae. Transmission of PepMV by these vectors remains to be confirmed.

Diagnostic A polyclonal antiserum raised against the original pepino isolate shows no reaction with Potato virus X (PVX) and Potato aucuba mosaic virus (PAMV), the only two other potexviruses known to infect tomato. Both in immuno-sorbent electron microscopy (IEM) and in ELISA it clearly reacted with different tomato isolates. Comparisons of polyclonal antisera raised against the pepino and tomato strains showed differences in heterologous titres between them, but the antisera show clear reactions with isolates from all known PepMV strains.

Virus Damage and Control Initially PepMV infections in commercial tomato crops were reported as relatively insignificant with no apparent plant or fruit symptoms and no or very limited yield reduction. Studies from the UK however, reported significant effects on fruit quality. Smaller sized fruits with different grades of uneven ripening and dis-coloration and occasionally mis-shaped fruits, led to production unsuited for the fresh UK market and hence economic damage.

544

Pepino Mosaic Virus (Alphaflexiviridae)

Studies now clearly show that symptoms induction by PepMV is highly dependent on environmental conditions and on the particular isolate involved. In Spain, PepMV infected tomato plants showed only symptoms from autumn through spring and symptoms disappeared with higher temperatures in late spring. Similar effects of high light conditions and high temperatures have been reported from other countries. However, symptoms of PepMV infected plants can be highly variable, ranging from very mild leaf symptoms to severe leaf and stem necrosis. Fruit symptoms may vary from mild discolorations to open fruits showing severe cracking. Mild, chlorotic and even necrotic PepMV isolates have been reported and the capacity to induce different symptoms is a property of the isolate. There are indications that severity of symptoms may be associated with increased virus accumulation. Distinct single nucleotide mutations resulting in specific amino acid mutations in the viral CP and TGB3 genes were shown to be involved in the development of yellowing and necrotic symptoms, respectively. Also the highly conserved ‘palm’ domain in the Polymerase domain of the viral RdRp encoded by ORF1 of plays a role in the induction of plant necrosis which is apparently modulated by the interplay between the TGB3 protein, plant genotype and environmental conditions. Currently no resistance genes to PepMV are available in commercial tomato cultivars. Extensive screening in wild Solanum sp. has been identified only a very limited number of potential resistance sources. Some accessions of S. lycopersicoides and S. chilense were resistant to isolates from the EU strain. Whether this resistance holds against the other PepMV strains remains to be investigated. One method currently applied in various countries to control negative effects of PepMV infections is cross-protection i.e., the use of attenuated virus isolates. A mild isolate is inoculated onto plants in an early growth stage and the resulting systemic infection protects the plant against more severe isolates of the virus. Cross-protection is based on the sequence specific activation of the RNA-induced silencing complex (RISC). This general antiviral defense mechanism is sequence homology-dependent and is therefore not virus isolate- but strain-specific. Protection against severe EU-strain isolates requires an attenuated EU isolate while an attenuated CH2 isolate is required to protect against aggressive CH2 isolates. In many instances isolates of the CH2 and EU strains occur in mixed infections so for adequate protection mild isolates of both strains are required. PepMV is a mechanically transmitted virus. The most important transmission routes are through contaminated tools, clothes and surfaces. The virus is relatively stable at room temperature and can survive and stay infectious for several weeks in plant debris and on contaminated surfaces. Fruits from infected plants can contain high concentrations of virus and do not necessarily show symptoms. Long distance transmission of the virus in this way is relatively likely. Implementation of strict hygiene protocols during the growing season and thorough cleaning of greenhouses at the end of the growing season have shown to effectively control the introduction and spread of the virus. Effective control of the virus in commercial tomato culture should come from the use of virus-free seeds and planting material. For this reliable and sensitive methods, both serology and molecular based, for the detection of the virus in seed, plants and fruits are available. Since no adequate plant resistance is available yet strict hygiene measures and a constant monitoring for possible infections are required to avoid unwanted spread of the virus.

See also: Alphaflexiviruses (Alphaflexiviridae). Potexviruses (Alphaflexiviridae)

Further Reading Agüero, J., Gómez-Aix, C., Sempere, R.N., et al., 2018. Stable and broad spectrum cross-protection against Pepino mosaic virus attained by mixed infection. Frontiers in Plant Sciences 9, 1810. doi:10.3389/fpls.2018.01810. Blystad, D.-R., van der Vlugt, R., Alfaro-Fernández, A., et al., 2015. Host range and symptomatology of Pepino mosaic virus strains occurring in Europe. European Journal of Plant Pathology 143, 43–56. Hanssen, I., Thomma, B., 2010. Pepino mosaic virus: a successful pathogen that rapidly evolved from emerging to endemic in tomato crops. Molecular Plant Pathology 11, 179–189. doi:10.1111/j.1364–3703.2009.00600.x. Jones, R.A.C., Koenig, R., Lesemann, D.-E., 1980. Pepino mosaic virus, a new potexvirus from pepino (Solanum muricatum). Annals of Applied Biology 94, 61. Moreno-Pérez, M.-G., Pagán, I., Aragón-Caballero, L., et al., 2014. Ecological and genetic determinants of Pepino mosaic virus emergence. Journal of Virology 88 (6), 3359. Van der Vlugt, R.A.A., Cuperus, C., Vink, J., et al., 2002. Identification and characterization of Pepino mosaic virus in tomato. EPPO Bulletin 32, 503.

Relevant Websites https://gd.eppo.int/taxon/PEPMV0/documents EPPO Global Datbase Pepino mosaic virus. https://gd.eppo.int/taxon/PEPMV0/distribution EPFO Global Database Pepino mosaic virus Distribution.

Plant Reoviruses (Reoviridae) Yu Huang and Yi Li, Peking University, Beijing, China r 2021 Elsevier Ltd. All rights reserved. This article is an update of R.J. Geijskes, R.M. Harding, Plant Reoviruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00468-4.

Nomenclature aa Amino acid(s) cDNA Complementary deoxyribonucleic acid dsDNA Double-stranded deoxyribonucleic acid dsRNA Double-stranded ribonucleic acid ELISA Enzyme-linked immuno-sorbent assay mRNA Messenger RNA

Glossary Capsid The protein shell that surrounds a virus particle. Cell-to-cell movement protein Virally encoded protein that helps viral virions or genome RNA (s)/protein complexes to move from cell to cell. Dicotyledon A flowering plant that has two cotyledons in the seed. Icosahedron Having 20 equal sides or faces. Monocotyledon A flowering plant that has only one cotyledon, or seed leaf, in the seed.

ORF Open reading frame RdRp RNA-dependent RNA polymerase RT-LAMP Reverse transcription loop-mediated isothermal amplification RT-PCR Reverse transcription polymerase chain reaction VSR Viral suppressor of RNA silencing

RNA-dependent RNA polymerase Virally encoded RNA-dependent RNA polymerase that replicates viral genome(s). Transovarial transmission Transmission from one generation to another through eggs. Viral suppressor of RNA silencing Virally encoded protein(s) that suppress(es) host RNA silencing pathways. Viroplasm Cytoplasmic, non-membranous structure in which plant reoviruses are thought to replicate and assemble.

Introduction The family Reoviridae comprises a diverse group of viruses that can infect vertebrates, invertebrates, and plants. Despite their large host range, all members of the family Reoviridae share common properties, including an icosahedral-shaped virion and a segmented double-stranded RNA (dsRNA) genome. The family Reoviridae consists of 15 genera divided between two subfamilies, Spinareovirinae and Sedoreovirinae. Plant-infecting reoviruses are members of the genera Oryzavirus and Fijivirus, which belong to the Spinareovirinae subfamily, and the genus Phytoreovirus, part of the Sedoreovirinae subfamily. These reoviruses generally replicate both in their plant hosts and in their insect vectors. Infection of the insect vector is non-cytopathic and often persists throughout the life of the insect. Infection of the host plant is tissue specific (except for Rice dwarf virus [RDV]) and can cause severe disease. Fiji leaf gall disease, caused by Fiji disease virus (FDV), has caused yield losses of up to 90% in susceptible varieties of sugarcane in Australia. Rice ragged stunt virus (RRSV) is reported to reduce yield of rice by up to 100% in severe infections (although reductions of 10%–20% are more common). Rice common dwarf disease, caused by RDV, can also cause significant losses as infected plants often fail to set seed. Southern rice black-streaked dwarf disease, caused by Southern rice black-streaked dwarf virus (SRBSDV), was first discovered in Yangxi County, Guangdong Province, China, in 2001, and its distribution was limited to southern China until the end of 2008. By 2010, however, it had spread to 29 provinces of Vietnam and 13 provinces of China, and it has also now been found in some areas in Japan. The rapid spread of SRBSDV has caused serious rice losses in East and Southeast Asia.

Taxonomy and Classification Currently, three genera of the family Reoviridae are classified as plant-infecting reoviruses: Fijivirus, Oryzavirus, and Phytoreovirus. The genera of plant-infecting reoviruses are differentiated according to the number of genomic dsRNA segments and their electrophoretic profile, hosts, serological relationships, and capsid morphology (Table 1). These reoviruses replicate both in plant hosts (except for one fijivirus: Nilaparvata lugens reovirus) and in their insect vectors (Table 1). Infection of the host plant is species specific, although the host range can often be extended under experimental conditions and can produce a variety of symptoms, including severe disease. None genome sequence information is now available for PaSV, GDV and ERSV. Partial genome sequence information is now available for WTV and OSDV. Complete genome sequence information have been obtained for all the other plant reoviruses. This has allowed detailed comparisons to be made within these genera and across all of the family Reoviridae,

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546

Table 1

Plant Reoviruses (Reoviridae)

Characteristics of plant reoviruses

Subfamily, Genus Species name

Abbr.

Spinareovirinae Fijivirus Fiji disease virus (FDV) Rice black-streaked dwarf virus (RBSDV)

dsRNA segments

Host

Vector (s)

10 10

Gramineae Gramineae

Planthoppers: Perkinsiella saccharicida, P. vastatrix, P. vitiensis Planthoppers: Laodelphax striatellus, Ribautodelphax albafascia, Unkanodes sapporona Planthopper: Ribautodelphax notabilis Planthoppers: Sogatella furcifera, S. kolophon Planthopper: Delphacodes kuscheli Planthopper: Sogatella furcifera

Maize rough dwarf virus Pangola stunt virus Mal del Rio Cuarto virus Southern rice black-streaked dwarf virus Oat sterile dwarf virus

(MRDV) (PaSV) (MRCV) (SRBSDV)

10 10 10 10

Gramineae Gramineae Gramineae Gramineae

(OSDV)

10

Gramineae

Garlic dwarf virus Nilaparvata lugens reovirus

(GDV) (NLRV)

10 10

Liliaceae No plant host reported

Spinareovirinae Oryzavirus Rice ragged stunt virus (RRSV) Echinochloa ragged stunt virus (ERSV)

10 10

Gramineae Gramineae

Planthopper: Nilaparvata lugens Planthoppers: Sogatella longifurcifera and S. vibix

Spinareovirinae Phytoreovirus Wound tumor virus Rice dwarf virus Rice gall dwarf virus

12 12 12

Dicot Gramineae Gramineae

Leafhoppers: Agallia constricta, A. quadripunctata, Agalliopsis novella Leafhoppers: Nephotettix cincticeps, N. nigropictus, Recillia dorsalis Leafhoppers: Nephotettix cincticeps, N. nigropictus, N. virescens, N. malayanus, Recillia dorsalis

(WTV) (RDV) (RGDV)

Planthoppers: Javesella pellucidia, J. discolour, J. dubia, J. obscurella, Dicranotropis hamata Planthopper: Unknown Planthoppers: Nilaparvata lugens, Laodelphax striatellus

providing a basis for their classification into species and genera. One general feature is the presence of inverted repeats several bases long adjacent to the conserved terminal sequences in all plant-infecting reoviruses; the sequences of the repeats differ among the three genera. Within the genus Fijivirus, individual species have considerable similarities to FDV, the type species. Their classification into separate species is based on unique characteristics, such as capacity to exchange genome segments, relatively high aa sequence similarity, serological cross-reaction, cross-hybridization of RNA or cDNA probes, host species, and insect vector species. Analysis of the available genome sequences has further assisted the identification of Fijivirus species. The gross genome characteristics of fijiviruses includes a genome size of approximately 29 kbp and a characteristically low G þ C content of 34%–36%. The 30 terminal trinucleotide in the positive strand is unique and highly conserved in all RNA segments among all species within the genus Fijivirus (Table 2). Members of the genera Oryzavirus and Phytoreovirus have significant similarity to their type species, RRSV and Wound tumor virus (WTV), respectively. Demarcation of species within the oryzaviruses and phytoreoviruses is primarily based on the ability to exchange genome segments although other characteristics, as mentioned for fijiviruses above, are also used. When available, genomic sequences are examined to reveal distinguishing features to support the classifications. Oryzaviruses have a total genome size of approximately 26 kbp and specific 50 - and 30 terminal sequences in all RNA segments, resulting in the canonical structure 0 50 GAUAAA–GUGC3 . Phytoreoviruses have a total genome size of approximately 25 kbp, a G þ C content between 38% and 48%, 0 0 0 and specific 5 - and 30 -terminal sequences in all RNA segments [canonical structure 5GG(U/C)A–(U/C)GAU3 ].

Virion Structure and Genome Organization Fijivirus Fijivirus virions have a complex double icosahedral capsid construction and consist of a capsid, a core, and a nucleoprotein complex. Virions are fragile structures and readily break down in vitro to give cores unless pretreated with fixative. The outer capsid is 65–70 nm in diameter with 12 “A”-type spikes located at the vertices of the icosahedron. The inner core is about 55 nm in diameter, with 12 “B”-type spikes located at the vertices. The viral nucleic acid is located at the center of the virus particle, within the inner core capsid (Fig. 1(A)). Each virion contains a single full-length copy of the genome. Fijivirus genomes contain 10 dsRNA genomic segments varying from approximately 1.8 to 4.5 kbp (Fig. 2). The total genome is approximately 29 kbp, with a low G þ C content of 34%–36%. Within the genus, the only conserved sequence is found at the 30 -terminus (canonical structure …GUC-30 ; Table 2). Segment-specific inverted repeats are found adjacent to these terminal sequences. Segments 1–6, 8, and 10 are monocistronic, containing one open reading frame (ORF), while segments 7 and 9 each contain two ORFs, although the expression of the second ORF has not been certified in vivo in either plant cells or insect vectors. NLRV is the only fijivirus identified to date that differs from this standard structure, with only one ORF on segment 10, which is equivalent to segment

Plant Reoviruses (Reoviridae)

Table 2

547

Conserved terminal sequences (positive strand) in fijiviruses

Virus Fiji disease virus Rice black-streaked dwarf virus Maize rough dwarf virus Mal del Rio Cuarto virus Oat sterile dwarf virus Nilaparvata lugens reovirus Southern rice black-streaked dwarf virus

5 0 conserved sequence

3 0 conserved sequencea

50

50

0

AAGUUUUU–3 0 AAGUUUUU–3 0 50 AAGUUUUUU–3 50 30 AAGUUUUU– 0 50 AACGAAAAAAA–3 50 30 AGU– 0 50 AAGUUUUU–3 50

0

–CAGCNNNNGUC3 0 –AGCUNN(C/U)GUC3 50 30 –UGUC 0 50 –CAGCUNNNGUC3 0 0 5 –UUUUUUUUAGUC3 50 30 –GUUGUC 0 50 –CAGCUGAUGUC3 50

Italicized trinucleotides are conserved in all fijivirus sequences reported to date.

a

Fig. 1 Diagrammatic representation of the particle structures of Fiji disease virus (A), Rice ragged stunt virus (B), and Rice dwarf virus (C).

Fig. 2 Genome organization of Fiji disease virus (FDV) containing 10 dsRNA segments. Each segment contains one ORF except for Seg 7 and 0 0 Seg 9 which contain two ORFs. The green arrows indicate the conserved 50 terminal sequence (5 AAGUUUUU–3 ) while the red arrows indicate the 0 0 5 3 conserved 30 terminal sequence ( –CAGCNNNNGUC ). The red segment encodes the RdRp. The green segments encode the structural proteins. The yellow segments encode non-structural proteins or proteins with unknown function.

7 of FDV. The functions of the proteins encoded by some of the ORFs are still unconfirmed; the functions of segments 1–3 and 7–10 have been predicted based on protein expression studies or deduced from those of the equivalent segments of other virus species within the genus (Table 3).

Oryzavirus Oryzavirus virions have a double-shelled icosahedral capsid and consist of an outer capsid, an inner capsid, and a core. Virions are fragile and readily break down in vitro to give subviral core particles unless pretreated with fixative. The outer capsid is 75–80 nm in diameter with 12 “A”-type spikes located at the 5-fold axis of the icosahedron. The core capsid is about 57–65 nm in diameter,

548

Table 3

Plant Reoviruses (Reoviridae)

Genome organization of FDV and predicted gene function

Segment

Size (bp)

Protein nomenclature

Predicted MW (kDa)

Predicted function (location)

S1 S2 S3 S4 S5 S6 S7

4532 3820 3623 3568 3150 2831 2194

VP1 VP2 VP3 VP4 VP5 VP6 VP7a VP7b

170.6 137.0 135.5 133.2 115.3 96.8 41.7 36.7

RdRp (core) Major core (core) Possible B spike (outer capsid) Unknown Unknown Unknown Possible tubule protein (nonstructural) Unknown

S8 S9

1959 1843

VP8 VP9a VP9b

68.9 38.6 23.8

Possible NTP-binding (core) Structural protein (viroplasm) Unknown

S10

1819

VP10

63.0

Outer capsid protein (major outer capsid)

Fig. 3 Genome organization of Rice ragged stunt virus (RRSV) containing 10 dsRNA segments. Each segment contains one ORF except for 0 0 Seg 4 which contains two ORFs. The green arrows indicate the conserved 50 terminal sequence (5 GAUAAA–3 ) while the red arrows indicate the 50 30 0 conserved 3 terminal sequence ( –GUGC ). The red segment encodes the RdRp. The green segments encode the structural proteins. The yellow segments encode non-structural proteins or proteins with unknown function.

with 12 “B”-type spikes. The viral nucleic acid is located at the center of the viral particle, within the core capsid (Fig. 1(B)). The virus genome consists of 10 linear dsRNA segments ranging in size from 1162 to 3849 bp (RRSV) with a total length of 26 kbp (Fig. 3). Genome segments 1–3 and 5–10 of RRSV each contain a single ORF, while segment 4 contains two ORFs. Segment 8 encodes a polyprotein that is autocatalytically cleaved into at least two polypeptides. Table 4 summarizes the organization of the RRSV dsRNA segments and the predicted function of the encoded proteins.

Phytoreovirus Phytoreovirus virions have a double-shelled icosahedral capsid construction and consist of an outer capsid, a core capsid, and a smooth core. Virions are approximately 70 nm in diameter, with 12 spikes located at the 5-fold vertices of the icosahedron, and generally remain intact when purified. WTV, the type member, has three protein shells: an outer amorphous layer made up of two proteins, an inner capsid made up of two proteins, and a smooth core made up of three proteins that is about 50 nm in diameter (Fig. 1(C)). Each virion contains a single full-length copy of the genome. Phytoreoviruses have 12 segments of linear dsRNA ranging in size from approximately 1 to 4.5 kbp, with a total genome length of approximately 25 kbp (Fig. 4) and a G þ C content of 41%–48%. Each segment of RDV contains a single ORF except for segments 11 and 12, which contains two ORFs. Table 5 summarizes the organization of the RDV dsRNA segments and the putative function of the encoded proteins.

Plant Reoviruses (Reoviridae)

Table 4

549

Genome organization of RRSV and predicted gene function

Segment Size (bp) Protein nomenclature

Predicted MW (kDa)

Apparent MW (kDa)

Predicted function (location)

S1 S2 S3 S4

3849 3810 3699 3823

P1 P2 P3 P4a P4b

137.7 133.1 130.8 141.4 36.9

137 118 130 145

Virion core associated (B spike) Inner core capsid, putative guanylyl-transferase (core capsid) Major core capsid (core capsid) RdRp Unknown

S5 S6 S7 S8

2682 2157 1938 1814

P5 Pns6 Pns7 P8 P8a P8b

91.4 65.6 68 67.3 25.6 41.7

90

Capping enzyme Non-structural protein, VSR, cell-to-cell movement Non-structural protein, NTP-binding protein (unknown) Precursor polyprotein/protease Spike Major capsid protein

S9 S10

1132 1162

P9 Pns10

38.6 32.3

37 32

66 67 47 44

Vector transmission, mild silencing suppressor activity (spike) Non-structural protein associating with a mitochondrial membrane component, oligomycin-sensitivity conferral protein (OSCP), to facilitate virus propagation in the vector

Fig. 4 Genome organization of Rice dwarf virus (RDV) containing 12 dsRNA segments. Each segment contains one ORF except for Seg 11 and 0 0 Seg 12 which contain two ORFs. The green arrows indicate the conserved 50 terminal sequence (5 GG(U/C)A–3 ) while the red arrows indicate the 50 30 0 conserved 3 terminal sequence ( –(U/C)GAU ). The red segment encodes the RdRp. The green segments encode the structural proteins while the yellow segments encode non-structural proteins.

Replication and Gene Expression The replication and gene expression of plant-infecting reoviruses is thought to be similar to that of other reoviruses. The best described of these is Bluetongue virus (BTV), the type member of the genus Orbivirus. If the BTV model is accurate for the plantinfecting reoviruses, replication occurs after virions (or viral cores) are delivered into the host cell. Replication is initiated when the viral capsid layer is removed and the transcriptionally active core enters the cytoplasm. The viral genome (10–12 segments) remains packaged in the central cavity of the viral core to ensure that host cell defense responses to dsRNA are not activated. The core is biochemically active with an RNA-dependent RNA polymerase (RdRp), a capping enzyme, and a helicase. The viral core contains a number of channels, the largest of which is at the 5-fold axis of the icosahedral structure. Smaller channels allow the entry of nucleotides, which are required for transcription, into the core. The large channel is located adjacent to the replicase/ transcriptase complex, which through its helicase activity executes the unwinding and rewinding of the dsRNA genome during transcription of the negative (antisense) RNA strand. The newly formed positive-strand (sense) mRNA molecules are modified to form a Cap 1 structure [7mGpppG(2-Om)…] by the guanylyl-transferase, nucleotide phospho-hydrolase, and transmethylase

550

Table 5

Plant Reoviruses (Reoviridae)

Genome organization of RDV and gene function

Segment

Size (bp)

Protein nomenclature

MWa

Function (location)

S1 S2

4423 3512

P1 P2

170 130

S3 S4

3195 2468

P3 Pns4

110 83

S5 S6

2570 1699

P5 Pns6

89 56

S7 S8 S9 S10

1696 1427 1305 1321

P7 P8 P9 Pns10

58 43 49 35

S11

1067

Pns11a

23

Pns11b

24

RdRp (core) Capsid structural protein, essential for vector transmission and symptom formation in plant host (outer capsid) Major core protein (core capsid) Phosphorylated nonstructural protein, associated with large cytoplasmic fibrils to form novel mini-tubules in infected cultured leafhopper cells, essential for viral infection in its insect vector (viroplasm, at the early stages of infection; amorphous or fibrillar inclusions, at the later stages of infection) Guanylyltransferase (core) Nonstructural protein, viroplasm matrix protein in insect vector cells, involved in cell-to-cell movement and nuclear acid binding in plant host RNA-binding protein (core) Major outer capsid protein Structural protein (outer capsid) Nonstructural protein, VSR in plant host, involved in intercellular transport in insect vector cells Nonstructural protein, viroplasm matrix protein, involved in nuclear acid binding protein Non-structural protein, enhancing the enzymatic activity of rice S-adenosyl-Lmethionine synthetase, triggering ethylene production in rice in order to achieve efficient infection

Pns12

34

Pns12OPa Pns12OPb

8 7

S12

1066

Nonstructural protein, essential for viroplasm formation, viral replication, and infection in insect vector

a

MW determined by SDS-PAGE (kDa).

activities of the capping enzyme and are then extruded through the major pore into the cytoplasm, where they can be translated to produce viral proteins. Non-structural viral proteins form viroplasm. The viroplasm is the site of most of the mRNA synthesis, genome replication, and assembly of core proteins. The various species of mRNAs synthesized in the viroplasm display in nonequimolar ratios. However, the dsRNA genome segments are usually packaged in exactly equimolar ratios (i.e., one copy of each genome segment per particle), which indicates that the selection of viral mRNAs for packaging is highly specific; recognition signals on each mRNA species may be related to this process. Once a copy of an mRNA is inside a new viral core, the corresponding negative strand is synthesized, completing the replication of the dsRNA genome. The progeny viral core containing dsRNA then moves to the periphery of the viroplasm, where capsid proteins are assembled to form the complete new viral particle and are thought to stop further mRNA synthesis. Like the parental-virus-derived smooth cores (without a capsid layer), progeny viral cores also synthesize mRNAs, providing an amplification step during replication. In recent years, researchers have proposed a new model for the genesis and maturation of viroplasms induced by SRBSDV in insect vector cells. The SRBSDV viroplasm consists of a granular matrix formed by the non-structural protein P9 and a filamentous matrix formed by the non-structural protein P5. The progeny viral cores and mature virions assemble in the filamentous matrix. The new viral mRNAs synthesized by the progeny viral cores and virions within the filamentous matrix are then transported to the granular matrix by association with SRBSDV non-structural protein P6. As a result, viral mRNAs accumulate in the granular matrix, and these viral mRNAs may either migrate to the adjacent cytoplasm to synthesize viral proteins that promote further maturation of viroplasms or migrate back to the filamentous viroplasm matrix and be packaged into empty core particles, which are assembled and aligned along the filaments within the filamentous matrix to form viral-RNA-containing core particles. The subsequent minusstrand RNA synthesis and outer capsid protein assembly processes also take place in the filamentous matrix, leading to the maturation of a new virion. Thus, the P9 formed granular matrix plays important roles in viral RNA and protein production and filamentous matrix maturation. Compared to the globular structures of phytoreovirus and oryzavirus viroplasms, the filamentous matrix of the fijivirus (SRBSDV) viroplasm gives it a much larger surface-area-to-volume ratio, which leads to produce more progeny virions in the insect vectors. The control of gene expression for a multi-segmented genome is complex and not fully understood. Each genome segment contained within the viral core is associated with a single replicase/transcription complex, located adjacent to the major pore in the vertices of the icosahedron, and is transcribed separately to make full-length positive-strand RNA copies. The location of the replicase/transcription complex also restricts the number of genome segments to a maximum of 12. These 10–12 mRNAs are produced in different molar amounts based largely on segment size, resulting in more copies of the smaller mRNAs. The conserved 50 - and 30 terminal sequences of the RNA segments may function as recognition signals for the replicase/transcription complex and be involved in the efficient initiation of viral mRNAs translation, providing some control over the expression levels of individual virus genes. In addition, the conserved and non-hybridized 30 -terminal sequences have been shown to be required for efficient negative-strand synthesis. A third level of control

Plant Reoviruses (Reoviridae)

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results from the presence of multiple or overlapping ORFs on one mRNA strand that are translated at different efficiencies. Lastly, some ORFs encode a polyprotein that must be processed to form functional proteins.

Distribution Plant-infecting reoviruses are seasonally distributed as a result of plant host/crop cycles and abundance of insect vectors. They can overwinter in certain weed crops, in autumn-sown cereals, and in diapausing insect hosts. With SRBSDV, for example, the distribution of the virus is consistent with the migration and existing ranges of its main host vector, the white back planthopper (WBPH; Sogatella furcifera). In Southeast Asia, WBPH is a long-distance migratory rice pest. In most rice-growing areas in China, no winter rice is sown, with the exceptions being the southwestern province, Yunnan and Hainan. These two areas have therefore become the major WBPH overwintering regions in China. In early spring, viruliferous WBPH begin their migration northward with the help of the northeastward air flow, and by late June or early July, they can reach as far as Japan. As a result, viruliferous WBPH transmit the virus throughout its journey, in regions including Guangdong, Guangxi, Hunan, Jiangxi, and Fujian provinces in China. Moreover, viruliferous WBPH also lay eggs in the newly colonized areas, and the next generation of WBPH propagate on the infected rice seedlings, acquire viruses, become viruliferous, disperse, and induce new disease outbreaks. In the autumn, the viruliferous WBPHs then return to their ideal overwintering region, Yunnan and Hainan provinces, when the monsoon winds shift direction. Plant reoviruses have been identified from every continent, but some genera are more widespread than others. Fijiviruses are the most widely distributed, which is not surprising given that they are the most numerous, although they are absent from North America and India. Fijiviruses have been reported to occur in Africa, Europe, South America, Asia, Australia, and the South Pacific Islands. Oryzaviruses have only been isolated from the Indian subcontinent and Asia, while phytoreoviruses have been isolated from North America, Asia, and Africa.

Host Range and Virus Transmission Fijiviruses The genus Fijivirus contains nine species whose members infect a range of monocotyledonous plants of the families Gramineae and Liliaceae. Common natural plant hosts include Avena sativa, Oryza sativa, Saccharum officinarum, and Zea mays among the Gramineae and Allium sativum among the Liliaceae, although this host range can be extended significantly by experimental virus infection. Viruses are transmitted by delphacid planthoppers (Table 1). Viruses can be acquired in juvenile plant growth stages, replicate in the vector, and, following a latent period of at most 2 weeks, are transmitted to plants in a persistent manner. No transovarial or seed transmission of viruses has been reported. In addition to transmission by insect vectors, mechanical transmission of the virus to susceptible hosts has been achieved for some fijiviruses with difficulty. In 2017, researchers reported a novel method for transmitting SRBSDV to rice, the bud-cutting method. With this approach, the 1.0-cm pregerminated rice bud was cut at a 451 angle approximately 0.5 cm from the base of the bud. The seedlings were then soaked in a crude SRBSDV extract obtained from SRBSDV-infected rice samples with a phosphate-buffered saline and then incubated in the dark at 28–301C for 3 days. Lastly, the rice seedlings were planted in a soil matrix and incubated under growing conditions appropriate for rice. After 25 days, the rice seedlings displayed disease symptoms similar to those of rice seedlings infected with WBPH, and intact SRBSDV virions were isolated from the seedlings. The transmission efficiency with this method was more than 80%.

Oryzaviruses The genus Oryzavirus contains two species whose members infect monocotyledonous plants of the family Gramineae. Common plant hosts include O. sativa and Echinochloa crus-galli. However, this natural host range can be extended by experimental virus infection to include other economically important species such as Hordeum vulgare, Triticum aestivum, and Zea mays. Virus is transmitted by delphacid planthoppers (Table 1). An acquisition period of 3 h is required followed by a 9-day latent period prior to transmission at all life stages in an intermittent manner. No transovarial transmission or mechanical transmission of virus has been reported.

Phytoreoviruses The genus Phytoreovirus contains two species whose members infect monocotyledonous plants of the family Gramineae and one species that infects dicotyledonous plants. Common plant hosts include the Gramineae O. sativa and the dicot Melilotus officinalis. However, the natural host range of the dicot-infecting WTV can be extended significantly by experimental virus infection. Virus is transmitted by cicadellid leafhoppers (Table 1). Virus can be acquired after a short feeding period, replicates in the vector, and, following a 10- to 20-day latent period, is transmitted to plants throughout the life of the vector. Transovarial transmission of virus has been reported. Attempts to transmit the virus to susceptible hosts by mechanical methods have been unsuccessful.

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Pathogenicity The pathogenicity of plant reoviruses is particularly interesting as most viruses replicate in both insects and plant hosts. Most do not appear to cause any disease in the insect host, so that their pathogenicity is restricted to the plant host. The pathogenicity of fijiviruses varies considerably. Fiji leaf gall disease (caused by FDV) is reported to cause losses of up to 90% in susceptible sugarcane varieties, while NLRV has no known plant host and, therefore, no known pathogenicity. Oryzaviruses can also cause important yield losses. Rice ragged stunt disease (caused by RRSV) is reported to cause losses of generally 10%–20%, but sometimes as high as 100% in severe infections of susceptible varieties. The pathogenicity of phytoreoviruses is much milder, although rice dwarf disease (caused by RDV) can have severe effects, as infected plants often fail to bear seeds. Furthermore, different strains of RDV–O, D84, and S–cause diseases of varying severity: RDV-S causes the severest symptoms and RDV-O the mildest. Although many aspects of the molecular bases for pathogenicity remain opaque, we now know a small part of the processes leading to rice dwarf disease symptoms. As mentioned before, RDV-infected plants exhibit disease symptoms that include dwarfing, short crown root, and increased tiller number. Some researchers have found this phenomenon similar to the phenotype observed when plants lose their sensitivity to auxin signaling. After meticulous experimentation, researchers finally determined that the symptoms are caused by the RDV capsid protein P2, which disrupts interactions within the auxin coreceptor complex, leading to the plant’s failing to perceive auxin. Besides this, viruses have evolved many other mechanisms to counteract plant antiviral strategies, most notably through the expression of viral suppressors of RNA silencing (VSRs). VSRs take advantage of RNA silencing and plant hormone pathways to destroy plant fortifications. As another example, the RDV nonstructural protein Pns11 can manipulate the plant ethylene biosynthesis process by enhancing the enzymatic activity of S-adenosyl-L-methionine synthetases, resulting in higher ethylene level in plants and making plants more susceptible to RDV infection. It is also has been demonstrated that RRSV infection mediates disease symptoms via suppression of jasmonic acid-associated antimicrobe signal through the microRNA319-TCP21 regulatory module. Furthermore, the pathogenic effects of different plant-infecting reoviruses can influence each other. RRSV is widely distributed in Eastern and Southeastern Asia, but until recently had not caused serious crop losses in China owing to its low incidence. Lately, as SRBSDV has spread, RRSV has become more and more common in China and has been frequently found coinfecting plants with SRBSDV. Researchers propose that the recent increase in RRSV incidence in China is partially due to synergism between SRBSDV and RRSV because they have found that not only do rice plants coinfected with both viruses exhibit higher virus titer and enhanced and earlier disease symptoms, but also the respective insect vectors of SRBSDV and RRSV acquired the viruses from the coinfected plants at higher rates than from singly infected plants.

Diagnosis and Control Diagnosis of plant reovirus infections can be done on the basis of disease symptoms or through the use of molecular tests. Symptoms vary between virus/host complexes but commonly include dark-green coloration, suppression of flowering, plant stunting, increased numbers of side shoots, and vein swelling or gall formation derived from phloem tissue on the backs of leaves. Given the variability in time to symptom expression and symptom severity, alternative tests are often used. Molecular and serological tests have been developed to assist in the diagnosis of viral infection in non-symptomatic plant material and vector insects. Serological tests are usually in enzyme-linked immunosorbent assay (ELISA) format and rely on polyclonal antisera raised against virions or expressed viral proteins. For example, antibodies against glutathione-S-transferase–NS7 fusion protein are useful in ELISAs for the detection of RRSV in infected plants and insect vectors, and a rapid, highly reliable, sensitive, and specific dotformat ELISA has been developed for SRBSDV detection using anti-SRBSDV rabbit antiserum. Recently, molecular tests such as reverse transcription–polymerase chain reaction (RT-PCR), which are faster and more specific than serology, have become the most common diagnostic methods. As increasing numbers of plant reovirus genomes are sequenced, species-specific primers are now becoming commonly available. A simple, rapid, sensitive, so-called multiple (m) RT-PCR method, which uses a mixed set of primers specific for Rice stripe virus (RSV), RBDSV, and SRBSDV together, can efficiently detect all three viruses simultaneously, and a one-step mRT-PCR assay has been established for simultaneous detection of RSV, RBSDV, and RDV. Meanwhile, a reverse transcription loop-mediated isothermal amplification (RT-LAMP) assay has been developed for the detection of RBSDV, RDV, RGDV, and RRSV, which can obtain a diagnosis within 2 h when combined with RNA rapid extraction. Control strategies for plant reoviruses can be focused on either the host plant or the insect vector. Plant-based control through breeding to develop resistant plant species is most commonly utilized in combination with the removal of susceptible varieties and infected plants that provide a source of inoculum. This approach has provided robust control of RDV in rice and FDV in sugarcane. Genetic engineering of plant hosts has also been explored as an alternative control strategy. Pathogen-derived resistance approaches directed at either coat proteins or other viral genes to control RDV, RRSV, and FDV have proved less successful than those used to control other RNA plant viruses. Control of insect vector populations with insecticide has provided some additional disease control. Unfortunately, chemical control appears to be of limited use in cases of high vector pressure. The current combination of diagnosis and control measures is already relatively effective and has resulted in reduced disease incidence and impact. Control measures based on protecting plants from WBPH, including seedbed coverage, chemical seed treatments, and chemical spraying of seedlings, have been demonstrated to be useful in controlling SRBSDV in China.

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Despite breakthroughs over the years, such as the discovery that RDV can use actin-based tubule motility to overcome transmission barriers and exploit the ancient oocyte entry path of the bacterial symbiont in leafhopper, many gaps in our understanding of transmission mechanisms remain. However, this situation may improve in the future as advances in genome sequencing, reverse genetics systems, genome editing, and molecular technologies help provide more information on virus infection and replication and, thus, new resistance targets.

Future Although our understanding of plant reoviruses is expanding, many of the molecular and biological properties of these viruses are still unknown. The complete sequence information now available for a number of these viruses will allow the production of cDNA probes to further elucidate the viral infection and replication processes in both plant and insect cells. The potential to produce infectious clones also holds promise for detailed studies of both plant and insect host ranges and of the methods of resistance employed by non-host species. This information, combined with knowledge gained from comparisons with animal reoviruses, may assist the further development of control strategies for diseases caused by these viruses in plants.

Further Reading Bamford, D.H., 2000. Virus structures: Those magnificent molecular machines. Current Biology 10, R558–R561. King, A.M., Adams, M.J., Carstens, E.B., Lefkowitz, E.J., 2011. Virus Taxonomy – Classification and Nomenclature of Viruses: Ninth Report of the International Committee on the Taxonomy of Viruses. San Diego, CA: Elsevier Academic Press. Hull, R., 2002. Matthews' Plant Virology. London: Academic Press. Mertens, P., 2004. The dsRNA viruses. Virus Research 101, 3–13. Mertens, P.P.C., Diprose, J., 2004. The bluetongue virus core: A nano-scale transcription machine. Virus Research 101, 29–43. Wei, T., Li, Y., 2016. Rice reoviruses in insect vectors. Annual Review of Phytopathology 54, 99–120. Miyazaki, N., Nakagawa, A., Iwasaki, K., 2013. Life cycle of phytoreoviruses visualized by electron microscopy and tomography. Frontiers in Microbiology 4, 306.

Relevant Websites http://www.dpvweb.net/ DPVWeb Home Page. https://talk.ictvonline.org/ International Committee on Taxonomy of Viruses (ICTV). https://viralzone.expasy.org/ ViralZone root - ExPASy.

Plant Resistance to Geminiviruses Basavaprabhu L Patil, ICAR–Indian Institute of Horticultural Research, Bengaluru, India Supriya Chakraborty, Jawaharlal Nehru University, New Delhi, India Henryk Czosnek, The Hebrew University of Jerusalem, Rehovot, Israel Elvira Fiallo-Olivé, Institute for Mediterranean and Subtropical Horticulture “La Mayora”–Spanish National Research Council–University of Malaga, Algarrobo-Costa, Málaga, Spain Robert L Gilbertson, University of California, Davis, CA, United States James Legg, International Institute of Tropical Agriculture, Dar-es-Salaam, Tanzania Shahid Mansoor, National Institute for Biotechnology and Genetic Engineering, Faisalabad, Pakistan Jesús Navas-Castillo, Institute for Mediterranean and Subtropical Horticulture “La Mayora”–Spanish National Research Council–University of Malaga, Algarrobo-Costa, Málaga, Spain Rubab Z Naqvi and Saleem U Rahman, National Institute for Biotechnology and Genetic Engineering, Faisalabad, Pakistan Francisco M Zerbini, Federal University of Viçosa, Viçosa, Brazil r 2021 Elsevier Ltd. All rights reserved.

Nomenclature AGO Argonaute protein CAS CRISPR-associated proteins CMD Cassava mosaic disease CMG Cassava mosaic geminiviruses CRISPR Clustered regularly interspaced short palindromic repeats DCL Dicer-like dsRNA Double-stranded ribonucleic acid GM Genetically modified crops gRNA Genomic RNA HR Hypersensitive response LRR Leucine-rich repeat mRNA Messenger RNA

Glossary Allele Allele is a distinct form of a gene at a particular locus, that shows different effect on the phenotype from the other forms of alleles. Betasatellite A type of circular single-stranded satellite DNA associated with begomoviruses. CRISPR-Cas The acronym for Clustered Regularly Interspaced Short Palindromic Repeat – CRISPRassociated protein, an adaptive immune system used by bacteria and archaea against viruses and mobile genetic elements, which has been adapted for genome editing in eukaryotes. Dominant allele An allele expressing its phenotypic effect even when having a recessive allele as its counterpart in a heterozygous system. NB-LRR protein Plant disease resistance proteins containing a central nucleotide-binding domain (NB) and C-terminal leucine-rich repeat (LRR) domain. Also known as NBS-LRR or NB-ARC-LRR proteins. Quantitative trait loci Corresponds to genomic regions that is associated with the phenotypic variation of a quantitative trait. Recessive allele An allele whose phenotypic effect is not expressed in a heterozygote.

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NGS Next generation sequencing nt Nucleotide(s) QTL Quantitative trait loci R Dominant resistance RDR RNA-dependent RNA directed polymerase RdRp RNA-dependent RNA polymerase RGAs Resistance gene analogs RIL Recombinant inbred line RISC RNA-induced silencing complex RNAi RNA interference SNP Single-nucleotide polymorphism ssRNA Single-stranded ribonucleic acid VIGS Virus-induced gene silencing

Resistance gene Plant gene conferring resistance to a specific pathogen, that possess a corresponding avirulence gene. RNA interference A broad antiviral defense system used by plants, animals, and fungi. It recognizes double-stranded RNA as a cue of viral presence and induces degradation or silencing of viral transcripts or genomes. Transcriptional gene silencing A mechanism of gene silencing involving decreased RNA synthesis because of promotor methylation. Virus immunity Genetic mechanism by which plants are unable to support replication of a virus, and consequently where the virus cannot be isolated by any mean from the infected plant. Virus resistance Genetic mechanism by which plants are able to defend themselves against a viral invasion (by one or the other mechanism) that leads to reduced viral replication levels. Virus tolerance Genetic mechanism by which plants are infected and viral replication/infection occurs at wild type levels but without any visual symptoms. Virus-induced gene silencing It is a technology that exploits an RNA-mediated antiviral defense mechanism.

Encyclopedia of Virology, 4th Edition, Volume 3

doi:10.1016/B978-0-12-809633-8.21565-3

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Introduction Domestication of crops from their wild relatives has resulted in the loss of many resistance genes because these are often genetically linked to unwanted traits (such as low yield, poor flavor, small size). With the commercialization of agriculture and increasing production of crop plants in monoculture, genetic diversity has dwindled, which has made crops susceptible to a variety of pathogens. Therefore, breeding for resistance has consisted in reintroducing these resistance genes in the susceptible domesticated crop by crossing and selection. The wild relatives or ancestors of these crop plants present in their centers of origin or diversity are the best sources of resistance genes (R genes). Pathogen resistance in crop plants can also be generated by either natural or induced mutations. For example, Rice yellow mottle virus (RYMV) resistance in cultivated rice spontaneously appeared under RYMV pressure. Cassava varieties such as TME3 are also probably mutants which were selected under cassava mosaic disease pressure. Geminiviruses are single-stranded, circular DNA viruses that cause economically important diseases in a wide range of crop plants across the globe. Geminiviruses are characterized by small geminate particles (18
30 nm) containing either one (A component, monopartite) or two (A and B components, bipartite) single-stranded circular DNA molecules of B2.7 kb and sometimes associated with satellite DNA molecules of B1.4 kb. Around 460 distinct species and a larger number of strains belong to the family Geminiviridae. In this article, we briefly discuss the resistance genes and the sources of resistance against the most important geminiviral diseases in cassava, tomato, bean, maize, and cotton.

An Overview of Natural Resistance to Viruses in Plants Depending on the number of genes involved in providing the resistance phenotype, the host resistance can be classified into monogenic, digenic, or polygenic. Many of the monogenic and digenic types of resistance are qualitative in nature, with phenotypes showing complete absence of pathogen in the host plant. Whereas the polygenic resistance is manifested by several genes. A quantitative trait locus (QTL) is a region of DNA that is associated with a specific phenotypic trait, that varies in degree and can be attributed to polygenic effects, i.e., the product of two or more genes, and their interaction with the environment. Several such QTLs could be associated with one single trait. Such QTLs are mapped by genetic markers by using recombinant inbred lines (RILs) that are derived by crosses between parents with two contrasting resistance phenotypes. The inheritance pattern of the resistance trait can be dominant or recessive in nature and half of the virus resistance traits known hitherto are recessive in nature. Plant R genes confer resistance to plant pathogens including plant viruses and each R gene confers resistance to a specific pathogen triggered by a specific avirulence (avr) gene or an effector protein. More than 80% of the R genes studied so far are monogenic in nature and one third of the R genes are tagged by molecular genetic markers. R-genes can be either dominant or recessive in nature. Studies done so far have shown that dominant R genes are of the NB-LRR (Nucleotide-Binding-Leucine-Rich Repeat) type and that recessive ones encode for translation initiation factors. Dominant gene resistance is usually associated with HR while recessive genes function at single cell level. R genes are known to show two unique patterns of clustering: in one case R genes with similar inheritance and resistance phenotypes are clustered at one locus, while in another case the R genes for virus resistance are clustered with unrelated R genes. Most of the recessive R genes identified to date are against potyviruses. Although there are no reports of R genes showing resistance to geminiviruses, there are reports of genetic mapping of the resistance loci through molecular markers. There are also reports demonstrating the interaction of geminivirus proteins with the trans-membrane receptor kinases and components of brassinosteroid signaling. It was recently shown that silencing of a Permease I-like protein gene transmembrane transporter in tomato using the virus-induced gene silencing (VIGS) vector system, rendered it susceptible to Tomato yellow leaf curl virus. In response to the pathogen recognition by the R gene, changes in ion fluxes are observed, leading to activation of signaling pathways, alteration in transcriptional profiles, production of reactive oxygen species (ROS) and generation of nitric oxide (NO). Gene silencing is another important mechanism which plants have evolved to defend against viruses. In plants, posttranscriptional gene silencing (PTGS) acts as a natural anti-viral defense system and is considered as part of the plant’s innate immune system. RNA silencing involves suppression of gene expression by sequence-specific interaction at the transcriptional or post-transcriptional level in diverse eukaryotes. Double stranded RNA (dsRNA) is the trigger for gene silencing, which involves a chain of events to produce small interfering RNA (siRNA) through a diverse range of enzymes and its complexes, including the RISC-complex (RNA induced silencing complex) as well as DCL (Dicer-Like) and RDR (RNA dependent RNA polymerase) proteins. In virus-infected plants, RNA silencing is initiated by the double-stranded (ds) RNA that can be a viral replication intermediate or by ‘aberrant’ RNAs, or single-stranded RNA (ssRNA) that is converted to dsRNA by host-encoded RNA-dependent RNA polymerase (RdRP). The dsRNAs can also be formed because of bi-directional transcription of these viruses with transcripts occurring from opposite polarity overlapping at their 30 -ends, which partly explains how these geminiviruses induce PTGS in infected plants. Alternatively, the early and abundant transcripts of the AC1 gene of these geminiviruses can serve as the template for the host RdRP to produce dsRNAs. Yet another possibility is the fold-back structures of geminivirus transcripts which can serve as a template for DICERs to cleave at specific locations and produce siRNAs. In plants, some virus-host interactions naturally lead to host recovery that are similar to RNA-mediated virus resistance. This symptom recovery phenomenon is also reported for some of the geminiviruses (Fig. 1). This recovery phenomenon is associated with the production of virus-derived siRNAs, that later become abundant in the newly developed symptom-less recovered leaves. This increase in virus-derived siRNA accumulation is accompanied by a reduction in the levels of both viral DNA and mRNA accumulation.

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Fig. 1 Symptom recovery in Nicotiana benthamiana plants infected by two distinct cassava mosaic geminiviruses (ACMV and EACMKV), as a result of gene silencing.

Since RNA silencing and R gene signaling are both components of the plants innate immunity, it will be interesting to know if there is any cross-talk between these two pathways through one of the systemic signaling systems such as the salicylic acid pathway. It has been shown that the RdRPs NbRdRp1m, NtRdRp1 and RDR1 are all inducible by both salicylic acid and certain viruses, although recent studies failed to find any overlap between the DCL endoribonucleases involved in silencing and the SAinduced resistance to positive-sense RNA viruses. More recently it has been shown that the virus resistance induced by NB-LRR proteins involved Argonaute4-dependent translational control. It has also been shown that the viral silencing suppressor HC-Pro enhances R-gene mediated resistance to viruses and this resistance is weak at higher temperatures. Geminiviruses are known to be associated with DNA satellites which modulate the viral symptoms and the betasatellites in particular are known to enhance the symptoms through their PTGS suppressor proteins (ßC1). In this article we discuss in greater detail and on a case by case basis the principal sources of resistance for the major geminiviral diseases of the world (Table 1).

Resistance to Cassava Mosaic Geminiviruses It was recognized from the earliest period of research on cassava mosaic geminiviruses (CMGs) (Family Geminiviridae; Genus Begomovirus) that resistance would be the most effective method for controlling the cassava mosaic disease (CMD) caused by these viruses. The team responsible for the first biological characterization of CMGs in the former Tanganyika in the 1930s realised that wild relatives of cassava (Manihot esculenta Crantz) would provide an important potential source of resistance genes. Resistant progeny from Manihot esculenta
Manihot glaziovii (Müll-Arg.) crosses were backcrossed with cultivated cassava to produce the first CMD-resistant cassava varieties. Material from this initiative was subsequently used in the 1970s to extend the development of CMD resistance within the cassava breeding program of the International Institute of Tropical Agriculture (IITA). A recurrent selection approach was taken with the primary source of CMD resistance being the cassava clone 58308. It had been suggested since the 1950s that the resistance source derived from the M. glaziovii cross was polygenic, but this was also shown to be recessive in the early 1970s. During this period, evaluations of resistant varieties were undertaken in West, Central and East Africa, as well as in south India. Resistance was shown to be effective in all of these regions, providing a strong demonstration of the broad activity of the resistance source as well as its likely durability. We now know that there is significant virus diversity across these regions (ACMV, EACMV-like viruses, ICMV and SLCMV), which confirms the validity of the earlier conclusions about the broad-based activity of the M. glaziovii-derived resistance. This polygenic or quantitative recessive resistance source is now referred to as CMD1. In the 1980s and 1990s in Africa there were extensive trial-based epidemiology studies of CMGs, several of which compared the responses of varieties with contrasting levels of resistance. Different components of resistance were identified, including: resistance to infection, resistance to virus multiplication, resistance to virus movement (leading to incomplete systemicity), and resistance of normal plant function to the effects of virus infection. These were assessed through measurements of virus titre, incidence of infected plants, severity of symptoms of infection, distribution of symptoms through the plant, and yield of healthy versus infected plants. Strong correlations were demonstrated between each of these variables. Resistant varieties were therefore infected less, had

Plant Resistance to Geminiviruses

Table 1

List of geminivirus resistant genes/loci and their characteristics for the major geminivirus diseases

Target virus (es)

Target host Gene/Locus

CMGs

Cassava

CMD1 CMD2 CMD3

TYLCVs

Tomato

Ty-1 Ty-2 Ty-3 Ty-4 ty-5 Ty-6 –

ToLCV

tgr-1

ToCMoV

tcm-1

S. lycopersicum

Nature/Location of resistance gene/locus

Source of resistance

Type of resistance

Manihot glaziovii West African Cassava landraces Cassava genotype TMS97/2205

Polygenic recessive Single dominant gene Polygenic or quantitative

RdRp NB-LRR RdRp – Pelota – –

Solanum chilense S. habrochaites S. chilense S. chilense S. peruvianum? S. chilense S. habrochaites

Partial dominant Dominant Partial dominant Partial dominant Recessive Partial dominant Oligogenic (Complete resistance)

Cell-to-cell movement

S. chilense

ToLCNDV

Cucurbits

CmoCh08G001490 BZIP

Cucurbita moschata

Quantitative

PYMV

Potato

–

S. pimpinellifolium

Bigenic

BCTV

Common bean

Bct-1

Red Mexican common bean varieties Arabidopsis thaliana (3 ecotypes, Ms-0, Pr-0, and Cen-O)

Single dominant gene

–

– CaLCuV and TYLCV BGYMV

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Common Bean

Monogenic

gip-1

Arabidopsis thaliana (Pla-1 ecotype)

bgm-1

Phaseolus vulgaris (Red S)

Recessive

bgm-2 bgm-3 Bgp-2

Phaseolus coccineus (scarlet runner Bean)

Dominant

Non-TIR-NBS-LRR LRR-RP (on chromosome 18)

Vigna mungo Glycine max

Dominant Dominant

Chromosome-1 SNP on: Chromosome-3 Chromosome-3 Chromosome-7 Chromosome-9

Zea mays (CML206
CML312) Zea mays (TZIL07A01005
TZIL07A01322)

Partial dominant effect Dominant Additive Dominant Additive & dominant

MYMIV

Black gram CYR-1 Soybean Glyma18g02850

MSV

Maize

msv1 Q1 - PHM5502_31 Q2 - PZA02616_1 Q3 - PZA02872_1 Q4 - PHM1766_1

Abbreviations: CMGs, cassava mosaic geminiviruses; TYLCVs, tomato yellow leaf curl viruses; ToLCNDV, tomato leaf curl New Delhi virus; ToLCV, tomato leaf curl virus; ToCMoV, tomato chlorotic mottle virus; PYMV, potato yellow mosaic virus; BCTV, beet curly top virus; CaLCuV, cabbage leaf curl virus; BGYMV, bean golden yellow mosaic virus; MYMIV, mungbean yellow mosaic India virus; MSV, maize streak virus; NBS-LRR, nucleotide-binding site Leucine-rich repeat; LRR-RP, leucine-rich repeat receptor-like protein kinase.

less severe symptoms which were spatially limited in the plant, had lower virus titers and yielded more, demonstrating that each of these were expressions of the same resistance. Following the increasing accessibility of PCR and sequencing analyses from the late 1990s onwards, it became possible to examine the molecular characteristics of CMD-resistant cassava varieties. A key development came in 2002 when a novel source of resistance to CMGs was identified from West African cassava landraces. Molecular genetic mapping was used to identify a single dominant gene referred to as CMD2, and this development made it possible for breeding programs to speed up their resistance breeding work through marker-assisted selection (MAS). Several markers were identified which could pick out clones carrying CMD2 with a high degree of reliability. Moreover, similar approaches also led to the identification of a third resistance source – CMD3 – in the genotype TMS97/2205. This clone also carried CMD2, and it was observed that the combination of CMD2 and CMD3 gave the plants very high levels of resistance to CMD. An additional spin-off from the development of MAS was that the technique could be applied in regions where CMGs did not even occur, but where there was an interest in having strategic stocks of CMD-resistant germplasm. This would allow these regions to respond rapidly to any future invasive spread of CMGs. This activity was pioneered by the International Center for Tropical Agriculture (CIAT) in Colombia, to prepare for the potential future introduction of CMGs to Latin America.

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Fig. 2 (Top) CMD resistant cassava variety TME3 showing recovery from ACMV (2 weeks post inoculation) and EACMV (4 weeks post inoculation) infections in glasshouse experiments. Reproduced from Patil, B.L., Fauquet, C.M., 2015. Studies on differential behavior of cassava mosaic geminivirus DNA components, symptom recovery patterns, and their siRNA profiles. Virus Genes. 50 (3), 474–486. doi:10.1007/s11262-015-1184-y. (Bottom) CMD resistant and susceptible varieties of cassava cultivated in fields of Burundi.

In addition to extensive field evaluations for CMD resistance, targeted approaches such as microparticle bombardment using infectious clones have been used to screen for resistance to CMGs. In comparisons of ACMV and EACMV resistance, putative resistant accessions with CMD1, CMD2, or CMD3 were shown to have high levels of resistance to ACMV yet were severely affected by EACMV. The only exception to this was TMS97/2205 which was known to combine CMD1, CMD2, and CMD3. As a consequence of these and related findings, the primary objective of most cassava geminivirus resistance breeding programs is to combine these different resistance sources into varieties that also have favorable yield, organoleptic, and processing qualities. There has been limited progress to date in identifying specific genes associated with CMD1, CMD2, and CMD3. However, transcriptome data have revealed differences between CMG-susceptible (T200) and CMG-resistant (TME3; Fig. 2) cassava varieties. Moreover, resistance gene analogs (RGAs) were also shown to be expressed differentially during the process of symptom recovery (from CMD infection), suggesting that they may provide a complementary resistance mechanism to RNA silencing. Methylation of viral DNA has been shown in several systems to be an important resistance strategy employed by plants against geminivirus infection and has been demonstrated specifically for ICMV. However, although resistance sources such as CMD2 have been pinpointed down to chromosome and linkage group level through genome-wide sequencing and mapping studies, further research is required before a more comprehensive understanding can be obtained of the genetics of the responses of cassava plants to virus infection. While studies continue to expand knowledge about natural resistance to CMGs, there are widespread and highly successful programs being implemented to deploy CMD-resistant varieties (Fig. 2). These were disseminated throughout East and Central Africa to control the severe CMD pandemic of the 1990s/2000s with great success, and similar programs are now being used to address the more recent CMD pandemic in South-East Asia. Africa-derived CMD resistance has proven to be effective in controlling virus infection caused by the South Asian CMGs – ICMV and SLCMV – so they should also provide effective control for the currently spreading South-East Asian pandemic associated with SLCMV.

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Fig. 3 Resistant (green plants) and susceptible tomato plants (dead or yellow plants) from a tomato breeding field of a program for begomovirus resistance in Guatemala, tomato experimental field in Sanarate Guatemala, 2005. Funded by USAID-CDR. Courtesy: Luis Mejia, Douglas P Maxwell, Favi Vidavski, Henryk Czosnek.

Geminivirus Resistance in Tomato The Case of Resistance to Tomato Yellow Leaf Curl Virus Tomato crops can be infected by a large number of geminiviruses (family Geminiviridae) mainly belonging to the genus Begomovirus. Tomato yellow leaf curl disease, one of the most devastating virus disease affecting tomato worldwide, is caused by a complex of begomoviruses originated in the Middle East but expanded initially to the Mediterranean Basin and then to the Far East, the Caribbean, North America, and Australia. Tomato yellow leaf curl virus (TYLCV) was the first monopartite begomovirus characterized and the most successful in spreading the disease worldwide.

Breeding for TYLCV Resistance Breeding programs aimed at producing tomato varieties resistant to TYLCV started in the late 1960s and have expanded since (Fig. 3). These programs are based on the introgression of resistance found in some accessions of wild tomato species into the domesticated tomato (Solanum esculentum). Certain accessions of the wild tomato species S. chesmanii, S. chilense, S. habrochaites, S. peruvianum, and S. pimpinellifolium are resistant (with mild or no symptoms, but containing virus) to whitefly-mediated inoculation. The discovery of loci, and later on of genes, associated with TYLCV resistance, was facilitated by the development of saturated maps based on DNA polymorphism distinguishing resistance from susceptibility (RFLP, AFLP, SSR, SCAR, etc.). To date, six loci, coined Ty-1 to Ty-6, have been found to be associated with TYLCV resistance (Table 1). They have been mapped on different tomato chromosomes and five of them are independently inherited. The Ty-1 and Ty-3 loci originated from S. chilense accessions LA1969 and LA2779, respectively, and they were mapped to the long arm of tomato chromosome 6. Later, Ty-3 was shown to be allelic to Ty-1. Ty-2 was introgressed from S. habrochaites accession B6013 and was located on the long arm of chromosome 11. Ty-4 was introgressed from S. chilense accession LA1932 and mapped to the long arm of chromosome 3. Compared to other Ty genes, Ty-4 is less effective against TYLCV. ty-5, the only recessively inherited Ty gene known to date, was introgressed from S. peruvianum and mapped on chromosome 4. ty-5 has been reported from different sources including line TY172 (derived from four different accessions of S. peruvianum which were crossed with S. arcanum) and tomato cultivar Tyking. Ty-6 originated from S. chilense accessions LA1938 and LA2779 and was located on the long arm of chromosome 10. It has been shown that Ty-6 is effective, in addition to TYLCV, against Tomato mottle virus, a New World begomovirus. Pyramiding resistance from various wild tomato species in a single tomato line often broadens resistance. Today, most commercial tomato varieties resistant to TYLCV contain the Ty-1 gene. Upon whitefly-mediated infection in the field and greenhouse, the plants remain symptomless and their yield compares with that of non-infected plants. However, reports in 2008 indicated that Ty-1-mediated resistance could be broken under high virus pressure. Worse, it was recorded in 2016 in Jordan and Israel that Ty-1 resistant varieties suddenly became susceptible to TYLCV. Molecular cloning and sequencing indicated that the symptomatic plants were contaminated with the cotton leaf curl Gezira betasatellite, which in association with TYLCV, overcame the resistance based on Ty-1. These finding were later confirmed in the laboratory using cloned virus and betasatellite.

Characterization of Ty Genes Three of the Ty loci have also been associated to genes which have been recently cloned and characterized. Ty-1/Ty-3 were identified as coding for an RNA-dependent RNA polymerase (RdRp) belonging to the RdRp type which has an atypical DFDGD motif in the catalytic domain. It has been shown that Ty-1 confers resistance to TYLCV and other begomoviruses by enhancing transcriptional gene silencing through an increase in cytosine methylation of viral genomes.

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Ty-2 encodes a nucleotide-binding domain and leucine-rich repeat-containing (NB-LRR) gene, TYNBS1. Introduction of genomic fragments containing the TYNBS1 gene into susceptible tomato plants resulted in transgenic lines resistant to TYLCV. ty-5 is a loss-of-function mutant allele of the gene that encodes a messenger RNA surveillance factor Pelota (Pelo) located on chromosome 4. Pelota is involved in ribosome recycling phase of protein synthesis. The mutation in ty-5 is caused by a T-to-G transversion in the coding region. Ty-6 is located on chromosome 10, that compliments resistance conferred by both Ty-3 and ty-5 genes, thus providing resistance to both monopartite and bipartite begomoviruses.

Genetic Engineering for TYLCV Resistance In some cases, as mentioned above for Ty-2, host-derived resistance has been engineered into susceptible tomato plants. Nonetheless, most of the tomato transgenic plants with incorporated begomovirus resistance were produced by expressing viral genes. TYLCV genes or gene segments, whether functional or not, have been used to confer a narrow as well as a broad-range virus-specific resistance (pathogen-derived resistance). The expression of the TYLCV CP gene in an interspecific S. lycopersicum x S. pennellii hybrid was characterized by the late development of disease symptoms, often followed by their disappearance and recovery. The expression of the TYLCV Rep gene in tomato cultivars provided a virus-specific resistance. For example, tomato plants transformed with a truncated TYLCV Rep gene exhibited various degrees of resistance to TYLCV infection in the field in Florida. However, these TYLCV-resistant plants were susceptible to the bipartite begomovirus Tomato mottle virus conspicuous in Florida tomato fields. Similarly, tomato plants transformed with a 30 end-lacking Rep gene of Tomato yellow leaf curl Sardinia virus (TYLCSV, a monopartite begomovirus closely related to TYLCV) were resistant to TYLCSV from Italy but were susceptible to a cognate TYLCSV isolate from Spain. Resistance was also achieved by mimicking the plant RNA interference (RNAi) antiviral mechanisms relying on pathogen transcriptional (TGS) and post-transcriptional (PTGS) gene silencing. Small interfering RNAs (siRNA) directed against TYLCV genes were designed, based on hairpin RNAi (hpRNAi), and used to produce transgenic tomato plants. It was found that hpRNAibased resistance was highly sequence-specific. The first set of experiments using PTGS hpRNAi demonstrated that TYLCV and TYLCSV non-coding regions could trigger the accumulation of virus-specific siRNAs, followed by high levels of resistance against these two viruses. TYLCV Rep was also the target of hpRNAi. Homozygous transgenic tomatoes expressing hpRNAi TYLCV Rep were resistant to TYLCV in greenhouses and fields and carried large quantities of small RNAs. In addition, resistance was associated with changes in the general plant host transcriptome, mainly genes involved in biological regulation, metabolic and cellular processes such as catalytic and binding activities. The hpRNAi strategy has also been used against the IR, CP, V2, and Rep genes of TYLCV from Oman. The transgenic lines obtained exhibited various levels of resistance upon TYLCV agro-inoculation. The CRISPR-Cas system, which protects prokaryotes from potentially deleterious foreign DNA, is the latest tool in the molecular breeder’s panoply aimed at conferring TYLCV resistance. CRISPR-Cas was recently implemented to the making of virus-free crops, demonstrating that this technology can be applied to the molecular breeding of virus-immune tomatoes. Indeed, TYLCV resistance was conferred by editing the tomato genome. Targeting the TYLCV CP and Rep genes with Cas9 single RNA-guided DNA endonuclease resulted in stable resistance to the virus. The Cas9-treated tomato plants carried reduced TYLCV amounts and their progeny was resistant to TYLCV. Therefore, antiviral strategies based on genome editing have the potential to constitute an additional, maybe preferred, tool in the breeding of TYLCV-resistant tomato varieties.

Resistance to Beet Curly Top Virus Beet curly top virus (BCTV, genus Curtovirus, family Geminiviridae), is a phloem-limited virus that causes curly top disease in several economically important crops such as common bean, pepper, sugar beet, and tomato. This disease is more prevalent and is known to cause heavy economic losses in the western USA, north-central Mexico and countries of the Mediterranean Basin and the Middle East (e.g., Iran and Turkey). Under favorable environmental conditions the disease outbreaks are driven by populations of the beet leafhopper vector (Circulifer tenellus). Sources of curly top disease resistance have been identified in germplasm of common bean, flax, squash and sugar beet. However, the availability of commercially available resistant varieties that can be used in a management program for BCTV depends on the crop plant. Resistant varieties have played a major role in management of curly top in sugar beets and somewhat of a role in common bean, whereas there are no commercially available resistant varieties for pepper or tomato. In the early 1900s in the western USA, curly top disease emerged as a major constraint on sugar beet production, in some cases resulting in sugar refineries having to close. To search for resistance, mass selection from heavily infested fields led to the identification and release of the first curly top resistant sugar beet variety, US1, in 1933. This variety became widely grown in the western USA, despite some shortcomings. The identification and selection of resistant sugar beet varieties has been facilitated by the use of breeding plots, known as curly top nurseries, in which high populations of leafhoppers viruliferous for the most prevalent and virulent strains of BCTV are released. As a result of the high selection pressure in these nurseries, combined with the development of commercial hybrids and molecular breeding technologies, curly top resistance in sugar beet has advanced tremendously, despite this being a complex quantitatively inherited trait. Modern curly top resistant varieties have greater uniformity, higher yield and improved

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resistance. However, plants of these resistant sugar beet varieties will develop curly top symptoms and may suffer yield losses when infected at an early stage of development (two to four leaf stage). Therefore, resistant varieties need to be part of an IPM program that addresses the susceptibility of young plants, e.g., seed treatment with systemic insecticides. Common bean is another crop that can be heavily impacted by curly top in the western USA. Curly top resistance was identified in some common bean varieties (e.g., Red Mexican type). Genetic studies identified a single dominant gene (Bct-1) associated with curly top resistance in common bean. Breeding efforts led to the release of the commercial curly top-resistant varieties Great Northern UI 15 and Red Mexican UI 34. Subsequently, snap bean varieties with curly top resistance were developed and released. These varieties have provided acceptable yields under high curly top pressure. Now there is a need to develop curly top-resistant varieties for other classes of common bean. In terms of tomato and pepper, curly top resistance was not found in commercial cultivars. However, high levels of resistance were found in accessions of some wild species of tomato, including Solanum chilense, S. habrochiates, S. lycopersicoides and S. peruvianum. Unfortunately, introgression of this resistance into commercial tomato varieties has been difficult. However, as described in the case of resistance to TYLCV, a series of 6 loci (genes) conferring some degree of resistance to the virus, have been introgressed into tomato breeding lines. These lines have been used to generate commercial varieties with effective resistance against monopartite begomoviruses such as TYLCV. The mechanisms of Ty gene products determined to date, such as the Ty-1/Ty3 enhancing transcriptional gene silencing (TGS) and cysteine methylation of the viral genome, appear to be common to all geminiviruses rather than genus-specific. Therefore, BCTV agro-inoculation screening was used to assess the response of 15 breeding lines, having different combinations of Ty genes (e.g., Ty-1, Ty-2, Ty-3, ty-5, and Ty-6), kindly provided by the World Vegetable Center. The response was assessed based upon a 0–4 rating scale, with 0 ¼no symptoms to 4 ¼severe stunting and leaf curling and vein purpling. In screens with the BCTV-Svr-[US: SVR:Cfh], the susceptible control received a rating of 4.0, whereas 11 of the breeding lines had ratings o2.5, with three lines with rating of 1.5–1.7 (lines with Ty3 only, Ty3 and Ty2 and Ty5 and Ty6). In an equivalent screen with BCTV strain BCTV-LH-71-[US: Cal:10], which was associated with the 2013 curly top outbreak in processing tomatoes in California, only 6 of the Ty breeding lines had ratings o2.5, with two lines having ratings of 0.9 and 1.2 (both having the Ty2 and Ty3 combination). Together, these results indicate that pyramiding Ty genes in tomato can provide resistance to BCTV, consistent with these gene products targeting a general geminivirus property, such as replication or transcription. Importantly, the effectiveness of the resistance was not the same for two strains of BCTV. A similar result was reported in the reverse scenario in common bean, where the Bct-1 gene that confers qualitative (strong) resistance to BCTV, provided a high level of quantitative resistance to the bipartite begomovirus, Bean dwarf mosaic virus (BDMV). These results suggest the possibility of generating broad spectrum geminivirus resistance. To assess the potential for using Ty genes to generate a curly top resistant tomato variety, a breeding line was generated by screening progeny of crosses aimed at pyramiding the Ty-1, Ty-2, and Ty-3 genes by agro-inoculation. Plants with a resistance phenotype were then selected and further selection performed by selfing plants and screening progeny by agro-inoculation. The breeding line that was generated, named Line 20#12, carries the Ty-1, Ty-2, and Ty-3 genes and has moderate to strong resistance to curly top (Fig. 4). Moreover, the resistance in this line involves initial development of mild up-curling and vein purpling in the first true leaves, followed by a recovery phenotype in which all subsequently emerging leaves show no symptoms (although virus can be detected in these leaves). This resistance phenotype is consistent with TGS targeting the BCTV genome for methylation, reducing transcription and replication. Thus, these results suggest that pyramiding Ty genes in a commercial tomato background could be a strategy to generate commercial curly top-resistant varieties.

Geminivirus Resistance in Beans Geminiviruses cause severe diseases in common bean (Phaseolus vulgaris), and also in other species of Phaseolus such as lima bean (P. lunatus). In the Americas, the viral species involved are Bean golden mosaic virus (BGMV), Bean golden yellow mosaic virus (BGYMV) and Macroptilium yellow spot virus (MaYSV), all of which cause similar symptoms of severe yellow ("golden") mosaic. BGMV and MaYSV occur in South America (MaYSV has so far been only reported in Brazil), while BGYMV is present throughout Central America and the Caribbean. Many institutions in the Americas had breeding programs for resistance to golden mosaic disease in Phaseolus. The most successful program was conducted at the International Center for Tropical Agriculture (CIAT) in Cali, Colombia, starting in the mid-1970s. This program targeted BGYMV, although most of the relevant literature refers to BGMV, since the distinction between the two viruses was not completely clear at the time. Indeed, the fact that resistance to BGYMV was not successful against BGMV provided additional evidence for the distinction between the two viruses. A first source of resistance to BGYMV was identified in a black bean landrace named Porrillo Sintetico, originated from El Salvador (Central America). The analysis of segregating populations derived from crosses with this source indicated that resistance was quantitative. Resistance was partial and often broke down under high inoculum pressure. The red-seeded line DOR 364 provided additional quantitative resistance genes, and a number of cultivars containing these two sources, with resistance that was markedly superior to that conferred by either source alone, were released during the 1980s. During that decade, virtually all resistant cultivars that were deployed in Central America and the Caribbean derived from these two sources. These cultivars were largely responsible for the recovery of bean yields in that region. The main limitation with these sources of resistance was that all resistant cultivars produced black seeds, and many of the commercial common bean cultivars grown in disease-prone regions had different seed colors that were changed when crossed with

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Fig. 4 Advancement and characterization of tomato breeding lines for beet curly top virus resistance, by agro-inoculation of BCTV in glasshouse conditions at the University of California, Davis, CA, USA.

the black-seeded parental sources of resistance. Moreover, resistance was not complete and could still be overcome in years when conditions were highly favorable to the disease. An improved source of resistance was identified in the early 1990s in Garrapato, a landrace from the Durango group from Mexico. A breeding line derived from Garrapato, named A429, provided the best level of resistance. Inheritance studies indicated that the resistance in A429 was controlled by a single recessive gene, named bgm-1. The presence of the bgm-1 gene strongly reduces the mosaic and yellowing symptoms caused by BGYMV. The bgm-1 containing DOR lines developed by the breeding program at CIAT remain the best source of durable and stable BGYMV resistance (Fig. 5). A co-dominant sequence-characterized amplified region (SCAR) marker, SR2, which is tightly linked to bgm-1, was developed by genetic mapping. The position of the bgm-1 gene was estimated to be close to that of bc-1, which confers resistance to the potyvirus Bean common mosaic virus (BCMV), on chromosome 5. Interestingly, the mapping study indicated that bgm-1 may be sub-telomeric. Sub-telomeric regions tend to be highly recombinogenic, a characteristic that is advantageous in the context of hostpathogen co-evolution. The success at obtaining resistant varieties to BGYMV was not, unfortunately, replicated in South America for BGMV (a different virus with a low degree of sequence identity with BGYMV). Breeding for resistance to BGMV initiated in the 1970s at the Campinas Agronomical Institute (IAC in the Portuguese acronym) in Campinas, Brazil, and also at center for bean research at Embrapa, in Goiânia. Although a number of varieties with delayed incidence of the disease were obtained, inheritance was complex, yields were

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Fig. 5 The "Mosaico Dorado Amarillo de Frijol" (Bean golden yellow mosaic virus) nursery established at San Andrés Experimental Station of the Centro Nacional de Tecnología Agropecuaria y Forestal "Enriques Álvarez Córdova" in El Salvador (Central America), as part of the breeding program conducted by CIAT. The nursery contains a large number of landraces and breeding lines of black and red beans with different levels of resistance to Bean golden yellow mosaic virus (BGYMV). Courtesy of M.A. Grajales.

unremarkable, and their agronomical traits were not generally favored by growers. The most promising varieties were released during the 1990s by the breeding program at the Paraná Agronomical Institute (IAPAR) in Londrina. However, the quantitative, partial resistance provided by these lines would break down under the high BGMV incidence levels that often occurred in southern Brazil. The lack of success at obtaining cultivars with good natural resistance to BGMV led to the development of a genetically modified common bean, based on RNAi technology, which provided excellent resistance.

Resistance to Mungbean Yellow Mosaic Viruses In black gram (Vigna mungo) Mungbean yellow mosaic India virus (MYMIV) resistance is governed by a single recessive gene having NB-ARC domain with normal functional group. A resistant gene (CYR1) for MYMIV has been characterized, similarly its truncated allele (cyr1) was also found in susceptible black gram. Moreover, a SNP in LRR-like protein kinase gene responsible for a G/C transversion was identified in soybean. The yellow mosaic disease (YMD) was found to be monogenic. The comparative transcriptome analysis showed an upregulation in NAC transcription factor, Argonaute, NB-LRR, Ankyrin and WRKY33 genes. Full length cDNA of black gram MAPK (Mitogen-activated protein kinases) designated as VmMAPK1 was found increased during MYMIV infection, which helps in autophosphorylation and restricting the multiplication of MYMIV by mediating salicylic acid signaling pathway. MAPK16 and MAPKK3 genes are down regulated during MYMIV infection. Moreover, the G2/mitotic specific cyclin-1-like and Cyclin-B1 (CYCB1) were found up-regulated. MYMIV interacts with proliferative cell nuclear antigen, recombination-dependent replication (RDR51, RDR54), minichromosome maintenance protein subunit 2 (MCM2) and replication protein A, respectively. Micro RNAs (miRNAs) also confer resistance against MYMIV such as gma-miR5785 found in soybean, that helps in cleavage of BC1 gene of MYMIV. In Mungbean RAPD markers OPP 07895 and OPP 07900 are associated with Mungbean yellow mosaic virus (MYMV) resistance, while OPP 07730 was found in French bean. A resistance gene analog (RGA) marker has been associated with MYMV resistance in mungbean and black gram. Moreover, the MYMIV-resistant markers YR4 and CYR1 were found in urdbean and mungbean. In black gram the SSR marker (CEDG180) was found closely linked with YMD resistance. The QTLs (qTMIV1 to qYMIV5), qMYMIV2 and qMYMIV were identified for MYMIV resistance in Vigna radiata. The SSR marker CEDG044 was found highly linked with MYMIV resistance. The MYMIV resistance gene was mapped on chr 6 (LG C2), which is present 3.5-cM on genomic region between two SSR markers GMAC7L and Satt322. Similarly, in black gram the ISSR8111357 marker was developed and confirmed for MYMV resistance. Fig. 6 illustrates the screening of soybean germplasm for resistance to Mungbean yellow mosaic virus.

Resistance to Cotton Leaf Curl Virus in Cotton Livelihoods of many rely on cotton in many countries across the world. Cotton leaf curl viruses (CLCuVs), members of the genus Begomovirus, in the family Geminiviridae, and their natural vector whitefly (Bemisia tabaci) pose serious threat to cotton production. In the Indian subcontinent, whitefly-mediated cotton leaf curl disease (CLCuD) stands a devastating factor that limits overall production of cotton. Multiple strategies based on pathogen-derived resistance have been tried but huge diversity of geminiviruses has resulted in limited success under field conditions. Gossypium hirsutum, tetraploid cotton cultivars produce high quality lint and fiber but are susceptible to CLCuD. On the contrary, one of the diploid progenitors of cotton, G. arboreum (A genome) is resistant to CLCuD, making it a valued source for novel genes to improve CLCuD resistance. Whole transcriptome analysis of G. arboreum under graft-mediated CLCuD infection revealed the genetic basis and underlying mechanism of resistance. The genes related to oxidative stress, transcription factors, R-gene family, phytohormone signaling, membrane transporters and channel proteins were found differentially expressed and may be involved in resistance to CLCuD. Correlation

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Fig. 6 Mungbean yellow mosaic disease resistant germplasm. The panels (A) show soybean (Glycine max) line V-47 showing susceptibility to Mungbean yellow mosaic virus. The panels (B) show soybean line SG-16, which is resistant to Mungbean yellow mosaic virus.

Fig. 7 Cotton germplasm showing resistance to CLCuD. Mac7 plants grown in field condition showing resistance to Cotton leaf curl disease (CLCuD), while all the susceptible varieties grown around are showing typical symptoms of CLCuD. The black panel represents plant of CLCuD resistant Mac7 accession and susceptible varieties grown in glasshouse conditions. On the left side of panel is the closer look of asymptomatic leaf of Mac7 (Upper) and diseased or symptomatic leaf of susceptible (lower) cotton varieties.

of these identified genes with available QTLs (associated with CLCuD resistance) can help in picking the desirable genes further to impart CLCuD resistance in cotton. Several efforts have been made for the identification of resistance sources among large collections of available cotton germplasm (Fig. 7). Around 5000 cotton USDA germplasm accessions from their different breeding programs were screened against a high pressure of CLCuD. This massive screening revealed a disease resistant accession Mac7. The understanding of mechanisms underlying the interaction of virus, whitefly and host cotton plant can help in devising robust strategies for controlling the virus and vector (Fig. 8). Therefore, transcriptome analysis of Mac7 under whitefly-mediated CLCuV infestation has shed light on the molecular insights of CLCuD and whitefly resistance in cotton. This study has pinpointed the involvement of heat shock proteins and geminivirus‐interacting genes in Mac7 resistance to CLCuD. Virus-induced gene silencing (VIGS) based silencing of HSC80, E3 ligase and serine threonine kinase (STK) genes exhibited increased susceptibility to whitefly and CLCuD in Mac7 cotton. These identified genes in resistant cotton accession could be used as useful sources of resistance to CLCuD and can help in improving the whitefly and virus resistance in cotton germplasm. Understanding and exploitation of host plant resistance against whitefly can also help in control of geminiviruses. In this context, a study in cotton infested with whitefly showed differential gene expression of WRKY40, GhMPK3 and copper transport protein. Furthermore, VIGS of GhMPK3, validated the suppression of Jasmonic acid (JA) and Ethylene (ET) signaling related genes leading to the whitefly susceptibility. Small RNA-seq and genome-wide miRNA analysis of whitefly resistant and susceptible cultivars of G. hirsutum has been done recently, that revealed the involvement of miRNAs related to leucine-rich repeat (LRR) protein, MYB transcription factors and auxin response factor (ARF). VIGS of ARF8 from whitefly resistant cotton targeting miRNA390 enhanced the auxin and JA accumulation that increased the whitefly tolerance. Susceptible cotton variety of G. hirsutum when infected with CLCuD viruliferous whiteflies indicated the differential gene expression of genes including transcription factors (NAC, bHLH, MYB), heat shock proteins, methyl-transferases and metabolism related genes like cytochrome p450. Thus the genes identified and highlighted in these studies can be used in developing and enhancing cotton resistance to whitefly. Now in the era of new plant breeding technologies, CLCuD/whitefly resistant cotton cultivars harboring ample genetic diversity can be achieved using CRISPR/Cas9 system. CRISPR/Cas9 based gene targeting is now becoming a method of choice with high reliability and efficiency for cotton genome editing. It would help in targeting CLCuD related susceptibility genes in cotton to

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Fig. 8 Geminivirus (CLCuV)/whitefly – cotton interactions and mechanisms of host plant resistance. Cotton leaf curl geminivirus disease complex entry into cotton through whitefly mediates different layers of mechanisms in cotton to deal with pest and virus including interaction of geminivirus proteins with plant genes, activation of differential gene expression, miRNA machinery, metabolic changes and phytohormone signaling pathways.

Fig. 9 Maize streak virus (MSV) induced stunting in a susceptible maize variety Gusau pool 16 (bottom) and resistant variety TZL Composite 4 CL (top). Source: Taiwo et al., 2006. Plant Disease 90, 199–202.

counter the whitefly and fast evolving viral genomes. Moreover, development of improved resistant crop varieties by bridging the conventional and new genetic approaches along with pest/disease awareness to the farmers, capacity building of researchers would be helpful in controlling the geminiviruses and related diseases effectively.

Resistance to Maize Streak Virus Maize streak virus (MSV, genus Mastrevirus, family Geminiviridae) causes maize streak disease (MSD), one of the most important viral diseases of maize in sub-Saharan Africa. MSV largely remains uncontrolled in most of the African continent and during epidemic years it can lead to widespread yield losses and famine. Fig. 9 depicts field performance of a MSV resistant maize variety “TZL composite 4L” along with a MSV susceptible maize variety “Gusau Pool 16” in Nigeria. Resistance to MSV is associated with up to five different QTLs located on chromosome 1, 3, 7, and 9, that are either dominant or additive in nature (Table 1). None of these QTLs in isolation can prevent MSV infections, however combinations of them can resist the MSV infections. Although maize

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genotypes that tolerate MSV infection without compromising much with yield losses have been developed, there has been only limited success in the field. The MSV resistance seems to be specific to certain geographic locations, selection made of MSV resistance at one location may fail to tolerate MSV infections in another location. Thus the breeding efforts and screening for MSV resistance have to be specific to different geographical areas or the agro-climatic zones of Africa. One of the major concern is that the improved MSV tolerant maize genotypes don’t have the desirable agronomic traits, such as good yields. Thus developing maize genotypes that possess both MSV tolerance and high yields remains a big challenge to the maize breeders. Further involvement of multiple MSV tolerant QTLs makes it difficult to introgress each of them in one single maize genotype and development of such a variety may take several years of breeding. Despite the humungous efforts made in development of MSV resistant maize varieties, most farmers prefer to use high-yielding maize varieties.

Conclusion The current knowledge on the genes that confer resistance to geminiviruses is scarce when compared to the information available for plant infecting RNA viruses. Although the geminivirus resistant locus/loci are mapped in several cultivated plant species, the actual genes associated with the resistance are yet to be cloned and characterized. The few resistant genes that have been identified, the underlying resistant mechanism is yet to be understood. In this article we have summarized the sources and genes for geminivirus resistance in the most important crop plants that are heavily infested by geminiviruses. The geminivirus resistant genes identified so far are diverse in their characteristics, they are monogenic and polygenic, dominant and recessive, and specific as well as broad spectrum in nature. Introgression of these resistant genes in cultivated crop varieties has not been very successful since some of these genes are associated with poor agronomic traits. Loci or genes conferring resistance to the whitefly vectors are not yet identified and identification of such genes may boost the breeding for begomovirus resistance. Revolution in genome editing technologies and employment of CRISPR-Cas based technologies for precise editing of host genes are promising for manipulating the recessive resistant genes that makes them immune to geminivirus infections. Transgenics by employing CRISPR-Cas technology that destroys the geminiviral DNA genome have been developed, however their durability remains uncertain with reference to the fast mutating geminiviruses which may evade the targeting by sgRNAs that are key for Cas9 based cleavage. However, these are considered as transgenics and their social acceptance is questionable. Thus emphasis on identification and characterization of novel geminiviral resistant genes offers more promise for development of improved geminivirus resistant crop varieties.

Further Reading Beebe, S.E., Ochoa, I., Skroch, P., Nienhuis, J., Tivang, J., 1995. Genetic diversity among common bean breeding lines developed for Central America. Crop Science 35, 1178–1183. Bianchini, A., 1999. Resistance to Bean golden mosaic virus in bean genotypes. Plant Disease 83, 615–620. Blair, M.M., Rodriguez, L.M., Pedraza, F., Morales, F., Beebe, S., 2007. Genetic mapping of the Bean golden yellow mosaic geminivirus resistance gene bgm-1 and linkage with potyvirus resistance in common bean (Phaseolus vulgaris L.). Theoretical and Applied Genetics 114, 261–271. Chellappan, P., Vanitharani, R., Fauquet, C.M., 2004. Short interfering RNA accumulation correlates with host recovery in DNA virus-infected hosts and gene silencing targets specific viral sequences. Journal of Virology 78, 7465–7477. Dharajiya, D.T., Ravindrababu, Y., 2019. Identification of molecular marker associated with Mungbean yellow mosaic virus resistance in mungbean [Vigna radiata (L.) Wilczek]. Vegetos 32, 532–539. Ji, Y., Schuster, D.J., Scott, J.W., et al., 2007. Sources of resistance, inheritance, and location of genetic loci conferring resistance to members of the tomato-infecting begomoviruses. In: Czosnek, H. (Ed.), The Tomato Yellow Leaf Curl Virus Disease: Management, Molecular Biology, Breeding for Resistance. Dordrecht: Springer, pp. 343–362. Kundu, A., Singh, P.K., Dey, A., Ganguli, S., Pal, A., 2019. Complex molecular mechanisms underlying MYMIV-resistance in Vigna mungo revealed by comparative transcriptome profiling. Scientific Reports 9, 1–13. Lapidot, M., Friedmann, M., 2002. Breeding for resistance to whitefly-transmitted geminiviruses. Annals of Applied Biology 140, 109–127. Li, J., Zhu, L., Hull, J.J., et al., 2016. Transcriptome analysis reveals a comprehensive insect resistance response mechanism in cotton to infestation by the phloem feeding insect Bemisia tabaci (whitefly). Plant Biotechnology Journal 14, 1956–1975. Naqvi, R.Z., Shan-e-Ali Zaidi, S., Akhtar, K.P., et al., 2017. Transcriptomics reveals multiple resistance mechanisms against cotton leaf curl disease in a naturally immune cotton species, Gossypium arboreum. Scientific Reports 7, 1–15. Navas-Castillo, J., Fiallo-Olivé, E., Sánchez-Campos, S., 2011. Emerging virus diseases transmitted by whiteflies. Annual Review of Phytopathology 49, 219–248. Patil, B.L., 2018. Genes, Genetics and Transgenics for Virus Resistance in Plants. Norfolk: Caister Academic Press, doi:10.21775/9781910190814. Patil, B.L., Fauquet, C.M., 2011. Chapter 11. Ecology of plant viruses, with special reference to geminiviruses. In: Hurst, C. (Ed.), Studies in Viral Ecology. Volume 1. Hoboken, NJ: John Wiley & Sons, Inc, pp. 273–306. Patil, B.L., Fauquet, C.M., 2015. Studies on differential behavior of cassava mosaic geminivirus DNA components, symptom recovery patterns, and their siRNA profiles. Virus Genes. 50 (3), 474–486. doi:10.1007/s11262-015-1184-y. Rojas, M.R., Macedo, M.A., Maliano, M.R., et al., 2018. World management of geminiviruses. Annual Review of Phytopathology 56, 637–677. Singh, N., Mallick, J., Sagolsem, D., Mandal, N., Bhattacharyya, S., 2018. Mapping of molecular markers linked with MYMIV and yield attributing traits in mungbean. Indian Journal of Genetics 78, 118–126. Strausbaugh, C.A., Eujayl, I.A., Wintermantel, W.M., 2017. Beet curly top virus strains associated with sugar beet in Idaho, Oregon, and a Western U.S. Collection. Plant Disease 101, 1373–1382. Zaidi, S.S.e.A., Naqvi, R.Z., Asif, M., et al., 2019. Molecular insight into cotton leaf curl geminivirus disease resistance in cultivated cotton (Gossypium hirsutum). Plant Biotechnology Journal 18, 691–706.

Plant Rhabdoviruses (Rhabdoviridae) Ralf G Dietzgen, The University of Queensland, St. Lucia, QLD, Australia Michael M Goodin, University of Kentucky, Lexington, KY, United States Zhenghe Li, Zhejang University, Hangzhou, China r 2021 Elsevier Ltd. All rights reserved. This is an update of A.O. Jackson, R.G. Dietzgen, R.-X. Fang, M.M. Goodin, S.A. Hogenhout, M. Deng, J.N. Bragg, Plant Rhabdoviruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00492-1.

Nomenclature aa Amino acid(s) AGO Argonaute ER Endoplasmic reticulum G Glycoprotein IGSs Intergenic or “gene-junction” sequences kb Kilobase kDa Kilo Dalton ldr Leader M Matrix protein

Glossary Chytrid fungi Obligate parasitic root-infecting fungi of the Chytridiomycetes that rarely appear to affect the host plants but act as vectors for plant viruses, for example Olpidium virulentus. L protein Main component of the viral polymerase complex, responsible for replication and transcription with RNA-dependent RNA polymerase, mRNA 50 capping, 30 polyadenylation and protein kinase activities. Leader and trailer Regulatory regions at the 30 and 50 termini of rhabdovirus genomes that show a high degree of sequence complementarity and are predicted to form a panhandle structure during replication.

MP Movement protein N Nucleocapsid protein NC Condensed nucleocapsid NE Nuclear envelope nt Nucleotide(s) ORF Open reading frame P Phosphoprotein RdRp RNA-dependent RNA polymerase trl Trailer UTR Untranslated region

Negative-sense RNA viruses Have negative-sense singlestranded RNA as their genetic material. Genomic RNA of negative polarity is complementary to the mRNA that is transcribed from it for translation of encoded proteins. Purified negative-sense RNA by itself is not infectious but requires viral core proteins to commence replication and transcription. Nucleocapsid core The transcriptionally-competent ribonucleoprotein that represents the minimal infectious unit of rhabdoviruses. Viroplasm The site of viral RNA replication, transcription and nucleocapsid formation.

Introduction ‘Classic’ plant rhabdoviruses have distinctive enveloped bacilliform or bullet-shaped particles and are distinguished based on whether they replicate and undergo morphogenesis in the cytoplasm or in the nucleus. Consequently, they have been classified in two genera, Cytorhabdovirus or Nucleorhabdovirus. More than 70 putative plant rhabdoviruses have been described over the years based on their morphology although, in many cases, molecular characterizations necessary for unambiguous classification are incomplete or lacking. More recently, an increasing number of novel plant rhabdovirus genomes have been assembled from high throughput sequencing data of diverse plant and insect species. All sequenced plant rhabdoviruses have the same general genome organization as other members of the Rhabdoviridae, and each encodes at least six open reading frames (ORFs), one of which facilitates cell-to-cell movement of the virus. Thus, plant rhabdoviruses have a number of similarities to members of other rhabdovirus genera, but they differ in several respects from rhabdoviruses infecting vertebrates. Two new genera of bisegmented plant viruses were recently classified as rhabdoviruses, the dichorhaviruses that infect horticultural and ornamental plant species and are transmitted by Brevipalpus mites, and the varicosaviruses that are transmitted by chytrid fungi. Rhabdoviruses infect plant species from a large number of different families, including numerous weed hosts and several major horticultural and cereal crops. Symptoms of infection vary substantially and range from stunting, vein clearing, mosaic, and mottling of leaf tissue, to tissue necrosis. The most serious pathogens include Maize mosaic virus (MMV), Lettuce necrotic yellows virus (LNYV), Rice yellow stunt virus (RYSV), Rice stripe mosaic virus (RSMV), Eggplant mottled dwarf virus (EMDV), Strawberry crinkle virus (SCV), Potato yellow dwarf virus (PYDV), Barley yellow striate mosaic virus (BYSMV), Northern cereal mosaic virus (NCMV), Orchid fleck virus (OFV), and Coffee ringspot virus (CoRSV). A number of other rhabdoviruses also have disease potential that can be affected by agronomic practices, incorporation of genes for disease resistance, and control of insect vectors.

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The transmission of most plant rhabdoviruses is dependent on specific phytophagous insects that support replication of the virus. Therefore, viral prevalence and distribution is influenced to a large extent by the ecology and host preferences of their vectors. Although some rhabdoviruses can be transmitted mechanically by abrasion of leaves, this mode of transmission does not contribute significantly to their natural spread due to the labile nature of the virion. Moreover, seed or pollen transmission of plant rhabdoviruses has not been described; thus, aside from vegetative propagation, direct plant-to-plant transmission is unlikely to be a major factor in the ecology or epidemiology of these pathogens. This article focuses on recent findings concerning the taxonomy, structure, replication, and vector relationships of the expanded group of plant rhabdoviruses. More extensive aspects of plant rhabdovirus biology, specifically ecology, disease development and control, can be found in earlier reviews.

Taxonomy and Classification Plant rhabdoviruses are taxonomically classified in the order Mononegavirales, family Rhabdoviridae. The International Committee on Taxonomy of Viruses (ICTV) in its 9th Report (2011) listed two genera containing plant rhabdovirus species, Cytorhabdovirus and Nucleorhabdovirus, the members of which have unsegmented genomes and are discriminated by their replication and morphogenesis in the cytoplasm and nucleus, respectively of infected cells. For the 10th Report (2018), the genera Dichorhavirus and Varicosavirus were added to the family, based on phylogenetic relatedness of its members to the rhabdoviruses. The genus Dichorhavirus was newly created, whereas Varicosavirus had not previously been assigned to a family. Dichorhaviruses are most closely related to nucleorhabdoviruses, but have a bisegmented genome and virions do not appear to be enveloped. Varicosaviruses will not be discussed further in this chapter. In 2019, in the family Rhabdoviridae, out of the 19 recognized genera 4 contain plant viruses. The ICTV has listed eleven accepted plant virus species in the genus Cytorhabdovirus, ten in the genus Nucleorhabdovirus, five in the genus Dichorhavirus and one in the genus Varicosavirus, a total of 27 plant virus species out of 289. Phylogenetic analyses of these rhabdoviruses have confirmed their taxonomic classification, although it has become evident that following inclusion of the dichorhaviruses, the genus Nucleorhabdovirus no longer has a monophyletic origin (Fig. 1) and will need to be split. The genomes of more than 20 novel plant rhabdoviruses have been recently identified by high throughput sequencing and await official classification. However, many plant rhabdoviruses that were identified before next generation sequencing technologies were available have not been investigated in much detail beyond cursory infectivity studies, crude physicochemical analyses of virus particles, and electron microscopic observations of morphogenesis. Consequently, more than 70 putative rhabdoviruses await assignment to a species and genus once genome sequences can be obtained. Viruses characterized in greater detail, with coding sequences deposited in GenBank, that have approved or pending taxonomic assignments are listed in Table 1.

Particle Morphology and Composition In comparison to rhabdoviruses restricted to vertebrate hosts, plant-adapted rhabdoviruses are typically bacilliform in shape (Fig. 2). The modal lengths of particles, measured by electron microscopy of thin sections, range from 45 to 100 nm wide by 130–350 nm in length. Suggestive of packaging genomic RNAs in separate particles, particles of dichorhaviruses and varicosaviruses tend to be approximately half the length of rhabdovirus particles. While dichorhavirus particles lack a lipid envelope, those of the majority of rhabdoviruses consist of an outer layer of host-derived membranes, the lipid composition of which is commensurate with that of the membrane environment where morphogenesis occurs, be it on the nuclear envelope (NE) or endoplasmic reticulum (ER) (Fig. 2(b)). Protruding from the envelope is the spike glycoprotein (G) that is distributed more or less uniformly around the virion (Fig. 2(a)). Serving to both stabilize the particle and connect the G-containing envelope to the condensed nucleocapsid (NC) is the matrix protein (M). However, the biological functions of M may extend far beyond that of its structural role. The NC contained within the virion is a ribonucleoprotein complex composed of single-stranded negative-sense RNA, encapsidated over its entire length by the nucleocapsid protein (N) (Fig. 2(a)). Associated with this complex are small amounts of the viral phosphoprotein (P) and large (L) RNA-dependent RNA polymerase (RdRp). This “core” NC is transcriptionally competent for synthesizing viral mRNAs once it gains entry to cells compatible for replication. Collectively, virions of plant rhabdoviruses have a composition that is approximately 70% protein, 2% RNA, 20%–25% lipid, with a minimal fraction of carbohydrate associated with the G protein. Genomic RNAs of plant rhabdoviruses tend to range from 10 to 15 kb, while those of dichorhaviruses, and varicosaviruses are 5–6 kb for each segment.

Genome Structure and Organization The general genomic structure of plant rhabdoviruses is conserved in a manner similar to that of the prototypical rhabdoviruses like Vesicular stomatitis virus (VSV) or Rabies virus. As such, the protein-encoding open reading frames (ORFs) are flanked by short untranslated leader (ldr), and untranscribed trailer (trl) regions, at the 30 and 50 ends of the genomic RNA. These regions show a high degree of sequence complementarity (Fig. 3(a)) and are thought to potentially form a panhandle structure of the genome during

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Fig. 1 Maximum likelihood (ML) phylogeny of plant rhabdoviruses. L protein amino acid sequences of selected plant rhabdoviruses (cytorhabdoviruses, nucleorhabdoviruses, dichorhaviruses, and varicosaviruses) and two well-known animal rhabdoviruses (as outgroup) were multiple aligned using MAFFT version 7. Subsequently, ambiguously aligned regions were trimmed with Gblocks. A ML tree was constructed based on the sequence alignment using PhyML 3.0 with the best fit model LG with þ I þ G þ F. The types of plant host and arthropod vectors are differentiated by circles filled with different colors (monocot and dicot) or shaded cartoons, respectively. An asterisk following the virus name indicates that the vector has not yet been determined. Numbers at the nodes indicate bootstrap values 450%. Reprinted with permission from Whitfield, A.E., Huot, O.B., Martin, K.M., Kondo, H., Dietzgen, R.G., 2018. Plant rhabdoviruses – Their origins and vector interactions. Current Opinion in Virology 33, 198–207, Copyright 2018, Elsevier B.V.

replication. Located between each ORF are so-called intergenic or “gene-junction” sequences (IGSs) that provide cis-regulatory elements for transcription and replication. IGSs have a modular structure composed of three elements, (1) Module 1; a poly(U) tract at the 30 end of each gene located after the un-translated region (UTR) of the preceding ORF, (2) Module 2; a di- to poly-nucleotide spacer typically starting with a guanyl residue that is not templated in mRNAs, (3) Module 3; a transcriptional “start” sequence that is typically a tri- to penta-nucleotide sequence found in each of the viral mRNA transcripts (Fig. 3(b)).

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Table 1 Characteristics of plant rhabdoviruses, members and/or tentative members of 4 genera of the family Rhabdoviridae. Only accessions of complete or coding-complete genome sequences are provided. Type species are in bold Genus/Species

Acronym

Genbank accession

Host plant

Arthropod vector

Cytorhabdovirus Alfalfa dwarf cytorhabdovirus Barley yellow striate mosaic cytorhabdovirus Broccoli necrotic yellows cytorhabdovirus Colocasia bobone disease-associated cytorhabdovirus Festuca leaf streak cytorhabdovirus Lettuce necrotic yellows cytorhabdovirus Lettuce yellow mottle cytorhabdovirus Northern cereal mosaic cytorhabdovirus Sonchus cytorhabdovirus Strawberry crinkle cytorhabdovirus Wheat American striate mosaic cytorhabdovirus

ADV BYSMV BNYV CBDaV FLSV LNYV LYMoV NCMV SonV SCV WASMV

KP205452 KM213865

Dicot Monocot Dicot Monocot Monocot Dicot Dicot Monocot Dicot Dicot Monocot

Aphid? Planthopper Aphid Planthopper? Planthopper Aphid Unknown Planthopper Unknown Aphid Leafhopper

Tentative species Bean-associated cytorhabdovirus Cabbage cytorhabdovirus 1 Maize-associated cytorhabdovirus Maize yellow striate virus Papaya virus E Persimmon virus A Raspberry vein chlorosis virus Rice stripe mosaic virus Strawberry cytorhabdovirus 1 Tomato yellow mottle-associated virus Trifolium pratense cytorhabdovirus A Trifolium pretense cytorhabdovirus B Yerba mate chlorosis-associated virus

BaCV CCyV  1 MaCV MYSV PpVE PeVA RVCV RSMV StrV  1 TYMaV TpVA TpVB YmCaV

MK202584 KY810772 KY965147 KY884303 MH282832 AB735628 MK240091 KX525586 MK211271 KY075646 MH982250 MH982249 KY366322

Dicot Dicot Monocot Monocot Dicot Dicot Dicot Monocot Dicot Dicot Dicot Dicot Dicot

Unknown Unknown Unknown Unknown Unknown Unknown Aphid Leafhopper Aphid Aphid? Unknown Unknown Unknown

Nucleorhabdovirus Datura yellow vein nucleorhabdovirus Eggplant mottled dwarf nucleorhabdovirus Maize fine steak nucleorhabdovirus Maize Iranian mosaic nucleorhabdovirus Maize mosaic nucleorhabdovirus Potato yellow dwarf nucleorhabdovirus Rice yellow stunt nucleorhabdovirus Sonchus yellow net nucleorhabdovirus Sow thistle yellow vein nucleorhabdovirus Taro vein chlorosis nucleorhabdovirus

DYVV EMDV MFSV MIMV MMV PYDV RYSV SYNV SYVV TaVCV

KM823531 KC905081 AY618417 MF102281 MK828539 GU734660 AB011257 L32603 AY674964

Dicot Dicot Monocot Monocot Monocot Dicot Monocot Dicots Dicot Monocot

Unknown Leafhopper Leafhopper Planthopper Planthopper Leafhopper Leafhopper Aphid Aphid Unknown

Tentative species Alfalfa-associated nucleorhabdovirus Apple rootstock virus A Bird’s-foot trefoil-associated virus 1 Black currant-associated rhabdovirus 1 Green Sichuan pepper nucleorhabdovirus 1 Morogoro maize-associated virus Physostegia chlorotic mottle virus Wheat yellow striate virus

AaNV ApRVA BFTV  1 BCaRV  1 GSPNuV MMaV PCMV WYSV

MG948563 MH778545 MH614262 MF543022 MH323437 MK112501 KX636164 MG604920

Dicot Dicot Dicot Dicot Dicot Monocot Dicot Monocot

Unknown Unknown Unknown Unknown Unknown Unknown Unknown Leafhopper

Dichorhavirus Citrus chlorotic spot dichorhavirus Citrus leprosis dichorhavirus N Clerodendrum chlorotic spot dichorhavirus Coffee ringspot dichorhavirus Orchid fleck dichorhavirus

CiCSV CiLV-N ClCSV CoRSV OFV

RNA1 KY700685 KX982176 MG983790 KF812525 AB244417

RNA2 KY700686 KX982179 MG983791 KF812526 AB244418

Dicot Dicot Dicot Dicot Dicot

Brevipalpus Brevipalpus Brevipalpus Brevipalpus Brevipalpus

Varicosavirus Lettuce big-vein associated varicosavirus

LBVaV

AB075039

AB114138

Dicot

Olpidium fungi

KT381973 AJ867584 EF687738 AB030277

GU985153

MH129615

MK257717

KY549567 AB516283

mites mites mites mites mites

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Fig. 2 Virion structure and morphology of enveloped plant rhabdoviruses. (a) Stylized virion structure of an enveloped, unsegmented plant rhabdovirus. The left side of the figure shows the approximate orientation and shape of the glycoprotein (G) and lipid bilayer envelope (orange). The matrix protein (M; blue hexagons) connects the envelope to the condensed nucleocapsid core, which is composed of the genomic-length single-stranded RNA, encapsidated by the nucleocapsid protein (N; red circles). Associated with this RNA-N complex are the phosphoprotein (P; yellow circles), and the large (L) RNA-dependent RNA polymerase (blue circle). (b) Transmission electron micrograph of a thin-section of sonchus yellow net virus-infected leaf tissue of Nicotiana benthamiana. Note accumulation of bacilliform shaped virus particles (V) in the perinuclear space between the outer (ONM) and inner (INM) membranes of the nuclear envelope (NE). Nucleocapsids are condensed in the viroplasm (VP) that is highly enriched with P protein prior to migrating to the NE where morphogenesis takes place. EM image courtesy of Dr. Tea Meulia, Molecular and Cellular Imaging Center, The Ohio State University (https://mcic.osu.edu/home).

Fig. 3 Intergenic and terminal noncoding regions of plant rhabdovirus genomes. (a) Complementary sequences at the 30 and 50 termini of plant rhabdovirus genomic RNAs; complementary nucleotides are shown in bold font. Dichorhavirus OFV, Orchid fleck virus; nucleorhabdovirus PYDV, Potato yellow dwarf virus; cytorhabdovirus LNYV, Lettuce necrotic yellows virus. (b) Conserved modular intergenic sequences separating each gene.

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Fig. 4 Schematic diagrams of the genome organization of representative plant rhabdoviruses classified in the genera Nucleorhabdovirus, Cytorhabdovirus, Dichorhavirus, and Varicosavirus. See Table 1 for full virus names. Each open reading frame (ORF) is indicated by an arrow labelled with the name of the encoded protein (N ¼ nucleocapsid protein; P ¼ phosphoprotein; M ¼ matrix protein; G ¼ glycoprotein; L ¼ Large polymerase). Accessory genes are shaded in grey and numbered. In some cases, ORF 3 has been shown to encode a cell-to-cell movement protein.

In all genomes characterized to date the N ORF follows immediately after the leader, and the large (L) RdRp ORF precedes the trailer region. Situated between these ORFs are three additional protein coding regions conserved in all rhabdoviruses, namely the P protein, M protein, and G protein. Interspersed between these conserved ORFs are additional coding regions peculiar to plant-adapted viruses generally, or unique to a particular virus. These regions, typically designated as ‘P30 or ‘P60 are located in genomes in the general order 30 -ldr-N-P-P3-M-G-P6-L-trl-50 (Fig. 4). The number of ORFs between the P and M genes may be variable. Overall, the plant rhabdovirus genomes analyzed so far contain between 6 and 10 ORFs (Fig. 4). Proteins encoded in position P3 (Fig. 4) have long been predicted, and recently experimentally demonstrated, to be required for the cell-to-cell movement functions essential for all plant viruses. The general gene order and genome structure of plant rhabdoviruses are also conserved in the dichorhaviruses and varicosaviruses. Key distinctions are that the N to G ORFs are encoded on RNA1 in dichorhaviruses, and on RNA2 in varicosaviruses, while the cognate L polymerases are encoded on RNA2 and RNA1, respectively (Fig. 4).

Properties of Viral Proteins Comprehensive biochemical, molecular, subcellular localization, and protein-protein interaction analyses of plant rhabdovirusencoded proteins have been carried out mostly with the Sonchus yellow net virus (SYNV), PYDV, LNYV, and OFV models. Putative functions have been assigned to proteins encoded by less-well studied plant rhabdoviruses on the basis of their genomic locations and sequence relatedness to plant and animal model rhabdoviruses. Overall, amongst the five structural proteins of plant

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Fig. 5 Confocal micrographs of leaf epidermal cells of transgenic N. benthamiana expressing RFP targeted to endomembranes. (a) in the absence of virus, RFP accumulates in the envelope of the nucleus (N). (b) Infection by the nucleorhabdovirus SYNV results in intranuclear accumulation of the inner nuclear membrane. The resulting expansion of perinuclear volume fills with the RFP-ER marker. In the image planes shown here, the reticulate pattern of the ER is not visible in the cytoplasm (C). (c) Within SYNV-infected nuclei, nucleocapsid formation takes place in viroplasm “rings” that are highly enriched in P protein (expressed here as a fusion to green fluorescent protein, in transgenic plants expressing the nucleolar marker protein RFP-Fibrillarin 1). Scale bar ¼ 2 mm.

rhabdoviruses, the L protein has conserved polymerase motifs common to all mononegaviruses, whereas the N, P, M, and G proteins are less conserved and have only limited sequence relatedness to analogous proteins of animal rhabdoviruses.

The Nucleocapsid Protein (N) The N protein functions to encapsidate the entire viral RNA genome and tightly packages it to form NC that are RNase-resistant (Fig. 2(a)). VSV N protein has a tendency to form large aggregates and binds cellular RNAs non-specifically. Interactions of P protein with free N protein form a soluble complex, designated N(0)-P, which selectively encapsidates viral RNA. The availability of N(0)-P complexes regulates a critical switch from transcription to replication in the viral infection cycle, since genome replication entails encapsidation of nascent RNAs and only the encapsidated form of the genome can serve as template for viral polymerase. Biochemical experiments have shown that the solubility of SYNV N protein is increased upon co-expression and interactions with the P protein, so the SYNV P protein appears to have a similar chaperone function to keep the N protein in a biologically active form. The N protein is also a major component of the viroplasm, the site of viral RNA replication, transcription and NC formation (Fig. 5). Confocal microscopic studies have revealed the subcellular localization of N proteins of selected cyto- and nucleorhabdoviruses consistent with their role in viroplasm formation. For instance, the N protein of the cytorhabdovirus LNYV localizes to the cytoplasm, whereas the homologs from the nucleorhabdoviruses SYNV, PYDV, MMV, Maize fine streak virus (MFSV), and RYSV are targeted to subnuclear foci. When the N proteins were co-expressed with their respective cognate P proteins, the N-P complexes co-localized to distinct cytoplasmic (in the case of LYNV) or subnuclear (SYNV, PYDV, OFV, etc.) inclusions characteristic of viroplasms, which are often different from the localization of either protein alone. In SYNV, formation of these subnuclear foci requires self-interactions of the N protein that are mediated by a helix-loop-helix motif near the amino (N)-terminus, as well as interactions of the P protein with the N-terminus of the N protein. However, heterologous combinations of SYNV and MFSV N and P proteins fail to form subnuclear foci. These co-localization studies suggest that cognate N-N and N-P protein interactions are essential for viroplasm formation. The N proteins of nucleorhabdoviruses and dichorhaviruses contain a nuclear localization signal (NLS) for nuclear import. Experiments conducted in plant and yeast cells have shown that SYNV and PYDV N proteins contain a bipartite NLS near the carboxy (C)-terminus that is required for nuclear localization, and biochemical studies have shown that the N proteins interact in vitro with importin a homologs. Related bipartite NLS sequences are also present in the C-terminus of MFSV and RYSV N proteins, while the analogous regions of the MMV and Taro vein chlorosis virus N proteins contain a monopartite NLS.

The Phosphoprotein (P) In addition to the chaperone role in maintaining the N protein in soluble form, rhabdovirus P protein is also a co-factor of the L polymerase and a component of the viral NC core (Fig. 2(a)). The SYNV P protein forms complexes in vivo with the N and L proteins that are analogous to N-P and P-L complexes found in VSV-infected cells, and also engages in self-interactions that are mediated by the N-terminus of the P protein. Likewise, pairwise N-P and P-P interactions have also been documented for several other plant rhabdoviruses including LYNV, PYDV, and OFV. Rhabdoviral P proteins are highly phosphorylated as their name suggests, and phosphorylation of specific residues play important roles in regulating the P protein activities in transcription and

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replication. Direct evidence showing phosphorylation of plant rhabdovirus P protein is available for SYNV. In SYNV, the P protein is phosphorylated in vivo at threonine residues, which differs from the VSV P protein that is phosphorylated at serine residues. Hence, the P proteins of plant rhabdoviruses probably have replication-related functions similar to their well-characterized vertebrate counterparts. Sequence analyses show that the P proteins of the nucleorhabdoviruses PYDV, RYSV, and MMV carry a NLS, which is consistent with their pronounced nuclear localization patterns. Although similar NLS sequences have not been identified for the SYNV and MFSV P proteins, these proteins localize to the nucleus as well as the cytoplasm when transiently expressed in plant cells. Furthermore, the SYNV P protein binds directly to human importin b derivatives in vitro and the central third of the P protein is required for nuclear import, but other regions of the protein affect the import efficiency. However, co-expression with cognate N proteins relocalizes the SYNV and MFSV P proteins to subnuclear foci exclusively, suggesting that the P proteins may contain a nuclear export signal whose function is masked by the N protein-binding. Apart from functioning in replication, the P proteins of two cytorhabdoviruses have been shown to exhibit relative weak, yet evident RNA silencing suppressor activities. LYNV P suppresses local RNA silencing and delays systemic silencing. Further mechanistic studies have shown that LYNV P inhibits microRNA-mediated mRNA slicing and transitive RNA silencing likely by interactions with Argonaute (AGO) proteins and the RdRp 6/ Suppressor of Gene Silencing 3 (RDR6/SGS3) complex. Similarly, Alfalfa dwarf virus (ADV) P suppresses local silencing weakly but systemic silencing strongly. ADV P interacts with AGO1 and AGO4, inhibits miRNA-guided AGO1 cleavage and prevents transitive RNA silencing.

The P3 Movement Protein Like other plant viruses, plant rhabdoviruses encode proteins to assist in cell-to-cell movement of viral infection entities through the plasmodesmata and their systemic transport through the vascular system. Considerable evidence for a movement protein (MP) function has been accumulated for proteins encoded by the gene located between the P and M genes designated P3, whose name varies with viruses, including SYNV sc4, LNYV 4b, MFSV P4, PYDV Y, and P3 in most other viruses (Fig. 4). The predicted secondary structures of several plant rhabdovirus P3 homologs have a distant relatedness to structural motifs of the Tobacco mosaic virus 30K superfamily of plant viral MP. Additional evidence for a role of the P3 homologs in cell-to-cell movement is their localizations to the cell periphery or the plasmodesmata and associations with host membranes during transient expression. The movement function for the P3 proteins has been reinforced by trans-complementation experiments, in which P3 homologs encoded by various cyto-, and nucleorhabdoviruses, and dichorhaviruses have been shown to support cell-to-cell movement of MP-defective positive-strand RNA viruses. In addition, a recombinant SYNV sc4 deletion mutant is unable to move from initially infected cells to adjacent tissues, but the movement defect is rescued by transient expression of the sc4 protein in trans. Protein interaction experiments revealed that the P3 homologs from SYNV, PYDV, RYSV, and Tomato yellow mottle-associated virus (TYMaV) interact with their respective N and/or P proteins, and in the case of SYNV sc4, the specific sc4-N and sc4-P interactions appear to facilitate inter- and intracellular NC trafficking. Together, these lines of evidence collectively support a MP role for the plant rhabdovirus P3 proteins. Protein localization and cellular fractionation assays indicate that several rhabdovirus P3 proteins are membrane-associated, although membrane protein prediction algorithms generally fail to identify any transmembrane regions in these proteins. In addition, SYNV sc4 protein was found to be present in purified virions, and after sucrose gradient fractionation, sc4 co-sedimented with the G protein-containing membrane fractions of the virion preparations. Similarly, immunogold labelling has shown that Rice transitory yellowing virus (a strain of RYSV) P3 is also associated with virus particles, but in this case the P3 appears to localize to the interior of the virion in the NC. Thus, these findings provide evidence that plant rhabdovirus MP can also function as structural proteins, but the actual structural role remains obscure.

The Matrix Protein (M) Within the rhabdovirus particle, the M protein forms a layer connecting the helical NC core and the transmembrane G protein (Fig. 2(a)). During infections, the M protein plays critical roles in virus budding by interacting with the NC core, membrane lipid and the G protein. Studies with vertebrate rhabdoviruses have shown that late in replication, the M protein interacts with genomic NC and directs the condensed core to membrane budding sites that are enriched in viral G protein. Similar functions for plant rhabdovirus M protein are supported by protein localization and interactions studies. The N-M interactions appears to be common among several plant rhabdoviruses including SYNV and PYDV. Consistent with the morphogenesis sites of cyto- and nucleorhabdoviruses, the LYNV M protein localizes to the cytoplasm near the cell periphery, but the SYNV and PYDV M proteins are targeted to the nucleus. Notably, when transiently expressed in uninfected cells, the SYNV and PYDV M proteins distribute uniformly throughout the nucleoplasm; however, in virus-infected cells, the M proteins appear as subnuclear foci bridging the periphery of the inner nuclear membrane (INM) and the viroplasms. These foci likely represent virion maturation sites where coiled NC cores bud through the nuclear membranes. As with VSV M, the PYDV and SYNV M proteins also interact with and remodel host membranes. Infection by SYNV, and to a lesser extent by PYDV, results in the formation of intranuclear spherules derived from invagination of the INM (Fig. 5). The spherule formation is also induced when PYDV M is ectopically expressed by itself, and requires a di-leucine motif located at aa

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residues 223 and 224 of the M protein. In contrast, expression of SYNV M is insufficient to trigger nuclear membrane rearrangement, and even infection by a recombinant SYNV M deletion mutant fails to induce intranuclear spherule formation. The SYNV M protein requires interactions with the G protein’s C-terminal tail to associate with INM and induce membrane remodelling. In addition to their roles in viral morphogenesis, preliminary experiments indicate that rhabdovirus M proteins have important roles in host–virus interactions because they appear to be able to inhibit host gene expression. The nucleorhabdovirus M protein may also play a role in NC egress from the nucleus. SYNV M protein forms motile foci liberating from the nuclear envelope and moving towards the cytoplasm, whereas PYDV M contains a putative nuclear export signal that may be involved in transport of NC cores into the cytoplasm.

The Glycoprotein (G) The G protein forms the glycoprotein spikes of rhabdovirus virions (Fig. 2(a)). The plant rhabdovirus G proteins are typical type I membrane proteins, consisting of a signal peptide sequence of B20 aa, an N-terminal luminal (or virion surface) domain that contains potential glycosylation sites, a transmembrane region followed by a short C-terminal tail. The signal peptide is cleaved by signal peptidases during co-translational translocation to the ER lumen to yield a “mature” transmembrane G protein. In addition, the SYNV G protein contains a putative NLS near the C-terminus that could be involved in transit to the inner nuclear membrane prior to virion morphogenesis. Several glycosylation inhibitors interfere with N-glycosylation of the SYNV G protein and also block SYNV morphogenesis. A prominent role of G protein in morphogenesis is also supported by SYNV reverse genetics studies. A SYNV G protein deletion mutant is abortive in virion maturation, leading to accumulation of striking arrays of naked NC cores that fail to bud through the INM. The C-terminal tail of SYNV G protein is critical for morphogenesis, since this domain is required for relocalization of the M protein to INM surfaces and subsequently membrane remodelling. The N-terminal domain of SYNV G also contributes to efficient budding, but is dispensable for systemic infections. Although SYNV G protein is not absolutely required for infections of Nicotiana benthamiana plant host, a more critical role can be envisioned during infections of insect vectors by analogy to animal rhabdoviruses. The plant rhabdovirus G protein may be crucial for binding to proposed insect midgut receptors and host cell entry. Indeed, antibody blockage or enzymatic digestion of PYDV G protein drastically reduces virus infectivity in insect vector cells.

The Polymerase Protein (L) The L proteins are the most closely related of the rhabdovirus-encoded proteins and contain conserved mononegavirus polymerase domains and RNA-binding motifs. Alignment of the L protein sequences of non-segmented negative-strand RNA viruses reveals conservation within 12 motifs. Phylogenetic trees derived from L protein alignments indicate that the nucleorhabdoviruses, dichorhaviruses and cytorhabdoviruses cluster together in two major clades separated from the vertebrate rhabdoviruses (Fig. 1). The L proteins of plant rhabdoviruses are present in low abundance within virus particles and in infected cells (Fig. 2(a)). A viral RdRp is activated after treatment of virions of the cytorhabdoviruses LNYV and Broccoli necrotic yellows virus with mild nonionic detergents, and this activity co-sediments with loosely coiled NC filaments that are released from virions. The transcribed products are complementary to the genome, as expected of mRNAs. Thus, the described polymerases of these plant cytorhabdoviruses appear to be similar to the extensively studied polymerases of the vesiculoviruses. In contrast, no appreciable polymerase activity is evident in dissociated preparations of SYNV or other nucleorhabdovirus virions that have been analyzed. However, an active polymerase can be recovered from the nuclei of plants infected with SYNV. Polymerase activity is associated with a complex consisting of the N, P, and L proteins that co-sediments with SYNV NC cores. The polymerase complex can be precipitated with P protein antibodies, but the activity of the complex is not inhibited by these antibodies. However, antibody inhibition experiments demonstrate that the L protein is required for polymerase activity. Kinetic analysis of transcription products also reveals that the complex is capable of sequentially transcribing a polyadenylated plus-sense leader RNA and polyadenylated mRNAs corresponding to each of the six SYNV-encoded proteins. Potential replication intermediates consisting of short incomplete minus-strand products homologous to the genomic RNA are also transcribed. These results thus provide a model whereby nucleorhabdovirus particles require polymerase activation by host components early in infection. In contrast, the polymerases of the cytorhabdoviruses appear to be present in an active form in virions and the released cores are capable of initiating primary transcription immediately upon uncoating in vitro.

Other Accessory Proteins Besides the canonical genes, plant rhabdovirus genomes may also contain 1–4 additional ORFs encoding putative accessory proteins (Fig. 4). For example, an X ORF is present in PYDV and EMDV between the N and P genes, and within the P and M genes, there are 4 ORFs in the two cereal cytorhabdoviruses, i.e., NCMV and BYSMV, instead of only one ORF (P3) in most of the other plant rhabdoviruses. In addition, a short transcriptional units was found in several cyto- and nucleorhabdoviruses, designated P6 in RYSV, RSMV, ADV, and SCV, and P9 in NCMV and BYSMV. Small ORFs preceding the L gene were also found in the genomes of some animal rhabdoviruses, such as ephemero-, hapa-, and tibroviruses. These ORFs encode small, highly hydrophobic, and basic proteins named viroporins, which function to modify the cell membrane permeability to ions or other small molecules and are

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involved in virion assembly and release. The plant rhabdovirus P6 and P9 proteins do not share obvious sequence identity with each other or with the viroporins of animal rhabdoviruses, but some share a characteristic transmembrane domain. Only very limited information is available regarding the functions of plant rhabdovirus accessory genes. RYSV P6 protein is present in virion preparations purified from infected rice plants and in protein samples from viruliferous leafhoppers, suggesting that P6 has a structural role in infection. RYSV P6 was also shown to suppress systemic RNA silencing by affecting RDR6-mediated secondary siRNA amplification, but did not affect local RNA silencing. However, ADV P6 does not appear to have similar RNA silencing suppressor activities, suggesting that this function is not conserved amongst plant rhabdoviruses.

Cytopathology and Replication Entry Little information is available about early entry and uncoating events during plant infections. It is hypothesized that the rigid plant cell wall is breached through insect vector feeding or experimentally by mechanical abrasion, and that the introduced enveloped virions may fuse to cellular membranes to release NC cores into the cytoplasm, followed by M protein disassembly to liberate the flexuous NC filaments. However, during insect vector infections, the surface G proteins of plant rhabdoviruses presumably play a role in binding to proposed midgut receptors that mediate virion uptake into epithelial cells by endocytosis, although such receptors have so far not been identified.

Transcription and Genome Replication The fundamental replication and transcription processes of plant rhabdoviruses are similar to their animal-infecting counterparts. Cytorhabdoviruses closely resemble animal rhabdoviruses in that both replicate in the cytoplasm, whereas nucleorhabdoviruses and dichorhaviruses have exclusive nuclear replication phases (Fig. 6). The latter two groups of viruses utilize the host nucleocytoplasmic transport machinery to recognize karyophylic signals present on the N and P proteins to facilitate nuclear targeting of viral core proteins and NC. The plant rhabdovirus replication cycles described below were developed largely by analogy to the VSV model and supported by biochemical characterization of viral polymerase fractions, analyses of viral mRNAs, electron microscopic observations, protein localization and interaction assays, and recent reverse genetics studies. After NC release from virions, primary rounds of transcription are initiated by using the negative-stranded genome template to produce capped and polyadenylated leader RNA and mRNAs for each of the viral genes. Transcription occurs progressively and sequentially from the genomic 30 end, yielding discrete mRNA species in a decreasing molar gradient, i.e., the mRNA levels follow the order N4P4P34M4G4L. This sequential and polar transcription is mediated by interactions of the viral polymerase complex with cis-regulatory elements located in each gene junction. As primary transcription proceeds, newly synthesized viral proteins increase in abundance. Accumulation of the N and P proteins triggers a switch of the polymerase function from mRNA transcription to genome replication. Upon replication, the viral polymerase initiates antigenomic RNA synthesis at the exact 30 terminus of the genomic RNA and ignores all internal regulatory signals located in the gene junctions. During genome replication, the nascent RNAs are encapsidated by the N-P complex to form antigenomic NC, which then serve as template of the P-L replicase complexes for synthesis of genomic RNAs that are subsequently encapsidated to form genomic NC. Continuous NC formation depletes free N, P, and L proteins, and lower core protein abundances signal a transition from replication to secondary rounds of mRNA transcription to increase the pool of viral mRNAs for replenishment of viral proteins. This cyclic phase of replication and transcription is coordinated by a feedback mechanism that measures the abundance of the core proteins by the levels of nascent leader RNA encapsidation. Later in cytorhabdovirus replication, accumulation of viral core proteins and progeny NC result in formation of thread-like viroplasms in the cytoplasm that are located in close proximity to dense networks of the ER. Likewise, replication of nucleorhabdoviruses and dichorhaviruses induces formation of subnuclear viroplasms in greatly enlarged nuclei that consist of large masses of granular material containing viral RNA and the N, P, and L proteins (Fig. 2(b)).

Morphogenesis The plant rhabdoviruses vary profoundly in their sites of morphogenesis and differ substantially from vertebrate rhabdoviruses that undergo virion maturation at the host plasma membrane. In plant cells, cytorhabdoviruses bud through ER membranes, and mature particles accumulated in ER-associated cytoplasmic cisternae that are greatly proliferated during virus infections. In contrast, the budding of nucleorhabdoviruses, such as SYNV and PYDV, occurs in association with the INM, and enveloped virions accumulate in enlarged perinuclear spaces formed between the inner and outer nuclear envelopes (Fig. 2(b); Fig. 5). However, dichorhaviruses that also replicate in the nucleus do not have an enveloped form of virions. Rather, rod-like non-enveloped particles are present in nuclei of infected cells, either scattered through the nucleoplasm, forming crystalline arrays or arranged perpendicularly onto the INM. Occasionally, these particles are also present in the cytoplasm and appear to be enwrapped by membranous tubular structures. In this case, the virions are arranged radially and perpendicularly to the membrane and form a “spoke wheel” configuration.

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Fig. 6 Models for cyto- and nucleorhabdovirus infection cycles in plant cells. Plant rhabdoviruses are initially introduced into plant cells through mechanical breach of the cell wall (CW) during insect vector feeding. Upon entry into the cell, viral nucleocapsids (NC) are liberated from the virions and replication takes place in the cytoplasm (cytorhabdoviruses) or nucleus (nucleorhabdoviruses). For cytorhabdoviruses, the NC initiate primary rounds of transcription in the cytoplasm to yield capped, polyadenylated mRNAs for translation of each viral protein. Primary transcription is followed by genome replication to produce viral antigenomic and genomic RNAs that are encapsidated by the newly synthesized nucleocapsid proteins to form progeny NC, and then by secondary rounds of transcription to increase the pool of mRNAs. As cyclic waves of transcription and replication proceed, accumulation of viral core proteins and NC form cytoplasmic viroplasms (VP). During the later stages of replication, matrix (M) proteins coil NC and likely interact with ER-anchored viral glycoproteins, and morphogenesis occurs by budding of condensed NC into the proliferated ER membranes. Matured bacilliform virions accumulate in pronounced ER vesicles. The replication cycles of nucleorhabdoviruses are fundamentally similar to those of cytorhabdoviruses, but their NC are transported into the nucleus for transcription and replication cycles. The viral mRNAs are exported to the cytoplasm and translated, and the core proteins and M protein are then transported through the nuclear pore complex into the nucleus. As replication progresses, subnuclear VP appear and the nuclei become greatly enlarged. Morphogenesis occurs by budding of coiled NC into inner nuclear membrane (INM), and enveloped virions accumulate between INM and outer nuclear membrane (ONM) of the nuclear envelope (NE). During cell-to-cell movement, the NC exit the nuclei either by a nuclear export activity of a viral core protein or by a “bud-out” process of the enveloped virions. Once in the cytoplasm, the NC of cyto- and nucleorhabdoviruses interact with membrane-associated P3 movement proteins (MP) in a species-specific manner. The MP then direct intracellular NC trafficking to the cell periphery and intercellular movement through gated plasmodesmata (PD) across the CW by mechanisms that may involve microtubule (MF)- and microfilament (MF)-associated host proteins.

Morphogenesis of cyto- and nucleorhabdoviruses occurs during the late stages of replication in which the M protein accumulates and reaches concentrations sufficient to initiate coiling of the genomic NC. The M protein condenses the NC into a compact bacilliform core, which leads to downregulation of RNA synthesis and a transition from replication to virion assembly. The coiled cores are brought to the vicinity of endomembrane surfaces where the transmembrane G proteins are enriched, and then bud through the membranes to acquire lipid envelope and the G proteins. Infections with SYNV or PYDV induce massive INM invagination into the nucleoplasm, forming intranuclear spherule-like structures (Fig. 5). This structural rearrangement may function to bring the INM surfaces in close proximity to viroplasms and to provide readily accessible sites for virion maturation.

Cell-To-Cell Movement A MP function has been assigned to the P3 accessory gene located between the P and M genes for several cyto- and nucleorhabdoviruses, as well as dichorhaviruses, since the P3 homologs of these diverse plant rhabdoviruses display crossfamily movement trans-complementation with positive-strand RNA viruses. Reverse genetics analyses using recombinant SYNV

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further reinforce that the sc4 protein, a P3 homolog, is essential for SYNV cell-to-cell movement. The plant rhabdovirus movement complex has been postulated to involve viral NC, since NC constitutes the minimal infectious unit. This hypothesis has been verified by SYNV reverse genetics studies, which show that SYNV minigenomes comprising the N, P, sc4, and L genes are capable of autonomous localized movement. Reverse genetics analyses have also revealed a highly specific cell-to-cell movement mechanism for cyto- and nucleorhabdoviruses. The movement defects of MP-deletion mutants of SYNV and TYMaV can only be rescued by their respective cognate MP, but not by MP from related rhabdoviruses, distantly related or unrelated plant RNA viruses. The movement specificities of SYNV and TYMaV are explained by the species-specific interactions between NC core proteins and MP. Available evidence suggests that during replication, some of the newly synthesized NC in the cytoplasm interact specifically with membrane-anchored MP, which facilitate intracellular NC trafficking to the cell periphery, followed by intercellular transport through the plasmodesmata (Fig. 6). Furthermore, NC transport may also involve host cytoskeleton components. Microtubuleassociated factors have been identified to interact with MP of SYNV and LNYV, and actin filaments are required for BYSMV P protein trafficking (Fig. 6). For nucleorhabdoviruses and dichorhaviruses that employ nuclear phases of replication, their NC have to be exported from the nuclear replication sites to the cytoplasm before cell-to-cell movement can occur. How the NC nuclear egress is achieved has not been resolved, but this process may involve nuclear exporting function of a viral NC core protein or deenvelopment of intraluminal virions.

Vector Relationships, Distribution and Evolution Phylogenetic studies support the general hypothesis that natural host ranges of plant rhabdoviruses are determined by their arthropod vectors more so than by host plant species (Fig. 1). Moreover, where studied, plant rhabdoviruses replicate in cells of their insect vectors as well as in their plant hosts. Taken together, the adaptive evolutionary trajectory of plant rhabdoviruses is likely to be from insects to plants. Plant rhabdoviruses are most frequently transmitted by aphids (Aphididae), leafhoppers (Cicadellidae), or planthoppers (Delphacidae), while dichorhaviruses are transmitted by false spider mites (Brevipalpus sp.) (Table 1, Fig. 1). The relationship between virus and vector appears to be tightly regulated, given that closely related rhabdoviruses are transmitted by different insect species. This is exemplified by the sanguinolenta and constricta strains of PYDV that are distinguished by their differential transmission by the leafhopper vectors, Aceratagallia sanguinolenta and Agallia constricta. This relationship occurs in cytorhabdoviruses as well, where Soybean blotchy mosaic virus, for example, has been shown to replicate and persist in its vector, the leafhopper, Peragallia caboverdensis. In some cases, plant rhabdoviruses have no known vector, however, phylogenetic clustering of viruses that group according to vector can inform prediction of what type of insect maybe involved in the epidemiology of such viruses (Fig. 1). Importantly, while the virus-insect relationship is constrained tightly, the same rhabdovirus may occur naturally in multiple plant host species. For example, whereas PYDV was once agronomically important on potato, it still occurs in reservoirs of red clover (Trifolium incarnatum). Although hampered by the inability to mechanically transmit many plant rhabdoviruses, experimental host ranges have been shown to be substantially broader than those reported from field data. However, in most cases the reservoir species are unknown for this group of viruses, which limits our understanding of their true ecology. Unbiased detection methods, such as RNA-Seq, in wild plant communities are likely to provide significant insight into the population dynamics of rhabdoviruses, and related plant viruses. Once acquired by the insect vector, it appears that cyto- and nucleorhabdoviruses are persistently transmitted in a propagative manner and can be transmitted to vector progeny, in those cases where this aspect of rhabdovirus biology has been examined. Typical of such relationships, long latent periods (hours to days) are required for virus acquisition before vectors are competent for transmission. Insects remain viruliferous throughout their lives, and transovarial passage through eggs and nymphs has been determined experimentally in some cases. Evidence for tissue tropism of rhabdoviruses is derived from studies with MMV in its planthopper vector Peregrinus maidis. Following acquisition from plant tissues, MMV accumulates in the epithelial cells of the anterior midgut, followed by infection of nerve cells. Subsequent infection of tracheal cells, hemocytes, muscles, salivary glands, fat cells, mycetocytes, and, finally, epidermal tissues follows, thus completing the circulative infection cycle. Delivery of MMV by microinjection of P. maidis results in efficient virus infection and transmission, facilitating functional virus-vector interaction studies in combination with RNA silencing. Akin to cyto- and nucleorhabdoviruses, following long periods of acquisition (up to two weeks), dichorhaviruses are transmitted in a circulative-propagative manner by Brevipalpus mites. Furthermore, transovarial transmission has been reported for a citrus leprosis-associated dichorhavirus. However, to-date vertical transmission of dichorhaviruses has not been confirmed. Cytological studies have demonstrated the presence of both viroplasm and particle formation in mites that have fed on plants infected with several dichorhaviruses including OFV, Citrus leprosis virus N, Clerodendrum chlorotic spot virus and CoRSV (Fig. 7), suggesting that the ability to replicate in vector tissues is conserved within this group of viruses. As with cyto- and nucleorhabdoviruses, tropism for specific cell types is evident with dichorhaviruses, and cells from midgut epithelium and other cell types were shown to have similar cytopathology to that observed in infected plant cells. There have only been limited studies on the population structures of rhabdoviruses but certain generalities are evident such as the low degree of sequence divergence within populations. Genetic variability studies have been conducted for LNYV, type member of the genus Cytorhabdovirus, using the N gene of isolates sampled from both Australia and New Zealand, as well as analyzing symptom expression on Nicotiana glutinosa. Sequence analyses supported separation of LNYV isolates into two

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Fig. 7 Transmission electron micrograph of a thin section of the midgut of the mite Brevipalpus phoenicis collected from bleeding heart (Clerodendrum x speciosum) infected by Clerodendrum chlorotic spot virus. Rod-like particles (V) can be seen in the periphery of the nucleus (N), some arranged perpendicularly onto the nuclear envelope (NE). Nearby, a cell of the symbiotic bacterium Candidatus Cardinium (Car) can be seen. Courtesy of Prof. E.W. Kitajima, University of São Paulo, Brazil.

subgroups (I and II), with subgroup I predicted to be an older lineage than subgroup II. Interestingly, these subgroups could not be differentiated based on symptom expression in the experimental host N. glutinosa. Sub-group I has apparently gone extinct in Australia, presumably due to its less-efficient dispersal in plant reservoir and/or vectors. Similarly, the dichorhavirus CoRSV was found at one hundred percent of locations (n ¼ 45) sampled in an area spanning 75% of the coffee growing region of Brazil. All cultivars, regardless of cherry color or growth habit, were found to serve as hosts, suggesting that there is limited resistance in commercially deployed germplasm. Phylogenetic analysis based on the N gene identified a strong geo-spatial relationship among isolates, which clustered into three clades, although the overall sequence diversity between isolates was very low. In contrast to LNYV, and despite low genetic diversity among isolates, variation in symptom expression of CoRSV infection was observed in the experimental host Chenopodium quinoa. Based on these studies the hypothesis that the spread of CoRSV is constrained by the clonal expansion of thelytokous populations of B. papayensis has been proposed. Further studies, particularly in reservoir host species, once identified, are required to determine if population structure observed in the agronomic hosts holds across a broader range of hosts. While the distribution of most dichorhaviruses appears to be limited to South America, OFV has a global distribution.

Further Reading Dietzgen, R.G., Freitas-Astúa, J., Chabi-Jesus, C., et al., 2018. Dichorhaviruses in their host plants and mite vectors. Advances in Virus Research 102, 119–148. Dietzgen, R.G., Kondo, H., Goodin, M.M., Kurath, G., Vasilakis, N., 2017. The family Rhabdoviridae: Mono- and bipartite negative-sense RNA viruses with diverse genome organization and common evolutionary origins. Virus Research 227, 158–170. Fang, X.-D., Yan, T., Gao, Q., et al., 2019. A cytorhabdovirus phosphoprotein forms mobile inclusions trafficked on the actin/ER network for viral RNA synthesis. Journal of Experimental Botany 70, 4049–4062. doi:10.1093/jxb/erz195. Gao, Q., Xu, W.-Y., Yan, T., et al., 2019. Rescue of a plant cytorhabdovirus as versatile expression platforms for planthopper and cereal genomic studies. New Phytologist 223, 2120–2133. doi:10.1111/nph.15889. Goodin, M.M., Chakrabarty, R., Yelton, S., et al., 2007. Membrane and protein dynamics in live plant nuclei infected with Sonchus yellow net virus, a plant-adapted rhabdovirus. Journal of General Virology 88, 1810–1820. Jackson, A.O., Dietzgen, R.G., Goodin, M.M., Li, Z., 2018. Development of model systems for plant rhabdovirus research. Advances in Virus Research 102, 23–57. Jang, C., Wang, R., Wells, J., et al., 2017. Genome sequence variation in the constricta strain dramatically alters the protein interaction and localization map of Potato yellow dwarf virus. Journal of General Virology 98, 1526–1536. Mann, K.S., Bejerman, N., Johnson, K.N., Dietzgen, R.G., 2016. Cytorhabdovirus P3 genes encode 30K-like cell-to-cell movement proteins. Virology 489, 20–33. Mann, K.S., Johnson, K.N., Carroll, B.J., Dietzgen, R.G., 2016. Cytorhabdovirus P protein suppresses RISC-mediated cleavage and RNA silencing amplification in planta. Virology 490, 27–40. Martin, K.M., Whitfield, A.E., 2018. Cellular localization and interactions of nucleorhabdovirus proteins are conserved between insect and plant cells. Virology 523, 6–14. Meulia, T., Stewart, L., Goodin, M.M., 2018. Sonchus yellow net virus core particles form on ring-like nuclear structure enriched in viral phosphoprotein. Virus Research 258, 64–67. Sun, K., Zhou, X., Lin, W., et al., 2018. Matrix-glycoprotein interactions required for budding of a plant nucleorhabdovirus and induction of inner nuclear membrane invagination. Molecular Plant Pathology 19, 2288–2301. Wang, Q., Ma, X., Qian, S., et al., 2015. Rescue of a plant negative-strand RNA virus from cloned cDNA: Insights into enveloped plant virus movement and morphogenesis. PLoS Pathogens 11, e1005223. Whitfield, A.E., Huot, O.B., Martin, K.M., Kondo, H., Dietzgen, R.G., 2018. Plant rhabdoviruses–their origins and vector interactions. Current Opinion in Virology 33, 198–207. Zhou, X., Lin, W., Sun, K., et al., 2019. Specificity of plant rhabdovirus cell-to-cell movement. Journal of Virology 93. e00296-19. doi:10.1128/JVI.00296-19.

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Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/negative-sense-rna-viruses/mononegavirales/w/rhabdoviridae Family: Rhabdoviridae Mononegavirales ICTV.

Plant Satellite Viruses (Albetovirus, Aumaivirus, Papanivirus, Virtovirus) Mart Krupovic, Archaeal Virology Unit, Institut Pasteur, Paris, France r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

ORF Open reading frame satRNA Satellite RNA ssRNA Single-stranded ribonucleic acid UTR Untranslated region

aa Amino acid(s) CP Coat protein or capsid protein nt Nucleotide(s)

Glossary Albetovirus Al- for alphanecrovirus [helper virus], be- for betanecrovirus [helper virus], to- for tobacco. Aumaivirus Au- for aureusvirus [helper virus], mai- for maize. Hyperparasite A parasite whose host (in this article, a helper virus) is also a parasite. Papanivirus Pa- for panicovirus [helper virus], pani- for panicum.

Satellite RNA A subviral RNA genome dependent on a helper virus for both replication and encapsidation. Satellite virus A polyphyletic group of viruses encoding structural components of their virions, but incapable of completing the infection cycle without the assistance of a helper virus. Virtovirus Vir- for virgavirus [helper virus], to- for tobacco.

Introduction Viruses span an entire range of morphological, genomic, and functional complexity. Some viruses are exceedingly complex, surpassing many unicellular organisms in terms of physical dimensions and genome sizes. For instance, members of the family Mimiviridae carry megabase-sized genomes and encode over 1000 proteins, including many molecular machineries required for virus multiplication. On the other side of the complexity spectrum are viruses carrying just one or two genes, suggesting a stronger reliance on the host compared to the bigger viruses. Notably, some viruses depend not only on the host cells but also on other viruses for reproduction. The phenomenon whereby one virus depends for its propagation on another virus has been first described in a plant virus system in the early 1960s. Certain preparations of tobacco necrosis viruses (TNV; genera Alphanecrovirus and Betanecrovirus, family Tombusviridae) contained two types of spherical particles that differed in size and antigenic properties. The larger, TNV, particles could propagate autonomously, whereas the smaller ones were unable to replicate in the absence of TNV. The smaller particles became known as particles of satellite tobacco necrosis virus (STNV), whereas TNV is referred to as STNV’s helper virus. Subsequently, several other plant viruses having features similar to those of STNV have been discovered. Furthermore, viruses which depend on another virus for reproduction have been described in protists and animals. Overall, satellite viruses vary considerably in terms of complexity and evolutionary origins. Also, the functions supplemented by the helper viruses vary from one virus system to another. For instance, satellite viruses of plants rely on helper viruses for genome replication, whereas members of the Lavidaviridae depend on giant mimiviruses for transcription, clearly indicating that satellite viruses have evolved on several independent occasions. The tripartite relationship between satellite viruses, helper viruses and host cells is an example of hyperparasitism, which is not uncommon in complex natural ecosystems. Indeed, there are other genetic elements which parasitize viruses, most notably, including satellite nucleic acids. The key difference between satellite nucleic acids and satellite viruses is that the latter necessarily encode components of their virions, whereas the former do not. Instead, satellite nucleic acids commonly encode proteins responsible for the replication of the cognate nucleic acid, whereas the components of the virion for horizontal transfer between the hosts are being provided by the helper viruses. Due to their dependency on other, quasi-autonomous viruses for reproduction, most satellite viruses, with a notable exception of members of the genera Dependoparvovirus and Deltavirus, have been for a long time considered as sub-viral agents and not officially classified by the International Committee on Taxonomy of Viruses (ICTV). However, in 2016, the arbitrary distinction between satellite viruses and other viruses has been abandoned and satellite viruses were incorporated into the ICTV classification scheme, resulting in establishment of several genera and families. Plant satellite viruses have been classified into genera Albetovirus, Aumaivirus, Papanivirus, and Virtovirus; family Sarthroviridae has been created for classification of positive-sense ( þ )RNA satellite viruses that infect arthropods, and Lavidaviridae includes double-stranded DNA viruses replicating in protist cells and parasitizing giant viruses of the family Mimiviridae. This article describes the diversity and properties of plant satellite viruses belonging to the genera Albetovirus, Aumaivirus, Papanivirus and Virtovirus.

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Fig. 1 Plant satellite viruses. Structural similarity between the virions and jelly-roll capsid proteins of satellite tobacco necrosis virus (STNV; PDB ID: 2BUK), satellite panicum mosaic virus (SPMV; PDB ID: 1STM) and satellite tobacco mosaic virus (STMV; PDB ID: 4OQ8). All three virions have T ¼ 1 icosahedral symmetry. Images of the depicted virions were downloaded from the VIPERdb database (viperdb.scripps.edu/). Genome maps of the three viruses are shown at the bottom with the gene encoding the capsid protein shown in blue.

Albetoviruses: Viruses Related to Satellite Tobacco Necrosis Virus STNV is one of the most extensively studied satellite viruses. Over the years, many properties of this virus, including its genome sequence, virion structure and mechanism of capsid assembly, as well as details of the interaction with the helper virus, have been elucidated. The linear positive-sense ( þ )RNA genome of STNV consists of 1239 nt and encodes a single jelly-roll-fold capsid protein (CP) of 195 aa, which is necessary and sufficient for formation of icosahedral (T ¼ 1) virions (Fig. 1). Virion assembly proceeds cooperatively via interactions between the packaging signals, degenerated stem-loop structures distributed throughout the genome, and multiple CP copies. In the natural habitat, STNV is transmitted through the soil the same way as its helper virus, i.e., by zoospores of a plantpathogenic fungus Olpidium brassicae, suggesting that both viruses share certain transmission determinants. For both TNV and STNV, the 50 ends of the genomes are phosphorylated and lack a 7-methylguanylate cap or a genome-linked protein, whereas the 30 -termini lack a polyadenylation sequence. Several cis-acting elements located within the 50 and 30 untranslated regions (UTRs) are responsible for efficient translation and replication of the STNV genome. The 30 and 50 UTRs of STNV and TNV can be exchanged without abolishing RNA accumulation, although the translation elements at the 30 -terminal regions appear to be unrelated in the two viruses. Three serotypes of STNV have been described: STNV-1 (or STNV), STNV-2, and STNV-C (Table 1). Different STNV strains are activated by different helper viruses. The replication of STNV-1 and STNV-2 is supported by isolates of TNV-A, the sole member of the Tobacco necrosis virus A species (genus Alphanecrovirus), whereas TNV-D, the sole member of the Tobacco necrosis virus D species (genus Betanecrovirus), supports the replication of STNV-C. Genome sequences and organizations for the three STNV strains are overall similar (Table 1). CPs are E50%–63% identical in sequence. Whereas the 50 UTRs of STNV-1 and STNV-2 are 30 nt in length and are nearly identical, the corresponding region in STNV-C is considerably longer (101 nt). Similarly, whereas the 30 UTRs of STNV-1 and STNV-2 approximately 64% similar to each other, they share only 40% and 38% identity to that of STNV-C. STNV-like viruses STNV-1, STNV-2, and STNV-C are classified into three species, Tobacco albetovirus 1, 2, and 3, respectively, within a genus Albetovirus (sigil: Al- for alphanecrovirus [helper virus], be- for betanecrovirus [helper virus], to- for tobacco).

Aumaivirus: Satellite Maize White Line Mosaic Virus The fourth, more divergent member of the STNV-like virus group is satellite maize white line mosaic virus (SMWLMV; Table 1). SMWLMV depends on maize white line mosaic virus (MWLMV; species Maize white line mosaic virus, genus Aureusvirus, family Tombusviridae) for multiplication. MWLMV can infect maize in the absence of SMWLMV, whereas the same is not true for SMWLMV, which replicates in maize only when co-inoculated with MWLMV. The ssRNA genome of SMWLMV is 1168 nt in length and encodes one CP (Table 1). Similar to STNV-like viruses, the SMWLMV virion is 17 nm in diameter. SMWLMV CP displays 32% identity to the CP of STNV-1, strongly suggesting that SMWLMV and the STNV-like viruses have diverged from a common ancestor. However, due to lower sequence similarity between the CPs of STNV-like viruses and SMWLMV, the latter virus has been classified into a species, Maize aumaivirus 1, within a separate genus Aumaivirus (sigil: Au- for aureusvirus [helper virus], mai- for maize).

Papaniviruses: Viruses Related to Satellite Panicum Mosaic Virus Satellite panicum mosaic virus (SPMV) depends on Panicum mosaic virus (PMV; species Panicum mosaic virus, genus Panicovirus, family Tombusviridae) for replication as well as systemic spread in plants. The 50 -terminus of the SPMV genome is phosphorylated and lacks a 7-methylguanylate cap. Several secondary structure elements implicated in the replication of the SPMV genome were

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Satellite virus

Species

Helper virus

Accession #Genome, ntCapsid Ø, nmComments

Albetovirus Satellite tobacco necrosis Tobacco tobacco necrosis virus A V01468 albetovirus 1 (Tombusviridae) virus (STNV-1)

1239

17

Satellite tobacco necrosis Tobacco tobacco necrosis virus A M64479 albetovirus 2 (Tombusviridae) virus 2 (STNV-2)

1245

17

Satellite tobacco necrosis Tobacco tobacco necrosis virus D AJ000898 albetovirus 3 (Tombusviridae) virus C (STNV-C)

1221

17

1168

17

CP is 32% identical to that of STNV-1

826

16

Besides virion formation, CP of SPMV has several other biological functions, including systemic accumulation, maintenance and movement of the SPMV RNA SSADV is a strain of SPMV (36 nt substitutions; 5 aa changes)

Aumaivirus Satellite maize white line Maize maize white line mosaic M55012 aumaivirus 1 virus (Tombusviridae) mosaic virus (SMWLMV) Papanivirus Satellite panicum mosaic Panicum Panicum mosaic virus papanivirus 1 (Tombusviridae) virus (SPMV)

M17182

Panicum Satellite St. Augustine St. Augustine decline virusL10083 824 papanivirus 1 strain of PMV decline virus (SSADV) (Tombusviridae) Satellite grapevine virus unclassified grapevine virus F KC149510 1060 (SGVV) (Betaflexiviridae)? Virtovirus Satellite tobacco mosaic Tobacco tobacco mosaic virus virtovirus 1 virus (STMV) (Virgaviridae)

M25782

1059

ND

STNV suppresses the replication of its helper virus and ameliorates the TNV-induced symptoms in different hosts STNV and STNV-2 coat protein genes share 55% nucleotide sequence identity, whereas the UTRs are more similar STNV and STNV-C coat proteins share 62% sequence identity, whereas the 30 UTRs are only 40% identical

ND

SGVV and GVF share stem-loop structures at the 50 ends of the genomes

17

The 30 UTR is similar to that of tobamoviruses, with a clear sequence similarity between STMV and TMV

predicted in the 50 and 30 UTRs. The 826 nt-long ssRNA genome of SPMV contains two open reading frames (ORF; Table 1). However, only one of them (the one encoding CP) is expressed in in vitro translation assays. The sequence of SPMV CP is not appreciably similar to those of STNV-like viruses (below 15% identity). However, X-ray structure analysis of the SPMV particle revealed that the protein has a similar jelly-roll fold as the corresponding protein of STNV (Fig. 1). Beside its structural role, SPMV CP has several other biological functions, most notably, systemic accumulation, maintenance, and movement of the cognate SPMV RNA in plants. Interestingly, the latter activities of SPMV CP apparently extend to the helper virus RNA, assisting in its maintenance or stabilization. It is noteworthy that PMV and SPMV are involved in a peculiar tripartite association with a 350 nt-long satellite RNA (satRNA), whereby PMV provides necessary factors for satRNA replication and SPMV provides CPs for satRNA encapsidation. Two other viruses encoding SPMV-like CPs have been reported (Table 1). The first one, satellite St. Augustine decline virus (SSADV), is associated with the St. Augustine decline strain of PMV. SSADV is 95% identical to SPMV over the entire genome length (36 nt changes, 5 aa changes) and is considered as a different strain of SPMV rather than a different species. Notably, SSADV genome is 2 nucleotides shorter than that of SPMV, making it the shortest known satellite virus genome (Table 1). The second putative satellite virus, satellite grapevine virus (SGVV), has been discovered by deep sequencing of total genomic RNA from grapevine. However, neither the viral particles nor the associated helper virus have been characterized. SGVV CP shares E24% sequence identity with SPMV CP. SPMV and SSADV have been assigned into a genus Papanivirus (sigil: Pa- for panicovirus [helper virus], pani- for panicum), within a species Panicum papanivirus 1. By contrast, SGVV remains unclassified due to lack of information on its ability to form virions as well as on the identity of its helper virus. Interestingly, homologs of SPMV CP are encoded by certain satRNAs. In particular, bamboo mosaic virus satellite RNA (satBaMV; 836 nt) encodes a protein (P20) which is 44% identical to the CP of SPMV. P20 plays a role in the accumulation and movement of the satBaMV but does not participate in satRNA encapsidation. Instead, satBaMV is packaged into rod-shaped particles by the CP of the helper bamboo mosaic virus, a member of the family Alphaflexiviridae. In addition, a sequence of olive viral satellite RNA (OVsatRNA) has been deposited to GenBank that encodes a protein 35% identical to P20 of satBaMV. Considering the conservation of the SPMV-like CPs, it appears likely that satBaMV and OVsatRNA have evolved from genuine

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satellite viruses, once again emphasizing the apparent ease with which transitions between different types of mobile elements (i.e., parasitic nucleic acids and viruses) occur.

Virtovirus: Satellite Tobacco Mosaic Virus Satellite tobacco mosaic virus (STMV) has been isolated from Nicotiana glauca (tree tobacco) and is naturally associated with and dependent on tobacco mild green mosaic virus (TMGMV; species Tobacco mild green mosaic virus, genus Tobamovirus, family Virgaviridae). However, under experimental settings, STMV can adapt and replicate in many plant hosts (e.g., tobacco, pepper, tomato) in association with other tobamoviruses, including tobacco mosaic virus (TMV). The adaptation of STMV to other helper viruses involves the deletion of a single G residue in a stretch of five G residues (positions 61–65) in the 50 UTR, together with a change in the 5 0 nucleotide. Thus far, STMV is the only known satellite virus that uses rod-shaped viruses as helpers. The STMV genome is a linear ssRNA molecule of 1059 nt that contains two ORFs (Fig. 1), both of which are functional in the in vitro translation assay. The first ORF encodes a protein of 58 aa which lacks similarity to proteins in the public databases and appears to be dispensable for STMV multiplication. Indeed, certain naturally occurring isolates of STMV contain a deletion within ORF1 and do not produce the corresponding product. The second ORF encodes STMV CP, which also has no identifiable homologs in sequence databases. However, structural analysis shows that the STMV CP has a jelly-roll fold similar to those of STNV and SPMV (Fig. 1), suggesting that the three satellite viruses might be evolutionarily related. As in the case of STNV and SPMV, but different from the helper TMV virus, the genome of STMV lacks a 7-methylguanylate cap and the first 6 nt of the STMV RNA are identical to those of the STNV genome. However, in contrast to STNV and SPMV, the 50 end of STMV genome is not phosphorylated. The 30 UTR is predicted to contain a series of pseudoknots followed by a tRNA-like structure which can be amino acylated with histidine. The latter features are strikingly similar to those of the genome of helper TMV and other tobamoviruses, with 40–50 nt-long regions of near identity among the STMV and TMV 30 UTRs. These secondary structure elements play critical roles in STMV genome replication, translation, and initiation of virion assembly. STMV is currently a sole representative of the species Tobacco virtovirus 1 within the genus Virtovirus (sigil: Vir- for virgavirus [helper virus], to- for tobacco).

Effect of Mixed Infections on the Host and the Helper Viruses The parasitic lifestyle of satellite viruses is usually associated with negative effects on the fitness of the helper virus, whereas the effect on the host cell during a co-infection might differ drastically between viruses. The negative effect of STNV on the propagation of its helper virus manifests as a decrease in (1) the number of necrotic lesions formed in co-inoculated plant leaves, (2) diameter of the lesions, and (3) the amount of TNV produced in inoculated leaves. However, several factors influence the extent to which TNV multiplication is affected. In particular, these include the relative concentrations of the satellite and helper viruses in the inoculum, the physiological state of the host plants before and during the infection, and the type of plant used for the assay. An excess of STNV present during co-infection reduces the yield of TNV to non-detectable levels. This reduction is associated with a selective decrease in the synthesis of TNV capsid protein, suggesting that STNV affects TNV virion production. By contrast, co-infection of PMV with SPMV exacerbates PMV disease phenotype in millet plants resulting in a severe chlorosis and stunting, reduced seed production, and often death of the infected plant. The synergism between PMV and SPMV is also characterized by the more rapid systemic accumulation of PMV and SPMV CPs and genomic RNAs than during infection by PMV alone. Expression of the SPMV CP from a potato virus X-based vector was shown to induce the formation of chlorotic spots on the host as well as non-host plants, confirming that SPMV CP is a pathogenicity determinant. The effects of STMV on the multiplication of its helper virus, as well as on the helper virus-induced symptoms, are dependent on the host. In tobacco plants, STMV does not change the mild mosaic symptom caused by TMGMV, whereas in jalapeno pepper severe leaf blistering induced by TMGMV is attenuated by STMV infection. Furthermore, tobamovirus titers are greatly decreased by STMV in pepper compared to other hosts.

Common Ancestry of Plant Satellite Viruses? Plant satellite viruses from the four groups described above all propagate in flowering plants (angiosperms) and share several genomic and structural characteristics that distinguish them from other known viruses. Namely, representatives of all four satellite virus groups form capsids with T ¼ 1 icosahedral symmetry and composed of 60 CP subunits (Fig. 1). Although common among ssDNA viruses (e.g., nanoviruses and circoviruses), T ¼ 1 capsids are not used by other known ssRNA viruses, which typically have larger T ¼ 3 or T ¼ 4 capsids. Furthermore, the CPs of all described plant satellite viruses have the same jelly-roll fold (Fig. 1), despite negligible sequence similarity. The similarities also extend to the genomic characteristics. In particular, in all plant satellite viruses, the linear ssRNA genomes lack the 50 7-methylguanylate caps and polyadenylation sequences in their 30 UTRs. Furthermore, 50 ends of the STNV and SPMV genomes are phosphorylated, whereas the first six

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nucleotides at the 50 -terminus of the STMV genome are identical to those in STNV. Considering these similarities, it is conceivable that all plant satellite viruses have evolved from a common ancestor. Notably, in structural comparison of the satellite virus CPs with the jelly-roll CPs of other viruses, satellite viruses formed a monophyletic clade, indicating that they indeed might share a common ancestor, despite high sequence divergence.

Concluding Remarks Due to their simplicity, satellite viruses have served (and continue to serve) as useful models for elucidating the molecular details underpinning various processes, including mechanisms of cap-independent protein translation, virion assembly and viral genome encapsidation. For instance, it has been recently demonstrated that assembly of the icosahedral STNV virion proceeds through recognition of the packaging signals on the viral RNA by the CP, which promotes the protein-protein interactions needed to build the capsid. Similar virion assembly mechanism has been also validated for more complex (T ¼ 3) bacterial and animal ssRNA viruses, underscoring the relevance of satellite viruses as model systems. Plant satellite viruses also provide a window into the complexity of natural ecosystems, where “one-on-one” virus-host encounters are likely to be rare, but rather involve multiple viruses and other types of genetic parasites interacting in an antagonistic or synergistic manner depending on environmental as well as virus- and host-specific cues. Thus, further studies on tripartite host-virus-satellite virus systems are likely to reveal new facets of virus-host and virus-virus interactions. Finally, despite their minimalism, the ultimate origins of satellite viruses remain enigmatic. On the one hand, it is possible that plant satellite viruses resemble the simplest, potentially ancestral, RNA viruses, which could have existed at the early stages of the evolution of the virosphere. On the other hand, it cannot be excluded that they are a result of reductive evolution from more complex quasi-autonomous RNA viruses. Further exploration of satellite virus diversity in other plants and animals should shed light on the evolutionary origins of this remarkable group of viruses.

Further Reading Ban, N., Larson, S.B., McPherson, A., 1995. Structural comparison of the plant satellite viruses. Virology 214 (2), 571–583. Bringloe, D.H., Pleij, C.W., Coutts, R.H., 1999. Mutation analysis of cis-elements in the 30 - and 50 -untranslated regions of satellite tobacco necrosis virus strain C RNA. Virology 264 (1), 76–84. Dodds, J.A., 1999. Satellite tobacco mosaic virus. Current Topics in Microbiology and Immunology 239, 145–157. Duponchel, S., Fischer, M.G., 2019. Viva lavidaviruses! Five features of virophages that parasitize giant DNA viruses. PLoS Pathogens 15 (3), e1007592. Ford, R.J., Barker, A.M., Bakker, S.E., et al., 2013. Sequence-specific, RNA-protein interactions overcome electrostatic barriers preventing assembly of satellite tobacco necrosis virus coat protein. Journal of Molecular Biology 425 (6), 1050–1064. Francki, R.I., 1985. Plant virus satellites. Annual Review of Microbiology 39, 151–174. Gnanasekaran, P., Chakraborty, S., 2018. Biology of viral satellites and their role in pathogenesis. Current Opinion in Virology 33, 96–105. Jones, I.M., Reichmann, M.E., 1973. The proteins synthesized in tobacco leaves infected with tobacco necrosis virus and satellite tobacco necrosis virus. Virology 52 (1), 49–56. Kassanis, B., 1962. Properties and behaviour of a virus depending for its multiplication on another. Journal of General Microbiology 27, 477–488. Kassanis, B., Nixon, H.L., 1960. Activation of one plant virus by another. Nature 187, 713–714. Kneller, E.L., Rakotondrafara, A.M., Miller, W.A., 2006. Cap-independent translation of plant viral RNAs. Virus Research 119 (1), 63–75. Krupovic, M., Cvirkaite-Krupovic, V., 2011. Virophages or satellite viruses? Nature Reviews Microbiology 9 (11), 762–763. Krupovic, M., Kuhn, J.H., Fischer, M.G., 2016. A classification system for virophages and satellite viruses. Archives of Virology 161 (1), 233–247. Larson, S.B., McPherson, A., 2001. Satellite tobacco mosaic virus RNA: Structure and implications for assembly. Current Opinion in Structural Biology 11 (1), 59–65. Mougari, S., Sahmi-Bounsiar, D., Levasseur, A., Colson, P., La Scola, B., 2019. Virophages of giant viruses: An update at eleven. Viruses 11 (8), E733. Murant, A.F., Mayo, M.A., 1982. Satellites of plant viruses. Annual Review of Phytopathology 20, 49–70. Patel, N., Dykeman, E.C., Coutts, R.H.A., et al., 2015. Revealing the density of encoded functions in a viral RNA. Proceedings of the National Academy of Sciences of the United States of America 112 (7), 2227–2232. Patel, N., Wroblewski, E., Leonov, G., et al., 2017. Rewriting nature's assembly manual for a ssRNA virus. Proceedings of the National Academy of Sciences of the United States of America 114 (46), 12255–12260. Scholthof, K.B., Jones, R.W., Jackson, A.O., 1999. Biology and structure of plant satellite viruses activated by icosahedral helper viruses. Current Topics in Microbiology and Immunology 239, 123–143. van Lipzig, R., Gultyaev, A.P., Pleij, C.W.A., et al., 2002. The 50 and 30 extremities of the satellite tobacco necrosis virus translational enhancer domain contribute differentially to stimulation of translation. RNA 8 (2), 229–236.

Plum Pox Virus (Potyviridae) Miroslav Glasa, Biomedical Research Center, Slovak Academy of Sciences, Bratislava, Slovakia Thierry Candresse, The National Research Institute for Agriculture, Food and the Environment, University of Bordeaux, Villenave d′Ornon, France r 2021 Elsevier Ltd. All rights reserved.

Glossary Asia Minor The westernmost extent of Western Asia, comprises most of the Asian part of modern Turkey and the Armenian highland. Cross-protection Protection of plant against the symptoms of a severe virus isolate by the pre-inoculation with a less severe isolate.

Interactomics Systematic survey of all interactions between viral proteins and plant cellular proteins Polyacrylamide gel electrophoresis (PAGE) An analytical method for separating biological macromolecules (proteins or nucleic acids) by size and charge. Sharka disease Sharka means pox in Bulgarian, as such symptoms on fruits were first observed in this country.

Introduction Plum pox virus (PPV), the agent responsible for the Sharka disease, belongs to the genus Potyvirus (family Potyviridae). The natural host range of this virus is restricted to Prunus spp. (stone fruits and ornamental trees). The infection of susceptible genotypes results in characteristic foliar and fruit symptoms and premature fruit drop. The geographical distribution covers the vast majority of Prunus producing area (except Australia, New Zealand, South Africa and the USA), although with widely different incidence levels in different countries. PPV is transmitted non-persistently by more than 20 aphid species, by grafting and vegetative multiplication of infected plants, but is not seed-borne. To date, at least 10 strains/molecular groups of PPV have been identified based on biological, serological, and molecular properties (M, D, Rec, EA, T, W, C, CR, CV and An). Although many diagnostic tools are available for the sensitive and/or specific detection of PPV, its huge variability, uneven distribution in infected woody hosts and its low titre outside of the active growth period significantly complicate its detection. In regions free of PPV, strict quarantine measures are usually enforced. With the limited availability of resistant or tolerant fruit tree varieties, a mix of prophylactic approaches including the use of virus-free propagation material, eradication of diseased trees, and vector control is generally used in an effort to control the virus in regions where it has not reached an endemic status.

Economical Importance PPV is considered as the most detrimental viral pathogen of stone fruit Prunus crops (peach, apricot, plum, Japanese plum). The disease is named according to the characteristic symptoms on fruits (sharka means pox in Bulgarian, as it was first observed in this country). During approximately a century of recognized existence, PPV has had a devastating impact on the global stone-fruit industry and nursery production, mainly in the central and south European countries. The damages are linked not only to the direct fruit yield and quality losses, but include also quarantine, eradication, prevention and compensatory measures. Since the 1970s the global negative impact of PPV has been estimated to exceed 10 billion euros. In the field, Prunus trees infected with PPV cannot be cured and are often eliminated as a consequence of disease eradication or containment efforts. PPV infected trees are usually not stunted and do not die, however, the infection is manifested by pronounced symptoms negatively affecting fruit yield and quality. The fruits of susceptible Prunus hosts are in some cases unsuitable for consumption or processing because of decreased weight and sugar content, overall lower gustative quality and visual defects (depressions, flesh browning, necrosis or gumming). The most spectacular symptom is the premature fruit drop, which may reach 80%–100% in the most susceptible cultivars. Consequently, traditional susceptible cultivars have in many places been replaced by less susceptible or tolerant cultivars, which are often of lower gustatory quality. The economic losses are not only associated with fruit production, but PPV also negatively affect the production of propagation materials (rootstock, budwood, scions) as in case of contamination these cannot be commercialized. Because of its high potential impact on stone fruit crops, PPV has been included in a list of the top ten plant viruses for impact or scientific interest and is listed as a quarantine pathogen in many parts of the world. In Europe for example, PPV is listed in the EC Plant Health Directive (Annex II of the European Union council directive 2000/29/EEC).

History and Geographical Distribution The sharka disease was observed for the first time around 1917 on plum cv. Kyustendil in the village of Zemen in western Bulgaria, near the Yugoslavian border. The viral character of the disease on plum and on apricot was described in 1933. Later, PPV was recognized on peach (in the 1960’s) and on cherries (in the 1980’s). Mainly after World War II, sharka was gradually reported in

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various Prunus species from different regions across the European continent and Mediterranean basin and later from other continents. Although these reports and their timescale do not necessarily represent the actual dissemination events, they suggest a progressive spread of PPV, probably through infected propagation material. Apart from the widespread distribution in Europe, the virus has been reported from South American (Chile, Argentina), the North American continent (USA, Canada), Asia (China, Japan, Asia Minor) and North Africa (Egypt, Tunisia). However, its prevalence varies from region to region from the endemic occurrence observed in central and eastern European countries, where the virus is well established, to local and more limited incidence observed in other countries where the virus was only introduced recently or where strict phytosanitary control measures have been enforced and have successfully kept the virus under some level of control (e.g., USA, where eradication was recently acheived).

Host Range and Symptomatology The natural host range of PPV is restricted to species of the genus Prunus (family Rosaceae), including cultivated stone fruits – plum (P. domestica), Japanese plum (P. salicina), apricot (P. armeniaca), peach and nectarine (P. persica), almond (P. dulcis) and myrobalan (P. cerasifera). Sweet cherry (P. avium), sour cherry (P. cerasus) and mahaleb cherry (P. mahaleb) can also be naturally infected by cherry-adapted PPV isolates. PPV also infects wild and ornamental species, such as nanking cherry (P. tomentosa), Japanese apricot (P. mume), Canadian plum (P. nigra), American plum (P. americana), dwarf flowering almond (P. glandulosa), and blackthorn (P. spinosa), which may thus act as local reservoirs of the virus. Symptoms on Prunus trees may be observed on leaves, flowers, fruits, and/or stones, however, they vary depending on host/cultivar susceptibility, virus isolate, physiological status and age of the host and environmental conditions (Fig. 1). The presence of other viruses in the trees, such as Prunus necrotic ringspot virus (PNRSV), prune dwarf virus (PDV) or apple chlorotic leafspot virus (ACLSV) may further increase the severity of symptoms. Under field conditions, the symptoms on Prunus plants are often masked late in the season or during the warm period of the growing season. Also, early infections are often characterized by symptoms restricted to only some parts of tree canopy and full systemic invasion of a tree may require several years. These factors may complicate in some cases the identification of the disease by visual inspection. Moreover, the irregular distribution and translocation of the virus in the trees and the low titre outside the active growth period may complicate the detection of the virus when methods of insufficient sensitivity are used. Typical foliar symptoms on plum consist generally of pale green chlorotic rings, spots, or patterns. Susceptible cultivars develop shallow ring or arabesque depressions on fruits, sometimes with brown or reddish necrotic flesh and gumming. Tolerant plum cultivars show no symptoms on fruits. Infected apricots develop chlorotic or pale-green rings and lines on leaves, light-colored depressed rings on fruits, which may be severely deformed. Stones are marked with typical discolored rings. Symptoms on susceptible peach genotypes are pronounced vein clearing, small chlorotic blotches, and distortions of the leaves. Color-breaking symptoms on the petals are observed in some varieties. Pale rings or diffuse band are visible on the skin of the fruits. In general, the symptoms on peach tend to be less visible in comparison to those on plums and apricots. Infection of almond is often symptomless or with limited foliar symptoms. Characteristic symptoms on cherries consist of pale green patterns and rings on leaves. Fruits may be slightly deformed, with chlorotic and necrotic rings and notched marks. Premature drop of fruits (reaching up to 100% in the most susceptible cultivars) is frequently observed, particularly in plum and apricot.

Virion Structure and Genome Properties The flexuous filamentous viral particles are approximately 750
15 nm. Viral particles are composed of the single-stranded genomic RNA encapsidated by a single type of capsid protein subunit. The genome is of positive polarity and is 9741–9795 nt in length. It has a polyadenylated 30 end and a virus-encoded protein (VPg) covalently bound at its 50 end. The genomic organization is typical of potyviruses, with a single open reading frame encoding a large polyprotein precursor (3125–3143 amino acids, c. 355 kDa) that is proteolytically processed by three virus encoded proteinases (P1, HC-Pro, and NIa-Pro) to yield as many as ten mature functional proteins. As in other potyviruses, an additional product, P3N-PIPO, produced by a polymerase slippage mechanism from an alternative short ORF embedded within the P3 coding sequence, has been predicted (Fig. 2).

Strains/Molecular Groups Initial attempts to study PPV variability involved the description of symptoms observed following mechanical inoculation of susceptible experimental herbaceous hosts. The most widely used biological classification system resulting from these efforts was based on the reaction of an experimental local-lesion host, Chenopodium foetidum, and PPV isolates were assigned as yellow, intermediate, and necrotic types depending on the type of local lesions they induced. It was also observed that different isolates

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Fig. 1 Typical leaf and fruit symptoms caused by Plum pox virus infection on susceptible Prunus genotypes, i.e., plum (P. domestica) (a, b, f), apricot (P. armeniaca) (c, d) and peach (P. persica) (e).

can induce variously severe symptoms on some woody indicators (Prunus persica GF305, P. tomentosa) after chip budding or controlled aphid transmission. However, such phenotypic characterization efforts remained often unreproducible because of difficulties to standardize the assays and of the influence, at least to some extent, of environmental conditions. Moreover, these experiments did not establish the link between these biological properties and the serological/molecular properties of PPV isolates. In the late 1970s, the existence of two serogroups of PPV was demonstrated using agar double diffusion assays employing polyclonal antibodies and a purified, formaldehyde-treated, suspension of undegraded viral particles. These two serogroups were named M and D, after the first inoculum sources used in these experiments (Marcus from peach in Greece and Dideron from apricot in south-eastern France, respectively).

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Fig. 2 Schematic representation of the Plum pox virus genome (A). The viral polyprotein and the alternative PIPO small ORF are represented by oval rectangles divided into boxes representing the respective functional products with their deduced length in amino acids (based on the AY028309 isolate). Polyprotein processing (B) by the three viral-encoded proteinases to yield the mature viral proteins. PIPO is expressed by a frameshifting resulting from a polymerase slippage mechanism.

It was later shown that different PPV isolates, belonging to these two serogroups (recognized as strains), show also reproducible coat protein (CP) mobility differences in denaturing polyacrylamide gel electrophoresis (PAGE) as well as a strain-specific Rsa I restriction site polymorphism in the region encoding the CP carboxy-terminus (CP C-terminus). However, some rare isolates behaved unusually using this RFLP typing assay, indicating that the serological and/or molecular variability of PPV is wider than that observed in these two D and M strains. Indeed, on the basis of recent molecular analysis, at least 10 strains of PPV, grouping isolates sharing common molecular (and, partially, serological) properties are now recognized. This classification has been validated by comparisons of complete genomic sequences of representative isolates for each of these strains. Although isolates of each strains were initially thought to have also in common some biological and epidemiological properties, such shared characteristics are now understood to exist only in some strains, and to be limited by the high intra-strain biological variability of PPV. Most PPV isolates can be assigned to one of the three major strains (PPV-M, D and Rec). Other strains are geographically limited PPV-W, EA and T, cherry adapted strains PPV-C, CR and CV and the atypical PPV-An strain. PPV-M (named after the type isolate Marcus). The isolates of this strain are present in many European countries, but absent from the Americas and China. They are often associated with rapidly spreading epidemics in peach but are less frequently found on plums. Usually, PPV-M isolates are transmitted efficiently by aphids. A high intra-strain molecular diversity was recently confirmed in PPV-M by the identification of divergent PPV-M-Ist isolates from Turkey. PPV-D (named after the type isolate Dideron). Based on geographical distribution data, PPV-D seems to be the most successful strain as PPV-D isolates are widespread in all areas where PPV has been reported, including the recent outbreaks in Asia and South and North America. PPV-D isolates naturally infect all the susceptible Prunus species excluding cherries, but are less frequently associated with spreading epidemics in peach. PPV-Rec (from “recombinant”). This group of isolates was recognized through the use of improved strain-typing methods targeting different genome portions. PPV-Rec genomes are characterized by a conserved homologous recombination event between PPV-M and PPV-D-like sequences, with a cross-over located in the 30 terminal part of the NIb coding region. PPV-Rec is therefore unrecognisable from PPV-M when using CP-based serological tests. Therefore, a number of isolates originally described as PPV-M were later retyped as belonging in fact to the PPV-Rec strain. PPV-Rec isolates are widespread in several central and eastern European countries, frequently associated with plums and efficiently transmitted by aphids. Interestingly, natural infections of peach by PPV-Rec are rare and mostly symptomless. PPV-W (from “Winona”). Although the first PPV-W report is from a few infected plum trees in Canada, the probable origin of this strain can be placed in Eastern Europe. PPV-W isolates were found naturally spreading and disseminated in different Prunus in Russia, Latvia and Ukraine. The Canadian PPV-W isolate shows multiple recombination events in its genome, one of which is absent in the genomes of East European PPV-W. PPV-W shows the highest intra-strain variability. PPV-W isolates are efficiently transmitted by aphids. Although not reported from central or western Europe, PPV-W isolates are competitive with the main strains in their regions of occurrence. PPV-EA (named after the type isolate El Amar) was originally isolated from apricots in the El Amar region of the Nile delta in Egypt in the late 1980s. It has also been observed in peach and in other regions of Egypt but has thus far not been reported outside this country. PPV-T (from “Turkey”). Found widespread in Turkey and also reported from Albania, PPV-T isolates are characterized by a unique recombination event affecting the 50 part of their genome. PPV-C (from “Cherry”) was first reported on sour cherry in Moldova and on sweet cherry in Italy. This group of cherry-adapted isolates has until now been sporadically reported from some central and eastern European countries, apparently without an epidemiological significance. PPV-C isolates are aphid transmissible and systemically infect sweet and sour cherry and P. mahaleb, however, they can also infect other Prunus species under experimental conditions. PPV-CR (from “Cherry Russian”) is the second strain naturally infecting cherries, although molecularly distinct from PPV-C. The PPV-CR isolates were found in two geographically distant areas of European Russia (Moscow and Samara regions). Similarly, to PPV-C, the isolates of this strain can be experimentally inoculated to other Prunus species and are transmitted by aphids.

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Fig. 3 Phylogenetic relationships of strains of PPV. (A) Unrooted phylogenetic tree computed using full-length genome sequences (available in the GenBank database, accessed on March 2019) showing the relationships between isolates of all ten recognized PPV strains. (B) Pairwise genetic distances calculated on the full-length genomes of representative isolates of all ten recognized PPV strains. The representative isolates are identified by their Genbank accession numbers.

PPV-CV (from “Cherry Volga”) is the third cherry-adapted strain of PPV, identified recently on sour cherries in the Tatarstan region (middle Volga river region of Russia). The sequence identity between PPV-CV isolates and those of other cherry-adapted strains (C, CR) reaches only ca. 83%, clearly separating them in phylogenetic analyzes into distinct monophyletic groups. Due to their restricted geographical dissemination and recent discovery, the epidemiological importance of these strains for cherry production remains to be determined. Cherry-adapted strains calculated intra-strain diversity are low, probably reflecting some epidemiological constraints limiting their genetic divergence, a possible recent emergence or insufficient sampling to date. PPV-An (from “ancestral”). A putative PPV strain represented by a recently identified isolate from eastern Albania. The full-length genomic sequence of this isolate fulfils the features of an ancestral PPV-M isolate previously hypothesized in the PPV evolutionary scenario. Full-length genomic sequences for more than 200 PPV isolates, representing each of the ten recognized strains, are available in public databases (Fig. 3(A)). Moreover, partial sequences, focusing mainly on the 3´ terminal part of genome or the P3–6K1 region, are available for a large number of isolates, making PPV one of the genetically best studied potyviruses. The comparison of complete genomic sequences revealed from 4.7% (PPV-D versus Rec) up to 28% (cherry adapted strains versus EA) nucleotide divergence between PPV strains (Fig. 3(B)). Recombination seems to have played a significant role in the evolutionary history of PPV. The PPV-Rec strain derives from a single recombination event involving isolates belonging to the PPV-D and PPV-M strains, with a recombination breakpoint in the C-terminus of the NIb gene. In addition, an analysis of complete genomic sequences has recently demonstrated that the PPV-D and PPV-M strains themselves share an ancestrally recombined 50 part of their genome (50 non-coding region, P1, HC-Pro, and N-terminus of P3). Other recently identified recombination events involve the W3174 isolate of PPV-W, which shows a mosaic structure as a consequence of recombination with PPV-M and PPV-D in the P1-HC region. The genome of PPV-T

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Fig. 4 Neighbor joining phylogenetic tree of the full-length genome sequences of selected potyviruses, including Plum pox virus (in bold). The scale bar indicates branch lengths in substitutions per nucleotide.

isolates shows a recombination in the HC-Pro-P3 region (with PPV-D as a minor parent). In addition, a recent analysis involving a larger set of PPV-T isolates identified other recombination breakpoints in the central part of the genomes of divergent isolates of this strain (Fig. 4). Thus, a growing number of available full-length genomes and of bioinformatics tools has enabled to show that besides the well-established recombinant strains (PPV-Rec, PPV-T), some other strains have recombination events in their ancient or more recent evolutionary history. Moreover, although less recognisable, it seems that at least for some PPV-W isolates, an intra-strain recombination event has also been documented. On the other hand, several strain (typical PPV-W, PPV-C, PPV-CR, PPV-CV and PPV-EA) appear to represent independent evolutionary lineages not affected by recombination. Although the complexity of PPV diversity is unlikely to have yet been fully uncovered, phylogenetic relationships based on complete genomes suggest that PPV evolved as two major branches or groups. One group involves the M, D, Rec, T, An and EA strains, that share partially or completely a common 50 genomic region as a result of an ancestral recombination. The second group is represented by the W, C, CR and CV strains. As most isolates of these latter strains were, up to now, detected in Russia and its historically linked East European countries, an influence of such geographical separation could be connected to their limited spread or absence in other countries.

Detection The recommended detection methods include biological indexing as well as serological and molecular assays. The use of woody biological indicators (such as peach GF305 or P. tomentosa) for diagnostic purposes is, however, restricted due to the labor intensive nature of such assays as well as to their length. Moreover, particular PPV strains do not seem to produce symptoms on these indicators with equal effectiveness (as in the case of GF305 and PPV-Rec). Specific monoclonal antibodies (MAbs) against several PPV serotypes (D, M/Rec, EA, W, C) have been prepared, although some isolates may show abnormal typing properties, escaping also to the recognition by broad-spectrum antibodies. A panel of molecular methods based on RT-PCR, real-time PCR or loop-mediated isothermal amplification can be used for the accurate identification or characterization of PPV strains. Some of these methods are part of the International Plant Protection ConventionFood and Agriculture Organization (IPPC-FAO) protocol for PPV diagnosis. Proper identification of recombinant isolates requires the use of several techniques targeting different parts of the viral genome or the use of specific primers with binding sites located on both sides of the targeted recombination breakpoint. In addition, whole genome sequencing, or sequencing of several strain-informative genome portions can provide unambiguous PPV strain identification. An additional level in our understanding of PPV diversity derives from the analysis of PPV-infected plants by high-throughput sequencing, enabling an unbiased identification of new sequence variants. Moreover, the analysis of mixed infections involving different PPV variants/strains and their persistence, until now not well documented, is another challenge in PPV research.

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Virus Spread Natural spread of PPV occurs through aphid vectors from neighboring infected reservoirs. The aphids can acquire the virus from leaves or young shoots of cultivated, wild, or ornamental Prunus trees, but also from contaminated fruits, indicating that fruit shipments could also represent a minor pathway for international virus movement. The experimental host range of PPV is large, with over 60 reported host species (such as Chenopodium foetidum, Nicotiana benthamiana and N. clevelandii, Pisum sativum, Ranunculus arvensis, Senecio vulgaris, Stellaria media, etc.) in eight families. However, no epidemiologically significant contribution of weeds and annual herbaceous plants to the spread of PPV has been reported. The most efficient long-range movement of the virus is linked to international exchange of contaminated Prunus propagation materials (rootstocks, budwood, nursery plants). PPV is thus usually introduced in a new location by infected propagation material. Once established in the plantation, the virus is rapidly spread by aphid vectors when making test probes and migrating between plants. PPV is naturally transmitted by over 20 different aphid species (Myzus persicae, M. varians, Aphis gossypii, A. craccivora, A. hederae, A. spiraecola, Brachycaudus cardui, B. helichrysi, B. persicae, Phorodon humuli, Rhopalosiphum padi, etc.). The virus is transmitted in a non-persistent manner so that aphids may acquire it very rapidly when probing infected plants with their stylets (piercing–sucking mouthparts). However, aphids can only transmit the virus for a short time after virus acquisition as it is retained in a viable state in the aphid’s mouthparts for a period of only a few minutes to a few hours. The virus appears to be lost by the aphids the first time they probe on the next plant and aphids do not remain infectious after molting nor do they pass PPV onto their progeny. The efficiency of transmission depends on the particular aphid species, on the virus isolate and on the host species from which the virus is acquired and to which it is transmitted. As for other potyviruses, two viral proteins are known to be involved in the transmission mechanism, the HC-Pro (helper component) and the CP. The DAG and PTK amino acid motifs are highly conserved in the N-terminus of the CP and HC protein of aphid-transmissible PPV isolates, respectively, and loss of transmissibility is correlated with their deletion or modification. Although the virus was detected in fruits and seed coat and some conflicting results have been reported in the past concerning seed transmission of PPV in some Prunus species, recent studies have failed to demonstrate seed transmission of PPV in any of its woody hosts.

Ecology and Control In the countries and regions where the virus is not yet present (e.g., Australia), strict quarantine measures need to be established to prevent the introduction of PPV through legal or illegal importation of infected fruit tree propagation material or fruits. In regions where the disease is present but still localized and under control, a mix of prophylactic approaches is usually implemented, including the application of programmes for production and use of virus-free propagation material (rootstocks, budwoods), tight aphid vector control in nurseries (e.g., by application of mineral oil), regular inspection of orchards, and prompt eradication of infected plants. In regions of endemic PPV occurrence, where eradication brings no solution to the spread of the disease, the practical measures to pursue stone fruit production rely on the use of less susceptible or tolerant genotypes. This practice, however, does not restrict the virus spread and, paradoxically, contributed in the past to the destruction of traditional high-quality genotypes in the central and south-eastern European countries (e.g., Pozegaca or Bistrita plums). Early attempts to use cross-protection with attenuated virus isolates, a technique successfully used to control some other potyviruses (Zucchini yellow mosaic virus, Papaya ringspot virus), have not met with success in the case of PPV. In theory, a long term PPV control could be achieved using PPV-resistant varieties. Therefore, many breeding programmes aiming at the development of such varieties have been engaged. They were based on the use of natural resistance sources identified mainly within the P. armeniaca, P. davidiana and P. domestica germplasms. Efficient breeding progress is, however, complicated by the high degree of heterozygosis, the existence of incompatibility barriers between species, the long generation times and the length and non-standardisation of the resistance tests. Some progress has been achieved in plum and apricot, with the deployment of a few resistant varieties in some countries. However, the durability of these resistances remains an open question and so is their stability if confronted to the high diversity of PPV strains. Development of genotypes with a hypersensitive response as an active defense mechanism against PPV is another promising way to produce resistant fruit trees or rootstocks, as was demonstrated for some plum varieties (e.g., Jojo), but no hypersensitivity sources are so far known in apricot and peach. The pathogen-derived resistance strategy and the use of sequence-specific gene silencing offer a complementary approach for the development of PPV-resistant stone fruit cultivars. Experimental herbaceous hosts or Prunus plants transformed with different regions of the PPV genome (i.e., P1, HCPro, CI, NIa, NIb, or CP) or with post-transcriptional gene silencing (PTGS)-inducing hairpin-constructs containing viral transgenes have been developed and shown to provide partial or complete resistance to PPV. The transgenic plum “HoneySweet”, transformed with a sense PPV CP gene construct, has been shown to be highly resistant to PPV infection and was recently approved for commercial cultivation in the USA, although its use is so far local and limited. The most recently explored strategies rely on interactomics or genetic studies aimed at the identification and inactivation of host susceptibility factors, such as the translation initiation factor eIF4E and its isoforms.

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Further Reading Cirilli, M., Geuna, F., Babini, A.R., et al., 2016. Fighting sharka in pach: Current limitations and future perspectives. Frontiers in Plant Science 7, 1290. doi:10.3389/fpls.2016.01290. Clemente-Moreno, M.J., Hernández, J.A., Diaz-Vivancos, P., 2015. Sharka: How do plants respond to plum pox virus infection? Journal of Experimental Botany 66, 25–35. García, J.A., Glasa, M., Cambra, M., Candresse, T., 2014. Plum pox virus and sharka: A model potyvirus and a major disease. Molecular Plant Pathology 15, 226–241. Illardi, V., Tavazza, M., 2015. Biotechnological strategies and tools for plum pox virus resistance: Trans-, intra-, cis-genesis, and beyond. Frontiers in Plant Science 6, 379. doi:10.3389/fpls.2015.00379. Rimbaud, L., Dallot, S., Gottwald, T., et al., 2015. Sharka epidemiology and worldwide management strategies: Learning lessons to optimize disease control in perennial plants. Annual Review of Phytopathology 53, 357–378.

Relevant Websites https://www.ippc.int/en/publications/dp-2-2012-plum-pox-virus/ DP 02: Plum pox virus. https://gd.eppo.int/taxon/PPV000 Plum pox virus (PPV000). https://www.cabi.org/isc/datasheet/42203 Plum pox virus (sharka). https://www.sharco.eu/ SharCo. Accueil.

Poleroviruses (Luteoviridae) Hernan Garcia-Ruiz, Natalie M Holste, and Katherine LaTourrette, University of Nebraska–Lincoln, Lincoln, NE, United States r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

mRNA messenger RNA nt Nucleotide(s) ORF Open Reading Frame Rap1 Replication associated protein 1 RdRp RNA-dependent RNA polymerase RISC RNA-induced silencing complex satRNA satellite RNA UTR Untranslated region VIGS Virus-induced gene silencing VPg Viral protein genome-linked

AGO Argonaute proteins CP Coat protein or capsid protein ELISA Enzyme-linked immunological assays ER Endoplasmic reticulum HC-Pro Helper component-protease IRES Internal ribosomal entry site kb Kilobase kDa KiloDalton MP Movement protein

Glossary Aphid-borne Is a term used to describe aphid-transmitted viruses that infect plants. Argonaute Are the catalytic component of RNA-induced silencing complexes. They associate with cellular or virusderived small interfering RNAs. Together, this complex targets cellular or viral RNA for degradation or translational repression, resulting in gene silencing. F-box Is a conserved region of P0 that mediates interaction with Argonaute protein 1, leading to Argonaute protein 1 ubiquitination and degradation. Gene silencing Also known as RNA interference, is a sequence-specific gene inactivation system mediated by small interfering RNAs (siRNAs). It downregulates RNA accumulation without affecting DNA sequence. Leaky scanning Happens during initiation of translation when a start codon is skipped over in preference for another start codon located downstream on the mRNA. This process allows translation of several proteins from the same mRNA. P0 Is the silencing suppressor encoded for by poleroviruses.

Phloem-limited Refers to viruses that can only replicate in the phloem of the host plant. These viruses cannot infect mesophyll cells and are generally not transmitted mechanically. Poleroviruses Are viruses in the genus Polerovirus within the family Luteoviridae. They are obligatorily transmitted by aphids, phloem-limited, and encode a strong silencing suppressor (P0). Ribosomal frameshift Occurs during translation of an mRNA with two overlapping open reading frames. During the shift, the ribosome slips one base backward towards the 50 -end or one base forward towards the 30 -end. The result is the fusion of two proteins encoded for by different open reading frames on the same mRNA. Ribosomal read-through Occurs when ribosomes overlook a stop codon and continue translation. Silencing suppressors Are proteins encoded by viruses, which interfere with gene silencing. They prevent degradation of viral RNA, enhance transcription of viral RNA, and alter silencing of cellular genes. Combined, these effects are in part responsible for symptom development in virus- infected plants.

Introduction The family Luteoviridae consists exclusively of plant infecting viruses divided into three genera: Luteovirus, Polerovirus, and Enamovirus (Table 1). Luteoviruses diversified into three genera approximately 1500 years ago in correlation with the expansion of agriculture. Members of the genus Luteovirus (luteoviruses) contain the standard genome of the entire Luteoviridae family: a single, positive single-stranded RNA (ssRNA) encoding the seven proteins P1 through P7 with multiple overlapping open reading frames (ORFs). Barley yellow dwarf virus (BYDV) is the type species of the genus Luteovirus, which encode a weak silencing suppressor (P4). Members of the genus Polerovirus (poleroviruses) are similar to luteoviruses and contain an additional protein (P0) that is a strong silencing suppressor. Luteoviruses and poleroviruses shared a common ancestor approximately 900 years ago with P0 deriving separately in the polerovirus lineage. The type species of the genus Polerovirus is Potato leafroll virus (PLRV). Pea enation mosaic virus (PEMV) is the type species of the genus Enamovirus (enamoviruses), which encode P0 but not P4. Within the family Luteoviridae, BYDV and PLRV infect important staple crops and cause major economic damage. However, poleroviruses are the most damaging and diverse genus, and have a wide host range. Poleroviruses have a single, positive ssRNA genome of 5.3–5.7 kb, encapsidated in an icosahedral non-enveloped virion (Fig. 1). Unique features of poleroviruses include obligate transmission by aphids in a circulative, non-propagative manner, infection restricted to the phloem, and lack of mechanical transmission. Symptoms induced by poleroviruses include stunting,

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Table 1

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Taxonomic organization of the family Luteoviridae. Species are grouped by genus

Genus

Species

Abbreviation

Accession numbera

Luteovirus

Barley yellow dwarf virus KerII Barley yellow dwarf virus KerIII Barley yellow dwarf virus – MAV Barley yellow dwarf virus – PAS Barley yellow dwarf virus – PAV Bean leafroll virus Nectarine stem pitting-associated virus Rose spring dwarf-associated virus Soybean dwarf virus

BYDV-KerII BYDV-KerIII BYDV-MAV BYDV-PAS BYDV-PAV BLRV NSPaV RSDaV SbDV

NC_021481.1 KC559092.1 NC_003680.1 NC_002160.2 NC_004750.1 NC_003369.1 NC_027211.1 NC_010806.1 NC_003056.1

Polerovirus

Beet chlorosis virus Beet mild yellowing virus Beet western yellows virus Carrot red leaf virus Cereal yellow dwarf virus – RPS Cereal yellow dwarf virus – RPV Chickpea chlorotic stunt virus Cotton leafroll dwarf virus Cucurbit aphid-borne yellows virus Maize yellow dwarf virus RMV Maize yellow mosaic virus Melon aphid-borne yellows virus Pepo aphid-borne yellows virus Pepper vein yellows virus Pepper vein yellows virus 5 Potato leafroll virus Suakwa aphid-borne yellows virus Sugarcane yellow leaf virus Tobacco vein distorting virus Turnip yellows virus

BChV BMYV BWYV CRLV CYDV-RPS CYDV-RPV CpCSV CLRV CABYV MYDV-RMV MYMV MABYV PABYV PVYV PVYV  5 PLRV SABYV ScYLV TVDV TuYV

NC_002766.1 NC_003491.1 NC_004756.1 NC_006265.1 NC_002198.2 NC_004751.1 NC_008249.1 NC_014545.1 NC_003688.1 NC_021484.1 KU248489.1 NC_010809.1 NC_030225.1 NC_015050.1 NC_036803.1 NC_001747.1 NC_018571.2 NC_000874.1 NC_010732.1 NC_003743.1

Enamovirus

Alfalfa enamovirus 1 Citrus vein enation virus Grapevine enamovirus 1 Pea enation mosaic virus 1

AEV  1 CVEV GVEV  1 PEMV  1

NC_029993.1 NC_021564.1 NC_034836.1 NC_003629.1

Unassigned

Barley yellow dwarf virus – GPV Barley yellow dwarf virus – SGV Chickpea stunt disease associated virus Groundnut rosette assistor virus Indonesian soybean dwarf virus Sweet potato leaf speckling virus Tobacco necrotic dwarf virus

BYDV-GPV BYDV-SGV CpSDaV GRAV ISDV SPLSV TNDV

NC_039035.1 AY541039.1 Y11530.1 NC_038509.1 NC_038510.1

a

Accession numbers in GenBank. Accessions beginning with NC_ are the reference for a particular species.

yellowing, a streaking pattern, and stiff leaves. These symptoms are often confused with adverse environmental factors. Most poleroviruses can be present in seed, tubers, and plant parts used for vegetative propagation. Furthermore, some plant-virus combinations remain asymptomatic. Currently, there are 32 species in the genus Polerovirus infecting both monocots and dicots (Table 1). These include economically important staple crops including maize, wheat, sugarcane, and potato. Poleroviruses with the most economic importance are PLRV, Sugarcane yellow leaf virus, and three beet-infecting poleroviruses.

Polerovirus Physical Properties The polerovirus virion has a T3 icosohedral symmetry with an average diameter of 23 nm (Fig. 1). The capsid is formed by 180 monomers that consist mainly of the coat protein (CP) (approximately 23 kDa) and also contain minor amounts of a readthrough protein (approximately 80 kDa). The readthrough protein substitutes one coat protein monomer when assembling the virion. The ratio of CP to readthrough protein varies from 4:1 to 100:1. The thermal inactivation point is between 501C and 651C, with a dilution endpoint between 10–3 and 10–4. Polerovirus virions withstand deep-freeze and thaw, and withstand chloroform and detergents. The longevity in sap at 21C is between 5 and 10 days.

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Fig. 1 Illustration of polerovirus virion structure based on potato leafroll virus (PLRV). (A) Transmission electron microscope picture of PLRV virions. The bar represents 100 nm. Photograph courtesy of I.M. Roberts. (B) The coat protein (CP) creates a virion that with T¼3 icosahedral symmetry composed of 180 capsid proteins organized into 60 asymmetric units. colored according to CP quasi-conformers, where subunit A is blue, subunit B is green, and subunit C is red. There is no envelope and the diameter averages 23 nm. (C) Cryo-electron microscopy of PLRV-like particles. Section of representative density and molecular model, slice through unsharpened maps, depicting density for packaged RNA and/or disordered R domain. (D) cryo-EM maps of whole virus capsid. Structure refinement was carried out with icosahedral symmetry imposed, yielding density maps at a resolution of 3.4 Å . Reproduced with permission from Byrne, M.J., Steele, J.F.C., Hesketh, E.L., et al., 2019. Combining transient expression and cryo-EM to obtain high-resolution structures of luteovirid particles. Structure 27, 1761–1770. doi:10.1016/j.str.2019.09.010.

Polerovirus Genome Organization and Gene Expression The polerovirus genome consists of a single, positive-strand ssRNA encoding P0 through P7 organized in overlapping ORFs (Fig. 2(A)). The 50 -end is protected by the genome-linked protein VPg. The 30 -end contains an -OH group and lacks a poly-A tail. Two sub-genomic RNAs are formed during replication. Translation of polerovirus proteins involves a combination of strategies: leaky scanning, internal ribosomal entry, frameshift, and ribosomal read-through (Fig. 2(B)). Additionally, VPg is released from protein P1 by protease processing. Proteins P0 and P1 are translated from the genomic RNA using leaky scanning and alternate translation initiation codons (Fig. 2(B)). P1 can be expressed either individually or fused with P2. When P1 is expressed by itself, it contains two putative domains: VPg and a protease that releases VPg. A ribosomal frameshift produces a P1–P2 fusion protein that generates the RNAdependent RNA polymerase (RdRp) responsible for viral RNA replication and sub-genomic RNA synthesis. P2 is never expressed by itself. Replication associated protein 1 (Rap1) is translated from genomic RNA through an internal ribosome entry site (IRES). Protein 3a and the movement protein (MP, P4) are translated by leaky scanning from sub-genomic RNA1 and both are involved in virus movement along with the CP. A ribosomal read-through is required for the translation of the CP read-through (P3–P5), which is less abundant than the CP (P3). An amber stop codon (UAG) separates these ORFs in sub-genomic RNA1 (Fig. 2(B)). The CP readthrough is not necessary for virion formation, but it is essential for aphid transmission and virus movement in plants. The N-terminal half of the CP read-through determines vector specificity by regulating the efficiency of virus movement through the salivary tissues and gut. Accordingly, mutants lacking the CP read-through accumulate to low levels and are not transmitted by vectors. The C terminal half of the CP read-through is involved in efficient virus movement, tissue tropism, and symptom development in plants. Several proteins in the genome are currently not well understood. P3a is newly discovered part of the genome. It sits directly upstream of the CP ORF (P3) and is translated by a non-AUG start codon. P3a is required for long-distance movement of poleroviruses. Proteins P6 and P7 are translated by leaky scanning from sub-genomic RNA2. P7 has nucleic acid binding properties. However, the biological role of P6 and P7 remains to be determined.

Polerovirus Phylogenetic Diversity The evolutionary relationship and phylogenetic diversity of poleroviruses is just beginning to be elucidated. Published studies used the CP and read-through domains based on the assumption that they are highly conserved (Fig. 3). Based on the CP

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Fig. 2 Schematic representation of polerovirus genome organization and gene expression. Single lines represent non-coding regions and labeled boxes represent cistrons. Sub-genomic RNAs, their formation and proteins translated from them are indicated. (A) Generalized polerovirus genome organization. Coordinates are based on Potato leafroll virus accession number KY856831. (B) Polerovirus gene expression strategies include formation of sub-genomic RNAs, translation by IRES-mediated internal initiation, leaky scanning, ribosomal frameshift, and ribosomal read-through. Protein 1 is processed into mature VPg by proteolysis. Pro: Putative protease. VPg: Viral protein genome-linked. RdRp: RNA-dependent RNA polymerase. Rap1: Replication associated protein. CP: Capsid protein, major and minor. MP: Putative movement protein. p3a: Protein essential for systemic virus movement. IRES: Internal ribosomal entry site.

(Fig. 3(A)), PLRV is an out-group, while Pepper yellow leaf curl virus (PYLCV), Pepper vein yellows virus (PeVYV), and Pepper yellows virus (PepYV) clustered on the same branch, and probably evolved from TVDV. However, the N-terminus of the CP read-through (Fig. 3(B)) separates Tobacco vein-distorting virus (TVDV) and places Cucurbit aphid-borne yellows virus (CABYV) close to pepper-infecting poleroviruses. In contrast, the C-terminus of the CP read-through (Fig. 3(C)) separates pepper-infecting poleroviruses and place TVDV close to PYLCV. Differences in the arrangements of poleroviruses based on CP, N, or C terminal parts of the CP read-through suggest that RNA recombination occurs frequently and is an important contributor to the evolution of poleroviruses.

Polerovirus Transmission Polerovirus species have evolved to be efficiently transmitted by particular aphid species. PLRV is efficiently transmitted by Myzus persicae, while maize poleroviruses are efficiently transmitted by the corn leaf aphid (Rhopalosiphum maidis). Other aphids that vector poleroviruses include R. padi, Stiobion avenae, and Aphis gossypi. Aphids vector poleroviruses in a circulative, non-propagative manner. The cycle begins when an aphid feeds on a polerovirus-infected plant. The virus first will reach the salivary glands of the aphid. It has been found that the read-through domain is not required for the virion to cross the salivary gland, but it does improve the success of the transport. If the species is from a yellow dwarf lineage, the virion then moves through the hindgut. If the species

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Fig. 3 Phylogenetic analysis of selected poleroviruses based on the following proteins: (A) CP (ORF3), (B) the N-terminus of the RTD (ORF5, first 233 aa), (C) the C-terminus of the RTD (ORF5, last 262 aa) of Potato leaf curl virus [PLRV (Y07496)], Tobacco vein-distorting virus [TVDV (EF529624)], Cucurbit aphid-borne yellows virus [CABYV (X76931)], Pepper vein yellows virus [PeVYV (AB5948280)], Pepper yellow leaf curl virus [PYLCV (HM439608)], and Pepper yellows virus [(PepYV) FN600344]. Adapted with permission from Dombrovsky, A., Glanz, E., Lachman, O., et al., 2013. The complete genomics of Pepper yellow leaf curl virus (PYLCV) and its implications for our understanding of evolution dynamics in the genus Polerovirus. PLoS One 8, e70722. doi:10.1371/journal.pone.0070722.

mainly uses dicots as their host, such as PLRV or Beet western yellows virus (BWYV), the virion instead moves through the midgut. Once inside the gut, the virus normally moves between the cytoplasm and the epithelial cells. It then fuses with the plasmalemma and is released between the basal lamina and the membrane. Aphids can then release the virion into the phloem parenchyma and/ or the companion cells to initiate local infection. Cell-to-cell and systemic infection may occur and require the combined activity of the movement protein, capsid protein, capsid protein read-through, and P0. Phloem-limited viruses cannot normally be mechanically transmitted. However, using particle bombardment, infection has been achieved with PLRV and BWYV. The high number of aphid vectors allows poleroviruses to infect a wide range of hosts. Potatoes, sugarcane, and beets are important species infected by poleroviruses. These all propagate in a vegetative manner. Poleroviruses can spread through infected contaminated plants parts used for propagation, such as tubers and sugarcane cuttings.

Virus-Virus Interactions Co-infections of PLRV with Potato virus X (PVX) (Potexvirus) or Potato virus Y (PVY) (Potyvirus) result in an enhancement in symptom severity and yield loss. Similarly, co-infection of Brassica yellows virus (BrYV) (unassigned Polerovirus) and Pea enation mosaic virus 2 (PEMV-2) (Umbravirus) results in similar synergism. PEMV 2 is an umbravirus that accompanies PEMV as a satellite virus. PEMV-2 can only infect plants when a member of the Luteoviridae is present. Co-infection of BrYV and PEMV-2 results in higher accumulation of BrYV, more severe symptoms, and the acquisition of mechanical transmission. Co-infection of BWYV and the potyvirus Beet mosaic virus (BtMV) causes faster systemic virus movement and earlier, more severe symptoms. The combination of the polerovirus and the potyvirus disrupts photosynthesis and vascular transport, and both

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viruses accumulate to high levels when co-infected. Experimentally, the potyvirus helper-component proteinase (HC-Pro), a strong silencing suppressor, increased the accumulation of PLRV. It also allowed the virus to spread into mesophyll cells. Maize lethal necrosis is a re-emerging disease of current epidemic proportions in Sub-Saharan Africa. Maize lethal necrosis disease was discovered in 1976 in Nebraska and Kansas, USA. It was caused by Maize chlorotic mottle virus (MCMV) in combination with a potyvirus, Sugarcane mosaic virus (SCMV). Several recent studies have found poleroviruses in maize in combination with MCMV, SCMV, or both. The poleroviruses detected include the Maize yellow dwarf virus-RMV (MYDV-RMV), Maize yellow mosaic virus (MaYMV), and Barley virus G (BVG). However, MYDV-RMV was the most common. Plants with a combination of a polerovirus, SCMV, and MCMV have atypical symptoms. These and other observations suggest that poleroviruses, in combination with MCMV, could also cause maize lethal necrosis disease.

Silencing Suppression by Poleroviruses In plants, gene silencing is an essential component of antiviral defense. Virus infection induces antiviral gene silencing. Argonaute (AGO) proteins are the catalytic components of the RNA-induced silencing complex (RISC) and associate with cellular or virusderived small interfering RNAs (siRNA). Binary complexes formed between argonaute proteins and siRNAs specifically target RNA, including viral RNA, complementary to the siRNA. They have also been implicated in cell-to-cell and systemic movement of gene silencing signals. This results in amplification of gene silencing in areas beyond the initial activation site, thereby conferring virus immunity. In order to establish infection and move within plants, viruses encode specialized proteins that suppress gene silencing. In poleroviruses, P0 is a silencing suppressor. P0 silencing suppression activity has been demonstrated for 10 of the 32 poleroviruses (Table 1). For Beet chlorosis virus (BChV), and some strains of Beet mild yellowing virus (BMYV), no silencing suppression activity was found for P0. For all other poleroviruses, no information is available or the suppression activity of P0 has not been determined. In standard experimental assays, for some poleroviruses, P0 suppresses either local or both local and systemic gene silencing. P0 suppresses gene silencing by targeting argonaute 1 (AGO1) protein for degradation by ubiquitination. Through the F-boxlike motif, P0 interacts with the DUF1785 motif in AGO1 to mark it for degradation. In Arabidopsis thaliana, the P0 F-box-like motif interacts with the F-box of the S phase kinase-associated protein 1 (SKP1) and with the ASK1 and ASK2 orthologues. AGO1 is the primary interaction partner of microRNAs and is a crucial for normal plant development. Thus, symptoms induced by polerovirus infection are in part due to the effect on AGO1, and potentially other AGO proteins being tagged for degraded by P0 which in turn affects normal plant development.

Potato Leafroll Virus PLRV is the first polerovirus discovered, one of the most damaging poleroviruses worldwide, and the most damaging potato virus. It is highly prevalent and has been found on every continent except Antarctica. PLRV was first detected in the 1770s, causes 50%–60% yield loss, and costs the United States 100 million-dollars yearly. PLRV is transmitted by infected tubers and by aphids. When the virus is transmitted by aphids, symptoms begin in young, top leaves that roll and turn pale. When grown from an infected tuber, the plants may be pale or dwarfed, and the leaves may be upright, rolled, yellow, or brittle (Fig. 4(A)). However, the appearance of water-soaked leaves is usually the first symptom. In the stem and the tuber sieve tubes, abnormal amounts of callose accumulates. The carbohydrates in the leaves reach high levels causing the phloem transport to be impaired, which results in tuber reduction. This could occur because photo-assimilation is reduced, sucrose is unable to enter the phloem, or a combination of the two. These factors result in leaves with an upright and rolled appearance. In some cultivars, the margins of the leaves may turn purple or red and develop necrosis in later stages. This necrosis starts in the phloem of the petioles and stems.

Sugarcane Yellow Leaf Virus Worldwide, damage by poleroviruses in sugarcane is a close second to PLRV. ScYLV is a good representation of the yellow leaf or yellow dwarf viruses amongst poleroviruses. ScYLV was first discovered in Hawaii in 1989 and is distributed world-wide. Currently, it is primarily detected in South America, Asia, and the Pacific islands. ScYLV affects sugarcane production in over 90 countries, which grow sugarcane for sugar, biofuel, and fibers. Crops affected by ScYLV have losses that reach up to 43%, and it is spread by infected seed canes and aphids. ScYLV infection reduces the cane thickness, the number of canes produced, and the rate of photosynthesis in the plant. It causes yellowing at the midrib (Fig. 4(B)), and, at 6–8 months, the yellowing spreads laterally to the leaf lamina and causes necrosis at the tip. Because of the short internode spacing, this causes the plant to be dwarfed. Interestingly, when co-infected with a certain bacterium, Leifsonia xyli sub species xyli, it increases the severity of the disease. Even when the virus is latent in the plant, the yield is decreased, especially in non-resistant varieties. Unfortunately, all yellow leaf viruses are hard to distinguish from normal environmental damage.

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Fig. 4 Representative symptoms, in leaves and whole plants, caused by the top three poleroviruses. Other features of the symptoms are described below the images. (A) Symptoms caused by Potato leafroll virus in potato plants and leaves. (B) Symptoms caused by Sugarcane yellow leaf virus in sugarcane. (C) Symptoms caused by Beet western yellows virus is sugar beet. Reproduced with permission from (A) Jack Kelly Clark, University of California Statewide IPM Program. (B) CIRAD: The French Agricultural Research Organization working for the sustainable development of tropical and Mediterranean regions. (C) G.J. Holmes, California Polytechnic State University at San Luis Obispo.

Beet Poleroviruses The three main poleroviruses infecting beet are BWYV, BMYV, and BChV (Table 1). Most of the strains are different isolates of BWYV originating from Europe. Within the last decade, the virus has spread to Australia and resulted in a 26% yield loss in pluses, canola, and various vegetables. In canola, 59% of the 65% yield loss was due to BWYV alone. BMYV is also known to result in about 22% crop loss if it appears in June. However, it has been found that these viruses have a much broader host range within the whole Amaranthacae family, temperate legume crops, and brassicas. They also widely infect the weeds that grow around these crops. Of the beet-infecting poleroviruses, BChV has a smaller a host range because it only infects sugar beets. These persistent viruses are transmitted by a wide variety of aphids, with the highest being the genus Myzus. Normally, sugar beets are durable crops that are tolerant to drought and can withstand intense wilting with no yield loss. However, poleroviruses cause stunting, chlorosis, rolling leaves, stiff followed by brittle leaves, and yellowing in the veins (Fig. 4(C)). Some leaves may turn orange rather than the typical yellow, which starts at the tip of the leaves. Beet-infecting poleroviruses remain problematic because of their impacts on crops, wide host range, and difficulty to diagnosis.

Diagnosis There are several methods of detecting poleroviruses. The first test designed was the Ingel–Lange test. This test stained callose with a resorcin blue dye in the tubers of a potato infected with PLRV. Currently, poleroviruses are detected using a variety of RNA-based

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and protein-based approaches. Protein-based approaches use antibodies that recognize the capsid protein. The most common protein-based approach is enzyme-linked immunosorbent assay (ELISA). A variation is the double antibody sandwich ELISA. Other protein-based approaches include immune-electron microscopy, immune-electrophoresis, and double diffusion agar tests. RNA-based approaches include reverse-transcription polymerase chain reaction (RT-PCR), deep sequencing of the siRNAs, and high-throughput sequencing of transcript RNA. There are universal polerovirus primers for detection by RT-PCR. Universal primers Pol-G-F and Pol-G-R amplify a 1.4 kb PCR product spanning part of the RdRp gene, the intergenic region, and the complete CP gene. Additionally, northern blotting may be used to detect genomic and sub-genomic RNAs. Electron microscopy is commonly used to visually detect poleroviruses. Using the transmission electron microscope, Cereal yellow dwarf virus (CYDV), BWYV, PLRV, and CABYV have been observed being transported through the gut and epithelial cells into the aphid. Electron microscopy was also used to track the readthrough domain to determine its role inside the aphid.

Disease Management Polerovirus resistant plants are not common. In crops of economic importance, there are no polerovirus-resistant varieties. Varieties that do exist are only resistant to one or a few strains of the virus. Because resistant cultivars impose selection pressure and viruses mutate quickly, viruses break genetic resistance within a few years. Since poleroviruses are vectored by aphids, physical barriers have been implemented to prevent polerovirus spread. Plastic reflective mulch can be placed around the crops. UV wavelengths will be reflected, thus repelling the aphids. Floating row covers with a fine mesh can also physically block the aphids from reaching the plant. Topical approaches include mineral oil and aphicides. Mineral oil can be applied to crops to smother aphids. Aphicides can be used on plants similar to chemical pesticides. Carefully planned planting can reduce virus transmission by aphids. Planting when aphids are low, like after a short rain season, could potentially reduce the number of aphids in the field. This strategy can then be combined with timed pesticide use. Government restricted closed seasons for planting certain crops will also reduce polerovirus transmission. Several countries already restrict crop planting during certain times, but the approach could be further expanded around the globe. Farmers should also practice good crop rotation and diversification. A maize and soybean rotation will likely have different aphid vectors, so there is a decreased likelihood of continual crop infection. A less practiced method is to control the weeds in the field. Weeds that are not removed between crop planting could still be infected, thus leading to crop infection in the next season. It is also good practice to burn any infected plant material because it is the only surefire method to destroy the virus. This method also eliminates the possibility aphids may feed on infected plant material and spread the infection to healthy plants. Aphids are attracted to bare soil, so farmers should work to plant crops closer, include cover crops, and have untilled soil. Preventative measures provide options to limit poleroviruses exposure, but the possibility of resistant plants remains the only option to fight infection directly.

Further Reading Byrne, M.J., Steele, J.F.C., Hesketh, E.L., et al., 2019. Combining transient expression and cryo-EM to obtain high-resolution structures of luteovirid particles. Structure 27, 1761–1770. doi:10.1016/j.str.2019.09.010. Chen, S., Jiang, G., Wu, J., et al., 2016. Characterization of a novel polerovirus infecting maize in China. Viruses 8 (5), doi:10.3390/v8050120. DeBlasio, S.L., Chavez, J.D., Alexander, M.M., et al., 2015. Visualization of host-polerovirus interaction topologies using protein interaction reporter technology. Journal of Virology 90 (4), 1973–1987. doi:10.1128/JVI.01706-15. DeBlasio, S.L., Xu, Y., Johnson, R.S., et al., 2018. The interaction dynamics of two Potato leafroll virus movement proteins affects their localization to the outer membranes of mitochondria and plastids. Viruses 10. doi:10.3390/v10110585. Dombrovsky, A., Glanz, E., Lachman, O., et al., 2013. The complete genomics of Pepper yellow leaf curl virus (PYLCV) and its implications for our understanding of evolution dynamics in the genus Polerovirus. PLoS One 8, e70722. doi:10.1371/journal.pone.0070722. Gray, S., Cilia, M., Ghanim, M., 2014. Circulative, non-propagative virus transmission: An orchestra of virus-, insect-, and plant-derived instruments. In: Maramorosch, K., Murphy, F.A. (Eds.), Advances in Virus Research 89. Elsevier, pp. 141–199. doi:10.1016/B978-0-12-800172-1.00004-5. Kaplan, I.B., Lee, L., Ripoll, D.R., et al., 2007. Point mutations in the Potato leafroll virus major capsid protein alter virion stability and aphid transmission. Journal of General Virology 88 (6), 1821–1830. Knierim, D., Deng, T.C., Tsai, W.S., Green, S.K., Kenyon, L., 2010. Molecular identification of three distinct polerovirus species and a recombinant Cucurbit aphid-borne yellows virus strain infecting cucurbit crops in Taiwan. Plant Pathology 59, 991–1002. Krueger, E.N., Beckett, R.J., Gray, S.M., Miller, W.A., 2013. The complete nucleotide sequence of the genome of Barley yellow dwarf virus-RMV reveals it to be a new polerovirus distantly related to other yellow dwarf viruses. Frontiers in Microbiology 4, 205. doi:10.3389/fmicb.2013.00205. Lee, L., Palukaitis, P., Gray, S.M., 2002. Host-dependent requirement for the Potato leafroll virus 17kda protein in virus movement. Molecular Plant-Microbe Interactions 15, 1086–1094. doi:10.1094/MPMI.2002.15.10.1086. Massawe, D.P., Stewart, L.R., Kamatenesi, J., Asiimwe, T., Redinbaugh, M.G., 2018. Complete sequence and diversity of a maize-associated polerovirus in East Africa. Virus Genes 54, 432–437. doi:10.1007/s11262-018-1560-5. Pagan, I., Holmes, E.C., 2010. Long-term evolution of the Luteoviridae: Time scale and mode of virus speciation. Journal of Virology 84 (12), 6177–6187. doi:10.1128/ JVI.02160-09. Peter, K.A., Liang, D., Palukaitis, P., Gray, S.M., 2008. Small deletions in the Potato leafroll virus readthrough protein affect particle morphology, aphid transmission, virus movement and accumulation. Journal of General Virology 89, 2037–2045. doi:10.1099/vir.0.83625-0. Rashid, M.-O., Zhang, X.-Y., Wang, Y., et al., 2019. The three essential motifs in P0 for suppression of RNA silencing activity of Potato leafroll virus are required for virus systemic infection. Viruses 11 (2), 170. doi:10.3390/v11020170.

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Wintermantel, W.M., 2005. Co-infection of Beet mosaic virus with Beet yellowing viruses leads to increased symptom expression on sugar beet. Plant Disease 89 (3), 325–331. doi:10.1094/pd-89-0325. Xu, Y., Da Silva, W.L., Qian, Y., Gray, S.M., 2018. An aromatic amino acid and associated helix in the C-terminus of the Potato leafroll virus minor capsid protein regulate systemic infection and symptom expression. PLoS Pathog 14, e1007451. doi:10.1371/journal.ppat.1007451. Zhou, C.-J., Zhang, X.-Y., Liu, S.-Y., et al., 2017. Synergistic infection of BrYV and PEMV-2 increases the accumulations of both BrYV and BrYV-derived siRNAs in Nicotiana benthamiana. Scientific Reports 7, 45132. doi:10.1038/srep45132.

Pomoviruses (Virgaviridae) Eugene I Savenkov, Swedish University of Agricultural Sciences, Uppsala, Sweden and Linnean Center for Plant Biology, Uppsala, Sweden r 2021 Elsevier Ltd. All rights reserved. This is an update of L. Torrance, Pomovirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00543-4.

Nomenclature aa Amino acid(s) AGO Argonaute CP Coat protein or capsid protein ELISA Enzyme-linked immunosorbent assay ER Endoplasmic reticulum GFP Green fluorescent protein ID Internal domain kb Kilobase kDa Kilodalton MP Movement protein mRFP Monomeric red fluorescent protein NGS Next generation sequencing nt Nucleotide(s) NTD N-terminal domain

Glossary Spore balls Resting spores (cystosori) of Spongospora subterranea found in lesions or pustules (powdery scabs) on potato tubers; a single cystosorus comprises 500–1000

ORF Open Reading Frame RdRp RNA-dependent RNA polymerase RISC RNA-induced silencing complex RTD Read-through domain S Sedimentation coefficient; Svedberg units SEL Size exclusion limit siRNA Small interfering RNA ssRNA Single-stranded ribonucleic acid TGB Triple gene block tRNA Transfer RNA UTR Un-translated region vRNA Viral RNA vRNP Viral ribonucleoprotein VSR Viral suppressor of RNA silencing

resting spores aggregated to form in a ball that is partially hollow, traversed by irregular channels. Spraing Virus disease symptoms of internal brown lines and arcs in potato tuber flesh.

Introduction Pomovirus is a genus of plant viruses assigned to the family Virgaviridae. Pomoviruses have tubular rod-shaped particles and tripartite genomes; they are transmitted by soil-borne zoosporic organisms belonging to two genera (Polymyxa and Spongospora) in the family Plasmodiophoraceae. Pomoviruses have limited host ranges, infecting species in a few families of dicotyledonous plants. Agriculturally important hosts include potato and sugar beet. There are six member species in the genus Pomovirus: Potato mop-top virus (PMTV), Beet soil-borne virus (BSBV), Beet virus Q (BVQ), Broad bean necrosis virus (BBNV), Colombian potato soil-borne virus (CPSbV) and a tentative member Soil-borne virus 2 (SbV2). BVQ and BSBV can occur in mixed infections with the benyvirus Beet necrotic yellow vein virus (BNYVV).

Taxonomy and Classification The classification of tubular rod-shaped viruses transmitted by soil-borne plasmodiophorid vectors was revised in 1998 to establish four genera: Furovirus, Pomovirus, Benyvirus, and Pecluvirus. The revision was prompted by new virus sequence information that revealed major differences in genome properties (number of RNA segments, sequence, and genome organization). The genus Pomovirus is assigned to the family Virgaviridae. Pomovirus is a siglum from Potato mop-top virus, the type species.

Physical Properties of Particles Pomovirus particles are hollow, helical rods, 18–20 nm in diameter, comprising multiple copies of a single major coat protein (CP; c. 19–20 kDa). The CP gene is terminated by a UAG (or UAA in BBNV) stop codon that is thought to be suppressible, readthrough of which would produce a fusion protein with a read-through domain (RTD) of variable mass (54–104 kDa), CP-RTD. One or a few copies of the CP-RTD fusion protein are present at the extremity of PMTV particles and thought to contain the 50 end of the virus RNA. Pomovirus particles are fragile and particle size distribution measurements are variable; PMTV

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Fig. 1 Diagram of the PMTV genome organization; boxes indicate ORFs with the molecular masses of predicted protein products (kDa) indicated within.

particles have predominant lengths of 125, 137, and 283 nm. PMTV particles sediment as three components with sedimentation coefficients (S20,W) of 126, 171, and 236S.

Genome Properties and Functions of the Encoded Proteins Complete genome sequences are available for four member species. An almost complete sequence (without the 50 and 30 untranslated regions (UTRs)) is available for BBNV. Complete sequences of two genomic components, namely, RNA-rep and RNATGB are available for SbV2. Pomovirus genomes comprise three species of positive-sense single-stranded RNA (ssRNA) of c. 5.8–6.2, 2.8–3.4, and 2.3–3.1 kb (Fig. 1). RNA-rep encodes the replicase proteins. It contains a large open reading frame (ORF) that is interrupted by a UGA stop codon (ORF1) (or UAA in BVQ and BSBV; UAG in SbV2); the sequence continues in-phase to encode a readthrough protein (204–207 kDa). The protein encoded by ORF1 (145–149 kDa) contains methyl-transferase and helicase motifs while the RTD contains the GDD RNA-dependent RNA polymerase (RdRp) motif. Phylogenetic analysis reveals that the RdRps of the pomoviruses and the furovirus Soil-borne wheat mosaic virus share between 50% and 60% sequence identity. Phylogenetic analysis of available PMTV RNA-rep sequences reveals two major clades. The isolates from Peru, Colombia, Europe, China, Canada and USA group together in clade I, whereas clade II is exclusively represented by Colombian isolates. The 50 UTR of pomovirus RNAs contains the starting sequence GU(A)1–4(U)n (except BVQ RNA-rep which begins with AUA, CPSbV RNA-rep and RNA-CP which begin with ACAT(G)4–5TATT, and CPSbV RNA-TGB as well as RNA-rep and RNA-TGB of SbV2 have a different sequence at the 50 end). The RNAs are probably capped at the 50 end since the RNA-rep-encoded viral replicase contains methyl-transferase motifs associated with capping activity. The terminal 80 nt of the 30 UTR can be folded into a tRNAlike structure that contains an anticodon for valine. Both BSBV and PMTV RNAs were shown to be valylated experimentally. Different from other five pomoviruses, a tRNA-like structure of CPSbV genomic components contains an anticodone for leucine. The virus movement proteins (MPs) are encoded on RNA-TGB of pomoviruses. Three 50 -proximal overlapping ORFs encode a conserved module of movement proteins known as the triple gene block (TGB). TGB movement proteins are found in the genomes of other rod-shaped viruses in the families Virgaviridae (hordei-, gora- and pecluviruses) and Benyviridae (benyvirus) and in some genera of monopartite filamentous viruses in the families Alphaflexiviridae and Betaflexiviridae. The TGB proteins (TGB1, TGB2, and TGB3) are named according to the position of their genes on the RNA and have molecular masses of 48–54, 13, and 20–22 kDa, respectively. The TGB1 contains three structurally distinct domains: an unstructured intrinsically disordered N-terminal domain (NTD), an ordered internal domain (ID) and a helicase domain (belonging to the viral superfamily 1 RNA helicases). TGB1 NTDs of different pomoviruses do not share obvious sequence similarity. TGB1 binds RNA and is thought to interact with genomic RNAs to facilitate movement. The NTD contains two nucleolar localization signals, interacts with nuclear transport receptors of Importin-a family and accumulates in the nucleoplasm and in the nucleolus. The nucleolar passage of PMTV TGB1 is needed for the virus systemic movement. The half of ID of PMTV TGB1 is predicted to form an a-helix that seems to mediate oligomerisation of TGB1 through self-interaction. The ability for self-interaction (TGB1 oligomerisation) is crucial for the cell-to-cell movement of PMTV. PMTV TGB1 also interacts (through its helicase domain) with a read-through domain (RTD) of CP-RTD at one extremity of the virions assisting in the systemic movement of the virus particles. The sequence of the second TGB protein is the most conserved with BSBV, BVQ, and PMTV sharing 63%–75% nt sequence identity and 49% identity with that of BBNV; there is little sequence identity among the TGB3 sequences. Analysis shows that TGB2 and TGB3 proteins contain two hydrophobic regions (predicted transmembrane domains) separated by a hydrophilic domain and these proteins are associated with intracellular membranes in infected plants. PMTV TGB2 associates with the membranes of endoplasmic reticulum (ER), mobile granules in the cytoplasm, components of endosomal network, and the outer

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membrane of chloroplasts. PMTV TGB2 also binds RNA in a sequence non-specific manner, but is not required for RNA replication, although it was suggested that TGB2 might play a role in delivering the viral RNA to the chloroplast membranes for replication. PMTV TGB2 can increase permeability of plasmodesmata (so-called the size exclusion limit, SEL) and together with TGB3 (which also increases SEL of plasmodesmata) is thought to facilitate movement of the viral ribonucleoprotein (vRNP) complexes through plasmodesmata to the neighboring cells (cell-to-cell movement). At the subcellular level, TGB3 associates with the ER, motile granules, stationary aggregates at the periphery of the cell, termed “peripheral bodies”, and plasmodesmata. Similar to TGB2, the TGB3 protein of pomoviruses contains two trans-membrane domains and embedded into the membranes in U-shaped orientation with its central hydrophilic domain protruding into the ER lumen and N-terminus located in the cytosol. TGB3 contains a conserved tyrosine-based motif YQDLN involved in targeting TGB3 to plasmodesmata. In PMTV and BBNV, a fourth small ORF of RNA-TGB is predicted to encode an 8 kDa cysteine-rich (8K) or 6 kDa glycine-rich protein, respectively, whereas no such ORF is present in BSBV, BVQ, CPSbV or SbV2. The 8K protein is classified as a zinc-finger protein with a putative SWIM zinc-finger motif characterized by a consensus sequence CxCxnCxC, in which C stands for cysteine residues and x refers to any aa residue. The 8K protein of PMTV is not needed for the virus movement or infection of Nicotiana benthamiana. However, 8K is essential for the efficient virus accumulation and the appearance of the viral symptoms is delayed in the absence of the 8K expression. The 8K protein functions as a viral suppressor of RNA silencing (VSR). Mutations resulting in substitution of cysteine residues with alanines within a putative SWIM zinc-finger motif abolish 8K VSR activity. Comparison of the 8K sequences from more than 80 isolates reveals that the 8K protein sequences show an extraordinary variability with 23 variable sites per 68-aa-residues-long protein. Moreover, the 8K ORF is under positive selection, whereas other PMTV ORFs are under neutral or negative selection. The extraordinary variability and positive selection seem to drive efficient RNA silencing suppression activity of the 8K protein as the 8K protein encoded by certain PMTV isolates has stronger RNA silencing suppression activity compared those encoded by other isolates. Site-directed mutagenesis identified a serine residue (Ser-50) as being essential for enhanced VSR activity of one natural variant of 8K. Next generation sequencing (NGS) of small RNA populations in the presence of the 8K proteins with contrasting abilities to suppress RNA silencing (strong versus weak) reveals a lower abundance of 21 nt and 22 nt small interfering RNAs (siRNAs) with uracil (U) residue at the 50 end after expression of relatively strong 8K natural variant compared to that of a weak 8K allele. As accurate sorting of siRNAs into ARGONAUTE (AGO)-containing antiviral RNAinduced silencing complex (RISC) – with AGOs preferentially binding to siRNA with certain residue at the 50 terminus – is important for antiviral defense, it has been speculated that the natural variant of 8K with relatively strong RNA silencing suppression activity may destabilize either siRNAs with U residue at the 50 end or complexes into which such siRNAs are sorted, i.e., AGO1-conatining RISC that play a primary role (along with AGO2) in antiviral RNA silencing. The sequence of the 8K ORF along with the beginning of the sequence of TGB1 ORF is involved in biogenesis of a defective interfering (DI) RNA. DI RNA is readily detectable, accumulates at high levels in the virus-infected plants, and interferes with efficient virus accumulation. The disruption of a predicted minus-strand stem-loop secondary structure comprising complementary sequences of the 50 TGB1 ORF and the 30 8K ORF results in inhibition of DI RNA production suggesting intramolecular template switching during positive-strand synthesis as a possible mechanism for the DI RNA biogenesis. Phylogenetic analysis of available PMTV RNA-TGB sequences reveals two major clades. The isolates from Peru, Colombia, Europe, China, Canada and USA group together in clade I, whereas clade II is represented by a Peruvian isolate. The CP and CP-RTD proteins are encoded on RNA-CP. The PMTV RNA-CP is of variable size, from 2315 to 3252 nt. The longest RNA-CP (3252 nt) is found in PMTV isolate J20 from Cajamarca, Peru. Compared to other PMTV isolates, isolate J20 has additional sequences in 30 -UTR. In most PMTV isolates, RNA-CP is 3134 nt in length. Analysis shows that RTD of PMTV and BVQ contains two transmembrane domains, TM1 and TM2, suggesting that CP-RTD is a membrane protein integrated into the lipid bilayer in a U-shaped orientation. Indeed, PMTV CP-RTD localizes to cell periphery probably through association with plasma membrane. Analysis of naturally occurring and glasshouse-propagated isolates revealed that deletions of c. 500–1000 nt occur in both field and laboratory isolates and that the deletions occur predominantly in RTD, particularly in the region toward the 30 end of the RTD ORF. Deletions in this region are correlated with loss of transmission by the natural vector Spongospora subterranea. Hydrophobicity analysis shows that these in-frame deletions result in the loss of the transmembrane domain TM2. The occurrence of encapsidated deleted forms of RNA-CP and RNA-TGB in natural and laboratory isolates of PMTV has been described and sequence analysis as well as mutagenesis (for RNA-TGB) suggests that these PMTV RNAs contain sites that are susceptible to recombination possibly through a template switching mechanism. Variable base composition is found in natural isolates of BSBV in the sequence at the 30 end of RNA-TGB between the stop codon of the third TGB and the terminal tRNA-like structure. Phylogenetic analysis of available PMTV RNA-CP sequences reveals three major clades. The isolates from Peru, Colombia, Europe, China, Canada and USA group together in clade I, whereas clade II and clade III are exclusively represented by Peruvian isolates.

Virus–Host Interactions and Movement Cytoplasmic inclusions of enlarged ER and the accumulation of distorted membranes and small virion bundles can be seen by electron microscope examination of thin sections of BSBV- and BVQ-infected leaves. In PMTV-infected potato leaves, abnormal chloroplasts with cytoplasmic invaginations were seen in thin sections as well as tubular structures in the cytoplasm associated with the ER and tonoplast, and in the vacuole.

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PMTV does not require CP for movement and it is thought that TGB1 interacts with viral RNA forming a movement competent ribonucleoprotein (RNP) complex. Studies of transiently expressed PMTV TGB proteins fused to marker proteins such as green fluorescent protein (GFP) or monomeric red fluorescent protein (mRFP) in epidermal cells of N. benthamiana have helped to elucidate events in intracellular trafficking and indicate that PMTV interacts with the cellular membrane recycling system. PMTV TGB2 and TGB3 were shown to co-localize on the ER (Fig. 2(a)–(c)) and in small motile granules that utilize the actin–ER network to reach the cell periphery and plasmodesmata and TGB3 contains a putative tyrosine sorting signal (YQDLN), mutation of which inhibits TGB3 localization to plasmodesmata. TGB2 and TGB3 act together to transport GFP-TGB1 to the plasmodesmata for movement into neighboring cells and TGB2 and TGB3 have the capacity to gate the plasmodesmata pore. TGB2 co-localizes in endocytic vesicles with the Rab 5 ortholog Ara 7 (AtRabF2b) that marks the early endosomal compartment. Also, proteininteraction analysis revealed that recombinant TGB2 interacted with a tobacco protein belonging to the highly conserved RME-8 (receptor mediated endocytosis-8) family of J-domain chaperones, essential for endocytic trafficking. The systemic movement of many plant viruses occurs as virions that require expression and accumulation of CP in the infected cells. Yet, a small group of viruses including PMTV can move systemically in plants in the absence of CP as an RNP complex formed by interaction of the TGB1 protein with viral RNA. Moreover, mutational analysis revealed an essential role of CP-RTD in the long-distance movement of RNA-CP because in the absence of the CP-RTD expression PMTV CP inhibits systemic movement of RNA-CP, whereas the movement of RNA-TGB and RNA-rep is not affected. CP-RTD is needed for PMTV transmission in soil by its soil-borne vector Spongospora subterranea and minor amounts of CP-RTD molecules are incorporated into one terminus of the virus particles forming polar virions. Mutational analysis identified a CP-RTD domain consisting of 140 aa residues near the C-terminus as required for RNA-CP long-distance transport. Moreover, this particular domain is involved in the interaction of CP-RTD with TGB1 suggesting long-distance movement of polar virions that contain CP-RTD and TGB1 at the tip. Thus, the structure of movement-competent PMTV virions appears to be quite complex and involves at least two supplementary proteins at the particle tip. Since PMTV TGB1 plays indispensable role in the movement of both viral RNPs (vRNPs) and polar virions identification of the host proteins as well as sub-cellular domains that interact with TGB1 has been crucial to understanding the mechanism of PMTV movement in plants. At early stages of virus infection (at the very front of infection foci) PMTV TGB1 is localized to plasmodesmata consistent with the central role of the protein in the virus movement. In addition to localization in plasmodesmata TGB1 is observed in the nucleoplasm, nucleolus and in association with microtubules (Fig. 2(d)) at later stages of virus infection (two-four cell behind the infection front). The nuclear/nucleolar localization of PMTV TGB1 is mediated by interaction of the protein N-terminus with a nuclear transport receptor Importin-a. Disruption of TGB1 nucleolar localization (through mutagenesis) or knock down of Importin-a expression or disruption of microtubules by chemical inhibitors does not affect the virus cell-tocell movement but inhibits its systemic spread. As the N terminus of TGB1 appears to be important for the virus systemic movement and such movement correlates with the association of TGB1 with the nucleolus and microtubules, Nicotiana benthamiana cDNA library was panned using a yeast twohybrid screen to identify host proteins that interact with the TGB1 protein. The screen yielded HIPP26, a metallochaperone, as TGB1 interacting partner. The HIPP26 gene belongs to the large gene family that is unique in vascular plants. The HIPP proteins act in the heavy metal homeostasis (hence the name: Heavy metal-associated Isoprenylated Plant Protein, HIPP), involved in the regulation of transcriptional response to the biotic and abiotic stress and acts in a signal transduction as a plasma membrane-tonucleus relay during abiotic stress. HIPP26 is post-translationally modified through lipidation (S-acylation and prenylation). Knockdown of HIPP26 expression inhibits virus long-distance movement of both virions (when all three RNA genomic components are used as inoculum) and vRNPs (produced by inoculation of RNA-rep and RNA-TGB only) but does not affect cell-tocell movement of the virus. HIPP26 is upregulated in plants subjected to drought as well as in PMTV-infected plants. Moreover, PMTV infection protects plants from drought stress. Analysis of the spatial expression pattern of HIPP26 using HIPP26 promoterreporter fusions reveals its vascular tissue-specific expression consistent with the role of HIPP26 in systemic movement of the virus. Biochemical and mutational analyses show that HIPP26 subcellular localization at the plasmodesmata and plasma membrane is mediated by post-translational modifications through lipidation (S-acylation and prenylation), as nonlipidated HIPP26 is mostly found in the nucleus. Furthermore, HIPP26 is predominantly accumulates in the nucleus when co-expressed with TGB1. TGB1 interacts with the C-terminal 152CVVM155 (prenyl) domain of HIPP26, and the TGB1-HIPP26 complex decorates microtubules and accumulates in the nucleolus. Overall, these data support a mechanism in which interaction TGB1 with HIPP26 results in the inhibition of the HIPP26 protein lipidation or/and de-lipidation of already modified HIPP26, thus releasing membrane-associated HIPP26 from plasma membrane into the cytosol. The HIPP26/TGB1 complex, than, localizes to microtubules and subsequently directed to the nucleus and nucleolus in Importin-a-dependent manner. In the nucleus HIPP26 activates the drought stress response to facilitate virus long-distance movement.

Host Range, Geographical Distribution, Transmission by Vector and Variability Pomoviruses have a limited host range and are transmitted in soil by zoosporic plasmodiophorid vectors that have been classified as protists (Fig. 3). Viruses that are transmitted by plasmodiophorid vectors include pomo-, peclu-, furo-, beny- and bymoviruses and they are thought to be carried within the zoospores. The vector life cycle includes production of environmentally resistant thick-walled resting spores. The viruliferous resting spores can survive in soil for decades.

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Fig. 2 Confocal laser scanning microscope images of PMTV TGB2 and TGB3 (a–c) and TGB1 (d) fluorescent fusion proteins in epidermal cells of Nicotiana benthamiana. Monomeric red fluorescent protein (mRFP) tagged PMTV-TGB2 was transiently expressed with green fluorescent protein (GFP) tagged PMTV-TGB3, the proteins co-localized on membranes of the ER and in motile granules seen moving on the ER network. (a) Expression of mRFPTGB2 (red channel); (b) expression of GFP-TGB3 (green channel); (c) merged image of (a) and (b). (d) Merged image of the expression of yellow fluorescent (YFP) tagged TGB1 (red channel) from modified PMTV in N. benthamiana plants expressing ER-tagged GFP (green channel). At later stages of virus infection TGB1 does not associate with ER (green channel) anymore and mostly localizes on microtubules (red channel). Scale ¼ 10 nm. (a–c) Courtesy Lesley Torrance, The James Hutton Institute, UK.

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Fig. 3 (a) Biflagellate zoopore of Spongospora subterranea. (b) Bright field and (c) fluorescence microscope images of zoosporangia in tomato root hair. Courtesy Lesley Torrance, The James Hutton Institute, UK.

BBNV has been reported only from Japan; it causes necrosis and stunting in broad beans and peas. It is mechanically transmitted by inoculation of sap to a few species including Vicia faba, Pisum sativum, and Chenopodium quinoa. BVQ and BSBV are often found associated with BNYVV; BVQ is reported only from Europe whereas BSBV is found in sugar-beetgrowing areas worldwide. No symptoms have been attributed to BSBV or BVQ alone in sugar beet. BSBV is ubiquitous virus and is often

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found in the soils in sugar-beet growing areas, but the virus alone does not cause any disease in sugar beet. The viruses are thought to be transmitted in soil by Polymyxa betae. BVQ is mechanically transmissible only to C. quinoa and BSBV to members of the Chenopodiaceae. PMTV is reported to have originated in the center of potato domestication – the Andean region of South America – and has now found in potato-growing regions of Europe, North and South America, Asia and recently in New Zealand; virus incidence is favored by cool, wet growing conditions. It is transmitted in soil by S. subterranea, also a potato pathogen which causes the tuber blemish disease powdery scab. PMTV can be transmitted mechanically to members of the Solanaceae and Chenopodiaceae. CPSbV and SbV2 has been reported only from Colombia; the viruses seem to cause symptomless infections in potato. The viruses are thought to be transmitted in soil by S. subterranea. CPSbV and SbV2 are mechanically transmitted by inoculation of sap to members of the Solanaceae and Chenopodiaceae. The complete and partial genomic sequences are available for numerous PMTV isolates from around the globe. Based on the CP-RTD sequence, two major strains of PMTV are recognized and coined as S (for “severe”) and M (for “mild”). Seven signature aa residues differ between the S (RRSIGAV) and the M (KKNVREM) strains in the CP-RTD protein encoded by RNA-CP. The M strain is only reported from Peru, whereas the S strain is found worldwide including Peru. Phylogenetic analysis identified two major genotypes (clades) for RNA-rep (I and II), two major genotypes for RNA-TGB (I and II), and three genotypes for RNA-CP (I, II and III). As PMTV has a tripartite genome, a classification of PMTV isolates based on genotype constellations has been suggested. In this classification, the genotype of each genomic RNA is taken into consideration. Four genetic constellation has been found so far. Two genetic constellations – I þ I þ II and UN þ II þ III (RNA-rep þ RNA-TGB þ RNA-CP, “UN”, indicates unknown genotype of the RNA-rep segment, for which sequence is not available) – are exclusively found in Peru. Another genetic constellation (II þ I þ I) is found in Colombia, whereas the constellation I þ I þ I is found worldwide including Peru and Colombia. These observations suggest that this particular constellation (I þ I þ I) originates from the Andean region of Peru, the center of potato domestication, and was spread to the other parts of the world. In general, greatest diversity of PMTV is found in the Andean region of South America, whereas the virus has very little variability in the rest of the world.

Serological Relationships, Diagnosis, and Control The viruses are serologically distinct. Distant relationships have been reported between BSBV and BVQ; PMTV and Soil-borne wheat mosaic virus (Furovirus); and PMTV, BBNV, and tobamoviruses. PMTV, BSBV, and BVQ CPs contain a conserved sequence (SALNVAHQL) that reacts in Western blots with a monoclonal antibody (SCR70) produced against PMTV, but that is not exposed on intact particles. PMTV particles contain an immunodominant epitope at the N-terminus of the CP that is exposed at the surface along the sides of the particles and can be detected by monoclonal antibody SCR69 (Fig. 4). The viruses can be detected by serological tests (immunosorbent electron microscopy, enzyme-linked immunosorbent assay), by assays based on the reverse transcriptase-polymerase chain reaction (RT-PCR) and by quantitative real-time RT-PCR (RT-qPCR) in leaves, roots, or tubers from naturally infected plants. PMTV is known to be erratically distributed in potato leaves and tubers and can move systemically in the absence of CP in potato leaves which raises a risk of false negative diagnosis. The problem can be overcome by using RT-PCR or RT-qPCR for detection of RNA-TGB and/or RNA-rep as RNA-CP might be absent in those samples. PMTV causes an economically important disease affecting the quality of tubers grown for the fresh and processing markets. Tubers of susceptible potato cultivars that are infected from soil during the growing season develop “spraing” symptoms that include unsightly brown lines, arcs, or marks in the flesh sometimes accompanied by slightly raised external lines and rings (Fig. 5 (a)). Potato plants grown from infected tubers display V-shaped yellow markings or chevrons on the leaves (Fig. 5(b)) and may have shortened internodes (mop-top) producing cracked or malformed tubers; both tuber quality and yield can be affected. However, the virus does not infect all plants grown from infected tubers. Haulm and tuber symptoms vary markedly with cultivar, and some cultivars are symptomlessly infected. Environmental conditions also affect disease incidence and severity, and PMTV

Fig. 4 Electron micrograph of PMTV particles labeled with monoclonal antibody SCR69/gold conjugate. Courtesy Lesley Torrance, The James Hutton Institute, UK.

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Fig. 5 (a) Symptoms of PMTV in potato tubers cv. Saturna. (b) Chevron-like symptoms of PMTV on leaves of potato cv. Qompis.

incidence was shown to increase with annual rainfall. Soil temperatures of 12–171C and high soil moisture at tuber initiation favor powdery scab incidence. PMTV can be established at new sites by planting infected tubers and once established, PMTV is a persistent problem, as the resting spore balls of the vector S. subterranea are long-lived and resistant to drought and agrochemicals. Viruliferous spore balls are spread readily to new sites by farm vehicles, contaminated seed tubers, wind-blown surface soil; motile zoospores can be spread through contaminated irrigation or drainage water. In certain areas, potatoes have become infected by PMTV 18 years after potatoes were last grown (the longest period recorded). There is no effective practicable means to control S. subterranea but decreased severity of powdery scab can be achieved by application of fluazinam to soil. In addition, disinfection of tubers with chemicals such as formaldehyde decreases virus incidence, although the efficacy of this treatment depends on the level of soil infestation where the tubers are planted and a combination of tuber and soil treatments may be more effective. The best prospect for virus disease control is development of resistant cultivars but there are no known sources of PMTV resistance in commercial potato cultivars and most of the commercially grown cultivars are also susceptible to S. subterranea. However, plants transformed with virus-derived transgenes (CP and a mutated form of TGB2) have exhibited resistance to PMTV with decreased virus accumulation and incidence in tubers of plants grown in infested soil.

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Further Reading Cowan, G.H., Roberts, A.G., Jones, S., et al., 2018. Potato mop-top virus co-opts the stress sensor HIPP26 for long-distance movement. Plant Physiology 176, 2052–2070. Gil, J.F., Adams, I., Boonham, N., Nielsen, S.L., Nicolaisen, M., 2016. Molecular and biological characterization of two novel pomo-like viruses associated with potato (Solanum tuberosum) fields in Colombia. Archives of Virology 161, 1601–1610. Haupt, S., Cowan, G.H., Ziegler, A., et al., 2005. Two plant-viral movement proteins traffic in the endocytic recycling pathway. Plant Cell 17, 164–181. Kalyandurg, P., Gil, J.F., Lukhovitskaya, N.I., et al., 2017. Molecular and pathobiological characterization of 61 Potato mop-top virus full-length cDNAs reveals great variability of the virus in the center of potato domestication, novel genotypes and evidence for recombination. Molecular Plant Pathology 18, 864–877. Kalyandurg, P.B., Tahmasebi, A., Vetukuri, R.R., et al., 2019. Efficient RNA silencing suppression activity of Potato Mop-Top Virus 8K protein is driven by variability and positive selection. Virology 535, 111–121. Lukhovitskaya, N.I., Cowan, G.H., Vetukuri, R.R., et al., 2015. Importin-a mediated nucleolar localization of Potato mop-top virus TRIPLE GENE BLOCK1 (TGB1) protein facilitates virus systemic movement, whereas TGB1 self-interaction is required for cell-to-cell movement in Nicotiana benthamiana. Plant Physiology 167, 738–752. Lukhovitskaya, N.I., Thaduri, S., Garushyants, S.K., Torrance, L., Savenkov, E.I., 2013. Deciphering the mechanism of defective interfering RNA (DI RNA) biogenesis reveals that a viral protein and the DI RNA act antagonistically in virus infection. Journal of Virology 87, 6091–6103. Pereira, L.G., Torrance, L., Roberts, I.M., Harrison, B.D., 1994. Antigenic structure of the coat protein of Potato mop-top virus. Virology 203, 277–285. Reavy, B., Arif, M., Cowan, G.H., Torrance, L., 1998. Association of sequences in the coat protein/read-through domain of Potato mop-top virus with transmission by Spongospora subterranea. Journal of General Virology 79, 2343–2347. Rochon, D., Kakani, K., Robbins, M., Reade, R., 2004. Molecular aspects of plant virus transmission by olpidium and plasmodiophorid vectors. Annual Review of Phytopathology 42, 211–241. Sandgren, M., Savenkov, E.I., Valkonen, J.P.T., 2001. The readthrough region of Potato mop-top virus (PMTV) coat protein encoding RNA, the second largest RNA of PMTV genome, undergoes structural changes in naturally infected and experimentally inoculated plants. Archives of Virology 146, 467–477. Solovyev, A.G., Savenkov, E.I., 2014. Factors involved in systemic transport of plant RNA viruses, the emerging role of the nucleus. Journal of Experimental Botany 65, 1689–1697. Torrance, L., Lukhovitskaya, N.I., Schepetilnikov, M.V., et al., 2009. Unusual long-distance movement strategies of Potato mop-top virus RNAs in Nicotiana benthamiana. Molecular Plant-Microbe Interactions 22, 381–390. Torrance, L., Wright, K.M., Crutzen, F., et al., 2011. Unusual features of pomoviral RNA movement. Frontiers in Microbiology 2, 1–7. (article 259). Zamyatnin, A.A., Solovyev, A.G., Savenkov, E.I., et al., 2004. Transient co-expression of individual genes encoded by the triple gene block of Potato mop-top virus reveals requirements for TGBp1 trafficking. Molecular Plant Microbe Interactions 17, 921–930.

Relevant Websites https://www.cabi.org/isc/datasheet/42827#toDistributionMaps Invasive Species Compendium. https://pestdisplace.org/ PestDisPlace.

Potato Virus Y (Potyviridae) Laurent Glais, French Federation of Seed Potato Growers/Research, Development, Promotion of Seed Potato, Paris, France and Institute for Genetics, Environment and Plant Protection, Agrocampus West, French National Institute for Agriculture, Food, and Environment, University of Rennes 1, Le Rheu, France Benoît Moury, Plant Pathology Unit, INRAE – French National Institute for Agriculture, Food and Environment, Montfavet, France r 2021 Elsevier Ltd. All rights reserved. This is an update of C. Kerlan, B. Moury, Potato Virus Y, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00737-8.

Nomenclature

MP Movement protein NLR Nucleotide-binding and leucine-rich nt Nucleotide(s) NTR Non-translated region ORF Open reading frame PIPO Pretty interesting Potyviridae ORF PTNRD Potato tuber necrosis ringspot disease RdRp RNA-dependent RNA polymerase RISC RNA-induced silencing complex satRNA Satellite RNA UTR Untranslated region VIGS Virus-induced gene silencing VLP Virus-like particles VPg Viral protein genome-linked VRC Virus replication complexes vRNA virion RNA

aa Amino acid(s) AGO Argonaute 1 CI Cylindrical inclusion CITE Cap-independent translation enhancer Co-Pro Protease cofactor CP Coat protein or capsid protein ELISA Enzyme-linked immunological assays ER Endoplasmic reticulum HC-Pro Helper component-protease HR Hypersensitive reaction IRES Internal ribosomal entry site kb Kilobase kDa Kilo dalton LAMP Loop mediated amplification MAb Monoclonal antibody

Glossary Hypersensitive Reaction A specific virus strain plant defense reaction mediated by N genes associated with an

infection limited to tissue surrounding the initially infected cells with visible local or systemic necrosis.

Introduction Potato virus Y (PVY) was first recognized in 1931 as an aphid-transmitted member within a group of viruses associated with potato degeneration, a disorder known since the eighteenth century. PVY is the type species of the genus Potyvirus, one of the six genera in the family Potyviridae. PVY is naturally spread by vegetatively propagated material and by aphids in non-persistent and noncirculative manners. Horizontal transmission by contact between plants and vertical transmission by true seeds have also been reported. PVY has a wide host range and is highly variable with some host specificity. Genome sequences and reliable tools for detection and strain differentiation are available. Bioassays and serology have been largely developed. PVY is one of the most damaging plant pathogens causing significant losses in four main crops around the world: potato, pepper, tomato, and tobacco. In surveys of viruses with worldwide economic importance, PVY was listed in the top-ten viruses affecting field-grown vegetables. PVY was also found responsible for damages in petunias in Europe, in eggplant crops in India and poha (or Cape gooseberry; Physalis peruviana) in Hawaii. Efficient control strategies depending on the crop have been developed. However, none of them seems capable to contain entirely the risks of PVY epidemics linked to virus evolution or changes of epidemiological conditions.

Viral Particle, Genome and Cytopathology PVY particles are non-enveloped, filamentous, flexuous rods, 730–740 nm long, 11–12 nm in diameter, with an axial canal 2–3 nm in diameter and helical symmetry (Fig. 1). Each virion is composed of circa 2000 units of viral coat protein (CP) of 30 kDa, encapsidating a single-stranded, linear, positive-sense RNA molecule (3.1–3.2
106 Da) of approximately 10 kb long. The PVY genome has two distal non-translated regions (NTR) at the 50 - (from 183 to 195 nt) and the 30 - (from 326 to 407 nt) ends, with a poly-A tail at the 30 -terminus and a viral protein covalently linked (VPg) at the 50 -end. The genomic RNA contains one single open reading frame (ORF), and is translated into a large polyprotein (3061–3062 aa) cleaved in cis or trans by three virus-

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Fig. 1 Purified suspension of PVY. Magnification 22,000
. Photograph: D. Thomas, CNRS-Rennes 1 University, France. The bar represents 250 nm.

Fig. 2 Schematic representation of genomic organization of PVY and polyprotein processing. The viral genome-linked protein (VPg) is illustrated by a circle at the 50 -end of the genome. The poly-A tail is presented at the 30 -end. The large open reading frame encoded by the viral genome is presented with the names of corresponding proteins (P1, HC-Pro, P3,P3N-PIPO, 6K1, CI, 6K2, VPg, NIa-Pro, NIb, CP). The short PIPO ORF overlapping the P3 protein is presented. The nucleotide and amino acids are numbered according to isolate N605 (accession number X97895). NTR: non-translated region.

encoded proteases (P1, HC-Pro, NIa-Pro) into ten functional proteins: P1 (275 aa), HC-Pro (465 aa), P3 (364–365 aa), 6K1 (52 aa), CI (634 aa), 6K2 (52 aa), VPg (188 aa), NIa-Pro (244 aa), NIb (519 aa), and CP (267 aa). An eleventh protein, P3N-PIPO, is produced by slippage of the viral polymerase at a specific motif embedded in the P3-coding region called PIPO (Pretty Interesting Potyviridae ORF) (Fig. 2). These 11 viral proteins are mainly multi-functional. Some of them interact with some other viral proteins and/or with some host proteins to determine plant infection or virus plant-to-plant transmission. In the process of PVY infection, viral proteins are often associated with specific subcellular compartments (chloroplasts, endoplasmic reticulum and Golgi apparatus) where they perform their functions (Fig. 3). After PVY particles are introduced into the cytoplasm of plant cells by mechanical or vectormediated inoculation, the PVY infection cycle is initiated by uncoating of virions and release of the viral RNA genome which is then recruited by the host translation machinery (eukaryotic initiation factors, ribosomes, etc.) to synthesize the polyprotein. After its processing, the 11 viral proteins are released. Nuclear inclusion proteins (NIa, NIb) migrate to the nucleus and cylindrical inclusion (CI) proteins accumulate in the cytoplasm by forming pinwheel-shaped inclusions. Viral replication complexes associate to the membrane of host organelles (endoplasmic reticulum, Golgi apparatus and vesicles) through the 6K2 protein. During replication, the positive-sense single-stranded RNA (ssRNA) is copied into a complementary negative-sense RNA and vice versa by

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Fig. 3 Schematic representation of PVY infection cycle in infected plant cell, with the location of viral proteins and their mutual interactions. Extracted from Lacomme, C., Jacquot, E., 2017. General characteristics of potato virus Y (PVY) and its impact on potato production: An overview. In: Lacomme, C., Glais, L., Bellstedt, D.U., et al. (Eds.), Potato Virus Y: Biodiversity,Pathogenicity, Epidemiology and Management. Switzerland: Springer, pp. 1–19.

the RNA-dependent RNA polymerase (RdRp) (NIb protein) and the RNA helicase (CI protein). The obtained positive-sense ssRNA are either encapsidated to produce new virions or recruited by viral proteins (CI, CP, HC-Pro, P3N-PIPO) and host proteins to form ribonucleic complexes that move from cell to cell through plasmodesmata and systemically in phloem vessels.

Host Range The natural host range of PVY comprises plants in nine families. It includes potato (Solanum tuberosum ssp. tuberosum), several species of native potatoes in the Andes including Solanum andigena and numerous Solanum species, pepper (Capsicum spp.), tobacco (Nicotiana spp.), tomato (Solanum lycopersicum), eggplant (Solanum melongena), Cape gooseberry (Physalis peruviana), ornementals (Petunia spp., Dahlia spp.), perennial plants (Physalis virginiana, Physalis heterophyla), and a number of annual plants. Many region-specific hosts were recorded such as Cotula australis in New Zealand or Sorbaria tomentosa in Himalaya. Weeds in the family Solanaceae are often potential inoculum sources for PVY infections in tomato and pepper crops: Solanum nigrum and Solanum dulcamara in many countries; Solanum chacoense and Physalis viscosa in Southern America; Solanum gracile, Solanum aculeatissimum, Physalis angulata, Physalis ciliosa, and Physalis floridana in Florida. Physalis virginiana and Physalis heterophyla (perennial ground cherries) are overwintering hosts in Northern America. Solanum dulcamara in Western Europe or Datura spp. in Mediterranean countries are potential inoculum sources in potato crops as well as volunteer potatoes. Infected pepper and tomato plants, and seed and volunteer potatoes were also identified as potential sources of inoculum in tobacco crops in the USA, Canada, and Italy. PVY is easily sap-transmitted and can be transmitted by stem- and tuber-grafting. Its experimental host range comprises plants in 495 species and in 72 genera of 31 families. It includes 287 species in the family Solanaceae (among which 141 Solanum species and 70 Nicotiana species), 28 species of Amaranthaceae, 25 species of Fabaceae, 20 species of Chenopodiaceae, and 11 species of Asteraceae. A large part of these plants show only local lesions upon PVY inoculation. Datura stramonium, formerly reported as a host plant, was demonstrated to be totally immune to all strains tested in 1980–1990. Some PVY isolates can infect the model plant Arabidopsis thaliana. Nicotiana tabacum, Nicotiana benthamiana, Nicotiana occidentalis can be used as diagnostic species susceptible to all PVY strains. N. tabacum plantlets are often used as source and test plants for aphid transmission experiments. N. tabacum is a suitable host for

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Fig. 4 Winged form of Myzus persicae, an insect vector of PVY. Photograph: B. Chaubet, INRAE, France.

virus purification. Yields of purification including a final step of cesium chloride gradient centrifugation usually vary from 10 to 25 mg kg1 of tobacco leaves. Leaves of N. tabacum are the best virus-infected material to store. Antigenic properties can be retained for 1 year in freeze-dried crude extracts. Infectivity was reported to be preserved in freeze-dried material stored over calcium chloride at 41C for 15 years. However from our own experience numerous long-term stored isolates of PVY available in international collections are no longer infectious. Solanum demissum Y, S. demissum PI 230579, and the hybrid S. demissum A6 are local lesion hosts. The ‘A6 test’ on detached leaves was commonly used in the past as a diagnostic tool for differentiating PVY from Potato virus A (PVA), another potyvirus. Cultivars of N. tabacum, potato and pepper, Chenopodium amaranticolor, P. floridana, some accessions of Solanum brachycarpum and Solanum sparsipilum are useful for distinguishing among PVY strains and pathotypes. Potato cultivars such as Bintje or Saco can be used to separate PVY from plants co-infected with PVA or Potato virus X (PVX). The capacity of PVY isolates from the C1 group to infect pepper (C. annuum) was shown to require mutations in both the CI and the P3 and/or P3N-PIPO coding regions, and seem to have evolved several times independently.

Transmission In natural conditions, different transmission routes are used by PVY. The first one is vertical transmission from infected mother plants to their progeny. Potato plants are propagated vegetatively by planting tubers from one generation to the next. When PVY is inoculated to a potato plant, after a multiplication phase in the first infected cells, the virus migrates from cell to cell, cross different cell layers of the inoculated organ and reach the phloem. Once in the phloem, PVY is rapidly translocated to the whole plant including to daughter tubers, where it multiplies. If these infected tubers (‘seed’ potatoes) are planted the year after, the germinated seedlings will also be infected and usually initiate a rapid PVY dissemination in fields through secondary aphid transmission. Transmission of PVY from mother plants to its ‘true’ seeds has been reported from Solanum nigrum, Nicandra physaloides and pepper (Capsicum annuum). The second transmission route corresponds to horizontal transmission from plant to plant by aphids (Fig. 4). PVY displays a large range of aphid vectors. Aphids in 65 species, all in the family Aphidinae, colonizing or not the source plants, were demonstrated to be able to transmit PVY. Myzus persicae is the most efficient PVY vector and most of the other aphid species have much lower transmission efficiencies comparatively. Both alate and wingless aphids are vectors of PVY. PVY is transmitted by aphids in a nonpersistent manner, meaning that acquisition and inoculation periods are brief (a few seconds or minutes) and that the virus is not retained on the long term by the aphids. Aphid stylets penetrate into the epidermal cell layer of the plants and puncture plant cell membranes, allowing insects to suck the cellular content. In the case of infected cell, this probe or feeding is accompanied by the sucking of virus particles that bind to specific receptors located in the extreme tip of the aphid’s stylet, called the acrostyle. Only particles attached to these receptors are involved in aphid transmission. PVY transmission by aphids is also noncirculative. There is no discernible latent period and the virus does not need to circulate into the aphid body to allow transmission. PVY does not pass through insect moults and retention of the virus in the aphid usually lasts not more than 1 or 2 h. However, longer retention periods (up to 17 h in Aphis nasturtii) were reported. Prior starvation of the aphids increases the efficiency of transmission though it does not affect the number of electrically-recorded membrane punctures during acquisition periods. PVY transmissibility is determined by both HC-Pro and CP proteins. The non-structural HC-Pro protein corresponds to the helper component of the aphid transmission, which interacts in its homodimer active form as a bridge between the virion and the aphid stylet. It has been recently proved with other potyviruses that this HC-Pro dimerization is a key step for an efficient transmission. Studies performed on LMV (Lettuce mosaic virus) and TuMV (Turnip mosaic virus) demonstrated that viruses activate their own transmission by dimerization of HC-Pro protein from the moment they perceive an unknown plant signal announcing the penetration of infected cells by aphid vector. Without the presence of this active form of HC-Pro protein, the transmission is impossible. This phenomenon is called the transmission activation (AT). Otherwise, two domains in HC-Pro protein are crucial for PVY transmission: the KITC motif (Lysine/Isoleucine/Threonine/Cysteine) located in its N-terminal region,

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which is involved in the binding to a specific receptor in aphid’s mouthparts, and the PTK motif (Proline/Threonine/Lysine) in its C-terminal region, which interacts with the DAG motif (Aspartic acid/Alanine/Glycine) at the CP N-terminus. With these two motifs, HC-pro provides the link between the aphid’s stylet and the virion. PVY transmission by pollen has never been proved for any host plant. In experimental conditions, other methods are also used to transmit PVY from plant to plant. The most commonly used is mechanical inoculation. A healthy plant becomes infected by manually rubbing leaves or cotyledons with a mixture containing PVY-infected plant sap and a fine abrasive powder. Transmission by grafting a scion from a virus-infected plant to a healthy stock plant (or vice versa) is also efficient. Transmission of PVY by woundings caused by squashing, squeezing or bouncing of healthy plants in the vicinity of infected plants can happen but is not frequent in field conditions. Transmission of PVY from an infected to a healthy tuber by seed cutting was not reported.

Serology PVY is considered to be strongly immunogenic. The PVY antigenic structures are carried by the CP. Its C-terminal region contains epitopes specific to PVY, whereas its highly variable N-terminal region carries epitopes allowing the differentiation of PVY in two main serogroups: serotype O (PVYO, PVYC, PVYN-Wi strain groups), and serotype N (PVYN, PVYNTN, PVYZ, PVYE). Antisera and monoclonal antibodies (MAbs) have been produced in rabbits or mice immunized with purified virus preparations or synthetic peptides, or by using DNA-based immunization, or phage display antibody technology. Numerous sources of antibodies and serological detection kits are available. Polyclonal antibodies do not discriminate among PVY strain groups, whereas monoclonal antibodies, triggering a single epitope, allow the serotyping of PVY isolates. These serological reagents have been mainly used in ELISA (Enzyme-Linked Immunosorbent Assay) and related protocols (dot-blot immunobinding assay, immunosorbent electron microscopy, latex and viro-bacterial agglutination tests, immunodiffusion in agar gels) for the detection and the identification of PVY from plant material.

Phylogeny and Evolution Outside recombinant isolates, three major PVY phylogenetic groups with worldwide distributions, namely C, N, and O, can be distinguished based on genome sequences (Fig. 5). Group C can be further divided into two subgroups, C1 and C2. These phylogenetic groups partly correlate with PVY host specificity. In potato, most isolates belonged to groups C2, N, and O, whereas in pepper (Capsicum spp.), most isolates belonged to group C1. All kinds of isolates could be found in tomato or tobacco. A large number of isolates are recombinants between these main phylogenetic groups. Thirty-six different recombination patterns have been identified, mainly between the N and O groups. Such recombinants are now becoming predominant in potato crops worldwide. N
O recombinants are also becoming widespread in pepper crops, whereas isolates from their parental groups are poorly infectious in this crop plant. Two additional PVY phylogenetic groups were recently identified and have more restricted distributions: a “Chilean” group on tobacco and pepper (Capsicum baccatum) and a Brazilian U group on tobacco (Fig. 5). Phylogenetic reconstruction of host specificity in PVY suggests that infection in pepper (Capsicum spp.) is a recently-acquired trait and that it was acquired at least twice independently. In contrast, it is uncertain if infection in potato is an ancestral or derived trait in PVY. PVY belongs to a cluster of 19 potyvirus species, the closest to PVY being Pepper severe mosaic virus. Most of these potyviruses infect plants in the family Solanaceae but others infect plants of the Amaranthaceae, Asteraceae, Liliaceae, Amaryllidaceae, or Verbenaceae.

Fig. 5 Phylogenetic relationships between PVY groups and inference of adaptation to pepper (pep) (A) and potato (pot) (B). (A) The most parsimonious scenario infers that the most recent common ancestor (MRCA) of PVY was not infectious in pepper and get adapted to pepper at least two times independently. (B) There are two equally parsimonious scenarios for PVY adaptation to potato, considering that the MRCA was either infectious (in gray) or not infectious (in black) in potato. Both scenarios involve at least three changes in PVY adaptation to potato. In green: extant PVY clades that are infectious in the respective crop plant. In red: extant PVY clades that are not infectious in the respective crop plant.

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Fig. 6 (A) Natural secondary infection by PVY in a potato field: crinkling, yellowing, and growth reduction of the leaflets. Symptoms associated with the potato tuber necrotic ringspot disease (PTNRD). (B) Yellowing and necrosis on a basal leaf. (C) Necrotic oak-leaf pattern on an intermediate leaf. (D) Typical symptoms on tubers of the cv. Monalisa. Photographs: Y. Le Hingrat, FN3PT, France (A), K. Charlet-Ramage, GNIS, France (B) and (C), L. Glais, FN3PT, France (D).

PVY in Potato For almost 50 years, PVY has become the most important virus in most growing areas for seed, ware, and processed potatoes. Serious outbreaks were reported in the 2000s. Tuber quality can be severely affected due to necrosis or defects for processing potatoes. Reduction in size and number of harvested tubers can result in losses up to 80% if no control of PVY spreading was used. Symptoms consist of mild to severe mottle, often associated with crinkling of the leaves (Fig. 6(A)). Yellowing and necrosis (vein necrosis and necrotic spots) frequently occur in the lower leaves (Fig. 6(B)). Symptoms also include collapse and dropping of intermediate leaves, which remain clinging to the stem (leaf drop). Secondarily-infected plants (when infected tubers are planted) are stunted with crinkled and smaller leaflets. Necrosis on and around leaf veins, on petioles, stems, and tubers may occur in numerous cultivars. Some necroses on stems and petioles are called stipple-streak disease. The potato tuber necrotic ringspot disease (PTNRD) is characterized by particular patterns on leaves (sometimes oak-leaf necrosis) (Fig. 6(C)) and stems, and superficial annular and arched necroses on tubers, first slightly protruding from tuber skin, then becoming dark brown with sometimes crackings of tuber skin (Fig. 6(D)). Potato PVY isolates are classified according to three criteria: their capacity to induce necrotic symptoms on N. tabacum, to overcome different resistance genes in Solanum tuberosum and to produce necrosis on potato tubers. Based on these biological features, PVY isolates are divided into seven strain groups: PVYO, PVYC, PVYN, PVYNTN, PVYN-Wi, PVYZ, PVYE. Beside these strain groups, several individual isolates sharing particular biological properties are till now unclassified. PVYO and PVYC induce mosaic symptoms on tobacco leaves. They are mainly separated on the basis of hypersensitive reactions (HR) in potato cultivars. PVYO induces a HR reaction in cultivars bearing the resistance genes Nytbr, Ny-1 and Ny-2 (Table 1), whereas PVYC elicits HR response in cultivars carrying the Nc gene. PVYN differs from PVYO and PVYC in causing a veinal necrosis in N. tabacum cv. Samsun or cv. Xanthi and eliciting a HR in cultivars carrying the Ny-1 and Ny-2 resistance genes (Table 1). Two aa at positions 400 and 419 in the HC-Pro protein are involved in the induction of the necrotic reaction in tobacco. Isolates of the PVYNTN group express the same biological feature as PVYN, except that they are able to induce tuber necrosis on susceptible cultivars. One of the viral molecular determinants inducing PTNRD was identified at aa position 419 in the HC-Pro protein, but some others are still unknown. Whole genome analyses of PVYNTN isolates revealed that most of them result from three to four recombination events between PVYO and PVYN isolates, with breakpoints being not exactly at the same nucleotide position. Some PVYNTN isolates however are not recombinants (Fig. 7).

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Table 1

Molecular determinants of pathogenicity and transmission identified in PVY

Phenotype

Plant

Pathogenicity Veinal necrosis Veinal necrosis

Genome region

Mutation

Nicotiana tabacum Hc-Pro Nicotiana tabacum with the Rk nematode resistance Nib gene Solanum tuberosum HC-Pro Physalis floridana CP

N339D, D400K, D419E –

Host range Systemic infection

Capsicum annuum

N131Y, N131F

Systemic infection

Capsicum annuum

P3 and/or P3NPIPO CI Hc-Pro Hc-Pro VPg VPg and CI NIb VPg

K269R and/or R270K (local necrosis induction) K269R and/or R270K (necrosis induction) Different mutations in codons 101–123 – K472E R119H

VPg NIa-Pro

105E, 105T, 105M, 119A H46Y, D81A, C151F or H167P

Hc-Pro CP

K59E E68K

Tuber necrosis Leaf necrosis

Interaction with plant resistance Resistance induction Solanum tuberosum with the Nc resistance gene Resistance Resistance Resistance Resistance Resistance

induction breakdown breakdown breakdown breakdown

Resistance breakdown Loss of resistance induction Aphid transmission Loss of transmissibility Increase of transmissibility

Solanum tuberosum with the Ny resistance gene Capsicum annuum with the pvr2 resistance gene Capsicum annuum with the pvr23 resistance allele Capsicum annuum with the Pvr4 resistance gene Solanum lycopersicum with the pot  1 resistance gene Nicotiana tabacum with the va resistance gene Solanum tuberosum with the Ry resistance gene

D419E –

–

Note: “–”: unavailable data.

Isolates of the PVYN-Wi group, similar to PVYN:O in North America, are differentiated from PVYN isolates by their O/C (instead of N) serotype, and by their capacity to induce PTNRD on specific cultivars. The fact that these isolates share both PVYN and PVYO properties is due to the presence of two to four recombination breakpoints in their genome leading to a succession of O- and N-type nucleotide sequences (Fig. 7). PVYZ and PVYE isolates are closely related to PVYNTN isolates because they cause PTNRD in tubers and display a similar recombinant genome, but they differ by their capacity to induce mosaic symptoms in tobacco leaves, as PVYO isolates. Moreover, PVYZ isolates elicit a HR reaction in potato cultivars carrying the Nz resistance gene, which is not the case of PVYE isolates. The first PVY groups identified in the early 1930s were PVYC and PVYO, and PVYN a few years later. In contrast to PVYC which became less frequent in potato, PVYO remained the dominant group in potato fields up to the 2000s. However, from the 1960s, infections caused by PVYN isolates increased progressively in Europe and became dominant 40 years later. This change was associated to the emergence of PVYNTN isolates, which were first described in Hungary in 1982 and soon spread in most potato growing countries, except in North America where they were classified as quarantine pathogens. So far, PVYNTN isolates have a worldwide distribution and are predominant notably in Europe. In Switzerland and Estonia, a significant decrease in the occurrence of PVYNTN isolates was recently observed concurrently with an increase of PVYN-Wi infections, probably due to a change in potato varieties. PVYN-Wi isolates were identified for the first time in Poland in 1984 and they are currently present throughout the world with varying frequencies. Some surveys performed these last two decade reported that PVYNTN and PVYN-Wi groups have almost replaced the PVYO and PVYN ancestral groups in many countries. PVYZ and PVYE groups are not frequent. Differences in aphid transmission between strains were reported. PVYN isolates seem better transmitted than other PVY isolates with longer retention periods in aphids. Whatever the strain, Myzus persicae is clearly the most important vector in most potatogrowing areas. However, despite their low transmission efficiency, other species colonizing potatoes such as Aphis nasturtii, or visiting potatoes can also contribute to PVY spread. Some of these species, notably cereal aphids or the pea aphid, seem to be involved in PVY epidemics in potato crops in Europe and the USA due to their high abundance.

Control Methods Control methods are first based on sanitary control of seed potatoes and breeding for resistance, especially regarding PVYN. Sophisticated schemes of seed production include monitoring of aphid vectors, treatments by mineral oils against aphid transmission, eradication of weeds, and post-harvest detection tests using large-scale ELISA. Numerous molecular assays were also developed for PVY detection in leaves, potato tubers, and aphids, including numerous RT-PCR protocols, microarray technology, and real-time RT-PCR methods. Resistance based on hypersensitive reaction (HR) and extreme resistance (ER) against PVY are

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Fig. 7 Schematic representation of the main molecular diversity within potato PVY strain groups. Boxes in white, brown and black correspond to PVYO, PVYC and PVYN nucleotide sequences, respectively. RJ1, RJ2, RJ3 and RJ4 correspond to the position of the hot spots of recombination described in PVY genome. “Other” refers to some PVY isolates displaying particular biological properties. The arrow indicates the presence of single nucleotide mutation leading to a serological misidentification. Adapted from Glais, L., Bellstedt, D.U., Lacomme, C., 2017. Diversity, characterization and classification. In: Lacomme, C., Glais, L., Bellstedt, D.U., et al. (Eds.), Potato Virus Y: Biodiversity, Pathogenicity, Epidemiology and Management. Switzerland: Springer, pp. 43–76.

largely used. HR is based on the single dominant genes Nc, Nz and Ny and ER is based on the Ry genes. The Ry genes (Ryadg, Rychc, Ry(o)phu, Rysto), from wild Solanum species, map on chromosomes 11, 9, 9, and 12, respectively, and confer broad-spectrum resistance, whereas HR genes Nc, Nytbr (which map on chromosome 4) and Nz found in old potato cultivars, protect against PVYC, PVYO, and PVYZ, respectively. More recently, Ny-1 and Ny-2 genes eliciting hypersensitive response against both PVY1 and PVYN isolates were mapped on potato chromosomes 9 and 11, respectively. Beside these well documented resistance genes, new putative resistance genes were reported according to local symptoms observed on leaves inoculated with a set of differential PVY strains. Nw, Ne, Nn, Nna seem to confer resistance against PVYN-Wi (or PVYN:O), PVYE, PVYN, and PVYN from North America, respectively (Table 2). Sources of resistance against aphids were identified in wild Solanum species but not introgressed in potato cultivars yet.

PVY in Pepper PVY is the causal agent of major diseases and production losses in pepper crops. PVY affects pepper production worldwide, while other pepper-infecting potyviruses, like Tobacco etch virus, Pepper severe mosaic virus, Pepper yellow mosaic virus, Pepper veinal mottle virus or Chilli veinal mottle virus are mostly restricted to particular continents and are not present in Europe. Symptoms are mostly visible on the vegetative parts of the plants but rarely on fruits. Depending on the pepper genotype and on the virus isolate, they consist of mosaic and vein banding on leaves or necrotic symptoms on leaves, petioles, and stems (Fig. 8(A) and (B)). The most widespread and efficient way to control PVY in pepper is through the growing of cultivars carrying resistance genes. The first resistance genes used to control PVY were the pvr21 and pvr22 alleles, which control high-level resistance since no virus can be detected in inoculated organs of the plants. These genes have been exploited for decades in a large number of pepper cultivars and proved to be durable. However, PVY isolates virulent to the pvr21 allele were occasionally observed, especially in the Mediterranean basin and in tropical regions, while isolates virulent to the pvr22 allele are exceptional.

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List of identified PVY resistance genes in various hosts

Carrier chromosome

Gene or allele name

Nature of encoded protein

Source of resistance

Potato Chr. 11 Chr. 12 Chr. 9 Chr. 9 Chr. 4 Chr. 4 Chr. 9 Chr. 11 – – – –

Ryadg Rysto Rychc Ry(o)phu Nytbr Nc Ny  1 Ny  2 Nz Nw Ne Nn

– NLR – NLR – – – – – – – –

S. S. S. S. S. S. S. S. S. S. S. S.

Tobacco Chr. 21

va

eIF4E

N. tabacum ‘Virgin A Mutant’

pvr2 (430 resistance alleles) Pvr4 Pvr7

eIF4E

430 accessions in C. annuum, C. frutescens, C. chinense, C. baccatum,

NLR –

C. annuum ‘CM3340 C. chinense ‘PI1592360

pot  1 pot  12

eIF4E eIF4E

Solanum habrochaites ‘PI2470870 Solanum pimpinellifolium ‘LA04110

Pepper P4 P10 P10 Tomato Chr. 3 Chr. 3

andigena stoloniferum chacoense phureja tuberosum tuberosum tuberosum tuberosum tuberosum tuberosum tuberosum tuberosum

Note: eIF4E: eukaryotic initiation factor 4E; NLR: nucleotide-binding leucine-rich repeats. “–” corresponds to unknown data.

These two alleles and the wild-type susceptibility allele pvr2 þ encode different copies of the eukaryotic translation initiation factor 4E1 (eIF4E1) that differ by a few amino acid substitutions. Reciprocally, amino acid substitutions in the central part of the VPg of PVY determine the virulence towards pvr2 alleles. In addition to pvr21 and pvr22, more than 30 pvr2 resistance alleles are found in locallygrown pepper accessions of almost all Capsicum species analyzed. Most of these alleles have a specific resistance spectrum that match PVY VPg diversity. The large amino acid diversity in both pepper eIF4E1 and PVY VPg and evidence of positive selection during the evolution of these two proteins suggest a co-evolution between pepper and PVY. Pepper infection and resistance based on eIF4E1/VPg interaction obey a lock-key model: direct binding between eIF4E1 and VPg proteins allows PVY to replicate and translate its genome and to infect the plant. In contrast, amino acid substitutions in either the eIF4E1 or VPg that abolish binding are responsible for plant resistance. There are great differences in durability for the resistance conferred by different pvr2 alleles. From the 1990s, the dominant Pvr4 gene, originating from a Mexican C. annuum population, was introgressed into bell-pepper cultivars. This gene confers a high-level resistance to all tested isolates of PVY in addition to five other potyviruses belonging to the PVY phylogenetic group: EcRV (Ecuadorian rocoto virus), PepMoV (Pepper mottle virus), PepSMV (Pepper severe mosaic virus), PepYMV (Pepper yellow mosaic virus), and PTV (Peru tomato mosaic virus). This resistance shares common properties with hypersensitivitybased resistances. Only a few laboratory-selected PVY isolates are able to break down the Pvr4 resistance, indicating a high durability of the resistance. The Pvr4 gene was shown to encode a nucleotide-binding and leucine-rich (NLR) domain protein. It shares common structure and sequence with the Tsw gene that confers resistance to Tomato spotted wilt virus, suggesting a common origin. The ability of laboratory isolates to break the Pvr4 resistance was shown to be due to a single substitution in the NIb coding region. This mutation induces a high competitiveness cost to PVY in plants devoid of the Pvr4 gene, which may explain the high durability of this resistance. Other pepper resistances to PVY have been described but not used in commercial cultivars. A polygenic resistance from the C. annuum accession Perennial was analyzed and several QTLs (quantitative trait loci) have been mapped in the plant genome. These QTLs can be used in combination to a pvr2 resistance allele to improve its durability. The Pvr7 resistance gene from the Capsicum chinense accession PI159236 confers a high level of resistance to PVY and PepMoV and is allelic to Pvr4. Other control measures such as the use of oil sprays or physical barriers (polyethylene sheets and coarse nets) were also used for controlling spread of PVY in pepper crops in Israel and Florida.

PVY in Tomato PVY can induce severe diseases in tomato crops but has long been considered a pathogen of secondary importance, inducing mild mosaic on the foliage only. Since the 1980s, new strains of PVY have arisen in Mediterranean countries, causing serious

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Fig. 8 PVY infection in a pepper field: (A) severe crinkling and distortion of the leaves. (B) Necrotic patterns on a leaf. Most pepper isolates of PVY belong to the C1 phylogenetic group, but more and more N/O recombinants are observed. An O/C recombinant strain inducing veinal necrosis was also isolated in pepper in Italy. PVY infection on tobacco cv. Burley in field trials in Southwestern France. (C) Chlorotic oak-leaf and halo patterns. (D) Severe systemic necrosis. Photographs: P. Gognalons, (A) and M. Pepelnjak Slovenia (B), Blancard (C) and (D), INRAE, France.

yield and quality loss of tomato fruits. These strains induce necrotic lesions on leaves and necrotic streaks on stems in all tomato varieties and frequently affected 100% of the plants in greenhouses as well as in open fields. Importantly, combination of PVY infections with other viruses such as Cucumber mosaic virus can induce very severe diseases in tomato crops. Data about the genetic diversity of PVY isolates from tomato are scarce. Isolates belonging to the C1, N, O, and N/O groups have been observed in tomato crops. In laboratory tests, most of the isolates belonging to all PVY phylogenetic groups induce systemic infections in tomato. There are few reports about PVY resistance in tomato and related species. Accession PI247087 of Solanum habrochaites was described highly resistant to PVY. The resistance is efficient toward all tested PVY isolates, inhibits the multiplication of the virus in the inoculated organs and is controlled by a single recessive gene (named pot-1) which is homologous to the pvr2 gene in pepper and encodes the eIF4E1. Strains of PVY virulent to the pot-1 resistance can be selected in laboratory tests, a property that is controlled by a single aa substitution in the VPg. The pot-1 gene was introduced into genotypes of S. lycopersicum, but varieties carrying this gene are not widespread, possibly because PI247087 is an accession of the wild species S. habrochaites and introgression of the resistance into cultivated tomato is difficult or is deleterious for yield and/or quality. Accession LA0411 of Solanum pimpinellifolium carries also a pot-1 mutant allele (pot-12) that confers resistance to PVY, but the spectrum of the resistance is narrow (only a few PVY isolates) and its durability potential is low. Engineered resistance was also developed in tomato (S. lycopersicum) using a TILLING (targeting induced local lesions in genomes) approach, which combines ethyl methane sulfonate mutagenesis and gene-specific detection of mutations. An eIF4E1 KO (knock out) mutant showed a narrow spectrum of resistance against PVY. This spectrum was enlarged to all tested PVY isolates (and to Tobacco etch virus isolates) when the eIF4E1 KO mutation was combined with a KO mutation in another gene of the eIF4E family, eIF4E2. However, this double mutant showed a stunted growth.

PVY in Tobacco PVY was reported to be the most damaging virus in tobacco crops. It causes height reductions and modifies the chemical composition of cured leaves, especially the nicotine content. Yield losses of 14%–59% were reported with incidence of up to 100%. Symptoms on leaves are usually mild mottling but particular chlorotic patterns and necroses, notably the veinal necrosis disease, may also occur (Fig. 8(C) and (D)). PVY isolates are split into two groups according to their capacity to induce (PVYN) or

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not (PVYO and PVYC) veinal necrosis on tobacco leaves. As said previously, the HC-Pro was shown to carry molecular determinants involved in this phenotype. Recently, the RPP8-like gene in tobacco, called NtTPN1 (Nicotiana tabacum Tolerance to PVY-induced Necrosis1), was shown to determine the expression of the necrotic symptoms, presumably through direct or indirect interaction with viral HC-Pro protein, because the Glycine to Arginine mutation at codon position 497 in NtTPN1 is sufficient to suppress PVYN-induced veinal necrosis. Genome analyses of PVY isolates collected in tobacco in different countries reported that O, N, and C1 isolates as well as N/O recombinants (PVYNTN, PVYN-Wi) are found on tobacco. Breeding for resistance is the main control method against PVY in tobacco production. Many European and American commercial cultivars carry recessive resistances of varying efficiency and durability. The va gene of the Virgin A Mutant (VAM) line, corresponding to a large deletion around the eIF4E-1 gene on chromosome 21 after X-ray irradiation, has been extensively used, though its introgression is often associated with a decrease of yield and cured leaf quality. Nevertheless, some va-breaking necrotic PVY isolates were observed in open fields. Recently, it was reported that the durability of this resistance is enhanced by a complex genetic locus on chromosome 14 containing three other eIF4E genes. The viral VPg protein is involved in the breakdown of the resistance conferred by va (Table 1). In countries where early infection occurs frequently, additional measures consist in protecting seedbeds by fleece, eradication of weeds and isolation from potato, tomato and pepper fields. Pathogen-derived resistance has been extensively studied in tobacco, although more for investigating the mechanisms of protection than for use in commercial tobacco cultivars.

Prospects Since the first identification of PVY, a significant amount of information was collected by researchers. However, some gray areas remain in understanding the complexity of the interaction between PVY and its hosts and its vectors: How to explain the emergence and prevalence of PVY recombinants in crops? How is PVY aphid transmission activated? What are the plant and virus molecular determinants involved in pathogenicity (symptoms, resistance breakdown or induction)? How to better understand and exploit resistance genes against PVY? Are aphids the only insects to transmit PVY? A better understanding of the plant/PVY/vector pathosystem would allow us to control better PVY epidemics.

See also: Bean Common Mosaic Virus and Bean Common Mosaic Necrosis Virus (Potyviridae). Papaya Ringspot Virus (Potyviridae). Plum Pox Virus (Potyviridae). Potyviruses (Potyviridae)

Further Reading Faurez, F., Baldwin, T., Tribodet, M., Jacquot, E., 2012. Identification of new Potato virus Y (PVY) molecular determinants for the induction of vein necrosis in tobacco. Molecular Plant Pathology 13, 948–959. Glais, L., Bellstedt, D.U., Lacomme, C., 2017. Diversity, characterization and classification. In: Lacomme, C., Glais, L., Bellstedt, D.U., et al. (Eds.), Potato Virus Y: Biodiversity, Pathogenicity, Epidemiology and Management. Switzerland: Springer, pp. 43–76. Glais, L., Faurez, F., Tribodet, M., Boulard, F., Jacquot, E., 2015. The amino acid 419 in HC-Pro is involved in the ability of PVY isolate N605 to induce necrotic symptoms on potato tubers. Virus Research 208, 110–119. Janzac, B., Tribodet, M., Lacroix, C., et al., 2014. Evolutionary pathways to break down the resistance of allelic versions of the PVY resistance gene va. Plant Disease 98, 1521–1529. Janzac, B., Willemsen, A., Cuevas, J.M., et al., 2015. Brazilian Potato virus Y isolates identified as members of a new clade facilitate the reconstruction of evolutionary traits within this species. Plant Pathology 64, 799–807. Kim, S.B., Kang, W.H., Huy, H.N., et al., 2017. Divergent evolution of multiple virus-resistance genes from a progenitor in Capsicum spp. New Phytologist 213, 886–899. Lacomme, C., Jacquot, E., 2017. General characteristics of potato virus Y (PVY) and its impact on potato production: An overview. In: Lacomme, C., Glais, L., Bellstedt, D.U., et al. (Eds.), Potato virus Y: Biodiversity, Pathogenicity, Epidemiology and Management. Switzerland: Springer, pp. 1–19. Michel, V., Julio, E., Candresse, T., et al., 2018. NtTPN1: A RPP8-like R gene required for Potato virus Y-induced veinal necrosis in tobacco. Plant Journal 95, 700–714. Michel, V., Julio, E., Candresse, T., et al., 2019. A complex eIF4E locus impacts the durability of va resistance to Potato virus Y in tobacco. Molecular Plant Pathology 20, 1051–1066. Moury, B., 2010. A new lineage sheds light on the evolutionary history of Potato virus Y. Molecular Plant Pathology 11, 161–168. Quenouille, J., Vassilakos, N., Moury, B., 2013. Potato virus Y: A major crop pathogen that has provided major insights into the evolution of viral pathogenicity. Molecular Plant Pathology 14, 439–452. Rowley, J.S., Gray, S.M., Karasev, A.V., 2015. Screening potato cultivars for new sources of resistance to Potato virus Y. American Journal of Potato Research 92, 38–48. Ruffel, S., Gallois, J.L., Lesage, M.L., Caranta, C., 2005. The recessive potyvirus resistance gene pot-1 is the tomato orthologue of the pepper pvr2-eIF4E gene. Molecular Genetics and Genomics 274 (4), 346–353. Uzest, M., Gargani, D., Drucker, M., et al., 2007. A protein key to plant virus transmission at the tip of the insect vector style. Proceedings of the National Academy of Sciences of the United States of America 104, 17959–17964. White, K.A., 2015. The polymerase slips and PIPO exists. EMBO Reports 16, 885–886.

Potexviruses (Alphaflexiviridae) Ki H Ryu and Eun G Song, Seoul Women’s University, Seoul, South Korea Jin S Hong, Kangwon National University, Chunchon, South Korea r 2021 Elsevier Ltd. All rights reserved. This is an update of K.H. Ryu, J.S. Hong, Potexvirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00738-X.

Nomenclature

MP Movement protein nt Nucleotide(s) NTR Non-translated region ORF Open reading frame PCR Polymerase chain reaction Pol RNA-dependent RNA polymerase Poly(A) Polyadenylated RdRp RnA-dependent RNA polymerase RT-PCR Reverse transcription polymerase chain reaction SL Stem loop ssRNA Single-stranded RNA TEM Transmission electron microscopy TGB Triple gene block

aa Amino acid(s) CP Coat protein or capsid protein EM Electron microscopy FD1 Ferredoxin 1 Hel Helicase ISH In situ hybridization JAX1 Jacalin-type lectin gene kb Kilobase kDa Kilo Dalton LAMP Loop mediated amplification LFIA Lateral flow immunoassay Met Methyl-transferase

Glossary Jacalin-type lectin 1 (JAX1) Antiviral resistance gene to Potato virus X related to lectin – mediated resistance (LMR) confers plant innate immunity against viruses. RNA silencing suppressor Plant viruses have encoding protein to counteract the plant antiviral system known as RNA silencing.

Rx protein A highly similar CC-NBS-LRR protein, which prevents virus accumulation, mediates extreme resistance against PVX. Triple gene block The protein comprises of a substantially similar element of three partially overlapping ORFs and is contained within their genomes. The TGB proteins are correlated with cell-to-cell movement and potentially influence on host antiviral defenses.

Introduction A genus Potexvirus is one of seven genera in the family Alphaflexiviridae. Most potexviruses are found wherever their hosts are grown. The genus Potexvirus contains 38 definitive species including 35 species determined complete genome sequences. The genomes of Cassava virus X (CsVX) and Lagenaria mild mosaic virus (LaMMoV) have been partially sequenced and that of Plantain virus X (PlVX) has yet been reported. There are also 5 tentative species for which genomic sequences are available, but they have not yet been accepted by ICTV (Table 1). The members of the genus Potexvirus can be divided into three main phylogenetic groups based on their genome sequences. The phylogenetic relationships of the genus Potexvirus are not associated with their natural host other than cactus-infecting viruses. The virions of the genus Potexvirus have nonenveloped and flexible particles of 470–1000 nm in length and 12–14 nm in diameter. Their genomes of 5.8–7.2 kb in length consist of a positive-sense single-stranded RNA (ssRNA) with 5 0 terminal cap structure and a 30 terminal poly(A) tail. The genomes encode for an RNA-dependent RNA polymerase (RdRp) for viral replication, a triple gene block (TGBp1,2, and 3) for viral movement, and a coat protein (CP). The RdRp is associated with jacalin‐type lectin required for potexvirus resistance 1 (JAX1)‐mediated resistance of host plant. The TGBp1, known as viral suppressors of RNA silencing, acts an avr protein associated with an HR‐like response. Host plants naturally infected with potexviruses may show mosaic, mottle, necrosis, chlorosis, spots, or dwarf symptoms or may be symptomless. The natural host range of individual viruses is usually restricted to a few plant species, although a few of the viruses can infect a wide range of plant species. Potexviruses are transmitted mechanically through contact with virus-infected plant sap, or with contaminated agricultural equipment. Most potexviruses are not transmitted by vertebrate, invertebrate, or fungal vectors; Strawberry mild yellow edge virus (SMYEV) and Potato aucuba mosaic virus (PAMV) are transmitted by aphids. The type species of the genus Potexvirus is Potato virus X (PVX). During the last few decades, PVX has greatly contributed to our understanding of host resistance and gene silencing mechanisms, in particular because of the use of viral vectors in studying gene expression and RNA silencing in plants.

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Table 1 Virus species in the genus Potexvirus of the family Alphaflexiviridae. Type species in the genus is written in bold. Complete sequences accession number and length (nt) of the genomes are indicated (ICTV, 2019) Species name

Acronym

Accession number

Genome length (nt)

Actinidia virus X Allium virus X Alstroemeria virus X Alternanthera mosaic virus Asparagus virus 3 Babaco mosaic virus Bamboo mosaic virus Cactus virus X Cassava common mosaic virus Cassava virus X Clover yellow mosaic virus Cymbidium mosaic virus Foxtail mosaic virus Hosta virus X Hydrangea ringspot virus Lagenaria mild mosaic virus Lettuce virus X Lily virus X Malva mosaic virus Mint virus X Narcissus mosaic virus Nerine virus X Opuntia virus X Papaya mosaic virus Pepino mosaic virus Phaius virus X Pitaya virus X Plantago asiatica mosaic virus Plantain virus X Potato aucuba mosaic virus Potato virus X Schlumbergera virus X Strawberry mild yellow edge virus Tamus red mosaic virus Tulip virus X Vanilla virus X White clover mosaic virus Yam virus X Zygocactus virus X

AVX AlVX AlsVX AltMV AV3 BabMV BaMV CVX CsCMV CsVX ClYMV CymMV FoMV HVX HdRSV LaMMoV LeVX LVX MalMV MVX NMV NVX OpVX PapMV PepMV PhVX PiVX PlAMV PlVX PAMV PVX SchVX SMYEV TRMV TVX VVX WClMV YVX ZyVX

NC028649 NC012211 NC007408 NC007731 NC003400 NC036587 NC001642 NC002815 NC001658 NC034375a NC001753 NC001812 NC001483 NC011544 NC006943 NC043079a NC010832 NC007192 NC008251 NC006948 NC001441 NC007679 NC006060 NC001748 NC004067 NC010295 NC024458 NC003849 ND NC003632 NC011620 NC011659 NC003794 NC016003 NC004322 NC035205 NC003820 NC025252 NC006059

6888 7118 7009 6607 6985 6692 6366 6614 6376 5879 7015 6227 6151 6528 6185 3600 7212 5823 6858 5914 6955 6582 6653 6656 6450 5816 6677 6128

EuYVV

NC035190 NC040842 NC030746 NC040644

7279 5839 6775 6278

Tentative species Euonymus yellow vein virus Potexvirus sp. Senna mosaic virus Turtle grass virus X

SeMV TuGVX

7059 6435 6633 5966 6495 6056 6295 5845 6158 6624

a

Incomplete genome sequence.

Taxonomy, Phylogeny, and Evolution The genus Potexvirus is one of seven genera in the family Alphaflexiviridae. Its name is derived from Potato virus X (the type species of potexviruses). The genus contains 39 definitive species, of which 36 have complete genome sequences (Table 1). The compete genome sequences of some members have recently been determined. The genomes of Cassava virus X (CsVX) and LaMMoV have been partially sequenced. The genome sequences of PlVX has yet been reported. Another 4 tentative species have genomic sequences available, but they have not yet been accepted by ICTV (Table 1). Phylogenetic analysis using the complete genome sequences of the 36 potexviruses of their RNA-dependent RNA polymerase (RdRp), has revealed that they can be divided in seven major branches from I to VII (Fig. 1). Phylogenetic analysis of the complete sequences of the same 36 potexviruses of their coat protein (CP), (Fig. 1) and their triple gene block (TGB) proteins (Fig. 2), are showing that the seven clusters, with only a few exceptions, can be identified for all the proteins with the except for the TGBp3, which is showing a major reorganization of the potexvirus diversity. However it is also important to note that the TGBp3 phylogenetic tree is not very stable as only 5 branches have a bootstrap value above 50%. This result seems to indicate that there has been a co-evolution of the genes/proteins composing the potexvirus genome, with perhaps the exception of TGBp3. These

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Fig. 1 Phylogenetic analysis of genus Potexvirus based on the amino acid sequences of the RdRP and CP. Multiple sequence alignments were generated using DNAMAN software version 5.1 (Lynnon Biosoft, San Ramon, Ca, USA) and phylogenetic trees were constructed by the neighborjoining algorithm, based on calculations from pairwise amino acid sequence distances for protein analyses derived from the multiple alignment format. The horizontal branch lengths are proportional to the genetic distance, and the numbers at each point indicate bootstrap values. The dataset was subjected to 10,000 bootstrap replicates and values above 50% are indicated on the branches. Virus names are indicated by their abbreviation and their full names are in Table 1. NCBI database accession numbers follow the virus abbreviation name. The black vertival bars identify the seven major clusters (from I to VII) as defined per the RdRp phylogenetic tree and differential colored boxes are shown in each tree.

phylogenetic trees do not show a correlation of the viruses with their type of host with the exception of the cactus-infecting viruses such as Cactus virus X (CVX), Pitaya virus X (PiVX), Opuntia virus X (OpVX), Schlumbergera virus X (SchVX), and Zygocactus virus X (ZyVX) that are grouped together in each tree (cluster III). The species demarcation criteria in the genus Potexvirus are the followings: (1) members of distinct species have less than 78% nt or 61% aa identity of their RdRp, less than 73% nt or 73% aa identity of their TGBp1, less than 65% nt or 64% aa identity of their TGBp2, less than 66% nt or 59% aa identity of their TGBp3, and less than 57% nt or 49% aa identity of their CP; (2) members of distinct species are readily differentiated by serology, and some virus strains can be differentiated with monoclonal antibodies; and (3) members of distinct species fail to cross-protect in common host plant species, and they usually have distinguishable experimental host ranges.

Virion Structure Virions are non-enveloped and flexible filaments of 470–1000 nm in length and 12–14 nm in diameter (Fig. 3). They have helical symmetry with a pitch of 3.3–3.6 nm. The number of protein subunits per turn of the primary and final symmetries is slightly less than 9.0. The viral RNA is at a radius of 3.0–3.5 nm. Virion Mr is about 3.5  106 with 5%–7% nucleic acid content. PVX virion is a filamentous particle of about 515 nm in length and 13.5 nm in diameter and forms a helical array (3.5 nm pitch) of about 1300 identical CP subunits with the viral RNA packed between turns of the helix. The virion has 8.9 CP subunits, each consisting of 236 aa residues, per primary helix and five nucleotides are related to each protein subunit. The viral RNA molecule have about 6% by weight of the virion. Narcissus mosaic virus (NWV) helix has eight subunits per turn and an outer radius of 5.5 nm. The assembly of potexvirus ribonucleoproteins does not depend on the RNA nucleotide sequences.

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Fig. 2 Phylogenetic analysis of genus Potexvirus based on the amino acid sequences of the TGBp1, TGBp2, and TGBp3. Multiple sequence alignments were generated using DNAMAN software version 5.1 (Lynnon Biosoft, San Ramon, Ca, USA) and phylogenetic trees were constructed by the neighbor-joining algorithm, based on calculations from pairwise amino acid sequence distances for protein analyses derived from the multiple alignment format. The horizontal branch lengths are proportional to the genetic distance, and the numbers at each point indicate bootstrap values. The dataset was subjected to 10,000 bootstrap replicates and values above 50% are indicated on the branches. Virus names are indicated by their abbreviation and their full names are in Table 1. NCBI database accession numbers follow the virus abbreviation name. The black vertival bars identify the seven major clusters (from I to VII) as defined per the RdRp phylogenetic tree and differential colored boxes are shown in each tree.

Fig. 3 Electron microscopy of Potato virus X (PVX) particles negatively stained. The scale bar is 100 nm. Picture courtesy of Lesemann, D.E.

Genome Organization The genome of potexviruses is a sense single-stranded positive-sense RNA (ssRNA) with 50 terminal cap structure and a 30 terminal poly (A) tail. The genome of 5.8–7.2 kb in size contains five ORFs (encoding an RdRP), three movement proteins called TGB, and the CP from the 50 to the 30 end (Fig. 5). The genome organization resembles that of genera Foveavirus and Robigovirus in the family Betaflexiviridae, but is distinguishable from them by the size of genome (8–9.3 kb). The complete genomic sequences of almost all potexviruses has been determined (Table 1). The genome structure of PVX is showed in Fig. 4. The genome consists of 6435 nt and contained five ORFs coding for the proteins of an RdRp of 166 kDa (ORF1; 4371 nt), a TGBp1 of 25 kDa (ORF2; 681 nt), a TGBp2 of 12 kDa (ORF3; 348 nt), a TGBp3 of 8 kDa (ORF4; 213 nt), and a CP of 25 kDa (ORF5; 714 nt) from the 50 to 30 end. Both terminal regions of ORF3 (12 kDa) shows overlap with ORF2 and ORF4, respectively. The 50 terminal region of ORF5 shows an overlap with ORF3. The lengths of the 50 non-translated region (NTR) and the 30 NTR have 84 and 73 nt. The 30 NTR region contains the poly(A) tail. Some potexviruses eg, Actinidia virus X (AVX); Alstroemeria virus X (AlsVX); Cymbidium mosaic virus (CymMV); NMV; PVX; SMYEV; White clover mosaic virus (WClMV), have a putative ORF6 located within the ORF5 or over the ORF5 and 30 NTR. Bamboo mosaic virus (BaMV) is frequently associated with a satellite RNA (satRNA). The satRNA is a linear RNA of 836 nt and comprises a single ORF encoding a 20 kDa protein.

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Fig. 4 Genome organization of Potato virus X, the type species of the genus Potexvirus. RNA-dependent RNA polymerase (166 kDa) (green box) includes a methyl-transferase (Met), a helicase domain (Hel), a polymerase (Pol) domains. Purple boxes indicate the triple gene block corresponding to three overlapping ORFs (TGBp1, TGBp2, and TGBp3), and ORF5 encodes the CP (orange box).

Fig. 5 Symptoms of potexvirus-infected plants. (A) Malva spp. infected with Malva mosaic virus (MalMV) exhibiting symptom of mosaic on leaves. (B) Hydrangea spp. infected with Hydrangea ringspot virus (HdRSV) exhibiting symptoms of chlorotic and necrotic ringspots on leaves. (C) Lilium spp. infected with Plantago asiatica mosaic virus (PlAMV) exhibiting symptoms of brown necrotic streaking and irregular chlorosis on leave. (D) Solanum lycopersicum infected with Pepino mosaic virus (PepMV) exhibiting symptoms of open fruit. (E) Bambusa vulgaris infected with Bamboo mosaic virus (BaMV) exhibiting symptoms of interveinal chlorotic mosaic and stripes on leaves. (F) Dendrobium phalaenopsis infected with Cymbidium mosaic virus (CymMV) exhibiting symptoms of chlorotic and necrotic ringspots on leaves.

Properties and Functions of Gene Products The RdRP of potexviruses contains three conserved domains: a methyl-transferase (Met) domain; an RNA helicase (Hel) domain; a polymerase (Pol) domain. The RdRP is related to jacalin-type lectin required for potexvirus resistance 1 (JAX1)-mediated resistance of host plant. The cell-to-cell movement of potexviruses requires the poxviruses TGB and CP. TGBp1 and CP form a complex with viral RNA that transfers to the next cell via the plasmodesmata. The TGBp1 also acts as viral suppressors of RNA silencing and an avr protein

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associated with an HR-like response. It has been recently demonstrated that PVX TGBp1 interacts with the chloroplast protein ferredoxin 1 (FD1) related to its role in the chloroplast electron transport chain. The TGBp1 of Pepino mosaic virus (PepMV) is associated with a thioredoxin domain-containing protein that is related to virus movement. TGBp2 and TGBp3 are endoplasmic reticulum-binding proteins for virus movement. The TGBp2 is also associated with a functional role in PVX replication. BaMV TGBp2 interacts with a Nicotiana benthamiana-expressed thioredoxin type h protein (NbTRXh2), which acts as a negative regulator of viral movement. CP is required for virion assembly. PVX CP is the elicitor for Rx-mediated resistance that impairs virus accumulation in host plant. The CP of Potato aucuba mosaic virus (PAMV) has a DAG motif associated with aphid transmissibility, located in the aa residues 14–16 from the N‐terminus of the CP, which is also found in the CP of potyviruses. Satellite RNA (satRNA), with crucial roles in modulating viral replication and symptom development, is associated with BaMV and it was found to be the only satRNA of the genus Potexvirus. The RNA-binding domain of PaMV CP, located between 90 and 130 aa, is associated with virion assembly.

Replication and Propagation The 50 NTR regulates plus-strand RNA synthesis, sub-genomic RNA synthesis, cell-to-cell movement, and virion assembly. Two RNA stem-loop (SL) structures within the 50 NTR, named 50 SL1 and 50 SL2, are required for viral replication. The 50 SL1 is a multifunctional element associated with virus replication, gene expression, cell-to-cell movement, and virion assembly. RdRP can be translated directed from the genomic RNA and then three sub-genomic RNAs are generated for TGB and CP synthesis. The 30 NTR including three RNA stem-loop structures, named 30 SL1, 30 SL2, and 30 SL3, controls minus-strand and plus-strand RNA accumulations. The 30 NTR has multifunctional activities, related to viral replication and host-factor interaction. The RNA replication and accumulation of PVX interact synergistically with potyviruses (Pepper mottle virus, Plum pox virus, Potato virus Y, Tobacco vein mottling virus, and Tobacco etch virus). The PVX/potyviral-associated synergism is related to the 50 proximal potyviral sequence and microRNAs derived from co-infected plants. The 50 NTR of satRNA, which associated with BaMV, acts as a regulator of BaMV replication. The transgene induced from potexvirus-based vector can be stably expressed as a N-terminal translational CP fusion linked by the autocatalytic 2A peptide, which is derived from foot-and-mouth disease virus.

Transmission, Host Range Potexviruses can be naturally transmitted to various horticultural crops. Most potexviruses can be transmitted mechanically via contact with virus-infected plant sap, or with contaminated agricultural equipment. PAMV and PVX cause common symptoms in Solanum tuberosum grown worldwide. PAMV exhibiting slight leaf mottling on S. jasminoides causes yellow leaf flecking, deformation, and stunting on susceptible potato plants. PVX cause mild mosaic, necrosis, or is symptomless on potato plants and can be co-infected with Potato virus Y and Candidatus phytoplasma trifolii on S. melongena. Five viruses, CVX, OpVX, PiVX, SchVX, and ZyVX, cause mild mosaic, mosaic, necrosis, ringspots or are symptomless on Cactaceae. The CVX, OpVX, SchVX, and ZyVX can be transmitted by mechanical inoculation. LVX, PlAMV, and Tulip virus X (TVX) can be naturally transmitted to Lilium spp. The PlAMV can cause mosaic and necrosis on Plantago asiatica and Solanaceae, as well as Lilium spp. The TVX can cause chlorotic and necrotic spots of leaves and streaks of petals in Lilium spp. and Tulipa spp. The LVX appears to be asymptomatic in Lilium spp. Cassava common mosaic virus (CsCMV) and CsVX can be naturally transmitted to Manihot esculenta and these viruses can be mechanically transmitted to N. benthanmiana. The CsCMV causes mosaic and chlorosis on M. esculenta while the CsVX exhibits no symptoms. Clover yellow mosaic virus (ClYMV) and WClMV can be naturally transmitted to Trifolium spp. The ClYMV causes chlorotic streaks of leaves and color breaking of flowers on T. gesneriana and it causes yellow mosaic or is symptomless on T. repens. CymMV causes chlorotic or necrotic spots on the leaves and flowers of orchid plants. The flowers infected with the CymMV exhibit deformation and color breaking. CymMV has also been associated with flecking on leaves, necrotic spots on the stem, and vine decline of vanilla plants. Papaya mosaic virus (PapMV) causes mosaic and stunting on Carica papaya while it exhibits no symptoms on Ullucus tuberosus. PepMV can be naturally transmitted to greenhouse tomato species (S. lycopersicum) and wild tomato species (S. chilense, S. chmielewskii, S. parviflorum, and S. peruvianum) and it causes fruit marbling and discoloration, open fruit, nettle-heads, leaf blistering or bubbling, leaf chlorosis and yellow angular leaf spots, leaf mosaic and leaf or stem necrosis in susceptible tomato plants. SMYEV causes marginal yellowing symptoms on leaves of commonly cultivated strawberry crops in worldwide. LVX, SMYEV, PAMV, and PepMV can be transmitted by aphids. Hosta virus X (HVX) can be naturally transmitted to Hosta spp. and cannot be transmitted mechanically to N. benthanmiana and N. clevelandii. HVX and PepMV can be transmitted by seeds. BaMV can be transmitted by dipterans (Gastrozona fasciventris and Atherigona orientalis). Some viruses are not transmitted by mechanical inoculation. Allium virus X (AlVX) causes diffuse yellow stripes or mottle on leaves of Allium spp. Yam virus X (YVX) causes mild chlorosis on leaves of Dioscorea trifida. The methods for transmission of three potexviruses, such as AlVX, PiVX, and YVX, are presently not known.

Epidemiology and Control Most potexviruses can cause usually mild mosaic, mosaic, chlorosis, spots, or symptomless in their host plants. Some food and ornamental cops sustain economic damage caused by potexviruses that require suitable disease management. Transgenic cactus,

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orchid, potato, and tobacco plants, which are resistant to BaMV, CymMV, CVX, PVX, and SMYEV respectively, have been developed. PVX can cause yield losses reaching 20% of productivity depending upon the virus stain. New potato varieties carrying the Rx1 and/ or Rx2 genes derived from the wild species Solanum tuberosum ssp. andigena and S. acaule, respectively, has been developed to control the virus. Salicylic acid has been demonstrated to mitigate physiological and proteomic changes caused by PVX (SPCP1 strain) in tomato. CsCMV has been found in cassava, which grows in the Americas. The CsCMV can cause a reduction of up to 60% of root productivity. CymMV has been reported in most cultivated orchid species grown in worldwide since it was first described on Cymbidium in California. The virus has also been found in vanilla from most growing areas since it was first detected in vanilla (Vanilla spp.) in the Society Islands (1987). Skimmed milk represented inactivation of CymMV on local host, but has not been evaluated on systemic host. RNA silencing is an important defense mechanism against plant viruses and it has been applied to control potexviruses. The potexvirus TGBp1, is known as RNA silencing suppressor, and it has been used frequently for producing virus-derived small interfering RNAs. PlAMV has been isolated from various species worldwide (eg, Achyranthes bidentate; Lilium spp.; Nandina domestica; Primula sieboldii; Rehmannia glutinosa; Stellaria media; Urtica urens; Viola grypoceras), since it was first reported from Plantago asiatica grown in Russia (1976). This viral disease is especially known to reduce considerably the commercial value of lilies. Early bulb test of the viral infection should be carried out in order to control the virus in greenhouse. PepMV causes a severe disease in greenhouse tomato crops worldwide, since it was found in the Netherlands. The factor associated with the epidemiology of the PepMV can be dependent on the properties of the different virus strains. The virus is currently controlled using virus-free seeds. SMYEV has often been found in co-infection with other viruses, such as Arabis mosaic virus (ArMV, genus Nepovirus), Strawberry crinkle virus (unclassified within the family Rhabdoviridae), Strawberry mottle virus (unclassified within the family Secoviridae), Strawberry polerovirus 1 (unclassified within family Luteoviridae), Strawberry pallidosis-associated virus (unclassified within the family Closteroviridae), or Tomato ringspot virus (ToRSV, genus Nepovirus), and the viral co-infection can significantly reduce the yield of cultivated strawberry crops grown in greenhouses. Chemotherapy with ribavirin has recently been tested to control of the ArMV, SMYEV, and ToRSV.

Diagnosis Potexviruses can be detected by polymerase chain reaction (PCR)-based methods, such as reverse transcription (RT)-PCR, immunocapture-RT-PCR, loop-mediated isothermal amplification (RT-LAMP) assay, multiplex RT-PCR, and real-time PCR. Some commercial antisera are used for immunological detection of potexviruses. Lateral flow immunoassay (LFIA) used with alkaline phosphatase has been developed for PVX detection and it has a lower limit of detection (0.3 ng/mL1) than that of conventional LFIA. Electron microscopy (EM) is an essential tool for virus detection and transmission electron microscopy (TEM) is especially used to observe the virus particles in virus-infected plants. Amorphous inclusion bodies, formed by virus particles, can be observed in PVX-infected plant cells. In situ hybridization (ISH) technique detecting the mixed-infection of two PepMV isolates has been recently developed. Digoxigenin-labeled RNA probes, which targeted PepMoV genomic regions exhibiting a highly specificity and sensitivity for the two isolates, was used in the ISH technique.

Concluding Remarks The complete genome sequences of AVX, YVX, and VVX have been determined recently. Novel potexviruses have been isolated from various plants. The genome of three definitive species, CsVX, LaMMoV, and PlVX, have not yet been completely sequenced. The phylogenetic relationship based on their genome sequences are not significantly associated with their natural host other than cactus-infecting viruses containing CVX, PiVX, OpVX, SchVX, and ZyVX. Some potexviruses have a putative ORF 6 within CP cording region or CP-30 NTR regions. Potexviruses are commonly known to be transmitted by aphids. Additionally, BaMV can be transmitted by diptera insects. Rx and JAX1 genes regulate plant innate immunity against PVX. TGBp1 of PVX is a suppressor of RNA silencing that protects host cells against the pathogen. Potexviruses are currently mostly controlled by using virus-free seed production and early test of viral infection. Immunological and PCR-based techniques are most commonly used for detection of potexviruses.

Further Reading Aguilar, E., del Toro, F.J., Brosseau, C., et al., 2019. Cell death triggered by the P25 protein in Potato virus X-associated synergisms results from endoplasmic reticulum stress in Nicotiana benthamiana. Molecular Plant Pathology 20, 194–210. Baulcombe, D.C., Lloyd, J., Manoussopoulos, I.N., Robert, I.M., Harrison, B.D., 1993. Signal for potyvirus-dependent aphid transmission of Potato aucuba mosaic virus and the effect of its transfer to Potato virus X. Journal of General Virology 74, 1245–1253. Chang, K., Chang, L., Huang, Y., et al., 2017. Transmission of Bamboo mosaic virus in bamboo mediated by insects in the Order Diptera. Frontier in Microbiology Virology 8, 870. Cruz, S.Z., Chapman, S., Roberts, A.G., et al., 1996. Assembly and movement of a plant virus carrying a green fluorescent protein overcoat. Proceedings of the National Academy of Sciences of the United States of America 93, 6286–6290. Hanssen, I.M., Thomma, B.P.H.J., 2010. Pepino mosaic virus: A successful pathogen that rapidly evolved from emerging to endemic in tomato crops. Molecular Plant Pathology 11, 179–198.

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Pacheco, R., Garcia-Marcos, A., Barajas, D., Martianez, J., Tenllado, F., 2012. PVX-potyvirus synergistic infections differentially alter microRNA accumulation in Nicotiana benthamiana. Virus Research 165, 231–235. Petrova, E.K., Trifonova, E.A., Nikitin, N.A., et al., 2016. Study of the potexvirus ribonucleoproteins signal of assembly. Virology 71, 45–49. Ruiz-Ramon, F., Sempere, R.N., Mendez-Lopez, E., Sanchez-Pina, M.A., Aranda, M.A., 2019. Second generation of Pepino mosaic virus vectors: Improved stability in tomato and wide range of reporter genes. Plant Methods 15, 58. Sugawara, K., Shiraishi, T., Yoshida, T., et al., 2013. A replicase of Potato virus X acts as the resistance-breaking determinant for JAX1-mediated resistance. Molecular PlantMicrobe Interactions 26, 1106–1112. Verchot-Lubicz, J., Ye, C., Bamunusinghe, D., 2007. Molecular biology of potexviruses: Recent advances. Journal of General Virology 88, 1643–1655.

Potyviruses (Potyviridae) Adrián Valli and Juan A García, National Center for Biotechnology-Spanish National Research Council, Madrid, Spain Juan J López-Moya, Center for Research in Agricultural Genomics and Spanish National Research Council, Barcelona, Spain r 2021 Elsevier Ltd. All rights reserved. This is an update of J.J. López-Moya, J.A. García, Potyviruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00475-1.

Glossary Pinwheel Characteristically shaped cylindrical inclusions present in the cytoplasm of cells infected by potyvirids. PIPO (Pretty Interesting Potyviral ORF) Conserved ORF (Open Reading Frame) embedded within the genome of potyvirids in a frame different of the polyprotein, and expressed through polymerase slippage occurring in a conserved motif.

Polyprotein Protein, generally big in size, made of several smaller proteins. It is the gene expression strategy of potyvirids, as they are translated from large ORFs further processed by virus-encoded proteases. Potyvirid Member of the family Potyviridae. Virion Viral particle.

Introduction The existence of potyvirids as a specific group of plant pathogens with numerous members was recognized soon after the onset of virology as a scientific discipline. Historical evidence of symptoms caused by potyvirids in plants includes the color-breaking of infected tulip flowers that were reproduced in old Dutch paintings. Some members of this family were among the first plant viruses to be identified due to the importance of the diseases they cause, and it happened relatively soon after potyvirids were grouped according to common characteristics. Electron microscopy provided a clear taxonomic criterion to allocate viruses into the group by showing flexuous rod-shaped viral particles, and the consistent presence of pinwheel-shaped cytoplasmic inclusions in infected plant cells. Biological and serological studies, including vector organisms, genome composition and the deciphering of their fulllength genome sequences, have contributed to taxonomically differentiate potyvirids into several genera. Currently, a still evolving classification recognizes at least ten genera and a few unclassified viruses. The large number of viral species within the family Potyviridae reveals its evolutionary success. More than 210 species are included in the family, with approximately 175 in the aphid-transmitted genus Potyvirus. Members of the family are distributed throughout the world, with each particular virus being geographically limited to the area potentially occupied by susceptible host plants. Generally speaking, host ranges are constrained to a limited number of natural and experimental hosts, although some potyvirids can infect a considerable number of plant species in many botanical families. Members of the family can infect economically important crops, including grain, legumes, forages, vegetables, fruits and ornamentals, causing severe losses and consequently fostering the interest of researchers, plant breeders and agronomists seeking practical solutions to the diseases that they cause. The present entry focuses on current knowledge about viruses in the family Potyviridae, dealing with taxonomy, evolution, diagnostics, molecular biology, functional and structural aspects of viral proteins, vector transmission and host-virus interactions, including characterization of resistance genes and defense responses, as well as biotechnological applications.

Taxonomy, Phylogeny and Evolution Taxonomic standards for classification into the Potyviridae family include: (i) properties of virus particles: long flexuous filaments (Figs. 1(a) and 2); (ii) cytopathological manifestations: presence of pinwheel or scroll-shaped cylindrical cytoplasmic inclusions in infected plant cells (Fig. 1(b)), and (iii) genome structure and expression strategy: positive sense ssRNA genomes, with 50 terminal proteins and 30 poly-A tails, translated as large polyprotein precursors, and at least one embedded out-of-frame ORF (see Fig. 4). The family Potyviridae is related to the picornavirus supergroup, having an equivalent main strategy for genome expression through polyprotein synthesis and further processing, and a well conserved set of replication-related proteins being present at equivalent positions in the different genomes, in particular a viral RNA-dependent RNA polymerase (RdRp). Within the family, ten different genera are differentiated so far according to biological and molecular criteria. Potyviruses, for instance, are aphid-transmissible and possess a monopartite genome, characteristics that they share with macluraviruses, although particles of the latter are shorter in length and their first gene product in the polyprotein is missing. Other monopartite genera include whitefly-transmitted ipomoviruses, and eryophid mite-transmitted poacevirus, rymoviruses and tritimoviruses, with vector organisms belonging to genera Abacarus and Aceria. The remaining genera (Bevemovirus, Brambyvirus and Roymovirus) include few species and, along with unclassified viruses, they still have no assigned vector. Bymoviruses, in turn, have bipartite genomes being encapsidated separately, and plasmodiophoraceous fungus-like organisms of the genus Polymyxa act as their transmission vectors. Phylogenetic analysis based on the alignment of genomic regions has established standard thresholds of around 50% and 75% of

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Fig. 1 (a) Negative stain preparation of purified TEV particles. (b) Thin section showing the typical pinwheel shaped cytoplasmic inclusions present in cells of a Nicotiana benthamiana plant infected with PPV (bars equal 200 nm). Pictures are courtesy of D. López-Abella, CIB, CSIC.

nucleotide identities (variable depending on the considered segments) as demarcation criteria for assigning viruses to genera and species, respectively. The current classification is shown in Table 1. An example of phylogenetic analysis of potyvirid genomes is shown in Fig. 3. All genera are differentiated in well-defined branches, with rymoviruses being close to potyviruses, although their distinctive biological characteristics justify their placement in different genera. Viral subgroups, particularly in members of the large genus Potyvirus, are also used for classification, following for instance host range restrictions as biological demarcation criteria. Information about taxonomy and sequence of members of the family Potyviridae is available at several web-based databases (Table 2). Recombination, switching events, radiation, as well as host and geographical adaptation are proposed as drivers of evolution. Sub-populations able to differentiate and evolve independently within infected perennial plants illustrate the versatility and capacity of adaptation, and similar phenomena can happen in epidemics of viruses affecting annual hosts. Recombination of virus genomes can give rise to new virus entities with novel phenotypes in virulence, induction of symptoms, and even host range. Therefore, recombination events, partial duplications, indels and point mutations conferring different responses in infected host, as well as other factors, might help to explain the extraordinary variability observed among potyvirids.

Virion Structure Virions of viruses belonging to the family Potyviridae are flexuous rods made by protein (95%) and RNA (5%) (Figs. 1(a) and 2). They are about 11–15 nm wide, with lengths ranging from less than 700 nm, as in the case of macluraviruses, to up to 900 nm, as in ipomoviruses (Table 1). Available data on particle structure show a helical assembly of identical coat protein (CP) subunits

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Fig. 2 Structure of potyvirus virions as determined at near-atomic resolution by cryoEM for WMV. (a) The diameter and pitch dimensions corresponding to the helical arrangement of 8.8 CP subunits per turn is shown in 3D density map (resolution 4.0 Å ), with one of the subunit highlighted in blue. (b and c) The different parts are indicated according to the color code in the model and a cutaway section shows the inner side of the helix that forms a 70 Å chanel. Images are courtesy of M. Valle, CIC bioGUNE, Spain. Reproduced from Zamora, M., Méndez-López, E., Agirrezabala, X., et al., 2017. Potyvirus virion structure shows conserved protein fold and RNA binding site in ssRNA viruses. Science Advances 3. (eaao2182).

(about 2000) surrounding the nucleic acid, and a distribution of 8, 8 subunits per turn in the case of Watermelon mosaic virus (WMV). Molecular weight of CP subunits ranges from 30 to 40 kDa, with differences mainly due to the variable length of the N-terminal region. A more conserved internal CP core (about 220 amino acids) defines particle architecture. The N-terminus appears to be surface-exposed, while the C-terminal part is located in the interior. Superficially located residues might interact with other proteins during essential processes of the virus life cycle. As an example, the conserved DAG motif near the N-terminus of the CP is required for aphid transmission. Posttranslational modifications, including phosphorylations and glycosylations, have been found in the CP of potyviruses, having potential regulatory functions during the viral cycle. Also, peculiar structures at one of the virion ends might represent the presence of VPg, HCpro and CI proteins, as observed in particles of purified Potato virus Y (PVY) and Potato virus A (PVA).

Properties of the Genome The genome of monopartite potyvirids is an ssRNA molecule of þ (messenger) sense. In all members, genomic RNA presents a 50 terminal protein (VPg) and a 30 poly-A tail of variable length. The genome size ranges from 8.2 to 11.5 kb, and it comprises an ORF that spans the whole genome and codes for a long polyprotein (340–370 kDa), which generates different mature products after ongoing an autoproteolytic processing cascade. An extra ORF, embedded in the P3 cistron and transcribed by viral RNA polymerase slippage events,

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Table 1 Classification of virus species of the family Potyviridae into genera, with indication of type species, number of virus species, number of genomic RNAs, and taxonomically relevant characteristics like transmission vectors and particle morphology Genus

Type species

Abbreviation

Numbera

Genome

Particleb

Vectorc

Bevemovirus Brambyvirus Bymovirus Ipomovirus Macluravirus Poacevirus Potyvirus Roymovirus Rymovirus Tritimovirus Unclassifiedd

Bellflower veinal mottle virus Blackberry virus Y Barley yellow mosaic virus Sweet potato mild mottle virus Maclura mosaic virus Triticum mosaic virus Potato virus Y Rose yellow mosaic virus Ryegrass mosaic virus Wheat streak mosaic virus –

(BVMoV) (BVY) (BYMV) (SPMMV) (MacMV) (TriMV) (PVY) (RoYMV) (RyMV) (WSMV)

1 1 6 7 8 3 175 1 3 6 (5)

Monopartite Monopartite Bipartite Monopartite Monopartite Monopartite Monopartite Monopartite Monopartite Monopartite –

760 nm 800 nm 250–300 and 500–600 nm 900 nm 650–675 nm 680–750 nm 700–900 nm Unreported 700 nm 690–700 nm –

Unknown Unknown Fungus-like Whiteflies Aphids Mites Aphids Unknown Mites Mites –

a

Number of species according to the International Committee on Taxonomy of Viruses (Master Species List 2018b.v1). Approximate length of virus particles. c Vector organisms responsible for transmission. d Among viruses still unclassified into genera there are members of two potentially new genera Arepavirus and Celavirus. b

codes for the Pretty Interesting Potyviral ORF (PIPO) and generates the P3N-PIPO product. The bipartite genome of bymoviruses is divided into two RNAs: a long RNA-1 homologous to the 30 three-quarters portion of potyvirid monopartite genomes, and a short RNA-2 with partial similarity to the 50 region of the genome of other potyvirids. The proteolytic processing of the viral polyprotein depends on the action of either two (P1 and NIapro, or HCpro and NIapro) or three (P1, HCpro and NIapro) virus-encoded proteinases, depending on the species. In some cases, two divergent copies of P1 or HCpro could be part of the proteinase set of a single potyvirid. P1 serine- and HCpro cysteine-proteinases cleave in cis at their respective C-ends, whereas the NIapro serine-proteinase cleaves all the other sites, both in cis and trans. Information on processing cleavage sites for all known viruses of the family is available on a web-based database (Table 2). The dynamics of the proteolytic processes have been suggested to have regulatory implications for the sequential appearance and accumulation of intermediate and final products. A typical potyvirid genome starts with a 50 non-coding region (NCR), with a variable length (less than 300 bp). This leader might act as a translational enhancer, and although the general mechanism is not fully understood, regulatory elements were found in this region. Studies in Plum pox virus (PPV), for instance, showed that (i) most of this region is dispensable for infectivity, although it contributes to viral competitiveness and pathogenesis, and (ii) translation takes place by a cap-independent leaky scanning mechanism. In the case of the 50 -leader of Tobacco etch virus (TEV), the existence of an internal initiation site has been suggested, with the presence of an RNA pseudoknot that provides cap-independent translation. Another NCR variable in size (generally up to 200 bp) is located at the 30 end of the genome, just upstream of the poly-A tail. Putative RNA structures in this region confer pathogenic properties in Tobacco vein mottling virus (TVMV), and studies with TEV and Clover yellow vein virus (ClYVV) demonstrated the existence of cis-acting elements necessary for infectivity.

Properties and Functions of Gene Products Derived From the Polyprotein The polyprotein encoded in the genome of most potyvirids comprises the following gene products (from N-terminus to C-terminus): P1, HCpro, P3 and P3N-PIPO (with PIPO in a different reading frame), 6K1, CI, 6K2, VPg, NIapro, NIb and CP. For bymoviruses, RNA-2 encodes two proteins P2–1 and P2–2, while RNA-1 encodes the remaining products starting with P3. Besides the information provided in Fig. 4, some characteristics of these products are given in this section. The P1 protein is a serine protease, extremely variable in size (from 30 to 60 kDa), considered as an accessory factor for genome amplification. This region contains important sequence differences among potyvirids, and the whole cistron is even absent in macluraviruses and some unclassified potyvirids. In some ipomoviruses, such as Cucumber vein yellowing virus (CVYV), two P1-like serine proteinases are found in tandem, with the second one acting as a suppressor of gene silencing, likely compensating the absence of the HCpro cistron in its genome. The helper component HCpro is a multifunctional protein named after its participation as a helper during aphid transmission. HCpro of tritimoviruses is also implicated in mite transmission. HCpro is smaller in macluraviruses, and sequences encoding a tandem of similar small HCpros are found in the genome of two potyvirids infecting areca palm, which has been proposed to become a novel genus tentatively named Arepavirus. A modular distribution of functions has been proposed for the HCpro of potyviruses: the N-terminus is essential for vector transmission, the central region is involved in suppression of RNA silencing, and the C-terminal part contains the papain-like cysteine protease activity. HCpro can interact with RNA, including double-stranded small interfering RNA molecules, a common feature of many other viral-derived RNA silencing suppressors. Structural studies with Lettuce mosaic virus (LMV) and TEV showed that HCpro form pairwise oligomeric structures. Relevant interactions of HCpro with several host factors have been described.

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Fig. 3 Phylogenetic analysis derived from the comparison of full-length viral genome sequences of 38 members of the family Potyviridae belonging to different genera (a), and 122 members of the genus Potyvirus (b). Figures depict neighbour joining trees derived from Mega v6.06 after alignment by Muscle with 1000 bootstrap replicates, with branch lengths being proportional to genetic distance (substitutions per site). The bootstrap support (40.7) for branches is indicated by proportional dots. For bymoviruses, the comparison was performed with a concatamer of RNA2 followed by RNA1. Viruses are identified by their official acronyms available at the ICTV webpage, and the sequences were obtained using the GenBank accession numbers of the corresponding type isolates, as found in the latest release (MSL34, from March 2019) of the Virus Metadata Repository (see Table 2). Grouping by genera and by number of genome segments are indicated as sectors in (a), where only the type member of potyviruses (PVY) is included, while the rest of selected members in the genus are shown in (b).

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Potyviruses (Potyviridae) Selection of resources available on the Internet (World Wide Web) about the family Potyviridae

Web Page Address (URL)

Contents and characteristics

https://talk.ictvonline.org/ictv-reports/ictv_9th_report/positivesense-rna-viruses-2011/w/posrna_viruses/271/potyviridae https://talk.ictvonline.org/taxonomy/vmr/

ICTV (International Committe on Taxonomy of Viruses). Taxonomy structure, list of species Virus Metadata Resource (VMR) spreadsheet, with the latest release containing data on new virus species ratified by ICTV Web-resource providing general molecular and epidemiological information, along with virion and genome figures, and access to UniProtKB/Swiss-Prot viral protein entries. VIDE (Virus Identification Data Exchange) project. Nomenclature, host range, virion properties Analysis of the polyprotein cleavage sites

https://viralzone.expasy.org/48?outline=all_by_species

http://bio-mirror.im.ac.cn/mirrors/pvo/vide/refs.htm http://www.dpvweb.net/potycleavage/index.html

In the case of bymoviruses, very limited information is available about the role of P2–1 and P2–2. It has been postulated that P2–2 might participate in vector transmission, since non-transmissible variants exhibited deletions in this region. Intriguingly, sequence alignments indicate that P2–1, a putative cysteine proteinase, is indeed more closely related to HCpro than P2–2. With the exception of P1 and HCpro, potyvirid gene products are excised from the viral polyprotein by the action of the NIapro proteinase. The third gene product is P3, a protein of unknown functions presumably implicated in RNA replication. An intrinsic difficulty to establish the actual role of P3 lays on the presence of the overlapping P3N-PIPO gene, which is directly involved in cell-to-cell movement. P3 is followed by the 6K1 peptide. Remarkably, separation of 6K1 from P3 in TEV does not occur in vitro and, moreover, this excision seems to be not required for the viability of PPV. Immunology studies determined the presence of both P3 and P3 þ 6K1 in infected cells, and the latter is perhaps the functional product. Proteins encoded in the P3–6K1 region have been shown to play roles in host range definition and pathogenicity of several potyvirids. The CI protein is the largest potyvirid gene product. It forms very distinctive pinwheel-shaped cylindrical inclusions in the cytoplasm of infected cells, with high taxonomic value because they are unique to members of the Potyviridae family (Fig. 1(b)). CI exhibits RNA helicase activity, and is supposed to act during RNA replication. Along with P3N-PIPO, CI forms conical structures at the cell plasmodesmata to allow viral cell-to-cell movement. 6K2 is a trans-membrane protein that induces the membrane remodeling needed for viral RNA replication and anchors the replication complex to virus-induced membrane structures. Studies with PVA 6K2 revealed that this protein might be also involved in movement and symptom induction. The whole NIa (nuclear inclusion a) protein has about 49 kDa, but it is subjected to an internal suboptimal cleavage that gives rise to VPg (21 kDa), the protein covalently attached to the 50 end of viral RNA, and a proteinase fragment (NIapro, 28 kDa). VPg can be uridylated in vitro by the RNA replicase NIb, suggesting that it is used as a primer during RNA replication, as in the case of picornaviruses. The VPg from diverse potyvirids, such as the one from Turnip mosaic virus (TuMV), can also counteract antiviral RNA silencing, a function involving direct interaction with, and promoting the degradation of, components of the host silencing machinery. In PVA, VPg is translocated as a 'phloem protein' that specifically acts in companion cells to facilitate virus unloading. ATPase activity was found in the VPg of Pepper vein banding virus (PVBV), being functionally autoinhibited by its N-terminal region, and stimulated by NIapro. The main role of NIapro is to act in cis and trans as the protease responsible for most cleavages in the polyprotein, processing sites mainly defined by specific heptapeptide sequences. NIapro shares structural motifs with host serine proteases, but it has a cysteine residue at the catalytic site. NIapro has non-specific RNA binding activity, and was shown to exhibit non-specific double-stranded DNA degradation activity in PVBV. The complete NIa can form inclusion bodies in the nucleus, where it is translocated responding to a bipartite nuclear localization signal. The crystal structure of TEV NIapro has been solved, providing clues on its mode of action. NIb (nuclear inclusion b) is the RdRp responsible for viral replication. The recruitment of this protein towards the replication complex is postulated to occur via interaction with NIa. All NIb functions essential for RNA amplification can be provided in trans, as knockout TEV mutants lacking NIb infect transgenic plants expressing this protein. NIb is the second component of nuclear inclusions, being directed to the nucleus by specific signals in its sequence. Recent results with TuMV have shown that sumoylation of NIb occurs in the nucleus, and this modification retargets it to the cytoplasm where viral replication takes place. Moreover, sumoylation of NIb contributes to the suppression of the host NPR1-mediated immune response. Phosphorylation of TuMV NIb has been described, although it is still not known the significance of this post-translational modification. The last product of the viral ORF is CP, a protein with many roles apart from coating and protecting the RNA genome. It is implicated in aphid transmission through interaction with HCpro, and in both cell-to-cell and long-distance movement. The importance of maintaining the net charge of the CP N-terminus for infectivity has been shown in TVMV and Zucchini yellow mosaic virus (ZYMV).

Expression of Overlapping Gene Products The presence of PIPO within the P3 cistron was predicted by bioinformatics analyses, including alignments of potyvirid genomes and synonymous sequence variability. This ORF is preceded by a conserved GA6 nucleotide sequence, a motif not present in other viral regions except in the case of a subgroup of sweet potato-infecting potyviruses, which harbor an additional GA6 within the P1 region that precedes a larger ORF named PISPO. Interestingly, the predicted P1N-PISPO gene product was found to be a suppressor of gene

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silencing. Experimental evidence deriving from high-throughput sequencing of different potyvirid progenies support the idea that the changes in the reading frame required to produce these overlapping products occur during positive or negative strand (it is not yet defined) RNA synthesis by polymerase slippage. Interestingly, in another set of RNA species, an apparently equivalent mechanism causes the deletion of one nucleotide at the same GA6 motif in Clover yellow vein virus (ClYVV), with the consequent change in the reading frame. This deletion introduces a premature stop codon, and therefore a truncated P3, termed P3N-ALT, is produced. As P3N-PIPO, ClYVV P3N-ALT seems to participate in cell-to-cell movement.

Replication and Propagation Potyvirids replicate in the cytoplasm of infected cells, as schematically shown in Fig. 5. After entering the cell, the viral genomic RNA must be first translated in the cytoplasm. In general, the early events during infection are still poorly understood. Disassembly of particles might occur co-translationally, and after translation the replication complex must be formed with participation of several viral products and probably host factors. This complex uses the genomic RNA ( þ sense) as template to generate a complementary chain (- sense) through a dsRNA intermediate, and proceed with the asymmetric synthesis of numerous molecules of genomic RNA. Viral-derived siRNAs have been shown to accumulate during potyvirid infections, revealing the action of host RNA silencing-mediated antiviral defenses. Therefore, potyvirids need to overcome this barrier, and for that they express RNA silencing suppressors. In the case of potyviruses, this role is mainly exerted by HCpro, although this function might be shared or displaced to other products in different genera. Studies performed with Pea seed-borne mosaic virus (PSbMV) served to identify a translation shut-off affecting many host proteins during infection. In the case of PVA, the shift for translation to replication appears to be controlled by phosphorylation-dependent RNA binding of non-assembled viral CP. The potyvirid expression strategy implies production of equimolar amounts of all gene products, giving rise to large excesses of some proteins, which might remain soluble, be degraded or secreted, or end up in inclusion bodies. It is intriguing that, whereas CI pinwheels are always formed, HCpro amorphous or NIa/NIb crystalline inclusions are present in some, but not all, viral infections. Potyvirid RNA replication takes place in membranous vesicles derived from the endoplasmic reticulum that target chloroplasts and aggregate into large perinuclear globular structures. NIb forms the core of the replication machinery, with involvement of CI, VPg, 6K2, and NIapro. Other factors, viral- and host-derived, might also be involved. For instance, the VPg of TuMV can interact with initiation factors eIF4E and eIF(iso)4E, and also with the Poly-A binding protein. The fact that several resistance genes, identified in host plants as potyvirid-specific, are related to translation initiation factors points to the importance of these interactions for the infection cycle. Another critical process for a successful virus infection is the genomic RNA transport from the initially infected cell towards adjacent cells and distant tissues. In principle, the virus can invade the whole plant, with the probable exception of meristems, although a few seed-transmissible potyvirids might be capable of invading these particular tissues. The CP, but probably not virions, is essential for virus movement. RNA replication and virus movement appear to be closely linked. It has been proposed that replication vesicles are able to move through the cytoplasm along microfilaments to reach plasmodesmata and cross them in order invade the neighboring uninfected cell, and that these vesicles are even able to traffic through the vascular system. In the process of local and systemic spread, host defenses must be confronted by the virus in a race to avoid its blockage by the action of both general and specific long-distance defensive signals. The final step of the infective cycle is the virus spread to new plants, which rely on the acquisition of virions by the corresponding vector organisms and further transmission. Two viral proteins, CP and HCpro, are involved in the non-persistent aphid transmission of potyviruses. Available data supports a “bridge” hypothesis, in which HCpro serves as a reversible linker to retain virus particles in aphid mouthparts. A conserved PTK motif in the central portion of HCpro could participate in binding to the DAG motif at the N-terminus of CP, while a KITC domain at the N-terminal region of HCpro would be involved in binding to unknown structures on the aphid stylet. HCpro is also implicated in the semipersistent transmission of tritimoviruses by eryophid mites. Less information is available about the transmission processes of macluraviruses by aphids, bymoviruses by plasmodiophorids, rymoviruses and poaceviruses by eryophid mites, and ipomoviruses by whiteflies. Multifunctionality is observed in most potyviral proteins, illustrated for instance by HCpro. This capacity to participate in multiple processes suggests that potyvirus infection is not a consecutive succession of independent events, but a tightly regulated network of complex interactions, still to be elucidated, between viral and host factors.

Pathogenicity The information about how potyvirids cause diseases in their host plants is very limited. Several studies have served to identify sequences and/or products in the genome of potyvirids directly implicated in the production of symptoms. As mentioned above, symptom determinants are even present in the 50 - and 30 -NCR. In addition, HCpro, P3, CI, 6K2 and VPg of different viruses have been described as determinants of pathogenicity, although single products were not always responsible for the different pathogenic responses. As already mentioned, the P1/HCpro region was found to be responsible for the synergistic effect in mixed infections of potyviruses with unrelated viruses, perhaps reflecting the capacity of HCpro to counteract the RNA silencing-mediated defense. Interestingly, HCpro also interferes with the action of microRNAs, a particular type of regulatory host-derived small RNAs,

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Fig. 4 Genome maps of representative viruses in the family Potyviridae. A monopartite virus characteristic of a potyvirus such as the type member of the genus PVY is shown in (a), together with SPFMV as representative of the sweet-potato infecting subgroup. Maps of representatives of other genera are shown for the macluravirus CdMV (b), three different ipomoviruses SPMMV, CVYV and CBSV (c), and the two genomic RNAs of the bipartite bymovirus BYMV (d). ssRNA genomes are shown as solid horizontal lines with VPg represented by solid circles at 50 end, and poly-A tails at 30 end. Viral ORFs are depicted as boxes divided in the different gene products, with their names indicated. The partially out-of-frame products deriving from polymerase slippage mechanisms are shown below. The protease-specific cleavages sites are indicated by arrows above the first map, with the matching symbol shown in the gene products responsible of the proteolytic process. Additional genome structures of other viruses (not shown) include the potyvirus EuRSV, containing a HAM1-like gene between NIb and CP coding sequences, and the still unclassified viruses of the proposed new genus Arepavirus, with presence of two HCPro-related cistrons, or the proposed new genus Celavirus, with a signal peptide preceding P1 and a very long 30 NCR.

providing a molecular explanation for some virus-induced symptoms. However, although it is tempting to regard interference with small RNAs metabolism in susceptible plants as a major element of pathogenicity, many other processes may also be affected. For example, specific features of the soybean Rsv1, a Soybean mosaic virus (SMV) resistance gene, appear to be responsible for the induction of either systemic mosaic or lethal systemic hypersensitive response in infected plants. The fact that the HCpro protein of LMV targets, and affects the functions of, the 26S proteasome might exemplify another way by which these viruses can cause disease symptoms. Despite the fact that in most cases the mechanisms of pathogenicity remain uncertain, it is reasonable to conclude that the final macroscopic effects of potyvirids are caused by a combination of additive interferences with several host functions and processes.

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Fig. 5 Schematic representation of events during potyvirid infection of a plant cell. Colors of gene products match those displayed in Fig. 4. For simplicity, most organelles have been omitted, as well as the participation of host factors during the cycle. The accumulation of viral proteins as inclusions in different compartments is indicated. The cycle begins (left up) when the virus enters the cell, either from an adjacent infected cell or by inoculation from a vector organism. Then, the genomic RNA undergoes des-encapsidation, translation and processing to originate mature products. The replication complex involves participation of at least NIb, CI, VPg, 6K2, and NIa, together with host-derived factors, and uses the genomic RNA ( þ sense) to generate a complementary chain (- sense), serving as template for the synthesis of numerous genomic RNAs. Also, during polymerization, variants with slippage at GA6 motifs could be generated and translated to yield partially out-of-frame products. dsRNA replicative intermediates or regions with secondary structure produce viral siRNAs by action of the host silencing machinery, which is counterattacked by suppressors of RNA silencing, such as HCpro or other viral products. After replication, the RNA progeny can move to adjacent cells through plasmodesmata, with involvement of P3NPIPO, CI, VPg and CP, or it can be encapsidated and acquired to be transmitted again by a vector organism like aphids, in a process requiring HC-Pro. See text for further details.

Epidemiology and Control Strategies currently applicable for the control of potyvirids are diverse, from cultural practices to the use of genetic resistance. Insecticide or other treatments against vectors are frequently considered unsatisfactory and have limited use because of the transmission type. Severity of outbreaks is commonly related to abundance of initial foci of infection, dynamics of vector populations and other factors, such as the presence of weeds acting as reservoirs of viruses. The human intervention is responsible in many cases for the introduction of emerging diseases into new territories, while vector organisms are mainly involved in propagation within particular regions. For a few potyviruses, seed transmission is also an important means of dissemination. A typical example of well documented spread of a potyvirus over a territory and over time is the progressive emergence of Sharka disease, caused by PPV, in European countries during the XX century, and its recent diffusion to other continents to become nowadays a global pandemic.

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Potyviruses (Potyviridae)

The exploitation of pathogen-derived resistance and RNA silencing is yielding promising results for virus control. Transgenic plants incorporating viral sequences were found to be resistant to several potyvirids. An example of success is provided by the engineered papaya varieties, which are resistant to Papaya ringspot virus due to the expression of the cognate viral CP as a transgene. Nowadays, RNA silencing is being further exploited for resistance by designing specific transgenes with hairpin structures as sources of viral-derived siRNAs, or even by direct application of dsRNAs. Other approaches to achieve resistance include the expression of ribozymes, plantibodies or proteinase inhibitors. Cross-protection has also been related to RNA silencing mechanisms, although other processes could be operating, as suggested by the involvement of specific proteins, such as NIapro and CP, in superinfection exclusion. Stability, durability and biosafety issues of the different strategies are under constant evaluation. The identification of potyvirid-specific resistance genes in host plant is also leading to important advances. These studies indicate that canonical resistance genes might be operating, for instance TIR-NBS-LRR class in PVY, or NBS-LRR class in SMV. Other examples of dominant resistance genes acting against different potyviruses are the three Restricted TEV Movement (RTM) genes, which code for a jacalin-type protein, a small heat shock protein and a MATH domain-containing protein. As mentioned, mutations in eukaryotic initiation factors are frequently found as responsible for recessive resistance against potyvirids, but particular features of other host factors, as the products of PROTEIN DISULFIDE ISOMERASE LIKE 5–1 of barley or two MATH domain-containing genes of apricot, has been also found to be associated with natural recessive resistance against potyvirids. Therefore, the generation of variability in candidate genes, using TILLING or equivalent platforms, must be a promising strategy for the generation of resistant plants.

Diagnosis Serological tools have been the preferred diagnostic system for potyvirids. While the non-structural viral proteins have limited use in serological diagnosis, the virus particles are in general strongly immunogenic. However, serological relationships among potyvirids are complex, with unexpected and inconsistent cross-reactivities hindering their application in taxonomy and specific virus identification. The cause of this phenomenon is the presence of common epitopes in the conserved internal CP core, while specific epitopes map in the variable N-terminus, a surface-exposed region prone to degradation. New species-specific antibodies targeting the immunodominant N-terminal region of CP provide excellent tools for the serological detection of many potyvirids. RT-PCR is another way to deal with diagnosis. Degenerated primers matching conserved genomic regions might allow the amplification of virtually any potyvirid. For instance, primers flanking the variable CP N-terminal region have been developed and successfully used for both detection and identification of highly divergent potyvirids. Interestingly, the combination of serological capture of virions followed by highly sensitive RT-PCR has resulted in robust detection systems. Together, these molecular tools have been extensively used to identify new viruses infecting new hosts. The application of new technologies and modern molecular methodologies to diagnostics is being continually updated, which results in more sensitive and more specific approaches.

Concluding Remarks The abundant information on potyvirids have reached a level that allows to develop biotechnological applications, including the production of foreign proteins in plants using virus-based expression vectors, and also the exploration of viruses as direct sources of genetic elements and functionally active products, such as the NIapro proteases used to remove affinity tags from fusion proteins. Another example is the use of HCpro to block the host RNA silencing machinery with the consequent boost in the expression of products in plants. Several potyvirids have been engineered and tested as expression vectors. Small peptides can be expressed as CP-fusion products, allowing the protein chimeras to be used as antigen presentation systems for immunization or diagnosis. Complete foreign genes have been expressed in vectors based on a quite large list of potyvirids, which covers an important range of potential hosts. The seminal work with TEV pointed to the P1/HCpro junction as an adequate insertion site for foreign sequences, but additional sites were exploited later, such as the NIb/CP junction or the P1 region. Moreover, more recent reports demonstrated the huge versatility of potyvirids for the expression of foreign genes, as they can even express different products simultaneously from independent insertion sites. To summarize, it can be concluded that the family Potyviridae is a fascinating group of viruses. Keeping key genomic features, potyvirids have diverged in a multitude of lineages, which we have yet to finish to discover, and one of them, the genus Potyvirus, can be considered one of the most successful in the world of viruses. The currently available knowledge about potyvirid replication, movement and transmission is allowing the development of new control strategies aiming to interfere with key-steps in the virus life cycle. Structure resolution of viral components might also serve to explore new resistance strategies. Diagnostic tools are continually under improvement, and future developments will certainly supply specific and sensitive means of virus identification. Our current insight about the selective participation of different host factors in resistance and pathogenesis is fueling works aiming to understand the complex virus-plant crosstalk that occurs during the infection. Besides, peculiarities of certain viruses are being unraveled, and these findings will help to discover new generic features and specificities of members of the Potyviridae family. All in all, the study of potyvirid infections at the molecular level has been extraordinarily successful over the last years, and is likely to continue providing relevant data on the viruses, as well as in terms of understanding many plant processes.

Potyviruses (Potyviridae)

641

Further Reading Adams, M.J., Antoniw, J.F., Fauquet, C.M., 2005. Molecular criteria for genus and species discrimination within the family Potyviridae. Archives of Virology 150, 459–479. Cheng, X., Xiong, R., Li, Y., et al., 2017. Sumoylation of Turnip mosaic virus RNA polymerase promotes viral infection by counteracting the host NPR1-mediated immune response. Plant Cell 29, 508–525. Elena, S.F., Rodrigo, G., 2012. Towards an integrated molecular model of plant-virus interactions. Current Opinion in Virology 2, 719–724. Ivanov, K.I., Eskelin, K., Lohmus, A., Mäkinen, K., 2014. Molecular and cellular mechanisms underlying potyvirus infection. Journal of General Virology 95, 1415–1429. Jiang, J., Laliberté, J.F., 2011. The genome-linked protein VPg of plant viruses-a protein with many partners. Current Opinion in Virology 1, 347–354. Lauber, C., Seifert, M., Bartenschlager, R., Seitz, S., 2019. Discovery of highly divergent lineages of plant-associated astro-like viruses sheds light on the emergence of potyviruses. Virus Research 260, 38–48. Movahed, N., Patarroyo, C., Sun, J., et al., 2017. Cylindrical Inclusion protein of Turnip mosaic virus serves as a docking point for the intercellular movement of viral replication vesicles. Plant Physiology 175, 1732–1744. Olspert, A., Chung, B.Y.-W., Atkins, J.F., Carr, J.P., Firth, A.E., 2015. Transcriptional slippage in the positive-sense RNA virus family Potyviridae. EMBO Reports 16, 995–1004. Revers, F., García, J.A., 2015. Molecular biology of potyviruses. Advances in Virus Research 92, 101–199. Rodamilans, B., Shan, H., Pasin, F., García, J.A., 2018. Plant viral proteases: Beyond the role of peptide cutters. Frontiers in Plant Science 9, 666. Rodamilans, B., Valli, A., Mingot, A., et al., 2015. RNA polymerase slippage as a mechanism for the production of frameshift gene products in plant viruses of the Potyviridae family. Journal of Virology 89, 6965–6967. Sorel, M., Garcia, J.A., German-Retana, S., 2014. The Potyviridae cylindrical inclusion helicase: A key multipartner and multifunctional protein. Molecular Plant-Microbe Interactions 27, 215–226. Truniger, V., Aranda, M.A., 2009. Recessive resistance to plant viruses. Advances in Virus Research 75, 119–159. Valli, A.A., Gallo, A., Rodamilans, B., López-Moya, J.J., García, J.A., 2018. The HCPro from the Potyviridae family: An enviable multitasking Helper Component that every virus would like to have. Molecular Plant Pathology 19, 744–763. Wolf, Y.I., Kazlauskas, D., Iranzo, J., et al., 2018. Origins and evolution of the global RNA virome. mBio 9, e02329. 18. Wylie, S.J., Adams, M., Chalam, C., et al., 2017. ICTV Virus taxonomy profile: Potyviridae. Journal of General Virology 98, 352–354. Zamora, M., Méndez-López, E., Agirrezabala, X., et al., 2017. Potyvirus virion structure shows conserved protein fold and RNA binding site in ssRNA viruses. Science Advances 3. (eaao2182).

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_9th_report/positive-sense-rna-viruses-2011/w/posrna_viruses/271/potyviridae ICTV 9th Report (2011). http://www.dpvweb.net/potycleavage/index.html Potyviridae cleavage sites Descriptions of Plant Viruses. http://bio-mirror.im.ac.cn/mirrors/pvo/vide/refs.htm VIDE (Virus Identification Data Exchange) project. Nomenclature, host range, virion properties. https://viralzone.expasy.org/48?outline=all_by_species ViralZone. https://talk.ictvonline.org/taxonomy/vmr/ VMR - International Committee on Taxonomy of Viruses (ICTV).

Quinviruses (Betaflexiviridae) Ki H Ryu and Eun G Song, Seoul Women’s University, Seoul, South Korea r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

ORF Open Reading Frame PCR Polymerase chain reaction Pol RNA-dependent RNA polymerase Poly(A) Polyadenylated P-Pro Papain-like protease RdRp RNA-dependent RNA polymerase RT-PCR Reverse transcription polymerase chain reaction S20,w Corrected sedimentation coefficient; extrapolated in water at 201C and infinitely diluted SSR Simple sequence repeats ssRNA Single-stranded RNA TGB Triple gene block ZF Zinc finger motif

aa Amino acid(s) AlkB Alpha-ketoglutarate-dependent dioxygenase CP Coat protein or capsid protein CRP Cysteine-rich protein cSSR Compound simple sequence repeat Hel Helicase kb Kilobase kDa Kilo dalton LAMP Loop mediated amplification MP Movement protein NLS Nuclear localization signal nt Nucleotide(s) NTR Non-translated region

Glossary AlkB protein The protein found in bacteria, fungi, and mammalian genomes is iron (II)- and 2-oxoglutaratedependent dioxygenases that remove alkyl adducts from oxidative dealkylation DNA repair reverse methylation damage, such as 1-methyl-adenine and 3-methyl-cytosine. Viral AlkB proteins can repair alkylated bases in DNA or RNA. Cysteine-rich protein The protein related to viral pathogenicity determinants. The CRPs can be divided into two groups according to their domain structures. The CRPs with a conserved CGxxH motif (C, cysteine; G, glycine;

H, histidine; x, any aa residue). The CRPs with an argininerich nuclear localization signal (NLS) in the central part of the protein. Carlavirus CRPs contain a zinc finger motif (ZF) located adjacent to the NLS. RNA silencing suppressor Plant viruses are encoding proteins to counteract the plant antiviral system known as RNA silencing. Triple gene block The protein consists of a strikingly similar element of three partially overlapping ORFs and is included in their genomes. The TGB proteins are related to cell-to-cell movement and potentially influence on host antiviral defenses.

Introduction The subfamily Quinvirinae, family Betaflexiviridae, contains three genera (Carlavirus, Foveavirus, and Robigovirus) and three unassigned species. The complete genome sequences of most of the members of the subfamily Quinvirinae have been determined recently. The members of the genus Carlavirus contain 58 species including Carnation latent virus (CLV) as the type species for the genus Carlavirus. The members of the genus Foveavirus contain 8 species including Apple stem pitting virus (APSV) as the type species for the genus Foveavirus. The members of the genus Robigovirus contain 5 species including Cherry necrotic rusty mottle virus (CNRMV) as the type species for the genus Robigovirus. The genus Robigovirus has been recently established as a new member of the subfamily Quinvirinae by the International Committee on Taxonomy of Viruses (ICTV). The members of the subfamily Quinvirinae can be divided into two or three main phylogenetic groups based on their genome sequences. The two genera, Foveavirus and Robigovirus, and unassigned species can be clustered in one branch. The phylogenetic relationships of the subfamily Quinvirinae are not associated with their natural host. The virions of the subfamily Quinvirinae have non-enveloped and flexible filaments of 640–1000 nm in length and 12–15 nm in diameter. Their genomes are composed of a positive-sense single-stranded RNA (ssRNA) with 50 terminal cap structure and a 30 terminal poly(A) tail, and contain RNA-dependent RNA polymerase (RdRp) for viral replication, triple gene block (TGBp1, 2, and 3) for viral movement (MP), and coat protein (CP). The RdRp contains five conserved protein domains as follows: an AlkB domain; a papain-like protease (p-Pro) domain; an RNA helicase (HEL) domain; a polymerase (POL) domain. The sizes of the genomes have 7.4–9.3 kbp. The genomes of the carlaviruses contain mostly a cysteine-rich protein (CRP) including a nuclear localization signal (NLS) and a zinc finger motif (ZF). Most viruses have a relatively restricted host range, which may exhibit mosaic, mottle, ringspot, necrosis, or pitting symptoms or may be symptomless. Most viruses are spread by mechanical transmission and some viruses can be transmitted by insect vectors, such as aphid and whitefly. Some viruses are often transmitted by seed and grafting. Virus diseases caused by some viruses can seriously damage major crops and the viruses require suitable disease managements.

642

Encyclopedia of Virology, 4th Edition, Volume 3

doi:10.1016/B978-0-12-809633-8.21531-8

Quinviruses (Betaflexiviridae)

643

Taxonomy and Phylogeny Three genera, Carlavius, Foveavirus, and Robigovirus, is members of the subfamily Quinvirinae within family Betaflexiviridae. The genus Carlavirus contains 53 definitive species, of which 41 are determined completed genome sequences (Table 1). Its name is derived from Carnation latent virus (CLV, the type species of Carlavirus). The CLV genome has not been completely sequenced. The genus Foveavirus contains 8 definitive species, these viruses have complete genome sequences (see Table 1). Its name is derived from Latin fovea for “pit” or “hole”, which is a type of symptom induced by the type species (Apple stem pitting virus, ASPV). The genus Robigovirus, a newly created genus within the subfamily Quinvirinae, contains 5 definitive species, these viruses have complete genome sequences (Table 1). Its name is derived from Latin robigo for “rust”, which is a type of symptoms induced by the type species (Cherry necrotic rusty mottle virus, CNRMV). Additionally, the subfamily Quinvirinae contains three unassigned species Banana mild mosaic virus (BanMMV), Banana virus X (BanVX), and Sugarcane striate mosaic-associated virus (SCSMaV). Phylogenetic tree of members of the subfamily Quinvirinae has revealed a clustering into two major groups of their RNA-dependent RNA polymerase (RdRp) aa sequences (Fig. 1(A)). The major group I consists of members of the genus Carlavirus whereas the major group II can be divided into three distinct subgroups (carlavirus, foveavirus, and robigovirus groups). The carlavirus group within the major group II contains Sweet potato chlorotic fleck virus (SPCFV), Melon yellowing-associated virus (MYaV), and Nerine latent virus (NeLV). The foveavirus group has divided into three branches and ASPV, the type species of Foveavirus, is closely grouped with Apricot latent virus (ApLV). The two unassigned species, BanMMV and SCSMaV, are supported as members of the robigovirus group within the major group II. The robigovirus group is divided into two branches and CNRMV, the type species of Robigovirus, is more closely related to Cherry twisted leaf associated virus (CTLAV). In the phylogenetic tree of the coat protein (CP), Foveavirus, Robigovirus, and unassigned species are clustered in one branch (Fig. 1(B)). The foveavirus group could be divided into three branches and the ASPV is more closely grouped with Apple green crinkle associated virus (AGCaV) than the ApLV. The robigovirus group is divided into two branches and the CNRMV is more closely grouped to Cherry rusty mottle associated virus (CRMAV) than the CTLAV. Of three sambucus–infecting viruses in the phylogenetic trees of the RdRp and CP, Sambucus virus C and Sambucus virus E that are grouped together are more closely related than Sambucus virus D. Four cherry-infecting robigoviruses in the phylogenetic trees, Cherry green ring mottle virus (CGRMV), CNRMV, CTLAV, and CRMAV, are clustered in one branch. The aa sequence identities of their RdRp and CP proteins of the subfamily Quinvirinae are less than 67.6% and 85.5%. The phylogenetic trees of the TGBp1, TGBp2, and TGBp3 of the subfamily Quinvirinae could be divided into three major groups (Fig. 2). The two genera, Foveavirus and Robigovirus, and unassigned species in the phylogenetic tree of The TGBp1 are clustered in one branch, similar to those of the RdRp and CP. The two genera in the TGBp2 and TGBp3 are clustered in distinct branches. In the tree of the TGBp3, the members of the genus Foveavirus are clustered in distinct branches. Two foveaviruses, Grapevine rupestris stem pitting-associated virus (GRSPaV) and Grapevine virus T (GVT), clustered together are more closely related to three carlaviruses, Potato latent virus (PotLV), Hippeastrum latent virus (HLV), and Cowpea mild mottle virus (CPMMV), than to other foveaviruses. The cheery-infecting viruses except for CGRMV, which is missing the TGB aa sequences, are clustered in one branch. The aa sequence identities of their TGBp1, TGBp2, and TGBp3 proteins of the subfamily Quinvirinae are less than 72.2%, 79.0%, and 63.9%, respectively. Phylogenetic tree based on a cysteine-rich protein (CRP) aa sequences of members of the genus Carlavirus is revealed that divided into five groups. Of three sambucus-infecting viruses, SVC and SVE are grouped together, whereas SVD is grouped with Pea streak virus (PSV) in a distinct branch. The aa sequence identities of their CRP proteins is less than 77.1%.

Virion Structure Virions are non-enveloped and flexible filaments of 640–1000 nm in length and 12–15 nm in diameter. Carlavirus and foveavirus particles are 610–700 nm and 800–1000 nm in length, respectively. Carlaviruses have helical symmetry with a pitch of 3.4–3.6 nm. Carlaviruses virion Mr is about 60  106 with 5.4%–6% nucleic acid content. Carlaviruses have the sedimentation coefficient (S20, w) of 147–176S, the buoyant density in CsCl of 1.3 g cm3. Potato virus M (PVM), Shallot latent virus (SLV), and Daphne virus S (DVS) have the sedimentation coefficient (S20, w) of 157S, 147.5S, and 174 S, respectively. A carlavirus isolated from elderberry (Sambucus spp.) has the sedimentation coefficient (S20, w) of 155S and molecular weight of CP subunits of 31 kDa. The virus is an average normal length of 678 nm in length and 12 nm in diameter. The virus particles can be observed in the cytoplasm of their infected plant. The virus particles of Helloborus net necrosis virus (HeNNV) is approximately 800 nm in length and 17 nm in diameter. The virions of ASPV, Grapevine rupestris stem pitting-associated virus (GRSPaV), and CGRMV apparently do not contain lipids or carbohydrates.

Genome Organization The genome is a positive-sense single-stranded RNA (ssRNA) with 50 terminal cap structure and a 30 terminal poly(A) tail. The genome is 7.4–9.3 kb in size and contains five or more ORFs. ORF1 encodes a large polypeptide that is RdRp including domains of methyl-transferase, helicase, and polymerase. ORFs 2, 3, and 4 constitute the TGB that facilitate virus movement. ORF5 encodes the CP and ORF6 encodes a cysteine-rich protein (CRP). The genome structures of the distinguished genera in the subfamily Quinvirinae is showed in Fig. 3. The genome of PVM, a member of the genus Carlavirus, consists of 8533 nt and contains six ORFs

CapLV

CLV

CVB CoLV CVNV

Caper latent virus

Carnation latent virus

Chrysanthemum virus B Cole latent virus Coleus vein necrosis virus Cowpea mild mottle virus

Humulus lupulus

675  14

HpLV

HpMV

HdCMV

Hop mosaic virus

Hydrangea chlorotic mottle virus

650

Hydrangea macrophylla

Humulus lupulus

Mosaic/Chlorotic spots

Hippeastrum hybridum

706

HLV

675  14

Black streaks/Black spots (sepals)

800  17

MT (G. globosa, H. niger)

MT/Aphid (M. persicae)

Aphid/MT (C. murale)

Aphid (M. persicae)

MT/Whitefly (B. tabaci)

NC038966

NC017859

NC002795

Access #

Worldwide Brazil USA

Europe

Italy

USA Germany USA/Japan/New Zealand Netherlands/Taiwan

New Zealand/South Korea Germany Worldwide

Tanzania

NC011540

NC038323 NC043082 NC012038

NC023892 NC016440

NC008020

NC038322

NC014730

NC009087 NC038322 NC009764

NC038865

NC043080

North America/ Europe NC003499 Japan NC013527 Ukraine ND

Whitefly (Bemisia tabaci) MT Worldwide Exp Hosts

MT Exp Hosts/Aphid (Myzus persicae) Aphid (M. persicae) MT/Aphid (M. persicae) MT Exp Hosts

MT Exp Hosts

MT MT (C. quinoa)/Aphid

South Korea

North America/Europe

Japan/Israel

Distribution

Aphid (Phorodon humuli)/ Europe/USA/Australia/ NC002552 MT China Europe/Australia/ North NC010538 Chlorotic vein-banding/Leaf distortion/Stunting Aphid (M. persicae, Marcrosiphum euphorbiae, America/China/Japan P. humuli) Leaf blistering/Reddening/Chlorosis/Chlorotic MT Exp Hosts USA/New Zealand/ NC012869 mottle/Leaf distortion/Ringspot South Korea Symptomless

Spots/Color breaking/Flower distortion

Impatiens holstii Helleborus niger Helleborus spp.

Chlorotic mosaic/Distortion

Daphne spp.

HVS HeMV HeNNV

Severe mosaic/Mottle/Deformation

Chlorotic spots/Yellow stripes/Symptomless

Mottle/Leaf malformation/Vein chlorosis/ Chlorotic ringspot and blotch/Leaf rolling

Arachis hypogaea/ Glycine max/ Phaseolus vulgaris/ Solanum lycopersium Citrullus lanatus

Gaillardia aristata Allium sativum

Mosaic/Vein banding/Mottle/Vein chlorosis Symptomless Mottle/Necrosis

GaILV GarCLV

676–683

685  12 650 650

Chlorotic spots/Mosaic/Mottle

Symptomless

Blossom blight/Necrosis/Chlorosis Necrotic ringspot

Chrysanthemum spp. Brassica oleracea Verena x hybrida

Dianthus caryophyllus

650  14

662

Vaccinium corymbosum Petasites japonicus Echinopsis sp./ Mammillaria elongata Capparis spinosa

690  14 670  13 650  12

Gaillardia latent virus Garlic common latent virus Helenium virus S Helleborus mosaic virus Helleborus net necrosis virus Hippeastrum latent virus Hop latent virus

Cucumber vein-clearing CuVCV virus Daphne virus S DVS

CPMV

BlScV ButMV CV  2

Silver mottle

Atractylodes macrocephala

American hop latent virus Atractylodes mottle virus Blueberry scorch virus Butterbur mosaic virus Cactus virus 2

AtrMoV

Transmission

Humulus lupulus

Symptoms

AHLV

640

Natural Hosts Mosaic/Spots on petals/Symptomless/ Mottle MT Exp Hosts/Aphid-(A. napellus) Symptomless Aphid (Phorodon humuli)

AcLV

Carlavirus Aconitum latent virus

Particle size (nm) Aconitum napellus/ Delphinium sp.

FYI

Genus/Virus name

Table 1 Virus species of three genera (Carlavirus, Foveavirus, and Robigovirus) of the subfamily Quinvirinae, family Betaflexiviridae. Type species in the genera are written in bold. Complete sequences of the genomes are indicated

644 Quinviruses (Betaflexiviridae)

Foveavirus Apple stem pitting virus

Sweet potato C6 virus Sweet potato chlorotic fleck virus Verbena latent virus Yam latent virus

PVH PVM PVP PVS RCVMV

Potato virus H Potato virus M Potato virus P Potato virus S Red clover vein mosaic virus Sambucus virus C Sambucus virus D Sambucus virus E Shallot latent virus Sint-Jan onion latent virus Strawberry pseudo mild yellow edge virus

650–652

610–700 645

570

685 670

800  15

650

VeLV YLV

ASPV

750–800 800

SPC6V SPCFV

SPMYEV 625  12

SVC SVD SVE SLV SJOLV

PeSV PhlVB PhlVM PhlVS PopMV PotLV

Pea streak virus Phlox virus B Phlox virus M Phlox virus S Poplar mosaic virus Potato latent virus

Symptomless/Mild mottle

Verbena spp./ Tropaeolum majus Dioscorea opposita

Malus spp./ Pyrus spp./ Prunus spp. Yellow banding/Red mottling/ flecking

Chlorotic spots/Vein clearing Chlorotic flecks/Symptomless

Chlorotic mottle/Reddening/Necrosis

Ipomoea batatas Ipomoea batatas

Fragaria spp.

Solanum tuberosum Solanum tuberosum Solanum tuberosum Solanum tuberosum Pisum sativum/ Cicer arietinum/ Lens culinaris Sambucus spp. Sambucus spp. Sambucus spp. Allium spp. Allium spp.

Severe necrosis Virus-like symptoms Virus-like symptoms Virus-like symptoms Severe mosaic/Mild mosaic Symptomless/Slight vein clearing/Chlorotic spots Mild symptoms Mottle/Mosaic/Crinkling/Rolling/Stunting Symptomless Symptomless Chlorosis/Stunting/Rosetting/Vein clearing/ Plant death Mosaic/Leaf yellowing/Malformation/Dwarfing Mosaic/Leaf yellowing/Malformation/Dwarfing Mosaic/Leaf yellowing/Malformation/Dwarfing Symptomless Symptomless

Chlorotic streaking Mosaic

Hippeastrum spp. Passiflora spp.

NeLVPLV Pisum sativum/ Trifolium spp. Phlox spp. Phlox spp. Phlox spp. Populus trichocarpa Solanum tuberosum

Yellow stripes/Mosaic

Narcissus tazetta

NCLV

619

Slight leaf mottle/Leaf wrinkling

Mirabilis jalapa

MiMV

Stunting/Leaf malformation Symptomless/Very mild mosaic

Symptomless Necrotic ringspot/Line patterns

Yellowing/Mottle

Kalanchoë blossfeldiana Ligustrum japonicum/ Mazus reptans Ligustrum obtusifolium Liliaceae Cucumis melo

660

688 674

MYaV

LVA LSV

KLV LNRSV

Melon yellowingassociated virus Mirabilis jalapa mottle virus Narcissus common latent virus Nerine latent virus Passiflora latent virus

Kalanchoe latent virus Ligustrum necrotic ringspot virus Ligustrum virus A Lily symptomless virus

(M. persicae) (M. persicae) (M. persicae) (M. persicae) (Acyrthosiphon

MT/GT

MT

Aphid (Chaetosiphon frageaefolii, Aphis gossypii) MT/GT MT

MT/Aphid (M. ascalonicus) MT/Aphid

MT/Aphid MT/Aphid MT/Aphid MT/Aphid MT/Aphid pisum)

MT MT (N. megalosiphon) MT/Aphid (M. persicae)

MT MT

MT

MT

Whitefly/GT

Aphid

MT (C. quinoa) MT

Europe/Asia

Israel/New Zealand China

Dominican Republic Worldwide

Japan

Australia Germany/Australia/ USA/Israel North America/Europe USA USA USA Germany Canada/USA/ Netherlands Netherlands/China Worldwide Brazil/Russia/Argentina Worldwide USA/New Zealand/ Netherlands USA USA USA Worldwide Netherlands

China

USA

South Korea Netherlands/South Korea/China/USA/ India/Brazil Brazil

Denmark USA

(Continued )

NC003462

NC043085 NC026248

NC018448 NC006550

ND

NC029087 NC029088 NC029089 MG657357 NC043084

NC018175 NC001361 NC009759 NC007289 NC012210

NC009383 NC005343 NC011525

NC027527 NC009991

NC028111 NC008292

NC008266

NC016080

NC038324

NC031089 NC005138

NC013006 NC010305

Quinviruses (Betaflexiviridae) 645

BVX SCSMaV 950  15

BaMMV

Footnotes: MT; mechanical transmission, GT, graft transmission.

Unassigned species Banana mild mosaic virus Banana virus X Sugarcane striate mosaic-associated virus Mild chlorotic streaks

Short chlorotic streaks

Musa spp. Saccharum spp

Twisting/Ring pox/Pit pox

Musa spp.

Prunus avium

CTLaV

MT

Seed

MT (N. occidentialis)/GT

Leaves: Brown necrotic spots/Rusty Chlorosis/ GT Shot holes Bark: Blisters/Gum pockets/ Necrosis Rusty mottle GT

Prunus avium/ Prunus persica Prunus avium

Yellow mottle/Ringspots

Prunus spp.

CRMaV

Ringspots/Yellowing/Necrosis GT

MT

Elaeis guineensis

Cherry rusty mottle associated virus Cherry twisted leaf associated virus

800

800

CNRMV

CGRMV

AOPRV

Cherry necrotic rusty mottle virus

Robigovirus African oil palm ringspot virus Cherry green ring mottle virus

Mild chlorosis/Vein clearing

Rubus canadensis

RuCV1

MT

Fanleaf symptoms/Symptomless Chlorotic mottle/Chlorotic ringspots

Vitis vinifera Prunus persica

Prunus mume

GVT PCMV

Transmission

Vitis spp.

Grapevine rupestris stem pittingassociated virus Grapevine virus T Peach chlorotic mottle virus Rubus canadensis virus 1

Symptoms

GRSPaV

APV2

Asian prunus virus 2

Natural Hosts Symptomless/Chlorotic spots GT Chlorotic spots/Fruit Deformation/Vein MT (N. occidentalis)/GT enlargement & discoloration/Necrotic spots Chlorotic spots/Fruit deformation/Vein GT enlargement & discoloration Pitting/Grooving/Distortion GT

ApLV APV1

Apricot latent virus Asian prunus virus 1

Particle size (nm) Prunus spp. Prunus nume/ Prunus persica

FYI

Continued

Genus/Virus name

Table 1

France Australia

India

North America

North America/Europe/ New Zealand/South Korea North America

South America/South Africa North America/Europe

USA

Europe Canada

USA/Uruguay/China

USA

Europe/Africa USA

Distribution

NC04308 NC003870

NC002729

NC024449

NC020996

NC002468

NC001946

NC012519

NC019025

NC035203 NC009892

NC001948

NC028868

NC014821 NC025388

Access #

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Fig. 1 Phylogenetic analysis of subfamily Quinvirinae based on the amino acid sequences of the RdRP (A) and CP (B). Multiple sequence alignments were generated using DNAMAN software version 5.1 (Lynnon Biosoft, San Ramon, CA, USA) and phylogenetic trees were constructed by the neighbor-joining algorithm, based on calculations from pairwise amino acid sequence distances for protein analyses derived from the multiple alignment format. The horizontal branch lengths are proportional to the genetic distance, and the numbers at each point indicate bootstrap values. The dataset was subjected to 10,000 bootstrap replicates. Square brackets denote accession numbers in the NCBI database.

coding for proteins of 223 kDa RdRp (5907 nt), 25 kDa TGBp1 (690 nt), 12 kDa TGBp2 (330 nt), 7 kDa TGBp3 (192 nt), 34 kDa CP (915 nt), and 11 kDa CRP (327 nt) from the 50 to 30 -end. The both terminal regions of TGBp2 (12 kDa) shows overlap with TGBp1 and TGBp3, respectively. The 50 -terminal region of ORF6 shows an overlap with ORF5 (CP). The lengths of the 50 nontranslated region (NTR) and the 30 -NTR have 73 and 72 nt. The 30 -NTR region contains the poly (A) tail. The genome of ASPV, a member of the genus Foveavirus, consists of 9332 nt and contains five ORFs encoding 247 kDa RdRp (6552 nt), 25 kDa TGBp1 (672 nt), 13 kDa TGBp2 (363 nt), 7 kDa TGBp3 (213 nt), and 44 kDa CP (1245 nt) from the 50 to 30 -end. The 30 -terminal region of TGBp2 shows an overlap with TGBp3. The lengths of the 50 -NTR and the 30 -NTR have 59 and 133 nt. The 30 -NTR region contains the poly(A) tail. The genome of CNRMV, a member of the genus Robigovirus, has 8432 nt and contains five ORFs encoding 232 kDa RdRp (6117 nt), 25 kDa TGBp1 (669 nt), 12 kDa TGBp2 (348 nt), 7 kDa TGBp3 (204 nt), and 30 kDa CP (804 nt) from

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Fig. 2 Phylogenetic analysis of subfamily Quinvirinae based on the amino acid sequences of the TGBp1 (A), TGBp2 (B), and TGBp3 (C). Green and yellow boxes indicate respectively members of the genus Foveovirus and members of the genus Robigovirus. Multiple sequence alignments were generated using DNAMAN software version 5.1 (Lynnon Biosoft, San Ramon, CA, USA) and phylogenetic trees were constructed by the neighbor-joining algorithm, based on calculations from pairwise amino acid sequence distances for protein analyses derived from the multiple alignment format. The horizontal branch lengths are proportional to the genetic distance, and the numbers at each point indicate bootstrap values. The dataset was subjected to 10,000 bootstrap replicates. Square brackets denote accession numbers in the NCBI database.

the 50 to 30 -end. The 30 -terminal region of TGBp2 shows an overlap with TGBp3. The genome has putative 13 kDa and 18 kDa within the TGBp1 and the CP, respectively. The lengths of the 50 -NTR and the 30 -NTR have 117 and 192 nt. The 30 -NTR region contains the poly(A) tail.

Properties and Functions of Gene Products The RdRp of quinviruses contains five conserved domains: a methyl-transferase (MT) domain; an AlkB domain; a papain-like protease (p-Pro) domain; a RNA helicase (HEL) domain; a polymerase (POL) domain. The AlkB domain that repairs alkylated RNA is located in the RdRp of quinviruses, potexviruses, and potyviruses. The papain-like protease (p-Pro) is related to polyprotein processing in viral replication. TGB resembling potex-like TGB that is essential for their cell-to-cell and long-distance movement.

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Fig. 3 Comparison of the genome organizations of the different genera (Carlavirus, Foveavirus, Robigovirus) in the subfamily Quinvirinae. RNAdependent RNA polymerase (RdRp) includes a methyl-transferase (MT), an AlkB domain, a papain-like protease (p-Pro) domain, a helicase (HEL) domain, and a polymerase (POL) domain. Gray box indicates triple gene block corresponding to three overlapping ORFs (TGBp1, TGBp2, and TGBp3). ORF5 encodes coat protein (CP). The ORF6 of PVM, genus Carlavirus, encodes a cysteine-rich protein (CRP).

TGBp1 is associated with ATPase, RNA-binding, and RNA-helicase activities. TGBp2 and TGBp3 are integral membrane proteins required for virus movement. The TGBp1 and CP form a complex with viral RNA that transfers via the plasmodesmata. TGBp1 acts as viral suppressors of RNA silencing. The TGBp1 of PVM demonstrates suppressor activity on systemic silencing. The CP is required for virion assembly. The CRP (ORF6) of carlaviruses contains two motifs: a nuclear localization signal (NLS) and a zinc finger motif (ZF). The CRP is implicated as silencing suppressors for a pathogenicity determinant. The CRPs of PVM, DVS, Chrysanthemum virus B (CVB), Lily symptomless virus (LSV), HeNNV, and Narcissus common latent virus (NLV) have the ability to enhance the pathogenicity of a heterologous virus (Potato virus X) and effects on virus accumulation levels. The CRPs of genera Benyvirus, Tobravirus, Furovirus, and Hordeiviruses have a similar role to that of the carlavirus CRP. The analysis of simple sequence repeats (SSRs) and compound simple sequence repeat (cSSR) have been demonstrated in 32 species of carlaviruses. Mononucleoide A/T is most commonly followed by dinucleotide GT/TG and trinucleotide AAG/GAA in their genome. The SSR and cSSR are mainly located in the RdRp and CRP independent of their genome size.

Replication and Propagation ORF1 (RdRp) is translated directed from the genomic RNA. The ORF1 of Blueberry scorch virus (BlScV), a member of the Carlavirus genus, is proteolytically processed by a papain-like proteinase. The largest product of 190 kDa can be synthesized in vitro using the viral RNA of Potato virus S (PVS) isolated from infected potato plant. The proteolytic processing of the 190 kDa protein is associated with accumulation of 150 kDa peptide after 60 min incubation. L-canavanine is related to specifically reduce the quantity of 34 kDa for PVS CP. Carlavirus ORF1 and TGBp3 contain a heptanucleotide motif (C/UUUAGGU) for sub-genomic RNA promoter. Two sub-genomic RNAs for TGB, CP, and CRP synthesis have been demonstrated in some carlaviruses. These subgenomic RNAs might be encapsidated into their viral particles. Virus particles are usually found in the cytoplasm in infected plant.

Transmission and Host Range Most carlaviruses are spread by mechanical transmission in horticultural crops and herbaceous plants. Some carlaviruses, American hop latent virus (AHLV), BlScV, DVS, LSV, Hop latent virus (HpLV), Hop mosaic virus (HpMV), Potato virus S (PVS), PVM, and PotLV, can be transmitted non-persistently by aphid. AHpLV, HpLV, and HpMV are known to infect to hop plants and known to occur in mixed infection. The three viruses are transmitted by hop aphid (Phorodon humuli). HpLV and HpMV are

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Fig. 4 Symptoms of diseases induced by carlaviruses (a). Helleborus net necrosis virus (HeNNV; Carlavirus, Quinvirinae, Betaflexiviridae) on Helleborus spp (b). Hydrangea chlorotic mottle virus (HdCMoV; Carlavirus, Quinvirinae, Betaflexiviridae) on Hydrangea macrophylla (c). Melon yellowing associated virus (MYaV; Carlavirus, Quinvirinae, Betaflexiviridae) on watermelon and on cucumber (d). Source of photographs: (a) Shiraishi, T., Hoshi, H., Eimori, K., et al., 2011. First report of Helleborus net necrosis virus isolated from hellebores with black death syndrome in Japan. Journal of General Plant Pathology 77, 269–272. doi:10.1007/s10327-011-0321-2. (b) Tang, J., Harper, S.J., Wei, T., et al., 2010. Characterization of hydrangea chlorotic mottle virus, a new member of the genus Carlavirus. Archives of Virology 155, 7–12. doi:10.1007/s00705009-0541-3. (c) & (d) Nagata, T., Dutra, L.S., Oliveira, P.A., et al., 2010. Analysis of the triple gene block sequence in an important melon pathogen, Melon yellowing-associated virus. Journal of General Plant Pathology 76, 268–272. doi:10.1007/s10327-010-0241-6.

transmitted by potato aphid (Macrosiphum euphorbiae) and green peach aphid (Myzus persicae), and can also be transmitted by mechanical inoculation. AHLV and HpLV cause commonly symptomless on any commercial hop plants. HpMV causes chlorotic mosaic mottling on susceptible hop plants. BlScV can cause necrosis of flower and young leaves, shoot blight, and chlorosis in highbush blueberry (Vaccinium corymbosum L.) whereas it appears to be asymptomatic in cranberry (Vaccinium macrocarpon L.). PVS is mechanically transmissible and causes severe disease in potato and localized infection of Chenopodium spp. The virus recently has been detected from arracacha (Arracacia xanthorrhiza) in Peru. Two purified carlaviruses, Pea streak virus and Red clover vein mosaic virus (RCVMV) causing severe streaking symptoms on pea, can be transmitted by Acrythosiphon pisum (pea aphid). The RCVMV can be mechanically transmitted to chickpea (Cicer arietinum L.), faba bean (Vicia faba L.) and lentil (Lens culinaris Medik.). Ligustrum necrotic ringspot virus (LNRSV) isolated from Ligustrum japonicum Thunb showing necrotic ringspots and line patterns is mechanically transmissible to Chenopodium amaranticolor. CPMMV is transmitted non-persistently by whitefly and is the possibility of seed transmission. The virus can cause systemic mottling, chlorotic blotches, and leaf malformations in cowpea (Vigna unguiculata L.) and can be naturally transmitted to peanut (Arachis hypogaea L.), soybean (Glycine max L.), and common bean (Phaseolus vulgaris L.). Cucumber vein-clearing virus (CuVCV) can transmitted by mechanical inoculation and by whitefly. Melon yellowing-associated virus also is transmitted by whitefly but is not transmitted by mechanical inoculation. Rose virus A has been detected recently in a memorial rose (Rosa wichuraiana Crep.) showing leaf deformation and mosaic symptoms. Foveaviruses and robigoviruses naturally infect woody plants (Fig. 5). These viruses can be transmitted by grafting and sap inoculation. ASPV causes systemic infection on Nicotiana occidentalis and local infection on Chopodium murale. ApLV, a member of the genus Foveavirus, causes chlorotic spots on Prunus persica and P. cerasifera, red to purple rings and mottling on P. avum, and chlorotic blotching and malformation on P. armeniaca cv. Tirynthos and Haward but not on flowers and fruits. The ApLV can cause asymptomatic infection on cherry and plum cultivars. CGRMV can be sap transmitted from cherry to cherry. Novel foveavirus has been detected in stems and leaves of ginseng (Panax ginseng) exhibiting mosaic and ringspot symptoms. GRSPaV can be transmitted by grafting but not

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Fig. 5 (a) Apple cultivar ‘Hongro’ with mixed infection of Apple stem pitting virus (ASPV; Foveavirus, Quinvirinae, Betaflexiviridae), Apple chlorotic leaf spot virus (ACLSV, Trichovirus, Trivirinae, Betaflexiviridae), and Apple stem grooving virus (ASGV; Capillovirus, Trivirinae, Betaflexiviridae), showing chlorosis along the leaf veins. (b) Symptoms of Grapevine rupestris stem pitting-associated virus (GRSPaV; Foveavirus, Betaflexiviridae) of vein necrosis in the grapevine indicator 110 R: necrosis of leaf veinlets and petioles (left) and of the pith (right). (Photograph courtesy of G. Martelli). (c) Necrotic symptoms from Cherry necrotic rusty mottle virus (CNRMoV; Robigovirus, Quinvirinae, Betaflexiviridae) on Sweetheart cherries tree (Photo from Iain MacSwann) and (d) detail of necrosis on Corum cherry leaves (Photograph from Jay W. Pscheidt). Source of photographs: (a) Yoon, J.Y., Joa, J.H., Choi, K.S., et al., 2014. Genetic diversity of a natural population of Apple stem pitting virus isolated from apple in Korea. The Plant Pathology Journal 30 (2), 195–199. doi:10.5423/PPJ.NT.02.2014.0015. (b) Bouyahia, H., Boscia, D., Savino, V.N., et al., 2015. Grapevine rupestris stem pitting-associated virus is linked with grapevine vein necrosis. Vitis 44 (3), 133–137. (d) Pacific Northwest Plant Disease Management Handbook. https://pnwhandbooks.org/node/2463.

transmitted by mechanical inoculation. The five members of the genus Robigovirus can be transmitted to cherry and other stone fruit tree and cause serious diseases (Figs. 4 and 5; Table 1).

Epidemiology and Control Quinviruses can cause usually mosaic, mottle, and ringspot symptoms in their host plants. PVS and PVM, members of the genus Carlavirus, are essentially symptomless in most potato cultivars. The two viruses are easily transmitted during the cutting of tubers to make seed pieces. The PVS can cause yield losses of up to 20% in potato field. The primary method for producing virus-free potato tuber seeds has been used to control the viruses. The Ns gene derived from potato (Solanum tuberosum spp. andigena) induces hypersensitive resistance to PVS and have been used in potato breeding programs. Orchards and vegetables sustain economic damage caused by some quinviruses that require suitable disease management. MYaV, a member of the genus Carlavirus, causes severe yellowing disease in melon crops and could transmitted by whitefly. New melon plants containing a dominant MYaV resistance gene recently have been developed for the virus disease management. A whitefly-transmitted CPMMV, a member of the genus Carlavirus, causes substantial losses in the yield and quality of cowpea. The whitefly-transmitted CPMMV can be controlled with insecticides such as Nuprid 2F and Dimethoate. The SSR markers associated with CPMMV resistance in cowpea have been recently identified and will be used for new cowpea variety resistance to CPMMV. HeNNV, a member of the genus Carlavirus, associated with black death of Hellleborus spp. has been reported in New Zealand, Japan, UK, and the USA. LSV, a member of the genus Carlavirus, which is generally symptomless in field-grown plants may cause problems under the insufficient light and

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growing conditions of year-round cut-flower production in greenhouses. In the Netherlands, an aphid-transmitted LSV spreads mainly in June–July, considerably less in May, and least in August/September. It is curtailed by a mixture of mineral oil and pyrethroid insecticide. Three hop viruses, AHpLV, HpLV, and HpMV, can be controlled by the use of certified virus-free planting stock and insecticides for reducing aphids. HpMV has been found infecting hops in Europe, Australia, the United States, New Zealand, China, Japan, and South Africa. Blueberry scorch disease caused by aphid-transmitted BlScV is a severe disease of highbush blueberry (Vaccinium corymbosum L.) in Canada, Europe, and the USA. The viral disease can be controlled by heat therapy and apical meristem culture. Transgenic chrysanthemum, tobacco, and potato, which are resistant to CPMMV, Chrysanthemum virus B, PVM, and PVS have been developed.

Diagnosis Quinviruses can be detected by polymerase chain reaction (PCR)-based methods, such as reverse transcription (RT)-PCR, immunocapture-RT-PCR, multiplex RT-PCR, and real-time PCR. Universal primers for detection of potato carlaviruses (PVS, PVM, and PotLV) have been developed based on the conserved aa sequences “SNNMA” and “GLGVPTE” in the CP. Other universal primer, a Carla-CP, has been designed based on the conserved nt sequences of the GLGVPTE motif in the CPs derived from 17 carlavirus species (Aconitum latent virus, BlScV, Carnation latent virus, Cole latent virus, CPMMV, CVB, DVS, Helenium virus S, HpMV, LSV, Narcissus common latent virus, Passiflora latent virus, PotLV, PVM, Potato virus P, PVS). Carlaviruses can be detected by RT-PCR using the Carla-CP and oligo-d(T21) primers. Some commercial and non-commercial antisera are be used for immunological detection of quinviruses. Four carlaviruses, CLV, HpLV, HpMV, and PVS are antigenically similar.

Concluding Remarks Two genera, Carlavirus and Foveavrus, belong to subfamily Quinvirinae and a newly created genus Robigovirus has been recently added to that subfamily. The completed genome sequences of novel carlaviruses (LAV, MYaV, and SPCFV) and foveavirus (GVT) have been determined recently. African oil palm ringspot virus, Cherry green ring mottle virus, CNRMV, Cherry rusty mottle associated virus, and Cherry twisted leaf associated virus belong to newly established genus Robigovirus. In phylogenetic trees based on their RdRp and CP proteins of members of the subfamily Quinvirinae, the genera Carlavirus and Foveavirus have not been grouped according to their natural host species. Cowpea, cherry, grapevine, melon, and potato cause severe symptoms by some quinviruses. Some carlaviruses can be transmitted by aphid and whitefly. Quinviruses have TGB resembling potex-like TGB for viral movement. Carlaviruses have two proteins (TGBp1 and CRP) identified as RNA silencing suppressor. Immunological and PCR-based techniques are most commonly used for detection of quinviruses.

Further Reading Alam, C.M., Singh, A.K., Sharfuddin, C., Ali, S., 2014. Genome-wide scan for analysis of simple and imperfect microsatellites in diverse carlaviruses. Infection, Genetics and Evolution 21, 287–294. Belay, D.K., Huckaba, R.M., Ramirez, A.M., Rodrigues, C.V., Foster, J.E., 2012. Insecticidal control of Bemisia tabaci (Hemiptera: Aleyrodidae) transmitting Carlavirus on soybeans and detection of the virus in alternate hosts. Crop Protection 35, 53–57. Bhattarai, G., Shi, A., Qin, J., et al., 2017. Association analysis of Cowpea mosaic virus (CPMV) resistance in the USDA cowpea germplasm collection. Euphytica 213, 230. Costa, T.M., Blawid, R., da Costa Junior, A.C., et al., 2017. Complete genome sequence of Melon yellowing-associated virus from melon plants with the severe yellowing disease in Brazil. Archives of Virology 162, 3899–3901. De Souza, J., Gamarra, H., Müller, G., Kreuze, J., 2018. First report of Potato virus S naturally infecting arracacha (Arracacia xanthorrhiza) in Peru. Plant Disease 102, 460. Diaz-Lara, A., Mollov, D., Golino, D., Rwahnih, M.A., 2020. Complete genome sequence of Rose virus A, the first carlavirus identified in rose. Archives of Virology 165, 241–244. Fujita, N., Komatsu, K., Ayukawa, Y., et al., 2016. N-terminal region of cysteine-rich protein (CRP) in carlaviruses is involved in the determination of symptom types. Molecular Plant Pathology 19, 180–190. Ho, T., Quito-Avila, D., Keller, K.E., et al., 2016. Evidence of sympatric speciation of elderberry carlaviruses. Virus Research 215, 72–75. Park, D., Zhang, M., Hahn, Y., 2019. Novel foveavirus (family Betaflexiviridae) species identified in geinseng. Acta Virologica 63, 155–161. Rodamilans, B., Shan, H., Pasin, F., Garcia, J.A., 2018. Plant viral protease: Beyond the role of peptide cutters. Frontiers in Plant Science 9, 666. Williamson, M.A., Scott, S.W., 2018. First report of hellebore black death of Lenten rose, caused by Helleborus net necrosis virus, in South Carolina. Plant Disease 102, 2667. Wu, L., Liu, H., Bateman, H., Komorowska, B., Li, R., 2019. First identification and molecular characterization of a novel cherry robiovirus. Archives of Virology 164, 3103–3106.

Reverse-Transcribing Viruses (Belpaoviridae, Metaviridae, and Pseudoviridae) Carlos Llorens, Biotechvana, Scientific Park University of Valencia, Valencia, Spain Beatriz Soriano, Biotechvana, Scientific Park University of Valencia and Institute for Integrative Systems Biology (I2SysBio), University of Valencia–Spanish National Research Council, Valencia, Spain Maria A Navarrete-Muñoz, Biotechvana, Madrid, Spain; Institute of Health Research-Jiménez Díaz Foundation, Autonomous University of Madrid; and Rey Juan Carlos University Hospital, Móstoles, Spain Ahmed Hafez, Biotechvana, Valencia, Spain; Pompeu Fabra University, Barcelona, Spain; and Minia University, Minya, Egypt Vicente Arnau, Institute for Integrative Systems Biology (I2SysBio), University of Valencia–Spanish National Research Council, Valencia, Spain Jose Miguel Benito, Health Research Institute of the Jiménez Díaz Foundation, Autonomous University of Madrid and Rey Juan Carlos University Hospital, Móstoles, Spain Toni Gabaldon, Barcelona Supercomputing Center-National Center for Supercomputing, Institute of Research in Biomedicine, and Catalan Institution for Research and Advanced Studies, Barcelona, Spain Norma Rallon, Institute of Health Research-Jiménez Díaz Foundation, Autonomous University of Madrid and Rey Juan Carlos University Hospital, Móstoles, Spain Jaume Pérez-Sánchez, Institute of Aquaculture Torre de la Sal, Spanish National Research Council, Castellon, Spain Mart Krupovic, Archaeal Virology Unit, Institut Pasteur, Paris, France r 2021 Elsevier Ltd. All rights reserved.

Nomenclature CP Capsid protein ENV Envelope ERV Endogenous retrovirus Gag Group-specific antigen GYA Billion (giga) years ago HERV Human endogenous retrovirus ICTV International Committee on Taxonomy of Viruses INT Integrase Int-Chr Chromodomain-containing Integrase kb Kilobase LTR Long terminal repeat MYA Million (mega) years ago NC Nucleocapsid

Glossary Chromoviruses In reference to metaviruses coding for chrodomain-like integrases. Errantivirus Erranti comes from the Latin errans, meaning to wander. LTR-retroelement Term used to collectively refer to retroviruses and LTR-retrotransposons, regardless of whether they possess env or env-like proteins. Metavirus Meta comes from the Greek metathesis for transposition. Monophyletic Phylogenetic group collecting the full set of operational taxonomic units descending from a single ancestor. Operational taxonomic unit Termed used to classify groups of closely related individuals. Ortervirales Orter is an inversion of retro, which was derived from reverse transcription; and virales - which is the suffix for an order in virology.

Encyclopedia of Virology, 4th Edition, Volume 3

ORF Open reading frame OTU Operative taxonomical unit PBS Primer binding site PPT Polypurine track PR Protease RH Ribonuclease H RNAi RNA interference RNP Ribonucleoprotein RT Reverse transcriptase ssRNA single-stranded ribonucleic acid SU Surface TM Transmembrane VLP Virus like particle

Paraphyletic clade Phylogenetic group collecting some but not all operational taxonomic units descending from a single ancestor. Polyphyletic Phylogenetic group composed of OTUs descending from more than one ancestor. Pro-retroviruses Term used here to describe LTR retrotransposons that have acquired a third ORF, env, and are thus potential retrovirus but with no evidence of infectivity. Pseudovirus Pseudo comes from Greek pseudo, “false” to connote some uncertainty as to whether these are true viruses. Semotivirus Semoti comes from the Latin semotus, meaning distant or removed (it was formerly used to classify Belpao elements).

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Introduction: The Paradigm of LTR Retrotransposons and Reverse-Transcribing Viruses Metaviridae, Pseudoviridae, and Belpaoviridae are the three families officially introduced by the International Committee on Taxonomy of Viruses (ICTV) for classification of the eukaryotic Ty3/Gypsy, Ty1/Copia, and Bel/Pao LTR retroelements. The members of the families Metaviridae, Pseudoviridae, and Belpaoviridae are widely (but differentially) distributed in plants, fungi and animals where they may exist as LTR retrotransposons or as potential or true reverse-transcribing viruses. Due to multiple distinct similarities in the life cycles, sequence and architecture of genes, and functional features, it is commonly accepted that members of the families Metaviridae, Pseudoviridae, and Belpaoviridae share origins and evolutionary history with viruses of the Retroviridae family, which is thus considered as a member of the same phylogenetic lineage of reverse-transcribing viruses and LTR retrotransposons. It is worth for the reader to distinguish simple reverse-transcribing viruses from the complex ones. In the family Retroviridae, simple retroviruses are those that present a core genomic organization of gag, pol, and env genes flanked by LTRs, with no accessory genes (i.e., LTR-gag-pol-env-LTR), while complex retroviruses incorporate one or more accessory genes (vif, vpx, rev, nef, tat, or others) needed to complete the life cycle and infect other cells (Fig. 1). In their most current form, the members of the families Metaviridae, Pseudoviridae, and Belpaoviridae are LTR retrotransposons presenting a “LTR-gag-pol-LTR” genomic architecture (Fig. 1). Thus, the major difference between LTR retrotransposons and simple retroviruses is that the later include a third gene (env) for an envelope protein which confers infectious abilities to the retroviruses. However, some members of the families Metaviridae, Pseudoviridae, and Belpaoviridae also carry the env gene, suggesting that an LTR retrotransposon might become a retrovirus upon acquiring a third env gene. Among metaviruses, pseudoviruses and belpaoviruses with env, only those from the Errantivirus genus in the family Metaviridae have been demonstrated to generate infectious virions. Interestingly, the LTR retrotransposon-virus paradigm gets support also in the reverse path because a reverse-transcribing virus may become or act as an LTR retrotransposon when the env gene is lost and/or the machinery required for infecting other cells is neutralized. This is the case for most endogenous retroviruses (ERVs) of vertebrates, which usually derive from exogenous retroviruses that after colonizing the germ cells became unable to infect other cells due to accumulation of multiple mutations. Interestingly, the replication system of LTR retroelements (i.e., reverse-transcribing viruses and LTR retrotransposons) constituted by the members of the families Metaviridae, Belpaoviridae, Pseudoviridae, and Retroviridae is also related to that of reverse-transcribing viruses of the family Caulimoviridae which infect plants. Caulimoviruses are dsDNA viruses whose genome usually harbors two ORFs encoding for coat (gag) and pol polyproteins (Fig. 1), with domain features closely similar to those of metaviruses, pseudoviruses and belpaoviruses. These similarities suggest that caulimoviruses share a common evolutionary history with LTR retrotransposons and that caulimoviruses would be chimeric viruses resulting from recombination between a metavirus and another RNA virus or LTR retrotransposon that evolved to the status of a virus by recruiting new genes through horizontal transfer. In recognition of all the structural, functional and evolutionary relationships among the members of the families Caulimoviridae, Belpaoviridae, Metaviridae, Pseudoviridae, and Retroviridae, the ICTV has recently unified the five families into a single viral order, the Ortervirales.

Genome Organization and Functions of Gene Products and Other Structural Features The full-lenght genome sequence of canonical LTR retroelements of the Metaviridae, Pseudoviridae, and Belpaoviridae families normally present two LTRs that flank an internal region of variable size (from 3 to 15 kb) that contains open reading frames (ORFs) for the encoded Gag, Pol, and/or Env protein domains (Fig. 1). The LTRs are two normally homologous non-coding sequences that usually begin and end in dinucleotide (50 -TG … CA-30 ) inverted repeats. A typical LTR displays three regions: a U3 region of 200–1200 nt that contains the promoters; a region called “R” because it is repeated on each end of the transcript; and a U5 region of 75–250 nt that constitutes the first portion of the reverse-transcribed genome. Normally, the LTRs do not contain coding genes but present regulatory elements such as “enhancers” and “promoters” that regulate the expression of the genes in the internal region of the genome and also, in certain cases, the host genes. The internal region is usually delimited by two small motifs: a 18 nt sequence downstream to the 50 LTR called Primer Binding Site (PBS) and by another small region of B10 A/G located upstream of the 30 LTR called “Polypurine Tract” (PPT). The internal region normally includes ORFs encoding for two polyproteins, Gag and Pol. The Gag polyprotein presents domains for the structural proteins including the capsid protein (CP), which forms the core of the Virus-Like Particle (VLP), and a nucleocapsid (NC) involved in RNA packaging through recognition of a specific region in the viral genome called C. Curiously, C is a frequent site of deletions in the proviral genome, giving defective VLPs which could generate non-infectious virions in some reverse-transcribing viruses with the entire LTR-gag-pol-env-LTR structure. On the other hand, Pol presents domains for distinct enzymes such as a protease (PR) belonging to the clan AA of aspartic peptidases that is required for proteolytic processing of the two Gag and Pol polyproteins; a reverse transcriptase (RT) involved in the reverse transcription synthesis of cDNA from the single-stranded RNA (ssRNA) template; a ribonuclease H (RH) responsible for the hydrolysis of the original RNA template that is part of the RNA/DNA hybrid generated during reverse transcription; an integrase (INT) needed for the integration of the proviral cDNA into the host genome. As also shown in Fig. 1, viruses in the Metaviridae and Belpaoviridae families usually present a common organization of gag-pol domains (CP-NC-PR-RTRH-INT), which are also observed in retroviruses. The only difference between the two groups is that retroviruses additionally contain the matrix domain (MA) in the Gag polyprotein, upstream of the CP, which is not present (or at least not preserved at the domain level) in metaviruses and belpaoviruses (nor in pseudoviruses). The order of the RT and INT domains in the internal region of members of the Pseudoviridae family is different from that of metaviruses, belpaoviruses and retroviruses. While

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Fig. 1 Representative elements of each family within the order Ortervirales. Simian inmunodeficiency virus (SIV) of Macaca mulatta is taken as a complex retrovirus example of the family Retroviridae, where the typical accessory genes are colored green. Koala retrovirus (KoRV) is used as an example of a simple retrovirus in the family Retroviridae. Legolas and Gypsy are used as representatives for the family Metaviridae. Copia and Sire are shown as representatives of the family Pseudoviridae. Pao and Tas represent the family Belpaoviridae. The Cauliflower mosaic virus (CaMV) is provided as a representative of the family Caulimoviridae. Domain features of each gag, pol, and env region are indicated below each represented genome description. PBS and PPT acronyms mean Primer Binding Site and Polypurine Tract, respectively. MA, CP and NC acronyms in gag refer to Matrix, Capsid and Nucleocapsid domains, respectively. PR, RT, RH and INT acronyms in pol refer to Protease, Reverse Transcriptase, Ribonuclease H and Integrase domains, respectively. SU and TM acronyms at env refer to Surface and Transmebrane domains, respectively. P1, P2 P3 and P6 in caulimoviruses refer to Movement protein, Aphid transmission factor, Virion associated protein, and Transactivator/ Viroplasmin, respectively. For more examples of orterviral representative full-length genomes, visit the GyDB (http://gydb.org). In addition, Table 1 provides a summary of all members of the Metaviridae, Belpaoviridae, and Pseudoviridae families currently classified by the ICTV.

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Reverse-Transcribing Viruses (Belpaoviridae, Metaviridae, and Pseudoviridae)

metaviruses and belpaoviruses usually present RT-RH domains upstream of INT, pseudoviruses present a distinct organization with the INT domain being interspersed between PR and RT domains. Besides the “LTR-gag-pol-LTR” domain architecture, many lineages of the members of the Metaviridae, Pseudoviridae and Belpaoviridae families additionally contain a third env ORF which potentially endows them with infectivity. In retroviruses, Env protein binds to a receptor on the target cell surface and this event triggers a conformational change which leads to the fusion of viral and cellular membranes. Furthermore, at least some Metaviridae members, such as those belonging to the Tat clade in plants, encode accessory proteins (similar to complex retroviruses), whose function, however, remains unclear.

Taxonomy, Phylogeny and Members of the Families Classification of the distinct LTR retroelements or viruses in the order Ortervirales is performed by phylogenetic methods. In Fig. 2, we show the phylogeny of the Ortevirales inferred based on the Pol polypeptide domains. For simplicity, branches of caulimoviruses and retroviruses are collapsed. Each family splits into distinct phylogenetic lineages, herein referred to as clades or branches (they are bigger in terms of both OTUs - Operative Taxonomical Units - and host distribution). We use popular nomenclature to refer to each clade or branch in the tree. In the following section the taxonomic structure of each family is discussed in detail. As an additional information, Table 1 provides a summary of all members of the Metaviridae, Belpaoviridae, and Pseudoviridae families currently classified by the ICTV. The Metaviridae family is composed of metaviruses (LTR retrotransposons) and pro-retroviruses (LTR retroelements with env gene but with no evidence of an infective behavior). Originally, the distinct elements belonging to the Metaviridae family were classified into three major genera: Errantivirus (Ty3/Gypsy elements with env gene), Metavirus (without env), and Semotivirus (Bel/ Pao elements). This classification derives from the early discovery of Ty3-like LTR retrotransposons of fungi and their Gypsy-like Env-encoding relatives in drosophilids, followed by the characterization of other elements (including Bel/Pao elements) in multiple other eukaryotes. More recently, Bel/Pao elements (genus Semotivirus) were removed from the family Metaviridae because they do not form a monophyletic group with other Metaviridae members. The family Metaviridae needs further revision because the diversity of Ty3/Gypsy elements is much greater and more complex than previously thought. The genus Errantivirus includes Gypsy-like viruses of drosophila, which as shown in Fig. 2, forms a monophyletic clade splitting into at least two sub-clades (denoted as Gypsy and 17.6). Other Ty3/Gypsy elements form distinct clades, including Athila, Osvaldo and Tor4a, and encode for a third ORF showing structural features typical of Env polyproteins, such as the large Surface (SU) domain providing antigenic sites rich in cysteines, the transmembrane (TM) domain near their carboxyl terminus, multiple putative N-glycosylation sites, and putative protease cleavage sites at conserved positions. These lineages are pro-retroviruses because they carry an env gene, albeit they do not cluster phylogenetically with errantiviruses but more properly with other Ty3/Gypsy lineages of LTR retrotransposons (thus, contradicting the errantivirus-metavirus classification). This is for instance the case of Athila clade of pro-retroviruses of plants that are a sister clade of LTR retrotransposons called Tat. Athila and Tat are supported as a monophyletic branch in all Ty3/Gypsy phylogenies (although Athila is a pro-retrovirus and Tat a metavirus). On the other hand, the phylogenetic relationships of all the Ty3/Gypsy lineages originally considered to constitute the genus Metavirus cannot be resolved based on current data due to the high sequence divergence in that family. Morever, members of one of the largest and best supported branches of Ty3/Gypsy LTR retrotransposons, labeled Chromovirus in Fig. 2, differ from other Ty3/Gypsy elements in that (with very few exceptions, such as Ty3, TF1 and TF2) their INTs have a conserved chromodomain at the C-terminus. This branch of chromodomain-Ty3/Gypsy LTR retrotransposons are formally classified as representative members of the genus Metavirus but is widely referred to as the genus “Chromovirus” (because of their chromodomain INTs) and splits into at least three major branches widely distributed in plants, fungi and vertebrates herein referred to as “Plants”, Algae” and “Fungi/Vertebrates” according to the host distribution of the corresponding elements. Another lineage traditionally classified within the genus Metavirus includes Mag – a large branch collecting different clades of Ty3/Gypsy LTR retrotransposons of metazoans. The remaining diversity of elements originally assigned to the genus Metavirus is distributed across a number of smaller clades (Cer-like, Mdg3/Micropia, 412/Mdg1, Osvaldo/Gmr1, CsRN1, Tor-like and others) (Table 1). The Belpaoviridae family groups all the Bel/Pao LTR retrotransposons and pro-retroviruses of the belpao family formerly classified by the ICTV as the genus Semotivirus within the family Metaviridae because they present the same gag-pol organization as metaviruses (Fig. 1). However, belpaoviruses are significantly divergent on the sequence level from metaviruses and they constitute a well supported monophyletic branch that falls outside of the Metaviridae family. Taking this into consideration, the ICTV has recently moved the Semotivirus genus from the family Metaviridae into a separate family, the Belpaoviridae, within the order Ortervirales. Based on current data, belpaoviruses are less diverse than their metavirus counterparts, albeit their currently known diversity is also greater than previously supposed. Based on the Pol tree shown in Fig. 2, five clades can be defined: Pao, Sinbad, Bel, Tas, and Suzu. Of these, Pao, Sinbad, and Suzu clades group together in a well supported branch. The relationship of this branch with the two other clades (Bel and Tas) remains unresolved. Bel and Tas clades mainly include LTR retrotransposons. However, within Tas clade, some elements, such as Tas (the element that gives the clade its name), carry an ORF encoding for a putative Env polyprotein, suggesting that at least some belpaoviruses might be able to form infectious virions. The Pseudoviridae family is also highly diverse and includes all Ty1/Copia LTR retrotransposons and pro-retroviruses. At present, the ICTV recognizes three genera within the Pseudoviridae – Pseudovirus, Hemivirus, and Sirevirus. Although these three genera are well supported in the current phylogeny, they are not sufficient to cover the whole diversity of Ty1/Copia LTR retroelements. Based

Reverse-Transcribing Viruses (Belpaoviridae, Metaviridae, and Pseudoviridae)

657

Fig. 2 Phylogeny of members of the order Ortervirales based on an aligment concatenation of the most conserved part or cores of the four pol polyprotein domains (PR-RT-RH-INT) of 268 OTUs belonging to the Retroviridae, Caulimoviridae, Metaviridae, Pseudoviridae, and Belpaoviridae families. OTUs are collapsed in clades except for those elements providing clusters of a single OTU, which are represented with their name in gray and italicized. Families and genera currently recognized by the ICTV are provided in bold italics. The absence of an INT domain in the alignment is represented with a gap in the pol alignment. For simplicity, bootstrap information is provided only for the most internal nodes of the tree and only when boostrap values are higher than 50. Families collecting true or potential retroviruses or LTR retrotransposons are highlighted in red. The figure is adapted from Fig. 1 in Llorens C., et al. 2009. The full expanded version of this tree in web format is available as Suplemental file 1 in Llorens C., et al., 2009. The web version of this tree provides information about names, Genbank accessions and hosts of all LTR retroelement taxa used. By clicking on each OTU in the pol tree, the reader can access the accession of the requested element from GyDB or NCBI, respectively. Although our current knowledge about the diversity of Ortervirales is still incomplete, this Pol tree provides an acceptable and realistic perspective on their evolutionary history, mainly because of the strong phylogenetic signal of the RT, which is the pol domain most frequently used to infer retroelement phylogenies in general. With reasonable variations due to differences in rates of evolution, this evolutionary history is also supported by gag (the other protein domain common to all Ortervirales) and by other proteins encoded by these LTR retroelements. Readers interested in more specific details about the evolutionary history of the order Ortervirales can visit GyDB and access a comprehensive collection of phylogenetic trees inferred based on all encoded products of these LTR retroelements. The figure is adapted from Fig. 1 in Llorens, C., Muñoz-Pomer, A., Bernad, L., Botella, H., Moya, A., 2009. Network dynamics of eukaryotic LTR retroelements beyond phylogenetic trees. Biology Direct 4, 41. The full expanded version of this tree in web format is also provided as Suplemental file 1 in that article.

658

Reverse-Transcribing Viruses (Belpaoviridae, Metaviridae, and Pseudoviridae)

Table 1

Members in the families Belpaoviridae, Metaviridae, and Pseudoviridae (order Ortervirales). Genera type species are written in bold

Family/Genus /Phylogenetic GyDB cladea Belpaoviridae Semotivirus Tas Pao

Bel

Sinbad Suzu Undefined at GyDB Metaviridae Errantivirus Gypsy

17.6

Undefined At GyDB Metavirus Athila/Tat Mag Cer1 Osvaldo

412/Mdg1

Micropia/Mdg3

Chromovirusb

Undefined At GyDB

Species

Exemplar virus

Acronym

Literature name GenBank accession

Ascaris lumbricoides Tas virus Antheraea Tamy virus Bombyx mori Pao virus Drosophila simulans Ninja virus Drosophila melanogaster Bel virus Drosophila melanogaster Roo virus Drosophila melanogaster Max virus Anopheles gambiae Moose virus Schistosoma Sinbad virus Takifugu rubripes Suzu virus Caenorhabditis elegans Cer13 virus

Ascaris lumbricoides Tas virus Antheraea mylitta Tamy virus Bombyx mori Pao virus Drosophila simulans Ninja virus Drosophila melanogaster Bel virus Drosophila melanogaster Roo virus Drosophila melanogaster Max virus Anopheles gambiae Moose virus Schistosoma mansoni Sinbad virus Fugu rubripes Suzu virus Caenorhabditis elegans Cer13 virus

AluTasV TamyV BmoPaoV DsiNinV DmeBelV DmeRooV DmeMaxV AgaMooV FruSuzV CelCer13V

Tas Tamy Pao Ninja Bel Roo Max Moose Sinbad Suzu Cer13

Z29712 AF530470 L09635 D83207 U23420 AY180917 AJ487856 AF060859 AY506538 AF537216 Z81510

Ceratitis capitata Yoyo virus Drosophila melanogaster Gypsy virus Drosophila melanogaster 17–6 virus Drosophila melanogaster 297 virus Drosophila melanogaster Idefix virus Drosophila ananassae Tom virus Drosophila melanogaster Zam virus Drosophila virilis Tv1 virus Trichoplusia ni TED virus Drosophila melanogaster Tirant virus

Ceratitis capitata Yoyo virus Drosophila melanogaster Gypsy virus Drosophila melanogaster 17.6 virus Drosophila melanogaster 297 virus Drosophila melanogaster Idefix virus Drosophila ananassae Tom virus Drosophila melanogaster Zam virus Drosophila virilis Tv1 virus Trichoplusia ni TED virus Drosophila melanogaster Tirant virus

CcaYoyV DmeGypV

Yoyo Gypsy

U60529 M12927

Dme176V Dme297V DmeIdeV DanTomV DmeZamV DviTv1V TniTedV DmeTirV

17.6 297 Idefix Tom Zam Tv1 Ted Tirant

X01472 X03431 AJ009736 Z24451 AJ000387 AF056940 M32662 X93507

Arabidopsis thaliana Athila virus Arabidopsis thaliana Tat4 virus Bombyx mori Mag virus Tripneustis gratilla SURL virus Caenorhabditis elegans Cer1 virus Drosophila virilis Ulysses virus Tribolium castaneum Woot virus Drosophila buzzatii Osvaldo virus Drosophila melanogaster 412 virus Drosophila melanogaster Mdg1 virus Drosophila melanogaster Mdg3 virus Drosophila melanogaster Micropia virus Drosophila melanogaster Blastopia virus Fusarium oxysporum Skippy virus Lilium henryi Del1 virus Saccharomyces cerevisiae Ty3 virus Schizosaccharomyces pombe Tf1 virus Schizosaccharomyces pombe Tf2 virus Takifugu rubripes Sushi virus Dictyostelium discoideum Skipper virus Cladosporium fulvum T  1 virus

Arabidopsis thaliana Athila virus Arabidopsis thaliana Tat4 virus Bombyx mori Mag virus Tripneustis gratilla SURL virus Caenorhabditis elegans Cer1 virus Drosophila virilis Ulysses virus Tribolium castaneum Woot virus Drosophila buzzatii Osvaldo virus Drosophila melanogaster 412 virus Drosophila melanogaster Mdg1 virus Drosophila melanogaster Mdg3 virus Drosophila melanogaster Micropia virus Drosophila melanogaster Blastopia virus Fusarium oxysporum Skippy virus Lilium henryi Del1 virus Saccharomyces cerevisiae Ty3 virus

AthAthV AthTatV BmoMagV TgrSurV CelCer1V DviUllV TcaWooV DbuOsvV Dme412V DmeMdg1V

Athila Tat Mag SURL Cer1 Ulysses Woot Osvaldo 412 Mdg1

AC007209 AB005247 X17219 M75723 U15406 X56645 U09586 AJ133521 X04132 X59545

Schizosaccharomyces pombe Tf1 virus Schizosaccharomyces pombe Tf2 virus Takifugu rubripes Sushi virus Dictyostelium discoideum Skipper virus Cladosporium fulvum T  1 virus

DmeMdg3V Mdg3

X95908

DmeMicV

Micropia

X14037

DmeBlaV

Blastopia

Z27119

FoxSkiV LheDel1V ScerTy3V

Skyppy Del Ty3

L34658 X13886 M34549

SpoTf1V

Tf1

M38526

SpoTf2V

Tf2

L10324

TruSusV DdiSkiV

Sushi Skipper

AF030881 AF049230

CfuT1V

T1

Z11866

Reverse-Transcribing Viruses (Belpaoviridae, Metaviridae, and Pseudoviridae) Table 1

659

Continued Species

Exemplar virus

Acronym

Literature name GenBank accession

Drosophila melanogaster 1731 virus Drosophila melanogaster copia virus Volvox carteri Osser virus Aedes aegypti Mosqcopia virus Candida albicans Tca2 virus Saccharomyces cerevisiae Ty5 virus Volvox carteri Lueckenbuesser virus Candida albicans Tca5 virus

Drosophila melanogaster 1731 virus Drosophila melanogaster copia virus Volvox carteri Osser virus Aedes aegypti Mosqcopia virus Candida albicans Tca2 virus Saccharomyces cerevisiae Ty5 virus Volvox carteri Lueckenbuesser virus Candida albicans Tca5 virus

Dme1731V DmecopiaV VcaOssV AaeMosV CalTca2V SceTy5V VcaLeuV CalTca5V

1731 Copia Osser Mosqcopia Tca2 Ty5 Lueckenbuesser Tca5

X07656 X04456 X69552 AF134899 AF050215 U19263 U90320 AF065434

Zea mays Sto4 virus Nicotiana tabacum Tto1 virus Saccharomyces cerevisiae Ty1 virus Saccharomyces cerevisiae Ty2 virus Saccharomyces cerevisiae Ty4 virus Brassica oleracea Melmoth virus Oryza longistaminata Retrofit virus Zea mays Hopscotch virus Cajanus cajan Panzee virus Glycine max Tgmr virus Hordeum vulgare BARE  1 virus Nicotiana tabacum Tnt1 virus Arabidopsis thaliana Art1 virus Arabidopsis thaliana AtRE1 virus Arabidopsis thaliana evelknievel virus Arabidopsis thaliana Ta1 virus Oryza australiensis RIRE1 virus Physarum polycephalum Tp1 virus Solanum tuberosum Tst1 virus Triticum aestivum WIS2 virus

Zea mays Sto  4 virus ZmaSto4V Nicotiana tabacum Tto1 virus NtaTto1V Saccharomyces cerevisiae Ty1 virus SceTy1V

Sto  4 Tto1 Ty1

AF082133 D83003 M18706

Saccharomyces cerevisiae Ty2 virus Saccharomyces cerevisiae Ty4 virus Brassica oleracea Melmoth virus Oryza longistaminata Retrofit virus Zea mays Hopscotch virus Cajanus cajan Panzee virus Glycine max Tgmr virus Hordeum vulgare BARE  1 virus Nicotiana tabacum Tnt1 virus Arabidopsis thaliana Art1 virus Arabidopsis thaliana AtRE1 virus Arabidopsis thaliana evelknievel virus Arabidopsis thaliana Ta1 virus Oryza australiensis RIRE1 virus Physarum polycephalum Tp1 virus Solanum tuberosum Tst1 virus Triticum aestivum WIS  2 virus

SceTy2V SceTy4V BolMelV OloRetrofitV ZmaHopV CcaPanV GmaTgmrV HvuBar1V NtaTnt1V AthArt1V AthAtRE1V AthEveV AthTA1V OauRIRE1V PpoTp1V StuTst1V TaeWis2V

Ty2 Ty4 Melmoth REtrofit Hopscotch Panzee Tgmr BARE  1 Tnt1 Art1 atRE1 Evelknievel Ta1 RIRE1 Tp1 Tst1 Wis-2–1A

X03840 M94164 Y12321 AH005614 U12626 AJ000893 U96748 Z17327 X13777 Y08010 AB021265 AF039373 X13291 D85597 X53558 X52387 X63184

Arabidopsis thaliana Endovir virus Glycine max SIRE1 virus Lycopersicon esculentum ToRTL1 virus Zea mays Opie  2 virus Zea mays Prem  2 virus

AthEndV GmaSIRV LesToRV

Endovir SIRE1 TorRTL1

AY016208 AF053008 U68072

Undefined At GyDB

Arabidopsis thaliana Endovir virus Glycine max SIRE1 virus Lycopersicon esculentum ToRTL1 virus Zea mays Opie2 virus Zea mays Prem2 virus

ZmaOp2V ZmaPr2V

Opie  2 Prem  2

U68408 U41000

Unassigned Undefined At GyDB

Phaseolus vulgaris Tpv2–6 virus

Phaseolus vulgaris Tpv2–6 virus

PvuTpvV

Tpv2–6

AJ005762

Family/Genus /Phylogenetic GyDB cladea Pseudoviridae Hemivirus 1731 Copia Osser Undefined At GyDB

Pseudovirus Tork Pseudovirus

Retrofit

Undefined At GyDB

Sirevirus Sire

a

GyDB [http://gydb.org]. The clade “Chromovirus” is often considered a different genus.

b

on the Pol tree shown in Fig. 2, Ty4 and Ty2 clades form a branch which is representative of the Pseudovirus genus, Copia clade represents the genus Hemivirus, whereas Oryco and Sire clades form a group constituting the Sirevirus genus. The remaining lineages constitute small clades whose relationships remain unresolved. Finally, the clades designated as CoDi-like include four different Ty1/Copia lineages described in the diatom Phaeodactylum tricornutum. It is worth mentioning that Ty1/Copia elements in the CoDi-I clade, similar to Ty3/Gypsy chromoviruses, encode INTs containing a chromodomain at their C-terminus.

Life Cycle and Replication The life cycle of all LTR retroelements consists of four steps (Fig. 3): (1) transcription and protein synthesis, (2) RNA packaging and VLP formation, (3) reverse transcription, and (4) integration. In the first step, integrated copies of the LTR retroelement are transcribed by the cellular RNA polymerase II. The transcription initiates from a promoter located in the 50 LTR and then, the full-

660

Reverse-Transcribing Viruses (Belpaoviridae, Metaviridae, and Pseudoviridae)

Fig. 3 Life cycle of LTR retrotransposons and retroviruses. The integrated LTR retroelement is transcribed into mRNA and exported to the cytoplasm where it is translated into the Gag and Pol polyproteins that are later processed by PR into the distinct Gag and Pol domains. These proteins are subsequently assembled into a VLP together with the full LTR retroelement transcript and host tRNAs. The reverse transcription from RNA to cDNA occurs in the VLP mediated by the RT and RH. The cycle is completed when the cDNA is transferred into the nucleus and integrated as a new copy into the host genome by the INT. In general lines, the life cycle of retroviruses differs from that of LTR retrotransposons in the capability of the retroviruses to fuse the viral and host membranes in an Env-dependent manner and infect other cells. During the reverse transcription step (box to the right and below) the first intermediate of cDNA produced is the –SSS (note that plus-strand and minus-strand cDNAs are shown in blue and red, respectively). Then, RH degrades the R and U5 regions from the 50 end of the transcript and the –SSS fragment is transferred and annealed to the 30 end of the RNA. The bulk of the minus-strand cDNA is then produced while RH degrades the rest of the transcript except for the PPT. The plus-strand strong-stop is produced and RH removes the tRNA primer from the 50 end of the minus-strand cDNA. Once circularized, the plus-strand and minus-strand cDNAs are extended to generate their full-length forms. Regarding the integration examples, in the box to the left, we show: (A) The self-priming mechanism of TF1 which requires an RNA structure consisting of three duplexes (labeled as 5 bp duplex, PBS-duplex, and 11 bp duplex); (B) Additional base pairing exists between nucleotides (nt 124–130 and nt 162–156) within a 49 nt loop. The dashed line indicates that the nucleotides shown connect directly to the indicated regions of the diagram. (C) The U5-IR stem–loop of RSV. The three regions of duplex structure in the transcript of RSV are U5-IR, PBS, and U5-leader stem. These structures show considerable similarity to the self-priming RNA of TF1. Reproduced from Lin, J.H., Levin, H.L., 1998. Reverse transcription of a self-primed retrotransposon requires an RNA structure similar to the U5-IR stem-loop of retroviruses. Molecular and Cell Biology 18(11), 6859–6869. with permission from American Society for Microbiology.

Reverse-Transcribing Viruses (Belpaoviridae, Metaviridae, and Pseudoviridae)

661

retroelement mRNA is translated into the Gag and Pol polyproteins that are proteolitically processed by PR to release mature CP, NC, INT, and RT-RH proteins (Pol can be expressed as a translational fusion with Gag or through one or more ribosomal frameshifting). During the second step of RNA packaging and VLP formation, translated Pol peptides and two copies of full-length mRNA are typically encapsidated together with a specific cellular tRNA involved in priming into a VLP assembled using Gag peptides. The CP oligomerizes to form the immature capsid, while NC packages the genome. The initial VLP assembly is driven by the uncleaved Gag polyprotein, ensuring that the full-length retroelement mRNA is recruited and encapsidated, allowing the Gag-RNA-Gag interactions, contributing to the assembly of an immature Gag/ribonucleoprotein (Gag-RNP) particle. The immature VLP is subsequently maturated by proteolytic cleavage of PR between CP and NC domains. Current evidence suggests that the VLPs assembled by metaviruses, pseudoviruses and belpaoviruses might not be infectious when applied extracellularly (except for errantiviruses) but we cannot dismiss further evidence to change this interpretation. Note for instance that the vast majority of fungal RNA viruses are not infectious when applied extracellularly. Yet, they are still considered viruses. The capsids of LTR retroelements play a protective role against defense systems of the host cell during the subsequent steps of the life cycle. Note that in contrast to retroviruses which are released from the host cell to the environment, LTR retrotransposons do not have an extracellular stage in their life cycle, similar to most fungal and some plant viruses. However, high-resolution structures of recently determined immature and mature Ty3 VLPs have shown a close structural similarity between the capsids of metaviruses and retroviruses. Based on the sequence conservation of the CP and NC domains, this similarity can be also extended to all other members of the order Ortervirales. In the third step of reverse transcription, the full-length retrolement mRNA serves as a template for the synthesis of cDNA within the VLP. As also shown in Fig. 3, the cellular tRNA (packaged in the VLP) anneals to the retroelement mRNA at the PBS region that is complementary to the 30 end of the tRNA and is used by RT as a primer to synthesize the (  ) DNA chain complementary to the R-U5 zone of 50 LTR. The tRNA primes the reverse transcription of a short minus-strand species of DNA referred to as the minus-strand-strong-stop (  SSS). As this cDNA is being synthesized, RH degrades the RNA template once it is reverse transcribed. Degradation of the 50 end of the RNA releases the  SSS from the 50 end of the RNA and allows it to anneal to the R sequence at the 30 end of the same RNA or other co-packaged genome. This positions the  SSS to be extended by RT to generate the minus-strand of the LTR-retroelement. The remainder of the RNA is degraded by RH with the exception of the PPT adjacent to the 30 LTR, sequence that confers to RNA fragment resistance to RH cleavage. This 30 PPT RNA serves as the plus-strand primer for reverse transcription. Plus strand cDNA is synthetized until 50 -end of the minus strans, and the PBS sequence of tRNA is employed as a template before producing a plus strand strong-stop DNA ( þ SSS). The þ SSS is transferred to the 30 end of the minus strand previously generated and then, the tRNA primer is removed by the RH activity. The complementary sequences of PBS at the 30 ends of the þ SSS and of the minus strand generated mate with each other and a circular structure is formed. Each strand serves as a template for RT extension until the double strand DNA is full-length synthetized and LTR ends are duplicated. During minus strand synthesis most recombination events take place, when the –SSS fragment is transferred from 50 end of an RNA to R sequence at 30 end of a genetically distinct RNA. This process, together with several mutations accumulated by the low fidelity of RT, promote viral evolution and confer to the virus a better capacity to escape from host system. In contrast, other non beneficial mutations for the virus are also produced. For example, in retrovirus such as HIV several mutations in PPT sequence have been observed, what has been correlated to low infectious levels. It is worth returning to the priming mechanism to highlight that the tRNA complementarity of the PBS varies within each family and depending on the element. For instance, the PBS for errantiviruses is predicted to normally be complementary to a tRNASer, with a notable exception of the Gypsy element that presents a PBS with complementarity to a tRNALys. In contrast, plant chromoviruses normally use a tRNAMet as a primer, whereas chromoviruses of fungi and vertebrates use a selfpriming mechanism (also depicted in Fig. 3), with some exceptions such as Ty3 which is though to be primed by the tRNAMet. The self-priming mechanism of fungal and vertebrate chromoviruses has been extensively studied in elements such as TF1. According to this model, the first 11 nt of the mRNA are complementary to its PBS and act as a primer of the -SSS cDNA synthesis. The fourth step of integration takes place when the reverse transcription is completed (as also illustrated in Fig. 3(A)). During this step, the double-stranded cDNA is imported from the VLP into the nucleus and is inserted into a chromosomal target site, usually generating a 5 bp duplication of host target DNA. Integration is catalized by the retroelement INT (which belong to the superfamily of DDE INTs) in two reactions: 30 processing and strand transfer. During the 30 processing reaction, the INT removes two or three nucleotides from the 30 ends of the cDNA to position the highly conserved CP dinucleotide at the 30 end of the cDNA. In retroviruses and some LTR-retrotransposons, the tRNA primers initiate reverse transcription two or three nucleotides from the LTR and this results in the addition of deoxynucleotides at the cDNA termini that the INT subsequently removes during this reaction. In other cases, such as the TF1 LTR retrotransposons, the minus-strand primer initiates reverse transcription with no extra nucleotides added before the LTR. With respect to the strand transfer, reverse-transcribing viruses seem to direct the integration of their genoma in specific locations, suggesting that it is not a random process. This event could be in benefit of viruses, when cDNA integration takes place into particular active genes. This leads to viral clonal expansion and persistence in the infected cells. Active genes are usually located in the nuclear periphery where chromatin regions are in an open conformation. Activity of integrated LTR promoters depends on the open chromatin to make it accessible for the binding of the different transcription factors. On the other hand, integration of cDNA is able to disrupt or alter gene functions, with potential detrimental effects on the viability of host cells and, by extension, viability of the inserted LTR retroelement (which depends entirely on the well-being of the host cell). However, in

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Reverse-Transcribing Viruses (Belpaoviridae, Metaviridae, and Pseudoviridae)

the course of evolution, LTR retroelements have developed mechanisms to specifically target integration without altering the gene functions. This is primarily achieved by integration into noncoding regions, preferential targeting of heterochromatin regions (not permissive for transcription) or by association with centromeric regions. For example, Ty1 and Ty3 retrotransposons of S. cerevisiae integrate near sites of RNA polymerase III (pol III) transcription by recognizing pol III transcription complexes or chromatin states associated with pol III transcription. By contrast, TF1 recognizes certain RNA polymerase II (pol II) promoters. The great majority of Gypsy elements are located in the pericentromeric regions of the D. melanogaster genome and this location is conserved in strains from different geographic origins thus suggesting that a high number of Gypsy elements are non-mobile and were already present in the pericentromeric sequences of the common ancestor of the D. melanogaster strains. The accumulation of metaviruses such as Athila/Tat and chromoviruses in centromeric sequences is also observed in plants. Recent studies have shown that the chromodomain specifies the target site preference of chromoviral LTR retrotransposons by the recognition of characteristic chromatin modifications. These mechanisms for silencing of the viral genomes are also used to contribute to viral latency and to escape from the immune system. For instance, human immunodeficiency virus (HIV), a retrovirus that targets integration into heterochromatin regions, makes association with alphoid pericentromeric regions in different chromosomes, or integrates its genome in an orientation opposite to the host gene transcriptional readthrough.

Host Range and Evolutionary Dynamics Phylogenies tend to separate metaviruses, pseudoviruses, belpaoviruses, caulimoviruses, and retroviruses into five well defined phylogenetic clusters to which we refer as families within the order Ortervirales. Such perspective is consistent with the idea of gradual evolution and, although accurate, it is based solely on the phylogenetic signal coming from the gag and Pol polyproteins and, inherently, does not consider other biological and evolutionary phenomena that probably played a relevant role in shaping the diversity of the Ortervirales. These include: (1) modularity, the distinct genes and/or genomic regions of a virus/LTR retrotransposon usually present different rates of evolution, (2) chimerism, different events of gene recruitment, genome rearrangement and recombination that can be traced in the evolution of the Ortervirales at different evolutionary times, (3) alternance of horizontal and vertical means of transmission; while eukaryotes usually transmit their LTR retroelement communities via germ lines (i.e., by vertical means), exogenous retroviruses and caulimoviruses spread via infection (i.e., horizontally) and there is evidence that several LTR retrotransposons might have been able to protagonize some episodes of horizontal transmission, (4) clonal interference and competition with other viruses and other self genetic elements, and (5) co-evolution with eukaryotes; different cellular and molecular host environments that arose at distinct evolutionary times in plants, fungi and animals have forced the distinct orterviral communities to adapt depending on the changes in host environment. All these considerations lead to a scenario of reticulate evolution suggesting that our comprehension of the history of the Ortervirales is far from complete and could be more accurately traced as a phylogenetic network shaped by a range of changes in the cellular and molecular environments provided by eukaryotes to their retrotransposable parasites that reciprocally were also key for eukaryotic evolution. In Fig. 4, we superimpose the Pol tree previously shown in Fig. 2 over the evolutionary history of eukaryotes. For simplicity, we divide the resulting tree into three major temporal ranges to provide the reader with a system biology macroevolutionary perspective of the origin and evolution of members of the order Ortervirales undergoing three major evolutionary transitions in their eukarotic hosts. The first transition covers events from the earliest eubacterial organisms around 3500 million years ago (MYA) and the first unicellular algae to the diversification of eukaryotes into the major supergroups (approximately 2170–1950 MYA) between the Archean and the Proterozoic periods (as shown in Fig. 4). Current sequencing projects show that Metaviridae chromoviruses are present in multiple distinct algae, amoebae, chromalveolates, fungi, plants and animals. The most parsimonious hypothesis for such a wide distribution is thus that early members of the family Metaviridae have already existed before the split between plants and unikonts (1550 MYA) and that chromoviruses might constitute the oldest branch within this family. Sequencing projects also show pseudoviruses to be widespread among algae, plants, fungi and animals, suggesting that pseudoviruses co-existed with metaviruses during this transition. Thus, the ultimate origins of the ortervirads likely took place at the onset of eukaryogenesis and involved combination of gag and pol genes with the LTR features to raise an ancestral LTR retrotransposons from which pseudoviruses and metaviruses diverged into two independent but stable lineages following the genomic relocation of the int gene that distinguishes the two families. The second transition covers a period of 1330–380 MYA from the Proterozoic to the Phanerozoic, during which eukaryotes underwent key events of diversification, such as the divergence between primitive animal phyla (e.g., Porifera, Cnidaria, Ctenophora) and the more complex animals, including the emergence of eumetazoans, the Cambrian explosion, the divergence between arthropods and priapulids, the rise of vertebrates, the emergence of land plants and the establishment of plant-fungal interactions. Respecting the Metaviridae, although chromoviruses are present in vertebrates, such as fish, in amphibians and in amniotes, they are absent in the known genomes of cnidarians, mollusks, protostomes and basal deuterostomes. However, we find others metavirus lineages. For example, Mag and other elements, from which we have apparently emerged before the split of bilaterians into protostomes and deuterostomes (761–531 MYA). It is currently unclear whether chromoviruses were driven to extinction in protostomes, echinoderms and urochordates or were horizontally transmitted from plants or fungi to vertebrates. Horizontal transmission seems to be a good explanation for vertebrate chromoviruses, taking into account the strong similarity to their fungal relatives, but if so, such an event might be as ancient as the split of osteichthyes into actinopterygians and sarcopterygians

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Fig. 4 Evolutionary history of LTR retroelements superimposed over the evolutionary history of eukaryotes. The tree of Ty3/Gypsy LTR retroelements is traced in blue; Ty1/Copia elements yellow; Bel/Pao black; Retroviridae violet; and Caulimoviridae green. This representation defines the natural origin and distribution of the Ortervirales constrained by the evolutionary history of eukaryotes and also includes information about diverse reticulate events in the evolutionary history of LTR retroelements. Host ranges are indicated in red. The framework has been divided into three major eukaryotic transitions (colored in gradient of gray) to interpret the distribution of the distinct LTR retroelement families and communities on the basis of the most likely ages of their hosts. GYA refers to billions (giga) years ago. The distinct evolutionary paths traced in this figure only reflect the host distributions of the distinct orterviral families and their likely times in those hosts, not extinction events. For example, if a host range is represented in the second transition but not in the third one, it indicates that that orterviral family is inhabiting the genomes of those hosts from that age to the present time. Adapted from Llorens, C., Muñoz-Pomer, A., Bernad, L., Botella, H., Moya, A., 2009. Network dynamics of eukaryotic LTR retroelements beyond phylogenetic trees. Biology Direct 4, 41. with permission from BioMed Central, part of Springer Nature.

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(410–415 MYA). Regarding the Pseudoviridae, sequencing projects reveal pseudoviruses to be also widespread in cnidarians, arthropods and deuterostomes. However, to our knowledge, there is no evidence of pseudoviruses in platyhelminthes and nematodes, suggesting that metaviruses underwent more explosive diversification in metazoans compared to pseudoviruses. As for the Belpaoviridae, their wide distribution in metazoans and their absence in fungi and plants suggests that the origin of belpaoviruses probably predates the split between primitive animals and higher eumetazoans (1300–380 MYA). With respect to the emergence of bona fide retroviruses (family Retroviridae), sequencing projects reveal three Retroviridae lineages including gammaretroviruses, epsilonretroviruses and spumaretroviruses to overlap with chromoviruses in fishes, amphibians and sauropsids thus suggesting that the first Retroviridae emerged in marine vertebrates before the Osteichthyes split into the actinopterygians and the sarcopterygians. We can thus assume that metaviruses and pseudoviruses have radiated into a variety of lineages, with belpaoviruses and the Retroviridae emerging during this second transition. The third eukaryotic transition (range in light gray within Fig. 4) covers a period from the Paleozoic era (330 MYA) to the present, a timeframe which included the origin of first gymnosperms (360–248 MYA), the split of amniotes into sauropsids and synapsids (330–312 MYA), the diversification of winged insects (380–325 MYA) and the origin of flowering plants (160–90 MYA). New levels of complexity and ecosystems probably accompanied the aforementioned events in the evolutionary history of eukaryotes, activating parallel radiations of LTR retrotransposons as well as the emergence of new viral and retroviral forms to colonize the new host enviroments such as those of synapsids (mammals and their extinct relatives). Consistent with this perspective, sequencing projects revealed that metaviruses and retroviruses overlap in sauropsids but there is no trace of functional metaviruses, pseudoviruses or belpaoviruses in mammals. The only orterviral family known to spread in mammals is the family Retroviridae, albeit several host genes, most notably including the neuronal gene Arc which evolved from the CP domain of domesticated metaviruses, can also be detected in the mammalian genomes. This suggests that while metaviruses were co-opted and driven to extinction by synapsids, the retroviruses co-evolved with their hosts, recruiting new accessory genes to adapt their infective means of transmission to the new and more complex mammalian host environments. In parallel, mammals evolved more sophisticated mechanisms of innate immunity to combat retrovirues. Retrovirus accessory genes and mammalian mechanisms of innate immunity will be best understood when considered as the join products of macro-evolutionary conflicts played out over a geological scale. During this period, other retroviral forms, such as errantiviruses, also emerged from the Metaviridae family in winged insects such as Drosophila, albeit the lower biological complexity of insects (in comparison to that of mammals) allowed the diversification of the metaviruses in other lineages as well as the expansion of pseudoviruses and belpaoviruses in these and other insects. A very similar perspective is found in plants. In this particular, gymnosperms and angiosperms show massive radiations of Athila/Tat-like metaviruses and LTR retrotranposons to overlap with their chromoviral counterparts and with pseudoviruses which also show some examples of protoretroviral forms, such us some elements belonging to Sire clade. Thus, pro-retroviruses might have emerged in plants before the divergence of gymnosperms and angiosperms. On the other hand, phylogenetic and comparative genomic analyses of gag-pol regions also support the evolution of caulimoviruses from the Metaviridae (note in Fig. 2 how the Caulimoviridae are rooted within the Metaviridae). However, the genomes of caulimoviruses additionally present other genes, such as the one encoding for the movement protein, which are necessary for the viral life cycle and transmission of these viruses in plants. While it is not yet clear whether the ancestors of caulimoviruses were inhabitants of plants or insects (the latter act as mechanical vectors), the origin of caulimoviruses was probably concomitant with the emergence of insects and flowering plants (130–90 MYA). These two events probably gave an opportunity to metaviruses to explore other gene combinations and viral strategies from which caulimoviruses probably emerged (recruiting new genes or recombining with other viruses).

Impact and Relationships of LTR retrotransposons and Retroviruses With Their Hosts Until recently, LTR retrotransposons were considered to be merely molecular parasites whose sole aim is to self-replicate in the host genomes and be transmitted from one host generation to another. Current evidence suggests, however, that the co-evolutionary history of LTR retrotransposons and their hosts is much more complex than previously though. Retroelements with and withouth LTRs are mutagens, whose activities might lead to diverse genetic alterations, including: (1) gene knockouts, (2) modifications of gene expression, (3) epigenetic reprogramming, (4) insertions and deletions, (5) structural rearrangements, and (6) segmental duplications. To avoid potential damages to their hosts, LTR retroelements have developed specific mechanisms to integrate the retrotranscribed cDNAs into non-coding genomic regions, as discussed above. The mechanisms ensuring specific insertion of LTR retrotransposons are the basis of a co-evolutionary history that has permitted these elements to successfully proliferate in their eukaryotic hosts. In sharp contrast, the host may benefit from the new insertions of retroelements because they are a constant source of new sequence substrate. In plants, for instance, the metaviruses and the pseudoviruses constitute the major fraction of all plant transposons. This fraction is, in fact, correlated with the genome size in some groups of plants, such as angiosperms, thus providing a potential explanation why the amount of DNA in a haploid genome does not match with the complexity of these taxa (i.e., the C-value paradox). The situation is similar in other plant, fungal and animal taxa across the Tree of Life. In humans, for example, endogenous retroviruses (HERVs) constitute approximately 8% of the human genome. If we also consider the rest of retrotranscribing elements (with and without LTRs) and related sequences, the fraction reaches up to 45% of the human genome.

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The contribution of new sequence susbstrate as a consequence of the mobilization of retrotransposable elements can help the host organism to explore new levels of complexity. Integration of multiple copies of LTR retroelements and other mobile genetic elements into a particular chromosomal region may result in repeat clusters, which are hotspots for chromosomal rearrangements and segmental duplications due to erroneous homologous recombinations during mitosis and meiosis. Understanding these clusters is still lacking but they appear to provide for genome plasticity to their hosts. Notably, several fungal pathogens, such as Leptosphaeria maculans or Verticillium dahlia, have been able to take advantage of these plastic loci for evolving increased virulence and pathogenesis, and adapt better to changing environments than members of these species lacking the corresponding clusters. The sequences contributed by retroelements as a consequence of their mobilization is not only a source of new substrate but also of variability from which new genetic combinations and functions may emerge and be co-opted by the host, giving rise to new genes and regulatory elements, able to rewire the gene expression networks of eukaryotes. In parallel, a functional equilibrium must exist to regulate the activitities of retrotransposable elements, to avoid the disruption of the normal regulatory mechanisms of gene expression and pathways. The loss of this equilibrium may have severe consequences, leading to developmental disorders and diseases. In humans, for example, the activation and mobilization of HERVs and other reverse-transcribing elements have been associated with cancer, autoimmune diseases, genomic instability and neurodegenerative diseases, including Tau pathology and Alzheimer. In D. melanogaster, the mobilization of Gypsy and Copia elements (Metaviridae and Pseudoviridae, respectively) has also been correlated with the activity of Tau, rendering D. melanogaster an excellent model to explore the molecular mechanisms behind diseases associated not only with neurodegenerative processes but also with tumorigenesis and tumor immunity. The relationship between cancer and retroelements is, in fact, object of current research both in humans and in other taxa. For instance, some reverse-transcribing viruses, such as retroviruses, have shown strong preference to integrate into genes encoding cellular transcription factors involved in cell growth. BACH2 and MKL2 are two of the proteins encoded by the genes where retrovirus integration has been observed. Integration into these genes alters their functions and increases the life-time and survival of the host cell. In particular, rearrangements of these genes have been associated to proto-oncogenic activities. In other taxa, like Mya arenaria, a soft-shelled clam characteristic of many areas in the North Atlantic, a fatal leukemia-like disease of unknown origin has been reported to be catalized by the expression of Steamer, an LTR retrotransposon of the Metaviridae family. Therefore, the proper regulation of gene activation and silencing has an extreme importance for the co-existence of LTR retroelements and the hosts they inhabit. To this end, host genomes and transposable elements may both exist in variable DNA methylation states whose differential transcription is modulated by distinct types of regulatory elements, such as enhancers, silencers and chromatin insulators. Of these, chromatin insulators play a central role in the proper regulation of gene expression and organization of chromosome architecture. A well studied example of chromatin insulator in D. melanogaster is the insulator of Gypsy-like errantiviruses, that is able to modulate enhancers and promoters. Uncontrolled expression of Gypsy may result in insertions of this retrovirus with negative consequences on the normal expression patterns of the host genes. To keep Gypsy under control, Drosophila implements an RNA interference (RNAi) system consituted by Piwi and flamenco, a locus that is the source of antisense small RNAs interacting with Piwi to repress the expression and infective properties of Gypsy. The represive role of flamenco in Drosophila also extends to other metaviral and pseudoviral elements whose expression is also modulated by this system.

Concluding Remarks Viruses and mobile genetic elements oftentimes represent different faces of the same coin. This is the case of the members of the families Metaviridae, Pseudoviridae, and Belpaoviridae which may exist as LTR retrotransposons and as potential or even true viruses that share origins and evolutionary history with other reverse-transcribing viruses, such as vertebrate retroviruses and plant caulimoviruses. In recognition of this evolutionary connection, the ICTV has recently grouped the Belpaoviridae, Metaviridae, Pseudoviridae, Retroviridae, and Caulimoviridae families into a single viral order, the Ortervirales. In this article, we have reviewed what is currently known about metaviruses, pseudoviruses and belpaoviruses, with the aim to provide the reader with a systems biology perspective on these viruses and discuss the deep evolutionary history of the Ortervirales. Although retroviruses and caulimoviruses have been extensively studied, our understading of the real impact of reverse-transcribing viruses in eukaryotes is still in its infancy. Due to this reason, the members of the families Metaviridae, Pseudoviridae, and Belpaoviridae present excellent opportunities not only to investigate the origin and evolution of reverse-transcribing viruses in plants, fungi and animals but also to investigate their roles as inductors of disease, regulatory elements and vectors of complexity and evolution in plants, fungi and animals.

Further Reading Burns, K.H., 2017. Transposable elements in cancer. Nature Reviews Cancer 17 (7), 415–424. de la Chaux, N., Wagner, A., 2011. BEL/Pao retrotransposons in metazoan genomes. BMC Evolutionary Biology 11, 154. Dodonova, S.O., Prinz, S., Bilanchone, V., Sandmeyer, S., Briggs, J., 2019. Structure of the Ty3/Gypsy retrotransposon capsid and the evolution of retroviruses. Proceedings of the National Academy of Sciences of the United States of America 116 (20), 10048–10057. Eickbush, T., Boeke, J.D., Sandmeyer, S.B., Voytas, D.F., 2011. Metaviridae. In: King, A.M.Q., Adams, M.J., Carstens, E.B., Lefkowitz, E.J. (Eds.), Virus Taxonomy: Classification and Nomenclature of Viruses: Ninth Report of the International Committee on Taxonomy of Viruses. San Diego: Elsevier Academic Press, pp. 457–466. Gao, X., Hou, Y., Ebina, H., Levin, H.L., Voytas, D.F., 2008. Chromodomains direct integration of retrotransposons to heterochromatin. Genome Research 18 (3), 359–369.

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Geering, A.D.W., 2014. Caulimoviridae (plant pararetroviruses). eLS. Chichester, United Kingdom: John Wiley & Sons, Ltd. Greenig, M., 2019. HERVs, immunity, and autoimmunity: Understanding the connection. PeerJ 7, e6711. Grzebelus, D., 2018. The functional impact of transposable elements on the diversity of plant genomes. Diversity 10 (2), 18. Guo, C., Jeong, H.H., Hsieh, Y.C., et al., 2018. Tau activates transposable elements in Alzheimer’s disease. Cell Reports 23 (10), 2874–2880. Havecker, E.R., Gao, X., Voytas, D.F., 2004. The diversity of LTR retrotransposons. Genome biology 5 (6), 225. Howlett, B.L., Lowe, R.G.T., Marcroft, S.J., van de Wouw, A.P., 2015. Evolution of virulence in fungal plant pathogens: Exploiting fungal genomics to control plant disease. Mycologia 107 (3), 441–451. Katzourakis, A., Gifford, R.J., Tristem, M., Gilbert, M.T., Pybus, O.G., 2009. Macroevolution of complex retroviruses. Science 325 (5947), 1512. Krupovic, M., Blomberg, J., Coffin, J.M., et al., 2018. Ortervirales: New Virus Order Unifying Five Families of Reverse-Transcribing Viruses. Journal of Virology 92 (12), e00515–e00518. Krupovic, M., Dolja, V.V., Koonin, E.V., 2019. Origin of viruses: Primordial replicators recruiting capsids from hosts. Nature Reviews Microbiology 17 (7), 449–458. Krupovic, M., Koonin, E.V., 2017. Homologous capsid proteins testify to the common ancestry of retroviruses, caulimoviruses, pseudoviruses, and metaviruses. Journal of Virology 91 (12), e00210–e00217. Lin, J.H., Levin, H.L., 1998. Reverse transcription of a self-primed retrotransposon requires an RNA structure similar to the U5-IR stem-loop of retroviruses. Molecular and Cell Biology 18 (11), 6859–6869. Llorens, C., Muñoz-Pomer, A., Bernad, L., Botella, H., Moya, A., 2009. Network dynamics of eukaryotic LTR retroelements beyond phylogenetic trees. Biology Direct 4, 41. Metzger, M.J., Reinisch, C., Sherry, J., Goff, S.P., 2015. Horizontal transmission of clonal cancer cells causes leukemia in soft-shell clams. Cell 161 (2), 255–263. Pelisson, A., Mejlumian, L., Robert, V., Terzian, C., Bucheton, A., 2002. Drosophila germline invasion by the endogenous retrovirus gypsy: Involvement of the viral env gene. Insect Biochemistry and Molecular Biology 32 (10), 1249–1256. Sarot, E., Payen-Groschêne, G., Bucheton, A., Pélisson, A., 2004. Evidence for a piwi-dependent RNA silencing of the gypsy endogenous retrovirus by the Drosophila melanogaster flamenco gene. Genetics 166 (3), 1313–1321. Wei, W., Brennan, M.D., 2001. The gypsy insulator can act as a promoter-specific transcriptional stimulator. Molecular and Cellular Biology 21 (22), 7714–7720.

Relevant Websites http://gydb.org Gypsy Database 2.0. https://www.girinst.org/repbase Repbase.GIRI.Genetic Information Research Institute. http://urgi.versailles.inra.fr/repetdb RepetDB: Home. INRA-URGI. http://repeatexplorer.org RepeatExplorer. https://talk.ictvonline.org/ictv-reports/ictv_9th_report/reverse-transcribing-dna-and-rna-viruses-2011/w/rt_viruses Reverse Transcribing DNA and RNA Viruses. ICTV 9th Report (2011).

Rice Tungro Disease (Secoviridae, Caulimoviridae) Gaurav Kumar, Fauzia Zarreen, and Indranil Dasgupta, University of Delhi, New Delhi, India r 2021 Elsevier Ltd. All rights reserved. This is an update of R. Hull, Rice Tungro Disease, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00714-7.

Glossary Agro-inoculation Infection of a plant by injecting Agrobacterium containing a modified Ti plasmid into which an infectious copy of the viral genome has been inserted. In the case of pararetroviruses, this infectious copy comprises the sequence that can express the more-than-full-length RNA transcript required for the reverse transcription stage of viral replication. Pararetrovirus A virus that replicates by reverse transcription but differs from retroviruses in that it encapsidates the DNA phase of replication and that virus replication does not require integration of the viral genome into the host chromosome. RNA Interference (RNAi) RNAi is an evolutionarily conserved sequence-specific natural defense mechanism in plants against transposable elements and intracellular

pathogens such as viruses, targeting the cellular or viral double-stranded RNA (dsRNA). In this process non-coding small RNAs degrade viral and cellular transcripts, resulting in post-transcriptional gene silencing. These small RNAs are formed as a result of cleavage or slicing of dsRNA or RNA having strong secondary structures by host ribonucleases and are called small interfering RNA and microRNA. Semi-persistent transmission Virus-vector interaction in which the vector acquires the virus rapidly upon feeding and the acquisition period is relatively short, typically less than a week. The virus is believed to interact with components of the vectors’ mouthparts or foregut. Silencing suppressor Viral proteins that counteract host RNAi defense by inhibiting key steps of cellular RNAi pathways.

Introduction Rice tungro disease is caused by a complex of two viruses, Rice tungro bacilliform virus (RTBV), which is a member of the family Caulimoviridae and Rice tungro spherical virus (RTSV), a member of the family Secoviridae. The viral complex is transmitted by several species of green leafhoppers in a semi-persistent manner. The genomes of RTBV and RTSV are double-stranded DNA and single-stranded RNA respectively and are replicated using very different mechanisms. Genetic resistance against RTBV has not been characterized, but for RTSV, recessive resistance has been mapped to a gene encoding a translational initiation factor. We are reviewing here the status of knowledge for the disease and for both viruses.

Rice Tungro Disease The characteristic symptoms of rice tungro disease had been reported from several South-East Asian countries for many years before they were recognized in 1963 to be of viral etiology. During and since the 1960s, there have been several severe outbreaks of the disease mainly associated with the major agronomic improvements in rice cultivation resulting from the ‘green revolution’. The disease is found in all countries of South and South-East Asia and in the South and South-East of China (Fig. 1(A)). The importance and distribution of the disease are evidenced by the variety of local names for it: accep na pula (Philippines), mentek (Indonesia), penyakit merah (Malaysia), yellow-orange leaf (Thailand), leaf yellowing (India). In the early 1990s, it was estimated that tungro caused annual losses in excess of $1.5 billion without taking account of the cost of insecticide to control the vector. In parts of India, the disease causes losses of up to 50% of the expected crop yield. In most of the rice cultivars, tungro infection is manifested by stunting and reduced tillering, yellow to orange-yellow leaf discoloration beginning from the leaf tip and extending towards leaf base, delay in flowering, failure of panicle emergence and rust-colored patches or twisted leaves in some cases. Infection to a young rice plant is more severe with mottled appearance and interveinal chlorosis as compared to an old rice plant having rust-colored specks of varying sizes. A typical view of a paddy field showing tungro symptoms is shown in Fig. 2. The disease symptoms are often mild or almost absent in tolerant rice varieties. The disease is transmitted in a semi-persistent manner by several leafhopper species with the rice green leafhopper (GLH), Nephotettix virescens (Fig. 3), being the most important vector. It is also transmitted by Nephotettix cincticeps, Nephotettix nigropictus, Nephotettix malayanus and Recilia dorsalis; these other vectors may be of relative importance in certain situations. The distribution of the disease is circumscribed by the distribution of N. virescens (Fig. 1(A)). It was not until 1978 that it was recognized that tungro is caused by a complex of two viruses, Rice tungro bacilliform virus (RTBV) and Rice tungro spherical virus (RTSV). RTBV alone causes severe disease symptoms in the host but it cannot be transmitted by GLH in the absence of RTSV. RTSV, on the other hand, can be transmitted independently by GLH (Fig. 4) but causes only mild stunting

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Fig. 1 (A) Distribution of rice tungro disease in Asia and the circle shows the distribution of the main rice tungro disease leafhopper vector. (B) Phylogenetic trees of complete sequences of RTBV (top) and RTSV (bottom). The percentage of identity relative to the first sequence is indicated.

Fig. 2 Field symptoms of rice tungro disease.

Fig. 3 The rice green leafhopper, Nephotettix virescens, major vector of the rice tungro disease.

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Fig. 4 Rice tungro virus transmission. (A) RTBV cannot be transmitted on its own by GLH; (B) RTSV can be transmitted on its own by GLH but cause very less or negligible symptoms to appear; (C) An RTSV harboring GLH is able to transfer RTBV alone and the presence of both RTBV and RTSV causes visible symptoms.

symptoms. The dependence of RTBV upon RTSV for transmission suggests the presence of a “helper component” in RTSV or RTSV-infected cells. The origin, nature and functional properties of the above “helper component” remains elusive.

Rice Tungro Bacilliform Virus Taxonomy RTBV is a dsDNA pararetrovirus belonging to a mono-specific genus, Tungrovirus, in the family Caulimoviridae.

Transmission and Host Range GLH transmits RTBV in a semi-persistent manner in rice only in the presence of RTSV. However, cloned RTBV DNA can be transmitted to rice plants if inoculated using agrobacterium, even in the absence of RTSV. This process is known as agro-inoculation. Till now, this method has not been used to investigate the host range of RTBV.

Structure RTBV consists of bacilliform particles of about 130 nm length (though there can be longer ones in some isolates) and 30 nm diameter (Fig. 5). The particle structure is based on a T ¼ 3 icosahedron cut across its threefold axes, the tubular portion being made up of rings of hexamer subunits with a repeat distance of about 10 nm.

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100 nm

Fig. 5 Particles of RTBV (long particles) and RTSV (spherical particles) negatively stained in uranyl acetate. The bar indicates 100 nm.

Fig. 6 The genome organization of RTBV and the different transcripts associated with it. The innermost circle represents the double-stranded 8 kb DNA. The numbers inside the circle represent the position of the nucleotide (  1000) on the genome. The approximate sites of the two discontinuities D1 and D2 are marked. The position of the four open reading frames, ORF I, ORF II, ORF III and ORF IV, are represented as arcs. The proteins encoded by ORF III polyprotein have been marked. MP: Movement protein; CP: Coat protein; PR: Protease and RT/RNase H: Reverse transcriptase/ribonuclease H. The arc in orange represents pre-genomic RNA. The blue-green arc represents the spliced ORF IV RNA. The dotted arcs represent the intronic region of the transcript.

Genome The RTBV genome is a double-stranded circular DNA of an approximate size of 8 kb (Fig. 6). It has high resemblance with badnaviruses and caulimoviruses, in having site-specific discontinuities one on each strand of the double stranded circular genome, a poly-protein analogous to retroviral gag-pol and involvement of an RNA intermediate in replication. The RTBV genome potentially has four Open Reading Frames (ORFs) viz., ORF I (P24), ORF II (P12), ORF III (P194) and ORF IV (P46) named according to the sizes of the proteins they encode. The transcription of RTBV genome is asymmetric with one strand (containing the minus-strand priming site) possessing all the coding capacity. The ORF I, II and III are compactly arranged and have a coinciding start-stop signal (ATGA). The ATG of this signal acts as a start codon for downstream ORFs and TGA of this signal codes for stop codon for the upstream ORF. The third ORF encodes for a poly-protein, which is cleaved subsequently to produce four different proteins. A short non-coding intergenic region separates the ORF III from ORF IV, which is expressed from a

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spliced RNA. ORF I and ORF IV are also separated by a long intergenic region which includes the single RTBV promoter, the signal for poly-adenylation, various short ORFs and the splice donor site.

Functions of Proteins The 24 KDa protein (P24) encoded by ORF I is poorly characterized till date although it is reported to play some role in viral particle assembly. P24 interacts with the host D1 protein, associated with photosystem II, which may affect the host photosynthetic machinery, leading to the appearance of tungro-associated symptoms. The ORF II encodes for a 12 KDa protein (P12), which is shown to interact both in vivo and in vitro with the coat protein domain of the ORF III encoded poly-protein and plays a role as a protein scaffold in the assembly of the virus capsid. The longest ORF of RTBV is the ORF III, which encodes for a polyprotein having four clear-cut domains for movement protein (MP), coat protein (CP), aspartate protease (PR) and reverse transcriptase/ribonuclease H (RT/RNase H) arranged clockwise on the genome map. The presence of an ORF IV downstream of the gag-pol analog of retroviruses is the single feature that differentiates RTBV from other badnaviruses. ORF IV possesses an unconventional isoleucine (ATT) start codon rather the canonical methionine ATG codon and is expressed as a spliced product in association with small ORF I. ORF IV encodes for a 46.2 kDa (P46) protein, which acts as a viral suppressor of RNA silencing, affecting the cell-to-cell spread of the RNA silencing signal. The viral coat protein comprises one major species of Mr 37 kDa; this may be processed to give a product of Mr about 33 kDa and possibly others. The molecular mass of the coat protein has been measured by mass spectrometry and the N- and C-terminal sequences have been determined. This protein contains two specific amino acid sequence motifs, CXCX2CX4HX4C characteristic of reverse transcribing virus gag proteins and CX2CX11CX2CX4CX2C unique to badnaviruses and tungroviruses.

Genome Variability There are 13 complete nucleotide sequences of RTBV available in databases till date, four from India and nine from South-East Asia. The genome sizes of the South-East Asian isolates are slightly more (B100 bp) as compared to the Indian isolates. The molecular as well as the nucleotide and amino acid sequence based phylogenetic analyzes of these RTBV isolates have now grouped them into several clusters; those from India forming a distinct cluster (Indian strains), whereas the rest are more diverse (Fig. 1(B)). The Indian strains of RTBV showed divergence from the rest due to several insertions, deletions and substitutions, mostly in the intergenic regions. A replication mechanism with the absence of proof-reading may be a plausible reason for the slight variations among these isolates. The sequence variability of RTBV genomes is similar to those of badnaviruses, such as banana streak virus, dioscorea bacilliform virus and cacao swollen shoot virus. For badnaviruses, virologists consider 80% as the similarity threshold to differentiate species. If this threshold is applied to RTBV, 7 species would exist among the 13 complete genome sequences (Fig. 1(B)), including three in the Philippines alone. Therefore, we would have a single disease, the Rice Tungro Disease, associated with several different viruses, a situation very similar to badnaviruses. The situation is completely different for RTSV, where all the six complete sequences from three countries (Philippines, Malaysia and India) are within 90% similarity or above (Fig. 1(B)).

Virus-Host Relationships Host plants, upon infection with viruses, launch an RNA-based defense response against the virus, which leads to degradation of the viral transcript into 21–24 nucleotide fragments, which are called viral si (small-interfering) RNAs. This degradation is sequence-specific and viral siRNAs initially formed by the recognition of RTBV transcript for degradation, leads to an amplification of further degradation in a sequence-specific manner, resulting in the accumulation of copious amounts of siRNAs from RTBV in infected rice plants. This finally results in a gradual fall in RTBV levels in a few week’s time. Certain regions of the RTBV genome, namely the portion immediately prior to ORF I, the 50 -untranslated region (50 UTR), gives rise to a large amount of siRNA, as compared to other regions of the genome. Despite the production of large quantities of siRNAs, the RTBV transcripts largely escape degradation because the 50 UTR folds into a strong degradation-resistant secondary structure. This could be a new type of escape employed by RTBV against host defense, not generally seen in other viruses.

Control There are three major approaches to controlling tungro; by the manipulation of various cultural practices such as planting time, having a crop-free period, and seedbed protection; invasion of young susceptible rice plants by viruliferous GLH can be avoided. Spraying insecticide can control the leafhoppers but this has not proved to be particularly effective. There have been various programs to breed resistance to tungro but most have proved of limited success. The International Rice Research Institute Philippines, has screened more than 40,000 accessions of rice for resistance to tungro viruses and have found two basic types of resistance, that to the vector and that to the virus. Since RTBV and RTSV occur together in the plant, breeding efforts have not been directed against exclusively RTBV or RTSV, but both together. Resistance to the vector has also been used, but has proved to be not very durable.

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In addition to the breeding-based approach, transgenic strategies have also been employed, targeting RTBV. With the development of the concept of pathogen derived resistance (PDR), in 1980s, there has been several attempts to confer resistance against RTBV through generation of transgenic plants containing virus-derived genome fragments or full-length genes. Both proteinmediated and RNA-mediated strategies of PDR have been used for generating virus resistant transgenic plants. The movement protein gene, the coat protein (CP) genes, protease or the viral replicase genes have been exploited for initiating successful PDR. The proteins or their parts act as a dominant negative competitor during the infection of virus. RNA-based approaches have been used, using the concept of RNA-interference (RNAi). Rice plants expressing an RNAi construct against RTBV ORF IV have been produced, which show only mild symptoms upon GLH-mediated inoculation of both RTBV and RTSV. This resistance has been recently diversified into several popular rice varieties of southern India by marker-assisted breeding and many of the progenies have acquired the resistance along with the transgene, which is characterized by almost 1000-fold fall in the titers of RTBV upon GLH-mediated inoculation, when compared to the titers seen in non-transgenic rice plants of the same variety. The above plants do not show any fall in RTSV titers. A similar approach has also been used to generate transgenic rice plants expressing RNAi constructs targeting both RTBV and RTSV simultaneously, which, as expected, show resistance against both the viruses. However, it needs to be emphasized that all the work involving transgenic rice plants described above, have been conducted under controlled laboratory or green house conditions and are yet to be validated in the field.

Biotechnological Applications Viral promoters, such as CaMV 35S, find wide application as expression modules for heterologous genes in plants. The potential of RTBV promoter, which has a similar overall location in the viral genome as CaMV 35S, has been explored. RTBV promoter shows tissue-specific activity, specifically in the phloem, in transgenic rice plants generated with a construct wherein a reporter gene is driven by the upstream sequences of the RTBV ORF I taken from a Philippine isolate. Further work with an Indian isolate of RTBV identified a complex arrangement of positive and negative regulatory elements in the promoter region, some of which were located downstream to the transcription start site. These constructs have potential use in driving heterologous gene expression in transgenic rice plants, and possibly in other monocot species. Transient gene silencing in plants can be achieved by the use of viral vectors, utilizing the phenomenon of RNAi. This phenomenon is termed Virus Induced Gene Silencing or VIGS. RTBV genome has been modified suitably to develop a VIGS system for rice. Using this system, any rice gene can be silenced in a transient manner and its effect can be studied, giving the investigator a glimpse of its function in the plant. This approach has important applications in revealing gene functions in the rice plant.

Rice Tungro Spherical Virus Taxonomy RTSV belongs to the genus Waikavirus, in the family Secoviridae, in the order Picornavirales.

Transmission and Host Range RTSV appears to have a restricted host range limited to members of the Poaceae and Cyperaceae (Echinochloa crus-galli, E. glabrescens, E. colona, Leptochloa chinensis, Leersia hexandra, Oryza sativa, Panicum repens, Cyperus rotundus); RTSV causes few, if any, symptoms in these hosts.

Structure RTSV has isometric particles having a diameter of 30 nm (Fig. 5). The virus is limited to vascular tissues where it is restricted to the phloem cells. Virus particles are scattered in the cytoplasm or are embedded in non-enveloped electron-dense granular inclusions. Particles also occur as crystalline aggregates often in vacuoles. Small vesicles containing fibers are found in the cytoplasm, usually along the wall of infected cells.

Gene Functions The capsid is made up of three coat protein species (CP1, CP2, and CP3) that are apparently in equimolar amounts, and is thought to have a similar T ¼ 1 icosahedral architecture to that of picornaviruses. Identification of cleavage sites by N-terminal sequencing shows that CP1 is 22.9 kDa and CP2 is 22.3 kDa; the C-terminus of CP3 has not been identified. In Western blots of crude extracts from infected plants, an antiserum to CP3 identifies a band at 33 kDa and several in the range 40–42 kDa; the nature of these larger bands is unknown. The genome is translated into a poly-protein, which is processed to give various products including the three coat protein species, a putative type III helicase, a cysteine or 3C-type protease, and an RNA-dependent RNA polymerase at its 30 end. Functions of other products have not yet been determined. No functions have been ascribed to the products of the 30 short ORFs and there is no evidence that these are expressed.

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Genome Diversity Unfortunately, there is not a lot of sequence information available for RTSV, there are however 6 complete genome sequences originating from three countries and all sequences are 90% similar or greater (Fig. 1(B)), indicating that they most probably all belong to the same species.

Genome Each RTSV particle contains one molecule of single stranded positive-sense RNA of about 12.2 kbp. The RNA encodes a large poly-protein of about 393 kDa and has two short ORFs at the 30 end (Fig. 7). A total of six nucleotide sequences, representing the full RTSV genome are available in the NCBI Nucleotide database (June 2019) of which three are from India, two from Philippines and one from Malaysia (Fig. 1(B)). RTSV strain Vt6 overcomes the resistance to the type strain in rice cultivar TKM6. Several serological variants of RTSV have been reported. Molecular techniques reveal much micro-variation in the sequences encoding CP1 and CP2 but there is no evidence for major geographic strains that reflect those of RTBV. Comparatively speaking, RTSV is much more conserved than RTBV, when sequences available from various regions of South and South-East Asia are compared.

Virus-Host Relationships Recently, the virus-derived siRNAs have been studied in rice infected with both RTBV and RTSV using high throughput sequencing. Despite a reasonably large quantity of such siRNAs accumulating in the infected plants against RTBV, the accumulation of the corresponding siRNAs against RTSV is extremely low. This study points towards possibly an uncharacterized mechanism of escape from the siRNA-mediated resistance response of the plant by RTSV.

Control Natural resistance in rice against RTSV is conferred as a recessive trait controlled by the translation initiation factor 4-gamma gene (eIF4G). Three residues of eIF4G, Y1059 V1060 V1061 are known to be associated with resistance against RTSV. Using CRISPR/Cas9 system in the RTSV-susceptible variety IR64, mutations in eIF4G, have been generated to confer resistance against RTSV. These mutations were located adjacent to the YVV residues and were successfully transmitted to the next generations. However, these approaches have yet to be used at the field level to effectively control RTSV.

Concluding Remarks RTBV and RTSV are a unique combination of unrelated viruses, which have come together to cause the Tungro Disease, one of the most important crop diseases. Tungro has the potential to cause major disruptions in the production of rice in south and South-East Asia, thereby posing a threat to food security in this part of the World. The functions of the proteins encoded by the two viruses are yet to be worked out in detail. Recent research has revealed a novel role for RTBV P24 as an RNAi suppressor, hence possibly acting as a virulence factor for the virus. Despite the above RNAi suppressor being active, RNAi-mediated resistance has been demonstrated against RTBV and RTSV in rice under experimental conditions. Such studies can have

Fig. 7 The genome organization of RTSV. The polyadenylated RNA is indicated as a single line and the coding regions as boxes. The nucleotide and amino acid positions of the different coding regions are indicated above. The amino acids present at the cleavage sites are indicated below the boxes. The position of the TGA stop codon in sORF II has been indicated by red dotted line. The 3473 amino acids long polyprotein is indicated by spiraled line below the RTSV genome. VPg: Viral protein of the genome; P1: Leader protein 1; CP1–3: Coat protein 1–3; NTP: Nucleotide triphosphate binding protein; Pro: Protease; Pol: RNA dependent RNA polymerase; sORF: short ORF.

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important applications in controlling the viruses and hence the Tungro Disease. Other important aspects of the viruses, namely the semi-persistent transmission of the viral complex by the vector GLH and the potential helper function mediated by RTSV for the transmission of RTBV are, however, still shrouded in mystery.

See also: Caulimoviruses (Caulimoviridae). Plant Antiviral Defense: Gene-Silencing Pathways. Plant Resistance to Viruses: Engineered Resistance. Secoviruses (Secoviridae)

Further Reading Anjaneyulu, A., Satapathy, M.K., Shukla, V.D., 1994. Rice Tungro. New Delhi: Oxford and IBH Publishing Co. Pvt. Ltd. Hull, R., 1996. Molecular biology of rice tungro viruses. Annual Review of Phytopathology 34, 275–297. Hull, R., 2014. Matthews’ Plant Virology, fifth ed. London: Academic Press. Kumar, G., Dasgupta, I., 2017. Molecular biology of rice tungro viruses and strategies for their control. In: Shamim, M., Singh, K.N. (Eds.), Biotic Stress Management in Rice. Apple Academic Press, pp. 1–15.

Rice Yellow Mottle Virus (Solemoviridae) Eugénie Hébrard and Nils Poulicard, Interactions Plantes Microorganismes Environnement, Institut de Recherche pour le Développement, Centre de coopération internationale en recherche agronomique pour le développement, University of Montpellier, Montpellier, France Mbolarinosy Rakotomalala, FOFIFA, Antananarivo, Madagascar r 2021 Elsevier Ltd. All rights reserved. This is an update of E. Hebrard, D. Fargette, G. Konate, Rice Yellow Mottle Virus. In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00715-9.

Introduction Rice yellow mottle virus (RYMV) is a member of the genus Sobemovirus in the family Solemoviridae. RYMV is present only on the African continent. It was first reported in Kenya in 1966 and in Côte d0 Ivoire in 1974 in irrigated rice fields. Since the early 1990s, RYMV is present everywhere where rice is grown in sub-saharan Africa and in Madagascar. It affected all types of rice cultivations, including lowland, upland rainfed, floating and mangrove rice. RYMV regularly induces severe yield losses to rice production ranging from 20% to 100%. In some regions, when epidemics are recurrently very severe, farmers abandoned their fields and planted new rice fields. Highly susceptible cultivars have been eliminated by the disease. RYMV is now ranked as the main biotic threat to rice production in Africa.

Virion Properties RYMV has icosahedral particles of 25 nm in diameter (Fig. 1). The virion particles are composed of a single coat protein (CP) of 29 kDa, a genomic RNA (gRNA), and one subgenomic RNA (sgRNA) molecule. The capsid contains 180 copies of the coat protein subunit arranged with T ¼ 3 symmetry. The structure of RYMV was determined by X-ray crystallography at 2.8 Å resolution and compared to the structure of Southern cowpea mosaic virus (SCPMV) and Sesbania mosaic virus (SeMV). As for other sobemoviruses, RYMV CP subunits are chemically identical but structurally not equivalent. Three types of CP subunits termed A, B, C are related by quasi three-fold axes of symmetry and are involved in different inter-subunit contacts. In the C-type subunits, a longer part of the N-terminus is ordered (residues 27–49), forming an additional b-strand named bA arm. Sobemovirus particles are stabilized by divalent cations, pH-dependent protein-protein interactions, and salt bridges between protein and RNA. Upon alkali treatment in presence of chelators, the capsid shell swells and becomes sensitive to enzymes and denaturants. In these conditions, RYMV is more stable than SCPMV. This property is likely due to the 3D swapping of the RYMV bA arm around the distal quasi-six-fold axes as found with SeMV whereas the SCPMV bA arm makes a U-turn and is localized around the nearby quasi-six-fold axes. Another reason for the better stability of RYMV is likely to be the strong RNA-protein interactions resulting from the presence of ordered RNA within the capsid shell. RYMV particles exist in three forms with different stability (1) an unstable swollen form dependent on basic pH but lacking Ca2 þ , (2) a transitional compact form dependent on acidic pH, but also lacking Ca2 þ and (3) a stable compact form that is pH independent and contains Ca2 þ . The compact form is highly infectious and probably required for virus movement and transport by vector. Molecular diversity of the CP was analysed in relation to the capsid three-dimensional structure in order to identify which amino acids are involved in the differential recognition by certain monoclonal antibodies. The residues in position 178 and 180, despite their internal localisation in the capsid, can modify the antigenic reactivity, with Mabs G and E allowing serotypes Sr3 and Sr5 to be differentiated.

Organization of the Genome The genomic RNA is one single-stranded messenger-sense molecule, ca. 4450 nt in size. The 50 -terminus of the RNAs has a genome-linked protein (VPg), and lacks a poly(A) tail. RYMV often encapsidates, in addition to its genomic and subgenomic RNA, a small circular satellite RNA (satRNA), that is dependent on a helper virus for replication. RYMV satRNA has 220 nt and it is a viroid-like RNA. No link between the RYMV satRNA and its pathogenicity has been demonstrated. Like other sobemoviruses, the RYMV genome is organized in 5 overlapping open reading frames (ORF) (Fig. 2). Two noncoding regions at the RNA extremities contain 80 and 289 nt, respectively. The ORF1 and the ORF3 are localized at the 50 and 30 -extremities, respectively. An intergenic region of 12 nt separates ORF1 and ORFx. All the following coding regions overlapped two by two (ORFx and ORF2a, ORF2a and ORF2b, ORF2b and ORF3). The ORF1 encodes a protein P1 which is dispensable for replication but is required for systemic movement in plants. Moreover, RYMV P1 displays multiple functions in silencing by locally suppressing transgenic silencing but enhancing the local and systemic spread of silencing in the non-host plant Nicotiana benthamiana and inhibiting the DCL4-dependent endogenous siRNA pathway when transgenically expressed in rice. The putative protein encodes by ORFx is predicted to be translated from a leaky scanning and a non-AUG initiation. As for all the sobemoviruses, the protein Px has not been detected in infected plants and its function is unknown.

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Fig. 1 RYMV capsid structure. The capsid comprised 180 copies of one single type of polypeptide arranged in T ¼ 3 quasi-equivalent symmetry. The icosahedral asymmetric unit contains three subunits: A (in blue), B (in red) and C (in green). Each subunit is involved in different inter-subunit contacts. This image was automatically generated from ViPER virus capsid PDB file 1F2N using the MultiScale extension to the Chimera interactive molecular graphics package. Reproduce from Goddard, T.D., Huang, C.C., Ferrin, T.E., 2005. Software Extensions to UCSF Chimera for Interactive Visualization of Large Molecular Assemblies, Structure 13 (3), 473–482. Shepherd, C.M., Borelli, I.A., Lander, G., et al., 2006. 3rd, Reddy VS., VIPERdb: A relational database for structural virology. Nucleic Acids Research 34, (Database issue), D386-9.

Fig. 2 RYMV genomic organization. Positions of the open reading frames (ORFs) are indicated in nucleotides. P1, protein X (Px), proteinase (Pro), VPg, RNA-dependent RNA-polymerase (Pol) and coat protein (CP) are labelled. The dotted line at nucleotide 1979 represents the frameshifting signal. The fusion point of the polyprotein P2a þ b is unknown, and an AUG codon present at the beginning of the ORF2b (nucleotide 2093) is indicated by the vertical line.

The polyproteins encoded by the RYMV ORF2 are presumably translated from leaky translation. The start codon of ORF2 appeared to be in a more favorable context than ORF1. The ORF2b is translated as a polyprotein fused with ORF2a after a  1 programmed ribosomal frameshifting mechanism. The presence of a heptanucleotide slippery sequence UUUAAAC and a predicted stem-loop structure are necessary. The ORF2a encodes the proteinase, the VPg and two putative proteins P10 and P8, the ORF2b encodes the RNA-dependent RNA-polymerase. RYMV proteinase contain the consensus sequence H(X32–35)[D/E](X61–62) TXXGXSG characteristic of serine proteinases, they cleave between E-T and E-S amino acid residues. RYMV VPg is covalently linked to a serine, is characterized by the presence of a conserved W[A/G]D sequence followed by a D- and E-rich region and can be phosphorylated. The interaction between the VPg and the rice eukaryotic translation initiation factor eIF4(iso)G1 is needed for a successful infection. The VPg and the P8 protein are intrinsically disordered. The localisation of the polymerase was predicted from the presence of the GDD motif and surrounding conserved motifs characteristics of RdRp. The ORF3 is translated from the subgenomic RNA and encodes the coat protein (CP). RYMV CP is dispensable for replication but required for movement to long-distance likely through encapsidation. This movement was suggested to occur during the cell differentiation to vessels. The N-terminal sequence contains a putative bipartite nuclear localisation signal (NLS). This highly basic region is thought to interact with viral RNA and to stabilize the virion but virus like-particles (VLP) can be formed.

Relationships of the Species With Other Taxa The species Rice yellow mottle virus belongs to the genus Sobemovirus in the family Solemoviridae. The genus contains 19 definitive species including Southern bean mosaic virus, the type species. The sobemovirus CPs are related to those of necroviruses (Tombusviridae) whereas the proteinase, VPg and polymerase are related to those of poleroviruses and enamoviruses (Luteoviridae).

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Biological Properties The natural host range of RYMV is restricted to a few members of the Poaceae family, principally Eragrostidae and Oryzae sp. The most commonly cited species are Oryza sativa, O. glaberrima, O. longistaminata, O. barthii and Echinochloa colona. A few additional Poaceae plants have been infected experimentally. RYMV particles are present in all plant parts including roots and seeds. Particles are present in large amount in mesophyll cells, and in vascular tissues mainly in xylem and associated parenchyma cells. Particles were found individually, as aggregates, or as crystalline forms in the cytoplasm and the vacuoles of infected cells. In addition, the cytoplasm of infected cells contains fibrillar material located either in vesicles or distributed in diffused patches. Virus particles are also found in mature xylem, as well as inside the primary wall. Cytological changes were found in the chloroplasts of infected mesophyll cells where the starch grains decreased in size and number. The chloroplasts form invaginations containing mitochondria, peroxisomes and virus particles. Electron dense materials were observed in the nuclei and associated with fibrillar elements in cytoplasmic vesicles. The most dramatic changes induced by RYMV occurred in the cell walls of parenchyma and mature xylem cells causing disorganisation of the middle lamellae wall. From infected cells, a viral ribonucleoprotein complex moves from cell-to-cell to reach the vascular bundle sheath. Viral replication, encapsidation and storage in vacuoles occur mainly in vascular parenchyma cells. The calcium linkage from pit membranes to virus particles during xylem differentiation could contribute to disruption of these membranes and facilitate systemic virus transport. In the upper leaves, virions are suspected to cross the pit membrane to infect new vascular cells and virions spread out by cell-to-cell movement through plasmodesmata. At this stage of infection, most replication occurs in mesophyll and vascular cells, and most of the virions accumulate in large crystalline patches. Vector transmission is mainly due to coleoptera vectors such as Chaetocnema spp., Sesselia pusilla et Trichispa sericea, but other biting insects including the grasshopper Conocephalus have been reported to occasionally transmit the virus. Transmission is not persistent. The virus is present within the seed, although RYMV is not seed-transmissible. Other transmission modes have been also reported through animals like rats, common in rice fields, or cows and donkeys, used for field work or during grazing. Abiotic transmissions through plant residues, irrigation water, guttation water, wind, direct contact between plants and contamination by infected agricultural tools have been observed. Experimentally, mechanical transmission is easily achieved.

Diagnostic and Identification RYMV diagnosis was first based on symptoms. However, many factors such as mineral deficiencies can induce symptoms of yellow mottle on rice. Therefore, several tools for specific detection of RYMV were developed. First, polyclonal antibodies (PAbs) directed against different isolates were used in double-diffusion tests and in direct antibody sandwich enzyme linked immunosorbent assay (DAS-ELISA). No cross reactions with other sobemoviruses was observed, demonstrating the high specificity of these antibodies. Western-blot analysis can be used to detect RYMV with PAbs. These antibodies were also used in immunolocalisation techniques. Immuno-printing tests derived from the direct tissue blotting technique were developed to analyse the distribution of the virus in entire leaves. Moreover, cytological detection of RYMV CP in rice tissues can be performed by immunofluorescence microscopy after glutaraldehyde/paraformaldehyde fixation and labelling with secondary antibodies linked to fluorescein. Polyclonal antibodies directed against P1 produced in E. coli are also available, and were used in Western-blot analyses. Monoclonal antibodies were also developed to study RYMV diversity. Indirect triple antibody ELISA (TAS-ELISA) with strain-specific MAbs allows the major serotypes to be differentiated. In parallel to the serological methods, molecular tools were developed. The first full-length sequence was obtained after RNA extraction from purified virus. Several specific primers were defined to perform RT-PCR amplification and to produce infectious transcripts. Northern and Southern blot methods are available using a DNA probe obtained by PCR amplification of the CP gene (ORF4) from the infectious clone CIa. In situ hybridisation tests of the RNA in rice tissues were also performed. In this case, the probe was DNA, obtained from the full-length genome of the clone CIa labelled randomly with digoxigenin-UTP. Real time-RT-PCR assay is also available using primers in the conserved ORF2a. RT-PCR amplification of the CP gene followed by direct DNA sequencing progressively has replaced other molecular methods of analyses.

Epidemiological Aspects Typical symptoms of RYMV are a mottle and a yellowing of the leaves. However, streaks, necrosis, and whitening are sometimes observed with some cultivars and under specific growing conditions. Infection resulted in stunting of the plant, reduced tillering, poor panicle exertion and sterility. Death of the plants of susceptible cultivars occurs after early infection. Severe yield losses ranging from 20% to 100% after RYMV infection have been reported in several countries. Most cultivars currently grown are susceptible to RYMV. Heavy infection is associated with reduced fertility. RYMV is an emergent disease. It is thought that the virus was originally harboured in wild Poaceae and transferred only recently to cultivated rice. The rapid and intense spread of the virus was associated to the change of agricultural practices in response to increasing food demand. Very productive but susceptible O. sativa cultivars have been introduced from Asia which allows two crop

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Fig. 3 Descriptive model of RYMV epidemiology. (1). Environment. RYMV is present in the environment in rice stubble and regrowths and in wild Poaceae. (2). Rice seedbeds. Primary infection occurred via biotic transmission (beetles, rats, cows) whereas cultural practices contribute to secondary infection in seedbeds. (3). Rice fields. Primary infection occurred via biotic transmission from the environment, and also when transplanting infected seedlings from seedbeds. Both biotic and abiotic transmissions contribute to secondary spread in rice fields. After harvesting, RYMV is perpetuated in contra-season in the environment through rice stubbles and regrowths and transmitted to wild Poaceae. From Hébrard, E., Fargette, D., Konaté, G., 2008. Rice Yellow Mottle Virus. In: Mahy, B.W.J., Van Regenmortel, M.H.V. (Eds.), Encyclopedia of Virology third ed. Oxford: Academic Press, pp. 485–490.

cycles a year resulting in the maintenance of the inoculum all over the year. RYMV propagation probably occurs as follows: (1) a virus source made of volunteers, of wild Poaceae or plant residues act as primary inoculum; (2) a direct contamination of the new rice fields occurs after mechanical transmission by man or biological infection by insects; (3) rice beds can also be infected by insect vector; (4) the new rice field originating from these infected seed-beds are contaminated by man during replanting; (5) further propagation of the virus is done by man animals, wind and irrigation water. The epidemiology has been little studied and there are even uncertainties about the respective importance of biotic and abiotic transmission (Fig. 3). Under these conditions, precise forecasting of RYMV infection is not possible although growing intensification of rice cultivation in Africa will no doubt favour RYMV spread, unless durable resistant cultivars are introduced. Phytosanitary measures are sometimes advised although they are often economically not practicable and their impact to reduce virus spread is unknown. They include protection of seed-beds by nets, disinfection of tools used at replanting, destruction of volunteers and rice residues. Chemical control of the vectors is not economically feasible, is ecologically dangerous, and is most unlikely to be successful considering the large number of species involved in transmission. Several breeding programs have been conducted in order to select and breed resistant cultivars. The genotype and the phenotype of two kinds of natural resistance have been characterized. Partial resistance is encountered in cultivars of Oryza japonica species with a delayed virus multiplication and symptom expression. Partial resistance is polygenic and a major quantitative trait locus has been identified on chromosome 12. High resistance has been identified in a limited number of accessions of Oryza glaberrima and two O. sativa indica species. No symptoms are apparent and virus content is most often undetectable. Three resistance genes have been identified. The recessive gene RYMV1 that maps on chromosome 4 encodes the translation initiation factor eIF(iso)4G1. Compared to susceptible varieties, the resistance alleles are characterized by a point mutation and/or a small deletion in the conserved domain of the gene. The VPg was identified as the virulence factor, it interacts directly with eIF(iso)4G1 in susceptible plants. The RYMV1 resistance was efficient against some of the representative isolates of the main virus strains. However, resistance-breakdown was reported in experimental conditions depending on the viral strain and the resistance allele. A single point mutation was sufficient to break the resistance but negative epistasis can inhibit the virulence emergence. The amino acid located at position 49 of the VPg reflects inter-species adaptation of the viral strains and has indirect effects on resistance durability. The recessive resistance gene RYMV2 is homologous to the Arabidopsis thaliana gene for Constitutive expressor of Pathogenesis-Related genes-5 (CPR5), a nucleoporin controlling defense mechanisms. RYMV2-mediated resistance is conferred by a null allele. The virulence factor is the polyprotein P2a. The dominant resistance gene RYMV3 was mapped in a 15 kb interval in which a CC-NBS-LRR domain-containing protein was annotated. Recently, a hypervirulent pathotype able to overcome all natural sources of high resistance has been identified in West-Central Africa. Transgenic plants with portions of the ORF2 expressing partial resistance have also been produced. This transgenic resistance is thought to involve a gene silencing mechanism. However, the transgenic lines showed a less effective, partial and temporary resistance compared to natural resistances.

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Fig. 4 Phylogeny of RYMV. Phylogenetic tree inferred by maximum likelihood from the full sequences of 48 isolates representative of the genetic diversity and geographic distribution of RYMV in Africa. The different strains identified in West Africa (S1ca, S1wa, S2, S3, Sa, Sg) and East Africa (S4et, S4lm, S4lv, S4mg, S4ug) are differentiated by color. Strong bootstrap support of the nodes (480%) is indicated.

Diversity and Evolution The diversity of RYMV was assessed by studying isolates from several countries were the disease has been observed. The sequences of more than 500 CP genes are now available. Forty-eight isolates representative of the geographic and molecular diversity were fully sequenced. RYMV is a highly variable virus and analyses of the geographic distribution of the genetic diversity elucidated the process of evolution and of dispersal of the virus. RYMV showed a high level of population structure marked at the continental scale with two subdivisions: East-Africa and West-Africa (Fig. 4). The highest diversity was observed in East-Africa, with a pronounced peak in Eastern Tanzania, and a decrease from the east to the west of the continent. This pattern suggests a westward expansion with a succession of founder effects and subsequent diversification phases. Accordingly, genetic diversity would be adversely affected by recurrent bottlenecks occurring along the route of colonization, resulting in the lowest diversity in the extreme west. This, together with accumulation of de novo mutations postdating population separation, provides an explanation for the genetic differences among strains across Africa. The geographical overlaps between several strains would explain the recent detection of some recombination strains.

Further Reading Albar, L., Bangratz-Reyser, M., Hébrard, E., et al., 2006. Mutations in the eIF(iso)4G translation initiation factor confer high resistance of rice to Rice yellow mottle virus. The Plant Journal 47, 417. Bonneau, C., Brugidou, C., Chen, L., et al., 1998. Expression of the Rice yellow mottle virus P1 protein in vitro and in vivo and its involvement in virus spread. Virology 244, 79. Brugidou, C., Holt, C., Yassi, et al., 1995. Synthesis of an infectious full-length cDNA clone of Rice yellow mottle virus and mutagenesis of the coat protein. Virology 206, 108. Brugidou, C., Opalka, N., Yeager, M., et al., 2002. Stability of Rice yellow mottle virus and cellular compartmentalization during the infection process in Oryza sativa (L.). Virology 297, 98. Hébrard, E., Pinel-Galzi, A., Oludare, A., et al., 2018. Identification of a hypervirulent pathotype of Rice yellow mottle virus: A threat to genetic resistance deployment in West-Central Africa. Phytopathology 108 (2), 299–307.

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Pidon, H., Ghesquière, A., Chéron, S., et al., 2017. Fine mapping of RYMV3: A new resistance gene to Rice yellow mottle virus from Oryza glaberrima. Theoretical and Applied Genetics 130 (4), 807–818. Pinel-Galzi, A., Dubreuil-Tranchant, C., Hébrard, E., et al., 2016. Mutations in Rice yellow mottle virus polyprotein P2a involved in RYMV2 gene resistance breakdown. Frontiers in Plant Science 7, 1779. Pinel-Galzi, A., Rakotomalala, M., Sangu, E., et al., 2007. Theme and variations in the evolutionary pathways to virulence of an RNA plant virus species. PLOS Pathogens 3, e180. Pinel-Galzi, A., Traoré, O., Séré, Y., Hébrard, E., Fargette, D., 2015. The biogeography of viral emergence: Rice yellow mottle virus as a case study. Current Opinion in Virology 10, 7–13. Pinel, A., Abubakar, Z., Traore, O., et al., 2003. Molecular epidemiology of the RNA satellite of Rice yellow mottle virus in Africa. Archives of Virology 148, 1721. Poulicard, N., Pinel-Galzi, A., Traoré, O., et al., 2012. Historical contingencies modulate the adaptability of Rice yellow mottle virus. PLoS Pathogens 8 (1), e1002482. Qu, C., Liljas, L., Opalka, N., et al., 2000. 3D domain swapping modulates the stability of members of an icosahedral virus group. Structure 8, 1095. Sorho, F., Pinel, A., Traoré, O., et al., 2005. Durability of natural and transgenic resistances in rice to Rice yellow mottle virus. European Journal of Plant Pathology 112, 349. Traoré, O., Sorho, F., Pinel, A., et al., 2005. Processes of diversification and dispersion of Rice yellow mottle virus inferred from large-scale and high resolution phylogeographic studies. Molecular Ecology 14, 2097. Traoré, O., Traoré, M., Fargette, D., Konaté, G., 2006. Rice seedbed as a source of primary infection by Rice yellow mottle virus. European Journal of Plant Pathology 115, 181.

Satellite Nucleic Acids and Viruses Olufemi J Alabi, Texas A&M AgriLife Research and Extension Center, Weslaco, TX, United States Alfredo Diaz-Lara and Maher Al Rwahnih, University of California, Davis, CA, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of P. Palukaitis, A. Rezaian, F. Garcia-Arenal, Satellite Nucleic Acids and Viruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00500-8.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein diRNAs Defective interfering RNAs dsDNA Double-stranded deoxyribonucleic acid gRNA Genomic RNA HEL Helicase HV Helper virus kb Kilobases; the size of a ssDNA or ssRNA molecule kbp Kilobase pairs; the size of a dsDNA or dsRNA molecule kDa Kilodaltons; the size of a protein

Glossary Satellite RNA or satellite DNA A subviral genome dependent on a helper virus for both replication and encapsidation. Satellite virus A subviral genome dependent on a helper virus for its replication but encoding its own capsid protein. Satellite-like RNA A subviral genome dependent upon another virus for replication and encapsidation but is required for vector transmission of the helper virus.

MCP Major capsid protein mCP Minor capsid protein NTR Non-translated region ORF Open reading frame Poly(A) Polyadenylated PRO Protease sgRNA Sub-genomic RNA ssDNA Single-stranded deoxyribonucleic acid ssRNA Single-stranded ribonucleic acid tRNA Transfer RNA Vpg Genome-linked protein

Virophages Double-stranded DNA that depend on giant viruses for their replicate in co-infected eukaryotic cells. Virus-associated nucleic acid A subviral genome that depends on another virus for encapsidation and transmission, but not for its replication.

Introduction Satellites are a diverse collection of subviral agents that are capable of attenuating or exacerbating disease symptoms caused by their cognate helper viruses (HVs). Satellites can be differentiated from other subviral nucleic acids, such as sub-genomic (sg) RNAs, defective (d) or defective interfering (di) RNAs, and viroids, by their molecular, biological, and genetic nature. Unlike sgRNAs, dRNAs, and diRNAs, satellites and viroids have little or no sequence similarity to any known virus, except for the terminal regions. Whereas viroids are replicated by host polymerases, RNA satellites rely on the HV’s polymerase for their replication. Satellites have been found associated with DNA and RNA viruses, and in the latter case, with both ssRNA and dsRNA viral genomes, the satellites being of the same nucleic acid type as the HV. While, in general, the HV can exist independent of the satellite, there are exceptions where a satellite contributes to the transmission of the HV and thus is referred to as being satellitelike. In addition, some viral RNAs and DNAs are found in association with other viral genomes on which they depend for their encapsidation and transmission, but not for replication. These viral RNAs and DNAs are referred to here as virus-associated nucleic acids. True satellites are divided basically into two main groups; those that encode their own capsid protein (CP) are called satellite viruses, while those that rely on their HV for both replication and encapsidation are referred to as satellite RNAs or DNAs (satellite nucleic acids). There are also satellites of satellites, in which some satellite RNAs are replicated by the HV but are encapsidated by the CP of a satellite virus. In addition, in the case of turnip crinkle virus (TCV, genus Betacarmovirus, family Tombusviridae), which replicates both satellite RNAs and diRNAs, it also produces a chimeric RNA (referred to as satC) that is in part a diRNA and in part a satellite RNA (satD). While most satellites are found in association with plant viruses, a few have been found in association with animal, fungal or protist viral genomes. Most plant viruses are not associated with satellites. However, the few that does have attracted significant attention due to the disease modulating roles of the associated satellites. With the advent of metagenomics, additional satellites have been identified prompting a revisit of their taxonomic classification to align them with that of bona fide viruses.

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History The use of the term satellite as a subviral agent was first conceptualized in 1962, to describe the relationship of a 17 nm diameter viral particle found in association with some isolates of the 26 nm diameter tobacco necrosis virus (TNV; genus Alphanecrovirus, family Tombusviridae). The smaller particle was dependent on TNV for its accumulation but was serologically unrelated to TNV. A few other satellite viruses have been described since then, but, in general, they are rare. By contrast, satellite nucleic acids are more common. The first satellite RNA was described in 1969, in association with tobacco ringspot virus (TRSV, genus Nepovirus, family Secoviridae), the satellite RNA being encapsidated by the CP of TRSV. The symptoms induced by TRSV were attenuated dramatically by the presence of the satellite RNA; but this is not always the case. Some satellite RNAs have no effect on either the accumulation or pathogenesis of the HV while a few may exacerbate the HV-induced disease. Most of these satellite RNAs contain ssRNA genomes, while a few contain genomes of dsRNA. Some of the satellite RNAs with ssRNA genomes are translated to produce proteins, which may or may not be required for their replication, depending on the specific satellite RNA. The first demonstration that satellite molecules are not limited to RNA viral systems was made in 1997 when a tomato leaf curl virus satellite (TLCV-sat-DNA) was characterized from disease-affected tomato plants in Australia. TLCV-sat-DNA is dependent on its TLCV HV for replication and encapsidation. In addition, it was shown that TLCV-sat-DNA replication was also supported by other taxonomically distinct geminiviruses, including the begomoviruses tomato yellow leaf curl virus (TYLCV) and African cassava mosaic virus (ACMV), and the curtovirus beet curly top virus (BCTV). Since then, additional classes of satellite DNAs have been characterized and described.

Geographical Distribution The host range and geographical distribution of satellites mirror those of their HVs owing to the dependence of the former on the latter for genome replication and encapsidation. Similarly, satellite nucleic acids depend on the HV’s transmission mechanism (including the dependence on arthropod vectors) for short and long-distance spread. Thus, the worldwide prevalence and distribution of satellite molecules draws from the global distribution of their HVs. Interestingly, alphasatellites and betasatellites are mostly associated with Old World monopartite geminiviruses, except for some alphasatellites that have been recently reported to occur in association with New World bipartite begomovirus infections.

Classification Satellite viruses, satellite nucleic acids, virus-associated nucleic acids, and other satellite-like agents are very diverse in terms of their nucleic acid type, size, sequence, structure, or translatability. Satellite genomes may be composed of single or double stranded RNA or DNA molecules, which may be linear or circular. Whereas satellite viruses encode a CP for genome encapsidation, satellite nucleic acids depend on the CP of their HV for genome encapsidation. True satellites such as satellite viruses and satellite nucleic acids are dependent on the HV for their multiplication in co-infected host cells since they lack genes that encode functions needed for replication. Most or all portions of the genomes of true satellite, except for the terminal regions, are distinct from the genomes of their cognate HV. Satellite-like nucleic acids are another class of subviral agents that differ from true satellites in that they encode a function necessary for the biological success of the associated virus. However, like the true satellites, they also are incapable of autonomous replication because they do not encode a replicase, with the exception of alphasatellites. Thus, they function primarily to remedy a deficiency in a defective virus and have sometimes been classified as part of the genome of the virus they assist. For instance, satellite-like RNAs associated with groundnut rosette virus (genus Umbravirus, family Tombusviridae) or beet necrotic yellow vein virus (BNYVV, genus Benyvirus, family Benyviridae) contribute to vector transmissibility of their associated viruses. Other classes of autonomously replicating nucleic acids (virus-associated nucleic acids) such as alphasatellites are dependent on their HV for encapsidation, cell-to-cell and long-distance movement and vector transmission. Thus, satellites do not constitute a homogeneous group and are traditionally not formally classified into species and higher taxa by the International Committee on Taxonomy of Viruses (ICTV). However, because several satellite molecules encode functional proteins, their taxonomy has been revisited to bring it in alignment with that of bona fide viruses. A list of currently known satellites categories is provided in Table 1.

General Properties and Effects of Satellites Satellite Viruses Category 1: Chronic bee-paralysis associated satellite virus Chronic bee-paralysis associated satellite virus (CBPVA; unassigned) is the only member of this group characterized to date (Table 2). The isometric particles of this satellite virus are found in bees infected with the HV, chronic bee-paralysis virus (CBPV), with CBPVA capable of interfering with CBPV replication.

Satellite Nucleic Acids and Viruses Table 1

Categories of satellites

Satellite viruses: 1 2 3 4 5

Chronic bee-paralysis virus-associated satellite virus Satellites that resemble tobacco necrosis satellite virus Nodavirus-associated satellite virus Adenovirus-associated satellite virus (Dependovirus) Mimivirus-associated satellite virus (Sputnik, virophage)

Virus-dependent nucleic acids: Single stranded DNAs 6a 6b Double stranded RNAs 7 Single stranded RNAs 8a 8b 8c 8d 8e

Table 2

Alphasatellites Betasatellites and deltasatellites Satellites dsRNAs Large linear single stranded satellite RNAs Small linear single stranded satellite and satellite-like RNAs Small circular single stranded satellite RNAs Hepadnavirus-associated satellite-like RNAs (Deltavirus) Polerovirus-associated RNAs

Classification of satellites viruses a

Category Family

Subfamily

Genus

Species

Type speciesb accession #

Genome/Host typesc

Helper virus genus

N/A

ssRNA/ Arthropod ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/ Arthropod

Unclassified

Albetovirus Albetovirus Albetovirus Aumaivirus Papanivirus Virtovirus Macronovirus

Chronic bee-paralysis associated satellite virus (CBPVA) Tobacco albetovirus 1 Tobacco albetovirus 2 Tobacco albetovirus 3 Maize aumaivirus 1 Panicum papanivirus 1 Tobacco virtovirus 1 Macrobrachium satellite virus 1

AF063497

Adenovirus

AY186198

ssDNA/Humans, Vertebrates ssDNA/Humans, Vertebrates ssDNA/Humans, Vertebrates ssDNA/Humans, Vertebrates ssDNA/Humans, Vertebrates ssDNA/Humans, Vertebrates ssDNA/Humans, Vertebrates ssDNA/Humans, Vertebrates dsDNA/Protists dsDNA/Protists dsDNA/Protists

Mimivirus

RNA Satellite viruses 1 2

3

Sarthroviridae

V01468 M64479 AJ000898 M55012 M17182 M25782 AY247793

DNA Satellite viruses 4 Parvoviridae

Parvovirinae Dependoparvovirus

Parvoviridae

Parvovirinae Dependoparvovirus

Parvoviridae

Parvovirinae Dependoparvovirus

Adeno-associated dependoparvovirus A Adeno-associated dependoparvovirus B Anseriform dependoparvovirus 1

Parvoviridae

Parvovirinae Dependoparvovirus

Avian dependoparvovirus 1

Parvoviridae

Parvovirinae Dependoparvovirus

Chiropteran dependoparvovirus 1 GU226971

Parvoviridae

Parvovirinae Dependoparvovirus

Pinniped dependoparvovirus 1

JN420372

Parvoviridae

Parvovirinae Dependoparvovirus

Squamate dependoparvovirus 1

AY349010

Parvoviridae

Parvovirinae Dependoparvovirus

Squamate dependoparvovirus 2

KP733794

5

a

Lavidaviridae Lavidaviridae

Mavirus Sputnikvirus

Lavidaviridae

Sputnikvirus

AF085716 U22967

Cafeteriavirus-dependent mavirus HQ712116 Mimivirus-dependent virus EU606015 Sputnik Mimivirus-dependent virus HG531932 Zamilon

Category adopted from the 9th ICTV Report. N/A, unavailable. c ssRNA, single-stranded RNA; ssDNA, single-stranded DNA; dsRNA, double-stranded RNA; dsDNA, double-stranded DNA. b

683

Alphanecrovirus Alphanecrovirus Alphanecrovirus Aureusvirus Panicovirus Tobamovirus Nodavirus

Adenovirus Adenovirus Adenovirus Adenovirus Adenovirus Adenovirus Adenovirus Mimivirus Mimivirus

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Category 2: Satellites that resemble tobacco necrosis satellite virus (genera Albetovirus, Aumaivirus, Papanivirus and Virtovirus) The group consists of several plant-infecting satellite viruses that are found in association with taxonomically distinct HVs that used to be collectively known as the tobacco necrosis satellite virus or satellite tobacco necrosis virus (STNV) group. Recently, the ICTV recognized species belonging to this group and assigned them to the genera Albetovirus, Aumaivirus, Papanivirus, and Virtovirus. Member belonging to these unassigned genera are found in association with ss( þ )RNA plant HVs belonging to the genera Alphanecrovirus, Aureusvirus, Panicovirus (Tombusviridae), and Tobamovirus, (Virgaviridae) respectively (Table 2). These satellite viruses depend on their HVs for replication. With the exception of the rod-shaped Tobacco virtovirus 1 (unassigned genus Virtovirus), particles of the plant satellite viruses are isometric, 17 nm in diameter, with a T ¼ 1 symmetry, built of 60 protein subunits, as determined at high resolution by X-ray diffraction for Tobacco albetovirus 1 (genus Albetovirus) and Panicum papanivirus 1 (genus Papanivirus). Thus, the particle morphology of most satellite viruses differs from that of their respective HVs. In spite of the different structure of CP and virus particle, the particles of both the satellite and HV may share important properties; for example, the particles of Tobacco albetovirus 1 can bind specifically to the zoospores of the vector fungus Olpidium brassicae. The ca. 1200 nt RNA of Tobacco albetovirus 1 does not have a methylated cap structure or a genome-linked protein (VPg) in its 50 end. Unlike most plant virus RNAs, it has a phosphorylated 50 end. The 30 termini may share a structure with that of the HV genomic RNA (gRNA) as is the case for both tobacco mosaic virus and Tobacco virtovirus 1 where their 30 termini form tRNA-like structures that are aminoacylable with histidine. Cis-acting sequences necessary for RNA amplification have been mapped at the 50 and 30 nontranslated regions (NTRs) of Panicum papanivirus 1 RNA as well as within the CP open reading frame (ORF), and are conserved in a defective RNA, which is maintained by this satellite virus. A translational enhancer domain has been mapped in the 30 NTR of albetoviruses. Like other satellite viruses, the isometric particle of Maize aumaivirus 1 (genus Aumaivirus, unassigned) is 17 nm in diameter, but its CP has limited sequence similarity with orthologous proteins of other satellite viruses. Albetoviruses have been reported to interfere with each other’s replication and with the accumulation of the HV. Satellite viruses can also modify the symptoms induced by the HV, as is the case with Panicum papanivirus 1, which in co-infection with Panicum mosaic virus (PMV) enhances the mild symptoms of the HV to cause severe mosaic and chlorosis in millet plants. Symptom induction is due to the Panicum papanivirus 1 CP, and a chlorosis-inducing domain has been mapped. In addition to its structural and symptom-inducing functions, the CP of Panicum papanivirus 1 is involved in systemic movement, binds PMV particles, and counters the effects of posttranscriptional gene silencing suppressors. Furthermore, the particles of albetoviruses may contain non-coding satellite RNA of about 620 nt, which depends on TNV for replication and on the satellite virus for encapsidation. A satellite RNA with similar dependence relationships has been described for PMV and Panicum papanivirus 1. These systems are good examples of the complexity of dependence relationships in satellitism. Category 3: Nodavirus-associated satellite virus (genus Macronovirus, family Sarthroviridae) Macrobrachium satellite virus 1 (genus Macronovirus; family Sarthroviridae) is the only currently recognized species of this group (Table 2), with extra small virus (XSV) as its representative member. XSV and its HV Macrobrachium rosenbergii nodavirus (MrNV) infect giant freshwater prawns and cause the debilitating and fatal white tail disease or white muscle disease in several countries. XSV is consistently found in co-infection with MrNV on which it depends for its replication. The linear, positive sense ssRNA genome of XSV is 796 nt in length and contains a short poly(A) tail of 15–20 nt at the 30 end. The non-enveloped virions of XSV are spherical (ca. 15 nm in diameter) and they are serologically and structurally different to those of MrNV. It encodes a single ORF that is translated into two CPs, one an N-terminally truncated version of the other. XSV and MrNV have also been detected in several species of aquatic insects that were sampled from infected freshwater prawn nursery ponds. In addition, the satellite and HVs can replicate in mosquito cell lines, suggesting that aquatic insects serve as vectors for their spread. Category 4: Adenovirus-associated satellite virus (family Parvoviridae; genus Dependoparvovirus) The genus Dependoparvovirus (family Parvoviridae) currently consists of eight ICTV approved species that are collectively known as adenovirus-associated satellite viruses (AAVs; Table 2). They are: Adeno-associated dependoparvovirus A (the type species), Adenoassociated dependoparvovirus B, Anseriform dependoparvovirus 1, Avian dependoparvovirus 1, Chiropteran dependoparvovirus 1, Pinniped dependoparvovirus 1, Squamate dependoparvovirus 1, and Squamate dependoparvovirus 2. Their non-enveloped icosahedral virion is 18–26 nm in diameter with the capsid consisting of 60 copies of CPs, encapsidating a linear, ssDNA( þ /  ) genome of about 4.7 kb in size. As their genus name implies, the AAVs are mostly (or thought to be likely) dependent on helper DNA viruses, most commonly an adenovirus or herpevirus, for replication in co-infected cells of their mammalian, avian or reptilian hosts. Though ubiquitous in humans and most other vertebrates, AAVs are by themselves not associated with any known pathology. Category 5: Mimivirus-associated satellite virus (genera Mavirus, Sputnikvirus, family Lavidaviridae) The discovery of giant viruses (extremely large genomes) infecting protists also resulted in the identification of new satellite viruses. The family Lavidaviridae includes the Sputnik and Zamilon viruses and the Maverick-related virus (genus Mavirus); also called giant satellite viruses (17–30 kbp linear or circular dsDNA genomes). “Lavida-” stands for large virus-dependent or -associated virus and refers to the property of Sputnik and other satellite viruses of depending on or associating with large dsDNA viruses. Sputnik virus was identified in an amoebal host in co-infection with its HV, Acanthamoeba polyphaga mimivirus (genus Mimivirus, family Mimiviridae). Though the icosahedral virus particle of Sputnik virus is only a tenth of the size of Acanthamoeba polyphaga mimivirus, the satellite virus may target the cytoplasmic replication factory of the giant virus and cause aberrant capsid phenotypes. As a result, the Sputnik virus was considered a viral parasite of a virus and named as “virophage”. This terminology has also been used to describe other satellites associated with giant viruses. Another typical feature of virophages is the presence of core genes in their genomes, a conserved set of six proteins or domains: major capsid protein (MCP), minor capsid protein (mCP),

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ATPase, protease (PRO), helicase (HEL), and a zinc-ribbon-containing domain. Currently, the family Lavidaviridae is divided in the genera Sputnikvirus and Mavirus with two and one species, respectively (Table 2).

Satellite Nucleic Acids and Satellite-Related Nucleic Acids Category 6a: Satellite ssDNA (family Alphasatellitidae; subfamilies Nanoalphasatellitinae and Geminialphasatellitinae) The recently created family Alphasatellitidae consists of circular single stranded DNA (ssDNA) molecules of approximately 1–1.4 kb that are associated with nanoviruses (subfamily Nanoalphasatellitinae) and some geminiviruses that belong to the genera Begomovirus and Mastrevirus (subfamily Geminialphasatellitinae). Currently, the subfamily Geminialphasatellitinae consists of four genera (Ageyesisatellite, Clecrusatellite, Colecusatellite, and Gosmusatellite) and 43 established species (Table 3) based on genus and species demarcation thresholds of E70% and E88%, respectively of their complete genomes. The subfamily Nanoalphasatellitinae is more variable with at least seven genera (Babusatellite, Clostunsatellite, Fabenesatellite, Milvetsatellite, Mivedwarsatellite, Sophoyesatellite, and Subclovsatellite) and 19 species (Table 3) using complete genome genus and species demarcation thresholds of E67% and E80%, respectively. Collectively referred to as alphasatellites, these plant-infecting molecules resemble the DNA-R component of nanoviruses in that they encode a replication-associated protein (Rep) and are capable of autonomous replication in host cells. They also possess a stem-loop region containing the nonanucleotide TAA/GTATTAC. However, they depend on their ssDNA HVs for encapsidation, movement, and insect transmission. The specific role of alphasatellites in the pathogenesis of their HV is unclear but some published studies have implicated their co-infections in reduced or exacerbated symptoms of the HV, reduced titer of the HV or betasatellite, or suppression of host defenses based on RNA interference. Category 6b: Satellite ssDNA (family Tolecusatellitidae; genera Betasatellite and Deltasatellite) Different satellite ssDNAs have been classified under the family Tolecusatellitidae with two genera (Betasatellite and Deltasatellite) based on their nt sequences. Like members of the Geminialphasatellitinae subfamily, the circular ssDNA molecules of betasatellites are about half the sizes (E1.3 kb) of their helper Old World monopartite begomoviruses, possess a stem-loop region containing the nonanucleotide TAATATTAC, and code for a single ORF (bC1), whose product has been shown to function as a pathogenicity determinant and suppressor of host gene silencing. Deltasatellites, however, are usually about one quarter (E0.7 kb) of the size of the genome of their monopartite Old World begomoviruses and bipartite New World begomoviruses. Unlike other geminivirusassociated satellite ssDNA molecules, deltasatellites do not encode for any protein. Consequently, like betasatellites, they also rely on their HV for encapsidation, replication, movement, and vector transmission. Whereas betasatellites are known to enhance the virulence of the helper begomovirus and may be essential for the maintenance of disease in the field, deltasatellites may cause a reduction on begomovirus accumulation and/or symptom severity in certain satellites-virus-host interactions. Current species demarcation thresholds within the family stands at o91% complete nt sequence identity for both genera, leading to the recognition of 61 betasatellite and 11 deltasatellite species (Table 3). Category 7: Satellite dsRNAs Most of the satellite dsRNAs (Table 4) occur in association with viruses in the families Totiviridae and Partitiviridae, on which they depend for the encapsidation and replication of their 0.5–1.8 kbp genomes. The totivirus-associated satellite dsRNAs encode a secreted protein toxin (the killer toxin) that is lethal to virus-free sensitive cells and to the HV that lacks the satellite dsRNA molecule. In turn, the HV that carry the toxin-encoding satellite dsRNA has immunity to that toxin; thus, the satellite dsRNA confers ecological advantage to its HV by killing competing virus- or satellite-free fungi. Unlike their Totiviridae counterparts, the satellite dsRNAs associated with partitiviruses do not code for functional proteins and their biological significance is not known. Category 8a: Large linear satellite ssRNAs Members of the large linear satellite ssRNAs have messenger RNA properties. They range in size between 0.8 and 1.5 kb and encode a nonstructural protein that may be essential for satellite RNA multiplication. All but two of the currently listed members are satellites of viruses in the family Secoviridae, the exceptions being bamboo mosaic virus (BaMV) satellite RNA and BNYVV RNA5 that are associated with a potexvirus and a benyvirus, respectively (Table 4). Several members of the Secoviridae-associated large linear satellite ssRNAs have been studied. They share 50 and 30 structural features of the gRNAs of the HV (a 30 terminal poly(A) sequence and a 50 terminal VPg). The encoded non-structural protein of different satellite RNAs are basic proteins of 36–48 kDa. For the satellite RNAs of tomato black ring virus, grapevine fanleaf virus, and Arabis mosaic virus (ArMV) (all three: genus Nepovirus, family Secoviridae), it has been shown that the encoded proteins are needed for the replication of the satellite RNA. Most large satellite RNAs of nepoviruses do not seem to have an effect on the accumulation or pathogenicity of the HV. However, this may depend on the experimental system, as the large satellite RNA of ArMV was shown to modulate the symptoms of the HV depending on the species of host plant. The BaMV satellite RNA is the only member that is encapsidated into rod-shaped particles. It encodes a 20 kDa protein that is expressed in vivo but is not needed for satellite RNA replication. The presence of BaMV satellite RNA significantly reduces the accumulation of the HV’s gRNA. Cis-acting sequences required for BaMV satellite RNA replication have been mapped at the 50 NTR and comprise a stem-loop structure that is conserved among BaMV satellite RNA variants. The BNYVV RNA5 encode a protein of 26 kDa, which is responsible for intensification of the rhizomania disease induced by infection of BNYVV RNAs 1 and 2, plus its two satellite-like RNAs 3 and 4. Category 8b: Small linear satellite ssRNAs The group consists of small (o0.7 kb), linear satellite RNA molecules that are associated with plant-infecting HVs in the families Tombusviridae, Bromoviridae, and the genus Umbravirus (Table 4). Some of the Umbravirus-associated members such as groundnut rosette virus satellite RNA and tobacco bushy top virus satellite RNA may be regarded as satellite-like RNAs due to their

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Table 3

Satellite Nucleic Acids and Viruses

Classification of ssDNA nucleic acid satellites

Family/Subfamily

Genus

Species

Type species Genome/Host typesa Helper virus genus accession #

Alphasatellitidae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Geminialphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae

Ageyesisatellite Ageyesisatellite Clecrusatellite Clecrusatellite Clecrusatellite Clecrusatellite Clecrusatellite Clecrusatellite Clecrusatellite Clecrusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Colecusatellite Gosmusatellite Gosmusatellite Gosmusatellite Gosmusatellite Gosmusatellite Gosmusatellite Unassigned Unassigned Babusatellite Babusatellite Babusatellite Babusatellite Clostunsatellite Clostunsatellite Clostunsatellite Clostunsatellite Clostunsatellite Fabenesatellite Milvetsatellite Mivedwarsatellite Mivedwarsatellite Mivedwarsatellite Mivedwarsatellite Sophoyesatellite

Ageratum yellow vein Singapore alphasatellite Cotton leaf curl Saudi Arabia alphasatellite Cleome leaf crumple alphasatellite Croton yellow vein mosaic alphasatellite Euphorbia yellow mosaic alphasatellite Melon chlorotic mosaic alphasatellite Sida Cuba alphasatellite Tomato yellow spot alphasatellite Whitefly associated Guatemala alphasatellite 2 Whitefly associated Puerto Rico alphasatellite 1 Ageratum enation alphasatellite Ageratum yellow vein alphasatellite Ageratum yellow vein China alphasatellite Ageratum yellow vein India alphasatellite Bhendi yellow vein alphasatellite Cassava mosaic Madagascar alphasatellite Chili leaf curl alphasatellite Cotton leaf curl Egypt alphasatellite Cotton leaf curl Gezira alphasatellite Cotton leaf curl Lucknow alphasatellite Cotton leaf curl Multan alphasatellite Gossypium darwinii symptomless alphasatellite Malvastrum yellow mosaic alphasatellite Malvastrum yellow mosaic Cameroon alphasatellite Pedilanthus leaf curl alphasatellite Sida leaf curl alphasatellite Sida yellow vein Vietnam alphasatellite Sunflower leaf curl Karnataka alphasatellite Synedrella leaf curl alphasatellite Tobacco curly shoot alphasatellite Tomato leaf curl Buea alphasatellite Tomato leaf curl Cameroon alphasatellite Tomato yellow leaf curl China alphasatellite Tomato yellow leaf curl Thailand alphasatellite Tomato yellow leaf curl Yunnan alphasatellite Gossypium mustelinum symptomless alphasatellite Hollyhock yellow vein alphasatellite Mesta yellow vein mosaic alphasatellite Okra enation leaf curl alphasatellite Okra yellow crinkle Cameroon alphasatellite Vernonia yellow vein Fujian alphasatellite Dragonfly associated alphasatellite Whitefly associated Guatemala alphasatellite 1 Banana bunchy top alphasatellite 1 Banana bunchy top alphasatellite 2 Banana bunchy top alphasatellite 3 Cardamom bushy dwarf alphasatellite Milk vetch dwarf alphasatellite 2 Pea necrotic yellow dwarf alphasatellite 2 Sophora yellow stunt alphasatellite 4 Sophora yellow stunt alphasatellite 5 Subterranean clover stunt alphasatellite 2 Faba bean necrotic yellows alphasatellite 2 Milk vetch dwarf alphasatellite 3 Faba bean necrotic stunt alphasatellite Milk vetch dwarf alphasatellite 1 Pea necrotic yellow dwarf alphasatellite 1 Sophora yellow stunt alphasatellite 2 Sophora yellow stunt alphasatellite 3

AJ416153 HG530543 FN436007 FN658711 FN436008 HM163578 HE806451 KX348228 KT099170 KT099173 JX913532 AJ238493 KF785752 JX570736 FN658716 HE984148 KF471043 AJ512960 EU589450 AJ132344 HQ343234 EU384623 FN675297 AM236765 KX168428 FR772088 DQ641718 JX569789 KJ939346 HQ407396 FN675299 FN675296 AJ579359 AM749493 KX759649 EU384656 FR772086 JX183090 HF546575 FN675285 KC959931 JX458742 KT099172 HQ616080 L32167 AF416471 KF435148 AB000922 KC979052 KX534409 KX534398 U16735 AJ005966 AB009047 KC978990 AB000920 KC979051 KX534408 KX534400

ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant

Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Babuvirus Babuvirus Babuvirus Babuvirus Nanovirus Nanovirus Nanovirus Nanovirus Nanovirus Nanovirus Nanovirus Nanovirus Nanovirus Nanovirus Nanovirus Nanovirus

Satellite Nucleic Acids and Viruses

Table 3

687

Continued

Family/Subfamily

Genus

Species

Type species Genome/Host typesa Helper virus genus accession #

Nanoalphasatellitinae Nanoalphasatellitinae Nanoalphasatellitinae Unassigned

Subclovsatellite Subclovsatellite Subclovsatellite Unassigned

Faba bean necrotic yellows alphasatellite 1 Sophora yellow stunt alphasatellite 1 Subterranean clover stunt alphasatellite 1 Coconut foliar decay alphasatellite

X80879 KX534399 U16731 M29963

ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant

Nanovirus Nanovirus Nanovirus Nanovirus

Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite

Ageratum leaf curl Buea betasatellite Ageratum leaf curl Cameroon betasatellite Ageratum yellow leaf curl betasatellite Ageratum yellow vein betasatellite Ageratum yellow vein India betasatellite Ageratum yellow vein Sri Lanka betasatellite Alternanthera yellow vein betasatellite Andrographis yellow vein leaf curl betasatellite Bhendi yellow vein mosaic betasatellite Cardiospermum yellow leaf curl betasatellite Chili leaf curl betasatellite Chili leaf curl Jaunpur betasatellite Chili leaf curl Sri Lanka betasatellite Cotton leaf curl Gezira betasatellite Cotton leaf curl Multan betasatellite Croton yellow vein mosaic betasatellite Eupatorium yellow vein betasatellite Eupatorium yellow vein mosaic betasatellite French bean leaf curl betasatellite Hedyotis yellow mosaic betasatellite Honeysuckle yellow vein betasatellite Honeysuckle yellow vein mosaic betasatellite Malvastrum leaf curl betasatellite Malvastrum leaf curl Guangdong betasatellite Mirabilis leaf curl betasatellite Momordica yellow mosaic betasatellite Mungbean yellow mosaic betasatellite Okra leaf curl Oman betasatellite Papaya leaf curl betasatellite Papaya leaf curl China betasatellite Papaya leaf curl India betasatellite Rhynchosia yellow mosaic betasatellite Rose leaf curl betasatellite Siegesbeckia yellow vein betasatellite Tobacco curly shoot betasatellite Tobacco leaf curl betasatellite Tobacco leaf curl Japan betasatellite Tobacco leaf curl Patna betasatellite Tomato leaf curl Bangalore betasatellite Tomato leaf curl Bangladesh betasatellite Tomato leaf curl betasatellite Tomato leaf curl China betasatellite Tomato leaf curl Gandhinagar betasatellite Tomato leaf curl Java betasatellite Tomato leaf curl Joydebpur betasatellite Tomato leaf curl Laguna betasatellite Tomato leaf curl Laos betasatellite Tomato leaf curl Malaysia betasatellite Tomato leaf curl Nepal betasatellite Tomato leaf curl Patna betasatellite Tomato leaf curl Philippine betasatellite Tomato leaf curl Sri Lanka betasatellite Tomato leaf curl Yemen betasatellite Tomato yellow leaf curl China betasatellite

FR717140 FM164737 AJ316026 AJ252072 AJ557441 AJ542498 DQ641716 KC967282 AJ308425 AM933578 AJ316032 HM007103 JN638445 DQ644564 AJ298903 AM410551 AJ438938 AB300464 JQ866298 KF641186 AJ316040 AB182263 AM072289 KF912951 KT454829 LK054803 JX443646 KF267444 AY244706 KJ642219 HM143906 GQ478344 KP752092 KF499590 AJ421484 AM260465 HQ180394 AJ316036 AY428768 AB236324 AJ542489 AJ704609 KC952006 KC282642 AJ966244 AB307732 AJ542491 KM051528 AJ542492 EU862324 AB308071 AJ542493 JF919717 AJ420313

ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant

Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus (Continued )

Tolecusatellitidae

688

Table 3

Satellite Nucleic Acids and Viruses

Continued

Family/Subfamily

Genus

Species

Type species Genome/Host typesa Helper virus genus accession #

Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Betasatellite Deltasatellite Deltasatellite Deltasatellite Deltasatellite Deltasatellite Deltasatellite Deltasatellite Deltasatellite Deltasatellite Deltasatellite Deltasatellite

Tomato yellow leaf curl Rajasthan betasatellite Tomato yellow leaf curl Shandong betasatellite Tomato yellow leaf curl Thailand betasatellite Tomato yellow leaf curl Vietnam betasatellite Tomato yellow leaf curl Yunnan betasatellite Vernonia yellow vein betasatellite Vernonia yellow vein Fujian betasatellite Croton yellow vein deltasatellite Malvastrum leaf curl deltasatellite Sida golden yellow vein deltasatellite 1 Sida golden yellow vein deltasatellite 2 Sida golden yellow vein deltasatellite 3 Sweet potato leaf curl deltasatellite 1 Sweet potato leaf curl deltasatellite 2 Sweet potato leaf curl deltasatellite 3 Tomato leaf curl deltasatellite Tomato yellow leaf distortion deltasatellite 1 Tomato yellow leaf distortion deltasatellite 2

AY438558 KP322555 AJ566746 DQ641714 KF640694 JF733779 FN435836 AJ968684 KF433066 JN986808 JN819490 JN819498 FJ914390 KF716173 KT099179 U74627 JN819495 KU232893

ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant ssDNA/Plant

Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus Begomovirus

a

ssDNA, single-stranded DNA.

essential roles as components of disease complexes. Although they do not encode functional proteins, some members can attenuate or exacerbate the symptoms induced by HV infection. Category 8c: Small circular satellite ssRNAs Members have small (0.35 kb) genomes that may be present in both circular and linear forms in infected cells. They include satellite RNAs associated with viruses in the plant virus families Secoviridae, Luteoviridae, and Solemoviridae; and possibly a viroidlike satellite RNA (Table 4). There is no mRNA activity associated with these various satellite RNAs, and they all have a high degree of secondary structure. The satellite RNAs either have no effect on (Solemoviridae-associated satellites) or reduce the accumulation of (Secoviridae- and Luteoviridae-associated satellites) their HV. Category 8d: Hepadnavirus-associated satellite-like RNAs (genus Deltavirus) Hepatitis delta virus (HDV, unassigned genus Deltavirus) is a subviral agent of the human pathogen, hepatitis B virus (HBV, genus Orthohepadnavirus, family Hepadnaviridae). HDV is replicated in the nucleus by the host DNA-dependent RNA polymerase II, but is dependent upon the envelope proteins of HBV for its encapsidation and transmission, and is thus not a satellite. The HDV RNA (1679 nt) folds into an unbranched rod-like structure similar to that of viroids, and in common with the circular ssRNA satellites, contains ribozymes that are involved in the processing and maturation of this RNA during replication. The HDV RNA encodes two forms of a protein designated the delta antigen (δAg). The smaller, 22 kDa form (δAg-S), which is a nuclear phosphoprotein, is required for replication of HDV RNA, while the 24 kDa larger form (δAg-L), which contains an additional 19 aa at the C-terminus and is produced late in infection, is required along with δAg-S for assembly of the HDV RNA into HBV envelope particles. δAg-L also acts as an inhibitor of HDV replication. HDV intensifies the severity of liver disease caused by HBV. Category 8e: Polerovirus-associated RNAs Some isolates of three poleroviruses (genus Polerovirus, family Luteoviridae), Beet western yellows virus, carrot red leaf virus (CRLV), and Tobacco vein distorting virus were found to have associated with them additional RNAs of size 2.8–3 kb. These associated RNAs encoded their own viral replicases and are thus capable of autonomous replication. However, they each depend on their respective helper poleroviruses for movement, encapsidation, and transmission. Some members, such as CRLV-associated RNA, contribute to the disease caused by their HV.

Replication and Structure Satellite RNAs A general property of satellite RNAs is that their replication depends on the replication machinery of the HV, which involves virusand host-encoded factors. Hence, replication of satellite RNAs depends on interactions with both the HV and the host plant. Satellite RNAs may or may not share structural features at their 50 and 30 termini in common with the HV RNA. For instance, the large satellite RNAs of the nepoviruses have an HV-encoded VPg at the 50 end and a poly(A) tail at the 30 end, as do their HV RNAs, but the satellite RNAs of cucumber mosaic virus (CMV) have a 50 cap structure, as do the CMV RNAs. But unlike the HV, the satellite RNAs do not have a tRNA-like structure that can be validated at the 30 end. Structural differences and, often, differences in the replication process between the satellite RNA and the HV RNA, indicate that the replication machinery of the HV may need to

Satellite Nucleic Acids and Viruses

Table 4

689

Classification of satellites RNA nucleic acids

Category/Species and Tentative Species

Type isolatea accession # Genome/Host typesb Helper virus genus

7. Satellite dsRNAs Bombyx mori cypovirus 1 satellite RNA AB183384 Satellite of Atkinsonella hypoxylon virus L39127 Satellite of Gremmeniella abietina virus MS1 AY089995 Satellite of Penicillium stoloniferum virus F AY738338 Satellite of Zygosaccharomyces bailii virus AF515592 Satellites of Amasya cherry disease-associated virus: Sat A, Sat B AM085138, AM085139 Satellites of cherry chlorotic rusty spot -associated virus: Sat A, Sat B, Sat C N/A Satellites of Discula destructiva virus 1: dsRNA 3, dsRNA4 AF316994, AF316995 U78817, X56987, N/A Satellites of Saccharomyces cerevisiae L-A virus: M1, M2, M28 Satellites of Trichomonas vaginalis T1 virus U15991 Satellites of Ustilago maydis virus H: M-P1, M-P4, M-P6 M63149, L12226, P16948

dsRNA/Fungi dsRNA/Fungi dsRNA/Fungi dsRNA/Fungi dsRNA/Fungi dsRNA/Fungi dsRNA/Fungi dsRNA/Fungi dsRNA/Fungi dsRNA/Fungi dsRNA/Fungi

Cypovirus Betapartitivirus Gammapartitivirus Gammapartitivirus Zybavirus Alphachrysovirus Alphapartitivirus Gammapartitivirus Totivirus Trichomonasvirus Totivirus

8a. Large linear satellite ssRNAs Arabis mosaic virus large satellite RNA Bamboo mosaic virus satellite RNA Beet necrotic yellow vein virus RNA5* Beet ringspot virus satellite RNA Blackcurrant reversion virus satellite RNA Chicory yellow mottle virus large satellite RNA Grapevine Bulgarian latent virus satellite RNA Grapevine fanleaf virus satellite RNA Myrobalan latent ringspot virus satellite RNA Strawberry latent ringspot virus satellite RNA Tomato black ring virus satellite RNA

D00664 L22762 D63759 KX033801 AF112119 D00686 N/A D00442 N/A X69826 X00978

ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant

Nepovirus Potexvirus Benyvirus Nepovirus Nepovirus Nepovirus Nepovirus Nepovirus Nepovirus Nepovirus Nepovirus

8b. Small linear satellite ssRNAs Artichoke mottled crinkle virus satellite RNA Beet black scorch virus satellite RNA Carnation Italian ringspot virus satellite RNA Carrot mottle mimic virus satellite RNA Cucumber mosaic virus satellite RNA (several types) Cymbidium ringspot virus satellite RNA Groundnut rosette virus satellite RNA* Panicum mosaic virus satellite RNA Pea enation mosaic virus satellite RNA Peanut stunt virus satellite RNA Pelargonium leaf curl virus satellite RNA Petunia asteroid mosaic virus satellite RNA Tobacco bushy top virus satellite RNA* Tobacco necrosis virus small satellite RNA Tomato bushy stunt virus satellite RNA (several types)

N/A AY394497 N/A EU914919 X69136 D00720 Z29702 M17182 U03564 Z98197 N/A N/A N/A E03054 AF022787–8, FJ666076

ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant

Tombusvirus Betanecrovirus Tombusvirus Umbravirus Cucumovirus Tombusvirus Umbravirus Panicovirus Umbravirus Cucumovirus Tombusvirus Tombusvirus Umbravirus Alphanecrovirus Tombusvirus

8c. Small circular satellite ssRNAs Arabis mosaic virus small satellite RNA Cereal yellow dwarf virus-RPV satellite RNA Cherry small circular viroid-like RNA Chicory yellow mottle virus satellite RNA Lucerne transient streak virus satellite RNA Rice yellow mottle virus satellite Solanum nodiflorum mottle virus satellite RNA Subterranean clover mottle virus satellite RNA (2 types) Tobacco ringspot virus satellite RNA Velvet tobacco mottle virus satellite RNA

M21212 M63666 Y12833 D00721 X01984 AF039909 J02386 M33000, M33001 M14879 J02439

ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant ssRNA/Plant

Nepovirus Polerovirus Nepovirus Sobemovirus Sobemovirus Sobemovirus Sobemovirus Nepovirus Sobemovirus

8d. Hepadnavirus-associated RNAs Hepatitis delta virus

M21012

ssRNA/Animal

Orthohepadnavirus

8e. Polerovirus-associated RNAs Beet western yellows virus ST9-associated RNA Carrot red leaf virus-associated RNA Tobacco vein distorting virus-associated RNA

L04281 AF020617 EF529625

ssRNA/Plant ssRNA/Plant ssRNA/Plant

Polerovirus Polerovirus Polerovirus

a

N/A, unavailable. ssRNA, single-stranded RNA; dsRNA, double-stranded RNA.

b

690

Satellite Nucleic Acids and Viruses

be adapted to replicate the satellite RNA, in ways which are not completely understood. The HV replication complex could be modified by satellite-encoded factors, as has been hypothesized for the large satellite RNAs of nepoviruses, or by unidentified host factors. In the best characterized systems, the efficiency of replication depends on the host plant. For CMV satellite RNAs, replication efficiency also depends on the strain of HV. Similarly, while the expression of the HV RNA-dependent RNA polymerase was enough for the replication of cereal yellow dwarf virus (CYDV) satellite RNA in the homologous host, this was not the case for Cymbidium mosaic virus satellite RNA in a heterologous host system, in which, however, a diRNA was amplified. Replication has been studied best in the small noncoding satellite RNAs. For the small linear satellite RNAs of TCV and CMV, multimeric forms of both positive (arbitrarily defined as the encapsidated sense) and negative sense are found in infected tissues. The junction between monomers can be perfect or have deletions. Circular forms are not found, and replication does not proceed through a rolling-circle mechanism. Regulatory sequences for RNA replication have been mapped in detail in the satellite RNAs of TCV. In the hybrid satC molecule, a hairpin structure is a replication enhancer and has a role in depressing the accumulation of the HV. Also, structural motifs favoring recombination have been analyzed extensively in satD and satC of TCV. The replication of small, circular satellite RNAs is by a rolling-circle mechanism; upon infection, linear, multimeric forms of the negative strand are synthesized. The multimeric plus strand is then synthesized on this template or on negative strand circular monomers, depending on the satellite RNA. The positive strand multimer is cleaved autocatalytically and ligation occurs for those satellite RNAs that are encapsidated as a circular form. Hammerhead and hairpin ribozyme structures are found in these satellite RNAs monomers or dimers, which catalyze hydrolysis or hydrolysis and ligation, respectively. Other structures required for replication and encapsidation have been mapped in CYDV satellite RNA. Structural analyses have been done mostly with small satellite RNAs. Models for the in vitro secondary structure have been proposed for several satellite RNAs (CMV satellite RNA, TCV satellite RNA, TRSV satellite RNA, and CYDV satellite RNA) based on nuclease sensitivity and chemical modification of bases data. For CMV satellite RNA, in vivo models have also been proposed. All analyzed satellite RNAs have a high degree of secondary structure, with more than 50% and up to 70% of bases involved in pairing. The high degree of secondary structure could explain the high stability of these molecules as well as the very high infectivity reported for some of them, such as CMV satellite RNA and TCV satellite RNA. The high degree of secondary structure may also be related to their biological activity, which for these noncoding molecules must depend on structural features. As has been detailed in other sections, sequences, structural domains, and tertiary interactions involved in pathogenicity and replication have been well characterized for some small satellite RNAs.

Satellite DNAs The replication of satellite ssDNA mimics that of the HV. Unlike RNA viruses, geminiviruses do not encode a polymerase and depend on host DNA polymerases for replication. The only geminivirus-encoded protein essential for replication is Rep. Satellite DNAs utilize the HV Rep and contain the conserved sequence for DNA nicking by the Rep. Replication takes place by a combination of rolling circle and recombination dependent mechanisms and the DNA is encapsidated in the HV virion. The replication is independent of bC1 expression. Sequence comparisons at the nucleotide and aa levels reveal several conserved structural features, despite a significant level of sequence divergence. These include an A-rich region, a sequence of about 80 nt referred to as the satellite conserved region, and putative stem-loop structure containing a mononucleotide motif, also present in geminivirus genomes.

Sequence Variation and Evolution Sequence variation and evolution has been analyzed most for CMV satellite RNA. Early experiments under controlled conditions showed a high variability and genetic plasticity. Populations started from cDNA clones rapidly evolved to a swarm of sequences whose master sequence changed upon passage on different hosts. Analysis of genetic variants from natural populations showed again a high diversity for CMV satellite RNA. In natural populations, satellite RNA diversification and evolution proceeded by mutation accumulation and by recombination. Constraints to genetic variation were analyzed, and were related to the maintenance of base pairing in secondary structure elements. Population diversity of CMV satellite RNA was higher than for the HV, and the population structures of CMV and the satellite RNA were not correlated. These analyses also showed that CMV satellite RNA behaved in nature as a molecular hyperparasite spreading epidemically on the CMV populations. The incidence of CMV satellite RNA in CMV populations has been shown to be low, except during episodes of epidemics of tomato necrosis. High diversity of natural populations has also been shown for panicum mosaic satellite virus RNA and for BaMV satellite RNA. For BaMV satellite RNA, the diversity was highest in the NTRs. Conversely, the population of rice yellow mottle virus satellite RNAs across sub-Saharan Africa showed little sequence diversity. The incidence of these three satellite RNAs in the analyzed populations of the respective HV was, in all cases, high. Phylogenetic analyses divide geminivirus-related satellites broadly into two categories, those isolated from host plants of the family Malvaceae and the rest isolated from other plants, mostly in the Solanaceae. Compared to satellite RNAs, satellite DNAs do not exhibit a wide diversity. They are structurally similar and lack strict specificity for HV species. Geminivirus-related satellite DNA sequences analyzed exhibit a minimum overall similarity of about 47% and 37%, at the nt level and aa level, respectively. There has been much speculation on the origin of satellite RNAs. For CMV satellite RNA and TCV satellite RNA, it has been suggested that they could have been generated out of small sequences synthesized upon the HV RNA as a template by the HV

Satellite Nucleic Acids and Viruses

691

polymerase. This hypothesis is no longer sustained since attempts to locate the sequences with significant similarity to satellite RNA in the genomes of hosts, HVs and vectors have in general been unsuccessful. Whatever the mechanism of generation of the satellite RNAs, it seems that the satellitism as a phenomenon, that is, the dependence on a certain virus for replication, has evolved independently several times. This is suggested by the lack of correlation between satellite and HV taxonomy, and by the existence of subviral nucleic acids with different degrees of dependence on an HV. For instance, an evolutionary line from nondependent viruses to satellites such as BaMV satellite RNA through satellite viruses could have proceeded by size and information content reduction. A phylogenetic relationship between viroids and small circular satellite RNAs has also been proposed.

Expression of Foreign Sequences From Satellite Vectors The satellite RNAs of BaMV and BNYVV, as well as the satellite DNA of tomato yellow leaf curl China virus isolate Y10 (TYLCCNV-Y10), all of which express proteins not required for the replication or spread of the satellite or HV, have been used as expression vectors of foreign sequences. In the case of the satellite RNA of BaMV, the expression of such sequences and the accumulation of the satellite RNA were reduced considerably in systemically infected leaves. Nevertheless, this satellite expression system has been useful for studying cis- and trans-acting replication signals. The 26 kDa protein encoded by RNA5 of BNYYV has been replaced with the sequence encoding the green fluorescent protein, which was expressed in both the inoculated leaves and systemically infected leaves. The small size of the satellite DNA of TYLCCNV-Y10 precludes use of this system for expression of most genes, but the system has been used to express plant gene segments inducing RNA silencing in several plants species. The STMSV system also has been used to express plant gene sequences, in place of its CP, resulting in RNA silencing of those plant genes, but not of the satellite virus or its HV. The modified STMSV expression vector was able to spread through the plant in the absence of its CP.

Satellite-Mediated Control of Viruses The ability of some satellite RNAs to interfere with the replication of, and attenuate the symptoms induced by, their HV has been exploited to develop strategies for disease control. Two prophylactic strategies have been employed for this biocontrol approach. In the first strategy (cross-protection), the target plants are pre-inoculated with a combination of the satellite RNA and a mild strain of its HV to “vaccinate” the plant against a virulent strain of the HV. The second strategy utilizes the transgenic approach wherein the satellite RNA have been expressed in transgenic plants to provide either resistance to infection by the respective HV, or tolerance to the disease induced by the HV and/or related pathogenic satellites. Multiple mechanisms like competition for viral replicase, gene silencing and possibly other immune actions of the host may be involved in resistance. A recent study evaluated the effects of post‐inoculation with CMV satellite RNA on symptoms derived from pre‐infection by its HV and demonstrated that this new therapeutic strategy could be useful for combating plant viruses using properly timed treatment with attenuation satellite RNAs.

See also: Alphasatellites (Alphasatellitidae). Beet Necrotic Yellow Vein Virus (Benyviridae). Betasatellites and Deltasatelliles (Tolecusatellitidae). Geminiviruses (Geminiviridae). Luteoviruses (Luteoviridae). Machlomovirus and Panicoviruses (Tombusviridae). Nanoviruses (Nanoviridae). Plant Satellite Viruses (Albetovirus, Aumaivirus, Papanivirus, Virtovirus). Secoviruses (Secoviridae). Tombusviruses (Tombusviridae)

Further Reading Bonami, J.-R., Sri Widada, J., 2011. Viral diseases of the giant freshwater prawn Macrobrachium rosenbergii: A review. Journal of Invertebrate Pathology 106, 131–142. Briddon, R.W., Martin, D.P., Roumagnac, P., et al., 2018. Alphasatellitidae: A new family with two subfamilies for the classification of geminivirus- and nanovirus-associated alphasatellites. Archives of Virology 163, 2587–2600. Cao, X., Liu, S., Yu, C., Li, X., Yuan, X., 2019. A new strategy of using satellite RNA to control viral plant diseases: Post-inoculation with satellite RNA attenuates symptoms derived from pre-infection with its helper virus. Plant Biotechnology Journal 17, 1856–1858. Cui, H.G., Liu, H.Z., Chen, J., et al., 2015. Genetic diversity of prunus necrotic ringspot virus infecting stone fruit trees grown at seven regions in China and differentiation of three phylogroups by multiplex RT-PCR. Crop Protection 74, 30–36. Dodds, J.A., 1998. Satellite tobacco mosaic virus. Annual Review of Phytopathology 36, 295–310. Dry, I.B., Krake, L.R., Rigden, J.E., Rezaian, M.A., 1997. A novel subviral agent associated with a geminivirus: The first report of a DNA satellite. PNAS 94, 7088–7093. Gosselé, V., Metzlaff, M., 2006. Using satellite Tobacco mosaic virus vectors for gene silencing. Current Protocols in Microbiology 00 (1), 16I.5.1–16I.5.17. Idris, A.M., Shahid, M.S., Briddon, R.W., et al., 2011. An unusual alphasatellite associated with monopartite begomoviruses attenuates symptoms and reduces betasatellite accumulation. Journal of General Virology 92, 706–717. Kassanis, B., 1962. Properties and behaviour of a virus depending for its multiplication on another. Microbiology 27, 477–488. Kassanis, B., White, R.F., 1972. Interference between two satellite viruses of tobacco necrosis virus. Journal of General Virology 17, 177–183. Krupovic, M., Kuhn, J.H., Fischer, M.G., 2016. A classification system for virophages and satellite viruses. Archives of Virology 161, 233–247. Ribière, M., Olivier, V., Blanchard, P., 2010. Chronic bee paralysis: A disease and a virus like no other? Journal of Invertebrate Pathology 103, S120–S131. Schneider, I.R., 1969. Satellite-like particle of Tobacco ringspot virus that resembles Tobacco ringspot virus. Science 166, 1627–1629. Zhou, X., 2013. Advances in understanding begomovirus satellites. Annual Review of Phytopathology 51, 357–381. Zinn, E., Vandenberghe, L.H., 2014. Adeno-associated virus: Fit to serve. Current Opinion in Virology 8, 90–97.

Secoviruses (Secoviridae) Jeremy R Thompson, Cornell University, Ithaca, NY, United States r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

MP Movement protein nt Nucleotide(s) ORF Open reading frame RdRp RNA-dependent RNA polymerase RISC RNA-induced silencing complex satRNA Satellite RNA UTR Un-translated region VIGS Virus-induced gene silencing VLPs Virus-like particles VPg Viral protein genome-linked VRC Virus replication complexes vRNA Virion RNA

aa Amino acid(s) AGO Argonaute 1 CITE Cap-independent translation enhancer Co-Pro Protease cofactor CP Coat protein or capsid protein ELISA Enzyme-linked immunological assays ER Endoplasmic reticulum HC-Pro Helper component-protease IRES Internal ribosomal entry site kb Kilobase kDa Kilo dalton LAMP Loop mediated amplification

Glossary Affimers Small proteins that mimic antibodies. Argonaute 1 Member of the Argonaute family of proteins involved in RNA silencing and a component of the RNA-induced silencing complex. Glutamic protease A proteolytic enzyme that contains a glutamic acid residue within the active site. Internal ribosomal entry site An RNA element that facilitates cap-independent translation. Satellite RNA RNA that depends on the helper virus for replication and encapsidation, and can in some cases modulate symptoms.

Viral protein genome-linked A protein covalently attached to the 50 end of positive strand viral RNA and acting as a primer during RNA synthesis. Virus re-assortment The mixing of the genome segments of a virus into new combinations. Virus replication complexes Intracellular complexes formed by association with intracellular membranes that are sites for stages in the replication cycle of a virus. b-barrel Beta-sheet composed of tandem repeats forming a closed structure.

Introduction Secoviruses are the plant-infecting members of the taxonomic order Picornavirales, a group of viruses that includes more broadly known members such as poliovirus, rhinovirus and foot-and-mouth disease virus. Symptoms of Grapevine fanleaf virus (GFLV) were some of the first to be described independently across a number of European countries at the end of the nineteenth century. In the early 1960s the plant virus group to which GFLV belongs, the nematode-transmissible nepoviruses, was one of the first to be recognized, followed by, in subsequent decades, the related comovirus and fabavirus groups. Unlike all other members of the Picornavirales order which contain their genetic information on one positive sense strand of RNA (monopartite), the comoviruses, fabaviruses and nepoviruses are segmented, meaning they have their genome split in two. This organization is typical for the vast majority of secoviruses that have been characterized so far. However, members of the sequivirus and waikavirus groups – the latter represented by the type member Rice tungro spherical virus (RTSV), a causal component of the eponymous disease that devastated South East Asia in the 1960s and 70s – are the exceptions with an unsegmented monopartite genome. The secoviruses, like all members of the Picornavirales have a number of common features. Their genomic segments produce polyproteins (contiguous clusters of functional domains) which are proteolytically cleaved into smaller functional proteins by a virus protease which is encoded within a commonly conserved helicase-protease-polymerase replication block. They are all predicted to encode a small protein (VPg) that binds to the 50 end of their genome. Virus particles are non-enveloped and approximately 30 nm in diameter. The structure of Cowpea mosaic virus, one of the frontrunners in the development of nanobiotechnological tools, is particularly well-characterized. Since the 2000s, exceptions to the general rule that genomic RNA segments encode single polyproteins within the Picornavirales order have been identified. For secoviruses, evidence has come with the emergence and characterization of the torradoviruses which are apparently bicistronic for one of their genome segments, encoding a predicted protein of as yet unknown function. Another exception to assumed commonalities has been the recent discovery that Strawberry mottle virus (SMoV) encodes for a glutamic protease, in addition to the 3C-like protease conserved across all other secoviruses. The former protease is the first of its kind reported for any positive single stranded RNA virus.

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Taxonomy, Phylogeny, and Evolution The International Committee for the Taxonomy of Viruses (ICTV) (see “Relevant Websites section”) lists specific demarcation criteria for the recognition of taxa which are to be used as guidelines on a case-by-case basis for determining taxonomies. The family Secoviridae, is classified within the Picornavirales order (a name derived from fusing pico meaning ‘small’prefixed to ‘RNA’), a group of viruses which encode their genome on single stranded positive sense RNA. The Secoviridae is the only plantinfecting member of this group. Other family members (along with host and typical species) include; Dicistroviridae (arthropods – Cricket paralysis virus), Iflaviridae (arthropods – Infectious flacherie virus), Marnaviridae (algae – Heterosigma akashiwo RNA virus), Picornaviridae (vertebrates – Enterovirus C), Polycipiviridae (arthropods – Solenopsis invicta virus 2). There are also two floating genera; Bacillarnavirus and Labyrnavirus infecting algae and protists, respectively. At present, the family Secoviridae contains eighty-six species, eighty-one of which are grouped into eight genera. The vast majority (sixty-two) of species fall into three related genera (Comovirus, Fabavirus and Nepovirus) that form the subfamily Comovirinae. Nineteen species fall within the remaining five genera; Cheravirus, Sadwavirus, Sequivirus, Torradovirus and Waikavirus. There are five unassigned species Chocolate lily virus A (CLVA), Black raspberry necrosis virus (BRNV), Dioscorea mosaic associated virus (DMaV), Strawberry mottle virus (SMoV), and Strawberry latent ringspot virus (SLRSV). Recent taxonomical changes, not yet ratified by the ICTV, will place four of these species within the Sadwavirus genus leaving only SLRSV as unassigned. Absorption of these four species to the Sadwavirus genus was achieved through the generation of three new subgenera, Cholivirus, Satsumavirus and Stramovirus. The Nepovirus genus is usually informally split into three subgroups A, B and C depending on phylogenetic relationships (Fig. 1). These relationships for all members of the order Picornavirales have been traditionally defined by a contiguous region (Pro-Pol) spanning the functional domains of the protease (Pro) and polymerase (Pol), characterized at each extreme by the amino acids CG and GDD, respectively (Fig. 2). Trees produced from alignments of other functional domains on the whole are less structured but are not incongruent with the relationships defined by Pro-Pol trees. Notably, monophyly of the genera Sequivirus, Torradovirus and Waikavirus has been detected for the helicase and coat protein domains. There is substantial evidence for the modular nature of virus genomes and despite the overall lack of phylogenetic discordance across their functional domains the secoviruses possess genetic characteristics that are modular in nature as is suggested by the detection of distant, yet significantly closer affinities with functional domains of invertebrate-infecting members of the Picornavirales than with fellow secoviruses. The experimentally demonstrated existence of a completely novel glutamic protease encoded by SMoV, is in line with this paradigm. For most secoviruses, functional domains have been ascribed based on their similarity in sequence to proteins of known function. Because of the differences in their hosts and modes of infection, within the order Picornavirales, MPs are unique to the secoviruses. For plant viruses in general MPs are believed to be ultimately host in origin, for secoviruses these origins are possibly multiple as illustrated in tomato-infecting and non-tomato-infecting torradoviruses where putative MPs have their closest similarities to either umbraviruses and bromoviruses, respectively. Current models using comparative genomics suggest secoviruses are likely invertebrate in origin. For this to be the case there must have been one or many host-switching events (invertebrate to plant) leading to a period of in planta adaptation followed by virus emergence. The intimate relationship between virus, host and vector provides ample opportunity for host-switching in either direction (invertebrate to plant or plant to invertebrate) to have occurred. The mechanisms necessary for such a significant (interkingdom) evolutionary step to occur are still to be discovered, but unlike members of other virus families (e.g., Reoviridae, Bunyaviridae) which replicate in both plant and vector, the secoviruses have apparently lost their capacity to infect the latter. Attempts to establish when members of the Secoviridae might have evolved from ancestral viruses has been done using coalescent analyses on ninety-four coat protein sequences of the subfamily Comovirinae. An estimated time of emergence of less than a 1000 years

Fig. 1 Schematic of the taxonomic organization of the family Secoviridae showing typical features for each taxon. Horizontal numbers – present number of species per taxon. Vertical numbers – year taxon was recognized (irrespective of name given). Rounded rectangles (gray) – subfamily; (colored) – genera. Ellipses – subgenus groupings. Note – listed features may not apply to all members of group. * – presented subgenera organization of the Sadwavirus genus has yet to be ratified by the ICTV.

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Fig. 2 Maximum likelihood inferred phylogenetic unrooted tree of members of the family Secoviridae based on an alignment of amino acid sequences of the conserved domains between the “CG” motif of the 3C-proteinase and the “GDD” motif of the polymerase (Pro-Pol region). The bar represents the genetic distance. Rounded rectangles (gray) – subfamily; (colored) – genera. For each species, the sequence of the exemplar isolate was used for the alignments (see species tables for the sequence accession numbers). Percent bootstrap values (1000 replicates) are shown at each branch node. Branches with bootstrap values below 70% were collapsed. Virus names and abbreviations are listed in Table 1.

ago was calculated for the whole group with individual species emerging at various times between 50 and 250 years ago a period coinciding with an intensification in agricultural practices. Such estimates overlook possible variations in mutation rates and the birth and death of virus lineages over time. Nevertheless, this information provides a foundation to work from as computational phylogenetic methods become more sophisticated and the number of secovirus sequences deposited in the database grows. The bipartite nature of most secoviruses is a likely adaptation from monopartite ancestors. In the virosphere those segmented viral genomes so far discovered are predominantly plant-infecting. Segmentation results in an uncoupling of the structural block of proteins from the replication–associated protein block allowing a more refined control of translation. It also facilitates genetic shuffling of each RNA to quickly produce novel combinations (reassortants) that may or may not have a selective advantage. Genome splitting might also allow for more efficient virion packaging. A bipartite version of foot and mouth disease picornavirus, produced spontaneously by multiple passages, was demonstrated to produce more stable virions than its monopartite counterpart, with presumably a longer viability outside of the cell. Nevertheless, a bipartite genome strategy means that both segments (and virions containing each segment) have to infect the same cell for successful infection, a theoretically more probable scenario for plants viruses, as their entry into the first cells of the host will be mechanically-mediated rather than receptor-mediated as for animal viruses. Genetic mixing, whether reassortment or recombination, an important driver of evolution, requires a physical proximity of divergent viruses achieved by mixed infections which will be prolonged in perennial hosts. Evidence for the reassortment of secovirus RNA1 and 2 has been found for a raspberry isolate of Tomato ringspot virus (ToRSV) and a number of Korean isolates of Broad bean wilt virus 2 (BBWV2). Recombination is more frequently detected, at times leading to the putative emergence of new species as in the case of grapevine chrome mosaic and grapevine deformation nepoviruses.

Members of the Family There are presently eighty-six recognized species of secoviruses distributed across eight genera, with one unassigned species SLRSV. For the majority (62) full-length genomic sequences are available including a NCBI RefSeq assembly number. For others (24) there is still no sequence information available (Fig. 1; Table 1).

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Table 1 List of all ICTV classified members of the Secoviridae, their taxonomic order, acronyms and GenBank sequence information. # Seq – number of sequences for each virus registered in NCBI Taxonomy Browser data from which a complete genome has been assembled to give a RefSeq assembly number (GCF_number). The last column to the right provides explanation if no RefSeq assembly number is available in the form of accession numbers for incomplete viral sequences or failing that, when no sequence information is available at all, the year, (in parentheses) of the species ratification by the International Committee for the Taxonomy of Viruses (ICTV) Subfamily

Genus

Comovirinae Comovirus

Subgenus

Species

Abbreviation # Seq RefSeq assembly

–

Andean potato mottle virus Bean pod mottle virus Bean rugose mosaic virus Broad bean stain virus Broad bean true mosaic virus Cowpea mosaic virus Cowpea severe mosaic virus Glycine mosaic virus Pea green mottle virus Pea mild mosaic virus Quail pea mosaic virus Radish mosaic virus Red clover mottle virus Squash mosaic virus Ullucus virus C Broad bean wilt virus 1 Broad bean wilt virus 2 Cucurbit mild mosaic virus Gentian mosaic virus Grapevine fabavirus Lamium mild mosaic virus Prunus virus F Aeonium ringspot virus Apricot latent ringspot virus Arabis mosaic virus Arracacha virus A Artichoke Aegean ringspot virus Artichoke Italian latent virus Artichoke yellow ringspot virus Beet ringspot virus Blackcurrant reversion virus Blueberry latent spherical virus Blueberry leaf mottle virus Cassava American latent virus Cassava green mottle virus Cherry leaf roll virus Chicory yellow mottle virus Cocoa necrosis virus Crimson clover latent virus Cycas necrotic stunt virus Grapevine Anatolian ringspot virus Grapevine Bulgarian latent virus Grapevine chrome mosaic virus Grapevine deformation virus Grapevine fanleaf virus Grapevine Tunisian ringspot virus Hibiscus latent ringspot virus Lucerne Australian latent virus Melon mild mottle virus Mulberry mosaic leaf roll associated virus Mulberry ringspot virus Myrobalan latent ringspot virus Olive latent ringspot virus Peach rosette mosaic virus Potato black ringspot virus

APMV BPMV BRMV BBSVa BBTMVb CPMV CPSMV GMV PGMV PMiMV QPMV RaMV RCMV SqMV UVC BBWV1 BBWV2 CuMMV GeMV GFabV LMMV PrVF AeRSV ALRSV ArMV AVA AARSV AILV AYRSV BRSV BRV BLSV BLMV CsALV CGMV CLRV ChYMV CoNV CCLV CNSV GARSV GBLV GCMV GDeV GFLV GTuRSV HLRSV LALV MMMoV MMLRaV

8 46 10 2 7 33 27 0 0 0 0 18 13 33 0 123 187 5 12 18 4 18 4 2 150 2 0 9 6 12 113 4 5 0 2 395 0 2 0 13 5 5 8 5 788 0 0 0 4 4

GCF_002833585.1 GCF_000862925.1 GCF_001430295.1 – GCF_000910755.1 GCF_000860385.1 GCF_000861225.1 – – – – GCF_000879335.1 GCF_000860425.1 GCF_000861625.1 – GCF_000853465.1 GCF_000861605.1 GCF_002867105.1 GCF_000880435.1 GCF_003033815.1 GCF_000916715.1 GCF_003033845.1 GCF_002867145.1 – GCF_000855205.1 – – GCF_005410585.1 GCF_002986065.1 GCF_000860405.1 GCF_000849565.1 GCF_002867165.1 – – – GCF_000893515.1 – – – GCF_000860925.1 GCF_000897495.1 GCF_000892035.1 GCF_000862025.1 GCF_000898115.1 GCF_000860305.1 – – – GCF_002867185.1 GCF_002867205.1

MRSV MLRSV OLRSV PRMV PBRSV

0 0 2 7 16

– (1979) – (1979) GCF_002986085.1 GCF_002029615.1 GCF_000913055.1

Fabavirus

–

Nepovirus

–

GenBank accession (ICTV entry)

FJ028650.2, KJ746622.1

(1982) (1995) (1991) (1976)

(1995)

AJ278875 KY569301.1, KY569302.1 (1998)

U20622.1, U20621.1 (1991) MG581962.1, MG581963.1 D00685.1, D00686.1c EU741694 (1982)

(1991) (1982) (1982)

(Continued )

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Table 1

n/a

Secoviruses (Secoviridae)

Continued Potato virus B Potato virus U Raspberry ringspot virus Soybean latent spherical virus Tobacco ringspot virus Tomato black ring virus Tomato ringspot virus Cheravirus – Apple latent spherical virus Arracacha virus B Cherry rasp leaf virus Currant latent virus Stocky prune virus Sadwavirusd Satsumavirus Satsuma dwarf virus Stramovirus Strawberry mottle virus Black raspberry necrosis virus Choliavirus Chocolate lily virus A Dioscorea mosaic associated virus Sequivirus – Carrot necrotic dieback virus Dandelion yellow mosaic virus Parsnip yellow fleck virus Torradovirus – Carrot torradovirus 1 Lettuce necrotic leaf curl virus Motherwort yellow mottle virus Squash chlorotic leaf spot virus Tomato marchitez virus Tomato torrado virus Waikavirus – Anthriscus yellows virus Bellflower vein chlorosis virus Maize chlorotic dwarf virus Rice tungro spherical virus Unassigned – Strawberry latent ringspot virus

PVB PVU RpRSV SLSV TRSV TBRV ToRSV ALSV AVB CRLV CuLV StPV SDV SMoV BRNV CLVA DMaV CNDV DaYMV PYFV CaTV1 LNLCV MYMoV SCLSV ToMarV ToTV AYV BVCV MCDV RTSV SLRSV

5 4 35 4 115 86 138 4 14 10 6 4 33 83 26 4 4 2 2 2 10 4 4 35 25 148 0 2 4 114 32

– GCF_004117715.1 GCF_000854425.1 GCF_001923735.1 GCF_000856105.1 GCF_000853325.1 GCF_000860465.1 GCF_000861545.1 GCF_000907115.1 GCF_000859565.1 GCF_001503195.1 – GCF_000860985.1 GCF_000850445.1 GCF_000867325.1 GCF_000895075.1 GCF_001876835.1 GCF_002817415.1 – GCF_000862945.1 GCF_000927615.2 GCF_002219685.1 GCF_002220005.1 GCF_002219665.1 GCF_000879575.1 GCF_000872965.1 – GCF_001308735.1 GCF_000861445.1 GCF_000860625.1 GCF_000857165.1

KX656670.1. KX656671.1

DQ143874.1, DQ143875.1

DQ675189

(1995)

a

BBSV acronym has been confused with BBTMV. BBSV has highest identity (65%) with RNA2 of RCMV. Original acronym of BBTMV was TBBMV when ratified by the ICTV in 1972. c Accession numbers listed for chicory yellow mottle virus are satellite sequences. d Subgenera organization of the Sadwavirus genus has yet to be ratified by the ICTV. b

Virion Structure Virions are non-enveloped particles (ca. 30 nm diameter) with icosahedral symmetry (T ¼ 1, pseudo T ¼ 3, Fig. 3). Virions can either be RNA containing or empty. In the case of those viruses with segmented (bipartite) genomes virions can contain either RNA1 or RNA2, but not both. Thus, for successful completion of the virus infection cycle inoculum must consist of both RNAcontaining particles. For bipartite viruses, virions can be grouped into three components based on their buoyant densities; the top (T) component which is empty, and the middle (M) and bottom (B) components which contain RNA2 and RNA1, respectively. Irrespective of the number of CPs proteolytically cleaved the CP unit as a whole comprises three b-barrel domains each one containing two anti-parallel wedge-shaped b-sheets in a jelly roll fold conformation. Each sheet is made up of four b strands. The three b-barrel domains are either contained within one CP (genus Nepovirus) or cleaved to generate either three individual CPs each with one domain (genera Waıkavirus, Cheravirus, Sequivirus and Torradovirus), or unevenly to produce a large (L) and small (S) CP with two and one domain, respectively (genera Sadwavirus, Fabavirus and Comovirus). The virion is formed by the assembly of 60 CP units (whether comprised of one, two or three cleaved products), and as such is said to have pseudo T ¼ 3 symmetry as compared to viruses that use 180 identical CP copies to assemble virus particles with canonical T ¼ 3 symmetry. Pseudo T ¼ 3 symmetry is typical of all members of the Picornavirales and has been suggested in the case of secoviruses to be an evolutionary vestige that in animalinfecting counterparts permits improved evasion from the immune system when compared to true T ¼ 3 capsids. High resolution structures have been determined for a number of secoviruses, including, Cowpea mosaic virus (CPMV), Bean pod mosaic virus (BPMV), Red clover mottle virus (RCMV) (Comovirus); Arabis mosaic virus (ArMV), Blackcurrant reversion virus (BRV), GFLV and Tobacco ringspot virus (TRSV) (Nepovirus). The existence of empty (T) particles demonstrates RNA is not required for the maintenance of virus particles, but does not preclude its involvement in assembly. Nevertheless, there appears to be some specificity to the interactions of viral RNA in the assembled particles, electron densities indicating a consensus base composition of the interacting RNA. At the three-fold axis of symmetry of the BPMV virion, viral RNA folds into a trefoil shape implying a potential function for this structure in packaging and particle assembly. More defined interacting amino acids have been identified for the CPs; in CPMV M-virions the amino acid N174 in the large CP subunit is intimately associated with and shown to be essential for genomic RNA encapsidation. Attempts to develop an in vitro

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Fig. 3 Structure of Cowpea mosaic virus (CPMV) (a) Genome organization of CPMV. (b) Gradient centrifugation of wild-type CPMV permits separation into three components; empty CPMV particles sediment at the top (CPMV-T), CPMV containing RNA2 in the middle (CPMV-M) and RNA1 containing CPMV particles at the bottom (CPMV-B). (c) An asymmetric unit of CPMV empty virus-like particle (CPMV-T). The large CP subunit (L subunit, green) and the small coat protein subunit (S subunit, blue). The C terminal extension, only visualized in CPMV-T particles is colored pink. (d) Icosahedral organization of CPMV using an EM derived map of CPMV-T particles. Each of the 60 asymmetric units comprises one copy of the L subunit and the S subunit (colored as in 3C). A view down the two-fold axis is shown. (e) Electron micrographs of CPMV-M (top) and CPMV-T (bottom) particles. Courtesy of Hesketh, E.L., Meshcheriakova, Y., Thompson, R.F., Lomonossoff, G.P., Ranson, N.A., 2017. The structures of a naturally empty cowpea mosaic virus particle and its genome-containing counterpart by cryo-electron microscopy. Scientific Reports 7.

assembly assay, similar to other icosahedral viruses such as Cowpea chlorotic mottle virus, for CPMV which could help shed light on the mechanisms and requirements for assembly have so far proved unsuccessful due to the insolubility of the coat proteins.

Genome Organization Whether monopartite or bipartite, the genome of secoviruses shows an overall similar organization, with two distinct blocks for replication-associated and structural functional domains. For bipartite viruses in which these blocks are segregated the adopted standard nomenclature is RNA1 and RNA2, respectively. Typically, viral proteins are expressed as large polyproteins, one per RNA, that are eventually cleaved into smaller functional mature proteins by viral encoded proteases (Fig. 4). However, comoviruses have been demonstrated to produce two different sized polyproteins from the same RNA2 ORF derived from two different initiation codons. And unlike any other genus in the family, torradoviruses are predicted to encode a protein 50 and overlapping to the longer more conserved long polyprotein containing the predicted CPs. The lengths of the 50 and 30 untranslated region vary considerably from species to species. For some viruses (nepoviruses and sadwaviruses), there is high sequence identity between RNA1 and/or RNA2 in the 50 and/or 30 untranslated regions (UTRs). For example, the 30 UTRs of Strawberry mottle virus are ca. 1150 nt long and 96% identical. Protein domain organization, within the polyprotein(s) is similar to that of other members of the order Picornavirales. The structural block, which in monopartite genomes is located upstream of the replication block encodes the CP domain, upstream of which is the MP domain. The number of CP domain proteolytic derivatives that assemble to form the capsid varies from one to three depending on the genus (see Section “Virion Structure”). Upstream of the MP, nepoviruses encode a domain that is cleaved into one (2a protein in GFLV) or two (X3 and X4 proteins in ToRSV) proteins. The 2a protein fused to the green fluorescent protein of Aequorea victoria has been shown to associate with the replication complex assembled from RNA1-encoded proteins. The replication block contains domains with signature motifs typical of Helicases (NTP-binding domain proteins) (Hel), 3C-like protease (Pro) and RNA-dependent RNA polymerase (Pol). Between the Hel and Pro domains lies a small domain predicted to encode the VPg (viral protein genome-linked) (Fig. 4). Upstream of the Hel domain there exists for many secoviruses a divergent region encoding a predicted hydrophobic domain of unknown function. In CPMV the domain is encompassed by a single cleaved protein derivative that has been shown to affect protease activity, and thus was assigned the name of ‘protease cofactor’ (Co-Pro). In Tomato ringspot virus, there are two protein cleavage derivatives upstream of the Hel domain one (X2) sharing a conserved motif with the Co-Pro of CPMV. Genome sizes of secoviruses is variable; the smallest are the como- and fabaviruses (RNA1 B5.9 kb and RNA2 3.3–4 kb), while the related nepoviruses are some of the biggest (e.g., ToRSV, RNA1 B8.2 kb and RNA2 7.3 kb). In the latter, this size includes a long almost identical 30 UTR in both RNAs. In monopartite secoviruses the genome sizes range from 9.9 kb (sequiviruses) to 12.2 kb (RTSV). All genome segments are assumed to be 30 polyadenylated although experiments carried out on the sequivirus Parsnip yellow fleck virus (PYFV) found no evidence of a poly(A) tail. The only other fully sequenced sequivirus, Carrot necrotic dieback virus, is polyadenylated. Based on the genome organization of secoviruses and comparisons with the related animal-infecting genera (Picornaviridae, Discistroviridae and Iflaviridae) translation is plausibly mediated through a 50 internal ribosomal entry site (IRES), a highly structured RNA element that facilitates cap-independent translation. Experimental evidence is limited to date but the presence

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Fig. 4 Genome organization of representative members of the family Secoviridae. Boxed depiction at top is the generic secovirus organization. Only regions that notably deviate from this for each taxon are highlighted in the structures below. Each RNA is shown with the open reading frames (ORFs) represented by boxes (not to scale). Ovals depict VPg molecules covalently attached at the 50 end of the RNAs. Poly(A) tails are represented at the 30 end of the RNAs by AAA. Black vertical lines indicate evidence of cleavage sites. Movement protein – MP, coat protein(s) – CP, Helicase NTP-binding protein – Hel, protease – Pro, RNA-dependent RNA polymerase - Pol. Note – indicated subgenera organization of Sadwavirus genus not yet approved by the ICTV.

of a functional 50 IRES has been identified in the 50 UTR of Blackcurrant reversion virus, although sequences in the 30 UTR were also shown to be required for translation.

Properties and Functions of Gene Products Once inside the cell the viral RNA can function as a messenger RNA resulting in the translation of the viral polyprotein(s). IRES have been identified for a number of members of the Picornavirales. The only experimental evidence for a putative IRES in secoviruses comes from Blackcurrant reversion virus for which both a candidate stem-loop (SL) in the 50 leader sequence, a capindependent translation enhancer (CITE) at the 30 UTR and a potential kissing-loop interaction between them have been identified. Concordant with this translational model, mutational analyses of nucleotide stretches in the 50 UTR of BRV complementary to the 18S rRNA imply a potential recruitment of the 40S ribosomal subunit (Fig. 5) at the leader sequence. Similar conserved 50 UTR sequence stretches complementary to the 18S rRNA have been found for a number of other nepoviruses. After translation, cleavage by the 3C protease (encoded on RNA1 in bipartite viruses) of the polyprotein into mature functional proteins occurs. Although cleavage site specificity of the protease varies across genera, the mechanism involved is assumed to be the same; an amino acid in the substrate-binding pocket of the protease interacts directly with the amino acid directly upstream and flanking the cleavage site. Although structurally similar to serine proteases the catalytic amino acid of secovirus proteases is a cysteine (C) – except in the Pro of Blueberry latent spherical virus for which it is a serine (S). Specificity of 3C proteases depends on the substrate binding pocket which for the majority of secoviruses is a histidine (H), but can be a leucine (nepovirus subgroups A

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Fig. 5 Proposed model describing the possible mechanisms of translation for BRV RNAs. Step 1, the 40S ribosomal subunit (small oval) is recruited to the 50 untranslated region (UTR) internal ribosomal entry site (IRES), with base pairing between plant 18S rRNA and the corresponding complementary sequences of the BRV 50 leader (two small gray boxes). Step 2, after the start codon recognition, the 60S ribosomal subunit (large oval) joins to form the 80S initiation complex. Step 3, during translation elongation, the polyprotein (thin curly line) is synthesized. Step 4, after translation termination, the 80S complex is disassembled and the polypeptide is released. Step 5, long distance RNA–RNA base pairing between the 50 SL of the BRV 50 leader and the SL-1 of the 30 UTR holds two mRNA ends in close proximity, facilitating recycling of the ribosomal subunits again to the 50 UTR. An alternative mode of mRNA circularization could involve an interaction (double-headed arrow) of the poly(A)-binding protein (PABP; gray ovals) with a yet unidentified factor X, which in turn interacts (directly or through other factors) with the 50 UTR. Curved box – open reading frame (ORF); thick lines, UTRs; (A)n – poly(A) tail. From Karetnikov, A., Lehto, K., 2008. Translation mechanisms involving long-distance base pairing interactions between the 50 and 30 non-translated regions and internal ribosomal entry are conserved for both genomic RNAs of Blackcurrant reversion nepovirus. Virology 371, 292–308.

and B and PYFV) or valine (Stocky prune virus, TRSV). When the binding pocket contains an H the cleavage site is typically, though not always, a glutamine (Q). The efficiency of cleavage depends on the cleavage site composition itself, the context of the amino acids flanking the cleavage site and the overall protein conformation, thus allowing for the existence of proteolytically intermediate products with functions different to that of their individual components. For example, in nepoviruses the structural block polyprotein derived from RNA2 is less efficiently cleaved by the VPg-Pro intermediate than by the fully released and mature Pro implying a possible mechanism for sequential processing and maturation of the individual functional domains in line with the infection cycle. In addition to viral protein maturation the secovirus Pro may also interact with and cleave host translation factors as has been demonstrated for the Pro of picornaviruses. Once the main replication associated proteins (Pol, Hel and VPg) are functional, replication of the virus genome can occur within virus replication complexes (VRC) (see next section). The Hel domain, because of homology to known RNA helicases is assumed to be involved in binding and unwinding double stranded RNA structures that may form during replication. By analogy with picornaviruses the VPg is essential for viral replication acting as a primer for RNA synthesis but it may be removed from virion RNA by a host component (similar to the poliovirus unlinking enzyme). VPg is also absent on viral mRNA undergoing translation and has no effect on the infectivity of naked RNA. In nepoviruses full maturation of the Pro would shift proteolysis of the viral polyprotein and its intermediates to releasing the structural functional domains (CP and MP) of RNA2 both of which are required for virus movement. In the unique case of the newly proposed subgenus Stramovirus full release of the CP would also require proteolytic removal of the downstream glutamic protease. There is plenty of evidence to suggest secoviruses move cell-to-cell in the form of virions (rather than ribonucleoproteins), como- and nepoviral MPs having been shown to be a component of tubular structures that act as a conduit for virions passing through the cell wall. Besides having an ultimate role in RNA encapsidation, some CP(s) also function as suppressors of gene silencing. Although not as potent as other classic plant virus silencing repressors (VSR) in that it does not prevent targeted mRNA degradation by the plant silencing machinery the CP of ToRSV has been shown to interact with AGO1 and relieve a translation repression mechanism. VSR activity has also been demonstrated for the small CP (CP-S) of CPMV and the smallest CP (Vp20) of Apple latent spherical virus (ALSV).

Replication and Propagation Purified total viral RNA is highly infectious but for viruses with a bipartite genome, neither RNA species by itself can infect plants systemically. RNA1 encodes for the replication-associated protein block and this is able to replicate in individual cells, but can neither move cell-to-cell or assemble into virus particles without the complementary functions of those proteins present on RNA2.

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Replication occurs in VRCs, intracellular complexes formed by association with intracellular membranes. For como- and nepoviruses the Hel domain and upstream encoded proteins have been shown to interact with the endoplasmic reticulum (ER). Replication is reduced upon treatment with an inhibitor of type II fatty acid synthase, an observation that suggests VRC proliferation is not a simple reorganization of existing intracellular membranes. The Hel protein and the protein directly upstream of the Hel (Co-Pro in CPMV and X2 in ToRSV) are implicated in the accumulation of VRCs. In CPMV, VRCs in the form of irregularly shaped vesicles develop first in the cytoplasm and then coalesce around the nucleus. Electron tomography of picornavirus VRCs shows structures that during the exponential phase of infection are predominantly closed smooth single-membrane tubules that as infection progresses develop into doublemembrane multilamellar structures. The ToRSV X2, Hel and Hel–VPg fusion protein and the CPMV Co-Pro and Hel–VPg fusion protein associate with the ER while other RNA1 derived mature proteins (VPg, Pro, and Pol) are soluble when expressed ectopically and are only peripherally associated with the VRCs in infected cells. This localization can be explained by the predicted hydrophobicity of the X2 and Co-Pro and equivalent domains in other secoviruses. Both molecules have predicted transmembrane helices and an amphipathic helix suggesting they may act as membrane anchors in the VRC. How viral RNA is recruited to the VRC in the first place is still unclear but replication and translation of RNA1 are tightly linked. Consistent with other virus replication-associated proteins, BPMV infection RNA1 derived proteins are exclusively cis-acting supporting the notion of a tight spatial molecular relationship. The RNA2 derived N-terminal proteins of BPMV (p58) and GFLV (2A) also function in cis and potentially serve to recruit RNA2 to the VRC.

Transmission, Host Range In nature the majority of secoviruses are transmitted in a specific manner by insects or nematodes. Some (e.g., nepoviruses, cheraviruses, Satsuma dwarf virus (SDV), SLRSV) can be transmitted by seed and/or pollen. For members of the subgroup Comovirinae there is some predictability between the virus species within a genus and the vector (e.g., nepoviruses-nematodes, comoviruses-beetles, fabaviruses-aphids), while for other genera vector-virus relationships are more variable. For example, the torradoviruses; Tomato torrado virus (ToTV) and Tomato marchitez virus (ToMarV) are transmitted by whitefly species in a semipersistent manner. However, detectable spread (horizontal transmission) of Carrot torradovirus 1 (CaTV1) and Lettuce necrotic leaf curl virus (LNLCV) in regions where whitefly numbers are limited (United Kingdom and Netherlands, respectively) has led researchers to identify aphid species as the most likely vectors. The type species of the Cheravirus genus Cherry rasp leaf virus (CRLV), is transmitted by the nematode Xiphinema americanum in the field and also through seeds, whereas ALSV has no known vector; transmission through seed and via pollen from infected apple trees occurring at a rate of 4.4% and 0.4%, respectively. Such is the relationship sometimes between virus and their vector that their names have become intertwined, as in the case of nepoviruses and nematodes – nematode‐transmitted viruses with polyhedral particles. This relationship can be very specific, exemplified by GFLV, for which a single amino acid change (Gly297Asp) on a surface loop of the CP is sufficient to significantly reduce efficiency of transmission by its only known vector Xiphinema index. This mutation does not seem to alter virion structure but has been proposed to inhibit the physical interaction between virus and the nematode vector receptor(s) necessary for successful transmission. However, broader nepovirus-vector relationships do exist: TRSV and ToRSV are known to be vectored by least five nematode species; and BRV, consistent with its serological uniqueness, is transmitted by the eriophyid gall mite Cecidophyopsis ribis. In some rare cases, secoviruses assist in the transmission of other viruses. Rice tungro, a disease that has periodically hit rice production in South East Asia is found associated with two distinct viruses; RTSV and Rice tungro bacilliform virus (RTBV) from the family Caulimoviridae. RTSV is transmitted by the leafhopper Nephotettix virescens but causes mild to no symptoms. RTBV is the main causal agent of the disease but depends on RTSV for transmission. Similarly, another waikavirus, Anthriscus yellows acts a helper virus in the aphid transmission of PYFV. Secoviruses can either have broad or narrow host ranges, infecting predominantly dicotyledonous plants. Most nepoviruses and fabaviruses have wide host ranges, while comovirus infection is limited to a few species in the family Leguminosae. The whitefly transmissible ToTV, initially considered limited in host range to tomato, has been detected in over twenty weed species from eight families. SDV and related viruses have no known vector and natural infection appears restricted to species of citrus via grafting and mechanical transmission. Nevertheless, experimentally SDV has been shown to have a potentially broad host range infecting cowpea, cucumber, sesame and tobacco in a strain specific manner. A small number of secoviruses (some nepoviruses and Strawberry latent ringspot virus) are found associated with a satellite RNA (satRNA); RNA that depends on the helper virus for replication and encapsidation, and can in some cases modulate symptoms. TRSV satellite RNA was the first satRNA ever described, and consists like other satRNAs in its class of a small (200–500 nt) RNA molecule lacking a functional open reading frame (ORF), an attached VPg and a poly(A)tail. In contrast, the other class of secoviral satRNAs are larger (1.1–1.5 kb), polyadenylated and associated with a 50 terminus VPg, and encode a protein essential for replication. Sequence similarities between different satRNAs is very limited although the predicted ORF of the large satRNAs show some structural similarities indicative of a shared function.

Epidemiology and Control Dispersal of viruses occurs either through the infected host or the vector. In the past, plant virus spread would be local and limited to where virus and vector coincided geographically. In the wake of human globalization, virus movement across continents has

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accelerated; for example, human beings are likely responsible for the present global distribution of GFLV and its nematode vector X. Index. And despite each country’s legal measures to minimize the introduction of new plant materials the implementation of up-to-date certification schemes is essential in the monitoring and mitigation of disease outbreaks. Nematodes are slow moving, as will be the spread of any virus they vector in the field. Nevertheless, in the case of woody plants like grapevine that are cropped for multiple years, if phytosanitation practices are inadequate, virus-infected budwood can be inadvertently introduced at the very beginning of the vineyards lifetime. Unlike nematodes; whiteflies and aphids are highly mobile and can move long distances rapidly extending the range and spread of the viruses they vector. Surveys made in the 2000s in Spain showed that ToTV was often found in mixed infections in tomato with another emerging non-whitefly-vectored potexvirus, Pepino mosaic virus (PepMV). The reason for the co-existence of both viruses has been hypothesized to be in part due to the conspicuous presence of whitefly populations for the former and a lack of genetic resistance against the latter. No increase in severity of torrado symptoms was observed in mixed infections. In 1970s Japan a disease in rice termed waika caused by infection of RTSV emerged to induce stunting and a reduction in grain quality. Generally RTSV alone causes mild symptoms but its ability to complement the transmission by leafhoppers of RTBV allowed for the emergence of tungro disease in the late 60s to mid-70s. Both viruses are transmitted in a semi-persistent manner by at least six leafhopper species that can remain viruliferous for up to a week. Inoculated plants can become sources of inoculum within one week. These conditions meant rapid spread of tungro in susceptible varieties. Attempts to control the disease has mainly alternated between the application of pesticides to reduce vector populations and the introduction of resistant varieties. Resistance principally took the form of resistance to the vector which over time broke down forcing growers to return to chemical control. In more recent years transgenic approaches that exploit RNA interference (RNAi) against both viruses have been explored along with RTSV resistance generated by CRISPR/Cas9 targeted mutagenesis of the eukaryotic translation initiation factor 4G (eIF4G). RNAi technologies have also been applied to other nepoviruses (GFLV, ArMV, TRSV and ToRSV) mainly in herbaceous hosts, although a number of grapevine rootstocks transgenic for the CP have been developed. Constitutive expression of specific camelid‐ derived heavy‐chain‐only antibodies, or nanobodies, in Nicotiana benthamiana conferred strong resistance to GFLV, neutralizing the virus before cell-to-cell movement. What control methods are employed and deployed against any plant pathogen will eventually depend on public perception of the associated risks and benefits.

Biotechnological Applications CPMV has been at the vanguard nanobiotechnological developments and applications in part because of its high yield, simple purification and virion stability. Like other “nanotech” plant virus systems CPMV has the advantages of scalability, low-cost, lowtoxicity and biodegradability. These factors, in combination with an early knowledge of its particle structure, resulted in it becoming the first plant virus to be successfully engineered to present foreign peptides on its surface with a 25 aa insertion of part of the VP1 domain of Foot and mouth disease virus (FMDV) into the bB-bC loop of the CP-S. Expression of other epitopes inserted in the bB-bC and other surface loops have also been effective, provided the insert is no greater than 40 aa with an isoelectric point (pI) of less than 9. Employing these virus-like particles (VLPs) to elicit an immunogenic response to the inserted epitope has been shown for Human rhinovirus 14, Mink enteritis virus, Human immunodeficiency virus, the bacteria Pseudomonas aeruginosa and Staphylococcus aureus, and for a number of human cancers (e.g., lung, melanoma, ovarian). The main motivation for these studies is the development of vaccines, and their stable expression in plants offers the appealing possibility of their delivery as oral vaccines provided conditions can be optimized so that CPMV VLPs tolerate the extreme physio-chemical conditions of the gastrointestinal tract long enough to be absorbed into the bloodstream. Uptake of CPMV particles into mammalian endothelial cells has been shown to be mediated by binding of the virus to the cell surface protein vimentin, a known receptor for some mammalian picornaviruses. This unexpected property could be put to use in the development of CPMV as a delivery system for medical therapeutics and imaging - RNA-containing virions of CPMV being readily infusible with a range of functional molecules. Aside from nanotechnological applications CPMV containing vectors exploiting the hyper-translatability of a modified RNA2 50 UTR sequence have been engineered for the rapid expression of large amounts (1 g per kilogram of tissue) of pharmaceutical proteins in plants via agroinfiltration. Other secoviruses too have been cloned for biotechnological applications. In particular ALSV vectors have been used for functional genomic studies in a number of plants by means of virus-induced gene silencing (VIGS). One of the benefits of the ALSV system over other VIGS vectors is the virus’ broad experimental host range which includes nightshades, cucurbits and legumes. Additionally its asymptomatic infection of most plants also means that the influence of virus infection on phenotype, as encountered with other VIGS vectors, is minimized. The ALSV system has been effectively used to “vaccinate” plants against infection from agronomically important viruses like Cucumber mosaic virus and potyviruses.

Diagnosis One of the classical characteristics of nepovirus infection is symptom recovery, where the youngest emerging leaves of an infected plant are symptomless. Although not typical (or unique) for all nepoviruses, the cause for this recovery phenomenon has its origins in the gene silencing mechanism of the plant.

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Minimizing the introduction of secoviruses into new planting material is best achieved by effective diagnostic screens and certification of seeds and propagative materials. Since the advent of molecular and serological methods diagnosis is commonly achieved by enzyme-linked immunological assays (ELISA) and/or the polymerase chain reaction. Despite its robustness, relative low cost and high throughput, ELISA or any serological method is sometimes impractical or intractable due to the technical challenges of virion purification or alternative methods of heterologous protein expression in the development of antibodies. These can be specific to each virus. Commercial ELISA kits available for the following secovirus species – Andean potato mottle virus (APMV), ArMV, Arracacha virus B (ABV), Broad bean true mosaic virus (BBTMV), BBWV1, BBWV2, Carrot necrotic dieback virus (CNDV), CPMV, Cowpea severe mosaic virus (CPSMV), GFLV, Potato black ringspot virus (PBRSV), Radish mosaic virus (RaMV), Raspberry ringspot virus (RpRSV), Squash mosaic virus (SqMV), SLRSV, Tomato black ring virus (TBRV), TRSV and ToRSV. A subset of these viruses can also be detected using commercial lateral flow devices. The utility of an interesting new technology that employs small proteins, called affimers, that mimic antibodies, has been shown to detect CPMV in crude extracts. Affimers are synthetically produced on a cystatin consensus scaffold and identified by phage display screening. Their alleged benefits over antibodies are their low cost and thermostability. Multiplex detection methods where multiple viruses are detected in a single test are normally crop specific. Multiplex RT-PCR assays have been developed for citrus that include Satsuma dwarf viruses, and for grapevine that include ArMV and GFLV. Microand macro-arrays too have been employed for the multiplex detection of multiple grapevine infecting secoviruses. With the advent of high throughput sequencing methods and their continuing drop in cost and turnover time the ability to detect multiple pathogens in multiple samples simultaneously is becoming routine and is rapidly superseding previous methodologies. Nevertheless there is a burgeoning need for “quick and dirty” tests that can be carried out by anyone anywhere. Lateral flow assays have addressed some of this need but more sensitive isothermal nucleic acid based methods like loop-mediated amplification (LAMP) and recombinase polymerase amplification (RPA) are rapidly becoming the methods of choice for point-of-use diagnostics.

Concluding Remarks Research into secoviruses has so far provided a fascinating insight into a diverse group of viruses that share many features with their animal- and algal-infecting relatives. As human agricultural activities continue expanding and globalizing, the threat of novel disease outbreaks from existing and emergent secoviruses will remain. A clearer understanding of the biology and evolution of this group of viruses will undoubtedly better prepare us for combating their negative impact, and enlighten us as to how they and their picornavirus relatives adapt and exploit new environments and hosts. Which methods are used to control secovirus outbreaks will ultimately depend on the public’s level of acceptance and perception of the associated risks and benefits. It is certainly plausible that the promise CPMV holds as a biotechnological and biomedical tool will help the secovirus name reach a broader more informed audience.

Further Reading Azzam, O., Chancellor, T.C.B., 2002. The biology, epidemiology, and management of rice tungro disease in Asia. Plant Disease 86, 88–100. Fuchs, M., Schmitt-Keichinger, C., Sanfaçon, H., 2017. A Renaissance in nepovirus research provides new insights into their molecular interface with hosts and vectors. Advances in Virus Research 97, 61–105. Lin, T.W., Johnson, J.E., 2003. Structures of picorna-like plant viruses: Implications and applications. Advances in Virus Research 62, 167–239. Sainsbury, F., Canizares, M.C., Lomonossoff, G.P., 2010. Cowpea mosaic virus: The plant virus-based biotechnology workhorse. Annual Review of Phytopathology 48, 437–455. Sanfaçon, H., 2015. Secoviridae: A family of plant picorna-like viruses with monopartite or bipartite genomes. In: eLS. Chichester: John Wiley & Sons, Ltd. doi:10.1002/ 9780470015902.a0000764.pub3. Sanfacon, H., Wellink, J., Le Gall, O., et al., 2009. Secoviridae: A proposed family of plant viruses within the order Picornavirales that combines the families Sequiviridae and Comoviridae, the unassigned genera Cheravirus and Sadwavirus, and the proposed genus Torradovirus. Archives of Virology 154, 899–907. Thompson, J.R., Dasgupta, I., Fuchs, M., et al., 2017. ICTV virus taxonomy profile: Secoviridae. Journal of General Virology 98, 529–531. Thompson, J.R., Kamath, N., Perry, K.L., 2014. An evolutionary analysis of the Secoviridae family of viruses. PLoS One 9 (9), e106305. doi:10.1371/journal.pone.0106305. van der Vlugt, R.A.A., Verbeek, M., Dullemans, A.M., et al., 2015. Torradoviruses. Annual Review of Phytopathology 53, 485–512.

Relevant Websites https://talk.ictvonline.org/ International Committee on Taxonomy of Viruses.

Sequiviruses and Waikaviruses (Secoviridae) Lucy Rae Stewart, Agricultural Research Service, US Department of Agriculture, Wooster, OH, United States Published by Elsevier Inc. This is an update of I.-R. Choi, Sequiviruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00462-3.

Nomenclature

nt Nucleotide(s) ORF Open reading frame RdRp RNA-dependent RNA polymerase UTR Untranslated region VPg Viral protein genome-linked

aa Amino acid(s) CP Coat protein or capsid protein kb Kilobase kDa Kilo dalton nm Nanometer

Glossary 3C-like protease Viral enzymes for polyprotein cleavage with similarity to the picornavirus 3C proteases, with similarity to chymotrypsin. Helper component A viral-encoded protein not part of the capsid that mediates the transmission of viruses by vectors. Monopartite Adjective describing a virus with a single particle containing a single complete genomic nucleic acid. Polyprotein Large precursor polypeptide that is post- or co-translationally cleaved into mature protein components,

often with dynamics reflecting changing functions over time. Semipersistent transmission A mode of vector-mediated transmission of plant viruses in which viruses are usually acquired within few minutes, and retained up to several days by vectors. Viruses do not multiply or circulate in vectors, and require no latent period between acquisition and transmission. Synergy Co-infection of two or more viruses where disease is greater than additive and/or titer of one or more virus is enhanced.

Introduction Viruses in the genera Sequivirus and Waikavirus are monopartite members of the family Secoviridae, comprising the former family Sequiviridae. Viruses belonging to these groups are often referred to as ‘picorna-like’ due to their similarities to picornaviruses in virion morphology and genome structure, and polyprotein expression strategy. They have monopartite single-stranded RNA (ssRNA) genomes encapsidated in isometric particles composed of three capsid proteins. They infect plants and are usually transmitted by insect vectors in nature. The genus Waikavirus includes pathogens of important cereal crops: Rice tungro spherical virus (RTSV), and Maize chlorotic dwarf virus (MCDV). Rice tungro disease is caused by a synergistic co-infection of RTSV with an unrelated caulimovirus, Rice tungro bacilliform virus (RTBV), and impacts rice production in South and South-East Asia. Maize chlorotic dwarf disease has been problematic in the MidWest and Southeastern United States. Members of the genus Waikavirus with described symptomology are phloem-limited and cause stunting, yellowing, and sometimes veinal chlorosis (Fig. 1). The genus Sequivirus shares similar genome organization but is sequence-divergent from the waikaviruses and fewer members are described, with the type species being Parsnip yellow fleck virus (PYFV). Members of the genus Sequivirus cause vein yellowing, yellow flecks, mosaic, and necrosis in infected hosts, can be mechanically transmitted experimentally, and are not phloem-limited. A unique biological feature of viruses in these genera is the ability of some members to act as (waikaviruses) or require (sequiviruses) a helper virus for transmission by insect vectors. Viruses belonging to the genus Sequivirus infect mesophyll and epidermal cells and are able to be transmitted by mechanical inoculation and insect vectors, while those described in the genus Waikavirus are limited in phloem tissue and transmitted only by insect vectors or direct mechanical inoculation to phloem, as by vascular puncture inoculation in maize. Sequiviruses are dependent on a helper virus for their transmission by insect vectors. Waikaviruses are independently transmitted by insect vectors and presumably encode a helper component in their genomes. The viruses in each genus are primarily classified on the basis of their biological and physical characteristics, but conspicuous differences are also found in their genome features. The genomes of MCDV and RTSV are approximately 12 kb and polyadenylated at the 30 end, whereas that of PYFV is about 10 kb and lacks 30 polyadenylation.

Classification: Taxonomy, Phylogeny, and Evolution The family Secoviridae is a combination of former families Comoviridae and Sequiviridae. The family is composed of eight genera: Sequivirus, Waikavirus, Torradovirus, Sadwavirus, Cheravirus, Nepovirus, Fabavirus, and Comovirus, as well as some unassigned members

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Fig. 1 Waikavirus symptoms and particles. (A) RTSV virions, ca. 30 nm diameter. Bar ¼ 100 nm. (Photograph courtesy of Dr. Filomena Sta Cruz, University of the Philippines-Los Banos). (B) Rice plants with rice tungro disease synergistic disease from coinfection with RTSV and RTBV (left) compared to single infections with RTBV (center left), RTSV (center right), and uninfected plant (right). (Photograph courtesy of Dr. Filomena Sta Cruz, University of the Philippines-Los Banos). (C) MCDV-S vein chlorosis in corn. (D) The black-faced leafhopper, Graminella nigrifrons, vector of MCDV. (Photograph courtesy of Jane Todd, USDA-ARS). (E) MCDV-S-infected stunted plants in a pot with a single uninfected plant. Reproduced, by permission, from Stewart, L.R., 2011. Waikaviruses: Studied but not understood. APS Features. doi:10.1094/APSFeature-2011-11. https://www.apsnet.org/edcenter/apsnetfeatures/Pages/waikavirus.aspx.

(Fig. 2). Most of these are bipartite viruses, but the genera Waikavirus and Sequivirus are monopartite. The 50 end of the monopartite viruses corresponds to RNA2 and the 30 end to RNA1 of bipartite family members (Fig. 3). Currently there are three species described in the genus Sequivirus: Parsnip yellow fleck virus (the type species); Dandelion yellow mosaic virus; and Carrot necrotic dieback virus, formerly named Parsnip yellow fleck virus, Anthriscus strain (Table 1). In the genus Waikavirus, four species are described: Rice tungro spherical virus (type species); Maize chlorotic dwarf virus (MCDV); Anthriscus yellows virus (AYV); and Bellflower vein chlorosis virus (BVCV; Table 1). Other putative waikaviruses have been found in deep sequencing studies, including Blackcurrant waikavirus A, Red clover associated virus 1, and Brassica napus RNA virus 1. Sequiviruses and waikaviruses, like other members of the family Secoviridae, share properties of plant picorna-like viruses, and are classified in the order Picornavirales. Comparison of amino acid sequences revealed that the sequence identity of the MCDV (isolate TN) polyprotein to those of RTSV (strain A) and PYFV (isolate P-121) were 51% and 35%, respectively. The NTP-binding domain in the polyprotein of MCDV (TN isolate) showed significant sequence identity to those of viruses such as RTSV (strain A, 79%), PYFV (isolate P-121, 55%), CPMV (46%), Hepatitis A virus (HAV, 47%), and Poliovirus (45%). The RdRp also shows significant similarities among members of the family Secoviridae. For instance, the sequence similarities in the RdRp region of MCDV (TN isolate) to those of related viruses are 75% for RTSV, 50% for PYFV, 48% for CPMV, 44% for TBRV, and 40% for HAV. Phylogenetic analysis based on the sequences of NTP-binding domains indicates that PYFV and RTSV are not noticeably similar to each other compared to their relatedness to picornaviruses and comoviruses. However, phylogenetic analysis based on the regions of RdRp suggests that PYFV and RTSV are more closely related to each other than to picornaviruses and comoviruses. The 26 kDa CP of PYFV and the 22 kDa CP of RTSV contain amino acid sequences resembling those in VP3 of Encephalomyocarditis virus and Human rhinovirus 14. Like other members of the Secoviridae, members of the genera Waikavirus and Sequivirus encode CPs with three jelly-roll domain in each of the 20 faces of the icosahedral virus particle. For the waika- and sequiviruses, each jellyroll domain is on a separate CP, in contrast to family members with one jellyroll domain on a small CP and two on a large CP.

Variation of Isolates and Strains Complete sequences of sequiviruses PYFV and CNDV but not DaYMV are available. Although CNDV was formerly considered the Anthriscus strain of PYFV, strain variation in the currently classified species is not described for members of the genus Sequivirus. Complete sequences of the waikaviruses RTSV, MCDV, and BVCV, and putative waikaviruses Red clover associated virus 1 and Brassica napus RNA virus 1 are available, as are partial sequences of the putative member Blackcurrant virus A, but no sequences

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Fig. 2 Maximum likelihood inferred phylogenetic unrooted tree of members of the family Secoviridae based on an alignment of amino acid sequences of the conserved domains between the “CG” motif of the 3C-proteinase and the “GDD” motif of the polymerase (Pro-Pol region). The bar represents the genetic distance. Rounded rectangles (gray) – subfamily; (colored) – genera. For each species, the sequence of the exemplar isolate was used for the alignments (see species tables for the sequence accession numbers). Percent bootstrap values (1000 replicates) are shown at each branch node. Branches with bootstrap values below 70% were collapsed. Virus names, abbreviations, and GenBank accession numbers of sequences used to create the tree are: Strawberry latent ringspot virus (SLRSV), GCF_000857165; Stocky prune virus (StPV), DQ143874 and DQ143875; Arracacha virus B (AVB), GCF_000907115; Currant latent virus (CuLV), GCF_001503195; Cherry rasp leaf virus (CRLV), GCF_000859565; Apple latent spherical virus (ALSV), GCF_000861545; Parsnip yellow fleck virus (PYFV), GCF_000862945; Carrot necrotic dieback virus (CNDV), GCF_002817415; Maize chlorotic dwarf virus (MCDV), GCF_000861445; Rice tungro spherical virus (RTSV), GCF_000860625; Bellflower vein chlorosis virus (BVCV), GCF_001308735; Squash chlorotic leaf spot virus (SCLSV), GCF_002219665; Tomato torrado virus (ToTV), GCF_000872965; Tomato marchitez virus (ToMarV), GCF_000879575; Carrot torradovirus 1 (CaTV1), GCF_000927615; Motherwort yellow mottle virus (MYMoV), GCF_002220005; Lettuce necrotic leaf curl virus (LNLCV), GCF_002219685; Strawberry mottle virus (SMoV), GCF_000850445; Black raspberry necrosis virus (BRNV), GCF_000867325; Satsuma dwarf virus (SDV), GCF_000860985; Chocolate lily virus A (CLVA), GCF_000895075; Dioscorea mosaic associated virus (DMaV), GCF_001876835; Grapevine fabavirus (GFabV), GCF_003033815; Prunus virus F (PrVF), GCF_003033845; Cucurbit mild mosaic virus (CuMMV), GCF_002867105; Broad bean wilt virus 1 (BBWV1), GCF_000853465; Broad bean wilt virus 2 (BBWV2), GCF_000861605; Gentian mosaic virus (GeMV), GCF_000880435; Cowpea severe mosaic virus (CPSMV), GCF_000861225; Squash mosaic virus (SqMV), GCF_000861625; Radish mosaic virus (RaMV), GCF_000879335; Bean pod mottle virus (BPVM), GCF_000862925; Red clover mottle virus (RCMV), GCF_000860425; Cowpea mosaic virus (CPMV), GCF_000860385; Raspberry ringspot virus (RpRSV), GCF_000854425; Tomato ringspot virus (ToRSV), GCF_000860465; Cherry leaf roll virus (CLRV), GCF_000893515; Soybean latent spherical virus (SLSV), GCF_001923735; Peach rosette mosaic virus (PRMV), GCF_002029615; Blueberry latent spherical virus (BLSV), GCF_002867165; Grapevine fanleaf virus (GFLV), GCF_000860305; Arabis mosaic virus (ArMV), GCF_000855205; Cycas necrotic stunt virus (CNSV), GCF_000860925; Potato virus B (PVB), KX656670.1 and KX656671; Grapevine chrome mosaic virus (GCMV), GCF_000862025; Tomato black ring virus (TBRV), GCF_000853325; Beet ringspot virus (BRSV), GCF_000849565; Artichoke yellow ringspot virus (AYRSV), GCF_002986065; Grapevine Bulgarian latent virus (GBLV), GCF_000892035; Blackcurrant reversion virus (BRV), GCF_000860405; Tobacco ringspot virus (TRSV), GCF_000856105; Aeonium ringspot virus (AeRSV), GCF_002867145; Melon mild mottle virus (MMMoV), GCF_002867185; Mulberry mosaic leaf roll associated virus (MMLRaV), GCF_002867205. Created by Thompson, J.R. for Encyclopedia of Virology and reproduced with permission.

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Fig. 3 Comparative genome organizations of PYFV, RTSV-A, MCDV-S, and BPMV. ORFs are indicated with rectangles. Horizontal lines at both ends of the ORFs represent the 50 - and 30 -UTRs of the genome. Solid vertical lines dividing the ORF correspond to the positions of verified proteolytic cleavage sites, while the positions corresponding to predicted cleavage sites are indicated with dotted lines. Nucleotide coordinates of translation start and stop codons for the polyproteins and the length of the entire genomes are indicated with numbers above the genomes. Numbers below the regions encoding CP1–CP3 are the predicted molecular weight (kDa) for the respective CP. Shadowed areas represent the approximate region encoding NTP-binding domain. The regions in the ORF are shown with Hel for putative NTPase/helicase, Pro for 3C-like protease, and RdRp for RNA-dependent RNA polymerase. A(n) indicates polyadenylation.

Table 1

Virus members in the genera Sequivirus and Waikavirus in the family Secoviridae. Type species are written in bold fonts

Genus/virus Sequivirus Parsnip yellow fleck virus Carrot necrotic dieback virus Dandelion yellow mosaic virus Waikavirus Rice tungro spherical virus Anthriscus yellows virus Bellflower vein chlorosis virus Maize chlorotic dwarf virus

AcronymAccession#

Genome size (kb)

Major natural host

Transmission vector

PYFV

GCF_000862945.19.9

Europe

CNDV

GCF_002817415.19.9

Europe

Parsnip, hogweed, Aphids (Cavariella aegopodii, C. pastinacae) cow parsley Carrot, cow parsley Aphids (Cavariella aegopodii)

DaYMV GCF_002817435.110.0a

Europe

Lettuce, dandelion

Aphids (Acyrthosiphon solani, Myzus ornatus, M. ascalonicus, M. persicae)

RTSV

GCF_000860625.112.2

Asia

Rice, Oryza spp.

AYV

–

Eurasia, UK

Cow parsley

Green leafhoppers (Nephotettix virescens and four other species) Aphids (Cavariella aegopodii)

BVCV

GCF_001308735.111.6

Korea

Bellflower

Unknown

MCDV

GCF_000861445.111.8

USA

Maize, Johnsongrass Deltocephaline leafhopper (Graminella nigrifrons)

USA Korea

Blackcurrant Rapeseed

10.6a

Tentative members Blackcurrant virus ABcVA – 11.0a Brassica napus BnRV1 GCF_00413465.1 12.3 RNA virus 1 Red clover RCaV1 MH325329 11.9 associated virus 1 a

Geographical distribution

Estimated size (nucleotide sequence not reported).

Czech Republic Red clover

Unknown Unknown Unknown

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are available for AYV. Multiple isolates of RTSV and MCDV have been completely sequenced and strains are described based on sequence and biological differences. Complete sequences are available for RTSV isolates RTSV-A (Philippines), Seberang Perai (Malaysia), and Vt6, a resistance-breaking Philippine isolate, and Andrha Pradesh, West Bengal, and Orissa collected in India. Nucleotide sequences of these isolates share 90%–97% identity, with Indian isolates clustering separately from isolates from the Philippines and Malaysia. Most diversity analyses for RTSV focus on the CP. Examination of nucleotide sequences of the RTSV CP revealed broad genotypic variation among and within geographic isolates, although the relationship of genotypic variation with the pathogenicity is not understood yet. CPs of isolates of RTSV from the Philippines and Malaysia show about 95% sequence similarity, while those from Bangladesh and India differ from the Philippine isolate by about 15%. The CP3 of the Indian isolate was also distinguishable from that of the Philippine isolate in electrophoretic mobility and the response to cellulolytic enzyme. Genotypic surveys on the CP sequences of RTSV field isolates collected from various sites in the Philippines and Indonesia indicated that a high degree of genetic diversity exists among the field isolates, and that infections with mixed genotypes in single sites are not uncommon. Phylogenetic analysis based on the CP sequences of the RTSV field isolate suggested that the clustering of genotypes found in Philippine sites was significantly different from that found in Indonesia sites, indicating some geographic isolation of RTSV populations. Strain Vt6 of RTSV was found to possess infectivity to some rice cultivars which the type strain A may not be able to infect. Amino acid sequence of strain Vt6 was approximately 96% identical to that of strain A, with greater dissimilarity in the leader protein region and the putative small ORF found near the 30 end. Four isolates of MCDV have been reported. The S isolate of MCDV shares 99% nucleotide and polyprotein sequence identity to the type (T) isolate and both have bright veinal chorosis and strong symptoms, particularly the S isolate. The sequences and origin of isolates described as S and T in earliest pre-sequencing literature are somewhat unclear, but S was probably a lab-selected subset of T collected in Ohio, and both are considered the same strain. The mild (M1) isolate exhibits very mild or no obvious symptoms by itself including no measurable stunting, but leaf texture changes and infected plant weight is reduced. M1 may develop transient severe symptoms in co-infection with S. M1 has only 61% polyprotein amino acid identity with isolate T. Antisera raised against isolate T react strongly with isolate S, but not with isolate M1. Isolate Tennessee (TN) of MCDV is also significantly divergent from isolate T, showing only 60% of amino acid sequence identity, but comparative TN symptom severity is unreported. In a survey of Ohio, where MCDV-M1 and MCDV-T isolates were first collected and identified, M1 and T sequences but no intermediate, recombinant, or divergent sequences were found. A more divergent partial MCDV sequence was identified in an Oklahoma tallgrass survey for viruses, but other MCDV isolates have yet to be identified. The low levels of amino acid sequence identity among the isolates of MCDV has raised the suggestion that they may represent distinct virus species. Within the family, species are demarcated by CP aa sequences o75% identical, Pro-Pol aa sequences o80% identical, antigenic differences, host range differences, vector specificity differences, and absence of cross-protection. MCDV-M1 CP polyprotein amino acid sequence is 85% identical to that of MCDV-TN, but only 65% identical to that of MCDV-S or MCDV-T. MCDV Pro-Pol amino acid sequences (based on RTSV-A protease start annotation) from MCDV-M1 are 80% identical to MCDV-TN, but each is only 67%–68% identical to MCDV-S or MCDV-T. Thus, MCDV-TN and MCDV-M1 meet sequence criteria for classification as a distinct species from MCDV-S or MCDV-T, but biological differences other than symptom severity have not been described.

Properties of Virions Sequiviruses and waikaviruses have non-enveloped isometric particles approximately 30 nm in diameter (Fig. 1). The sedimentation coefficients of the virions are 153–159S for sequiviruses and 175–183S for waikaviruses. The buoyant density of PYFV virions in CsCl is 1.49 g ml1, and that of waikaviruses is 1.51–1.55 g ml1. Genome sequences and immunodetection of the viruses indicate that the virions consist of three capsid proteins (CPs). The sizes of CPs range from 22 to 35 kDa, depending on virus species and isolates. The virion of RTSV (Philippine-type strain A) consists of three proteins of 22.5 (CP1), 22 (CP2), and 33 (CP3) kDa. The predicted molecular masses for the CP of MCDV TN are 22, 23, and 31 kDa, and those of PYFV (P-121 isolate) are 22.5, 26, and 31 kDa.

Genome Organization The genomes of sequiviruses and waikaviruses are positive-sense, monopartite ssRNA. The genome sizes range from approximately 10 kb for sequiviruses to 12 kb for waikaviruses. The genomes contain a large open reading frame (ORF) encoding a polyprotein presumably proteolytically processed by viral-encoded protease(s) during translation (Fig. 3). The large ORF in the genome of PYFV putatively encodes a polyprotein consisting of 3027 aa residues with a predicted molecular mass of about 336 kb. The large ORF in the genomes of waikaviruses encodes a polyprotein of approximately 400 kb, which has about 3440–3470 aa residues. In addition to the ORF for the polyprotein, short ORFs were also identified near the 30 and the 50 ends of MCDV and RTSV genomes. Two RNA species which seemingly correspond to sub-genomic transcripts from the short ORFs locating near the 30 end of the RTSV genome (strain A) were detected from infected plants. However, the lengths and locations of the short ORF in the genomes of waikaviruses vary considerably among viruses and isolates, and the presence of products translated from these ORFs has not been confirmed in plants. Another small ORF overlapping the N-terminal leader protein, ORFX, was identified by bioinformatic analysis. ORFX sequence is conserved across waikaviruses, but is not found in sequiviruses. ORFX expression is not experimentally

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confirmed and mode of expression is not known. An AUG is present, but upstream sequence is also in frame. Waikaviruses and sequiviruses are likely to have a 50 virus-linked protein (VPg) and not a 5-methyl guanosine cap, but covalent linkage of a 50 VPg has not been demonstrated. The 50 -untranslated region (UTR) in the genome of PYFV is about 0.28 kb in length, while those of waikaviruses are longer, about 0.43 kb in MCDV and 0.52 kb in RTSV. The 50 -UTR in the genomes of MCDV and RTSV has several AUG sequences upstream of the putative polyprotein start site, and appear to form extensive secondary structures. It is possible that such secondary structures may serve as the internal ribosomal entry site, as observed in the genomes of picornaviruses, to avoid the interference on translation by upstream AUG sequences, but no experimental evidence for this is yet available. No stable secondary structures are recognized in the 50 UTR of the PYFV genome; however, several short stretches of pyrimidines (UCUCUY) are present in the region. The 30 UTR in the genomes of waikaviruses are polyadenylated and unusually long. The length of 30 UTR in the genomes of MCDV and RTSV are approximately 1.0 and 1.2 kb, respectively, provided that the short ORFs near the 30 end of genomes are not translated. The 30 UTR in the genome of PYFV is approximately 0.5 kb in length. Unlike the genomes of waikaviruses, that of PYFV is not polyadenylated, but a part of 30 UTR is likely to form a stem–loop structure which is similar to that found in the genomes of flaviviruses.

Properties of Viral Proteins Polyproteins of sequiviruses and waikaviruses are predicted to be cleaved into seven or more mature proteins through proteolytic maturation. Polyproteins of sequiviruses and waikaviruses contain a protease region similar to the 3C cysteine protease of picornaviruses and other viral proteases resembling the 3C protease such as the 24 kb protease of family member cowpea mosaic virus (CPMV) and the NIa protease of the potyvirus tobacco etch virus. For example, considerable similarity was found between the region delimited by the aa residues 2643 and 2853 in the polyprotein of RTSV (strain A), and a region in the 24 kb protease of CPMV. RTSV protease cleavage sites are not clearly conserved in MCDV, and the sequence context is not well-defined. 3C-like protease cleavage sites are not reliably predicted, but usually cleavage occurs between a glutamine (Q) residue and a small amino acid residue (e.g., M, S, V, A, L, I, G). Possible cleavage sites within polyproteins have only been validated for a subset of mature proteins. Whether other protease activity besides the 3C-like protease is encoded by sequiviruses or waikaviruses is also not experimentally determined. The N-terminal part of the polyprotein is highly divergent between waikaviruses and sequiviruses, and among members of the family Secoviridae. Within the family, proteins encoded in this position function in virus movement (e.g., Bean pod mottle virus; Fig. 3) or have unknown function. Although the predicted sizes of the putative leader proteins are about 40 kDa in PYFV and 70–78 kDa in waikaviruses, the sizes of proteins detected with the antisera specific to the putative leader proteins are significantly smaller than predicted. An antiserum specific to the putative RTSV leader protein detected a protein of about 32 kDa in extracts from infected plants. An antiserum specific to the ca. 78 kDa MCDV leader protein detected proteins with of ca. 50, 35, and 25 kDa in extracts from plants infected with MCDV-S. The MCDV 78 kDa polyprotein was demonstrated to be cleaved by the virusencoded 3C-like protease into p51 and p27 proteins. The precursor 78 kDa protein and p51 have silencing suppressor activity. Silencing suppressor function is conserved in BVCV and other waikaviruses but appears to have variable strength in the agroinfiltration assay among the orthologs. No evidence for silencing suppressor activity or in vitro cleavage by the cognate viral protease was observed for the corresponding N-terminal ca. 43 kDa PYFV protein, which is sequence divergent from similarly-positioned waikavirus proteins and may not be proteolytically processed further. A leader protein found in the polyprotein of aphthoviruses in the family Picornaviridae has a protease activity which cleaves itself autocatalytically from the polyprotein. It was proposed that the ca. 35 kDa protein of MCDV is generated from the leader protein region through autoproteolysis of p51 at the putative cleavage site ALVRLFHGSAE (aa residues 150–160), while p27 results from the cleavage at Q445/S446 and Q686/S687 by the viral 3C-like protease. Autocatalytic cleavage has not been experimentally demonstrated, however. The p27 product of the MCDV 78 kDa protein has been implicated as a helper component since a protein of this size was detectable in leafhoppers fed on infected plant using antiserum against the MCDV leader polyprotein, but function has not been definitively demonstrated. This postulated role of p27 is consistent with the result from a serological blocking experiment showing that the helper component is not the virion or CPs. Three consecutive CPs are encoded after the N-terminal protein region of the sequi- and waikaviruses. Protease cleavage sites of the N-termini of CPs are defined by amino acid sequencing of purified virion proteins for PYFV, MCDV-S and MCDV-TN, and RTSV. The proteolytic cleavage sites for the RTSV CPs determined by N-terminal amino acid sequencing were mapped to Q644/ A645, Q852/S853, and Q1055/D1056 for the junctions between leader protein/CP1, CP1/CP2, and CP2/CP3, respectively. Similarly, MCDV-S CP junctions were mapped to Q686/S687, Q896/L897, and Q1098/V1099 from N to C-terminus. The context of these cleavage sites suggests that the CPs are processed in trans by the viral 3C-like protease. However, cleavage at these sites with the RTSV protease was not detected in vitro. Based on N-terminal sequencing of CPs, PYFV sequenced CP junctions were mapped to S394/P395, N588/A589, and Q810/A811. The difference in the reactivity among the CPs of RTSV with the antibody raised against the virus particles indicated that the largest 33 kDa CP (CP3) is the major antigenic determinant on the surface of particles, although the structural details of virus particles have not been elucidated yet. The size of CP3 of RTSV (Philippine isolate) in the crude extract from infected plants detected by the antibody specific to CP3 appeared to be 40–42 kDa, markedly larger than that detected from purified virus

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preparation or that expected from the genome sequence. It is likely that the size of CP3 in infected cells is larger due to posttranslational modification, but the modified moiety appeared to be cleaved off probably by the treatment with cellulolytic enzymes during virion purification. For MCDV, the antisera against the purified virion detects the largest two coat proteins, CP1 and CP2, most strongly, and CP3 weakly or not at all, suggesting that this coat protein is internal or less antigenic. (For MCDV, CPs are numbered by size rather than ORF location as for RTSV, and the most antigenic CPs correspond to the largest ones in both.) A slower-migrating protein was also observed with MCDV virion antisera, which was also hypothesized to be a post-translationally modified largest CP. The central region flanked by the CPs and the protease regions in the polyproteins of viruses belonging to the family Secoviridae contains sequences with similarities to helicase, protease, and RNA-dependent RNA polymerase (RdRp) of picornaviruses and comoviruses. The NTP-binding domain of the central polyprotein regions shows extensive similarity to the corresponding domains in the 58 kDa protein of CPMV and the 2C proteins of picornaviruses and contains motifs GX4GKS and DD, which are conserved among proteins with NTP-binding domains. The 58 kDa protein and the 2C protein presumably function as a nucleoside triphosphatase (NTPase)/helicase required for the initiation of negative-strand RNA synthesis. Antibodies specific to protein segments from the central region of the MCDV polyprotein reacted with three protein species in extracts from infected plants, indicating that the central region containing NTP-binding and helicase domains is likely also processed into smaller proteins. The 3C-like protease of sequi- and waikaviruses follows the central region and precedes the polymerase. Immunodetection using an antiserum raised against a protein containing the putative protease region of RTSV indicated that the size of mature protease in infected plants is approximately 35 kDa. The sequence context of cleavage sites for the 3C-like cysteine protease and the results from in vitro translation and autocatalytic cleavage assays from partial genomic RNA templates defined the region of the protease in the RTSV polyprotein to be from aa 2527–2852. An internal cleavage site of the RTSV protease was also identified in vitro in a later study, but its biological significance, if any, is not known. The RTSV protease presumably acts in cis to cleave itself from the adjacent protein regions, but trans-cleavage by the RTSV protease was also observed to occur between the protease and the putative helicase regions in vitro. Based on the sequence alignment among the 3C-type proteases, aa Glu2717 and Cys2811 in the RTSV polyprotein were predicted to constitute part of the catalytic triad with His2680 and His2830 as contributing residues. Substitutions of these aa abolished or drastically reduced the proteolytic activity, substantiating their important roles in the protease. These aa are conserved in the protease regions of other waikaviruses. The C-terminal region in the polyproteins of sequi- and waikaviruses is a predicted RNA-dependent RNA polymerase (RdRp). The RdRp region of RTSV (strain A) was defined in the region between aa 2853 and 3473 with the conserved YGDD motif at aa 3270–3273. The RdRp region of MCDV (isolate TN) has the conserved YGDD motif at aa 3238–3241. In addition, motifs such as DYSXFDG (aa 3129–3135 in the MCDV polyprotein) and PSGX3TX3NS (aa3189–3200) were identified to be conserved among the RdRp regions of MCDV, CPMV, and Tomato black ring virus (TBRV). Multiple alignment of the RdRp region of PYFV (isolate P-121) with those of CPMV, TBRV, and Poliovirus showed that they share several conserved motifs including sequences YGDD (aa 2629–2632 in the PYFV polyprotein), PSGX3TX3NS (aa 2580–2591), and FLKR (aa 2684–2687).

Transmission Several waikaviruses exhibit transmission ‘helper’ phenomena, as observed in cow parsley infected with PYFV and the waikavirus Anthriscus yellows virus (AYV), and rice plants infected with Rice tungro bacilliform tungrovirus (RTBV) and the waikavirus RTSV. In these cases mixed infections are the outcome of a waikavirus ‘helping’ insect transmission of another virus (AYV helping PYFV or RTSV helping RTBV). Due to transmission helping, both the waikavirus and the virus it helps are also transmitted by aphid or leafhopper vectors in a semipersistent manner, and in both cases the non-waikavirus cannot otherwise be transmitted by the insect vector. PYFV is transmitted by aphids from plants infected with both viruses or by aphids previously fed on plants infected with AYV. Aphids seem to retain PYFV and AYV for up to 4 days. Even though AYV cannot infect parsnip, PYFV can be transmitted to parsnip by aphids fed on other plants infected with AYV and PYFV. Therefore, AYV apparently acts only as the helper virus for aphids to acquire PYFV, and is not necessary for the infection process of PYFV. The name of the genus Sequivirus comes from the Latin word ‘sequi’, which means to follow or accompany, in reference to the dependent insect transmission of PYFV. Incidences of mixed infection in lettuce with dandelion yellow mosaic virus and lettuce mosaic virus were also reported, but their relationships in aphid-mediated transmission are still unclear. Green leafhoppers (Nephotettix spp.) transmit RTSV and RTBV simultaneously or singly from source plants infected with both viruses. RTSV is independently transmitted by leafhoppers, but RTBV can only be transmitted by leafhoppers that have fed on plants infected with RTSV. Although leafhoppers retain RTSV for only 3–4 days, their ability to acquire and transmit RTBV may persist for up to 7 days. Neutralization of RTSV-viruliferous green leafhoppers with anti-RTSV immunoglobulin markedly reduced the ability to transmit RTSV but still retained the ability to acquire and transmit RTBV. These observations indicate that while RTSV is the helper virus for the leafhopper-mediated transmission of RTBV, the helper function is associated with factors other than RTSV virions or CP. Evidence for a helper component in the waikavirus MCDV is based on experiments showing requirements for its own transmission, which include a component not purified with virions in membrane feeding experiments. Purified MCDV-S virions fed to

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blackfaced leafhopper vectors by membrane feeding were not transmitted unless leafhoppers were previously fed on plants infected with the MCDV mild strain MCDV-M1. Leafoppers fed on MCDV show an electron-dense “matrix” in the foreguts observed by transmission electron microscopy, which has been proposed to be composed of helper component protein. The ca. 25 kDa proteolysis product of MCDV R78 (likely p27) was postulated to be the helper component protein based on detected retention in virus-exposed leafhoppers. However, the helper component for MCDV or other waikaviruses has not yet been definitively demonstrated. For most more recently discovered members of the genera Sequivirus and Waikavirus discovered by deep sequencing, the vectors and transmission modes are not yet described.

Diseases and Management Important diseases are described for the most-studied waika- and sequiviruses, but not known for some of the viruses identified by sequences. Major diseases caused by waikaviruses include rice tungro disease and maize chlorotic dwarf disease. Rice tungro disease is caused by the synergistic coinfection of RTSV and RTBV. Both viruses are transmitted by green leafhoppers in a semipersistent manner. RTSV alone does not cause conspicuous symptoms except occasional slight stunted growth, but severe disease results when plants are co-infected with RTSV and RTBV (Fig. 1). Such synergistic effects of RTSV on symptom development are also observed when RTSV co-infects plants with viruses such as Rice grassy stunt virus and Rice ragged stunt virus. Rice plants infected with RTBV alone exhibit symptoms such as stunted growth, yellow to yellow-orange discoloration of leaves, and reduced tillering. Severity of rice tungro disease outbreaks vary from year to year with vector populations. It is managed with vector control using insecticides or crop rotation as well as genetic resistance in rice to leafhoppers and RTSV. MCDV is present in the U.S. and is managed by removal of its overwintering host, Johnsongrass (Sorghum halepense) in and around fields. Other unidentified changes in cultivation practices may have contributed to its incidence reduction since the 1970s. Serological and amplification-based assays are effective for diagnostics of both RTSV and MCDV. PYFV and CNDV are pathogens of vegetable crops, with PYFV causing yellowing and mosaic of parsnip and celery, and CNDV causing mosaic, necrosis and dieback in carrot, chervil, coriander, and dill. These are managed by planting seed in areas with no virus reservoir and limited vector aphid populations.

Concluding Remarks Waikaviruses and Sequiviruses are unique monopartite viruses that include important pathogens and have unusual molecular biology and transmission features. Only a few viruses are well-described from each genus, although additional viruses with sequences similarity continue to be identified by deep sequencing, and tools continue to add information about the biology and molecular biology of these viruses in the family Secoviridae.

Further Reading Ammar, E.D., Nault, L.R., 1991. Maize chlorotic dwarf viruslike particles associated with the foregut in vector and nonvector leafhopper species. Phytopathology 81, 444–448. Azzam, O., Chancellor, T.C.B., 2002. The biology, epidemiology, and management of rice tungro disease in Asia. Plant Disease 86, 88–100. Azzam, O., Yambao, M.L.M., Muhsin, M., McNally, K.L., Umadhay, K.M.L., 2000. Genetic diversity of Rice tungro spherical virus in tungro-endemic provinces of the Philippines and Indonesia. Archives of Virology 145, 1183–1197. Chaouch-Hamada, R., Redinbaugh, M.G., Gingery, R.E., Willie, K., Hogenhout, S.A., 2004. Accumulation of maize chlorotic dwarf virus proteins in its plant host and leafhopper vector. Virology 325, 379–388. Elnagar, S., Murant, A.F., 1976. Relations of the semi-persistent viruses, parsnip yellow fleck and anthriscus yellows, with their vector, Cavariella aegopodii. Annals of Applied Biology 84, 153–167. Firth, A., Atkins, J., 2008. Bioinformatic analysis suggests that a conserved ORF in the waikaviruses encodes an overlapping genes. Archives of Virology 153, 1379–1383. Galvez, G.E., 1971. Rice tungro virus. Description of Plant Viruses 67. Available at: http://www.dpvweb.net/dpv/showadpv.php?dpvno¼ 67. Gingery, R.E., Bradfute, O.E., Gordon, D.T., Nault, L.R., 1978. Maize chlorotic dwarf virus. Description of Plant Viruses 67. Available at: http://www.dpvweb.net/dpv/showdpv. php?dpvno ¼ 194. Hibino, H., 1983. Transmission of two Rice tungro-associated viruses and Rice waika virus from doubly or singly infected source plants by leafhopper vectors. Plant Disease 67, 774–777. Hull, R., 1996. Molecular biology of rice tungro viruses. Annual Review of Phytopathology 34, 275–297. Hunt, R.E., Nault, L.R., Gingery, R.E., 1988. Evidence for infectivity of Maize chlorotic dwarf virus and for a helper component in its leafhopper transmission. Phytopathology 78, 499–504. Isogai, M., Cabauatan, P.Q., Masuta, C., Uyeda, I., Azzam, O., 2000. Complete nucleotide sequence of the Rice tungro spherical virus genome of the highly virulent strain Vt6. Virus Genes 20, 79–85. Murant, A.F., 2003. Parsnip yellow fleck virus. Description of Plant Viruses 394. Available at: http://www.dpvweb.net/dpv/showadpv.php?dpvno ¼ 394. Reddick, B.B., Habera, L.F., Law, M.D., 1997. Nucleotide sequence and taxonomy of Maize chlorotic dwarf virus within the family Sequiviridae. Journal of General Virology 78, 1165–1174. Shen, P., Kaniewska, M., Smith, C., Beachy, R.N., 1993. Nucleotide sequence and genomic organization of Rice tungro spherical virus. Virology 193, 621–630. Stewart, L.R., 2011. Waikaviruses: Studied but not understood. APS Features. doi:10.1094/APSFeature-2011-11. Stewart, L.R., Jarugula, S., Zhao, Y., Qu, F., Marty, D., 2016. Identification of a Maize chlorotic dwarf virus silencing suppressor protein. Virology 504, 88–95. Thole, V., Hull, R., 1998. Rice tungro spherical virus polyprotein processing: Identification of a virus-encoded protease and mutational analysis of putative cleavage sites. Virology 247, 106–114. Turnbull-Ross, A.D., Mayo, M.A., Reavy, B., Murant, A.F., 1993. Sequence analysis of the Parsnip yellow fleck virus polyprotein: Evidence of affinities with picornaviruses. Journal of General Virology 74, 555–561.

Sequiviruses and Waikaviruses (Secoviridae)

Relevant Websites https://viralzone.expasy.org/660?outline=all_by_species Sequivirus B ViralZone page. https://viralzone.expasy.org/661?outline=all_by_species Waikavirus B ViralZone page.

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Solemoviruses (Solemoviridae) Cecilia Sarmiento, Merike Sõmera, and Erkki Truve†, Tallinn University of Technology, Tallinn, Estonia r 2021 Published by Elsevier Ltd. This is an update of M. Meier, A. Olspert, C. Sarmiento, E. Truve, Sobemovirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00507-0.

Nomenclature

nt Nucleotide(s) ORF Open reading frame RdRp RNA-dependent RNA polymerase satRNA Satellite RNA sgRNA Sub-genomic RNA TM Trans-membrane anchor UTR Untranslated region VPg Viral protein genome-linked

aa Amino acid(s) ARM Arginine-rich motif CP Coat protein or capsid protein diRNA Defective interfering RNA kb Kilobase kDa Kilo dalton MP Movement protein NLS Nuclear localization signal

Glossary Icosahedral particle Spherical viral particle that is a polyhedron having 20 faces. Leaky scanning of ribosomes Mechanism during translation initiation for escaping the first start codon, which occurs when the first AUG resides in a very poor context and therefore only some ribosomes initiate translation at that point.

Ribosomal frameshifting
1 Event occurring during translation elongation when the ribosome shifts its frame for reading the mRNA exactly one position in the upstream direction. Satellite RNA Subviral agent consisting of RNA that becomes packaged in protein shells made from coat protein of the helper virus and whose replication is dependent on that virus.

Introduction Solemoviruses are plant viruses with single-stranded RNA (ssRNA) genome of positive polarity with small icosahedral virion. Some solemoviruses are distributed throughout the world; others are limited to one continent or even to one endemic region. A number of solemoviruses are the causative agents of viral diseases of important crops. Rice yellow mottle virus (RYMV) is economically the most important virus causing 20%–100% yield loss of rice production in Africa. Subterranean clover mottle virus (SCMoV) damages seriously Australian grasslands, and Papaya lethal yellowing virus (PLYV) is responsible for a severe disease in Brazilian papaya plantations. The first isolated solemovirus was Southern bean mosaic virus (SBMV), described in 1943. It was proposed in 1969 that singlecomponent-RNA beetle-transmitted viruses be placed into a Southern bean mosaic virus group. Since 1995, this group was recognized by the International Committee on Taxonomy of Viruses (ICTV) as the genus Sobemovirus, unassigned to any family. The family Solemoviridae was created in 2017 to group together the genera Sobemovirus (including 19 species) and Polemovirus (with only one member; Table 1). During the last years, the list of members has increased and is expected to keep growing, especially due to new available metagenomic data. From the name Southern bean mosaic virus derives the genus name Sobemovirus. The name of the genus Polemovirus comes from polerovirus and sobemovirus to indicate the relationship of polemoviruses to those genera, as explained in the genome section.

Virion Structure Solemovirus particles are very stable icosahedral virions of 25–34 nm in diameter displaying T¼3 symmetry. The particle is built of 180 coat protein (CP) molecules (B30 kDa) that are translated from sub-genomic RNA (sgRNA). The single-stranded genomic RNA and sgRNA together with a viral genome-linked protein (VPg) are packaged inside the virion. The three-dimensional structures of Cocksfoot mottle virus (CfMV), Sesbania mosaic virus (SeMV), Southern cowpea mosaic virus (SCPMV), RYMV, and Ryegrass mottle virus (RGMoV) virions have been determined utilizing X-ray crystallography. CP monomers are chemically identical and can adopt three different conformations (A, B, and C). The A subunits group at 5-fold axes to form 12 pentamers, while pairs of B and C subunits meet at 3-fold axes to form 20 hexamers. The combination of hexamers and pentamers gives the particle its characteristic shape. The atomic structure of SBMV particle drawn with computer graphics is shown in Fig. 1 (A subunits in red, B and C in blue and green). †

Deceased.

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Solemoviruses (Solemoviridae)

Table 1

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Members of the family Solemoviridae

Genus/Species

Acronym

Accession #

Sobemovirus Artemisia virus A Blueberry shoestring virus Cocksfoot mottle virus Cymbidium chlorotic mosaic virus Imperata yellow mottle virus Lucerne transient streak virus Papaya lethal yellowing virus Rice yellow mottle virus Rottboellia yellow mottle virus Ryegrass mottle virus Sesbania mosaic virus Solanum nodiflorum mottle virus

ArtVA BSSV CfMV CyCMV IYMV LTSV PLYV RYMV RoMoV RGMoV SeMV SNMoV

NC_017914.1 NC_029578.1 NC_002618.2 NC_027123.1 NC_011536.1 NC_001696.2 NC_018449.1 NC_001575.2 NC_027198.1 NC_003747.2 NC_002568.2 NC_033706.1

Southern bean mosaic virus Southern cowpea mosaic virus Sowbane mosaic virus Soybean yellow common mosaic virus Subterranean clover mottle virus Turnip rosette virus Velvet tobacco mottle virus

SBMV SCPMV SoMV SYCMV SCMoV TRoV VTMoV

NC_004060.2 NC_001625.2 NC_011187.1 NC_016033.1 NC_004346.1 NC_004553.3 NC_014509.2

Polemovirus Poinsettia latent virus

PnLV

NC_011543.1

Vector

aphid beetle

beetle

beetle, mirid beetle beetle aphid, leafminer, leafhopper, mirid beetle beetle, mirid beetle, mirid

Fig. 1 Atomic structure of the entire virus particle of SBMV shown with computer graphics. Each subunit is shown as the Ca tracing in different colors to depict the different geometric environments within the icosahedral shell. Red: Icosahedral 5-fold; Blue–Green: icosahedral 3-fold or quasi6-fold clusters; Green at the center of the viral particle: icosahedral 2-fold. From Abad-Zapatero., C., 2002. Crystals and Life: A Personal Journey. International University Line, ISBN: 0-9720774-0-5. plate XVIII, with permission.

The root mean square deviations between superimposed coordinates of Ca atoms of solemoviral CP residues are in general 1.4–1.5 Å . Interestingly, the virion of RGMoV is smaller and slightly more similar to the virion of Tobacco necrosis virus (family Tombusviridae) than to other solemovirus virions. Homology modeling of the 3D structures of the Rottboellia yellow mottle virus (RoMoV) and Artemisia virus A (ArtVA) CPs confirm with 100% confidence that they adopt the overall fold shown for the RGMoV CP and tombusvirus-like viral CPs. In addition to protein–protein and protein–RNA interactions, each icosahedral unit is stabilized by three calcium-binding sites located between subunits AB, BC, and CA. Studies with SCPMV, SeMV, and RYMV particles

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demonstrate that the stability of the virions depends greatly on pH and the availability of calcium ions. Upon alkaline pH or removal of the cations, the virus particles swell and become less stable. The primary sequences of CPs among solemoviruses are quite different. However, the three-dimensional structures are nearly identical. The CP is divided into two domains: C-terminal S (shell) domain, which has an eight-strand jellyroll b-sandwich topology, common to non-enveloped icosahedral viruses, and N-terminal R (random) domain, which is disordered in subunits A and B, but is partially structured in subunit C. The disordered beginning of R domain seems to interact with the genomic RNA in the interior of the virus particle. The S domain is the building block of the virion. The R domain is rich in basic amino acid residues and contains at its first half an arginine-rich motif (ARM). The ARM comprises a bipartite nuclear targeting signal (NLS). In the case of CfMV, abolishment of the NLS does not affect virus infectivity. Studies with SCPMV and SeMV CPs demonstrate that ARM is essential for RNA encapsidation but not for particle formation. However, the presence of RNA enhances the overall stability of capsids. Packing of viral nucleic acid is supposed to happen when CP recognizes a specific region inside the virus genome. In the case of SCPMV, a putative stem-loop sequence within a conserved region encoding serine protease has been reported to bind CP. Nevertheless, this sequence has not been demonstrated to nucleate SCPMV assembly. According to a number of studies, solemovirus particles assemble as follows: CP subunits with disordered amino termini form a pentamer of AB dimers. Then, ARM interacts with RNA and CC dimers are formed. More CC dimers are added and a swollen T¼3 particle is built that becomes compacted when calcium ions enter the subunit interfaces. Virion disassembly occurs when cations are removed. In the case of SBMV virions, pentamers have been suggested as exit ports for viral RNA. Particles of this virus disassemble completely after initiation of RNA translation. In addition to the genomic and sgRNA, some solemoviruses (Lucerne transient streak virus (LTSV), Solanum nodiflorum mottle virus (SNMoV), SCMoV, Velvet tobacco mottle virus (VTMoV), and RYMV) encapsidate a circular viroid-like satellite RNA (satRNA). The sizes of these circular satRNAs range from 220 to 390 nt, RYMV satRNA being the smallest (220 nt) known naturally occurring circular RNA.

Genome Solemoviral genomes are 4.0–4.6 kb in length. The polycistronic positive-sense ssRNA genome consists of five open reading frames (ORFs) in the case of sobemoviruses and four in the case of polemoviruses (Fig. 2). All ORFs, except the most 30 proximal, are expressed from the genomic template. The last ORF, encoding CP, is translated from the sgRNA. The genomic and sub-genomic RNAs have a VPg covalently bound to its 50 end. The 30 terminus of the genomic RNA is nonpolyadenylated. An approximately 100 nt long 50 UTR and up to 289 nt long 30 UTR are supposed to compensate the absence of cap and poly(A) tail. VPg may attach to the genome via phosphodiester bond formed through tyrosine, serine, or threonine residues, depending on the viral species. The 50 UTR of sobemoviruses comprise a polypurine tract and 30 UTR are more diverse. In the case of CfMV, 50 UTR acts as an enhancer of translation and 30 UTR seems to contain sequences or structures important for the transport of viral RNA. For this virus, as well as for RYMV, a potential tRNA-like structure has been predicted at the 30 UTR. SeMV 30 UTR has a stem-loop 28–55 nt upstream of the 30 end that is important for RNA-dependent RNA polymerase (RdRp) template recognition. Interestingly, also the polemovirus Poinsettia latent virus (PnLV) has a stable hairpin at its 30 UTR, which buries the last nucleotide. Not all, but most of the solemoviral genomes – including PnLV – have a conserved 50 sequence ACAA(AA), considered to play a role in viral RNA replication. This motif is also present upstream of the translation initiation codon of CPs, as a possible 50 terminus of sgRNA. In addition to the approximately 1 kb sgRNA encoding the CP, PnLV has a larger putative sgRNA of 2.60 kb. Further studies are needed to find out whether this is a real sgRNA or a defective RNA. In fact, sobemovirus CfMV particles may contain different defective interfering RNAs (diRNA).

Fig. 2 Genome organization of the genera Solemovirus and Polemovirus in the family Solemoviridae. Comparison of the genome organization of Southern bean mosaic virus (SBMV) and Poinsettia latent virus (PnLV), type species of the corresponding genera. ORFs encoding proteins sharing sequence homology are indicated in similar colors: silencing suppressor (violet), serine protease-VPg-s (red), RNA-dependent RNA polymerases (yellow), coat proteins (green). VPg-s are depicted as red dots at the 50 -end of the genomic RNA. Proteins with unknown function and no sequence homology are shown in gray. Non-canonical AUG start codon is marked with asterisk (*).
1 ribosomal frameshifting is marked by long vertical dashed line.

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Fig. 3 Maximum-likelihood trees showing phylogenetic relationships between solemoviruses and viruses belonging to related families. Phylogenetic relationships between members of the genera Sobemovirus (family Solemoviridae, shown in green), Polemovirus (family Solemoviridae, shown in teal), Polerovirus (family Luteoviridae, shown in blue), Enamovirus (family Luteoviridae, shown in turquoise), Luteovirus (family Luteoviridae, shown in orange), Alphanecrovirus and Betanecrovirus (family Tombusviridae, shown in red), Barnavirus (family Barnaviridae, shown in violet), and Dinornavirus (family Alvernaviridae, shown in gray) demonstrate the discordant placement of these genera on the unrooted maximum-likelihood (ML) trees. ML trees based on amino acid sequence alignments were predicted for RNA-dependent RNA polymerases (RdRp-s, panel A) and for coat proteins (CPs, panel B). Genera Polerovirus and Luteovirus are represented by a reduced number of species. Species officially recognized by ICTV are shown in italics. The ML trees were calculated with 1000 bootstrap replicates using the Whelan and Goldman (WAG) model for RdRps and Le Gascuel (LG) model for CPs. The percentage of trees in which the associated taxa clustered together is shown next to the branches if 470. Branch lengths are proportional to genetic divergence. Scale indicates the number of amino acid substitutions per site.

Solemoviruses differ in the 50 terminal part of their genomes. This is part of the differences between the members of the genera Polemovirus and Sobemovirus. Indeed, the name Polemovirus comes from polerovirus and sobemovirus to indicate the chimeric nature of this genus: the 50 -three quarters of the genome are similar in organization and sequence to poleroviruses (polemoviral RdRp groups together with poleroviral RdRps, Fig. 3(A)) and the CP shares the highest homology with sobemoviruses (Fig. 3(B)). The most 50 proximal ORF in the case of polemovirus is ORF0 that encodes a protein of 30 kDa of unknown function. Sobemoviruses first ORF is called ORF1 and encodes the protein P1, a suppressor of RNA silencing. P1 is dispensable for replication and is suspected to be the viral movement protein, as it has been shown to be required for systemic infection. Moreover, in the case of SeMV, P1 has been shown to interact with its genome-bound VPg and with P10, an ATPase that could

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supply the needed energy for an active transport. It remains to be elucidated, if P1 RNA silencing suppressor function is uncoupled from its role in viral movement. The only conserved element among sobemoviral P1 is a Zn-finger motif that is required for suppressing RNA silencing. ORF1 partially overlaps the small ORFx from which the Px protein is translated. The translation initiation codon of ORF1 is located in an unfavorable context and therefore a leaky scanning of ribosomes occurs. Since Px translation starts from a non-AUG codon, protein synthesis rates are very low. Px is a protein of unknown function and it has been shown to be needed for viral infection in the case of Turnip rosette virus (TRoV). ORFx and ORF1 are the most variable regions in sobemoviral genomes. No small ORF has been detected for polemovirus PnLV. All solemoviruses comprise at the central part of their genome two overlapping ORFs. The second ORF always encodes the RdRp that is translated via
1 ribosomal frameshifting. Polemoviral ORF1 contains putative helicase and protease domains as well as a predicted VPg sequence in-between (altogether the product is 72 kDa), while ORF2 encodes the RdRp (122 kDa). In the case of sobemoviruses, the ineffective initiation of translation of P1 and Px proteins allows the expression of ORF2a that encodes the most abundant polyprotein P2a. ORF2a encodes an N-terminal trans-membrane anchor (TM), a virus serine protease (Pro) and the VPg. The exact functions of the proteins encoded by the C-terminal portion of ORF2a are unclear, but this part has been shown to have ATPase and NTP-binding properties and is further processed to P10 and P8, at least in the case of SeMV. Sobemoviral ORF2b is expressed as a fusion protein with ORF2a through
1 ribosomal frameshifting shortly after VPg domain. For CfMV it is known that in 10%–20% of the cases the longer version polyprotein P2ab (TM–Pro–VPg–RdRP) is translated instead of the regular polyprotein P2a (TM-–Pro–VPg–P10–P8). The
1 ribosomal frameshift is needed to regulate the production of sobemoviral RdRp, which is encoded by ORF2b and involves two elements: a slippery sequence conserved among sobemoviruses (UUUAAAC) and a stem-loop located 7 or 8 nt downstream of it. PnLV slippery sequence for RdRp translation seems to be GGGAAAC, shared by some poleroviruses. Proteolytic cleavages in the polyproteins are carried out by the viral serine protease that cuts at E/T residues for SBMV, at E/N and E/T residues for CfMV, at E/S and E/T residues for RYMV and at E/S residues in the case of RGMoV. The C-terminal cleavage of VPg takes place upstream of the slippery sequence. SeMV is the only solemovirus for which all four cleavage sites have been characterized and its protease domain has been crystallized. The structure exhibits the characteristic features of trypsin fold. Mutations of glutamate-binding site residues H298, T279, and N308 render the protease inactive. Pro–VPg is crucial for protease activity and this precursor protein accumulates in membranes of infected cells. Notably, Pro–VPg is released from P2a, while Pro is detected when P2ab is expressed. TM putative function is to anchor the polyprotein into cellular membranes to facilitate proteolytic processing and probably minus strand synthesis. If no VPg processing occurs, the virus is not viable. VPg is an intrinsically disordered protein that is able to bind different partners. SeMV VPg has been shown to interact with P1 but not with RdRp. Importantly, RYMV VPg interacts with eukaryotic translation initiation factor eIF(iso)4G and it seems that also P8 is involved in this binding. SeMV P8 contains an RNA binding region and is needed for the ATPase activity of P10 in the precursor P10–P8. SeMV P10 partners are P1, as mentioned before, and RdRp, meaning that this protein may be related to virus movement and replication. According to their sequences, RdRp-s of solemoviruses are grouped together with the RdRps of polero-, enamo-, and barnaviruses (Fig. 3(A)). This is the so-called “sobemo-lineage” in the supergroup I of plus-sense plant RNA viruses. RdRps of this group seem to lack the NTP-binding region conserved in viral helicases. The GDD motif (SGSYCTSSTNX19–35GDD) that characterizes RdRps of RNA viruses with ssRNA and positive polarity genomes is essential. Studies on SeMV RdRp have shown that it can synthesize RNA in a primer-independent manner. This polymerase interacts at its C-terminal region with P10 that apparently catalyzes RdRp activity. Some solemoviral CPs are related to processes different to capsid formation. CP of SeMV has been reported to interact with P1, protein related to viral movement, as explained before. Interestingly, CP of CfMV has been shown to act as suppressor of RNA silencing, in addition to P1.

Phylogenetic Relationships The viruses belonging to the families Solemoviridae, Luteoviridae, and Tombusviridae seem to have evolved via recombinational shuffling of the genes encoding the protease, RdRp and CP. The sequences of solemoviral long polyproteins are similar to the polyproteins of polero- and enamoviruses (genera belonging to the family Luteoviridae), whereas solemoviral CPs share more similarity to CPs of alpha- and betanecroviruses (belonging to the family Tombusviridae). The only known member of the genus Polemovirus is the product of a recombination between a polerovirus and a sobemovirus. As mentioned before and as shown in Fig. 3, RdRp of polemovirus PnLV is closely related to poleroviral RdRps, but its CP sequence is highly similar to sobemoviral CPs. It has been proposed that the recombination happened in the presence of a coinfection by switching templates close to the sgRNA start site during replication. Although PnLV shares similar sequences with poleroviruses and sobemoviruses, it is most closely related to the genus Sobemovirus than to any other genus and therefore, polemo- and sobemoviruses are grouped into one family. The phylogenetic trees based on RdRp and on CP sequences show discordant placement of the analyzed species. This corroborates the supposition of possible recombinational events between ancestral solemo-, luteo-, and tombusviruses (Fig. 3). In addition, representatives of other genera (Barnavirus and Dinornavirus) seem to be involved in this evolutionary process.

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Among sobemoviral CPs, sequences from RGMoV, RoMoV, and ArtVA are slightly more similar to CP sequences of several tombusviruses than to other sobemoviral CPs (Fig. 3(B)). As explained above, also the 3D structures of the CPs of these three sobemoviruses are predicted to be closely related to tombusviral CPs. Studies comparing related viruses at the level of their RdRp sequences have concluded that sobemo-, barna-, polero-, and luteoviruses divergence time is around 9000 years. If luteoviruses are not considered, then the divergence is reduced to 5000 years, and leaving apart poleroviruses means going down to 4000 years. Based on this kind of calculations, a hypothesis correlating the development of agriculture to the appearance of different viruses has been raised.

Replication and Movement The replication of solemoviruses is poorly understood. The genomic RNA of incoming virus particles is probably uncoated by the co-translational disassembly mechanism that is followed by RNA replication once the RdRp has been synthesized. Little is known about the signals needed for viral replication. As mentioned before, solemoviral genomes have a conserved 50 sequence ACAA(AA) that is also characteristic for 50 -termini of polero-, diantho-, and barnaviruses. This motif is also present at the 50 -terminus of sgRNA. Due to the conservation of this region, the complementary sequence in minus-strand has been predicted to function in replication by promoting or enhancing the binding of RdRp to both genomic and sgRNA. The stem-loop structure found at the 30 -end of some solemoviral genomes is important for template recognition by RdRp and thus, for the synthesis of minus-strand. It is not yet clear, whether solemoviral VPg primes the synthesis of viral RNA. SeMV VPg has been shown to be dispensable for in vitro synthesis of minus-strand RNA. At least five diRNA molecules have been cloned from CfMV-infected plants. Their sequences consist of the 50 -terminal 35–40 nt linked to the 850–950 nt of the 30 -terminus. This indicates the possibility that RdRp is able to switch templates and create recombinant RNA molecules. In addition to the viral genome, RdRps of some solemoviruses also replicate satRNAs. These are 220–390 nt circular molecules that are replicated by a helper virus. satRNAs have several interesting interactions with the replication of the helper viruses. For example, LTSV supports the replication of satRNA of SNMoV but SNMoV does not replicate LTSV satRNA. At the same time, LTSV satRNA replication is supported by CfMV, SBMV, Sowbane mosaic virus (SoMV), and TRoV, whereas this support is host dependent. Importantly, satRNAs influence sometimes the symptoms caused by the helper virus. satRNAs are predicted to have viroid-like secondary structures and replicate via rolling circle mechanism. After replication, monomers are self-cut by endogenous hammerhead ribozyme structures. RYMV satRNA is the smallest described so far (220 nt long). Remarkably, LTSV satRNA encodes a protein of 16 kDa with unknown function. Since the length of the circular genome is not a number divisible by three, translations happen one after another continuing from different codons in each round. Solemoviral particles are detected mainly in mesophyll and vascular tissues, but also in epidermal, bundle sheath, guard cells, and even in meristem cells, depending on the viral species. Concerning vascular tissues, virus particles have been reported in both xylem and phloem: CfMV, SCPMV, and SBMV particles have been found mainly in the phloem, whereas RYMV, SoMV, and Blueberry shoestring virus (BSSV) virions have been detected predominantly in the xylem. CfMV particles have been reported in phloem parenchyma and bundle sheath cells, when infection starts. In later stages of infection, CfMV spreads in mesophyll cells surrounding vascular bundles and is only seldom present in xylem parenchyma. At the subcellular level, virus particles are found in cytoplasm, vacuoles, and nuclei. Some sobemoviruses have NLS signals in their CP sequences that make a nuclear localization of the virion feasible. At late stages of infection, particles accumulate forming crystalline structures in the cytoplasm or vacuoles. No particles have been detected in mitochondria and chloroplasts, but the latter are noted to form finger-like extrusions in infected cells. Studies with solemoviruses emphasize that cell-to-cell and long-distance (systemic) movement are two distinct processes. For the latter movement, correct particle formation may be essential. This is the case for SCPMV and RYMV. Indeed, studies with fulllength clones of these two viruses have shown that CP is dispensable for virus replication but systemic virus movement is completely abolished in the absence of CP. In the case of SeMV, it has been proven that its virion and CP itself interact with P1, the protein related to virus movement. On the contrary, experiments with CfMV lacking CP demonstrated that this virus does not need CP for cell-to-cell or for systemic movement, implying that it moves as a ribonucleoprotein complex and not as viral particle. However, CP-deficient CfMV was not able to be transmitted mechanically using sap-inoculation. An efficient transmission of solemoviruses by vectors is facilitated by a correct virion formation that is resistant to RNases present in the regurgitant or in the hemolymph and even in feces of insects.

Host Range and Transmission The host range of individual solemoviruses is narrow – limited to one plant family – with the exception of SoMV that naturally infects plants from the families Chenopodiaceae, Vitaceae, and Rosaceae. Genetic relationships are stronger between virus species infecting plants from the same family. Most of the so far sequenced solemoviruses infect dicotyledonous plants. According to phylogenetic data, commelinid (Poales)-infecting sobemoviruses have emerged at least twice during diversification of sobemovirus species.

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Solemoviruses (Solemoviridae)

The main route of transmission for solemoviruses is mechanical wounding of plants. In addition, transmission from soil has been suggested for many solemoviruses. The polemovirus PnLV is distributed by grafting and vegetative propagation of the host plant; however, the natural means of transmission remains unknown. Viruses in the genus Sobemovirus are transmitted by vectors like beetles, aphids, bugs, and thrips. Interestingly, phylogenetically related sobemoviruses may have vectors that belong to different orders. In the case of RYMV, it has been shown that the virus is transmitted over short distances by herbivore mammals and trampling livestock. Importantly, several sobemoviruses are seed-transmissible.

Pathological and Epidemiological Aspects The polemovirus PnLV is spread worldwide in cultivated poinsettia (Euphorbia pulcherrima) plants without causing symptoms. Equally, some sobemoviruses do not provoke symptoms in their host plants. However, in most cases, solemoviral infections induce symptoms like chlorosis at different levels, mosaic or mottling patterns on leaves. In addition, stunting, necrosis, vein clearing, and even sterility have been reported. The outcomes depend on the specific virus, the host and other factors like environmental conditions and the number of viral particle present in the infected plant. As said, solemoviruses form crystalline arrays and tubules in the cytoplasm. Some of these structures are enveloped in endoplasmic reticulum-derived vesicles. Viral particles from some solemoviruses are associated with chloroplast membranes and thus, induce changes in the structure of this organelle. In CfMV- and SoMV-infected cells, a proliferation of tonoplast membranes that bulge into the vacuole takes place. Many cytological changes have been observed in RYMV-infected cells. Probably the most dramatic change induced by this virus occurs in the cell walls of parenchyma and mature xylem cells, where middle lamellae of the wall are disorganized. RYMV infection also causes important changes in the abundance of host proteins. For instance, the expression levels of several defense- and stress-related proteins like superoxide dismutase and different heat shock proteins increase several times. The changes in the levels of antioxidant enzymes and production of reactive oxygen species has been analyzed in cocksfoot plants infected with CfMV. Susceptible plants and plants with acquired resistance showed completely different regulation of the levels of antioxidant enzymes and reactive oxygen species after inoculation with CfMV. Natural resistance to solemoviruses has been detected at least for CfMV in cocksfoot, for SBMV in beans, for SCMoV in subterranean clover, for SCPMV in cowpea and for RYMV in different Oryza species including rice. The molecular mechanisms conferring resistance have only been described for RYMV in Oryza species. Pathogen-derived transgenic resistance against RYMV has been achieved by transforming plants with constructs expressing parts of ORF2. However, this RNA silencing-based resistance is low compared to natural resistance that is being exploited to obtain RYMV-resistant rice varieties.

Further Reading Adams, M.J., Jeske, H., 2012. Genus polemovirus. In: King, A.M.Q., Adams, M.J., Carstens, E.B., Lefkowitz, E.J. (Eds.), Virus taxonomy: Classification and Nomenclature of Viruses. United States: Elsevier Academic Press, pp. 1181–1184. Albar, L., Bangratz-Reyser, M., Hébrard, E., et al., 2006. Mutations in the eIF(iso)4G translation initiation factor confer high resistance of rice to Rice yellow mottle virus. The Plant Journal 47, 417–426. aus dem Siepen, M., Pohl, J.O., Koo, B.J., Wege, C., Jeske, H., 2005. Poinsettia latent virus is not a cryptic virus, but a natural polerovirus-sobemovirus hybrid. Virology 336, 240–250. Gayathri, P., Sateshkumar, P.S., Prasad, K., et al., 2006. Crystal structure of the serine protease domain of Sesbania mosaic virus polyprotein and mutational analysis of residues forming the S1-binding pocket. Virology 346, 440–451. Ling, R., Pate, A.E., Carr, J.P., Firth, A.E., 2013. An essential fifth coding ORF in the sobemoviruses. Virology 446, 397–408. Mäkelainen, K., Mäkinen, K., 2005. Factors affecting translation at the programmed
1 ribosomal frameshifting site of Cocksfoot mottle virus RNA in vivo. Nucleic Acids Research 33, 2239–2247. Meier, M., Truve, E., 2007. Sobemoviruses possess a common CfMV-like genomic organization. Archives of Virology 152, 635–640. Nair, S., Savithri, H.S., 2010. Natively unfolded nucleic acid binding P8 domain of SeMV polyprotein 2a affects the novel ATPase activity of the preceding P10 domain. FEBS Letters 584, 571–576. Olspert, A., Peil, L., Hébrard, E., Fargette, D., Truve, E., 2011. Protein-RNA linkage and post-translational modifications of two sobemovirus VPgs. Journal of General Virology 92, 445–452. Plevka, P., Tars, K., Zeltin¸š, A., et al., 2007. The three-dimensional structure of Ryegrass mottle virus at 2.9 Å resolution. Virology 369, 364–374. Sõmera, M., Truve, E., 2013. The genome organization of lucerne transient streak and turnip rosette sobemoviruses revisited. Archives of Virology 158, 673–678. Sõmera, M., Truve, E., 2015. Rottboellia yellow mottle virus is a distinct species in the genus Sobemovirus. Archives of Virology 160, 857–863. Sõmera, M., Sarmiento, C., Truve, E., 2015. Overview on sobemoviruses and a proposal for the creation of the family Sobemoviridae. Viruses 7, 3076–3115. Tamm, T., Suurväli, J., Lucchesi, Y., Olspert, A., Truve, E., 2009. Stem-loop structure of Cocksfoot mottle virus RNA is indispensable for programmed
1 ribosomal frameshifting. Virus Research 146, 73–80. Tars, K., Zeltins, A., Liljas, L., 2003. The three-dimensional structure of Cocksfoot mottle virus at 2.7 Å resolution. Virology 310, 288–289.

Tenuiviruses (Phenuiviridae) Bertha Cecilia Ramirez, The Institute for Integrative Biology of the Cell, The French Alternative Energies and Atomic Energy Commission, French National Center for Scientific Research, University of Paris-Sud, University of Paris-Saclay, Gif-sur-Yvette, France Anne-Lise Haenni, Institut Jacques Monod, French National Center for Scientific Research, Paris Diderot University, Paris, France r 2021 Elsevier Ltd. All rights reserved. This is an update of B.C. Ramirez, Tenuivirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00510-0.

Nomenclature

RdRp RNA-dependent RNA polymerase RNP Ribonucleoprotein sgRNA sub-genomic RNA ssRNA single-stranded RNA vcRNA viral complementary RNA vRNA virion RNA VSR Viral suppressor of RNA silencing

aa Amino acid(s) IR Intergenic region. kb Kilo base kDa Kilo Dalton MP Movement protein NC Nucleocapsid protein ORF Open reading frame

Glossary Ambisense polarity The presence of open reading frames (ORFs) in both the vRNA and its vcRNA, means that both strands encode proteins. This expression strategy requires strong transcription termination signals to avoid the synthesis of complementary RNA and the formation of double stranded RNA that may induce RNA silencing. Cap-snatching A mechanism of viral mRNA synthesis by the viral RdRp that cleaves off several nucleotides from the 50 terminus of host capped mRNAs and used them as primers to initiate transcription. Movement protein This refers to a viral protein that in plants assists in cell-to-cell movement of viral complexes through the plasmodesmata before systemic transport through the plant vascular system.

Persistent viral transmission This is a mode of transmission of a plant virus by an insect vector in which the virus after entering the body of the vector, replicates and circulates for systemic invasion of vector insect tissues during days to weeks. The virus passes through the insect’s midgut and salivary gland barriers for transmission to new host plants. RNA silencing An evolutionarily conserved mechanism that is active in many eukaryotic organisms (plants and insects) as antiviral defense. It is a sequence-specific genesilencing process induced by double-stranded RNA that is either targeted for degradation by the RNA-induced silencing complex or for translational repression. RNA silencing suppressor A viral factor or protein that counteracts the host plant RNA silencing defense response.

Introduction The tenuiviruses were first described in the Fifth Report of the International Committee on Taxonomy of Viruses in 1982 as nonenveloped plant viruses possibly possessing a negative ssRNA genome. Their name comes from the Latin ‘tenuis’, (thin, fine, weak) that refers to the structure of the viral particle as seen by electron microscopy (Fig. 2). Epidemics of Rice stripe virus (RStV) and Rice hoja blanca virus (RHBV), two members of the genus Tenuivirus, cause important yield losses in rice-growing areas of Asia and the former USSR, and of tropical South America, respectively. Tenuiviruses exhibit unique properties that make them different from other plant viruses. Some properties of tenuiviruses are the following: (1) The peculiar flexuous viral particles have a thread-like morphology and can adopt circular forms (Fig. 2 Left). (2) A non-structural protein (NS4) also designated disease specific (DS) protein (16–22 kDa) accumulates in large amounts in infected plants, forming large inclusions (Fig. 2 Right). (3) The viruses are persistently transmitted by a species of planthopper (Fig. 5 Bottom) in a propagative manner. For some of the members of the genus it has been demonstrated that the virus multiplies both in the host plant and in the insect vector. Multiplication of the virus in the vector may have deleterious effects on the insect. The viruses can be transmitted transovarially by viruliferous female planthoppers to their offspring, and through sperm from viruliferous males. The ability to disseminate and to infect reproductive tissues is a prerequisite for the transovarial transmission of tenuiviruses.

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(4) The genome of tenuiviruses is multi-segmented and composed of ssRNAs that have either a negative or an ambisense polarity (Fig. 3). (5) An RdRp is associated with the viral particle. (6) It has been observed for some tenuiviruses that the mRNAs are synthesized via cap-snatching (Fig. 4). (7) The presence of a highly conserved octanucleotide at the 50 (50 -ACACAAAG) and 30 (CUUUGUGU-30 ) termini of their genomic segments. (8) Tenuiviruses infect crop plants of the family Poaceae.

Taxonomy In 2016 the ICTV grouped under the order Bunyavirales all the negative stranded RNA viruses previously classified in the family Bunyaviridae and the unassigned “bunyavirus-like” viruses such as the tenuiviruses, in order to better reflect their evolutionary relationships. A total of 12 families were created in the order Bunyavirales and the family Phenuiviridae was created to group the related phleboviruses and tenuiviruses. Among these 12 families, 3 of them comprise viruses infecting plants; Fimoviridae, Tospoviridae, and Phenuiviridae. The Phenuiviridae family clusters 15 genera, and only the genus Tenuivirus hosts plant viruses (Table 1). Rice stripe tenuivirus (RStV) is the type species of the genus Tenuivirus. Other species in the genus are Echinochloa hoja blanca tenuivirus (EHBV), Iranian wheat stripe tenuivirus (IWSV), Maize stripe tenuivirus (MStV), Rice grassy stunt tenuivirus (RGSV), Rice hoja blanca tenuivirus (RHBV), and Urochloa hoja blanca tenuivirus (UHBV). Tentative species are European wheat striate mosaic virus (EWStMV), Maize yellow stripe virus (MYSV), and Melon chlorotic spot virus (MChSV) (Table 2).

Classification The criteria for species demarcation are given as follows: (1) (2) (3) (4) (5)

Vector specificity: transmission by different vector species. Different sizes and/or numbers of RNA segments. Host range: ability to infect different key plant species. Amino acid (aa) sequence identity of less than 85% between corresponding gene products. Nucleotide (nt) sequence identity of less than 60% between corresponding non-coding intergenic regions.

An example of species discrimination is that between RStV and MStV. RStV is transmitted by L. striatellus and infects 37 graminaceous species including rice and wheat. MStV is transmitted by P. maidis and infects maize, occasionally sorghum, and a few other graminaceous plants but not rice or wheat. Finally, RStV isolates have genomes of four RNA segments and the MStV genome consists of five segments. An example of difficult species demarcation is the group of the hoja blanca viruses (RHBV, EHBV, and UHBV). They have different vectors, different sizes and numbers of RNA segments, different hosts, and the nt sequence identity of their intergenic regions is less than 60%. However, the aa sequences of the four proteins on RNAs 3 and 4 are about 90% identical between RHBV, EHBV, and UHBV. Since four of five criteria are met, these viruses could be considered as distinct species that possibly separated recently and are now diverging with little field contact between them. Phylogenetic analysis of the sequence data from RNA3 and RNA4 of the tenuiviruses shows that RHBV, EHBV, and UHBV are related and form a group distinct from RStV and MStV (Fig. 1). IWSV was recently included as a member of the tenuivirus genus, it is related but distinct from RHBV and EHBV based on the phylogenetic analysis of RNA 3 and RNA 4 (Fig. 1) as well as on serology.

Virion Structure Morphology The particles also referred to as ribonucleoproteins (RNPs) are thin filaments that appear circular (Fig. 2 Left) or spiral shaped. The RNPs are 3–10 nm in diameter, with lengths proportional to the sizes of the RNAs they contain. No envelope has been observed.

Physical and Physicochemical Properties RNP preparations can be separated into four to six components by sucrose density gradient centrifugation; they correspond to the four to six RNA segments coated in the nucleocapsid protein (NC). The buoyant density of the RNP in CsCl when centrifuged to equilibrium is 1.282–1.288 g cm–3. The RNA constitutes 5%–12% of the particle weight.

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Table 1 in bold

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List of families and genera in the order Bunyavirales. Viruses infecting plants are surlined in light gray, and type species are written

Family

Genus

Species

Arenaviridae Cruliviridae Fimoviridae

Emaravirus

Actinidia chlorotic ringspot-associated emaravirus European mountain ash ringspot-associated emaravirus Fig mosaic emaravirus High Plains wheat mosaic emaravirus Pigeonpea sterility mosaic emaravirus 1 Pigeonpea sterility mosaic emaravirus 2 Raspberry leaf blotch emaravirus Redbud yellow ringspot-associated emaravirus Rose rosette emaravirus

Hantaviridae Leishbuviridae Mypoviridae Nairoviridae Peribunyaviridae Phasmaviridae Phenuiviridae

Banyangvirus Beidivirus Goukovirus Horwuvirus Hudivirus Hudovirus Kabutovirus Laulavirus Mobuvirus Phasivirus Phlebovirus Pidchovirus Tenuivirus

Tospoviridae

Wenrivirus Wubeivirus Orthotospovirus

Wupedeviridae Unassigned

Wumivirus Coguvirus

Echinochloa hoja blanca tenuivirus Iranian wheat stripe tenuivirus Maize stripe tenuivirus Rice grassy stunt tenuivirus Rice hoja blanca tenuivirus Rice stripe tenuivirus Urochloa hoja blanca tenuivirus Bean necrotic mosaic orthotospovirus Calla lily chlorotic spot orthotospovirus Capsicum chlorosis orthotospovirus Chrysanthemum stem necrosis orthotospovirus Groundnut bud necrosis orthotospovirus Groundnut ringspot orthotospovirus Groundnut yellow spot orthotospovirus Impatiens necrotic spot orthotospovirus Iris yellow spot orthotospovirus Melon severe mosaic orthotospovirus Melon yellow spot orthotospovirus Polygonum ringspot orthotospovirus Soybean vein necrosis orthotospovirus Tomato chlorotic spot orthotospovirus Tomato spotted wilt orthotospovirus Watermelon bud necrosis orthotospovirus Watermelon silver mottle orthotospovirus Zucchini lethal chlorosis orthotospovirus

Citrus coguvirus

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Table 2

Tenuiviruses (Phenuiviridae) List of member species of the genus Tenuivirus, family Phenuiviridae. The type species is written in bold

Species/Virus name

Acronym

RNA #

RefSeq

Size (Kb)

Echinochloa hoja blanca tenuivirus Echinochloa hoja blanca virus

(EHBV)

RNA 3 RNA 4 RNA 5

NC_038934.1 NC_038935.1 NC_038936.1

2.34 1.87 1.33

Iranian wheat stripe tenuivirus Iranian wheat stripe virus

(IWStV)

RNA 2 RNA 3 RNA 4

NC_038748.1 NC_038750.1 NC_038749.1

3.47 2.34 1.83

Maize stripe tenuivirus Maize stripe virus

(MStV)

RNA RNA RNA RNA

2 3 4 5

NC_038751.1 NC_038754.1 NC_038752.1 NC_038753.1

3.34 2.36 2.23 1.32

Rice grassy stunt tenuivirus Rice grassy stunt virus

(RGSV)

RNA RNA RNA RNA RNA RNA

1 2 3 4 5 6

AB009656.1 AB010376.1 AB010377.1 AB010378.1 AB000403.1 AB000404.1

9.76 4.06 3.12 2.92 2.70 2.58

Rice hoja blanca tenuivirus Rice hoja blanca virus

(RHBV)

RNA RNA RNA RNA

1 2 3 4

NC_036597.1 NC_036598.1 NC_036602.1 NC_036599.1

9.00 3.62 2.30 2.00

Rice stripe tenuivirus Rice stripe virus

(RStV)

RNA RNA RNA RNA

1 2 3 4

NC_003755.1 NC_003754.1 NC_003776.1 NC_003753.1

8.97 3.51 2.50 2.16

Urochloa hoja blanca tenuivirus Urochloa hoja blanca virus

(UHBV)

RNA 3 RNA 4

NC_038757.1 NC_038758.1

2.27 1.86

Unassigned European wheat striate mosaic virus

(EWStMV)

RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA RNA

MN160329.1 MN160345.1 MN160360.1 MN160376.1 AJ969412.1a AJ969413.1a AJ969414.1a AJ969416.1a NC_040450.1 NC_040451.1 NC_040448.1 NC_040454.1 NC_040449.1 NC_040452.1 NC_040453.1 NC_040455.1

9.50 3.45 2.06 1.89 1.15 1.09 1.21 1.56 9.10 1.85 1.60 1.59 1.55 1.51 1.49 1.41

Maize yellow stripe virus

(MYStV)

Melon chlorotic spot virus

(MChSV)

1 2 3 4 1 2 3 5 1 2 3 4 5 6 7 8

a

Partial sequence.

Components The RNPs contain RNA coated with the NC protein of 34–35 kDa, and small amounts of the RdRp of 230–330 kDa. The RdRp associated with the RNPs of RStV and RHBV is capable of replicating and transcribing the RNA segments in vitro. The tenuivirus genome is composed of four to six non-capped ssRNA segments. The approximate size of the genome of the type member RStV is 17 kb (Fig. 3). The largest RNA segment (9.0 kb) of RStV, MStV, and RHBV is of negative polarity and encodes the RdRp. RNA2 (3.3–3.6 kb), RNA3 (2.2–2.5 kb) and RNA4 (1.9–2.2 kb) of RStV, MStV, IWSV, EHBV, UHBV, and RHBV are ambisense. RNA5 (1.3 kb), detected in virions of MStV and of EHBV is of negative polarity. A fifth RNA segment has also been reported for some

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Fig. 1 Phylogenetic relationships of tenuiviruses. Phylogenetic tree showing the relationships between tenuiviruses on the basis of the nucleotide sequences of the coding regions of RNAs 3 and 4 for all tenuiviruses except RGSV for which the coding regions of RNAs 5 and 6 were used. Shown is the percentage bootstrap support at each node. Echinochloa hoja blanca virus (EHBV), Iranian wheat stripe virus (IWSV), Maize stripe virus (MSpV), Rice grassy stunt virus (RGSV), Rice hoja blanca virus (RHBV), Rice stripe virus (RStV) and Urochloa hoja blanca virus (UHBV). The following sequences were used: EHBV (L48441, L75930), IWSV (AY312436, AY312435), MSpV-US (L13438, M57426), RGSV-PH (AB000404, AB000403), RGSV-CN (AF287949, AF290947), RHBV-CO (L14952, AF004658), RHBV-CR (AF004657, L07940), RSV-JP (D10979, X53563), RSV-CN (Y11096, Y11095) and UHBV (U82446, U82447). The MUSCLE multiple alignments and the phylogenetic tree were performed using Phylogeny. fr (http://www.phylogeny.fr/index.cgi), as described in Dereeper, A., Guignon, V., Blanc, G., et al., 2008. Phylogeny. fr: Robust phylogenetic analysis for the non-specialist. Nucleic Acids Research. 36 (Web Server issue), W465–W469.

Fig. 2 Structure of tenuivirions. (Left) Electron micrograph of sucrose density gradient purified RNP of Rice hoja blanca virus (RHBV) The bar represents 100 nm. A circular RNP is indicated by the arrow. (Right) Ultrathin section of an RHBV-infected rice leaf cell. The arrow indicates viral inclusion bodies inside a nucleus showing cytopathic effects. The bar represents 500 nm. Courtesy of Francisco Morales.

isolates of RStV. RGSV RNA1, 2, 5, and 6 are homologous to RNA1, 2, 3, and 4, respectively, of other tenuiviruses. RGSV RNA3 (3.1 kb) and RNA4 (2.9 kb) are ambisense and unique to RGSV.

Genome Organization The 50 and 30 terminal sequences (about 20 nt) are complementary to each other and can base-pair to give rise to circular RNPs. The terminal 8 nt (50 ACACAAAG) and their complement are conserved between tenuiviruses and viruses of the genus Phlebovirus. Several RNA segments encode two proteins in an ambisense arrangement (Fig. 3). The RStV gene products of each RNA segment are: - The RdRp (337 kDa) is encoded by the viral complementary vcRNA1, - The p2 (23 kDa) is present in the RNA2 and the pC2 (94 kDa) in the vcRNA2, - The p3 (24 kDa), also designated NS3, is encoded by the vRNA3. NS3 is a viral suppressor of RNA silencing (VSR) as demonstrated for RStV and for RHBV in both plant and insect hosts. The NC protein (35 kDa), also designated N or pC3, is encoded by the vcRNA3. The major non-structural protein NS4 (21 kDa), also designated p4 or disease-specific protein, is encoded by vRNA4. NS4 accumulates in infected plant tissues in large inclusion bodies. RStV NS4 facilitates tissue tropism in the insect vector. Finally, the pC4 (32 kDa), also designated movement protein (MP) or NSvc4, is encoded by the vcRNA4. The MP is implicated in the passage of viruses from cell-to-cell.

Replication and Transcription The RdRp synthesizes full-length vRNA and vcRNA that are not capped and sgRNAs of different sizes and of either polarity that serve as mRNA for the synthesis of the corresponding viral proteins. For MStV, RHBV, and RStV, it has been shown that

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Fig. 3 Schematic representation of the genome organization of RStV. v, viral sense RNA. vc, viral complementary RNA. RdRp, RNA-dependent RNA polymerase. NS3, non-structural protein 3 that acts as viral suppressor of RNA silencing (VSR). NC, nucleocapsid protein. NS4, nonstructural protein that accumulates in viral inclusion bodies in infected plant tissues; it is also designated disease-specific (DS) or non-capsid protein (NCP). MP, movement protein.

Fig. 4 Schematic representation of the replication, transcription, and translation strategies of the ambisense RNA segments of tenuiviruses. The cap and the non-viral nucleotides of host origin used as primers by the RdRp as a result of cap-snatching are represented at the 5 0 end of the mRNA. IR, intergenic region.

these mRNAs are synthesized via cap-snatching (Fig. 4). The 50 end of the mRNAs contains 10–17 non-viral nt and are capped; these extra sequences are derived from host cell mRNAs that are taken or snatched by the RdRp and are used as primers to initiate mRNA synthesis. Intergenic non-coding regions located between the ORFs can in certain cases adopt hairpin structures. The cap-snatching mechanism has been observed for mRNA synthesis of influenza viruses and viruses of the order Bunyavirales.

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Biological Properties Host Range and Geographic Distribution Plant hosts of tenuiviruses all belong to the family Poaceae. Members of the genus Tenuivirus cause severe disease in important food cereal crops such as rice and maize and their planthopper vectors themselves are also serious crops pests. Tenuiviruses-induced diseases are economically important resulting in serious yield losses. RStV was estimated in the 1960s to affect around 19% of the total Japan’s rice area. RHBV was reported to be responsible of 25%–75% yield losses in Latin American rice, and MSpV-associated losses in maize of 80% were reported from Mauritius. The geographic distribution of tenuiviruses is determined by the presence of their vector. RStV and RGSV are limited to the Far East, especially in China, Japan, and Korea; RHBV and EHBV to tropical and subtropical regions of the Western Hemisphere, Central and South America, the Caribbean, and the Southern United States; and MSpV has the most widespread geographic distribution around the world but only in subtropical and tropical regions.

Cytopathology In infected plants large inclusion bodies (Fig. 2 Right) are observed that contain the major non-structural protein NS4.

Disease Symptoms and Control Typical disease symptoms induced by tenuivirus on host plants are chlorotic stripes and yellow stipplings on the leaf blade between the veins, as well as plant dwarfing and panicle sterility. Young infected plants often exhibit complete chlorosis on the emerging whorl leaf and symptom severity and subsequent yield losses vary with age of the plant at the time of infection. RStV-infected rice leaves show yellow strips and dead tissue streaks. RHBV (Fig. 5 Top) and EHBV induce chlorotic leaf striping and mottling - called white leaf or hoja blanca -, plant dwarfing, panicle sterility, premature wilting, and necrosis, causing serious yield losses. MSpV-infected maize plants show early leaf chlorosis between the veins that later develop into continuous stripes, often with a “brush-out” appearance toward the tip of stripes. RStV-induced disease has been successfully controlled in Japan and China during the last few decades by using naturally resistant rice. RGSV causes significant economic losses in rice production in South, Southeast, and East Asian countries and

Fig. 5 Disease symptoms and insect vector. (Top Left) RHBV-infected chlorotic rice leaf with hoja blanca striping and mottling. (Top Right) RHBV-infected rice panicle. Healthy leaves and panicle are shown for comparison. (Bottom) Tagosodes orizicolus vector of RHBV.

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suitable resistance genes against RGSV have not yet been found in natural rice resources. A promising method to confer resistance against tenuiviruses is the use of RNA interference to target viral genes NC and MP that play important roles in viral infection and proliferation at an early stage of viral replication. Transgenic plants have been reported to show resistance against RStV and RGSV infection and inhibited the proliferation of the virus. In the 1980s, the identification of RHBV as the causal agent of the hoja blanca disease of rice together with genetic studies on the interaction of this virus with its planthopper vector, confirmed the pathogenic nature of the virus to its vector and helped to explain the cyclical nature of the epidemics. This disease was first controlled by hybridization of susceptible indica and resistant japonica rice genotypes and the adoption of integrated disease control practices. Recently, new RHBV-resistance sources belonging to the indica type have been identified; making it feasible to use these potential sources in crosses targeting the tropics.

Insect Vector and Type of Transmission Each virus species is persistently transmitted by a particular species of planthopper in a propagative manner and can be transmitted transovarially by viruliferous female planthoppers to their offspring, and through sperm from viruliferous males. The principal vectors of the species are Laodelphax striatellus for RStV, Tagosodes cubanus for EHBV, Peregrinus maidis for MStV, Nilaparvata lugens for RGSV, Tagosodes orizicolus for RHBV (Fig. 5 Bottom), Ukanoes tanasijevici for IWSV and Caenodelphax teapae for UHBV. Known vector of the tentative species is Javesella pellucida for EWStMV.

Relation to Other Taxa Viruses of the genus Tenuivirus share several properties with those of the genus Phlebovirus. The multi-segmented genomes of tenuiviruses and phleboviruses contain negative-sense and ambisense components. The presence of the highly conserved octanucleotide at 50 (50 -ACACAAAG) and 30 (CUUUGUGU-30 ) termini of their genomic segments, these terminal complementary sequences can base-pair and may give rise to circular RNPs. The synthesis of mRNAs by the viral RdRp follows a cap-snatching mechanism for viruses of the two genera. In addition, they encode functionally related proteins at equivalent genome positions: the tenuivirus RdRp and 94 kDa proteins are homologous to the RdRp and glycoproteins of other members of the family Phenuiviridae. Finally, their infection cycle includes arthropod transmission and the ability to replicate in the arthropod vector. The different number of genome components and the apparent lack of an enveloped viral particle distinguish tenuiviruses from other viruses of the family Phenuiviridae. Tenuiviruses are more complex in terms of the number of genome segments and generally encode more proteins than phleboviruses. However, differences in number of genome segments are not uncommon traits among genera of viruses belonging to the same family (e.g., Reoviridae, Closteroviridae, Secoviridae, Potyviridae, Rhabdoviridae), or even between members of the same genus (e.g., Emaravirus). Although enveloped tenuivirions have not been found, tenuiviruses do encode a glycoprotein closely related to those of phleboviruses (a similar situation can be found in the family Rhabdoviridae among members of the genus Dichorhavirus).

Further Reading de Miranda, J.R., Muñoz, M., Wu, R., Espinoza, A.M., 2001. Phylogenetic placement of a novel tenuivirus from the grass Urochloa plantaginea. Virus Genes 22, 329–333. de Miranda, J.R., Ramirez, B.C., Muñoz, M., et al., 1997. Comparison of Colombian and Costa Rican strains of Rice hoja blanca tenuivirus. Virus Genes 15, 191–193. Estabrook, E.M., Suyenaga, K., Tsai, J.H., Falk, B.W., 1996. Maize stripe tenuivirus RNA 2 transcripts in plant and insect hosts and analysis of pvc2, a protein similar to the phlebovirus virion membrane glycoproteins. Virus Genes 12, 239–247. Falk, B.W., Tsai, J.H., 1998. Biology and molecular biology of viruses in the genus tenuivirus. Annual Review of Phytopathology 36, 139–163. Huiet, L., Feldstein, P.A., Tsai, J.H., Falk, B.W., 1993. The Maize stripe virus major non-capsid protein messenger RNA transcripts contain heterogeneous leader sequences at their 50 termini. Virology 197, 808–812. Kormelink, R., Garcia, M.L., Goodin, M., Sasaya, T., Haenni, A.-L., 2011. Negative-strand RNA viruses: The plant-infecting counterparts. Virus Research 162, 184–202. Liu, X., Jin, J., Qiu, P., et al., 2018. Rice stripe tenuivirus has a greater tendency to use the prime-and-realign mechanism in transcription of genomic than in transcription of antigenomic template RNAs. Journal of Virology 92 (92), e01414–e01417. Nguyen, M., Ramirez, B.C., Golbach, R., Haenni, A.-L., 1997. Characterization of the in vitro activity of the RNA-dependent RNA polymerase associated with the ribonucleoproteins of Rice hoja blanca tenuivirus. Journal of Virology 71, 2621–2627. Ramirez, B.C., Garcin, D., Calvert, L.A., Kolakofsky, D., Haenni, A.-L., 1995. Capped non-viral sequences at the 50 end of the mRNA of Rice hoja blanca virus RNA4. Journal of Virology 69, 1951–1954. Ramirez, B.C., Haenni, A.-L., 1994. Molecular biology of tenuiviruses, A remarkable group of plant viruses. Journal of General Virology 75, 467–475. Ramirez, B.C., Lozano, I., Constantino, L.-M., Haenni, A.-L., Calvert, L.A., 1993. Complete nucleotide sequence and coding strategy of Rice hoja blanca virus RNA4. Journal of General Virology 74, 2463–2468. Ramirez, B.C., Macaya, G., Calvert, L.A., Haenni, A.-L., 1992. Rice hoja blanca virus genome characterization and expression in vitro. Journal of General Virology 73, 1457–1464. Shimizu, T., Toriyama, S., Takahashi, M., Akutsu, K., Yoneyama, K., 1996. Non-viral sequences at the 50 -termini of mRNAs derived from virus-sense and virus-complementary sequences of the ambisense RNA segments of Rice stripe tenuivirus. Journal of General Virology 77, 541–546. Toriyama, S., Kimishima, T., Takahashi, M., 1997. The proteins encoded by Rice grassy stunt virus RNA5 and RNA6 are only distantly related to the corresponding proteins of other members of the genus tenuivirus. Journal of General Virology 78, 2355–2363. Toriyama, S., Kimishima, T., Takahashi, M., et al., 1998. The complete nucleotide sequence of the Rice grassy stunt virus genome and genomic comparisons with viruses of the genus Tenuivirus. Journal of General Virology 79, 2051–2058.

Tobacco Mosaic Virus (Virgaviridae) Marc HV Van Regenmortel, University of Strasbourg, Strasbourg, France r 2021 Elsevier Ltd. All rights reserved.

Glossary Co-translational disassembly A biological phenomenon by which translation of the tobacco mosaic virus (TMV) occurs simultaneously as the disassembly of the TMV particle. Cryptotope A determinant (or epitope) of an immunological antigen or immunogen which is initially hidden and becomes functional only when the molecule is broken or degraded. Heterospecific antibodies Antibodies that are unable to react with the immunogen used to produce them but are capable to recognize immunogens from other species or mutants. Mapping of epitopes Epitope mapping is the process of experimentally identifying the binding sites, or “epitopes”, of antibodies on their target antigens. Movement protein A movement protein is a non-structural protein which is encoded by some plant viruses to enable their movement from one infected cell to neighboring cells.

Neotope Types of epitopes which depend on the protein quaternary structure and are absent in dissociated protein subunits. Segmental mobility in proteins The mobility of an antigenic determinant may make it easier to adjust to a pre-existing antibody site not fashioned to fit the exact geometry of a protein. Serological differentiation index Serological cross-reactivity between two viruses expressed as the number of two-fold dilutions steps separating homologous and heterologous titers. Viral mutants The term mutant is also applied to a virus with an alteration in its nucleotide sequence. Virus disassembly The disassembly of TMV particles is initiated when the end of the particles containing the 50 terminus of the RNA becomes associated with ribosomes. Virus self-assembly Biological phenomenon by which the different components of a virus particles are capable, under certain conditions, to auto-assemble to form complete and functional virus particles.

The Beginnings of Virology Tobacco mosaic virus occupies a unique place in the history of virology and was in the forefront of virus research since the end of the nineteenth century. It was the German Adolf Mayer, working in the Netherlands, who in 1882 first described a severe disease of tobacco which he called tobacco mosaic disease. He showed that the disease was infectious and could be transmitted to healthy tobacco plants by inoculation with capillary glass tubes containing sap from diseased plants. Although Mayer was not able to isolate a germ as the cause of the disease, he did not question the then prevailing view that all infectious diseases were caused by microbes and he remained convinced that he was dealing with a bacterial disease. About the same time, in St. Petersburg, Dmitri Ivanovsky was studying the same disease and he reported in 1892 that when sap from a diseased tobacco plant was passed through a bacteria-retaining Chamberland filter, the filtrate remained infectious and could be used to infect healthy tobacco plants. Ivanovsky was the first person to show that the agent causing the tobacco mosaic disease passed through a sterilizing filter and this gave rise to the subsequent characterization of viruses as filterable agents. A virology conference was held in 1992 in St. Petersburg to celebrate the centenary of this discovery. Although Ivanovsky is often considered one of the fathers of virology, the significance of his work for the development of virology remains somewhat controversial because all his publications show that he did not really grasp the significance of his filtration experiments. He remained convinced that he was dealing with either a small bacterium or with bacterial spores and never appreciated that he had discovered a new type of infectious agent. Following in the footsteps of Mayer, Beijerinck in Delft, Holland, showed in 1898 that sap from tobacco plants infected with the mosaic disease was still infectious after filtration through porcelain filters. He also demonstrated that the causative agent was able to diffuse through several millimeters of an agar gel and he concluded that the infection was not caused by a microbe. Beijerinck called the agent causing the tobacco mosaic disease a contagium vivum fluidum (a contagious living liquid), in opposition to a contagium vivum fixum. In those days the term contagium was used to refer to any contagious, disease-causing agent, while the term fixum meant that the agent was a solid particle or a cellular microbe. On the basis of his filtration and agar diffusion experiments, Beijerinck was convinced that the agent causing tobacco mosaic was neither a microbe nor a small particle or corpuscle (meaning a small body or particle from the Latin corpus for body). He proposed instead that the disease-causing agent, which he called a virus, was a living liquid containing a dissolved, non-particular and non-corpuscular entity. Lute Bos has claimed that Beijerinck's introduction in 1898 of the unorthodox and rather odd concept of a contagium vivum fluidum marked the historic moment when virology was conceived conceptually. However, it is clear that his definition of a virus as a soluble, living agent not consisting of particles certainly does not correspond to our modern view of what a virus is. It is Beijerinck's insistence that a virus is not a microbe and his willingness to challenge the then widely held view that all infectious diseases are caused by germs that make many people regard his contribution to the development of virology as more important than that of Ivanovsky. In 1998, a meeting held in the Netherlands commemorated the centenary of the work of Beijerinck on TMV. About the same time a meeting was held in Germany to honor the contribution of Loeffler and Frosch who, also in 1898, had shown that the agent causing the foot-and-mouth disease of cattle was able to pass through a Chamberland-type filter, in the

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same way as TMV. In addition the German workers also established that their disease-causing agent was not able to go through a finer grain Kitasato filter, from which they concluded that the agent, which was multiplying within the host, was corpuscular and not soluble as claimed by Beijerinck. Loeffler, however, continued to believe that his pathogen was a very small germ or spore, invisible in the light microscope, that was unable to cross the small pores of a Kitasato filter. There is no doubt that none of these ‘fathers’ of virology realized the nature of the filterable pathogenic agents they were investigating and that the term virus which they used did not have the meaning it has today. It took another 30 years before chemical analysis eventually revealed what viruses are actually made of protein and RNA.

Physical and Chemical Properties of TMV A major change in our perception of what viruses are occurred in 1935 when Stanley, working at the Rockefeller Institute in Princeton, obtained needle-like crystals of TMV that were infectious and consisted of protein. He used methods that were being developed at the time for purifying enzymes and was greatly helped by a bioassay developed by Holmes in 1929 that made it possible to quantify the amount of TMV present in plant sap. In this assay, extracts containing the virus are rubbed on the leaves of Nicotiana glutinosa tobacco plants which leads to a number of necrotic lesions proportional to the amount of virus present. Stanley's demonstration that TMV was a crystallizable chemical substance rather than a microorganism, a discovery that earned him the Nobel Prize in 1946 and had a profound impact on the thinking of biologists because it suggested that viruses were actually living molecules able to reproduce themselves. For a while, it seemed that viruses closed the gap between chemistry and biology and might even hold the key to the origin of life. Stanley had originally described TMV as a pure protein, but in 1936 Bawden and Pirie showed that the virus contained phosphorus and carbohydrate and actually consisted of 95% protein and 5% RNA. TMV thus became the first virus to be purified and shown to be a complex of protein and nucleic acid. Following the development of ultracentrifuges in the 1940s, ultracentrifugation became the standard method for purifying TMV and many other viruses. In 1939, TMV became the first virus to be visualized in the electron microscope and in 1941, TMV particles were shown to be rods about 280 nm long and 15 nm wide. Subsequently, X-ray analysis established that the rods were hollow tubes consisting of a helical array of 2130 identical protein subunits with 16(1/3) subunits per turn and containing one molecule of RNA deeply embedded in the protein subunits at a radius of 4 nm (Fig. 1). The length of the TMV particle is controlled by the length of the RNA molecule which becomes fully coated with protein and is thereby protected from nuclease attack. The viral protein subunits can also aggregate on their own, without incorporating RNA, to form rods very similar to TMV particles but in this case the rods are of variable length (Fig. 2). It is possible to degrade the virus particles with acetic acid or weak alkali and to obtain in this way dissociated protein subunits and viral RNA, the latter being rapidly degraded by nucleases. In 1956 Gierer and Schramm in Tübingen, Germany, and Fraenkel-Conrat in Berkeley, California, showed that if the viral RNA was obtained by degrading the particles with phenol or detergent, intact RNA molecules were obtained that were infectious and could

RNA

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Protein subunit

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Fig. 1 (a) Electron micrograph of negatively stained TMV particles. (b) Diagram of a TMV particle showing about one-sixth of the length of a complete particle. The protein subunits form a helical array with 16(1/3) units per turn and the RNA is packed at a radius of about 4 nm from the helix axis. From Klug, A., 1999. The tobacco mosaic virus particle: Structure and assembly. Philosophical Transactions of the Royal Society: Biological Sciences 354, 531–535.

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Single helix

Virus A-protein

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etc.

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High ionic strength Limited stacks of disks Stacked disk rod Irreversible Crystal

Fig. 2 Diagram showing the different polymorphic aggregates of TMV coat protein obtained at different pH values and ionic strengths. Note that both polar and nonpolar stacked disks can be obtained. The “lock-washer” is not well defined and represents a metastable transitory state. Adapted from Durham, A.C.H., Klug, A., 1972. Structures and roles of the polymorphic forms of tobacco mosaic virus protein. Journal of Molecular Biology 67, 315–332, with permission from Elsevier.

produce the same disease as intact virus. This was the first demonstration that the nucleic acid component of a virus was the carrier of viral infectivity and that it possessed the genetic information that coded for the viral coat protein. In 1960 the TMV coat protein was the first viral protein to have its primary structure elucidated when its sequence was determined simultaneously in Tübingen and in Berkeley and found to consist of 158 amino acids. Subsequently, the coat protein sequence of numerous TMV mutants obtained by treating the virus with the mutagenic agent nitrous acid was also determined and these sequence data helped to establish the validity of the genetic code that was being elucidated in the early 1960s in Nirenberg's and Ochoa's laboratories. One of the changes induced by the action of nitrous acid on viral RNA is the nucleotide conversion of cytosine (C) to uracil (U). This changes the codon CCC for the amino acid proline to the codons UCC or CUC for serine and leucine, respectively. In addition, these two codons can be further converted to UUU which codes for phenylalanine. When the coat protein sequences of various TMV mutants were determined, it was found that most of the exchanged amino acids could be attributed to C-U nucleotide conversions, which confirmed the validity and universality of the proposed genetic code.

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A particularly interesting mutation present in mutant Ni 1927 was the exchange proline to leucine at position 156 in the sequence which was found to greatly decrease the chemical stability of the virus. It had been known for many years that the virus was resistant to degradation by the enzyme carboxypeptidase and that this enzyme released only about 2000 threonine residues from each virus particle. The C-terminal residue in the coat protein is in fact threonine and the enzyme degradation data were interpreted at the time to mean that the virus particle contained 2000 subunits. However, when the proline at position 156 was replaced by leucine in the mutated protein, the enzyme was able to degrade the protein far beyond the C-terminal residue. The location of a proline near the C-terminus together with the presence of an acetyl group on the N-terminus are actually responsible for the remarkable resistance of the virus to exopeptidase degradation. The same exchange at position 156 in mutant Ni 1927 also led to an interesting discovery in immunology (see below).

Self-Assembly of TMV Particles Dissociated coat protein molecules of TMV are able to assemble into different types of aggregates depending on the pH and ionic strength (Fig. 2). The disk aggregate is a two-layer cylindrical structure with a sedimentation coefficient of 20S, where each layer consists of a ring of 17 subunits compared with the 16(1/3) molecules present in each turn of the assembled helix. The disks are able to form stacks which can be either polar or nonpolar, depending on the relative orientation of adjacent disks (Fig. 2). Short stacks of disks were shown to be nonpolar by immunoelectron microscopy, using monoclonal antibodies that reacted with only one end of the viral helix but were able to bind to both ends of the stacked disks. Whereas short polar stacks of disks can be transformed via lock-washer intermediates into helices (Fig. 2), this is not possible for nonpolar stacks which cannot be incorporated as such during the elongation process that leads to the formation of RNA-containing virus particles. It is generally accepted that initiation of TMV assembly from its RNA and protein components requires a 20S protein aggregate, which could be either a disk or a short, lock-washer type proto-helix. The surface of this aggregate constitutes a template which is recognized by a specific viral RNA sequence. The nucleation of the assembly reaction was found to occur by a rather complex process involving a hairpin structure on the viral RNA, known as the origin of assembly sequence (OAS) located about 1000 nt from the 30 end, which is inserted through the central hole of the 20 S aggregate. The nucleotide sequence of the stem–loop OAS hairpin is responsible for the preferential incorporation of viral RNA rather than foreign RNA. Elongation occurs by addition of more protein subunits which pull up more RNA through the central hole. This process requires that the 50 tail of the RNA loops back down the central hole and is slowly “swallowed up” while the 30 tail continues to protrude freely from the other end. This unexpected mechanism was visualized by electron micrographs obtained in Hirth's laboratory in Strasbourg which revealed how particles were growing with both 30 and 50 RNA tails protruding initially from the same end of the particle. The elongation is thus bidirectional and occurs faster along the longer 50 tail by incorporation of 20S aggregates and more slowly toward the 30 end through incorporation of 4S protein. The elucidation of the assembly process was greatly facilitated by the sequencing of the viral RNA which was completed in 1982. TMV RNA consists of 6395 nt and was the first genome of a plant virus to be sequenced completely.

Virus Disassembly Since TMV particles are extremely stable in vitro, it was not at all obvious how the RNA managed to be released from the particles in order to start the virus replication cycle. Using a cell-free translation system, Wilson discovered that the disassembly of TMV particles is initiated when the end of the particles containing the 50 terminus of the RNA becomes associated with ribosomes. This leads to viral subunits becoming dislodged from the particles while the 50 terminal open reading frame in the RNA is being translated by the ribosomes, a process known as co-translational disassembly. This mechanism allows the coat protein subunits to protect the RNA from enzymatic degradation until the particle has reached a site in the infected cell where translation can be initiated.

Antigenicity of TMV The antigenic properties of TMV have been studied extensively for more than 60 years and these studies have given us much information on how antibodies recognize proteins and viruses. TMV is an excellent immunogen and antibodies to the virus are readily obtained by immunization of experimental animals. When the sequence of TMV coat protein became available, it was possible for the first time to locate the antigenic sites or epitopes of a viral protein at the molecular level. Initial studies focused on two antigenic regions of the coat protein, the C-terminal region (residues 153–158) situated at the surface of virus particles and the disordered loop region (residues 103–112) located in the central hole of the particles which is accessible to antibodies only in dissociated protein subunits (Fig. 3). The C-terminal hexapeptide coupled to bovine serum albumin was used to raise antibodies and the resulting antiserum was found to precipitate the virus and neutralize its infectivity. Since both natural peptide fragments and synthetic peptides were used in this work, Anderer and his colleagues should be credited with the discovery that synthetic peptides can elicit antibodies that neutralize the infectivity of a virus. Only when similar results were obtained with animal viruses 15 years later, did the potential of peptides for developing synthetic vaccines become clear.

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112 90 34

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Fig. 3 Backbone of the TMV coat protein subunit based on crystallographic data. Residues 94–106 have been omitted because this region, located on the inside of the particle, is disordered. The N and C termini of the protein are located on the outer surface of TMV particles. The position of 7 continuous epitopes (residues 1–10, 34–39, 55–61, 62–68, 80–90, 108–112, and 153–158) is indicated by solid lines. Reproduced from Al Moudallal, Z., Briand, J.P., Van Regenmortel, M.H.V., 1985. A major part of the polypeptide chain of tobacco mosaic virus is antigenic. EMBO Journal 4, 1231–1235, with permission from Nature Publishing Group.

It has been known since the 1950s that intact TMV particles and dissociated coat protein subunits harbor different types of epitopes recognized by specific antibodies. Certain epitopes present only on virions are constituted by residues from neighboring subunits that are recognized as a single entity by certain antibodies; other epitopes arise from conformational changes in the protein that result from inter-subunit bonds. Both these types of epitopes which depend on the protein quaternary structure and are absent in dissociated protein subunits have been called neotopes. Another type of epitope known as cryptotope occurs on the portion of the protein surface that is buried in the polymerized rod and becomes accessible to antibodies only in the dissociated subunits. The mapping of neotopes and cryptotopes on the surface of TMV coat protein was greatly simplified once monoclonal antibodies (Mabs) became available. Many continuous epitopes were identified at the surface of dissociated coat protein subunits by measuring the ability of peptides to react with Mabs or with antisera (Fig. 3). These epitopes were found to correspond to regions of the protein shown by X-ray crystallography to possess a high segmental mobility. This correlation between antigenicity and mobility along the peptide chain was also found to exist in other proteins and was used to develop algorithms for predicting the location of certain epitopes in proteins from their primary structure. The surface of the protein subunits accessible at either end of the virus particle is different and the one located near the 50 terminus end of the RNA harbors the two helices corresponding to residues 73–89 and 115–135 (Fig. 3). Many Mabs specific for this surface have been obtained and in addition to reacting with both ends of nonpolar stacked disks, they were found to block TMV disassembly by sterically preventing the interaction between RNA and ribosomes. It has been suggested that if such antibodies could be expressed in plants, they might be able to control viral infection by preventing the disassembly of virions. Another interesting immunological phenomenon was discovered when TMV antibodies were analyzed for their ability to react with certain TMV mutants. It was found that all rabbits immunized with TMV induced the formation of heterospecific antibodies, that is, antibodies that were unable to react with the TMV immunogen but recognized the mutant Ni 1927 quite well. When all the antibodies in a TMV antiserum capable of reacting with TMV were removed by cross-absorption with the virus, it was found that the depleted antiserum still reacted strongly with this mutant which had a single proline-leucine exchange. Apparently the removal of this proline at position 156 exposes binding sites for both peptidases and antibodies that are normally out of reach in the wild-type protein structure. The induction of heterospecific antibodies by immunization with TMV is another illustration of the difference between the antigenicity and immunogenicity of proteins. TMV particles possess the immunogenic capacity of eliciting heterospecific antibodies that react with the Ni 1927 mutant although they do not have the antigenic capacity of reacting with these antibodies. The reverse situation where an antigenic peptide or protein is able to react with a particular antibody but is unable to induce the same type of antibody when used as immunogen is a more commonly observed phenomenon which greatly hampers the development of synthetic peptide vaccines.

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Taxonomy and Classification Many of the viruses that are currently classified in the genus Tobamovirus were initially considered to be strains of TMV on the basis of similar particle morphology and ability to cross-react with TMV antibodies. Antigenic relationships between different tobamoviruses have been quantified using a parameter known as the serological differentiation index (SDI) which is the average number of twofold dilution steps separating homologous from heterologous antiserum titers. A close correlation exists between the antigenic distance of two viruses expressed as SDI values and the degree of sequence difference in their coat proteins. It is often accepted that when two viruses differ antigenically by an SDI value larger than 4, they are considered to belong to separate species. This is valid only if relatedness is measured with polyclonal antisera containing antibodies to a range of different epitopes since comparisons made with Mabs specific for a single epitope will emphasize antigenic similarities or differences depending on which region of the protein is recognized by the particular Mab that is used. Relationships between individual tobamoviruses are nowadays usually assessed by comparisons between viral genomes. Phylogenetic studies have shown that tobamoviruses are very ancient and co-evolved with their angiosperm hosts, which means that they are at least 120 million years old. The Tobamovirus genus is one of seven recognized genera in the family Virgaviridae and it comprises 37 species.

Replication and Cell-to-Cell Movement of TMV TMV-infected tissues contain four viral proteins: the 126 kDa and 183 kDa proteins of the replicase complex, the 30 kDa movement protein, and the 17.6 kDa coat protein. TMV replication is initiated by the translation of the viral RNA to produce the replicase proteins and this leads to the synthesis of minus-sense and plus-sense copies of the RNA. Translation of the viral coat protein gene then occurs leading to the assembly of progeny virus particles from full-length genomic RNA and coat protein. The translation of the coat protein and movement protein is controlled by the production of two separate subgenomic mRNAs, a mechanism later found to occur in many other viruses but first demonstrated with TMV. The mechanism that allows plant viruses to move from cell to cell in their hosts has remained a mystery for many years. The rigid cellulose-rich walls of plant cells impede intercellular communication which occurs only through tubular connections known as plasmodesmata. However, plasmodesmata are too narrow to allow the passage of virus particles. Studies with the 30 kDa TMV movement protein showed that this protein changed the size exclusion limit of the plasmodesmata, allowing the virus to move through them in the form of a thin, less than 2 nm wide, ribonucleoprotein complex consisting of the genomic RNA and the movement protein. Movement proteins that possess RNA -binding properties have subsequently been found in many other plant viruses. Studies with TMV also revealed how plant viruses encode a suppressor to combat the post-transcriptional gene silencing reaction that plants use to fight virus infection.

Biotechnology Applications of TMV TMV-resistant plants have been obtained by transforming them with a DNA copy of the TMV coat protein gene. This coat proteinmediated resistance is due to the ability of transgenically expressed coat protein to interfere in transgenic cells with the disassembly of TMV particles. Coat protein-mediated resistance has subsequently been obtained with many positive-sense RNA plant viruses. A portion of the TMV RNA leader sequence, called omega, was shown to enhance the translation of foreign genes introduced into transgenic plants. This translational enhancer has been incorporated successfully in many gene vectors for a variety of applications. In another type of application, TMV particles have been used as surface carriers of foreign peptide epitopes for constructing recombinant vaccines and producing them in plants. It was found that the N- and C-termini of the TMV coat protein as well as the surface loop area corresponding to residues 59–65 were able to accept foreign peptide fusions without impairing the ability of the resulting chimeric virus to infect plants systemically. Several experimental vaccines against viral and parasitic infections that are based on genetically engineered TMV particles produced in tobacco are currently under evaluation. In conclusion, a series of contingent discoveries made at the beginning of the 20th century focused on TMV as a new type of pathogen which was extremely stable and obtainable in large quantities from infected plants, and this allowed this virus to play a major role in the development of virology. Furthermore, studies of TMV also contributed significantly to the advancement of molecular biology and to our understanding of the physicochemical and antigenic properties of macromolecules.

Bionanotechnology Applications of TMV Our considerable knowledge of the three-dimensional structure of TMV particles combined with our understanding of the mechanism of virion self- assembly gave rise to numerous applications in the new field of bionanotechnology. This has led, for instance, to the development of efficient vectors for virus-induced gene silencing in plants, the use of virus-derived sequences for the creation of virus-resistant lines of plants and to the fusion of peptides and various metals on the exterior surface of the virus

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for creating a variety of TMV -derived nanostructures. It is also possible to encapsidate any RNA molecule provided it contains the OAS of TMV which then leads to the protection of labile RNA against degradation. TMV -like particles can also been coupled to various organic molecules, including chromophores or polymers, in order to create classes of effective colloids that have potential biomedical applications inside living organisms. TMV-based nanorods of various sizes have been fabricated in vitro and some of them have been administered via the bloodstream for developing tumor-targeting agents. It is quite remarkable that a simple viral structure consisting of a single type of protein subunit together with a single molecule of RNA has led to such an enormous number of possible biotechnological applications. There is of course always another side to a coin, which in this case is the finding that the stability of TMV particles allows the virus to retain its infectivity for several years during the manufacturing process of cigarettes and cigars. TMV RNA has been shown to be present in the saliva of cigarette smokers (but not of non-smokers) and these smokers were also found to have a higher level of anti-TMV IgG antibodies in their blood than non-smokers. TMV antibodies were also found to cross react with the human mitochondrial protein TOMM40L which shares a six residue sequence (FGTGGA) with residues 36–41 of the TMV coat protein which is also a continuous epitope of the viral protein. It cannot be excluded that these antibodies resulting from an antigenic mimicry phenomenon between TOMM40L and TMV may play role in mitochondrial dysfunction implicated in the pathogenesis of Parkinson disease, Alzheimer disease and other degenerative disorders (Liu et al., 2013).

See also: Tobamoviruses (Virgaviridae). Virgaviruses (Virgaviridae)

Further Reading Al Moudallal, Z., Briand, J.P., Van Regenmortel, M.H.V., 1985. A major part of the polypeptide chain of tobacco mosaic virus is antigenic. EMBO Journal 4, 1231–1235. Balique, F., Colson, P., Raoult, D., 2012. Tobacco mosaic virus in cigarettes and saliva of smokers. Journal of Clinical Virology 55, 374. Bos, L., 1999. Beijerinck's work on tobacco mosaic virus: Historical context and legacy. Philosophical Transactions of the Royal Society, London, Series B 534, 675–685. Calisher, C.H., Horzinek, M.C., 1999. 100 Years of Virology. The Birth and Growth of a Discipline. Vienna: Springer, pp. 1-220. Durham, A.C.H., Klug, A., 1972. Structures and roles of the polymorphic forms of tobacco mosaic virus protein. Journal of Molecular Biology 67, 315–332. Harrison, B.D., Wilson, T.M.A., 1999. Tobacco mosaic virus: Pioneering research for a century. Philosophical Transactions of the Royal Society, London, Series B 354, 517–685. Hirth, L., Richards, K.E., 1981. Tobacco mosaic virus: Model for structure and function of a simple virus. Advances in Virus Research 26, 145–199. Liu, R., Vaishnav, R.A., Roberts, A.M., Friedland, R.P., 2013. Humans have antibodies against a plant virus: Evidence from tobacco mosaic virus. PLoS One 8 (4), e60621. doi:10.1371/journal.pone.0060621. Lomonossoff, G.P., 2018. So what have plant viruses ever done for virology and molecular biology? Advances in Virus Research 100, 145–162. Lomonossoff, G.P., Wege, C., 2018. TMV particles: The journey from fundamental studies to bionanotechnology applications. Advances in Virus Research 102, 149–176. Mahy, B.W.J., Lvov, D.K., 1993. Concepts in Virology: From Ivanovsky to the Present. Langhorne, PA: Harwood Academic Publishers. Scholthof, K.-B.G., 2004. Tobacco mosaic virus: A model system for plant biology. Annual Review of Phytopathology 42, 13–34. Scholthof, K.-B.G., Shaw, J.G., Zaitlin, M., 1999. Tobacco Mosaic Virus: One Hundred Years of Contributions to Virology. St. Paul, MN: American Phytopathologial Society Press, pp. 1–256. The life of a virus. In: Creager, A.N.H. (Ed.), Tobacco Mosaic Virus as an Experimental Model. Chicago, IL: University of Chicago Press. Van Helvoort, T., 1991. What is a virus? The case of tobacco mosaic disease. Studies in History and Philosophy of Science 22, 557–588. Van Regenmortel, M.H.V., 1999. The antigenicity of TMV. Philosophical Transactions of the Royal Society, London, Series B 354, 559–568. Van Regenmortel, M.H.V., Fraenkel-Conrat, H., 1986. The Plant Viruses: The Rod-Shaped Plant Viruses. vol. 2. New York, NY: Plenum, pp. 1–180.

Tobamoviruses (Virgaviridae) Ulrich Melcher, Oklaoma State University, Stillwater, OK, United States Dennis J Lewandowski, University of Florida, Lake Alfred, FL, United States William O Dawson, Citrus Research and Education Center, Lake Alfred, FL, United States and University of Florida, Lake Alfred, FL, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of D.J. Lewandowski, Tobamovirus, In Encyclopedia of Virology (Third Edition), Edited by Brian W.J. Mahy, Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00514-8.

Nomenclature

nt Nucleotide(s) OAS Origin of assembly ORF Open reading frame RdRp RNA-dependent RNA polymerase RT-PCR Reverse transcription-polymerase chain reaction satRNA Satellite RNA sgRNA Sub-genomic RNA siRNAs Small interfering RNAs TILLING Targeting induced local lesions in genomes tRNA Transfer RNA UTR Un-translated region

aa Amino acid(s) CP Coat protein or capsid protein ELISA Enzyme-linked immunological assays ER Endoplasmic reticulum kb Kilobase kDa Kilo dalton LAMP Loop mediated amplification MET Methyl-transferase miRNA MicroRNA MP Movement protein mRNA Messenger RNA

Glossary Origin of assembly Stem–loop structure that is the site of initiation of virion assembly. Protohelix Contain approximately 40 CP TMV subunits arranged in a spiral around a central hollow core, similar to the arrangement within the virion.

Pseudoknot RNA structure with base pairing between a loop and other regions of the RNA.

History of Tobamovirus Research Research in the late 1800s on the causal agent of the mosaic disease of tobacco led to the discovery of viruses as new infectious agents. Thus Tobacco mosaic virus (TMV), the type species of the genus Tobamovirus, became the first virus to be discovered, and subsequently has had a significant role in many fundamental discoveries in virology. The first quantitative biological assay for plant viruses was the use of Nicotiana glutinosa plants, which produce necrotic local lesions when inoculated with TMV or other tobamoviruses. The resistance gene, N, that confers this hypersensitive response-type resistance to TMV was the first resistance gene against a plant virus to be cloned and characterized. TMV was also the first virus to be purified and crystallized, which led to the discovery of the nucleoprotein nature of viruses and determination of the atomic structure of the coat protein (CP) and the virion. TMV was the first virus to be visualized in the electron microscope, confirming the predicted rigid rod-shaped virions. The genetic material of TMV was shown to be RNA, a property previously thought to be restricted to DNA. The first viral protein for which an aa sequence was determined was the TMV CP. TMV was also the first virus to be mutagenized. The subsequent determination of the CP sequences from a number of strains and mutants helped establish the universality of the genetic code. Methods of infecting plant protoplasts with viruses were developed with the tobacco–TMV system, creating a synchronous system to study events in the infection cycle. The TMV 30 kDa protein was the first viral protein shown to be required for virus movement. Finally, TMV was the first plant virus genome to be completely sequenced in 1975.

Taxonomy and Classification The genus Tobamovirus has been assigned to the family Virgaviridae. Currently, there are 38 recognized species within the genus (Table 1). Several recently sequenced viruses are tentative members of new species in the genus (Table 1). Although tobamoviruses comprise one of the more intensively studied plant virus genera, taxonomy has often been confusing. Historically, plant viruses with rigid virions of approximately 18
300 nm and causing various diseases were all designated strains of TMV. Thus, many viruses originally referred to

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Table 1

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Definitive and tentative species members of the genus Tobamovirus, family Virgaviridae

Species name

Acronym

Accession #

Length (nt)

Status

Natural host family

Bell pepper mottle virus Brugmansia mild mottle virus Cactus mild mottle virus Clitoria yellow mottle virus Cucumber fruit mottle mosaic virus Cucumber green mottle mosaic virus Cucumber mottle virus Frangipani mosaic virus Hibiscus latent Fort Pierce virus Hibiscus latent Singapore virus Kyuri green mottle mosaic virus Maracuja mosaic virus Obuda pepper virus Odontoglossum ringspot virus Opuntia virus 2 Paprika mild mottle virus Passion fruit mosaic virus Pepper mild mottle virus Plumeria mosaic virus Rattail cactus necrosis-associated virus Rehmannia mosaic virus Ribgrass mosaic virus Streptocarpus flower break virus Sunn-hemp mosaic virus Tobacco latent virus Tobacco mild green mosaic virus Tobacco mosaic virus Tomato brown rugose fruit virus Tomato mosaic virus Tomato mottle mosaic virus Tropical soda apple mosaic virus Turnip vein-clearing virus Ullucus mild mottle virus Wasabi mottle virus Yellow tailflower mild mottle virus Youcai mosaic virus Zucchini green mottle mosaic virus

BPMoV BrMMoV CMMoV ClYMoV CGMoMV CFMoMV CuMoV FrMV HLFPV HLSGV KGMoMV MaMV ObPV ORSV OV2 PaMMoV PfMV PeMMoV PlMV RCNaV ReMV RMV SFBV SHMV TLV TMGMV TMV ToBRFV ToMV ToMoMV TSAMV TuVCV UMMoV WMoV YTMMoV YMoV ZGMoV

NC009642 NC010944 NC011803 NC016519 NC002633 NC001801 NC008614 NC014546 NC025381 NC008310 NC003610 NC008716 NC003852 NC001728 NC040685 NC004106 NC015552 NC003630 NC026816 NC016442 NC009041 NC002792 NC008365 NC043383 NC038703 NC001556 NC001367 NC028478 NC002692 NC022230 NC030229 NC001873 ND NC003355 NC022801 NC004422 NC003878

6375 6381 6449 6514 6562 6424 6485 6643 6431 6485 6514 6794 6507 6618 6453 6524 6791 6357 6688 6506 6395 6311 6279 4683 1415 6355 6395 6393 6383 6398 6350 6311

Complete Complete Complete Complete Complete Complete Complete Complete Complete Complete Complete Complete Ccomplete Complete Complete Complete Complete Complete Complete Complete Complete Complete Complete Incomplete Incomplete Complete Complete Complete Complete Complete Complete Complete

6298 6379 6303 6513

Complete Complete Complete Complete

Solanaceae Solanaceae Cactaceae Fabaceae Cucurbitaceae Cucurbitaceae Cucurbitaceae Apocynaceae Malvaceae Malvaceae Cucurbitaceae Passifloraceae Solanaceae Orchidaceae Solanaceae Solanaceae Passifloraceae Solanaceae Apocynaceae Cactaceae Orobanchacee Plantaginaceae Gesneriaceae Leguminosae Solanaceae Solanaceae Solanaceae Solanaceae Solanaceae Solanaceae Solanaceae Brassicaceae Brassicaceae Brassicaceae Taxaceae Brassicaceae Cucurbitaceae

Tentative species Chara corallina Canada virus Chara corallina Australia virus Hoya chlorotic spot virus

CCCAV CCAUV HCSV

MK521928 AEJ33768 NC034509

9593 9065 6386

Complete complete complete

Characeae Characeae Apocynaceae

as TMV are now recognized as belonging to separate species. For example, the tobamovirus that was referred to as the tomato strain of TMV, and is approximately 80% identical to TMV at the nt sequence level, is now recognized as a different species, Tomato mosaic virus (ToMV). One major criterion for distinguishing the members of separate tobamovirus species is a nt sequence difference of at least 10%.

Evolution Both nt and aa sequences have been used to suggest evolutionary relationships of these viruses. Phylogenetic trees of the MP, CP, and RdRp replicase ORFs have similar topologies, indicating that this block has remained more or less constant as the viruses evolved. Additionally, the trees appear to identify clusters of viruses based on the family of the hosts from which they were identified. Comparison of phylogenetic trees for different coding regions of the genome suggest the clustering of tobamoviruses based on the host family. For example, viruses from solanaceous plants separate from those of cruciferous plant species in all coding regions. Similar congruence of phylogenetic positions is also seen for viruses whose principal hosts are in the Cucurbitaceae family. The observations suggest that the viruses were present in plant ancestors to both plant families. The observation that an orchid virus, ORSV, clustered with viruses of solanaceous plants in all but the methyl-transferase region of the replicase, where it branches with viruses from cruciferous plants, challenges that view. Such placement is suggestive that RNA–RNA recombination played a role in evolution of some tobamoviruses (Fig. 1).

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EF375551 RheMV V01408 TMV KT383474 TBRFV 100 AF332868 ToMV 100 86 KF477193 TMoMV 99 DQ355023 BPMV M81413 PMMoV 75 KU659022 TSAMV 100 99 AM398436 BrMMV KF495564 YTMMY 99 D13438 ObPV 100 100 AB089381 PaMMV 100 M34077 TMGMV AM040955 SFBV X82130 ORSV 99 HQ667979 RMV 100 U03387 TVCV 100 AB017503 WMoV 99 U30944 YoMV JN566124 CIYMV 100 U47034 SHMV AB917427 HLFPV 100 AF395898 HLSV DQ356949 MarMV 100 HQ389540 PFMV D12505 CGMMV AB261167 CMoV 100 AF321057 CFMMV AJ295948 KGMMV 100 100 AJ295949 ZGMMV EU043335 CMMoV JF729471 RCNaV 100 HM026454 FrMV 100 KJ395757 PluMV X14006 PEBV 91 100 L23972 PepRSV AF166084 TRV X99149 IPCV 100 X78602 PCV 99 J04342 BSMV Hordeivirus Goravirus AB976029 GORV AJ012005 CWMV 91 100 L07937 SBWMV AJ132576 SBCMV 100 100 AB033689 JSBWMV 100 AJ132578 OGSV 85 AB033691 SrCSV 100 D86636 BBNV Z97873 BSBV 87 AJ223596 BVQ 99 100 KT225277 SBV2 KT225271 CPSBV 70 AJ238607 PMTV 100

100

0.2

EF375551 RheMV V01408 TMV KT383474 TBRFV AF332868 ToMV 80 KF477193 TMoMV DQ355023 BPMV 73 X82130 ORSV 68 M81413 PMMoV KU659022 TSAMV 98 M34077 TMGMV KF495564 YTMMV AY137775 TLV 98 D13438 ObPV AB089381 PaMMV 97 71 AM398436 BrMMV AM040955 SFBV U03387 TVCV 88 HQ667979 RMV 99 AB017503 WMoV 97 U30944 YoMV HM026454 FrMV 100 KJ395757 PluMV JN566124 CIYMV 99 J02413 SHMV AB917427 HLFPV 63 AF395898 HLSV 98 D12505 CGMMV 71 AB261167 CMoV AJ295949 ZGMMV 82 AF321057 CFMMV 99 AJ295948 KGMMV 72 DQ356949 MarMV HQ389540 PFMV 100 EU043335 CMMoV JF729471 RCNaV 93 X15883 PEBV 70 92 X03241 PepRSV Z36974 TRV Goravirus KP760462 DrVA AF447397 IPCV 99 L07269 PCV 94 AB976030 GORSV Goravirus Z46351 LRSV X03854 BSMV M81486 PSLV 65 D00906 NVMV unclassified KT225272 CPSBV 97 AJ243719 PMTV 87 U64512 BSBV AJ223597 BVQ D86637 BBNV AB033692 SrCSV 92 AJ012006 CWMV 76 AJ132579 OGSV 94 L07938 SBWMV AB033690 JSBWMV 67 99 AJ132577 SBCMV 98

100 99

94

99

75

0.2

Fig. 1 Phylogenetic trees of tobamoviruses based on RdRP (left) and CP (right) gene sequences. The analyses were conducted in MEGA7 using the Maximum Likelihood method based on the Tamura-Nei model and 1000 bootstrap replicates. The tree with the highest log likelihood is shown. The percentage of trees in which the associated taxa clustered together is shown next to the branches (where 460%). The scale indicates the number of substitutions per site. All positions containing gaps and missing data were eliminated. Virus abbreviations are explained in the tables of member species and other viruses. Reproduced from: https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/w/virgaviridae.

The tobamovirus members are divided into those having the origin of assembly in the MP coding region (subgroup 1) and those for which the origin of assembly is within the CP coding region (subgroup 2). This division is strongly supported by phylogenetic analysis that shows the tree with all available sequences to consist of two major branches, one corresponding exactly to subgroup 1 and the other to subgroup 2 (Fig. 1). In these phylogenetic analyses, the closest neighbors that were not designated members of the tobamoviruses were a group of viruses typified by Beet necrotic yellow vein virus, thus defining the closest family of the Tombusviridae as the Benyvirus genus. The assembly of evolutionary relationships in phylogenetic trees allows some thoughts on the evolution of virus genera. The tobamoviruses have sequence neighbors in other genera of ssRNA viruses. They include genera Benyvirus (as mentioned above and consisting of plant viruses) and Orthohepevirus (which are animal viruses). Most closely related to the genus Tobamovirus are the charaviruses which have been isolated from the algal genus Chara (Fig. 2).

Virus Structure and Composition Tobamovirus virions are straight particles of approximately 18
300 nm with a central hollow core 4 nm in diameter (Fig. 3(A) and (B)). Virion composition is approximately 95% protein and 5% RNA. For TMV, approximately 2100 subunits of a single CP are arranged in a right-handed helix around a single genomic RNA molecule, with each subunit associated with three adjacent nucleotides. Protein–protein associations are the essential first event of virion assembly. Coat protein subunits assemble into several types of aggregates. Coat protein monomers and small heterogeneous aggregates of a few subunits are collectively referred to as ‘A-protein’. The equilibrium between A-protein and larger aggregates is primarily dependent upon pH and ionic strength. Purified CP and viral RNA can assemble into

Tobamoviruses (Virgaviridae)

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Fig. 2 Maximum-likelihood phylogeny of the amino acid sequences of the RdRp-2 regions of the replicase proteins of the charaviruses, benyviruses and selected tobamoviruses and relatives. Acronyms: ABV13, agaricus bisporus virus 13 (AQM49942); BastlV-VN, bastrovirus-like virus-VietNam Bat (YP009333174); BastV-Braz, bastrovirus Brazil/sewage (ASM79505); BMoV, burdock mottle virus (YP008219063); BNYVV, beet necrotic yellow vein virus (NP612615); BSbMV, beet soil borne mosaic virus (NP612601); CGMMV, cucumber green mottle mosaic virus (NP044577); CMMtV, cactus mild mottle virus (YP002455590); CTV, cutthroat trout piscihepevirus (YP004464917); CuMtV, cucumber mottle virus (YP908760); CV-Aus, charavirus australis (AEJ33768); CV-Can, charavirus canadensis (MK521928); HBlV1, hubei Beny-like virus 1 (APG77690); HHlV1, hubei hepe-like virus 1 (YP009336840); HLSV, hibiscus latent Singapore virus 178 (YP719997); HVlV16, hubei virga-like virus 16 (YP009336677); KGMMV, kyuri green mottle mosaic 179 virus (YP908760); MILV, mangifera indica latent virus (AMQ23297); OHV-A, orthohepevirus A (ABB88699, AGE83293, AGE83340, AGT38396, ANW09725, BAE86910); OHV-B, orthohepevirus B (AEX93357, CAQ16023, YP_009001465; OHV-C, orthohepevirus C (ADB96199, AFL69932, ANJ02843, BAO47898, BAT70058); OHV-D, orthohepevirus D (AIF74285, YP006576507); RCNaV, rattail cactus necrosis-associated virus (YP0044936166); RMV, ribgrass mosaic virus (YP005476600); RStNV, rice stripe necrosis virus (ABU94739); SanBV, san bernardo virus (AQM55436); TbTlV, tick borne tetravirus-like virus (AII01815); TMV, tobacco mosaic virus (NP597746). The green disks mark nodes with 40.9 SH support; two thirds of the nodes in the Orthohepevirus cluster have 40.9 SH support but, for clarity, are not marked.

infectious particles in vitro. Larger aggregates are disks composed of two individual stacked rings of CP subunits, and protohelices. Protohelices contain approximately 40 CP subunits arranged in a spiral around a central hollow core, similar to the arrangement within the virion. A sequence-specific stem–loop structure in the RNA, the origin of assembly (OAS), initiates encapsidation and prevents defective packaging that could result from multiple independent initiation events on a single RNA molecule. Virion assembly initiates as the primary loop of the OAS is threaded through a CP disk or protohelix with both ends of the RNA trailing from one side. The conformation of the CP protohelix changes as the RNA becomes embedded within the groove between the two layers of subunits. Elongation is bidirectional, proceeding rapidly toward the 50 end of the RNA as the RNA loop is extruded through the elongating virion and additional CP disks are added. There is disagreement about the mechanism of elongation toward the 30 terminus of the RNA, but it appears that this slower process involves the addition of smaller protein aggregates (Fig. 3(C)). Sub-genomic mRNAs containing the OAS are encapsidated into shorter particles that are not required for infectivity. The OAS is located within the open reading frame (ORF) for the movement protein (MP) of most tobamoviruses. The level of accumulation of a particular sub-genomic mRNA containing the OAS determines the relative proportion of that particular virion species. Thus, all tobamovirus virion populations contain a small percentage of MP sub-genomic mRNAs (sgRNA). In some tobamoviruses, including Cucumber green mottle mosaic virus (CGMMV), Hibiscus latent Singapore virus (HLSV), Kyuri green mottle mosaic virus (KGMoMV), Maracuja mosaic virus (MaMV), Sunn-hemp mosaic virus (SHMV), Zucchini green mottle mosaic virus (ZMoMV), and Cactus mild mottle virus (CMMoV), the OAS is located within the CP ORF. Thus, these so-called subgroup 2 tobamoviruses produce a significant proportion of small virions that contain CP sgRNA. Hybrid non-viral RNAs containing an OAS will also assemble with CP into virus-like particles of length proportional to that of the RNA.

Genome Organization The genomes of tobamoviruses consist of one single-stranded positive-sense 6.3–6.8 kb RNA (Fig. 4). A methyl-guanosine cap at the 50 terminus, precedes a 55–75 nt AU-rich leader. The 30 end of the RNA can be folded into a series of pseudoknot structures including a tRNA-like terminus. The hibiscus-infecting tobamoviruses Hibiscus Latent Singapore virus (HLSV) and Hibiscus latent Fort Pierce virus (HLFPV) contain, in addition, a poly(A) stretch between the 30 end of the CP ORF and the tRNA-like structure. The tRNA-like termini can be amino-acylated in vitro, in most cases specifically accepting histidine. The 30 terminus of SHMV is an

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Fig. 3 Structure of the Tobacco mosaic virus (Tobamovirus, Virgaviridae). (A) Model of three-dimensional structure of a hollow tube that is assembled from TMV CP subunits. (B) Electron micrograph of purified TMV virions showing tubular structures. (C) Diagram of proposed steps in the assembly of TMV virions from CP subunits and RNA. Scheme of the assembly stages of TMV nanoparticles from OAs‐containing RNA and CP subunits. Refer to the text for a thorough description. (a) Shows the successive assembly intermediates and the overall movement of the RNA scaffold upon its encapsidation. (b) Visualizes the organization of the final particle and its representation as applied in subsequent figures, with the RNA helix sandwiched between the helically arranged CP subunits. Images are not to scale. CP, coat protein; OAs, origin of assembly; TMV, tobacco mosaic virus. (A) Courtesy of J.Y. Sgro, VirusWorld, Institute for Molecular Virology, University of Wisconsin-Madison, USA. (B) Photograph of J.T. Finch reproduced from Fauquet, C.M., et al., 2005. VIIIth ICTV Report, Academic Press. (C) Adapted according to the Creative Commons Attribution 2.0 International Public License from Koch, C., Eber, F.J., Azucena, C., et al., 2016. Beilstein J. Nanotechnol. 7, 613–629. doi:10.3762/bjnano.7.54.

exception, accepting valine and appears to have arisen by a recombination event between a tobamovirus and a tymovirus. RNA extracted from infected plant material can be used to make transcription plasmids, whose transcription will produce transcriptable RNA molecules used to allow production of specific proteins in recipient plants.

Tobamoviruses (Virgaviridae)

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Fig. 4 Tobamovirus genome organization and gene expression strategy. (a) Tobamovirus genome organization. ORFs designated as open boxes. Non-translated sequences designated as lines; positions of sub-genomic promoters are marked. (b) Non-structural proteins involved in tobamovirus replication. Functional domains shared with other viruses within the ‘alphavirus supergroup’ are designated as hatched boxes. (c) Sub-genomic mRNAs with 50 proximal ORF labeled. MP, movement protein; CP, coat protein; MT, methyl-transferase; HEL, helicase; POL, polymerase; MPsg, MP sub-genomic mRNA; CPsg, CP sub-genomic mRNA.

The four ORFs that are contained within all tobamovirus genomes (Fig. 4) correspond to the proteins found in infected tissue. Two overlapping ORFs begin at the 50 proximal start codon. Termination at the first in-frame stop codon produces a 125–130 kDa protein. A 180–190 kDa protein is produced by readthrough of this leaky termination codon approximately 5–10% of the time. The remaining proteins are expressed from individual 30 co-terminal sub-genomic mRNAs, from which only the 50 proximal ORF is expressed (Fig. 4). The next ORF encodes the 28–34 kDa movement protein, which has RNA-binding activity and is required for cell-to-cell movement. The 30 proximal ORF encodes a 17–18 kDa CP. A sub-genomic mRNA containing an ORF for a 54 kDa protein that encompasses the readthrough domain of the 180–190 kDa ORF has been isolated from infected tissue, although no protein has been detected. Within the protein-coding regions of the genome, there are nt sequences that also function as cis-acting elements for sub-genomic mRNA synthesis, virion assembly, and replication. Gene expression from sub-genomic mRNAs is regulated both temporally and quantitatively. The MP is produced early and accumulates to low levels, whereas the CP is produced late and accumulates to high levels. The regulatory elements for sub-genomic mRNA synthesis are located on the genome-length complementary RNA overlapping the upstream ORF (Fig. 4). There is limited (40%) sequence identity between the TMV MP and CP sub-genomic promoters. The TMV MP sub-genomic promoter is located upstream of the MP ORF, flanking the transcription initiation site. Unlike the MP sub-genomic promoter, full activity of the CP sub-genomic promoter requires sequences within the CP ORF.

Viral Proteins The tobamovirus 125–130/180–190 kDa proteins are involved in viral replication, gene expression, and movement. Both are contained in crude replicase preparations, and temperature-sensitive replication-deficient mutants map to these ORFs. The 125–130/180–190 kDa proteins contain two functional domains common to replicase proteins of many positive-stranded RNA plant and animal viruses (Fig. 4). The N-terminal domain has methyl-transferase and guanylyl-transferase activities associated with capping of viral RNA. The second common domain is a proposed helicase, based upon conserved sequence motifs. The readthrough domain of the 180–190 kDa protein has sequence motifs characteristic of RNA-dependent RNA polymerases (RdRp). Both proteins are necessary for efficient replication, although the TMV 126 kDa protein is dispensable for replication and gene expression in protoplasts. The 125–130 kDa protein (or sequences within this region) of the 180–190 kDa protein are required for cell-to-cell movement. Additionally, these multifunctional proteins are symptom determinants, as mutations in mild strains map to these ORFs. The 28–34 kDa MP has a plasmodesmatal binding function associated with its C-terminus and a single-stranded nucleic acidbinding domain associated with the N-terminus. The MP–host interaction determines whether the virus can systemically infect some plant species. Although principally a structural protein, the CP is also involved in other host interactions. Coat protein is required for efficient long-distance movement of the virus. Coat protein is also a symptom determinant in some susceptible plant species and an elicitor of plant defense mechanisms in other plant species.

Interactions Between Viral and Host Proteins Available evidence suggests that the interactions of viral proteins with host factors are important determinants of viral movement and host ranges. Amino acid substitutions in the movement protein and 125–130/180–190 kDa proteins can alter the movement function in different hosts. Some viruses, including tobamoviruses, can assist movement of other viruses that are incapable of

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movement in a particular plant species. These interactions suggest that there are more precise associations of viral proteins with host factor(s) than with viral RNA. Additionally, precise CP–plant interactions are required for movement to distal positions within the plant. The helicase domain of the 130–190 kDa proteins elicits the N gene-mediated resistance in N. glutinosa.

Virus Replication Virions or free viral RNA will infect plants or protoplasts. Because tobamoviruses have a genome consisting of messenger-sense RNA that is infectious, one of the first events is translation of the 50 proximal ORFs to produce the proteins required for replication of the genomic RNA and transcription of sgRNAs. When virions are the infecting agent, the first event is thought to be co-translational disassembly, in which the CP subunits at the end of the virion surrounding the 50 end of the RNA loosen, making the RNA available for translation. Ribosomes then associate with the RNA, and translation of the 126/183 kDa ORFs is thought to displace CP subunits from the viral RNA. After the formation of an active replicase complex, a complementary negative-strand RNA is synthesized from the genomic positive-strand RNA template. Negative-strand RNA serves as template for both genomic and sgRNAs. Negative-strand RNA synthesis ceases early in infection, while positive-strand RNA synthesis continues. This results in an asymmetric positive- to negative-strand RNA ratio. Early in infection, genomic RNA functions as template for negative-strand RNA synthesis and as mRNA for production of the 126/183 kDa proteins. Later in the infection cycle, most of the newly synthesized genomic RNA is encapsidated into virions. Subgenomic mRNAs transcribed during infection function as mRNA for the 30 ORFs. Within cells of an infected leaf, replication proceeds rapidly between approximately 16 and 96 h post infection within a cell, then ceases. Even though the infected cells become packed with virions, these cells remain metabolically active for long periods. During the early stages of infection of an individual cell, the infection spreads through plasmodesmatal connections to adjacent cells. This event requires the viral MP that modifies plasmodesmata to accommodate larger molecules and the 126/183 kDa proteins. Movement through plasmodesmata does not require the CP. A second function of the MP appears to be binding to the viral RNA to assist its movement through the small plasmodesmatal openings. The MP also appears to associate with the cytoskeleton. As the virus spreads from cell to cell throughout a leaf, it enters the phloem for rapid long-distance movement to other leaves and organs of the plant, a complex process that requires the CP.

The Cis-Acting Sequences The 50 non-translated region contains sequences that are required for replication. This region is an efficient translational leader. The 30 non-translated region contains cis-acting sequences that are involved in replication. Certain deletions within the pseudoknots are not lethal, but result in reduced levels of replication. Exchange of 30 non-translated elements between cloned tobamovirus species has resulted in some lethal and nonlethal hybrids, suggesting a requirement for sequence specificity and/or secondary structure. The 30 non-translated region appears to be a translational enhancer, both in the viral genome and when fused to heterologous reporter mRNAs. Sequences encoding the internal ORFs for the MP and CP are dispensable for replication. Duplication of the sub-genomic promoters results in transcription of an additional new sub-genomic mRNA. Heterologous tobamovirus sub-genomic promoters inserted into the viral genome are recognized by the replicase complex and transcribed. Foreign sequences inserted behind tobamovirus sub-genomic mRNA promoters have been expressed to high levels in plants and protoplasts. Placement of the RNA sequence required for initiation of RNA encapsidation determines the size of the resulting virion accounting for the major differences in particle sizes within the tobamoviruses.

Satellite Tobacco Mosaic Virus Satellite tobacco mosaic virus (STMV), a tobamovirus-dependent satellite virus, was isolated from Nicotiana glauca plants infected with Tobacco mild green mosaic virus (TMGMV). The STMV genome consists of one single-stranded positive-sense RNA of 1059 nt. The 240 30 nt share approximately 65% sequence identity with TMGMV and TMV, contain pseudoknot structures, and have a tRNAlike terminus. No sequence similarity with any tobamovirus exists over the remainder of the genome. Two overlapping ORFs that are expressed in in vitro translation reactions are present in the genomic RNA of most STMV isolates. The 50 proximal ORF encodes a 6.8 kDa protein that has not been detected in vivo and is not present in all STMV isolates. The second ORF encodes a 17.5 kDa CP that is not serologically related to any tobamovirus CP. The 17 nm icosahedral virions are composed of a single STMV genomic RNA encapsidated within 60 STMV CP subunits. Replication of natural populations of STMV is supported by other tobamoviruses, but at lower levels than with the natural helper virus, TMGMV. The host range of STMV parallels that of the helper virus.

Host Range and Symptomology Tobamoviruses have a fairly large host range. All-in-all, dicotyledonous and monocotyledonous have been identified as hosts for the members of the 36 tobamovirus species and the 3 tentative members. TMV alone is known to infect members of nine plant

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Fig. 5 Symptoms of Tobacco mosaic virus on tobacco White Burley (A. left) (Courtesy of Dominique Blancart, INRA, France) on tomato leaves (B. top right) and fruits (C. bottom right) (Courtesy T.A. Zitter, Vegetablemdonline, Cornell University, Ithaca, NY).

families, and at least 125 individual species, including tobacco, tomato, pepper (many members of the family Solanaceae), cucumbers, and a number of ornamental flowers, including orchids. Many tobamoviruses infect plants producing a variety of virus-specific leaf pigment patterns, described by terms such as mosaics, mottles and yellows, depending on the virus tested. The first symptom of this virus disease is a light green coloration between the veins of young leaves. This is followed quickly by the development of a mosaic symptom of light and dark green areas on the leaves. “Rugosity” (crinkling of leaves) is observed in several cases of plant infection by tobamoviruses, as have other leaf deformations. These symptoms are more pronounced on younger leaves. TMV infection does not result in plant death, but if infection occurs early in the season, plants could be stunted. On tobacco, lower older leaves could be subject to "mosaic burn" especially during periods of hot and dry weather. In these cases, large dead areas develop in the leaves. This constitutes one of the most destructive phases of TMV infection. However, if TMV infects crops like grape and apple, it is almost symptomless. TMV over the last century has been known to cause a relatively stable production loss for tobacco of up to two percent per year in North Carolina, which is financially important for the tobacco industry. TMV and ToMV are worldwide considered as major diseases for vegetable production (Fig. 5) and cause tremendous losses in many solanaceous crops, especially in greenhouses where a lot of movement and passages of workers happen.

Transmission TMV and related tobamoviruses are transmissible to new plants only by contact of a plant with an inoculum. Therefore plants may become infected by rub inoculation of virions containing substances like Insect, animals, and humans visiting the fields. Plant infection begins when healthy plants contact virus contaminated sources. Particularly effective are rubbing with an aqueous suspension of virions. Animals moving through a field of infectious plants can, by brushing, pick up virions and transfer them to new host plants. Insects feeding on infected plants can also transmit tobamoviruses into their next host. Bee hives from an infected field can initiate an infection in another field. It is not clear whether the hive contains infectious particles and must itself be considered as the vector or not. TMV is known as one of the most stable viruses. It has a very wide survival range. As long as the surrounding temperature remains below approximately 401C, TMV can maintain its infectious capacity and all it needs is a host to infect. Greenhouses and botanical gardens provide the most favorable condition for TMV spreading, due to the high population density of possible hosts, the number of workers going through the plants and the constant temperature throughout the year. One of the common and useful control methods for TMV is sanitation, which includes removing infected plants and washing hands and tools in between each planting. Crop rotation is also employed to avoid infected soil/seed beds for some time. As for any plant disease, looking for resistant varieties against TMV has been done for tobacco, tomato and other plants. Furthermore, the cross protection method has been used, whereby a more severe strain of TMV is inhibited by previously infecting the host plant with a mild strain of TMV. This method has been used for decades in the vegetable industry in Europe. Breeders have selected genes of resistance in tobacco, tomato and other crops, and those genes have been used in the industry. Precautionary measures taken in the use of tools and other objects has also been proven to decrease TMV spread in crops.

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Further Reading Fraile, A., Garcia-Arenal, F., 2013. Tobamoviruses as models for the study of virus evolution. Advances in Virus Research 102, 117. Gibbs, A.J., 1977. Tobamovirus Group. CMI/AAB Descriptions of Plant Viruses, No. 184. Available at: http://www.dpvweb.net/dpv/showadpv.php?Dpvno184. Gibbs, A.J., Torronen, M., Mackenzie, A.M., et al., 2011. The enigmatic genome of Chara australis virus. Journal of General Virology 92, 2679. Hull, R., 2002. Matthews' Plant Virology, fourth ed. New York: Academic Press. Ishibashi, K., Ishikawa, M., 2016. Replication of tobamovirus RNA. Annual Review of Phytopathology 54, 55–78. Lewandowski, D.J., 2005. Tobamovirus genus. In: Fauquet, C.M., Mayo, M.A., Maniloff, J., Desselburger, U., Ball, L.A. (Eds.), Virus Taxonomy, Classification and Nomenclature of Viruses: Eighth Report of the International Committee on Taxonomy of Viruses. San Diego, CA: Elsevier Academic Press, pp. 1009–1014. Malpica-López, N., Rajeswaran, R., Beknazariants, D., et al., 2018. Revisiting the roles of tobamovirus replicase complex proteins in viral replication and silencing suppression. Molecular Plant-Microbe Interactions 31, 125. Pogue, G.P., Lindbo, J.A., Garger, S.J., Fitzmaurice, W.P., 2002. Making an ally from an enemy: Plant virology and the new agriculture. Annual Review of Phytopathology 40, 45–74. Scholthof, K.B., 2005. Tobacco mosaic virus: A model system for plant virology. Annual Review of Phytopathology 42, 13–34. Scholthof, K.B., Shaw, J.G., Zaitlin, M., 1999. Tobacco Mosaic Virus: One Hundred Years of Contributions to Virology. St. Paul, MN: APS Press. van Regenmortel, M.H.V., Fraenkel-Conrat, H., 1986. The Plant Viruses: The Rod-Shaped Plant Viruses. 2. New York: Plenum. Vlok, M., Gibbs, A.J., Suttle, C.A., 2019. Metagenomes of a freshwater Charavirus from British Columbia provide a window into ancient lineages of viruses. Viruses 11, 299.

Tobraviruses (Virgaviridae) Stuart A MacFarlane, The James Hutton Institute, Invergowrie, United Kingdom r 2021 Elsevier Ltd. All rights reserved. This is an update of S.A. MacFarlane, Tobravirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00515-X.

Glossary Agroinfiltration Infiltration into air spaces within the plant leaves of cultured Agrobacterium tumefaciens bacteria leading to (usually) transient expression of functional genes or virus infectious clones in plants by transcription from a binary plasmid carried within the bacteria. NM infection Infection of some plant species (primarily potato) by only the RNA1 molecule of TRV.

Initially erroneously thought to be “non-multiplying” hence NM. Spraing Symptoms of dark flecking and arc-formation in tubers of some potato cultivars resulting from TRV infection. Systemic infection Movement of virus from the point of inoculation to more distantly located leaves or roots via the plant vascular system (phloem).

Taxonomy and Characteristics The genus Tobravirus is comprised of three species, the type species Tobacco rattle virus (TRV) together with Pea early-browning virus (PEBV) and Pepper ringspot virus (PepRSV), which was initially referred to as the CAM strain of TRV. The genus has been assigned to the Virgaviridae family. Tobraviruses have a genome of two, positive-sense, single-strand RNAs that are packaged separately into rod-shaped particles. In some situations, the larger genomic RNA (RNA1) can cause a systemic infection in the absence of the second, smaller RNA (RNA2) and without the formation of virus particles. Tobraviruses are transmitted between plants by rootfeeding nematodes of the genera Trichodorus and Paratrichodorus, and in some plant species are also seed transmitted.

Virus Particle Production and Structure Tobravirus RNA1 is encapsidated into the L (long) particle with a length of between 180 and 215 nm, depending on virus species, and RNA2 is encapsidated into S (short) particles which range in length from 46 to 115 nm, depending on virus isolate (Fig. 1). Both L and S particles have an apparent diameter of 20–23 nm, depending on the technique used to examine them. In vitro translation experiments using RNA extracted from purified virus preparations, as well as studies with the TRV SYM isolate, which has an unusual genome structure, showed that some, if not all, tobravirus sub-genomic (sg)RNAs are also encapsidated, in particles of various lengths. The tobraviruses encode a single coat protein (CP), molecules of which assemble in a helical arrangement around a central cavity with a diameter of 4–5 nm, and with a distance of 2.5 nm between successive turns of the helix. In vitro reconstitution experiments suggested that virus particle formation initiates at the 50 end of the viral RNA, although the encapsidated sgRNAs do not carry the 50 terminal part of the virus genomic RNAs. No specific “origin-of-assembly” sequence has been identified for any of the different tobravirus RNAs. Peptide mapping showed that the major antigenic regions of the CP are, in descending order of strength, the C-terminal 20 amino acids (aa), 5 aa in the central region of the protein and 5 aa at the N-terminus. This and other spectroscopic analysis suggest that the N- and C-termini are exposed on the outer surface of the particle, while the central region is exposed in the central canal (where interactions of the CP with the viral RNA take place). The C-terminal domain appears to be unstructured and plays a role in interactions with other virus proteins that are involved in nematode transmission of the virus. Particularly with TRV, there is significant amino acid sequence difference between the CP of different virus isolates, resulting in many different serotypes of this virus.

M-Type and NM-Type Infections In early studies with tobraviruses (usually with TRV) the infectivity of fractionated and purified L and S particles was examined, showing that L particles were infectious (producing local and systemic symptoms on particular hosts) but that S particles were not. However, plants infected with L particles did not contain virus particles, whereas, plants infected with both L and S particles did contain (both) virus particles. This was explained by sequencing, which revealed that RNA1 encodes proteins for RNA replication and movement, whereas, RNA2 encodes the CP. Because infections with RNA1 do not produce virus particles they were very difficult to maintain by repeat inoculation of sap extracts, and were referred to as non-multiplying (NM-type) infections. Infections derived from both RNA1 and RNA2 and producing virus particles were easily passaged and were referred to as multiplying (M-type) infections. Subsequently it was found that extraction with phenol of RNA from NM-infected plants in fact produced highly infectious preparations.

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Fig. 1 Electron micrograph of long and short particles of TRV isolate RH. Bar is 100 nm.

Early observations also suggested that NM-type infections caused more severe symptoms than M-type infections and moved only slowly up the plant, giving rise to the theory that un-encapsidated RNA1 could spread systemically only from cell to cell via plasmodesmata. However, experiments with TRV and PEBV carrying defined mutations in the CP gene demonstrated unequivocally that un-encapsidated virus RNAs moved very rapidly via the vascular system in both Nicotiana benthamiana and N. clevelandii. In addition, with wild type (encapsidated) virus systemic infection always included both RNA1 and RNA2 but with the (un-encapsidated) CP mutants RNA2 occasionally became separated from RNA1 and was not detected in some systemic leaves. One PEBV mutant carried only a small (28 aa) deletion at the C-terminus of the CP but did not produce virus particles or indeed any detectable CP in infected plants. Nevertheless, this mutant moved systemically as quickly as did wild type virus and in this case RNA2 did not become separated from RNA1. This suggests, perhaps, that some form of CP, which can be present at very low levels but is not incorporated into particles, enables the co-ordinated transport of RNA1 and RNA2 through the phloem. In another study, plants were infected with TRV RNA1 and two types of RNA2, one wild type and the second encoding a CP in which the 15 aa at the C-terminus were replaced with 3 non-viral residues. Both RNA2 species were encapsidated by the CP that they encoded but neither appeared to be encapsidated by the CP from the other RNA2. Apparently, although RNA1 is encapsidated in trans, RNA2 is only encapsidated in cis. The mechanism for this is not known but may be linked to the role of CP in the co-ordinated systemic movement of RNA1 and RNA2.

Genome Structure and Expression The viral RNAs have a 50 methylated guanosine cap. The 30 end of the RNAs is not polyadenylated but folds into a tRNA-like pseudoknot structure that, in contrast to some other virus RNAs, cannot be aminoacylated. RNA1 is from 6.8 to 7 kb in size, and RNA2 varies between isolates from 1.8 to 4.2 kb (Fig. 2). By February 2019 GenBank contained complete sequences for 17 TRV RNA1 and 36 TRV RNA2 molecules. However, for both PEBV and PepRSV only a single RNA1 and three RNA2 sequences were reported.

RNA Sequences The larger RNA (RNA1) is highly conserved in nucleotide sequence between different isolates of the same virus (e.g., 99% identity between RNA1 of TRV isolates SYM and ORY) but there is much less identity between isolates of the different tobravirus species (e.g., 62% identity between RNA1 of TRV isolate SYM and PEBV isolate SP5). The smaller RNA (RNA2) varies considerably both in terms of overall nucleotide sequence identity as well as protein coding capacity between isolates of the same virus, although phylogenetic analysis shows that the coat proteins (CPs) of the different tobraviruses, which is encoded by RNA2, are more closely related to each other than to CPs from viruses in other genera. The tobraviruses are most closely related to tobamoviruses, in terms of particle structure, overall gene organization and viral protein sequence homologies. Depending on the specific virus and isolate, RNA1 has a 50 non-coding region (NCR) of between 126 and 202 nt, and a 30 NCR of 459 to 255 nt. RNA2 has a much larger 50 non-coding region (NCR) of between 470 and 710 nt, and a 30 NCR of 780 to 392 nt. There is almost no sequence homology between the 50 NCRs of the tobraviruses, however, the 25 nt at the 30 terminus of the TRV

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Fig. 2 Genome diagram and expression strategy of TRV isolates PpK20 and SYM. Open boxes denote virus genes. The solid lines above the RNA1 genes show the location of the two, overlapping replicase genes. The 134K protein contains methyltransferase (MT) and helicase (HEL) motifs. The C-terminal part of the 194K protein contains an RNA-dependent RNA polymerase (RDRP) motif. The asterisk denotes the 134K gene termination codon where readthrough translation to produce the 194K protein occurs. The movement protein (MP) gene is also known as the 29K or 1a gene. The cysteine-rich 16K gene is also known as the 1b gene. For RNA2, CP denotes the coat protein gene. N1 to N3 denote novel open reading frames present in isolate SYM RNA2. 2bD is a truncated 2b gene. The dashed lines beneath each RNA denote the 30 co-terminal sub-genomic RNAs that are known or suspected (?) to exist.

and PEBV RNAs are identical and there is 70% identity over the next 140 nt. Consequently, when recombinants were made between the RNA2 of TRV PpK20 and PEBV SP5, the RNA1-encoded replicase complex of both viruses could replicate RNA2 molecules carrying the 30 NCR from either virus. However, RNA2 carrying the TRV 50 NCR could only be replicated by a TRV RNA1, and RNA2 carrying the PEBV 50 NCR could only be replicated by a PEBV RNA1.

Expression of Virus Genes The 50 proximal gene of RNA1, encoding viral replicase proteins, is expressed by direct translation of the genomic RNA. The two other RNA1-encoded genes (1a/29K and 1b/16K) are expressed from sgRNAs that start with the sequence AUA (within a conserved motif GCAUA) and that are co-terminal with the 30 end of the genomic RNA. The 50 proximal gene of RNA2 of almost all tobravirus isolates encodes the CP. However, the CP gene is located at least 470 nt downstream of the 50 terminus of RNA2, and this region contains numerous (at least six) AUG codons upstream of the translation initiation codon of the CP gene, that are inhibitory to CP expression. Experiments to delete parts of the 50 NCR of PEBV RNA2 showed that only a very small amount of CP is translated in vitro from full-length (genomic) RNA2 but that translation is increased 25-fold when the 50 NCR is removed. Thus, even though the CP gene is usually the 50 proximal gene on RNA2, it (and the other genes on RNA2), is expressed from a sgRNA. However, the CP gene of TRV SYM is located near to the 30 end of RNA2, downstream of other genes. The sgRNA for this CP is much shorter than genomic RNA2 and is encapsidated into a clearly identifiable VS (very short) virus particle. A stem-loop sequence is found upstream of tobravirus sgRNA start sites and is particularly conserved for the CP sgRNA. The introduction of mutations into this structure (8 bp stem, 4 nt loop) upstream of the PEBV CP gene showed that the stem-loop forms an essential part of the sgRNA promoter, and that the structure of the stem rather than its actual sequence is important for sgRNA synthesis in planta. The promoter regions for the TRV and PEBV CP genes could be used as a cassette and moved to other locations in RNA2 to express non-viral genes.

Recombination and Sequence Deletions in Tobravirus RNAs The large variation in the overall length of RNA2 from different tobravirus isolates reflects the fact that most are recombinant molecules, where the 30 part of RNA2 has been replaced by sequences from the 30 part of RNA1. The recombination junction can occur at any position in RNA2 that is downstream of the CP gene, and the recombinant RNA2 molecule may retain none, some or even both of the other genes (2b and 2c) that might be considered RNA2-specific. The region of RNA1 that is transferred to RNA2 always includes the 30 NCR, often includes the 1b/16K/12K gene immediately upstream of the 30 NCR and may occasionally include part of the 1a/29K gene that is located upstream of the 1b gene. This means that many of these recombinant isolates carry two, probably both functional, copies of the 1b gene, one on RNA1 and the second on RNA2. In one study, a single Alstroemeria plant was found to contain seven different recombinant TRV RNA2 molecules, three with PEBV RNA1-like 30 NCR sequences and four with different TRV RNA1 30 coding and NCR sequences. The mechanism for such recombination is not known but is speculated to be caused by template switching by viral replicase from RNA1 to RNA2 during minus strand RNA synthesis. Sequences that resemble the 50 terminus of tobravirus genomic RNAs (rich in A and U residues) are often found at or near the recombination junction, and conceivably could facilitate the recombination event. It is also not completely clear when tobravirus recombination occurs, though the feeling is that repeated passage of virus by mechanical inoculation in glasshouse-grown plants, bypassing the nematode

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transmission process, either encourages recombination or reduces selection against the survival of recombinants. Nevertheless, at least one known recombinant isolate, TRV PaY4, which was cloned after only a very limited period of multiplication in the glasshouse, did retain its ability to be nematode transmitted. When a TRV-infected potato plant was propagated by bud cuttings, a small (80 bp) deletion in RNA2 was detected only 5 days after planting, and by 33 days post planting an RNA2 was found lacking nearly 50% of its original sequence. By 55 days post planting no RNA2 could be detected in the plant, even though an RNA1 infection was maintained. A different recombination process also occurs in which the CP and/or 2b gene in RNA2 of some TRV isolates appears to have been derived from PEBV. For this reason, serological analysis is not always successful in discriminating between the different tobraviruses. Attempts to reproduce this recombination by co-inoculating TRV and PEBV to plants in the glasshouse were not successful, suggesting that this may be only a rare occurrence, or may be stimulated by particular environmental conditions. Nevertheless, one report showed that 30% of the TRV isolates recovered from fields in the coastal bulb growing region in the Netherlands were TRV/PEBV recombinants. For PepRSV RNA2, the three sequenced molecules all have RNA1-derived sequences at their 30 end and do not retain any 2b or 2c sequences. However, it is assumed in nature that this virus is nematode transmitted and, thus, that isolates carrying the 2b and 2c genes must exist.

Viral Proteins RNA1 RNA1 encodes proteins for virus RNA replication and intra-plant movement (local and systemic). The 5 0 half of RNA1 encodes a large (134–141 kDa) protein that contains methyltransferase and helicase domains and is expected to be part of the viral replicase complex. Readthrough translation of the stop codon of this protein produces a larger protein (194–201 kDa) that contains motifs associated with RNA-dependent RNA polymerase proteins. In vitro translation experiments showed that the opal (UGA) translation termination codon of the TRV 134 kDa protein is suppressed by tRNAs that incorporate tryptophan or cysteine at this position. Downstream of the replicase genes is the 1a gene encoding a 29–30 kDa cell-to-cell movement protein (MP) belonging to the “30K” superfamily of plant virus movement proteins. Disruption of this gene prevents accumulation of TRV in inoculated leaves but can be overcome by co-inoculation with tobacco mosaic virus (TMV) or in transgenic plants expressing the TMV 30K movement protein. The C-terminal 1b gene encodes a 12 kDa (PEBV; PepRSV) or 16 kDa (TRV) cysteine-rich protein that is involved in seed transmission of PEBV in pea, and pathogenicity of TRV and PEBV. The TRV 16K protein is a suppressor of RNA silencing that is predominantly cytoplasmic but can also be found in the nucleus and nucleolus. The nuclear/nucleolar localization is important for the interaction of 16K with the plant protein coilin and the modulation of host plant anti-viral defense pathways. The TRV 16K protein is required to enable the virus to transiently enter the plant meristem. In N. benthamiana plants, a TRV 16K mutant virus does not enter the meristem, causes more severe infection symptoms and persists in the plant, whereas, with wild type TRV the plant recovers as the infection subsides and symptoms disappear. One study reported that the TRV 29K protein could also suppress RNA silencing when expressed during virus replication but not when expressed transiently via agroinfiltration. The PepRSV 12K protein also is a silencing suppressor and was found to enable potato virus X (PVX) to infect Arabidopsis when co-inoculated with PepRSV.

RNA2 RNA2 encodes proteins for virus particle formation and transmission by nematodes. Generally, the CP gene is the most 50 proximal gene on RNA2, even though it is expressed from a sgRNA rather than from the full-length, genomic RNA. As mentioned above TRV SYM has an unusual gene organization, with three potential genes located upstream of the CP gene. Several other TRV isolates from potato have also been identified in which the CP gene is preceded on RNA2 by novel genes that have no known function. Several studies have identified the CP as being involved in the nematode transmission process, and deletion of part of the C-terminal domain of the CP prevented the transmission of PEBV without affecting virus particle formation. Infectious, nematode-transmissible clones of three TRV isolates and one PEBV isolate have been constructed. In addition to the CP, these RNAs all carry two other genes encoding the 2b and 2c proteins, both of which (for PEBV) have been detected by western blotting in extracts of virus-infected plants. Mutation studies showed that the 2b gene is necessary for transmission of TRV and PEBV, whereas the 2c protein was only required for transmission of PEBV. This difference may reflect the different species of nematode that transmitted the particular virus isolates used in these studies, rather than a clear mechanistic difference between TRV and PEBV transmission. A further, small open reading frame, encoding a putative 9 kDa protein immediately downstream of and in-frame with the CP gene, is present in PEBV-TpA56 and TRV TpO1. For PEBV, mutation of this gene greatly reduced nematode transmission frequency, although the protein was not detected by western blotting. The 2b proteins of the different tobravirus isolates share some amino acid sequence homology with each other, ranging from only 11% identity (TRV Umt1 v. TRV PaY4) to 99% identity (TRV Umt1 v. TRV OR2). They also range in size from 238 aa

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(TRV PaY4) to 354 aa (TRV PpK20). The TRV PaY4 2b protein has been shown to act in trans, with nematode transmission of a 2b-mutant virus being complemented by co-infection with wild type virus. However, this appears to be isolate specific, as the TRV PaY4 2b mutant was complemented only by wild type TRV PaY4 and not by wild type TRV PpK20. The 2b protein may influence nematode transmission by more than one mechanism. In a microscopy study of the roots of N. benthamiana plants infected by PEBV, large numbers of particles of the wild type virus were found in all regions of the root tip, which is where the vector nematodes preferentially feed, whereas virus particles of a mutant deleted for the 2b and 2c genes were present in roots only in much lower numbers. A similar enhancement was found in the efficiency of invasion of roots in N. benthamiana, as well as in leaves and roots of Arabidopsis thaliana, when TRV carrying the 2b gene was compared with TRV lacking the 2b gene. Chemical analysis studies indicate that TRV infection increases the production of specific root volatiles and that TRV-infected plants are more attractive to vector nematodes. Furthermore, it seems that the TRV 2b gene plays some, as yet unclear, role in this process. The 2b protein also physically interacts with the TRV CP (as examined by yeast two-hybrid (Y2H) analysis) and the virus particle (as examined by immuno-electron microscopy). One suggestion is that the 2b protein acts as a bridge to trap virus particles to specific sites of retention on the nematode esophageal cuticle. Aggregates of virus particles have been observed using the electron microscope to collect in this part of the nematode but the co-location of the 2b protein has not yet been demonstrated. It appears that interaction between the CP and 2b protein stabilizes the 2b protein, as in plants infected with wild type TRV PaY4, carrying both PaY4 CP and PaY42b genes, the 2b protein was detected by western blotting. However, with a recombinant TRV carrying the TRV PpK20 CP gene and the TRV PaY4 2b gene, the 2b protein could not be detected, possibly because these proteins from two different isolates cannot interact with one another. Although the 2c protein is involved in transmission of PEBV, very little else is known about this protein. Amino acid sequence homologies between the 2c proteins of different TRV isolates range from almost none to over 95% identity. In a Y2H assay the TRV PpK20 2c protein interacted with the PpK20CP, and removal of the CP C-terminal flexible domain did not affect the interaction.

Tobraviruses as Gene Expression/Silencing Vectors As RNA1 encodes all the proteins necessary for tobravirus replication and movement, the RNA2 can be modified to carry non-viral sequences without greatly affecting virus infection. Expression vectors have been constructed from all three tobraviruses, in which a duplicate CP promoter sequence is inserted downstream of the native CP gene followed by restriction sites to allow the cloning of other sequences. Together, the tobraviruses can infect a wide range of plant species, often without causing particularly severe symptoms, features which increase their utility as expression vectors. Plants infected with tobraviruses often undergo a rapid recovery in which infection symptoms and virus levels fall dramatically, although the plants do not become free of virus, most particularly in the meristem regions. However, these plants have developed a strong resistance to further infection by the same virus, most likely by an RNA silencing-based mechanism. Although not well understood, it seems that tobraviruses are potent triggers of RNA silencing but do not encode a strong silencing suppressor protein to counteract this host defense activity. A consequence of this is that when a host plant sequence is inserted into the virus genome, very strong silencing is initiated that targets expression of the host gene itself, a process known as virus-induced gene silencing (VIGS). TRV has become one of the most widely used VIGS vectors for studies of plant gene function in many different plant species, and PEBV is also a very effective VIGS vector for studies in pea.

Diseases Caused by Tobraviruses TRV is found in many countries (Europe, North America, Japan, and Brazil) and has a particularly wide host range, infecting more than 100 species in nature and more than 400 species when tested in the glasshouse, although not all of these infections are systemic. As tobraviruses are transmitted by soil-inhabiting nematodes, in the field infection may be limited to the roots. Weeds may play an important role in the maintenance and spread of tobraviruses, with Capsella bursa-pastoris, Senecio vulgaris, Stellaria media, and Viola arvensis being the most commonly found (in the UK) weed hosts of TRV. Virus transmission in seed of these plants was also reported. Many crop plants are infected by TRV, the major diseases being those of potato and ornamental bulbs (narcissus, gladiolus, tulip, lily, and crocus). The symptoms of TRV infection in potato are the formation of arcs and flecks of brown corky tissue in the tuber which is referred to as spraing, and which can make the tuber unfit for sale. Both M-type and NM-type infections can produce spraing symptoms, which have been shown to be a hypersensitive (defense) response by the plant to the virus. It was thought that potato cultivars that did not show spraing symptoms were resistant to TRV, however, some infections can be symptomless in particular cultivars. Nevertheless, even these symptomless infections lead to significant reductions in tuber yield and tuber quality. Inclusion of tubers carrying symptomless infection could have major consequences for the production and distribution of seed potatoes (Fig. 3).

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Fig. 3 Cross-section of a potato tuber showing spraing symptoms caused by TRV infection.

PEBV has been reported in several European countries (UK, Netherlands, Italy, Belgium, Sweden) as well as Algeria and Morocco. In the field it infects mainly legumes, including pea, faba bean, French bean, lupin, and alfalfa. Several weeds and other crop plants may be infected, although often only in the roots. Early studies referred to Broad bean yellow band virus (BBYBV) as a defined virus infecting various legumes. This is now known to be a distinct serotype of PEBV. PepRSV has only been reported from Brazil, where it infects pepper, tomato, and artichoke, as well as local weed species.

Further Reading Harrison, B.D., Robinson, D.J., 1978. The Tobraviruses. Advances in Virus Research 23, 25–77. MacFarlane, S.A., 1999. Molecular biology of the Tobraviruses. Journal of General Virology 80, 2799–2807. MacFarlane, S.A., 2010. Tobraviruses – Plant pathogens and tools for biotechnology. Molecular Plant Pathology 11, 577–583. MacFarlane, S.A., Robinson, D.J.R., 2004. Transmission of plant viruses by nematodes. In: Gillespie, S.H., Smith, G.L., Osbourne, A. (Eds.), Proceedings of the SGM Symposium 63 on Microbe-Vector Interactions in Vector-Borne Diseases. Cambridge University Press.

Tomato Leaf Curl New Delhi Virus (Geminiviridae) Supriya Chakraborty and Manish Kumar, Jawaharlal Nehru University, New Delhi, India r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) AGO Argonaute protein amiRNA Artificial microRNA CAS CRISPR-associated protein CP Coat protein or capsid protein CRISPR Clustered regularly interspaced short palindromic repeats dsRNA Double-stranded ribonucleic acid ERF4 Ethylene response factor-4 HR Hypersensitive response kb Kilobases kDa Kilo Daltons LAMP Loop mediated amplification LTP Lipid transfer protein MEAM1 Middle-East Asia minor I MED1 Mediterranean 1 miRNA microRNA MP Movement protein

Glossary Agro-inoculation A method enabling the transfer of DNA (such as cloned begomoviral molecules) into a host plant using Agrobacterium tumefaciens. Rolling circle amplification A technique that allows amplification of circular DNA molecules (such as geminivirus genomes) with f29 DNA polymerase and random primers under isothermal conditions.

NASH Nucleic acids spot hybridization NBS-LRR Nucleotide-binding site–leucine-rich repeat nm Nanometer(s) NSP Nuclear shuttle protein nt Nucleotide(s) NtRDR1 RNA-Dependent RNA Polymerase 1 from N. tabacum PCR Polymerase chain reaction PTGS Post-transcriptional gene silencing RDR6 RNA-dependent RNA polymerase 6 SCR Satellite conserved region SGS3 Suppressor of gene silencing 3 siRNA Small interfering RNA SRAP Sequence-related amplified polymorphism sRNA Small RNA ssDNA Single-stranded deoxyribonucleic acid TGS Transcriptional gene silencing TrAP Transcriptional activator protein VSR Viral suppressor of RNA silencing

Tolerance Defined as an interaction in which viruses accumulate to some degree without causing significant loss of vigor or fitness to their hosts. Viral titer detection Radiolabelled nucleic acid molecules (begomoviral DNA) detection by using virus gene-specific labeled probe through Southern blot experiments. Viruliferous whitefly A whitefly insect-vector population carrying an infectious begomoviral molecule.

Introduction Tomato (Solanum lycopersicum) belongs to the family Solanaceae and is one of the most important vegetable crops grown in India. India is the second-largest tomato producing country in the world. Tomato is originating from America, probably in Mexico, and the center of diversification was in South America, probably in Peru. Tomato leaf curl New Delhi virus (ToLCNDV) is a bipartite begomovirus species (genus Begomovirus, family Geminiviridae) and is a well-known plant DNA virus responsible for the disease called Tomato leaf curl disease (ToLCD). The epidemic might be linked to the increased population of viruliferous whitefly (Bemisia tabaci), the insect vector accountable for the spread of the pathogen of ToLCD, around the world including India. The first reported incidence of this disease was from northern India in 1948. The occurrence was also documented from central India during the 1950s, followed by disease incidence in tomato-growing regions of South India. The disease has since begun to emerge as a major threat to the production of the tomato crop in India. The total yield loss in tomato production by begomoviruses ranges from 18% to 100%. Factors such as age of the plant, viruliferous whitefly populations, aggressivity of the virus isolate and susceptibility of the tomato variety influence the production and may cause up to 100% yield loss. Tomato plants affected by the leaf curl disease exhibit typical symptoms of upward and downward leaf curling, leaf wrinkling, leaf blistering, vein clearing, distortion of leaves, and stunted plant growth that might result in a complete yield loss (Fig. 1). Tomato leaf curl New Delhi virus (ToLCNDV) has been recorded in recent decades to cause devastating damage to tomato yield and other vegetable crop productions in the Indian agriculture sector, notably the disease in the northern part of India is more widespread. Using infected tissue samples from two separate locations, ToLCNDV was first sequenced and characterized in India in 1992. Until recently, the epidemic of ToLCNDV was confined to South-East Asian countries like India, Bangladesh, Thailand, Indonesia, Sri-Lanka and Pakistan. Recent reports are indicative of its presence in other countries such as Spain, Morocco, Italy, and

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Fig. 1 Plants infected with ToLCNDV displaying typical yellow mosaic symptoms on leaves (A) tomato, (B) chilli, (C) egg plant (D) pumpkin, (E) zucchini plants, (F) An infected zucchini plants at the field. Zucchini plants pictures courtesy of E. Moriones, Institute of Subtropical and Mediterranean Horticulture La Mayora, Algarrobo-Costa, Málaga, Spain. Picture of pumpkin crop courtesy of B. Mandal, Indian Agricultural Research Institute, New Delhi, India and picture of eggplant reproduced from the research article by Pratap, D., Kashikar, A.R., Mukherjee, S.K., 2011. Molecular characterization and infectivity of a Tomato leaf curl New Delhi virus variant associated with newly emerging yellow mosaic disease of eggplant in India. Virology Journal 8 (1), 305.

Israel. However, ToLCNDV infectivity studies have not been performed yet on tomato, or in any other solanaceous or cucurbitaceous crops, in other countries and therefore the virus may be more widespread than anticipated.

Tomato Leaf Curl Disease and Virus Biodiversity on Tomato in the Indian Subcontinent Members of the Geminiviridae family infect various monocots and dicots plants across the continents causing severe losses. These insect-transmissible geminiviruses are encapsulated in 2.7–5.2 kb twinned icosahedral particles with one or two (mono or bipartite) circular, non-enveloped single-stranded DNA (ssDNA) genomes. The members of the family Geminiviridae are further classified into nine genera such as Becurtovirus, Begomovirus, Capulavirus, Curtovirus, Eragrovirus, Grablovirus, Mastrevirus, Topocuvirus, and Turncurtovirus based on their genome organization, nucleotide sequence identities, and mode of insect transmission. Furthermore, Begomovirus, the largest genus of the family, comprises approximately 450 species of geminiviruses that infect many economically important crops, ornamental plants as well as weed plants, around the world, and they are all transmitted by whiteflies (Bemisia tabaci Genn). As per the current International Committee on Virus Taxonomy (ICTV) recommendation and revision of taxonomy from the Xth ICTV Report, based on pairwise nucleotide alignment of the begomovirus genome sequences, the name of ToLCNDV covers in fact four geminivirus species. These four virus species are Tomato leaf curl New Delhi virus (ToLCNDV DNA-A and DNA-B, U15015 and U15017), Tomato leaf curl New Delhi virus 2 (ToLCVNDV2, JQ897969), Tomato leaf curl New Delhi virus 4 (ToLCVNDV4, KF551592) and Tomato leaf curl New Delhi virus 5 (ToLCVNDV5, EF450316). The taxonomic classification also depends upon other factors such as relatedness with the major or minor parental sequences, the frequency and extent of recombination and the biological properties of the viruses. More than 280 ToLCNDV’s complete full-length sequences have been reported till date at NCBI GenBank public database and out of that number at least 80 of them are reported from India infecting different crop plants but tomato. This virus seems to be very successful at invading space and new host niches, in the Indian subcontinent and also in several regions of the world. This information suggests that the emergence of ToLCNDV, which is already underway, could be the major agricultural threat for vegetable crops to India and other countries in the very near future. Other begomovirus species such as Tomato leaf curl Gujarat virus, Tomato leaf curl Bangalore virus, Tomato leaf curl Bangladesh virus, Tomato leaf curl Pune virus, Chilli leaf curl virus, Chilli leaf curl India virus, Pepper leaf curl Lahore virus, Ageratum enation virus, and Tobacco curly shoot virus are also known to be associated with ToLCD, affecting the tomato production in the Indian subcontinent. The fact that ToLCD remains yet to be fully characterized even after 70 years of its first report, might be due to the diversity of the viruses causing the disease, broader host range, and high level of host-virus adaptability.

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The full-length DNA sequence of an Indian isolate of ToLCNDV DNA-A (U15015) and DNA-B (U15017) was first elucidated in the early 1990s from a begomovirus infected tomato (Solanum lycopersicum) from New Delhi (hence the name of the virus species). The size of the ToLCNDV-[India: New Delhi: Severe:1992] DNA-A and DNA-B are 2739 and 2696 bp, respectively. Similarly, ToLCNDV2-[India: IANDS1:2011] (JQ897969) isolated from tomato in New Delhi, is 2735 bp, ToLCNDV4-[India: Junagadh: TC306:2011] (KF551592)isolated from tomato in Gujarat is 2739 bp and ToLCNDV5-[Bengladesh: cucumber:2006] (EF450316) isolated from Cucumis sativus (cucumber) infected leaves, from Bangladeshis 2738 bp. Although all of the above-mentioned ToLCNDV species are nearly similar in size, they share sequence-similarities of less than 90% and have been assigned so far into the four separate ToLCNDV species listed above.

Phylogeny and Molecular Diversity Based on phylogeny, genome organization, and geographical distribution, begomoviruses are divided into two super groups namely the Old-World (OW) and New-World (NW) begomoviruses. Among the Old World viruses more than 57 begomovirus species have been identified on tomato and there are many other begomoviruses capable of infecting tomato as well. In the Indian subcontinent alone more than 17 species of begomoviruses have been identified from tomato. However, among all these viruses ToLCNDV has emerged in the recent years as the major virus infecting tomato in the subcontinent, as already mentioned four species bear the name of ToLCNDV and as of 2019, there was a total of 282 records at NCBI under the name of ToLCNDV. In other words there is a lot of begomovirus variability at large associated with the general syndrome of tomato leaf curl disease (ToLCD) in the world and in India particularly, but ToLCNDV has prevailed in the recent years on tomato and other crops in the Indian subcontinent. When a detailed study comparison of all the 282 ToLCNDV isolates is performed, it is obvious that there are no strictly defined demarcation strains among all these isolates, with a few exception (see below) indicating that there have been a lot of recombination events among the individuals of this population. The pairwise multiple sequence alignment of all records bearing the name of ToLCNDV spans a great variability between 72% and 100%. Officially, four viruses, ToLCNDV (U15015), ToLCNDV2 (JQ897969), ToLCNDV4 (KF551592) and ToLCNDV5 (EF450316), have been identified by their sequence as representatives of four species. The original reference of ToLCNDV (U15015) scores 74% identity with ToLCNDV2, 87% with ToLCNDV5 and 89% with ToLCNDV4, however ToLCNDV2 is sharing 94% identity with the members of the species PeaLDV (identified since ToLCNDV2 was sequenced) indicating that ToLCNDV2 is an isolate of this species now called PeaLDV. For simplicity and clarity it is therefore proposed to drop the name ToLCNDV2. ToLCNDV5 is borderline and ToLCNDV4 is even more border line for species demarcation in the ToLCNDV population and it is believed it is best for the time being to consider them as unassigned isolates of the species ToLCNDV. In addition another isolate from Capsicum annuum, from New Delhi, is showing less than 75% identity with the original ToLCNDV reference (U15015), and 91% with members of the species ToLCNDV making it an isolate of that species. Finally a last isolate from a weed from Pakistan named ToLCNDV-[PK: Chiniot:ZI27: G11V4:12] (HG937523), is in fact 94–98% identical to PaLCV isolates and should be renamed as PaLCV–[PK: Chiniot:ZI27: G11V4:12]. Within the ToLCNDV cluster of the remaining 276 isolates spanning pairwise identities between 86% and 100%, one can recognize several sub-clusters that we recorded for simplicity from Strain A to P plus an additional 13 unassigned isolates that could each become a node for new strains when more samples will be sequenced. Among these 15 Strains some are very well defined with no overlap with others like the strain M and G for the Thai-Indonesian isolates, the strain C for the Spain-Italy isolates, or some Indian-Pakistan strains such as Strains L, A, B. The strains D, E, F, J, K, L and H are more overlapped and consequently more difficult to define. The original first isolate of ToLCNDV (U15015) belongs to the Strain H and it is the largest cluster within the ToLCNDV population with 86 isolates (Fig. 2). Apparently for all the isolates collected so far there is some level of correlation with the geographical distribution for Europe and South East Asia, a correlation with the hosts has not been found. The same virus can infect a range of different host in the same region. Recently it is of interest to note that ToLCNDV has been identified on cotton (Gossypium hirsutum) in Pakistan along with the betasatellite Cotton leaf curl Multan betasatellite. These 8 cotton isolates cluster in the middle of the Strain H which is the cluster of the original ToLCNDV isolate (U15015).

Genome Organization and Function of Gene Products Four families of ssDNA viruses (Circoviridae, Parvoviridae, Spiraviridae and Geminiviridae) infect a range of hosts such as plants, hyper-thermophilic archaebacteria, vertebrates (mammals, birds, reptiles, fish) and invertebrates (arthropods, insects, crustacean, echinoderms). Among these four families, only the members of the family Geminiviridae are known to infect plants and the members are further categorized into nine genera and among them, only the genus Begomovirus contains either mono or bipartite genome. ToLCNDV DNA-A comprises six open reading frames (ORFs), two in virion or sense strand (AV1/CP and AV2/Pre-coat) and four in complementary or antisense strand (AC1/Rep, AC2/Trap, AC3/Ren, and AC4). AC1 encodes a multifunctional and most conserved protein, Rep or replication-associated protein that initiates the viral replication. AC2 encodes Trap or transcriptionalactivator protein and is involved in the modulation of transcriptional gene silencing (TGS) as well as in the post-transcriptional

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Fig. 2 Maximum-likelihood dendrogram of ToLCNDV isolates, available at the NCBI GenBank database. Accession numbers for all these sequences are mentioned in the phylogenetic tree. The nucleotide sequences are aligned using Muscle algorithm with bootstrap values of 1000 replicates. The scale bar indicates the nucleotide substitution rate per site, estimated by the Jukes-Cantor model. The branches of the individuals belonging to the same cluster/strain have been collapsed for simplification, The orange color indicate isolates from India-Pakistan, dark blue indicates isolates from Iran, Green from Thailand, Indonesia, pink from Bengladesh, pale blue from Spain-Italy-Morocco. The Unassigned isolates are written in black. The putative other ToLCNDV species are written in red and their corrected name in dark blue (other species) or black (unassigned ToLCNDV isolates).

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gene silencing (PTGS) pathways. The viral DNA replication is facilitated by another early transcriptional gene, AC3 or REn or replication enhancer protein. AC4 is the least conserved ORF and structurally overlapped with the AC1 ORF. The AC4 functions as an RNA-silencing suppressor and also the suppressor of PTGS pathway. AC4 encoded protein of ToLCNDV interacts with tomato AGO4. Non-functional mutant of ToLCNDV AC4 remained however infectious on solanaceous crops. The AV1 ORF encodes coat protein (CP) which is a highly conserved protein facilitating the assembly and packaging of the viral genome. ToLCNDVAV2 encoded pre-coat protein acts as host silencing suppressor and pathogenicity determinant. In nature a severe and a mild strain of ToLCNDV (U15015 and U15016, respectively) isolated from tomato in India, the viruses were cloned and sequenced, and infectious clones were produced. It was demonstrated on Nicotiana benthamiana and S. lycopersicum plants, that the severe symptom was associated with the higher capacity of the Rep of the severe isolate to replicate components A and B. The infectivity data suggested that the asparagine at the 10th position (Asn10) of the Rep protein (AC1) of the severe strain specifically recognizes the 3rd base of the conserved iteron sequences although both severe and mild strain shares 94% pairwise nt identity on their Rep genes. It is therefore possible that in nature such mutation may be able to trans-replicate more molecules that could benefit the ToLCNDV DNA-A and DNA-B to impact its host range and ultimately its epidemiology. Bipartite begomoviruses encode two additional ORFs (BV1 and BC1) in their DNA-B component. BV1 ORF encodes nuclear shuttle protein (NSP) in the virion or sense strand and BC1 encodes movement protein (MP) in the complementary or antisense strand of the viral genome. According to recent studies, transient overexpression of ToLCNDV-NSP induces the hypersensitive response (HR) or necrosis in N. tabacum plant. Similarly, ToLCNDV-BV1 N-or C-terminal mutant failed to show HR in S. lycopersicum. It is therefore postulated that the NSP may act as a symptom determinant and could serve as a silencing suppressor. Movement protein encoded by BC1 plays an important role in the intercellular movement of the viral DNA through the plasmodesmata. In the context of the Indian subcontinent, one characteristic feature of ToLCNDV is its frequent association with either alpha- or betasatellites, despite being a bipartite begomovirus. Betasatellites are approximate B1.5 kb in size and are half of the size of their helper DNA-A viruses, which are necessary for their replication and transportation. These molecules contain a satellite conserved region (SCR), an adenine-rich (A-rich) region and a single ORF in complementary sense strand named bC1. bC1 is a multifunctional protein that acts as a symptom-determinant in the host plant during begomoviral pathogenesis. Another category of ssDNA satellite molecules of B1.5 kb was also identified and named as alphasatellites, which require a helper begomoviruses for inter- and intra-cellular movements, but is independent for replication. Alphasatellites were shown to be responsible for the reduced accumulation of betasatellite levels in begomovirus-beta satellite complexes and to act as a suppressor of RNA silencing. Under laboratory conditions, maintenance of betasatellites (Cotton leaf curl Multan betasatellite, CLCuMB and Radish leaf curl betasatellite, RaLCuB) by ToLCNDV has been well documented. N. benthamiana plants agro-inoculated with ToLCNDV DNA-A contained lesser viral titer as compare to when co-inoculated with either CLCuMB or RaLCuB. Co-inoculated plants also exhibited severe symptoms indicating role of betasatellites in leaf curl disease development. However, it is important to note that DNA-A level was higher in plants co-inoculated with DNA-B as compared with betasatellites co-inoculated plants, which emphasized the important role of DNA-B over betasatellite in facilitating the movement of DNA-A. Furthermore, reduced level of CLCuMB in plants co-inoculated along with DNA-A and DNA-B as compared to DNA-A þ CLCuMB indicates the role of DNA-B in modulating betasatellite levels in infected plants, precise mechanism of which is not yet known. Likewise, three alphasatellites were found to be associated with ToLCNDV in chilli, cotton and cucurbits plants during mixed infection (Table 1). However, the role of alphasatellites on ToLCNDV pathogenesis is yet to be ascertained (Fig. 3).

Synergism and Genetic Changes Leading to Begomovirus Emergence and Evolution There are plenty of non-genetic based factors such as mixed infection, synergism, vector and vegetative dispersal, cultural practices and above all human movement of geminiviruses in new environments, which are promoting geminivirus infection in noncultivated plants and/or other new cultivated crops. However, among all plant viruses, geminiviruses have a particularly high emergence potential because they can use each one or the combination of 4 important genetic-based mechanisms to rapidly change their genetic make-up and capacity, and thereby evolve within a concurrently changing environment to thrive and survive. In addition to having access to these molecular mechanisms, geminiviruses also enjoy a very high rate of multiplication, producing billions of new genomic molecules in a very short span of time, therefore they can afford a large fraction of non-functional molecules to explore many avenues at the same time. These four mechanisms are: the mutations, the recombination process, the pseudo-recombination and the capture of satellites. Recent studies suggest that geminiviruses and nanoviruses, do show a high level of genetic variation and nucleotide substitution rate (103–104 substitution/site/year), and the mechanism is quite similar to RNA viruses. It is believed that these plant DNA viruses may use low-fidelity DNA polymerases for their genome replication which might lead to sequence variation in their genome fairly rapidly. Although, other factors which cause spontaneous mutation in the begomoviral genome (oxidation, transition, transversion, deamination and methylation of nitrogenous bases) might lead to their genetic variability and variation within the species as well. Recombination plays an important role during geminivirus evolution and it can be achieved by two mechanisms, the rolling circle replication and the recombination dependent replication, which leads to the formation of complete new viral genomes. Studies suggest that inter and intra-species recombination events are very frequent in the geminivirus genomes. The impact of recombination has been studied and documented for several viruses such as the Tomato yellow leaf curl viruses, the East African cassava mosaic

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Table 1

Association of ToLCNDV components with other begomoviruses and DNA satellites, in natural and experimental hosts

ToLCNDV genome

Accession # viral genomic components and satellites associated with ToLCNDV

Associated betasatellite

Access# Betasatellite

Natural host

Experimental host

B and HM345979 HM803117 EF095958

CLCuMuB

S. lycopersicum

Nb, Sl

B and EF068246

LuLDB

LN845926, HG934394 KR957354

CLCuMuB PaLCuB

AY083590 EF043234

LuLDB

AY728262

DNA-A ToLCNDV-A, b-sat ToLCNDV-A, b-sat ToLCNDV-A, ToLCNDV-A,

DNA-B

EF408038

B B

LN845962 LN845963 EF043231 EF043232

ToLCNDV-A, B

HM989845 HM989846

Associated BGs

b-satellite

EF068245 AY438562 CLCKoV

AY9399 26 AY9399 24 ToLCNDV-A, B ToLCNDV-A, B

U15015 U15017 HQ141673 HQ141674

ToLCNDV-A, ToLCNDV-A, ToLCNDV-A, ToLCNDV-A, ToLCNDV-A, ToLCNDV-A ToLCNDV-A, ToLCNDV-A, ToLCNDV-A

U15015 HQ141673 HQ141673 U15015 U15015 U15015 U15015 U15015 KU196750

ToLCNDV-A ToLCNDV-A ToLCNDV-B ToLCNDV-B ToLCNDV-B ToLCNDV-B ToLCNDV-B ToLCNDV-B ToLCNDV-B ToLCNDV-B ToLCNDV-B ToLCNDV-B ToLCNDV-B ToLCNDV-B

B B B B B B B

U15017 HQ141674 HQ141674 U15017 U15017 U15017 U15017 U15017

U15015 KP235541 GU112083 LN845955 HQ141674 U15017 U15017 HM803117 U15017 JN663871 JN663848 KP235543 JN663867 KF471060

DQ020491 RaLCB ToLCPalV-A ToLCPalV‐B ToLCPalV-A ToLCPalV-A ToLCPalV-B RaLCV CYVMV CYVMV CYVMB ToLCRnV ToLCRnB ChLCV-A ToLCBDB ToLCGV‐B ToLCGV-B CroYVMB BYVMV ToLCGV-A ToLCGV-A PepLCLV ToLCGV-A ToLCGV-A TYLCTHB PepLCLV ChLCB ChiLCV-PK ToLCRnB ToLCV-Ban ToLCBDB PePLCV-Pal CroYVMB ToLCV-Kar ToLCBDB ChiLCV RaLCB

EF175734

FJ593630 GQ994096 KR957354

FJ593630

HM989847 AM279673 JN663872 JN663849 JN663878 JN663868 JN663873

S. lycopersicum G. hirsutum S. lycopersicum S. tuberosum L. cylindrica M. charantia C. sativus L. siceraria S. lycopersicum S. lycopersicum C.melo S. lycopersicum S. lycopersicum S. lycopersicum S. lycopersicum S. lycopersicum S. lycopersicum S. lycopersicum C. annuum C. chinense S. lycopersicum C. annuum A. esculentus G. hirsutum S. lycopersicum S. lycopersicum S. lycopersicum S. lycopersicum C.annuum C. annuum C. annuum C. annuum C. annuum C. annuum

Nb

Nb, Sl Sl Nb Sl,Cs Sl, Cs Nb, Sl Nb, Sl Nb, Sl Nb, Sl Nb Nb, Ca Nb, Sl,Nt

Nb, Nb, Nb, Nb, Nb,

Sl Nt, Ca Nt, Sl Sl Nt, Ca

Three alphasatellites have been reported to be associated with ToLCNDV DNA-A and other DNA-B in chilli leaf curl disease (ChiLCD) infected samples. These are: ChiLCA (KF471051) with ToLCNDV (DNA-A, KU196750), associated with ChiLCD from New Delhi; ChiLCA (KF471052) with ToLCNDV (DNA-A and B, respectively, KP235540 and JN663871), associated with ChiLCD, Himachal Pradesh and ChiLCA (KF471046) with ToLCNDV (DNA-B, JN663848) associated with ChiLCD, from Karnataka. In cucurbit plants, association of cucurbit yellow mosaic alphasatellite (CYMA, KT948075), ToLCNDV (DNA-A, HM98984; DNA-B, AM286435) and Papaya leaf curl betasatellite, KT948074). In addition, CLCuMA (LN845922, LN845923, LN845924, HG934391, HG934392, and HG934393) was also found to be associated with ToLCNDV A in cotton leaf curl disease (CuLCD) plants.

viruses and the Cotton leaf curl viruses. Although there has not been such detailed recombination analysis done for ToLCNDV, there is no reason to believe that ToLCNDV, does not benefit from the recombination strategy that many begomoviruses explored extensively. Furthermore, there are reports suggesting that ToLCNDV serves as a major parent donor helping in the production of other recombinant begomoviruses, such as Radish leaf curl virus (RaLCuV), Tomato leaf curl Rajasthan virus (ToLCRaV), Papaya leaf crumple virus (PaLCrV), Tomato leaf curl Patna virus (ToLCPaV) and Tomato leaf curl Palampur virus (ToLCPalV). Interestingly pseudo-recombination or, re-assortment of genomic components of DNA-A and DNA-B for the bipartite begomoviruses results in exchange of the genetic materials similar but different to what’s happening during intra-molecular recombination. It is similar because the bipartite begomovirus unit will acquire new proteins with new functions, but it is different because it is the whole set of two proteins acquired instantly and together with the DNA-B re-assortment. There is no creation of de novo chimeric proteins. This event has been reported to be more frequent for Old World begomoviruses, like that of between African cassava mosaic virus (ACMV) and East African Cassava mosaic virus (EACMV) but it has also been reported for New World begomoviruses such as the Pepper golden mosaic virus (PePGMV) and Pepper Huasteco yellow vein virus (PHYVV).

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Fig. 3 Genome organization of ToLCNDV (DNA-A and DNA-B), a betasatellite and an alphasatellite. ToLCNDV is a bipartite geminivirus with components known as DNA-A and DNA-B. The proteins encoded by the components are indicated as the replication associated protein (Rep), the transcriptional activator protein (TrAP), the replication enhancer protein (REn), the C4 protein (C4), the coat protein (CP) and the V2 protein (V2) on the genomes of the DNA A component. The DNA-B component encodes the nuclear shuttle protein (NSP) and the movement protein (MP). A sequence conserved between the DNA-A and DNA-B components of bipartite begomoviruses is known as the conserved region (CR). The betasatellites have a single gene (bC1) in the complementary-sense and a region of sequence highly conserved between all betasatellites [known as the satellite conserved region (SCR)] and a region of sequence rich in adenine (A-rich). The alphasatellites have a single large protein (Rep) and also contain an A-rich sequence. For each component the conserved hairpin structure, containing the nonanucleotide sequence TAATATTAC (TAGTATTAC for most alphasatellites) within the loop structure, is shown at position zero.

Besides, in a laboratory-based experiment done with a pair of highly virulent pseudo-recombinant viruses between the DNA-A from Tomato leaf curl New Delhi virus (ToLCNDV, # U15015) and the DNA-B from Tomato leaf curl Gujarat virus DNA-B (ToLCGuV, # AY190291) it was shown that re-assortment was perfectly functional. This genetic re-assortment resulted in causing a very severe disease on tomato and other solanaceous crops. In those pseudo-recombinant infected plants, the levels of DNA-A and DNA-B were several-fold enhanced. Subsequently, these two molecules were observed under natural condition in the diverse agroclimatic regions of India in chilli, tomato and other solanaceous crops (Table 1). Furthermore, this association also resulted in the breakdown of geminivirus resistance in chilli indicating the relevance of fitness of viral molecules in the host plant. This super compatible integration of two different molecules in an innate condition may affect the host range and the epidemiology of the new virus disease. The fourth possibility for genetic changes for begomoviruses is the capture of ssDNA satellites that are particularly abundant and variable (Table 1).

Host Range of ToLCNDV The members of the ToLCNDV species share today a broad spectrum of host range, and they are known to infect 13 families of vegetable crops, flowering plants and weeds (Table 2). It is not known if the host range of ToLCNDV has effectively changed over

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Table 2

List of plants, and their families, infected with ToLCNDV

Family

Crops

Acanthaceae Amaranthaceae

Crossandra infundibuliformis Chenopodium album Convolvulus arvensis Daucus carota Calotropisprocera Catharanthus roseus Calyptocarpus vialis Chrysanthemum indicum Parthenium hysterophorus Carica papaya Benincasa hispida Citrullus lanatus Coccinia grandis Cucumis melo Cucumis anguinus Cucumis sativus Cucurbita moschata Cucurbita pepo Lagenaria siceraria Luffa aegyptiaca Luffa acutangula Luffa cylindrical Momordica charantia Momordica dioica Sechiumedule Glycine max Lens culinaris Abelmoschusesculentus Gossypium hirsutum Papaver somniferum Sauropus androgynus Rumex dentatus Capsicum annuum Lycopersicon esculentum Nicotiana tabacum Solanum lycoperscicum Solanum nigrum Solanum tuberosum

Apiaceae Apocynaceae Asteraceae Caricaceae Cucurbitaceae

Fabaceae Malvaceae Papaveraceae Phyllanthaceae Polygonaceae Solanaceae

time, but we can only take note that this virus has been isolated over time from a growing number of natural hosts, cotton being the last one. Because of the great capacity for this virus to capture other B components in mixed infections, acquiring thereby the capacity to move more efficiently both components in these hosts, or capturing satellites that could play an important role to suppress gene silencing of the hosts, or a combination of both strategies, it is possible that ToLCNDV has de facto expanded its host range. It is also noticeable, that the new epidemic of ToLCNDV in Europe is mostly on cucurbitaceae, where the symptoms are very severe compared to tomato (Fig. 1). The sequence of both DNA-A and DNA-B components, although having both a large fraction of the typical ToLCNDV components, are novel and unique in the whole ToLCNDV molecular diversity for both components. A detail recombination study is needed to better characterize these molecules and understand the epidemiology of this new ToCNDV strain. Interestingly, co-infection of ToLCNDV is not limited to bipartite begomoviruses but it is also observed in association with monopartite begomoviruses such as Bhendi yellow vein mosaic virus (BYVMV), Chilli leaf curl virus (ChiLCV), Croton yellow vein mosaic virus (CYVMV), Squash leaf curl China virus (SLCCNV), Tomato leaf curl Ranchi virus (ToLCRnV) and Cotton leaf curl Kokhran virus (CLCKoV). A recent study demonstrates the presence of the monopartite begomovirus (ChiLCV) along with ToLCNDV (a bipartite begomovirus) and a betasatellite leading to a severe chilli leaf curl disease (ChiLCD). This interaction between the betasatellite and the geminiviruses led to a synergistic interaction benefitting the geminiviruses, and as a consequence enhancing the disease severity on chilli. Most surprisingly, in Pakistan, ToLCNDV (LN845962) was recently detected in cotton (Gossypium hirsutum) plants showing a typical cotton leaf curl disease. The association of ToLCNDV together with Cotton leaf curl Kokhran virus strain Burewala (CLCKoV-Bur) and Cotton leaf curl Multan betasatellite (CLCMuB) also indicate the possibility of synergistic interactions that might lead to the breakdown of CLCuD resistance in cotton.

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Fig. 4 Geographical distribution of ToLCNDV isolates in the world. ToLCNDV has been isolated from thirteen different plant families that are shown in different colors. The dates for the world spread are roughly indicated with the color circles.

Transmission and Epidemiology The occurrence of ToLCNDV in several countries in Asia, South East Asia, Middle East or Europe indicates that it is not limited by any geographical barrier and it must be efficiently transmitted by a range of whiteflies wherever the virus is present (Fig. 4). ToLCNDV is transmitted by whiteflies (Bemisia tabaci, Genn., Aleyrodidae) under natural conditions and the role of seed transmission has not been evidenced yet but is most probably not relevant. Several biotypes/species of B. tabaci are reported to be transmitting begomoviruses to the Solanaceae, Cucurbitaceae, Malvaceae and many other crops. Middle-East Asia Minor I (MEAM1) (formally known as biotype B) is the most predominant whitefly population in India, especially in the southern part. Initially, the role of MEAM1 whiteflies as vector for transmission of begomoviruses onto solanaceous crops was not considered to be common, despite its presence along with other whitefly biotypes. There is a possibility that the invasive species population favors the emergence of a particular breed because of climate change. The other invasive species of whitefly known as Mediterranean 1 (MED1) (formerly known as biotype Q) was recently identified in Spain to cause ToLCNDV epidemics. This could be an alarming situation and a strategic plan to combat the disease needs to be developed. ToLCNDV has emerged as an Old World bipartite begomovirus that threatens several economically important crops in the world. The presence of ToLCNDV has been reported from thirteen different families of crop plants (Table 2) affected worldwide. The epidemic outbreaks of ToLCNDV in India might be dependent on several factors such as the use of susceptible varieties for crop cultivation, the co-existence of different agro-climatic zones favoring the vector population survival, average rainfall, and the almost constant availability of the whitefly vector population that are swapping from one crop to another. Surprisingly, ToLCNDV is now more widespread in members of the Cucurbitaceae family as compared to Solanaceae or any other crops. In India, farmers generally cultivate cucurbits (bottle gourd, bitter gourd, muskmelon, squash, pumpkin and water melon) in almost every climatic season. The population of whitefly vector remains available throughout the year and the alternate hosts could act as a reservoir for ToLCNDV. In addition, the solanaceous crops such as tomato and chilli, are mostly grown in summer seasons. The higher temperature, relative humidity and availability of various whitefly biotypes/species may therefore provide an advantage for the insect vector to grow more and transmit the virus to several plant species.

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Virus-Host Interactions The geminiviruses utilize the host machinery to effectively function within the host. The plant-virus interplay differs along the spectrum of antagonism and mutualism. Begomoviruses have relatively small genomes that encode multitasking proteins required for the viral replication, encapsidation, transport, viral suppressor of RNA silencing and vector transmission. Many studies suggest plant viruses interact with many host factors and also manipulate several basic mechanisms of the hosts to perform their essential functions. As a counter defense mechanism, plants have developed several antiviral approaches. In a broader perspective, plants utilize resistance gene (R-gene) mediated defense and small non-coding RNA based defense mechanisms against the viral pathogen. In the next few paragraphs these two defense approaches have been discussed.

Small Non-Coding RNAs Geminivirus infection triggers the production of two major classes of small non-coding RNA (sRNA): -small interfering RNAs (siRNAs) and -micro RNAs (miRNAs). These sRNA plays vital roles in the RNA silencing pathways and also control the expression of host genes. As an example, tomato plants infected with ToLCNDV showed upregulation of conserved miR319 and miR172 sRNA expression. These 21 nt long miRNAs play crucial role in regulating antiviral mechanism. Microarray data from ToLCNDV's agro-inoculated tomato plants suggested the differential regulation of miR398, miR399, miR162 and miR168. The miR162 and miR168 are known to regulate the expression of Dicer-like protein-1 (DCL1) and Argonaute 1 (AGO1) protein, respectively. The miR398 is involved in the abiotic-stress related response in plants, and it was observed that the level was higher when infected with the ToLCNDV. These data suggest ToLCNDV infection results in the transcriptional activation and suppression of specific miRNAs in host plant. Therefore, begomoviruses use this distressed gene expression to their purpose, resulting in building a suitable environment for survival inside the host cells. The functional characterization of siRNAs in susceptible, tolerant, and resistant tomato varieties during ToLCNDV infection also has been studied. In addition, the virus-specific siRNAs (Rep specific siRNAs) accumulation at 21 days post inoculation was higher in the tolerant varieties as compared to the susceptible varieties, as showed by northern blot hybridization experiments. The presence of virus specific siRNA were also noted in the systemic tomato leaves. These data further suggest that the virus specific siRNA might be inducing the gene silencing pathways in the tolerant varieties and it failed or is compromised to do so in the susceptible varieties. The higher ToLCNDV DNA-A level was effectively detected in the susceptible variety by Southern experiment.

Counter Defense RNA Silencing Plants have an RNA silencing mechanism to control virus infection and as counter defense, viruses have viral suppressor of RNA silencing (VSRs) which regulate pathogenesis in the host cell. A bipartite begomovirus, such as ToLCNDV, encodes different proteins such as AC2/TraP, AC4, AV2/Pre-coat, and BV1/NSP to attenuate the gene silencing defense mechanism of the host by different means. ToLCNDV-AV2 inhibits the symptom remission on N. tabacum plants by blocking the RNA-Dependent RNA Polymerase 1 from N. tabacum (NtRDR1). NtRDR1 mediated antiviral silencing and the spread of virus specific siRNAs. Mutation in the AC4 ORF of ToLCNDV N50H (Asn position 50) led to the changes in b-sheet structure in SH3 domain resulting in affecting its ability to suppress the RNA silencing pathway. Although it was observed that these mutations do not affect the physical interaction of AC4 with AG04. Hence, ToLCNDV-AC4 might have a role in targeting SH3 domain containing proteins to further modulate the begomoviral pathogenesis in a host cell.

Basal Level of Defense It has been shown that ToLCNDV infections (in tomato and potato) can potentially induce the expression of plant-innate immunity and hormone signaling genes including ethylene-signaling genes such as ethylene response factor-4 (ERF4), WRKY, and NAC transcription factors. The ToLCNDV infection prompts the hypersensitivity response (HR) in the infected cells (in tomato plants) as a result of accumulation of higher reactive oxygen species (ROS) level. Oxidative burst at the infected sites eventually kills both the host and the invaded pathogen. Fortunately, during the course of evolution, a very efficient ROS-scavenging mechanism has also evolved in the plants.

Resistance Gene or R-Gene Mediated Defense Plants have different strategies to combat the pathogens. One of the mechanisms of active plant defense is the resistance gene or R-gene mediated response. The R-gene can identify a specific effector molecule which is produced by the pathogen, and then initiate the HR, resulting in programmed cell death to inhibit the further amplification or spread of the pathogen. Over many decades, breeders have manipulated a likely host-target-loci that is liable with natural sources of resistance for viral infection. In some solanaceous (tomato) and cucurbitaceous plant genotypes, different resistance or tolerant genes have been identified to develop resistance against the ToLCNDV. Particularly, in tomato (wild type species) six TYLCV resistance loci (Ty-1, Ty-2, Ty-3, Ty-4, Ty-5. and Ty-6) have been identified, in which Ty-2 alone failed to show resistance against ToLCNDV. Interestingly, the combination of Ty-2

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(in homozygous or heterozygous condition) and Ty-3 (heterozygous) through marker-assisted selection and breeding, is showing enhanced resistance against ToLCNDV and associated betasatellite. The loci Ty-1 and Ty-3 encode RNA-dependent RNA polymerase (RDR6). RDR6 is involved in the amplification of virus- specific siRNA which leads to the activation of the RNA silencing pathway. Similarly, ToLCNDV infection in solanaceous crop (Capsicum annum, N. tabacum, N. benthamiana and S. lycoperscicum) showed differential expression of defense-related genes such as Suppressor of Gene Silencing 3 (SGS3), RNA-Dependent RNA Polymerase 6 (RDR6) (genes which are involved in PTGS pathways), and the resistance gene (R-gene) such as nt-binding site and leucine-rich repeat (NBS-LRR), Lipid transfer protein (LTP) and pathogenesis-related (PR) genes. A recent study suggests that some of the cultivated species of cucurbits (genus, C. pepo), wild type germplasm lines and F1-hybrid (subspecies C. pepo x C. pepo and subspecies C. pepo x C. ovifera) accessions exhibit resistance against ToLCNDV. Furthermore, the cucurbits accessions used for this study showed very mild symptoms, resulting in low viral titer even after several rounds of mechanical and whitefly inoculations. Similarly, inheritance study of Luffa cylindrica (sponge gourd) conducted during both rainy and spring seasons revealed presence of two sequence-related amplified polymorphism (SRAP) markers closely associated with dominant monogenic inheritance of ToLCNDV-resistance. Therefore these studies further indicate the existence of a genetic basis of resistance in cucurbit plants against ToLCNDV. The continuous evolution of geminiviruses and augmented host range, call for the need of resistance gene that should be investigated continuously to confer useful resistance against ToLCD to benefit farmers.

Diagnosis and Management It is essential to confirm that a given disease is actually caused by a geminivirus, before developing a management program. Currently, the ToLCNDV detection is done by using polymerase chain reaction (PCR)-based amplification of the viral genome and loop mediated isothermal amplification assay (LAMP). Another recent technique is developed is the use of gold nanoparticle conjugated to a probe as biosensors. Furthermore, there are other methods of ToLCNDV detection such as coat protein specific antibody and non-radioactive nucleic acids spot hybridization (NASH) assay. Because of its ability to infect and adapt in various plants, ToLCNDV is exceptionally tricky to manage and control. Some progress on the identification of resistance genes both in tomato and cucurbits is providing hopes that ToLCNDV could be controlled efficiently. Genetic engineering technique has been a key promising approach against the pathogen in a wide variety of crops around the world. One study suggests targeting of AV1/AV2 common overlapping regions by artificial miRNA (amiRNA), conferring tolerance against the ToLCNDV in transgenic tomato crops. However, as begomoviruses are continuously evolving it is very difficult to generate stable resistant or tolerant transgenic lines. The siRNA profiling in tomato and other solanaceous crop gives an idea about the novel resistance host genes. Their characterization can further provide a potential target to develop virus resistance/tolerance plants. Identifying the immune sources and implementing the specific gene in a targeted location to address the challenges of the specific virus, might lead to the sustainable management of the disease. Various other approaches such as sense/antisense technology, RNAi techniques and application of double-stranded RNA (dsRNA) can also be used as potential method to develop resistance against the plant affecting begomoviruses. Recent advancement in the genetic manipulation of the DNA leads to the development of another system termed as clustered regularly interspaced short palindromic repeats-CRISPR associated 9 (CRISPER-Cas9) and is used to engineer broad-spectrum geminivirus resistance in plants. Other management practices like whitefly vector control, breading of resistance varieties and micro-propagation of virus-free plants could be the alternative and complementary approaches to control ToLCNDV.

See also: Betasatellites and Deltasatelliles (Tolecusatellitidae). Emerging Geminiviruses (Geminiviridae). Geminiviruses (Geminiviridae). Plant Resistance to Geminiviruses. Tomato Yellow Leaf Curl Viruses (Geminiviridae)

Further Reading Basu, S., Kushwaha, N., Singh, A.K., et al., 2018. Dynamics of a geminivirus-encoded pre-coat protein and host RNA-dependent RNA polymerase 1 in regulating symptom recovery in tobacco. Journal of Experimental Botany 69 (8), 2085–2102. Chatterji, A., Padidam, M., Beachy, R.N., Fauquet, C.M., 1999. Identification of replication specificity determinations in two strains of tomato leaf curl virus from New Delhi. Journal of Virology 73 (7), 5481–5489. Fortes, I.M., Sánchez-Campos, S., Fiallo-Olivé, E., et al., 2016. A novel strain of Tomato leaf curl New Delhi virus has spread to the Mediterranean basin. Viruses 8 (11), 307. García-Arenal, F., Zerbini, F.M., 2019. Life on the edge: Geminiviruses at the interface between crops and wild plant hosts. Annual Review of Virology 6, 411–433. Hussain, M., Mansoor, S., Iram, S., Fatima, A.N., Zafar, Y., 2005. The nuclear shuttle protein of Tomato leaf curl New Delhi virus is a pathogenicity determinant. Journal of Virology 79 (7), 4434–4439. Islam, S., Anilabh, D.M., Verma, M., et al., 2011. Screening of Luffa cylindrica Roem. for resistance against Tomato leaf curl New Delhi virus, inheritance of resistance, and identification of SRAP markers linked to the single dominant resistance gene. The Journal of Horticultural Science and Biotechnology 86 (6), 661–667. Kumar, R.V., Singh, A.K., Singh, A.K., et al., 2015. Complexity of begomovirus and betasatellite populations associated with chilli leaf curl disease in India. Journal of General Virology 96 (10), 3143–3158. Kushwaha, N., Singh, A., Basu, S., Chakraborty, S., 2015. Differential response of diverse solanaceous hosts to Tomato leaf curl New Delhi virus infection indicates coordinated action of NBS-LRR and RNAi-mediated host defense. Archives of Virology 160 (6), 1499–1509.

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Malik, A.H., Briddon, R.W., Mansoor, S., 2011. Infectious clones of Tomato leaf curl Palampur virus with a defective DNA-B and their pseudo-recombination with Tomato leaf curl New Delhi virus. Virology Journal 8 (1), 173. Moriones, E., Praveen, S., Chakraborty, S., 2017. Tomato leaf curl New Delhi virus: An emerging virus complex threatening vegetable and fiber crops. Viruses 9 (10), 264. Naqvi, A.R., Haq, Q.M., Mukherjee, S.K., 2010. MicroRNA profiling of Tomato leaf curl New Delhi virus (ToLCNDV) infected tomato leaves indicates that deregulation of mir159/319 and mir172 might be linked with leaf curl disease. Virology journal 7 (1), 281. Padidam, M., Beachy, R.N., Fauquet, C.M., 1995. Tomato leaf curl geminivirus from India has a bipartite genome and coat protein is not essential for infectivity. Journal of General Virology 76, 25–35. Pratap, D., Kashikar, A.R., Mukherjee, S.K., 2011. Molecular characterization and infectivity of a Tomato leaf curl New Delhi virus variant associated with newly emerging yellow mosaic disease of eggplant in India. Virology Journal 8 (1), 305. Sáez, C., Martínez, C., Ferriol, M., et al., 2019. Resistance to Tomato leaf curl New Delhi virus in Cucurbita spp. Annals of Applied Biology 169 (1), 91–105. Sahu, P.P., Rai, N.K., Chakraborty, S., et al., 2010. Tomato cultivar tolerant to Tomato leaf curl New Delhi virus infection induces virus-specific siRNA accumulation and defense associated host gene expression. Molecular Plant Pathology 11 (4), 531–544. Singh, A.K., Kushwaha, N., Chakraborty, S., 2016. Synergistic interaction among begomoviruses leads to the suppression of host defense-related gene expression and breakdown of resistance in chilli. Applied Microbiology and Biotechnology 100 (9), 4035–4049. Van, Vu T., Choudhury, N.R., Mukherjee, S.K., 2013. Transgenic tomato plants expressing artificial microRNAs for silencing the pre-coat and coat proteins of a begomovirus, Tomato leaf curl New Delhi virus, show tolerance to virus infection. Virus Research 172 (1–2), 35–45.

Tomato Spotted Wilt Virus (Tospoviridae) Hanu R Pappu, Washington State University, Pullman, WA, United States Anna E Whitfield, North Carolina State University, Raleigh, NC, United States Athos S de Oliveira, University of Brasília, Brasília, Brazil r 2021 Elsevier Ltd. All rights reserved. This is an update of H.R. Pappu, Tomato Spotted Wilt Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00648-8.

Nomenclature aa Amino acid(s) Avr Avirulence CC Coiled coil ELISA Enzyme-linked immunological assays GRSV Groundnut ringspot virus HR Hypersensitive response IDM Integrated disease management JA Jasmonate LRR Leucine-rich repeat MP Movement protein N Nucleocapsid NB Nucleotide-binding

Glossary Avirulence determinant Microorganism molecular patterns, e.g., a viral protein, that activate plant resistance proteins, inhibiting infection/virulence. Circulative, propagative transmission Virus transmission process in which the virus circulates

NLR Nucleotide-binding, leucine-rich repeat receptor ORF Open reading frame PB Processing body RB Resistance-breaking RdRp RNA-dependent RNA polymerase RI Resistance-inducing RNAi RNA interference RNP Ribonucleoprotein SA Salicylic acid SD Solanaceae domain TEM Transmission electron microscopy TSG Tubular salivary glands TSWV Tomato spotted wilt virus

through insect body and replicates in various tissue systems. Processing bodies RNA-protein complexes located in the cytoplasm where RNA decay pathways occur.

Classification Tomato spotted wilt orthotospovirus is the founding virus species in the genus Orthotospovirus, family Tospoviridae, order Bunyavirales. Tomato spotted wilt virus (TSWV) was first described in 1915 and was recognized as a bunyavirus in the 1990s based on virion morphology, genome sequence, and replication in the insect vector, thrips. TSWV was the first phytovirus classified as a bunyavirus. At present, there are 26 accepted species and 5 tentative species in the genus Orthotospovirus and there are three other phytovirus genera (Tenuivirus, Emaravirus, and Coguvirus) within different families in the order Bunyavirales.

Virion Structure TSWV has the classic bunyavirus particle structure that is composed of a membrane that encapsulates the three genome segments (L, M, and S) that are coated in nucleocapsid (N) proteins. The genome is composed of negative-sense RNAs and the 50 and 30 ends of the viral RNAs are highly conserved and complementary. The complementary ends of the genome segments enable the formation of panhandle structures and these sequences serve as promoters for transcription of genes and genome replication. The N proteins arrange as asymmetric trimeric rings, creating an inner cavity where the viral RNAs bind. The virus particle also contains several copies of the L protein, an RNA-dependent RNA polymerase (RdRp), that is essential for infectivity of the virus upon delivery into the host plant or vector. The two glycoproteins, Gn and Gc, decorate the surface of the virion and are the attachment and entry proteins involved in the infection cycle in the insect vector (Fig. 1). Among tospoviral proteins, the glycoproteins, and the N protein of TSWV and groundnut ringspot virus (GRSV) and the NSs have been predicted by folding prediction, and the native structure of the N protein was determined by crystallization, and the RNA binding sites were mapped in the simulated 3D structure. The N protein structure is composed of three main parts: N-terminal arm, C-terminal arm, and core domain. To form the trimeric ring, the N- and C-terminal arms of each N protein

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Fig. 1 (Left) Virus particle diagram of TSWV. (Right) Transmission electron microscope picture of TSWV particles. The bar represents 100 nm. Virion cartoon used with permission from Rotenberg, D., Jacobson, A.L., Schneweis, D.J., Whitfield, A.E., 2015. Thrips transmission of tospoviruses. Current Opinion in Virology 15, 80–89. Photo courtesy of Dr Jan Van Lent.

interact with the core domain of the other two adjacent proteins. The folding pattern consisted of a globular core shape containing two b-sheets and 12–17 a-helix, and two terminal chains corresponding to the N and the C termini exposed on the surface of the protein. The N protein structure has important aa for binding to RNA and protein folding such as R94/R95 and K183/Y184, which are mapped onto a charged surface cleft of the three-dimensional structure. Interestingly R95 is conserved among all orthotospoviruses and is consistently located in a a-helix.

Genome The three negative-sense RNAs of TSWV encode six proteins on five genes, and some functions of each TSWV protein have been elucidated. The L protein (RdRp) works as a transcriptase, synthesizing mRNAs to be translated into functional proteins, and as a replicase, synthesizing new genomic RNAs which are then covered by N proteins. Interestingly, the L protein also snatches 50 capped ends from host mRNAs to cap-protect the viral mRNAs, the so-called ‘cap-snatching’ phenomenon. The L protein prefers cap-donors with multiple base complementarity to the viral RNA templates and seems to select host mRNAs which are destined for decapping in processing bodies (PBs). It has been shown that alterations in protein components of PBs, where mRNA decay pathways occur, interfere on TSWV cap-snatching and infection in Arabidopsis mutants. The nucleoprotein (N) functions as a protective layer encapsidating the three viral genomic RNA segments, but also plays an important role in viral RNA transcription and replication. The N in the RNP complex interacts with the cytoplasmic tails of the glycoproteins which collectively form the mature virion or can interact with the NSm protein for cell to cell movement through the plasmodesmata. The glycoproteins of TSWV are the viral determinants of insect transmission. The glycoproteins are encoded on single ORF and post-translationally cleaved into mature independent Gn and Gc proteins. The glycoproteins associate together and move from the ER to the Golgi where virus budding occurs. The glycoproteins in the Golgi form the external core that encapsidates the RNPs through interactions with the N protein and the cytoplasmic tails of the glycoproteins. The resulting double enveloped virion interacts with the endoplasmic reticulum cisternae, clustering several virions on vesicles. Gn also functions as an attachment protein for virus acquisition and is a determinant for virus transmissibility by thrips, while Gc is a viral fusion-like protein. The upstream sequence of Gn/Gc was found to contain promoter elements that are light and auxin inducible when expressed in Arabidopsis. The NSm, a non-structural protein, serves as the cell to cell and long-distance movement protein (MP), through interaction with the N protein in the RNP complex. For cell to cell movement, the NSm forms tubular structures and modifies the size exclusion of the plasmodesmata with neighboring cells. Additionally, the NSm is associated with symptom expression and was shown to be an avirulence (avr) determinant in resistant plant varieties harboring the gene Sw-5b. The NSs, another non-structural protein, has been shown to serve as suppressor of antiviral RNA silencing in host plants and insect vector cells. The NSs also plays a major role in TSWV transmission by suppressing the biosynthesis of volatile monoterpenes, plant repellents of thrips. The NSs targets MYC transcription factors, regulators of the jasmonate(JA)-signaling pathway that controls the expression of terpene synthase genes. Therefore, TSWV-infected plants become more attractive to thrips, which favors virus acquisition by these insect vectors. The NSs is also associated with symptom development and supports systemic viral movement by interfering with antiviral RNA silencing in plants.

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Fig. 2 The tomato spotted wilt virus transmission cycle. Thrips eggs are oviposited into plant tissue and within a few days the first instar larvae emerge. Virus acquisition occurs during the larval stages after which the virus persists in the vector and is passed transstadially to the adult. The pupal insects may occur in the soil or leaf tissue and these stages are nonfeeding. Adults emerge and may disperse widely. Only adult thrips (male and female) that acquired the virus during their larval stages can transmit tospoviruses. Used with permission from Rotenberg, D., Jacobson, A.L., Schneweis, D.J., Whitfield, A.E., 2015. Thrips transmission of tospoviruses. Current Opinion in Virology 15, 80–89.

Transmission Insects in the order Thysanoptera are commonly called ‘thrips’ and they transmit TSWV and other orthotospoviruses in a circulative, propagative manner. There are over 7000 thrips species, but only nine species have been shown to transmit TSWV (Frankliniella bispinosa, F. cephalica, F. fusca, F. gemina, F. intonsa, F. occidentalis, F. schultzei, Thrips setosus, T. tabaci). All thrips species that transmit plant viruses are classified in the family Thripidae and they are phytophagous insects that feed in a piercing-sucking manner. Thrips feeding behaviors and high reproductive capacity make them efficient vectors of TSWV. TSWV has a unique insect developmental-stage-dependent transmission cycle (Fig. 2). Virus is acquired when larval thrips feed on TSWV-infected tissue. As thrips mature, they lose their ability to become efficient vectors and adult thrips cannot effectively acquire virus and become virus transmitters. In the larval thrips that acquire TSWV, the virus disseminates from the initial site of infection, the midgut, to the salivary gland tissues. The pupal stages do not feed and thus do not acquire or inoculate virus. Both male and female adults are efficient virus transmitters and they can effectively disperse the virus over short and long distances. The small size of thrips makes them an efficient transport mechanism for viruses and thrips are often found in shipments of diverse types of plant materials. The TSWV dissemination pathway in thrips has been described in detail. During larval thrips acquisition, the virus is ingested and enters the anterior midgut epithelial cells. This is the primary site of virus replication and virus infection appears to spread to surrounding cells. From the gut epithelial cells, virions traverse the basal membrane and basal labyrinth to then infect the muscle

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cells surrounding the gut. Confocal and TEM immunolabeling studies have shown that the cells of the tubular salivary glands (TSG) contain detectable amounts of TSWV. The TSGs connect the midgut to the principal salivary glands and may serve as a conduit for virus movement between insect tissue systems. Alternatively, virus may move between gut tissues and principal salivary glands during the larval stage when these tissues are in direct contact. The principal salivary glands are the final site for virus replication prior to delivery of virus with feeding and salivation into the plant host. The extent of principal salivary gland infection has been shown to be associated with transmission efficiency. The virus abundance in larval thrips is positively correlated with infection status as adults and transmission efficiency. Interestingly, TSWV has not been found in the hemolymph of infected insects but this could be due to technical difficulties associated with virus detection in this small insect.

Epidemiology Numerous experiments have confirmed that TSWV has a beneficial effect on F. occidentalis. The infection of several different plant hosts has resulted in faster developmental times for this species of vector. In addition, female thrips are attracted to the virus infected plants and this synergistic interaction facilitates transmission of the virus because larval thrips acquire TSWV. F. occidentalis is the most widespread vector of TSWV, but other vectors have regional importance and more varied interactions with the virus. For example, in the South-Eastern U.S. F. fusca is an important vector of TSWV to peanut but insect exposure to virus-infected peanuts negatively impacted vector development time. In addition to variation between TSWV-vector species interactions, there can be virus and insect isolate differences in interactions. Examination of TSWV-infected plants shows that they have higher levels of free amino acids and may have modified volatile organic compound profiles.

Pathogenesis TSWV infects over a thousand plant species and infection results in significant yield losses for many agronomically important crops. Typical virus symptoms include stunting, wilting, and necrotic or chlorotic concentric spots on susceptible plants (Fig. 3). Many plants have an age-dependent susceptibility to TSWV. Physiological measurements of TSWV-infected peanut revealed that photosynthesis, transpiration, and water‐use efficiency were significantly decreased in virus‐affected plants. In TSWV-infected plants, the phytohormone salicylic acid is present at higher levels and SA-related genes are up-regulated indicating a plant defense response to virus infection. Several TSWV resistance genes have been identified and studied at the ecological and molecular level. In plants and insects, the TSWV NSs protein is a suppressor of RNA silencing and therefore a counter defense to one of the major antiviral responses in eukaryotic hosts. Also, the NSs of TSWV has been reported as an avr determinant in pepper (Capsicum annuum). This suggests an additional role for the NSs besides the well-defined RNAi suppressor activity. Likewise, it was recently suggested that the NSs of TSWV has a role in translation, and persistent infection and transmission by F. occidentalis. Furthermore, accumulation of high levels of NSs in the salivary glands of thrips could be indicative of NSs protein being co-injected into plants during thrips feeding. The NSs protein interferes with the RNA silencing response in plants by affinity for different types of dsRNA molecules. Progress has been made in characterizing the thrips response to virus infection. There are diverse thrips-TSWV interactions that have been documented ranging from symbiotic to pathogenic and for the best characterized system, TSWV and F. occidentalis, the interaction is beneficial to the insect in most experiments. Gene expression analysis of western flower thrips, F. occidentalis, response to TSWV revealed that there is a developmental-stage-dependent transcriptome response to virus infection. The number and identity of responsive transcripts was unique to each thrips stage of development (larval, pupal, and adult). Only three sequences were differential in all life stages tested. The diverse response to virus infection over thrips development is likely due to the unique tissue tropism over the lifespan of the insect and dynamics of the virus replication cycle at the distinct infection stages. Similar experiments with TSWV-infected F. fusca yielded a greater number of differential transcripts but again a unique suite of responsive genes in larval, pupal, and adult thrips with only eight being responsive at all developmental stages.

Diagnosis TSWV causes distinct foliar symptoms on some susceptible crops such as groundnut (peanut), potato, tobacco, and tomato that are of diagnostic value. Confirmatory, laboratory-based assays include ELISA-based serological test using antibodies specific to the N protein. The genome of several isolates of TSWV was sequenced and RT-PCR-based molecular detection methods are available.

Prevention In case of perennials and vegetatively propagated plants and crops, virus can spread from infected mother plants through cuttings and tubers. Use of virus-free material is very effective in preventing disease outbreaks. TSWV is not seed-transmitted, and the primary source of infection in annual crops is mainly due to feeding by viruliferous thrips. Reducing yield losses due to TSWV infection is mainly through an integrated disease management (IDM) strategy that involves vector management by biological and

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Fig. 3 Symptoms caused by TSWV in plants and the thrips Frankliniella occidentalis. Pepper fruit (A) and (B) leave photographed by Gerald Holmes, California Polytechnic State University at San Luis Obispo. (C) Tomatoes photographed by Elizabeth Bush, Virginia Polytechnic Institute and State University (VT). (D) Tobacco leaves photographed by Mary Ann Hansen, VT. (E) Frankliniella occidentalis photographed by Jack T. Reed, Mississippi State University.

chemical means, cultural practices, and growing TSWV resistant cultivars. A set of IDM tactics was developed in peanut based on the relative importance of risk factors (TSWV Risk Index) that result in disease epidemics.

Plant Resistance Several natural resistance sources have been reported to halt TSWV infection. Among them, only the tomato Sw-5b gene-mediated resistance has been deeply studied. Originally, the Sw-5b gene is from Solanum peruvianum, a wild species of tomato naturally occurring in northern Chile and southern Peru. The Sw-5b shares homology with other four Sw-5 paralogs, named Sw-5a, Sw-5c, Sw-5d, and Sw-5e, encoding nucleotide-binding (N) leucine-rich repeat (L) receptors (R). The NLR proteins are usually activated by direct or indirect interaction with avr determinants, which eventually result in the appearance of necrotic spots on inoculated leaves, phenomenon known as hypersensitive cell death response (HR). Although many Sw-5 paralogs and orthologs have been reported, only the Sw-5b protein has been proved to activate resistance to orthotospoviruses (Fig. 4). Due to its durability and broad-spectrum resistance, the Sw-5b gene has been introgressed into many commercial tomato cultivars, becoming the main natural resistance source against TSWV for this crop. This broad-spectrum resistance is not just observed for TSWV isolates, but also for orthotospoviruses classified in other species phylogenetically-related to TSWV. The NSm protein is the cognate avr determinant of Sw-5b. Single expression of NSm in Sw-5b-transformed N. benthamiana and in Sw-5b-resistant tomato lines triggers HR in agro-infiltrated leaves. The recognition of NSm as avr occurs by its direct association with Sw-5b. Two domains of Sw-5b, NB-ARC and LRR, directly interact with NSm by recognizing an epitope of 21 aa, which is conserved among TSWV-related orthotospoviruses. Most NLR proteins, however, are thought to indirectly recognize pathogens since they monitor host proteins that are targeted and modified by avr determinants.

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Fig. 4 Sw-5b-mediated resistance to TSWV in tomatoes. Although there are several Sw-5 paralogs in tomato plants bred with S. peruvianum, only Sw-5b senses an epitope of 21 aa in NSm. This protein-protein interaction activates resistance to TSWV. Some point mutations in NSm, however, can turn resistance-inducing (RI) TSWV isolates into resistance-breaking (RB) isolates. The Sw-5 proteins contain the domains SD (Solanaceae Domain), CC (Coiled-Coil), NB (Nucleotide-Binding), and LRR (Leucine-Rich-Repeat).

Other resistance sources to TSWV (Sw-1a, Sw-1b, Sw-2, Sw-3, Sw-4, Sw-6, and Sw-7), either recessive or dominant, have been reported and introgressed into commercial tomato cultivars. In comparison to Sw-5b, they commonly have a narrower-spectrum resistance against TSWV isolates and have not been characterized molecularly. In pepper, the Tsw gene activates resistance against TSWV isolates, inducing a clear HR on inoculated leaves. Besides some similarities with Sw-5b, the Tsw-mediated HR is triggered by the NSs protein of TSWV. Many other plant species and accessions have been reported to be resistant to TSWV isolates but, contrasting with plants harboring Sw-5b or Tsw, there is none or little genetic and molecular information available. Overtime, the constant use of a monogenic resistance source on a virus population selects virus mutants that are not sensed by the resistance gene product or that suppress the resistance mechanism. In this context, TSWV resistance-breaking (RB) isolates have been reported for both Sw-5b and Tsw genes in tomato and pepper growing fields, respectively. Two aa substitutions in NSm proteins, C118Y or T120N, overcome Sw-5b-mediated resistance by TSWV isolates. A single mutation in the NSs protein, T104A, overcomes Tsw-mediated resistance. Evidences of reassortment events have also been reported for TSWV. This evolutionary mechanism is an additional advantage of TSWV in adapting to new plant hosts and thrips vectors, and from escaping selection pressures as resistance plant genes. The durability of a resistance gene in the field, nevertheless, can be spanned with proper management of resistant cultivars. Actions such as alternating resistant and susceptible cultivars or planting susceptible cultivars alongside production fields may reduce the selection and arising of RB isolates.

Further Reading Abudurexiti, A., Adkins, S., Alioto, D., et al., 2019. Taxonomy of the order Bunyavirales: Update 2019. Archives of Virology 164. doi:10.1007/s00705-019–04253-6. Brown, Steve L., Culbreath, A.K., Todd, J.W., et al., 2005. Development of a method of risk assessment to facilitate integrated management of spotted wilt of peanut. Plant Disease 89, 348–356. Culbreath, A.K., Todd, J.W., Brown, S.L., 2003. Epidemiology and management of Tomato spotted wilt virus in peanut. Annual Review of Phytopathology 2003 (41), 53–75. de Oliveira, A.S., Boiteux, L.S., Kormelink, R., Resende, R.O., 2018. The Sw-5 gene cluster: tomato breeding and research toward orthotospovirus disease control. Frontiers Plant Science 9, 1055. doi:10.3389/fpls.2018.01055. Ma, X., Zhou, Y., Moffett, P., 2019. Alterations in cellular RNA decapping dynamics affect Tomato spotted wilt virus (TSWV) cap snatching and infection in Arabidopsis. New Phytologist. doi:10.1111/nph.16049. Montero-Astúa, M., Rotenberg, D., Leach-Kieffaber, A., et al., 2014. Disruption of vector transmission by a plant-expressed viral glycoprotein. Molecular Plant Microbe Interactions 27 (3), 296–304. doi:10.1094/MPMI-09–13-0287-FI. Montero-Astúa, M., Stafford-Banks, C.A., Badillo-Vargas, I.E., et al., 2016. Chapter 20: Tospovirus–thrips biology. In: Brown, J.K. (Ed.), Vector-Mediated Transmission of Plant Pathogens. APS Press. doi:10.1094/9780890545355.020. Montero-Astúa, Mauricio, Ullman, Diane E., Whitfield, Anna E., 2016. Salivary gland morphology, tissue tropism and the progression of tospovirus infection in Frankliniella occidentalis. Virology 493, 39–51. doi:10.1016/j.virol.2016.03.003. Olaya, C., Adhikari, B., Raikhy, G., Cheng, J., Pappu, H.R., 2019. Identification and localization of Tospoviridae family-wide conserved residues in 3D models of the nucleocapsid and the silencing suppressor proteins. Virology Journal 16, 7. doi:10.1186/s12985-018–1106-4. Rothenberg, D., Whitfield, A.E., 2018. Molecular interactions between tospoviruses and thrips vectors. Current Opinion in Virology 33, 191–197. doi:10.1016/j. coviro.2018.11.007.

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Schneweis, D.J., Whitfield, A.E., Rotenberg, D., 2017. Thrips developmental stage-specific transcriptome response to Tomato spotted wilt virus during the virus infection cycle in Frankliniella occidentalis, the primary vector. Virology 500, 226–237. doi:10.1016/j.virol.2016.10.009. Srinivasan, R., Abney, M., Culbreath, A., et al., 2017. Three decades of managing Tomato spotted wilt virus in peanut in southeastern United States. Virus Research 241, 203–212. Zhai, Y., Peng, H., Neff, M.M., Pappu, H.R., 2019. Putative auxin and light responsive promoter elements from the Tomato spotted wilt virus genome, when expressed as cDNA, are functional in Arabidopsis. Frontiers in Plant Science 10. doi:10.3389/fpls.2019.00804.

Relevant Websites https://site.extension.uga.edu/mitchellag/files/2016/06/Peanut-Rx-and-Various-Fungicide-Programs.pdf Peanut Rx and Various Fungicide Programs. University of Georgia. http://www.ento.csiro.au/thysanoptera/worldthrips.php Thysanoptera (Thrips) World Checklist. CSIRO Entomology. http://climate.ncsu.edu/thrips Tobacco Thrips Flight and TSWV Intensity Predictor North Carolina Climate Office. https://www.hgsc.bcm.edu/arthropods/western-flower-thrips-genome-project Western Flower Thrips Genome Project. BCM-HGSC.

Tomato Yellow Leaf Curl Viruses (Geminiviridae) Henryk Czosnek, The Hebrew University of Jerusalem, Rehovot, Israel r 2021 Elsevier Ltd. All rights reserved. This is an update of H. Czosnek, Tomato Yellow Leaf Curl Virus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00717-2.

Glossary Agroinoculation, agroinfection An alternative route for viral infection; the viral genome is cloned, usually as a headto-tail dimer, in the T-DNA of Agrobacterium tumefaciens and is delivered to plants by inoculation. Introgression Incorporation of chromosomal segments of wild tomato species in the domesticated tomato by crosses and selection; introgression lines are used to localize genes to the tomato chromosomes

Rolling circle Mechanism of replication followed by many viral DNA and by begomoviruses in particular. Viruliferous whitefly Insect that has acquired virus from an infected plant and is ready to infect other host plants. Whitefly Insect that belongs to the order Homoptera; they cause damage to plants by feeding and by vectoring plant viruses.

Introduction Tomato yellow leaf curl virus (TYLCV) was coined in the late 1950s to describe a virus infecting tomato plants in the Middle East. TYLCV causes one of the most devastating diseases of tomato worldwide. The virus (genus Begomovirus, family Geminiviridae) is transmitted by the whitefly Bemisia tabaci. (family Aleyrodidae, order Hemiptera) in circulative manner. The virus was isolated, cloned and sequenced in the late 1980s. It has a circular single-stranded DNA genomic molecule of 2787 nt encapsidated in a 20
30 nm geminate particle. The viral strand encodes four proteins while the viral complementary strand encodes two. The TYLCV genome replicates in host cell nuclei according to the rolling circle model. From the early 1960s, TYLCV has quickly spread from the Eastern Mediterranean Basin to the entire Middle East, Central and Southeast Asia, China, North and West Africa, Southeast Europe, the Caribbean islands, Southeast USA, the Southern Indian Ocean islands, Australia and Japan. Global warming may be instrumental to the expansion of the whitefly-transmitted virus to presently temperate regions. Sequence comparisons of TYLCV isolates from different regions revealed that the name TYLCV encompasses a complex of closely, as well as distantly, related begomovirus species affecting tomato. Over the last 40 years, TYLCVs have been the subject of numerous investigations, including epidemiology, phylogenetics, plant-virus-vector interactions, diagnosis, and breeding for resistance.

Classification and Taxonomy TYLCV is a member of the genus Begomovirus of the family Geminiviridae. Begomoviruses are transmitted by the whitefly Bemisia tabaci (family Aleyrodidae, order Hemiptera). B. tabaci comprises many species (or biotypes), distinguished by their DNA sequence. Among them, the Middle East – Asia Minor 1 (MEAM1, or B) and the Mediterranean (MED, or Q) species are efficient vectors of TYLCV. TYLCV was isolated and its genome sequenced in the late 1980s. Begomoviruses have a genome either split between two circular single-stranded DNA (ssDNA) molecules of approximately 2700 nt each, named DNA A and DNA B (bipartite), or with a single genomic DNA A-like molecule (monopartite). TYLCVs are monopartite, except for two viruses from Thailand. Sequence comparisons revealed that the name TYLCV encompasses a complex of closely, as well as distantly, related begomoviruses affecting tomato. Guidelines for the classification of begomoviruses state that genome-wide pairwise identities of 91% and 94% constitute the demarcation threshold between species and strains, respectively. Accordingly, TYLCVs are classified into several species and strains, listed in Table 1 (updated as of January 2015). This classification is rendered more complicated by the discovery that recombination between different TYLCVs happens relatively frequently (see below). Additional begomoviruses that infect tomato cultures are described in other chapters. These viruses clearly differ from TYLCVs in the symptoms they induce on tomato, in their host range, and in their nucleotide sequence.

Virion Structure Like all geminiviruses, TYLCV has a particle of twinned morphology of approximately 20
30 nm in size (Fig. 1(a)). The virus capsid consists of two joined, incomplete icosahedra, with a T ¼ 1 surface lattice containing a total of 22 capsomeres each containing five units of a 260 amino acid coat protein (30.3 kDa). The geminate particle contains a single 2787 nt covalently closed genomic circular ssDNA molecule. A model of the structure of Ageratum yellow vein virus, obtained by cryo-EM at 3.3 Å , indicates that the DNA fills the entire available space within the particle.

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Table 1 List of Tomato yellow leaf curl virus species, as of January 2015. Species names are shown in bold; isolate names are given in regular font. For species that do not have any known strains, only one isolate is listed, and that isolate is recognized as the “type” isolate. For species that have known strains, one isolate from each strain is shown, and the type isolate is the first one listed. Abbreviations are in the second column and the sequence accession numbers in the third TYLCV species and isolates

Abbreviation of name of virus isolates Accession number

Tomato yellow leaf curl Axarquia virus Tomato yellow leaf curl Axarquia virus – [Spain-Algarrobo-ES-mh800–2000]

TYLCAxV-[ES-Alg-ES-mh800-00]

AY227892

Tomato Tomato Tomato Tomato Tomato Tomato Tomato Tomato

TYLCCNV-BS1[CN-Yn10-Tob-00] TYLCCNV-[CN-Yn25-Tom-00] TYLCCNV-BS3[CN-Yn278-Mal07] TYLCCNV-Bea[CN-Yn-Bea-04] TYLCCNV-DL[CN-Yn5-Tob-99] TYLCCNV-Dat[CN-Yn72-Dat-05] TYLCCNV-HH[CN-Gx]

AJ319675 AJ457985 AM980509 DQ256460 AJ319674 EF011559 AF311734

Tomato yellow leaf curl Indonesia virus Tomato yellow leaf curl Indonesia virus – [Indonesia-Lembang-2005]

TYLCIDV-[ID-Lem-05]

AF189018

Tomato yellow leaf curl Kanchanaburi virus Tomato yellow leaf curl Kanchanaburi virus – [Thailand-Kanchanaburi 1-2001]

TYLCKaV-[TH-Kan1-01]

AF511528 AF511529

Tomato yellow leaf curl Malaga virus Tomato yellow leaf curl Malaga virus – [Spain-421-1999]

TYLCMaV-[ES-421-99]

AF271234

Tomato yellow leaf curl Mali virus Tomato yellow leaf curl Mali virus – Ethiopia [Ethiopia-Melkassa-2005] Tomato yellow leaf curl Mali virus – Mali [Mali-2003]

TYLCMLV-ET[ET-Mel-05] TYLCMLV-ML[ML-03]

DQ358913 AY502934

Tomato yellow leaf curl Sardinia virus Tomato yellow leaf curl Sardinia virus – [Italy-Sardinia-1988]

TYLCSV-[IT-Sar-88]

X61153

Tomato Tomato Tomato Tomato Tomato Tomato

yellow yellow yellow yellow yellow yellow

leaf leaf leaf leaf leaf leaf

curl curl curl curl curl curl

Thailand Thailand Thailand Thailand Thailand Thailand

TYLCTHV-A[TH-1] TYLCTHV-B[TH-ChMai] TYLCTHV-C[CN-Yn72-02] TYLCTHV-D[MY-Yan-99] TYLCTHV-E[TH-SaNa]

X63015
63016 AY514630 AY514633 AJ495812 AF206674 AY514632 AY514635

Tomato Tomato Tomato Tomato Tomato Tomato Tomato Tomato

yellow yellow yellow yellow yellow yellow yellow yellow

leaf leaf leaf leaf leaf leaf leaf leaf

curl curl curl curl curl curl curl curl

virus virus virus virus virus virus virus virus

TYLCV-[IL-Reo-86] TYLCV-Bou[IR-Gen29-06] TYLCV-IR[IR-Ira-98] TYLCV-Kah[IR-Kah-07] TYLCV-Ker[IR-Hor32-06] TYLCV-Mld[IL-93] TYLCV-OM[OM-Alb22-05]

X15656 GU076454 AJ132711 EU635776 GU076442 X76319 FJ956700

TYLCYnV-[CN-YN2013-11]

KC686705

yellow leaf curl China virus yellow leaf curl China virus – Baoshan1 [China-Yunnan 10-Tobacco-2000] yellow leaf curl China virus – [China-Yunnan 25-Tomato-2000] yellow leaf curl China virus – Baoshan3 [China-Yunnan 278-Malvastrum-2007] yellow leaf curl China virus – Bean [China-Yunnan-Bean-2004] yellow leaf curl China virus – Dali [China-Yunnan 5-Tobacco-1999] yellow leaf curl China virus – Datura [China-Yunnan 72- Datura-2005] yellow leaf curl China virus – Honghe [China-Guangxi]

– – – – – – –

virus virus virus virus virus virus

– – – – –

A [Thailand-1] B [Thailand-Chiang Mai] C [China-Yunnan 72-2002] D [Myanmar-Yangon-1999] E [Thailand-Sakon Nakhon]

[Israel-Rehovot-1986] Boushehr [Iran-Genaveh 29-2006] Iran [Iran-Iranshahr-1998] Kahnoo [Iran-Kahnooj-2007] Kerman [Iran-Hormozgan 32-2006] Mild [Israel-1993] Oman [Oman-Al-batinah 22-2005]

Tomato yellow leaf curl Yunnan virus Tomato yellow leaf curl Yunnan virus – [China-YN2013-2011]

Note: Adapted from Brown, J.K., Zerbini, F.M., Navas-Castillo, J., et al., 2015. Revision of Begomovirus taxonomy based on pairwise sequence comparisons. Archives of Virology 160, 1593–1619.

Genome TYLCVs have a monopartite genome, except for two viruses from Thailand (Tomato yellow leaf curl Thailand virus – TYLCTHV, and Tomato yellow leaf curl Kanchanaburi virus – TYLCKaV), which have bipartite genomes.

Monopartite TYLCVs A schematic drawing of the monopartite TYLCV genome is shown in Fig. 2. The TYLCV DNA-A-like viral genome encodes two large open reading frames (ORF) on the viral strand (V1 and V2), and four on the complementary strand (C1-C4). A 313 nt long intergenic region (IR) contains a 29 nt-long stem-loop structure with the conserved nanonucleotide TAATATTAC, which is the origin of replication (Ori) of the virus. The IR also contains the promoters of the V1, V2, C1 and C4 genes.

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Fig. 1 (a) Electron micrograph of TYLCV particles with twinned morphology of approximately 20
30 nm in size. (b) Non-infected and (b0 ) TYLCV-infected tomato plant with young leaves showing typical yellowing and curling. (c) Tomato leaflet infested with B. tabaci whiteflies; insert: close up of adult whitefly.

Fig. 2 Genome organization of Tomato yellow leaf curl virus. The single-stranded virion DNA comprises 2787 nt. Open reading frames (ORFs) of virion-sense and complementary-sense strand polarity are designated (V) and (C), respectively. ORFs are represented by an arrow; numbers indicate first and last nucleotide of each ORF. V1 encodes the capsid protein (CP), V2 a movement protein, C1 the replication initiator protein (Rep), C2 a transcriptional activator protein (TrAP), C3 a replication enhancer protein (REn), and C4 a symptom and movement determinant. IR: intergenic region. The conserved inverted repeat flanking the conserved nanonucleotide sequence TAATATTAC is symbolized by a stem-loop; an arrow head indicates the cleaving position of Rep in the TAATATT/AC loop; A at the cutting site (/) is nucleotide number one, by definition.

• • • • • •

V1 encodes the capsid protein (CP). The CP is multifunctional: monomers interact to form the capsid. It has a nuclear localization signal (NLS) and it is able to shuttle the viral genomic DNA in and out of the nucleus. It is essential for infectivity, and it is the only viral protein recognized by the insect vector. V2 has properties analogous to those of a movement protein. C1 encodes the replication initiator protein (Rep). The functional protein is an oligomer. Rep recognizes the Ori and specifically cleaves the nanonucleotide TAATATTAC between nucleotides 7 and 8. Together with plant host polymerase(s) it initiates viral DNA replication according to the rolling circle model (RCR) (see paragraph “Life cycle”). C2 encodes a transcription activator protein (TrAp). TrAp enhances transcription of the viral strand. It is able to bind to viral genomic DNA and possesses an NLS. In addition, it may act as silencing suppressors. C3 encodes the replication enhancer protein (REn). This protein interacts with itself to form oligomers. REn interacts with Rep and with cell-cycle associated host proteins to increase viral DNA amounts (genomic and double-stranded) in the infected plant. C4 encodes a protein not essential for infectivity but contributes to the spread of the virus in the plant and induction of symptoms. The protein may also act as a silencing suppressor.

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Bipartite TYLCVs Bipartite TYLCVs have a second genomic 2.7 kb component (coined DNA-B) with an ORF on the viral strand (BV1, encoding a nuclear shuttle protein) and another on the complementary strand (BC1, encoding a movement protein). Usually the DNA-A is enough to produce disease symptoms.

Satellite DNAs Associated With TYLCVs Monopartite begomoviruses, among them TYLCVs, are often associated with alphasatellites or betasatellites, two types of 1.3 kb circular ssDNA molecules. The only sequence homology between begomovirus and satellite DNAs is the nanonucleotide within the conserved stem-loop structure of begomovirus genomes required for RCR. Alphasatellites encode a RCR initiator protein (Rep). Betasatellite encodes the multifunctional protein bC1, which may be involved in virus movement, and suppression of gene silencing. bC1 may also enhance symptoms, virus amounts and whitefly transmission. Begomoviruses assist satellites for replication, and encapsidation. The introduction of betasatellites into western Mediterranean countries (e.g., Jordan, Israel, Egypt, and Arabian Peninsula) where satellites have never been reported before is a potential threat to tomato cultivation. Indeed, it was reported that TYLCV and an associate betasatellite overcome resistance provided by the Ty-1 gene.

TYLCV Genome Evolution Epidemiological recordings and phylogenetic analyses suggest that TYLCVs most probably arose in the Middle East between the 1930s and 1950s. The global spread of TYLCV only began in the 1980s after the evolution of the TYLCV and TYLCV-Mild strains. Other centers of TYLCV evolution have been shown to occur, for example in Iran; however because this region is geographically isolated, these TYLCV variants have not spread. TYLCV species often recombine. For example, TYLCV is a recombinant of TYLCV-Mild and Tomato leaf curl Karnataka virus (Table 1), while other begomoviruses from the Mediterranean basin are recombinants of TYLCV and TYLCV-Mild, and of TYLCV and Tomato yellow leaf curl Sardinia virus (TYLCSV). In Italy, Morocco and Spain, recombinations between TYLCV and TYLCSV occurred in tomato-growing areas. Recombinants were obtained in the laboratory by inoculating tomato plants with infective clones of two TYLCV species. Indeed, upon mixed infections, TYLCV and TYLCSV were detected in the same nucleus, a prerequisite for recombination.

Life Cycle During feeding on the host plant, whiteflies introduce TYLCV in the phloem with their stylets. The particles penetrate phloemassociated cells. The viral ssDNA genome is uncoated in the cell cytoplasm. Then, it is imported into the nucleus where it is converted to dsDNA (or replicative form RF) thanks to the host cell replication machinery. The dsDNA acts as a substrate for transcription of the viral mRNAs, which are exported to the cytoplasm for translation of the viral proteins required in the nucleus to carry out further replication and transcription. In the nucleus, the RF is duplicated several times according to the RCR model. Several host factors are required for progression and termination of the RCR fork. The IR has a hairpin structure with repeat sequences known as ‘iterons’. Replication is initiated by Rep binding the iterons followed by the specific nicking of the nanonucleotide TAATATT/AC (/: site of nicking). Rep binds covalently at the 5' end of nick, while the 3' end is used for extension of RCR. REn oligomers interact with Rep and enhance its ATPase activity. TYLCV Rep also acts as DNA helicase and as a site-specific type I topoisomerase, unwinding the DNA double helix in the IR for progression of the RCR fork. One round of RCR gives rise to a full viral dsDNA RF along with a genomelong viral ssDNA, which is released following Rep nicking. The new viral strand ssDNA is circularized using the closing activity of Rep and can be further converted to several dsDNA RF that serve as template for transcription/replication. From each RF molecule, several viral ssDNAs are produced. The replication process stops, perhaps with the increasing concentration of viral CP. The ssDNAs are either packaged in virions, or are transported out of the cell as nucleo-protein complexes. With the help of MP encoded by the virus, the transport occurs through the plasmodesmata into the neighboring cells, eventually leading to the long-distance spread within the host plant using its vascular system. Geminiviruses activate the host cell replication machinery for their own multiplication. Several plant factors interact with viral Rep and Ren to modulate viral DNA replication. The retinoblastoma related (RBR) protein, a plant homolog of the tumor suppressor retinoblastoma (pRb), controls the plant cells entry into S phase. RBR interacts with Rep, inducing the accumulation of both viral and host DNA. Rep is also associated with other host factors such as the Replication factor C (RFC) that helps generate a primer with 3'-OH terminus during initiation of DNA replication. Posttranslational modification processes like acetylation and sumoylation regulate various cellular processes, such as nuclearcytosolic transport, transcriptional regulation, apoptosis, protein stability, response to stress, and progression through the cell cycle. As such, they are targets of viruses, which aim at modifying the host environment. The viral Rep interacts with the host sumoylation enzyme, alters sumoylation of the host Proliferative cell nuclear antigen (PCNA) that acts as a cofactor for DNA polymerase δ, which

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enhances processing of the leading strand DNA synthesis during replication. Rep activity is reduced following the interaction with PCNA. In contrast, other host proteins may enhance the replication by Rep. For example, the host ssDNA-binding protein, Replication protein A (RPA) 32 kDa subunit (RPA32) directly interacts with the Rep C-terminus. RPA32 modulates the functions of Rep by enhancing its ATPase and decreasing its nicking and closing activities. The 3-D configuration of the genomic viral DNA may by itself guide the assembly of the particle from a pool of CP monomers. Once in the phloem, TYLCV virions can be picked up by whiteflies, circulate in the insect and may be used to infect new plants during feeding.

Hosts The domesticated tomato Solanum lycopersicum is the primary host of TYLCV. Most of the wild tomato species include accessions that either are symptomless carriers or are immune to the virus. In addition to tomato, several cultivated plants are hosts of TYLCV and present severe symptoms upon whitefly-mediated inoculation: bean (Phaseolus vulgaris), petunia (Petunia hybrida), and lisianthus (Eustoma grandiflorum). Pepper (Capsicum annuum) is a symptomless host of the virus. Weeds, such as Datura stramonium and Cynanchum acutum, present distinct symptoms while others, such as Malva parviflora, are symptomless carriers. Plants used to rear whiteflies in the laboratory, such as cotton (Gossypium hirsutum) and eggplant (Solanum melongena), are immune to TYLCV. Experimental hosts of the virus include jimsonweed (Datura stramonium) and Arabidopsis (Arabidopsis thaliana). Some plants, such as Nicotiana benthamiana and N. tabacum, which are recalcitrant to whitefly-mediated inoculation, may be inoculated with infectious TYLCV DNA clones using agroinoculation or DNA-coated micro-projectile bombardment.

Epidemiology Geographical Distribution and Spread The name Tomato yellow leaf curl virus (TYLCV) was coined in the early 1960s to describe a virus transmitted by the whitefly B. tabaci that affected tomato cultures in Israel. At the time, diagnosis of TYLCV was based essentially on symptom observation: border curling, yellowing and thickening of leaves, and plant growth stunting. The whitefly B. tabaci, has been instrumental in the worldwide spread of TYLCV. Whiteflies can fly for several km. They are very resilient and their eggs can sustain low temperatures. Global warming may facilitate the expansion of B. tabaci to presently cold regions devoted of whiteflies. Perennial weeds such as Cynanchum acutum and the annual weed Malva parviflora constitute TYLCV reservoirs in fields, especially outside crop growing seasons. A recent report claimed that TYLCV from South Korea is also transmitted through tomato seeds. This report needs to be confirmed. DNA sequencing has become the tool of choice allowing accurate identification of TYLCV isolates and their relationship with others. The virus first reported in Israel in the 1960s was isolated and sequenced in the late 1980s. Another, different, TYLCV species from Israel was coined TYLCV Mild and was sequenced in the early 1990s. TYLCV was reported in the mid and late 1970s in Cyprus, Jordan, and Lebanon. It was identified in Egypt and Turkey in the early 1980s. The virus has spread to Turkey, Iran and the Asian republics of the former USSR, and to the Arabian Peninsula during the mid-late 1990s. Two TYLCV isolates closely related to the Middle Eastern virus were described in Japan in 1996, from where they spread to South Korea where TYLCV was reported in 2008. In China, TYLCV isolates were identified in Shanghai in 2006 and have rapidly spread since to most of the country, thanks to invasive whitefly species. A local species was coined Tomato yellow leaf curl China virus (TYLCCNV). It was shown to possess a betasatellite. In the early 1990s two virus isolates belonging to a new species related to the Middle Eastern TYLCV, named Tomato yellow leaf curl Sardinia virus (TYLCSV) were identified in Sardinia and in Sicily, Italy. The Sardinian isolate was discovered in Spain in the early 1990s. By the mid 1990s the Middle Eastern TYLCV was found in Portugal and in Spain. In the latter country, it tended to displace the previously established Italian virus. Recombinants between the two viruses are not rare. TYLCV has invaded North Africa, possibly from Spain and Italy. In Morocco and Tunisia, both the Italian and the Middle Eastern strain were discovered in the early 2000s. TYLCV appeared in Southern France in 1999. In 2000, the Middle Eastern strain of TYLCV was identified in Greece. In East Africa, TYLCV was present in Sudan as early as the late 1970s. In Tanzania, a virus distinct from the Mediterranean isolates was identified. In the Réunion Island, TYLCV was detected in the late 1990s. The Middle Eastern strain of TYLCV has appeared in the Western Hemisphere in the mid 1990s in the Caribbean Islands, first in the Dominican Republic, then Cuba, Jamaica, Puerto Rico and the Bahamas. From there, the virus has reached the USA, identified first in Virginia in the late 1990s, then in Florida, Georgia, Louisiana, North Carolina and Mississippi. TYLCV was reported in the late 2000s in several regions of Mexico. In 2006, TYLCV was reported in the Brisbane area, Australia, likely imported from Japan. Hence, TYLCV has spread extremely fast and at present, constitutes a major limiting factor to tomato cultivation, worldwide. In many regions, the invading Mediterranean strain of TYLCV co-exists with local TYLCV strains but in several cases, it has almost replaced them, as in Southern Spain. Several introduction of TYLCV seemed to have occurred to the American/Caribbean region from the East-Asian, Australian, and Western Mediterranean regions, and several introductions of TYLCV to the East-Asian region

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from the Eastern Mediterranean and Western Mediterranean regions. The detection of TYLCSV in Jordan and in Israel, indicate that TYLCV spread may occur from the Western to the Eastern Mediterranean region and to other parts of the world.

TYLCV-Whitefly Vector Relationship Path of the virus in the insect Like all begomoviruses, TYLCV follows a well-defined path in its B. tabaci vector (Fig. 1(c)). Whiteflies feed when their stylets reach the virus-rich phloem of infected leaves. TYLCV can be detected in the insect head approximately 10 min after the beginning of feeding (acquisition access period AAP) on infected tomato. From the esophagus, the virus reaches the midgut where it can be detected about 30 min after it was first found in the head. At this point, some of the virus might be excreted through the hindgut and rectum (honeydew). The virus reaches the hemolymph 30 min after it was first detected in the midgut, 90 min after the beginning of the AAP. The crossing of the gut is likely to be an active process involving specialized, unknown, receptors. To avoid degradation in the hemolymph, TYLCV interacts with a GroEL-like chaperonin produced by the insect endosymbiotic bacteria and excreted in the hemolymph. TYLCV can be detected in the salivary glands approximately 5.5 h after it was first detected in the hemolymph, 7 h after the beginning of the AAP, approximately 1 h before the insects are able to infect tomato plants. Once the virus reaches the salivary gland, crossing several cell walls that may constitute selective receptor-mediated barriers, it is almost immediately excreted into the salivary pump and from there into the plant, together with the saliva. TYLCV transits with the same velocity in males and females. The virus capsid is the only structure recognized by the would-be receptors in the gut and salivary gland barriers, and by endosymbiotic GroEL.

Retention of the virus in the insect vector Once TYLCV is acquired, viral DNA remains associated with B. tabaci for the entire adult life of the insect, although infectivity decreases with time, but not entirely. TYLCV concentrates mainly in the filter chamber and the midgut. Various TYLCV isolates may present different parameters of interactions with their whitefly host. TYLCV, TYLCSV and TYLCCNV were detected with very low incidence in eggs and in the adult progeny of viruliferous whiteflies. The CP was shown to be essential for virus invasion of the female reproductive system of B. tabaci B and Q. Entry of TYLCV into eggs developing in ovaries is mediated by the interaction of CP with vitellogenin. The CP region necessary for crossing the gut into the hemolymph is also necessary for salivary glands and ovaries penetration. TYLCV and TYLCCNV were shown to be transmitted with very low frequency among whiteflies during mating.

Replication of the virus in the insect vector Replication of begomoviruses in their whitefly vector is a controversial issue. To prove, or disprove replication, viral DNA (genomic and RF) and CP amounts were usually appraised in whiteflies following a short AAP on TYLCV-infected plants and transfer to a non-host such as cotton, or on an artificial diet. In one set of experiments involving TYLCV, TYLCCNV and TYLCV-Mild, the amount of viral genomic DNA and its complementary strand, and of viral transcripts (especially from the viral genome complementary strand) increased with time, and then decreased, as measured by qPCR. These molecules were visualized by FISH in the midgut epithelial cell nuclei and in cells of the salivary glands. In situ fluorescent immunodetection and western blots showed increases in CP amounts. These results supported TYLCV replication in whiteflies. In another set of experiments, TYLCV and TYLCV-Mild did not accumulate following the end of the AAP. Moreover, whiteflies fed with purified TYLCV and TYLCV-Mild virions through membranes did not show any evidence of replication. These results led to the conclusion that TYLCV did not replicate in its vector.

Effects of TYLCV on life expectancy and fertility of the whitefly vector Several studies indicated direct and/or indirect influence of begomoviruses on B. tabaci, which can be detrimental, neutral or beneficial, depending on the virus-whitefly-plant combinations. In experiments performed in China, TYLCV infection of tomato indirectly improved the overall performance (higher survival, greater fecundity, shorter development time) of the B. tabaci Q biotype, whereas it reduced the overall performance of the B biotype. Other studies performed in China indicated TYLCV has little effect on the reproduction and development of Q whiteflies. In Israel, 3-days-old B whiteflies raised on eggplants (a TYLCV non-host) following a 48 h AAP on TYLCV-infected tomato plants showed a significant reduction in their life expectancy and a B50% decrease in the number of eggs laid. TYLCCNV also reduces female longevity and fecundity of B whiteflies. The negative impact of TYLCCNV was explained by TYLCCNV’s suppression of the whitefly immune responses by down-regulation genes in the Toll-like signaling and mitogen-activated protein kinase (MAPK) pathways. The deleterious effects on its whitefly vector, in addition to the invasion of the reproductive system, suggest that TYLCV has some features reminiscent of an insect.

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Pathogenesis In the field, it is common that tomato seedlings are infected immediately after planting. Infection is always associated with large amounts of whiteflies (Fig. 1(c)). Young leaves and shoot apices are the best targets for whitefly-mediated virus inoculation. In the field as well as the greenhouse, symptoms of TYLCV infection include severe stunting, reduction of leaf size, upward cupping/curling of leaves, chlorosis on leaves and flowers, and reduction of fruit production (Fig. 1(b) and (b)’). TYLCV can cause up to 90%–100% yield loss. In nethouses and greenhouses, infected plants are found near the (double) doors and close to ventilation openings.

Diagnosis Symptom observation The first step of diagnosis is examining symptoms, and inspecting the lower leaf surface for the presence of whiteflies. Early symptoms include yellowing, curling and thickening of young leaves. Later on, the plant becomes stunted and presents a bushylike growth.

Microscopy Light microscope observation of leaves infected with geminiviruses, including TYLCV, reveals characteristic large blue-violet nuclear inclusions following azure-A staining. Typical geminate particles can be observed with the transmission electron microscope (TEM) following virion purification. The virions can be decorated with anti-CP antibodies (immune electron microscopy) prior to observation under TEM. Particles can be TEM-observed in thin sections of infected leaves, using CP antisera and immuno-gold labeling. These methods are not of practical use for the detection of the virus in large numbers of plant samples.

Immunodetection Immunodetection of TYLCV includes ELISA and western blots. Particles are also visualized in preparations of plant and insect tissues using labeled antibodies. However, the antibodies raised against the viral CP (overexpressed in bacteria or from purified virions) are usually unable to distinguish TYLCV from closely related begomoviruses, and unable to discriminate between TYLCV species and strains.

Nucleic acid hybridization TYLCV genomic DNA and replicative dsDNA are readily detectable in DNA isolated from plants subjected to Southern blot hybridization, using a labeled DNA probe (preferably not radioactive). Similarly, TYLCV genomic DNA is detectable in viruliferous whiteflies. TYLCV DNA sequences are detected by hybridization of infected plant tissues and viruliferous whiteflies squashed onto a nylon membrane (squash-blot, or tissue-blot) with a labeled specific DNA probe. No treatment of the sample is necessary prior to squashing and hybridization. Large numbers of plant and insect samples can be collected from a field and analyzed in the laboratory. Viral RNAs are detectable in plant and insect preparations using fluorescently labeled (usually by Cy3 or Cy5) synthetic primers complementary to the RNAs. The samples are observed with a fluorescence microscope or a laser confocal microscope. A microarray containing 279 40-mer oligonucleotide probes was constructed to detect ten important tomato viruses. It included sequences derived from TYLCV CP DNA. It allowed detecting TYLCV but was unable to discriminate between TYLCV and TYLCSV.

Nucleic acid amplification PCR is widely used for the diagnosis of TYLCV strains. The virus is detectable in very small amounts of infected plant and vectors. Subsequently, the amplicon can be sequenced to identify the TYLCV strain. Whitefly GroEL, whether extracted from B. tabaci or overexpressed in E. coli, binds begomoviruses with strong affinity. This property was exploited to trap TYLCV by incubating infected plant and viruliferous whitefly extracts with GroEL-coated PCR tubes. TYLCV was then detected by PCR. RCA allows amplifying full-length viral genomes, including TYLCVs, in a single-step and without the use of specific primers. The amplicons can be sequenced for identification and phylogenetic analyses. RCA was implemented to generate infectious genomic clones directly from plant and whitefly DNA, and from specimens sometimes stored for decades, including TYLCV.

Sequencing Mass sequencing (millions of 100–700 nt reads) using platforms such as Illumina combined with bioinformatic softwares was used to study the effects of TYLCV on the transcriptome, and on virus-related RNAi and microRNA populations of plants and whiteflies.

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Metagenomic analyses of whiteflies and infected plants allowed delivering fast and inexpensive information on the presence of TYLCV isolates and other viruses (some previously unknown). This is an exquisite tool to survey the diversity and movement of TYLCV in small as well as large environmental settings.

Treatment Prevention Planting in whitefly-free period, eradication of inoculum Disease management strategies include the elimination of possible inoculum (residual infected plants and weeds), and planting when whitefly population pressure is low. In all regions where TYLCV is endemic, seedlings to be planted in the field or greenhouse need to be grown under cover to prevent early infection.

Cultures under cover Growing tomato in net or plastic houses covered with insect-proof 50-mesh screens and with double-doors, constitutes the optimal solution for crop protection against the TYLCV whitefly vector. Sticky yellow polyethylene sheets are used to decrease the pressure of the whitefly population in greenhouses, with limited success. Ultraviolet (UV)-absorbing plastic sheets or UV-absorbing 50-mesh screens used as nethouse covers, greatly repel whiteflies and reduce TYLCV incidence.

Chemical control of whiteflies TYLCV is usually controlled by spraying insecticides. Many chemicals used in the past, such as organophosphates, carbamates and pyrethroids, effectively reduced whitefly population, until the insects developed resistance. More potent insecticides, with novel modes of action, such as imidacloprid, have been introduced in the 1990s. As anticipated, whitefly populations resistant to imidacloprid have emerged in many places.

Biological control of whiteflies Whitefly IPM control in greenhouse tomato production of Europe, and to a lesser extent of the USA, includes biological control with natural enemies such as the parasitic wasp Eretmocerus spp. and the predatory bug Nesidiocorus tenuis, used alone or in combination. These practices are not yet common in the open-field cropping systems.

Breeding for Resistance Certain accessions of the wild tomato species S. chesmanii, S. chilense, S. habrochaites, S. peruvianum, and S. pimpinellifolium are resistant/tolerant to whitefly-mediated inoculation (show mild or no symptoms, but contain virus). Breeding programs aimed at producing tomato cultivars resistant to TYLCV started in the late 1960s. They were based on the introgression of resistance from wild tomato species into the domesticated tomato Solanum esculentum. The discovery of loci, and later on of genes, associated with TYLCV resistance, was facilitated by the development of saturated maps based on DNA polymorphism distinguishing resistance from susceptibility (RFLP, AFLP, SSR, SCAR, etc.). Altogether six loci, coined Ty-1 to Ty-6, were found to be associated with TYLCV resistance. The Ty-1 locus originating from S. chilense was mapped to tomato chromosome 6. Ultimately, Ty-1 was identified as coding for an RNA-dependent RNA polymerase. Another TYLCV resistance locus, Ty-3, was shown to be allelic to Ty-1. Ty-2 was introgressed from S. habrochaites and was located on chromosome 11. Ty-2 encodes a nucleotide-binding domain and leucine-rich repeat-containing (NB-LRR) gene. Ty-4 was introgressed from S. chilense and mapped to chromosome 3. Ty-5 was introgressed from S. peruvianum and mapped on chromosome 4. Ty-5 encodes a messenger RNA surveillance factor Pelota, which is involved in ribosome recycling phase of protein synthesis. Ty-6 originated from S. chilense and was located on chromosome 10. Combination of resistance loci from different sources in a single tomato line often enhances resistance. Today, most commercial tomato varieties are resistant/tolerant to TYLCV; in most cases, they have incorporated Ty-1. They remain symptomless (or with mild symptoms) upon infection, and yield nearly as uninfected plants (Fig. 3). However, Ty-1mediated resistance collapses under high disease pressure. It was recently reported in Jordan and in Israel that a newly found betasatellite associated with TYLCV overcomes the resistance based on Ty-1.

Genetic Engineering Expression of viral genes A variety of strategies have been devised based on the pathogen-derived resistance concept, which involves the expression of functional as well as dysfunctional viral genes. A transgenic interspecific F1 hybrid S. lycopersicum x S. pennellii expressing the TYLCV CP under the Cauliflower mosaic virus (CaMV) 35S promoter showed delay and remission of symptoms upon whitefly-mediated inoculation.

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Fig. 3 TYLCV-infected tomato fields planted with susceptible and resistant tomato genotypes. Note that the susceptible plants are stunted; they will not produce fruits. For comparison, the resistant plants will remain symptomless, or with mild symptoms and will yield.

Tomato and tobacco plants transformed with a truncated (2/5) TYLCV Rep gene under the control of the 35S CaMV promoter. Following whitefly-mediated inoculation with TYLCV in the laboratory, T2 generation lines evaluated in the field in Florida showed variable rates of resistance. However, the TYLCV-resistant plants were susceptible to the bipartite begomovirus Tomato mottle virus (ToMoV) from Florida. Similarly, tomato plants transformed with the Rep gene of TYLCSV truncated in its 30 end exhibited resistance to TYLCSV but were susceptible to the related strain from Spain. Expression of antisense sequence of the TYLCSV Rep gene in transgenic Nicotiana benthamiana resulted in lesser virus replication. The incorporation of a ribozyme structure into the antisense construct did not increase the efficiency of the system.

RNAi The silencing-based RNA interference (RNAi) antiviral defense mechanism was exploited to express siRNA directed against TYLCV genes. Post-translational gene silencing (PTGS) was activated through the transgenic expression of hairpin dsRNA cognate to the target viral genome. PTGS was efficient but was highly-sequence dependent. In the early experiments, non-coding conserved regions from the genome of TYLCV, TYLCV-Mild, and several TYLCSVs from Spain, were used to design constructs aimed at triggering broad resistance against different strains of TYLCV. Upon whiteflymediated inoculation, a high level of resistance was obtained against all three viruses, which correlated with the accumulation of TYLCV-specific siRNAs, indicating that PTGS can be used to engineer geminivirus-resistant plants. A hairpin RNAi (hpRNAi) aimed at expressing dsRNA homologous to sequences of the IR, the CP, V2 and Rep genes of TYLCV from Oman was used to transform tomato plants. Nine transgenic lines were obtained, which showed various levels of resistance upon TYLCV agroinoculation. RNAi directed at TYLCV Rep gene was expressed in transgenic tomato plants. The homozygous F4 and F6 generations exhibited resistance to TYLCV in the greenhouse and in whitefly-infested fields, and contained high amounts of small RNAs.

CRISPR/Cas9 The clustered regularly interspaced short palindromic repeats system (CRISPR/Cas), which protects prokaryotes from potentially deleterious foreign DNA, was used to engineer the genome of tomato and N. benthamiana to confer immunity to TYLCV. Targeting the TYLCV CP and Rep sequences with Cas9-single guide RNA resulted in efficient virus interference. The treated plants had low TYLCV load and their T3 progeny was tolerant to TYLCV. Therefore, genome editing might be the tool of the future to obtain TYLCV-resistant tomato genotypes.

TYLCV Transmission to Tomato and Host Response Transmission by whiteflies In nature, TYLCV is transmitted by the whitefly B. tabaci in a persistent circulative manner. Transmission of TYLCV to tomato by B. tabaci B biotype has been studied thoroughly. The virus is transmitted to tomato plants after vector feeding on infected tomato plants or alternative hosts. The incidence of the disease is directly correlated with the pressure of the whitefly population. One to three viruliferous insect are able to infect a tomato plant. The efficient AAP is 15–30 min, the latent period is 8–24 h, and the efficient inoculation access period (IAP) is at least 15 min. Female B. tabaci are better vectors than males. The ability of the whiteflies to transmit TYLCV to tomato test plants decreases with age, from 100% to 10%–20% during their 4–5-week adult lifetime.

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Response of plants to TYLCV infection RNAi RNAi is a natural defense mechanism against viruses. It is triggered by the processing of virus-derived dsRNA by host Dicer-like (DCL) enzymes into 21–24 nt small interfering RNA (siRNAs). Viral siRNAs bind Argonaute (AGO) proteins and guide the resulting RNAinduced silencing complexes toward their cognate RNA molecules, leading to post-transcriptional or transcriptional silencing in the case of RNA and DNA viruses, respectively. Viral siRNAs can move to other cells and into the phloem. As a counter-defense, geminiviruses encode one or more silencing suppressor proteins. TYLCV C2 and V2 proteins were reported to supress RNA silencing by inhibiting the siRNA amplification stage. Infection depends on the balance between silencing and suppression. This balance can be shifted to virus tolerance by overexpressing antiviral dsRNA in the form of RNA hairpins in transgenic plants. Aggregates Upon tomato infection, TYLCV CP-DNA complexes accumulate in the phloem associated cells of tomato leaves. At the early stages of infection, punctate aggregates are detected in the cytoplasm; in the later stages aggregates of increasing size are localized in cytoplasm and nuclei. Nuclear aggregates contained infectious particles transmissible to test plants by whiteflies. The formation of large aggregates was delayed in TYLCV-resistant tomatoes, perhaps as part of the resistance mechanism. Transcriptome and metabolome TYLCV induces significant changes already during the first days after inoculation, changes that exacerbate as infection progresses. They include the activation of genes involved in general stress-response, hormone biosynthesis, signal transduction, RNA regulation and processing, induction of the ubiquitination pathway, and initiation of autophagy. Photosynthesis decline, is accompanied by release of hexose that activates defense response or, if failing, promote pathogen replication and disease expression. Upon TYLCV infections, tomato plants emit reactive oxygen species (ROS), pathogenesis-related (PR), and wound-induced proteins. Sources of carbon and nitrogen are highly affected, accompanied by changes in the abundance of various classes of metabolites such as amino acids and polyamines, phenolic and indolic metabolites. Hence, upon TYLCV infection, there is a coordinated reprogramming of phenylpropanoid, tryptophan/nicotinate, urea/polyamine, and salicylic acid biosynthesis pathways leading to the production of defense compounds. Involvement of heat shock and quality control proteins in the establishment of TYLCV infection TYLCV does not induce a hypersensitive response and cell death upon whitefly-mediated infection of susceptible tomato plants. Cell death can be induced by the inactivation of heat shock protein 90 (HSP90) as well as by silencing the genes Hsp90 and Sgt1 (HSP90 co-chaperone), causing the dissociation of the 26S proteasome and a decrease of its peptidase activity, which led to the accumulation of damaged ubiquitinated proteins. TYLCV alleviates cell death and these accompanying effects to create an environment suitable to its replication. The interactions between TYLCV and HSP70 and HSP90 chaperons in plants and vectors are essential for virus infection to proceed. In infected host cells, HSP70 and HSP90 are redistributed from a soluble to an aggregated state. These aggregates contain, together with viral DNA/proteins and virions, HSPs and components of the protein quality control system such as ubiquitin, 26S proteasome subunits, and the autophagy protein ATG8. TYLCV CP can form complexes with HSPs in tomato and whitefly. Nonetheless, HSP70 and HSP90 play different roles in the viral cell cycle in the plant host. In the infected host cell, HSP70, but not HSP90, participates in the translocation of CP from the cytoplasm into the nucleus. Viral amounts decrease when HSP70 is inhibited, but increase when HSP90 is downregulated. In the whitefly vector, HSP70 impairs the circulative transmission of TYLCV; its inhibition increases transmission.

Further Reading Brown, J.K., Zerbini, F.M., Navas-Castillo, J., et al., 2015. Revision of Begomovirus taxonomy based on pairwise sequence comparisons. Archives of Virology 160, 1593–1619. CABI, 2019. Invasion Species Compendium. Tomato Yellow Leaf Curl Virus (Leaf Curl). Available at: https://www.cabi.org/isc/datasheet/55402. Czosnek, H., 2007. Tomato Yellow Leaf Curl Virus Disease: Management, Molecular Biology, Breeding for Resistance. Dordrecht: Springer. Czosnek, H., Hariton-Shalev, A., Sobol, I., Gorovits, R., Ghanim, M., 2017. The incredible journey of begomoviruses in their whitefly vector. Viruses 9, 273. Gorovits, R., Czosnek, H., 2017. The involvement of heat shock proteins in the establishment of Tomato yellow leaf curl virus infection. Frontiers in Plant Science 8, 355. Hanley-Bowdoin, L., Bejarano, E.R., Robertson, D., Mansoor, S., 2013. Geminiviruses: Masters at redirecting and reprogramming plant processes. Nature Reviews Microbiology 11, 777–788. Mabvakure, B., Martin, D.P., Kraberger, S., et al., 2016. Ongoing geographical spread of Tomato yellow leaf curl virus. Virology 498, 257–264. Snehi, S.K., Raj, S.K., Prasad, V., Singh, V., 2015. Recent research findings related to management strategies of Begomoviruses. Journal of Plant Pathology and Microbiology 6, 273. Zhou, X., 2013. Advances in understanding begomovirus satellites. Annual Review of Phytopathology 51, 357–381.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/ssdna-viruses/w/geminiviridae/392/genus-begomovirus Genus: Begomovirus.

Tombusvirus-Like Viruses (Tombusviridae) K Andrew White, York University, Toronto, ON, Canada r 2021 Elsevier Ltd. All rights reserved.

Nomenclature ', Prime symbol c Lowercase gamma (Greek)

Å

Glossary Frameshifting A ribosome shifting its reading frame during protein translation. Higher-order RNA structures RNA secondary and tertiary structures, such stem-loops or pseudoknots, respectively. Monocot A class of angiosperm plant with a single embryonic leaf and parallel veins. Nanoparticle A microscopic particle in the approximate range of 1–100 nm.

Angstrom

Open reading frame A segment of RNA encoding a protein. T Triangulation number, which describes the icosahedral geometry of spherical viruses. Tombusvirid A member of the virus family Tombusviridae. Transcription Herein, describes the process by which viral sg mRNAs are synthesized. Type A designated member that represents a group.

Introduction Tombusvirus is the type genus in the family Tombusviridae, which currently includes a total of sixteen genera. The members of this extensive family possess linear, single-stranded, positive-sense RNA genomes that are packaged into spherical particles and, collectively, infect a wide variety of plant hosts. The overall grouping of these viruses is based on the presence of a conserved RNA-dependent RNA polymerase (RdRp). Within the family, members are further subdivided into subfamilies, genera, and species based on similarities in gene expression strategies and encoded proteins. In this article, the genus Tombusvirus and those genera that are most closely related to this genus, i.e., Aureusvirus and Zeavirus, are described.

Taxonomy, Phylogeny, and Evolution The family Tombusviridae resides within the realm Riboviria, and includes three subfamilies (Procedovirinae, Regressovirinae and Calvusvirinae) and sixteen genera (Alphacarmovirus, Alphanecrovirus, Aureusvirus, Avenavirus, Betacarmovirus, Betanecrovirus, Dianthovirus, Gallantivirus, Gammacarmovirus, Macanavirus, Machlomovirus, Panicovirus, Pelarspovirus, Tombusvirus, Umbravirus, and Zeavirus), each of which contain member species that range in number from one to seventeen. Most genera belong to the subfamily Procedovirinae, all of which express their RdRps via translational readthrough of a stop codon. In contrast, dianthoviruses are in Regressovirinae, because they translate their RdRps via frameshifting. Umbraviruses, which do not encode a capsid gene, occupy Calvusvirinae. The current taxonomy is the result of recent reorganization within the family, and additional modifications and refinements are anticipated in the near future. Consequently, updates should be monitored by visiting the International Committee on Taxonomy of Viruses website (see “Relevant Websites section”). Taxonomic classification of tombusvirids is based primarily on the structural relationship of their viral RdRps. All contain supergroup-II RdRps that share high levels of amino acid sequence identity, including several tombusvirid-specific motifs located in C-terminal regions. Indeed, taxonomic organization down to the level of species can be guided entirely by sequence comparisons of RdRps. Nonetheless, additional features distinct to specific genera, such as the type of movement protein or silencing suppressor encoded, also aid in appropriate grouping. The notable level of diversity observed within these two protein types suggests that they were acquired via recombination-mediated modular transfer of gene cassettes. Phylogenetic analysis of the RdRps of tombusvirids have revealed an evolutionary tree that is comprised of distinct lineages (Fig. 1). The largest two of these lineages can be defined roughly as either tombusvirus-like or carmovirus-like, with the latter further classified into numerous genera. Only four genera reside within the tombusvirus-like clade, Tombusvirus, Aureusvirus, Zeavirus and Betanecrovirus. However, the latter genus has movement and capsid proteins typical of carmoviruses. Consequently, betanecroviruses are not discussed here, and instead are presented in the article covering necrovirus-like viruses. Of the remaining three tombusvirus-like genera, all share high degrees of similarity with respect to gene order, protein relatedness, and expression strategies. Based on RdRp phylogenetic analysis, zeaviruses are predicted to be more closely related to tombusviruses than aureusviruses. However, the zeavirus host range of monocot plants (e.g., grasses) is more akin to certain aureusviruses. Thus, although these genera are closely related, they also possess features that distinguish them from one another. Currently, the genus

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Fig. 1 Phylogenetic tree depicting evolutionary relationships of genera in the family Tombusviridae based on the RdRp amino acid sequences of tombusvirids. The tombusvirus-like lineage is shaded in yellow and the genera discussed in this article are shown in red. Adapted from Gunawardene, C.D., Donaldson, L.W., White, K.A., 2017. Tombusvirus polymerase: Structure and function. Virus Research 234, 74–86.

Tombusvirus is composed of seventeen species, including Tomato bushy stunt virus (TBSV), which is the type virus of both the genus and family (Table 1). The less crowded Aureusvirus genus has five confirmed member, with Pothos latent virus (PoLV) designated as the type species. Presently, the genus Zeavirus contains only one member, its type species, Maize necrotic streak virus (MNeSV).

Virion Structure, Assembly, and Disassembly Members of the tombusvirus-like group form non-enveloped, spherical virions that encase a single viral RNA genome. The capsids are B30 nm in diameter, have T¼3 icosahedral symmetry, and are composed of 180 subunits of a capsid protein (Fig. 2). The capsid proteins of tombus- and aureusviruses are of similar molecular masses, 40 and 41 kDa, respectively. In contrast, that for zeaviruses is substantially smaller, 27 kDa, due to the absence of a C-terminal domain present in the other two capsid proteins. Among the three tombusvirus-like genera, the particles of tombusviruses have been studied most extensively. Going from N- to C-terminus, these capsids are composed of three distinct structural domains, RNA-binding (R), shell (S), and protruding (P). Orientation of the subunits within a capsid is such that C-terminal P-domains extend out from the particle surface formed by the S-domain, while the N-terminal R-domain, a highly basic region, is positioned internally and neutralizes the negatively charged phosphate groups in the packaged RNA genome. The single capsid subunit is able to assume three different conformations that allow for its packing into the 5- and 3-fold centers of particle symmetry. Only conformation A subunits reside at the 5-fold axes (blue), while alternating B and C subunits (red and green, respectively) fill the 3-fold centers (Fig. 2). The P-domains of two adjacent subunits interact in AB or CC dimers to form ninety projections that can be observed in electron micrographs. This latter feature is not present in zeaviruses, as their capsid proteins lack the C-terminal P-domain that forms these protrusions. Details of the process of virion assembly in these viruses remain largely unknown. For many viruses, key regions of their RNA genomes have been identified as packaging signals that bind to capsid subunits and initiate virion assembly. Currently, no corresponding RNA assembly signals in the genomes of tombusvirus-like viruses have been identified. However, studies on the capsid protein of the tombusvirus Cucumber necrosis virus (CuNV) have identified a determinant in the R-domain, a basic KGKKGK motif, that dictates the efficiency of capsid assembly, as well as particle size, i.e., T¼1 (60 subunits) versus T¼3 (180 subunits). Furthermore, a second basic

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Table 1 List of current tombusvirus-like viruses grouped by genera. The type species of each genus is indicated in bold and the corresponding abbreviation for each virus is provided Genus

Species

Abbreviation

Aureusvirus

Pothos latent virus Cucumber leaf spot virus Johnsongrass chlorotic stripe mosaic virus Maize white line mosaic virus Yam spherical virus

PoLV CLSV JCSMV MWLMV YSV

Tombusvirus

Tomato bushy stunt virus Artichoke mottled crinkle virus Carnation Italian ringspot virus Cucumber Bulgarian latent virus Cucumber necrosis virus Cymbidium ringspot virus Eggplant mottled crinkle virus Grapevine Algerian latent virus Havel River virus Lato River virus Limonium flower distortion virus Moroccan pepper virus Neckar River virus Pelargonium leaf curl virus Pelargonium necrotic spot virus Petunia asteroid mosaic virus Sitke waterborne virus

TBSV AMCV CIRV CBLV CuNV CymRSV EMCV GALV HaRV LRV LFDV MPV NRV PLCV PENSV PeAMV SWBV

Zeavirus

Maize necrotic streak virus

MNeSV

Fig. 2 TBSV virion structure. Image of TBSV particle at 2.9 Å , which is typical of tombusvirus and aureusvirus virions. Subunit conformations are color coded: A (blue), B (red) and C (green). Image reproduced from http://viperdb.scripps.edu. Ho, P.T., Montiel-Garcia, D.J., Wong, J.J., et al., 2018. VIPERdb: A tool for virus research Annual Review of Virology 5 (1), 477–488.

motif in the arm region connecting the R- and S-domains is important for determining the amount of full-length RNA that gets incorporated into virions and may contribute to the specificity of packaging viral genomes. Accordingly, interactions involving these positively-charged interior regions of the capsid and the negatively-charged viral RNA are proposed to mediate virion formation; the detail of which remain to be determined. An equally important aspect of virion function is its ability to disassemble and release its viral genome in newly infected cells. Studies with the tombusvirus CuNV suggest that plant Hsp70 protein chaperone homologs, Hsc70 or Hsc70-2, assist in CuNV disassembly. Hsc70-2 copurifies with CuNV particles, overexpression of Hsc70 in plants facilitates infections, and incubation of Hsc70 with particles destabilizes them. These observations are consistent with the involvement of Hsc70 or its homologs in

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Fig. 3 Cartoon representation of the plus-strand RNA genomes of the type species for tombusvirus-like genera: Tomato bushy stunt virus (TBSV), Pothos latent virus (PoLV) and Maize necrotic streak virus (MNeSV). Encoded proteins are represented by color-coded boxes along with the molecular masses of corresponding proteins (p), in kDa. The functions of proteins are labeled for the TBSV genome, with corresponding proteins in the other two genomes having equivalent functions. RT indicates the position of the stop codon for accessory replication proteins in the genomes where translation readthrough occurs to produce their RNA-dependent RNA polymerases (RdRps).

mediating disassembly of this tombusvirus. Moreover, since Hsp70 homologs also associate with virions of the aureusvirus Cucumber leafspot virus, and overexpression of Hsc70-2 in host plants mediates more robust infections, this proposed mode for unpackaging may also apply to this related genus.

Genome Structure The linear, plus-sense, single-stranded RNA genomes of tombusvirus-like viruses are neither 50 -capped nor 30 -polyadenylated. The sizes of these genomes are B4.1, B4.2, and B4.8 kb in length, respectively, for zea-, aureus-, and tombusviruses (Fig. 3). These genomes encode five proteins, and share a similar gene composition and organization that is unique among tombusvirids. Encoded 50 -proximally is an auxiliary RNA replication protein, homologs of which are present at comparable genomic positions in all members of the family. Translational readthrough of the stop codon of this protein leads to the production of a supergroup-II RdRp, also a common product in all tombusvirids. Encoded downstream from the polymerase is a single capsid protein, which contains a P-domain in tombus- and aureusviruses that is absent in zeaviruses. The feature that clearly distinguishes the tombusvirus-like group from other genera is their movement and suppressor of gene silencing proteins. These two proteins are encoded as overlapping open reading frames (ORFs) and are positioned 30 -proximally in viral genomes (Fig. 3). The coding regions of the suppressor proteins are smaller than, and are located entirely within, corresponding movement protein ORFs. However, because the two ORFs reside in different reading frames, each encodes a structurally unique protein. Homologs of these two viral products are not found in other tombusvirids, thereby defining these genera as a distinctly-related trio. In addition to their role in encoding viral proteins, these plus-strand RNA genomes also house multiple RNA elements that are involved in regulating different viral processes. Some of these elements are local, such as the promoter sequences located at the 30 -termini of these genomes. Interestingly, tombusviruses also utilize at least six different sets of intragenomic, long-range, base pairing RNA-RNA interactions for different viral functions. Some of these span almost the entire length of these 4.8 kb genomes, whereas others traverse shorter distances, while still exceeding a kilobase in length. The presence of these interactive RNA-RNA networks suggests that there must be a significant level of structural organization in these viral genomes to assist bringing distal partner regions together and to coordinate the different interactions and their corresponding functions. Indeed, analysis of the RNA secondary structure of the complete TBSV genome revealed that much of it is ordered and forms discrete RNA domains (Fig. 4). Notably, the predicted structure included only two of the six long-range interactions needed during viral infections (i.e., AS1-RS1 and DE-CE) (Fig. 4). This suggests that dynamic reorganization of the genome is required to achieve the six different long-range interactions (Fig. 5(A)). Such structural rearrangements could be facilitated by viral or host factors that are able to direct particular RNA folding pathways. In this regard, the auxiliary RNA replication protein p33 of TBSV has been found to possess RNA chaperone activities, and thus could participate in such genome reorganizing events.

Gene Function Studies exploring protein function in the group have been carried out largely in tombusviruses (e.g., TBSV), however the corresponding proteins in the other two genera are predicted to operate similarly. Both of the 50 -proximally encoded proteins in these viral genomes are essential for viral RNA synthesis. The auxiliary RNA replication protein of TBSV, p33, is a multifunctional master regulator of viral RNA replication that interacts with a myriad of host factors. This protein is also a major structural component of virus replication complexes (VRCs), the sites of viral RNA synthesis. VRCs also contain an RdRp, p92 for TBSV, that catalyzes both replication of the genome and transcription of viral subgenomic (sg) mRNAs. Additional information on these two processes are provided in subsequent sections.

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Fig. 4 RNA secondary structure model of the TBSV genome. RNA domains within the genome are labeled and color-coded according to their relative sizes: large (green), medium (red) or small (blue). Inter-domain regions are depicted in black. The RNA segments that participate in known functional long-range RNA–RNA interactions are labeled (refer to Fig. 5 for details). Only the AS1-RS1 and DE-CE interactions involved in sg mRNA transcription are base paired in this genome structure. Adapted from Wu, B., Grigull, J., Ore, M.O., Morin, S., White, K.A., 2013. Global organization of a positive-strand RNA virus genome. PLoS Pathogens 9 (5), e1003363.

Fig. 5 TBSV RNA genome and sg mRNAs. (A) TBSV genomic RNA, with encoded proteins represented by boxes. Proteins translated directly from the genome, p33 and p92, are represented by grey bars and sg mRNA transcription start sites within the genome are labeled sg1 and sg2. Long-range RNA-RNA interactions involved in different viral processes are indicated by double-headed arrows above the genome and are color coded: 50 UTR-30 CITE, translation initiation; PRTE-DRTE, ribosome readthrough; UL-DL, genome replication; AS1-RS1, sg mRNA1 transcription; AS2-RS2, sg mRNA2 transcription. DE-CE is a second long-range interaction needed for sg mRNA2 transcription. (B) TBSV sg mRNAs. The two sg mRNAs transcribed during infections are shown along with the proteins that are translated from them.

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Movement proteins are necessary for cell-to-cell movement of viral infections in plants. The local spread of an infection occurs through plasmodesmata, which are channels that connect adjacent plant cells. This process requires the assistance of a virally-encoded movement protein that facilitates transport of viral RNA to, and through, these intercellular junctions. This event requires modification of the structure of plasmodesmata to allow transit and, in tombusviruses, this action is mediated by p22. Properties of p22, such as binding to RNA and localization to cell membranes and walls, are consistent with this role. The viral genome is likely transported intra- and intercellularly as a p22-RNA complex, and may be shuttled by host factor HFi22, a homeodomain-containing protein, that interacts with p22. This concept is supported by the observation that some movement-deficient p22 mutants are no longer able to interact with HFi22. Gene silencing is an antiviral mechanism utilized by plants. The first step in this defence mechanism occurs when long doublestranded (ds) viral RNAs generated during genome replication are bound and cleaved by the host protein dicer into small dsRNAs, termed small interfering RNAs (siRNA). Subsequently, single-strands of these siRNAs are loaded into a RNA-induced silencing complex (RISC) that uses them as guides to base pair with and cleave viral genomes. To counteract this attack, plant viruses encode suppressors of gene silencing that are able to inhibit different steps in the process. The p19 suppressor of tombusviruses functions as a homodimer that binds size-specifically to siRNAs generated by dicer. In doing so, it prevents them from being loaded into RISC, thereby preventing virus genome-specific RNA targeting and cleavage by this complex. The smaller p14 homologue in aureusvirus functions in a similar manner but, due to its lack of size-specificity, it also binds to longer dsRNAs.

Gene Expression Gene expression in these viruses involves cap-independent translation, ribosome readthrough, sg mRNA transcription, and leaky scanning. Intriguingly, for function, all but the latter activity require RNA-RNA interactions that span large distances within the viral genomes (Fig. 5(A)). All tombusvirid RNA genomes lack a 50 -cap and a 30 -poly(A) tail. Instead, they contain one of several types of 30 -cap independent translational enhancer (30 -CITE) in their 30 -untranslated regions (30 -UTRs). 30 -CITEs in the tombusvirus-like group are higher-order RNA structures that bind to translation initiation factor 4F (eIF4F). These 30 -CITEs, while bound to eIF4F, also interact with the 50 -UTRs of their genomes, via a long-range RNA-RNA base pairing interactions (Fig. 5(A)). This positions the bound eIF4F near the 50 -end of the genomes, allowing for 43S subunit recruitment that mediates translation of the auxiliary replication proteins, p33 in the case of TBSV (and the p92 RdRp via readthrough). Though closely related, the three genera contain different types of 30 -CITEs; tombusviruses have Y-shaped versions (except for, Cucumber Bulgarian latent virus, CBLV, which has an I-shaped type), aureusviruses contain either a PTE or I-shaped class, while zeaviruses contain only I-shaped versions. Accordingly, modularity in these viral genomes also exists with respect to translational regulatory RNA elements. Viral RdRps, such as p92 in TBSV, are expressed by translational readthrough of the stop codon for the auxiliary replication protein. Programmed ribosome readthrough occurs when RNA elements in a message promote the use of near-cognate tRNAs for decoding of a termination codon. In tombusviruses, this process requires an extended stem-loop RNA structure, termed RTSL, positioned just 30 to the stop codon. However, in order for readthrough to occur, a sequence in RTSL (termed PRTE) must base pair, via a long-range RNA-RNA interaction, with a segment in the 30 -UTR of the genome (DRTE) (Fig. 5(A)). This interaction shuts down minus-strand RNA synthesis during readthrough, thereby preventing a conflict between these processes that occur in opposite directions on the viral genome. Similar local and long-distance RNA structural requirements for readthrough production of RdRp are also predicted for aureusviruses and zeaviruses. Within the context of tombusvirus-like genomes, the three ORFs encoded in their latter halves are translationally silent. Consequently, expression of these gene products requires the transcription of viral sg mRNAs. During infections, two sg mRNAs are synthesized that encode at their 50 -ends either the capsid protein ORF or the two overlapping ORFs. The capsid protein is translated from the larger sg mRNA1, while both the movement and silencing suppressor are translated from sg mRNA2 (Fig. 5(B)). The latter protein is translated when 43S subunits scan past the more 50 -proximally positioned movement protein start codon and initiate at the downstream suppressor protein start codon, a mechanism referred to as leaky scanning. Transcription of sg mRNAs involves premature termination of the RdRp while synthesizing complementary genomic minus-strands. RdRp termination occurs when the viral polymerase encounters certain higher-order RNA structures in a genome, called attenuation structures. This results in truncated minus-strands that include transcription initiation sites, for either sg mRNA1 or 2, at their 30 -ends. These sg mRNA-sized minus-strands are then used as templates to transcribe the two different size classes of sg mRNAs. Interestingly, in tombusviruses and zeaviruses, the attenuation RNA structures for both sg mRNA1 and 2 are formed by long-range, base pairing, RNA-RNA interactions (AS1-RS1 and AS2-RS2, respectively) (Fig. 5(A)). These genome-level interactions may provide an RNA-based mechanism to coordinate transcription with other viral processes. Aureusviruses have comparable long-distance AS2-RS2 interactions for sg mRNA2 production, but, for sg mRNA1, the attenuation structure is formed by a locally-folded section of RNA. Although variations in attenuation structures exist, the sg mRNAs generated are structurally similar in that all are 30 -coterminal with their cognate genomes. This feature ensures that these smaller viral messages contain 30 -CITEs, which enhance their translation via cognate 50 UTR-30 CITE interactions.

Genome Replication Tombusvirus genome replication has been investigated extensively in different systems, including in vitro cell extracts, plant protoplasts, whole plants, and surrogate yeast cells. These studies, particularly those carried out in yeast-based systems, have

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Fig. 6 Steps in VRC assembly and RNA synthesis in TBSV. (A) Peroxisome membrane targeting, (B) VRC assembly and (C) Viral RNA synthesis. Plus- and minus-strand genomes are represented as a solid and dotted lines, respectively. RdRps are shown at different steps in viral genome replication in (i) through (iv). See key for the identity of molecular components and refer to text for details. Diagram is not to scale. Adapted from Gunawardene, C.D., Donaldson, L.W., White, K.A., 2017. Tombusvirus polymerase: Structure and function. Virus Research 234, 74–86.

generated a large body of information that has allowed for the development of detailed models for various steps in this process. In general, the overall progression during infections can be roughly divided into three phases (1) membrane targeting, (2) VRC assembly, and (3) viral RNA synthesis. Here, these fundamental steps are summarized along with descriptions of the key viral and cellular factors involved, using TBSV as the example (Fig. 6). At the centre of this process are the viral proteins p33 and p92, with the former being responsible for orchestrating the majority of events required for productive RNA genome replication. p33 is a membrane-anchored protein that possesses multiple domains and motifs that allow it to interact with diverse host factors that are required at different steps in the process. Also important for p330 s activity is its ability to multimerize and interact with p92, and an early stage in RNA replication involves the targeting of these two viral proteins to specific intracellular membranes which, for TBSV, are peroxisomal (Fig. 6(A)). Membrane localization is dependent on N-proximal targeting sequences present in both of these proteins as well as interaction with peroxisomal biogenesis factor 19 (pex19). The viral genome also has to be shuttled to this membrane location, and this is accomplished by p92/p33 binding to an RNA structure, RII, located in the p92 coding region (Fig. 6(A)). Assembly of VRCs rely heavily upon p33 activity. VRCs are invaginations of intracellular membranes that maintain a narrow neck with the cytosol. p33 multimers are a key structural component of these so-called spherules, and their internal compartments house the

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RdRp and other factors necessary for viral RNA synthesis (Fig. 6(B)). A key step in forming these curved compartments is the hijacking of the host endosomal sorting complexes required for transport (ESCRT). p33 redirects the ESCRT system to form VRCs by interacting with different key components, such as Vps23p. The nature of the lipids within VRC membranes is also important for function. Phosphatidylethanolamine and phosphatidylcholine have stimulatory effects on VRC activity (Fig. 6(B)). To this end, p33 upregulates phospholipid production by interacting with and modulating regulatory proteins Opi1p and Scs2p. Other membrane components, such as sterols, which decrease membrane fluidity and directly bind to p33, also increase VRC function. Here again, p33 is involved in seizing control of these molecules by binding to and modifying the activity of oxysterol-binding proteins (ORPs), which are involved in controlling sterol distribution between closely associated membranes. Actin filaments also contribute to this process by stabilizing membrane contact sites, which allows for exchange of sterols from the endoplasmic reticulum (ER) to peroxisomal membranes that are destined for VRC formation. Stable actin filaments are required for this to occur, and an inhibitory interaction of p33 with cofilin, an actin depolymerization factor, ensures that a polymerized state is maintained. ER-peroxisomal intermembrane junctions thus represent critical cellular hubs for preparing appropriately composed lipid bilayers for VRC assembly. The p92 RdRp inside assembled VRCs synthesizes a full-length complementary negative-strand of a genome that is then used repeatedly as a template to produce multiple copies of progeny genomes (Fig. 6(C)). However, prior to any RNA synthesis, p92, in association with a phospholipid-enriched membrane, has to be activated by cellular protein chaperone Hsp70. Other host proteins are also needed during genome replication. Two translation elongation factors, eEF1A and eEF1Bg, are involved in the early stages of minus-strand synthesis (Fig. 6(C)(i)). Both bind to a promoter region in 30 -end of the genome and facilitate RdRp positioning and promoter access, respectively. Additionally, p92 that is bound to the RII RNA structure in RdRp-coding region of the viral genome is repositioned to the 30 -end by a long-range RNA–RNA interaction, UL-DL (Fig. 5(A)). p92 initiates RNA synthesis at a promoter sequence located at the 30 -end of the viral genome (Fig. 6(C)(ii)). Upon completion of minus-strand genome synthesis, the RNA helicase RH20 mediates release of p92 from the dsRNA intermediate, and facilitates its rebinding to the minus-strand template. This latter step is also assisted by two other RNA helicases, RH2 and RH5, as well as glyceraldehyde 3-phosphate dehydrogenase (GAPDH), all of which interact with the minus-strand (Fig. 6(C)(iii)). Notably, the activities of these helicases, as well as Hsp70, require ATP. This requirement is met by the recruitment of glycolytic enzymes, namely ATP-generating phosphoglycerate kinase and pyruvate kinase, to the vicinity of VRCs. The resulting activities of these enzymes leads to high local levels of ATP for components of the VRCs, thereby allowing for robust virus genome replication. Newly synthesized progeny viral genomes leave VRCs and enter the cytosol via the neck of spherules (Fig. 6(C)(iv) and (v)).

Subviral RNAs Tombusviruses are known to associate with two distinct types of subviral RNAs, satellite RNAs (satRNAs) and defective interfering RNAs (DI-RNAs). SatRNAs are naturally-occurring, noncoding, small linear RNAs that require coinfection with a standard virus, termed helper virus, for their replication and packaging. They are also dependent on their helper virus for gene silencing suppression and movement functions. Structurally, satRNAs do not share sequence identity with their helper virus. The discrete sequence of these RNAs is a paradoxical feature, because they utilize the same RdRp and capsid protein as their helpers. This suggests that they may engage viral and host factors in unique ways. SatRNAs have been found only in tombusvirus infections. Four distinct satRNAs have been identified, with the first being satRNA-Cym, which was discovered in association with Cymbidium ringspot virus (CymRSV). Subsequently, three others were identified in different TBSV infections, satRNA-B1, B10, and L. These satRNAs have lengths in the range of 600–800 nt and maintain 50 -ends that form higher-order RNA structures similar to those found in the 50 -UTRs of their helper genomes. When present in infections, satRNAs can increase, decrease, or have no effect on symptoms. Of the four tombusvirus satRNAs, only satRNA-B10 modulates infections; i.e., it attenuates symptoms and prevents death of infected plants. The other class of subviral RNA associated with tombusvirus infections are DI-RNAs. Similar to satRNAs, DI-RNAs are small, parasitic, noncoding, linear RNAs that depend on their helper virus for replication and packaging. However, unlike satRNAs, DI-RNAs are not normally found in natural infections, and they contain sequences that correspond directly to those in their helper virus genomes. DI-RNAs associated with tombusviruses were the first to be discovered for any plant virus. These 400–600 nt long RNAs contain sort genomic segments from both termini and two different internal regions (RI through RIV), and these segments are either essential (RI, RII and RIV) or greatly enhance (RII) replication of the DI RNA. This knowledge of DI-RNA structure and function also aided the identification of corresponding RNA replication elements in full-length viral genomes. Moreover, due to their small size, efficient accumulation in coinfections, and absence of coding functions, DI-RNAs have been excellent tools for studying virus RNA replication; particularly in the yeast-based system. They have also provided insights into viral RNA recombination, a process mediated by template hopping or switching by the viral RdRp during RNA synthesis. DI-RNA formation, via recombination, and their subsequent accumulation is promoted when infections are passaged at high virus titer; an artificial condition that favors coinfection of cells with helper genomes and nascently generated DI-RNAs. Although they are not found in infections in the wild, adding DI-RNAs to tombusvirus infections in laboratory experiments revealed their ability to suppress helper virus levels and the symptoms induced in plant hosts. This interference was initially believed to be due primarily to competition with the helper virus for RNA replication factors. However, tombusvirus DI-RNAs have also been shown to strongly activate gene silencing in coinfections, while being highly resistant to its effects. Thus, the disease attenuation observed could be caused by enhanced targeting and degradation of the helper genome.

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A third class of subviral RNA, a satellite virus (SV), is present in some infections of the aureusvirus Maize white line mosaic virus (MWLMV). SV have properties similar to those of satRNAs, except that they encode their own unique capsid protein, in which their RNA is packaged. The linear RNA genome of SV-MWLMV is B1.2 kb in length and encodes a 24 kDa capsid protein. Together, these components form spherical virions of B17 nm in diameter, which are about half the size of the B35 nm virions of the helper MWLMV. These particle sizes are consistent with corresponding T¼1 and T¼ 3 icosahedral symmetries. Although some SVs are known to modulate symptoms induced by their helper virus, the possible effects of SV-MWLMV on MWLMV infections have not been reported.

Symptoms and Transmission Symptoms induced by tombusviruses range from mild to severe, depending on the host. Local effects at sites of infection include chlorotic or necrotic lesions, or mosaic patterns. TBSV infections of tomato plants cause mottling and crinkling of infected leaves, along with stunted growth. Infections that go systemic result in necrosis of apical leaves and cause deformation and necrosis of fruit. Aureusvirus infections range from symptomless, such as infections of pothos plants by PoLV, to systemic mottling and fruit streaking in cucumber infections by Cucumber leaf spot virus (CLSV). Maize infected with the MNeSV zeavirus initially exhibit pale green or yellow spots and streaks on leaves. But, as the infection progresses, the chlorotic bandings expand and merge on leaves and stalks, with some areas becoming necrotic. Both tombusviruses and aureusviruses accumulate to high levels in infected plants and, as soil-borne viruses, are readily transmitted to healthy plants grown in virion-containing soil. Other means of natural transmission for certain members also include seed and pollen transmission. A biological vector has been identified that actively spreads certain members of both of these genera. The root-associated fungus Olpidium bornovanus is able to transmit the tombusvirus CuNV and the aureusvirus CLSV to healthy plants. This process has been studied extensively in CuNV and results revealed that the capsid protein is the viral determinant of transmissibility. Outer capsid residues in CuNV particles are able to interact with glycoproteins on the surface of the zoospore. Subsequent plant invasion, mediated by encystment of virus-tethered zoospores, results in virus transfer into root tissue. No biological vectors for zeaviruses have been identified to date, although different potential candidates have been tested; including select aphids, leafhopper, planthopper, and beetles. Mechanical laboratory infection of maize plants is accomplished by performing vascular puncture inoculation, because traditional rub-inoculation of leaves that works for most tombusvirids is not effective.

Host Range and Geographic Distribution Natural infections of different tombusvirus species are generally limited to a few hosts. Examples of such natural hosts can be gleaned from the names given to species which, for most viruses, correspond to the plant from which they were first isolated (Table 1). However, not evident from this list is the ability of some species, such as TBSV, to infect cherry and other woody hosts. Aureusviruses infect a range of herbaceous hosts, ranging from vegetable fruits, such as cucumber, to monocots, like wild grasses and cultivated maize. Zeaviruses also infect maize, however as a relatively newly discovered genus, the existence of additional natural hosts remains to be explored. The distribution of tombusviruses is wide and includes locations in North and South America, different regions in Europe, Algeria, as well as Mediterranean areas. Reports of aureusvirus infections include countries such as Italy, Great Britain, Greece, Jordan, Bulgaria, and India. Zeaviruses were first identified in infected maize in Arizona USA, and since then there have been no additional reports. Collectively, members of the tombusvirus-like group can be considered to be dispersed worldwide.

Applied Aspects Tombusviruses have been used as expression vectors for the efficient production of foreign proteins in plants. For example, antigens of HIV have been produced using different strategies. A complete protein was expressed by replacing the capsid protein ORF of TBSV with that of HIV p24 capsid protein. In an alternative approach, a peptide epitope of gp120 was displayed on the surface of TBSV particles by fusing its corresponding nucleotide sequence to the 30 -end of the capsid protein ORF. In both of these cases, the foreign protein and modified particle were produced in plants. A related tactic, using insect cells, was used to explore developing a potential vaccine for the poison ricin. Virus-like particles (VPLs) were produced using a baculoviral vector encoding the TBSV capsid protein containing a C-terminally fused ricin peptide. This position in the capsid, i.e., an extension of the P-domain, allowed for optimal presentation of 180 copies of this epitope on the surface of particles. For all cases described above, the proteins or particles were found to be immunogenic and elicited corresponding antibody production. Though still in the proof of concept stage, such products could potentially be useful for vaccine production and diagnostics. Whether protein expression occurs via virus vectors or agrobacterium-mediated schemes, in whole plants or plant cell suspension cultures, a common obstacle faced in producing optimal levels of a foreign protein is the plant gene silencing system. Overexpression of exogenous mRNAs often leads to targeting and degradation of these messages by this antiviral system. It has been shown, in different production schemes, that these negative effects can be mitigated by co-expressing p19. Depending on the system, doing this can increase yields of the foreign protein by a modest B20% to over fivefold. Accordingly, further exploration of the utility of this gene silencing suppressor in helping to maximize yield in different large-scale protein production schemes appears warranted.

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Another emerging field in which tombusviruses are being employed is nanoparticle technology. The existing nanoscale of selfordering virus particles makes them attractive candidates for further manipulation. Reengineering of their structural features, such as size and physicochemical properties, can convert them into nanoscaffolds, nanocontainers, or nanobuilding blocks. Genetic and chemical manipulations of TBSV capsids have generated virus-based nanoparticles capable of being chemically derivatized or encapsulating cargo. Additionally, skeletonized TBSV particles have been assembled using multiple copies of a peptide that is a key structural feature that interconnects full-sized capsid subunits in wild type particles. Such designer nanoparticles could by utilized for a variety of purposes, ranging from molecular medicine to biomaterial science. For human applications, safety is an obvious concern, and the effects of TBSV-based nanoparticles on different biological systems is currently being assessed.

Concluding Remarks Tombusvirus-like viruses are distributed throughout the world and can cause damaging diseases in important crops. Tombusviruses, in particular, have served as excellent model systems to understand different steps in their reproductive cycle. Indeed, members of this genus are among the best characterized plus-strand RNA viruses at the molecular level. Virion analysis has yielded novel information on particle structure and function. Studies of other viral proteins have revealed diverse and critical roles for these viral gene products. Functional analyses of viral RNA genomes have uncovered a remarkable network of intragenomic RNA-RNA interactions that regulate a variety of viral processes. This knowledge, combined with the identification of a multitude of host factors, both protein and lipid-based, has provided an extraordinarily detailed understanding of many steps in virus reproduction. Some infections have been found to contain subviral RNAs that can modulate helper virus levels and disease progression. Among these, DI-RNAs have been particularly useful tools for identifying and studying RNA elements and protein factors involved in viral RNA replication. At an applied level, tombusviruses are also being explored for use in different areas of biotechnology, such as vaccines and nanoparticle development. Future studies of this group of viruses, at both the basic and applied levels, will undoubtedly reveal other fundamental properties and uncover additional useful traits.

Further Reading Alam, S.B., Reade, R., Theilmann, J., Rochon, D., 2017. Evidence for the role of basic amino acids in the coat protein arm region of Cucumber necrosis virus in particle assembly and selective encapsidation of viral RNA. Virology 512, 83–94. Chkuaseli, T., White, K.A., 2018. Intragenomic long-distance RNA–RNA interactions in plus-strand RNA plant viruses. Frontiers in Microbiology 9, 529. Danielson, D.C., Pezacki, J.P., 2013. Studying the RNA silencing pathway with the p19 protein. FEBS Letters 587 (8), 1198–1205. Gunawardene, C.D., Donaldson, L.W., White, K.A., 2017. Tombusvirus polymerase: Structure and function. Virus Research 234, 74–86. Ho, P.T., Montiel-Garcia, D.J., Wong, J.J., et al., 2018. VIPERdb: A tool for virus research. Annual Review of Virology 5 (1), 477–488. Jiwan, S.D., White, K.A., 2011. Subgenomic mRNA transcription in Tombusviridae. RNA Biology 8 (2), 287–294. Matsuura, K., 2018. Synthetic approaches to construct viral capsid-like spherical nanomaterials. Chemistry Communication 54 (65), 8944–8959. Nagy, P.D., 2016. Tombusvirus-host interactions: Co-opted evolutionarily conserved host factors take center court. Annual Review of Virology 3 (1), 491–515. Nagy, P.D., Pogany, J., 2006. Yeast as a model host to dissect functions of viral and host factors in tombusvirus replication. Virology 344 (1), 211–220. Nagy, P.D., Pogany, J., Xu, K., 2016. Cell-free and cell-based approaches to explore the roles of host membranes and lipids in the formation of viral replication compartment induced by tombusviruses. Viruses 8 (3), 68. Palukaitis, P., 2016. Satellite RNAs and satellite viruses. Molecular Plant Microbe Interaction 29 (3), 181–186. Scholthof, H.B., 2006. The tombusvirus-encoded P19: From irrelevance to elegance. Nature Reviews in Microbiology 4 (5), 405–411. Truniger, V., Miras, M., Aranda, M.A., 2017. Structural and functional diversity of plant virus 30 -cap-independent translation enhancers (30 -CITEs). Frontiers in Plant Science 8, 2047. White, K.A., Nagy, P.D., 2004. Advances in the molecular biology of tombusviruses: Gene expression, genome replication, and recombination. Progress in Nucleic Acid Research and Molecular Biology 78, 187–226. Wu, B., Grigull, J., Ore, M.O., Morin, S., White, K.A., 2013. Global organization of a positive-strand RNA virus genome. PLoS Pathogens 9 (5), e1003363.

Relevant Websites https://talk.ictvonline.org/ International Committee on Taxonomy of Viruses.

Tombusviruses (Tombusviridae) Luisa Rubino, Institute for Sustainable Plant Protection, National Research Council, Bari, Italy Kay Scheets, Oklahoma State University, Stillwater, OK, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of S.A. Lommel, T.L. Sit, Tombusviruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy, Marc H.V. Van Regenmortel, Elsever Inc, 2008, doi:10.1016/B978-012374410-4.00382-4.

Nomenclature

nt Nucleotide(s) PTGS Post-transcriptional gene silencing RAP Replicase associated protein RdRp RNA dependent RNA polymerase S20,w Sedimentation coefficient of a biomolecule in a solution normalized to acceleration in pure water at 201C


1FS Minus one frameshift aa Amino acid(s) CP Coat protein DI RNA Defective interfering RNA kb Kilobase(s) MP Movement protein

Glossary Citopathology The study of the cellular alterations induced by infectious agents. Dicer Eukaryotic ribonuclease that cuts long dsRNAs and structured ssRNAs into B20–25 bp fragments with 2-nt 50 extensions that activate RISC (see below). Frameshift A change of the reading frame, which results in a different translation and a different protein product. Plasmodesmata (Singular: plasmodesma) Channels traversing the plant cell walls, allowing for the cell-to-cell transport of substances. Plasmodesmata are internally coated by the endoplasmic reticulum. PTSG Post-transcriptional gene silencing; rapid degradation of cellular mRNAs or vRNAs triggered by siRNAs or cellular microRNAs (miRNAs). Readthrough Readthrough consists of the “translation” of a stop codon, such that the synthesis of a larger protein is

allowed. It is a common phenomenon in viruses, and it has a regulatory function. Riboviria A higher taxonomic rank including all viruses possessing an RNA genome. RISC RNA-induced silencing complex; a eukaryotic multiprotein complex that recognizes single stranded microRNAs or double-stranded siRNAs to initiate the destruction of cellular mRNAs or double-stranded viral RNAs. Silencing suppressor (Suppressor of silencing): Virally encoded proteins from diverse families that interfere with host production or effectiveness of siRNAs. siRNAs Small interfering RNAs or silencing RNAs, 20–25 bp dsRNAs with two-nt 30 overhanging ends that target invading viral RNAs or cellular RNAs for degradation in eukaryotic cells. Tombusvirid Member of the family Tombusviridae.

Classification Realm Riboviria, Kingdom Orthornavirae, Phylum Kitrinoviricota, Class Tolucaviricetes, Order Tolivirales, Family Tombusviridae, Subfamilies Calvusvirinae, Procedovirinae, Regressovirinae.

Taxonomy, Phylogeny and Evolution The members of this family all have uncapped positive sense, single-stranded RNA [( þ )ssRNA] genomes that are encapsidated in icosahedral virions and are related by the similarities of their RNA-dependent RNA polymerases (RdRps). The number of genera doubled between 1998 and 2016 and currently consists of 16 genera assigned to three subfamilies (Table 1). The increase occurred by subdivision of two genera (Carmovirus and Necrovirus), incorporation of genus Umbravirus, discovery of new unique species, and the availability of complete genome sequences of earlier members. Genus Umbravirus is the only genus in the subfamily Calvusvirinae (from the Latin “calvus”, naked). Umbraviruses have monopartite genomes that use a
1FS mechanism to express their RdRps but lack a coat protein (CP) gene. The subfamily Procedovirinae (from the Latin “procedo”, go ahead) contains 14 genera and six species unassigned within the subfamily. These viruses possess a monopartite genome and use a stop codon readthrough (RT) mechanism for the expression of the RdRp. The subfamily Regressovirinae (from the Latin “regressus”, step back) contains the genus Dianthovirus, the members of which have a bipartite genome and use a
1FS mechanism for RdRp expression. The continued expansion of genera in the family is partly due to the apparent ability of unrelated tombusvirids infecting the same plant to recombine via exchange of coding regions and/or untranslated regions (UTRs) that may change the host range, leading to

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Table 1

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List of members of the family Tombusviridae organized by subfamilies and genera Species

Acronym

Accession No.

Length (nt)

No. sgRNAsa

Carrot mottle mimic virus Carrot mottle virus Ethiopian tobacco bushy top virus Groundnut rosette virus Lettuce speckles mottle virus Opium poppy mosaic virus Pea enation mosaic virus 2 Tobacco bushy top virus Tobacco mottle virus Ixeridium yellow mottle virus 2

CMoMV CMoV ETBTV GRV LSMV OPMV PEMV2 TBTV TMoV IYMoV2

NC001726 NC011515 NC024808 NC003603 ND NC027710 NC003853 NC004366 NC043206 NC034243

4201 4193 4236 4019

1

Angelonia flower break virus Calibrachoa mottle virus Carnation mottle virus Honeysuckle ringspot virus Nootka lupine vein clearing virus Pelargonium flower break virus Saguaro cactus virus Adonis mosaic virus

AFBV CbMV CarMV HoRSV NLVCV PFBV SgCV AdMV

NC007733 NC021926 NC001265 NC014967 NC009017 NC005286 NC001780 LC171345

3962 3919 4003 3956 4172 3923 3879 3991

2

Alphanecrovirus

Olive latent virus 1 Olive mild mosaic virus Potato necrosis virus Tobacco necrosis virus A

OLV1 OMMV PoNV TNVA

NC001721 NC006939 NC029900 NC001777

3699 3683 3674 3684

2

Aureusvirus

CLSV JCSMV MWLMV PoLV YSV ElAV1

NC007816 NC005287 NC009533 NC000939 NC022895 NC040713

4431 4421 4293 4415 4464 1134a

2

Unassigned aureusvirus

Cucumber leaf spot virus Johnsongrass chlorotic stripe mosaic virus Maize white line mosaic virus Pothos latent virus Yam spherical virus Elderberry aureusvirus 1

Avenavirus

Oat chlorotic stunt virus

OCSV

NC003633

4114

1

Betacarmovirus

Cardamine chlorotic fleck virus Hibiscus chlorotic ringspot virus Japanese iris necrotic ring virus Turnip crinkle virus

CCFV HCRSV JINRV TCV

NC001600 NC003608 NC002187 NC003821

4041 3911 4014 4054

2

Betanecrovirus

Beet black scorch virus Leek white stripe virus Tobacco necrosis virus D

BBSV LWSV TNVD

NC004452 NC001822 NC003487

3644 3662 3762

2

Gallantivirus

Galinsoga mosaic virus

GaMV

NC001818

3803

2

Gammacarmovirus

Cowpea mottle virus Melon necrotic spot virus Pea stem necrosis virus Soybean yellow mottle mosaic virus

CPMV MNSV PSNV SYMMV

NC003535 NC001504 NC004995 NC011643

4029 4266 4048 4009

2

Macanavirus

Furcraea necrotic streak virus

FNSV

NC020469

3966

1

Machlomovirus

Maize chlorotic mottle virus

MCMV

NC003627

4437

1

Panicovirus

Cocksfoot mild mosaic virus Panicum mosaic virus Thin paspalum asymptomatic virus Bermuda grass latent virus

CMMV PMV TPAV BGLV

NC011108 NC002598 NC021705 NC032405

4198 4326 4195 4044

1

Clematis chlorotic mottle virus Elderberry latent virus Pelargonium chlorotic ring pattern virus Pelargonium line pattern virus Pelargonium ringspot virus Rosa rugosa leaf distortion virus

CCMoV ELV PCRPV PLPV PelRSV RrLDV

NC033777 NC026239 NC005985 NC007017 NC026240 NC020415

3880 3892 3904 3883 3865 3971

1

Subfamily/Genus Calvusvirinae Umbravirus

Unassigned umbravirus Procedovirinae Alphacarmovirus

unassigned alphacarmovirus

Unassigned panicovirus Pelarspovirus

4230 4253 4152 2159a 4196

(Continued )

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Table 1

Tombusviruses (Tombusviridae)

Continued No. sgRNAsa

Subfamily/Genus

Species

Acronym

Accession No.

Length (nt)

unassigned pelarspovirus

Jasmine virus H Jasmine mosaic associated virus 1

JaVH1 JMaV

KX897157 MG958505

3867 3859

Tombusvirus

Artichoke mottled crinkle virus Carnation Italian ringspot virus Cucumber Bulgarian latent virus Cucumber necrosis virus Cymbidium ringspot virus Eggplant mottled crinkle virus Grapevine Algerian latent virus Havel River virus Lato River virus Limonium flower distortion virus Moroccan pepper virus Neckar River virus Pelargonium leaf curl virus Pelargonium necrotic spot virus Petunia asteroid mosaic virus Sitke waterborne virus Tomato bushy stunt virus Gentian virus A

AMCV CIRV CBLV CNV CyRV EMCV GALV HRV LRV LFDV MPV NRV PLCV PNSV PAMV SWBV TBSV GVA

NC001339 NC003500 NC004725 NC001469 NC003532 NC023339 NC011535 NC038690 ND NC038691 NC020073 NC038927 NC030452 NC005285 NC038692 NC038693 NC001554 LC373507

4789 4763 4576 4701 4733 4767 4731 2088a 1231a 4772 1305a 4789 4744 1238a 1194a 4776 4742

Maize necrotic streak virus Trailing lespedeza virus 1 Gompholobium virus A Rice virus A

MNeSV TLV1 GomVA RVA

NC007729 NC015227 NC030742 NC035452

4094 3929 3935 4832

1

Carnation ringspot virus

CRSV RCNMV

Sweet clover necrotic mosaic virus

SCNMV

Rice virus X

RVX

1403 3840 1448 3890 3876 1449 4486

1

Red clover necrotic mosaic virus

NC003531 NC003530 NC003775 NC003756 NC003806 NC003807 AB033715

unassigned tombusvirus Zeavirus unassiged Procedovirinae

Regressovirinae Dianthovirus

unassigned Regressovirinae

2

a

No. sgRNAs; genus demarcation criteria uses number of protein-coding sgRNAs only; protein-coding sgRNAs only for genus demarcation.

speciation. The members of the genus Luteovirus (current family Luteoviridae) have viruses that encode frameshifting replicases that are phylogenetically similar to dianthoviruses and are likely to be moved into the family Tombusviridae in the near future. This family serves as an excellent example supporting the concept of modular evolution of viruses. The RpRp is the sole shared module binding the family (Fig. 1). Within the limitations of the RdRp, the family is highly variable in the arrangement and expression of the various genes on its polycistronic RNA. While all members of the family have a T ¼ 3 icosahedral virion, members of different genera achieve this structure using two phylogenetically distinct, but related, CP modules. The family has acquired at least three phylogenetically different cell-to-cell movement modules. The sources of the virus suppressor of host gene silencing appear to be equally diverse. In addition, some species within a particular genus have acquired additional unique accessory modules, possibly for transmission by fungal vectors or for host-specific viral movement.

Virion Structure CP-coding tombusvirids produce non-enveloped virions, consisting of 180 CP subunits with a T ¼ 3 icosahedral symmetry. Virus infections produce virions containing a single copy of their genome ranging in size from 3.6 to 4.8 kb for monopartite species and 5.3 kb for dianthoviruses. Dianthoviruses package one each of the genomic RNAs into one virion via a trans-activator (TA) loop on RNA1 binding to a complementary trans-activator binding site (TABS) just upstream of the subgenomic RNA (sgRNA) start site of RNA2. Genomic RNA constitutes about 17% of the particle weight. Although the underlying size and structure of the virions are similar, the members can be broadly subdivided into two groups based on the presence or absence of a protruding (P) domain at the C-terminus of the CP. Viruses in genera containing P domains (Alphacarmovirus, Aureusvirus, Avenavirus, Betacarmovirus, Dianthovirus, Gallantivirus, Gammacarmovirus, Macanavirus, Pelarspovirus, and Tombusvirus) produce virions that are 32–38 nm in diameter and display a granular surface. Most of these CP subunits range in size from 35 to 42 kDa, while the avenavirus oat chlorotic stunt virus (OCSV) encodes a 48 kDa CP. The X-ray crystal structures of tomato bushy

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Fig. 1 Phylogenetic (distance) analysis of the complete RdRps for all sequenced tombusvirids plus selected viruses from the family Luteoviridae (turquoise box). Alignments of the 78 sequences were made using MUSCLE while trees were generated with the Maximum Likelihood (ML) algorithm of Mega7 using 1000 bootstrap replicates (showing values 450%). Positions with o50% site coverage were eliminated, leaving 809 positions in the final dataset. Condensed triangles mark monophyletic lineages. Brackets connecting tombusvirid branching, enamovirus plus polerovirus branching, and HCV RdRp (aa 2421–3011) were proportionately shortened. Tombusviridae subfamily names are on the right, and colored boxes enclose member genera. Species names are in Table 1.

stunt virus (TBSV) and turnip crinkle virus (TCV) revealed the discrete organization of each CP subunit, and virion structures for many additional species have confirmed the structures for both types of CPs. Generally, the CP subunit can be divided into four domains: the RNA-binding (R) domain is located at the N-terminus followed by the arm (a), shell (S), and P domains (66, 35, 67, and 110 aa, respectively, in TBSV). The R domain contains many basic residues and is found in the interior of the virion. Cryo-electron microscopy reconstructions have further revealed the presence of internal ordered cages of RNA intertwined with CP residues from the R domains beneath the virion surface. This internal scaffold may play a role in directing specific packaging of viral RNA and formation of the icosahedral virions. The virion structure is primarily formed by the globular S domains (stabilized by a pair of Ca2 þ per subunit) which are composed of two sets of four-stranded antiparallel b-sheets. The S domain is also the most highly conserved region of the CP subunit. There is a flexible hinge of five residues between the S and P domains that allows the CP subunit to adopt different configurations by varying the angle between the domains. This feature of the CP subunit overcomes the structural constraints imposed by the icosahedral morphology. P domains (containing antiparallel b-sheet structures in a jellyroll conformation, with one six-stranded b-sheet and one four-stranded b-sheet) of adjacent CP subunits dimerize to produce 90 projections leading to the granular surface texture. Genera lacking the P domain (Alphanecrovirus, Betanecrovirus, Machlomovirus, Panicovirus and Zeavirus) produce virions that are 28–32 nm in diameter with a smooth surface similar to viruses in the genus Sobemovirus. These smaller CP subunits range from 25 to 30 kDa in size. In natural infections, umbravirus RNA is encapsidated separately within their helper-virus CPs to produce virions of B30 nm, similar in structure and size to tombusvirids encoding small CPs. Due to the quasi-equivalent nature of the CP subunits when arranged in an icosahedron, each subunit can take on one of three distinct conformations, termed A, B, and C (respectively, blue, red, and green in Fig. 2). The A and B conformations differ in the angle between the S and P domains and their arrangement on the virion surface (A: fivefold axes, B: threefold axes). The C conformation

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Fig. 2 Virion images based on X-ray crystal structures. (Left) Representation of a particle of tobacco necrosis virus (TNV) at 2.25 Å . The capsid protein of this virus does not contain a protruding domain and is representative of the species in the genera Alphanecrovirus, Betanecrovirus, Machlomovirus, Panicovirus, Umbravirus, and Zeavirus. (Right) Representation of a particle of tomato bushy stunt virus (TBSV) at 2.9 Å . The capsid protein of this virus contains a protruding domain and is representative of species in the remaining genera. For both particles, the individual subunits are colored according to their various conformations: A (blue), B (red), and C (green). From Shepherd, C.M., Borelli, I.A., Lander, G., et al., 2006. VIPERdb: A relational database for structural virology. Nucleic Acids Research 34 (Database Issue), D386–D389.

differs from A and B in that their R/a domains intertwine to form an ordered internal structure around threefold axes of symmetry termed the b-annulus. The three different CP subunit conformations pack together as either AB or CC dimers in the virion particle. For TCV, virion assembly initiates with three CP dimers and viral RNA followed by formation of the virion shell according to structural constraints imposed by CP/RNA interactions. In TCV, the origin of assembly RNA sequence (which specifically initiates the interaction with CP subunits) is contained within a 186 nt region at the 30 end of the CP ORF. Assembly studies with TCV also showed that only RNAs equal to or smaller than 4.35 kb in size could be packaged, suggesting very strict size limitations for icosahedral virions. Virions of tombusvirids have a Mr of B 5.7–9.1  106 and produce a single, well-defined band upon centrifugation with sedimentation coefficients ranging from 118 to 140 S20,w. Virion densities range from 1.34 to 1.36 g cm
3 in CsCl gradients. Virus particles are stable at acidic pH, but expand above pH 7 and in the presence of ethylenediaminetetraacetic acid. They are resistant to elevated temperatures, although thermal inactivation usually occurs above 801C. Due to the lack of a lipid membrane, virions are insensitive to organic solvents and nonionic detergents with the exception of umbraviruses, which lack a CP ORF. In the absence of the assistor virus, plants manually inoculated with the umbravirus genome produce lipid-containing ribonucleoprotein structures, which are sensitive to organic solvents.

Genome Organization and Replication Strategy The unifying genomic feature is the presence of an interrupted ORF for the virally encoded replicase at the 50 end of the genome with identical aminoacyl termini: a high copy replicase-associated protein (RAP) and the larger RdRp, containing the GDD (glycine aspartate) active site motif of supergroup-II RdRps. Both proteins are indispensable for virus RNA (vRNA) replication. In all members but maize chlorotic mottle virus (MCMV), the replicase gene is the most 50 ORF. Thus, incoming vRNA serves as the mRNA for RAP and RdRp. All genes downstream of the RdRp ORF are expressed from 1 or 2 sgRNAs (see Table 1 and Fig. 3), which is one of the discriminating characteristics that define genera. These polymerase subunits share a high degree of sequence similarity within the family. Notably, they do not contain a helicase-type motif and the genomes do not encode any helicase-like ORF. ORFs downstream of the polymerase are expressed via 30 co-terminal sgRNAs with the resident ORFs arranged with an assortment of translational strategies such as ribosomal scanning and frameshifting. The single CP ORF is located either internally (tombusviruses, aureusviruses, and zeavirus) or in the 30 -terminal region of the genomic RNA (Fig. 3). Tombusvirid genomes also encode one or more movement proteins (MPs) that are involved in cell-to-cell spread of the virus. These range in size from the 34–35 kDa version produced by dianthoviruses to the 21–22 kDa proteins expressed by tombusviruses, aureusviruses and maize necrotic streak virus (MNeSV), and down to the 6–9 kDa pair of peptides found in carmo-like and necro-like viruses. Although no mutational analyses of OCSV have been reported, it is likely that the single 8 kDa protein gene overprinted in the CP ORF encodes an MP. In most cases, the MPs can potentiate the local movement (cell-to-cell) of unencapsidated viral RNA but systemic infection

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Fig. 3 Genome organizations for type species of family Tombusviridae. Genomes are arranged in the same order as in Fig. 2. Genus name (left) and acronym (right) of type species are shown. Genome lengths (heavy black lines) are proportional, and ORFs are color-coded for protein homology. Subgenomic RNAs are thin black lines below the genomes. Stop codon readthrough (arrowhead), frameshift (bent arrow), and noncanonical start codon (asterisk) locations are black symbols. Below the genomes, functions of gene products are identified by color-coded boxes. 30k supergroup MPs are related to tobacco mosaic virus 30 kDa movement protein.

generally requires encapsidation. The exception is for tombusviruses which can infect systemically without CP, albeit at a reduced rate. The other prominent protein product with a clearly defined function is the tombusvirus p19 protein which is a potent suppressor of PTGS. p19 binds siRNAs that are generated by a Dicer-like RNase as part of the host RNA-silencing response to viral infection. This binding prevents incorporation of the siRNAs into the RNA-induced silencing complex (RISC) to prevent further cleavage of viral RNAs. Similar to the function, structure, and mechanism of p19, aureusvirus p14 binds siRNAs, albeit less specifically. Although other members of the family may not encode a unique suppressor protein, it is known that other proteins can have suppressor activity aside from their primary functions. This is the case for TCV where the CP is the viral suppressor of PTGS. Various additional ORFs are also encoded in some tombusvirids with undefined functions. One other method utilized by tombusvirids for gene expression is genome segmentation which is only employed by dianthoviruses where RNA2 is monocistronic and encodes the MP. Additional ORFs have been identified in the genomes of panicoviruses and MCMV. For the panicovirus PMV, the unique p15 ORF overprinted within the CP ORF is not necessary for replication but is required for systemic infection along with the CP. For the closely related MCMV, the unrelated ORF overprinted within its CP ORF is required for efficient systemic infection.

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Since the genomic RNAs are positive sense, they are directly infectious as unencapsidated RNAs. vRNAs lack a 50 cap structure and the 30 ends are not polyadenylated nor do they form tRNA-like structures. Instead, tombusvirids and the closely related viruses in the luteovirus genus form several different types of complex structures at their 30 ends called cap-independent translational enhancers (30 CITEs) that interact with small or large stem loops (SLs) usually in the 50 UTRs of vRNA or sgRNAs to allow the circularization that is typically accomplished by recruitment of eukaryotic translation initiation factors (eIFs) that bind to poly(A), 50 caps, and ribosome subunits to initiate translation. Various 30 CITEs have been identified in tombusvirid genomes; the barley yellow dwarf virus (BYDV)-like 30 CITE (BTE) of tobacco necrosis virus (TNVD), the panicum mosaic virus (PMV)-like 30 CITE (PTE) of panicoviruses and cucumber leaf spot virus (CLSV), the Y-shaped 30 CITE of TBSV and carnation Italian ringspot virus (CIRV), the PTE variant of MCMV, and the smaller I-shaped 30 CITE of the zeavirus MNeSV which is also present in some aureusviruses and carmo-like viruses. The 30 CITEs have different structures and binding properties. The direct translation readthrough or frame-shift strategies ensure that the RdRp active site is present in only 5%–10% of the vRNA translation products since readthrough and frameshifting are inefficient. Translation of both RAP and RdRp subunits is followed by localization to and proliferation of cellular membranes. The location of viral replication varies depending on the particular virus and is specified by the pre-readthrough portion of the polymerase (see Cytopathology). In all cases studied to date, the polymerase anchors on membranes, causing proliferation. The advent of yeast-based replication systems for four tombuviruses i.e. CIRV, TBSV, cucumber necrosis virus (CNV) and cymbidium ringspot virus (CymRSV) has provided valuable insights into the host components utilized by the viral replication complex. Once the polymerase has been assembled, it binds to the 30 terminus to initiate minus-strand synthesis. If full-length copies are generated, these serve as templates for synthesis of progeny genomic RNAs. This synthesis is highly asymmetrical with ( þ ) RNA accumulating at much higher levels than (–) RNA. This is controlled by a small base-pairing interaction between a structure at the extreme 30 end of vRNAs and sgRNAs. These are the replication silencer element (RSE) and a 30 -complementary silencer sequence (30 CSS) present in a bulge B50–60 nt upstream which sequesters the 30 end from the RdRp. Occasionally, positive-strand synthesis will be terminated prematurely which leads to the formation of templates for plus-strand sgRNA synthesis. This premature termination is controlled by long-distance RNA–RNA interactions which occur in cis for monopartite tombusvirids such as TBSV, or in trans for dianthoviruses via the same TA/TABS base-pairing interaction used to ensure encapsidation of both RNAs inside one virion. Generally, sgRNAs are produced later in the infection cycle and behave as monocistronic mRNA or polycistronic RNAs, depending on the genus. The sgRNAs demonstrate various translational strategies such as ribosomal scanning to express the p19 ORF which is nested within the p22 ORF in TBSV. In this case, the p22 start codon is suboptimal and is occasionally read through by ribosomes before initiation at the optimal p19 start codon. MNeSV and aureusviruses express their silencing suppressor ORFs using the same mechanism. Similar ribosomal scanning past suboptimal start codons express one or more of the downstream ORFs of carmo-like viruses and necro-like viruses. Very small noncoding viral RNAs that are coterminal with the parent virus 30 end have been identified in infections of MCMV, TCV, CNV, TNVD, and the three dianthoviruses red clover necrotic mosaic virus (RCNMV), carnation ringspot virus (CRSV), and sweet clover necrotic mosaic virus (SCNMV). RNase analyses of RNA fragments from the 30 UTRs of RCNMV, MCMV and the umbravirus opium poppy mosaic virus (OPMV) using Xrn1 in vitro identified a conserved pseudoknot in the 30 UTR that protects the RNA from further 50 -30 degradation. For RCNMV and TNVD, these were shown to be generated by the host 50 -30 exoribonuclease Xrn4, which is stalled by RNA secondary structure. The function of these small RNAs is not yet fully clarified. Sequence searches for program-predicted exoribonuclease-resistant RNAs (xrRNAs) followed by in vitro analyses identified additional Xrnblocking structures in the 50 UTR of most umbravirus sgRNAs.

Life Cycle For most genera, infection is initiated either by mechanical inoculation (piercing of leaf or root tissues with virus-contaminated material) or via vector transmission by a fungal vector in roots. For umbraviruses natural infections occur via viruliferous aphids in the presence of the umbravirus helper virus. After the virus enters an epidermal cell, the virion disassembles, and vRNA engages ribosomes and translation factors to synthesize RAP and RdRp. When enough newly-synthesized vRNA accumulates in initially infected cells, vRNA is transmitted via their MP(s) through plasmodesmata to neighboring cells for further rounds of replication and spread cell-to-cell. At some point newly assembled virions can move throughout the plant via vascular tissue, to produce systemic infections. Viruses can survive in debris and soil for a long time before a new infection cycle starts.

Cytopathology Plants infected with tombusvirids display a distinctive cytopathology, observed by electron microscopy as dense-staining features. Depending on the genus, different virus species can replicate on, remodel, and proliferate membranes of peroxisomes, mitochondria and/ or the cortical ER. The hallmark of infections by members of the genus Tombusvirus are the cytopathic structures known as multivesicular bodies, which are generated after the virus-induced progressive vesiculation and invagination either of the limiting membrane of peroxisomes or the outer membrane of mitochondria. The vesicles are believed to be sites of viral replication. Exceedingly high concentrations of progeny virions can accumulate in the multivesicular bodies, but are also arranged to form crystals in the cytoplasm.

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Subviral Molecules Three different types of subviral entities are associated with the replication of members in the family Tombusviridae: satellite viruses, satellite RNAs (satRNAs) and defective interfering (DI) RNAs. All depend on the presence of a helper virus for their replication, movement, and transmission and may affect helper virus infections. Whereas DI RNA sequences are identical to those of the helper genomic RNAs, satellite viruses and satRNAs share no or limited sequence identity with their cognate viruses. Satellite viruses code for their own CP. In most cases satRNAs are encapsidated by the helper virus-encoded CP, but satC and satS, which are only found in plants infected with PMV and satPMV, are encapsidated by satPMV CP. Satellite viruses described so far are supported by viruses in the genera Alpha- and Betanecrovirus (Satellite tobacco necrosis virus, STNV), Panicovirus (Satellite panicum mosaic virus, SPMV) and Aureusvirus (Satellite maize white line mosaic virus, SMWLMV), and are classified in their own genera Albetovirus, Papanivirus and Aumaivirus, respectively. Satellite viruses associated with tombusvirids possess an uncapped ( þ ) ssRNA genome of B0.8–1.2 kb, lack a poly(A) tail, and harbor a single ORF that encodes their own CP. The subunits are arranged in a T ¼ 1 lattice, and some satellite viruses (SPMV) harbor signals for specific packaging of the satellite virus genome, while STNV1 packaging appears to be mostly related to the abundance of appropriate-sized RNAs. As mentioned above, satellite virus particles are readily distinguished from the helper virions, due to their different size. The STNV genome can form secondary structures for specific recognition by the helper virus RdRp, and contains a 30 -translational enhancer domain (TED) which enhances translation by interacting with a short nucleotide stretch in the 50 UTR. STNV can attenuate the symptoms induced by its helper virus, probably by competing for factors essential for replication. Conversely, SPMV results in a severe exacerbation of PMV-induced symptoms, often leading to plant death, probably due to synergy with the helper virus and to a cytotoxic effect exerted by the satellite CP. Whereas SPMV has the smallest known satellite virus genome, SMWLMV is characterized by the largest satellite virus CP. No effect on the helper virus symptomatology for SMWLMV has been reported. The origin of nonhomologous satRNAs is unclear at present, but they can be generated from recombination events with other satRNAs, the viral genome, DI RNAs, or host sequences. Generally, satRNAs associated with tombusvirids do not share sequence homology with their helper viruses, except for the four satRNAs associated with tombusviruses, which have short conserved nucleotide stretches at the 50 and 30 ends, in addition to a B50 nt sequence in the 50 -terminal region, in common with genomic and DI RNAs. Also the satRNAs (C, D and F) associated with the betacarmovirus TCV have a conserved heptanucleotide at the 30 end. In addition, whereas satRNA D and satRNA F are true satRNAs, satRNA C is a recombinant between satRNA D (50 region) and TCV (30 region). Other tombusvirids are also known to harbor satRNAs, like the 615-nt satRNA associated with the betanecrovirus beet black scorch virus (BBSV) and the satRNAs reported in association with umbraviruses. All the satRNAs described so far have high levels of secondary structure, which account for their stability and survival. Some predicted structures are conserved and are important for the recognition and replication of these molecules by the helper virus. The effect on virus-induced symptomatology is variable. SatRNA C enhances TCV symptoms, whereas satRNA D and F do not interfere with TCV symptomatology. BBSV satRNA exacerbates its cognate virus symptoms. The four 600–800 nt satRNAs associated with tombusviruses differently influence the viral symptoms: the satRNA associated with CyRSV and TBSV satRNAs B1 and L do not modulate the helper virus infection, whereas TBSV satRNA B10 attenuates symptom expression. The satRNAs associated with the umbraviruses groundnut rosette virus (GRV) and tobacco bushy top virus (TBTV) are essential for the encapsidation and transmission of their helpers by an assistor virus. Many tombusvirids generate DI RNAs that can affect pathogenesis. DI RNAs are essentially small, non-coding deletion mutants of the viral genome which are generated by errors in replication such as rearrangement or recombination. Recombination can occur between both homologous and nonhomologous sequences, probably by a copy-choice mechanism in which the viral RdRp jumps from one template to another during the replication process. They usually consist of genomic-derived 50 and 30 non-coding sequences and some internal regions, which are essential for their replication by the virus' RdRp. DI RNAs are more readily generated at high multiplicity of infection. Several species such as TBSV and TCV produce DI RNAs during viral infection. For TBSV the presence of DI RNAs results in attenuation of symptoms, whereas for TCV the presence of DI RNAs intensifies symptoms. The attenuation of symptoms is not only due to competition for host factors, but largely resides in the highly structured DI molecules, which can trigger post transcriptional gene silencing against the helper virus genome. Both DI RNAs and satRNAs have been useful molecular tools for studying viral replication and recombination. Since both satRNAs and DI RNAs are dependent on the host or parent virus for replication, they must retain any sequence or structural signals required for recognition by the viral polymerase, and therefore can help determine those signals. Recombination can facilitate viral evolution as well as repair of viral genomes, so it is an important factor in the virus life cycle, and it can have profound effects on pathogenesis. In the tombusvirus family, recombination in the carmo-like and tombus-like viruses has been shown to repair damaged or deleted 30 ends of virus-associated RNAs, as well as generate new satRNAs or DI RNAs.

Pathogenesis Depending on the host and on the genus, tombusvirid infections can be symptomless or consist of mottling, crinkling, mosaic, distortion, chlorotic and/or necrotic lesions on the leaves, and stunting of the plants. The natural host range of most species is relatively narrow; however, the experimental host range tends to be broad. Members can naturally infect either monocotyledonous or dicotyledonous plants. Fewer tombusvirids infect monocots: eight infect grasses (Family Poaceae) while four infect members of the Order Asparagales. Natural infections can be limited to or are often concentrated in the root system. Many species are found in the soil, where

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they survive for a long time. All species are readily transmitted by host vegetative propagation and mechanical inoculation (although some species cannot be inoculated by leaf-rub inoculation), and some may be transmitted by contact and through seeds. A number of species are readily detected in surface waters, rivers, lakes, and even the ocean. Transmission by chytrid fungi in the genus Olpidium and by beetles has also been reported for members of several genera. For viruses that can be transmitted by fungi, there is a considerable amount of specificity required between the virus and the fungal vector. For example, the tombusvirus CNV, the gammacarmovirus melon necrotic spot virus (MNSV), and the aureusvirus CLSV are transmitted by the root-inhabiting chytrid fungus Olpidium bornovanus, whereas the alphanecrovirus TNVA, and betanecroviruses TNVD and BBSV are transmitted exclusively by Olpidium brassicae. Umbraviruses are aphid-transmitted, but only when plants are co-infected with an “assistor” luteovirid. Many members can be transmitted through the soil either dependent on, or independent of, a biological vector. This rather unusual soil and water mode of transmission is unique to this plant virus family and is based on the unusually robust constitution of the virion. Geographical distribution of particular species varies from wide to restricted and is strongly linked to the presence of the specific host. Most species occur in temperate regions although legume-infecting gammacarmoviruses and furcraea necrotic spot virus (FNSV) have been found in tropical areas. Two alphacarmoviruses occupy more extreme environments: Nootka lupine vein clearing virus (NLVCV) grows in Alaska, USA (northern temperate climate above the arctic circle), and saguaro cactus virus (SgCV) was found in a hot desert in Arizona, USA. MCMV was initially found in South America, but it is now one of the most widespread family members due to the extreme increase in maize (Zea mays)-producing nations throughout tropical and temperate zones.

Further Reading Chkuaseli, T., White, K.A., 2018. Intragenomic long-distance RNA-RNA interactions in plus-strand RNA plant viruses. Frontiers in Microbiology 9, 529. doi:10.3389/ fmicb.2018.00529. Gunawardene, C.D., Newburn, L.R., White, K.A., 2019. A 212-nt long RNA structure in the Tobacco necrosis virus-D RNA genome is resistant to Xrn degradation. Nucleic Acids Research 47, 3929–9342. Kim, M., Roossinck, M.J., 2017. Small linear satellite RNAs. In: Hadidi, A., Flores, R., Randles, J.W., Palukaitis, P. (Eds.), Viroids and Satellites. London, UK: Elsevier Academic Press, pp. 567–575. Kotta-Loizou, I., Peyret, H., Saunders, K., Coutts, R.H.A., Lomonossoff, G.P., 2019. Investigating the biological relevance of in vitro-identified putative packaging signals at the 50 terminus of Satellite tobacco necrosis virus 1 genomic RNA. Journal of Virology 93, e02106–e02118. May, J.P., Yuan, X., Sawicki, E., Simon, A.E., 2018. RNA evasion of nonsense-mediated decay. PLoS Pathogens 14, e1007459. Miller, W.A., Shen, R., Staplin, W., Kanodia, P., 2016. Noncoding RNAs of plant viruses and viroids: Sponges of host translation and RNA interference machinery. Molecular Plant-Microbe Interactions 29, 156–164. Nagy, P.D., 2017. Exploitation of a surrogate host, Saccharomyces cerevisiae, to identify cellular targets and develop novel antiviral approaches. Current Opinion in Virology 26, 132–140. Nagy, P.D., Strating, J.R., van Kuppeveld, F.J., 2016. Building viral replication organelles: Close encounters of the membrane types. PLoS Pathogens 12, e1995912. Navarro, J.A., Pallas, V., 2017. An update on the intracellular and intercellular trafficking of carmoviruses. Frontiers in Plant Sciences 8, #1801. Pyle, J.D., Scholthof, K.-B.G., 2017. Biology and pathogenesis of satellite viruses. In: Hadidi, A., Flores, R., Randles, J.W., Palukaitis, P. (Eds.), Viroids and Satellites. London, UK: Elsevier Academic Press, pp. 627–636. Rubino, L., 2017. Biology of satellites. In: Hadidi, A., Flores, R., Randles, J.W., Palukaitis, P. (Eds.), Viroids and Satellites. London, UK: Elsevier Academic Press, pp. 649–658. Rubino, L., Russo, M., Martelli, G.P., 2014. Tombusvirus-induced multivesicular bodies: Origin and role in virus-host interaction. In: Gaur, R.K., Hohn, T., Sharma, P. (Eds.), Plant Virus-Host Interaction. Molecular Approaches and Evolution. Oxford, UK: Elsevier Academic Press, pp. 163–175. Scheets, K., Jordan, R., White, K.A., Hernandez, C., 2015. Pelarspovirus, a proposed new genus in the family Tombusviridae. Archives of Virology 160, 2385–2393. Serra-Soriano, M., Navarro, J.A., Pallas, V., 2017. Dissecting the multi-functional role of the N-terminal domain of the Melon necrotic spot virus coat protein in RNA packaging, viral movement and interference with antiviral plant defence. Molecular Plant Pathology 18, 837–849. Steckelberg, A.L., Vicens, Q., Kieft, J.S., 2018. Exoribonuclease-resistant RNAs exist within both coding and non-coding subgenomic RNAs. mBio. 9, e02461.

Relevant Websites https://talk.ictvonline.org/taxonomy/ Virus Taxonomy: 2019 Release. International Committee on Taxonomy of Viruses (ICTV).

Tritimoviruses and Rymoviruses (Potyviridae) Satyanarayana Tatineni, Agricultural Research Service, US Department of Agriculture, Lincoln, NE, United States and University of Nebraska–Lincoln, Lincoln, NE, United States Gary L Hein, University of Nebraska–Lincoln, Lincoln, NE, United States r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) CP Coat protein or capsid protein ELISA Enzyme-linked immunosorbent assays HC-Pro Helper component-protease kb Kilobase kDa Kilo dalton Mr Relative molecular mass nm Nanometer(s) nt Nucleotide(s) NTR Non-translated region

Glossary Cleavage Processing of large proteins into smaller peptides at specific amino acid sequences by proteases. Motif A short amino acid or nucleotide sequence that is presumed to have a biological function through sequencespecific binding.

ORF Open Reading Frame PAGE Polyacrylamide gel electrophoresis PCR Polymerase chain reaction qRT-PCR Quantitative reverse transcription polymerase chain reaction RdRp RNA dependent RNA polymerase SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis ssRNA single-stranded RNA VPg Viral protein genome-linked

Polyprotein A large protein with a chain of covalently joined smaller peptides that can be cleaved by virus-encoded proteases into smaller proteins with different biological functions. Virion The protective shell of genomic RNA/DNAs of viruses covered mostly with virus-encoded capsid and often with other virus-encoded accessory proteins.

Introduction The members of the genus Tritimovirus have previously been included in the genus Rymovirus based on biological properties such as monocotyledonous hosts and transmission by eriophyid mites. However, phylogenetic studies based on partial or complete genome sequences of Rymovirus members revealed that the eriophyid mite transmission characteristic was a paraphyletic trait within the genus Rymovirus. Hence, Wheat streak mosaic virus (WSMV) and Brome streak mosaic virus (BrSMV) were separated from the genus Rymovirus and classified into the new genus Tritimovirus with WSMV as the type species. Subsequently, four other definitive species were assigned to the genus Tritmovirus. Among the members of genera Tritimovirus and Rymovirus, WSMV is the most economically important virus on wheat in most wheat growing regions of the world, causing significant yield losses. WSMV was first reported from Nebraska, USA as wheat mosaic disease, and disease caused by WSMV was referred as wheat streak mosaic disease (Fig. 1).

Taxonomy and Classification The family Potyviridae contains a total of ten genera, plus three unassigned species, and includes Tritimovirus and Rymovirus as distinct genera. Some members of the Rymovirus and Tritimovirus genera are transmitted by eriophyid mites Abacarus hystrix (Nalepa) and Aceria tosichella (Keifer) (Fig. 2(A)), respectively; however, members of Rymovirus are phylogenetically more related to aphid-transmitted members of the genus Potyvirus. Recently, there was an unsuccessful proposal to assimilate the genus Rymovirus into the genus Potyvirus as a rymovirus subgroup based on phylogenetic relatedness. Currently, the genus Rymovirus consists of three known definitive members (Agropyron mosaic virus, Hordeum mosaic virus, and Ryegrass mosaic virus) with Ryegrass mosaic virus (RGMV) as the type species, while the genus Tritimovirus consists of six recognized species (Brome streak mosaic virus, Oat necrotic mottle virus, Tall oatgrass mosaic virus, Wheat eqlid mosaic virus, WSMV, Yellow oat grass mosaic virus) with Wheat streak mosaic virus as the type species (Table 1).

Virion Structure Virions of Tritimovirus and Rymovirus members are flexuous filaments of 690–720 nm in length and 11–15 nm in diameter (Fig. 2(B)) made up of 95% protein and 5% RNA. The molecular weight (MW) of rymo- and tritimoviral CP subunits range between 27 and

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Fig. 1 Wheat streak mosaic virus-infected wheat plants showing chlorotic streaks (A) and mosaic symptoms (B) under field conditions. Photographs are courtesy of Mary Burrows, Montana State University, USA and the Government of Western Australia, https://www.agric.wa.gov.au.

A

B

Fig. 2 Scanning electron micrograph of an adult wheat curl mite (Aceria tosichella Keifer) on wheat leaf (A). Wheat curl mite micrograph picture credit: G. Bauchan and R. Ochoa, USDA-ARS. Electron micrograph picture showing flexuous filamentous virion particles of Wheat streak mosaic virus. Bar indicates 500 nm (B).

Table 1

Genome features of members of Rymovirus and Tritimovirus genera in the family Potyviridae

Genus/Species

Acronym

Vector

Genome size (nt)

5’-NTRa (nt)

Polyprotein size; amino acids # and MW (kDa)

3'-NTR (nt)

Coat protein MW (kDa)

Accession #

Rymovirus Agropyron mosaic virus Hordeum mosaic virus Ryegrass mosaic virus*

AgMV HoMV RGMV

A. hystrix NVRb A. hystrix

9540 9463 9522

131 131 112

3079; 349 3051; 345 3087; 348

173 180 163

37.8 34.2 35.5

NC_00590 NC_005904 NC_001814

Tritimovirus Brome streak mosaic virus Oat necrotic mottle virus Tall oatgrass mosaic virus Wheat eqlid mosaic virus Wheat streak mosaic virus* Yellow oat grass mosaic virus

BrSMV ONMoV TOGMV WEMV WSMV YOGMV

A. tosichella NVR NVR NVR A. tosichella NVR

9672 9346 9359 9636 9384 9292

145 130 124 137 130 128

3094; 3024; 3030; 3110; 3036; 3007;

244 144 146 170 147 144

34.1 36.0 35.9 38.6 36.7 34.8

NC_003501 NC_005136 NC_022745 NC_009805 NC_001886 NC_024471

a

Non-translated region. No vector reported. Source: cexcluding polyA tailr.

b

348 344 345 352 344 341

Tritimoviruses and Rymoviruses (Potyviridae)

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39 kDa (Table 1) with differences mainly due to variable lengths at the N-terminal region. The internal core of the CP residues is likely involved in virion architecture. The N- and C-terminal aa are exposed on the surface of the viral particles, and these exposed aa most likely interact with other viral, host, and vector proteins to successfully complete the virus life-cycle. The conserved N-terminal DAG-motif found in potyviral CPs that is required for aphid transmission is not present in most rymo- and tritimoviral CPs. The helical virion assembly of WSMV suggests that each virion particle possesses B2000 CP subunits surrounding the RNA with 7–8 subunits per turn. The aa sequence suggests the MW of WSMV CP to be 37 kDa; however, SDS-PAGE predicts the molecular weight to be 45 kDa. Partially purified virions of WSMV resolved into a 45 kDa major CP, plus 32 and 29 kDa minor CP bands. These minor proteins are part of the 45 kDa CP. The exact nature of the production of these additional CP bands is not known. However, these bands are thought to be produced due to proteolytic activity in senescence leaves.

Genome Organization The genomic organization of Rymovirus and Tritimovirus members is similar to those in the genus Potyvirus. Rymovirus and Tritimovirus members possess a positive-sense single-stranded RNA (ssRNA) with sizes ranging from 9292 to 9672 nt (Table 1). The genomic RNAs of these viruses likely contain a viral protein genome-linked (VPg) at the 5’ terminus and a poly-A-tail at the 30 terminus. The genome encodes a single ORF coding for a polyprotein with 3007 to 3110 aa with a predicted MW of 341–352 kDa protein (Table 1). The polyprotein is processed into mature proteins by three virus-encoded proteinases: P1, HC-Pro and NIa-Pro. The serine proteinase P1 and cysteine proteinase HC-Pro cleave the polyprotein in cis at their C-terminal ends, while the serine proteinase NIa-Pro cleaves the polyprotein in trans at other heptapeptide cleavage sites. Thus, cleavage of the polyprotein results in 10 mature proteins. An additional small ORF, PIPO, is expressed by the polymerase slippage mechanism from the P3 cistron as P3N-PIPO. The 50 and 30 non-translated regions (NTRs) in all members range from 112–145 and 144–244 nt, respectively (Table 1). The NTRs are predicted to contain the cis-acting elements for replication and translation enhancers. The mechanism of genomic RNA translation in rymo- and tritimoviruses is not known. However, translation of genomic RNA potentially takes place through the cap-independent leaky scanning mechanism similar to members of potyviruses.

Functions of Viral Proteins The mature polyproteins encoded by members of the genera Rymovirus and Tritimovirus consist of the following proteins from the N-terminus to C-terminus: P1, HC-Pro, P3, P3N-PIPO, 6K1, CI, 6K2, NIa-VPg, NIa-Pro, NIb, and CP (Fig. 3). WSMV is the most extensively studied virus outside the genus Potyvirus in the family Potyviridae. Hence, functions described for viral proteins in this section are mostly based on studies on WSMV. The P1 protein is a serine proteinase and is the most divergent among WSMV isolates. The P1 protein of WSMV and Oat necrotic mottle virus (ONMoV) but not Agropyron mosaic virus (AgMV) and Hordeum mosaic virus (HoMV) is a suppressor of RNA silencing and a pathogenicity enhancer (Fig. 3). In contrast to the multi-functional helper component proteinase (HC-Pro) of potyviruses, the HC-Pro from WSMV is dispensable for systemic infection, but it is required for wheat curl mite transmission. WSMV HC-Pro is predicted to be a virus-encoded accessory protein of virion component similar to members of the genus Potyvirus. The HC-Pro proteins of AgMV and HoMV of the genus Rymovirus are suppressors of RNA silencing. The P3 protein is required for replication, while P3N-PIPO of WSMV is required for cell-to-cell movement. The P3N-PIPO is highly conserved among WSMV isolates and mutating P3N-PIPO without affecting the P3 coding sequence severely debilitates cell-to-cell movement of WSMV. The exact role of 6K1 in virus biology is not known though deletion of this cistron is lethal to WSMV. The CI protein is the largest of all virus-encoded proteins. CI proteins of rymo- and tritimoviruses possess RNA helicase activity and an ATP binding domain. These domains are predicted to form the viral replication complex. The CI is also predicted to be involved in cell-to-cell movement similar to potyviral CI proteins. The 6K2 is another small peptide present in all rymo- and tritimoviruses and is thought to be involved in viral replication, movement and pathogenicity. The B49 kDa NIa protein is cleaved into an N-terminal VPg (B23 kDa) and a C-terminal proteinase (Pro; B26 kDa) (Fig. 3). The VPg protein is covalently attached to the 50 end of viral genomic RNA and is predicted to act as a primer during replication. VPg is also predicted to interact with host factors for translation, long-distance movement, and symptom development. NIa-Pro, a serine protease, is responsible for the cleavage and release of mature proteins from the polypeptide except for P1 and HC-Pro. The NIa-Pro cleaves in cis at its C-terminus for its own release from the polypeptide, followed by in trans cleavage at specific heptapeptide cleavage sites. The NIa-Pro of WSMV was identified as an elicitor of superinfection exclusion. NIa and NIb proteins form inclusion bodies in the nucleus, hence the name ‘nuclear inclusion’ for these proteins. The NIb protein is an RNA-dependent RNA polymerase (RdRp) and is responsible for viral replication through interactions with other viral and host proteins. The C-terminal product of the polyprotein is the CP with multifunctional roles. The CP, in addition to virion encapsidation, is also involved in wheat curl mite transmission, cell-to-cell and long-distance movement, pathogenicity, and superinfection exclusion (Fig. 3). The C-terminal region of WSMV CP is dispensable for virion assembly, but it is required for cell-to-cell movement. The CP aa 36–84 are dispensable for virion assembly and systemic infection, but deletions comprising aa 57–84 elicited more severe symptoms compared with wild type virus. The CP aa 57–100 are identified as viral determinants along with HC-Pro for wheat curl mite transmission.

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Fig. 3 Schematic representation of genomic organization of Wheat streak mosaic virus (WSMV). The large rectangle indicates an open reading frame encoding a large polyprotein that is cleaved in cis by P1 and HC-Pro (indicated with curved arrows). The reminder of the polyprotein is predicted to be cleaved by NIa-Pro (indicated with upward arrows) in cis at its C-terminal end and in trans between P3/6K1, 6K1/CI, CI/6K2, 6K2/NIa-VPg, NIa-VPg/NIa-Pro, NIa-Pro/NIb, and NIb/CP at heptapeptide cleavage sites. Boxes below the genomic organization indicate proteins encoded by WSMV and their putative functions in virus biology.

Life Cycle and Epidemiology Host Range The known host range of rymo- and tritimoviruses is restricted to monocotyledonous plants including several cereals and other grass species. Among members of rymo- and tritimoviruses, the host range of WSMV has been most extensively examined. Wheat (Triticum aestivum) and corn (Zea mays) are the most important hosts for WSMV, but natural WSMV infection of corn has been mostly controlled with the introduction of hybrid maize cultivars resistant to WSMV. The host range of WSMV also includes oats (Avena sativa), barley (Hordeum vulgare), rye (Secale cereale), foxtail millet (Setaria italica), and proso millet (Panicum miliaceum). In addition to cereals, a number of annual and perennial grasses serve as hosts for WSMV. Many of these plants also serve as hosts for wheat curl mite, the vector of WSMV.

Vector Transmission Some members of both the rymo- and tritimoviruses are transmitted by eriophyid mites. RGMV is efficiently transmitted by the cereal rust mite (A. hystrix), but this vector also transmits AgMV inefficiently. No vector has been found for HoMV, the third member of the genus Rymovirus. The mechanisms and viral determinants of rymoviral transmission by A. hystrix are not known. Among members of the genus Tritimovirus, WSMV and BrSMV are the only viruses with known vectors, and these two are transmitted by the wheat curl mite Aceria tosichella (Keifer) (Fig. 2(A)). Transmission of WSMV has been extensively studied. The HC-Pro and CP of WSMV are identified as viral determinants of mite transmission. Only immature mites can acquire the virus in as little as 15 min with improved transmission efficiencies as acquisition time increases up to 16 h. Mites remain viruliferous through molting and as adults for at least seven days. Thus, WSMV transmission by wheat curl mites is not stylet or foregut borne but exhibits several characteristic features of persistent transmission. However, the short acquisition time and lack of a latent period for WSMV transmission by wheat curl mites are not typical of persistent transmission. This incongruence could be due to the microscopic morphologies of these mites that could impact feeding and internal mite-virus interactions. Overall, WSMV transmission by wheat curl mites does appear to fit the category of persistent transmission with some differences. The remaining viruses in these two genera have not been associated with a particular vector. Recent work has demonstrated that both A. tosichella and A. hystrix are each a complex of several genetic variants. Some variants within these two species have distinct, sometimes unique, host relationships within the grasses. Additionally, some of these variants have been shown to have differing virus transmission capacity with WSMV, but little is known about the virus capabilities of many of these variants. Further research is needed to delineate the vectors of several of the viruses in these two genera, and perhaps the genetic variants of these two mite species should be considered as potential vectors for these remaining viruses.

Tritimoviruses and Rymoviruses (Potyviridae)

801

Fig. 4 Wheat streak mosaic virus (WSMV) disease cycle in winter wheat. Wheat curl mites disperse from maturing wheat onto summer green bridge hosts; wheat curl mites move from green bridge hosts onto newly emerged wheat seedlings during the fall and transmit WSMV; mites and virus overwinter in winter wheat; as temperature warms up in the spring, virus symptoms develop and impact wheat yield.

Seed Transmission No information on seed transmission of rymo- and tritimoviruses is available except for WSMV. Seed transmission of WSMV was first reported at 0.1% level in maize in seed production lots from Iowa. WSMV transmission through seed in eight wheat genotypes was reported at 0.2%–0.5%. However, seed transmission of up to 1.5% was reported in individual wheat genotypes. Low seed transmission of WSMV may not be significant in establishing epidemics in areas where the virus is prevalent, but wheat plants infected through seed transmission can serve as a source of virus inoculum for the secondary spread through wheat curl mites. Seed transmission of WSMV is epidemiologically most significant with the possibility of virus spread through global trading and movement of infected seed.

Disease Cycle In eriophyid mite-transmitted diseases, the bridging of hosts between two cropping seasons plays an important role in continuing the disease cycle. The epidemiology and disease cycle for wheat curl mite-transmitted WSMV have been extensively studied. Knowledge and management of the WSMV disease cycle (Fig. 4) likely apply to other viruses in the genera Rymovirus and Tritimovirus. Due to the lack of well-developed legs, wheat curl mites rely on wind currents to disperse within and between the fields. Mites move from the leaves and stems to wheat spikes in headed wheat fields for further propagation and protection. As the wheat crop matures with the drying of heads and foliar tissue, the mites must find new hosts to survive during the summer. This is particularly critical as the mites cannot survive for more than a day or two off a living host. These hosts consist of volunteer wheat, corn and other cereal crops, and summer grasses that form the green bridge for mites and viruses between the summer wheat harvest and emergence of the fall-planted crop. Mites from WSMV-infected volunteer wheat and other cereal and grass hosts can move onto wheat seedlings during the fall season via wind. These mites propagate rapidly due to their rigorous reproductive capacity and transmit WSMV to wheat seedlings. In general, warmer fall conditions, early wheat planting, and presence of a green bridge to host the mites and virus between summer harvest and emergence of the fall crop increase virus epidemics in susceptible wheat cultivars. Overwintering of mites will occur as eggs, larvae, nymphs, and adults in wheat and other plant hosts, and WSMV will survive in

802

Tritimoviruses and Rymoviruses (Potyviridae)

live wheat leaves and crown of virus-infected plants. Serious epidemics result from extensive mite presence and virus transmission occurring through the fall with the most serious symptoms and impact developing with the advent of warm temperatures in the spring. As temperatures warm up in the spring season, mites become active, multiply rapidly, and eventually move to the summer green bridge just before harvest to continue the disease cycle (Fig. 4).

Pathogenicity Knowledge of how rymo- and tritimoviruses elicit disease is scant except for WSMV. The availability of reverse genetics system for WSMV facilitated the identification of viral determinants involved in disease development. The P1 of WSMV was identified as a suppressor of RNA silencing and a pathogenicity enhancer. Potato virus X infection of transgenic N. benthamiana plants expressing WSMV P1 protein resulted in more severe symptoms compared to non-transgenic N. benthamiana plants. Wheat plants infected by WSMV with complete deletion of HC-Pro elicited mild symptoms compared with wild-type virus. Surprisingly, WSMV harboring deletions comprising aa 57–84 in CP elicited more severe symptoms in multiple cereal hosts with increased accumulation of viral genomic RNA copies. In members of the genus Potyvirus, the 50 - and 30 NTRs have been identified as symptom determinants as mutations in these regions affected symptom severity. However, it is possible that these mutations might have affected viral cis-acting elements located in the 50 and 30 NTRs for replication. Additionally, P3, CI, 6K2, and NIa-VPg of different potyviruses have been identified as pathogenicity enhancers. However, no information is available on the role of these proteins in disease development of rymo- and tritimoviruses. Co-infection of wheat by unrelated WSMV and Triticum mosaic virus (TriMV), a Poacevirus in the family Potyviridae, exacerbates the disease phenotype with enhanced accumulation of both interacting viruses. Interactions between WSMV and TriMV during early stages are asymmetrical because prior infection of wheat by TriMV caused accelerated cell-to-cell movement and genomic RNA accumulation of WSMV but not vice-versa. The mechanisms and viral and host factors responsible for synergistic interaction between WSMV and TriMV are not known.

Diagnosis The signature pinwheel-shaped inclusion bodies and flexuous filamentous virion particles are diagnostic features of potyviral infections. However, the requirement of sophisticated methodologies and equipment limit the use of these characteristic features as diagnostic methods for potyviruses. Most of the purified virions of rymo- and tritimoviruses are immunogenic in rabbits for the production of polyclonal antibodies. The polyclonal antibodies raised against purified virions have been widely used for the detection of rymo- and tritimoviruses in direct antigen coating, double antibody sandwich, and triple antibody sandwich forms of enzyme-linked immunosorbent assays (ELISAs). ELISA is less expensive and can be used for large-scale detection of samples, but the sensitivities of ELISAs depend on the quality of antibodies. The availability of genome sequences of rymo- and tritimoviruses facilitated the development of RT-PCR and real-time RT-PCR (RT-qPCR) methods for sensitive detection of viruses to overcome the limitations of ELISA-based detection methods. Additionally, multiplex RT-PCR methods allow for the simultaneous detection of multiple viruses. RT-qPCR has been used for absolute quantification of WSMV genomic RNA copies to study virus biology and virus-host, virus-vector and virus-virus interactions.

Disease Management WSMV is the most economically important virus among eriophyid mite-transmitted rymo- and tritimoviruses. Hence, extensive research on the development of management practices for WSMV in wheat has been performed. Viruliferous mites from oversummering hosts are the primary source of infection. Hence, cultural practices targeted at managing mite populations and deployment of plants resistant to eriophyid mites, viruses, or both are needed for effective management of rymo- and tritimoviruses.

Cultural Practice Cultural practices developed for the management of WSMV are most likely applicable to other viruses in these two genera, though some fine tuning might be required. Due to the secluded nature of mite feeding, management of wheat curl mites through pesticide application is ineffective. The main cultural management practice for WSMV is the elimination of the green bridge by removing volunteer wheat and other cereals and grasses that host both virus and vector through the summer, thus preventing the disease cycle from continuing into the fall and the next wheat crop. Mite populations are at their peak just before wheat harvest, so any volunteer wheat growing at this time is rapidly infested with mites and if not controlled will serve as an optimal green bridge host for mites and virus. Severe WSMV outbreaks are likely if hailstorms occur prior to wheat harvest and increase volunteer wheat growth. The overlap of emerged winter wheat in the fall with adjacent late-maturing crops such as maize, sorghum and small grain cover crops should also be avoided. These late-maturing crops act as reservoirs for virus and vector. Additionally, avoiding early planting of winter wheat will reduce infestation potential and the period of favorable fall temperatures for mite and virus development. Thus, effective management of WSMV depends on an integrated disease management approach by optimizing cultural practices in combination with genetic resistance.

Tritimoviruses and Rymoviruses (Potyviridae)

803

Genetic Resistance Except for WSMV, little or no information is available on genetic resistance against rymo- and tritimoviruses. The deployment of genetically resistant wheat cultivars with nonallelic Wsm1, Wsm2, or Wsm3 genes has been used for WSMV disease management. Wheat cultivar Mace with the Wsm1 gene, a single dominant R gene, associated with chromosome 4D from intermediate wheatgrass [Thinopyrum intermedium (Host) Barkworth & D.R. Dewey] provides resistance to WSMV and TriMV. The Wsm2 gene, a single dominant gene, present in wheat cultivars RonL, Snowmass, Clara CL, and Oakley CL is resistant to WSMV but not to TriMV. The Wsm2 gene is associated with chromosome 3BS with unknown origin. Wheat cultivars with Wsm1 or Wsm2 genes provide resistance through temperature-dependent impairment of viral long-distance movement; however, for both of these genes, resistance begins to break down as temperatures increase above 201C. Wsm3 is a true resistance gene that is transferred from wheatgrass (T. intermedium) to provide resistance against WSMV at higher temperatures. However, commercial wheat cultivars with Wsm3 gene are not yet available. Eriophyid mites not only transmit rymo- and tritimoviruses, but they can also cause yield losses in crop plants due to their great reproductive and feeding capabilities. Thus, the development of crop plants resistant to eriophyid mites could minimize losses from viruses as well as feeding damage by eriophyid mites. Thus far, genetic resistance to wheat curl mite has been reported for four curl mite colonization (Cmc) genes. The Cmc1/Cmc4, Cmc2, or Cmc3 genes in wheat have resulted from DNA transfer from Aegilops tauschii, A. elongatum, and S. cereal, respectively. However, with the extensive deployment of wheat cv. TAM107 (Cmc3 gene), wheat curl mite populations were able to adapt to this resistance gene. Thus, the durability of genetic resistance against wheat curl mites has been questioned. Wheat cultivars with multiple lines of resistance through gene pyramiding may provide more durable resistance to wheat curl mites. Additionally, pyramiding mite-resistance with virus-resistance could provide an effective management tool. Mutation of recessive genes encoding the eukaryotic translation initiation factor 4E or its isomers to prevent interaction with viral proteins without being lethal to host plants provides resistance against several members in the genus Potyvirus. However, there is no information on recessive resistance against rymo- and tritimoviruses. With the advent of CRISPR-Cas9 technology, mutation of candidate recessive genes in crop plants can be achieved to obtain resistance against rymo- and tritimoviruses.

Biotechnological Application Plant viruses can be used for the expression of specialty products and for functional genomics of host genes through virus-induced gene silencing. WSMV has an excellent reverse genetics system that can be exploited for recombinant protein expression, vaccine production, and drug delivery as a nano-agent. WSMV has been used to express GUS, GFP, and RFP proteins in wheat to examine virus movement, virus-host, and virus-virus interactions. Lastly, small peptides can be fused to CP or HC-Pro to integrate into virions that can be used for biotechnological purposes. A major limitation of using plant viruses as expression vectors is the instability of inserted sequences since many plant viruses recombine by deleting all or parts of inserted sequences. However, inserted GFP or RFP in the WSMV genome was stably expressed for more than 120 days with no indication of deletions or reassortment in inserted sequences. Additionally, WSMV stably maintained and simultaneously expressed two foreign genes inserted between the P1/HC-Pro and NIb/CP cistrons for more than 30 days in wheat. WSMV has been used as an expression vector to express TriMV cistrons in wheat to examine virus-virus interactions. The P1 protein of WSMV is a strong suppressor of RNA silencing. Consequently, even though many biotechnological applications are impaired by the host defense mechanism, WSMV P1 can be used to attain consistent high-level expression of transgenes in monocotyledonous plants. Additionally, transgenic plants expressing WSMV P1 can be used to enhance the levels of expression of viral vectors.

Concluding Remarks Viral gene functions, viral proteins interacting with host and vector proteins, and mechanisms of suppression of host defense have been extensively studied for members of the genus Potyvirus. Among rymo- and tritmoviruses, extensive research has been performed particularly on WSMV regarding genetic diversity and the roles of HC-Pro and CP in vector transmission and disease development. However, knowledge of host and vector proteins required for disease development and transmission, respectively, is lacking. Knowledge of viral gene functions and interaction of viral proteins with host and vector factors could eventually facilitate the development of new management strategies. Identification of candidate host factors involved in disease development could be targeted via gene-editing technology to interrupt virus-host interactions. Ultimately, disruption of virus-host interactions could mitigate the viruses’ ability to infect plants. This could be an important development especially for WSMV, the most important virus found in these two genera.

Further Reading French, R., Stenger, D.C., 2003. Evolution of Wheat streak mosaic virus: Dynamics of population growth within plants may explain limited variation. Annual Review of Phytopathology 41, 199–214. French, R., Stenger, D.C., 2005. Genome sequences of Agropyron mosaic virus and Hordeum mosaic virus support reciprocal monophyly of the genera Potyvirus and Rymovirus in the family Potyviridae. Archives of Virology 150, 299–312.

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Tritimoviruses and Rymoviruses (Potyviridae)

Gibbs, A.J., Gibbs, M.J., 2018. Rymovirus: A cautionary tale. Archives of Virology 163, 815–817. Rabenstein, F., Seifers, D.L., Schubert, J., French, R., Stenger, D.C., 2002. Phylogenetic relationships, strain diversity and biogeography of tritimoviruses. Journal of General Virology 83, 895–906. Revers, F., Garcia, J.A., 2015. Molecular biology of potyviruses. Advances in Virus Research 92, 101–199. Sing, K., Wegulo, S.N., Skoracka, A., Kundu, J.K., 2018. Wheat streak mosaic virus: A century old virus with rising importance worldwide. Molecular Plant Pathology 19, 2193–2206. Skoracka, A., Dabert, M., 2010. The cereal rust mite Abacarus hystrix (Acari: Eriophyidea) is a complex of species: Evidence from mitochondrial and nuclear DNA sequences. Bulletin of Entomological Research 100, 263–272. Skoracka, A., Kuczynski, L., de Mendonca, R.S., et al., 2012. Cryptic species within the wheat curl mite Aceria tosichella (Keofer) (Avari: Eriophyoidea), revealed by mitochondrial, nuclear and morphometric data. Invertebrate Systematics 26, 417–433. Stenger, D.C., French, R., 2004. Complete nucleotide sequence of Oat necrotic mottle virus: A distinct Tritimovirus species (family Potyviridae) most closely related to Wheat streak mosaic virus. Archives of Virology 149, 633–640. Stenger, D.C., Hall, J.S., Choi, I., French, R., 1998. Phylogenetic relationships within the family Potyviridae: Wheat streak mosaic virus and Brome streak mosaic virus are not members of the genus Rymovirus. Phytopathology 88, 782–787. Stenger, D.C., Hein, G.L., Gildow, F.E., Horken, K.M., French, R., 2005a. Plant virus HC-Pro is a determinant of eriophyid mite transmission. Journal of Virology 79, 9054–9061. Szydło, W., Hein, G.L., Denizhan, E., Skoracka, A., 2015. Exceptionally high levels of genetic diversity in wheat curl mite (Acari: Eriophyidae) populations from Turkey. Journal of Economic Entomology 108, 2030–2039. Tatineni, S., Graybosch, R.A., Hein, G.L., Wegulo, S.N., French, R., 2010. Wheat cultivar-specific disease synergism and alteration of virus accumulation during co-infection with Wheat streak mosaic virus and Triticum mosaic virus. Phytopathology 100, 230–238. Tatineni, S., Hein, G.H., 2018. Genetics and mechanisms underlying transmission of Wheat streak mosaic virus by the wheat curl mite. Current Opinion in Virology 33, 47–54. Valli, A.A., Gallo, A., Rodamilans, B., Lopez-Moya, J.J., Garcia, J.A., 2018. The HC-Pro from Potyviridae family: An enviable multitasking helper component that every virus would like to have. Molecular Plant Pathology 19, 744–776. Wang, A.M., Krishnaswamy, S., 2012. Eukaryotic translation initiation factor 4E-mediated recessive resistance to plant viruses and its utility in crop improvement. Molecular Plant Pathology 13, 795–803. Ward, C.W., 2017. Is it time to retire the genus Rymovirus from the family Potyviridae? Archives of Virology 162, 2175–2179. Wosula, E.N., McMechan, A.J., Oliveira-Hofman, C., Wegulo, S.N., Hein, G.L., 2016. Differential transmission of two isolates of Wheat streak mosaic virus by five wheat curl mite populations. Plant Disease 100, 154–158.

Relevant Websites https://talk.ictvonline.org/taxonomy/ Taxonomy International Committee on Taxonomy of Viruses (ICTV).

Triviruses (Betaflexiviridae) Yahya ZA Gaafar and Heiko Ziebell, Julius Kühn Institute – Federal Research Center for Cultivated Plants, Braunschweig, Germany r 2021 Elsevier Ltd. All rights reserved.

Nomenclature aa Amino acid(s) AlkB Alpha-ketoglutarate-dependent dioxygenase CP Coat protein dsRNA Double stranded ribonucleic acid Hel Helicase HTS High-throughput sequencing IC-RT-PCR Immunocapture reverse transcription polymerase chain reaction kb Kilobase kbp Kilobase pair kDa Kilo Dalton LAMP Loop-mediated isothermal amplification ML Maximum-likelihood MP Movement protein Mtr Methyl-transferase

Glossary Capillovirus From Latin capillus (hair). Chordovirus From chord, the name of the strings on a harp or stringed instruments. Citrivirus From the type species Citrus leaf blotch virus. Divavirus From the type species Diuris virus. Fivivirus From Ficus – Vitis, the genera name of the natural hosts of the type species in this newly proposed virus genus.

nm Nanometer nt Nucleotide(s) ORF Open reading frame OTu-Pro Ovarian tumor-like protease Pol RNA-dependent RNA polymerase Poly(A) Polyadenylated P-Pro Papain-like protease qRT-PCR Quantitative reverse transcription polymerase chain reaction RP Replicase RT-PCR Reverse transcription polymerase chain reaction sgRNA Sub-genomic RNA ssRNA Single-stranded ribonucleic acid TGB Triple gene block V-region Variable region

Prunevirus From Prunus, genus name of the apricot (Prunus armeniaca), the natural host of the type species. Tepovirus From the type species Potato virus T. Trichovirus From Greek thrix (hair). Trivirus A virus member of the Trivirinae subfamily. Vitivirus From Vitis, generic name of the grapevine (Vitis vinifera) the host of the type species. Wamavirus From the type species Watermelon virus A.

Introduction The subfamily Trivirinae of the family Betaflexiviridae consists of viruses infecting predominantly woody plants such as fruit trees and grapevine but infection of herbaceous plants (in particular tuber crops) is also possible. Transmission of triviruses occurs through insect vectors, in particular pseudococcid mealybugs, aphids and erineum mites; transmission by grafting, seeds or mechanical means are also common. Triviruses are distributed worldwide; they often occur in mixed infections with other viruses (e.g., closteroviruses) so that it is difficult to correlate the symptomology with the identity of the virus causing these symptoms. Triviruses have flexuous, non-enveloped particles (ranging in size from 640 to 1500 nm) that contain a positive-sense singlestranded RNA (ssRNA) molecule. The genome lengths range from approximately 6.45 to 8.75 kb. The name of the subfamily reflects the three conserved proteins that all members’ genomes commonly encode, i.e., the replicase (RP), movement (MP) and coat proteins (CP). Currently, nine genera are recognized within the Trivirinae subfamily as well as thirty-eight assigned species although a range of newly proposed species is pending approval with constantly increasing numbers. Moreover, a new genus tentatively named “Fivivirus” was proposed recently.

Classification Trivirinae is a subfamily within the family Betaflexiviridae (order: Tymovirales, realm: Riboviria). Nine genera are currently assigned to this subfamily i.e., Capillovirus, Chordovirus, Citrivirus, Divavirus, Prunevirus, Tepovirus, Trichovirus, Vitivirus, and Wamavirus, with thirty-eight assigned virus species (Table 1). Several new members have been proposed recently, including Agave tequilana leaf virus (ATLV), Grapevine virus M (GVM) and Peach virus M (PeVM) (Table 1). Additionally, a new genus “Fivivirus” to include the tentative species i.e., Fig latent virus 1 (FLV1) and Grapevine Kizil Sapak virus (GKSV) was proposed recently. The name of the subfamily reflects the three conserved proteins that all members’ genomes encode in common i.e., the RP, MP, and CP (Fig. 1). This distinguishes them from the members of the Quinvirinae, which have five conserved genes encoding the RP, triple gene block

Encyclopedia of Virology, 4th Edition, Volume 3

doi:10.1016/B978-0-12-809633-8.21533-1

805

Divavirus Unknown

Citrivirus 960
12–15 nm

Chordovirus 640–760
10–12 nm

Capillovirus 670–700
12 nm

Foliar Chlorotic & Necrotic Ringspots/ Rough Vein/Flower Variegation/Mixed Infection Unknown/Mixed Infection Unknown Unknown

Camellia spp. Eriobotrya japonica Hevea brasiliensis Smallanthus sonchifolius

CRSaV3

LoVA RTV1

YVA

Diuris virus Ab Diuris virus B Hardenbergia virus A Ocimum basilicum RNA virus 1c

Unknown Unknown Unknown Unknown

Diuris pendunculata Diuris pendunculata Hardenbergia comptoniana Ocimum basilicum

ObRV1

Actinidia chinensis, Citrus spp., Nagami Abnormal Graft/Leaf Blotching kumquat, Paeonia lactiflora & Prunus avium Citrus tamuranua Leaf Chlorotic Blotching

Unknown Mosaic/Symptomless

Daucus carota Lactuca sativa & L. serriola

DVA DVB HarVA

Citrus leaf blotch CLBV virusb Citrus leaf blotch CLBV2 virus 2c

Unknown

Daucus carota

Symptomless/Mixed Infection Unknown Chlorotic Spots Symptomless/Mixed Infection

CVA CuVA MuVA BCV

CtChV1 Carrot Ch virus 1b Carrot Ch virus 2 CtChV2 LeCV1 Lettuce chordovirus 1c

Cherry virus A Currant virus A Mume virus A Birch capillovirusc Camellia ringspot associated virus 3c Loquat virus Ac Rubber tree virus 1c Yacon virus Ac

Stem grooving/Graft Necrosis & Pitting/ Tatter Leaf/Leaf Deformation/Chlorotic mottle/Leaf Epinasty

Symptoms

Actinidia chinensis, A. deliciosa, Bambusa nana, Bauhinia variegate, Citrus spp., Cnidium officinale, Dendrocalamus dianxiensis, D. hamiltonii, Fargesia somnigensis, Ficus palmata, Malus spp., Nandina domestica, Phyllostachys pubescens, Prunus spp., Pyrus communis, P. pyrifolia, Rosa spp. & Rubus ellipticus Prunus spp. Ribes rubrum Prunus mume Betula pubescen & B. pendula

ASGV Apple stem grooving virusb

Acronym Natural Hosts

Unknown

Italy

Australia Australia Australia

China

Unknown

UK France

UK

Poland

China China

USA

Worldwide Czech Republic Japan Finland & Germany

Worldwide

Unknown Unknown Mechanically-transmitted

Accessiona

NC035462

NC019029 NC019030 NC015395

MH144344

NC003877

NC025468 NC040627

NC025469

NC030657

MK936045 MN047299

MK050795

NC003689 NC029301 NC040568 MK402233

Worldwide in apple NC001749 & citrus producing countries

Distribution

Graft-transmitted

Unknown Unknown

Unknown

Unknown

Unknown Unknown

Seed-transmitted

Graft-transmitted Unknown Graft-transmitted Graft-transmitted

Graft-& seed-transmitted (lily, Chenopodium quinoa)

Transmission

Overview of the genera assigned of the subfamily Trivirinae (Betaflexiviridae) including type species and biological properties (ICTV, 2019)

Genus Particle size Virus name

Table 1

806 Triviruses (Betaflexiviridae)

Fig latent virus 1,b,d Grapevine Kizil Sapak virusc

Vein Chlorotic Spots/Leaf Reddening Foliar Chlorotic & Necrotic Ringspots/ Rough Vein/Symptomless/Mixed Infection Foliar Chlorotic & Necrotic Ringspots/ Rough Vein/Symptomless/Mixed Infection Vein Necrosis/Chlorotic Spots/Top Necrosis/Symptomless Unknown Unknown

Prunus amygdalus Camellia spp. Camellia spp.

Chenopodiaceae, Fabaceae, Solanaceae & other herbaceous hosts Prunus spp. Zostera muelleri

CPrV

CRSaV1

CRSaV2

PVT

Grapevine berry inner necrosis virus Grapevine Pinot gris virus Peach mosaic virus

Mechanically- & grafttransmitted

Mechanically- & seedtransmitted Unknown Unknown

Seed-transmitted

Seed-transmitted

Unknown

Unknown

Seed-transmitted

Mechanically & seedtransmitted Unknown

Chlorotic Mottle/Distortion & Puckering/ Late Ripening/Fruit Wart-like Outgrowths/Symptomless Leaf Mottle/Ring Spot/Mosaic/Necrotic Spots Mottle/Leaf Chlorosis/Leaf Deformation/ Symptomless Retarded Foliation/Leaf & Fruit Deformation/Stunting /Flower Breaking

Vitis spp. Vitis riparia & V. vinifera Prunus spp.

GINV

GPGV

PcMV

MN172165

FN377573

Asia, Australia & Europe

Worldwide

Azerbaijan & Italy Australia

South America

USA

USA

Azerbaijan

Australia, China, Europe

Mechanically-, graft-transmitted Canada, Mexico & USA & by peach bud mite (Eriophyes insidious)

(Continued )

NC011552

NC015782

NC015220

NC002500

NC006946

NC001409

NC024686 MK514426

NC011062

MK050793

MK050792

NC038325

NC023295

China, New Zealand NC040800 & South Korea

Turkmenistan

Worldwide

Mechanically-, graft-transmitted China, Canada & & by Eriophyes inaequalis USA eriophyid mites Mechanically-, graft-transmitted China & Japan & by erineum mite (Colomerus vitis) Erineum mite (Colomerus vitis) Worldwide

Stem Pitting/Stem Grooving/Symptomless Mechanically- & grafttransmitted

Prunus spp.

Apricot pseudochlorotic leaf spot virus Cherry mottle leaf CMLV virus

Chanenomeles japonica, Cydonia oblonga, Malus domestica, Prunus domestica, P. persica, Pyrus communis & Sorbus aucuparia APsCLSV Prunus spp.

Mottle/Ringspot/Line Patterns/Fruit Deformation/Symptomless

Unknown

Prunus spp.

AVCaV

PrVT ZVT

Vein Chlorosis/Mottle/Symptomless

Actinidia spp.

Symptomless

Vitis vinifera

GKSV

ASbLV

Mosaic/Stunting/Symptomless

Ficus carica

FLV1

Trichovirus ACLSV 680–780
9.5–12 Apple chlorotic nm leaf spot virusb

Prunus virus T Zostera virus Tc

Prunevirus 950–1500
13 nm Actinidia seed borne latent virus Apricot vein clearing associated virusb Caucasus prunus virus Camellia ringspot associated virus 1c Camellia ringspot associated virus 2c Tepovirus 640
12 nm Potato virus Tb

Fivivirusa B700 nm

Triviruses (Betaflexiviridae) 807

Continued

Vitivirus 725–825
12 nm

Unknown Yellow Mottle/Chlorotic Ringspot

Prunus persica Prunus spp.

PCLSV

PeVM

Arracacia xanthorrhiza Rubus spp. Vitis vinifera Vitis spp.

AcVB

AVV BVA

GVA

GVB

Actinidia virus B

Arracacha virus V Blackberry virus A Grapevine virus Ab Grapevine virus B

Mechanically- & grafttransmitted Mechanically- & grafttransmitted Unknown Unknown

Unknown

Unknown

Graft-transmitted

Mechanically-transmitted

Transmission

Unknown Green Mosaic

Unknown

Vitis vinifera Vitis spp. Vitis vinifera Vitis vinifera Vitis vinifera Vitis vinifera Vitis vinifera Apiaceae spp. Mentha x gracilis Agave tequilana Arracacia xanthorrhiza Vaccinium spp.

Vitis vinifera

Grapevine virus E GVE

GVF GVG GVH GVI GVJ HLV

MV2 ATLV

ALVV

BGMaV

GVK

Grapevine virus F Grapevine virus G Grapevine virus H Grapevine virus I Grapevine virus J Heracleum latent virus Mint virus 2 Agave tequilana leaf virusc Arracacha latent virus Vc Blueberry green mosaicassociated virusc Grapevine virus Kc

Unknown

Unknown

Unknown

Stem Pitting & Grooving/Mixed infections Mechanically-, graft-transmitted & by mealybugs Corky Bark/Symptomless Mechanically-, graft-transmitted & by mealybugs Corky Rugose Bark Mechanically- & grafttransmitted Unknown Mechanically- & grafttransmitted Unknown Unknown Unknown Unknown Symptomless Unknown Symptomless Unknown Symptomless Unknown Symptomless Mechanically- & aphid transmitted Symptomless Aphid transmitted Unknown Unknown

Grapevine virus D GVD

Actinidia spp.

AcVA

Vein Chlorosis/Flecking/Ringspots/ Symptomless Leaf Vein Chlorosis/Flecking/Ringspots/ Vein Clearing/Mottle/Symptomless Mild Mosaic Chlorosis/Leaf Deformation Unknown

Symptomless

Prunus spp.

CLV1

Actinidia spp.

Mosaic/Yellow Flecking/Stunting

Phlomis spp.

Symptoms

PhMV

Acronym Natural Hosts

Actinidia virus A

Phlomis mottle virus Cherry latent virus 1c Peach chlorotic leaf spot virusc Peach virus Mc

Genus Particle size Virus name

Table 1

MK012336

MH084695

MK770441

NC043412

Accessiona

NC011106

NC038326

NC003602

NC003604

Croatia & Italy

USA

Peru

USA Mexico

NC035202

MK460433

KY451036

NC043088 NC034833

Worldwide NC018458 New Zealand & USA NC040554 Worldwide NC040545 New Zealand & USA NC037058 Turkmenistan NC040564 Europe NC039087

Worldwide

Brazil & Europe

Worldwide

Worldwide

China, New Zealand NC043087 & South Korea China, New Zealand NC016404 & South Korea Brazil NC034264 USA NC040630

Mexico, USA

China

Georgia

Greece & Italy

Distribution

808 Triviruses (Betaflexiviridae)

GVL

Watermelon virus WVA Ab

Citrullus lanatus

Vitis Symptomless vinifera Vitis spp. Unknown Crinkling/Mosaic

Unknown

Unknown

b

proposed genus. type species. c tentative species. d it has been proposed to remove Fig latent virus 1 from the genus Trichovirus to the newly proposed genus Fivivirus.

a

Wamavirus Unknown

Grapevine virus Mc GVM

Grapevine virus Lc

Unknown

China, Croatia, New Zealand & USA USA China

MK492703

MH248020

NC034377

Triviruses (Betaflexiviridae) 809

810

Triviruses (Betaflexiviridae)

Fig. 1 Graphical representation of the genomic organization of selective members of the family Betaflexiviridae showing the relative positions of the ORFs and their gene products. The motifs are Mtr: methyltransferase; OTu-Pro: ovarian tumor-like protease; AlkB: alpha-ketoglutaratedependent dioxygenase; P-Pro: papain-like protease; Hel: helicase; Pol: RNA-dependent RNA polymerase; ?: cap structure has not been demonstrated yet. Trivirinae: ACLSV: Apple chlorotic leaf spot virus, ASGV: Apple stem grooving virus, AVCaV: Apricot vein clearing associated virus, CLBV: Citrus leaf blotch virus, DVA: Diuris virus A, GKSV: Grapevine Kizil Sapak virus, GVA: Grapevine virus A, LeCV1: Lettuce chordovirus 1, PVT: Potato virus T, WVA: Watermelon virus A. Quinvirinae: ASPV: Apple stem pitting virus, CNRMV: Cherry necrotic rusty mottle virus and RCVMV: Red clover vein mosaic virus.

(TGB) and CP. The RP of triviruses belongs to the “alphavirus-like” supergroup of RNA viruses. The RP proteins are most closely related to those of other virus families in the order Tymovirales whereas the MPs of the “30K” superfamily are more closely related to those in the family Virgaviridae. For taxonomic assignment, host range, serological specificity, nt (less than 72%) or aa (less than 80%) identity of CP or RP as well as vector specificity have been used to distinguish species.

Virion Structure The virions of triviruses are non-enveloped flexuous filaments with helical symmetry (with a pitch of approximately 3.3–3.8 nm). Depending on the negative contrast material used for electron microscopy, cross-banding, criss-cross or rope-like features can be visible. The virions are composed of a ssRNA molecule (representing approximately 5% of the virion weight) and a polypeptide (ranging in size from 18 to approximately 41 kDa). The length of the virions is genus-dependent and can vary from 640 to 1500 nm (Table 1) with a diameter ranging from 10 to 15 nm. Fig. 2 shows the virions of Potato virus T (PVT), a tepovirus. To maintain the integrity of the particles, divalent cations are required.

Triviruses (Betaflexiviridae)

811

Fig. 2 Electron micrograph of Potato virus T (PVT; Tepovirus) particles. The micrograph shows non-enveloped flexuous filaments virions with helical symmetry. Courtesy: Katja R. Richert-Pöggeler, Julius Kuehn Institute, Germany.

Genomic Organization and Properties of Encoded Proteins The genomes of triviruses are encoded by a linear positive-sense ssRNA with two to five open reading frames (ORF) (Fig. 1). The 50 terminus has a 7-methyl-guanylate cap (m7G) but this has not been confirmed for all genera. The 30 terminus is polyadenylated (poly(A)). The lengths range from approximately 6.45 to 8.75 kb excluding the poly(A) tail. Genomes of triviruses have three conical genes encoding the RP, MP, and CP. ORF1 encodes the RP with a viral methyltransferase (Mtr) motif at the N-terminal region, and viral helicase 1 (Hel) and RNA-dependent RNA polymerases (Pol) core motifs near the C-terminal region (Fig. 1). Additional motifs were reported in most members i.e., alpha-ketoglutarate-dependent dioxygenase (AlkB) and papain-like protease (p-Pro) motifs between the Mtr and Hel motifs. The ovarian tumor-like protease motif (OTu-Pro) could only be found on ORF1 of citriviruses (Fig. 1). A second ORF encodes a cell-to-cell MP of the “30K” superfamily (B31 to B50 kDa). The third conserved ORF of triviruses encodes a CP of approximately 21–40.5 kDa. The genomic organization of capilloviruses and divaviruses differs from the other genera in the subfamily Trivirinae. They have only two overlapping ORFs (Fig. 1). Their ORF1 encodes a polyprotein containing the RP and the CP. A “variable region (V-region)” can be found between the Pol motif (within the RP) and the CP where the aa sequence shows high variability among different virus isolates (Fig. 1). A second ORF, encoding the MP, is nested within the ORF1. An exception of capilloviruses is the recently identified Loquat virus A (LoVA). It possesses a genome composed of three ORFs, where the CP coding ORF3 is separated from the RP coding ORF1. Chordoviruses, pruneviruses, vitiviruses, and some trichoviruses have, in addition to the RP, MP, and CP, a 30 -terminal ORF encoding a putative nucleic acid binding protein (NABP) with a zinc binding finger motif (approximately 10.2–17.1 kDa) (Fig. 1). Citriviruses, tepoviruses, and some trichoviruses have only the three conical genes. In vitiviruses, there is an additional ORF encoding a hypothetical protein (B17.2 to B24.8 kDa) between the RP and the MP (Fig. 1). In wamaviruses, another ORF is located between the MP and the CP encoding a hypothetical protein of approximately 25 kDa. The genome organization of the two putative members of the proposed genus Fivivirus FLV1 and GKSV are different. Four predicted, slightly overlapping ORFs were identified in the FLV1 genome, the putative RP, MP (43 kDa), CP (46 kDa), and NABP (12 kDa). GKSV has five predicted ORFs where the RP (196.7 kDa) is followed by the MP (31.7 kDa) which is slightly overlapped by the CP (22 kDa) and a hypothetical protein (21.4 kDa) is overlapped with a predicted NABP which overlap (11.7 kDa).

Phylogenetic Relationships of the Different Triviruses Phylogenetic analysis based on RPs alignment of betaflexivirids shows that the triviruses of each genus are grouped together, mostly within well-supported branches (Fig. 3(A)). A slightly different picture emerges when using aa alignments of the MP: The MP of the citriviruses i.e., Citrus leaf blotch virus (CLBV) and Citrus leaf blotch virus 2 (CLBV2) form a group with pruneviruses. As triviruses have a “30K”-type MP while quinviruses have the TGB, quinviruses are not included in the tree. The closely related Furovirus genus (family: Virgaviridae) was used instead for the Maximum-likelihood (ML) tree (Fig. 3(B)). Fig. 3(C) shows an ML tree representing the phylogenetic relationship between the CP aa sequence of selective members of the subfamily Trivirinae and Quinvirinae. Interestingly, citriviruses and the wamavirus WVA are grouped with quinviruses (Fig. 3(C)).

812

Triviruses (Betaflexiviridae)

Triviruses (Betaflexiviridae)

813

Cellular Localization of Trivirus Particles and Viral Replication Trivirus replication and assembly occur in the cytoplasm. Virions can be found in the cytoplasm of infected mesophyll, parenchyma, and phloem cells of leaves and roots. The cellular localization however is virus- and host-dependent. For example, virions of Grapevine virus A (GVA), Grapevine virus B (GVB) and Heracleum latent virus (HLV) (vitiviruses), and Apple chlorotic leaf spot virus (ACLSV; a trichovirus) are strictly phloem-limited, but in herbaceous hosts, they can be found in the parenchyma. Moreover, virions of ACLSV could be located in the nucleus. Trivirus particles accumulate in discrete bundles or paracrystalline aggregates in the cytoplasm where multiplication occurs. Trivirus-infected cells were found to be damaged to a varying extent. Some triviruses were found to cause tonoplast vesicular evagination where fine fibrillar material (suggested to be the replicative forms of the viral RNA) can be observed. Other cellular damages include enlarged starch grains, swollen chloroplasts and damaged mitochondria. Moreover, a study showed that ASGV infection of apple trees extensively alters the gene expression patterns even with no disease symptoms observed. The expression of trivirus proteins relies on production of sub-genomic RNA (sgRNA) as suggested by the analyses of dsRNA patterns from infected plants. As an example, five dsRNA species were detected in plant tissues infected with apple stem grooving virus (ASGV; a capillovirus) i.e., 6.5, 5.5, 4.5, 2.0, and 1.0 kbp. The 6.5 kbp species represents the dsRNA of the full genome of ASGV. The 5.5 and 4.5 species are the 50 co-terminal with ASGV genomic RNA while the 2.0 and 1.0 kbp are suggested to be dsRNA of sgRNA of the MP and CP, respectively. Six dsRNAs species with sizes of about 7.5, 6.4, 5.4, 2.2, 1.1, and 1.0 kbp were detected in ACLSV-infected plant tissues. The largest dsRNA species here represent the full genome of the virus, the 6.4 and 5.4 dsRNA species are the 50 co-terminal with ACLSV genomic RNA. The dsRNA 2.2 and 1.1 kbp species are 30 co-terminal with ACLSV genomic RNA which could be representing the sgRNA of the MP and the CP, respectively. The 1.0 kbp species is internal to the genome. It is suggested that the MP of triviruses acts as a suppressor of plant RNA silencing pathways. For example, ACLSV-MP suppresses the systemic movement of the silencing signal whereas CLBV-MP suppresses the intracellular silencing without affecting the systemic spread of the signals.

Host Range and Distribution of Triviruses Triviruses infect predominantly grapevine, citrus fruit and fruit trees/shrubs and therefore they can be found worldwide. The host range of triviruses appears to be quite narrow under natural conditions, infecting only one or few related plants species. However, under experimental conditions it is possible to transmit many triviruses mechanically to herbaceous indicator plants such as Nicotiana or Chenopidium spp. (see Table 1 for details on natural hosts). In recent years, many new triviruses have been described that infect herbaceous plants. For a long time, the tepovirus PVT (identified from potato crops in South America and classified in many countries as quarantine pest) and Heracleum latent virus (HLV) that was described from hogweed (Heracleum sphondylium) Fig. 3 Maximum-likelihood (ML) trees of members of the subfamily Trivirinae (Betaflexiviridae) based on alignments of the amino acid sequences of (A) the replicase, (B) the movement, (C) the capsid proteins, and their homologs from representative members of subfamily Quinvirinae (Betaflexiviridae) and the genus Furovirus (Virgaviridae). MEGAX was used to perform MUSCLE alignments and for producing the ML trees (based on Jones-Taylor-Thornton (JTT) matrix-based model). The numbers on branches indicating the percentage of bootstraps (1000 bootstrap replications; only 450% are shown). The viruses and their accession numbers are: Betaflexiviridae: Trivirinae; Capillovirus: Apple stem grooving virus (ASGV; NC001749), Camellia ringspot associated virus 3 (CRSaV-3; MK050795), Cherry virus A (CVA; NC003689), Currant virus A (CuVA; NC029301), Loquat virus A (LoVA; MK936045), Mume virus A (MuVA; NC040568), Rubber tree virus 1 (RTV1; MN047299), Yacon virus A (YVA; NC030657), Chordovirus: Carrot Ch virus 1 (CtChV1; NC025469), Carrot Ch virus 2 (CtChV2; NC025468), Citrivirus: Citrus leaf blotch virus (CLBV; NC003877), Citrus leaf blotch virus 2 (CLBV2; MH144344), Divavirus: Diuris virus A (DVA; NC019029), Diuris virus B (DVB; NC019030), Hardenbergia virus A (HarVA; NC015395), Ocimum basilicum RNA virus 1 (ObRV1; NC035462), Fivivirus (*proposed genus): Grapevine Kizil Sapak virus (GKSV; MN172165), Prunevirus: Apricot vein clearing associated virus (AVCaV; NC023295), Camellia ringspot associated virus 1 (CRSaV-1; MK050792), Camellia ringspot associated virus 2 (CRSaV2; MK050793), Caucasus prunus virus (CPrV; NC038325), Tepovirus: Potato virus T (PVT; NC011062), Prunus virus T (PrVT; NC024686), Trichovirus: Apple chlorotic leaf spot virus (ACLSV; NC001409), Apricot pseudo-chlorotic leaf spot virus (APsCLSV; NC006946), Cherry mottle leaf virus (CMLV; NC002500), Grapevine berry inner necrosis virus (GINV; NC015220), Grapevine Pinot gris virus (GPGV; NC015782), Peach mosaic virus (PcMV; NC011552), Peach virus M (PeVM; MK012336), Vitivirus: Actinidia virus A (AcVA; LC491285), Actinidia virus B (AcVB; NC016404), Agave tequilana leaf virus (ATLV; NC034833), Arracacha virus V (AVV; NC034264), Blackberry virus A (BVA; NC040630), Blueberry green mosaic associated virus (BGMaV; MK460433), Grapevine virus A (GVA; NC003604), Grapevine virus B (GVB; NC003602), Grapevine virus D (GVD; MF774336), Grapevine virus E (GVE; NC011106), Grapevine virus F (GVF; NC018458), Grapevine virus G (GVG; NC040554), Grapevine virus H (GVH; NC040545), Grapevine virus I (GVI; NC037058), Grapevine virus J (GVJ; NC040564), Grapevine virus K (GVK; NC035202), Grapevine virus M (GVM; MK492703), Heracleum latent virus (HLV; MN314973), Wamavirus: Watermelon virus A (WVA; NC034377). Betaflexiviridae: Quinvirinae; Carlavirus: Potato virus M (PVM; NC001361), Hop latent virus (HpLV; NC002552), Garlic latent virus (GLV; NC003557), Red clover vein mosaic virus (RCVMV; NC012210), Foveavirus: Apple stem pitting virus (ASPV; NC003462), Peach chlorotic mottle virus (PCMV; NC009892), Apricot latent virus (ApLV; NC014821), Apple green crinkle associated virus (AGCaV; NC018714), Robigovirus: Cherry green ring mottle virus (CGRMV; NC001946), Cherry necrotic rusty mottle virus (CNRMV; NC002468), African oil palm ringspot virus (AOPRV; NC012519), Cherry rusty mottle associated virus (CRMaV; NC020996). Virgaviridae: Furovirus: Oat golden stripe virus (OGSV; NC002358), Chinese wheat mosaic virus (CWMV; NC002359), Sorghum chlorotic spot virus (SrCSV; NC004014), Japanese soil-borne wheat mosaic virus (JSBWMV; NC038850).

814

Triviruses (Betaflexiviridae)

in Europe were the only triviruses found naturally on herbaceous plants. New triviruses found on non-woody plants include Yacon virus A (YVA; a capillovirus) that infects tuberous plants (Smallanthus sonchifolius); Carrot chordovirus 1 and 2 (CtChV1 and CtChV2) as well as Lettuce chordovirus 1 (LeCV1; Chordovirus) that were identified from carrots (in the UK) and lettuce (from France), respectively; Diuris virus A and B (DVA and DVB) from donkey orchids (Diuris penduculata) and Hardenbergia virus A (HVA; Divavirus) from the legume Hardenbergia comptoniana, from Australia; Zosteria virus T (ZVT; Tepovirus) that infects Zostera muelleri, a seagrass species that can be found in regions of Australia, New Zealand, and Papua New Guinea; Phlomis mottle virus (PhMV; Trichovirus) infecting Phlomis ssp. in the Mediterranean region; Agave tequilana leaf virus (ATLV; Vitivirus) infecting Agave tequilana in Mexico, Arracacha virus V (AVV) and Arracacha latent virus (ALVV; Vitivirus) infecting the South American root crop Arracacia xanthorrhiza; Mint virus 2 (MV2; Vitivirus) from mint in USA and Watermelon virus A (WVA) infecting watermelon (Citrullus lanatus) in China. Recently discovered triviruses include also LoVA from loquat (Eriobotrya japonica) in China, Rubber tree virus 1 (RTV1; Capillovirus) infecting rubber trees (Hevea brasiliensis) in China, Camellia ringspot associated virus 1, 2 and 3 (CRSaV1, CRSaV2; Prunevirus, and CRSaV3; Capillovirus) from Camellia spp. in the USA, and Blueberry green mosaic-associated virus” (BGMaV; Vitivirus) from blueberries in the USA. Only limited information is available on alternative hosts or distribution of these viruses in other regions.

Symptomology, Impact and Vectors Common symptoms in fruit trees include graft incompatibilities, necrosis and pitting of the graft union and stem grooving of wood; leaf symptoms may include chlorotic spots, line patterns or ring spots, mottling, mosaic or necrotic spots (Fig. 4). For example, ACLSV causes symptoms like bark split and dark green sunken mottle (Fig. 4(A) and (B)), AVCaV causes symptoms of

Fig. 4 Symptoms induced by triviruses. (A) Symptoms of bark split caused by Apple chlorotic leaf spot virus (ACLSV) in plum (cv. Prune d0 Ente), (B) Typical dark green sunken mottle caused by an ACLSV isolates in the GF305 peach seedling indicator, (C) Symptoms of vein clearing and blotching observed on apricot leaf infected with Apricot vein clearing-associated virus (AVCaV), (D) Vitis berlandieri
V. riparia (indicator plants) expressed stem grooving disease associated with infection by Grapevine virus A (GVA); Bottom is healthy control, (E) Couderc 1613
V. berlandieri (indicator plants), developed corky-bark symptoms associated with infection by Grapevine virus B (GVB); Bottom is healthy control. Courtesy: (A) Jean Dunez, INRAE, France, (B) Thierry Candresse, INRAE, France, (C) Toufic Elbeaino, Istituto Agronomico Mediterraneo di Bari, Italy, (D) and (E) Maher Al Rwahnih, Foundation plant Services – University of California – Davis, USA.

Triviruses (Betaflexiviridae)

815

vein clearing and blotching on apricot leaves (Fig. 4(C)), and color breaking of petals was observed for PcMV infection. Fruits may be smaller, ripen late or may have wart-like outgrowths or injuries. The majority of grapevine-infecting triviruses does not cause visible symptoms on the plants; some infections may lead to necrotic spots, mosaic, mottle or ring spots on leaf but more dramatic is the effect on yield quantity and quality e.g., of Grapevine Pinot gris virus (GPGV) on berry quantity and quality that may be both reduced in these economically important crops. As another example, GVA is associated with pitting and grooving of wood but occurs very often in mixed infections with other viruses (Fig. 4(D)); corky bark disorder was described for GVB infection (Fig. 4(E)) whereas Grapevine virus D (GVD) is associated with corky rugose wood disease. Both GVA and GVB are associated with the Kober stem grooving syndrome. However, for many triviruses, no specific symptoms were observed, e.g., HLV may cause no symptoms at all. Many of the recently added triviruses where identified during virus ecology studies using high-throughput sequencing technologies (HTS) of pooled samples and therefore symptoms cannot be attributed to the identified viruses. Mixed infections of triviruses and in particular with closteroviruses (family: Closteroviridae) are quite common in nature; multiple infections of one plant with several triviruses have also been reported. An example is the occurrence of Grapevine leafroll-associated viruses (GLRaVs; closteroviruses) and the vitiviruses GVA, GVB, GVD, GVE, and GVF in mixed infections of vineyards. As another example, the Plum bark necrosis stem pitting-associated virus (PBNSPaV, Closterovirus) and Apricot vein clearing associated virus (AVCaV, Prunevirus) can be found in dual/mixed infections. Therefore, it is difficult to assign a certain symptomology to infection with a specific virus. Most, if not all, fruit tree- and grapevine-infecting triviruses are graft-transmissible; many triviruses could be transferred mechanically under experimental conditions to herbaceous indicator plants. Transmission via seeds was reported in few studies e.g., PVT, CRSaV1, CRSaV2, CRSaV3, and ASGV (for ASGV, seed-transmission was shown only for indicator plants e.g., lily and Malus platycarpa, but it is unclear if this dissemination route is of major importance under natural conditions). Insect vectors appear to have a more prominent role in transmission of triviruses; HLV and Mint virus 2 (MV2) can be transmitted by aphids; fruit tree and grapevine-infecting triviruses can be transmitted by eriophyid mites (Eriophyes inaequalis: transmitting CMLV; Eriophyes insidiosa: transmitting PcMV), erineum mites (Colomerus vitis: transmitting GINV and GPGV) and various mealybug species (Pseudococcus longispinus, Ps. affinis, Planococcus ficus, Pl. citri, Phenacoccus aceris) and scale insects (Neopulvinaria innumerabilis) that are involved in the transmission of GVA, predominantly. It appears that L1 stages of Ph. aceris are more efficient vectors than L2, L3 or adult females although the reasons are unknown. They are transmitted in semi-persistent manner. Moreover, HLV was found to require Heracleum virus 6 (HV6; synonym of Carrot yellow leaf virus [CYLV], a closterovirus) as a helper for vector transmission. For many of the novel triviruses, information on vectors or alternative routes of transmission is missing (Table 1).

Diagnosis Many of the early investigations on triviruses used test plants as bioassays for induction of symptoms and propagation of viruses. However, these bioassays cannot be used for identification and characterization of triviruses and are laborious. It is possible to identify virions by various electron microscopy techniques, i.e., transmission electron microscopy, immunogold labeling, or immunosorbent electron microscopy. Depending on the genus, trivirus virions are poor to moderate antigens. Several polyclonal antisera and monoclonal antibodies have been developed for the detection and identification of several triviruses. Cross-reaction of antibodies can occur; e.g., antibodies raised against CMLV cross-react with PcMV. Other monoclonal antibodies have been produced for the specific identification of GVD; those antibodies did not cross-react with other vitiviruses. Various PCR-based assays have been developed for the sensitive and reliable detection of triviruses. Depending on primer design, these assays were successfully implemented to detect specifically individual viral species but the use of degenerate primers such as dPR1 and dPR2 allowed the detection of several vitiviruses (GVA, GVB, GVD, HLV), a tepovirus (PVT) and a divergent ACLSV isolate (Trichovirus). Improved sensitivity compared to a conventional RT-PCR assay was achieved using an immunocapture RT‐PCR (IC-RT-PCR) assay for the detection of ASGV. In this assay, polyclonal antibodies were used to trap the virus particles followed by RT-PCR using ASGV specific primers. However, these assays have mainly been replaced with multiplex assays targeting multiple viruses and/or quantitative RT-PCR (qRT-PCR) protocols. As many triviruses are present in mixed infections, several multiplex detection assays have been developed, in particular for grapevine and fruit trees. For example, multiplex RT-PCR and qRT-PCR assays were designed for the simultaneous detection of several grapevine viruses including GVA and GVB. For a Canadian survey of grapevines, a multiplex RT-PCR assay targeting seventeen grapevine viruses (including GVA, GVB, GPGV) was generated and successfully applied in the detection of the triviruses in the Ontario region. For fruit trees, RT-PCR protocols for the simultaneous detection and identification of eight stone fruit viruses (including ACLSV), four cherry viruses (including CVA), and pear viruses (including ASGV or ACLSV) have been reported. A single qRT-PCR assay for simultaneous detection of GVA, GVB, and GVD was more sensitive and faster than a conventional RT-PCR assay that used degenerate primers to detect these viruses. Similar qRT-PCR protocols were for example developed for the detection of citrus viruses such as Citrus tristeza virus (Closteroviridae), Citrus psorosis virus (genus Ophiovirus, family Aspiviridae) and CLBV or the detection of ASGV, Apple stem pitting virus (Foveavirus) and Apple mosaic virus (Ilarvirus). Recently, loop-mediated isothermal amplification (LAMP) assays have also been established for fast and sensitive detection of triviruses and have been applied for the detection of ACLSV.

816

Triviruses (Betaflexiviridae)

The biggest challenge, however, for the development of specific and sensitive detection assays lies in the development of HTS technologies and the subsequent description of novel virus species and divergent isolates of known viruses. Many new triviruses have been detected recently using HTS (e.g., Grapevine virus M (GVM), WVA, AVCaV, etc.); diagnostic assays need to either be adapted to reflect the detection of these new species or have to be developed from scratch.

Prevention and Treatment Grapevine and fruit trees, the major host groups of viruses belonging to the Trivirinae, are crops of high economic value. They are also perennial plants, and often the highest yield or quality of fruits is only achieved after a few years of planting. The effect of infection with triviruses on these crops may not only be seen in quantitative yield reduction but also in qualitative damage to fruits or secondary metabolites influencing, e.g., the fruit aroma. Most of the preventative and curative measurements apply generally to viruses and viroids. Foremost, preventative measures such as quarantine programs and certification schemes have been adapted to provide healthy root stocks and bud wood as well as other planting material. Rules and recommendations for the provision of “virus-free” material are set by numerous organizations and networks world-wide for grapevine, stone and citrus fruits (e.g., International Council for the Study of Virus and Virus-like Disease of the Grapevine (ICVG), National Clean Plant Network (NCPN), Canadian Plant Protection Export Certification Program (PPECP), Commission of the European Union, North American Plant Protection Organization (NAPPO), European and Mediterranean Plant Protection Organization (EPPO), etc.). However, correct identification of (latent) infection is often hampered by the lack of sensitive detection methods or the presence of unknown viruses or divergent isolates for which a specific test has not been developed yet. Nevertheless, the prophylactic measures can provide healthy propagation material including seeds from which new orchards/vineyards can be planted. Eradication of infected plants might be successful if detected early and limited to a small geographic region. Secondary spread of the disease may be slowed down but in practice, it has been difficult to eradicate the pathogens completely, in particular if vectored by (fast moving) insects. Vector control using insecticides has become increasingly difficult to apply due to the detrimental effects on non-target organisms and the environment, and the development of insecticide-resistant vectors themselves. Additionally, in certain areas of the world there is a political desire to reduce the use of synthetic pesticides. Biological antagonists (such as predatory insects) of trivirus vectors (mites, mealy bugs and scale insects) are poorly studied, especially under field conditions. This knowledge gap whether antagonists can reduce vector populations thus reducing virus transmission needs to be closed in the future. Only few curative measurements for virus elimination of plant material have been described. Some of these have been successfully applied to eradicate trivirus infection; in particular, elimination of GVA seems to be quite successful. These methods include thermotherapy, meristem tip/shoot tip cultures, somatic embryogenesis, chemotherapy (using ombuin, glycyrrhizin/ quercetin, ribavirin, and quercetin/ribavirin), and electrotherapy, applied either alone or in combination. However, these methods may induce undesired by-effects such as phytotoxicity, putative mutagenic effects and the induction of somaclonal variation. To date, no natural resistance against triviruses has been described. Transgenic approaches showed some limited effects in the control of triviruses: In model plants (such as N. benthamiana) reduction of virus titer and absence of virus symptoms was observed in plants expressing GVA-CP or GVA/GVB-MPs. N. benthamiana plants expressing a GVA mini-replicon were extremely resistant to GVA infection but susceptible to GVB. Interestingly, N. occcidentalis plants that were expressing ACLSV-CP were resistant to GINV infection but not to ASGV or ASPV infection. Furthermore, partially functional deletion mutants of ACLSV also conferred resistance to GINV, probably due to interference with viral long-distance movement. An artificial microRNA derived from grapevine was engineered to target GVA genomic sequences in N. benthamiana thus inducing various degrees of resistance in that host. Although it was possible to transform grapevine embryonic tissue with MP genes from GVA or GVB, the resulting plants showed phenotypic abnormalities. Attempts to use the CP genes from GVA or GVB for transformation grapevine seems to be not successful. It therefore seems that due to practical and political reasons, virus-resistance introduced by transgenic approaches is currently not a short-term option.

Further Reading Al Rwahnih, M., Alabi, O.J., Hwang, M.S., Stevens, K., Golino, D., 2019. Identification and genomic characterization of grapevine Kizil Sapak virus, a novel grapevine-infecting member of the family Betaflexiviridae. Archives of Virology 164 (12), 3145–3149. doi:10.1007/s00705-019–04434-3. Barba, M., Ilardi, V., Paquini, G., 2015. Control of pome and stone fruit virus diseases. Advances in Virus Research 91, 47–83. doi:10.1016/bs.aivir.2014.11.001. Lee, R.F., 2015. Control of virus diseases of citrus. Advances in Virus Research 91, 143–173. doi:10.1016/bs.aivir.2014.10.002. Maliogkla, V.I., Martelli, G.P., Fuchs, M., Katis, N.I., 2015. Control of viruses infecting grapevine. Advances in Virus Research 91, 175–227. doi:10.1016/bs.aivir.2014.11.002. Marais, A., Faure, C., Mustafayev, E., Candresse, T., 2015. Characterization of new isolates of Apricot vein clearing-associated virus and of a new Prunus-infecting virus: Evidence for recombination as a driving force in Betaflexiviridae evolution. PLOS One 10 (6), e0129469. doi:10.1371/journal.pone.0129469. Maree, H.J., Blouin, A.G., Diaz-Lara, A., et al., 2020. Status of the current vitivirus taxonomy. Archives of Virology 165, 451–458. doi:10.1007/s00705-019-04500-w. Zherdev, A., Vinogradova, S., Byzova, N., et al., 2018. Methods for the diagnosis of grapevine viral infections: A review. Agriculture 8 (12), 195. doi:10.3390/agriculture8120195.

Triviruses (Betaflexiviridae)

Relevant Websites https://inspection.gc.ca/plant-health/plant-pests-invasive-species/directives/date/d-97-06/eng/1312330811581/1312331075782 Directive D-97-06: Plant Protection Export Certification Program for Grapevine Nursery Stock, Vitis spp. https://ec.europa.eu/info/index_en European Commission, official website. https://www.eppo.int/ European and Mediterranean Plant Protection Organization (EPPO). http://icvg.org/ International Council for the Study of Viruses and Virus-like Diseases of the Grapevine (ICVG). http://nationalcleanplantnetwork.org/ National Clean Plant Network. https://www.nappo.org/ North American Plant Protection Organization (NAPPO).

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Tymoviruses (Tymoviridae) Rosemarie W Hammond and Peter Abrahamian, Agricultural Research Service, US Department of Agriculture, Beltsville, MD, United States r 2021 Elsevier Ltd. All rights reserved. This is an update of A.-L. Haenni, T.W. Dreher, Tymoviruses, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00523-9.

Nomenclature

ORF Open reading frame RdRp RNA-dependent RNA polymerase RP Replication protein sgRNA Sub-genomic RNA ssRNA Single stranded RNA TLS tRNA-like structure UTR Un-translated region

aa Amino acid(s) ATC Artificial top component CP Coat protein or capsid protein EM Electron microscopy kb Kilobase kDa Kilo dalton nt Nucleotide(s)

Glossary Icosahedron A solid having 20 faces and 12 vertices. Phylogeny The complete evolutionary history of group of animals, plants, bacteria, viruses, etc. Pseudoknot RNA structure formed by base-pairing between nucleotides within the loop subtending a stem and nucleotides outside of this loop.

Sub-genomic RNA Derived from a viral RNA genome during replication, serves as mRNA for the expression of genes that are translationally silent in the genomic RNA.

The Family and Its Distinguishing Features The members of the family Tymoviridae, in the order Tymovirales, are presented in Table 1. Like the family itself, the genus Tymovirus derives its name from the type species, Turnip yellow mosaic virus. Turnip yellow mosaic virus (TYMV) was first isolated in 1946 and is by far the most intensively studied member of the family. Indeed, TYMV is one of the best-characterized plant viruses. The family was created in recognition of the close relationships between the genera Tymovirus, Marafivirus, and the founding member of newly created genus Maculavirus, Grapevine fleck virus (GFkV). Poinsettia mosaic virus (PnMV) and Bombyx mori latent virus (BmLV) are currently family members unassigned to a genus. Complete genome sequences are available for the type species of each of the three genera of the Tymoviridae and for PnMV and BmLV. Newly described tentative tymoviruses having sequence homologies and similar genome expression strategies, but not yet assigned to the family are listed at the end of Table 1. The members of the family Tymoviridae are characterized by their icosahedral, non-enveloped, B29 nm virions, that can readily be visualized by negative-staining electron microscopy (EM). Infections produce a characteristic mixture of filled, infectious virions, and empty or near-empty capsids. Regular surface features, representing prominent peaks formed by pentamers and hexamers of coat protein (CP) molecules, are evident by EM and by molecular structure and computer rendering (Fig. 1). The genomes of members of the Tymoviridae are composed of a single positive ssRNA generally 6.0–6.5 kb long, although the genome of GFkV is 7.5 kb long. Sub-genomic RNAs of 1 kb or less are associated with CP expression. All Tymoviridae genomes have a distinctive skewed nt composition that is rich in C residues (32%–50%). The unifying characteristic of the genome design of all Tymoviridae members is the presence of a long open reading frame (ORF) that covers most of the genome and encodes the replication polyprotein with identifiable domains: methyl-transferase, papain-like proteinase, helicase, and RNA-dependent RNA polymerase (RdRp) in order N- to C-terminal (Fig. 2). The CP ORF is situated downstream of the polyprotein ORF, to which it is fused in the marafivirus and PnMV genomes. Close familial relationships are easily discerned from alignments of the sequences of the RdRp and CP genes. Most viruses in the family have narrow host ranges. Many of the tymoviruses have been isolated from non-crop hosts and have thus far not presented major disease threats to crops. Marafiviruses are associated with significant crop losses, perhaps resulting from their more effective transmission by flying insects. Disease symptoms include bright yellow mosaic or mottling (tymoviruses and PnMV), chlorotic stripes, vein clearing, etched lines or dwarfing (marafiviruses), and leaf flecking (maculaviruses).

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Tymoviruses (Tymoviridae)

Table 1

Members of the family Tymoviridae

Species name

Virus acronym

Complete sequence accession no.

Genus Tymovirus Anagyris vein yellowing virus Andean potato latent virus Andean potato mild mosaic virus Belladonna mottle virus Cacao yellow mosaic virus Calopogonium yellow vein virus Chayote mosaic virus Clitoria yellow vein virus Desmodium yellow mottle virus Dulcamara mottle virus Eggplant mosaic virus Erysimum latent virus Kennedya yellow mosaic virus Melon rugose mosaic virus Nemesia ring necrosis virus Okra mosaic virus Ononis yellow mosaic virus Passion fruit yellow mosaic virus Peanut yellow mosaic virus Petunia vein banding virus Physalis mottle virus Plantago mottle virus Scrophularia mottle virus Tomato blistering mosaic virus Turnip yellow mosaic virus Voandzeia necrotic mosaic virus Wild cucumber mosaic virus

AVYV APLV APMMV BeMV CYMV CalYVV ChMV CYVV DYMoV DuMV EMV ErLV KYMV MRMV NeRNV OkMV OYMV PFYMV PeYMV PetVBV PhyMV PlMoV SrMV ToBMV TYMV VNMV WCMV

AY75180 JX508291 JX508290

Genus Marafivirus Bermuda grass etched-line virus Blackberry virus S Citrus sudden death-associated virus Grapevine Syrah virus 1 Maize rayado fino virus Nectarine virus M Oat blue dwarf virus Olive latent virus 3 Peach virus D Tentative species Grapevine asteroid mosaic-associated virus Grapevine rupestris vein feathering virus Genus Maculavirus Grapevine fleck virus Tentative member Grapevine red globe virus

AF195000

AY789137 J04374 AF098523 D00637 AY751778 EF554577 J04375 KY832429

Y16104 AY751779 AY751777 KC840043 J04373, X16378, X07441

BELV BVS CSDaV GSyV-1 MRFV NeVM OBDV OLV-3 PeVD

FJ915122 AY884005, DQ185573 FJ436028 AF265566 KT273411 U87832 FJ444852 KY084481

GAMaV GRVFV

KX354202 AY706994

GFkV

AJ309022

GRGV

KX171166

Unassigned viruses in the family Bombyx mori latent virus Poinsettia mosaic virus

BmMLV PnMV

AB186123 AJ271595

Tentative Tymoviruses Alfalfa virus F Grapevine virus Q Medicago sativa marafivirus 1 Narajilla chlorotic mosaic virus

AVF GVQ MsMV1 NarCMV

MG676465 FJ977041 MF443260 MG323924

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Fig. 1 Tymovirus particle structure. Rendering of a tymovirus particle based on experimental data deposited in the ViperDB database is shown on the left. The different colors denote the distance from the center of the virus, the brighter regions being more distant. Molecular structure courtesy of Antonio Šiber, available at: http://www.antoniosiber.org/publikacije_en.html. A negative-contrast electron micrograph of virions and ‘empty’ particles of belladonna mosaic tymovirus is shown at right, with the high-magnification inset showing the prominent surface structure. Scale ¼ 100 nm. Image courtesy Dr. D.-E. Lesemann.

Fig. 2 Genomes of the type members of the genera of the family Tymoviridae and of PnMV. All genomes have a 50 -m7GpppG cap, but the 30 terminal structures vary. TYMV has a valine-specific tRNA-like structure (cloverleaf), MRFV, GFkV and PnMV have a poly(A) tail. The known or predicted (?) expressed ORFs are indicated with the molecular weight (K, kDa) of the predicted protein. The replicases (RPs) all possess methyltransferase (Mtr), papain-like proteinase (Pro), helicase (Hel), and RdRp domains. The TYMV RP is cleaved as indicated, and similar cleavages are expected for the other viruses. The TYMV overlapping protein (OP) is the viral movement/RNAi suppressor protein. The TYMV CP is expressed from a sgRNA, and all other CPs are probably also expressed from sgRNAs (not shown). The 28 kDa MRFV CP (whose true size is likely to be closer to 25 kDa) may be produced by proteolytic cleavage from the RP–CP fusion protein, and an analogous event may occur with PnMV. Expression of the MRFV p43 and GFkV p31 and p16 has not yet been validated.

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Tymoviruses Properties and Distinguishing Characteristics Although virtually all information about tymoviruses has been derived from studies on TYMV, it is considered to be generally applicable to the other members of the genus. Virions of tymoviruses contain genomic RNAs 6.0–6.7 kb long and produce a single 30 colinear sgRNA less than 1 kb in length encoding the B20 kDa CP. Their genomes include a distinctive 16-nt-long ‘tymobox’ sequence, and the genomes of most species possess a 30 tRNA-like structure (TLS) that can be efficiently esterified with valine. The genomes encode three ORFs (Fig. 2), two of which almost completely overlap: the OP and RP ORFs, which begin at AUG codons separated by only 4 nt. Tymoviruses replicate in all major tissues of their host plants, and accumulate to high levels (more than 0.1 mg g1 leaf tissue). Both filled and empty or near-empty particles accumulate. They are readily transmitted mechanically under laboratory conditions, and are spread over limited distances by beetle vectors in nature. Infection produces mosaic symptoms and a distinctive cytopathy that is evident upon EM observation by the appearance of small vesicles on the surface of chloroplasts, together with vacuolation and clumping of chloroplasts.

Capsid Structure Virions of tymoviruses are highly stable B29 nm, T ¼ 3 icosahedra formed by 180 copies of the single CP, arranged as 12 pentamers and 20 hexamers. These groupings form the vertices with fivefold and sixfold symmetry that constitute the surface peaks visible upon high-resolution EM observation (Fig. 1). Inter-subunit stabilization is provided primarily by hydrophobic protein–protein contacts, allowing the formation of stable shells that appear to be devoid of RNA. These empty or near-empty capsids can account for about one-third of the particles present in infected tissues and are readily identifiable by internal staining in negative-contrast EM (Fig. 1). They sediment as the ‘top component’ at 45–55S in CsCl density gradients, and readily separate from the ‘bottom component’, the infectious 110–120S virions that contain the genomic RNA. Minor components of intermediate density contain a range of sub-genomic-size RNAs that can be translated to yield CP. Roughly equimolar amounts of genomic and sgRNAs are encapsidated, but the precise disposition of sgRNA in the various particles has not been determined. Like many other viral CPs, tymoviral CPs fold into eight-stranded, b-barrel, jelly-roll structures. However, unlike some other CPs, there is no positively charged domain at the N-terminus for interaction with RNA and charge neutralization during packaging. Charge neutralization is thought to be provided by polyamine molecules (mostly spermidine) associated with the RNA. The crystal structures of empty and infectious particles have been determined for TYMV, Desmodium yellow mottle virus (DYMV), and Physalis mottle virus. Consistent with the dominance of protein–protein interactions in particle stabilization, empty and infectious particles have very similar structures. Packaging signals in tymoviral RNAs have yet to be identified, though RNA recruitment may involve conserved hairpins in the 50 untranslated region (UTR) and interaction of C-rich segments of the RNA with CP at low pH. Localized areas of low pH (5  6) have been postulated to arise at the surfaces of photosynthesizing chloroplasts. Tymoviral replication occurs in characteristic vesicles that form at the chloroplast surfaces (see below), and capsid formation is most active near these vesicles, suggesting a possible coupling between replication and encapsidation during infection. Despite much effort, there has been no success in developing a cell-free packaging system. These efforts have focused on the possible role of ‘artificial top component’ (ATC) capsids as decapsidation and encapsidation intermediates. ATCs are protein shells devoid of RNA that can be made from infectious virions by treatments such as freeze–thawing and exposure to high pH and pressure. They are similar in structure to infectious virions, but lack a capsomere of six CP molecules that is ejected during the treatment, allowing RNA escape.

Genome Organization Tymoviral genomic RNAs possess a 50 m7GpppG cap and terminate at the 30 end in -CC(A) except for Dulcamara mottle virus (DuMV) that is in most cases part of a TLS. The typical tymoviral TLS is just over 80 nt long, has a distinctive pseudoknot close to the 30 terminus, and is a close structural mimic of cellular tRNAVal. A valine-specific anticodon is present, and the 30 terminus can be specifically amino-acylated with valine. The valylated RNAs of some, though not all tymoviruses, can form a tight complex in vitro with the GTP-bound form of translation elongation factor eEF1A. Most molecules of encapsidated RNA lack the terminal A residue in the -CC(A) end, which is thought to be added by the host tRNA-specific CCA-nucleotidyltransferase at the beginning of infection. The main roles of the TYMV TLS are thought to be (1) as a 30 enhancer of translation initiation, (2) as a regulator of the onset of RNA replication by modulating access by the polymerase to the 30 end, and (3) in maintaining an intact-CCA 30 end. A minority of tymoviral genomic RNAs have 30 UTRs that lack the typical valine-specific TLS (DuMV, Erysimum latent virus, and Nemesia ring necrosis virus, NeRNV). Tymoviral RNAs act as cap-dependent messenger RNAs. Of the three ORFs, two (OP and RP) are expressed directly from the genomic RNA, whereas the third (CP) is expressed from the 50 capped sgRNA (Fig. 2). Although the sequence contexts around the OP and RP initiation codons vary, 4 nt always separates these closely spaced AUGs. The RP ORF covers most of the viral genome, and encodes the B200 kDa RP, the only viral protein essential for supporting viral RNA replication in protoplasts. This ORF encodes discrete domains (Fig. 2) that are discernable by sequence relationships to similar domains encoded by a wide

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variety of positive-stranded RNA viral genomes. Particularly the RdRp domain is well conserved and appropriate for phylogenetic comparisons. Closely related, yet distinct, tymoviral genomes have RdRp domains with about 70% nt sequence identity. The RP is translated as a precursor protein that subsequently undergoes maturation cleavage catalyzed by the papain-like proteinase domain. The single known cleavage separates the RdRp domain from the remainder of the RP; thus, for TYMV, the precursor p206 is processed to yield p141 and p66. The OP ORF almost entirely overlaps the RP ORF. Its length and the sequence of the encoded protein are highly variable. OPs range between 49 and 82 kDa, and have only 25%–40% aa sequence identity between the most closely related pairs of viruses. OP expression is needed for establishment of infection and for spread of the virus in plants. The OP is a suppressor of the host RNA silencing antiviral response and is also believed to be the viral movement protein. It is a substrate for ubiquitin-dependent degradation by the proteasome. Long-distance movement of the virus in plants requires expression of the CP. The CP ORF initiates close to the end of the RP ORF, sometimes even a little upstream. CP sequences are variably conserved among tymoviruses, with 36%–86% aa sequence identity between the most closely related pairs of viruses. Tymoviral genomes possess a highly conserved sequence, the 16 nt tymobox (–GAGUCUGAAUUGCUUC) with small variations in some viruses, especially wild Cucumber mosaic virus, just upstream of the CP ORF. The tymobox and its associated ‘initiation box’ sequence CAA(U/G) positioned 8 or 9 nt downstream of the tymobox are believed to serve as the core elements of the sgRNA promoter. The tymobox overlaps with the 30 end of the RP ORF, resulting in the presence of the tripeptide – ELL – near the C terminus of all RPs.

Replication Cycle The ultrastructural changes that reflect viral replication activity and that result in the distinctive pathology of the chloroplasts have been particularly well described for TYMV. As is typical of positive-stranded RNA viruses, the replication cycle is completed entirely in the cytoplasm, and RNA replication occurs in association with membranes, specifically the chloroplast outer membrane. Vesicles 50–80 nm in diameter form as invaginations of the two outer membranes of chloroplasts. They form before the appearance of virions, which are initially found close to vesicle clusters and later throughout the cytoplasm; in some cases, empty capsids accumulate in nuclei. The RdRp has been localized to zones on chloroplasts that are rich in vesicles, and this localization depends on protein–protein interaction between the proteinase and RdRp domains of the mature RPs p141 and p66, respectively. RNA replication occurs via the production of full-length minus strands, whose synthesis in TYMV is directed by the 30 terminal –CCA serving as promoter and initiation site. No other minus-strand promoter elements have been identified in the 30 UTR. As mentioned above, the tymobox appears to function with the initiation box as the core of the promoter directing sgRNA synthesis by internal initiation on the minus strand. The sgRNA of TYMV has a m7GpppA 50 terminus. Studies on the functions of viral proteins and of cis-acting sequences in the genomic RNA have been greatly facilitated by the use of ‘infectious clones’, that is, molecularly cloned cDNA versions of the genome from which infectious RNA can be derived.

Infection and Transmission Tymoviruses cause chlorotic mosaic, vein-clearing, and mottling symptoms, generally without strong stunting. Host ranges are mostly narrow, and to date, no tymoviruses infecting monocot plants have been isolated. Tymoviruses are transmitted over limited distances by chrysomelid beetles, and some are weakly seed-transmissible. Tymoviruses and PnMV are readily transmitted mechanically and accumulate to high titers in infected plants.

Phylogenetic Relationships and Species Demarcation In the past, serology using antisera raised against intact virus was the main criterion in classifying tymoviruses and in distinguishing between species. This is no longer the most convenient approach, however, and it has in fact been shown that distinct viruses whose genomic and CP sequences have identities less than 80% and 90%, respectively, can appear to be serologically identical. This seems to be due to similar or identical dominant epitopes within otherwise distinct CPs. The study of relationships among tymoviruses and to other viruses currently primarily relies on the interpretation of sequences derived from the genomic RNA. The phylogenetic trees based on the sequences of the CP and the RP (Fig. 3) show similar relationships, both among the tymoviruses and to the other members of the family Tymoviridae. Genomes of the Tymoviridae are not under positive selection and recombination might be the major driving force of diversity within the family. For instance, the genome of NeRNV, which has a tobamoviral-type TLS capable of amino-acylation with histidine, indicates that recombination can occur and shape tymoviral genome evolution. Also, family-wide recombination analysis showed that Physalis mottle virus can be a donor for other marafivirus members. The complete genome sequences are available for 35 tymoviruses or tentative tymoviruses (Table 1). Alignments based on the replicase protein sequences indicate that the capilloviruses, carlaviruses, trichoviruses, and potexviruses, all members of the Flexiviridae, are the next most closely related virus groups. More distant sequence relationships indicate that members of the family Tymoviridae belong to the alpha-like virus group in the Tymovirales, the newly established order of viruses reported by the International Committee on Taxonomy of Viruses.

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Fig. 3 Maximum-likelihood phylogenetic trees showing the relationships between members of the family Tymoviridae based on amino acid sequence alignments of (a) RdRp and (b) coat protein (CP) sequences. Bootstrap support values (%) above 70% are indicated at supported nodes. Branch lengths reflect the difference between amino acid sequences. Viruses from each of the three genera are separately shaded as marked. Viruses with tentative taxonomic status are marked with an asterisk.

Marafiviruses Properties and Distinguishing Characteristics Three current members of the genus Marafivirus are viruses principally infect monocots (grasses), while others infect citrus, tree fruits, grapevine, and blackberry. Another tentative two marafiviruses were isolated from grapevine. Plant host ranges are narrow. The genomes are 6.3–6.8 kb long, with a single ORF encoding an RP/CP fusion protein covering most of the genome. Two forms of the same CP (B25 and B21 kDa) are found in virions. A single 30 -colinear sgRNA (o1 kb) encoding the smaller form of the CP is produced during infection. A slightly modified form of the tymobox sequence, termed ‘marafibox’, is present in the genome near the junction between the RdRp- and capsid-coding sequences. Marafiviruses are phloem-limited and not mechanically transmissible. The grass-infecting marafiviruses are transmitted by cicadellid leafhoppers in a persistent-propagative manner, involving virus replication in the insect. A distinctive feature of the marafiviruses is the presence of two CPs in the virions, a major CP B21 kDa form and a minor N-terminally extended 22–28 kDa form. The latter is thought to arise by proteolytic release from the polyprotein, whereas the former is expressed from the sgRNA. The production of this sgRNA is probably under the control of the marafibox, a 16 nt sequence (–CA(A/G)GGUGAAUUGCUUC–) very closely related to the tymobox, and the –CA(A/U)-initiation box located 8 nt downstream. The roles of the two forms of CP are unclear.

Virion Structure, Genome Organization, Replication Cycle, and Phylogenetic Relationships Marafiviruses produce virions and empty capsids that are similar to those produced by tymoviruses when observed by negative-staining EM. However, due to the phloem-limited replication of marafiviruses, the yield of virus is low, limiting the usefulness of EM for viral identification. Complete genome sequences are available for eight confirmed marafiviruses (Table 1). Marafivirus genomic RNAs appear to be 50 capped but they lack the 30 TLS that is distinctive of tymoviruses, and marafivirus genomes possess a 30 poly(A) tail. Almost the entire genome of marafiviruses is devoted to encoding a large 224–240 kDa polyprotein, which includes replication-associated and CP domains (Fig. 2). The relationships between the CP and RP genes of the marafiviruses and other members of the family Tymoviridae are indicated in Fig. 3. The polyprotein is believed to be proteolytically processed by the viral proteinase at two sites, between the helicase and RdRp domains as in tymoviruses, and between the RdRp and CP domains. Polyprotein cleavage sites were experimentally mapped in TYMV and OBDV, and aa sequence alignments have

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discerned probable cleavage sites occurring immediately downstream of Gly-Gly or Gly-Ala dipeptides at the helicase–RdRp and RdRp–CP junctions in marafivirus polyproteins. As the number of complete marafivirus genome sequences increases, the taxonomic implications of evident variation in genome organization may be clarified. MRFV possesses a predicted significant overlapping ORF, encoding a putative 43 kDa protein, while Grapevine syrah virus 1 has a predicted 26 kDa protein that shares 43% identify with the central portion of the 43 kDa of MRFV; expression in vivo has not been verified for either protein. Related but strongly interrupted coding sequences have been identified overlapping the methyl-transferase domains of other marafiviral genomes and, based on sequence similarities to tymoviral OP, it has been postulated that these sequences represent variously degenerate versions of the tymoviral OP. The loss of OP during evolution may explain the phloem limitation of marafiviruses. Citrus sudden death-associated virus (CSDaV) is the only marafivirus with an additional ORF near the 30 end of the genome. This ORF almost completely overlaps the CP ORF and could encode a 16 kDa protein. The three grass-infecting marafiviruses, MRFV, OBDV, and Bermuda grass etched-line marafivirus, are serologically related, but are not serologically cross-reactive with tymoviruses or more distantly related viruses. Despite the indications of genome variability mentioned above, the marafiviruses do form a phylogenetic group with interspecies relationships similarly close to those between the various tymoviruses (Fig. 3). Sequence relationships among the RP and CP ORFs clearly link the marafiviruses to the other members of the Tymoviridae. Like the tymoviral CPs, marafivirus CPs lack a cluster of basic aa at the N terminus.

Infection and Transmission Marafi- and maculaviruses are phloem limited. The best-studied marafiviruses are MRFV and OBDV. They cause stunting and chlorotic leaf spots or streaks, general leaf discoloration, or enations on leaf and stem veins. Cytological symptoms are restricted to the phloem and adjacent parenchyma cells, with clearly discernible hyperplasia and hypertrophy. These viruses are not mechanically transmissible or transmissible through seed, but are vectored in nature by leafhoppers. Significantly, MRFV and OBDV are maintained for long periods and through molts in leafhopper vectors, and transmission can occur from viruliferous vectors over a period of several weeks. OBDV replicates in the aster leafhopper (Macrosteles fascifrons), and the transmission behavior of MRFV is also consistent with replication in the cicadellid leafhopper (Dalbulus maidis). MRFV is commonly associated and co-transmitted with maize stunt spiroplasma or maize bushy stunt phytoplasma in causing economically significant outbreaks of corn stunt syndrome. OBDV can also be co-transmitted with a phytoplasma. The presence of virus in the leafhopper vectors is not associated with detectable symptoms or loss of reproductive fitness. The most economically significant marafivirus is probably CSDaV. This virus is believed to be the causative agent of the citrus sudden death syndrome, which has had devastating effects on orange trees in Brazil. Preliminary evidence suggests transmission by aphids. The grapevine-infecting tentative marafiviruses, Grapevine asteroid mosaic-associated virus and Grapevine rupestris veinfeathering virus, have no known insect vector.

Maculaviruses Properties and Distinguishing Characteristics Recent sequencing of the complete genome of GFkV supported the creation of the new genus Maculavirus. The distinguishing characteristics were the length of the GFkV genome, the absence of a tymobox/marafibox, the lack of a fused RP/CP ORF as present in marafivirus genomes, and the lack of phylogenetic clustering with either tymoviruses or marafiviruses. Grapevine red globe virus (GRGV) is a tentative member of the genus. Both viruses infect only Vitis species (grapevine and relatives). In common with marafiviruses, these maculaviruses are restricted to the phloem and are not sap transmissible. No insect vectors have been identified. As for all other members of the family Tymoviridae, maculaviruses produce B29 nm particles with prominent surface features observable by EM. Top component and infectious bottom component particles are produced. GFkV infections are cytologically distinct from marafivirus infections through the presence of severely modified mitochondria (‘multivesiculate bodies’). The genome of GFkV (7564 nt) is the longest among the members of the family Tymoviridae, and has the highest cytosine content (49.8%). The arrangement of RP and CP ORFs is like that of tymoviruses (Fig. 2). The RP ORF encodes a 215 kDa protein with replicationassociated domains that present sequence relatedness to the tymoviral and marafiviral RPs. The aa sequence of the RdRp domain shows the closest relationship to sister genera (59% and 67% identities with the RdRps of a tymovirus and marafivirus, respectively). The 24.5 kDa GFkV CP has 23%–31% sequence identity with CPs of tymoviruses and marafiviruses, and like other CPs of members of the family Tymoviridae it lacks a cluster of basic aa at the N-terminus. The GFkV genome encodes two additional proteins from ORFs that partially overlap the CP ORF in the 30 region of the genome. No functions for these putative proteins are predicted from sequence comparisons. Close relationship between GFkV and GRGV is rather weakly supported by sequence comparisons (Fig. 3). The GFkV genome is probably capped at the 50 end and has a poly(A) tail at the 30 end, in common with most marafiviruses and PnMV. It is the only known member of the family Tymoviridae to lack a tymobox- or marafibox-like sequence. Nevertheless, two or more sgRNAs, B1.3 and B1.0 kb in length, are present in infected tissues. These RNAs appear to be packaged in both top- and bottom-component particles, in the latter case apparently together with the genomic RNA.

Tymoviruses (Tymoviridae)

825

Poinsettia Mosaic Virus Properties and Distinguishing Characteristics PnMV possesses properties characteristic of both the tymoviruses and marafiviruses. The B28 nm virus particles have the EM structure typical of the Tymoviridae and separate into typical top components containing the sgRNA and infectious bottom components. Like tymoviruses, PnMV is produced in high yield during infection, is not phloem-limited, and can be mechanically transmitted in the laboratory. Replication is associated with chloroplast cytopathy, though not with the appearance of the replication-associated vesicles typical of tymovirus infection. The properties and coding arrangement of the 6.1 kb RNA are more closely aligned with those of the marafiviruses (Fig. 2). The genomic RNA is polyadenylated, contains a marafibox-related putative sgRNA promoter, and possesses a single long ORF that encodes an RP/CP fusion protein (p221). No OP ORF is present. The 21–24 kDa PnMV CP is presumably expressed from the 0.65 kbp sgRNA; the involvement of potential cleavage of the RP/CP fusion protein to produce a second CP variant is not known. Although PnMV sequences are more closely related to the marafiviruses than tymoviruses, the relationships appear to be insufficiently close to warrant inclusion in the genus Marafivirus (Fig. 3). In commercial poinsettia cultivation, PnMV is transmitted by vegetative propagation; the virus is not transmissible via seed or pollen. The natural host range is restricted to Euphorbia sp., especially poinsettia, E. pulcherrima. Symptoms appear seasonally, varying from unapparent to light mottling. PnMV is often associated with poinsettia cryptic virus (family Partitiviridae) and with a phytoplasma that is responsible for the desirable free-branching phenotype.

Bombyx Mori Latent Virus Properties and Distinguishing Characteristics Bombyx mori latent virus (BmLV) was initially identified in a cultured cell line (BmN) of the silk moth (Bombyx mori) by cDNA microarray. The full-length sequence is 6153 nt excluding the poly(A) tail. Putative isometric virions of B28–30 nm were observed and were found to be infections to insect Sf-9 cells and tissues of B. mori larvae. The genome encodes a protein of 1747 aa residues containing the conserved MTR, PRO, HEL, and POL regions of tymovirus replication polyprotein. It shares 44% aa identity to GFkV. The single CP is encoded in a second ORF and is express from a 1.25 kb sub-genomic RNA. A third putative ORF is located at the 3 0 terminus of the genome and encodes a putative 15.5 kDa protein. The genome lacks the ‘tymobox’ or ‘marafibox’ sub-genomic promoter. Phylogenetic analysis of the replicase and CP sequences suggests that BmLV is a putative member of the family Tymoviridae, distinct from tymo- and marafiviruses, but closely related to GFkV. In the absence of additional data, it has been designated as an unassigned species in the family.

Further Reading Bradel, B.G., W. Preil, W., Jeske, H., 2000. Sequence analysis and genome organization of Poinsettia mosaic virus (PnMV) reveal closer relationship to marafiviruses than to tymoviruses. Virology 271, 289–297. Canady, M.A., Larson, S.B., Day, J., McPherson, A., 1996. Crystal structure of Turnip yellow mosaic virus. Nature Structural & Molecular Biology 3, 771–781. Ding, S.W., Howe, J., Keese, P., et al., 1990. The tymobox, a sequence shared by most tymoviruses: Its use in molecular studies of tymoviruses. Nucleic Acids Research 18, 1181–1187. Dreher, T.W., 2004. Turnip yellow mosaic virus: Transfer RNA mimicry, chloroplasts and a C-rich genome. Molecular Plant Pathology 5, 367–375. Edwards, M.C., Zhang, Z., Weiland, J.J., 1997. Oat blue dwarf marafivirus resembles the tymoviruses in sequence, genome organization, and expression strategy. Virology 232, 217–229. Hirth, L., Givord, L., 1988. Tymoviruses. In: Koenig, R. (Ed.), The Plant Viruses. New York: Plenum, pp. 163–212. Jakubiec, A., Notaise, J., Tournier, V., et al., 2004. Assembly of Turnip yellow mosaic virus replication complexes: Interaction between the proteinase and polymerase domains of the replication proteins. Journal of Virology 78, 7945–7957. Kadaré, G., Rozanov, M., Haenni, A.-L., 1995. Expression of the Turnip yellow mosaic virus proteinase in Escherichia coli and determination of the cleavage site within the 206 kDa protein. Journal of General Virology 76, 2853–2857. Katsuma, S., Tanaka, S., Omuro, N., et al., 2005. Novel macula-like virus identified in Bombyx mori cultured cells. Journal of Virology 79, 5577–5584. Koenig, R., Pleij, C.W., Lesemann, D.E., Loss, S., Vetten, H.J., 2005. Molecular characterization of isolates of Anagyris vein yellowing virus, Plantago mottle virus and Scrophularia mottle virus: Comparison of various approaches for tymovirus classification. Archives of Virology 150, 2325–2338. Maccheroni, W., Alegria, M.C., Greggio, C.C., et al., 2005. Identification and genomic characterization of a new virus (Tymoviridae family) associated with citrus sudden death disease. Journal of Virology 79, 3028–3037. Martelli, G.P., Sabanadzovic, S., Abou-Ghanem Sabanadzovic, N., Edwards, M.C., Dreher, T., 2002. The family Tymoviridae. Archives of Virology 147, 1837–1846. Mayo, M.A., Dreher, T.W., Haenni, A.-L., 2000. Genus tymovirus. In: van Regenmortal, M.H.V., Fauquet, C.M., Bishop, D.H.L., et al. (Eds.), Seventh Report of the International Committee on Taxonomy of Viruses. New York: Academic Press. Morch, M.D., Boyer, J.C., Haenni, A.-L., 1988. Overlapping open reading frames revealed by complete nucleotide sequencing of Turnip yellow mosaic virus genomic RNA. Nucleic Acids Research 16, 6157–6173.

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Sabanadzovic, S., Ghanem-Sabanadzovic, N.A., Saldarelli, P., Martelli, G.P., 2001. Complete nucleotide sequence and genome organization of Grapevine fleck virus. Journal of General Virology 82, 2009–2015. Weiland, J.J., Dreher, T.W., 1989. Infectious TYMV RNA from cloned cDNA: Effects in vitro and in vivo of point substitutions in the initiation codons of two extensively overlapping ORFs. Nucleic Acids Research 17, 4675–4687.

Relevant Websites https://viralzone.expasy.org/54 Chordopoxvirinae ViralZone. https://talk.ictvonline.org/ictv-reports/ictv_9th_report/positive-sense-rna-viruses-2011/w/posrna_viruses/245/tymoviridae Tymoviridae Positive Sense RNA Viruses. https://link.springer.com/protocol/10.1385/0-89603-385-6:219 Tymovirus Isolation and Genomic RNA Extraction.

Umbraviruses (Tombusviridae) Eugene V Ryabov, USDA, Agricultural Research Service, Beltsville, MD, United States Michael E Taliansky, The James Hutton Institute, Dundee, United Kingdom r 2021 Elsevier Ltd. All rights reserved. This is an update of M. Taliansky, E. Ryabov, Umbravirus, In Encyclopedia of Virology (Third Edition), edited by Brian W.J. Mahy and Marc H.V. Van Regenmortel, Elsevier Ltd., 2008, doi:10.1016/B978-012374410-4.00524-0.

Glossary Fibrillarin An abundant nucleolar protein that participates in the processing and modification of rRNAs and a methyltransferase. Nucleolus A prominent subnuclear domain and the site of transcription of ribosomal RNA (rRNA), processing of the pre-rRNAs and biogenesis of pre-ribosomal particles, and

also participates in other aspects of RNA processing and cell function. Phloem Plant vascular system used for rapid long-distance transport of assimilates and macromolecules. Plasmodesmata Plant unique intercellular membranous channels that span plant cell walls linking the cytoplasm of contiguous cells.

Introduction Umbraviruses differ from other plant viruses in that they do not encode a coat protein (CP), and thus no conventional virus particles are formed in the infected plants. The name of the genus Umbravirus is derived from the Latin umbra, which means a shadow, or an uninvited guest who comes with an invited one. This name reflects the way in which umbraviruses depend for survival in nature on an assistor virus, which is always a member of the Enamovirus or Polerovirus genera of the family Luteoviridae. For transmission between plants, the CP of the assistor virus forms aphid-transmissible hybrid virus particles encapsidating umbraviral RNA. In nature, each umbravirus is associated with one particular member of the family Luteoviridae, although in experiments trans-capsidation is not needed for umbravirus infection accumulation within infected plants because functions such as protection and movement of the virus RNA, both cell-to-cell and long distance, do not require the presence of the assistor virus or its CP. Moreover, under experimental conditions, mechanical transmission of umbraviruses can take place without the aid of an assistor virus. This implies that umbraviruses encode some product(s) that functionally compensate for the lack of a CP (see below).

Classification Currently the genus Umbravirus includes nine distinct virus species which are accepted by ICTV taxonomy: Carrot mottle virus (CMoV) – the type species, Carrot mottle mimic virus (CMoMV), Groundnut rosette virus (GRV), Ethiopian tobacco bushy top virus (ETBTV), Lettuce speckles mottle virus (LSMV), Opium poppy mosaic virus (OPMV), Pea enation mosaic virus-2 (PEMV-2), Tobacco mottle virus (TMoV), and Tobacco bushy top virus (TBTV). These viruses together with the tentative members of the genus Umbravirus are listed in Table 1. Some of these viruses have been known since the early days of plant virology. The first to be described was TMoV, reported from Zimbabwe and Malawi in 1945. The most economically important umbravirus is GRV, which is endemic throughout sub-Saharan Africa. Sporadic, unpredictable outbreaks of groundnut rosette disease (actually caused by a satellite RNA of GRV) cause severe crop losses. Pea enation mosaic disease outbreaks are also sporadic and localized, but losses of nearly 90% in peas and up to 50% in field beans have been reported. One or both of the carrot-infecting umbraviruses probably occurs worldwide wherever carrots are grown, but they are uncommon in commercial crops because the insecticides used to control carrot fly also control the aphid vectors of CMoV and CMoMV. LSMV is reported only from California, USA.

Virion Structure Although in plants infected with umbraviruses unaccompanied by the assistor virus no virions are formed, the infectivity of CMoV and GRV in buffer extracts of infected leaves is surprisingly stable: survives for several hours at room temperature and is resistant to treatment with ribonuclease. It is however, sensitive to a treatment with organic solvents, suggesting that lipid-containing structures are involved in protection of umbravirus RNA. Indeed, in plants infected with a number of umbraviruses, including CMoV, enveloped structures B50 nm in diameter were observed. An infective fraction from GRV-infected tissue contained complexes with a buoyant density of 1.34–1.45 g/cm3 consisting of filamentous ribonucleoprotein particles, composed of the umbraviral ORF3 protein (see below) and virus RNA, embedded in a matrix (Fig. 1). The relationship between these two kinds of structure is unclear.

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Table 1

Umbraviruses (Tombusviridae) Virus members of the genus Umbravirus

Virus (isolate)

Abbreviation

Accession no.

Carrot mottle mimic virus Carrot mottle mimic virus-Australia

(CMoMV-AU)

U57305

Carrot mottle virus Carrot mottle virus-Weddel

(CmoV-We)

FJ188473

Ethiopian tobacco bushy top virus Ethiopian tobacco bushy top virus 18–2 Satellite RNA

(ETBTV18–2)

KJ918748 KJ918747

Groundnut rosette virus Groundnut rosette virus-MC1 Satellite RNAs

(GRV-MS1)

Z69910 Z29702-Z29711

Lettuce speckles mottle virus Lettuce speckles mottle virus-California Opium poppy mosaic virus Opium poppy mosaic virus-PHEL5235 Pea entaion mosaic virus-2 Pea entaion mosaic virus-2-WGS Bean yellow vein-banding virus Satellite RNA

Assistor virus Carrot red leaf virus Carrot red leaf virus Potato leafroll virus

Groundnut rosette assistor virus

Beet western yellows virus (LSMV-CA) Opium poppy mosaic-associated virus (OPMV-PHEL5235)

MG182693

(PEMV-2-WGS) (BYVBV)

U03563

Pea enation mosaic virus-1

U03564

Tobacco bushy top virus Tobacco bushy top virus-Baoshan

(TBTV-Bao)

AF402620

Tobacco vein distorting virus

Tobacco mottle virus Tobacco mottle virus-Zimbabwe

(TMoV-ZW)

AY007231

Tentative members in the genus Sunflower crinkle virus Sunflower yellow blotch virus Tobacco yellow vein virus

(SuCV) (SuYBV) (TYVV)

Tobacco vein distorting virus

Fig. 1 Electron microphotograph of a section of a Nicotiana benthamiana cell expressing GRV ORF3 protein. The section shows a complex of filamentous RNP particles embedded in an electron dense matrix. The section was labeled by in situ hybridization with an RNA probe specific for viral RNA. Bar, 100 mm.

Genome Organization and Expression The genome of umbraviruses consists of single linear segment of positive-sense single stranded RNA (ssRNA) (Fig. 2). The complete genomic RNA sequences of umbraviruses are of 4019–4253 nt in length. There is no polyadenylation at their 30 termini and there is no information about modifications of their 50 termini. At the 50 end, a very short non-coding sequence precedes ORF1, which encodes a putative 31–37 kDa protein. The ORF2, which overlaps the 30 end of ORF1, potentially encodes a

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Fig. 2 Organization and expression strategy of GRV genome. Translation of the ORF1 and the ORF2 with a frameshift event results in production of the ORF1-ORF2 frameshift protein; subgenomic RNA (sgRNA) synthesis is required for expression of the ORF3 and ORF4. Solid lines represent RNA molecules, boxes represent open reading frames (ORFs), corresponding translation products are shown below. RdRp, RNA-dependent RNA polymerase; FS,  1 frameshift signal.

63–65 kDa protein, but lacks an AUG initiation codon. It is likely that ORF1 and ORF2 are translated as a single 94–98 kDa protein by a –1 translational frameshifting mechanism; the sequence associated with frameshifting events in genomes of several animal and plant RNA viruses is found in the region at the 30 end of the ORF1. The predicted amino acid sequence of the ORF2-endoded product includes an RNA-dependent RNA polymerase (RdRp) domain typical to that of other members of the family Tombusviridae, containing all eight conserved motifs of RdRps of positive-strand RNA viruses. A short untranslated stretch separated ORF2 from ORF3 typical to that of and ORF4 which are almost completely overlap in different reading frames and each encode a 26–29 kDa products. The ORF4 contains motifs characteristic of the cell-to-cell movement proteins (MPs) of plant viruses, in particular with those of cucumoviruses. The ORF3 products possess up to 50% homology to each other but shows no significant similarity to any other viral or non-viral product. The ORF3 and OFR4 are translated from subgenomic RNA, the 30 terminal RN A of the appropriate size was detected in the GRV infected plant tissue. It should be pointed out that the sequenced genomes of umbraviruses do not encode a potential CP(s). The essential role of the ORF3-ORF4 block in movement of umbraviruses within plants has been experimentally demonstrated and is discussed below. The 30 untranslated region of PEMV2 contains three 30 proximal, cap-independent translation enhancers (30 CITEs), which are required for proper virus translation. Efficient translation of the ORF3 and ORF4 proteins from the sgRNA requires all three (30 CITEs), whereas translation of the genomic requires only two 30 CITEs.

Similarity With Other Taxa The only gene conserved across all RNA viruses is the RdRp which is cenral to virus replication and existence, therefore RdRp was used to investigate global phylogenetic relationships between viral taxonomic groups. Based on the RdRp phylogeny, umbraviruses were included in the family Tombusviridae, alongside with the members of the genera Alphacarmovirus, Alphanecrovirus, Aureusvirus, Avenavirus, Betacarmovirus, Dianthovirus, Gammacarmovirus, Macanavirus, Machlomovirus, Panicovirus, Pelarspovirus, Tombusvirus, and Zeavirus, all of which infect plant hosts. A recent analysis of the RdRp sequences of all known RNA viruses showed that the family Tombusviridae, together with the genus Luteovirus and the family Carmotetraviridae, with which it has highest RdRp homology, belongs so-called “Branch 3” group of global RdRp and reverse transcriptase phylogeny, which also includes subset of positive-sence RNA viruses of viruses, such as “alphavirus supergroup”, “flavivirus supergroup” and nodaviruses. Members of the Branch 3 infect hosts across all eukaryotic kingdoms.

Replication Replication of umbravirus RNA presumably involves the ORF2-encoded RdRp. Leaves of plants infected with umbraviruses contain abundant dsRNA including a major species of about 4.4–4.8 kbp corresponding in size to that expected for double stranded form of the viral genomic RNA which may be umbravirus RNA replication intermediate. No other details of the replication mechanism have been elucidated.

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Satellite RNA Satellite RNAs are associated with some umbraviruses. In the case of GRV, satellite RNA is found in all naturally occurring isolates, and is primarily responsible for the symptoms of groundnut rosette disease. GRV satellite RNA is a ssRNA of about 900 nt which relies on GRV for its replication and, more unusually, is also required for the Groundnut rosette assistor virus (GRAV)-dependent aphid transmission of GRV. Thus, unlike most virus satellite RNAs, it is essential for the biological survival (though not the replication) of its helper virus. The role of the satellite RNA in the transmission process is to mediate transcapsidation of GRV RNA by GRAV protein to form stable aphid-transmissible hybrid virus particles. Although different GRV satellite RNA variants contain up to five potential ORFs, none of the ORFs is essential for any of the functions and biological properties that have been ascribed to GRV satellites. In contrast, the satellite RNA that is associated with some isolates of PEMV-2 is not required for transcapsidation of PEMV-2 RNA by the CP of its assistor virus PEMV-1 or for aphid transmission of the hybrid particles, and other umbraviruses, such as CMoV, do not have satellite RNAs, yet are transcapsidated by their assistors and thus transmitted by aphids. The reasons for these differences have not been explained.

Cell-to-Cell Movement Function The highly conserved ORF4-encoded proteins of umbraviruses exhibit significant sequence similarity with cell-to-cell movement proteins (MPs) of other plant viruses, in particular the 3a proteins of cucumoviruses. Therefore it has been suggested that this protein involved in cell-to-cell trafficking of umbravirus RNA through plasmodesmata. This suggestion has been confirmed by a number of genetic, cytological and biochemical approaches. By using the gene replacement strategy, it was demonstrated that the GRV ORF4 protein could functionally replace the MPs of unrelated viruses, Potato virus X (PVX) (all the products encoded by the triple gene block and the CP) and Cucumber mosaic virus (CMV) (the 3a MP and the CP). Localization of the GRV ORF4 protein has much in common with localization of MPs of other plant virus groups. The green fluorescent protein (GFP)-tagged GRV ORF4 protein targeted to plasmodesmata (Fig. 3(A)). Also, this protein formed extending tubular structures on the surface of protoplasts infected either with GRV or with the heterologous virus expressing the GFP-tagged GRV ORF4. The GRV ORF4 protein binds to RNA in vitro in non-cooperative manner which may make the viral RNA of the ribonucleoprotein (RNP) complex accessible for translation and replication.

Fig. 3 Confocal laser scanning images showing the localization of (A) GRV ORF4 protein fused to GFP to plasmodesmata (shown by arrows) (bar, 25 mm) and (B) GRV ORF3 protein fused to GFP to the nucleolus (No) and cytoplasmic inclusions containing ORF3 protein RNP particles (cRNP) (bar, 10 mm).

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Phloem-Dependent Long-Distance Movement Function One of the striking features of umbraviruses is their ability to move long distance within a plant without having a conventional CP. Involvement of CPs in long distance movement of viral infection was shown for a number of plant viruses, including Tobacco mosaic virus (TMV) which utilizes the CP exclusively for its long-distance but not cell-to-cell movement function. By using the gene replacement strategy it was demonstrated that the ORF3 proteins of GRV, PEMV-2 and ToMV were able to functionally substitute the TMV CP in the long-distance movement process. The hybrid TMV mutants expressing the umbraviral ORF3 proteins were able to move rapidly within through the phloem. It was also shown that specific mutations in the ORF3 protein of PEMV-2 abolish ability of this virus to move long distance without affecting its cell-to cell spread within inoculated leaves. It should be noted that mechanisms of the long-distance movement facilitated by umbraviral ORF3 proteins is different from those mediated by suppressors of RNA silencing such as the 2b proteins of cucumber mosaic virus and the HC-Pro protein of potyviruses. Rather than supporting virus infection by suppressing the host RNA-mediated response, the ORF3 protein seems to protect RNA by binding to viral RNA. It was shown that the ORF3 proteins of GRV, PEMV-2 and ToMV increase stability of viral RNA and protect it from RNases. Immunogold labeling and in situ hybridization experiments showed that the GRV ORF3 protein accumulated in cytoplasmic granules consisted of filamentous RNP particles composed of the ORF3 protein and viral RNA (Fig. 1). It has been suggested that these particles may be a form in which viral RNA moves long-distance through the phloem. Also the ORF3-containing RNP complex may protect viral RNA from nucleases and RNA silencing machinery.

Involvement of Nucleolus in Umbravirus Systemic Infection In addition to cytoplasmic granules, the GRV ORF3 protein was also found in nuclei, preferably targeting nucleoli (Fig. 3(B)). Sequence analysis of the ORF3 proteins of unbraviruses revealed the presence of both nuclear localization signal (NLS) and nuclear export signal (NES), functional role of which in nuclear import and export of the GRV ORF3 has been confirmed experimentally by the genetic analysis. Functional analysis of ORF3 protein mutants revealed a correlation between the ORF3 protein nucleolar localization and its ability to form the RNP particles and transport viral RNA long distances. It was also shown that the ORF3 protein directly interacts with a nucleolar protein, fibrillarin, re-distributing it from the nucleolus to cytoplasm and such an interaction is absolutely essential for umbravirus long-distance movement through the phloem. The umbraviral ORF3 protein utilizes trafficking pathways involving Cajal bodies to enter the nucleolus and, along with fibrillarin, exit the nucleus to form viral ‘transport‐competent’ RNP particles in the cytoplasm.

Interaction With Assistor Virus Although umbraviruses accumulate and spread very efficiently within infected plants, they depend on assistance of luteoviruses for their survival in nature as they require encapsidation by luteovirus CP for horizontal transmission by aphids. In its turn, umbraviruses facilitate movement of phloem limited luteoviruses to and from the phloem, as well as cell-to-cell movement of luteoviruses between mesophyll and epidermal cells. It was shown that such an ability to promote cell-to-cell movement of luteoviruses is an unique feature of the umbraviral ORF4 MP. Moreover, the ORF4 MP of GRV can facilitate cell-to-cell movement of Potato leafroll luteovirus even when expressed from hetorologous PVX or CMV genomes. In most instances, the luteovirus partner does not depend on the umbravirus infection for survival in nature. A complex consisting of PEMV-1 (the genus Enamovirus, family Luteoviridae) and PEMV-2 is a notable exception. Unlike other members of the family Luteoviridae, PEMV-1 on its own lacks the ability to move, even trough the phloem; both long-distance and cell-to-cell movement functions of PEMV-1 are provided by the umbraviral component of the complex, PEMV-2. Such a strong mutual dependence and adaptation between umbraviruses and luteoviruses suggest a long co-evolution, which had resulted in establishing a range of complexes from facultative co-existence to complete dependence of PEMV-1.

Host Range Individual umbraviruses are confined in nature to one or two host plant species. For example, groundnut is the only known natural host of GRV and entire rosette disease complex (GRV, its satellite RNA and GRAV). Experimental host ranges of umbraviruses are broader but still restricted. The symptoms induced in infected plants are usually mottles and mosaics.

Transmission In nature, umbraviruses are transmitted by aphids, but only from plants that also infected with an assistor member of the family Luteoviridae. The mechanism of the dependent transmission is the encapsidation of the umbraviral RNA by the CP provided by the assistor virus. Hence, the transmission of the dependent umbravirus occurs in the same persistent (circulative, nonpropagative) manner as that of the luteovirus-assistor.

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Umbraviruses (Tombusviridae)

Prevention and Control For the avoidance of pea enation mosaic disease, it is recommended that pea or faba bean crops should be sited away from alfalfa and clover fields. Early sowing and close spacing can reduce groundnut rosette disease incidence, probably by inhibiting the landing response of the vector. This approach is also effective against tobacco rosette disease. However, the best control approach is to use resistant cultivars if they are available. Resistance to GRV controlled by two independent recessive genes has been found and groundnut lines possessing this resistance have been developed. A possibility for the future is the deployment of transgenic forms of resistance. Strategies for engineering transgenic resistance against umbraviruses include the transformation of plants with translatable or non-translatable sequences from the umbraviruses themselves or their satellite RNAs.

Further Reading Demler, S.A., Borkhsenious, O.N., Rucker, D.G., de Zoeten, G.A., 1994. Assessment of the autonomy of replicative and structural functions encoded by the luteo-phase of pea enation mosaic virus. Journal of General Virology 75, 997–1007. Gao, F., Alekhina, O.M., Vassilenko, K.S., Simon, A.E., 2018. Unusual dicistronic expression from closely spaced initiation codons in an umbravirus subgenomic RNA. Nucleic Acids Research 46, 11726–11742. Gao, F., Kasprzak, W.K., Szarko, C., Shapiro, B.A., Simon, A.S., 2014. The 30 Untranslated region of Pea enation mosaic virus contains two T-shaped, ribosome-binding, cap-independent translation enhancers. Journal of Virology 88, 11696–11712. Kim, S.H., Ryabov, E.V., Brown, J.W., Taliansky, M., 2004. Involvement of the nucleolus in plant virus systemic infection. Biochemical Society Transactions 32, 557–560. Kim, S.H., Ryabov, E.V., Kalinina, N.O., et al., 2007. Cajal bodies and the nucleolus are required for a plant virus systemic infection. The EMBO Journal 26, 2169–2179. Nurkiyanova, K.N., Ryabov, E.V., Kalinina, N.O., et al., 2001. Umbravirus-encoded movement protein induces tubule formation on the surface of protoplasts and binds RNA incompletely and non-cooperative. Journal of General Virology 82, 2579–2588. Ryabov, E.V., Robinson, D.J., Taliansky, M., 2001. Umbravirus-encoded proteins that both stabilize heterologous viral RNA in vivo and mediate its systemic movement in some plant species. Virology 288, 391–400. Ryabov, E.V., Robinson, D.J., Taliansky, M.E., 1999. A plant virus-encoded protein facilitates long distance movement of heterologous viral RNA. Proceedings of the National Academy of Sciences of the United States of America 96, 1212–1217. Taliansky, M., Roberts, I.M., Kalinina, N., et al., 2003. An umbraviral protein, involved in long-distance RNA movement, binds viral RNA and forms unique, protective ribonucleoprotein complexes. Journal of Virology 77, 3031–3040. Taliansky, M.E., Robinson, D.J., 2003. Molecular biology of umbraviruses: Phantom warriors. Journal of General Virology 84, 1951–1960. Wolf, Y.I., Kazlauskas, D., Iranzo, J., et al., 2018. Origins and evolution of the global RNA virome. mBio 9. (e02329-18).

Varicosaviruses (Rhabdoviridae) Takahide Sasaya, National Agriculture and Food Research Organization, Fukuyama, Japan r 2021 Elsevier Ltd. All rights reserved.

Glossary Big-vein disease of lettuce A lettuce disease that induces abnormal dilation or enlargement of a vein or the artery in a lettuce leaf, decreasing market value and reducing the proportion of harvestable lettuce. The name ‘big-vein disease’ refers to the appearance of the major symptoms. L protein The main subunit of the polymerase complex, responsible for most of the functions required for transcription and replication: RNA-dependent RNA polymerase, mRNA 50 -capping, 30 -poly(A) synthesis, and protein kinase activities. Necrotic syndrome of lettuce A lettuce disease that induces necrotic rings and spots in lettuce leaf and root

under a hydroponic culture of a nutrient film technique, decreasing market value and reducing the yield of lettuce. Olpidium virulentus A obligate parasitic root-infecting fungus and a member of the Chytridiomycetes that seldom appears to be deleterious to the host plants but functions as vector of plant viruses. Stop-and-start model A model for RNA synthesis of nonsegmented negative-sense RNA viruses that the viral polymerase complex transcribes a gene, polyadenylates and releases mRNA, and reinitiates transcription of the next gene under the direction of the gene-junction sequence.

Introduction The big-vein disease of lettuce (Lactuca sativa L.) was first reported in 1934. The causal agent was postulated to be a root-infecting virus, but the virus particles have remained unidentified for more than half a century. In 1983, a rod-shaped virus was found in field-grown lettuce showing big vein-band chlorosis (Fig. 1). Because the rod-shaped virus is always associated with big-vein-affected lettuce, the virus was regarded as the agent that induced big-vein disease, without it being confirmed that the virus alone actually induced big-vein symptoms in lettuce. The rod-shaped virus was named Lettuce big-vein virus and was classified as the type species of a new genus Varicosavirus, which was established by the International Committee on the Taxonomy of Viruses in 2000. A second virus, Mirafiori lettuce big-vein virus (MLBVV, genus Ophiovirus), has been found in commonly coinfected field-grown lettuce showing big-vein symptoms. Mechanical inoculation of partially purified MLBVV-preparation and transmission experiments using MLBVV- or Lettuce big-vein virus-carrying fungal vectors independently showed that MLBVV, but not Lettuce big-vein virus, was the causal agent of big-vein disease. For these reasons, lettuce big-vein virus was renamed lettuce big-vein associated virus (LBVaV) in 2005. LBVaV does not seem to induce vein-band chlorosis in lettuce, which is the characteristic symptom of the big-vein disease. However, LBVaV induces necrotic rings and spots in lettuce under a hydroponic culture of a nutrient film technique and it has become clear that the virus is the causal agent of the necrotic syndrome of lettuce. Stunt disease of tobacco (Nicotiana tabacum L.) reported so far only in Japan was first recorded in 1931 as a seed-bed disorder. Stunt disease has seriously damaged tobacco leaf production in Japan until the 1960s, when the disease was found to be transmitted by an obligate parasitic soil-borne fungus and tobacco growers paid more attention to tobacco seed-bed sanitation.

Fig. 1 Electron micrograph of Lettuce big-vein associated virus. Transmission electron micrograph of a negative-stained purified virus preparation. The bar represents 100 nm.

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Tobacco with stunt symptoms invariably has a varicosavirus, and mechanical inoculation of tobacco seedlings with partially purified virus-preparation leads to typical tobacco stunting; Tobacco stunt virus (TStV) is regarded as the causal agent of tobacco stunt disease. Because of marked differences in host reactions, LBVaV and TStV were considered to be distinct viruses. However, a comparison of partial sequences of LBVaV and TStV suggests now that TStV is a strain of LBVaV.

Taxonomy and Classification The floating genus Varicosavirus was first established by the International Committee on the Taxonomy of Viruses in 2000. Even though LBVaV is a two-segmented negative-sense RNA virus with a fragile non-enveloped rod shape, LBVaV shares several features with plant rhabdoviruses, which generally have linear non-segmented negative-sense RNA genomes and large enveloped particles with a prominent fringe of peplomers. On the base of genome structures and amino acid sequence comparisons between LBVaV and other plant rhabdoviruses, the taxonomic position was re-evaluated by the current committee in 2018, and the genus Varicosavirus is now classified in the family Rhabdoviridae. Lettuce big-vein associated varicosavirus is the type species of this genus, and LBVaV and three tentative viruses, have been tentatively placed into the genus; Alopecurus myosuroides varicosavirus 1, Tobacco stunt virus, and Red clover associated varicosavirus. LBVaV particles are fragile non-enveloped rods about 18 nm in diameter, with modal lengths of 320–360 nm. Virus particles have a central canal about 3 nm in diameter and an obvious helix with a pitch of about 5 nm, appearing similar to the inner striated nucleocapsid core of rhabdoviruses. Virus particles are very unstable in vitro, so the helix of particles, especially those in purified preparations, tends to loosen and become partially uncoiled, even in preparations fixed with glutaraldehyde. LBVaV was initially believed to have a divided genome consisting of two segments of double-stranded RNA, approximately 7.0 and 6.5 kbp. Once LBVaV purification was established and the complete nucleotide sequence of LBVaV determined, a precise reinvestigation of LBVaV genome components showed that LBVaV is not a double-stranded RNA virus but a single-stranded RNA virus with a bipartite genome. Negative-sense and positive-sense RNAs are separately encapsidated in the virions and the negativesense RNA is predominant. Criteria for membership in the genus are non-enveloped rod-shaped particles, a two-segmented negative-sense RNA genome, and transmission through moist soil by motile zoospores of the obligate parasitic soil-borne fungus, Olpidium virulentus.

Genetics and Evolution Nucleotide sequence and genome mapping studies showed that the LBVaV genome consists of 12878 nucleotides, divided into two segments (Fig. 2). The larger segment (RNA1) contains antisense information for a small hypothetical protein and a large gene that encodes the large protein (L) of 2040 amino acids. The smaller LBVaV genome segment (RNA2) contains antisense information for five major genes, which have coding capacities for 397, 333, 290, 164, and 368 amino acids, respectively.

L Protein Encoded on RNA1 As with other negative-sense RNA viruses, the L protein of LBVaV is positively charged and contains RNA-directed RNA polymerase (RdRp) and RNA binding domains. Alignment of the L protein sequence with those of several other negative-sense RNA viruses shows the conservation of functional RdRp motifs (Fig. 2). The L protein of LBVaV contains conserved premotif A, which presumably plays an important role in RNA template binding and positioning, and four motifs that correspond to motifs A, B, C, and D, thought to comprise the palm and finger regions of the polymerase active site in negative-sense RNA viruses. A putative ATP binding motif, Kx17GxGxG, proposed to be associated with polyadenylation or protein kinase activity, is present at position 1643–1665. LBVaV has been experimentally shown to transcribe 30 -polyadenylated mRNAs.

Fig. 2 Schematic representation of the genome organization of a negative-sense arrangement of genes encoded in the genome of Lettuce big-vein associated virus. Gene-junction regions, including the gene-end sequence (I), the intergenic sequence (II), and the gene-start sequence (III), are shown above negative-sense genomic RNAs.

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Fig. 3 Comparison of conserved motifs of the L protein among LBVaV, eight rhabdoviruses, and two plant segmented negative-sense RNA viruses. The amino acid sequence of the L protein of LBVaV is shown at the top followed by amino acid sequences of the cytorhabdoviruses Lettuce necrotic yellows virus (LNYV) and Northern cereal mosaic virus (NCMV), the nucleorhabdoviruses Sonchus yellow net virus (SYNV) and Rice yellow stunt virus (RYSV), and the vertebrate rhabdoviruses Vesicular stomatitis Indiana virus (VSIV)(now Indiana vesiculovirus), Rabies virus (RABV)(now Rabies lyssavirus), Bovine ephemeral fever virus (BEFV)(now Bovine fever ephemerovirus), Infectious hematopoietic necrosis virus (IHNV)(now Salmonid novirhabdovirus), the orthotospovirus Tomato spotted wilt virus (TSWV) and the tenuivirus Rice stripe tenuivirus (RSV). Numbers at the beginning of lines indicate the position of the first displayed amino acid. Numbers within brackets indicate the numbers of amino acids not represented in the figure. Conserved residues recognized previously for L proteins of negative-sense RNA viruses are shown in bold letters and residues characteristic of the division between nonsegmented and segmented negative-sense RNA viruses are underlined.

The L protein is the only region sufficiently conserved to enable evolutionary relationships to be established among negativesense RNA viruses. Even though LBVaV is segmented, most of the conserved motifs previously identified in the L proteins of non-segmented negative-sense RNA viruses are conserved in the L protein of LBVaV (Fig. 3), particularly the conserved sequence GDN in motif C, which likely constitutes the evolutionary and functional equivalent of the sequence GDD in polymerases of positive-sense RNA viruses. This is in striking contrast to segmented negative-sense RNA viruses whose polymerase has the conserved sequence SDD. Amino acids G at position 662 and W at position 671 in motif B, specific to the nonsegmented viruses, are also conserved in the L protein of LBVaV. Tetrapeptide E(F/Y)xS, located downstream from motif D, which is specifically conserved in polymerases of segmented negative-sense RNA viruses, is not found in the L protein of LBVaV. Phylogenetic analysis of the partial L proteins between LBVaV and rhabdoviruses shows that the L protein of LBVaV clusters within the family Rhabdoviridae, and that the L protein of LBVaV is more distantly related to those of plant rhabdoviruses than they are to each other, but is more closely related to those of plant rhabdoviruses than those of vertebrate rhabdoviruses (Fig. 4).

Proteins Encoded on RNA2 The first gene at the 30 end of genomic RNA encodes the coat protein (CP), which shows low but significant similarity to the nucleocapsid (N) proteins of plant rhabdoviruses. Most other genes encode hypothetical proteins with unknown functions, while the third encoded protein has several properties related to a viral cell-to-cell movement protein (MP). Gene Bank database homology searches using the amino acid sequence of the CP of LBVaV consistently retrieve N proteins of rhabdoviruses. Phylogenetic analysis between the CP of LBVaV and N proteins of rhabdoviruses shows that the CP clusters within the family Rhabdoviridae, and that the CP is most closely related to the N protein of a fish rhabdovirus, infectious hematopoietic necrosis virus (IHNV), and to a lesser extent to that of plant rhabdoviruses. In a PSI-BLAST search against the ProDom protein domain database, the carboxy-terminal part of the CP shows similarity to two domains, one is the homologous domain among N proteins of plant rhabdoviruses and another is the homologous domain among those of fish rhabdoviruses.

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Fig. 4 Computer-generated tree illustrating phylogenetic relationships between LBVaV and selected rhabdoviruses derived from aligned amino acid sequences of the partial L protein between premotif A and motif D. The tree was generated using the neighbor-joining method of the MEGA6 software. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) are shown. The evolutionary distances were computed using the Poisson correction method. Borna disease virus (BDV)(now Mammalian 1 orthobornavirus) is used as an out-group. See Fig. 3 for abbreviations of virus names.

The protein 2 of LBVaV is coded in the second gene that corresponds in position to the gene coding for the phosphoprotein (P) of rhabdoviruses, which interacts with the N protein and the L protein and constitutes the active polymerase complex. The predicted overall structure of the protein 2 and P proteins, assessed by amino acid composition and by relative hydropathicity distribution, is similar. Coiled-coil regions of P proteins of rhabdoviruses have been used to predict the regions for their oligomerization and interaction. Analysis using the COILS2 program predicts that the protein 2 contains three possible coiled-coil regions at the amino-terminal (aa 29-48), central (aa 122-135), and carboxy-terminal (aa 239-253) domains that are similar in position, to those in the P protein of Sonchus yellow net virus (SYNV). A detailed analysis of phosphorylation sites for cellular casein kinase II in the P protein of vesicular stomatitis Indiana virus (VSIV) confirmed five of ten potential phosphorylation sites of the P protein, which may regulate transcription and replication, are required to form the active polymerase complex. The protein 2 of LBVaV also has ten potential phosphorylation sites, and the five essential phosphorylation sites for the P protein of VSIV are conserved in positions similar to those of LBVaV. A predominantly basic 20-amino-acid-long region near the carboxy-terminus of the P protein of VSIV, which may serve as an attachment domain for combination with the L protein, also occurs in the carboxyterminal region of the protein 2 of LBVaV. The protein 3 of LBVaV is encoded in the third gene that corresponds in position to the gene coding for the MP of plant rhabdoviruses, which is essential for the plant viruses to assist in cell-to-cell movement through the host plant plasmodesmata. The protein 3 had a predicted structure similar to the 30K superfamily MP consensus structure and consisted of a series of b-elements flanked by an a-helix on each end. Our unpublished data of a transient expression using the GFP-fused P3 protein in epidermal cells of Nicotiana benthamiana showed the localization of the P3 protein to the plasmodesmata. The movement hypothesis for the protein 3 has been reinforced by our trans-complementation experiment with movement-defective tomato mosaic virus in N. benthamiana leaves: the protein 3 facilitated the spread of the movement-defective mutant from initially infected cells through two or more cell layers. The protein 5 of LBVaV, which corresponds in position to the gene coding for the glycoprotein (G) of plant rhabdoviruses, has also little direct relatedness to the envelope-associated glycoproteins of rhabdoviruses. The protein 5 sequence does not contain any canonical sites for N-linked glycosylation, and does do not contain the N-terminal signal peptide or carboxy-terminal hydrophobic transmembrane anchor domains, which are common among G proteins of rhabdoviruses. Conservation of cysteine residues is another common feature of G proteins of rhabdoviruses. Alignment using the Clustal W program for G proteins with plant rhabdoviruses shows that eleven cysteine residues are situated at conserved positions. In the case of LBVaV, six of ten cysteine residues are located at similar conserved positions in G proteins of plant rhabdoviruses. The protein 5 of LBVaV may thus result from the degeneration of G proteins of plant rhabdoviruses. The six cysteine residues in the protein 5 of LBVaV that may play an important role in activities, such as forming the secondary structure of G proteins of plant rhabdoviruses appear to have been stringently conserved among the protein 5 of LBVaV and G proteins of plant rhabdoviruses. The other five cysteine residues, however, appear to have evolved extensively to accommodate diverse host requirements and vector adaptations.

Transcription Termination/Initiation Strategy LBVaV transcribes capped and polyadenylated mono-cistronic mRNAs. In the gene-junction regions of LBVaV, transcription termination/polyadenylation and initiation signal sequences comparable to those of rhabdoviruses are recognized (Fig. 5), suggesting that LBVaV and rhabdoviruses may use a similar mechanism of transcription, a stop-and-start model, to differentially express individual genes from a contiguous virus genome.

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Fig. 5 Comparison of the gene-junction regions between LBVaV and selected rhabdoviruses. Gene-junction regions are separated into three sequences: I is the gene-end sequence that constitutes the poly(U) tract at the 30 end of each gene on negative-sense genomic RNAs and plays the role of a transcription termination/polyadenylation signal; II is the short intergenic sequence that is not transcribed during mRNA synthesis; and III is the gene-start sequence that constitutes the initiation site for transcription of each mRNA. Bold type in viral sequences indicates consensus nucleotides, R and W indicate A/G and A/U, and (N)x corresponds to a variable number of nucleotides. See Fig. 3 for abbreviations of virus names.

The gene-junction regions of LBVaV are grouped into three elements: a gene-end sequence (I) at the 30 end of each gene on the genomic template, which is required for transcription termination/polyadenylation; a short non-transcribed intergenic sequence (II), which is not represented in mRNAs and separates each gene; and a gene-start sequence (III) located at the beginning of each subsequent gene, which plays an important role in transcription initiation. Gene-end sequences of rhabdoviruses consist of an A-U rich region, a cytidylate, and a poly(U) tract. The poly(U) tract is thought to be a template for the poly(A) tail of mRNA due to reiterative transcription or slippage. The gene-end sequences of LBVaV are either 30 -UNCAUUUUUUU-50 (Type A) or 30 -AAUCUUUUUU-50 (Type B), reminiscent of those observed in other rhabdoviruses. When the gene-end sequence of LBVaV is Type A, transcription of the next gene is initiated immediately after a single nucleotide G, while for Type B, the two genes (genes 5 and L) are located at the 50 -most ends of each RNA (negative-sense) or transcription is reinitiated after a long 42 nt intergenic sequence located between genes 4 and 5, suggesting that the two variations of the gene-end sequences may influence signaling initiation of downstream mRNA synthesis as reported for VSIV in which the gene-end sequence plays an important role both in transcription termination/polyadenylation and in reinitiating transcription of downstream genes. In contrast to other rhabdoviruses, including nucleorhabdoviruses whose gene-start sequence in the gene-junction regions is 30 -UUGU-50 , the conserved gene-start sequence of LBVaV is 30 -CUCU-50 . Sequence 30 -CUCU-50 is also conserved in the genejunction regions of the cytorhabdoviruses lettuce necrotic yellows virus and northern cereal mosaic virus, suggesting that sequence 30 -CUCU-5 may be important in acting as a transcription initiation signal for both LBVaV and these cytorhabdoviruses, which replicate in the cytoplasm of plant cells.

Host Range and Geographic Distribution Fungal vector transmission experiments in the greenhouse have shown that LBVaV systemically infects seven plant species in four families, and the infected plants including lettuce do not show any symptoms. However, the virus induced necrotic rings and spots in lettuce in hydroponic culture. In contrast, the experimental host range of TStV encompasses 41 species in nine dicotyledonous families by mechanical inoculation and 35 plant species in 13 families through fungal vector transmission. The virus does not infect lettuce, but induces vein clearing, vein necrosis, mottling, and stunting in many Nicotiana species. LBVaV is widespread in cool to temperate regions and several parts of subtropical regions in Europe, Asia, Australia, and North and South America, but TStV has only been reported in Japan.

Transmission and Vectors LBVaV is transmitted through motile zoospores of an obligate parasitic soil-inhabiting fungus. LBVaV is very stable in soil within long-lived resting fungal spores for 20 years or more, but it is remains unclear whether the virus replicates inside fungal spores. In 1878, the natural vector of LBVaV was reported to be Olpidium brassicae (Wor.) Dang. O. brassicae was first found in a cabbage (Brassica oleracea) root and the specific name of the fungus is derived from the genus name of the cabbage. Several strains of O. brassicae that differ significantly in host specificity, morphology, and virus transmissibility have been recognized, particularly a crucifer strain, which readily multiplies in roots of crucifer plants such as cabbage and mustard, and a noncrucifer strain that can not multiply in roots of crucifer plants. The crucifer strain is host-specific and the non-crucifer strain is highly polyxenous. Morphologically, zoosporangia and exit tubes of the crucifer strain differ slightly from those of the noncrucifer strain. The noncrucifer strain is a vector of the soil-borne viruses MiLV and tobacco necrosis virus, but the crucifer strain fails to transmit these viruses. Furthermore, the crucifer strain shows sexual reproductivity, although the noncrucifer strain does not reproduce sexually. Zoospores from a single-sporangium isolate of the crucifer strain do not develop a resting spore, but after mating between different mating (sexual) types, a resting spore form. In contrast, zoospores from the noncrucifer strain develop a resting spore without mating. Based on these differences, the crucifer and noncrucifer strains have been divided into two distinct species, but this

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proposal has long been neglected. Recent rDNA-ITS analysis of O. brassicae strongly supported the hypothesis that crucifer and non-crucifer strains belong to separate species rather than being strains of the same Olpidium species, and a new species name, O. virulentus, has been proposed for the non-crucifer strain. The non-crucifer strain was confirmed to be a vector of LBVaV, although there is no reliable evidence that the crucifer strain (O. brassicae) is capable of transmitting LBVaV.

Further Reading Banerjee, A.K., Barik, S., De, B.P., 1991. Gene expression of non-segmented negative strand RNA viruses. Pharmacology & Therapeutics 51, 47. Jackson, A.O., Dietzgen, R.G., Goodin, M.M., Bragg, J.N., Deng, M., 2005. Biology of plant rhabdoviruses. Annual Review of Phytopathology 43, 623. Lot, H., Campbell, R.N., Souche, S., Milne, R.G., Roggero, P., 2002. Transmission by Olpidium brassicae of Mirafiori lettuce virus and Lettuce big-vein virus, and their roles in lettuce big-vein etiology. Phytopathology 92, 288. Maccarone, L.D., 2013. Relationships between the pathogen Olpidium virulentus and viruses associated with lettuce big-vein disease. Plant Disease 97, 700–707. Poch, O., Sauvaget, I., Delarue, M., Tordo, N., 1989. Identification of four conserved motifs among the RNA-dependent polymerase encoding elements. The EMBO Journal 8, 3867. Rose, J.K., Whitt, M.A., 2001. Rhabdoviridae: The viruses and their replication. In: Knipe, M., Howley, P.M. (Eds.), Field Virology, fourth ed. New York: Raven Press, p. 1221. Sasaya, T., Ishikawa, K., Koganezawa, H., 2002. The nucleotide sequence of RNA1 of Lettuce big-vein virus, genus Varicosavirus, reveals its relation to nonsegmented negativestrand RNA viruses. Virology 297, 289. Sasaya, T., Koganezawa, H., 2006. Molecular analysis and virus transmission tests place Olpidium virulentus, a vector of Mirafiori lettuce big-vein virus and Tobacco stunt virus, as a distinct species rather than a strain of Olpidium brassicae. Journal of General Plant Pathology 72, 20. Sasaya, T., Kusaba, S., Ishikawa, K., Koganezawa, H., 2004. Nucleotide sequence of RNA2 of Lettuce big-vein virus and evidence for a possible transcription termination/ initiation strategy similar to that of rhabdoviruses. Journal of General Virology 85, 2709. Verbeek, M., Dullemans, A.M., van Bekkum, P.J., van der Vlugt, R.A.A., 2013. Evidence for Lettuce big‐vein associated virus as the causal agent of a syndrome of necrotic rings and spots in lettuce. Plant Pathology 62, 444–451.

Virgaviruses (Virgaviridae) Eugene I Savenkov, Swedish University of Agricultural Sciences, Uppsala, Sweden and Linnean Center for Plant Biology, Uppsala, Sweden r 2021 Elsevier Ltd. All rights reserved.

Nomenclature

MP Movement protein NGS Next generation sequencing nt Nucleotide(s) ORF Open reading frame PCR Polymerase chain reaction RdRp RNA-dependent RNA polymerase RTD Read-through domain sgRNA Sub-genomic RNA ssRNA single-stranded RNA TGB Triple gene block UTR Untranslated region VSR Viral suppressor of RNA silencing

aa Amino acid(s) CP Coat protein or capsid protein CRP Cysteine-rich protein diRNA Defective interfering RNA EM Electron microscopy ER Endoplasmic reticulum Hel Helicase IPM Integrated pest management kb Kilobase kDa Kilo Dalton MET Methyl-transferase

Glossary 126-kDa protein The first protein expressed during tobamovirus infection, it has methyl-transferase and helicase domains, but lacks a polymerase domain. Readthrough of the stop-codon of the open reading frame (ORF) encoding 126 kDa protein results in translation of the larger the 183 kDa protein, which has a polymerase domain. Alpha-like viruses A group of viruses in the proposed classification of positive-stranded RNA viruses based on the sequence similarities and arrangement of domains of RNAdependent RNA polymerase. Three groups have been recognized: the picorna-like group, the flavi-like group and the alpha-like group. Alpha-like group is named after the genus Alphavirus of the family Togaviridae. Cross-protection A phenomenon whereby prior infection by one virus strain (the primary strain) can prevent or limit multiplication of another virus strain (the secondary strain) or a closely related virus. For this purpose, protecting strains (the primary strains) should cause attenuated symptoms and should reduce marketable yields only slightly. A few attenuated virus strains have been commercially used as a “vaccine”. Cross-protection has been applied in the fields to control poty-, tobamo- and closteroviruses. The phenomenon was first reported with Tobacco mosaic virus (TMV) in 1929. Defective interfering RNAs Subviral agents produced during replication of RNA viruses. Defective interfering (di) RNAs are parasitic RNAs because they take advantage of the parent virus-coded protein for their multiplication. These RNAs are called “defective” because they have lost the

capacity to code for the proteins needed for independent replication and thus are defective in the absence of the helper (parent) virus. diRNAs are referred as “interfering” because they can attenuate the symptoms caused by the helper virus, they multiply rapidly and eventually lead to slowing down of the helper virus multiplication. Lateral flow detection A chromatography method based on lateral flow using specific antibodies applied on strips. Once the strip is submerged with the “sample” side into the sample extract, the liquid migrates upwards and initiates the antigen-antibody reaction which results in visible lines. Both test and control lines become visible with positive extracts, whereas negative samples produce the upper control line only. Lines start developing after 1–2 mn and reach maximum intensity after 10–15 mn. Dried test strips can be kept as permanent records. Leaky scanning A mechanism used during the initiation steps of eukaryotic translation in which a “weak” initiation codon on mRNA is sometimes skipped by ribosome. During translation initiation, the small 40S ribosomal subunit “scans” or moves in a 50 ––4 30 direction along the 50 untranslated region to locate a start codon to commence elongation. Sometimes, the scanning ribosome may encounter an “unfavorable nucleotide context” around the start codon, bypasses this start codon and begins translation at further downstream AUG start codons. This way an mRNA can encode several different proteins if the AUG are not in frame or code for proteins with different N-termini if the AUG start codons are in the same frame.

Introduction The family Virgaviridae contains seven genera (Furovirus, Goravirus, Hordeivirus, Pecluvirus, Pomovirus, Tobamovirus, and Tobravirus) and twenty-seven species unassigned to a genus (Table 1). Virgaviruses possess positive-sense single-stranded RNA genomes encapsidated in rod-shaped particles. Virions are helically constructed from one type of coat protein (CP) with exception in members of the Furovirus and Pomovirus genera, which incorporate an additional larger minor CP into one extremity of the virus

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Table 1

Virgaviruses (Virgaviridae) List of members in the genera of the family Virgaviridae. Type species are in bold

Genus/Species

Acronym

# Segments

Furovirus Chinese wheat mosaic virus

CWMV

2

Japanese soil-borne wheat mosaic virus

Oat golden stripe virus

Soil-borne cereal mosaic virus

Soil-borne wheat mosaic virus

Sorghum chlorotic spot virus

Unassigned Furoviruses French barley mosaic virus

Goravirus Drakaea virus A

Gentian ovary ringspot virus

Hordeivirus Anthoxanthum latent blanching virus Barley stripe mosaic virus

Lychnis ringspot virus

Poa semilatent virus

Pecluvirus Indian peanut clump virus

Peanut clump virus

JSBWMV

OGSV

SBCMV

SBWMV

SCSV

FBMV

DraVA

GORSV

ALBV BSMV

LRSV

PSLV

IPCV

PCV

Access. #

Length (nt)

NC002359 NC002356

7147 3569

NC038850 NC038851

7226 3574

NC002357 NC002358

3232 7111

NC002351 NC002330

7025 3683

NC002041 NC002042

7099 3593

NC004015 NC004014

3418 6878

AJ749658 AJ749657

1507a 3291a

NC043398 NC043399

4490a 2905a

NC024501 NC024502

5519 3869

2

2

2

2

2

2

2

2

– 3

ND NC003469 NC003478 NC003481

3768 3164 3289

NC038933 NC038932 ND

1468a 3065a

MK377386 MK377387 MK377388

3873 3608 3169

NC004730 NC004729

4507 5841

NC003668 NC003672

4504 5897

3

3

2

2

Virgaviruses (Virgaviridae)

Table 1

841

Continued

Genus/Species

Acronym

# Segments

Pomovirus Beet soil-borne virus

BSBV

3

Beet virus Q

Broad bean necrosis virus

Colombian potato soil-borne virus

Potato mop-top virus

Unassigned Pomoviruses Soil-borne virus 2

BVQ

BBNV

CoPSNV

PMTV

SBV2

Access. #

Length (nt)

NC003520 NC003519 NC003518

5834 3005 3454

NC003512 NC003511 NC003510

2529 2913 6003

NC004424 NC004423 NC004425

2831 5600 2417

NC029037 NC029035 NC029034

3028 3164 6170

NC003723 NC003725 NC003724

6043 2964 3134

KT225277 KT225278

5961 2532 6375 6381 6449 6514 6562 6424 6485 6643 6431 6485 6514 6794 6507 6618 6453 6524 6791 6357 6688 6506 6395 6311 6279

3

3

3

3

2

Tobamovirus Bell pepper mottle virus Brugmansia mild mottle virus Cactus mild mottle virus Clitoria yellow mottle virus Cucumber fruit mottle mosaic virus Cucumber green mottle mosaic virus Cucumber mottle virus Frangipani mosaic virus Hibiscus latent Fort Pierce virus Hibiscus latent Singapore virus Kyuri green mottle mosaic virus Maracuja mosaic virus Obuda pepper virus Odontoglossum ringspot virus Opuntia virus 2 Paprika mild mottle virus Passion fruit mosaic virus Pepper mild mottle virus Plumeria mosaic virus Rattail cactus necrosis-associated virus Rehmannia mosaic virus Ribgrass mosaic virus Streptocarpus flower break virus Sunn-hemp mosaic virus Tobacco latent virus Tobacco mild green mosaic virus

BPMV BrMMoV CMMoV ClYMoV CGMoMV CFMoMV CuMoV FrMV HLFPV HLSV KGMoMV MaMV ObPV ORSV OV2 PaMMoV PfMV PeMMoV PlMV RCNaV ReMV RMV SFBV SHMV TLV TMGMV

1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

NC009642 NC010944 NC011803 NC016519 NC002633 NC001801 NC008614 NC014546 NC025381 NC008310 NC003610 NC008716 NC003852 NC001728 NC040685 NC004106 NC015552 NC003630 NC026816 NC016442 NC009041 NC002792 NC008365 ND NC038703 NC001556

Tobacco mosaic virus Tomato brown rugose fruit virus Tomato mosaic virus Tomato mottle mosaic virus Tropical soda apple mosaic virus

TMV ToBRFV ToMV ToMoMV TSAMV

1 1 1 1 1

NC001367 NC028478 NC002692 NC022230 NC030229

1415a 6355 6395 6393 6383 6398 6350 (Continued )

842

Table 1

Virgaviruses (Virgaviridae)

Continued

Genus/Species

Acronym

# Segments

Access. #

Length (nt)

Turnip vein-clearing virus Ullucus mild mottle virus Wasabi mottle virus Yellow tailflower mild mottle virus Youcai mosaic virus Zucchini green mottle mosaic virus

TuVCV UMMoV WMoV YTMMoV YMoV ZGMoV

1 1 1 1 1 1

NC001873 ND NC003355 NC022801 NC004422 NC003878

6311

Unassigned Tobamoviruses Actinidia tobamovirus Abutilon yellow mosaic virus Acidomyces richmondensis tobamovirus 1 Bottle gourd mottle virus Brugmansia latent virus Crucifer tobamovirus Hoya chlorotic spot virus Nigerian tobacco latent virus Penstemon ringspot virus Piper chlorosis virus Podosphaera prunicola tobamovirus Watermelon green mottle mosaic virus

AcTV AbYMV ArTV1 BGMoV BrLV CTV HCSV NTLV PRSV PiCV PpTV WGMoV

1 1 1 1 1 1 1 1 1 1 1 1

ND EU559678 MK279511 MF960828 NC010944 AB003936 NC034509 ND ND KX683424 ND MH837097

Tobravirus Pea early-browning virus

PEBV

2

Pepper ringspot virus

Tobacco rattle virus

PeRSV

TRV

6298 6379 6303 6513

5903 10291 6497 6381 2378a 6386

5686a 6482

NC001368 NC002036

3374 7073

NC003670 NC003669

1799 6828

NC003811 NC003805

3855 6791

MN661103 MN661104 MN661112 MN661113 MN661116 MN661118 MN661120 MH614308 MH614309 MH614318 MH614319 MK189194 MG195286 MH188019 MN513382 NC025674 KP900897 NC040613 NC040614 MN551100 MN551103 MN551104 MN551118 MN551125 MN551126 LC333735 KU236377

9006 8775 10309 9638 10862 10224 10266 1681a 962a 2265 1840a 3687a 2463a 10917 10200 9524 7499 8138 11919 6782 3486 3594 3407 3528 3560 877a 3574

2

2

Unassigned Virgaviruses Atrato virga-like virus 1 Atrato virga-like virus 2 Atrato virga-like virus 3 Atrato virga-like virus 4 Atrato virga-like virus 5 Atrato virga-like virus 6 Atrato virga-like virus 7 Bombus-associated virus 1 Bombus-associated virus 2 Bombus-associated virus 3 Bombus-associated virus 4 Botryosphaeria dothidea tobamo-like virus Cucumis melo virga-like virus Culex virga-like virus Hammarskog virga-like virus Macrophomina phaseolina tobamo-like virus Macrophomina phaseolina tobamo-like virus 1a Nephila clavipes virus 3 Nephila clavipes virus 4 Plasmopara viticola associated virga-like virus 1 Plasmopara viticola associated virga-like virus 2 Plasmopara viticola associated virga-like virus 3 Plasmopara viticola associated virga-like virus 5 Plasmopara viticola associated virga-like virus 6 Plasmopara viticola associated virga-like virus 7 Rosellinia necatrix virga-like-virus 1 Soil-borne barley mosaic virus a

Incomplete genomic sequence.

AVLV1 AVLV2 AVLV3 AVLV4 AVLV5 AVLV6 AVLV7 BaV1 BaV2 BaV3 BaV4 BdTLV CmVLV CuVLV HVLV MPTLV MpTLV1a NcV3 NcV4 PvaVLV1 PvaVLV2 PvaVLV3 PvaVLV5 PvaVLV6 PvaVLV7 RnVLV SBBMV

Virgaviruses (Virgaviridae)

843

Table 2 Classification of members of the family Virgaviridae into genera, with indication of type species, number of virus species, number of RNA genome components, type of movement protein, transmission vector and host type Genus

Type species

Furovirus Goravirus Hordeivirus Pecluvirus Pomovirus Tobamovirus Tobravirus

Soil-borne wheat mosaic virus Gentian ovary ringspot virus Barley stripe mosaic virus Peanut clump virus Potato mop-top virus Tobacco mosaic virus Tobacco rattle virus

Acronym

Numbera

SBWMV GORSV BSMV PCV PMTV TMV TRV

6þ1 2 4 2 5þ1 37 þ 12 3

Genome

MP

Bipartite Bipartite Tripartite Bipartite Tripartite Monopartite Bipartite

‘30K’ TGB TGB TGB TGB ‘30K’ ‘30K’

30 structure Val

tRNA tRNA tRNATyr tRNAVal tRNAVal/Leu tRNAHis tRNA

Transmission

Host type

P. graminis Pollen Pollen/seed P. graminis Plasmodiophorids Mechanical Nematodes

Monocots Dicots Monocots/dicots Monocots/dicots Dicots Dicots Dicots

a

Number of members according to the International Committee on Taxonomy of Viruses (ICTV) as of January 2020.

particles. This protein is produced through translational read-through of the stop codon of CP-encoding open reading frame (ORF) and is needed for acquisition and transmission of the viruses of these two genera by plasmodiophorid protist vectors. Many virgaviruses cause economically significant diseases in vegetable, fruit, cereal and fiber crops, and in ornamental plants worldwide. The members of Virgaviridae infect both monocot and dicots species and the viruses belonging to the Furovirus, Pecluvirus, Pomovirus and Tobravirus genera are soil-borne and transmitted in soil by plasmodiophorid and nematode vectors. Most Tobamoviruses are transmitted mechanically and seems not to have natural vectors. Tobacco mosaic virus (TMV; genus Tobamovirus) was the first virus to be discovered (in 1886) and crystallized (in 1935). The virus accumulates in extremely high concentrations in infected plants, extremely stable and is easily transmitted from plant to plant just through the contact of an infected plant with a healthy plant, e.g., through contacts facilitated by wind or air flow. This particular property of the virus, its various strains, and related viruses is used in agriculture for cross-protection.

Taxonomy, Phylogeny and Evolution Genus demarcation criteria in the family Virgaviridae are based on number of genome components (mono-, bi- or tripartite), genome organization (e.g., type of the encoded movement protein, MP), mode of transmission (pollen, seed, mechanical or by vector), type of vector (plasmodiophorid or nematode), and phylogenetic relationships. Currently, virgaviruses are classified into seven genera: Furovirus, Goravirus, Hordeivirus, Pecluvirus, Pomovirus, Tobamovirus, and Tobravirus, briefly summarized below (Table 2). The genus Furovirus comprises six recognized species: Chinese wheat mosaic virus (CWMV), Japanese soil-borne wheat mosaic virus (JSBWMV), Oat golden stripe virus (OGSV), Soil-borne cereal mosaic virus (SBCMV), Soil-borne wheat mosaic virus (SBWMV), and Sorghum chlorotic spot virus (SCSV); and one unassigned member: French barley mosaic virus (FBMV). The members of the genus are all bipartite soil-borne viruses transmitted by the plasmodiophorid Polymyxa graminis to graminaceous plants (monocots). The genus Goravirus comprises two recognized species, Drakaea virus A (DraVA) and Gentian ovary ringspot virus (GORSV). The members of the genus are bipartite pollen-transmitted viruses, which infect ornamental plants (an orchid and a clustered gentian). The genus Hordeivirus comprises four recognized species, Anthoxanthum latent blanching virus (ALBV), Barley stripe mosaic virus (BSMV), Lychnis ringspot virus (LRSV), and Poa semilatent virus (PSLV). The members of the genus are tripartite pollen- and seedtransmitted viruses, which infect monocot and dicot plants. The genus Pecluvirus includes two recognized species, Indian peanut clump virus (IPCV) and Peanut clump virus (PCV). The members of the genus are bipartite soil-borne viruses transmitted naturally by Polymyxa graminis or by seed (in groundnuts). The viruses are also mechanically transmissible and infect monocots and dicots. The genus Pomovirus comprises five recognized species and one unassigned member: Beet soil-borne virus (BSBV), Beet virus Q (BVQ), Broad bean necrosis virus (BBNV), Colombian potato soil-borne virus (CPSbV), Potato mop-top virus (PMTV) and Soil-borne virus 2 (a tentative member). The members of the genus are tripartite soil-borne viruses transmitted by the plasmodiophorids to dicot plants. The genus Tobamovirus, with 37 accepted species and 12 unassigned species, is the largest genus in the family Virgaviridae. Tobamoviruses are the only members of the family to have a non-segmented (monopartite) genome, are not vector-transmissible and when seed transmitted, the embryo is not affected. Tobamoviruses are efficiently transmitted mechanically and include TMV, the first virus to be discovered and crystallized, thus, extensively studied. The genus Tobravirus comprises three recognized species, Pea early-browning virus (PEBV), Pepper ringspot virus (PRSV), and Tobacco rattle virus (TRV). The members of the genus are bipartite soil-borne viruses transmitted by nematodes to dicot plants. Additionally, twenty-seven unassigned species in the family Virgaviridae have been recognized and include only two plant-infecting viruses: Cucumis melo virga-like virus and Soil-borne barley mosaic virus. The remaining twenty-five species were identified in metagenomics libraries and include viruses found in fungi, oomycetes, insects (mosquitoes and bumble bees), and spiders (Table 1, Table 3). Phylogenetic analysis of the sequences of replication protein of plant-infecting Virgaviridae shows that virgaviruses group in clusters corresponding to the seven existing genera (Fig. 1). Similar clustering is obtained when phylogenetic analysis is performed for the sequences of movement proteins, although ‘30K’-type and TGB1-type MPs are analyzed separately. There are also close relationships between the cysteine-rich proteins (CRPs) encoded by the viruses of the Furovirus, Hordeivirus, Pecluvirus, and

844

Table 3

Virgaviruses (Virgaviridae)

Unassigned virgaviruses, with indication of virus species, number of genes, host and country of origin

Species

Acronym

Number of genes

Atrato virga-like virus 1 Atrato virga-like virus 2 Atrato virga-like virus 3 Atrato virga-like virus 4 Atrato virga-like virus 5 Atrato virga-like virus 6 Atrato virga-like virus 7 Bombus-associated virus 1 Bombus-associated virus 2 Bombus-associated virus 3 Bombus-associated virus 4 Botryosphaeria dothidea tobamo-like virus Culex virga-like virus Hammarskog virga-like virus Macrophomina phaseolina tobamo-like virus Macrophomina phaseolina tobamo-like virus 1a Nephila clavipes virus 3 Nephila clavipes virus 4 Plasmopara viticola associated virga-like virus 1 Plasmopara viticola associated virga-like virus 2 Plasmopara viticola associated virga-like virus 3 Plasmopara viticola associated virga-like virus 5 Plasmopara viticola associated virga-like virus 6 Plasmopara viticola associated virga-like virus 7 Rosellinia necatrix virga-like-virus 1

AVLV1 AVLV2 AVLV3 AVLV4 AVLV5 AVLV6 AVLV7 BaV1 BaV2 BaV3 BaV4 BdTLV CuVLV HVLV MPTLV MpTLV1a NcV3 NcV4 PvaVLV1 PvaVLV2 PvaVLV3 PvaVLV5 PvaVLV6 PvaVLV7 RnVLV

2 2 2 2 4 4 4 ? ? 3 ? ? 3 4 4 4 2 3 1 3 2 1 1 2 ?

(PolyPra, CP) (PolyPr, CP) (PolyPr, CP) (PolyPr, CP) (PolyPr, CPs, UL36b) (PolyPr, HPsc) (PolyPr, HPs)

(?)

(Repd, HPs) (HPs) (Rep, MP, CP) (Rep, MP, CP) (Rep, gp2e) (Rep, gp1, gp2) (Rep) (Rep, HPs) (HP, Rep) (Rep) (Rep) (Rep, HP)

Host

Host species

Country

mosquito mosquito mosquito mosquito mosquito mosquito mosquito bumble bee bumble bee bumble bee bumble bee fungus mosquito mosquito fungus fungus spider spider oomycete oomycete oomycete oomycete oomycete oomycete fungus

Culex sp. Coquillettidia venezuelensis Mansonia titillans Psorophora albipes Mansonia titillans Psorophora sp. Anopheles darlingi Bombus sp. Bombus sp. Bombus sp. Bombus sp. Botryosphaeria dothidea Culex sp. Culex pipiens Macrophomina phaseolina Macrophomina phaseolina Nephila clavipes Nephila clavipes Plasmopara viticola Plasmopara viticola Plasmopara viticola Plasmopara viticola Plasmopara viticola Plasmopara viticola Rosellinia necatrix

Colombia Colombia Colombia Colombia Colombia Colombia Colombia UK: Scotland UK: Scotland UK: Scotland UK: Scotland China USA Sweden USA USA USA USA Spain Italy Italy Spain Italy Italy Spain

a

PolyPr, polyprotein. UL36, Large tegument protein UL36. c HP, hypothetical protein. d Rep, replicase. e gp, glycoprotein. b

Tobravirus genera, although CRP of a pomovirus, Potato mop-top virus (PMTV), do not align with them and has an entirely different sequence. In addition, within genera, closely-related tobamoviruses are grouped according to host they infect. Among 37 tobamovirus species, there are clearly groupings of closely-related viruses infecting similar hosts, namely those infecting cruciferous plants (Ribgrass mosaic virus, Turnip vein-clearing virus, Wasabi mottle virus, Youcai mosaic virus), cucurbits (Cucumber fruit mottle mosaic virus, Cucumber green mottle mosaic virus, Cucumber mottle virus, Kyuri green mottle mosaic virus, Zucchini green mottle mosaic virus), and solanaceous species (Brugmansia mild mottle virus, Obuda pepper virus, Paprika mild mottle virus, Pepper mild mottle virus, Rehmannia mosaic virus, Tobacco mild green mosaic virus, TMV, Tomato brown rugose fruit virus, Tomato mosaic virus, Tropical soda apple mosaic virus, Yellow tailflower mild mottle virus). Six other genera in the family Virgaviridae do not have enough diversity to distinguish particular sub-groupings because of the limited number virus species (two to six species depending on the genus) within the genus and very narrow host range (e.g., furoviruses, which infect graminaceous plants). Genetic diversity and evolution of virgaviruses seem to be driven by mutation, recombination and reassortment of genome components (for multipartite viruses) that significantly contribute to the emergence of new viral variants, increasing their potential of adaptation to different crop varieties, hosts and environmental conditions. These is nicely illustrated by the analysis of numerous isolates of PMTV. Phylogenetic analysis of available PMTV sequences for RNA-rep, RNA-TGB, and RNA-CP indicates that PMTV isolates around the world can be divided into two (RNA-rep and RNA-TGB) to three (RNA-CP) lineages depending on genomic segment. In all lineages, there are virus isolates from Andean region of South America demonstrating the grater variability of the virus in the Andes relative to the other parts of the world. Moreover, three of these lineages are exclusively represented by isolates from Andean region of Peru, the center of potato domestication; indicating that in Peru, PMTV has undergone great evolutionary divergence, resulting in the establishment of distinct phylogenetic clades, genotypes and genetic constellations (as PMTV has a tripartite genome).

Virion Structure Virgaviruses have non-enveloped, rod-shaped particles with helical symmetry with a pitch of 2.3–2.5 nm and an axial canal (Fig. 2). The virions are ca. 20 nm in diameter and up to 310 nm in length. No lipids have been reported in the virions of

Virgaviruses (Virgaviridae)

845

Fig. 1 Neighbor-joining phylogenetic tree of the codon-aligned nucleotide sequences of the replication proteins of plant-infecting viruses in the family Virgaviridae. The analyses were conducted in MEGA X using the Maximum Likelihood method, Tamura-Nei model and 1000 bootstrap replicates. The tree with the highest log likelihood (  185021,20) is shown. The percentage of trees in which the associated taxa clustered together is shown next to the branches. The tree is drawn to scale, with branch lengths measured in the number of substitutions per site. This analysis involved 54 nucleotide sequences. Virus abbreviations are explained in the Table 1. The bar below the tree indicates nucleotide substitutions per site.

846

Virgaviruses (Virgaviridae)

Fig. 2 (A) Negative stain electron micrograph of Barley stripe mosaic virus virions. Scale bar 500 Å . (B) Reconstruction of the Tobacco mosaic virus capsid by cryo-electron microscopy at 4.6 Å resolution. Molecular graphics were generated using data deposited in Protein Data Bank under the accession number 2XEA. (C) Reconstruction of the Barley stripe mosaic virus capsid by cryo-electron microscopy at 4.1 Å resolution. Molecular graphics were generated using data deposited in Protein Data Bank under the accession numbers: 5a79 for the wide structure (left panel) and 5a7a for the narrow structure (right panel). (D) Negative stained electron micrograph of Tobacco rattle virus virions. Scale bar 250 nm. Courtesy Elena V. Orlova, Institute of Structural and Molecular Biology, London, UK. (A) Reproduced from Kendall, A., Williams, D., Bian, W., Stewart, P.L., Stubbs, G., 2013. Barley stripe mosaic virus: Structure and relationship to the tobamoviruses. Virology 443, 265–270. Elsevier with permission. (D) Courtesy Elena V. Orlova, Institute of Structural and Molecular Biology, London, UK.

virgaviruses. The virions of all members of the family (except for members of the Furovirus and Pomovirus genera) are composed of identical CP subunits ranging in size from 17 to 24 kDa. In furoviruses and pomoviruses, the CP-RTD (coat protein – read-through domain) protein – produced by translational read-through of the CP ORF stop codon – is a minor virion component attached to one extremity of the virus particles. Currently, high-resolution capsid structures are available for five tobamoviruses including TMV, Cucumber green mottle mosaic virus (CGMMV), Tobacco mild green mosaic virus (TMGMV), Odontoglossum ringspot virus (ORSV), Ribgrass mosaic virus (RMV), and one hordeivirus – BSMV. The capsid structures of tobamoviruses are very similar to each other, whereas compared to the tobamovirus capsids, the BSMV virions are considerably more labile and, in general, the physico-chemical properties of BSMV virions resemble to certain extent those of flexuous filamentous viruses rather than tobamoviruses. The X-ray fiber diffraction analysis at 2.9 Å resolution and cryoelectron microscopy (EM) at 4.6 Å resolution has shown that the virions of TMV are built from one distinct conformation of a single CP. The inter-subunit contacts in tobamovirus virions include strong lateral and axial electrostatic interactions, and the hydrophobic girdle formed by lateral hydrophobic contacts between adjacent subunits at a high virion radius (Fig. 2(B)). These types of interactions are believed to predispose the exceptionally high stability of tobamovirus virions. Surprisingly, Cryo-EM structure at 4.1 Å resolution has shown that the capsids of the hordeivirus BSMV are built from two

Virgaviruses (Virgaviridae)

847

distinct conformations of a single CP, and that these conformational differences facilitate the formation two structurally distinct forms of BSMV virions, ‘wide’ and ‘narrow’ virions with diameters of 224 Å and 216 Å , respectively (Fig. 2(C)). Wide virions incorporate 111 subunits per helix period, while narrow virions have 106 subunits. Both virion forms co-exist in the populations of BSMV particles. The functional differences between the two forms of BSMV virions are not known. Structural data show that the narrow BSMV virions are more stable than the wide virions because of the stronger CP contacts with RNA and additional contacts between CP subunits. Each virion of virgaviruses contains a single copy of ssRNA. Thus, for viruses with bipartite or tripartite genomes, two or three virions, respectively, containing different genomic components are required for establishing infection. The differences in virion size are particularly evident for bipartite tobraviruses as they have virus particles of two predominant lengths, (L) 180–215 nm and (S) ranging from 46 to 115 nm, depending on the isolate (Fig. 2(D)). RNA1 is incapsidated into the L-particles, whereas the S-particles contain RNA2. Defective genome components (defective interfering RNA, diRNA) associated with pomoviruses have also been found to be encapsidated.

Genome Organization The 50 untranslated region (UTR) of many virgaviruses contain the starting sequence GU(A)1–4(U)n. The RNAs are probably capped at the 50 end since the viral replicases contains methyl-transferase motifs associated with capping activity. The terminal ca. 80–240 nt of the 30 UTR can be folded into a tRNA-like structure that contains an anticodon for valine (furo-, peclu- and pomoviruses), tyrosine (hordeiviruses) or histidine (tobamoviruses; Table 2). For some hordei-, pomo- and tobamoviruse the genomic RNAs were shown to be amino-acylated (tyrosilated, valylated and histidinated, respectively) experimentally. Different from other virgaviruses, tRNA-like structure of a pomovirus CPSBV contains an anticodone for leucine. Although genomic RNAs of virgaviruses are not polyadenylated, hordeiviruses BSMV and LRSV contain internal poly(A) tracts, which precede the tRNA-like structure. Genomic RNAs of virgaviruses typically have three to seven ORFs. The genome expression strategy is based on: (i) read-through of leaky stop codons - e.g., expression of replicases and CP-RTD proteins; (ii) expression of the downstream ORFs via the synthesis of a nested set of 30 co-terminal sub-genomic mRNAs (sgRNAs) – e.g., expression of MPs, cysteine-rich proteins and CP (in tobamoviruses); and (iii) leaky scanning – e.g., expression of TGB2/TGB3 (gora-, hordei- peclu- and pomoviruses), and CP/P39 (pecluviruses). The genome organization is conserved among species within the same genus, but not in the family in general (Fig. 3). However, the 50 proximal ORF of all largest genomic components invariably encodes the replication-associated proteins (replicases) with conserved domains for a methyltransferase (MET), a helicase (HEL), and an RNA-dependent RNA polymerase (RdRp), a domain arrangement typical of alpha-like viruses. The replicases are translated directly from the genomic RNAs. In viruses of all genera except the genus Hordeivirus, RdRp is expressed as the C-terminal part of the replicase by read-through of a leaky stop codon (Fig. 3). In the genus Hordeivirus RdRp is translated from a smaller genomic component RNAg. Other viral proteins are translated either directly from the smaller genomic RNAs or from sgRNAs, some of which may be bicistronic and are translated by leaky scanning mechanism. In viruses of all genera except the genus Tobamovirus, CP is translated directly from the smaller genomic RNAs. In the genus Tobamovirus, CP is expressed via formation of sgRNA. The viruses of the genera Furovirus, Tobamovirus and Tobravirus encode a single MP of the “30K” superfamily, whereas in other genera there is a triple gene block (TGB) of MPs (Fig. 3; Table 2). The coordinated expression and action of the proteins encoded by TGB mediates virus movement in the infected plant. The viruses of the genera Furovirus, Goravirus, Hordeivirus, Pecluvirus, Tobravirus, and PMTV, but not other pomoviruses, encode cysteine-rich proteins (CRPs, Fig. 3). These CRPs, except the one encoded by PMTV, share well-conserved C3H1-type (C, cysteine residue; H, histidine residue) zinc-finger domain. CRP encoded by PMTV has a SWIM-type zinc-finger characterized by a consensus sequence CxCxnCxC, in which C stands for cysteine residue and x refers to any aa residue. CRPs of virgaviruses are expressed from sgRNAs, function as suppressors of RNA silencing and are essential for virus infectivity or/and efficient virus accumulation in the infected plants. However, in tobamoviruses soluble replication proteins (not associated with membranes) are involved in suppression of RNA silencing, but do not participate in RNA replication (Fig. 3).

Replication After virus entry, the genomic RNA is translated to produce replication proteins. Replication occurs on virus-induced spherular invaginations of cytoplasmic membranes similar to RNA replication-linked spherules induced by many positive-sense RNA viruses. Full length negative-sense strand synthesis and subsequent positive-sense strand synthesis takes place inside the spherules. Replication proteins bind to the 30 terminal tRNA-like structure to initiate negative-strand RNA synthesis. In the spherule, the interior space of which is connected with the cytoplasm via a thin neck, viral positive-sense RNA strands are synthesized using negative-sense strand as a template. Subsequently, multiple copies of positive-sense RNA strands and sgRNAs are released into the cytoplasm through the neck. The replication proteins of virgaviruses, as those of other positive-sense RNA viruses, are tightly associated with membranes. Attempts to purify viral replicases from infected plants cells and to solubilize them resulted in drastic changes in their properties. sgRNAs are transcribed from negative-sense RNA through recognition of sub-genomic promoters by replication proteins. For some virgaviruses, comparison of the sequences of sub-genomic promoters reveals conservative consensus motifs. The membrane used to form the replication compartments varies and depends on the virus, for example, TMV, ToMV and Youcai mosaic virus (and probably other tobamiviruses) may forms active replication machinery on the vacuolar and endoplasmic

848

Virgaviruses (Virgaviridae)

Fig. 3 Diagram showing the genome organization of representative members of the seven genera in the family Virgaviridae. ORFs are color-coded according to the function of their protein products: replication proteins are in pink with the methyltransferase (M), helicase (H), and RdRp (P) domains; coat proteins are in green; ‘30K’-type movement proteins are in blue; triple gene block movement proteins are in light brown; cysteine-rich proteins are in yellow; and other ORFs are in white. Positions of ‘leaky’ stop codons are shown by black diamonds and tRNA-like structures with clover leaves. Val/Tyr/Leu/His/?/-: amino-acylation of tRNA-like structures, respectively. Brackets indicate ORFs that are missing from some strains.

reticulum (ER) membranes, whereas BSMV (a hordeivirus) uses the outer membranes of chloroplasts for replication. Evidence suggests that PMTV TGB2, a two-pass transmembrane protein, targets membranes of chloroplasts to establish the virus replication compartment in there. TRV viral factories are enriched in mitochondria-derived membranes. Tobamovirus replication proteins co-translationally bind to AC-rich region of the 5´ UTR of the genomic RNA to form so-called premembrane targeting complex. This represents the first step in tobamovirus RNA replication. Host factors involved in the tobamovirus replication were identified by forward genetic approaches and co-immunoprecipitation, and include TOM1, TOM2A, TOM2B, and ARL8 proteins. Tobamovirus replication complexes are anchored into membranes through interaction of the helicase domain of the replicase with TOM1, a seven-pass transmembrane protein. In turn, TOM2A, a four-pass transmebrane protein, interacts with TOM1. ARL8, an ADP-ribosylation factor-like small GTP-binding protein, interacts with both TOM1 and an N-terminal alpha-helix of the helicase domain of the replication proteins. The replication proteins first bind to membranes and then interact and form a complex with TOM1, TOM2A, and ARL8. Interaction with membranes and TOM1 and ARL8 proteins activates the RNA 5´ capping activity of the 126 kDa protein (a tobamovirus protein harboring methyl-transferase and helicase domains). The timing and the trigger of the activation of the tobamovirus RdRp activity and the mechanism of its template strand switch are unknown so far.

Virgaviruses (Virgaviridae)

849

When significant amount of positive-sense genomic RNA builds up in the infected cell, progeny rod-shaped virions are assembled in the cytoplasm. Mature virions accumulate in the infected cells and might be acquired by the vectors.

Infectious Cycle Infection with virgaviruses is initiated when a virus is delivered into the plant cell. Depending on the virus genus this is achieved either by rub-inoculation (mechanical transmission) – the mode of virus transmission possible for most virgaviruses and the only way of transmission for the viruses of the genus Tobamovirus – or by the vector (plasmodiophorid or nematode). In the plant cell, ssRNA of virgaviruses gets uncoated from virions and is translated to produce replication-associated proteins. The replication complex then initiates rearrangement and remodeling of host intracellular membranes to establish viral replication organelles or virus factories. Such compartments secure efficient RNA replication and synthesis of sgRNA, and protect viral RNA from nucleases and pattern recognition receptors that upon activation can trigger host innate immunity responses, e.g., in response to dsRNA accumulation. Later in infection, viral MPs are expressed from sgRNA (for most virgaviruses). MPs facilitate cell-to-cell and systemic movement of the virus in the infected plant. Some members of the family Virgaviridae require expression of CP for systemic movement. When sufficient amount of CP is expressed, ssRNA can then be encapsidated by CP into virions, which become available for vector acquisition and subsequent transmission. Virions of virgaviruses can aggregate forming inclusion bodies, virion arrays and other paracrystalline structures in the plant cell. An important step of the life cycle of the soil-borne viruses of the Furovirus, Pecluvirus and Pomovirus genera is the transmission between plants by a plasmodiophorid vector. The dormant resting spores of plasmodioporids carrying a virgavirus can persist in soil for more than fifteen years. Under high levels of soil moisture and cool temperatures, the resting spores germinate and release primary virus-carrying zoospores that attach to and penetrate root tissue of the host, mostly root hairs. Zoospores are responsible for transmission of virgaviruses to host plants and the viruses reside inside the zoospores. After penetration, the zoospore develops into a multinucleate plasmodium, which after growth, increasing in size and proliferating into segments forms zoosporangia, each harboring numerous secondary zoospores that can infect new roots. Although the exact mechanism of the virus acquisition by plasmodiophorids is not known, it has been speculated that it might occur at the plasmodium stage, probably, through fusion of membranes induced by CP-RTD proteins located at one extremity of virus particles. Intriguingly, CP-RTD proteins of virgaviruses have two transmembrane hydrophobic segments - the arrangement resembles membrane fusion proteins of enveloped animal viruses. Deletions in CP-RTD proteins resulting in the loss of at least one transmembrane segment prevent transmission of the virus by the vector. Although, whether virgaviruses replicate in the plasmodiophorid vector is still unknown.

Diagnosis The members of the family Virgaviridae incite various leaf symptoms including yellow mosaic, ring spots, stripes, streaks, necrotic spots, leaf malformations etc. However, identification of the virgaviruses based these symptoms might be misleading because similar symptoms may be induced by other viruses. Thus, diagnosis based on appearance of symptoms is not sufficient and specific diagnostic tests must be employed to confirm virgavirus infection. Brown arcs and rings in the potato tuber flesh are typical symptoms of ‘spraing’, a Scottish word referring to a dark, variegated streak or stripe. ‘Spraing’ can be caused by PMTV (genus Pomovirus) or TRV (genus Tobravirus). Reliable methods are needed to determine whether symptoms are caused by PMTV or TRV. Several techniques have been developed and successfully used to detect the presence of virgaviruses, including serological assays, nucleic-acid-based methodologies, and next generation sequencing (NGS). A number of serological diagnostic techniques such as an enzyme-linked immunosorbent assay (ELISA) have been developed for fast, simple, cost-effective and high-throughput testing. Serological reagents are available for many viruses in the family Virgaviridae, including reagents for ELISA (e.g., for PMTV, TMV, ToMV, TRV etc.) and strips for lateral flow detection (e.g., for detection of TMV and ToMV). Lateral flow assays provide very rapid diagnosis within minutes, are easily portable and highly robust, and have been adopted by agronomists and farm advisors as means of quickly confirming the presence of a specific virgavirus within a crop in the field. Nucleic acid-based diagnostic methods have become increasingly popular for detection and characterization of virgaviruses. Such techniques include various modifications of reverse transcription (RT) polymerase chain reaction (PCR): classical RT-PCR, immunocapture-RT-PCR, quantitative real-time RT-PCR (RT-qPCR), loop-mediated isothermal amplification, and oligonucleotide microarrays. These assays are often more robust and sensitive than serological assays. Recently, NGS, including RNA-seq and sequencing of small RNAs, has become increasingly accessible for discovery of new viruses, including virgaviruses. These holistic approaches provide an unbiased identification and characterization of a whole virome in a sample without any prior knowledge of the pathogens present.

Pathogenicity Despite significant advances in studies of the mechanisms of pathogenicity (the ability of a virus to cause disease on a particular host), the information about how virgaviruses cause disease in their host is surprisingly scant. Pathogenicity determinants include

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CRPs encoded by the viruses of the Furovirus, Goravirus, Hordeivirus, Pecluvirus, Pomovirus, and Tobravirus genera. For example, natural variants of CRPs from various BSMV strains have a number of specific effects on the virus pathogenesis including the ability of certain strains to establish systemic infection in the absence of the CRP expression. However, the failure of certain strains to establish systemic infection can be reversed by mutations (deletions of N-terminal duplication of the polymerase domain) that increase the abundance of viral polymerase, and, thus, increase the virus replication rate. The function of CP in pathogenicity and symptom modulation was shown for pomo- and tobamoviruses. PMTV causes symptomless infections in the absence of the CP expression. On the contrary, TMV mutants defective in CP expression still display mild mosaic symptoms. However, the appearance of bright chlorosis symptoms is linked to specific aa substitutions within the TMV CP and results from disruption of chloroplast development and homeostasis. TMV CP associates with and disrupts photosystem II complexes in the thylakoid membranes of chloroplasts.

Epidemiology and Control The viruses in the genera Furovirus, Pecluvirus and Pomovirus are transmitted in soil by plasmodiophorid vectors (Table 2). Polymyxa graminis and Spongospora subterranea are known vectors of the viruses in these three genera. Viruses transmitted by plasmodiophorids reside inside the zoospores or dormant resting spores of their vector and, thus, well protected from the environment. The virus-carrying resting spores also remain viruliferous for many years. These features make eradication of the diseases caused by plasmodiophorid-transmitted viruses very difficult, and infestations are usually permanent, ones the virus is introduced into the field. The viruses in the genus Tobravirus are transmitted in soil by nematode vectors. In general, management of nematodes is difficult. The existing infestations can be reduced through fallowing, crop rotation, and soil solarization. Transmission through seed and pollen has been documented for the viruses in the genera Goravirus and Hordeivirus. No vector is know for viruses in the genus Tobamovirus. Tobamoviruses are readily transmittable by mechanical inoculation. Most of the viruses in the family Virgaviridae are mechanically transmittable in lab conditions. The family Virgaviridae includes viruses (BSMV, PCV, PMTV, TMV, ToMV, TRV etc.) that are responsible for significant quantitative and qualitative yield losses in various crops, whereas the economic significance of other family members is limited, not well studied or not important. Many factors affect the occurrence and incidence of diseases caused by virgaviruses. Application of integrated pest management (IPM) strategies would be beneficial to control of virgavirus-induced diseases as IPM takes into account all those factors. For plasmodiophorid- and nematode-transmitted viruses disease management is mainly preventive through the use of virus-free seed material (e.g., virus-free seed tubers) and non-infested fields. Cross‐protection with mild viral strains has been used successfully to control some virus diseases caused by tobamoviruses (e.g., TMV and ToMV). Mild strain protection of tomato against ToMV is probably one of the most successful applications of cross-protection. It is successfully and routinely used in Europe, Canada, New Zealand, Israel, and Japan. One of the best ways to manage virgaviruses is to use varieties that are resistant to the viruses. Four dominant genes conferring resistance to virgaviruses have been cloned and sequenced. Three of these genes encode members of the nucleotide-binding site–leucine-rich repeat (NBS-LRR) class of R proteins, whereas Tm-1 gene encodes a TIM barrel structure protein. The R proteins localize to the cytoplasm consistent with the lifecycle of virgaviruses. ToMV was once one of the most important pathogens of tomato crop, but effective control has been established using resistant cultivars with three ToMV resistance genes: Tm-2/Tm-22 and Tm-1. N-gene-mediated resistance to TMV is another example of solving a practical problem and putting disease resistance in the field.

Unassigned Virgaviruses Recent advances in low-cost high-throughput NGS led to identification of a number of virga-like viruses in metagenomics data generated for invertebrates, fungi and oomycetes (Table 3). Most of these viruses seem to be monopartite with genomes ranging 6.8–11.9 kb. However, one cannot exclude the possibility that some of the viruses in this group might have multipartite genomes as currently only partial sequences are available.

Further Reading Adams, M.J., Adkins, S., Bragard, C., et al., 2017. ICTV virus taxonomy profile: Virgaviridae. Journal of General Virology 98, 1999–2000. Chujo, T., Ishibashi, K., Miyashita, S., Ishikawa, M., 2015. Functions of the 50 - and 30 -untranslated regions of tobamovirus RNA. Virus Research 206, 82–89. Gibbs, A.J., Wood, J., Garcia-Arenal, F., Ohshima, K., Armstrong, J.S., 2015. Tobamoviruses have probably co-diverged with their eudicotyledonous hosts for at least 110 million years. Virus evolution 1, vev019. Gil, J.F., Adams, I., Boonham, N., Nielsen, S.L., Nicolaisen, M., 2016. Molecular and biological characterisation of two novel pomo-like viruses associated with potato (Solanum tuberosum) fields in Colombia. Archives of Virology 161, 1601–1610. Ishibashi, K., Ishikawa, M., 2016. Replication of tobamovirus RNA. Annual Review of Phytopathology 54, 55–78. Jackson, A.O., Lim, H.S., Bragg, J., Ganesan, U., Lee, M.Y., 2009. Hordeivirus replication, movement, and pathogenesis. Annual Review of Phytopathology 47, 385–422.

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Kalyandurg, P., Gil, J.F., Lukhovitskaya, N.I., et al., 2017. Molecular and pathobiological characterization of 61 Potato mop-top virus full-length cDNAs reveals great variability of the virus in the centre of potato domestication, novel genotypes and evidence for recombination. Molecular Plant Pathology 18, 864–877. Koonin, E.V., Dolja, V.V., 1993. Evolution and taxonomy of positive-strand RNA viruses. Crit Rev Biochem Mol Bio 28, 375–430. Lecoq, H., 1998. Control of plant virus diseases by cross-protection. In: Hadidi, A., Khetarpal, R.K., Koganezawa, H. (Eds.), Plant Virus Disease Control. St. Paul, MN: APS Press, pp. 33–40. Melcher, U., 2000. The ‘30K’ superfamily of viral movement proteins. Journal of General Virology 81, 257–266. Rochon, D.’.A., Kakani, K., Robbins, M., Reade, R., 2004. Molecular aspects of plant virus transmission by olpidium and plasmodiophorid vectors. Annual Review of Phytopathology 42, 211–241. Solovyev, A.G., Makarov, V.V., 2016. Helical capsids of plant viruses: Architecture with structural lability. Journal of General Virology 97, 1739–1754. Solovyev, A.G., Savenkov, E.I., 2014. Factors involved in systemic transport of plant RNA viruses, the emerging role of the nucleus. Journal of Experimental Botany 65, 1689–1697. Zhang, K., Zhang, Y., Yang, M., et al., 2017. The Barley stripe mosaic virus gammab protein promotes chloroplast-targeted replication by enhancing unwinding of RNA duplexes. PLoS Pathogens 13, e1006319.

Relevant Websites https://talk.ictvonline.org/ictv-reports/ictv_online_report/positive-sense-rna-viruses/w/virgaviridae Virgaviridae-Positive-sense RNA Viruses-ICTV. https://talk.ictvonline.org/taxonomy/ Taxonomy-International Committee on Taxonomy of Viruses.

Viroids (Pospiviroidae and Avsunviroidae) Ricardo Flores, Polytechnic University of Valencia, Higher Council of Scientific Research, Valencia, Spain Robert A Owens, Beltsville Agricultural Research Center, Beltsville, MD 20705, United States r 2021 Elsevier Ltd. All rights reserved.

Glossary Catalytic RNA RNA molecules that are able to catalyze, in a protein-free medium, specific reactions involving the formation or breakage of covalent bonds. In nature, these reactions are usually transesterifications (self-cleavage and ligation) affecting the catalytic RNA itself. Hammerhead structure The conserved secondary/tertiary structure shared by the smallest class of natural ribozymes. Most were initially found in one or both strands of certain

viroid and viroid-like satellite RNAs where they mediate self-cleavage of multimeric intermediates arising from replication through a rolling-circle mechanism. More recently, the hammerhead ribozyme motif has been reported along the tree of life. Ribozyme RNA motif responsible for the catalytic activity of certain RNA molecules. In nature, they are found embedded within catalytic RNAs.

Taxonomy According with the presence of conserved sequence/structure motifs, and mode of replication and subcellular accumulation sites, viroids are allocated to the family Pospiviroidae (composed of genera Pospiviroid, Hostuviroid, Cocadviroid, Apsacaviroid and Coleviroid), type species Potato spindle tuber viroid, and family Avsunviroidae (composed of genera Avsunviroid, Pelamoviroid and Elaviroid), type species Avocado sublotch viroid.

Introduction Viroids are the smallest known agents of infectious disease – small (246–401 nt), highly structured, single-stranded, circular RNAs – that lack detectable messenger RNA activity. Whereas viruses have been described as “obligate parasites of the cell's translational system” and supply some or most of the genetic information required for their replication and movement, viroids can be regarded as “obligate parasites of the cell's transcriptional machinery”. Thus far, viroids are known to infect only plants. The first viroid disease to be studied by plant pathologists was potato spindle tuber. In 1923, its infectious nature and ability to spread in the field led Schultz and Folsom to group potato spindle tuber disease with several other “degeneration diseases” of potatoes. Nearly 50 years were to elapse before Diener's 1971 demonstration that the molecular properties of its causal agent, Potato spindle tuber viroid (PSTVd), were fundamentally different than those of conventional plant viruses.

Genome Structure Efforts to understand how viroids replicate and cause disease without the assistance of any viroid-encoded polypeptides has prompted detailed analysis of their structure. Viroids possess rather unusual properties for single-stranded RNAs (e.g., a pronounced resistance to digestion by ribonucleases and a highly cooperative thermal denaturation profile), leading to an early realization that they might have an unusual higher-order structure. To date, the complete sequences of 32 distinct viroid species plus a large number of sequence variants have been determined (Table 1). All are single-stranded circular RNAs with unmodified nucleotides. Theoretical calculations and physicochemical studies indicate that PSTVd and related viroids assume a highly base-paired, rod-like conformation in vitro (Fig. 1), with recent data supporting a similar secondary structure in vivo for PSTVd and Avocado sunblotch viroid (ASBVd), which appear to accumulate in planta as free RNAs unprotected by tightly bound proteins. Pair-wise sequence comparisons suggest that the series of short double helices and small internal loops that comprise this so-called “native” structure are organized into five domains whose boundaries are defined by sharp changes in sequence identity. The “central domain” is the most highly conserved viroid domain and contains the site where multimeric PSTVd RNAs are cleaved and ligated to form circular progeny. The “pathogenicity domain” contains one or more structural elements that modulate symptom expression, and the relatively small “variable domain” exhibits the greatest sequence variability between otherwise closely-related viroids. The two “terminal domains” appear to play an important role in viroid replication and evolution. Although these five domains were first identified in PSTVd, Apple scar skin viroid (ASSVd) and related viroids also contain a similar domain arrangement. Certain viroids, such as Columnea latent viroid (CLVd), Australian grapevine viroid (AGVd) and Coleus blumei viroid 2 (CbVd 2), appear to be “mosaic molecules” formed by exchange of domains between two or more viroids infecting the same cell. RNA rearrangement/

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Viroids (Pospiviroidae and Avsunviroidae) Table 1

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Classification of viroids of known nucleotide sequence

a

Family

Pospiviroidae

Genusa

Name

Abbreviation

Nucleotidesb

Pospiviroid

Chrysanthemum stunt viroid Citrus exocortis viroid Columnea latent viroid Iresine viroid Pepper chat fruit viroid Potato spindle tuber viroid Tomato apical stunt viroid Tomato chlorotic dwarf viroid Tomato planta macho viroid Citrus bark cracking viroid Coconut cadang-cadang viroid Coconut tinangaja viroid Hop latent viroid Dahlia latent viroid Hop stunt viroidc Apple dimple fruit viroid Apple scar skin viroidd Australian grapevine viroid Citrus bent leaf viroid Citrus dwarfing viroid Citrus viroid V Citrus viroid VI Grapevine yellow speckle viroid 1 Grapevine yellow speckle viroid 2e Pear blister canker viroid Coleus blumei viroid 1 Coleus blumei viroid 2 Coleus blumei viroid 3

CSVd CEVd CLVd IrVd PCFVd PSTVd TASVd TCDVd TPMVd CBCVd CCCVd CtiVd HLVd DLVd HSVd ADFVd ASSVd AGVd CBLVd CDVd CVd-V CVd-VI GVYSVd 1 GYSVd 2 PBCVd CbVd 1 CbVd 2 CbVd 3

354–356 368–375 370–373 370 348 356–361 360–363 360 359–360 282–286 246–247 254 256 342 294–303 306,307 329–334 369 315,318 294,297 284 330 366–368 363 315,316 248–251 301,302 361–364

Avocado sunblotch viroid Chrysanthemum chlorotic mottle viroid Peach latent mosaic viroid Eggplant latent viroid

ASBVd CchMVd PLMVd ELVd

246–251 398–401 335–339 (348–351) 332–335

Cocadviroid

Hostuviroid Apscaviroid

Coleviroid

Avsunviroidae Avsunviroid Pelamoviroid Elaviroid

(463–467)

(341)

(273–277) (287–301)

a Classification follows scheme proposed by Owens et al. 2012 (see Virus Taxonomy, IX Report of the International Committee on Taxonomy of Viruses) with minor modifications. Whether Apple fruit crinkle viroid, Apple hammerhead viroid, Coleus blumei viroids 4, 5 and 6, Grapevine hammerhead viroid-like RNA, Grapevine latent viroid, Grapevine yellow speckle viroid 3, Persimmon latent viroid, Persimmon viroid 2, and Portulaca latent viroid should be considered variants or new viroid species of a different genus is pending. b Sizes of variants containing insertions or deletions arising in vivo are shown in parentheses. c Includes Cucumber pale fruit viroid, Citrus cachexia viroid, Peach dapple viroid, and Plum dapple viroid. d Includes Pear rusty skin viroid and Dapple apple viroid. e Formerly termed Grapevine viroid 1B.

recombination can also occur within individual domains, leading, in Coconut cadang-cadang viroid (CCCVd) and Citrus exocortis viroid (CEVd), to duplications of the right terminal domain plus part of the variable domain. This domain model is not shared by ASBVd and related viroids. Much less is known about the tertiary structure of viroids, especially in vivo. UV-induced cross-linking of two nucleotides within a loop E motif in the central domain of PSTVd provided the first definitive evidence for such tertiary interactions. Similar UV-sensitive structural elements have also been discovered in a number of other RNAs including 5S eukaryotic rRNA, adenovirus VAI RNA, and the viroid-like domain of the hepatitis delta virus genome. Loop E forms during the conversion of multimeric PSTVd RNAs into monomers. The ability of ASBVd-related RNAs to undergo spontaneous selfcleavage mediated by hammerhead ribozymes, as well as the presence of kissing-loop interactions critical for infectivity in some other members of the Avsunviroidae (Fig. 1), provide additional evidence for the functional importance of viroid tertiary structure.

Classification Based upon differences in their structural and functional properties, viroids are assigned to one of two taxonomic families (Table 1). Members of the family Pospiviroidae (type species PSTVd) fold into a rod-like secondary structure with five structural-

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Fig. 1 (a) The rod-like secondary structure of PSTVd (intermediate strain) showing the five domains characteristic of members of the family Pospiviroidae: the Terminal Left (TL), Pathogenicity (P), Central (C), Variable (V), and Terminal Right (TR). The Central Conserved Region (CCR) is located within the C domain and contains a UV-sensitive loop E motif with non-canonical base-pairs (denoted by circles). The TL domains of genera Pospi- and Apscaviroid contain a Terminal Conserved Region (TCR), while those of genera Hostu- and Cocadviroid contain a Terminal Conserved Hairpin (not shown). The TR may also contain 1–2 copies of a protein-binding RY motif. (b) The branched secondary structure of PLMVd (reference variant). Plus and minus self-cleavage domains are indicated by flags, nucleotides conserved in most natural hammerhead structures by bars, and the self-cleavage sites by arrows. Black and white symbols refer to plus and minus polarities, respectively. Nucleotides involved in a kissing-loop interaction are indicated by broken lines. Redrawn with modifications from Gross, H.J., Domdey, H., Lossow, C., et al., 1978. Nucleotide sequence and secondary structure of potato spindle tuber viroid. Nature 273, 203–208. Hernández, C., Flores, R., 1992. Plus and minus RNAs of peach latent mosaic viroid self-cleave in vitro via hammerhead structures. Proceedings of the National Academy of Sciences of the United States of America 89, 3711–3715. Bussière, F., Ouellet, J., Côté, F., Lévesque, D., Perreault, J.P., 2000. Mapping in solution shows the peach latent mosaic viroid to possess a new pseudoknot in a complex, branched secondary structure. Journal of Virology 74, 2647–2654.

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functional domains and several conserved motifs. Most members of the family Avsunviroidae (type species ASBVd), in contrast, appear to adopt a branched conformation, and multimeric RNAs of all family species behave as catalytic RNAs and undergo spontaneous self-cleavage (Fig. 1). Differences in their sites of replication also support this classification scheme; i.e., PSTVd and ASBVd replicate in the nucleus and plastids (mostly chloroplasts), respectively, and the same appears to occur for other family species. Each family is subdivided into genera according to certain demarcating criteria. Groups of sequence variants that show 490% sequence identity in pair-wise comparisons and share some common biological property are arbitrarily defined as viroid species. In vivo, each viroid species is actually a “quasi-species”; i.e., a collection of closely-related sequence variants subject to a continuous process of mutation, competition, and selection. Phylogenetic evidence supports an evolutionary link between viroids and other viroid-like subviral RNAs (Fig. 2).

Host Range and Transmission All viroids are mechanically transmissible, and most are naturally transmitted from plant to plant by man and his tools. Individual viroids vary greatly in their ability to infect different plant species. PSTVd can replicate in about 160 primarily solanaceous hosts, while only two members of the family Lauraceae are known to support ASBVd replication. Hop stunt viroid (HSVd) has a particularly wide host range that includes herbaceous species as well as woody perennials. Many natural hosts are either vegetatively propagated or crops that are subjected to repeated grafting or pruning operations. PSTVd, ASBVd, and Coleus blumei viroid 1 (CbVd 1) are vertically transmitted through pollen and/or true seed, but the significance of this mode of transmission in their natural spread is unclear. PSTVd can be encapsidated by the coat protein of Potato leafroll virus (PLRV, a luteovirus) as well as Velvet tobacco mottle virus (VTMoV, a sobemovirus), and epidemiological surveys suggest that PLRV facilitates viroid spread under field conditions. Commonly used techniques for the experimental transmission of viroids include the standard leaf abrasion methods developed for conventional viruses, “razor slashing” methods in which phloem tissue in the stem or petiole is inoculated via cuts made with a razor blade previously dipped into the inoculum, and, in the case of CCCVd, high-pressure injection into folded apical leaves. Viroids can also be transmitted by either plant transformation or “agro-inoculation” during which a modified Agrobacterium tumefaciens Ti plasmid is used to introduce full-length viroid-complementary DNA into the potential host cell. Either technique can overcome the marked resistance of some hosts to mechanical inoculation. Identification of the molecular mechanism(s) that determine viroid host range remains an important research goal.

Symptomatology Viroids and conventional plant viruses induce a very similar range of macroscopic symptoms. Symptom expression is usually optimal at the same relatively high temperatures (30–331C) that promote viroid replication/accumulation. Stunting and leaf epinasty (a downward curling of the leaf lamina resulting from unbalanced growth within the various cell layers) are considered the classic symptoms of viroid infection. Other commonly observed symptoms include vein clearing, veinal discoloration or necrosis, and the appearance of localized chlorotic/necrotic spots or mottling in the foliage. Symptoms may also be expressed in flowers and bark, and fruits or tubers from viroid-infected plants may be abnormally shaped or discolored. Viroid infection of certain citrus rootstock/scion combinations may result in tree dwarfing (Fig. 3). Viroid infections are often latent and rarely kill the host. Viroid infections are also accompanied by a number of cytopathic effects – chloroplast and cell wall abnormalities, the formation of membranous structures in the cytoplasm, and the accumulation of electron-dense deposits in both chloroplasts and cytoplasm. Metabolic changes include dramatic alterations in growth regulator levels.

Geographic Distribution Although PSTVd, CEVd, HSVd and ASBVd are widely distributed throughout the world, other viroids have never been detected outside the areas where they were first reported. Several factors may contribute to this variation in distribution pattern. Among the crops most affected by viroid diseases are a number of valuable woody perennials such as grapes, citrus, various pome and stone fruits, and hops. Propagation and distribution of improved cultivars is highly commercialized, with the result that many cultivars are now grown worldwide. The international exchange of plant germplasm also continues to increase at a rapid rate. In both instances, the large number of latent (asymptomatic) hosts facilitates viroid spread.

Epidemiology and Control Viroid diseases represent a potential threat to agriculture, and several are of considerable economic importance. Ready transmission of PSTVd by vegetative propagation, foliar contact, and true seed or pollen continues to pose a serious threat to potato germplasm and breeding programs. Coconut cadang-cadang has killed over 30 million palm trees in the Philippines since it was first recognized in the early 1930s. While many viroids were first detected in ornamental or crop plants, most viroid diseases are

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Fig. 2 Neighbor-joining phylogenetic tree obtained from an alignment manually adjusted to take into account local similarities, insertions/deletions, and duplications/rearrangements described in the literature for viroid and viroid-like satellite RNAs. Bootstrap values were based on 1000 random replicates (only values 4 70% are reported). Viroid abbreviations as in Table 1. Viroid-like satellite RNAs: sLTSV (lucerne transient streak virus); sRYMV (rice yellow mottle virus); sSCMoV (subterranean clover mottle virus); sSNMoV (Solanum nodiflorum mottle virus); sVTMoV (velvet tobacco mottle virus); sTRSV (tobacco ringspot virus); sArMV (Arabis mosaic virus); sChYMV (chicory yellow mottle virus); sCYDV-RPV (cereal yellow dwarf virus-RPV). Adapted from Elena, S.F., Dopazo, J., de la Peña, M., et al., 2001. Phylogenetic analysis of viroid and viroid-like satellite RNAs from plants: A reassessment. Journal of Molecular Evolution 53, 155–159, with permission.

thought to result from chance transfer from endemically-infected wild species to susceptible cultivars (Diener, 1979). Several lines of circumstantial evidence are consistent with this hypothesis: (1) The experimental host ranges of several viroids include many wild species, and these wild species often tolerate viroid replication without the appearance of recognizable disease symptoms. (2) Although co-evolution of host and pathogen is often accompanied by appearance of gene-for-gene vertical resistance, no useful sources of resistance to PSTVd infection have been identified in the cultivated potato. (3) Viroids and/or viroid-related RNAs closely related to Tomato planta macho viroid (TPMVd) and CCCVd have been detected in weeds and other wild vegetation growing near fields containing viroid-infected plants.

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Fig. 3 (a) Symptoms of PSTVd infection in Rutgers tomato approximately four weeks after inoculation of cotyledons with PSTVd strains causing mild, intermediate, and severe symptoms. (b) Symptoms of ASBVd infection in avocado fruits and leaves. (c) CDVd-induced dwarfing of citrus growing on susceptible rootstocks. All trees in the block were graft-inoculated with CDVd shortly after transfer to the field; only one tree (right foreground) escaped infection. Note difference in height.

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Viroids (Pospiviroidae and Avsunviroidae)

Growers and plant pathologists are unlikely to have simply overlooked diseases with symptoms as severe as those of chrysanthemum stunt or cucumber pale fruit, two diseases first reported after World War II. Large-scale monoculture of genetically identical crops and the commercial propagation/distribution of many cultivars are two comparatively modern developments which would facilitate the development of serious disease problems following the chance transfer of viroids from wild hosts to cultivated plants. Viroid diseases may also arise by transfer between cultivated crop species. For example, pears provide a latent reservoir for ASSVd; likewise, while HSVd infections of grapes are often symptomless, this viroid causes severe disease in hops. In both instances, the two crops are often grown in close proximity. In other instances, however, such proximity does not apparently exist, as in a hop disease induced by Citrus bark cracking viroid (CBCVd) recently reported in Slovenia, where citrus is not grown. Because no useful sources of natural resistance to viroid diseases are known, diagnostic tests continue to play a key role in efforts to control these diseases. Since viroids lack a protein capsid, the antibody-based techniques used to detect many plant viruses are not applicable. Tests based upon their unique molecular properties have largely supplanted biological assays, which require extended periods (weeks to years) of time for completion and have difficulties in detecting mild or latent strains. Bioassays remain indispensable for assessing the autonomous replication of a candidate viroid-like RNA and, eventually, for fulfilling Koch’s postulates. Several rapid (1–2 day) protocols involving polyacrylamide gel electrophoresis under denaturing conditions take advantage of the circular nature of viroids. Using these protocols, nanogram amounts of viroid can be unambiguously detected without the use of radioactive isotopes. In recent years, diagnostic procedures based upon nucleic acid hybridization or the polymerase chain reaction (PCR) have become widely used. The simplest methods involve the hybridization of a nonradioactively labeled viroid-complementary DNA or RNA probe to viroid samples that have been bound to a solid support followed by colorimetric or chemiluminescent detection of the resulting DNA-RNA or RNA-RNA hybrids. Such conventional “dot blot” assays can detect picogram amounts of viroids using clarified plant sap or tissue prints rather than purified nucleic acid as the viroid source. PCR-based protocols are finding increasing acceptance in those cases where either this level of sensitivity is not reached or a number of closely-related viroids are present in the same sample.

Molecular Biology Although devoid of messenger RNA activity, viroids replicate autonomously and cause disease in a wide variety of plants. Much has been learned about the molecular biology of viroids and viroid-host interaction over the past 40 years, but the precise nature of the molecular signals involved remains elusive. A series of questions first posed by Diener summarizes the many gaps in our current understanding of the biological properties of these unusual molecules: (1) What molecular signals do viroids possess (and cellular RNAs evidently lack) that induce certain DNA-dependent RNA polymerases to accept them as templates for the synthesis of complementary RNA molecules? (2) What are the molecular mechanisms responsible for viroid replication? Are these mechanisms operative in uninfected cells? If so, what are their functions? (3) How do viroids induce disease? In the absence of viroid-specified proteins, disease must arise from direct interaction(s) of viroids (or viroid-derived RNA molecules) with host cell constituents. Infections by PSTVd or ASBVd induces RNA silencing (see below), but does this mechanism mediate pathogenesis for all viroids? (4) What determines viroid host range? Are viroids restricted to higher plants, or do they have counterparts in animals? (5) How did viroids originate?

Replication A variety of multimeric plus and minus strand RNAs have been detected by nucleic acid hybridization in viroid-infected tissues. Based on their analysis, viroid replication has been proposed to proceed via a “rolling circle” mechanism that involves reiterative transcription of the incoming plus circular RNA to produce a minus strand RNA template. ASBVd and related viroids utilize a symmetric replication cycle in which the multimeric minus strand is cleaved to unit-length molecules and circularized before serving as template for the synthesis of multimeric plus strands. PSTVd and related viroids utilize an asymmetric cycle in which the multimeric minus strand is directly transcribed into multimeric plus strands. In both cases, the multimeric plus strands are cleaved to unit-length molecules and circularized (Fig. 4). A diversity of host-encoded enzymes have been implicated in viroid replication. Low concentrations of a-amanitin specifically inhibit the synthesis of both PSTVd plus and minus strands in nuclei isolated from infected tomato, strongly suggesting the involvement of DNA-dependent RNA polymerase II, transcribing an RNA template, in the replication of PSTVd and related viroids. In nuclear extracts, synthesis of the PSTVd minus strand by RNA polymerase II starts in the left terminal loop of the plus strand template; furthermore, incubation of active replication complexes containing CEVd with a monoclonal antibody directed against the carboxy-terminal domain of RNA polymerase II results in the immunoprecipitation of both CEVd plus and minus strand RNAs. Mature PSTVd plus strands accumulate in the nucleolus and the nucleoplasm, while in situ hybridization indicates that minus strand RNAs are confined to the nucleoplasm. After some debate, the identity of the polymerase(s) responsible for replication of members of the family Avsunviroidae in the chloroplast appears now more certain. ASBVd synthesis is resistant to

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Fig. 4 Asymmetric and symmetric variants of the rolling-circle mechanism proposed for replication of members of the families Pospiviroidae and Avsunviroidae, respectively. Orange and blue colors refer to plus and minus polarities, respectively, with cleavage sites denoted by arrowheads. The enzymes and ribozymes that presumably catalyze the replication steps are indicated. Notice that RNA polymerase II (and NEP) is redirected to transcribe RNA templates, and DNA ligase 1 to circularize RNA substrates. Abbreviations: HHRz, hammerhead ribozyme; NEP, nuclear-encoded polymerase. Adapted from Flores, R., Gago-Zachert, S., Serra, P., Sanjuán, R., Elena, S.F., 2014. Viroids: survivors from the RNA world? Annual Review of Microbiology 68, 395–414, with permission.

tagetitoxin, strongly indicating the involvement of a nuclear-encoded chloroplastic DNA-dependent RNA polymerase (NEP) again transcribing an RNA template. Initiation sites for both ASBVd plus and minus strand synthesis have been mapped to the AU-rich terminal loops of their respective native structures. Furthermore, Peach latent mosaic viroid (PLMVd) replicates actively in albino leaf areas (where only NEP is active), with the initiation sites of plus and minus strands mapping at a short double-stranded RNA motif that also contains the self-cleavage sites of both polarity strands. In vitro and in vivo evidence indicates that specific cleavage of multimeric plus strand RNAs of PSTVd and related viroids requires: (1) rearrangement of the conserved central region, and (2) the action of one or more host-encoded nucleases. Other less efficient processing sites may also be used in vivo. Multimeric plus and minus strand RNAs of ASBVd and related viroids, in contrast, undergo spontaneous self-cleavage co-transcriptionally through hammerhead ribozymes to form linear monomers. Addition of certain chloroplast proteins acting as RNA chaperones facilitates this hammerhead ribozyme-mediated self-cleavage reaction. The final step in viroid replication, the ligation of linear monomers to form mature circular progeny, is catalyzed by DNA ligase 1 (redirected to accept RNA substrates) in the case of PSTVd, and by a plastid isoform of the tRNA ligase in members of the Avsunviroidae (Fig. 4).

Movement Upon entering a potential host cell, viroids must move to either the nucleus (Pospiviroidae) or chloroplast (Avsunviroidae) before beginning replication. Available data suggest that PSTVd enters the nucleus as a ribonucleoprotein complex formed by the interaction of cellular proteins with specific viroid sequence or structural motifs. VirP1, a bromodomain-containing protein isolated from tomato, has a nuclear localization signal and binds to the terminal right domain of PSTVd. Proteins such as TFIIIA and ribosomal protein L5 that bind to the loop E motif may also be involved in viroid transport into the nucleus. How ASBVd or other members of the family Avsunviroidae enter and exit the chloroplast is currently unknown. To establish a systemic infection, viroids must leave the initially infected cell – moving first from cell-to-cell through plasmodesmata and then long distances through the host vasculature. Upon injection into symplastically isolated guard cells in a mature tomato leaf, fluorescently-labeled PSTVd RNA does not move. Injection into interconnected mesophyll cells, in contrast, is followed by rapid cell-to-cell movement through the plasmodesmata. Long distance movement of viroids, like that of nearly all plant viruses, occurs in the phloem where it follows the typical source-to-sink pattern of photoassimilate transport. Viroid movement in the phloem almost certainly requires formation of a ribonucleoprotein complex, possibly involving a dimeric lectin known as Phloem Protein 2 (PP2), the most abundant protein in phloem exudate. Movement of PSTVd in the phloem appears to be sustained by replication in supporting cells and is tightly regulated by developmental and cellular factors. For example, in situ hybridization reveals the presence of PSTVd in vascular tissues underlying the shoot apical meristem of infected tomato, but entry into the shoot apical meristem itself appears to be blocked. Other important control points for PSTVd trafficking are the bundle sheath-mesophyll boundary and that between palisade and spongy mesophyll in the leaf. By disrupting normal pattern of viroid movement it may be possible to create a plant that is resistant/immune to viroid infection.

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Pathogenicity Sequence comparisons of naturally-occurring PSTVd and CEVd variants, as well as infectivity studies with chimeric viroids constructed by exchanging the pathogenicity domains of mild and severe strains of CEVd, have clearly shown that the pathogenicity domain in the family Pospiviroidae contains important determinants of symptom expression. In addition, symptom expression is also affected by the rate of viroid accumulation, and sequence changes in the variable domain have been shown to regulate progeny titers in infected plants. Studies with Tomato apical stunt viroid (TASVd) revealed the presence of a third pathogenicity determinant in the left terminal loop. Also, a single U/A change position 257 in the central domain of PSTVd results in the appearance of severe stunting and a “flat top” phenotype in tomato, and a single base pair in the right terminal domain of TPMVd is a virulence determinant factor. In the family Avsunviroidae, determinants of pathogenicity have been mapped to either a tetraloop capping a hairpin stem in Chrysanthemum chlorotic mottle viroid (CChMVd) or an insertion that folds into a hairpin also capped by a tetraloop in PLMVd. The ability of novel viroid chimeras to replicate and move normally from cell-to-cell implies certain basic similarities between their structures in vitro and in vivo but provides no information about the nature of the molecular interactions responsible for symptom development. Until recently, it was widely assumed that the mature viroid RNA was the direct pathogenic effector. Just like viruses, however, viroid replication is also accompanied by the appearance of a variety of small (21–24 nt) RNA molecules produced by the RNA silencing defensive response of their hosts. The role of these viroid-derived small RNAs (vd-sRNAs) in viroid pathogenicity is not yet clear, but the inverse relationship between accumulation levels of the mature viroid RNAs and the corresponding vd-sRNAs for members of the family Avsunviroidae suggests that the latter may regulate the titer of the former. In addition, vd-sRNAs containing the pathogenicity determinants of specific PLMVd variants that incite albinism or an intense yellow mosaic target and direct the cleavage (as predicted by RNA silencing) of host mRNAs coding for proteins involved in chloroplast biogenesis. The corresponding PLMVd-sRNAs have 50 -terminal Us, implicating Argonaute 1 in what likely are the initial alterations eliciting distinct chloroses. Also, recovery of tomato plants from the symptoms of severe PSTVd infections is preceded by the accumulation of vd-sRNAs (Fig. 5). Viroid infections are accompanied by quantitative changes in a variety of host-encoded proteins. Certain of these are “pathogenesis-related” proteins whose synthesis or activation is part of a general host reaction to biotic or abiotic stress, but others appear to be more specific. In tobacco, PSTVd infection results in the preferential phosphorylation of a host-encoded 68 kDa protein that is immunologically related to an interferon-inducible, dsRNA-dependent mammalian protein kinase of similar size. The human kinase is differentially activated by PSTVd strains of varying pathogenicity in vitro, while infection of tomato by intermediate or severe strains of PSTVd induces the synthesis of PKV, a dual-specificity, serine/threonine protein kinase. Broad changes in host gene expression following infection by PSTVd and CEVd have been detected by complementary DNA macroarray and proteomic analyzes, respectively.

Implications for Host Range Possibly as a result of its involvement in the cleavage/ligation of progeny RNA, nucleotides in the central domain of PSTVd and related viroids appears to play an important role in determining host range. For example, a single nucleotide substitution in the

Fig. 5 A model for the role of RNA silencing in the pathogenicity and evolution of viroids and viral satellite RNAs. Replication of the subviral RNAs generates dsRNA intermediates that are processed by Dicer into 21- to 24-nt siRNAs (or vd-sRNAs), one strand of which are then incorporated into the RNA-induced silencing complex (RISC). If significant sequence identity exists between a region in the subviral RNA genome and a region in host gene mRNA (shown in red), RISC will target the host mRNA for cleavage leading to symptoms development. RISC can also target the subviral genome for degradation, forcing the subviral RNA to evolve and to adopt and maintain an RNA silencing-resistant secondary structure. From Wang, M.B., Bian, X.Y., Wu, L.M., et al., 2004. On the role of RNA silencing in the pathogenicity and evolution of viroids and viral satellites. Proceedings of the National Academy of Sciences of the United States of America 101, 3275–3280, with permission, Copyright (2004) National Academy of Sciences, USA.

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loop E motif results in a dramatic increase in the rate of PSTVd accumulation (and possibly replication) in tobacco. The biological properties of CLVd also suggest that this domain contains one or more host range determinants. CLVd appears to be a natural mosaic of sequences present in other viroids; phylogenetic analysis (Fig. 2) suggests that it can be considered to be a PSTVd-related viroid whose conserved central domain has been replaced by that of HSVd. Like HSVd (but not PSTVd or related viroids), CLVd can replicate and cause disease in cucumber.

Origin and Evolution Much of the early speculation about viroid origin involved their possible origin as “escaped introns” (i.e., descent from normal host RNAs). More recently, however, viroids have been proposed to represent “living fossils” of a precellular RNA world that assumed an intracellular mode of existence sometime after the evolution of cellular organisms. The presence of ribozymes in members of the Avsunviroidae strongly supports this view, although the recent characterization of circular long non-coding RNAs transcribed from DNA templates has rekindled the idea that viroids might have a host origin. No viroid is known to code for protein, a feature that is consistent with the possibility that viroids are phylogenetically older than introns. The inherent stability of viroids and viroid-like satellite RNAs (structurally similar to viroids but functionally dependent on helper viruses) which arises from their small size and circularity would have enhanced the probability of their survival in primitive, error-prone RNA self-replicating systems and assured their complete replication without the need for initiation/termination signals. Some viroids (but not satellite RNAs or random sequences of the same base composition) also display structural periodicities with repeat units of 12, 60, or 80 nt. The high error rate of prebiotic replication systems may have favored the evolution of polyploid genomes, and the mechanism of viroid replication (i.e., rolling-circle transcription of a circular template) provides an effective means of genome duplication. Viroids and viroid-like satellite RNAs all possess efficient mechanisms for the precise cleavage of their oligomeric replication intermediates to form monomeric progeny. PSTVd and related viroids appear to require proteinaceous host factor(s) for cleavage, but others (members of the family Avsunviroidae and viroid-like satellite RNAs) contain ribozymes far smaller and simpler than those derived from introns. Thus, ASBVd and the other self-cleaving viroids may represent an evolutionary link between viroids and viroid-like satellite RNAs. Phylogenetic evidence for an evolutionary link between viroids and other viroid-like subviral RNAs has been presented (Fig. 2). Among several subviral RNAs possibly related to viroids is carnation small viroid-like RNA, a 275 nt circular molecule with selfcleaving hammerhead structures in both its plus and minus strands that has a DNA counterpart. This retroviroid-like element shares certain features with both viroids and viroid-like satellite RNAs on the one hand, and with small RNA transcripts from newt and many other species on the other hand.

References Diener, T.O., 1979. Viroids and Viroid Diseases. New York: Wiley Interscience.

Further Reading Diener, T.O., 2003. Discovering viroids: A personal perspective. Nature Reviews Microbiology 1, 75–80. Ding, B., 2009. The biology of viroid-host interactions. Annual Review of Phytopathology 47, 105–131. Ding, B., 2010. Viroids: Self-replicating, mobile, and fast-evolving noncoding regulatory RNAs. Wiley Interdisciplinary Reviews-RNA 1, 362–375. Flores., R., Gago-Zachert, S., Serra, P., Sanjuán, R., Elena, S.F., 2014. Viroids: Survivors from the RNA world? Annual Review of Microbiology 68, 395–414. Flores, R., Hernandez, C., Martinez de Alba, A.E., Daros, J.A., Di Serio, F., 2005. Viroids and viroid-host interactions. Annual Review of Phytopathology 43, 117–139. Hadidi, A., Flores, R., Randles, J.W., Palukaitis, P. (Eds.), 2017. Viroids and Satellites. London: Academic Press. Hull, R., 2013. Plant Virology, fifth ed. New York: Academic Press. Owens, R.A., Flores, R., Di Serio, F., et al., 2011. Viroids. In: King, A.M.Q., Adams, M.J., Carstens, E.B., Lefkowitz, E.J. (Eds.), Virus Taxonomy. IX Report of the International Committee on Taxonomy of Viruses. London: Elsevier/Academic Press, pp. 1221–1234.

Watermelon Mosaic Virus and Zucchini Yellow Mosaic Virus (Potyviridae) Cécile Desbiez and Hervé Lecoq, Plant Pathology Unit, INRAE – French National Research Institute for Agriculture, Food and Environment, Montfavet, France r 2021 Elsevier Ltd. All rights reserved.

Glossary Cross-protection A plant systemically infected by a mild virus strain will not develop additional symptoms when inoculated by a severe strain of the same virus. Most often, the severe strain does not multiply in the cross-protected plant. Filiform symptom A leaf deformation symptom in which the limb is drastically reduced but not the veins, giving a shoe strings aspect to the leaf.

Transmission propensity A measure of vector importance quantifying the natural ability of a species to inoculate a plant with a virus under conditions that allow vectors to move and feed freely.

History and Taxonomy Watermelon Mosaic Virus Mosaic diseases of cucurbit crops were first reported in the 1920s and the early literature contains a diversity of names for viruses or virus diseases that were only partially characterized. In 1940, a severe watermelon (Citrullus lanatus) mosaic disease was reported in California, but the nomenclature of the causal agent remained controversial until 1979. In 1965, ten isolates from Southern USA of what was then called the watermelon mosaic virus (WMV) complex were compared. Based on cross-protection experiments, serological relationships and host range reactions they were divided into two groups: WMV1 and WMV2. WMV1 and WMV2 were considered as different viruses, also different from a watermelon mosaic isolate reported from South Africa in 1960. Further work in 1969 increased the confusion by concluding that WMV1 and WMV2 were strains of a same virus. In 1979, the situation was definitively clarified by demonstrating that WMV1 and WMV2 were indeed serologically distinct entities, and that WMV1 was closely related to Papaya ringspot virus (PRSV). Now, WMV1 is considered as the W strain of PRSV (PRSV-W), while WMV2 is referred to as Watermelon mosaic virus (WMV). In addition, the same authors showed that a watermelon mosaic virus isolate from Morocco was a third serological entity. This isolate is considered as a distinct virus species, Moroccan watermelon mosaic virus (MWMV), to which probably also belongs the South African isolate. So, from the initial watermelon mosaic virus complex emerged three different virus species: Watermelon mosaic virus, Papaya ringspot virus and Moroccan watermelon mosaic virus.

Zucchini Yellow Mosaic Virus An apparently new cucurbit virus was isolated in 1973 from a zucchini squash (Cucurbita pepo) plant in Northern Italy, this virus was described as a new potyvirus species, Zucchini yellow mosaic virus (ZYMV). In 1979, many melon (Cucumis melo) crops were devastated in Southwestern France by an apparently new virus disease, whose causal agent was tentatively named muskmelon yellow stunt virus. Very rapidly it appeared that it was a strain of ZYMV. Within a few years, ZYMV was reported in many countries on the five continents. In this regards, ZYMV appears as a typical example of an emerging plant virus.

Classification Based on particle morphology, aphid transmissibility, serological relationships, ability to induce pinwheel cytoplasmic inclusions in host cells, genome organization and nucleotide sequences, WMV and ZYMV were identified as member species of the genus Potyvirus, family Potyviridae. Potyviruses belonging to several other species can infect cucurbit crops including PRSV-W and related viruses: MWMV, Zucchini yellow fleck virus (ZYFV), Zucchini tigre mosaic virus (ZTMV), Zucchini shoestring virus (ZSTV), Algerian watermelon mosaic virus (AWMV), Sudan watermelon mosaic virus (SuWMV), Wild melon vein banding virus (WMVBV), Melon vein banding mosaic virus (MVBMV), Telfairia mosaic virus (TeMV), Cucurbita vein-banding virus (CVBV), Turnip mosaic virus (TuMV), Clover yellow vein virus (ClYVV) and Bean yellow mosaic virus (BYMV). Except for PRSV-W, MWMV and ZTMV, these viruses have either only limited geographic distribution or minor economical incidence in cucurbit crops. Molecular analyses based on the coat protein (CP) coding sequence revealed that cucurbit-infecting potyviruses belong to several “clusters” of closely related species: ZYMV, WMV and MVBMV belong to a cluster that contains mostly legume-infecting potyviruses, whereas PRSV, MWMV and 6 other viruses are grouped in a “PRSV-like” cluster containing mostly cucurbit-restricted viruses (Fig. 1).

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Fig. 1 Taxonomy, based on coat protein amino-acid sequences, of potyviruses including cucurbit-infecting ones (in bold). Branch lengths indicate the molecular divergence of viruses (the scale bar represents 0.05 mutations per residue). Figures at some nodes represent bootstrap values (in %), indicating the robustness of each node. Only values above 75% are indicated.

Symptomatology Watermelon Mosaic Virus WMV induces a diversity of symptoms depending on the isolate and the host cultivar. On leaves, symptoms are mosaics, vein banding, more or less severe leaf deformations and filiform symptom. On fruits, some isolates induce discoloration and greening on yellow fruit zucchini squash cultivars, while they will not affect fruit yield and quality in green fruit cultivars. Mosaic and discoloration are also observed on leaves and fruits of some melon cultivars (Fig. 2).

Zucchini Yellow Mosaic Virus Since its first detection, ZYMV was recognized as a virus causing extremely severe symptoms leading to complete yield losses in the case of early contamination. This severe symptomatology was probably an important factor for the rapid identification of ZYMV soon after its first outbreaks in many countries. In melon, leaf symptoms include vein clearing, yellow mosaic, leaf deformation occasionally with blisters and enations, often with severe plant stunting. Some ZYMV isolates induce a rapid and complete wilt in

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Fig. 2 Mosaic symptoms on a leaf and fruit of a melon plant infected by a moderately severe WMV isolate.

Fig. 3 Severe mosaic and deformations on leaves and fruits of a zucchini squash plant infected by ZYMV.

cultivars possessing the Fn gene. On fruits, a diversity of symptoms is observed: external mosaic or necrotic cracks, internal marbling and hardening of the flesh. Seeds are occasionally severely deformed and have poor germination rates. In zucchini squash, symptoms are very severe on leaves with mosaic, yellowing, leaf distortion and sometimes very severe filiform symptom. Fruits are generally severely misshaped with prominent knobs and are of course unmarketable (Fig. 3). In cucumber (Cucumis sativus) and watermelon, mosaic and deformations are generally observed on leaves and fruits.

Synergism and Antagonism Synergism has been observed between WMV or ZYMV and other cucurbit-infecting viruses. This synergism can be expressed either by increase in virus multiplication rates or by more severe symptoms. Significant increase in Cucumber mosaic virus (CMV) multiplication rate was observed in cucumber plants co-infected by ZYMV. CMV could also partially overcome a resistance in cucumber when in mixed infection with ZYMV. In grafted cucumbers, double infection by CMV and ZYMV induces a severe and rapid wilting reaction, not observed in single infection by CMV or ZYMV. ZYMV was shown to facilitate the entry of CMV in cucumber xylem in co-infection. When WMV or ZYMV are in mixed infection with the polerovirus Cucurbit aphid-borne yellows virus (CABYV), CABYV multiplication rate and symptom intensity are increased. As for other potyviruses, these synergetic effects could be related to the strong silencing suppressor activity of WMV and ZYMV HC-Pro. In mixed infections of ZYMV and WMV, the multiplication rate of ZYMV is not affected whereas WMV accumulation is significantly reduced, but the virus is still readily transmitted by aphids. In field conditions, early epidemics of ZYMV seem to hinder WMV spread. Infection of ZYMV limits establishment and severity of powdery mildew infection in wild squash populations, as well as the incidence of bacterial wilt caused by Erwinia tracheiphila. The lower bacterial infection appears related to a reduction of floral volatile emissions in ZYMV-infected plants, resulting in a lower attractiveness for the cucumber beetles that transmit the bacteria.

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Geographic Distribution Both WMV and ZYMV are now widely distributed in the major cucurbit production areas worldwide. Their geographic distributions are broadly overlapping and frequent mixed infections are observed in the fields. For unknown reasons, WMV appear to be rare or absent in cucurbits in subtropical or tropical areas. For instance, in exhaustive surveys conducted in Nepal, Sudan and French Western Indies no WMV was detected, although ZYMV was relatively abundant. In Florida, WMV is frequent in Northern and Central counties but is not detected in Southern counties while ZYMV can be found throughout the state. This cannot be related to a lack of potential WMV vectors or reservoirs because both are abundant in tropical or subtropical areas. ZYMV is present worldwide in almost all countries where cucurbits are grown, under temperate, Mediterranean, subtropical and tropical climatic conditions. It affects highly mechanized cropping systems – such a glasshouse crop production in Northern Europe – as well as more traditional agroecosystems – such as flood irrigated crops on the Nile banks. ZYMV has been reported in very remote areas including semi-desertic regions or islands.

Host Range WMV has a relatively wide experimental host range for a potyvirus. It infects over 170 species in 26 mono- or dicotyledonous families. Besides cucurbits, WMV causes mosaic diseases in legumes (pea, bean) and orchids (vanilla, Habenaria radiata). It has been described on many cultivated and ornamental hosts, including Panax ginseng and Lagerstroemia indica. It also infects many weeds, either annual or perennial, that can serve as alternative hosts. Generally, naturally infected weeds do not present evident symptoms of viral infection. ZYMV has a relatively narrow host range. In natural conditions, it infects mostly cultivated or wild cucurbits but also a few flower species (Consolida ajacis, Althea rosea, Begonia sempervirens) or weeds.

Diagnostic Methods The confusion that was prevalent in the early descriptions of watermelon mosaic diseases was mainly due to the convergence of symptoms caused by WMV, PRSV and MWMV in cucurbits and to the lack of proper diagnostic tools. Symptomatology and virus particle morphology were clearly insufficient to differentiate these viruses. The production of specific polyclonal antisera, and the development of simple serological tests, such as the gel double-diffusion test in agar containing sodium dodecyl sulphate and sodium azide (SDS-ID), brought a major contribution to the proper diagnosis of cucurbit potyviruses. In particular, this method contributed to the rapid and unequivocal identification of ZYMV in several countries soon after its first observation. Now, DAS-ELISA is generalized and commercial kits are available for WMV and ZYMV. Commercial dip-stick serological tests based on the lateral flow technique have also been developed that allow an easy and rapid diagnosis of ZYMV in the fields. Monoclonal antibodies (MAbs) have been produced for ZYMV and WMV. They proved to be very useful to study the serological variability and to differentiate ZYMV and WMV subgroups. They have also been used successfully to analyze virus interactions, and in particular cross protection efficiency and specificity. Nanobodies, i.e., camel-derived variable domains of the heavy chain antibodies, have also been obtained against ZYMV CP. Many nucleotide sequences are now available for ZYMV and WMV, particularly in the coat protein (CP) coding region, what allowed the development of specific primers for each virus. Loop-mediated isothermal amplification (LAMP) has also been developed for the diagnostic of WMV and ZYMV.

Vector Relationships WMV is transmitted by at least 35 aphid species in 19 genera. Fewer aphid species were tested for their ability to transmit ZYMV, and 11 were identified as ZYMV vectors. Aphis craccivora, Aphis gossypii, Macrosiphum euphorbiae and Myzus persicae are efficient WMV and ZYMV vectors. Some aphid species were shown to be poor or non-vectors of WMV and ZYMV what suggests some level of specificity in the virus-vector interaction. WMV and ZYMV are transmitted on the non-persistent mode: they are acquired and transmitted during very short probes (a few seconds to minutes), and their retention period in the vector is relatively short (a few hours). WMV and ZYMV as typical potyviruses require the presence of a virus-encoded helper component (HC-Pro) protein for transmission. HC-Pro from WMV and ZYMV are interchangeable and both mediate efficiently the transmission of purified virions of both species. Several ZYMV isolates that have lost aphid transmissibility have been characterized, and a unique feature for this virus is that single amino acid mutants have been identified in the three domains important for transmission. ZYMV-NAT has a A to T substitution in the DAG motif in the CP, ZYMV-PAT a T to A substitution in the PTK motif and ZYMV-R1A a K to E substitution in

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the KLSC motif, both in the HC-Pro. These mutants led to the identification of an interaction between the HC-Pro and CP through their PTK and DAG domains. The non-transmissible isolate ZYMV-NAT (having the DTG motif in the CP) could be transmitted by aphids from plants infected concomitantly by a transmissible isolate of PRSV. This occurred through hetero-encapsidation, a phenomenon by which ZYMV RNA is completely or partially encapsidated by the functional PRSV CP. An aphid non-transmissible isolate deficient for the HC-Pro can also be transmitted by aphids when in mixed infection with an isolate that has a functional HC-Pro. The transmissible isolate provides its functional HC-Pro to mediate the transmission of the deficient isolate. These two mechanisms can contribute to the maintenance, in natural conditions, of variants which have lost their vector transmissibility. An interesting interaction has been observed between ZYMV and A. gossypii, an aphid vector colonizing cucurbit crops. A. gossypii lives longer and produce more offspring on ZYMV infected than on non-infected plants. In addition, more alatae are produced on infected plants, which may stimulate ZYMV spread. These phenomena might be related to the observed changes in phloem exudates composition (free amino acids, sugars) in virus infected plants.

Epidemiology Virus Sources WMV has not been described as seed borne in cucurbits or other crops, but may be transmitted through vegetative propagation in vanilla. ZYMV seed transmission has long remained controversial, with estimated rates ranging from 0% to 18%. Seed transmission was observed in naked seed pumpkin. Using RT-PCR detection, a vertical transmission rate of 1.6% was observed in C. pepo subsp. texana. The vertically-infected plants were symptomless and had low virus titers, but they could constitute sources for further horizontal transmission, either mechanically or by aphids. Pollen transmission occurred at a rate of only 0.13%, indicating that most seed transmission is via the ovule. Another possible way for long distance dissemination of ZYMV is through the globalization of vegetable production and trade. It has been shown that ZYMV infected fruits imported from Central America into Europe could be very efficient virus sources for aphids. Virus sources from which epidemics will initiate could be infected overwintering weeds or crops. In tropical and sub-tropical regions, cucurbit crops or weeds are growing all year round, and viruses could easily move from an old infected crop or cucurbit weed to a young planting. In more temperate regions non-cucurbit weeds were found to be efficient reservoirs for WMV but not for ZYMV. However, a few ornamental plants such as Begonia sempervirens were shown to be involved in ZYMV overwintering in Southern France. Winter protected crops could contribute to ZYMV overwintering in Mediterranean regions and residential gardens were found important sources of ZYMV in California.

Efficient Vectors Many potential aphid vector species have been identified for WMV and ZYMV. A study was conducted to compare the vector capacity of two aphid species, one colonizing cucurbits (A. gossypii), the other not (A. craccivora). Two parameters were used. Transmission efficiency was measured in the laboratory with single aphids exposed in sequence to an infected plant and then to four healthy plants. Transmission propensity was measured by arena tests (more representative of natural conditions) in which aphids could move between plants and feed without interference. It was shown that A. craccivora had both a higher efficiency and propensity to disseminate ZYMV than A. gossypii. This highlights the importance of non-colonizing transient vector species in the epidemiology of ZYMV. In addition to vector transmission, ZYMV transmission has been shown to occur in zucchini squash either through direct contact or trampling on leaves of neighboring plants or through tools used for collecting fruit.

Pattern of Spread The same general pattern is observed for WMV and ZYMV. The first contaminations occur generally shortly after planting, depending upon the availability of virus sources in the environment. Then, aphid flights, particularly of non-colonizing species, spread the virus from the primary infection foci to the rest of the crop, what corresponds to the secondary virus spread. The diseased plants are often distributed in large patches that rapidly extend and join each others, leading to the complete contamination of the crop within a few weeks. Simultaneously, the infected crop will serve as a source of virus to contaminate weeds or nearby young plantings. Epidemic development curves have an overall S shape, fitting generally well to the logistic model.

Control Methods Prophylactic Measures Prophylactic measures are intended to prevent or limit the contact of viruliferous aphids with cultivated plants. They are not specific for a particular virus and are generally efficient for all aphid-borne viruses. These include careful weeding near plantings

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and avoiding overlapping crops in the same area to reduce virus and aphid sources near new plantings. When possible, manipulation of planting dates can avoid exposing young plants to peaks of vector populations. Plastic mulches have a repelling action on aphids and significantly delay WMV and ZYMV spread. However, they confer only a temporary protection that is limited to the early stages of the crop, because their efficiency decreases when plant growth covers their surface. Row covers of different types (unwoven, perforated plastics…) can also be used; they physically prevent winged aphids from reaching the plants, but they must be removed to allow insect pollination necessary for cucurbits. Both methods have a major drawback: they require a lot of plastic material that farmers must dispose in an ecologically sound way after the crop cycle. Tall non-host barriers (millet, Pennisetum glaucum) planted upwind of potential infection sources also contribute to reducing virus infection. Insecticides applications have been generally found inefficient in limiting WMV and ZYMV spread. This is to relate to the large number of winged aphids that land on the plants and to the rapidity of the transmission process. Oil applications can delay virus spread when inoculum pressure is moderate. When applicable, a one-month crop-free period has been shown to be efficient in limiting ZYMV spread. On-farm hygiene practices such as decontamination of tools are also recommended to limit virus spread through contact transmission.

Cross-Protection Cross-protection has been developed at a commercial level to protect cucurbit crops against ZYMV. The principle is simple: when a mild virus isolate (i.e., that has no significant impact on commercial yield) is inoculated to young seedlings, it protects the plant from subsequent contaminations by severe isolates of the same virus. The mechanism appears related to virusinduced gene silencing. The mild strain ZYMV-WK is a natural variant of a severe aphid non-transmissible isolate. Although efficient against most ZYMV isolates (Fig. 4), ZYMV-WK does not protect against very divergent isolates such as those from Réunion Island indicating some specificity in the protection. A single amino acid change (R to I) in the FRNK conserved domain of the HC-Pro is responsible for symptom attenuation of ZYMV-WK. A complete technological package (mild strain production, quality control protocols, and inoculation machines) has been developed to implement commercially ZYMV cross-protection. Mild WMV isolates have been reported that could also have a potential for cross-protection but they have not been used commercially.

Resistant Cultivars The use of virus-resistant cultivars is probably the easiest and cheapest way to control plant viral diseases at the farmer’s level. Breeding for resistance still mainly relies upon search for resistance characters in germplasm collections and introgression of the resistance gene(s) into commercially acceptable cultivars. Considerable efforts have been made to look for resistance to WMV and ZYMV in genetic resources and some WMV- or ZYMV-resistant commercial cultivars are now available. Some resistance genes confer complete and durable resistance (such as the recessive zym gene in cucumber) while others confer only partial resistances or may be overcome by virus evolution (such as the dominant Zym gene in melon). An interesting situation is observed for ZYMV resistance in squash. Although the resistance level was high in the original accession of Cucurbita moschata in which the resistance was identified, when transferred through interspecific crosses to zucchini squash (C. pepo), the resistance phenotype was different: ZYMV multiplies but the plants present only very mild symptoms (this is called tolerance). However, tolerance appears not to be stable since aggressive variants of the virus (i.e., causing severe symptoms in tolerant plants) may emerge in these plants. A single amino acid change in the P3 gene is sufficient to confer this aggressive phenotype. However, the aggressive variants are counterselected when in competition with common ZYMV isolates in susceptible cucurbits. This genetic load associated with aggressiveness could be a factor that will make the tolerance durable. In melon, a resistance to WMV and ZYMV transmission by

Fig. 4 Cross-protected zucchini crop in greenhouse conditions. Left: unprotected plants infected with a severe strain of ZYMV; right: plants preventively inoculated with the mild strain ZYMV-WK, protected against infection with severe strains.

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A. gossypii was found to be governed by the single dominant gene Vat. This gene is present in many commercial cultivars, but confers only limited protection in the fields, probably because WMV and ZYMV are also transmitted by many other aphid species in natural conditions. In the last three decades, attempts were made to obtain WMV and ZYMV transgenic resistant plants using the pathogen derived resistance approach. Different constructs were tested to obtain resistant WMV or ZYMV plants: untranslatable or full-length CP gene, ribozymes. Freedom II, a transgenic squash hybrid containing the WMV and ZYMV CP genes was released in the USA in 1995, as the first virus-resistant transgenic crop to be commercially cultivated in the world. It proved to have a very efficient resistance to WMV and ZYMV in field conditions. Similar hybrids are presently grown mainly in South Eastern USA, particularly during late summer and fall when WMV and ZYMV inoculum pressure are high. Surprisingly, protection against ZYMV infection in transgenic squash presented indirect costs: higher predation by beetles and higher incidence of bacterial wilt. Exogenously applied dsRNA molecules deriving from the ZYMV genome conferred variable levels of resistance do ZYMV in experimental conditions: protection up to 82% was achieved in cucumber using a 588 bp fragment of the HC-pro. Inoculation of cucumber with an apple latent spherical virus vector harboring part of ZYMV genome prevented subsequent experimental infection by ZYMV, and also had a curative effect on ZYMV-infected plants. Recently, melons and cucumbers either silenced or mutated in the eukaryotic translation initiation factor 4E (eIF4E) that is required for potyvirus infection, were found to be resistant to several viruses including ZYMV.

Variability Only limited biological variability has been reported for WMV. This concerns mainly differences in symptom intensity, host range or aphid transmissibility. Among the few host range specificities, the ability or not to infect systemically Chenopodium quinoa is related to a point mutation in WMV CP. In contrast, from its first description, ZYMV appeared to have a very important biological diversity in host range and symptomatology on susceptible hosts, with isolates producing mild or atypical mosaic symptoms, necrosis or wilting reactions. Different pathotypes could be differentiated on melon or squash varieties possessing resistance genes. Important variability has also been observed in aphid transmissibility and several aphid non- or poorly-transmissible isolates have been described. Limited serological variability was observed in WMV and ZYMV when polyclonal antibodies were used. However, the development of monoclonal antibodies against WMV and ZYMV allowed the characterization of serotypes closely correlated to the molecular variability. The molecular variability of ZYMV and WMV was assessed first on the CP region, particularly the N-terminal part of the CP that is known for potyviruses to be highly variable and is frequently used for molecular studies. Besides, more than 50 complete sequences are now available for both ZYMV and WMV. For ZYMV, 3 major molecular groups were defined. Groups B and C contain isolates from specific geographic origins (Reunion Island, Singapore, Vietnam, Australia and East Timor for group B; China, Vietnam and Poland for group C) whereas group A contains worldwide isolates, including the first ZYMV isolates described in the 1970s in Europe. Within group A, 6–7 subgroups were defined without strong geographic structure (Fig. 5). Molecular analyses have allowed, in a few cases, tracking the putative origins of ZYMV strains emerging in a new area. For WMV, the first studies indicated the presence of 3 major molecular groups more or less correlated with the geographic origin of the isolates: Group 1 in Europe, the Mediterranean Basin and part of Asia, Group 2 in South America and the Pacific area and Group 3 in Eastern Asia. With the increase of the number of sequences available, the situation has become more complex (Fig. 5), and evidence for long-distance exchanges of genetic diversity was observed, sometimes followed by strain replacement as observed in several European countries in the 2000s.

Genetics and Evolution WMV and ZYMV genome organization is very similar to that of other potyviruses sequenced so far. The single-stranded, positivesense RNA genome (9.6 kb for ZYMV, 10 kb for WMV excluding the polyadenylated 30 extremity) is translated as a single polyprotein that is self-cleaved in 10 functional proteins, with an 11th protein expressed through replication slippage in the P3 coding region. The full-length sequence revealed that WMV is very closely related to the legume-infecting soybean mosaic virus (SMV); however, the P1 protein is 135 amino-acids longer than that of SMV, and the N-terminal half of the P1 shows no relation to SMV but is 85% identical to another legume-infecting potyvirus, bean common mosaic virus (BCMV). This suggests that WMV has emerged through an ancestral recombination event between a SMV-like and a BCMV-like potyvirus. SMV and BCMV have a narrow host range, mostly restricted to legumes, while WMV is one of the potyviruses with the broadest host range, including monocots and dicots. The impact of its recombinant nature on WMV biological properties remains unknown. Some SMV isolates from China have the same recombination with BCMV in the 50 part of the genome as WMV but they did not infect watermelon even though, contrary to regular SMV isolates, they could infect common bean. Partial and complete sequence data indicate the frequent presence of intraspecific recombinants in WMV. By sequencing the full-length genome of putative recombinants, recombination points were characterized in different parts of the genome, with a

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Fig. 5 Distance tree based on the nucleotide sequence of the coat protein coding region of ZYMV (left) and WMV (right), and showing the main molecular groups currently acknowledged for these viruses. Branch lengths indicate the molecular divergence between sequences (the scale bar represents 0.02 and 0.005 mutations per site for ZYMV and WMV respectively). Figures at some nodes represent bootstrap values (in %), indicating the robustness of each node.

hotspot in the P3-CI region. In France, recombinants between Groups 1 and 3 were observed in the few years following the introduction of Group 3, but they did not become prevalent in populations. In the case of ZYMV, sequence analysis software suggest that recombination might have taken place, but the situation is not as clear cut as for WMV. Besides recombination, viral genomes also evolve by mutations that take place during replication. In the case of ZYMV, the biological consequences of several point mutations that emerged in natural conditions were assessed by molecular studies: sequences of closely related strains with different biological properties were compared, and the mutations observed – particularly non-silent mutations located in conserved domains – were introduced by site-directed mutagenesis in an infectious cDNA of ZYMV in order to check if the mutation alone is sufficient to induce the difference in biological properties. Molecular studies have thus shown that single point mutations in HC-Pro or P3 protein of ZYMV were important for symptomatology. Similarly, single mutations in HC-Pro or CP greatly affect aphid transmissibility. For unknown reasons, some ZYMV isolates such as ZYMV-E15 seems to be particularly prone to mutations and many variants (mild, aphid non-transmissible, aggressive or virulent) have been derived from this isolate.

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Biotechnological Application The development of infectious cDNA clones of ZYMV brought the possibility of using ZYMV as a biotechnological tool to produce proteins of pharmaceutical or crop protection interest. The gene coding for the protein of interest can be inserted in the ZYMV genome either between the P1 and HC-Pro genes or between the NIb and CP genes. For optimal protein production, it is possible to use clones that have the mutation in the FRNK domain so that they produce mild symptoms and do not affect plant growth. The protein may be produced in the edible cucurbit fruits that could be used directly for oral administration. Several molecules of pharmaceutical interest have been efficiently produced through this technology: the human interferon-alpha 2, antiviral and antitumor proteins MAP30 and GAP31 and a mite allergen. This technology also provided a way to produce large amounts of nucleocapsid proteins of 5 tospoviruses that could be used for immunization and production of specific antibodies. Finally, the bar gene coding for a phosphinothricin acetyltransferase that confer resistance to glufosinate ammonium based herbicides has been inserted into the mild ZYMV expression vector. In weed infested plots, this construct efficiently protected inoculated zucchini squash plants from damage caused by an herbicide treatment that completely destroyed the weeds. In addition, these plants were protected against severe ZYMV isolates. It is interesting to see that ZYMV, which emerged as the most damaging cucurbit virus in the last decades, can be manipulated and used for the benefit of human health and agriculture.

Further Reading Arazi, T., Slutsky, S.G., Shiboleth, Y.M., et al., 2001. Engineering Zucchini yellow mosaic potyvirus as a non-pathogenic vector for expression of heterologous proteins in cucurbits. Journal of Biotechnology 87, 67–82. Coutts, B.A., Kehoe, M.A., Jones, R.A.C., 2011. Minimizing losses caused by Zucchini yellow mosaic virus in vegetable cucurbit crops in tropical, sub-tropical and Mediterranean environments through cultural methods and host resistance. Virus Research 159, 141–160. Coutts, B.A., Kehoe, M.A., Jones, R.A.C., 2013. Zucchini yellow mosaic virus: Contact transmission, stability on surfaces, and inactivation with disinfectants. Plant Disease 97, 765–771. Coutts, B.A., Kehoe, M.A., Webster, C.G., Wylie, S.J., Jones, R.A.C., 2011. Zucchini yellow mosaic virus: Biological properties, detection procedures and comparison of coat protein gene sequences. Archives of Virology 156, 2119–2131. Desbiez, C., Joannon, B., Wipf-Scheibel, C., Chandeysson, C., Lecoq, H., 2011. Recombination in natural populations of watermelon mosaic virus: New agronomic threat or damp squib? Journal of General Virology 92, 1939–1948. Desbiez, C., Lecoq, H., 1997. Zucchini yellow mosaic virus. Plant Pathology 46, 809–829. Desbiez, C., Lecoq, H., 2004. The nucleotide sequence of watermelon mosaic virus (WMV, Potyvirus) reveals interspecific recombination between two related potyviruses in the 50 part of the genome. Archives of Virology 149, 1619–1632. Jones, R.A.C., 2009. Plant virus emergence and evolution: Origins, new encounter scenarios, factors driving emergence, effects of changing world conditions, and prospects for control. Virus Research 141, 113–130. Lecoq, H., 1998. Control of plant virus diseases by cross protection. In: Hadidi, A., Kheterpal, R.K., Koganezawa, H. (Eds.), Plant Virus Disease Control. St Paul, Minnesota: APS Press, pp. 33–40. Lecoq, H., Desbiez, C., 2012. Viruses of cucurbit crops in the Mediterranean region: An ever-changing picture. Advances in Virus Research 84, 67–126. Lecoq, H., Wipf-Scheibel, C., Nozeran, K., Millot, P., Desbiez, C., 2014. Comparative molecular epidemiology provides new insights into Zucchini yellow mosaic virus occurrence in France. Virus Research 186, 135–143. Lisa, V., Lecoq, H., 1984. Zucchini yellow mosaic virus. CMI/AAB Descriptions of Plant Viruses No. 282. Kew, Surrey: Commonwealth Mycological Institute. Purcifull, D., Hiebert, E., Edwardson, J., 1984. Watermelon mosaic virus 2. CMI/AAB Descriptions of Plant Viruses No. 293. Kew, Surrey: Commonwealth Mycological Institute.