Advances in Virus Research, Vol. 92 [1st ed.] 0128021802, 9780128021804

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ADVISORY BOARD DAVID BALTIMORE PETER C. DOHERTY HANS J. GROSS BRYAN D. HARRISON BERNARD MOSS ERLING NORRBY PETER PALUKAITIS JOHN J. SKEHEL MARC H.V. VAN REGENMORTEL

Academic Press is an imprint of Elsevier 225 Wyman Street, Waltham, MA 02451, USA 525 B Street, Suite 1800, San Diego, CA 92101-4495, USA 125 London Wall, London, EC2Y 5AS, UK The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, UK First edition 2015 Copyright © 2015, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. ISBN: 978-0-12-802180-4 ISSN: 0065-3527 For information on all Academic Press publications visit our website at store.elsevier.com

DEDICATION Frederick A. Murphy, Editor of Advances in Virus Research Started in 1953, Advances in Virus Research was the first review series in virology. For much of its history, Frederick A. Murphy served as a distinguished Series Editor, editing over 60 volumes in the 31 years of his tenure. Fred has decided to retire following the publication of Volume 91, and we would like to take this opportunity to thank him for his hard work and dedication to the field of virology in general and to this publication more specifically. Fred continues to hold his post as a professor in the Department of Pathology at the University of Texas Medical Branch (UTMB), Galveston, Texas. At UTMB, he is a member of the Institute for Human Infections and Immunity, the Center for Biodefense and Emerging Infectious Diseases, the Galveston National Laboratory, and the McLaughlin Endowment for Infection and Immunity. He holds a BS and DVM from Cornell University and a PhD from the University of California, Davis. Fred served as Dean and Distinguished Professor, School of Veterinary Medicine, and Distinguished Professor, School of Medicine, University of California, Davis. Before that he served as Director of the Division of Viral and Rickettsial Diseases and then Director of the National Center for Infectious Diseases, Centers for Disease Control, Atlanta. He is a member of the Institute of Medicine of the U.S. National Academy of Sciences (where he is a Lifetime National Associate of the National Research Council and has served on 10 committees) and a member of the Deutsche Nationale Akademie der Wissenschaften (German National Academy of Sciences Leopoldina) and the Acade´mie Royale de Me´decine de Belgique (Belgian Royal Academy of Medicine). Over the course of his career, Fred has earned a long list of honors, including an honorary Doctor of Medicine and Surgery from the University of Turku, Turku, Finland; an honorary Doctor of Science from the University of Guelph, Ontario, Canada; an honorary Doctor of Veterinary Medicine from the Royal Veterinary College, University of London, United Kingdom; an honorary Doctor of Science from University College Dublin, Ireland; the Presidential Rank Award of the U.S. Government; the Penn Vet World Leadership Award from the University of Pennsylvania; the Distinguished Microbiologist Award from the American College of Veterinary Microbiologists; the Richard Moreland Taylor Award of the American v

vi

Dedication

Committee on Arthropod-Borne Viruses, American Society of Tropical Medicine and Hygiene; the University of California Davis Medal; the K.F. Meyer Gold Headed Cane of the American Veterinary Epidemiology Society and American Veterinary Medical Association; and an Honorary Fellowship from the John Curtin School of Medical Research, The Australian National University. Fred’s professional interests include the pathology and epidemiology of highly pathogenic viruses/viral diseases, with a focus on rabies, arboviruses, viral hemorrhagic fevers, and viral encephalitides. He has been a leader in advancing the concept of “new and emerging infectious diseases” and “new and emerging zoonoses,” which has reenergized the infectious disease research sciences. Most recently, Fred’s interests have included the threat posed by bioterrorism and national efforts in prevention of this threat. For the past three decades, Fred has been a crucial part of making Advances in Virus Research a continued success. His contributions and enthusiasm for moving research forward have enriched the pages of this book series for 31 years and we will miss him greatly. On behalf of all of the past, present, and future authors and editors of Advances in Virus Research, along with the thousands of researchers who continue to benefit from your work, we would like to extend a very heartfelt thank to you, Fred. Karl Maramorosch, Thomas Mettenleiter, and Mary Ann Zimmerman

John, Terence, Irene, Fred, Rick, and Tim Murphy, 1998.

CONTRIBUTORS Nina S. Atanasova Department of Biosciences and Institute of Biotechnology, University of Helsinki, Helsinki, Finland Dennis H. Bamford Department of Biosciences and Institute of Biotechnology, University of Helsinki, Helsinki, Finland Juan Antonio Garcı´a Centro Nacional de Biotecnologı´a (CNB-CSIC), Campus Universidad Auto´noma de Madrid, Madrid, Spain Stefanie Luecke Graduate School of Life Sciences, Universiteit Utrecht, Utrecht, The Netherlands Hanna M. Oksanen Department of Biosciences and Institute of Biotechnology, University of Helsinki, Helsinki, Finland Søren R. Paludan Department of Biomedicine, and Aarhus Research Center for Innate Immunology, Aarhus University, Aarhus, Denmark Maija K. Pietila¨ Department of Biosciences and Institute of Biotechnology, University of Helsinki, Helsinki, Finland Fre´de´ric Revers INRA, and Universite´ de Bordeaux, UMR 1332 de Biologie du fruit et Pathologie, Villenave d’Ornon, France Elina Roine Department of Biosciences and Institute of Biotechnology, University of Helsinki, Helsinki, Finland Ana Sencˇilo Department of Biosciences and Institute of Biotechnology, University of Helsinki, Helsinki, Finland Joanna L. Shisler Department of Microbiology, College of Medicine, University of Illinois, Urbana, Illinois, USA

ix

CHAPTER ONE

Comparison of Lipid-Containing Bacterial and Archaeal Viruses Nina S. Atanasova, Ana Senčilo, Maija K. Pietilä, Elina Roine, Hanna M. Oksanen, Dennis H. Bamford1 Department of Biosciences and Institute of Biotechnology, University of Helsinki, Helsinki, Finland 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Origin of lipids in prokaryotic viruses and their detection 2. Function and Significance of Lipids in Prokaryotic Virus Life Cycle 2.1 How prokaryotic viruses acquire their lipids 3. Currently Known Lipid-Containing Bacterial and Archaeal Viruses 3.1 Icosahedral viruses with an inner membrane 3.2 Enveloped icosahedral viruses: Phage ϕ6 and its relatives 3.3 Vesicular pleomorphic viruses 3.4 Prokaryotic viruses with helical symmetry: With or without a membrane 3.5 Lemon-shaped viruses are specific for archaea 3.6 Archaeal spherical viruses with helical NCs have an envelope 4. Conclusions Acknowledgments References

2 3 18 21 24 24 32 35 40 43 46 47 49 49

Abstract Lipid-containing bacteriophages were discovered late and considered to be rare. After further phage isolations and the establishment of the domain Archaea, several new prokaryotic viruses with lipids were observed. Consequently, the presence of lipids in prokaryotic viruses is reasonably common. The wealth of information about how prokaryotic viruses use their lipids comes from a few well-studied model viruses (PM2, PRD1, and ϕ6). These bacteriophages derive their lipid membranes selectively from the host during the virion assembly process which, in the case of PM2 and PRD1, culminates in the formation of protein capsid with an inner membrane, and for ϕ6 an outer envelope. Several inner membrane-containing viruses have been described for archaea, and their lipid acquisition models are reminiscent to those of PM2 and PRD1. Unselective acquisition of lipids has been observed for bacterial mycoplasmaviruses and archaeal pleolipoviruses, which resemble each other by size, morphology, and life style. In addition to these shared morphotypes of bacterial and archaeal viruses, archaea are infected by viruses with unique morphotypes, such as

Advances in Virus Research, Volume 92 ISSN 0065-3527 http://dx.doi.org/10.1016/bs.aivir.2014.11.005

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Nina S. Atanasova et al.

lemon-shaped, helical, and globular ones. It appears that structurally related viruses may or may not have a lipid component in the virion, suggesting that the significance of viral lipids might be to provide viruses extended means to interact with the host cell.

1. INTRODUCTION In principal, all cellular organisms are infected by viruses. In its simplest form, an infectious viral particle, the virion, is composed of a genome that is covered with a protective coat. The coat can be built using proteins or proteinaceous lipid membranes. To date, most of the studied prokaryotic viruses (bacteriophages and archaeal viruses) are of head-tailed icosahedral morphology with no lipids (Fig. 1). Among the described bacteriophages, lipid-containing phages are considered to be rare and represent less than 5% of the described isolates (Ackermann & Prangishvili, 2012). Although there is a wide variety of morphotypes in archaeal viruses, the number of known archaeal viruses is a small proportion of the known prokaryotic viruses (Ackermann & Prangishvili, 2012). Yet, many of them contain lipids (Table 1 and Fig. 1). The discovery of the first lipid-containing prokaryotic virus, PM2, occurred only in the late 1960s (Espejo & Canelo, 1968a). Bacteriophages had already been known for several decades, but the late discovery of lipid-containing prokaryotic viruses was partially due to the common protocol of treating phage stock solutions with chloroform in order to avoid bacterial growth (Adams, 1959). Soon after that ϕ6, an enveloped bacteriophage, was isolated (Vidaver et al., 1973) followed by the discovery of PRD1, an icosahedral virus with an inner membrane (Olsen et al., 1974). The first lipid-containing archaeal virus, the helical Thermoproteus tenax virus 1 (TTV1), was reported in the late 1980s (Zillig et al., 1988). The lipid membranes of the prokaryotic viruses usually have significant functions during viral life cycle. Although some generalizations can be made between the virion morphology and the membrane localization in relation to its functions, the type of the genome and its delivery during the infection, for example, may influence the specific roles the membrane plays during the viral life cycle. Although most of the information comes from the model bacteriophages such as PRD1, PM2, and ϕ6, new mass spectrometric techniques developed for lipid research may in the future bring additional information. This is important especially for the research of archaeal viruses due to difficulties in obtaining highly pure viral material.

Prokaryotic Viruses with Lipids

3

Figure 1 Morphotypes of known prokaryotic viruses: (A) bacteriophages and (B) archaeal viruses. The localization of the lipid membrane is indicated by red except for His1 for which the lipid-modified major structural protein is indicated by dashed red line. Numbers of the viruses that contain lipids are underlined. The scale bar represents 100 nm. The virions are in scale except for those marked with an asterisk are 0.5
their actual size. The representative morphotypes are exemplified by: 1. PRD1; 2. PM2, 3. ϕ6; 4. L2; 5. T4; 6. λ; 7. M13; 8. ϕX174; 9. MS2; 10. T7; 11. STIV; 12. PSV; 13. HRPV-1; 14. His1; 15. STSV1; 16. ATV; 17. AFV1; 18. SIRV-1; 19. Aeropyrum pernix bacilliform virus 1 (APBV1); 20. Acidianus bottle-shaped virus (ABV); 21. Sulfolobus neozealandicus droplet-shaped virus (SNDV); 22. Aerupyrum coilshaped virus (ACV); 23. Halorubrum sodomense tailed virus 2 (HSTV-2); 24. Haloarcula vallismortis tailed virus 1 (HVTV-1); and 25. Haloarcula sinaiiensis tailed virus 1 (HSTV-1). The lipid-containing viruses are presented in Table 1. For viruses without lipids, see King, Adams, Carstens, and Lefkowitz (2012) and Pietilä, Demina, Atanasova, Oksanen, and Bamford (2014).

1.1. Origin of lipids in prokaryotic viruses and their detection Since prokaryotic viruses obtain their membranes from the host, the foundations of prokaryotic viral lipids are laid by the host lipid synthesis. In this sense, bacteria and archaea are profoundly different. In general, the core structures of archaeal membrane lipids consist of two isoprenoid side chains that are linked to sn-glycerol-1-phosphate (G1P) backbone through ether bonds (Kates, 1993), whereas in bacteria, the core membrane lipids contain two fatty acid side chains linked to sn-glycerol-3-phosphate (G3P) through ester linkages (Fig. 2; Cronan, 2003; Kates, 1993). The two major core lipids of archaea are archaeol and caldarchaeol (Fig. 2). Archaeol, the

Table 1 Lipid-containing viruses infecting prokaryotes Virus

Isolation host Genome size

Life cycle

Lipids (position, acquisition)

Gram Domain staining References

Virus name

Morphotype

Family

Genome type

PRD1

Icosahedral

Tectiviridae

Linear dsDNA 14,927, ITR (110 bp)

Virulent

Internal membrane, selective

Salmonella typhimurium

B

Gram Olsen, Siak, and Gray (1974), Bamford et al. (1991), Cockburn et al. (2004), and Laurinavicˇius, Ka¨kela¨, Somerharju, and Bamford (2004)

PR3

Icosahedral

Tectiviridae

Linear dsDNA 14,937, ITR (111 bp)

Virulent

Internal membrane, ND

Pseudomonas aeruginosa PAO

B

Gram Stanisich (1974), Bamford, Rouhiainen, Takkinen, and S€ oderlund (1981), and Saren et al. (2005)

PR4

Icosahedral

Tectiviridae

Linear dsDNA 14,943, ITR (111 bp)

Virulent

Internal membrane, ND

P. aeruginosa PAO B

Gram Stanisich (1974), Bamford et al. (1981), and Saren et al. (2005)

PR5

Icosahedral

Tectiviridae

Linear dsDNA 14,939, ITR (110 bp)

Virulent

Internal membrane, selective

Escherichia coli K12 B

Gram Wong and Bryan (1978), Bamford et al. (1981), and Saren et al. (2005)

Strain name

PR722

Icosahedral

Tectiviridae

Linear dsDNA 14,942, ITR (111 bp)

Virulent

Internal membrane, ND

Proteus niirubilis PM5006

B

Gram Coetzee, Lecatsas, Coetzee, and Hedges (1979) and Saren et al. (2005)

L17

Icosahedral

Tectiviridae

Linear dsDNA 14,935, ITR (111 bp)

Virulent

Internal membrane, ND

Aeromonas B hydrophila (RP1) and E. coli W3110 (RP1)

Gram Bamford et al. (1981) and Saren et al. (2005)

P37-14

Icosahedral

Tectiviridae

dsDNA

ND

Internal membrane, ND

Thermus sp.

Gram Yu, Slater, and Ackermann (2006)

Bam35

Icosahedral

Tectiviridae

Linear dsDNA 14,935, ITR (74 bp)

Temperate Internal membrane, selective

Bacillus thuringiensis B

AP50

Icosahedral

Tectiviridae

Linear dsDNA 14,398

Temperate Internal membrane, ND

Bacillus anthracis

20,000

B

B

Gram+ Ackermann, Roy, Martin, Murthy, and Smirnoff (1978), Ravantti, Gaidelyte, Bamford, and Bamford (2003), Str€ omsten, Benson, Burnett, Bamford, and Bamford (2003), and Laurinavicˇius, Ka¨kela¨, Somerharju, and Bamford (2004) Gram+ Nagy, Pragai, and Ivanovics (1976), Nagy and Ivanovics (1977), and Sozhamannan et al. (2008) Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Isolation host Genome size

Gram Domain staining References

Virus name

Morphotype

Family

Genome type

GIL01

Icosahedral

Tectiviridae

Linear dsDNA 14,931, Temperate NDa imperfect ITR

B. thuringiensis serovar Israelensis strain AND508

B

Gram+ Verheust, Jensen, and Mahillon (2003)

GIL16

Icosahedral

Tectiviridae

Linear dsDNA 14,844, Temperate ND imperfect ITR

B. thuringiensis serovar Israelensis strain AND508

B

Gram+ Verheust, Fornelos, and Mahillon (2005)

ϕNS11

Icosahedral

Tectiviridae

Linear dsDNA ND

Temperate Internal Bacillus membrane, acidocaldarius TA6 nonselective

B

Gram+ Sakaki and Oshima (1976), Sakaki, Oshima, Yamada, and Oshima (1977), and Sakaki, Maeda, and Oshima (1979)

Wip1

Icosahedral

Unclassified

Linear dsDNA 14,319

Temperate Internal membrane, ND

B. anthracis

B

Gram+ Schuch, Pelzek, Kan, and Fischetti (2010) and Kan, Fornelos, Schuch, and Fischetti (2013)

P23-77

Icosahedral

Sphaerolipoviridae Circular dsDNA

Virulent

Thermus thermophilus ATCC 33923

B

Gram Yu et al. (2006), Jaatinen, Happonen, Laurinma¨ki, Butcher, and

17,036

Life cycle

Lipids (position, acquisition)

Internal membrane, selective

Strain name

Bamford (2008), Jalasvuori et al. (2009), and Pawlowski, Rissanen, Bamford, Krupovicˇ, and Jalasvuori (2014) KHP30

Icosahedral

Unclassified

Circular dsDNA

26,215

Temperate Internal membrane, ND

Helicobacter pylori NY43

B

Gram Uchiyama et al. (2012) and Uchiyama et al. (2013)

PM2

Icosahedral

Corticoviridae

Circular dsDNA

10,079

Virulent

Internal membrane, selective

Pseudoalteromonas espejiana BAL-31

B

Gram Espejo and Canelo (1968b), CameriniOtero and Franklin (1972), and Ma¨nnist€ o, Kivela¨, Paulin, Bamford, and Bamford (1999)

Salisaeta icosahedral phage 1 (SSIP-1)

Icosahedral

Unclassified

Circular dsDNA

43,788

Virulent

Internal membrane, selective

Salisaeta sp. SP9-1 B

Gram Aalto et al. (2012) and Atanasova, Roine, Oren, Bamford, and Oksanen (2012) Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Genome type

Isolation host Genome size

Life cycle

Lipids (position, acquisition)

Gram Domain staining References

Virus name

Morphotype

Family

SNJ1

Icosahedral

Sphaerolipoviridae Circular dsDNA

16,341

Temperate Internal membrane, selective

Natrinema sp. J7-1 A

–

Zhang et al. (2012) and Pawlowski et al. (2014)

Strain name

Sulfolobus Icosahedral turreted icosahedral virus (STIV)

Turriviridae

Circular dsDNA

17,663

Virulent

Internal membrane, selective

Sulfolobus solfataricus YNPRC179

A

–

Rice et al. (2004) and Maaty et al. (2006)

Sulfolobus Icosahedral turreted icosahedral virus 2 (STIV2)

Turriviridae

Circular dsDNA

16,622

Virulent

Internal membrane, ND

S. solfataricus 2-212

A

–

Happonen et al. (2010) and Happonen et al. (2013)

SH1

Icosahedral

Sphaerolipoviridae Linear dsDNA 30,898, ITR (309 bp)

Virulent

Internal membrane, selective

Haloarcula hispanica A ATCC 33960

–

Bamford et al. (2005), Porter et al. (2005), Kivela¨ et al. (2006), and Pawlowski et al. (2014)

H. hispanica Icosahedral icosahedral virus 2 (HHIV-2)

Sphaerolipoviridae Linear dsDNA 30,578, ITR (305 bp)

Virulent

Internal membrane, NDa,b

H. hispanica ATCC A 33960

–

Atanasova et al. (2012), Jaakkola et al. (2012), and Pawlowski et al. (2014)

PH1

Icosahedral

Sphaerolipoviridae Linear dsDNA 28,064, ITR (337 bp)

φ6

Spherical

Cystoviridae

Linear dsRNA

φ7

Spherical

Cystoviridae

φ8

Spherical

φ9 φ10

Virulent

Internal membrane, NDa,b

H. hispanica ATCC A 33960

–

Porter et al. (2013) and Pawlowski et al. (2014)

14,927 Virulent (in 3 (able to segments) form carrier state cells)

Outer membrane, selective

Pseudomonas syringae pv. phaseolicola HB1OY

B

Gram Vidaver, Koski, and Van Etten (1973), McGraw, Mindich, and Frangione (1986), Gottlieb et al. (1988), Mindich et al. (1988), and Laurinavicˇius, Ka¨kela¨, Bamford, and Somerharju (2004)

Linear dsRNA

13.4 Virulent (in 3 segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999)

Cystoviridae

Linear dsRNA

15 (in 3 Virulent segments)

Outer membrane, NDa

Pseudomonas pseudoalcaligenes ERA

B

Gram Mindich et al. (1999) and Sun, Qiao, Qiao, Onodera, and Mindich (2003)

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999)

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999) Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Isolation host Lipids (position, acquisition)

Gram Domain staining References

Virus name

Morphotype

Family

Genome type

Genome size

φ11

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999)

φ12

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. pseudoalcaligenes B ERA

Gram Mindich et al. (1999)

φ13

Spherical

Cystoviridae

Linear dsRNA

13.7 Virulent (in 3 (able to segments) form carrier state cells)

NDa

P. pseudoalcaligenes B ERA

Gram Mindich et al. (1999)

φ14

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999)

φ2954

Spherical

Cystoviridae

Linear dsRNA

12,685 Virulent (in 3 segments)

NDa

P. syringae LM2489

B

Gram Qiao, Sun, Qiao, Di Sanzo, and Mindich (2010)

L2

Pleomorphic Plasmaviridae

Circular dsDNA

11,965

Persistent

Lipid envelope, ND

Acholeplasma laidlawii

B

Gram Gourlay (1971), Putzrath and Maniloff (1977), and Maniloff, Kampo, and Dascher (1994)

L172

Globular

Circular ssDNA

14,000

Persistent

Lipid envelope, ND

A. laidlawii

B

Gram Dybvig, Nowak, Sladek, and Maniloff (1985)

Unclassified

Life cycle

Strain name

Halorubrum pleomorphic virus 1 (HRPV-1)

Pleomorphic Pleolipoviridae

Circular ssDNA

7048

Persistent

Lipid Halorubrum sp. PV6 envelope, nonselective

A

–

Pietila¨, Roine, Paulin, Kalkkinen, and Bamford (2009) and Pietila¨, Laurinavicˇius, Sund, Roine, and Bamford (2010)

H. hispanica pleomorphic virus 1 (HHPV-1)

Pleomorphic Pleolipoviridae

Circular dsDNA

8082

Persistent

Lipid H. hispanica ATCC A envelope, 33960 nonselective

–

Roine et al. (2010)

Halorubrum pleomorphic virus 3 (HRPV-3)

Pleomorphic Pleolipoviridae

Circular discontinuous dsDNA

8770

Persistent

Lipid Halorubrum sp. envelope, SP3-3 nonselective

A

–

Atanasova et al. (2012), Pietila¨ et al. (2012), and Sencˇilo, Paulin, Kellner, Helm, and Roine (2012)

Halorubrum pleomorphic virus 2 (HRPV-2)

Pleomorphic Pleolipoviridae

Circular ssDNA

10,656

Persistent

Lipid Halorubrum sp. envelope, SS5-4 nonselective

A

–

Atanasova et al. (2012), Pietila¨ et al. (2012), and Sencˇilo et al. (2012)

Halorubrum pleomorphic virus 6 (HRPV-6)

Pleomorphic Pleolipoviridae

Circular ssDNA

8549

Persistent

Lipid Halorubrum sp. envelope, SS7-4 nonselective

A

–

Pietila¨ et al. (2012) and Sencˇilo et al. (2012) Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Genome type

Gram Domain staining References

Genome size

Life cycle

9694

Persistent

Lipid Halogeometricum sp. A envelope, CG-9 nonselective

–

Atanasova et al. (2012), Pietila¨ et al. (2012), and Sencˇilo et al. (2012)

Persistent

Lipid H. hispanica ATCC A envelope, 33960 nonselective

–

Bath, Cukalac, Porter, and DyallSmith (2006) and Pietila¨ et al. (2012)

A

–

Janekovic et al. (1983), Zillig et al. (1988), and Neumann, Schwass, Eckerskorn, and Zillig (1989)

T. tenax Kra1

A

–

Janekovic et al. (1983)

Outer membrane, ND

T. tenax Kra1

A

–

Janekovic et al. (1983)

Outer membrane, selectived

Sulfolobus isolate HVE11/2

A

–

Arnold et al. (2000) and Peng et al. (2001)

Strain name

Virus name

Morphotype

Halogeometricum pleomorphic virus 1 (HGPV-1)

Pleomorphic Pleolipoviridae

Circular discontinuous dsDNA

His virus 2 (His2)

Pleomorphic Pleolipoviridae

Linear dsDNA 16,067, ITR (525 bp)

Thermoproteus tenax virus 1 (TTV1)

Filamentous Lipothrixviridae

Temperate Outer Linear dsDNA 13,669e (partial membrane, sequence) selective

T. tenax Kra1

T. tenax virus 2 Filamentous Lipothrixviridae (TTV2)

Linear dsDNA 16,000

Temperate Outer membrane, ND

T. tenax virus 3 Filamentous Lipothrixviridae (TTV3)

Linear dsDNA 27,000

ND

Filamentous Lipothrixviridae

Linear dsDNA 40,900e, ITR (at least 800 bp)

Persistent

Sulfolobus islandicus filamentous virus 1 (SIFV)

Family

Isolation host Lipids (position, acquisition)

Desulforolobus ambivalens filamentous virus (DAFV)

Filamentous Lipothrixviridae

Linear dsDNA 56,000

ND

Outer membrane, ND

Desulforolobus ambivalens

A

–

Zillig et al. (1994)

Acidianus filamentous virus 1 (AFV1)

Filamentous Lipothrixviridae

Linear dsDNA 20,869, ITR (11 bp)

Persistent

Outer membrane, selectived

Acidianus hospitalis YS6

A

–

Bettstetter, Peng, Garrett, and Prangishvili (2003)

Acidianus filamentous virus 2 (AFV2)

Filamentous Lipothrixviridae

Linear dsDNA 31,787e

Persistent

No lipids detected

Acidianus sp. strain A F28

–

Ha¨ring et al. (2005)

Acidianus filamentous virus 3 (AFV3)

Filamentous Lipothrixviridae

Linear dsDNA 40,449e, ITR

Persistent

Outer Acidianus sp. strain A membrane, Acii25 nonselective

–

Vestergaard et al. (2008)

Acidianus filamentous virus 6 (AFV6)

Filamentous Lipothrixviridae

Linear dsDNA 39,577e, ITR

Persistent

ND

Acidianus convivator A

–

Vestergaard et al. (2008)

Acidianus filamentous virus 7 (AFV7)

Filamentous Lipothrixviridae

Linear dsDNA 36,895e, ITR

Persistent

ND

A. convivator

A

–

Vestergaard et al. (2008)

Acidianus filamentous virus 8 (AFV8)

Filamentous Lipothrixviridae

Linear dsDNA 38,179e, ITR

Persistent

ND

A. convivator

A

–

Vestergaard et al. (2008)

Acidianus filamentous virus 9 (AFV9)

Filamentous Lipothrixviridae

Linear dsDNA 41,172, ITR (384 bp)

Persistent

Outer membrane, ND

Acidianus uzoniensis A

–

Bize et al. (2008)

Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Isolation host Genome size

Life cycle

Lipids (position, acquisition)

Gram Domain staining References

Virus name

Morphotype

Family

Genome type

His virus 1 (His1)

Spindle shaped

Fuselloviridae

Linear dsDNA 14,464, ITR (105 bp)

Persistent

No membrane, MCP is lipid modified

H. hispanica ATCC A 33960

–

Bath and DyallSmith (1998), Bath et al. (2006), and Pietila¨, Atanasova, Oksanen, and Bamford (2013)

Sulfolobus spindle-shaped virus 1 (SSV1)

Spindle shaped

Fuselloviridae

Circular dsDNA

15,465

Persistent

NDa

Sulfolobus shibatae B12

A

–

Reiter, Zillig, and Palm (1988), Palm et al. (1991), and Schleper, Kubo, and Zillig (1992)

Sulfolobus spindle-shaped virus 2 (SSV2)

Spindle shaped

Fuselloviridae

Circular dsDNA

14,796

ND

NDc

S. solfataricus P1

A

–

Stedman et al. (2003)

Sulfolobus spindle-shaped virus 4 (SSV4)

Spindle shaped

Fuselloviridae

Circular dsDNA

15,135

ND

NDc

S. islandicus ARN3/6

A

–

Peng (2008)

Sulfolobus spindle-shaped virus 5 (SSV5)

Spindle shaped

Fuselloviridae

Circular dsDNA

15,330

ND

ND

S. solfataricus P2

A

–

Redder et al. (2009)

Sulfolobus spindle-shaped virus 6 (SSV6)

Spindle shaped

Fuselloviridae

Circular dsDNA

15,684

ND

ND

S. islandicus G4ST- A T-11

–

Redder et al. (2009)

Strain name

Sulfolobus spindle-shaped virus 7 (SSV7)

Spindle shaped

Fuselloviridae

Circular dsDNA

17,602

ND

ND

S. islandicus G4T-1 A

–

Redder et al. (2009)

Sulfolobus spindle-shaped virus Ragged hills (SSVrh)

Spindle shaped

Fuselloviridae

Circular dsDNA

16,473

ND

ND

S. solfataricus

A

–

Wiedenheft et al. (2004)

Sulfolobus spindle-shaped virus Kamchatka1 (SSVk1)

Spindle shaped

Fuselloviridae

Circular dsDNA

17,384

ND

ND

S. solfataricus

A

–

Wiedenheft et al. (2004)

Acidianus spindle-shaped virus 1 (ASV1)

Spindle shaped

Fuselloviridae

Circular dsDNA

24,186

ND

ND

Acidianus brierleyi DSM1651

A

–

Redder et al. (2009)

Pyrococcus abyssi virus 1 (PAV1)

Spindle shaped

Unclassified

Circular dsDNA

18,098

Persistent

NDa

P. abyssi GE23

A

–

Geslin et al. (2003) and Geslin et al. (2007)

Thermococcus prieurii virus 1 (TPV1)

Spindle shaped

Unclassified

Circular dsDNA

21,591

Persistent

NDa

T. prieurii

A

–

Gorlas, Koonin, Bienvenu, Prieur, and Geslin (2012)

Sulfolobus Spindle tengchongensis shaped spindle-shaped virus 1 (STSV1)

Unclassified

Circular dsDNA

75,294

Persistent

Outer membrane, selective

S. tengchongensis RT8-4

A

–

Xiang et al. (2005)

Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Virus name

Lipids (position, acquisition)

Gram Domain staining References

Genome size

Life cycle

76,107

Persistent

Outer membrane, selectived

S. tengchongensis HB52

A

–

Erdmann et al. (2014)

Family

Genome type

S. tengchongensis Spindle spindle-shaped shaped virus 2 (STSV2)

Unclassified

Circular dsDNA

Pyrobaculum Spherical spherical virus 1 (PSV1)

Globuloviridae

Linear dsDNA 28,337, ITR (190 bp)

Persistent

Outer membrane, selectived

Pyrobaculum sp. D11

A

–

Ha¨ring et al. (2004)

T. tenax Spherical spherical virus 1 (TTSV1)

Globuloviridae

Linear dsDNA 20,933 and 700 bp terminie

Persistent

NDc

T. tenax YS44

A

–

Ahn et al. (2006)

a

Morphotype

Isolation host

Strain name

The presence of the lipids was assumed based on the sensitivity to organic solvents. The presence of the lipids was assumed based on the Sudan Black staining. The presence of the lipids was assumed based on the virus particle density. d The lipids are modified as determined by the thin-layer chromatography of the viral and host lipids. The viral lipids had different mobilities compared to the hosts’. e The nature of the genome termini is unclear. A, archaea; B, bacteria; ITR, inverted terminal repeats; ND, not determined. b c

Prokaryotic Viruses with Lipids

17

Figure 2 The basic structures of core lipids found in bacterial and archaeal membranes. Diacylglycerol is the backbone of bacterial membrane lipids, whereas archaeol and caldarchaeol are found in haloarchaeal or crenarchaeal membrane lipids, respectively.

diphytanylglycerol diether, and its variants can form a membrane bilayer and are the major constituents of the halophilic archaeal membranes. Caldarchaeol is one of the major components in the thermophilic crenarchaeal membranes (such as Sulfolobus spp). It is a dibiphytanyldiglycerol tetraether, an antiparallel arrangement of two glycerol units connected with two isoprenoid acyl chains through four ether linkages (Fig. 2). These lipids form monolayer membranes (Kates, 1993). The major components of bacterial and archaeal membranes are mostly polar phospholipids. In halophilic archaea, these include phosphatidylglycerol (PG), the methyl ester of phosphatidylglycerol (PGP-Me), and phosphatidylglycerosulfate (PGS). In extreme haloarchaea, the PGP-Me contributes approximately 50–80 mol% of the polar lipids (Tenchov, Vescio, Sprott, Zeidel, & Mathai, 2006). In addition, many different types of archaeal glycolipids and species of cardiolipins (CLs) are also components of the membranes. Neutral lipids constitute approximately 10% of haloarchaeal lipids (Corcelli & Lobasso, 2006). The cytoplasmic membrane of Gram-negative bacteria consists of 75% phosphatidylethanolamine (PE), 20% PG, and 5% CL (Cronan, 2003). However, environmental conditions have been reported to influence considerably the lipid composition of bacterial and archaeal membranes (Farrell & Rose, 1967; Lopalco et al., 2013). The fact that prokaryotic viruses obtain their lipids from host membranes can also cause problems in the identification of viral lipids. This is especially true for the enveloped viruses due to the often unselective manner of

18

Nina S. Atanasova et al.

membrane acquisition. Highly purified material is required for the characterization of viral lipids, because it is often difficult to separate virions from host membrane vesicles. This can, in some cases, be overcome by the analyses of membrane-containing subviral particles that have been obtained, for example, by quantitative biochemical dissociation of the virion (Vitale, Roine, Bamford, & Corcelli, 2013). The first indication of the presence of lipids in the virion is the sensitivity to organic solvents such as chloroform or the low buoyant density of the virion. Sudan Black staining of the highly purified virions analyzed in sodium dodecyl sulfate–polyacrylamide gel electrophoresis has also been used as the preliminary indication of membrane lipids or lipid-modified proteins (Pietila¨ et al., 2012; Prat, Lamy, & Weill, 1969). Further analyses of viral lipids have traditionally been conducted using organic solvent extraction followed by thin-layer chromatography (TLC), mass spectrometry (MS; e.g., electrospray ionization, ESI-MS), and/or nuclear magnetic resonance spectroscopy (NMR; Corcelli & Lobasso, 2006; London & Feigenson, 1979). Recently, a new method based on the direct detection of viral lipids by matrix-assisted laser desorption/ionization–time-offlight/mass spectrometer analysis using 9-aminoacridine as the matrix has been developed (Murphy & Gaskell, 2011; Vitale et al., 2013). This method avoids several drawbacks encountered in the traditional methods such as quantitative and systematic biases in the solvent extraction. The amount of viral material required for the analysis is miniscule compared to the traditional methods, and reliable detection of the minor components of viral lipids is possible (Vitale et al., 2013).

2. FUNCTION AND SIGNIFICANCE OF LIPIDS IN PROKARYOTIC VIRUS LIFE CYCLE Lipids have essential roles in different stages of the viral life cycle (entry, assembly, and exit) if the virus contains lipids as structural components of the virion. Although the exact mechanisms of bacteriophage lipid acquisition still require further research, the role of lipids in the life cycle is well established for PM2, PRD1, and ϕ6, the type species of the bestcharacterized families of lipid-containing bacteriophages, Corticoviridae, Tectiviridae (Section 3.1; Fig. 3), and Cystoviridae (Section 3.2; Fig. 3), respectively (Oksanen, Poranen, & Bamford, 2010). Compared to bacteria, archaea seem to have relatively more lipid-containing viruses, although to date, only approximately 100 archaeal viruses are known (Atanasova

Prokaryotic Viruses with Lipids

19

Figure 3 Sequential steps of the utilization of the viral membranes during the entry and assembly of dsDNA bacteriophage PRD1 with an internal membrane (A–G) and dsRNA phage ϕ6 with an external one (H–O). The host outer membrane (OM), peptidoglycan layer (PG), and cytoplasma membrane (CM) are colored with yellow, gray, and orange, respectively. The phage-specific membranes are in red. (A) PRD1 recognizes the cell envelope-associated DNA transfer complex via the receptor recognition protein P2 at the virion vertices. (B) After the binding, the particle is reorientated on the cell surface triggering the release of the vertex complexes (proteins P2, P5, P31, P16, and peripentonal P3) leading to the transformation of the inner membrane to a membraneous tail tube, which penetrates the cell envelope. The membrane-associated lytic transglycosylase protein P7 digests locally an opening to the peptidoglycan layer. The linear dsDNA genome with terminal proteins is transferred to the cytoplasm through the tail tube. Other proteins involved in the cell envelope penetration are at least P11, P14, P18, and P32. (C) Upon viral protein translation, the membrane-associated phage proteins are addressed to the host cytoplasmic membrane. (D) During the virion assembly, the phage-specific membrane batch at the CM is (Continued)

20

Nina S. Atanasova et al.

et al., 2012; Pietila¨ et al., 2014; Pina, Bize, Forterre, & Prangishvili, 2011). Consequently, detailed information about archaeal virus lipids is scarce, but represents an interesting field of research, as archaeal lipids are structurally very different from those of bacteria and eukaryotes (Section 1.1; Roine & Bamford, 2012; Sprott, 2011). In general, prokaryotic lipidcontaining viruses can be classified into three categories: those that contain an external or internal lipid bilayer, and those that have lipids as protein modifications. Entry is the first step in the virus life cycle in which viral membranes have an essential role. Viruses with an external membrane enter the cell by fusion of the viral membrane with that of the host (Cann, 2005). In the case of dsRNA bacteriophages with segmented genomes and an external membrane, the host membrane participating in the fusion is the outer membrane of the Gram-negative host bacterium (Poranen & Bamford, 2008). Membrane fusion with the cytoplasmic membrane of the host has been suggested for the pleomorphic mycoplasmaviruses (Section 3.3) as mycoplasmas lack the cell wall and the outer membrane (Putzrath & Maniloff, 1977). The entry mechanism is not known for any archaeal virus, although at least pleolipoviruses (Section 3.3) most probably use by membrane fusion (Pietila¨ et al., 2009; Roine & Oksanen, 2011). Some of the inner Figure 3—Cont'd pinched off with the help of phage-encoded nonstructural membrane-bound scaffolding protein P10. The particle assembly is also assisted by the host GroEL–GroES chaperonin complex and phage-encoded soluble assembly factors P17 and P33. (E) The virus capsid-associated proteins are added on the surface of the internal phage membrane enabling the formation of the PRD1 procapsid. (F) The phage genome is packaged by the packaging ATPase P9 through the membrane-bound unique vertex. (G) Mature virions are released upon host cell lysis. (H) The receptorbinding spike protein P3 of phage ϕ6 binds to the pilus receptor that retracts bringing the virion to face the outer membrane. (I) Translocation of P3 exposes the fusogenic protein P6 leading to the fusion of the viral membrane with the host outer membrane. Nucleocapsid (NC) surface protein P5 degrades a local opening to the peptidoglycan layer allowing the virus particle to face the cytoplasmic membrane of the host. (J) The NC surface protein P8 aids the acquisition of the cytoplasma membrane vesicle around the NC releasing the particle to the cell interior. (K) The internal polymerase complex (PC) particle is released from the membrane vesicle and starts transcription. (L) The ϕ6 particle assembly starts when the virus-specific membrane proteins are inserted to the cytoplasma membrane excluding locally the host proteins. (M) The NC associates with the virusspecific membrane, and the particle is release to the host cytoplasm with the aid of nonstructural assembly factor protein P12. (N) The P3 spike proteins are added to the virion surface anchored to protein P6 residing in the viral membrane. (O) Mature virions are released by cell lysis and ready to initiate the next life cycle.

Prokaryotic Viruses with Lipids

21

membrane-containing bacteriophages with linear dsDNA genomes use their membrane to form a tubular structure which serves as a DNA injection apparatus (Bamford & Mindich, 1982; Peralta et al., 2013). This has been well documented for the phages PRD1 and Bam35 which infect Gramnegative and Gram-positive hosts, respectively (Gaidelyte, CvirkaiteKrupovicˇ, Daugelavicˇius, Bamford, & Bamford, 2006; Peralta et al., 2013). Entry mechanisms of prokaryotic viruses with circular genomes are mostly unknown. However, it is considered that the entry of PM2, which has a circular supercoiled dsDNA genome, involves the fusion of the viral membrane with the host outer membrane (Cvirkaite-Krupovicˇ, Krupovicˇ, Daugelavicˇius, & Bamford, 2010; Kivela¨, Daugelavicˇius, Hankkio, Bamford, & Bamford, 2004). Inner membrane-containing viruses with either linear or circular genomes are known to infect archaea, but since the archaeal cell wall structure differs significantly from the bacterial one (Kandler & Konig, 1998), it remains to be seen how archaeal inner membrane-containing viruses enter the cell. Following entry and replication, lipids are incorporated into the new virions by yet mostly unrevealed ways during a complex assembly process or during egress if the virus exits the cell without lysis. This is introduced briefly below followed by more detailed descriptions about the different lipid-containing prokaryotic viruses.

2.1. How prokaryotic viruses acquire their lipids Prokaryotic viruses have no genes for lipid synthesis of their membranes. Thus, lipids are obtained from the host cytoplasmic membranes. This acquisition can result into a somewhat direct copy of the host lipid composition or be more or less selective. Although this selectivity depends on the particular virus, the localization of the lipid bilayer in the virion architecture might give insights into the mode of lipid acquisition. 2.1.1 Viruses with an external membrane Most of the known viruses with an external lipid bilayer (envelope) acquire their lipids as they bud out from the host cell. Thus, their lipid composition greatly resembles that of the host (Kuhn & Rossmann, 2005). Budding is a complex process, which is unique for different viruses, but always includes interactions between the viral integral membrane proteins and the host lipid bilayer proteins (Garoff, Hewson, & Opstelten, 1998). The maturation and release of this type of viruses is often a continuous process, and the viruses are

22

Nina S. Atanasova et al.

able to infect their host cells persistently (Cann, 2005). Among the known prokaryotic viruses, budding is suggested as the mode of exit for pleomorphic archaeal and bacterial viruses (pleolipoviruses, plasmaviruses, and the mycoplasma phage L172; Section 3.3; Dybvig et al., 1985; Pietila¨ et al., 2009; Putzrath & Maniloff, 1977). Because no established/confirmed models of prokaryotic virus budding are available at the moment, practically all the current knowledge considering this exit mechanism relies on eukaryotic virus research and is not discussed here. Another type of prokaryotic virus architecture involving an external lipid membrane is portrayed by ϕ6 and other viruses in the family Cystoviridae (Table 1; Section 3.2; Sarin et al., 2012). This virus morphology is characterized by an enveloped icosahedral nucleocapsid (NC) enclosing the dsRNA genome. So far, such viruses (or any viruses with an RNA genome) have not been observed for archaea. ϕ6-like viruses acquire their lipids after the assembly of the NC. The lipids forming the envelope are selected from the host cytoplasmic membrane (see Section 3.2 for details). 2.1.2 Viruses with a membrane underneath the icosahedral capsid Viruses with a membrane inside the protein capsid are common in the environment. Such bacteriophages infecting Gram-positive bacteria have temperate life styles, while the phages of the Gram-negative bacterial hosts are virulent (Oksanen & Bamford, 2012a). Archaeal inner membranecontaining viruses infect hosts belonging to the two major phyla Euryarchaeota or Crenarchaeaota. Both temperate and virulent life styles have been observed for these viruses (Pina et al., 2011). Virus life cycle has been studied in detail for PRD1. The membrane is derived from the host cytoplasmic membrane in a process where the major capsid protein (MCP) acquires the virus-specific lipid vesicle from the plasma membrane (Mindich, Bamford, McGraw, & Mackenzie, 1982; Rydman, Bamford, & Bamford, 2001). The virus has a unique vertex which is a conduit for the genome translocation into the empty procapsid (Str€ omsten, Bamford, & Bamford, 2003). Viruses are released by lysis of the host cell. The entry and lipid acquisition of PRD1-like viruses infecting Gram-positive bacteria are suggested to occur in a fashion similar to PRD1 (Ravantti et al., 2003). Lipid acquisition is thought to follow similar pathways also for the described archaeal inner membrane-containing viruses (Bamford et al., 2005; Brumfield et al., 2009; Maaty et al., 2006). The lipids of Sulfolobus turreted icosahedral virus (STIV) have been studied in detail and

Prokaryotic Viruses with Lipids

23

are known to be acidic species of crenarchaeal isoprenoid glycerol dialkyl tetraether lipids that are selectively acquired from the host (Maaty et al., 2006). Interestingly, the release of STIV, which infects a crenarchaeal host, involves the formation of the seven-sided pyramidal protrusions into the host cytoplasmic membrane causing cellular lysis (Brumfield et al., 2009). The lipid acquisition of inner membrane-containing viruses is usually selective, which might be due to features such as membrane curvature induced by the icosahedral capsid and/or specific lipid–protein interactions (see Section 3.1 for details; Bamford et al., 2005; Laurinavicˇius, Bamford, & Somerharju, 2007; Laurinma¨ki, Huiskonen, Bamford, & Butcher, 2005). 2.1.3 Viruses with lipids as structural protein modifications In some cases, lipids can be found in viruses only as structural protein modifications with no bilayer structure. Such viruses are rare, and these modifications are most probably incorporated into the viral proteins by host enzymes. These lipid modifications are considered to enable protein– protein interactions facilitating virion assembly (Hruby & Franke, 1993). The most commonly observed lipid modifications in the proteins of animal viruses are myristoylation and palmitoylation, which involve the co- or posttranslational additions of myristate or palmitate fatty acids to the target proteins (Hruby & Franke, 1993). Poliovirus is an example of a eukaryotic nonenveloped icosahedral virus, which contains lipids only as myristoylation of the internal capsid protein (Paul, Schultz, Pincus, Oroszlan, & Wimmer, 1987). In animal viruses, myristoylation has been suggested to be essential in the encapsidation of the viral genome and/or in assembly and is thought to be catalyzed by the host N-myristoyltransferase (Moscufo, Simons, & Chow, 1991). To our knowledge, virions that contain lipids only as modifications of structural proteins have not previously been described for bacteriophages. Archaeal lemon-shaped virus His1 is the only known prokaryotic virus, which lacks a bilayer but contains lipids as modification of the single major structural protein (Fig. 1; Section 3.5; Pietila¨ et al., 2013). However, the nature of the lipid modification is currently unknown (Pietila¨ et al., 2013). It is possible that the crenarchaeal fuselloviruses have similar lipid modifications and lack a bilayer, but it remains to be determined. In addition, lipid-modified spike proteins have been observed for two pleolipoviruses, HGPV-1 and His2, in which the virion is a membrane vesicle (Pietila¨ et al., 2012).

24

Nina S. Atanasova et al.

3. CURRENTLY KNOWN LIPID-CONTAINING BACTERIAL AND ARCHAEAL VIRUSES To date, the number of described bacteriophage isolates is above 6000, whereas there are only some 100 described archaeal viruses (Ackermann & Prangishvili, 2012; Pietila¨ et al., 2014). In spite of this, the observed diversity of virion morphotypes is higher for archaea predicting that novel viral architectures will be discovered in the future. Surprisingly, when taking into account the overall estimated abundance of viruses in the biosphere (above 1031; Suttle, 2005), we have only observed half-a-dozen morphotypes for bacteriophages and about a dozen for the archaeal ones (Fig. 1). The reasonably late discovery of lipids particularly in archaeal viruses has brought surprises and points toward additional diversity. In this chapter, we summarize the information on the lipids in bacterial and archaeal viruses.

3.1. Icosahedral viruses with an inner membrane Before the year 1968, there were no reports on bacteriophages having lipids as their structural component, and archaea were not even recognized as a distinct domain (Woese & Fox, 1977). Bacteriophage PM2 was the first prokaryotic virus for which lipids were observed as structural components of the virion (Camerini-Otero & Franklin, 1972; Espejo & Canelo, 1968b). These phage-infecting Pseudoalteromonas cells were fished out from sea water samples taken from the Pacific Ocean about 1 mile from Vina del Mar, Chile (Espejo & Canelo, 1968a). PM2 is an icosahedral virus with an internal lipid o, Kalkkinen, & Bamford, 1999). bilayer (Kivela¨ et al., 2004; Kivela¨, Ma¨nnist€ Later architecturally similar bacteriophages have been described. To date, it is known that archaeal cells are also infected by viruses that are structurally related to PM2. Currently, there are 23 known icosahedral viruses with an internal membrane of which 17 and 6 infect bacteria and archaea, respectively (Table 1). A common feature of these viruses is that their lipid composition differs from that of their host cell cytoplasmic membrane. All these viruses have a dsDNA genome, which is either linear or circular. The majority of inner membrane-containing viruses with a bacterial host has been classified into the family Tectiviridae, with PRD1 as the type species (Oksanen & Bamford, 2012a). These viruses infect either Gram-negative or Gram-positive bacteria (Table 1). In addition to bacteriophage PM2, which is still the only recognized member of the Corticoviridae family, a thermophilic bacteriophage P23-77 with similar morphology to PRD1 has been

Prokaryotic Viruses with Lipids

25

described for a Thermus host ( Jaatinen et al., 2008; Rissanen et al., 2012). Recently, a highly halophilic bacteriophage, Salisaeta icosahedral phage 1 (SSIP-1), was found from an experimental pond in Israel (unassigned family; Aalto et al., 2012). All these phages are virulent, except those infecting Gram-positive bacteria. The first described archaeal icosahedral virus with an internal membrane was STIV, isolated from an acidic high-temperature hot spring (Rice et al., 2004). A few years later, another crenarchaeal virus, STIV-2 was described (Happonen et al., 2010). STIV-2 is related to STIV and also infects Sulfolobus cells. To date, four viruses, SH1, HHIV-2, PH1, and SNJ1, similar to STIV and STIV-2 are known to infect euryarchaeal hosts ( Jaakkola et al., 2012; Porter et al., 2005, 2013; Zhang et al., 2012). These viruses infect either halophilic Haloarcula or Natrinema strains (Table 1). All the archaeal icosahedral viruses with an internal membrane are virulent, except SNJ1, which is temperate. The classification of these viruses is under discussion (Pawlowski et al., 2014). There is atomic-level information about two inner membranecontaining prokaryotic viruses. The best described is PRD1, isolated 40 years ago from sewage sample taken from Kalamazoo, MI, USA (Olsen et al., 1974). In addition to this, studies on bacteriophage PM2 have provided insights into the assembly of icosahedral viruses with an internal membrane. PRD1 and PM2 virus structures have been solved by X-ray ˚ resolution, respectively, allowing the detailed crystallography at 4 and 7 A analysis of the membranes and the membrane proteins (Abrescia et al., 2004, 2008; Cockburn et al., 2004). The difference in the genome type (linear vs. circular) dictates that the two viruses are bound to have different mechanisms for genome delivery, virus assembly, and genome packaging.

3.1.1 Lipids in the corticovirus PM2 with a circular supercoiled dsDNA genome PM2 is a virulent virus infecting two Gram-negative Pseudoalteromonas species (Kivela¨ et al., 1999; Oksanen & Bamford, 2012b). A significant portion of the available aquatic bacterial genomes contains PM2-like elements indicating that corticoviruses are wide-spread in marine environments (Krupovicˇ & Bamford, 2007). The genome is a highly negatively supercoiled, circular dsDNA molecule of 10,079 bp in length (Espejo, Canelo, & Sinsheimer, 1969; Ma¨nnist€ o et al., 1999). Other icosahedral viruses with an internal membrane and a circular dsDNA genome are STIV,

26

Nina S. Atanasova et al.

STIV-2, SNJ1, P23-77, and SSIP-1 (Table 1). Obviously, unique entry and assembly mechanisms for delivery and packaging of circular molecules have evolved. The viral membrane is most probably involved also in these processes which are rather poorly understood. The diameter of PM2 virion is 57 nm, and its mass is 47 MDa, which is divided to lipid (14%), nucleic acid (14%), and protein (72%; Franklin, Hinnen, Scha¨fer, & Tsukagoshi, 1976). The studies on PM2 virion structure have revealed that the membrane is well ordered following the icosahedral shape of the capsid (Abrescia et al., 2008; Huiskonen, Kivela¨, Bamford, & ˚ , which is Butcher, 2004). The thickness of the PM2 membrane is 29 A ˚ 3 A less than that of PRD1. This might be a consequence of the higher density of ordered transmembrane helices in PM2 (Abrescia et al., 2008; Cockburn et al., 2004). Most of the PM2 structural proteins are associated with the viral membrane (Kivela¨ et al., 2004). Only two proteins, the receptor-binding protein P1 and the MCP P2 form the capsid shell (Abrescia et al., 2008; Huiskonen et al., 2004; Kivela¨ et al., 2004). The other eight structural protein species (P3–P10) occupy about one-third of the viral membrane volume (Kivela¨, Kalkkinen, & Bamford, 2002; Laurinavicˇius et al., 2007; Scha¨fer, Hinnen, & Franklin, 1974). The virion lipid composition (64% PG, 36% PE, and traces of acyl-PG) deviates drastically from that of the host’s cytoplasmic membrane (25% PG, 75% PE; Laurinavicˇius et al., 2007). In addition, the different species of phospholipids are asymmetrically distributed in the membrane leaflets (Laurinavicˇius et al., 2007). Two transmembrane proteins P3 and P6, whose copy numbers are the highest among the PM2 membrane-associated proteins, form a well ordered, planar protein complex lying tangentially to the membrane under the capsid shell (Abrescia et al., 2008). This lattice of these membrane proteins mediates the interaction between the membrane and the icosahedral capsid shell. Similar to all membrane-containing viruses, also PM2 uses its membrane in genome delivery, but in a different way than PRD1 (Section 3.1.4), as the internal membrane of PM2 does not transform into a tail tube. After PM2 has recognized the cell surface receptor via the distal tip of the receptorbinding protein P1, the viral protein shell seemingly dissociates exposing the inner membrane surface (Abrescia et al., 2008; Kivela¨ et al., 2004). It has been proposed that the internal membrane mediates the translocation of the supercoiled genome across the host cell envelope via fusion of the viral membrane with host outer membrane (Kivela¨ et al., 2004). During this entry step, the integral membrane protein P10 most probably interacts with the

Prokaryotic Viruses with Lipids

27

host membrane (Kivela¨, Madonna, Krupovicˇ, Tutino, & Bamford, 2008). The genome penetration through the cytoplasmic membrane is dependent on calcium ions (Cvirkaite-Krupovicˇ, Krupovicˇ, et al., 2010). How is the supercoiled DNA incorporated into the membrane vesicle covered by the protein capsid? It is known that PM2 maturation does not proceed through empty membrane-containing procapsids as is the case for PRD1 (Section 3.1.5). It is established that PM2 DNA is replicated via the rolling circle mechanism in proximity to the host cytoplasmic membrane. However, it is not known, whether the replication is coupled to the o et al., 1999). The majority of DNA encapsidation (Brewer, 1978; Ma¨nnist€ the synthesized virus-specific proteins are associated with the host cell membrane (Datta, Braunstein, & Franklin, 1971). Results from transmission electron microscopy (TEM) experiments have proposed that during PM2 infection the maturation might occur via virus-sized empty vesicles (Dahlberg & Franklin, 1970). Empty vesicles have also been seen in cells infected with a temperature-sensitive PM2 mutant, in which the shift to permissive temperature induces the virion maturation process (Brewer, 1978, 1980) suggesting that empty vesicles are packaged. However, the virion architecture and the interactions seen in the virion X-ray structure (Abrescia et al., 2008) illustrate possible events leading to PM2 virion formation. The membrane proteins might act together with the condensed supercoiled genome nucleating virus assembly by initiating the bending of the membrane vesicle. This eventually would lead to the pinching off of DNA-filled vesicles resulting in coupled membrane morphogenesis, genome encapsidation, and capsid assembly. The highly organized transmembrane proteins (the lattice formed of P3 and P6) on the membrane surface can act as a scaffold for capsid assembly. The new virus particles always appear adjacent to or in association with the host cytoplasmic membrane, which apparently is the place for the virus assembly (Cota-Robles, Espejo, & Haywood, 1968). 3.1.2 Lipids in PRD1 and related viruses Viruses of the Tectiviridae family have a linear dsDNA genome and a proteinrich internal membrane enclosed in an icosahedral capsid shell. The genomes have inverted terminal repeats and a covalently linked 50 -terminal proteins used in the replication and packaging (Bamford, McGraw, MacKenzie, & Mindich, 1983; Ziedaite, Kivela¨, Bamford, & Bamford, 2009). Tectiviruses can be divided into two groups: viruses infecting Gram-negative bacteria and Gram-positive bacteria (Oksanen & Bamford, 2012a). The phages

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infecting Gram-negative bacteria include PRD1, PR3, PR4, PR5, PR722, and L17. They are very closely related having genome sequence identity between 91.9% and 99.8% (Saren et al., 2005; Table 1). These viruses have been isolated from distant locations all over the world, and the most different structural viral proteins are those responsible for host recognition (Saren et al., 2005). In addition, three phages, P37-14, P23-77, and ϕIN93, infecting Thermus strains have been reported, of which P23-77 is rather well described at genomic and structural levels ( Jaatinen et al., 2008; Jalasvuori et al., 2009; Rissanen et al., 2012, 2013). P23-77 and ϕIN93 share some sequence similarity, their genomes are colinear, and the virion proteins of P23-77 and ϕIN93 are 75% similar ( Jalasvuori et al., 2009). The recently isolated phage, KHP30, infecting Helicobacter pylori might also be a virus with an internal membrane (Uchiyama et al., 2013). The group of tectiviruses with Gram-positive Bacillus hosts currently includes Bam35, AP50, Wip1, GILO1, GIL16, and ϕNS11 (Table 1). These phages have also similarity at the nucleotide sequence level (Kan et al., 2013). The virion morphologies of PRD1 and Bam35 are indistinguishable, and the gene order is conserved in their genomes. In addition, their nucleotide identity across the genomes is 42% which, however, is very patchy (Ravantti et al., 2003). The placement of P23-77 (and maybe P37-14) in the family Tectiviridae is controversial, since the genome is circular, while the other tectiviruses have linear genomes (Oksanen & Bamford, 2012a). 3.1.3 Lipids of PRD1 form an icosahedrally ordered membrane The host range of PRD1 is broad including various Gram-negative bacteria such as Salmonella enterica, Escherichia coli, and Pseudomonas aeruginosa. However, the phage infects only strains which harbor an IncP-, IncW-, or IncNincompatibility group conjugative plasmid (Olsen et al., 1974). The genetic system available for PRD1 with a number of suppressor-sensitive virus mutants (Mindich, Cohen, & Weisburd, 1976) has been the key for success making PRD1 one of the best-described virus systems. The extensive genetic, biochemical, and structural approaches to study PRD1 have yielded invaluable knowledge about DNA delivery, virion assembly, and genome packaging. The structure of the PRD1 virus with a diameter of about 65 nm and a mass of about 66 MDa has been determined by X-ray crystallography at ˚ resolution (Abrescia et al., 2004; Cockburn et al., 2004). Fifteen per4 A cent of the virion mass is lipid, the rest being protein (70%) and DNA (15%; Bamford, Caldentey, & Bamford, 1995). The membrane constitutes

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of both protein and lipid in an approximate ratio of 1:1 (Davis, Muller, & Cronan, 1982). It is icosahedral and resides underneath the capsid shell enveloping the dsDNA genome. Half of the 18 PRD1 structural protein species are associated with the membrane. The vertex-stabilizing protein P16 at the fivefold vertices is the only icosahedrally ordered membrane protein which is visible in the electron density map (Abrescia et al., 2004). Laser Raman spectroscopy of PRD1 has shown that the lipids are in the liquid crystalline phase, and the membrane proteins are mainly α-helical (Bamford, Bamford, Towse, & Thomas, 1990; Tuma, Bamford, Bamford, Russell, & Thomas, 1996). The PRD1 membrane is well ordered, and it follows the icosahedral shape of the capsid (Cockburn et al., 2004). It is composed of 52% PE, 43% PG, and 5% CL (Laurinavicˇius, Ka¨kela¨, Somerharju, et al., 2004). The relative lipid composition in the PRD1 membrane is different from the outer or cytoplasmic membranes of the host bacterium S. enterica (for the host: 80% PE, 12% PG, and 8% CL; Laurinavicˇius, Ka¨kela¨, Somerharju, et al., 2004) showing that the virus acquires its lipids selectively upon assembly. The viral membrane comprises 26,000 lipid molecules asymmetrically distributed between the membrane leaflets (Cockburn et al., 2004; Laurinavicˇius et al., 2007). PG and CL are enriched in the outer membrane leaflet, whereas the zwitterionic PE is mainly found in the inner leaflet, facilitating close interactions with the genome (Cockburn et al., 2004; Laurinavicˇius et al., 2007). The enrichment of PG has also been observed in phage PM2 (Section 3.1.1; Laurinavicˇius et al., 2007). The asymmetric distribution may be a consequence of the shapes and charges of different phospholipid molecules and their interactions with the surrounding membrane proteins. In addition, the membrane proteins may affect the lipid composition locally driving the selective uptake of the lipids from the host pool. 3.1.4 PRD1 genome delivery occurs through a membranous tunneling nanotube The internal membrane of PRD1 is capable of forming a tubular structure, which has a central role in the delivery of DNA during infection (Bamford & Mindich, 1982; Grahn, Daugelavicˇius, & Bamford, 2002a, 2002b; Lundstr€ om, Bamford, Palva, & Lounatmaa, 1979; Peralta et al., 2013; Fig. 3A and B). The process is initiated by specific recognition of the cellular receptor (IncP plasmid-encoded DNA transfer complex) by the receptor recognizing fivefold spike protein P2 (Grahn, Caldentey, Bamford, &

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Bamford, 1999; Mindich et al., 1982). The irreversible binding to the cell surface receptor, which is possibly guided by the other fivefold spike protein P5, positions the virus particle to the cell surface and triggers structural changes in the virion. This leads to dissociation of the receptor recognizing fivefold vertex complexes composed of the receptor-binding protein P2, spike protein P5, penton protein P31, and vertex-stabilizing membrane protein P16, as well as the surrounding peripentonal capsid protein P3 trimers from the virion (Peralta et al., 2013). The openings formed at the vertices allow the capsid–membrane interactions to be destabilized, and the membrane vesicle is transformed into a tube-like structure that crosses the cell envelope (Peralta et al., 2013). The tube, which is 4.8 nm thick and 50 nm in length, is capable of penetrating the host envelope (the envelope of Salmonella is 15 nm thick) and protects the genome upon delivery to the cytoplasm. It has been proposed that the PRD1 membrane tube passes through the unique vertex also used for DNA packaging (Hong et al., 2014; Peralta et al., 2013; Str€ omsten, Bamford, et al., 2003). In the packaged virion, the putative interaction between the packaging ATPase P9 and the viral genome via the terminal protein P8 might serve as the nucleation point and facilitate tail tube formation through the vertex (Karhu, Ziedaite, Bamford, & Bamford, 2007; Peralta et al., 2013). However, the packaged genome is not a prerequisite for the tube formation, since the PRD1 procapsids devoid of the genome are still capable of forming the tube (Bamford & Mindich, 1982; Peralta et al., 2013). Cellular tomography of PRD1-infected cells allowed visualization of the entire tail tube penetrating the cell envelope. In most cases, the tail tube entered the cell surface almost orthogonally (Peralta et al., 2013). Several integral viral membrane proteins (P7, P14, P18, and P32) have been shown to be crucial for the formation of the membrane tube (Bamford & Mindich, 1982; Grahn et al., 2002a). In addition, the major membrane protein P11 (the adhesive factor of the membrane) most probably operates after receptor binding and interacts with the outer membrane of the host bacterium (Grahn et al., 2002a). For cell wall digestion, PRD1 uses a membraneassociated peptidoglycan degrading enzyme P7 which is a transglycosylase located in the viral membrane (Rydman & Bamford, 2000, 2002). The visualization of the tube by cryoelectron microscopy revealed different regions with either high or low densities indicating that the membrane proteins might act as a scaffold for the tube (Peralta et al., 2013).

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Once the tail tube has reached the cytoplasm, the viral genome is released most probably through the opening of the tip of the tail tube. The internal diameter of the tail tube is 4.5 nm, which allows one viral dsDNA chain to be translocated into the host cytoplasm (Peralta et al., 2013). The DNA release might be fuelled initially by the energy stored in the pressurized capsid and/or a change in the osmotic pressure (Cockburn et al., 2004; Peralta et al., 2013). 3.1.5 Assembly and packaging of internal membrane-containing bacteriophage PRD1 During PRD1 infection, no transcriptional activation of the host bacterium genes involved in the phospholipid biosynthesis has been recorded (Poranen et al., 2006). The overall changes in the host bacterium during virus infection are almost nonexistent and PRD1 utilizes only a small fraction (5–15%) of the synthesizing capacity of the host cell (Poranen et al., 2006). Fifteen minutes postinfection, a number of virus capsid-associated proteins, such as the MCP P3 trimers, receptor-binding protein P2 monomers, spike protein P5 trimers, and penton protein P31 pentamers are found soluble in the cytoplasm. The virus-encoded membrane proteins such as P7, P14, P11, and P18, on the other hand, are addressed to the host cytoplasmic membrane (Mindich et al., 1982; Fig. 3C). The correct folding of the viral proteins including several membrane proteins is mediated by the host GroEL/GroES chaperonins (Ha¨nninen et al., 1997). Most probably, the membrane proteins are clustered together in the host cytoplasmic membrane, where the lipid–protein interactions might serve as the nucleation point for procapsid assembly (Fig. 3D). The membrane-bound nonstructural scaffolding protein P10 guides the particle assembly, in which the virus-specific membrane area derived from the host cytoplasmic membrane is assembled together with the capsid-associated proteins (Rydman et al., 2001). Protein P30 is located on the surface of the viral membrane and is essential for particle assembly as it defines the size of the icosahedral particle by acting as a molecular tape measure protein (Abrescia et al., 2004; Rydman et al., 2001). In addition, phage-encoded nonstructural assembly factors P10, P17, and most probably P33 have roles in the procapsid formation (Mindich et al., 1982). It has been shown that protein P17, which is a soluble tetramer, is capable of binding to the positively charged lipid membranes (Caldentey et al., 1999; Holopainen, Saily, Caldentey, & Kinnunen, 2000).

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The empty membrane-containing procapsids, which are visible 40 min after infection, contain all structural proteins, except the packaging ATPase P9 and the genome terminal protein P8 (Fig. 3E; Mindich et al., 1982; Str€ omsten, Bamford, et al., 2003). PRD1 has a unique vertex through which the virus genome is packaged powered by the packaging ATPase P9 (Fig. 3F; Gowen, Bamford, Bamford, & Fuller, 2003; Karhu et al., 2007; Str€ omsten, Bamford, et al., 2003; Str€ omsten, Bamford, & Bamford, 2005; Ziedaite et al., 2009; Hong et al., 2014). Other unique vertex proteins are the packaging efficiency factor P6 and two small integral membrane proteins P20 and P22, which together form a conduit mediating the translocation of the viral DNA into the internal membrane vesicle of the procapsid (Hong et al., 2014). During packaging, the internal membrane slightly expands, and the interactions between the capsid and the underlying membrane increase (Butcher, Bamford, & Fuller, 1995; San Martin et al., 2002). In the virion, the curvature of the membrane is the highest underneath the fivefold vertices and at the edges of the facets (Cockburn et al., 2004). At these sites, the membrane is connected to the capsid shell either by the vertex-stabilizing membrane protein P16 at the fivefold vertices or by a series of interactions between the N-termini of the MCP P3 on the facet edges (Abrescia et al., 2004; Cockburn et al., 2004; Jaatinen et al., 2008). In addition to the membrane-anchored unique packaging vertex structure of PRD1, the only available structural information considering a packaging ATPase of an internal membrane-containing icosahedral virus is derived from studies of the archaeal virus STIV-2. The X-ray crystallographic structure of STIV-2 packaging ATPase B204 revealed that it belongs to the FtsK-HerA superfamily of P-loop ATPases, whose cellular and viral members have been suggested to have the same mechanism for DNA translocation (Happonen et al., 2010, 2013).

3.2. Enveloped icosahedral viruses: Phage ϕ6 and its relatives A relatively common bacterial virus morphotype is the one with an internal icosahedrally organized NC surrounded by a lipid envelope. There are some 10 isolates described so far (Table 1; Fig. 1). All this type of viruses have a segmented dsRNA genome (three segments), which resides inside the polymerase complex (PC). These viruses are classified into the family Cystoviridae. The PC is the innermost icosahedral structural element that is matured to a NC upon addition of a protein shell made of protein P8 around the PC. Due to the dsRNA genome, these viruses have to deliver

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their PC into the host cytoplasm as the host bacteria are not capable of replicating dsRNA. Within the cell, the PC particle translocates the transcribed ssRNA genomic segments from the interior of the PC to the cytoplasm where they are either used to guide protein synthesis or, later in the infection cycle, packaged to the newly formed PC particles where minus strand synthesis (replication) takes place. The hosts are Gram-negative bacteria with an outer and cytoplasmic membranes and a rigid peptidoglycan layer in between. The vast majority of information obtained for these viruses comes from the original isolate designated as ϕ6, which infects a plant pathogenic Pseudomonas syringae bacterium (Vidaver et al., 1973). The entire NC must be internalized for progeny production in a process where the viral and host membranes are crucial for penetrating the cell envelope. In this chapter, we follow the ϕ6 membrane throughout the infection cycle (Fig. 3H–O), the topic of this review. 3.2.1 Involvement of the membranes in ϕ6 entry There are receptor-binding spikes on the ϕ6 virion surface that are made of protein P3. These spikes are anchored to the virion through the integral membrane protein P6. For ϕ6, the receptor is the side of a type IV pilus (host pathogenesis factor) extending from the host cell surface (Fig. 3H; Mindich, Sinclair, & Cohen, 1976; Roine, Nunn, Paulin, & Romantschuk, 1996; Roine, Raineri, Romantschuk, Wilson, & Nunn, 1998; Romantschuk & Bamford, 1985, 1986). Other ϕ6-like viruses may use different cell surface structures for attachment. The retraction of the pilus filament brings the virion into contact with the cell surface (Romantschuk & Bamford, 1985; Romantschuk, Olkkonen, & Bamford, 1988). In this process, the P3-spike protein dislocates to expose protein P6 (Fig. 3I). Protein P6 has been shown to have fusogenic activity leading to the fusion of the viral membrane with the host outer membrane (Bamford, Palva, & Lounatmaa, 1976; Bamford, Romantschuk, & Somerharju, 1987; Lounatmaa & Bamford, 1978). This process locates the NC to face the peptidoglycan layer. There is a lytic enzyme, protein P5, associated with the NC surface digesting locally the peptidoglycan layer (Caldentey & Bamford, 1992; Hantula & Bamford, 1988; Kakitani, Iba, & Okada, 1980; Mindich, Lehman, & Huang, 1979; Romantschuk & Bamford, 1981). This positions the NC to face the host plasma membrane. The NC surface protein P8 is membrane active pushing the NC to acquire a plasma membranedriven envelope enclosing the NC in an endocytotic-like event (Fig. 3J; Ojala, Romantschuk, & Bamford, 1990; Olkkonen, Ojala, & Bamford,

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1991; Poranen, Daugelavicˇius, Ojala, Hess, & Bamford, 1999; Romantschuk et al., 1988). How the NC dissociates to release the P8 shell and how the PC escapes from the membrane vesicle and starts transcription are currently less understood than the early stages of entry. It is considered that the membrane-active nature of P8 and pH may be crucial factors in this process (Bamford, Bamford, Li, & Thomas, 1993; Cvirkaite-Krupovicˇ, Poranen, & Bamford, 2010; Huiskonen et al., 2006; Poranen et al., 1999; Tuma, Bamford, Bamford, & Thomas, 1999). 3.2.2 How ϕ6 acquires its membrane envelope The entering PC secretes the first transcripts into the cytoplasm-producing proteins needed for the empty PC assembly (Fig. 3K). Later in the life cycle, there is an increase of synthesis of envelope constituents and protein P8 assuring the maturation of the NC and formation of the viral envelope (Fig. 3L; Poranen, Tuma, & Bamford, 2008). The most crucial components for the membrane assembly are the integral membrane protein P9 that is inserted to the host plasma membrane and an assembly factor protein P12. P9 synthesis defines a virus-specific membrane region from where the host cytoplasmic membrane proteins are expelled (Fig. 3M). With the aid of nonstructural protein P12, a virus-specific vesicle that encloses the NC, is formed. These enveloped viral particles are released to the cell interior where the receptor-binding protein P3 is added to the particle finalizing the virion assembly process (Fig. 3N; Bamford et al., 1976; Ellis & Schlegel, 1974; Gonzalez, Langenberg, Van Etten, & Vidaver, 1977; Johnson & Mindich, 1994; Mindich et al., 1979; Stitt & Mindich, 1983a, 1983b). The acquirement of the membrane from the plasma membrane is an intriguing event due to the release of the virions to the cell interior instead of budding through the plasma membrane. The reason for such an assembly pathway obviously is the presence of the rigid peptidoglycan layer and the outer membrane of the host. The topology of the viral membrane in respect to the NC is particularly intriguing (Stitt & Mindich, 1983b). Either the NC (carrying the external P8 shell) recognizes the P9 batch in the cytoplasmic membrane and inverts to be released to the cell interior or a virus-specific vesicle with P9 and P8 is formed possibly attached to the host cytoplasmic membrane. P8 is in the context of the NC or alternatively, associated with the P9 in the virus-specific vesicle (Sarin et al., 2012). Finally, the NC is internalized to these virus-specific vesicles that are pinched off to release the viral particle to the cell interior. There is morphological electron microscopic support using mutant viruses for the presence of cytoplasmic

Prokaryotic Viruses with Lipids

35

membrane bags with internal NCs (Bamford, 1980). At the end of the life cycle, the virions are released by the host cell lysis (Kakitani et al., 1980; Mindich & Lehman, 1979; Romantschuk & Bamford, 1981). 3.2.3 Lipids of the ϕ6 virion and its host The phospholipid class and molecular species compositions of bacteriophage ϕ6 and its P. syringae host vary considerably pointing to specific mechanism to acquire the lipid (and protein) constituents for the virus. The phage contains significantly more PG and less PE than the cytoplasmic or outer membranes of its host. In addition, the phospholipid molecular species composition of the viral membrane also differs from those of the host membranes. However, it resembles more that of the cytoplasmic membrane than the outer membrane, which is in line with the observation that ϕ6 derives its phospholipids from the host cytoplasmic membrane (Laurinavicˇius, Ka¨kela¨, Bamford, & Somerharju, 2004). The outer leaflet contains approximately equal amounts of PE and PG. In the inner leaflet, PE is considerably enriched (65% PE and 31% PG; Laurinavicˇius et al., 2007). The phospholipid shape and charge as well as the interactions with the membrane proteins are the major selective phenomena. It should be noted that the membrane of ϕ6 is rich in proteins and the curvature and surface areas of the different leaflets are considerably different with some 40% more surface area in the outer leaflet (Laurinavicˇius, Ka¨kela¨, Bamford, et al., 2004). Obtaining and combining additional structural and biochemical data will allow to further refine the biogenesis of the ϕ6 membrane.

3.3. Vesicular pleomorphic viruses The word pleomorphic is commonly used to describe the morphology of enveloped viruses with asymmetric or variable virion architecture. Among the known prokaryotic viruses, L2 and L172 infecting mycoplasmas and pleolipoviruses infecting halophilic archaea, have a spherical, pleomorphic virion architecture (Table 1 and Fig. 1). Plasmavirus L2 has been classified into the family Plasmaviridae, but the other viruses are currently unclassified (Maniloff, 2012). The proposed viral family Pleolipoviridae includes the seven known pleolipoviruses (Pietila¨ et al., 2012) representing related viruses with different genome types (circular ssDNA and circular or linear dsDNA; Table 1 and Fig. 4; Sencˇilo et al., 2012). Interestingly, the genomes of the mycoplasma viruses L2 and L172 are ds and ssDNA, respectively, and the sizes correspond to those described for the genomes of pleolipoviruses (see below; Dybvig et al., 1985; Sencˇilo et al., 2012).

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Figure 4 Archaeal pleolipoviruses. The width of the transmission electron micrographs of the negatively stained virions represents 100 nm. The protein profile of the purified virions analyzed by SDS-PAGE with Coomassie blue and Sudan Black staining is shown on the right. The hypothetical positions of HGPV-1 VP2 and His2 VP32 proteins are marked with rectangles as they do not bind Coomassie Blue. The lipid-modified spike proteins of HGPV-2 and His2 are indicated by an asterisk. Molecular mass standards are shown on the far right.

Transmission electron micrographs of the prokaryotic pleomorphic viruses resemble each other, and the different viruses have roughly similar virion diameters. While pleolipoviruses are known to be decorated by spikes, so far such structural elements have not been observed for the mycoplasma viruses by the negative stain TEM (King et al., 2012; Maniloff, 2012; Pietila¨ et al., 2009, 2012; Roine et al., 2010). However, visualization of such virion surface structures may require additional methods and a higher resolution. The life cycles of all the prokaryotic pleomorphic viruses are nonlytic and persistent (Pietila¨ et al., 2012; Roine & Oksanen, 2011). The only significant difference in the life style is that L2 and L172 integrate in the host chromosome while pleolipoviruses have not been found to have integrases (Putzrath & Maniloff, 1977, 1978; Sencˇilo et al., 2012). However, several proviral pleolipovirus-like elements (plasmids and proviruses) in the genomes of some halophilic archaeal strains have been described (Sencˇilo et al., 2012), indicating that integration may be part of the life style of these viruses. Although the genomes of L172 and L2 are not related, the major structural proteins of both viruses are around 15–20 and 60–70 kDa (Dybvig et al., 1985), similar to those observed for pleolipoviruses. The genomes of pleolipoviruses are not related to plasmavirus L2 (E. Roine,

Prokaryotic Viruses with Lipids

37

personal communication), and no sequence data are currently available for L172. This, however, does not exclude relatedness at the level of virion architecture, which is considered to indicate common ancestry (Abrescia, Bamford, Grimes, & Stuart, 2012; Bamford, 2003). 3.3.1 Asymmetric lipid vesicles as viruses What makes pleolipoviruses special compared to other spherical pleomorphic viruses is the absence of a NC underneath the lipid membrane (Pietila¨ et al., 2009, 2010). In addition, the membrane bilayer of these viruses is covered with spike proteins, which are considered to partly contribute to the pleomorphic nature of the virions. These properties result in the characteristic “floppy,” membrane vesicle-like appearance. Like most of the known haloarchaeal viruses, pleolipoviruses require high NaCl concentrations for infectivity (Pietila¨ et al., 2010, 2012). The described pleolipoviruses (Fig. 4) infect halophilic archaea from the genera Halorubrum, Haloarcula, and Halogeometricum. Like their hosts, these viruses have been isolated from spatially distant hypersaline environments indicating worldwide distribution (Atanasova et al., 2012; Bath et al., 2006; Pietila¨ et al., 2009, 2012; Roine et al., 2010). The first and the best-described pleolipovirus, Halorubrum pleomorphic virus 1 (HRPV-1), was described in 2009 as the first archaeal virus with an ssDNA genome (circular 7048 nt; Pietila¨ et al., 2009). This virus and its host Halorubrum sp. were isolated from an Italian saltern in Trapani, Sicily. The major structural proteins of HRPV-1 are VP4 (spike protein, 53 kDa) and VP3 (integral membrane protein, 14 kDa; Pietila¨ et al., 2009). In addition, the putative ATPase (VP8) was detected as a minor protein component of the virion. The randomly distributed spikes are glycosylated and considered to serve as receptor binding and fusion proteins (Kandiba et al., 2012; Pietila¨ et al., 2010). TLC analysis of viral and host lipids showed that the virus acquires its lipids unselectively from the host (Pietila¨ et al., 2010). The viral lipids are composed of CL, PG, PG-Me, and PGS by MS. The absence of a NC was shown by quantitative biochemical dissociation studies (Pietila¨ et al., 2010). HRPV-1 is a nonlytic virus, which persists in the host cells allowing continuous virus production yielding high titers (Pietila¨ et al., 2009). In 2010, another virus, Haloarcula hispanica pleomorphic virus 1 (HHPV-1), with morphology similar to HRPV-1, was reported. Interestingly, the genome of this virus consisted of dsDNA (circular 8082 bp; Roine et al., 2010). Despite the different genome types, these two viruses were found to have gene synteny and sequence similarity proving their

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relatedness. In addition, HRPV-1 and HHPV-1 were found to be related to pHK2 plasmid of Haloferax sp. and to a proviral element in the genome of Haloferax volcanii (Roine & Oksanen, 2011). It is suggested that pHK2 might be a temperate virus resembling HRPV-1 and HHPV-1 (Roine et al., 2010). The viral lipids of HHPV-1 are similar to those observed for HRPV-1, and the virus acquires them unselectively from the host. The life cycle, protein profile, virion size, and morphology are similar to HRPV-1, but the spike of HHPV-1 is not glycosylated (Roine et al., 2010). A global survey of nine different hypersaline environments resulted in the isolation of four new pleomorphic viruses similar to HRPV-1 and HHPV-1 (Fig. 4; Atanasova et al., 2012). These six viruses as well as His2, which had previously been described as a spindle-shaped virus (Bath et al., 2006) but shown to have sequence similarity and morphological resemblance to HRPV-1 and HHPV-1, were included in comparative studies of virion architecture, life cycles, and genomics (Pietila¨ et al., 2012; Sencˇilo et al., 2012). Virion morphology was studied by cryoelectron microscopy and tomography revealing that all the viruses were roughly spherical and decorated with spikes having virion diameters from 40 to 70 nm (Fig. 4). Virion protein profiles of the new isolates were similar to the previously characterized ones, except that Halogeometricum pleomorphic virus 1 (HGPV-1) contained two membrane proteins and His2 contained two spike proteins. In addition, the spikes of these two viruses were found to be lipid modified (Pietila¨ et al., 2012). The lipid composition of the seven pleomorphic viruses resembles that of their hosts indicating unselective lipid acquisition. All viruses except HGPV-1 contained similar lipids than HRPV-1 and HHPV-1 (see above). In HGPV-1, as well as its host Halogeometricum sp. CG-9, the band corresponding to PGS in TLC was absent. Based on the presence of lipids and the nonlytic life style, it has been suggested that pleolipoviruses acquire their lipids and exit their host cells by budding (Pietila¨ et al., 2012). All the seven pleolipoviruses have a conserved cluster of five genes (including those encoding for the spike and internal membrane proteins) that are related to each other based on synteny and amino acid sequence similarity (Sencˇilo et al., 2012). In addition, it seems that pleomorphic viruses with circular genomes replicate via rolling circle replication, while protein-primed replication is suggested for His2 (Bath et al., 2006). The genomes of HRPV-3 and HGPV-1 have unusual, short single-stranded regions in their genomes suggesting that they might use a different replication strategy (Sencˇilo et al., 2012).

Prokaryotic Viruses with Lipids

39

3.3.2 Bacterial vesicular viruses Acholeplasma laidlawii virus L2 is the type species, and the only approved member, of the genus Plasmavirus in the family Plasmaviridae. Other candidates in the genus are the Acholeplasma phages M1, O1, v1, v2, v4, v5, and v7, but to date, these viruses remain unclassified (Maniloff, 2012). L2 virions are pleomorphic and approximately 80 nm in diameter. The viral genome is 11,965-bp circular dsDNA and contains a putative integrase. The virus is able to establish lysogeny with the host, A. laidlawii (strain Ja1; Dybvig & Maniloff, 1983). The productive virus life cycle is nonlytic and persistent retarding the host growth rate. After 6 h postinfection, the virus is able to persist in the host cells, which in turn can give rise to new carrier cell clones or cells that are resistant to the virus (Putzrath & Maniloff, 1977). In the lysogenic state, L2 integrates into a unique site in the host genome (Dybvig & Maniloff, 1983). L2 was isolated in England by washing the host cultures with phosphatebuffered saline (Gourlay, 1971). Virus production of infected cells is increased almost threefold by ultra violet radiation or mitomycin C (Putzrath & Maniloff, 1978). L2 acquires its lipids, which form the outer membrane of the virion, unselectively from A. laidlawii. It is considered that L2 obtains its lipid envelope during budding from the host cell (Al-Shammari & Smith, 1981). The L2 virion contains major structural proteins of 19, 58, 61, and 64 kDa, but additional minor protein species have been observed (Maniloff, 2012). L172 is the first described enveloped phage with an ssDNA genome (Dybvig et al., 1985). The virus was isolated from A. laidlawii strain S2 in the former Czechoslovakia (Liska, 1972). The presence of an envelope was first suggested based on sensitivity to detergents and organic solvents (Gourlay, Wyld, Garwes, & Pocock, 1979). Furthermore, the lipids were studied by TLC and confirmed to be unselectively acquired from the host (Al-Shammari & Smith, 1981). L172 viral lipids consist of phospho-, glyco-, and phosphoglycolipids and the fatty acid composition resembles that of the host (Al-Shammari & Smith, 1981). The virus is temperate and able to establish lysogeny. The productive infection cycle is nonlytic, and the virus exits the host presumably by budding. The 14.0-kbp circular ssDNA genome of L172 shows no homology to the dsDNA genome of L2 (Dybvig et al., 1985). However, the methods used at that time (hybridization) do not allow detailed comparison of sequence data. Virion size (60–80 nm), morphology, nonlytic life style, and structural protein profiles indicate that the two viruses may be related.

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3.4. Prokaryotic viruses with helical symmetry: With or without a membrane Viruses exhibiting helical virion symmetry are infecting both bacterial and archaeal hosts (Fig. 1 and Table 1). Bacterial helical viruses belong to the family Inoviridae, whereas archaeal ones are classified into two families: Lipothrixviridae and Rudiviridae, which are grouped into a single-order Ligamenvirales (Day, 2012; Prangishvili, 2012a, 2012b; Prangishvili & Krupovicˇ, 2012). Viruses belonging to these families have either filamentous or rod-shaped virions essentially containing a proteinaceous core and an ss or dsDNA genome (Day, 2012; Prangishvili, 2012a, 2012b). Of the three families, only the members of Lipothrixviridae have lipid constituents in the virion (Prangishvili, 2012b). 3.4.1 Lipothrixviruses: Helical viruses with a membrane envelope Up to date, a total of 12 lipothrixviruses infecting hyperthermophiles belonging to the genera Sulfolobus, Acidianus, and Thermoproteus have been reported (Bize et al., 2008; Ha¨ring, Vestergaard, Brugger, et al., 2005; Prangishvili, 2012b; Vestergaard et al., 2008; Table 1). Similarly to the majority of other hyperthermophilic archaeal viruses, lipothrixviruses of archaea in the genera Sulfolobus and Acidianus do not cause host cell lysis and are thought to establish a persistent infection (Arnold et al., 2000; Bettstetter et al., 2003; Bize et al., 2008; Ha¨ring, Vestergaard, Brugger, et al., 2005; Vestergaard et al., 2008). On the contrary, viruses infecting hosts belonging to the genus Thermoproteus have been suggested to be temperate (Zillig et al., 1988). Lipothrixvirus particles are flexible filaments minimally consisting of genomic DNA covered by a proteinaceous inner core and surrounded by an outer lipid membrane (Prangishvili, 2012b). Based on the details in virion organization, lipothrixviruses are divided into three established and one proposed genera: Alpha-, Beta-, Gamma-, and “Deltalipothrixviruses” (Prangishvili, 2012b). The genomes of lipothrixviruses are linear dsDNA molecules ranging from approximately 14 kb (TTV1) to 56 kb (DAFV; Neumann et al., 1989; Zillig et al., 1994). The nature of the genome termini is not clear, but it has been hypothesized that the termini may have an unknown chemical modification or may be associated with proteins (Arnold et al., 2000; Bettstetter et al., 2003; Bize et al., 2008; Ha¨ring, Vestergaard, Brugger, et al., 2005; Janekovic et al., 1983; Vestergaard et al., 2008). Lipothrixvirus

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genomes have modular organization shaped by horizontal gene transfer (HGT) between viruses within and outside the Lipothrixviridae family. Despite HGT obscuring the border between families and genera, a phylogenetic tree built based on the conserved hypothetical protein shared among all sequenced lipothrixviruses except for TTV1 verified the designation of lipothrixviruses into the existing genera (Vestergaard et al., 2008). 3.4.1.1 Alphalipothrixviruses

Alphalipothrixvirus Thermoproteus tenax virus 1 (TTV1) is one of the earliest and the best-characterized archaeal viruses ( Janekovic et al., 1983; Table 1). TTV1 particles are approximately 400 nm long and 40 nm wide filaments. Virion inner core is composed of genomic DNA superhelix wound around a central cavity and covered by DNA-binding proteins P1 and P2. Unlike other lipothrixviruses, TTV1 has an additional layer between the inner core and the outer lipid membrane. This layer consists of the helically arranged protein P3 delimited by the cap structures made of the protein P4. It has been suggested that this proteinaceous layer causes greater TTV1 virion rigidity compared to other lipothrixviruses (Arnold et al., 2000; Bize et al., 2008; Janekovic et al., 1983; Vestergaard et al., 2008; Zillig et al., 1988). TTV1 outer membrane was shown to have qualitatively the same, but quantitatively different lipid species than its host (Zillig et al., 1988). 3.4.1.2 Betalipothrixviruses

Betalipothrixvirus genus has eight members: T. tenax viruses 2 and 3 (TTV2 and TTV3), Desulforolobus ambivalens filamentous virus (DAFV), Acidianus filamentous viruses 3, 6, 7, 8, and 9 (AFV3, AFV6, AFV7, AFV8, and AFV9), and the type species Sulfolobus islandicus filamentous virus 1 (SIFV; Table 1). Virions are on average 2000 nm long and 25 nm wide filaments typically having tapered ends terminating with different number of fibers (Arnold et al., 2000; Bize et al., 2008; Janekovic et al., 1983; Vestergaard et al., 2008; Zillig et al., 1994). Particles of SIFV and AFV3 viruses have been studied in more detail, and the proposed models for their structures are in good agreement (Arnold et al., 2000; Vestergaard et al., 2008). According to the models, virion inner core has a nucleosome-like structure composed of two major structural proteins forming doughnutshaped subunits arranged in a zipper-like array with the genomic DNA wrapped around them. This model is fundamentally different from the suggested structure of TTV1 virion inner core. Both SIFV and AFV3 virions have fairly similar lipid constituents as their hosts (Arnold et al., 2000;

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Vestergaard et al., 2008). However, one lipid species found in S. islandicus is more abundant in the virus than in the host. In addition, there seems to be one modified host lipid species in the SIFV envelope, which is suggested to result from glycosylation of host lipids by virus-encoded glycosyl transferases (Arnold et al., 2000). 3.4.1.3 Gammalipothrixviruses and deltalipothrixviruses

Gammalipothrixvirus genus is represented by a single virus—Acidianus filamentous virus 1 (AFV1). AFV1 virions are 900 nm long and 24 nm wide filaments with claw-like structures at both termini (Table 1 and Fig. 5). The claw-like structures are connected to the main viral body (inner core and outer envelope) via narrow appendages with collars. As determined by TLC, one of the host lipids was absent from AFV1 particles, and the majority of the other lipids had different mobilities arguing that the viruses might modify the acquired host lipids (Bettstetter et al., 2003). One more proposed genus belonging to Lipothrixviridae is “Deltalipothrixvirus,” which includes a single isolate—Acidianus filamentous virus 2 (AFV2). Although AFV2 shares morphological similarity and

Figure 5 Lipothrixviruses. Electron micrographs of negatively stained virions of Acidianus filamentous virus 1 (AFV1) from the genus Gammalipothrixvirus. The bars represent 100 nm. Used with the permission from David Prangishvili and the International Committee on Taxonomy of Viruses (ICTV).

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a number of homologues with other lipothrixviruses, no lipids have been detected in AFV2 particles (Ha¨ring, Vestergaard, Brugger, et al., 2005).

3.5. Lemon-shaped viruses are specific for archaea Virus-like particles (VLPs) resembling lemons are abundant in both hyperthermic and hypersaline environments, where archaea typically dominate. Lemon- or spindle-shaped VLPs have also been observed in samples from freshwater sediments and an Antarctic lake (Borrel et al., 2012; Lo´pezBueno et al., 2009; Rachel et al., 2002; Santos et al., 2007; Sime-Ngando et al., 2011). So far, such viruses are archaea specific as no spindle-shaped virus has been described to infect bacteria or eukaryotes (King et al., 2012). Although spindle-shaped viruses share the lemon-shaped core, they can be divided into three groups based on the tail structure: (i) one short, (ii) long tail attached to one of the pointed ends, or (iii) one long tail attached to each end (King et al., 2012). 3.5.1 Viruses with one short tail His1 is so far the only spindle-shaped virus isolated from hypersaline environments (Bath & Dyall-Smith, 1998). His1 virions have a very short tail at one end, and the particles are flexible and their size may vary (Fig. 6A; Bath & Dyall-Smith, 1998; Pietila¨ et al., 2013). Although this flexibility may indicate the presence of a membrane, chloroform–methanol extraction and TLC have shown that the virions contain no free lipids. However, the MCP of His1, VP21, exists in two forms in the virion and one of them seems to be lipid modified (Fig. 6B). Thus, this modification may contribute to the plasticity of the virion (Pietila¨ et al., 2013). In addition to the MCP, a few minor structural proteins of His1 virions have been identified (Pietila¨ et al., 2013). Recently, the first DNA ejection study was performed for an archaeal virus using His1 as a model. It was concluded that the genome ejection from His1 virions to host cells is dependent on host factors as the ejection triggered by detergents was randomly paused and incomplete (Hanhija¨rvi, Ziedaite, Pietila¨, Haeggstr€ om, & Bamford, 2013). His1 resembles morphologically other short-tailed spindle-shaped viruses, but it is the only one described to have a linear dsDNA genome and to encode a putative, type-B DNA polymerase for protein-primed replication (Bath et al., 2006). Consequently, His1 is currently classified into a floating genus, Salterprovirus (King et al., 2012). The majority of short-tailed spindle-shaped viruses infect hyperthermophilic crenarchaea and are classified into the viral family Fuselloviridae, whose type species is Sulfolobus

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Figure 6 Morphology and protein composition of spindle-shaped virus His1. (A) Transmission electron micrographs of purified His1 virions which have been negatively stained with uranyl acetate (upper panel) and ammonium molybdate (lower panel). Arrows indicate large-sized particles. Ammonium molybdate results in highly elongated particles. Scale bar represents 100 nm. (B) Protein profiles of purified His1 virions in a tricine-SDS-polyacrylamide gel stained with Coomassie blue for proteins (left panel), with Sudan Black for lipids and lipoproteins (middle panel), or first with Sudan Black and then with Coomassie blue (right panel). St indicates molecular mass markers (VP, for virion protein).

spindle-shaped virus 1 (SSV1; King et al., 2012). Fuselloviruses have circular dsDNA genomes of similar size and they share gene synteny as well as significant nucleotide sequence similarity (Redder et al., 2009). SSV1 virions contain one major structural protein, VP1, and three minor ones (Menon et al., 2008; Reiter, 1987). SSV1 VP1 and His1 VP21 are similar at the amino acid level indicating that His1 may be related to fuselloviruses despite their different genome types (Pietila¨ et al., 2013). Interestingly, SSV1 VP1 has been reported to form two bands in a gel similar to His1 VP21 (Pietila¨ et al., 2013; Reiter, 1987). Like His1 virions, fuselloviral virions are rather flexible, but the presence of a lipid membrane is controversial (Martin et al., 1984; Redder et al., 2009). However, as is the case in His1 virions, the MCP of SSV1 may be lipid modified, explaining the two bands in the gel and the virion flexibility. A lemon-shaped VLP isolated from a hyperthermophilic euryarchaeon, Pyrococcus abyssi virus 1 (PAV1), resembles morphologically fuselloviruses

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and shares their genome type (Geslin et al., 2003, 2007; Gorlas et al., 2012). However, they have no significant sequence similarity, and thus PAV1 remains unclassified (Geslin et al., 2007; Gorlas et al., 2012). SSV1 and His1 virions and PAV1 particles are composed of one MCP with two hydrophobic domains (Geslin et al., 2003, 2007; Pietila¨ et al., 2013; Reiter, 1987). Thus, all these MCPs may have the same fold derived from a common ancestor and be lipid modified. 3.5.2 Viruses with one or two long tails In addition to spindle-shaped viruses with a short tail, virus isolates with either one or two long tails have been described. These viruses infect hyperthermophilic crenarchaea (Mochizuki, Sako, & Prangishvili, 2011; Prangishvili et al., 2006; Xiang et al., 2005). Sulfolobus tengchongensis spindle-shaped virus 1 (STSV1) has one long tail and is the largest spindle-shaped virus described so far (Xiang et al., 2005). However, the tail length varies and also two-tailed forms have occasionally been observed. TLC analysis indicates that the virion contains lipids which are selectively acquired from the host cell membrane (Xiang et al., 2005). Very recently, STSV2 was described, with 80% protein sequence identity to STSV1. Similarly to STSV1, the virus contains selectively acquired lipids, and the presence of a membrane has been suggested (Erdmann et al., 2014). So far, only one spindle-shaped virus with two long tails has been isolated. Acidianus two-tailed virus (ATV) is classified into the viral family Bicaudaviridae (King et al., 2012). The extraordinary feature of this virus is the capability to form the tails outside the host cells (Ha¨ring, Vestergaard, Rachel, et al., 2005; Prangishvili et al., 2006). It has been proposed that STSV1 may belong to the same viral family as ATV due to their similar appearance (Pina et al., 2011). Furthermore, ATV and STSV1 share nine homologous genes (Krupovicˇ, White, Forterre, & Prangishvili, 2012). In addition, the MCP of ATV with a unique four-helix bundle fold shares significant sequence similarity with STSV1 MCP for which no structure is available (Goulet et al., 2010; Krupovicˇ et al., 2012; Prangishvili et al., 2006; Xiang et al., 2005). However, ATV has several major structural proteins while STSV1 has only one, and no lipids were obtained from ATV virions after chloroform–methanol extraction (Prangishvili et al., 2006; Xiang et al., 2005). Like spindle-shaped viruses with a short tail, long-tailed isolates are nonlytic (Xiang et al., 2005). However, compared to short-tailed viruses,

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long-tailed ones have clearly larger genomes and virions (Mochizuki et al., 2011; Prangishvili et al., 2006; Xiang et al., 2005). Thus, both viruses seem to form their own groups.

3.6. Archaeal spherical viruses with helical NCs have an envelope In the beginning of the new millennium, several viruses with most intriguing, previously unknown morphologies were described infecting crenarchaeal hyperthermophiles. Not only did these viruses look unusual, but they also had gene sequences giving no matches in the public databases (Prangishvili & Garrett, 2004; Rachel et al., 2002). While some of these unique viral morphotypes (such as droplet- or bottle-shaped viruses) were never subjected to lipid analyses, Pyrobaculum spherical virus (PSV), the type species of the genus Globulovirus in the family Globuloviridae, is known to be surrounded by an outer lipid membrane which consists of selectively acquired host lipids (Ha¨ring et al., 2004). In addition to PSV, another globulovirus, Thermoproteus tenax spherical virus 1 (TTSV1) which morphologically resembles PSV, has been described (Ahn et al., 2006). The membrane envelope of globuloviruses encloses a helical nucleoprotein particle with linear dsDNA. The enveloped virions of PSV are 100 nm in diameter and contain variable numbers of spherical protrusions of about 15 nm in length. The buoyant density in CsCl is around 1.3 g ml1 which is typical for lipid-containing viruses. Interestingly, during isopycnic centrifugation, part of the PSV virions are disrupted which causes the release of the helical nucleoprotein core. The genome of PSV is a 28,337-bp long linear dsDNA molecule with inverted terminal repeats. The majority of the putative genes reside on one DNA strand (Ha¨ring et al., 2004). The major structural protein of the virus is 33 kDa in size, and the two minor ones have a mass of 16 and 20 kDa (King et al., 2012). The virion proteins show no matches to public data bases but have some degree of similarity to those of TTSV1. The infection of PSV is nonlytic and persistent. Viral lipids have been extracted by chloroform– methanol and analyzed by TCL. Two of the lipids have identical mobilities to those of the host and two are different indicating selective acquisition and modification of host lipids. PSV and its host Pyrobaculum sp. D11 were isolated from a neutral hot spring (85 °C) in Yellow Stone National Park (Ha¨ring et al., 2004). DNA metagenomic sequences related to PSV were detected in the same environment a few years later (Schoenfeld et al., 2008). In addition to Pyrobaculum sp., PSV infects a T. tenax strain, which

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is closely related to the original host. Regardless of the strict anaerobic nature of the hosts, the virions are stable when stored in the normal atmosphere (Ha¨ring et al., 2004). TTSV1 (Ahn et al., 2006) was isolated from the enrichment culture of the host T. tenax strain YS44 which was obtained from Indonesian hot springs in Siteri. No lipid analysis has been performed for TTSV1, but the morphology, which resembles that of PSV, indicates that the 70-nm virion is surrounded by a lipid envelope. Transmission electron micrographs of TTSV1 include virus particles which are aggregated as chains. The density of the virion is 1.29 g ml1 indicating presence of lipids. The genome of the virus is a linear dsDNA molecule of 21.6 kbp with no covalently attached proteins (Ahn et al., 2006). The viral life cycle is persistent and nonlytic, and the virus does not integrate into the host cell genome. The virus has one major structural protein of 10 kDa and two minor ones of 20 and 25 kDa in mass. The putative coding DNA sequences of TTSV1 are somewhat similar to those of PSV but have no matches in public databases.

4. CONCLUSIONS Prokaryotic viral lipids are derived from the cytoplasmic membranes of their host organism. Consequently, bacteriophage lipids are significantly different from those found in archaeal viruses. The main differences lay in the structure of the core lipids, which for bacteria (and eukaryotes) are based on fatty acid chains ester linked to glycerol and for archaea isoprenoid side chains ether linked to the glycerol backbone. Depending on the virion structure, derivatives of these core lipids are either incorporated into the virion during assembly of the icosahedral procapsids (internal membranecontaining viruses) or translocated over a preassembled icosahedrally symmetric particle (enveloped bacterial dsRNA viruses) as well as budding through the plasma membrane (enveloped bacterial and archaeal DNA viruses). Head-tailed icosahedral, icosahedral inner membrane-containing, and pleomorphic viral morphologies, of which the latter two contain lipids, are shared among bacterial and archaeal viruses (Fig. 1). Interestingly, the selectivity of lipid acquisition from the host seems to be dependent on virus architecture. All known icosahedral inner membrane-containing and enveloped dsRNA prokaryotic viruses take up their lipids selectively, while the pleomorphic viruses have close or almost identical lipid composition when compared to that of their host cytoplasmic membrane.

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Although lipid-containing bacteriophages have been studied already for several decades, the exact mechanism for lipid acquisition remains a challenge. The best-characterized model viruses are PM2, PRD1, and ϕ6. The viral membrane acquisition of these viruses has opposite end points (Fig. 3). The PM2 and PRD1 membranes lie underneath the protein capsid while in ϕ6 the membrane surrounds the NC (Fig. 3). This implies that the mechanisms of membrane acquirements must also be very different. In the case of these three viruses, the membrane is used as the genome delivery device. In PM2 (circular supercoiled dsDNA genome), the protein capsid dissociates and the membrane is suggested to fuse with the host outer membrane. In PRD1, which has a linear dsDNA genome, the membrane is converted into a tail tube that has all the activities needed to penetrate the host cell envelope. ϕ6 (segmented dsRNA genome inside the NC), on the other hand, uses a unique mechanism where all three membranes (viral membrane, OM, and CM) are involved. All these viruses translocate their progeny virions into the host cell cytoplasm prior to lysis. Due to the short history of archaeal virus research, no such models as above are available, and so far, a detailed comparison is not possible. In addition to the previously mentioned morphotypes that are shared by bacterial and archaeal viruses, archaea are infected by viruses with unique morphotypes, of which many contain lipids. However, for some of these viruses, the lipid status is unknown. An abundant group of archaeal viruses is the lemon-shaped ones with variable tail structures (exemplified by His1 with one short tail, STSV1 with one long tail, and ATV with two long tails; Fig. 1). Fuselloviruses and His1 seem to group together, whereas the long-tailed lemon-shaped viruses might form their own cluster. Currently, STSV1 is the only lemon-shaped virus for which lipids have been detected by TLC. ATV has been considered to be related to STSV1, but does not seem to contain lipids. His1 on the other hand contains lipids only as protein modifications. Similar observations have been made among the helical archaeal viruses. The lipid-containing lipothrixviruses are suggested to have a common origin with rudiviruses, for which no lipids have been detected. This is reminiscent to PRD1-type of viruses (double-beta barrel MCP fold), which either do or do not contain a lipid membrane. In the case of pleolipoviruses and plasmaviruses, for which no rigid protein capsid exists, the membrane also serves to protect the naked viral genome. This type of viruses thus depends on the presence of the membrane establishing the virion.

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Viruses can be classified into structure-based viral lineages according to the conserved MCP fold and the architectural principles of the virion. The core elements related to virion structure and assembly are conserved and thus can be applied to study virus evolution, while functions related to interactions with the host are in constant change. To date, four structure-based viral lineages have been established and several others are being proposed. These lineages contain structurally related viruses that infect organisms from all three domains of life and are considered to have a common origin. The presence of the membrane is not coupled to the virion architecture and viruses in the same structure-based lineage may or may not have lipids. However, viruses with the membrane vesicle architecture (pleolipoviruses and plasmaviruses) always contain lipids as they lack the proteinaceous capsid.

ACKNOWLEDGMENTS We thank Professor Angela Corcelli for critically reading the manuscript. This work was supported by Academy Professor (Academy of Finland) funding grants 255342 and 256518 (D. H. B.). We thank Academy of Finland (grants 271413 and 272853) and University of Helsinki for the support to EU ESFRI Instruct Centre for Virus Production. A. S. is a University of Helsinki Fellow in the Doctoral Program in Microbiology and Biotechnology.

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CHAPTER TWO

Innate Recognition of Alphaherpesvirus DNA Stefanie Luecke*, Søren R. Paludan†,{,1 *Graduate School of Life Sciences, Universiteit Utrecht, Utrecht, The Netherlands † Department of Biomedicine, Aarhus University, Aarhus, Denmark { Aarhus Research Center for Innate Immunology, Aarhus University, Aarhus, Denmark 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Alphaherpesviruses 1.2 Immunity to alphaherpesviruses 1.3 Innate DNA sensing 2. DNA Sensors 2.1 TLR9 2.2 Discovery of intracellular DNA sensors 2.3 DAI 2.4 AIM2 2.5 IFI16 2.6 cGAS 2.7 RNA Pol III and RIG-I 3. Accessibility of Viral DNA to DNA Sensors 4. Evasion of DNA-Induced Signaling 5. Relevance for Vaccine Design 6. Conclusions and Future Perspective References

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Abstract Alphaherpesviruses include human and animal pathogens, such as herpes simplex virus type 1, which establish life-long latent infections with episodes of recurrence. The immunocompetence of the infected host is an important determinant for the outcome of infections with alphaherpesviruses. Recognition of pathogen-associated molecular patterns by pattern recognition receptors is an essential, early step in the innate immune response to pathogens. In recent years, it has been discovered that herpesvirus DNA is a strong inducer of the innate immune system. The viral genome can be recognized in endosomes by TLR9, as well as intracellularly by a variety of DNA sensors, the best documented being cGAS, RNA Pol III, IFI16, and AIM2. These DNA sensors use converging signaling pathways to activate transcription factors, such as IRF3 and NF-κB, which induce the expression of type I interferons and other inflammatory cytokines Advances in Virus Research, Volume 92 ISSN 0065-3527 http://dx.doi.org/10.1016/bs.aivir.2014.11.003

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and activate the inflammasome. This review summarizes the recent literature on the innate sensing of alphaherpesvirus DNA, the mechanisms of activation of the different sensors, their mechanisms of signal transduction, their physiological role in defense against herpesvirus infection, and how alphaherpesviruses seek to evade these responses to allow establishment and maintenance of infection.

1. INTRODUCTION 1.1. Alphaherpesviruses The Herpesviridae family comprises more than 130 virus species, which infect mammals, birds, and reptiles. Herpesviruses are enveloped, double-stranded DNA viruses (Fig. 1A), which usually lyse productively infected cells and

Figure 1 Alphaherpesvirus structure and entry. (A) Structure of the alphaherpesvirus virion. The linear dsDNA genome is surrounded by an icosahedral capsid. Associated with the vertices of the capsid are the inner tegument proteins, which are surrounded by the outer tegument. The lipid bilayer envelope contains multiple viral glycoproteins. (B) Schematic illustration of alphaherpesviral cell entry, nuclear DNA delivery, and exposure to DNA sensors. The virion attaches to the cell surface through interaction of the glycoproteins with cellular receptors. The virus can enter the cell via two pathways: by direct fusion of the viral envelope with the plasma membrane or by endocytosis and subsequent fusion of envelope and endosomal membrane. The capsid is transported along microtubules toward the microtubule organizing center (MTOC) near the nucleus. The capsid then docks at a nuclear pore, and the viral genome is released into the nucleus. In the nucleus, the viral genome circularizes and lytic or latent infection takes place. Viral DNA is exposed to endosomal DNA sensors via endosomal entry or by autophagocytic delivery of cytosolic capsids to endosomes, to cytosolic sensors by proteasomal degradation of the capsid, and to nuclear sensors by nuclear DNA delivery.

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establish latent infections in their hosts. Eight herpesviruses are known to cause disease in humans, most notably in children and immunocompromised individuals. The family is divided into three subfamilies, alpha-, beta-, and gamma-herpesviruses. This division was originally based on biological similarities and later confirmed by genome sequencing (Pellett & Roizman, 2013). Alphaherpesviruses are characterized by a relatively broad host range and fast reproduction during lytic infection compared to other herpesviruses (Pellett & Roizman, 2013). They contain three virus species pathogenic to humans (herpes simplex virus type 1 (HSV-1, also called human herpesvirus 1), herpes simplex virus type 2 (HSV-2, also called human herpesvirus 2), and varicella zoster virus (VZV, also called human herpesvirus 3)) and many pathogens of veterinary importance, notably Marek’s disease virus (MDV, also called gallid herpesvirus 2) and pseudorabies virus (PRV, also called suid herpesvirus 1). Herpes simplex viruses are ubiquitous human pathogens, which lytically infect epithelial cells of mucosal surfaces and the skin and afterward establish latency in the cell bodies of peripheral sensory neurons innervating the infected area. During latency, the viral genome persists in the nucleus of the host while the majority of viral genes are silenced. It can be maintained for the lifetime of the host organism. Reactivation from latency can be induced by a number of factors, including tissue damage, UV radiation, immune status changes, and stress, but can also occur spontaneously. In reactivation, virus particles produced in the neuronal cell body are transported along the axons by anterograde transport and infect epithelial cells again. HSV-1 usually causes recurrent cold sores at the lips (herpes labialis), while HSV-2 is often responsible for genital herpes infections (herpes genitalis). HSV may also spread to the central nervous system, leading to herpes encephalitis, cause disseminating herpes infections in neonates and immunocompromised individuals, and infect the eyes, often leading to blindness (Roizman, Knipe, & Whitley, 2013). VZV is the causal agent of chicken pox after primary infection and of shingles (herpes zoster) upon reactivation. In primary infection, the virus usually infects epithelial cells in the upper respiratory tract followed by infection of T cells in lymphoid tissues. This allows for transport of the virus to skin areas over the whole body, where viral replications result in the characteristic rash. VZV then establishes latency in peripheral sensory neurons. The virus can reactivate to cause shingles, which may be associated with serious neurological complications (Arvin & Gilden, 2013).

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PRV first infects epithelial cells of the respiratory tract and then establishes latency in sensory neurons. The natural hosts for this virus are pigs, but it can also infect many other mammals such as cattle, sheep, and dogs. It is the causative agent of Aujeszky’s disease, which is characterized by symptoms ranging from high mortality and severe central nervous system defects in suckling piglets, via abortions and stillbirths in pregnant sows, to fever and respiratory symptoms in adult pigs (Pomeranz, Reynolds, & Hengartner, 2005). MDV establishes latency in and causes oncogenic transformation of immune cells, especially CD4+ T cells, and lytically infects epithelial cells of inner organs and the skin in chickens, leading to a variety of clinical symptoms including chronic polyneuritis and visceral lymphoma (Osterrieder, Kamil, Schumacher, Tischer, & Trapp, 2006). Herpesviruses enter their host cells by fusion of the viral lipid envelope with the plasma or endosomal membrane, which is initiated by interaction of viral glycoproteins in the virion envelope with cell surface receptors. Upon viral entry, the tegument proteins, located between the viral envelope and the capsid, are released into the cytoplasm and interact with host factors to create a favorable environment for the virus. In permissive cells, i.e., cells that allow productive replication of the virus, the icosahedral capsid containing the viral dsDNA genome moves along the microtubule network toward the nucleus, where the viral DNA is translocated through the nuclear pores. In the nucleus, the viral genome circularizes (Fig. 1). Silencing of viral gene expression can lead to the establishment of latent infection. During lytic infection, expression of viral genes takes place in three stages, immediate-early (responsible for subsequent viral gene expression and immune evasion), early (replication of viral DNA), and late (formation and release of progeny virions), and eventually leads to multiplication of the viral genome and to assembly and release of new virus particles (Arvin & Gilden, 2013; Roizman et al., 2013). However, some cells, such as macrophages in case of HSV-1, are nonpermissive for the virus, i.e., productive replication does not take place. Even in permissive cells, not all viral particles lead to productive infection. It is suspected that this is partly due to the early innate antiviral response of the cells (Paludan, Bowie, Horan, & Fitzgerald, 2011).

1.2. Immunity to alphaherpesviruses As with most infectious diseases, immunity to alphaherpesviruses relies on innate and adaptive immune responses. One of the first responses upon

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detection of a herpesvirus in an infected cell is the production and secretion of interferons (IFNs), especially type I (IFNα and IFNβ), but also type III (e.g., IFNλ), and other proinflammatory cytokines and chemokines (Ank et al., 2008; Egan, Wu, Wigdahl, & Jennings, 2013). By paracrine and autocrine signaling, IFNs mediate a number of antiviral activities in infected and neighboring cells by the induction and expression of a variety of genes (interferon-stimulated genes, ISGs) (Egan et al., 2013). IFNs induce the activation of a number of cells of the innate immune system, which are attracted to the site of infection by chemokines. Natural killer (NK) cells cause apoptotic cell death in infected cells, macrophages contribute to the inflammatory response by secretion of additional chemokines, and neutrophils clear infected dying cells by phagocytosis (Egan et al., 2013). Maturation of dendritic cells (DCs), induced upon detection of pathogens, allows for the activation of the T cell response, with the cytotoxic T cell and T helper 1 cell response being of particular importance in the attempt to achieve clearance of these intracellular pathogens and in long-term prevention of reactivation from latency (Arvin & Gilden, 2013; Roizman et al., 2013). The humoral immune response, particularly mucosal IgA production by B cells, also contributes to the immune response, albeit to a lesser extent (Cradock-Watson, Ridehalgh, & Bourne, 1979; El Falaky, Vestergaard, & Hornsleth, 1977). Herpesviruses encode a number of gene products that counteract the innate and the adaptive immune system, leading to a delicate balance between host defense and viral evasion (Arvin & Gilden, 2013; Paludan et al., 2011; Roizman et al., 2013).

1.3. Innate DNA sensing The innate immune system is responsible for distinguishing between self and nonself during early stages of infection with pathogens. For this, it makes use of pattern recognition receptors (PRRs), which recognize pathogenassociated molecular pattern (PAMP). PAMPs are pathogen-specific molecules or molecules with aberrant localization. Toll-like receptors (TLRs) recognize a variety of pathogen-derived proteins and lipids at the cell surface (e.g., TLR2 recognizes herpesviral glycoproteins) and foreign nucleic acids in endosomes (e.g., TLR3 recognizes dsRNA, which is a product of herpesviral replication, and TLR9 recognizes viral dsDNA). Recently, intracellular nucleic acids are emerging as potent PAMPs. RIG-I (retinoic acid-inducible gene 1) and MDA5 (melanoma differentiation-associated protein 5) recognize viral RNAs produced during HSV-1 replication and

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signal via the adaptor protein MAVS (mitochondrial antiviral signaling protein) to induce expression of inflammatory cytokines and type I IFNs. Over 10 intracellular DNA sensors have been proposed in recent years. They include DAI (DNA-dependent activator of IFN-regulatory factors), RNA Pol III (RNA polymerase III), AIM2 (absent in melanoma 2), IFI16 (interferon-inducible protein 16), cGAS (cyclic GMP–AMP synthase), DDX41, DHX9, DHX36, LRRFIP1 (leucine-rich repeat flightless-interacting protein 1), and two proteins involved in the DNA damage response, Mre11 and Ku70/DNA-PK (DNA-dependent protein kinase; Atianand & Fitzgerald, 2013; Goubau, Deddouche, & Reis e Sousa, 2013; Nie & Wang, 2013; Paludan et al., 2011). This review focuses on TLR9, AIM2, IFI16, RNA Pol III, and cGAS, which are best documented as DNA sensors (Fig. 2). As alphaherpesviruses contain a DNA genome, DNA sensing can take place soon after infection, once the viral DNA is exposed to sensors and without the need for viral replication taking place. Therefore, it is of particular interest to understand the early innate immune response to these viruses. TLR9, the first recognition system for herpesvirus DNA discovered (Lund, Sato, Akira, Medzhitov, & Iwasaki, 2003), senses viral DNA in endosomes and signals via the adaptor MyD88 (myeloid differentiation primary response gene 88) to activate the transcription factors NF-κB (nuclear factor kappa-light-chain-enhancer of activated B cells) via the IKK (IκB kinase) complex and IRF7 (interferon regulatory factor 7) via IKKα (Kawai & Akira, 2011). IFI16 and cGAS activate the signaling adaptor STING (stimulator of interferon genes), which is central to DNA-activated innate immune responses. The details of STING signaling have been reviewed in Ran, Shu, and Wang (2014) and Burdette and Vance (2013). Upon DNA binding, cGAS produces the secondary messenger, cyclic GMP– AMP (cGAMP), a cyclic dinucleotide (CDN) which binds to achieve STING activation (Sun, Wu, Du, Chen, & Chen, 2013; Wu et al., 2013), while IFI16 activates STING via an unknown mechanism. STING then interacts with the signaling kinase TBK1 (TANK-binding kinase 1) to induce the activation of IRF3 and NF-κB; activation of the latter also includes the IKK complex (Abe & Barber, 2014; Atianand & Fitzgerald, 2013; Tanaka & Chen, 2012; Zhong et al., 2008). Upon DNA stimulation, RNA Pol III synthesizes a 50 -ppp RNA product, which can bind to RIG-I to induce the MAVS adaptor-dependent signaling pathway, also induced by cytosolic RNA. MAVS mediates the activation of IRF3 and NF-κB via TBK1 and the IKK complex, respectively (Wu & Chen, 2014). While IFNα

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Figure 2 Model of sensing of alphaherpesvirus DNA. TLR9 senses herpesviral DNA in endosomes and signals via MyD88, which induces the activation of the IKK complex, consisting of IKKα, IKKβ, and the regulatory subunit NEMO. The IKK complex induces the degradation of IκB. Thus, NF-κB is released from inhibition by IκB and translocates to the nucleus. In pDCs, MyD88 additionally induces IKKα to activate IRF7. RNA Pol III produces a 50 ppp RNA when bound to AT-rich DNA. This RNA activates RIG-I, which signals via MAVS to activate the NF-κB and the IRF3 pathways. IRF3 homodimerizes or heterodimerizes with IRF7 and translocates to the nucleus. Upon cytosolic DNA stimulation, cGAS produces the cyclic dinucleotide cGAMP, which binds to STING to activate it. IFI16 senses DNA in the cytosol and in the nucleus and activates STING via an unknown mechanism that involves colocalization. Activated STING acts as a scaffold for TBK1 and IRF3 to induce IRF3 activation and it also activates the NF-κB pathway. While IFNα expression mostly requires the IRF pathway, IFNβ requires both the IRF and the NF-κB pathway. Many of the other cytokines are mainly under the control of NF-κB. IFI16 also induces inflammasome activation by interaction with ASC, which converts procaspase-1 to caspase-1, resulting in the maturation of the cytokines IL-1β and IL-18.

expression mostly requires the IRF pathway, IFNβ requires both the IRF and the NF-κB pathway. Many of the other cytokines are mainly under the control of NF-κB (Paludan et al., 2011). In addition to the induction of gene expression, AIM2 and IFI16 stimulate the formation of the inflammasome, a protein complex that mediates the proteolytic cleavage and maturation of the cytokines IL-1β (interleukin 1β) and IL-18, which have a broad proinflammatory effect and activate NK cells, respectively

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(Atianand & Fitzgerald, 2013; Egan et al., 2013; Strowig, Henao-Mejia, Elinav, & Flavell, 2012). Although the availability of mouse strains deficient in the proposed DNA sensors is still not complete, clear experimental evidence already exists showing that cytokine production based on endosomal and intracellular DNA sensing has a strong, nonredundant influence on the activation of the immune system in response to alphaherpesvirus infection (Ishii et al., 2008; Ishikawa, Ma, & Barber, 2009; Kis-Toth, Szanto, Thai, & Tsokos, 2011; Sorensen et al., 2008; Wuest et al., 2006). This review summarizes the recent literature on the innate recognition of alphaherpesvirus DNA, the mechanisms of activation of the different sensors, their methods of signal transduction, and their physiological role in defense against herpesviral infection. We discuss how the viral DNA becomes accessible to the sensors, how these recognition systems are evaded by the viruses, how DNA sensing can be applied to vaccine design, and what challenges await this field of research in the future.

2. DNA SENSORS 2.1. TLR9 TLR9 is the first discovered PRR recognizing DNA (Hemmi et al., 2000). Binding of TLR9 to DNA was confirmed among others in quantitative ligand-binding assays (Latz et al., 2007). It preferentially recognizes unmethylated CpG motifs in DNA (Kawai & Akira, 2011), which is abundant in the HSV-1 genome (Lund et al., 2003). TLR9 sensing takes place in the endosome (Ahmad-Nejad et al., 2002), where the sensor binds the DNA with its luminal ligand-binding domain containing leucine-rich repeats and signals via the cytoplasmic TIR (Toll/interleukin-1 receptor homology domain) signaling domain. Binding of DNA to TLR9 homodimers induces a conformational change in the cytosolic domain, resulting in signaling (Latz et al., 2007). In humans, TLR9 is expressed abundantly in plasmacytoid DCs (pDCs) and B cells, especially when the latter are activated, and in corneal endothelial cells (Bourke, Bosisio, Golay, Polentarutti, & Mantovani, 2003; Kadowaki et al., 2001; Takeda et al., 2011; Wagner et al., 2004). In mice, TLR9 expression is more ubiquitous, with TLR9 present in many cell types such as conventional DCs (cDCs), macrophages, B cells, and corneal epithelial cells (Sarangi, Kim, Kurt-Jones, & Rouse, 2007; Wagner, 2004). When activated by DNA binding, TLR9 signals via the adaptor protein MyD88 (Hacker et al., 2000; Schnare, Takeda, Akira, & Medzhitov, 2000) to induce a type I IFN, type III IFN, and inflammatory cytokine response

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(Fig. 2). In most cells, TLR9-induced MyD88 signaling leads to activation of TRAF6 (TNF-associated factor 6), which results in phosphorylation of IκB by the IKK complex, consisting of IKKα, IKKβ, and NEMO (NF-κB essential modulator)/IKKγ. Upon subsequent degradation of IκB, the uninhibited NF-κB can translocate to the nucleus to modulate the gene expression of inflammatory cytokines (Kawai & Akira, 2011; Takeuchi & Akira, 2010). In pDCs, TLR9 signals via an additional MyD88-dependent pathway which activates the type I IFN response by phosphorylation and activation of the transcription factor IRF7 via association of MyD88, TRAF6, TRAF3, IRAK1 (interleukin-1 receptor-associated kinase 1), and IKKα (Honda et al., 2005; Kawai & Akira, 2011; Takeuchi & Akira, 2010). While TLR9-DNA binding in early endosomes results in NF-κB signaling, later signaling from lysosome-related organelles preferentially results in IRF activation (Kawai & Akira, 2011). TLR9 mediates cytokine expression in response to HSV in a cell typeand tissue-dependent manner. pDCs are an important source of IFNα in response to DNA stimulation and HSV infection (Ank et al., 2008; Hochrein et al., 2004; Lund, Linehan, Iijima, & Iwasaki, 2006). Therefore, the first study that revealed a role for TLR9 in sensing of alphaherpesviruses focused on HSV-2-infected, bone marrow-derived pDCs. Cells derived from TLR9- or MyD88-deficient mice were incapable of secreting IFNα in response to UV-inactivated HSV-2 or CpG DNA (Lund et al., 2003). IFNα and IL-12 secretion in response to HSV-1 in pDCs was also TLR9 dependent (Krug et al., 2004). An intact endosomal pathway and viral DNA in the capsid were essential for the response, confirming the role of TLR9. Viral replication was not required, indicating that the incoming viral DNA was triggering the response (Krug et al., 2004; Lund et al., 2003; Rasmussen et al., 2007). While many cell types require multiple PRRs to induce the full response to HSV, TLR9 is sufficient in pDCs to induce IFNα/β, IL-6, IL-12, and RANTES (CCL5/chemokine (C–C) motif ligand 5) (Sato, Linehan, & Iwasaki, 2006; Sorensen et al., 2008). Additionally, pDCs induce expression of the inflammatory cytokines TNFα (tumor necrosis factor α), CCL2, CCL3, CCL4, and CXCL10 (chemokine (C–X–C) motif ligand 10) in response to HSV-1 (Megjugorac, Young, Amrute, Olshalsky, & Fitzgerald-Bocarsly, 2004). There are indications that the TLR9 dependency of the IFNα response in pDCs is time dependent, with later responses being TLR9 independent (Rasmussen et al., 2007), and dependent on the tissue from which the pDCs are derived, with bone marrowderived pDCs showing less dependency than spleen-derived pDCs

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(Hochrein et al., 2004). In vitro chemotaxis assays revealed that the chemokines produced by pDCs in a TLR9-dependent fashion in response to HSV-1 attract NK cells (Megjugorac et al., 2004). Although pDCs and their TLR9-dependent type I IFN secretion are very important for the innate immune response, a normal Th1 response was shown to develop in response to HSV-2 in the absence of this particular DC subset (Lund et al., 2006). However, activated T cells are attracted by TLR9-dependent chemokine secretion by HSV-1-infected pDC (Megjugorac et al., 2004). In murine cDCs, which are also important producers of IFN upon HSV infection (Ank et al., 2008), the production of type I and type III IFN in response to HSV infection is independent of TLR9 (Ank et al., 2008; Hochrein et al., 2004; Rasmussen et al., 2007; Sato et al., 2006). However, HSV-1-induced expression of TNFα, IL-6, and IL-12 is reduced in the absence of TLR9 (Hochrein et al., 2004; Sato et al., 2006), while HSV2-induced cytokine expression (CCL5, IL-6) in cDCs is independent of TLR9 (Sorensen et al., 2008). Similarly, in murine macrophage cell lines or primary macrophages, IFNα expression upon HSV-1 and HSV-2 is independent of TLR9, while the expression of inflammatory cytokines (TNFα, CCL5, IL-6, IL-12) in response to the viruses is partially TLR9 dependent (Hochrein et al., 2004; Lima et al., 2010; Malmgaard, Melchjorsen, Bowie, Mogensen, & Paludan, 2004; Rasmussen et al., 2007, 2009; Sorensen et al., 2008). In primary human macrophages, early sensing of HSV is TLR9 independent (Melchjorsen et al., 2010). Although B cells express TLR9, they are not major producers of IFNα in response to HSV-1 (Hochrein et al., 2004). In a human corneal endothelial cell line, a vast array of inflammatory cytokines, including CCL5, IL-6, and CXCL10, are expressed in a TLR9-dependent fashion via the NF-κB pathway in response to HSV-1 infection (Takeda et al., 2011). The production of the chemokines CXCL9 and CXCL10 in the cornea of TLR9-deficient mice infected with HSV-1 was reduced compared to wild-type mice. This was related to reduced infiltration of neutrophils into the cornea of these mice; however, macrophage infiltration was unaltered (Wuest et al., 2006). In human primary vaginal epithelial cells, a major target of HSV-2 infection, both IFNβ and IL-6 expression in response to live HSV-2 or purified HSV-2 DNA were reduced by TLR9 RNA interference (Triantafilou, Eryilmazlar, & Triantafilou, 2014). No reduction in expression of CXCL9 and CXCL10 was observed in the vaginal tissue of HSV-2-infected TLR9-deficient mice (Wuest et al., 2006).

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There are several studies that emphasize the importance of TLR9 in HSV-induced cytokine expression not only after local infection of cell lines, primary cells, and tissues but also after systemic infections. Upon intravenous injection of HSV-2, no IFNα could be detected in the serum of TLR9deficient mice (Lund et al., 2003). IFNα and CCL5 levels in the serum upon intraperitoneal infection with HSV-1 or HSV-2 were partially dependent on TLR9 at early time points after infection, while TNFα and IL-6 levels were not (Rasmussen et al., 2007; Sorensen et al., 2008). In the brain, TLR9-dependent TNFα and CXCL9 expression was detected after vaginal infection of mice with HSV-2, while type I IFN expression could not be detected (Sorensen et al., 2008). TLR9-deficient mice had altered cytokine profiles upon intranasal HSV-1 infection, both in brains and in trigeminal ganglia, with the expression of some cytokines increased and others decreased (Lima et al., 2010). Despite the important role of TLR9 in cytokine expression after HSV infection, studies on viral replication and disease outcomes in TLR9-deficient mice had mixed results. A first study using HSV-1 footpad injections showed no differences in viral replication and disease between TLR9-deficient and wild-type mice (Krug et al., 2004). The viral titers in the brains of mice infected intraperitoneally with HSV-2 were not altered by TLR9 deficiency (Sorensen et al., 2008). HSV-2-infected, TLR9- and TLR2-double-deficient mice showed impaired recruitment of NK cells to the spleen, but activation (measured by surface marker expression) and cytotoxic activity were unaltered (Sorensen et al., 2008). While TLR9deficient mice infected intraperitoneally with HSV-2 showed higher viral titers in the spleen, the survival was not affected (Rasmussen et al., 2007). TLR9-deficient mice were much more likely to succumb to lethal encephalitis caused by intranasal HSV-1 infection, and in this model, the viral titers in the brain were greatly increased (Lima et al., 2010). Ocular infection of MyD88 and TRIF (TIR-domain-containing adaptor-inducing IFNβ)double-deficient mice, which are incapable of any TLR signaling, did not result in higher viral titers in the cornea (Conrady, Zheng, Fitzgerald, Liu, & Carr, 2012). TLR9-deficient animals showed milder corneal lesions in response to HSV-1, but increased herpetic ocular disease scores. In this study, TLR9-deficient mice were not more susceptible to lethal infections, although they showed higher viral titers in brains and trigeminal ganglia (Sarangi et al., 2007). TLR9 deficiency also caused increased HSV-1 shedding in tears (Wuest et al., 2006). Interestingly, HSV-1-induced angiogenesis in a corneal micropocket assay and a murine macrophage cell line

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produced vascular endothelial growth factor upon CpG DNA stimulation. This process is speculated to contribute to the corneal scarring typical for HSV-1 infection (Zheng, Klinman, Gierynska, & Rouse, 2002). Interestingly, TLR9 signaling to induce NF-κB activation not only limits viral replication by activation of the immune system but is simultaneously required to some extent for initiation of HSV-1 gene expression, as the immediate-early infected cell protein (ICP)0 promoter contains NF-κB-responsive elements (Takeda et al., 2011). Less research has been performed on the involvement of TLR9 in the innate immune response to VZV. pDCs were shown to be responsible for IFNα production in a TLR9-dependent fashion during VZV infection, while IL-12 was not produced by human peripheral blood mononuclear cells in response to the virus. UV-inactivated virus led to less IFNα expression than infectious virus, indicating that an additional, TLR9-independent, replication-dependent process exists. In VZV infection, NF-κB was shown to be important in IFNα induction (Yu et al., 2011). In general, the TLR9 dependency of HSV recognition and the exact nature of downstream gene expression seems to depend very much on the cell and tissue type, the time course after infection, and the virus strain. However, it is likely that some of the observed differences are due to differences in experimental conditions and detection methods. Clearly, TLR9 has an important role in innate PAMP recognition, but several other mechanisms are in place that prevent too severe effects in animals lacking TLR9. For example, there are several indications that TLR9 acts in synergy with TLR2 in murine macrophages, in human vaginal epithelial cells, in the trigeminal ganglia, and in the brain (Lima et al., 2010; Sorensen et al., 2008; Triantafilou et al., 2014; Zolini et al., 2014). In humans, deficiency in IRAK4, which is required for TLR9 (and TLR7 and TLR8) signaling (Mogensen, 2009), does not lead to an increased susceptibility to alphaherpesvirus infections (Ku et al., 2007). Several attempts have been made to modulate the immune response to herpesviruses with synthetic TLR9 agonists and antagonists, for example, by Boivin and colleagues, who showed application of TLR9 agonists before infection could reduce the amount of virus in the brain, the production of inflammatory cytokines, and subsequently the mortality due to encephalitis (Boivin, Menasria, Piret, & Boivin, 2012). In the future, modulation of the TLR9 response may be used in combination with antiviral drugs to control HSV infection and resulting harmful inflammation.

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2.2. Discovery of intracellular DNA sensors Herpesvirus DNA is not only found in the lumen of endosomes. During the cause of the replication cycle, the virus migrates through the cytosol to the nucleus (Fig. 1B). Thus, the viral DNA is abundantly present in the cytoplasm and the nucleus. Soon after the discovery of TLR9 as a sensor of herpesvirus DNA, several studies demonstrated TLR-independent pathways for intracellular nucleic acid recognition. IFNα/β expression in murine macrophage-like cells in response to HSV-2 is maintained in the absence of MyD88 and under TLR9-inhibiting conditions (Malmgaard et al., 2004). Murine cDCs, macrophages, and bone marrow-derived pDCs were shown to produce IFNα and other cytokines largely independent of TLR9 upon HSV-1 stimulation (Hochrein et al., 2004). Ishii and colleagues were the first to stringently show that cytosolic DNA induces innate immune responses. Transfected genomic DNA derived from mammals, bacteria, and viruses, including HSV-1 and HSV-2, induced TLR- and RIG-Iindependent expression of IFNβ and inflammatory cytokines in MEFs (murine embryonic fibroblasts) (Ishii et al., 2005). It was then demonstrated that transfected DNA can induce a TLR-independent type I IFN response in a number of cell types, including murine pDCs, cDCs, macrophages, and MEFs. The sequence of the DNA was irrelevant to the response; however, a native DNA backbone composition was found to be essential (Stetson & Medzhitov, 2006). IRF3 is a key transcription factor in this DNA-induced type I IFN response (Ishii et al., 2005; Stetson & Medzhitov, 2006). The type I IFN expression in murine cDCs upon HSV-1 infection was then also shown to be independent of TLR9, to require viral entry and the presence of the viral genome in the virion, but to not require viral replication (Rasmussen et al., 2007). This indicated that recognition of intracellular DNA, rather than RNA, is taking place. Together, these studies suggested that additional, intracellular receptors exist for herpesviral DNA.

2.3. DAI In the search for the factor responsible for the innate signaling upon cytosolic DNA stimulation, the ISG DAI was the first protein proposed to bind cytosolic DNA and to initiate the innate immune response to it (Takaoka et al., 2007). DNA binding by DAI was demonstrated by fluorescence resonance energy transfer (FRET) and by coprecipitation of purified DAI with biotinlabeled DNA (Takaoka et al., 2007; Wang et al., 2008). DAI can bind both

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B- and Z- forms of DNA, independent of sequence (Kim et al., 2011; Takaoka et al., 2007; Wang et al., 2008). In fibroblasts, IRF3 was reported to be needed for DAI-mediated IFNβ response, while both IRF3 and IRF7 were needed for IFNα induction. Upon stimulation with cytosolic DNA, DAI associated with TBK1 and IRF3 (Takaoka et al., 2007). DAI was also found to be phosphorylated at multiple sites by TBK1, and this phosphorylation was necessary for IRF3-mediated gene expression (Wang et al., 2008). In addition to signaling to IRF3, an essential role for DAI in DNA-induced NF-κB activation was shown (Takaoka et al., 2007). DAI-induced activation of NF-κB was shown to depend on the interaction of DAI with RIP1 (receptor-interacting protein kinase 1) and to further increase upon interaction with RIP3 (Kaiser, Upton, & Mocarski, 2008). An initial study showed that overexpression of DAI in a mouse fibroblast cell line, L929 cells, increased the type I IFN response to transfected DNA species, including viral DNA. The innate response to HSV-1 infection of L929 cells was modestly reduced by RNA interference of DAI (Takaoka et al., 2007). With a focus on the neurotropism of HSV-1, it was shown that the innate response of microglia and astrocyte cells to B-DNA and HSV-1 was reduced by DAI knockdown (Furr, Chauhan, MoerdykSchauwecker, & Marriott, 2011). In vaginal cells with reduced DAI expression, the response to HSV-2 infection or HSV-2 DNA stimulation was found to be impaired (Triantafilou et al., 2014). However, soon after the proposal of DAI as a DNA sensor, several studies challenged the notion that DAI was an universal DNA sensor. While its role in the fibroblast-like L929 cell line was confirmed, MEFs and human alveolar epithelial cells did not require DAI for their innate response to DNA as shown by RNA interference (Lippmann et al., 2008; Wang et al., 2008). Additionally, MEFs, bone marrow-derived DCs, and immortalized bone marrow-derived macrophages from DAI-deficient mice responded normally to DNA, and the mice were not impaired in their immune response to a DNA vaccine (Ishii et al., 2008; Unterholzner et al., 2010). In vivo RNA interference of DAI in the cornea of mice did not result in increased viral titers after ocular HSV-1 infection (Conrady et al., 2012). Recently, a DNA sensing-independent function of DAI on HSV-1 replication in a hepatocellular carcinoma cell line was shown based on its ability to prevent expression of the immediate-early ICP0 gene. The cytokine response in these cells was not affected by modulation of DAI expression (Pham, Kwon, Kim, Kim, & Ahn, 2013).

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In conclusion, although DAI was convincingly shown to bind DNA, it is unlikely to be a central cytosolic sensor for the recognition of alphaherpesvirus DNA. However, it may be involved in DNA-induced signaling in some cell types, especially in fibroblasts. It is conceivable that DAI has a redundant function as a PRR for DNA and/or as a signaling element, which is more prominent in certain cell types. Independent of its DNAsensing properties, it is a restriction factor for HSV by targeting ICP0 expression.

2.4. AIM2 Transfected DNA of bacterial, viral, and mammalian origin causes formation of the inflammasome in murine macrophages and in the human monocyte cell line THP-1 (Muruve et al., 2008). The inflammasome is a protein complex formed around one of multiple sensor proteins, member of the NOD-like receptor (NLR) or PYHIN (pyrin and HIN domain-containing) protein families, which together recognize various viral PAMPs. The adaptor protein ASC (apoptosis-associated speck-like protein containing CARD), consisting of a pyrin and a CARD (caspase recruitment) domain, binds to the sensor proteins via its pyrin domain and to procaspase 1 via its CARD domain through homotypic interaction, resulting in caspase activation by autolytic cleavage. Caspase-1 mediates the maturation and secretion of the cytokines IL-1β and IL-18 (Strowig et al., 2012). These cytokines are thus regulated at a posttranslational level, in contrast to many other cytokines which are regulated at a transcriptional level. The search for the protein that induces inflammasome formation upon stimulation with cytosolic DNA yielded the ISG AIM2 as a cytosolic DNA urckst€ ummer et al., 2009; Fernandes-Alnemri, Yu, Datta, Wu, & sensor (B€ Alnemri, 2009; Hornung et al., 2009; Roberts et al., 2009). In studies with AIM2 RNA interference and AIM2-deficient mice, AIM2 was shown to be essential for inflammasome formation, caspase-1 activation, and processing of IL-1β and IL-18, but also for pyroptosis, which is an inflammasomeurckst€ ummer et al., 2009; Fernandesmediated form of cell death (B€ Alnemri et al., 2009; Hornung et al., 2009; Rathinam et al., 2010; Roberts et al., 2009; Sagulenko et al., 2013). Studies with AIM2-deficient mice showed that, in macrophages and DCs, AIM2 is the inflammasome formation-mediating DNA sensor in response to infection with various pathogens, including intracellular bacteria, vaccinia virus, and the betaherpesvirus murine cytomegalovirus

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(Hornung et al., 2009; Rathinam et al., 2010). HSV-1 induces strong caspase-1 activation in human and mouse macrophages and primary human fibroblasts ( Johnson, Chikoti, & Chandran, 2013; Muruve et al., 2008; Nour et al., 2011; Rathinam et al., 2010). However, this is independent of AIM2 ( Johnson et al., 2013; Rathinam et al., 2010). VZV also induces formation of the inflammasome complex in THP-1 cells and primary human lung fibroblasts. However, it also causes inflammasome activation in melanoma cells, which lack AIM2. Thus, the VZV-induced inflammasome is AIM2 independent; instead, it is dependent on NLRP3 (Nour et al., 2011). Since inflammasome formation upon transfected DNA was NLRP3 independent (Muruve et al., 2008), it is likely that VZV activates this inflammasome in a DNA-independent fashion. HSV-1 was also shown to transiently activate the NLRP3 inflammasome ( Johnson et al., 2013). Thus, although AIM2 is an important DNA sensor and mediator of the inflammasome response to some DNA viruses, this PRR is probably not involved in recognition of alphaherpesviruses.

2.5. IFI16 Similar to AIM2, IFI16 (and its mouse orthologue p204) belongs to the PYHIN family. IFI16 is also a receptor for intracellular DNA and mediates cytokine induction and inflammasome activation in response to a variety of dsDNA species (Fig. 2). Unterholzner and colleagues identified IFI16 as an intracellular DNA sensor by affinity purification from human monocyte cell extracts using vaccinia virus-derived DNA as bait (Unterholzner et al., 2010). IFI16 binds dsDNA directly, independent of sequence, and this binding is mediated by its two HIN domains, as shown by pull-down assays with tagged and untagged versions of IFI16, by electrophoretic mobility shift assay, by FRET, and by in vitro binding assays, including experiments using purified HIN domains, and by mutations studies with alanine substitution in the HIN domains (Conrady et al., 2012; Dawson & Trapani, 1995b; Morrone et al., 2014; Unterholzner et al., 2010). The crystal structures of the HIN domains of IFI16 and AIM2 in complex with DNA have been solved and reveal binding to the DNA backbone as the basis for the sequence-independent affinity to DNA ( Jin et al., 2012). Alanine substitution assay showed that the DNA-binding capacity of the HIN domains is essential for the DNA-induced immune response in IFI16-transfected HEK293 (human embryonic kidney) cells ( Jin et al., 2012). DNA binding by IFI16 is cooperative and length dependent, with dsDNA fragments

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shorter than 70 bp showing declining affinity (Morrone et al., 2014; Unterholzner et al., 2010). This can be explained by the fact that IFI16 oligomerizes on dsDNA to form filaments, through a process mediated by homotypic interaction between the IFI16 pyrin domains (Morrone et al., 2014). IFI16 (or p204) expression has been demonstrated in many primary and immortalized cell types including myeloid cells, fibroblasts, epithelial cells, and lymphoid cells (Caposio et al., 2007; Conrady et al., 2012; Cristea et al., 2010; Dawson & Trapani, 1995a, 1995b; Duan et al., 2011; Gariano et al., 2012; Hertel et al., 1999; Soby, Laursen, Ostergaard, & Melchjorsen, 2012; Triantafilou et al., 2014; Unterholzner et al., 2010; Veeranki, Duan, Panchanathan, Liu, & Choubey, 2011). In most cell types, e.g., fibroblasts, IFI16 is predominantly localized to the nucleus, but in some cell types, especially macrophage-like cells, a pool of the cellular IFI16 localizes to the cytoplasm ( Johnson et al., 2013; Orzalli, DeLuca, & Knipe, 2012; Unterholzner et al., 2010; Veeranki & Choubey, 2012; Veeranki et al., 2011). In cells commonly used for transfection experiments, such as HeLa and HEK293T, ectopic expression of IFI16 leads to localization of the protein to the nucleus (B€ urckst€ ummer et al., 2009; Hornung et al., 2009). The subcellular localization of IFI16 is regulated by a nonlinear nuclear localization signal (NLS), which can be acetylated by the acetyltransferase p300 in lymphocytes and macrophages, promoting cytosolic localization by inhibiting nuclear import (Li, Diner, Chen, & Cristea, 2012). IFI16-mediated sensing of HSV-1 DNA has been shown to take place independently of nuclear entry of viral DNA in THP-1 cells and monocyte-derived macrophages, in which IFI16 was found to colocalize with HSV-1 DNA in the cytosol (Horan et al., 2013; Unterholzner et al., 2010). However, DNA sensing by IFI16 can also take place in the nucleus. One study showed that in HFFs (human foreskin fibroblasts), recognition of the HSV-1 genome in the nucleus induced translocation of IFI16 to the cytosol where it mediated inflammasome activation ( Johnson et al., 2013). Another study, also using HFFs infected with HSV-1, showed that the nuclear localization of IFI16 did not change upon HSV-1 infection to mediate IFNβ expression via the STING pathway (Orzalli et al., 2012). The NLS of IFI16 was shown to be essential for IFNβ induction upon HSV-1 infection in IFI16-transfected U2OS cells (a human epithelial cell line) and partial relocalization of IFI16 to the cytosol upon infection was observed (Li et al., 2012).

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IFI16/p204 mediates induction of type I IFN and other inflammatory cytokines, such as CXCL10, IL-6, and TNFα, upon HSV-1 infection of human and murine cells (Conrady et al., 2012; Horan et al., 2013; Orzalli et al., 2012; Soby et al., 2012; Unterholzner et al., 2010; Zhu et al., 2014). An important role of IFI16/p204 in the innate immune response has also been proposed by in vivo RNA interference of p204 in mouse corneal epithelium and vaginal mucosa, which led to reduced IFNα induction, and increased viral titers in the cornea and the vaginal lumen after infection with HSV-1 and HSV-2, respectively (Conrady et al., 2012). IFI16-triggered cytokine expression is mediated via the STING/TBK1/ IRF3 pathway and the NF-κB pathway (Fig. 2) (Li, Chen, & Cristea, 2013; Unterholzner et al., 2010). IFI16 RNA interference prevented the nuclear translocation of IRF3 and NF-κB upon HSV-1 infection in murine macrophages cells (Unterholzner et al., 2010). Loss of IFI16 also prevented IRF3, but not IRF7 translocation in THCE (telomerase-immortalized human corneal epithelial) cells in response to HSV-1 (Conrady et al., 2012). The presence of STING was essential for IRF3 signaling in HFFs infected with HSV-1 (Orzalli et al., 2012). The strong maturation of DCs induced by DNA stimulation was partially dependent on IFI16 and STING (KisToth et al., 2011). Coimmunoprecipitations show that IFI16 and STING interact upon DNA stimulation (Unterholzner et al., 2010) and IFI16 and STING colocalize in the cytoplasm in human monocyte-derived macrophages after HSV-1 infection (Horan et al., 2013). However, it remains to be determined if this is a direct interaction and how exactly IFI16mediated STING activation takes place. In addition to its role in type I IFN induction, IFI16 has also been implicated as an activator of the inflammasome (Fig. 2); however, there are also studies that contradict this. Two initial studies that identified the role of AIM2 in inflammasome activation also analyzed IFI16 and failed to show association of IFI16 with the inflammasome adaptor ASC and IFI16mediated inflammasome activation (B€ urckst€ ummer et al., 2009; Hornung et al., 2009). Also, in THP-1 cells endogenous IFI16 did not associate with ASC (Hornung et al., 2009). However, IFI16 has been reported to recognize the HSV-1 genome in the nucleus after HSV-1 infection of HFFs and to subsequently mediate inflammasome assembly ( Johnson et al., 2013). In lytic and latent infections with gammaherpesviruses, IFI16 was also the inducer of inflammasome activation (Ansari et al., 2013; Kerur et al., 2011; Singh et al., 2013). IFI16, in addition to mediating innate immune signaling upon HSV infection, also acts as a restriction factor for

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immediate-early gene expression of HSV-1 by promoting the epigenetic silencing of the viral genome (Orzalli, Conwell, Berrios, DeCaprio, & Knipe, 2013). In conclusion, IFI16 is important for the early host response to HSV infection in many cell types. IFI16 has a DNA-binding domain and can recognize HSV-1 DNA both in the nucleus and in the cytoplasm to mediate type I IFN and inflammatory cytokine responses via the STING and inflammasome pathways. Several interesting questions remain open. It will be important to understand how IFI16 differentiates foreign from host DNA when sensing in the nucleus, despite its sequence-independent DNAbinding domain. Also, the exact mechanisms determining IFI16 localization in different cell types and upon various stimulation conditions await clarification. It needs to be determined which factors influence the type of inflammasome (AIM2 vs. IFI16) formed upon detection of DNA from different intracellular pathogens, but also from closely related viruses (betaherpesviruses vs. alpha- and gammaherpesviruses) and how the cellular context determines the exact role of IFI16 in inflammasome formation. Data on IFI16 sensing of VZV are lacking so far.

2.6. cGAS cGAS, a member of the nucleotidyltransferase family, is a recently discovered cytosolic DNA-sensing enzyme that catalyzes the cyclization reaction of ATP and GTP to form cGAMP, which acts as a secondary messenger to activate STING (Fig. 2; Sun et al., 2013; Wu et al., 2013). cGAMP as a secondary messenger activating STING was identified by mass spectrometry of components of heat-inactivated cytosolic extracts from cells transfected with various dsDNA that could mediate STING activation in permeabilized RAW264.7 and THP-1 cells (Wu et al., 2013). Subsequently, cGAMP was found to be synthesized by cGAS by biochemical fractionation and mass spectrometry of the cGAMP-producing fraction (Sun et al., 2013). Purified tagged cGAS could synthesize cGAMP from ATP and GTP in vitro in the presence of various dsDNA, but not without dsDNA stimulation (Sun et al., 2013). So far, cGAS expression has been confirmed in most cell types analyzed, but expression is low in some cell types, including MEFs (Sun et al., 2013). cGAS localizes to the cytosol as shown by subcellular fractionation of THP-1 cells and by confocal microscopy of L929 cells expressing epitopetagged cGAS, with low amounts also present in the nucleus in both cell types (Sun et al., 2013).

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cGAS directly binds dsDNA independent of sequence. Its N-terminal domain is required for DNA binding. Epitope-tagged cGAS, but not N-terminal truncation mutants, associated with biotinylated dsDNA in pull-down assays (Sun et al., 2013). cGAS colocalizes with transfected dsDNA in L929 cells (Sun et al., 2013). The crystal structures of human, murine, and porcine cGAS in combination with GTP and ATP have been solved (Civril et al., 2013; Gao et al., 2013; Kranzusch, Lee, Berger, & Doudna, 2013; Li, Shu, et al., 2013; Zhang et al., 2014). The DNA-binding domain consists of a zinc ribbon/thumb domain (Civril et al., 2013; Kranzusch et al., 2013). DNA binding is based on interactions with the DNA backbone across the minor groove, explaining the sequence independence (Civril et al., 2013). Although initial studies found no evidence that cGAS oligomerizes (Gao et al., 2013), later studies proposed that cGAS forms a 2:2 complex with DNA and that this dimerization is required for activation (Li, Shu, et al., 2013; Zhang et al., 2014). DNA binding to cGAS induces a conformational change, leading to the formation of a nucleotide binding pocket and reorganization of the catalytic site (Civril et al., 2013; Gao et al., 2013; Zhang et al., 2014). STING is essential for the cGAS-activated pathway (Fig. 2). DNA stimulation of L929 cells in the absence of STING did not lead to IRF3 activation (Wu et al., 2013). Overexpression of cGAS in HEK293T cells activated IRF3 and induced IFNβ expression in a manner dependent on STING (Sun et al., 2013). In contrast to other DNA sensors, the mechanism of STING activation by cGAS is now described in detail, with cGAMP binding to dimeric STING inducing a conformational change in the C-terminal domain in STING allowing for activation of TBK1 and downstream signaling to type I IFN expression (Wu et al., 2013; Zhang, Shi, et al., 2013). Interestingly, cGAMP can be transferred from infected cells to neighboring cells via GAP junctions to spread STING activation and IFNβ induction in response to a dsDNA virus (vaccinia virus) as shown in MEFs and transfected HEK293T cells (Ablasser et al., 2013). cGAS activity is regulated by the autophagy protein Beclin-1, which directly interacts with cGAS to stop production of cGAMP. This interaction also relieves Beclin-1 inhibition to activate autophagy of cytosolic DNA, restricting immune signaling after HSV-1 infection (Liang et al., 2014). Cytosolic DNA sensing by cGAS has a clear role in the innate immune response to HSV-1. Infection of THP-1 and L929 cells leads to production of cGAMP and IRF3 activation (Wu et al., 2013). RNA interference of cGAS prevents cGAMP synthesis, IRF3 activation, and the IFNβ response

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to HSV-1 infection in L929 cells and IRF3 activation in THP-1 cells (Sun et al., 2013). Cells from cGAS-deficient mice did not produce type I IFN, CXCL10, and IL-6 and failed to activate IRF3 in response to HSV-1 (Li, Wu, et al., 2013). The importance of cGAS was confirmed in vivo. In the absence of cGAS, IFNα and IFNβ serum levels were reduced in response to intravenous injection of HSV-1, the virus could spread efficiently to the brain, and the cGAS-deficient mice succumbed to the infection much earlier than wild-type mice (Li, Wu, et al., 2013). Data on the involvement of cGAS in immunity to HSV-2 and VZV are lacking so far.

2.7. RNA Pol III and RIG-I Soon after RIG-I was discovered as a cytosolic RNA sensor mediating antiviral innate signaling (Yoneyama et al., 2004), it was reported that RIG-I and its adaptor protein MAVS were also essential for expression of IFNβ in response to AT-rich dsDNA, and to HSV-1 infection in a human hepatoma cell line (Cheng, Zhong, Chung, & Chisari, 2007). Inhibition of the RIG-IMAVS pathway prevented poly(dA:dT)-induced activation of an IFNβ promoter reporter system in HEK293T cells (Ablasser et al., 2009; Chiu, MacMillan, & Chen, 2009). This indicated a role for RIG-I in cytosolic DNA sensing, but RIG-I was not found to bind to dsDNA directly (Cheng et al., 2007). This was followed by the demonstration that RNA extracted from poly(dA:dT)-transfected cells induced IFNβ in human peripheral blood mononuclear cells, indicating that RIG-I activation occurred via an RNA intermediate (Ablasser et al., 2009). The RNA intermediate was identified to be double stranded, to contain a 50 -triphosphate group, and to be synthesized by RNA Pol III (Fig. 2; Ablasser et al., 2009; Chiu et al., 2009). RNA Pol III DNA binding is likely to be dependent on the nucleotide composition of the DNA since GC-rich DNA could not be transcribed (Chiu et al., 2009). At the present stage, the physiological importance on the RNA Pol III pathway in recognition of HSV-1 remains unresolved, since the published data are somewhat contradictory. MEFs from RIG-I-deficient mice were impaired in their IFNβ, but not CCL5 and IL-6 response to HSV-1 DNA (Choi et al., 2009). Deactivation of RNA Pol III activity by a chemical inhibitor prevented IFNβ induction in response to HSV-1 in RAW264.7 cells (Chiu et al., 2009). However, another study found that HEK293T cells, which contain a functional RNA Pol III/RIG-I system, did not respond to HSV60mer DNA and that inhibition of RNA Pol III

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had no effect on HSV-derived DNA- and HSV-1 infection-induced IFNβ expression in RAW264.7 cells, nor on HSV-1-induced IFNβ and TNFα expression in human macrophages, although RNA Pol III inhibition did have an effect on the response to poly(dA:dT) (Melchjorsen et al., 2010; Unterholzner et al., 2010). There are no data yet on HSV-2, VZV, and the RNA Pol III pathway. Further research on the role of RNA Pol III in the antiviral defense in different cell types and especially in vivo is needed.

3. ACCESSIBILITY OF VIRAL DNA TO DNA SENSORS As discussed above, HSV-1 DNA is sensed in endosomes by TLR9, in the cytosol by AIM2, IFI16, cGAS, and RNA Pol III, and in the nucleus by IFI16 (and maybe also by RNA Pol III). The mechanisms through which the viral genome in the capsid is made accessible to the DNA sensors in these subcellular compartments are not fully understood. Since pDCs, in which TLR9 is preferentially expressed, are capable of phagocytosis (Tel et al., 2010), it is likely that virions are delivered to the endolysosomal pathway via this mechanism. Depending on the cell type, alphaherpesviruses enter cells via endocytosis or direct fusion with the plasma membrane (Fig. 1B; Arvin & Gilden, 2013; Roizman et al., 2013). The endocytic entry pathway followed by degradation of the virion would allow for exposure of the viral genome to TLR9 signaling in nonphagocytic cells (Fig. 1B). It is also possible that viral components including the genome are delivered to endosomes via autophagy (Fig. 1B), as autophagy is needed for sensing of ssRNA virus replication intermediates by TLR7 in endolysosomes and as pDCs lacking the autophagy protein ATG5 were severely impaired in their IFNα response to HSV-1 infection (Lee, Lund, Ramanathan, Mizushima, & Iwasaki, 2007). In macrophages, which are nonpermissive for productive HSV-1 replication, the HSV-1 capsid is ubiquitinated and degraded by the proteasome in the cytosol, exposing viral DNA to cytoplasmic sensors (Fig. 1B). However, this does not seem to occur in HFFs and U2OS cells, which are permissive for HSV-1 infection and in which the viral DNA efficiently reaches the nucleus (Horan et al., 2013). Proteasomal capsid degradation may thus be an important mechanism to protect certain cell types from viral infection (Paludan et al., 2011). At present, it is not known which cellular sensors detect the viral capsid, and which E3 ubiquitin ligases are involved; nor is it known whether herpesviruses seek to evade capsid targeting in the cytosol.

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In cells permissive for infection, the viral capsid is delivered to nuclear pores, where HSV DNA is transported into the nucleus (Roizman et al., 2013). In the nucleus, the viral DNA is not protected by the capsid and hence potentially exposed to nuclear PRRs (Fig. 1B). In this context, the question is not so much how the exposure occurs, but rather how nuclear DNA sensors can distinguish between self and foreign DNA. The observed recognition of viral DNA in the nucleus by IFI16 challenges the concept of the nucleus as an “immunoprivileged” subcellular compartment and the notion that intracellular DNA sensor reacts to DNA purely based on their aberrant localization in the cytoplasm. This question is especially intriguing, as IFI16 recognizes DNA sequence independently by interactions with the backbone ( Jin et al., 2012). It is possible that additional cofactors guide IFI16 specifically to viral DNA. This may involve components of the DNA damage response, some of which have been shown to have role in DNA sensing (Ferguson, Mansur, Peters, Ren, & Smith, 2012; Kondo et al., 2013; Zhang et al., 2011) and which are known to associate with alphaherpesviral DNA based on their ability to recognize dsDNA breaks (Weitzman, Lilley, & Chaurushiya, 2010). Interestingly, DNA damage can induce expression of IFN and inflammatory genes (Brzostek-Racine, Gordon, Van Scoy, & Reich, 2011). Additionally, histones may protect the host genome from recognition by PRRs. This may contribute to the fact that during latent infection, when viral DNA is circularized and associated with histones, it does not induce a strong innate immune response anymore.

4. EVASION OF DNA-INDUCED SIGNALING Evasion of the innate and adaptive immune system is very important for the ability of alphaherpesviruses to establish and maintain infection. Therefore, these viruses have evolved mechanisms to target all steps of the immune response. As discussed above, multiple PRRs recognize herpesviruses and converge at common signaling pathways, especially IRF3 and NF-κB signaling. So in addition to inhibiting certain PRRs directly, alphaherpesviruses target the downstream signaling events such as IRF3 and NF-κB activation (Fig. 3). STING as a crucial mediator of intracellular nucleic acid-induced signaling has been shown to be targeted by various RNA viruses, but not yet by DNA viruses (Ran et al., 2014). Since the intracellular DNA sensors have been discovered only in recent years, it can be expected that more viral immune evasion molecules inhibiting these specifically will be discovered soon. As the host–pathogen interaction very early in

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Figure 3 Evasion of innate DNA sensing by alphaherpesviruses. HSV ICP0 inhibits IFI16, NF-κB, and IRF3 signaling in the cytosol and nucleus. ICP0 also promotes cytoplasmic translocation of USP7, which inhibits NEMO when at this location. BoHV-1 ICP0 targets IRF3 for degradation. HSV ICP34.5 interferes with the action of TBK1 and the IKK complex and binds to Beclin-1 to inhibit autophagy, which may prevent delivery of PAMPs to the endosome for TLR9 sensing. HSV US3 blocks nuclear translocation of IRF3 and NF-κB and inhibits signaling from MyD88 to the IKK complex. HSV ICP27 binds to IκB, preventing its degradation. HSV VP16 interacts with a coactivator of IRF3 (CBP) and with NF-κB to prevent gene expression. HSV US11 interacts with RIG-I to prevent activation of MAVS. HSV UL36 impairs MAVS signaling. VZV IE62 blocks IRF3 activation by TBK1. VZV ORF47 kinase prevents nuclear translocation of IRF3, like HSV US3. Like ICP0, VZV ORF61 targets IRF3 for degradation.

infection determines the outcome of infection, many innate immune evasion molecules are tegument or immediate-early proteins. Herpesvirus proteins are often multifunctional and many viral inhibitors of innate signaling interfere through several mechanisms. More research needs to be done to elucidate in detail the mechanism of action of many of the immune evasion molecules. The HSV ICP0 is an immediate-early protein with multiple functions, including E3 ubiquitin ligase activity via its RING finger domain. ICP0 directly targets IFI16 for proteasomal degradation, and this interferes with IRF3-induced IFNβ expression and inflammasome-mediated secretion of mature IL-1β ( Johnson et al., 2013; Orzalli et al., 2012). ICP0 also

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sequesters activated IRF3 and its coactivators in the nucleus and enhances IRF3 degradation to prevent IFNβ gene induction (Melroe, Silva, Schaffer, & Knipe, 2007; Orzalli et al., 2012). Another study confirmed that ICP0 inhibits IRF3 signaling, but in this study cytosolic localization of ICP0 was essential and IRF3 was not degraded (Paladino, Collins, & Mossman, 2010). The bovine herpesvirus 1 (BoHV-1) ICP0 has been reported to target IRF3 for proteasomal degradation (Saira, Zhou, & Jones, 2007). USP7 is a host protein, which, when in the cytoplasm, downregulates NF-κB signaling by deubiquitinating TRAF6 and NEMO. This function is promoted by ICP0, which binds to nuclear USP7 and transfers it to the cytoplasm (Daubeuf et al., 2009). ICP0 also directly interacts with the NF-κB subunits p65 and p50 to prevent nuclear translocation and to induce degradation, respectively (Zhang, Wang, Wang, & Zheng, 2013). The HSV neurovirulence factor ICP34.5 interferes with the STINGTBK1-IRF3 signaling axis by binding to TBK1 to prevent complex formation with IRF3, thus inhibiting IRF3 activation. This requires the N-terminal part of the protein for this action (Ma et al., 2012; Verpooten, Ma, Hou, Yan, & He, 2009). ICP34.5 also binds to IKKα/β and protein phosphatase 1 to induce dephosphorylation of IKKα/β, hindering it from phosphorylating IκB, thus preventing activation of NF-κB, and ultimately impairing DC maturation, although this was studied in the context of TLR4 signaling ( Jin, Yan, Ma, Cao, & He, 2011). Interestingly, ICP34.5 also binds to Beclin-1 to inhibit autophagy, which may prevent delivery of PAMPs to the endosome for TLR9 sensing (Orvedahl et al., 2007). The HSV pUS3 kinase was also shown to inhibit IRF3 and NF-κB signaling (Peri et al., 2008; Sen, Liu, Roller, & Knipe, 2013; Wang, Ni, Wang, & Zheng, 2014; Wang, Wang, Lin, & Zheng, 2013). pUS3 interacts directly with IRF3 and NF-κB (p65 subunit) and mediates their hyperphosphorylation. This blocks dimerization in case of IRF3 and in both cases prevents nuclear translocation and gene induction (Wang et al., 2014; Wang, Wang, Lin, et al., 2013). pUS3 also reduces the polyubiquitination of TRAF6, which is needed for activation of the IKK complex and thus for NF-κB signaling (Sen et al., 2013). This study focused on TLR2 signaling, and it will be interesting to learn whether pUS3 also inhibits TLR9 signaling. The HSV immediate-early ICP27 protein is also involved in the suppression of IRF3 and NF-κB-mediated innate signaling, as an ICP27 deletion mutant of HSV-1 elicited higher cytokine responses than a wild-type virus

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in macrophages (Melchjorsen, Siren, Julkunen, Paludan, & Matikainen, 2006). ICP27 binds to the NF-κB inhibitor IκB, prevents its phosphorylation and ubiquitination, and stabilizes it in this way to prevent NF-κB signaling (Kim et al., 2008). It is currently unknown how ICP27-mediated inhibition of the IRF3-pathway occurs and what the physiological importance of this is. Although the HSV tegument protein VP16 was first reported not to have a direct role in evasion of cytokine expression, but rather to support the expression of evasion molecules, e.g., ICP27 (Mogensen, Melchjorsen, Malmgaard, Casola, & Paludan, 2004), a recent study shows that VP16 blocks both IRF3 and NF-κB signaling (Xing et al., 2013). VP16 was found not to block IRF3 activation directly, but rather to interact with the coactivator CBP (CREB-binding protein) to prevent gene expression. The HSV tegument protein pUS11 inhibits RNA virus-induced IFNβ signaling by interacting with RIG-I and MDA5 to prevent their association with the adaptor MAVS (Xing, Wang, Lin, Mossman, & Zheng, 2012). The tegument protein pUL36 impairs RIG-I-mediated IRF3 activation and IFNβ induction by deubiquitinating TRAF3, which is needed in ubiquitinated form for the recruitment of TBK1 (Wang, Wang, Li, & Zheng, 2013). Since the cytosolic DNA sensor RNA Pol III signals via RIG-I, it is likely that DNA-induced signaling is also inhibited via these mechanism. Although less well studied, VZV also encodes for proteins evading early innate signaling. The immediate-early protein IE62 blocks IRF3 phosphorylation by TBK1, which prevents its activation, although IE62 could not be shown to interact with IRF3 or TBK1 and the IRF3–TBK1 complex remained intact (Sen et al., 2010). Interestingly, the immediate-early ORF47 (open reading frame 47) kinase directly hyperphosphorylates IRF3, which prevents dimerization and thus activation (Zhu et al., 2011). The immediate-early protein ORF61, which contains an E3 ubiquitin ligase RING finger domain homologous to the ICP0 RING finger domain, targets activated IRF3 for proteasomal degradation (Vandevenne et al., 2011).

5. RELEVANCE FOR VACCINE DESIGN Finding efficient prophylactic and therapeutic vaccines against HSV-1 and HSV-2 has been an important but so far unachieved goal of vaccine research for many decades, while safe and useful vaccines against VZV are

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already in use (Coleman & Shukla, 2013; Gershon, 2013). The experimental HSV vaccines tried so far have not provided long-term protection and sterilizing immunity to prevent asymptomatic shedding of the virus (Coleman & Shukla, 2013). It is thought that optimization of DNA vaccines, on which much research effort has been focused in the past 20 years, could lead to progress toward the goal of achieving efficient and lasting humoral and cellular immune responses (Coban, Kobiyama, Jounai, Tozuka, & Ishii, 2013). Especially induction of a strong CD8+ T cell response is lacking in conventional vaccination (Rappuoli, 2007). A plasmid DNA vaccine against HSV-2 is currently in a phase I/II clinical trial (Awasthi & Friedman, 2014), and DNA vaccines are already in use against other viruses in veterinary medicine (Coban et al., 2013). However, the main issue with current DNA vaccines is low immunogenicity (Coban et al., 2013). The recent research on intracellular DNA sensing has revealed that the administered vaccine plasmid fulfills two functions, mediating expression of the antigen for (cross-)presentation by DCs and for activation of T and B cells and binding to cytosolic DNA sensors, thus inducing the cytokine response discussed above, resulting in an adjuvant effect of the vaccine plasmid (Coban et al., 2013). Components of the nucleic acid-sensing system, such as STING and TBK1, are important for successful generation of an efficient adaptive immune response after DNA vaccination (Ishii et al., 2008; Ishikawa et al., 2009). Several attempts have been made to increase this adjuvant effect based on the new knowledge about the DNA recognition system. For example, immunization of mice with plasmids encoding a model antigen and the first proposed DNA sensor DAI resulted in improved proliferation and activation (IFNγ production) of antigen-specific CD8+ T cells and the induction of memory. The Th1 response was also improved (Lladser et al., 2010). CDNs have been suggested as novel adjuvants, which mediate their function by binding to STING and inducing cytokine expression (Dubensky, Kanne, & Leong, 2013). In this context, the recent discovery that cGAS produces cGAMP is of great relevance. Mice immunized intramuscularly with a model antigen (purified OVA protein) in combination with cGAMP as an adjuvant displayed strong, STING-dependent antibody production that was not present in the OVA-only control and CD4+ and CD8+ T cells from these mice produced more cytokines (IFNγ and IL-2) (Li, Wu, et al., 2013). Thus, the new advances in the field of intracellular DNA sensing have already contributed to vaccine development and have potential to be applied in vaccines in the future, especially against intracellular pathogen, including

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alphaherpesviruses. However, further research on the exact mechanisms of action of DNA vaccines, especially in the context of DNA sensing, is needed to develop methods to accurately manipulate the resulting adaptive immune response.

6. CONCLUSIONS AND FUTURE PERSPECTIVE Great progress has been made in the past decade on the innate immune recognition of alphaherpesviruses. We have learned that viral nucleic acids, in particular DNA, are potent stimulators of the first line of defense against these viruses. This has broadened our perception of PAMPs and revealed of a number of new principles of the innate system. The identification of cGAS represents the single most important advance in this field, since the first discovery of innate DNA sensing. For the research that has formed the basis for the current knowledge, HSV-1 has often served as the model pathogen, which is why there is abundant data on DNA sensing of this virus in particular. The knowledge on innate immune sensing of HSV-1 DNA is of importance for vaccine design and development of immune modulation therapy, which in combination with antiviral therapy could lead to improved clinical efficacy of treatment of, e.g., herpes simplex encephalitis. Finally, with the great body of knowledge on immune activation and evasion by HSV1, experimental evidence regarding other alphaherpesviruses, including HSV-2, VZV, and alphaherpesviruses of veterinary relevance, is bound to follow soon. However, before this research can find medical application in humans, numerous questions remain to be investigated. A challenge for future research will be to find experimental procedures to differentiate between “true” DNA sensors primarily responsible for initiation of the immune response to DNA, and the numerous DNA-binding proteins also proposed to be bone fide DNA sensors, but more likely involved in the signaling network downstream in the pathways. Refinement of reconstitution systems will be useful for answering this question. Another central question is by what mechanisms STING can be activated and STING’s exact role within the innate signaling network. This is especially interesting as STING also seems to be involved in immune signaling following virus–cell membrane fusion (Holm et al., 2012) and ER stress (Petrasek et al., 2013), thus suggesting the existence of different types of STING activation. It is already clear that the signaling network employed by PPRs is much more complex and intertwined than previously thought and includes many possibilities for

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redundancy, which complicates research. Unraveling the interplay between the many PRRs and signaling adaptors will be crucial. Moreover, the discovery of nuclear DNA sensors such as IFI16 challenges the concept of an “immunoprivileged nucleus” and necessitates research on how host and pathogen DNA are distinguished in this compartment. It will be interesting to investigate how innate DNA recognition and its evasion by viral proteins are involved in the establishment and maintenance of alphaherpesviral latency. Importantly, the relevance of the novel DNA sensors needs to be confirmed in humans. The detailed study of primary immunodeficiencies which render humans more susceptible to diseases caused by alphaherpesvirus infections, as is known for TLR3 deficiency (Zhang et al., 2007), will play a key role in achieving this goal.

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CHAPTER THREE

Molecular Biology of Potyviruses Frédéric Revers*,†,1, Juan Antonio García{,1 *INRA, UMR 1332 de Biologie du fruit et Pathologie, Villenave d’Ornon, France † Universite´ de Bordeaux, UMR 1332 de Biologie du fruit et Pathologie, Villenave d’Ornon, France { Centro Nacional de Biotecnologı´a (CNB-CSIC), Campus Universidad Auto´noma de Madrid, Madrid, Spain 1 Corresponding authors: e-mail address: [email protected]; [email protected]

Contents 1. Introduction 2. Genera of the Family Potyviridae and the Main Differences in Genome Structures 3. Biological and Biochemical Features of Potyviral Proteins 3.1 P1 3.2 HCPro 3.3 P3, 6K1, and PIPO 3.4 CI 3.5 6K2 and NIa 3.6 NIb 3.7 CP 4. Virus Multiplication 4.1 Subcellular localization of potyvirus multiplication 4.2 Viral and plant factors involved in potyvirus multiplication 4.3 Putative functions of these factors during potyvirus multiplication 5. Virus Movement 5.1 Intracellular and cell-to-cell movements 5.2 Long-distance movement 6. Virus Transmission 6.1 Transmission by aphids 6.2 Seed transmission 7. Plant/Potyvirus Interactions in Compatible Pathosystems 7.1 Evolutionary abilities of potyviruses to adapt to their hosts 7.2 HCPro: A key pathogenicity determinant as suppressor of RNA silencing 7.3 Symptomatology 8. Biotechnological Applications of Potyviruses 9. Concluding Remarks Acknowledgments References

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Abstract Potyvirus is the largest genus of plant viruses causing significant losses in a wide range of crops. Potyviruses are aphid transmitted in a nonpersistent manner and some of them Advances in Virus Research, Volume 92 ISSN 0065-3527 http://dx.doi.org/10.1016/bs.aivir.2014.11.006

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are also seed transmitted. As important pathogens, potyviruses are much more studied than other plant viruses belonging to other genera and their study covers many aspects of plant virology, such as functional characterization of viral proteins, molecular interaction with hosts and vectors, structure, taxonomy, evolution, epidemiology, and diagnosis. Biotechnological applications of potyviruses are also being explored. During this last decade, substantial advances have been made in the understanding of the molecular biology of these viruses and the functions of their various proteins. After a general presentation on the family Potyviridae and the potyviral proteins, we present an update of the knowledge on potyvirus multiplication, movement, and transmission and on potyvirus/plant compatible interactions including pathogenicity and symptom determinants. We end the review providing information on biotechnological applications of potyviruses.

1. INTRODUCTION Potyvirus is the largest genus of plant viruses causing significant losses in a wide range of crops. Potyviruses are aphid transmitted in a nonpersistent manner and some of them are also seed transmitted. As important pathogens, potyviruses are much more studied than other plant viruses belonging to other genera and their study covers many aspects of plant virology, such as functional characterization of viral proteins, molecular interaction with hosts and vectors, structure, taxonomy, evolution, epidemiology, and diagnosis. Biotechnological applications of potyviruses are also being explored. Understanding the molecular biology of these viruses and the functions of their various proteins is a prerequisite to develop new resistance strategies. During this last decade, substantial advances have been made in this topic, since the last reviews written before 2004 (Revers, Le Gall, Candresse, & Maule, 1999; Rajama¨ki, Ma¨ki-Valkama, Ma¨kinen, & Valkonen, 2004; Urcuqui-Inchima, Haenni, & Bernardi, 2001). In particular, technical improvements have played important roles in producing new findings. This is the case for the production of potyvirus infectious clones from engineered viral cDNA, which is less arduous and allows identifying viral molecular determinants playing significant role during viral infection. In addition, breakthroughs in plant imaging techniques have highlighted in planta interactions between plant and potyviral proteins and their localization in cellular compartments. In addition, improvement of purification techniques of protein complexes from infected plants has revealed new host factors involved in virus infection.

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This review focuses on these new advances, particularly on the new host and viral determinants involved in the potyviral infection. We first provide general information on the family Potyviridae and the potyviral proteins. Then, we present an update of the knowledge concerning potyvirus multiplication, movement, and transmission with inexorable reminders of some classic data, and on potyvirus/plant compatible interactions (pathogenicity and symptom determinants, symptom development). We end the review with information on biotechnological applications of potyviruses.

2. GENERA OF THE FAMILY POTYVIRIDAE AND THE MAIN DIFFERENCES IN GENOME STRUCTURES The Potyviridae family comprises eight genera of viruses, the members of which are plant-infecting single-stranded positive-sense RNA viruses, with flexible and filamentous virus particles. These genera have been differentiated in terms of genome composition and structure, sequence similarity, and vector organisms responsible of their plant-to-plant transmission (Adams et al., 2011). In the latest report of the International Committee on Taxonomy of Viruses, the family Potyviridae includes 176 virus species (Adams et al., 2011). Most viruses of the family belong to the genus Potyvirus with 146 virus species. Potyviruses and viruses of the genera Brambyvirus, Ipomovirus, Macluravirus, Poacevirus, Rymovirus, and Tritimovirus have monopartite genomes, whereas viruses of the genus Bymovirus have a bipartite genome. The genomic RNAs of potyvirids (i.e., viruses belonging to the family Potyviridae) contain a single open reading frame (ORF) that codes for a major polyprotein, which is proteolytically processed by virus-encoded proteinases (Fig. 1). Potyvirid RNAs are flanked by a 50 -noncoding region (NCR), with a terminal protein (VPg) covalently linked to the 50 -terminal end (Siaw, Shahabuddin, Ballard, Shaw, & Rhoads, 1985), and a 30 -NCR followed by a polyadenylate tract whose length is variable (Laı´n, Riechmann, Me´ndez, & Garcı´a, 1988; Riechmann, Laı´n, & Garcı´a, 1990). The central and carboxy-terminal regions of the polyprotein in potyviruses have a conserved organization and encode the mature viral proteins P3–6K1–CI–6K2– VPg–NIaPro–NIb–CP, which is also the case with the polyprotein encoded by RNA1 of bymoviruses. Processing of this part of the polyprotein is carried out by the proteinase NIaPro (Adams, Antoniw, & Beaudoin, 2005). A universally conserved feature of this genomic region is the existence of a short ORF (pretty interesting potyvirus ORF, PIPO) embedded within

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Figure 1 Genomic maps of members of the family Potyviridae. The long open reading frame is represented as a box divided in final products by black lines. PIPO ORF is indicated as a striped area below the P3 region. The terminal protein (VPg) is represented as a black ellipse. Features that are not shared by all potyvirids are highlighted by different colors (gray shades in the print version): HCpro in blue (dark gray in the print version), potyvirus-type P1s in gray, P1b-type P1s in black, proteins encoded by the RNA 2 of bymoviruses in yellow (white in the print version) and light yellow (white in the print version), and the extra protein HAM and the AlkB domain of the brambyvirus Blackberry virus Y (Susaimuthu, Tzanetakis, Gergerich, & Martin, 2008), in pink (light gray in the print version).

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the P3-encoding region in a reading frame different from the polyprotein. It is used to express the P3N–PIPO protein using a +1 frameshift at a place defined by a GA6 sequence (Chung, Miller, Atkins, & Firth, 2008). Ipomoviruses are exceptional in that they can contain a Maf/HAM1-like gene sequence between NIb- and CP-coding regions (Mbanzibwa, Tian, Mukasa, & Valkonen, 2009). Two proteinases, P1 and HCPro, are produced from the amino-terminus of the potyviral polyprotein. They cleave themselves from the polyprotein. Homologous proteins are produced from the N-terminal regions of the polyproteins of rymoviruses, tritimoviruses, poaceviruses, brambyviruses, and at least one ipomovirus (Adams et al., 2011). However, the P1 proteins of potyviruses and rymoviruses appear to belong to a phylogenetic group distinct from that formed by the tritimovirus and poacevirus P1 proteins, and some ipomoviruses can have two P1 proteins, one of each family, and lack HCPro (Rodamilans, Valli, & Garcı´a, 2013; Valli, Lo´pez-Moya, & Garcı´a, 2007). Moreover, macluraviruses lack any P1 protein and have a smaller HCPro than other potyvirids (Kondo & Fujita, 2012). The polyprotein encoded by RNA2 of bymoviruses is unique for this genus and encodes two proteins, of which the first protein has domains with sequence similarities to HCPro (You & Shirako, 2010). It has been suggested that the emergence of the potyviral P1 ancestor in the evolutionary history of potyviruses might have been involved in the extraordinary radiation of this group of viruses (Rodamilans et al., 2013). As most of the data published these last years have come from studies on virus species belonging to the genus Potyvirus, we will concentrate here on the features of this genus.

3. BIOLOGICAL AND BIOCHEMICAL FEATURES OF POTYVIRAL PROTEINS Proteolytic processing of the potyviral polyprotein results in 11 mature proteins, including P3N–PIPO, but multiple partially processed intermediates are also produced, some of which are likely to be functionally relevant (Merits et al., 2002). The activities of the different potyviral proteins appear to be brought out in a coordinated and interdependent manner. Thirtythree interactions between potyviral proteins (including self-interactions) have been detected by testing 58 protein combinations in planta (Elena & Rodrigo, 2012; Zheng et al., 2011; Zilian & Maiss, 2011). The large network of connections with different viral (and host) proteins likely contributes to the multifunctional nature of many potyviral proteins. In this section,

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Figure 2 Relevant features and proposed functions of potyviral proteins. The long open reading frame is represented as a box divided in final products by black lines. PIPO ORF is indicated as a striped area below the P3 region. The terminal protein (VPg) is represented as a black ellipse. Arrows starting from the three proteases (P1, HCPro, and NIa) above the box indicate the cleavage sites in the polyprotein. All the known functions of each protein indicated with a blue (dark gray in the print version) dot are given at the end of the dotted lines starting from the given protein.

we provide a basic statement on the biochemical, structural, and interaction properties, posttranslational modification, and subcellular localization for each of the potyvirus proteins. Only basic information on protein function is given in this section as most of these data are detailed in the following sections and summarized in Fig. 2.

3.1. P1 The potyviral P1 protein is a serine protease that cleaves at its own C-terminus (Verchot, Koonin, & Carrington, 1991; Fig. 2). P1 functions in trans to stimulate genome amplification (Verchot & Carrington, 1995b). It is not essential for viral viability but the separation of P1 and HCPro is required (Pasin, Simo´n-Mateo, & Garcı´a, 2014; Verchot & Carrington, 1995a). P1 is a highly basic protein (Valli et al., 2007) and has the ability to interact with nucleic acids in vitro (Brantley & Hunt, 1993; Soumounou & Laliberte´, 1994), but the functional relevance of these properties is unknown. There are experimental data supporting the hypothesis that P1 stimulates the RNA silencing suppression activity of the protein HCPro (Anandalakshmi et al., 1998; Pruss et al., 2004; Rajama¨ki et al., 2005; Valli et al., 2007). However, other data suggest that the effect on silencing suppression might be due to enhancement of the synthesis of HCPro when it

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is preceded by P1 rather to a specific activity of the P1 protein (Tena Ferna´ndez et al., 2013). In addition, P1 enhances virus infection even in RNA silencing-deficient plants, which suggests that this protein plays a role independent of RNA silencing suppression (Pasin et al., 2014). Although its protease domain, which is placed at the C-terminal region of the protein, is well conserved, P1 is the most divergent potyviral protein in size (30–63 kDa) and sequence (Adams, Antoniw, & Fauquet, 2005; Valli et al., 2007; Yoshida, Shimura, Yamashita, Suzuki, & Masuta, 2012). This is a consequence of the large variability of its N-terminal region. This region, which appears to be highly disordered, negatively regulates P1 self-cleavage (Pasin et al., 2014). It is also interesting that a new small ORF, called PISPO, has been identified inside the coding sequence of the P1-coding sequence of some potyviruses that infect sweet potato. This ORF could be translated by a frameshift similar to that involved in the expression of the PIPO ORF (Clark et al., 2012; Li, Xu, Abad, & Li, 2012). It is unknown if PISPO can confer any specific advantage to infect sweet potato. The interaction of P1 with the host protein Rieske Fe/S (Table 1) has been described, but the role of this interaction has yet to be unraveled (Shi et al., 2007).

3.2. HCPro HCPro is probably the most studied potyviral protein. The name of this protein derives from its first discovered function: Helper Component (HC) for aphid transmission (Govier, Kassanis, & Pirone, 1977). HCPro is a cysteine protease that self-cleaves at its C-terminus (Carrington, Freed, & Sanders, 1989; Fig. 2) and is involved in multiple functions (Maia, Haenni, & Bernardi, 1996; Syller, 2005), some of them derived from its ability to suppress RNA silencing (Dunoyer, Lecellier, Parizotto, Himber, & Voinnet, 2004; Gonza´lez-Jara et al., 2005; Jay et al., 2011; Kasschau & Carrington, 2001; Kasschau et al., 2003; Mallory, Reinhart, Bartel, Vance, & Bowman, 2002; Soitamo, Jada, & Lehto, 2011). HCPro has been shown to aggregate in the form of cytoplasmic amorphous inclusions in some potyviral infections (De Mejia, Hiebert, Purcifull, Thornbury, & Pirone, 1985), although it can also accumulate as a soluble protein (Ravelonandro, Peyruchaud, Garrigue, de Marcillac, & Dunez, 1993), and there is evidence suggesting that it can play a role in the nucleus of infected cells (Sahana et al., 2014). Moreover, HCPro has been detected at

Table 1 Host factors interacting with the potyvirus proteins Potyvirus Origin of the viral and Putative function in viral protein Plant interactor Method plant partners infection

P1

HCPro

References

Shi, Chen, Hong, Chen, and Adams (2007)

Chloroplastic Rieske Fe/S protein

Y2H, Co-IP

SMV/Pinellia ternata Unknown

60S ribosomal subunit

Affinity purification

TEV/N. benthamiana Stimulation of translation Martı´nez and Daro´s (2014)

eIF4E/eIF(iso)4E

Y2H, BiFC

PVA-PVY-TEV/ tobacco-potato

Interaction associated with viral replication vesicles

Ala-Poikela, Goytia, Haikonen, Rajama¨ki, and Valkonen (2011)

Calmodulin-related Y2H, surface TEV-TuMVprotein plasmon resonance ClYVV/tobacco

Regulation of RNA silencing suppressor activity

Anandalakshmi et al. (2000) and Nakahara et al. (2012)

Ethylene-inducible Y2H, in vitro and transcription factor in vivo pull-down RAV2

TEV/tobacco; TuMV/A. thaliana

Mediating HCPro silencing suppressor activity

Anandalakshmi et al. (2000) and Endres et al. (2010)

RNA methyltransferase HEN1

ELISA-binding assay

ZYMV/A. thaliana

Inhibition of HEN1 Jamous et al. (2011) activity in RNA silencing

Proteasome proteins (PAE1, PAE2, PAA, PBB, and PBE)

Y2H, in vitro binding assays, BiFC

PVY/A. thaliana; LMV/lettuceA. thaliana; PRSV/ papaya

Inhibition of proteasome Ballut et al. (2005), Jin, Ma, Dong, Jin, et al. (2007), Jin, protease and RNAse Ma, Dong, Li, et al. (2007), activities by HCPro Dielen et al. (2011), and Sahana et al. (2012)

Microtubuleassociated protein HIP2

Y2H, BiFC

PVA/potato– tobacco

Reduced PVA accumulation in inoculated leaves of HIP2-silenced plants

Guo et al. (2003) and Haikonen, Rajama¨ki, Tian, and Valkonen (2013)

RING finger protein HIP1

Y2H

PVA/potato

Unknown

Guo, Spetz, Saarma, and Valkonen (2003)

Chloroplast precursor of ferredoxin-5

Y2H, BiFC

SCMV/maize

Symptom development? Cheng et al. (2008)

Calreticulin

Y2H, BiFC

PRSV/papaya

Plant calcium signaling pathways?

Shen et al. (2010)

Chloroplast division-related factor NtMinD

Y2H, BiFC

PVY/tobacco

Chloroplast division?

Jin, Ma, Dong, Jin, et al. (2007) and Jin, Ma, Dong, Li, et al. (2007)

P3

RubisCO subunits RbcL and RbcS

Y2H, Co-IP

SYSV-OYDVUnknown SMV-TuMV/onion

P3N– PIPO

Cation-binding protein PCaP1

Y2H, Co-IP, BiFC TuMV/A. thaliana

Cell-to-cell movement

Vijayapalani, Maeshima, Nagasaki-Takekuchi, and Miller (2012)

RubisCO subunits RbcL and RbcS

Y2H

SYSV-TuMV/ onion

Unknown

Lin et al. (2011)

eIF4E

ELISA-based assays, BiFC

LMV/lettuce

Cell-to-cell movement?

Tavert-Roudet et al. (2012)

CI

Lin et al. (2011)

Continued

Table 1 Host factors interacting with the potyvirus proteins—cont'd Potyvirus Origin of the viral and Putative function in viral protein Plant interactor Method plant partners infection

VPg

Antiviral defense?

References

Jime´nez et al. (2006)

Chloroplastic photosystem I, PSI-K

Y2H, pull-down

PPV-TVMV/ N. benthamiana– A. thaliana

P58IPK

Y2H, pull-down

TEV/N. benthamiana Virulence factor

Bilgin, Liu, Schiff, and Dinesh-Kumar (2003)

eIF4E/eIF(iso)4E

Y2H, in vitro binding assays, BiFC

Several potyvirus/ plant pathosystems

Wittmann, Chatel, Fortin, and Laliberte´ (1997), Leonard et al. (2000, 2004), Schaad, Anderberg, and Carrington (2000), Beauchemin, Boutet, and Laliberte´ (2007), and Charron et al. (2008)

Fibrillarin

Y2H, BiFC

PVA/N. benthamiana Unknown

Rajama¨ki and Valkonen (2009)

PABP

ELISA-based binding assay, Co-IP

TuMV/Brassica perviridis

Viral RNA translation/ replication

Leonard et al. (2004) and Beauchemin and Laliberte´ (2007)

eEF1A

TAP, ELISA-based TuMV/A. thaliana binding assay

Viral RNA translation/ replication

Thivierge et al. (2008)

Viral RNA translation/ replication

AtRH8

Y2H, BiFC

PPV/PeachA. thaliana

Viral RNA translation/ replication?

Huang, Wei, Laliberte´, and Wang (2010)

PVIP

Y2H

PSbMV–LMV– TuMV/peaN. benthamiana– A. thaliana

Movement?

Dunoyer, Thomas, et al. (2004)

NIaPro

Methionine sulfoxide reductase B1

Y2H

PRSV/papaya

Unknown

Gao et al. (2012)

NIb

eEF1A

TAP, ELISA-based TuMV/A. thaliana binding assay

Viral RNA translation/ replication

Thivierge et al. (2008)

PABP

Y2H, TAP

ZYMV/Cucumber; Viral RNA translation/ TuMV/A. thaliana replication

Wang, Ullah, and Grumet, (2000) and Dufresne, Thivierge, et al. (2008)

Hsc70-3

TAP

TuMV/A. thaliana

Dufresne, Thivierge, et al. (2008)

SUMOconjugating enzyme SCE1

Y2H, BiFC, FRET

TuMV-TEV-SMV/ SUMOylation of NIb A. thaliana– N. benthamiana

Xiong and Wang (2013)

RubisCO-LSU

ELISA-based binding assay, bacterial twohybrid system

PVY/Tobacco

Feki et al. (2005)

CP

Viral RNA translation/ replication

Continued

Table 1 Host factors interacting with the potyvirus proteins—cont'd Potyvirus Origin of the viral and Putative function in viral protein Plant interactor Method plant partners infection

CP degradation

References

Hafre´n et al. (2010) and Hofius et al. (2007)

CPIP

Co-IP, Y2H

PVA, PVY/ N. benthamianatobacco

HSP70

Co-IP

PVA/N. benthamiana CP degradation

Hafre´n, Hofius, Ronnholm, Sonnewald, and Ma¨kinen (2010)

Chloroplastic 37 kDa protein

ELISA-based binding assay

TuMV/lettuce

McClintock, Lamarre, Parsons, Laliberte´, and Fortin (1998)

Unknown

Y2H, yeast two hybrid; Co-IP, coimmunoprecipitation; BiFC, bimolecular fluorescence complementation; FRET, fluorescence resonance energy transfer; TAP, tandem affinity purification; ClYVV, Clover yellow vein virus; LMV, Lettuce mosaic virus; OYDV, Onion yellow dwarf virus; PPV, Plum pox virus; PRSV, Papaya ringspot virus; PSbMV, Pea Seedborne mosaic virus; PVA, Potato virus A; PVY, Potato virus Y; SMV, Soybean mosaic virus; SCMV, Sugarcane mosaic virus; SYSV, Shallot yellow stripe virus; TEV, Tobacco etch virus; TuMV, Turnip mosaic virus; TVMV, Tobacco vein mottling virus; ZYMV, Zucchini yellow mosaic virus.

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the ends of potyvirus virions (Manoussopoulos, Maiss, & Tsagris, 2000; Torrance et al., 2006). Recent results have revealed that HCPro is required to stabilize CP and for the proper yield and infectivity of potyviral progeny (Valli, Gallo, Calvo, Pe´rez, & Garcı´a, 2014). It has been suggested that HCPro acts as a dimer (Guo, Merits, & Saarma, 1999; Thornbury, Hellmann, Rhoads, & Pirone, 1985), and, probably, in the form of oligomers of higher order (Plisson et al., 2003; Ruiz-Ferrer et al., 2005). Three structural domains can be distinguished in the HCPro protein: the N- and C-terminal regions, of approximately 100 amino acids, and the central domain, of approximately 250 amino acids (Plisson et al., 2003). The C-terminal domain is responsible for the proteolytic activity of the protein. The atomic structure of this domain has been determined for the HCPro protein of Turnip mosaic virus (TuMV; Guo, Lin, & Ye, 2011). The N-terminal domain is required for aphid transmission of the virus, but most of the HCPro functions are based on the central region of the protein (Kasschau & Carrington, 2001; Varrelmann, Maiss, Pilot, & Palkovics, 2007). HCPro has been shown to interact with a large number of viral and host proteins, but, with the exception of the involvement in aphid transmission of the interaction with the CP protein (Andrejeva et al., 1999; Blanc et al., 1997; Roudet-Tavert et al., 2002), the functional relevance of these interactions is still to be characterized in detail. Other viral proteins, in addition to CP, with which HCPro can interact, are CI (Choi, Stenger, & French, 2000; Guo, Rajama¨ki, Saarma, & Valkonen, 2001; ulg, & Saarma, 1999), Zilian & Maiss, 2011), P1 (Merits, Guo, Ja¨rvek€ and VPg and its precursor NIa (Guo et al., 2001; Roudet-Tavert et al., 2007; Yambao, Masuta, Nakahara, & Uyeda, 2003). Several host proteins were shown to interact with HCPro (Table 1): a calmodulin-related protein (Anandalakshmi et al., 2000; Nakahara et al., 2012), the ethylene-inducible transcription factor RAV2 (Endres et al., 2010), and the small RNA methyltransferase HEN1 ( Jamous et al., 2011); several proteasome components (PAE1, PAA, PBB, and PBE; Dielen et al., 2011; Jin, Ma, Dong, Jin, et al., 2007; Jin, Ma, Dong, Li, et al., 2007), both eIF4E and eIF(iso)4E (Ala-Poikela et al., 2011), the microtubuleassociated protein HIP2 (Guo et al., 2003; Haikonen et al., 2013), the RING finger protein HIP1 (Guo et al., 2003), the chloroplast divisionrelated factor NtMinD ( Jin, Ma, Dong, Li, et al., 2007), the chloroplast precursor of ferredoxin-5 (Cheng et al., 2008), and calreticulin (Shen et al., 2010).

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3.3. P3, 6K1, and PIPO The protein P3 is one of the least well-characterized potyviral proteins. Unlike most potyviral proteins, P3, along with 6K1 and 6K2, does not bind to viral RNA (Merits, Guo, & Saarma, 1998). The protein P3 has been found associated with the cytoplasmic cylindrical inclusions formed by the viral protein CI (Rodrı´guez-Cerezo, Ammar, Pirone, & Shaw, 1993), and with nuclear inclusions formed by viral proteins NIa and NIb (Langenberg & Zhang, 1997). More recently, transient expression experiments in healthy and infected leaves showed that P3 is targeted to the membranes of the endoplasmic reticulum (ER) and forms inclusions associated with the Golgi apparatus that traffic along the actin filaments and colocalize with replication vesicles (Cui, Wei, Chowda-Reddy, Sun, & Wang, 2010; Eiamtanasate, Juricek, & Yap, 2007). Two hydrophobic regions were identified in P3 for several potyviruses and the one located in the C-terminal end of the protein was shown to be responsible for the ER targeting of P3 (Cui et al., 2010; Eiamtanasate et al., 2007). P3 interacts with the potyviral proteins CI, NIb, and NIa (Guo et al., 2001; Lin et al., 2009; Merits et al., 1999; Zilian & Maiss, 2011), but until now only one host factor corresponding to Rubisco subunits directly interacts with P3 (Lin et al., 2011; Table 1). Though a direct role of this host factor in potyviral infection has not been shown, the recent observation of the involvement of Rubisco in tobamovirus infections (Bhat et al., 2013; Zhao et al., 2013) may suggest a similar role with potyviruses. P3 is required for viral replication (Klein, Klein, Rodrı´guez-Cerezo, Hunt, & Shaw, 1994), and there is abundant information highlighting its relevance for viral pathogenicity and symptomatology (see Section 7); however, the precise role of P3 remains obscure. Based on the fact that the proteolytic splitting between P3 and 6K1 is not essential for virus infectivity, it has been hypothesized that P3–6K1, rather than P3 and 6K1, might be the main functional product, and its proteolytic processing would have a regulatory role (Riechmann, Cervera, & Garcı´a, 1995). However, processing at the P3–6K1 junction still affected symptom expression, suggesting that 6K1 alone may has a relevant role in potyviral infection. Supporting this possibility, Waltermann and Maiss (2006) clearly showed that 6K1 from Plum pox virus (PPV) was detected exclusively as a mature protein of 6 kDa in Nicotiana benthamiana, and Hong, Chen, Shi, and Chen (2007) showed that an antibody against Soybean mosaic virus

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(SMV) 6K1 labeled the cell periphery of Pinellia ternata which does not correspond to the P3 localization in ER. An undefined role in potyvirus multiplication has been proposed for this peptide (Kekarainen, Savilahti, & Valkonen, 2002; Merits et al., 2002). Before the discovery of P3N–PIPO, the functional relevance of its coding sequence was already noticed for the tritimovirus Wheat streak mosaic virus (WSMV; Choi, Horken, Stenger, & French, 2005). A role of P3N–PIPO in virus movement, in conjunction with CI and the host factor pCaP1 (Table 1), has been demonstrated (Vijayapalani et al., 2012; Wen & Hajimorad, 2010).

3.4. CI The CI protein forms the cylindrical inclusions in the form of pinwheels typical in the cytoplasm of cells infected with potyviruses (Edwardson & Christie, 1996). As mentioned above for HCPro, CI is a multipartner and multifunctional protein (Sorel, Garcia, & German-Retana, 2014). This protein has ATPase and RNA helicase activities (Eagles, Balmori-Melia´n, Beck, Gardner, & Forster, 1994; Laı´n, Martı´n, Riechmann, & Garcı´a, 1991; Laı´n, Riechmann, & Garcı´a, 1990), which are required for virus RNA replication (Ferna´ndez et al., 1997). Ultrastructural and temporal observations of the cylindrical inclusions as well as genetic analyses show that the CI protein acts in collaboration with P3N–PIPO in aiding virus movement (Carrington, Jensen, & Schaad, 1998; Go´mez de Cedro´n, Osaba, Lo´pez, & Garcı´a, 2006; Roberts, Wang, Findlay, & Maule, 1998; Rodrı´guez-Cerezo et al., 1997; Wei, Zhang, et al., 2010), but it is not known whether the enzymatic activities of the protein are required for this function. On the other hand, CI protein has been found associated with the ends of potyviral virions, and it has been suggested that it may provide a molecular motor function to help virus translocation through plasmodesmata (PD) and particle disassembly (Gabrenaite-Verkhovskaya et al., 2008). Furthermore, the CI protein acts as a virulence factor for different resistance genes (Sorel et al., 2014). Three host factors were shown to interact with CI, the translation initiation factor eIF4E (Tavert-Roudet et al., 2012), a component of the chloroplastic photosystem I (PSI-K; Jime´nez, Lo´pez, Alamillo, Valli, & Garcı´a, 2006) and a plant ortholog of a doublestranded RNA-dependent protein kinase inhibitor (P58IPK; Bilgin et al., 2003; Table 1).

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3.5. 6K2 and NIa NIa is the largest protein that forms a crystalline inclusion produced by many potyviruses (Kassanis, 1939; Knuhtsen, Hiebert, & Purcifull, 1974). These inclusions are mainly found in the nucleus but they can also be detected in the cytoplasm of the infected cells (Martı´n, Garcı´a, Cervera, Goldbach, & van Lent, 1992). NIa is partially processed to produce VPg and NIaPro (Dougherty & Dawn Parks, 1991). Different aspects of the protein VPg has been extensively studied ( Jiang & Laliberte´, 2011). VPg is intrinsically disordered, and this property confers upon this protein the flexibility required to interact with a variety of different partners, including itself (Oruetxebarria et al., 2001), allowing it to participate in diverse processes (Grzela et al., 2008; Rantalainen, Eskelin, Tompa, & Ma¨kinen, 2011; Rantalainen et al., 2008). The free VPg is the major form of protein linked to the 50 -end of the genomic RNA (Hari, 1981; Riechmann, Laı´n, & Garcı´a, 1989; Shahabuddin, Shaw, & Rhoads, 1988), although the complete NIa (VPg–NIaPro) has also been detected at the end of the RNA (Mathur & Savithri, 2012; Murphy, Rhoads, Hunt, & Shaw, 1990). When VPg is part of the NIa, it is localized both in the cytoplasm and in the nucleus of the infected cells (Beauchemin et al., 2007; Cotton et al., 2009; Rajama¨ki & Valkonen, 2009), whereas when it is part of the 6K2–VPg–NIaPro product, VPg is targeted to membranous factories induced by the virus where it plays a key role in viral RNA replication (Beauchemin et al., 2007; Wei & Wang, 2008). VPg contains a nucleotide-binding motif and, when it is bound to the NIaPro domain, preferably in cis, has NTPase activity (Mathur & Savithri, 2012). At the moment, it is only possible to speculate as to the functional relevance of this activity. It has been shown that VPg can be phosphorylated (Hafre´n & Ma¨kinen, 2008; Mathur et al., 2012; Puustinen, Rajama¨ki, Ivanov, Valkonen, & Ma¨kinen, 2002), and this posttranslational modification might be very important for the regulation of the multiple functions in which this protein is involved. VPg interacts with most of the potyviral proteins (Elena & Rodrigo, 2012; Jiang & Laliberte´, 2011; and references therein) and with several host factors: the eukaryotic initiation factor eIF4E (reviewed in recent papers: Robaglia & Caranta, 2006; Truniger & Aranda, 2009; Wang & Krishnaswamy, 2012), the nucleolar protein fibrillarin (Rajama¨ki & Valkonen, 2009), PABP (Beauchemin & Laliberte´, 2007), and a RNA helicase-like protein from peach and Arabidopsis (AtRH8) which is related to eIF4A (Huang et al., 2010; Table 1).

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NIaPro is the protease domain responsible for the proteolytic processing of the central and C-terminal regions of the potyviral polyprotein (Adams, Antoniw, & Beaudoin, 2005; Fig. 2). The functional features of the NIaPro domain have been characterized in detail in different experimental systems (Adams, Antoniw, & Beaudoin, 2005), and the structural basis for its activity and substrate specificity has been studied by X-ray crystallography (Nunn et al., 2005; Phan et al., 2002; Sun, Austin, Tozser, & Waugh, 2010). Variation in cleavage efficiency at the different NIaPro cleavage sites suggests that the maturation of potyviral proteins is highly regulated and plays a relevant role in the control of the potyviral infection. In addition to its proteinase activity, NIaPro has DNase activity. It has been speculated that degradation of host DNA by the protein NIa located in the nucleus might play some regulatory role in host gene expression relevant for the viral infection (Anindya & Savithri, 2004).

3.6. NIb NIb together with NIa forms the crystalline inclusions mentioned above (Kassanis, 1939; Knuhtsen et al., 1974). NIb is the RNA-dependent RNA polymerase, or RNA replicase, responsible for potyviral genome replication (Hong & Hunt, 1996). It is thought that NIb is targeted to the membranous structures where viral RNA replication takes place via its interaction with VPg and NIaPro domains of the 6K2–VPg–NIaPro product (Dufresne, Thivierge, et al., 2008). Interactions of NIb with the host proteins eEF1A, PABP, and Hsc70-3 should contribute to the formation of functional replication complexes (Dufresne, Thivierge, et al., 2008; Dufresne, Ubalijoro, Fortin, & Laliberte´, 2008; Thivierge et al., 2008; Wang et al., 2000). NIb uridylylates the protein VPg and uses the resulting product to prime viral RNA synthesis (Anindya, Chittori, & Savithri, 2005; Puustinen & Ma¨kinen, 2004). Very little is known about the role that NIb could play in the nucleus of the infected cells (Li, Valdez, Olvera, & Carrington, 1997; Restrepo, Freed, & Carrington, 1990). It has been recently reported that NIb interacts with the SUMO-conjugating enzyme SCE1 both in the nucleus and in the cytoplasm (Xiong & Wang, 2013; Table 1). It has been postulated that the nucleocytoplasmic transport of the complex NIb/SCE1 or the SUMOylated form of NIb may be important for potyviral infection. But the possibility also has been suggested that SUMOylation could directly regulate NIb activity, or NIb/SCE1 interaction disturbs the pattern of

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SUMOylation of cellular proteins, generating an environment more favorable for virus multiplication (Xiong & Wang, 2013).

3.7. CP The principal function of the CP protein is the encapsidation of the viral genome. About 2000 CP subunits helically arranged around the genomic RNA form the potyviral virions, which are flexuous rods of approximately 11–13 nm in diameter by 680–900 nm in length (Adams et al., 2011; Lo´pez-Moya, Valli, & Garcı´a, 2009). A first estimate of the structure of the potyvirus particles was obtained by combining electron microscopy and fiber diffraction (Kendall et al., 2008). SMV particles have approximately 8.9 subunits of CP per turn with a helical pitch of 33 A˚. The central region of the potyviral CP is highly conserved in potyviruses and forms the core of the virus particles (Dolja, Haldeman, Robertson, Dougherty, & Carrington, 1994; Jagadish, Huang, & Ward, 1993; Varrelmann & Maiss, 2000; Voloudakis et al., 2004). However, also the N- and C-terminal domains have been found to be crucial for CP intersubunit interactions involved in the initiation of virus assembly (Anindya & Savithri, 2003; Kang et al., 2006; Seo, Vo Phan, Kang, Choi, & Kim, 2013). The N-terminal region of CP is exposed at the surface of the viral particle, whereas the localization of its C-terminus seems to depend on the particular potyvirus species (Allison et al., 1985; Shukla, Strike, Tracy, Gough, & Ward, 1988). The primary sequence of the N-terminal region of CP is highly variable among potyviruses and was predicted to be disordered (Ksenofontov et al., 2013; Rybicki & Shukla, 1992; Ward, McKern, Frenkel, & Shukla, 1992). Several kinds of posttranslational modifications of the potyviral CP were described. The CP of Potato virus A (PVA) was shown to be phosphorylated, and it was reported that this modification reduces its affinity by the viral RNA (Ivanov et al., 2003; Ivanov, Puustinen, Merits, Saarma, & Ma¨kinen, 2001). The PPV CP is phosphorylated and O-GlcNAcylated (Chen et al., 2005; Ferna´ndez-Ferna´ndez, Camafeita, et al., 2002; Scott et al., 2006; Sˇubr, Rysˇlava´, & Kollerova´, 2007; Sˇubr et al., 2010). Several O-GlcNAc-modified sites (Kim et al., 2011; Pe´rez et al., 2013, 2006) and an amino acid whose mutation appears to alter the phosphorylation status of the protein (Sˇubr et al., 2010) have been mapped at the N-terminus of PPV CP, whereas a phosphorylated residue was mapped at the end of the core region of the PVA CP. O-GlcNAcylation of PPV CP enhances viral

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infection, but is not essential for virus viability (Chen et al., 2005; Pe´rez et al., 2013), and an important regulatory role was proposed for phosphorylation of CP in PVA infection (Ivanov et al., 2003). It has been hypothesized that these posttranslational modifications may be relevant control elements to regulate the fraction of genomic RNA allocated for translation, replication, and propagation during the different steps of the infection process. Cellular chaperones appear to play important roles in this regulation (Aparicio et al., 2005; Hafre´n et al., 2010; Hofius et al., 2007; Sugio, Dreos, Aparicio, & Maule, 2009). The potyviral CP has NTPase activity, and it is probable that this activity is also relevant in the regulatory mechanism (Rakitina et al., 2005). Besides encapsidation of the viral genome, other functions of CP in genome amplification, movement, and transmission have also been described (see the next sections). Similar to P3, CP has been reported to interact with the host Rubisco (Feki et al., 2005; Table 1), which, as mentioned above, appears to be an import host factor for viral infections (Bhat et al., 2013; Zhao et al., 2013). Thus, we cannot rule out the possibility that the interaction of the potyviral CP with Rubisco plays a role in viral infection and/or plant defensive responses.

4. VIRUS MULTIPLICATION Once potyviral virions enter plant cells, viral RNA is released in cytoplasm after a poorly understood step of virion disassembly and is directly translated. Then, thanks to the production of the viral proteins, RNA replication occurs to produce first minus-strand copies and then new positivestrand RNA molecules which are either involved in new replication steps, translated, or encapsidated. In what subcellular compartments these different processes occur, which cellular and viral factors are involved, and how these processes are regulated during early steps of the viral infection have been the main targets of many research efforts for plant viruses during the last decade or so, particularly for potyviruses, on which we focus here.

4.1. Subcellular localization of potyvirus multiplication Our understanding of the potyviral replication process has been overwhelmingly improved during the last few years. This has been made possible notably thanks to the combining of a number of techniques: the easy and efficient expression of viral proteins fused to fluorescent tags in plants (particularly by

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agroinfiltration in N. benthamiana), their tracking in cells using confocal microscopy, and availability of infectious cDNA clones of potyviruses, often expressing viral proteins tagged with fluorescent proteins. Thus, it was shown that, as for other plant and animal RNA viruses (den Boon & Ahlquist, 2010; Grangeon, Jiang, & Laliberte´, 2012; Laliberte´ & Sanfac¸on, 2010), potyviruses replicate in vesicles produced by host endomembrane remodeling (Grangeon, Jiang, et al., 2012; Schaad, Jensen, & Carrington, 1997). For several potyviruses [Tobacco etch virus (TEV) and TuMV particularly], it was confirmed that 6K2, which contains a central hydrophobic domain (Restrepo-Hartwig & Carrington, 1994), is associated with VPg–NIaPro in ER-derived membranes (Leonard et al., 2004; Schaad, Lellis, & Carrington, 1997) forming cytoplasmic vesicles distributed throughout the cortical and perinuclear ER membrane systems (Beauchemin et al., 2007; Beauchemin & Laliberte´, 2007). Using a recombinant TuMV expressing a fluorescent protein fused to 6K2, and antibodies directed against double-stranded RNA or neosynthesized 5-bromouridine-labeled RNA, Laliberte´ and colleagues definitively showed that these vesicles are viral replication sites (Cotton et al., 2009). Studies of the mechanism by which the vesicles proliferate and develop from the ER showed that the 6K2 protein colocalizes with ER exit sites (ERES), which are the ER export domains and are also associated with Golgi bodies (Lerich, Langhans, Sturm, & Robinson, 2011; Wei & Wang, 2008). In addition, the data indicate that the accumulation of 6K2 protein at the ERES occurs in a COPI- and COPII-dependent manner, and suggested that the vesicle biogenesis depends on retrograde and anterograde transport between ER and Golgi (Wei & Wang, 2008). The cytoplasmic motility of virus-induced or 6K2-induced vesicles along actin microfilaments was highlighted by time-lapse imaging (Cotton et al., 2009; Wei & Wang, 2008). In contrast, microtubules do not appear to be involved in vesicle motility. In experiments designed to investigate the biogenesis of the vesicles, Cotton et al. (2009) coinoculated plants with infectious cDNA clones of TuMV expressing 6K2 proteins tagged with either green fluorescent protein (GFP) or mCherry. Mainly green- and red-only vesicles were observed within the same cells, which suggests that each vesicle mostly derives from a single viral genome. Further work has shown that vesicles, whether they are induced by expression of 6K2 alone or from TuMV infection, target chloroplasts, where they amalgamate and induce membrane invaginations (Wei, Huang, et al., 2010). The presence of viral RNA in these chloroplast-associated vesicles strongly

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suggests that they are also sites of virus replication. Combined with previous data showing RNA, RNA-replicative intermediates, or negative-strand RNA of different potyviruses associated with chloroplasts (Gadh & Hari, 1986; Gunasinghe & Berger, 1991; Mayhew & Ford, 1974), the results of Wei, Huang, et al. (2010) suggest that chloroplasts may be the main location for potyvirus genome replication, whereas ER is the site where potyviruses initiate genome translation and form the 6K2 vesicles. More recently, it was shown that chloroplasts associated with 6K2 vesicles were present in a large perinuclear globular structure, which also contains compacted ER, Golgi apparatus, and COPII coatamers (Grangeon, Agbeci, et al., 2012). Virus infection triggers inhibition of protein secretion at the ER–Golgi interface, but the early secretory pathway at this interface is not required for the formation of the virus-induced perinuclear structure. The 6K2-tagged vesicles, the production of which is functionally linked to the perinuclear structure, move along microfilaments, and the transvascuolar and cortical ER, the structure of which appears not to be altered by viral infection (Grangeon, Agbeci, et al., 2012). The authors suggest that replication events take place within the globular structure and that 6K2 vesicles bud at ERES and traffic toward the plasma membrane and PD for delivery of the virus into neighboring cells. This team recently showed that the 6K2 replication-competent vesicles indeed move intracellularly to reach PD, but above all intercellularly crossing PD (Grangeon et al., 2013). In parallel, Wei, Zhang, Hou, Sanfac¸on, and Wang (2013) showed that the chloroplast–6K2 complex leads to the formation of chloroplast-bound 6K2 elongated tubular structures and chloroplast aggregates which are seen in perinuclear structures described by Grangeon, Agbeci, et al. (2012). These observations support the conclusions that potyviral replication takes place in chloroplasts forming tubular structures and aggregates included in ER perinuclear structures. From the data produced by Wei, Huang, et al. (2010) and Grangeon, Agbeci, et al. (2012), we can hypothesize that viral replication takes place in chloroplasts including in the perinuclear globular structure. However, at this stage we cannot rule out the possibility that replication occurs elsewhere in this large structure or in chloroplasts outside this body.

4.2. Viral and plant factors involved in potyvirus multiplication Almost all the potyviral proteins play a role in viral multiplication (Kekarainen et al., 2002; Klein et al., 1994; Revers et al., 1999; UrcuquiInchima et al., 2001). Over the last few years, P3, CI, CI–6K2,

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6K2–NIa, NIaPro, and NIb were identified in potyviral replication vesicles (Beauchemin et al., 2007; Cotton et al., 2009; Cui et al., 2010; Dufresne, Thivierge, et al., 2008; Merits et al., 2002). The core replication complex contains the NIb as the RNA replicase (Hong & Hunt, 1996), CI as the RNA helicase (Eagles et al., 1994; Laı´n et al., 1990), VPg and NIa as linked to the 50 -end of the viral RNA and interacting with the NIb (Fellers, Wan, Hong, Collins, & Hunt, 1998; Murphy, Klein, Hunt, & Shaw, 1996), and 6K2 as a membrane anchor (Restrepo-Hartwig & Carrington, 1994; Schaad, Jensen, et al., 1997). P3 and 6K1 also appear to be essential for virus multiplication (Kekarainen et al., 2002; Klein et al., 1994; Merits et al., 2002). Though it does not bind to viral RNA (Merits et al., 1998), P3 interacts with several viral proteins involved in replication, either in yeast (Guo et al., 2001; Lin et al., 2009; Merits et al., 1999) or in planta (Zilian & Maiss, 2011), and was found associated with the ER and 6K2 vesicles (Cui et al., 2010; Eiamtanasate et al., 2007), which strongly supports a role of P3 in RNA replication. However, the precise function of P3 in this process needs to be determined. The role of HCPro in potyvirus multiplication was explained, at least in part, by its RNA silencing suppression activity, which protects the replicative intermediates and the unencapsidated genomic strands of viral RNA (Burgya´n & Havelda, 2011; Kasschau & Carrington, 2001; Rajama¨ki et al., 2004). However, the recent observation of the interaction of the HCPro protein of three potyviruses with both eIF4E and eIF(iso)4E of two plant species and CI and VPg supports a putative direct function of HCPro in potyvirus multiplication (Ala-Poikela et al., 2011). P1 is not essential for potyvirus multiplication but acts as an accessory factor for genome amplification (Verchot & Carrington, 1995a). Only CP and P3N–PIPO have not been shown to play a role in this viral process (Mahajan, Dolja, & Carrington, 1996; Wen & Hajimorad, 2010). Although CP is not essential for RNA replication in potyviruses, translation until the CP codon 138 and a cis-acting RNA element placed between codons 211 and 246 are required for this process in the case of TEV (Mahajan et al., 1996). Using different biochemical approaches, several host factors interacting with viral proteins involved in potyvirus multiplication were identified and shown to play roles in this process. The first factors identified were the eukaryotic translation initiation factor eIF4E and its isoforms, which interact with VPg (Charron et al., 2008; Leonard et al., 2000, 2004; Schaad et al., 2000; Wittmann et al., 1997). In addition, it was shown that this VPg/eIF4E

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interaction is necessary for potyvirus infection (Charron et al., 2008; Leonard et al., 2000; Yeam, Cavatorta, Ripoll, Kang, & Jahn, 2007). The interaction between 6K2–VPg–NIaPro and eIF(iso)4E was highlighted in 6K2 vesicles from bimolecular fluorescence complementation (BiFC) experiments (Beauchemin et al., 2007). Using tandem affinity purification, ELISA binding, and coimmunoprecipitation assays, it was shown that NIb directly interacts with poly(A)-binding protein 2 (PABP2, a translation factor), heat-shock protein 70 (Hsc/HSP70-3, a cellular chaperone), and eEF1A, a translation elongation factor (Beauchemin & Laliberte´, 2007; Dufresne, Ubalijoro, et al., 2008; Thivierge et al., 2008). In TuMV-infected plant, PABP2 and Hsc70-3 levels are higher than in healthy plant (Beauchemin & Laliberte´, 2007; Dufresne, Thivierge, et al., 2008), which correlates with a higher expression of their corresponding genes (Aparicio et al., 2005; Dufresne, Thivierge, et al., 2008). Hsc70-3 relocalizes to 6K2–VPg–NIaPro-induced vesicles only when associated with NIb, but in the absence of 6K2–VPg–NIaPro, Hsc70-3/ NIb interaction is not sufficient for redistribution of Hsc70-3 to membranes (Dufresne, Thivierge, et al., 2008). Another study using a second potyvirus (PVA) also identified Hsc70-3 as a component of viral replication complexes associated with NIb and VPg (Hafre´n et al., 2010). PABP2 and eEF1A are also relocalized to vesicles when coexpressed with 6K2–VPg–NIaPro (Beauchemin & Laliberte´, 2007; Thivierge et al., 2008) as both proteins also directly interact with VPg–NIaPro (Leonard et al., 2004; Thivierge et al., 2008). In addition, biochemical treatments of membrane-enriched fractions derived from TuMV-infected plants suggest that NIb, PABP2, and VPg–NIaPro are luminal but not integral membrane proteins of the 6K2–VPg–Pro-induced vesicles (Beauchemin & Laliberte´, 2007). In vitro interaction experiments showed that VPg–NIaPro and NIb also interact with PABP8, though less strongly, and VPg–NIaPro also interacts with PABP4, the two other PABP class II proteins of Arabidopsis thaliana (Dufresne, Ubalijoro, et al., 2008). Using the yeast two-hybrid system and BiFC experiments, an RNA helicase-like protein from peach and Arabidopsis (AtRH8), which is related to eIF4A, has been shown to interact with VPg of PPV in 6K2 vesicles (Huang et al., 2010). Recently, it was shown that the SNARE protein Syp71 and the SNARE-like protein Vap27-1 are recruited to the TuMV 6K2-induced elongated tubular structures (Wei et al., 2013). The 6K2 protein interacted with Vap27-1 but not with Syp71. However, since Syp71 binds to Vap27-1, Vap27-1 may function as a linker between the 6K2 vesicle and Syp71.

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Mutation analyses were performed for some of these host factors to demonstrate their involvement in the potyviral infection. The first demonstration of the involvement of such factors in potyviral multiplication was shown for eIF4E factors using knockout (KO) mutant lines challenged with several potyviruses (Duprat et al., 2002; Lellis, Kasschau, Whitham, & Carrington, 2002). In the case of PABP, single KO mutants, pab2, pab4, and pab8, were all susceptible to TuMV similar to wild-type plants, whereas the double KO mutants, including pab2 (the triple mutant pab2pab4pab8 was not viable), showed a reduced viral RNA accumulation, which positively correlated with a reduced level of PABP in the membrane (Dufresne, Ubalijoro, et al., 2008). It can be concluded that PABP are important factors for virus accumulation and TuMV seems to be able to use the different PABP paralogs during infection. Downregulation of HSP70 in HSP70-silenced and quercetin (a flavonoid which inhibits HSP70 gene expression)-treated N. benthamiana plants disturbs PVA infection through inhibition of viral RNA translation (Hafre´n et al., 2010). HSP70-15deficient A. thaliana plants are also more tolerant to TuMV infection ( Jungkunz et al., 2011). In the case of AtRH8, the corresponding KO mutant was fully resistant to PPV and TuMV, demonstrating that this RNA helicase plays a crucial role in potyvirus multiplication (Huang et al., 2010). Downregulation of the expression of Syp71, but not that of Vap27-1, inhibited TuMV infection and the formation of the elongated tubular structures, suggesting an active role for Syp71 in potyvirus replication (Wei et al., 2013). Other host factors relevant for potyvirus infection, such as DBP1, appear not to be directly involved in virus replication but in regulating the interplay between plant and virus processes. DBP1 is a DNAbinding protein phosphatase 1 that participates in transcriptional regulation of gene expression in response to virus infection (Carrasco, Ancillo, Mayda, & Vera, 2003), and directly interacts with eIF(iso)4E to stabilize this factor limiting its proteasome-mediated degradation (Castello´, Carrasco, & Vera, 2010). More recently, a DBP1 interactor, named DBP1-interacting protein 2 (DIP2), which belongs to a novel family of conserved plant small polypeptides, was identified and might be a negative regulator of DBP1 function during potyvirus infection (Castello´ et al., 2011). However, DIP2 knockdown and overexpression plants did not cause any change in eIF(iso)4E accumulation, although these DIP2 gene expression changes affected potyvirus infection. In addition, a repression of DIP2 expression was observed during the course of PPV infection (Castello´ et al., 2011). Thus, the precise roles of DIP2 and DBP1 and their relationship in the potyvirus infection remain to be determined.

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Though not yet identified in potyvirus replication-associated membrane vesicles, eIF4G, a component along with eIF4E of the eIF4F translation complex, is another important plant factor for potyvirus multiplication. Indeed, Nicaise et al. (2007) showed that coordinated recruitment of a specific isoform of eIF4G, as also of eIF4E, is necessary for viral multiplication. As eIF4G forms a trimolecular complex with VPg and eIF4E in planta (Grzela et al., 2006; Michon, Estevez, Walter, German-Retana, & Le Gall, 2006), it is most likely that this factor is also associated with RNA replication complexes in membrane vesicles. Transcriptomic analyses revealed that several ribosomal proteins are induced during potyvirus infections (Dardick, 2007; Yang et al., 2007). Silencing of the ribosomal proteins RPS2, RPS6, RPL7, RPL13, and RPL19 leads to reduction of virus accumulation in potyvirus-infected leaves (Yang, Zhang, Dittman, & Whitham, 2009), which may suggest that these proteins are specifically required for potyvirus translation. More recently, the A. thaliana acidic ribosomal protein P0 was described to be a component of a membrane-associated PVA ribonucleoprotein (RNP) complex and was found to play an extraribosomal role in viral RNA translation, along with eIF(iso)4E and VPg (Hafre´n, Eskelin, & Ma¨kinen, 2013). Moreover, P1, which was shown to traffic to the nucleolus, specifically binds to the 60S ribosomal subunit and appears to stimulate translation of viral proteins (Martı´nez & Daro`s, 2014).

4.3. Putative functions of these factors during potyvirus multiplication The presence in membrane vesicles of virus double-strand RNA and viral proteins known to be necessary for viral multiplication (VPg–NIaPro, NIb, and CI), along with plant factors known or suspected to be involved in protein synthesis (eIF4E, PABP, eEF1A, AtRH8, and P0) or with chaperone activity (HSP70/HSC70), strongly indicate that viral RNA translation and replication are coupled events (Cotton et al., 2009; Hafre´n et al., 2010; Jiang & Laliberte´, 2011). Using replication-competent and replication-incompetent PVA cDNA clones, Hafre´n et al. (2010) presented evidence for replication-associated translation. However, whether translation occurs inside the vesicles or at the cytoplasmic side of the vesicle needs to be clearly demonstrated. It is also important to remark that even if a few translation factors have been detected in vesicles, the presence of other elements belonging to the translation machinery has not been investigated yet, and that translation-independent functions in RNA replication have been assigned to some host translation factors (Blumenthal, Landers, & Weber,

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1972; Sasvari, Izotova, Kinzy, & Nagy, 2011). Thus, it is plausible that the plant translation factors included in vesicles are directly involved in viral RNA synthesis in addition to their role in viral RNA translation, as some of them directly interact with the RNA replicase NIb (Beauchemin & Laliberte´, 2007; Dufresne, Thivierge, et al., 2008; Thivierge et al., 2008). Since viral RNA translation and synthesis are probably closely associated processes, the dynamic interactions between them and their regulation, as well as the precise roles of the plant and viral factors present in the virusinduced vesicles, need to be clarified. In particular, during the early steps of viral translation and RNA synthesis, the new positive RNA strands have to be mainly targeted for new rounds of translation and synthesis rather than for encapsidation in viral particles. This means that there should be a mechanism that prevents CP interaction with positive-strand RNA. CP, which is thought not to be directly involved in RNA replication, is not detected in 6K2-induced vesicles (Cotton et al., 2009). Hafre´n et al. (2010) showed that in trans expression of PVA CP inhibits viral translation. Moreover, expression of the CP-interacting protein (CPIP), a cochaperone that was shown to interact with the potyviral CP (Hofius et al., 2007), associated with the chaperone HSC70, induces CP degradation and enhances viral translation. As CP interacts with both CPIP and HSC70, it was suggested that CPIP delivers CP to HSC70 for ubiquitination-mediated CP degradation during the viral translation process, to prevent translation inhibition by binding of CP to the viral RNA (Hafre´n et al., 2010). These data illustrate that the early viral multiplication events are closely interconnected and are highly regulated to favor viral RNA translation and synthesis instead of virion assembly. Over the last few years, several studies have focused on the dynamic interactions between VPg and several host factors and their impact in virus RNA translation. Direct evidence for the involvement of VPg in viral translation comes from in vitro translation experiments in a wheat germ extract which showed that addition of TuMV VPg enhanced the translation efficiency of an uncapped RNA but inhibited that of a capped RNA (Khan, Yumak, Gallie, & Goss, 2008). The 50 -leader of the potyviral genomic RNA is sufficient to confer cap-independent translation and function as an internal ribosome entry site (IRES; Carrington & Freed, 1990; Khan et al., 2008; Khan, Yumak, & Goss, 2009; Ray et al., 2006; Yumak, Khan, & Goss, 2010). The VPg-mediated translation enhancement is dependent on the presence of the proposed IRES and on VPg/eIF4F–eIF(iso)4F interaction, with a stronger effect of VPg on the binding to the viral RNA of eIF4F than on that of eIF(iso)4F (Khan et al., 2008). Several in vitro binding

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studies showed that the eIF(iso)4F/VPg/IRES complex is more stable than the eIF(iso)4F/m7G cap complex (Khan, 2006) and that in the presence of PABP and/or eIF4B, two factors found in 6K2-induced vesicles (see Section 4.2), the binding affinity of VPg for eIF(iso)4F is higher (Khan & Goss, 2012). It is worth noting that in these studies, VPg enhances translation without being covalently linked to the viral RNA. In vivo evidence for the involvement of VPg in viral translation comes from experiments in which PVA VPg expressed in trans enhanced viral RNA translation but repressed cellular mRNA translation in N. benthamiana (Eskelin, Hafre´n, Rantalainen, & Ma¨kinen, 2011). Mutations in a VPg domain that has been suggested to be involved in the interaction with eIF4E (Grzela et al., 2006; Leonard et al., 2000), have a negative effect on the enhancement of the viral RNA translation, which suggests that VPg/eIF4E interaction is involved in the translation stimulus (Eskelin et al., 2011). The 50 -NCR, but not the 30 -NCR, of viral RNA, and eIF4E are required to observe this VPg-mediated translation enhancement (Eskelin et al., 2011). It has been suggested that the RNA-linked VPg when covalently linked to the 50 -end of the viral RNA mimics the cap present at the 50 -end of the cellular mRNA and interacts with several cellular factors involved in RNA translation to form a translation initiation complex (Robaglia & Caranta, 2006). However, in vitro and in vivo experiments described above suggest that free VPg or NIa would be involved during viral RNA translation, which might support that VPg covalently linked to the RNA may be only necessary for the first round of translation after disassembly of virus particles. Thus, until the presence or absence of VPg linked to the 50 -end of the viral RNA translated during the infection is demonstrated, the extent to which these translation experiments faithfully reproduce the conditions of natural infections is unknown. Another hypothesis would be that the RNA-linked VPg is involved in viral RNA synthesis rather than in RNA translation. VPg is expected to play a role not only in translation but also in RNA replication. Indeed, as VPg can be uridylylated by NIb in vitro, it has been suggested that it serves as a primer for RNA synthesis (Anindya et al., 2005; Puustinen & Ma¨kinen, 2004), as has been demonstrated for the poliovirus VPg (Murray & Barton, 2003). The putative involvement of VPg in viral RNA synthesis is also supported by its interaction with the RNA replicase NIb (Fellers et al., 1998; Zilian & Maiss, 2011). Thus, available evidence suggests that interactions between VPg, translation initiation factors, and other virus and host elements are highly relevant for both virus RNA translation and replication. The unraveling of the way in

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which these interactions are integrated in congruent machinery will be an important subject of research in the next years.

5. VIRUS MOVEMENT To carry out their propagation in the whole plant following replication in membrane vesicles described previously, viruses have first to move intracellularly toward PD, the symplasmic tunnels between cells that are the gateway for this movement, cross them to enter in the neighboring cells through a cell-to-cell movement process, and then enter into sieve elements (SE) after crossing successive borders, i.e., mesophyll cell/bundle sheath (BS), BS/vascular parenchyma cell (VP), VP/companion cell (CC), and CC/SE borders. Once in the SE, the virus is transported in the phloem sap to distant locations and exits from the SE to initiate new infection sites and to disseminate efficiently throughout the whole plant.

5.1. Intracellular and cell-to-cell movements The new infectious entities that are produced in cell sites where viral protein synthesis and RNA replication take place have to reach the neighboring cells as the first step for systemic invasion of the plant. Moreover, during the movement processes, viruses have to reinitiate multiplication steps in new infected cells in order to accumulate in sufficient amount to guarantee their survival in the infected plant and to be transmitted efficiently to other plants. Whether viral replication complexes move from cell to cell and reinitiate replication or/and virions or dedicated movement RNP complexes produced in initial infected cells move from cell to cell and disassemble to release viral RNA is not well understood yet. In the case of potyviruses, as shown above, replication complexes are mobile in cells, trafficking along actin microfilaments with involvement of myosin XI-K (Cotton et al., 2009; Cui et al., 2010; Grangeon, Agbeci, et al., 2012; Wei & Wang, 2008) and then move intercellularly across PD for delivery of the replication complexes into neighboring cells (Grangeon, Agbeci, et al., 2012; Grangeon, Jiang, & Laliberte´, 2012; Grangeon et al., 2013), as was suggested for other viruses (Harries, Schoelz, & Nelson, 2010; Schoelz, Harries, & Nelson, 2011). It was recently estimated for TuMV infection in N. benthamiana that the rate of cell-to-cell movement was one new infected cell every 3 h (Agbeci, Grangeon, Nelson, Zheng, & Laliberte´, 2013).

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During the 1990s, it was shown that the central part (core domain) of the CP and the N-terminal region of the CI are required for cell-to-cell movement (Carrington et al., 1998; Dolja, Haldeman-Cahill, Montgomery, Vandenbosch, & Carrington, 1995; Dolja et al., 1994). The involvement in cell-to-cell movement of the N-terminal region of the CI, a region necessary, and sufficient for self-interaction (Lo´pez, Urzainqui, Domı´nguez, & Garcı´a, 2001), was later confirmed for PPV (Go´mez de Cedro´n et al., 2006). Recently, it was also suggested that the surface-exposed CP C-terminal end of SMV has a role in the cell-to-cell movement process, probably acting in the CP intersubunit interaction and virion assembly (Seo et al., 2013). Previous immunoelectron microscopy studies showed that the presence of conical structures, corresponding to immature cylindrical inclusions, perpendicularly to the cell wall and often traversing it, over the apertures of PD connecting adjacent cells (Langenberg, 1986, 1993; Roberts et al., 1998; Rodrı´guez-Cerezo et al., 1997). CP-specific labels were clearly observed nearby or inside the cones and also in the PD cavities (Roberts et al., 1998; Rodrı´guez-Cerezo et al., 1997). P3 and viral RNA were also observed associated with the cell wall-bound conical structures (Rodrı´guez-Cerezo et al., 1997). Altogether, these data support a direct role of CI and CP for the transfer of the viral RNA genome from cell to cell through PD, in which CI could function to position virions or a CP-containing RNP for translocation across the cell wall. Microinjection experiments in N. benthamiana and lettuce of Escherichia coli-expressed proteins showed that CP and HCPro, but not CI, have properties of MP, as both are able to increase the PD size-exclusion limit (SEL), facilitate viral RNA cell-to-cell movement, bind to RNA, and move from cell to cell (Rojas, Zerbini, Allison, Gilbertson, & Lucas, 1997). It was not until 2008 that a crucial finding, i.e., the discovery of P3N– PIPO (Chung et al., 2008), was made to boost new discoveries to better understand how potyviruses move from cell to cell. Indeed, introduction of stop codon mutations in PIPO of the potyviruses SMV and TuMV and of the tritimovirus WSMV, which do not affect P3 amino acids, inhibited systemic infection, restricting viruses to small clusters of cells within the inoculated leaves (Choi et al., 2005; Chung et al., 2008; Wen & Hajimorad, 2010). In addition, it was shown that P3N–PIPO is able to move from cell to cell when expressed alone under the control of the 35S promoter in N. benthamiana leaves (Vijayapalani et al., 2012). All these data argued for a role of P3N–PIPO in potyvirus cell-to-cell movement, in addition to CI and CP.

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Agrobacterium-mediated transient expression coupled to confocal microscopy observations allowed a better understanding of the role of these potyviral MPs in the cell-to-cell movement process. Coexpression of CI and P3–PIPO resulted in cell wall-associated punctuate bodies of CI, whereas expression of CI alone or with another potyvirus protein leads to the formation of CI cytoplasmic aggregates (Wei, Zhang, et al., 2010). Transient expression of GFP–P3N–PIPO revealed that this protein localized at the cell wall with both PDLP1, a type I membrane PD protein (Thomas, Bayer, Ritzenthaler, Fernandez-Calvino, & Maule, 2008), and CI (Wei, Zhang, et al., 2010). In addition, BiFC experiments highlighted CI/P3N–PIPO interaction. CP was also observed as fibrillar structures (probably virions or RNP complexes) either in the cytoplasm, often associated with chloroplasts, or along the cell wall associated with CI (Wei, Zhang, et al., 2010). Using chemical and protein inhibitors, it was shown that the CI/P3N–PIPO delivery in PD is dependent on the secretory pathway but is independent of the actomyosin motility system (Wei, Zhang, et al., 2010). The same experiments performed with CI mutants defective in cell-to-cell movement activity (Carrington et al., 1998; Go´mez de Cedro´n et al., 2006; see above) revealed that mutated CI was observed in the nucleus and cell periphery but not associated with PD, although the CI self-interaction and interaction with P3N–PIPO were not abolished in spite of a significant reduction in the binding strength observed in yeast (Go´mez de Cedro´n et al., 2006; Wei, Zhang, et al., 2010). Thus, from these data, it is suggested that P3N–PIPO interacts directly with CI and directs CI to PD using the secretory pathway, and that proper self-interaction of CI is crucial in this process. However, as both CI and P3N–PIPO lack a typical transmembrane domain [the hydrophobic region identified in P3 responsible for the ER targeting is located at the C-terminal end of P3 and is therefore not included in P3N–PIPO (Cui et al., 2010; Eiamtanasate et al., 2007)], another factor seems necessary to anchor the CI/P3N–PIPO complex in PD. Such a factor has been recently identified by a yeast two-hybrid screening of an A. thaliana cDNA library using TuMV P3N–PIPO as bait (Vijayapalani et al., 2012). This factor, PCaP1 (AT4G20260), is a hydrophilic cation-binding protein anchored to the plasma membrane via myristoylation of a glycine residue [there is no transmembrane domain in PCaP1 (Nagasaki, Tomioka, & Maeshima, 2008)]. Direct P3N–PIPO/PCaP1 interaction in planta, via the PIPO domain, was confirmed by coimmunoprecipitation and BiFC (Vijayapalani et al., 2012). Colocalization studies revealed the presence of the P3N–PIPO/PCaP1 complex in PD but also in other locations. The

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functional relevance of PCaP1 was corroborated by showing that TuMV RNA accumulation was not affected in PCaP1 KO plants, but cell-to-cell movement was strongly reduced and viral infection of whole plants was strongly attenuated without being completely inhibited (Vijayapalani et al., 2012). More recently, it was also shown that the cell-to-cell movement process depends on early and late secretory pathways, as well as myosin XI motors, but not on the endocytic pathway (Agbeci et al., 2013; Grangeon, Agbeci, et al., 2012). From all these data, a model for the cell-to-cell movement of potyviruses can be suggested. The CI/P3N–PIPO complex, formed in ER membranes of the virus-induced large perinuclear structure (see above), is delivered to PD via the secretory system and anchored to PD thanks to the interaction of the PIPO domain with PCaP1. Then, more CI molecules bind to the CI/ P3N–PIPO/PCaP1 complex via CI self-interaction to form conical structures. CP and newly synthesized viral RNA molecules released from the replication vesicles moving along microfilaments toward the PD can form virions or RNP complexes, which bind to CI conical structures and move through PD to the neighboring cell. How these virus transport forms bind to CI remains to be determined. Using both atomic force microscopy and immunoelectron microscopy, it was shown in the case of PVA and Potato virus Y (PVY) that a small proportion of purified virus particles contain a protruding tip 40 nm long and 2.5 nm in diameter at one end in which HCPro and CI were detected (Gabrenaite-Verkhovskaya et al., 2008; Manoussopoulos et al., 2000; Torrance et al., 2006). This tip seems to be associated with the 50 -end of viral RNA as VPg was also detected. Indeed, VPg linked to the 50 -end of viral RNA was shown to be available for protein–protein interaction at the surface of virus particles and could be targeted with antibodies (Puustinen et al., 2002). Since PVA HCPro is able to interact with CI and VPg (Guo et al., 2001), CI might be associated with the tip indirectly by its binding to the HCPro/VPg complex, and then target the CI-associated virion/RNPs to the PD-associated CI conical structures via CI self-interaction. Alternatively, CI can also interact with virion/RNPs independent of HCPro as for some other potyviruses, since CI/VPg or CI/CP interactions have also been shown in vitro (Tavert-Roudet et al., 2012), in yeast (Lin et al., 2009), and in planta (Zilian & Maiss, 2011). Another question to be solved is which viral protein induces the modification of the PD that allows CI conical structures to traverse the cell wall and facilitate the virion/RNP complex to move to the neighboring cell?

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P3N–PIPO is the more likely candidate since it is directly associated with PD via its interaction with PCaP1 and is able to induce the increase of PD SEL when expressed alone (Vijayapalani et al., 2012). However, we cannot exclude an active role of CP and HCPro, which have also been shown to cause an increase of the PD SEL, traffic from cell to cell (Rojas et al., 1997), and are associated with possible viral transport forms (GabrenaiteVerkhovskaya et al., 2008; Torrance et al., 2006). In addition to PCaP1, two other host factors that might have a role in cellto-cell movement have been identified, both of them interacting with VPg. These are PVIP (for potyvirus VPg-interacting protein) and the aforementioned eIF4E and its isoform eIF(iso)4E. PVIP from pea, N. benthamiana and A. thaliana, exhibits the same ability to bind VPg proteins from several potyviruses (Dunoyer, Thomas, Harrison, Revers, & Maule, 2004). The PVIP/VPg interaction was shown to be important for virus movement, as mutations in the VPg sequence preventing its interaction with PVIPs or silencing of both Arabidopsis PVIP1 and PVIP2 strongly reduced TuMV local and systemic movement without affecting virus replication (Dunoyer, Thomas, et al., 2004). In A. thaliana, PVIP is part of a small gene family of proteins containing a plant homeodomain with the capacity to regulate gene expression through histone modifications (reviewed in Cosgrove, 2006). The Arabidopsis PVIP2 and PVIP1 genes correspond to OBERON1 and OBERON2 genes, respectively, which were described as having redundant functions in the establishment and/or maintenance of the shoot and root apical meristems (Saiga et al., 2008; Thomas, Schmidt, Bayer, Dreos, & Maule, 2009). They also act as central regulators in auxin-mediated control of development (Thomas et al., 2009). However, the nuclear localization of PVIP factors (Saiga et al., 2008) raises the possibility that PVIP/VPg interaction may modulate expression of host genes involved in virus movement, rather than having a direct role in the viral movement through PD. eIF4E isoforms have been shown to play a role in potyvirus multiplication (see the previous section). However, at least two studies have suggested a putative role for eIF4E in movement in the case of Pea seed-borne mosaic virus (PSbMV; Gao et al., 2004) and TEV (ContrerasParedes, Silva-Rosales, Daros, Alejandri-Ramirez, & Dinkova, 2013), but a clear demonstration of such a role is missing.

5.2. Long-distance movement Viral long-distance movement involves several steps starting from the virus entry into the phloem cells (BS, VP, or CC), delivery to the SE, transport

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along SE, and exit from the SE, following the source-to-sink transportation of carbohydrates. This process requires the crossing of successive borders, i.e., mesophyll cell/BS, BS/VP, VP/CC, and CC/SE borders, which requires the setting-up of specific interactions between virus and host factors (reviewed in Hipper, Brault, Ziegler-Graff, & Revers, 2013). However, spatial and kinetic analyses of long-distance movement of some viruses revealed that the direction and speed of movement may be different than those of photoassimilates. The slower rate of virus progression observed in some experimental cases, compared to the speed of photoassimilates, could be explained by additional virus unloading and amplification steps in CC before being reloaded into the SE. Indeed, Germundsson and Valkonen (2006) showed that N. benthamiana wild-type scions grafted above fully recovered leaves from VPg- or P1-transgenic rootstocks previously inoculated with PVA remained uninfected. As these recovered sections express RNA silencing-based resistance, the authors suggested that viral RNA has to be targeted in phloem cells. As during phloem transport, viral RNA is protected in the SE by viral proteins in the form of virions or RNP, they hypothesized that phloem movement of PVA proceeds in repeated movement cycles, each including virus loading to the SE for transport over a short distance and unloading to CCs and other phloem cells for replication, which would expose the virus RNA to RNA silencing. This hypothesis is in agreement with the fact that during infection with GFP-tagged virus, GFP is present in all the vasculature highlighting unloading and replication/ translation steps along the long-distance movement route (Germundsson & Valkonen, 2006). The different steps of the long-distance movement represent potential barriers for virus trafficking and examples of viruses blocked at one stage or another were described (Hipper et al., 2013). Thus, each of these steps may induce bottlenecks in a virus population. From a few studies, it has been suggested that the size of the virus population invading the sink organs from the vasculature depends on the concentration of virus in the sap and/or on barriers imposed by the host (Gutierrez et al., 2012). Regarding potyviruses, one study in the TEV/tobacco-pepper pathosystems indicates that a bottleneck is driven by the systemic transport of the virus (Zwart, Daro`s, & Elena, 2011). From a mechanistic point of view, potyviral transport in the vascular system is poorly understood. First, we do not know if potyviruses are transported over long distance as virions or in another RNP form. Second, no host susceptibility factor specifically involved in the long-distance

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movement process has been identified. Nevertheless, potyviral determinants promoting the systemic spread and some host factors restricting this process were characterized. 5.2.1 Viral determinants involved in potyviral long-distance movement During the 1990s, it was shown that the N- (CP-Nter) and C- (CP-Cter) terminal CP domains are dispensable for virus genome encapsidation, but essential for virus long-distance movement (Andersen & Johansen, 1998; Dolja et al., 1995, 1994; Lo´pez-Moya & Pirone, 1998). More recently, both CP-Nter and CP-Cter were shown to be host- and strain-specific longdistance movement determinants for Potyviridae family members (Carbonell et al., 2013; Decroocq et al., 2009; Desbiez, Chandeysson, & Lecoq, 2014; Salvador, Delgadillo, Sa´enz, Garcı´a, & Simo´n-Mateo, 2008; Tatineni & French, 2014; Tatineni, Van Winkle, & French, 2011). The CP-Nter appears to condition long-distance spread, not only playing a direct role in virus movement but also eliciting host-specific resistance mechanisms confining the virus in the inoculated leaves (Carbonell et al., 2013; Decroocq et al., 2009). As the CP-Nter sequence is highly variable among potyviruses (Shukla & Ward, 1989), the primary sequence seems not to be associated with the movement function. Several studies highlight the putative role of the global net charge (Arazi et al., 2001; Kimalov, Gal-On, Stav, Belausov, & Arazi, 2004) or posttranslational modifications such as O-glycosylation (Chen et al., 2005; Ferna´ndez-Ferna´ndez, Camafeita, et al., 2002; Pe´rez et al., 2013) and phosphorylation (Ivanov et al., 2003). However, none of these modifications have been shown to play a role in the long-distance movement function of CP-Nter. Recently, CP-Nter has been predicted to be a disordered domain (Chroboczek, He´brard, Ma¨kinen, Michon, & Rantalainen, 2012; Ksenofontov et al., 2013), raising the possibility that this domain may interact with multiple partners. Besides its role in virus replication (see the previous section; Jiang & Laliberte´, 2011), VPg of potyviruses is also involved in virus movement. Several studies showed that mutations in VPg overcome the resistance based on virus long-distance movement restriction. This was demonstrated for TEV in tobacco (Schaad, Lellis, et al., 1997) and for PVA in different plant species, such as Nicandra physaloides (Rajama¨ki & Valkonen, 1999), a diploid potato hybrid (Hamalainen, Kekarainen, Gebhardt, Watanabe, & Valkonen, 2000), and Solanum commersonii (Rajama¨ki & Valkonen, 2002). For PVA, one amino acid change in the central domain of the VPg is sufficient to restore viral long-distance movement, although this resistance bypass is host

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specific (Rajama¨ki & Valkonen, 1999, 2002). Using grafting experiments, it was also shown that the PVA long-distance movement restriction was likely due to the absence of virus loading into the SE (Hamalainen et al., 2000; Rajama¨ki & Valkonen, 2002). Rajama¨ki and Valkonen (2003) showed that VPg accumulated specifically in CC in veins of the sink leaves in S. commersonii, only at an early phase of PVA systemic infection, whereas neither other viral proteins nor viral RNA was detected. These findings have raised the hypothesis of a specific role of VPg as the vascular movement protein of potyviruses. Although the inability to detect viral RNA in distant CCs showing VPg accumulation suggests that VPg can act in long-distance movement as a free protein, a role of VPg linked to the viral RNA cannot be discarded, since it is exposed at one extremity of the virion and is accessible for interaction with other proteins (Puustinen et al., 2002), such as phloem host factors involved in virus movement. Regardless as to whether VPg is acting in a free or a RNA-linked form, incompatibility between VPg and specific host factors would abolish virus long-distance movement, thereby conferring resistance to the host, and mutations restoring productive interaction would confer resistance breaking. On the other hand, since VPg has been shown to have RNA silencing suppression activity, it has been suggested that the function of VPg in distant CC at early times of infection might be to suppress resistance barriers ahead of virus infection (Rajama¨ki & Valkonen, 2003, 2009). Rajama¨ki and Valkonen (1999) showed that, in addition to VPg, PVA 6K2 is a virulence determinant in N. physaloides enabling the virus to overcome the resistance that restricts PVA long-distance movement in this host. One amino acid change in the N-terminal sequence of 6K2 (6K2-N) was indeed sufficient to facilitate virus systemic spread. The involvement of PVA 6K2 in long-distance movement was also described in N. benthamiana and Nicotiana tabacum (Spetz & Valkonen, 2004). As detailed in Section 4, 6K2 is an integral membrane protein associated with VPg in ER-derived membranes (Leonard et al., 2004; Schaad, Jensen, et al., 1997) forming cytoplasmic vesicles that can be viral replication sites or vehicles for intracellular transport of viral replication products (Cotton et al., 2009; Grangeon, Agbeci, et al., 2012; Wei, Huang, et al., 2010). The 6K2-N is located on the cytoplasmic side of the membrane (Schaad, Jensen, et al., 1997) and then can potentially interact with viral or host factors implicated in potyvirus long-distance movement. In particular, a coordinated role for the VPg and 6K2 proteins in PVA vascular transport can be envisaged either as a 6K2–VPg unprocessed product or as a 6K2/VPg

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complex, since interaction between these two proteins has been shown for several potyviruses (Lin et al., 2009; Zilian & Maiss, 2011). However, whether the 6K2 protein from other potyviruses participates in virus long-distance movement needs to be addressed. 5.2.2 Host factors involved in the restriction of long-distance movement If no host factors assisting long-distance movement of potyviruses have been identified yet, several factors restricting this movement process were genetically identified. However, only the “Restriction to TEV Movement” (RTM) genes were cloned and characterized. These genes were identified at the end of the 1990s in several Arabidopsis accessions, particularly in Col-0 (Mahajan, Chisholm, Whitham, & Carrington, 1998). In the resistance controlled by these genes, viral replication, and cell-to-cell movement in the inoculated leaves are not affected, a hypersensitive response (HR), and systemic acquired resistance (SAR) are not triggered and salicylic acid (SA) is not involved (Mahajan et al., 1998). This dominant resistance is effective against at least three potyviruses, TEV, Lettuce mosaic virus (LMV), and PPV (Decroocq et al., 2006; Mahajan et al., 1998; Revers et al., 2003). Genetic characterization of natural A. thaliana accessions and A. thaliana mutants showed that at least five dominant genes, named RTM1, RTM2, RTM3, RTM4, and RTM5 are involved in this resistance (Cosson et al., 2012; Mahajan et al., 1998; Whitham, Yamamoto, & Carrington, 1999). A single mutation in one of the RTM genes is sufficient to abolish the resistance phenotype (Whitham et al., 1999). RTM1 encodes a protein belonging to the jacalin family (Chisholm, Mahajan, Whitham, Yamamoto, & Carrington, 2000). RTM2 expresses a protein with similarities to small heat-shock proteins and contains a transmembrane domain (Whitham, Anderberg, Chisholm, & Carrington, 2000). RTM3 belongs to a meprin and TRAF homology (MATH) domain protein family and possesses a coiled-coil domain at its C-terminus (Cosson et al., 2010). RTM4 and RTM5 have only been genetically characterized (Cosson et al., 2012). RTM1 and RTM2 are specifically expressed in phloem-associated tissues and the corresponding proteins localize to SE (Chisholm, Parra, Anderberg, & Carrington, 2001). Despite the fact that mutations in the CP of potyviruses overcome the RTM resistance (Decroocq et al., 2009), none of the RTM proteins have been found to physically interact with CP in yeast two-hybrid assays (Cosson et al., 2010). However, interaction of CP, free or in whole virions, with RTM proteins mediated by additional cellular or viral proteins is still conceivable. Indeed, self- and

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cross-interactions of RTM1 and RTM3 were observed, which suggest that these proteins may be part of a larger protein complex (Cosson et al., 2010). Several hypothesis can be proposed about the RTM resistance mechanism: (i) virus particles or RNPs, in the process of being loaded into SE, could be sequestered by the RTM complex; (ii) the RTM complex could reduce virus accessibility of cellular factor(s) or structure(s) required for potyvirus long-distance movement; and (iii) RTM complex could activate a movement-restricting response of the plant following virus infection. Interference with potyvirus long-distance movement has also been studied in Prunus (Ion-Nagy et al., 2006). This was genetically characterized and one major PPV resistance locus has been mapped in the upper part of apricot linkage group 1 and narrowed down to 196 kb according to the peach genome syntenic region (Soriano et al., 2012; Zuriaga et al., 2013). In this PPV resistance locus, Zuriaga et al. (2013) identified a cluster of genes coding for MATH domain-containing proteins, which appear to be appealing candidates, as Arabidopsis RTM3 codes for a MATH domain-containing protein included in a cluster similar to that of the Prunus PPV resistance locus (Cosson et al., 2010). However, whether the RTM resistance is really effective in Prunus against PPV remains to be determined. Recently, another PPV resistance locus, named SHA3, involved in the restriction of the long-distance movement in A. thaliana has been mapped in the RTM3 cluster (Pagny et al., 2012). However, unlike the RTM resistance, which is dominant, this resistance is recessive and SHA3 and RTM3 were shown to be distinct genes. The cloning of SHA3 will be an important breakthrough, as it could potentially represent the first identified susceptibility factor directly involved in potyvirus systemic movement. eIF(iso)4E has also been suggested to contribute to systemic spread of TEV (Contreras-Paredes et al., 2013); however, the data do not rule out the possibility that the disturbance of systemic spread observed in the absence of eIF(iso)4E could be a secondary effect of a defect in viral replication or cell-to-cell spread. Another recessive resistance gene named ra, blocking vascular transport of PVA, was genetically characterized in potato but has not been cloned yet, although it seems to be linked to a gene cluster including dominant resistance genes such as Ry and Na (Hamalainen et al., 2000). Other dominant genes involved in the HR to potyviruses were shown to block viral long-distance movement. In particular, this was observed in Na-, and Ny- and Nc-containing potato plants blocking long-distance movement of PVA and PVY, respectively (Hamalainen et al., 2000; Moury et al., 2011;

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Tian & Valkonen, 2013). The fact that PVY with a chimeric HCPro infects systemically Ncspl potato plants and induces necrotic lesions in noninoculated apical leaves (Moury et al., 2011) suggests that the necrotic reaction and the blocking of long-distance movement are controlled independently, as could be generalized for virus resistance associated with HR (Pallas & Garcı´a, 2011). The mechanism involved in movement restriction is still unknown.

6. VIRUS TRANSMISSION Potyviruses are naturally propagated mainly by aphids, but some of them are also transmitted by seeds. In spite of the high relevance of the transmission process, quite little information has been published on potyvirus transmission during the last decade.

6.1. Transmission by aphids Since the beginning of the studies of potyvirus transmission by aphids more than 40 years ago, important progress has been made for a better understanding of the molecular mechanism governing this mode of transmission, particularly during the 1980s and 1990s. Several excellent reviews on virus transmission by aphids were published since that time (Blanc, Uzest, & Drucker, 2011; Brault, Uzest, Monsion, Jacquot, & Blanc, 2010; Lo´pezMoya, 2002; Ng & Falk, 2006; Pirone & Blanc, 1996; Stafford, Walker, & Ullman, 2012). Aphids transmit potyviruses in a nonpersistent manner. In contrast with the capsid-only strategy, in which only the viral CP interacts with the vector, the helper strategy adopted by potyviruses to interact with aphids involves an additional nonstructural protein that collaborates with CP in the virus–vector interaction (Brault et al., 2010). The concept of the helper strategy for the aphid transmission of potyviruses was developed from the works of Govier and Kassanis in the 1960s and early the 1970s. These authors showed that the transmission of nontransmissible potyvirus isolates could be complemented either by mixed infection with a transmissible potyvirus or by previously feeding of aphids on plants infected with a transmissible potyvirus (Kassanis, 1961; Kassanis & Govier, 1971a, 1971b). Complementation experiments using feeding through membrane with sap extracted from infected plants highlighted the existence of an HC which was shown to be a protein of viral origin (Govier & Kassanis, 1974a, 1974b; Govier et al., 1977; Thornbury & Pirone, 1983). The bridge hypothesis was formulated to explain the role of HC in aphid transmission: two independent domains of this protein would attach to the virion and to

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the aphid stylet, providing a bridge to link the virus particles to the aphid (Pirone & Blanc, 1996). Since a proteinase activity was identified in the C-terminal domain of the HC of potyviruses, this protein was then referred to as HCPro (Carrington, Cary, Parks, & Dougherty, 1989; Carrington, Freed, et al., 1989). Different sequence comparisons and experimental analyses revealed the relevance of a highly conserved KITC motif in the N-terminal domain of HCPro, which was necessary to retain virions in the food canal and foregut of aphids and facilitate virus transmission (Atreya, Atreya, Thornbury, & Pirone, 1992; Blanc et al., 1998; Huet, Gal-On, Meir, Lecoq, & Raccah, 1994; Thornbury, Patterson, Dessens, & Pirone, 1990). A second highly conserved motif located in the HCPro C-terminal domain, the PTK motif, was shown to be also necessary for the HC activity and was required for efficient interaction with virion (Huet et al., 1994; Peng et al., 1998). Comparison of conserved sequences at the CP N-ter of several aphidtransmissible and nontransmissible potyviruses associated with mutagenesis analyses showed that an N-terminal highly conserved motif, mainly composed of Asp-Ala-Gly and thus referred as the DAG motif, is necessary for potyvirus transmission by aphids (Atreya, Atreya, & Pirone, 1991; Atreya, Lopez-Moya, Chu, Atreya, & Pirone, 1995; Atreya, Raccah, & Pirone, 1990; Gal-On, Antignus, Rosner, & Raccah, 1992; Harrison & Robinson, 1988), although the amino acid context in which this DAG motif is placed is also important for the transmission process (Lo´pez-Moya, Wang, & Pirone, 1999). Direct interaction of HCPro and CP involving the DAG motif was shown in an in vitro protein blotting-overlay assay (Blanc et al., 1997). All these data support a bridge model in which PTK and DAG motifs determine the HCPro–CP interaction, whereas the KITC motif of HCPro mediates interaction with the aphid stylet. Several structural analyses showed that active HCPro is in different oligomeric form (Plisson et al., 2003; RuizFerrer et al., 2005; Thornbury et al., 1985) where the KITC and PTK motifs might be sufficiently separated to fulfill their proposed role in the bridge mechanism (Plisson et al., 2003; Ruiz-Ferrer et al., 2005). However, even though these conserved motifs are present, aphid transmission demands species-specific interactions, as not all aphid species can transmit all aphid-transmissible potyviruses and some HCPro proteins cannot act as helper factors for some heterologous potyvirus particles. That suggests that the amino acid contexts around these motifs in HCPro and CP, and perhaps other protein domains, play important roles for efficient

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interactions (Andrejeva et al., 1999; Dombrovsky, Huet, Chejanovsky, & Raccah, 2005; Flasinski & Cassidy, 1998; Pirone & Blanc, 1996). Recent studies identified other CP and HCPro amino acids involved in aphid transmission. Thus, a highly conserved histidine in the SMV CP-C-ter and an arginine located near the cleavage site at the C-terminal end of HCPro were shown to play a role in CP/HCPro interaction and efficiency of aphid transmission (Seo, Kang, Seo, Jung, & Kim, 2010). On the other hand, a glutamic to lysine change at position 68 in PVY CP induces a twofold increase of aphid transmission (Moury & Simon, 2011). However, further work is needed to clarify the involvement of these amino acids in the aphid transmission of SMV, PVY, and other potyviruses. The receptor of HCPro in the aphid stylet is another important element for the specificity of the transmission. However, this receptor has not yet been identified. Nevertheless, in recent years, studies using in vitro binding assays between aphid proteins and HCPro have revealed several candidates. Dombrovsky, Gollop, Chen, Chejanovsky, and Raccah (2007) extracted proteins from whole aphids and tested interaction with Zucchini yellow mosaic virus (ZYMV) HCPro using an overlay assay after 1D or 2D separations. Several proteins, including cuticle proteins, were identified to interact with an HCPro form active for transmission but not with a defective HCPro. In addition, interaction of the aphid proteins with ZYMV CP was revealed only in the presence of HCPro, which further supports the bridge hypothesis. Using proteins extracted from aphid heads and a far-Western blotting strategy, Ferna´ndez-Calvino et al. (2010) identified at least nine proteins, different from those found by Dombrovsky et al. (2007), which bind to TEV HCPro. Specific interaction was confirmed with purified RPS2, a ribosomal protein homologous to the laminin receptor precursor, known to act as the receptor of several viruses. Whether this protein or the other proteins identified in either study are located in aphid stylet remains to be determined. An intriguing point highlighted by these studies is that many aphid proteins interacted with HCPro, which suggests that more than one aphid protein could be involved in the transmission process. Studies on Cauliflower mosaic virus (CaMV, Caulimovirus genus) showed that aphid transmission complexes might be assembled inside the infected plant cell forming specific structures denoted transmission bodies, which, in contact with aphids, are disintegrated releasing transmissible virions (Blanc et al., 2011; and references therein). In the case of potyviruses, HCPro aggregates in various inclusions (Riedel, Lesemann, & Maiss, 1998). However, these inclusions do not contain virions or CP and therefore

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likely do not correspond to the CaMV transmission bodies. Whether potyviruses use an alternative strategy to prepare for aphid transmission is still an open question. Some pieces of evidence support the view that potyviruses are acquired by aphids via sap ingestion and that salivary secretions rather than sap egestion may be involved in inoculation of new plants (Fereres, 2007; Martin, Collar, Tjallingii, & Fereres, 1997; Pirone & Perry, 2002). Acquisition is associated with the third subphase (II-3) of the intracellular stylet puncture, whereas inoculation occurs during the first phase (II-1; Martin et al., 1997; Moreno, Tjallingii, Fernandez-Mata, & Fereres, 2012). Moreno, Fereres, and Cambra (2009) estimated the viral charge acquired and inoculated/transmitted by single aphids using an electrical penetration graph and quantitative real-time RT-PCR. A number ranging from 305 to 216,589 (average of 26,750) PPV RNA targets are inoculated in a single probe, which was shown to correspond to about half of the number of the acquired targets. Several stylet punctures did not affect the efficiency of virus acquisition but increased the infection rate. These numbers of viral RNA targets inoculated by an aphid are higher than those estimated by Pirone and Thornbury (1988) for Tobacco vein mottling virus (TVMV; between 15 and 20,760) and much higher than those estimated by Moury, Fabre, and Senoussi (2007) for PVY (between 0.5 and 3.2); both studies used different experimental approaches for transmission, based on artificial media containing purified virus particles. Efficient potyvirus transmission by aphids is also related to aphid colonizing and probing behavior (Fereres & Moreno, 2009; Powell, Tosh, & Hardie, 2006; Stafford et al., 2012). Noncolonizer aphid species contribute more to potyvirus spread than colonizer species (Raccah, Gal-On, & Eastop, 1985; Yuan & Ullman, 1996). Other studies have shown that aphids making the first probes, coupled to frequent intracellular punctures made during the first minutes after landing, transmit potyviruses with high efficiency (Ferna´ndez-Calvino, Lo´pez-Abella, Lo´pez-Moya, & Fereres, 2006; Kalleshwaraswamy & Kumar, 2008; Yuan & Ullman, 1996). Virus infections also influence vector behavior and performance. Fereres, Kampmeier, and Irwin (1999) showed that the aphid Rhopalosiphum maidis remains on healthy soybean plants for a longer time period than on SMVinfected plants, thus increasing the chance to inoculate virus to healthy plants. More recently, two independent studies showed that PVY infection on potato plants influenced differentially the aphid feeding behavior of Myzus persicae and Macrosiphum euphorbiae (Boquel, Giordanengo, & Ameline, 2010) and, in mixed infection with a polerovirus, increased the

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fecundity and settling of both aphid species (Srinivasan & Alvarez, 2007). Salvaudon, De Moraes, and Mescher (2013) showed that squash plants infected with ZYMV emitted more organic volatile compounds, exhibited significant changes in leaf coloration, and were more attractive for aphids compared to healthy plants. An increase of M. persicae fecundity and settling was also described in N. benthamiana plants infected with TuMV, which seems to be related to a higher free amino acid content, reduced aphid specific plant defense gene expression, and reduced aphid-induced callose deposition in plant leaves (Casteel et al., 2014). In this last study, it was also shown that the expression of NIaPro alone in N. benthamiana is sufficient to increase M. persicae fecundity and settling (Casteel et al., 2014). Although the aphid transmission of potyviruses has been studied for more than 40 years, the precise mechanism of this process is only poorly understood and more studies will be necessary to decipher it. Some important remaining questions concern the identification of the aphid receptor(s) interacting with HCPro, the precise interaction between HCPro and virus particles in the stylet, the existence of dedicated transmission bodies in infected cells to prepare for aphid transmission, and the precise role of NIaPro in aphid transmission.

6.2. Seed transmission Seed transmission of plant viruses plays an important role in virus disease epidemiology as it provides a means for virus spread over time and distance and it allows the settlement of new foci of vector dispersal. Thus, a low rate of seed transmission is sufficient to enhance significantly the spread of virus disease. From just over 100 plant viruses which are known to be seed transmitted, about 20 viruses belong to the genus Potyvirus, including SMV, LMV, ZYMV, PVY, PSbMV, and Watermelon mosaic virus (Mink, 1993). Many descriptive works were published from the 1950s to the 1980s that resulted in the biological characterization of seed transmission of plant viruses (reviewed in Johansen, Edwards, & Hampton, 1994; Maule & Wang, 1996; Mink, 1993). However, despite its biological importance, very few works have been published on this topic since 1996. Seed transmission appears to be a complex mechanism for which most of the knowledge regarding potyviruses comes from studies on two viruses, PSbMV and SMV. Seed transmission of PSbMV in pea results in direct invasion of the immature embryo from the maternal tissues (Wang & Maule, 1992). Carrying out immunocytochemical and in situ hybridization studies, Wang

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and Maule (1994) detected PSbMV in and around the vascular strand located in the testa of immature seeds of pea cultivars able (Vedette) and unable (Progreta) to transmit PSbMV. However, virus invaded the neighboring tissues and the micropylar region only in Vedette to finally infect the suspensor, a transient structure which is composed of elongated and globular cells associated with the developing embryo. From ultrastructural analysis of the micropylar region combined with immunogold labeling of virus proteins, Roberts, Wang, Thomas, and Maule (2003) suggested the presence in this micropylar region of PD connecting testa and endosperm, which are used by the seed-transmitted virus to enter in the endosperm and invade the suspensor. Thus, the major point which governs the efficiency of seed transmission for a given virus isolate is its capacity to reach and to cross the testa/ endosperm boundary and to invade the suspensor before its degeneration. And it was shown that the genetic composition of both the host and the virus, environmental factors, and the age of the parent plant at the time of infection strongly influence this process ( Johansen et al., 1994; Maule & Wang, 1996). Analyzing seed transmission in reciprocal cross-pollination experiments between cultivars with opposite phenotype behavior showed that seed transmission is incompletely dominant and controlled by a few genes of maternal origin (Wang & Maule, 1994). However, none of these genes have been cloned. On the virus side, it was shown for PSbMV that several regions of the genome are involved in seed transmission, particularly, the 50 -NCR, HCPro, and a 30 -region encompassing CP ( Johansen, Dougherty, Keller, Wang, & Hampton, 1996). Altogether, these data show that seed transmission of PSbMV involves complex host/virus interactions. As for transmission of PSbMV, SMV transmission by seeds is also dependent on the virus and the soybean genotype (Bowers & Goodman, 1991; Domier et al., 2007). However, the mechanism that prevents transmission of defective isolates appears to be different for SMV, as both seedtransmitted and nonseed-transmitted SMV isolates are able to infect the embryo, but only the seed-transmissible isolate is able to remain after seed maturation (Bowers & Goodman, 1979; Iwai & Wakimoto, 1990). In addition, SMV is able to infect the embryo not only directly, as can PSbMV, but also indirectly by pollen (Iizuka, 1973). Recently, a quantitative trait loci analysis on SMV seed transmission showed that this trait is controlled by several genes (Domier et al., 2011). In particular, two genomic regions associated with SMV seed transmission, and which contain soybean homologues of Arabidopsis genes DCL3 and RDR6 involved in RNA silencing, have

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been mapped. Whether RNA silencing might be associated with movement of viruses into the embryo during the seed-transmission process remains to be clarified. Regarding virus determinants, seed transmission of SMV has been shown to be influenced by P1, HCPro, and CP. Mutations in the DAG motif of CP, which is required for HCPro/CP interactions during aphid transmission (see above), significantly reduced the efficiency of transmission by seeds, suggesting that HCPro/CP interactions might be important for multiple functions in the potyviral infection ( Jossey, Hobbs, & Domier, 2013). Recent results with PSbMV suggest that vertical transmission by seeds could represent a narrow bottleneck causing a drastic genetic drift that might slow down virus adaptation and decrease virus fitness (Fabre et al., 2014). However, this severe bottleneck was not observed in seed transmission of another potyvirus, ZYMV (Simmons et al., 2013). Curiously, seed transmission of ZYMV always selected symptomless virus for reasons still unknown; the authors point out that the cryptic nature of vertical infection may enhance its contribution to virus spread.

7. PLANT/POTYVIRUS INTERACTIONS IN COMPATIBLE PATHOSYSTEMS Compatible interactions between viruses and their hosts depend on the capacity of viruses (i) to recruit efficiently host factors necessary for each of their infection steps and (ii) to counteract plant defense responses or overcome resistances. Though less studied, this also depends on not only the host genetic background but also the developmental and physiological situation of the plant, which appears to condition the output of the plant–virus interaction. As an example, a drastic transient loss of TuMV from roots was observed during the period of bud formation in A. thaliana (Lunello, Mansilla, Sanchez, & Ponz, 2007). In the previous sections, we have described most of the known host factors recruited by viruses for multiplication, movement, and transmission. In this section, we discuss the viral determinants involved in pathogenicity and host range as well as viral determinants involved in symptom development. We have dedicated a specific section on HCPro as a key pathogenicity factor and its function as a suppressor of gene silencing. Finally, we also produce a synthesis on transcriptomic studies performed over recent years on different compatible plant/potyvirus pathosystems.

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7.1. Evolutionary abilities of potyviruses to adapt to their hosts Numerous studies have been published in recent years not only on the identification of host-range determinants of several potyviruses but also on the evolutionary capacities of potyviruses to adapt to new hosts. The ability to adapt to new hosts is an important biological property of most RNA viruses. This is particularly important for plant viruses that infect annual crop species and therefore need alternative host species to be maintained in nature. RNA viruses are characterized by high mutation rates, short generation times, and large population sizes and consequently have a high evolutionary potential, which make them major pathogens responsible for emerging diseases (Elena, Bedhomme, et al., 2011; Elena & Sanjua´n, 2007). However, because of the small size of RNA virus genomes, which makes the production of multifunctional proteins necessary, evolutionary constraints exist for the host switching processes based on epistatic and pleiotropic effects of viral genome mutations, which generate trade-offs for the host adaptation (Elena, Bedhomme, et al., 2011; Elena & Lalic´, 2013; Elena & Sanjua´n, 2007; Garcı´a-Arenal & Fraile, 2013). Therefore, if the virus is completely unable to replicate in the new host, the success for host switching will depend on the preexistence of mutants in the viral quasispecies of the reservoir host able to initiate its replication in the new host and to further evolve to gain fitness. Experimental assays allowed assessing the genome plasticity and the evolutionary constraints of potyviruses (Elena et al., 2008), whereas site-directed mutagenesis or chimeric viral infectious clones were used to identify hostrange determinants. Many studies have focused on the adaptive capacity of TEV. One key parameter identified was the TEV genome mutation rate, which has been estimated from 1.2 to 3  105 substitution per nucleotide and generation, a rate similar to the few determined for other plant RNA viruses (Sanjua´n, Agudelo-Romero, & Elena, 2009; Sanjua´n, Nebot, Chirico, Mansky, & Belshaw, 2010; Tromas & Elena, 2010). Several studies revealed the strong adaptive capacity of TEV (Bedhomme, Lafforgue, & Elena, 2012, 2013; Lalic´, Cuevas, & Elena, 2011). Using a cDNA clone of TEV isolated from N. tabacum, Bedhomme et al. (2012) studied its adaptation to four Solanaceae species following different strategies of plant-to-plant passages. This produced TEV lineages well adapted to a particular host, in many cases without a fitness cost in the alternative hosts. Sequencing of the different lineages showed that the mutation accumulation rate is similar among lineages but the mutations are not scattered along the genome and are specific for each

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evolutionary history. Different lineages showed overrepresentation of mutations in different proteins, P1, HCPro, P3, 6K2, or NIa, but lineages sharing hosts in their evolutionary histories also shared common mutations. In a more recent study and using the TEV lineages previously produced (Bedhomme et al., 2012), Bedhomme et al. (2013) showed that TEV was able to readapt rapidly to N. tabacum whatever its evolutionary history, further increasing divergence, which means that TEV can reach the same fitness through various evolutionary pathways. There is however one exception for TEV lineages having evolved first on Capsicum annuum that was unable to infect N. tabacum. The high ability of TEV to broaden its host range and improve fitness in new hosts was further supported by the large amount of mutations showing a beneficial effect in partially susceptible nonSolanaceae host species (Lalic´ et al., 2011). In the case of TuMV, serial passaging of a Brassica-host type isolate (UK1) on Raphanus sativus (almost nonsusceptible) and Brassica rapa (susceptible) allowed isolation of variants able to systemically infect both plant species and identification of virus factors involved in the adaptation (Ohshima, Akaishi, Kajiyama, Koga, & Gibbs, 2010; Tan et al., 2005). Several nonsynonymous common mutations were found among the different adapted variants which were located in P1, P3, CI, and VPg. Whether all these mutations are involved in host adaptation of TuMV and how they act have not been determined yet. The C-terminal end of P3 was previously shown to be the viral determinant involved in the ability of TuMV to infect R. sativus (Suehiro, Natsuaki, Watanabe, & Okuda, 2004), which suggests that mutations in P3 alone might be sufficient to adapt to this plant species. Several studies were published in recent years on the identification of potyvirus host-range determinants using site-directed mutagenesis or chimeric viral infectious clones. Thus, the ability for Papaya ringspot virus (PRSV) to infect papaya was related to one mutation in NIaPro (Chen et al., 2008). A lysine at position 27 led to infection in papaya, whereas an aspartate led to the absence of infection in this host. However, although this amino acid change involves a reversal of the charge at this position, this mutation does not seem to modify the structure of the protein. Thus, the mechanism that blocks the papaya infectivity of PRSV with a NIaPro containing Asp27 is yet to be elucidated. Attempts to identify PPV determinants involved in differential infections of Prunus/herbaceous host reveal that determinants were located in several regions of the viral genome. Amino acid changes in the CP-Nter of PPV are responsible for host adaptation in Nicotiana spp. and Prunus spp.

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(Carbonell et al., 2013; Salvador, Delgadillo, et al., 2008). However, the adaptation of PPV to Pisum sativum is associated with a definite mutation in the NIb-coding sequence (Wallis et al., 2007). The ability to infect Prunus is associated with several determinants located in the 50 -terminal third of the genome, from the P1- to 6K1-coding sequences, suggesting complex virus– plant interactions in the ability of PPV to infect Prunus (Dallot et al., 2001; Salvador, Delgadillo, et al., 2008). Amino acid sequence comparison analysis suggested that P1, HCPro, and P3 might be involved together in adaptation to Prunus. Several studies support an important role for P1 in host adaptation of PPV and other potyviruses (Bejerman, Giolitti, de Breuil, & Lenardon, 2010; Chiang et al., 2007; Maliogka, Salvador, et al., 2012; Salvador, Sa´enz, et al., 2008; Valli et al., 2007; Yang et al., 2011). Valli et al. (2007) highlighted the role of natural intra- and interspecies recombinations in the P1 sequence upstream of the protease domain in potyvirus host adaptation. The fact that PPV clones with P1-coding sequence of TVMV were still able to infect PPV herbaceous hosts but not a Prunus host also revealed evidence for a role of P1 in host adaptation (Salvador, Sa´enz, et al., 2008). Recently, two amino acids in PPV P1 (at positions 29 and 139) were identified to be responsible for the loss of infectivity in Prunus (Maliogka, Salvador, et al., 2012) and single amino acid alterations in 6K1 and CI (corresponding to the cleavage site recognized by NIaPro at the 6K1/CI junction) have been shown to be involved in alternative host adaptation of atypical PPV isolates to N. benthamiana and Prunus avium (Calvo, Malinowski, & Garcı´a, 2014). In this last case, the authors hypothesized that fine regulation of the polyprotein processing might depend on specific host factors and contribute to the adaptation to particular hosts. Virus adaptation to the host does not necessarily consist in optimization of virus replication, which might have negative trade-offs for the virus. It has been speculated that host-dependent regulation of P1/HCPro processing might have evolved to attenuate virus virulence in order to alleviate antiviral responses (Pasin et al., 2014). This could account for the relationship of P1 protein with specific adaptation of potyviruses to particular hosts. At the plant intraspecies level, potyviruses are also able to adapt to resistant cultivars and several determinants allowing the virus to overcome resistance have been identified in recent years. One of the most studied determinants is VPg for overcoming eIF4E-mediated recessive resistances. Several excellent reviews have been recently published on eIF4E/VPg interaction (Le Gall, Aranda, & Caranta, 2011; Truniger & Aranda, 2009; Wang & Krishnaswamy, 2012) and we will present here only the most

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recent data. Briefly, for several potyviruses, mutations in the central region of VPg allow overcoming of eIF4E-mediated resistance by restoring the VPg/eIF4E interaction. As the interaction is necessary for the potyvirus infection cycle, mutated isolates are able to infect resistant cultivars. In the case of the TEV/pepper pathosystem, using a 3D interaction model, a recent study suggests that the different eIF4E/VPg interaction outcomes were due to conformational changes caused by amino acid sequence variation at critical positions in the central region of VPg and in the eIF4E predicted cap-binding region (Perez et al., 2012). Virus evolution studies showed that both VPg and eIF4E are highly constrained, but identified a small number of positively selected amino acid positions, mostly involved in potyvirus adaptation to their hosts, supporting coevolution between potyviral VPg and eIF4E (Ayme, Petit-Pierre, Souche, Palloix, & Moury, 2007; Charron et al., 2008; Moury et al., 2014). The ability of some potyvirus strains to use different eIF4E/eIF(iso)4E isoforms, especially in plants coding for several copies of these translation factors, enhances their ability to overcome eIF4E-mediated resistances and adapt to different hosts (Hwang et al., 2009; Jenner, Nellist, Barker, & Walsh, 2010; Piron et al., 2010; Ruffel et al., 2006). Recently, it was observed that eIF4E/eIF(iso) 4E genes that were not able to support the infection of a particular potyvirus in its natural genomic background, enabling broad-spectrum resistance, could be used by the same virus when these genes are expressed ectopically in another plant ( Jenner et al., 2010; Nellist et al., 2014). Differences in the cell expression patterns and in the production of mis-spliced variants have been suggested as possible causes of this discrepancy. The ability of TuMV mutants with single amino acid changes in VPg to infect A. thaliana mutants knocked out for eIF(iso)4E and eIF(iso)4F has led to the hypothesis that an additional strategy for potyviruses to escape eIF4E-mediated resistance might be to use an eIF4F-independent infection pathway (Gallois et al., 2010). The outcome of the VPg/eIF4E interaction is not the sole determinant of viral infectivity in eIF4E-mediated resistant plants. In the case of LMV, it was demonstrated that CI determines virulence against two eIF4E alleles in lettuce which correspond to the two mo1 recessive resistance genes (AbdulRazzak et al., 2009). Amino acid changes either in the VPg or in the C-terminal end of the CI are associated with resistance breaking of mo11, whereas only the CI domain is associated with resistance breaking of mo12. One amino acid change at position 621 in the CI seems to play an important role in this process (Abdul-Razzak et al., 2009). Recently, the

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LMV CI C-terminal domain has been shown to interact with both eIF4E and VPg but in a stronger manner in the case of the LMV isolate overcoming resistance compared to the avirulent isolate (Tavert-Roudet et al., 2012). These data could mean that a minimal interaction threshold is needed for infection success in resistant lettuce. Whether a ternary complex is formed between these three partners and whether these interactions are involved in LMV replication or cell-to-cell movement remains to be determined. For Clover yellow vein virus (ClYVV), the P1-coding region was shown to be the determinant to overcome the pea cyv2 resistance gene, which encodes eIF4E (Nakahara et al., 2010). Sequence comparison and site-directed mutagenesis allowed the identification of one amino acid at position 24 in P1 that seems necessary and sufficient to overcome the cyv2 resistance, although the compatibility with cyv2 pea is not fully restored. These data are intriguing, since cyv2 is the same eIF4E allele as sbm1 and wlm, which confer resistance to PSbMV and Bean yellow mosaic virus (BYMV), respectively, and which are only overcome by mutations in viral VPg (Andrade, Abe, Nakahara, & Uyeda, 2009; Bruun-Rasmussen et al., 2007; Gao et al., 2004; Keller, Johansen, Martin, & Hampton, 1998). Moreover, a role of HCPro in eIF4E-mediated resistance has been suggested on the basis of interactions of HCPro with VPg and eIF4E/eIF(iso)4E (Ala-Poikela et al., 2011; Roudet-Tavert et al., 2007). Altogether, these data suggest that the overcoming of eIF4E-mediated resistance by potyviruses is conditioned not only by the outcome of the binary interaction between eIF4E and VPg but also by interactions involving other viral proteins, such as CI, P1, and HCPro, which might form multiprotein complexes together with VPg and eIF4E. The pea cyv1, sbm2, and mo recessive resistance genes, conferring resistance to ClYVV, PSbMV, and BYMV, respectively, have not been cloned yet. Even if they map close to an eIF(iso)4E gene, no sequence differences have been detected in eIF(iso)4E between susceptible and resistant pea lines, suggesting that this gene is not involved in the resistance (Choi, Nakahara, Andrade, & Uyeda, 2012; Gao et al., 2004). In addition, the determinants allowing the virus to overcome these resistance responses were characterized for PSbMV and ClYVV and do not correspond to VPg but to P3. For PSbMV, the determinant to overcome sbm2 was mapped to the N-terminal part of P3 (Hjulsager, Lund, & Johansen, 2002; Hjulsager et al., 2006). Remarkably, not only are three P3 amino acids important for virulence in pea sbm2 lines but also amino acid insertion or deletion (depending on the virulent PSbMV isolates) in the same P3 region were

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associated with the virulence process (Hjulsager et al., 2006). For ClYVV, the genomic region responsible for overcoming cyv1-mediated resistance was also mapped in the P3 cistron for which both P3 and P3N–PIPO proteins are involved in breaking the resistance (Choi et al., 2013). P3N–PIPO showed a quantitative involvement: the more P3N–PIPO that was produced, the higher was the ClYVV virulence in cyv1 pea. It is worth mentioning that recent reports have shown that defects in interactions between translation initiation factors and different potyviral proteins might not only prevent infection in resistant varieties of susceptible host species but also contribute to nonhost resistance (Calvo, Martı´nez-Turin˜o, & Garcı´a, 2014; Estevan et al., 2014; SvanellaDumas et al., 2014). Appropriate mutations in viral genes conferring to the proteins coded by them compatibility with translation initiation factors of nonhost plants can contribute to broaden the host range of a particular potyvirus. Potyvirus infections can be also restricted by dominant resistance genes belonging to the NB-LRR gene family, and several potyviral avirulence (avr) genes have been identified. In soybean, three resistance genes, Rsv1, Rsv3, and Rsv4, were characterized against SMV with strain specificities (Hayes et al., 2004; Saghai Maroof et al., 2010; Suh et al., 2011). However, virulent SMV isolates were isolated for all three resistance genes. Several studies have reported the identification of SMV resistance breaking determinants. At least two SMV proteins are involved in the breaking of each of the three Rsv resistances. P3 is involved in virulence in soybean genotypes carrying each of three Rsv resistances, but it is associated with CI for virulence in the Rsv3- and Rsv4-genotype soybeans, and to HCPro in the Rsv1genotype soybeans. At least three mutations located within or near the C-terminal membrane domain of P3 (at positions 1033, 1053, and 1054) were essential for overcoming the Rsv4 resistance (Ahangaran, Habibi, Mohammadi, Winter, & Garcı´a-Arenal, 2013; Chowda-Reddy, Sun, Chen, et al., 2011; Khatabi, Fajolu, Wen, & Hajimorad, 2012), whereas the P3 N-terminal domain, but not the P3N–PIPO protein, was shown to be involved in overcoming the Rsv1 resistance (Chowda-Reddy, Sun, Hill, Poysa, & Wang, 2011; Eggenberger, Hajimorad, & Hill, 2008; Hajimorad, Eggenberger, & Hill, 2005, 2006; Wen, Saghai Maroof, & Hajimorad, 2011). The involvement of CI in overcoming Rsv3-mediated resistance is associated with either a single amino acid substitution at position 1754 in the C-terminus (Seo, Lee, & Kim, 2009) or both the N- and C-termini (Zhang et al., 2009). For HCPro, the C-terminal domain is

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involved in overcoming the Rsv1 resistance (Chowda-Reddy, Sun, Hill, et al., 2011; Eggenberger et al., 2008; Hajimorad, Eggenberger, & Hill, 2008; Hajimorad, Wen, Eggenberger, Hill, & Saghai Maroof, 2011; Khatabi, Wen, & Hajimorad, 2013). However, as none of the Rsv genes has been cloned, interaction between Rsv and SMV proteins (HCPro, P3, and CI) cannot be tested. Another point to be elucidated is whether each of the three potyviral proteins involved in overcoming Rsv resistances acts separately or in a complex with distinct roles. Recently, it was demonstrated that recognition of HCPro and P3 in Rsv1 genotypes was associated with distinct resistance genes at the Rsv1 locus (Wen, Khatabi, Ashfield, Saghai Maroof, & Hajimorad, 2013). These same potyviral proteins are virulence determinants for overcoming R genes against TuMV in Brassica. Indeed, P3 and CI are the virulence determinants for either TuRB03 and TuRB04 or TuRB01 and TuRB05, respectively ( Jenner et al., 2000; Jenner, Tomimura, Ohshima, Hughes, & Walsh, 2002; Jenner et al., 2003). A mutation at position 153 in P3 and the P3 C-terminus are involved in overcoming of TuRB03 and TuRB04, respectively ( Jenner et al., 2002, 2003). A mutation in the CI C-terminal domain is essential for overcoming TuRB05 ( Jenner et al., 2002), whereas two different mutations in the same CI domain are necessary for overcoming TuRB01 ( Jenner et al., 2000). Several viral determinants were also identified in other regions of the PVY genome. In the case of the potato resistance gene Ry, which confers extreme resistance to PVY, the protease domain of PVY NIa was identified as the elicitor of the resistance (Mestre, Brigneti, & Baulcombe, 2000). It was later shown that the protease activity of NIa might be necessary but not sufficient for elicitation (Mestre, Brigneti, Durrant, & Baulcombe, 2003). In tobacco plants carrying the resistance gene Rk, NIb is the elicitor of a veinal necrosis-HR to PVY infection (Fellers, Tremblay, Handest, & Lommel, 2002), and a point mutation in NIb confers PVY virulence toward pepper plants harboring the resistance gene Pvr4 ( Janzac, Montarry, Palloix, Navaud, & Moury, 2010). Two recent independent studies showed that the genetic determinants required to overcome or trigger the potato HR-associated dominant resistance to PVY controlled by the genes Nytbr, Nctbr, and Ncspl reside within HCPro (Moury et al., 2011; Tian & Valkonen, 2013). Fine mapping in HCPro identified regions in the central part of the protein between amino acids 227 and 327 and between amino acids 326 and 355 sufficient to overcome Nytbr and Ncspl, respectively (Moury et al., 2011; Tian & Valkonen,

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2013). Structure modeling of HCPro suggested conformational difference between virulent (PVYN) and avirulent (PVYO) PVY isolates that might explain HCPro functional differences in the recognition of the virus by the resistance system (Tian & Valkonen, 2013). These studies also highlighted that the PVY isolates able to overcome the Nytbr resistance gene have emerged by recombination instead of by successive accumulation of nucleotide substitutions, which has not been described before for potyviruses although numerous recombinant potyvirus isolates have been identified (Desbiez, Joannon, Wipf-Scheibel, Chandeysson, & Lecoq, 2011; Gagarinova, Babu, Stromvik, & Wang, 2008; Glasa et al., 2011; Mangrauthia, Parameswari, Jain, & Praveen, 2008; Ohshima et al., 2007; Valli et al., 2007; Visser, Bellstedt, & Pirie, 2012). Plant–potyvirus confrontation appears to be conditioned by other not well-characterized antiviral responses both induced and counteracted by viral factors. Thus, the CI protein of PPV interacts with a component of the chloroplastic photosystem I, PSI-K. Downregulation of the PSI-K expression led to higher virus accumulation, suggesting that PSI-K is involved in antiviral defense ( Jimenez et al., 2006). The fact that coexpression of the CI caused a decrease in the accumulation level of PSI-K transiently expressed in plant leaves suggests that CI might be counteracting the defensive role of PSI-K. Interaction of the CI protein with the host protein P58IPK also appears to be related with defensive responses of the plant and viral pathogenesis (Bilgin et al., 2003).

7.2. HCPro: A key pathogenicity determinant as suppressor of RNA silencing In addition to antiviral defense mediated by NBS-LRR genes, which respond to specific features of each virus, plants deploy innate immunity defenses that recognize general patterns shared by all viruses. This is the case of RNA silencing, a key player in the posttranscriptional regulation of physiological processes of the host, which provides virus-specific defense in response to the detection of a general elicitor, dsRNA (Voinnet, 2001). The relevance of RNA silencing for potyviruses is highlighted by the strict dependence of potyviral infections on functional RNA silencing suppressors (Garcia-Ruiz et al., 2010; Maliogka, Calvo, Carbonell, Garcia, & Valli, 2012), which can contribute to host specificity (Carbonell, Dujovny, Garcı´a, & Valli, 2012). The host RNA silencing machinery produces viral small RNAs that target effector complexes to viral genomic RNAs (Pantaleo, Szittya, &

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Burgya´n, 2007). The vast majority of virus-derived small RNAs in Arabidopsis plants infected with the potyvirus TuMV were dependent on DCL4 and RDR1, although full antiviral defense also required DCL2 and RDR6 (Garcia-Ruiz et al., 2010), and RNA silencing induced by PPV was shown to be compromised in RDR6-defective plants (Vaistij & Jones, 2009). The silencing suppressor is not always able to block completely the antiviral RNA silencing response, and a dynamic process of systemic silencing and silencing suppression can be established (Gammelg˚ard, Mohan, & Valkonen, 2007). RNA silencing and its viral suppressors appear to interact with other antiviral defenses of the plant, resulting in a complex interplay, in which the final result could depend on specific circumstances of each plant–virus combination (Alamillo, Sae´nz, & Garcı´a, 2006; Ji & Ding, 2001; Pruss et al., 2004). For instance, RDR1, which is upregulated by a potyviral infection in Nicotiana glutinosa (Liu, Gao, Wu, Ai, & Guo, 2009), contributes to SA-mediated antiviral defense (Xie, Fan, Chen, & Chen, 2001) and it is thought to be involved in the amplification phase of antiviral RNA silencing (Diaz-Pendon, Li, Li, & Ding, 2007; Garcia-Ruiz et al., 2010). In addition, a defect in RDR1 has been proposed to be the cause of the high susceptibility of N. benthamiana to many viruses (Yang, Carter, Cole, Cheng, & Nelson, 2004). However, transgenic expression of the N. tabacum RDR1 gene in N. benthamiana causes hypersusceptibility to the potyviruses PPV and PVY and to other viruses, resembling the effect of RDR6 deficiency. The authors suggest that RDR1 might have a dual role, on one hand contributing to SA-mediated antiviral defense, and on the other hand suppressing the RDR6-mediated antiviral RNA silencing (Ying et al., 2010). Although it has been demonstrated that both the genomic RNA and its complementary strand can be targeted by miRNA-guided processing, evidence for functional effects of RNA silencing mediated by endogenous small RNAs on potyviral infections is still lacking (Simo´n-Mateo & Garcı´a, 2006). It has been proposed that RNA silencing could be exploited not only by the plant to limit actual viral infection but also for the infecting virus to avoid the initiation of infections of competing viruses (Ratcliff, MacFarlane, & Baulcombe, 1999). Partial restrictions preventing two closely related potyviruses infecting the same cell have been revealed (Dietrich & Maiss, 2003; Zwart et al., 2011). Moreover, infection with mild natural isolates (Lecoq, Lemaire, & Wipf-Scheibel, 1991) or engineered virus mutants

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can protect plants against potyviruses causing severe diseases (Lin, Wu, Jan, Hou, & Yeh, 2007; Nakazono-Nagaoka et al., 2009). However, this crossprotection only works efficiently between virus species showing high levels of genetic similarity (Capote et al., 2006; Nakazono-Nagaoka et al., 2009). A direct role of RNA silencing in cross-protection has not been demonstrated yet. Mixed infections are usual in nature, and in many cases the interaction of the infecting viruses enhances pathogenicity in a synergistic way (Roossinck, 2005). Usually synergistic diseases involve a potyvirus and a virus of a different family. Although sometimes the potyvirus benefits from the interaction (Karyeija, Kreuze, Gibson, & Valkonen, 2000; Mukasa, Rubaihayo, & Valkonen, 2006), it is common that the replication of the second virus, but not that of the potyvirus, is enhanced (Lim et al., 2007; Mascia et al., 2010; Rochow & Ross, 1955; Vance, 1991; Wang, Turina, Medina, & Falk, 2009; Wang, Gaba, Yang, Palukaitis & Gal-On, 2002). Different sources of evidence demonstrate that HCPro is the potyviral factor that enhances pathogenicity and viral replication of the partner virus in most of these infections (Fukuzawa et al., 2010; Gonza´lez-Jara et al., 2005; Pruss, Ge, Shi, Carrington, & Vance, 1997; Sa´enz, Quiot, Quiot, Candresse, & Garcı´a, 2001; Savenkov & Valkonen, 2001; Sonoda et al., 2000; Yang & Ravelonandro, 2002). Other potyviral proteins can also contribute to the synergistic effect, as it is the case of P3N–PIPO of ClYVV. However, whereas both P3N–PIPO and HCPro of ClYVV exacerbated symptom severity of the potexvirus White clover mosaic virus, only HCPro enhanced the accumulation of the potexvirus partner (Hisa et al., 2014). HCPro is the protein of potyviruses involved in suppressing antiviral RNA silencing. Although there is not a perfect correlation between the ability of HCPro to suppress silencing and virus virulence (Lin et al., 2007; Torres-Barcelo´, Martı´n, Daro`s, & Elena, 2008), silencing suppression appears to be directly related with the effect of HCPro in enhancement of pathogenicity both in single potyviral infections and in mixed infections including a potyvirus (Atsumi, Kagaya, Kitazawa, Nakahara, & Uyeda, 2009; Atsumi et al., 2012; Gonza´lez-Jara et al., 2005; Yambao et al., 2008). For some members of the family Potyviridae, in which suppression of RNA silencing is carried out by a protein different from HCPro, this other protein is the pathogenicity enhancer (Stenger, Young, Qu, Morris, & French, 2007; Tatineni, Qu, Li, Morris, & French, 2012; Valli, Dujovny, & Garcı´a, 2008; Young et al., 2012).

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HCPro is able to bind small RNAs, and it has been proposed that it suppresses silencing by sequestering viral small RNAs (Lakatos et al., 2006). It is interesting that a mutation in the FRNK motif of HCPro abolishes small RNA binding but does not affect the silencing suppression activity of the protein (Shiboleth et al., 2007). This conflict can be due to differences in in vitro and in vivo small RNA binding, as has been previously noted (Valli, Oliveros, Molnar, Baulcombe, & Garcia, 2011), as well as to differences in affinity of the mutant protein for different small RNA populations (Wu, Lin, Chen, Yeh, & Chua, 2010). However, it is also possible that HCPro has an additional, small RNA binding-independent, method to suppress silencing. In fact, it has been suggested that interaction of HCPro with the transcription factor RAV2 might induce the expression of host factors that interfere with antiviral silencing. On the other hand, another protein that interacts with HCPro, the calmodulin-like protein rgs-CaM has been proposed to be an endogenous suppressor of RNA silencing that mediates the HCPro activity (Anandalakshmi et al., 2000). However, more recently, it has been shown that rgs-CaM counteracts RNA silencing suppression by sequestering HCPro and facilitating its degradation by an autophagy-like mechanism (Nakahara et al., 2012). HCPro also interacts with another important component of the RNA silencing machinery, the protein HEN1, inhibits its methyltransferase activity in vitro ( Jamous et al., 2011), and blocks methylation of small RNAs in vivo (Lozsa, Csorba, Lakatos, & Burgya´n, 2008). It has been shown that whereas HCPro suppresses antiviral silencing, it enhances other defensive responses of the plant (Pruss et al., 2004; ShamsBakhsh, Canto, & Palukaitis, 2007), which also can cause notable pathogenic effects. Thus, interaction of HCPro with the microtubule-associated protein HIP2 appears to regulate an SA-unrelated host defense (Haikonen et al., 2013). Moreover, the fact that a recombinant Potato virus X expressing HCPro causes a systemic necrosis that might be related to a proteasome dysfunction (Pacheco et al., 2012) suggests that the interaction of HCPro with proteasome subunits, which interferes with nuclease and protease activities, (Ballut et al., 2005; Dielen et al., 2011; Jin, Ma, Dong, Jin, et al., 2007; Sahana et al., 2012) can also contribute to the potyvirus pathogenicity.

7.3. Symptomatology Compatible or partially compatible virus/host interactions affect host physiology and in some cases induce disease symptoms. However, complexities

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of the virus/host interactions make it difficult to decipher mechanisms of symptom induction. The nature and extent of symptoms depend upon the virus/host pathosystem as well as upon environmental conditions that influence plant physiology and development. In addition, temporal and spatial variations during the time course of virus infection add another level of complexity to identify causal events in symptom development (Culver & Padmanabhan, 2007; Maule, Leh, & Lederer, 2002; Pallas & Garcı´a, 2011). Between the two models proposed to explain symptom development, the data are in favor of the interaction model in which disruptions of host processes are related to specific interactions between viral and host components rather than the competitive disease model in which viruses usurp a substantial amount of plant metabolic resources for their own molecular synthesis (Culver & Padmanabhan, 2007). Potyviruses cause a wide range of symptoms and many of them induce severe diseases causing important economical losses on crops. Potyviruses usually induce longitudinal chlorotic or necrotic streaks in the leaves of monocotyledonous species, and chlorotic vein banding, mosaic mottling, necrosis, or/and distortion of leaves in dicotyledonous species as well as alterations on flowers, seeds, and fruits (Shukla, Ward, & Brunt, 1994). For many years, numerous studies have been published not only on viral determinants but also on host changes during infection, particularly gene expression changes thanks to highthroughput transcript profiling techniques, to identify viral and host factors triggering symptom development. HCPro is probably the most important potyvirus symptom determinant as demonstrated in several studies. Mutations in HCPro cause drastic alterations in the disease symptoms (Atreya et al., 1992; Chiang et al., 2007; Desbiez, Girard, & Lecoq, 2010; Faurez, Baldwin, Tribodet, & Jacquot, 2012; Gal-On, 2000; Haikonen et al., 2013; Hu, Karasev, Brown, & Lorenzen, 2009; Klein et al., 1994; Lin et al., 2007; Sa´enz et al., 2001; Seo, Sohn, & Kim, 2011; Shiboleth et al., 2007; Tribodet, Glais, Kerlan, & Jacquot, 2005; Yambao et al., 2008). Moreover, expression of HCPro out of the context of viral infection produces developmental abnormalities and significantly alters the gene expression profile of the plant (Chapman, Prokhnevsky, Gopinath, Dolja, & Carrington, 2004; Dunoyer, Lecellier, et al., 2004; Kasschau et al., 2003; Mallory et al., 2001; Mangrauthia, Singh, & Praveen, 2010; Siddiqui, Sarmiento, Truve, Lehto, & Lehto, 2008; Soitamo et al., 2011). HCPro affects the accumulation and activity of endogenous microRNAs (Mallory et al., 2002). It has not been demonstrated if this is a collateral effect

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of the activity against the viral small RNAs or if it contributes to optimize viral infection. In any case, the misregulation of endogenous RNA silencing by HCPro is a main cause of disease symptoms, which resemble developmental defects (Chapman et al., 2004; Dunoyer, Lecellier, et al., 2004; Kasschau et al., 2003). In fact, it has been demonstrated that upregulation of the miR167 target AUXIN RESPONSE FACTOR 8 underlies developmental aberrations exhibited by transgenic plants expressing HCPro, and disease symptoms caused by the potyvirus TuMV ( Jay et al., 2011). However, HCPro appears also to contribute to pathogenicity by other mechanisms not directly related with RNA silencing suppression. Thus, the SA-unrelated host defense to PVA infection mediated by HCPro– HIP2 interaction mentioned above causes necrotic symptoms (Haikonen et al., 2013). Nevertheless, other determinants of symptom induction scattered throughout the viral genome have been identified and some of them were already described in previous reviews on potyviruses (Rajama¨ki et al., 2004; Revers et al., 1999; Urcuqui-Inchima et al., 2001; Table 2). Thus, the 50 -NCR [PPV; chlorotic mosaic symptoms in Nicotiana clevelandii (Simo´n-Buela et al., 1997)], the C-terminal part of P1 [PPV; mild/severe symptoms on Nicotiana species and Prunus persica (Maliogka, Salvador, et al., 2012; Nagyova´ et al., 2012)], P3 [SMV; systemic necrosis in Rsv1-genotype soybean (Hajimorad et al., 2005); TuMV; systemic HR in A. thaliana (Kim et al., 2010)], the N- and C-terminal part of P3 [ZYMV; symptom severity in zucchini squash (Desbiez et al., 2003); TuMV; yellow mosaic symptoms in Brassica juncea ( Jenner et al., 2003)], the C-proximal part of P3 and 6K1 [PPV; chlorotic mottle symptoms in N. benthamiana (Sa´enz et al., 2000)], the P3/6K1 junction [PPV; symptom severity in N. clevelandii (Riechmann et al., 1995)], the P3 and CI–6K2–VPg encompassing regions [TEV; wilting response in Tabasco pepper (Chu et al., 1997)], VPg associated with P3 [TEV; severe symptom in A. thaliana (Agudelo-Romero et al., 2008)], the N-terminal region of CI [SMV; severe symptom on soybean (Zhang et al., 2009)], a genomic segment encoding NIa and NIb [PSbMV; vein-clearing symptoms in pea ( Johansen et al., 1996)], and the 30 -NCR [TVMV; vein-mottling and blotch symptoms in tobacco (Rodrı´guez-Cerezo et al., 1991)] were all involved in symptom development. However, the mechanisms of symptom induction involving these different potyvirus determinants remain to be elucidated. Only a few host genes playing a direct role in symptom development have been identified so far. TuNI was shown to be involved in the systemic

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Table 2 Potyvirus symptom determinants Potyvirus/host Genomic region Symptom pathosystems

References

50 -NCR

Chlorotic mosaic symptoms

PPV/ N. clevelandii

Simo´n-Buela, Guo, and Garcı´a (1997)

P1

Mild/severe symptoms

PPV/Nicotiana– Prunus persica

Maliogka, Salvador, et al. (2012) and Nagyova´, Kamencayova´, Glasa, and Sˇubr (2012)

HCPro

Severe symptoms

Several pathosystems

See in the text

P3

Systemic necrosis

SMV/soybean

Hajimorad et al. (2005)

Systemic HR

TuMV/ A. thaliana

Kim, Suehiro, Natsuaki, Inukai, and Masuta (2010)

P3 N- and Severe C-terminal parts symptoms

ZYMV/zucchini Desbiez, Gal-On, Girard, squash Wipf-Scheibel, and Lecoq (2003)

Yellow mosaic

TuMV/Brassica juncea

Jenner et al. (2003)

P3 C-terminal part and 6K1

Chlorotic mottle symptoms

PPV/ N. benthamiana

Sa´enz et al. (2000)

P3/6K1 junction

Severe symptoms

PPV/ N. clevelandii

Riechmann et al. (1995)

P3 and CI– 6K2–VPg encompassing regions

Wilting response

TEV/tabasco pepper

Chu, Lopez-Moya, LlaveCorreas, and Pirone (1997)

VPg associated with P3

Severe symptoms

TEV/A. thaliana Agudelo-Romero et al. (2008)

CI N-terminal part

Severe symptoms

SMV/soybean

Zhang et al. (2009)

NIa and NIb

Vein-clearing symptoms

PSbMV/pea

Johansen et al. (1996)

30 -NCR

Vein-mottling and blotch symptoms

TVMV/tobacco Rodrı´guez-Cerezo, Klein, and Shaw (1991)

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necrosis induced by TuMV infection in the ecotype Ler of A. thaliana (Kaneko, Inukai, Suehiro, Natsuaki, & Masuta, 2004). This necrosis phenotype is similar to the HR and was considered a form of defense response accompanying an HR-like cell death in the veinal area inhibiting TuMV movement from phloem to mesophyll cells (Kim, Masuta, Matsuura, Takahashi, & Inukai, 2008). Induction of the necrosis is mediated by SA and ethylene and is also regulated by light intensity (Kim et al., 2008). A few studies have described ultrastructural, biochemical, and metabolic changes in potyvirus-infected cells. As mentioned above, the cytoplasm usually reveals typical features of infection by potyviruses, such as cylindrical inclusions and proliferated ER, but in most instances, no changes were observed in nuclei, mitochondria, or peroxysomes (Shukla et al., 1994). In contrast, chloroplasts were notably affected during potyvirus infection. Pompe-Novak, Wrischer, and Ravnikar (2001) described chloroplast alterations in and around necrotic spots in PVY-infected potato. Zechmann, M€ uller, and Zellnig (2003) showed less chloroplasts in pumpkins cells infected with ZYMV than in healthy plants, with a reduced amount of thylakoids, but with more starch and plastoglobuli. In the case of PPV in peach and pea, infection caused an increase in the number and size of plastoglobuli in chloroplasts, which showed dilated thylakoids and lower starch content associated with alteration of chloroplast metabolism and PSII electron transport (Dı´az-Vivancos et al., 2008; Herna´ndez, Rubio, Olmos, Ros-Barcelo´, & Martı´nez-Go´mez, 2004). PPV infection in susceptible Prunus species and pea also produced an oxidative stress associated with an increase in lipid peroxidation, protein oxidation, electrolyte leakage, and H2O2 levels, and an alteration in the levels of antioxidant enzymes in soluble chloroplastic fractions of PPV-infected leaves (Dı´az-Vivancos et al., 2008, 2006; Herna´ndez et al., 2006, 2004). Oxidative stress associated with increase of peroxidase activities were also described in the case of Cucumis sativus and Cucurbita pepo infected with ZYMV (Riedle-Bauer, 2000). In tobacco infected with either PVY or PVA, it was shown that photosynthesis and anaplerotic metabolic pathways were also altered (Rysˇlava´, ˇ erˇovska´, 2003). These effects were M€ uller, Semora´dova´, Synkova´, & C NTN notably observed with PVY , which induces stronger symptoms and accumulates at higher level than PVYO or PVA (Doubnerova´ et al., 2009; Rysˇlava´ et al., 2003). Symptom expression in sunflower infected with Sunflower chlorotic mottle virus (SuCMoV) has been also studied. SuCMoV infection caused a decrease in the chlorophyll content, photosynthetic proteins, and apoplastic ROS; an increase in soluble sugars, starch content, and

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antioxidant enzyme activities; and inhibition of photosynthesis (Arias, Luna, Rodrı´guez, Lenardon, & Taleisnik, 2005; Rodrı´guez, Mun˜oz, Lenardon, & Lascano, 2012; Rodrı´guez, Taleisnik, Lenardon, & Lascano, 2010). Several chloroplast genes were downregulated from the early to the late phases of the symptom development process (Rodrı´guez et al., 2012). Changes in the cellular redox homeostasis, probably caused by higher sugar availability, were also revealed associated with chlorotic symptom development during SuCMoV infection (Rodrı´guez et al., 2012; Rodrı´guez, Mun˜oz, Lenardon, & Lascano, 2013). During the last decade, high-throughput approaches for transcriptome, proteome, and metabolome analyses have allowed more comprehensive pictures of the global effects of virus infection, particularly for potyviruses, to be obtained. Four proteomic analyses upon potyvirus infection have been published during this time (Dı´az-Vivancos et al., 2008, 2006; Wu, Han, et al., 2013; Wu, Wang, et al., 2013). Analysis of changes in the leaf apoplast proteome of P. persica associated with PPV infection revealed the induction of a pathogenesis-related thaumatin-like protein and the decrease of mandelonitrile lyase, but mostly identified unknown proteins (Dı´azVivancos et al., 2006). In a second study by the same research team that focused on the pea/PPV pathosystem, most of the changes of protein expression observed were related to photosynthesis and carbohydrate metabolism, with 12 and 17 proteins differentially expressed in the soluble and in the chloroplast fraction, respectively (Dı´az-Vivancos et al., 2008). Recently, two proteomic studies have been published for the maize/ Sugarcane mosaic virus pathosystem, one analyzing changes at 6 days postinoculation (dpi), whereas the other one analyzed apical leaves at 12 dpi (Wu, Han, et al., 2013; Wu, Wang, et al., 2013). In inoculated leaves, 47 proteins were identified as differentially expressed. More than 40% of these proteins are related to energy and metabolism and 25% are related to stress and defense, and almost 50% of these proteins are located in the chloroplast (Wu, Wang, et al., 2013). In apical leaves, the study detected changes in expression of 44 proteins, which were classified as functionally related to energy and metabolism, stress and defense response, carbon fixation, photosynthesis, protein synthesis and folding, or signal transduction and transcription, structural or unknown. These differentially expressed proteins are also mostly located in chloroplast (Wu, Han, et al., 2013). Similar proteome changes have been also observed in infections caused by viruses of other genera (Di Carli, Benvenuto, & Donini, 2012).

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A larger number of transcriptomic studies than proteomic ones have been published during this last decade on host/potyvirus pathosystems. Most of them were carried out on A. thaliana infected with TuMV (Whitham et al., 2003; Yang et al., 2007), PPV (Babu, Griffiths, Huang, & Wang, 2008), and TEV (Agudelo-Romero et al., 2008; Hillung, Cuevas, & Elena, 2012), but a few other studies focused on other plant species, such as pea infected with PSbMV (Aranda, Escaler, Wang, & Maule, 1996; Wang & Maule, 1995), N. benthamiana (Dardick, 2007), peach (Rubio et al., 2014), and plum (Rodamilans et al., 2014) infected with PPV, soybean infected with SMV (Babu, Gagarinova, Brandle, & Wang, 2008; Bilgin et al., 2008), tomato infected with Pepper yellow mosaic virus (AlfenasZerbini et al., 2009), potato infected with PVY (Baebler et al., 2009; Kogovsˇek et al., 2010; Pompe-Novak et al., 2006), or sunflower infected with SuCMoV (Rodrı´guez et al., 2012, 2013). The first studies on changes on host gene expression during potyvirus infection were carried out by the Maule group in the 1990s. In situ hybridization was used to follow expression of a few genes involved in metabolism, particularly in starch synthesis, or coding for chaperones/ubiquitin in cotyledon cells at the infection front where virus replication is the most active (Aranda et al., 1996; Wang & Maule, 1995). They highlighted downregulation [termed shutoff (Aranda & Maule, 1998)] of metabolism-related genes and upregulation of chaperons/ubiquitin genes. There was complete overlap between the host transcript changes and the location of PSbMV replication. Behind the infection front, host gene expression was restored, showing that regulation of host genes (shutoff and induction) is strictly associated with active viral replication (reviewed in Aranda & Maule, 1998; Maule, Escaler, & Aranda, 2000). Subsequently, microarray technologies allowed thousands of host genes to be analyzed during potyviruses infection (AgudeloRomero et al., 2008; Babu, Gagarinova, et al., 2008; Babu, Griffiths, et al., 2008; Baebler et al., 2009; Bilgin et al., 2008; Dardick, 2007; Hillung et al., 2012; Kogovsˇek et al., 2010; Rodamilans et al., 2014; Rubio et al., 2014; Whitham et al., 2003; Yang et al., 2007). Even if direct comparisons across experiments are not straightforward because of differences in profiling techniques and platforms, plant ecotypes and species, sampling schemes, as well as inoculation and growth environmental conditions, some generalities can be drawn from these studies. Several developmental functions, hormone signaling, biosynthesis of lipids, alcohols and polysaccharides, and secondary metabolism constitute the principal downregulated processes. Plastid genes and genes involved in chloroplast functions are also

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preferentially underexpressed. Overexpressed genes are associated with cell rescue, defense, apoptosis and cell death plus aging, including several defense- and stress-associated genes. Heat-shock proteins, ribosomal proteins, and protein turnover genes are also overexpressed after infection with viruses (reviewed in Elena, Carrera, & Rodrigo, 2011; Whitham, Yang, & Goodin, 2006). However, downregulation of photosynthesis genes and upregulation of host defense genes are not specific to potyviruses and even to other virus genera as they are usually observed in the case of biotic stresses (Bilgin et al., 2010) and even under abiotic stresses, regarding photosynthesis genes (Saibo, Lourenc¸o, & Oliveira, 2009). Among the transcriptomic analyses performed with potyviruses, a very interesting illustration of gene expression changes in relation to symptom development comes from several studies comparing gene expression profiles of hosts infected with the same potyvirus but with isolates differing by the severity of the induced symptoms. Agudelo-Romero et al. (2008) compared gene expression profiles of TEV-infected A. thaliana plants infected with either a nonadapted ancestral TEV isolate or an adapted TEV isolate (TEV-At17). The ancestral virus, although systemically infecting the A. thaliana Ler ecotype, does not induce symptoms, whereas the adapted TEV isolate not only accumulates at higher level but also induces severe symptoms. The transcriptomic analyses showed a set of differentially expressed genes almost three times larger for TEVAt17. In particular, among the underexpressed genes targeted by TEVAt17, the proportion of genes involved in metabolic processes is much higher, which might be related to the symptoms induced by this isolate. In addition, whereas genes involved in SAR and in activation of innate immune responses were overexpressed in plants infected with the ancestral TEV, these genes were not expressed in plants infected with TEV-At17. This suggests that the evolved virus acquired the ability to evade the plant defense responses more efficiently, which might explain the observed increase of viral load and the development of severe symptoms. In another study, the same group compared the gene expression profiles of a panel of six A. thaliana ecotypes infected with TEV-At17 for which they observed a correlation between infectivity, virus accumulation, or symptom severity (Hillung et al., 2012; Lalic´, Agudelo-Romero, Carrasco, & Elena, 2010). From this comparison, neighbor-joining dendrograms revealed two groups of ecotypes, which presented particular phenotypes upon infection with TEV-At17 and which shared several hundreds of altered genes. In one ecotype group, genes involved in abiotic stresses and cell wall construction were upregulated, whereas genes involved in secondary metabolism and some

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hormone-regulated pathways were downregulated, and in the second group, defense genes were upregulated, whereas genes involved in the production of cell wall components were shut down. All these data revealed the large heterogeneity of host/virus interactions resulting in diverse host responses at the molecular level. The fact that many of the assayed pathosystems were developed in the laboratory and did not have a history of host–virus coevolution might have contributed to their heterogeneity. However, even in natural host/ potyvirus pathosystems, host responses vary depending on the specific host cultivars and virus strains analyzed. Kogovsˇek et al. (2010) compared infections caused by two PVY strains (an aggressive strain, PVYNTN and a mild strain, PVYN) in two potato cultivars at very early times in inoculated leaves. Regardless of the plant cultivar, whereas immediately after inoculation the genes involved in photosynthesis were more highly expressed in plants inoculated with PVYNTN than in those inoculated with PVYN, a lower expression of photosynthesis-related genes was observed in PVYNTN- than in PVYN-inoculated plants at 12 and 48 hpi. An earlier accumulation of sugars was also observed in cultivars inoculated with PVYNTN. Antioxidant metabolism-associated genes were differentially expressed between both PVY strains but with opposite effects depending on the potato cultivar. A very recent study compared two strains of TuMV infecting A. thaliana focusing on host symptoms development and senescence progression (Manacorda et al., 2013). Both strains (UK1 and JPN1) accumulated at similar levels, but JPN1 induced milder symptoms and developmental effects. Fresh weight as well as chlorophyll and anthocyanin contents differed significantly between plants infected with each virus isolate. UK1, but not JPN1, induced ROS accumulation in systemically infected leaves, which might be responsible for the growth arrest observed in UK1-infected plants. Both viruses induced overexpression of senescence-associated genes, but alterations induced by UK1 were more pronounced. In addition, it was shown that most of these differential responses between UK1- and JPN1-infected plants were controlled by SA. In several studies, SA was also shown to play a role in symptom development (Atsumi et al., 2009; Baebler et al., 2011; Krecˇicˇ-Stres, Vucˇak, Ravnikar, & Kovacˇ, 2005; Nie, 2006). For instance, in the case of the PVY/potato pathosystem, NahG potato plants, which express a bacterial enzyme that hydrolyzes SA, showed more severe symptom upon PVY infection than wild plants (Baebler et al., 2011). These data, together with others, suggest that mechanisms which govern senescence-like symptoms

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induced by viruses are interconnected with those which control natural senescence and involve SA. Comparing the gene expression profiles of N. benthamiana plants infected with three viruses differing in aggressiveness in this host, Dardick (2007) observed that the number of genes in which expression was altered by the infection correlated with the severity of symptoms. However, this seemingly logical correlation appears not to be general, since in the SMV/soybean pathosystem, Babu, Gagarinova, et al. (2008) showed that the highest number of virus-regulated genes was found at 14 dpi in leaves with moderate mosaic symptoms and not later in leaves with more severe mosaic symptoms. In an attempt to connect expression profiles with protein–protein interaction networks, Rodrigo et al. (2012) carried out a meta-analysis of published transcriptomic data of A. thaliana/virus pathosystems and revealed that viruses alter the expression of master transcription factors and hub proteins. In addition, this analysis showed that phylogenetically related viruses significantly alter the expression of similar genes and that viruses naturally infecting Brassicaceae display a greater overlap in the plant response (Elena, Carrera, et al., 2011; Rodrigo et al., 2012). Virus infections displaying disease symptoms can undergo phenotypic alterations when plants are subjected to additional stresses. A few studies on this topic were published in the case of potyviruses. Soybean plants exposed to elevated levels of ozone (O3) exhibited a delay in systemic infection with SMV associated with a reduced negative effect on photosynthesis (Bilgin et al., 2008). Ozone treatment, which mimics pathogen infection, induced defense-related gene expression, and enhanced SMV resistance. A comparison of the gene expression profiles of SMV-infected plants, O3-treated plants, and plants exposed to both stresses showed specific profiles for each treatment, even if a number of differentially regulated soybean genes were common in response to all treatments (Bilgin et al., 2008). Recently, a study was carried out to assess the effect of combining heat and drought stresses with TuMV infection on A. thaliana (Prasch & Sonnewald, 2013). Plants were exposed to these three stresses in single, double, or triple combinations, and transcriptomic and metabolic changes were analyzed. The number of differentially regulated features increased with the complexity and severity of the stress, but the comparative analysis revealed a small group of stress-regulated genes specifically expressed in one or more situations. Abiotic stress significantly altered TuMV-specific signaling networks, and, in the case of heat stress, enhanced virus accumulation twoto threefold.

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8. BIOTECHNOLOGICAL APPLICATIONS OF POTYVIRUSES Currently, viruses are not considered only as pathogens. They can be also useful biotechnological tools (Hefferon, 2012). Different potyviruses have been engineered to be used as expression vectors for the production of foreign products in plants. Historically, the first potyviral vectors had the insertion site between the P1- and HCPro-coding sequences (Dolja, McBride, & Carrington, 1992; German-Retana, Candresse, Alias, Delbos, & Le Gall, 2000; Guo, Lo´pez-Moya, & Garcı´a, 1998). Then, vectors with foreign sequences inserted at the NIb/CP junction were also constructed (Arazi et al., 2001; Ferna´ndez-Ferna´ndez et al., 2001). More recently, it has been shown that the borders of other protein-coding sequences can also be used as cloning sites (Bedoya, Martı´nez, Rubio, & Daro`s, 2010; Chen et al., 2007). In addition, insertion sites inside viral cistrons have also been engineered in potyviral vectors to produce free foreign proteins or foreign peptides fused to viral proteins (Ferna´ndez-Ferna´ndez, Martı´nez-Torrecuadrada, Casal, & Garcı´a, 1998; Ferna´ndez-Ferna´ndez, Martı´nez-Torrecuadrada, Roncal, Domı´nguez, & Garcı´a, 2002; Rajama¨ki et al., 2005). The cloning capacity of potyviral expression vectors has been enlarged by using several insertion sites (Beauchemin, Bougie, & Laliberte´, 2005; Kelloniemi, Ma¨kinen, & Valkonen, 2008) or removing essential viral genes, which are then supplied in trans by a transgene (Bedoya et al., 2010; Chen et al., 2007). Potyviral amplicon systems, which combine the genetic stability of transgenic plants with the high-amplification potential of viruses, have also been constructed (Calvo et al., 2010; Dujovny, Valli, Calvo, & Garcı´a, 2009). Although most of the reports about vectors derived from potyviruses mainly describe its design and its performance to express reporter genes (Bedoya, Martı´nez, Orza´ez, & Daro`s, 2012; Gao et al., 2012; Kelloniemi et al., 2008; Naderpour & Johansen, 2011), the use of these expression vectors to produce proteins of interest has also been published (Arazi et al., 2002; Ferna´ndez-Ferna´ndez et al., 1998, 2001; Hsu, Lin, Liu, Su, & Yeh, 2004; Kelloniemi, Ma¨kinen, & Valkonen, 2006; Shiboleth, Arazi, Wang, & Gal-On, 2001). Not only the complete potyviruses but also individual potyviral factors can have practical applications. The high specificity and efficiency of the

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protease NIaPro confer upon it a remarkable biotechnological interest. It is being used both to remove tags added to proteins to facilitate their detection and purification (Parks, Leuther, Howard, Johnston, & Dougherty, 1994; Zheng et al., 2008) and for the simultaneous expression of multiple proteins from a single self-processing cassette in transgenic plants (Marcos & Beachy, 1997). NIaPro has been exploited also for the expression of recombinant proteins in E. coli (Pe´rez-Martı´n, Cases, & de Lorenzo, 1997) and for the development of an effective system for detecting protein–protein interactions (Zheng, Huang, Yin, Wang, & Xie, 2012).

9. CONCLUDING REMARKS During this last decade, our knowledge on the molecular biology of potyviruses has advanced considerably. After characterizing the structure and strategy of expression of the potyviral genome, and identifying the basic functions of most potyviral protein, recent research has boosted our understanding of potyvirus replication and cell-to-cell movement processes, for which not only viral and host factors directly involved have been identified, but also their subcellular localizations have been determined. Several host factors interacting with potyvirus proteins have also been identified, and for some of them, a direct role in potyvirus infection has been described (Table 1). Another aspect well characterized during the last decade is the disturbance of the host gene expression during potyvirus infection thanks to the high-throughput approaches for transcriptomic and proteomic analyses available to the research community. All these data now available allow considering the building of gene networks associated with transcriptomic metaanalyses to get a global picture of the plant/potyvirus interactions that can help identify new candidate genes for resistance strategies. Nevertheless, despite this new knowledge, many aspects of plant/potyvirus interactions are not well understood. Very little information is available on how RNA molecules are specifically sorted for translation, replication and encapsidation, and how these processes are coupled in precise locations within the infected cells. Long-distance movement is still a black box. No host factor necessary for movement of potyvirus in the SE has been identified and the precise nature of the viral transport form remains to be elucidated. The dynamic processes which govern aphid and seed transmission are not well defined and the aphid and host factors involved are still unknown. Despite the high number of viral determinants identified in many plant/ potyvirus pathosystems that control viral pathogenicity or symptom

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development, the host targets of such determinants have to be found and functionally characterized. Thus, numerous efforts are still necessary to decipher the molecular mechanisms which govern plant/potyvirus interaction.

ACKNOWLEDGMENTS Helpful comments and criticism on the manuscript from Jari Valkonen and Peter Palukaitis are gratefully acknowledged. The research of the authors was supported by grants ANR-08GENM-016-001 (Viromouv) of the ‘ge´nomique des plantes’ program of the French Agence Nationale de la Recherche to FR, and BIO2013-49053-R and Plant KBBE PCIN-2013056 of Spanish Ministerio de Economı´a y Competitividad to JAG.

Note Added in Proof Ivanov, K. I., Eskelin, K., Lo˜hmus, A., and Ma¨kinen, K. (2014). Molecular and cellular mechanisms underlying potyvirus infection. Journal of General Virology 95, 1415–1429. Ma¨kinen, K., and Hafren, A. (2014). Intracellular coordination of potyviral RNA functions in infection. Frontiers in Plant Science 5, 110.

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CHAPTER ONE

Comparison of Lipid-Containing Bacterial and Archaeal Viruses Nina S. Atanasova, Ana Senčilo, Maija K. Pietilä, Elina Roine, Hanna M. Oksanen, Dennis H. Bamford1 Department of Biosciences and Institute of Biotechnology, University of Helsinki, Helsinki, Finland 1 Corresponding author: e-mail address: [email protected]

Contents 1. Introduction 1.1 Origin of lipids in prokaryotic viruses and their detection 2. Function and Significance of Lipids in Prokaryotic Virus Life Cycle 2.1 How prokaryotic viruses acquire their lipids 3. Currently Known Lipid-Containing Bacterial and Archaeal Viruses 3.1 Icosahedral viruses with an inner membrane 3.2 Enveloped icosahedral viruses: Phage ϕ6 and its relatives 3.3 Vesicular pleomorphic viruses 3.4 Prokaryotic viruses with helical symmetry: With or without a membrane 3.5 Lemon-shaped viruses are specific for archaea 3.6 Archaeal spherical viruses with helical NCs have an envelope 4. Conclusions Acknowledgments References

2 3 18 21 24 24 32 35 40 43 46 47 49 49

Abstract Lipid-containing bacteriophages were discovered late and considered to be rare. After further phage isolations and the establishment of the domain Archaea, several new prokaryotic viruses with lipids were observed. Consequently, the presence of lipids in prokaryotic viruses is reasonably common. The wealth of information about how prokaryotic viruses use their lipids comes from a few well-studied model viruses (PM2, PRD1, and ϕ6). These bacteriophages derive their lipid membranes selectively from the host during the virion assembly process which, in the case of PM2 and PRD1, culminates in the formation of protein capsid with an inner membrane, and for ϕ6 an outer envelope. Several inner membrane-containing viruses have been described for archaea, and their lipid acquisition models are reminiscent to those of PM2 and PRD1. Unselective acquisition of lipids has been observed for bacterial mycoplasmaviruses and archaeal pleolipoviruses, which resemble each other by size, morphology, and life style. In addition to these shared morphotypes of bacterial and archaeal viruses, archaea are infected by viruses with unique morphotypes, such as

Advances in Virus Research, Volume 92 ISSN 0065-3527 http://dx.doi.org/10.1016/bs.aivir.2014.11.005

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2015 Elsevier Inc. All rights reserved.

1

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Nina S. Atanasova et al.

lemon-shaped, helical, and globular ones. It appears that structurally related viruses may or may not have a lipid component in the virion, suggesting that the significance of viral lipids might be to provide viruses extended means to interact with the host cell.

1. INTRODUCTION In principal, all cellular organisms are infected by viruses. In its simplest form, an infectious viral particle, the virion, is composed of a genome that is covered with a protective coat. The coat can be built using proteins or proteinaceous lipid membranes. To date, most of the studied prokaryotic viruses (bacteriophages and archaeal viruses) are of head-tailed icosahedral morphology with no lipids (Fig. 1). Among the described bacteriophages, lipid-containing phages are considered to be rare and represent less than 5% of the described isolates (Ackermann & Prangishvili, 2012). Although there is a wide variety of morphotypes in archaeal viruses, the number of known archaeal viruses is a small proportion of the known prokaryotic viruses (Ackermann & Prangishvili, 2012). Yet, many of them contain lipids (Table 1 and Fig. 1). The discovery of the first lipid-containing prokaryotic virus, PM2, occurred only in the late 1960s (Espejo & Canelo, 1968a). Bacteriophages had already been known for several decades, but the late discovery of lipid-containing prokaryotic viruses was partially due to the common protocol of treating phage stock solutions with chloroform in order to avoid bacterial growth (Adams, 1959). Soon after that ϕ6, an enveloped bacteriophage, was isolated (Vidaver et al., 1973) followed by the discovery of PRD1, an icosahedral virus with an inner membrane (Olsen et al., 1974). The first lipid-containing archaeal virus, the helical Thermoproteus tenax virus 1 (TTV1), was reported in the late 1980s (Zillig et al., 1988). The lipid membranes of the prokaryotic viruses usually have significant functions during viral life cycle. Although some generalizations can be made between the virion morphology and the membrane localization in relation to its functions, the type of the genome and its delivery during the infection, for example, may influence the specific roles the membrane plays during the viral life cycle. Although most of the information comes from the model bacteriophages such as PRD1, PM2, and ϕ6, new mass spectrometric techniques developed for lipid research may in the future bring additional information. This is important especially for the research of archaeal viruses due to difficulties in obtaining highly pure viral material.

Prokaryotic Viruses with Lipids

3

Figure 1 Morphotypes of known prokaryotic viruses: (A) bacteriophages and (B) archaeal viruses. The localization of the lipid membrane is indicated by red except for His1 for which the lipid-modified major structural protein is indicated by dashed red line. Numbers of the viruses that contain lipids are underlined. The scale bar represents 100 nm. The virions are in scale except for those marked with an asterisk are 0.5
their actual size. The representative morphotypes are exemplified by: 1. PRD1; 2. PM2, 3. ϕ6; 4. L2; 5. T4; 6. λ; 7. M13; 8. ϕX174; 9. MS2; 10. T7; 11. STIV; 12. PSV; 13. HRPV-1; 14. His1; 15. STSV1; 16. ATV; 17. AFV1; 18. SIRV-1; 19. Aeropyrum pernix bacilliform virus 1 (APBV1); 20. Acidianus bottle-shaped virus (ABV); 21. Sulfolobus neozealandicus droplet-shaped virus (SNDV); 22. Aerupyrum coilshaped virus (ACV); 23. Halorubrum sodomense tailed virus 2 (HSTV-2); 24. Haloarcula vallismortis tailed virus 1 (HVTV-1); and 25. Haloarcula sinaiiensis tailed virus 1 (HSTV-1). The lipid-containing viruses are presented in Table 1. For viruses without lipids, see King, Adams, Carstens, and Lefkowitz (2012) and Pietilä, Demina, Atanasova, Oksanen, and Bamford (2014).

1.1. Origin of lipids in prokaryotic viruses and their detection Since prokaryotic viruses obtain their membranes from the host, the foundations of prokaryotic viral lipids are laid by the host lipid synthesis. In this sense, bacteria and archaea are profoundly different. In general, the core structures of archaeal membrane lipids consist of two isoprenoid side chains that are linked to sn-glycerol-1-phosphate (G1P) backbone through ether bonds (Kates, 1993), whereas in bacteria, the core membrane lipids contain two fatty acid side chains linked to sn-glycerol-3-phosphate (G3P) through ester linkages (Fig. 2; Cronan, 2003; Kates, 1993). The two major core lipids of archaea are archaeol and caldarchaeol (Fig. 2). Archaeol, the

Table 1 Lipid-containing viruses infecting prokaryotes Virus

Isolation host Genome size

Life cycle

Lipids (position, acquisition)

Gram Domain staining References

Virus name

Morphotype

Family

Genome type

PRD1

Icosahedral

Tectiviridae

Linear dsDNA 14,927, ITR (110 bp)

Virulent

Internal membrane, selective

Salmonella typhimurium

B

Gram Olsen, Siak, and Gray (1974), Bamford et al. (1991), Cockburn et al. (2004), and Laurinavicˇius, Ka¨kela¨, Somerharju, and Bamford (2004)

PR3

Icosahedral

Tectiviridae

Linear dsDNA 14,937, ITR (111 bp)

Virulent

Internal membrane, ND

Pseudomonas aeruginosa PAO

B

Gram Stanisich (1974), Bamford, Rouhiainen, Takkinen, and S€ oderlund (1981), and Saren et al. (2005)

PR4

Icosahedral

Tectiviridae

Linear dsDNA 14,943, ITR (111 bp)

Virulent

Internal membrane, ND

P. aeruginosa PAO B

Gram Stanisich (1974), Bamford et al. (1981), and Saren et al. (2005)

PR5

Icosahedral

Tectiviridae

Linear dsDNA 14,939, ITR (110 bp)

Virulent

Internal membrane, selective

Escherichia coli K12 B

Gram Wong and Bryan (1978), Bamford et al. (1981), and Saren et al. (2005)

Strain name

PR722

Icosahedral

Tectiviridae

Linear dsDNA 14,942, ITR (111 bp)

Virulent

Internal membrane, ND

Proteus niirubilis PM5006

B

Gram Coetzee, Lecatsas, Coetzee, and Hedges (1979) and Saren et al. (2005)

L17

Icosahedral

Tectiviridae

Linear dsDNA 14,935, ITR (111 bp)

Virulent

Internal membrane, ND

Aeromonas B hydrophila (RP1) and E. coli W3110 (RP1)

Gram Bamford et al. (1981) and Saren et al. (2005)

P37-14

Icosahedral

Tectiviridae

dsDNA

ND

Internal membrane, ND

Thermus sp.

Gram Yu, Slater, and Ackermann (2006)

Bam35

Icosahedral

Tectiviridae

Linear dsDNA 14,935, ITR (74 bp)

Temperate Internal membrane, selective

Bacillus thuringiensis B

AP50

Icosahedral

Tectiviridae

Linear dsDNA 14,398

Temperate Internal membrane, ND

Bacillus anthracis

20,000

B

B

Gram+ Ackermann, Roy, Martin, Murthy, and Smirnoff (1978), Ravantti, Gaidelyte, Bamford, and Bamford (2003), Str€ omsten, Benson, Burnett, Bamford, and Bamford (2003), and Laurinavicˇius, Ka¨kela¨, Somerharju, and Bamford (2004) Gram+ Nagy, Pragai, and Ivanovics (1976), Nagy and Ivanovics (1977), and Sozhamannan et al. (2008) Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Isolation host Genome size

Gram Domain staining References

Virus name

Morphotype

Family

Genome type

GIL01

Icosahedral

Tectiviridae

Linear dsDNA 14,931, Temperate NDa imperfect ITR

B. thuringiensis serovar Israelensis strain AND508

B

Gram+ Verheust, Jensen, and Mahillon (2003)

GIL16

Icosahedral

Tectiviridae

Linear dsDNA 14,844, Temperate ND imperfect ITR

B. thuringiensis serovar Israelensis strain AND508

B

Gram+ Verheust, Fornelos, and Mahillon (2005)

ϕNS11

Icosahedral

Tectiviridae

Linear dsDNA ND

Temperate Internal Bacillus membrane, acidocaldarius TA6 nonselective

B

Gram+ Sakaki and Oshima (1976), Sakaki, Oshima, Yamada, and Oshima (1977), and Sakaki, Maeda, and Oshima (1979)

Wip1

Icosahedral

Unclassified

Linear dsDNA 14,319

Temperate Internal membrane, ND

B. anthracis

B

Gram+ Schuch, Pelzek, Kan, and Fischetti (2010) and Kan, Fornelos, Schuch, and Fischetti (2013)

P23-77

Icosahedral

Sphaerolipoviridae Circular dsDNA

Virulent

Thermus thermophilus ATCC 33923

B

Gram Yu et al. (2006), Jaatinen, Happonen, Laurinma¨ki, Butcher, and

17,036

Life cycle

Lipids (position, acquisition)

Internal membrane, selective

Strain name

Bamford (2008), Jalasvuori et al. (2009), and Pawlowski, Rissanen, Bamford, Krupovicˇ, and Jalasvuori (2014) KHP30

Icosahedral

Unclassified

Circular dsDNA

26,215

Temperate Internal membrane, ND

Helicobacter pylori NY43

B

Gram Uchiyama et al. (2012) and Uchiyama et al. (2013)

PM2

Icosahedral

Corticoviridae

Circular dsDNA

10,079

Virulent

Internal membrane, selective

Pseudoalteromonas espejiana BAL-31

B

Gram Espejo and Canelo (1968b), CameriniOtero and Franklin (1972), and Ma¨nnist€ o, Kivela¨, Paulin, Bamford, and Bamford (1999)

Salisaeta icosahedral phage 1 (SSIP-1)

Icosahedral

Unclassified

Circular dsDNA

43,788

Virulent

Internal membrane, selective

Salisaeta sp. SP9-1 B

Gram Aalto et al. (2012) and Atanasova, Roine, Oren, Bamford, and Oksanen (2012) Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Genome type

Isolation host Genome size

Life cycle

Lipids (position, acquisition)

Gram Domain staining References

Virus name

Morphotype

Family

SNJ1

Icosahedral

Sphaerolipoviridae Circular dsDNA

16,341

Temperate Internal membrane, selective

Natrinema sp. J7-1 A

–

Zhang et al. (2012) and Pawlowski et al. (2014)

Strain name

Sulfolobus Icosahedral turreted icosahedral virus (STIV)

Turriviridae

Circular dsDNA

17,663

Virulent

Internal membrane, selective

Sulfolobus solfataricus YNPRC179

A

–

Rice et al. (2004) and Maaty et al. (2006)

Sulfolobus Icosahedral turreted icosahedral virus 2 (STIV2)

Turriviridae

Circular dsDNA

16,622

Virulent

Internal membrane, ND

S. solfataricus 2-212

A

–

Happonen et al. (2010) and Happonen et al. (2013)

SH1

Icosahedral

Sphaerolipoviridae Linear dsDNA 30,898, ITR (309 bp)

Virulent

Internal membrane, selective

Haloarcula hispanica A ATCC 33960

–

Bamford et al. (2005), Porter et al. (2005), Kivela¨ et al. (2006), and Pawlowski et al. (2014)

H. hispanica Icosahedral icosahedral virus 2 (HHIV-2)

Sphaerolipoviridae Linear dsDNA 30,578, ITR (305 bp)

Virulent

Internal membrane, NDa,b

H. hispanica ATCC A 33960

–

Atanasova et al. (2012), Jaakkola et al. (2012), and Pawlowski et al. (2014)

PH1

Icosahedral

Sphaerolipoviridae Linear dsDNA 28,064, ITR (337 bp)

φ6

Spherical

Cystoviridae

Linear dsRNA

φ7

Spherical

Cystoviridae

φ8

Spherical

φ9 φ10

Virulent

Internal membrane, NDa,b

H. hispanica ATCC A 33960

–

Porter et al. (2013) and Pawlowski et al. (2014)

14,927 Virulent (in 3 (able to segments) form carrier state cells)

Outer membrane, selective

Pseudomonas syringae pv. phaseolicola HB1OY

B

Gram Vidaver, Koski, and Van Etten (1973), McGraw, Mindich, and Frangione (1986), Gottlieb et al. (1988), Mindich et al. (1988), and Laurinavicˇius, Ka¨kela¨, Bamford, and Somerharju (2004)

Linear dsRNA

13.4 Virulent (in 3 segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999)

Cystoviridae

Linear dsRNA

15 (in 3 Virulent segments)

Outer membrane, NDa

Pseudomonas pseudoalcaligenes ERA

B

Gram Mindich et al. (1999) and Sun, Qiao, Qiao, Onodera, and Mindich (2003)

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999)

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999) Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Isolation host Lipids (position, acquisition)

Gram Domain staining References

Virus name

Morphotype

Family

Genome type

Genome size

φ11

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999)

φ12

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. pseudoalcaligenes B ERA

Gram Mindich et al. (1999)

φ13

Spherical

Cystoviridae

Linear dsRNA

13.7 Virulent (in 3 (able to segments) form carrier state cells)

NDa

P. pseudoalcaligenes B ERA

Gram Mindich et al. (1999)

φ14

Spherical

Cystoviridae

Linear dsRNA

14 (in 3 Virulent segments)

NDa

P. syringae pv. phaseolicola

B

Gram Mindich et al. (1999)

φ2954

Spherical

Cystoviridae

Linear dsRNA

12,685 Virulent (in 3 segments)

NDa

P. syringae LM2489

B

Gram Qiao, Sun, Qiao, Di Sanzo, and Mindich (2010)

L2

Pleomorphic Plasmaviridae

Circular dsDNA

11,965

Persistent

Lipid envelope, ND

Acholeplasma laidlawii

B

Gram Gourlay (1971), Putzrath and Maniloff (1977), and Maniloff, Kampo, and Dascher (1994)

L172

Globular

Circular ssDNA

14,000

Persistent

Lipid envelope, ND

A. laidlawii

B

Gram Dybvig, Nowak, Sladek, and Maniloff (1985)

Unclassified

Life cycle

Strain name

Halorubrum pleomorphic virus 1 (HRPV-1)

Pleomorphic Pleolipoviridae

Circular ssDNA

7048

Persistent

Lipid Halorubrum sp. PV6 envelope, nonselective

A

–

Pietila¨, Roine, Paulin, Kalkkinen, and Bamford (2009) and Pietila¨, Laurinavicˇius, Sund, Roine, and Bamford (2010)

H. hispanica pleomorphic virus 1 (HHPV-1)

Pleomorphic Pleolipoviridae

Circular dsDNA

8082

Persistent

Lipid H. hispanica ATCC A envelope, 33960 nonselective

–

Roine et al. (2010)

Halorubrum pleomorphic virus 3 (HRPV-3)

Pleomorphic Pleolipoviridae

Circular discontinuous dsDNA

8770

Persistent

Lipid Halorubrum sp. envelope, SP3-3 nonselective

A

–

Atanasova et al. (2012), Pietila¨ et al. (2012), and Sencˇilo, Paulin, Kellner, Helm, and Roine (2012)

Halorubrum pleomorphic virus 2 (HRPV-2)

Pleomorphic Pleolipoviridae

Circular ssDNA

10,656

Persistent

Lipid Halorubrum sp. envelope, SS5-4 nonselective

A

–

Atanasova et al. (2012), Pietila¨ et al. (2012), and Sencˇilo et al. (2012)

Halorubrum pleomorphic virus 6 (HRPV-6)

Pleomorphic Pleolipoviridae

Circular ssDNA

8549

Persistent

Lipid Halorubrum sp. envelope, SS7-4 nonselective

A

–

Pietila¨ et al. (2012) and Sencˇilo et al. (2012) Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Genome type

Gram Domain staining References

Genome size

Life cycle

9694

Persistent

Lipid Halogeometricum sp. A envelope, CG-9 nonselective

–

Atanasova et al. (2012), Pietila¨ et al. (2012), and Sencˇilo et al. (2012)

Persistent

Lipid H. hispanica ATCC A envelope, 33960 nonselective

–

Bath, Cukalac, Porter, and DyallSmith (2006) and Pietila¨ et al. (2012)

A

–

Janekovic et al. (1983), Zillig et al. (1988), and Neumann, Schwass, Eckerskorn, and Zillig (1989)

T. tenax Kra1

A

–

Janekovic et al. (1983)

Outer membrane, ND

T. tenax Kra1

A

–

Janekovic et al. (1983)

Outer membrane, selectived

Sulfolobus isolate HVE11/2

A

–

Arnold et al. (2000) and Peng et al. (2001)

Strain name

Virus name

Morphotype

Halogeometricum pleomorphic virus 1 (HGPV-1)

Pleomorphic Pleolipoviridae

Circular discontinuous dsDNA

His virus 2 (His2)

Pleomorphic Pleolipoviridae

Linear dsDNA 16,067, ITR (525 bp)

Thermoproteus tenax virus 1 (TTV1)

Filamentous Lipothrixviridae

Temperate Outer Linear dsDNA 13,669e (partial membrane, sequence) selective

T. tenax Kra1

T. tenax virus 2 Filamentous Lipothrixviridae (TTV2)

Linear dsDNA 16,000

Temperate Outer membrane, ND

T. tenax virus 3 Filamentous Lipothrixviridae (TTV3)

Linear dsDNA 27,000

ND

Filamentous Lipothrixviridae

Linear dsDNA 40,900e, ITR (at least 800 bp)

Persistent

Sulfolobus islandicus filamentous virus 1 (SIFV)

Family

Isolation host Lipids (position, acquisition)

Desulforolobus ambivalens filamentous virus (DAFV)

Filamentous Lipothrixviridae

Linear dsDNA 56,000

ND

Outer membrane, ND

Desulforolobus ambivalens

A

–

Zillig et al. (1994)

Acidianus filamentous virus 1 (AFV1)

Filamentous Lipothrixviridae

Linear dsDNA 20,869, ITR (11 bp)

Persistent

Outer membrane, selectived

Acidianus hospitalis YS6

A

–

Bettstetter, Peng, Garrett, and Prangishvili (2003)

Acidianus filamentous virus 2 (AFV2)

Filamentous Lipothrixviridae

Linear dsDNA 31,787e

Persistent

No lipids detected

Acidianus sp. strain A F28

–

Ha¨ring et al. (2005)

Acidianus filamentous virus 3 (AFV3)

Filamentous Lipothrixviridae

Linear dsDNA 40,449e, ITR

Persistent

Outer Acidianus sp. strain A membrane, Acii25 nonselective

–

Vestergaard et al. (2008)

Acidianus filamentous virus 6 (AFV6)

Filamentous Lipothrixviridae

Linear dsDNA 39,577e, ITR

Persistent

ND

Acidianus convivator A

–

Vestergaard et al. (2008)

Acidianus filamentous virus 7 (AFV7)

Filamentous Lipothrixviridae

Linear dsDNA 36,895e, ITR

Persistent

ND

A. convivator

A

–

Vestergaard et al. (2008)

Acidianus filamentous virus 8 (AFV8)

Filamentous Lipothrixviridae

Linear dsDNA 38,179e, ITR

Persistent

ND

A. convivator

A

–

Vestergaard et al. (2008)

Acidianus filamentous virus 9 (AFV9)

Filamentous Lipothrixviridae

Linear dsDNA 41,172, ITR (384 bp)

Persistent

Outer membrane, ND

Acidianus uzoniensis A

–

Bize et al. (2008)

Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Isolation host Genome size

Life cycle

Lipids (position, acquisition)

Gram Domain staining References

Virus name

Morphotype

Family

Genome type

His virus 1 (His1)

Spindle shaped

Fuselloviridae

Linear dsDNA 14,464, ITR (105 bp)

Persistent

No membrane, MCP is lipid modified

H. hispanica ATCC A 33960

–

Bath and DyallSmith (1998), Bath et al. (2006), and Pietila¨, Atanasova, Oksanen, and Bamford (2013)

Sulfolobus spindle-shaped virus 1 (SSV1)

Spindle shaped

Fuselloviridae

Circular dsDNA

15,465

Persistent

NDa

Sulfolobus shibatae B12

A

–

Reiter, Zillig, and Palm (1988), Palm et al. (1991), and Schleper, Kubo, and Zillig (1992)

Sulfolobus spindle-shaped virus 2 (SSV2)

Spindle shaped

Fuselloviridae

Circular dsDNA

14,796

ND

NDc

S. solfataricus P1

A

–

Stedman et al. (2003)

Sulfolobus spindle-shaped virus 4 (SSV4)

Spindle shaped

Fuselloviridae

Circular dsDNA

15,135

ND

NDc

S. islandicus ARN3/6

A

–

Peng (2008)

Sulfolobus spindle-shaped virus 5 (SSV5)

Spindle shaped

Fuselloviridae

Circular dsDNA

15,330

ND

ND

S. solfataricus P2

A

–

Redder et al. (2009)

Sulfolobus spindle-shaped virus 6 (SSV6)

Spindle shaped

Fuselloviridae

Circular dsDNA

15,684

ND

ND

S. islandicus G4ST- A T-11

–

Redder et al. (2009)

Strain name

Sulfolobus spindle-shaped virus 7 (SSV7)

Spindle shaped

Fuselloviridae

Circular dsDNA

17,602

ND

ND

S. islandicus G4T-1 A

–

Redder et al. (2009)

Sulfolobus spindle-shaped virus Ragged hills (SSVrh)

Spindle shaped

Fuselloviridae

Circular dsDNA

16,473

ND

ND

S. solfataricus

A

–

Wiedenheft et al. (2004)

Sulfolobus spindle-shaped virus Kamchatka1 (SSVk1)

Spindle shaped

Fuselloviridae

Circular dsDNA

17,384

ND

ND

S. solfataricus

A

–

Wiedenheft et al. (2004)

Acidianus spindle-shaped virus 1 (ASV1)

Spindle shaped

Fuselloviridae

Circular dsDNA

24,186

ND

ND

Acidianus brierleyi DSM1651

A

–

Redder et al. (2009)

Pyrococcus abyssi virus 1 (PAV1)

Spindle shaped

Unclassified

Circular dsDNA

18,098

Persistent

NDa

P. abyssi GE23

A

–

Geslin et al. (2003) and Geslin et al. (2007)

Thermococcus prieurii virus 1 (TPV1)

Spindle shaped

Unclassified

Circular dsDNA

21,591

Persistent

NDa

T. prieurii

A

–

Gorlas, Koonin, Bienvenu, Prieur, and Geslin (2012)

Sulfolobus Spindle tengchongensis shaped spindle-shaped virus 1 (STSV1)

Unclassified

Circular dsDNA

75,294

Persistent

Outer membrane, selective

S. tengchongensis RT8-4

A

–

Xiang et al. (2005)

Continued

Table 1 Lipid-containing viruses infecting prokaryotes—cont'd Virus

Virus name

Lipids (position, acquisition)

Gram Domain staining References

Genome size

Life cycle

76,107

Persistent

Outer membrane, selectived

S. tengchongensis HB52

A

–

Erdmann et al. (2014)

Family

Genome type

S. tengchongensis Spindle spindle-shaped shaped virus 2 (STSV2)

Unclassified

Circular dsDNA

Pyrobaculum Spherical spherical virus 1 (PSV1)

Globuloviridae

Linear dsDNA 28,337, ITR (190 bp)

Persistent

Outer membrane, selectived

Pyrobaculum sp. D11

A

–

Ha¨ring et al. (2004)

T. tenax Spherical spherical virus 1 (TTSV1)

Globuloviridae

Linear dsDNA 20,933 and 700 bp terminie

Persistent

NDc

T. tenax YS44

A

–

Ahn et al. (2006)

a

Morphotype

Isolation host

Strain name

The presence of the lipids was assumed based on the sensitivity to organic solvents. The presence of the lipids was assumed based on the Sudan Black staining. The presence of the lipids was assumed based on the virus particle density. d The lipids are modified as determined by the thin-layer chromatography of the viral and host lipids. The viral lipids had different mobilities compared to the hosts’. e The nature of the genome termini is unclear. A, archaea; B, bacteria; ITR, inverted terminal repeats; ND, not determined. b c

Prokaryotic Viruses with Lipids

17

Figure 2 The basic structures of core lipids found in bacterial and archaeal membranes. Diacylglycerol is the backbone of bacterial membrane lipids, whereas archaeol and caldarchaeol are found in haloarchaeal or crenarchaeal membrane lipids, respectively.

diphytanylglycerol diether, and its variants can form a membrane bilayer and are the major constituents of the halophilic archaeal membranes. Caldarchaeol is one of the major components in the thermophilic crenarchaeal membranes (such as Sulfolobus spp). It is a dibiphytanyldiglycerol tetraether, an antiparallel arrangement of two glycerol units connected with two isoprenoid acyl chains through four ether linkages (Fig. 2). These lipids form monolayer membranes (Kates, 1993). The major components of bacterial and archaeal membranes are mostly polar phospholipids. In halophilic archaea, these include phosphatidylglycerol (PG), the methyl ester of phosphatidylglycerol (PGP-Me), and phosphatidylglycerosulfate (PGS). In extreme haloarchaea, the PGP-Me contributes approximately 50–80 mol% of the polar lipids (Tenchov, Vescio, Sprott, Zeidel, & Mathai, 2006). In addition, many different types of archaeal glycolipids and species of cardiolipins (CLs) are also components of the membranes. Neutral lipids constitute approximately 10% of haloarchaeal lipids (Corcelli & Lobasso, 2006). The cytoplasmic membrane of Gram-negative bacteria consists of 75% phosphatidylethanolamine (PE), 20% PG, and 5% CL (Cronan, 2003). However, environmental conditions have been reported to influence considerably the lipid composition of bacterial and archaeal membranes (Farrell & Rose, 1967; Lopalco et al., 2013). The fact that prokaryotic viruses obtain their lipids from host membranes can also cause problems in the identification of viral lipids. This is especially true for the enveloped viruses due to the often unselective manner of

18

Nina S. Atanasova et al.

membrane acquisition. Highly purified material is required for the characterization of viral lipids, because it is often difficult to separate virions from host membrane vesicles. This can, in some cases, be overcome by the analyses of membrane-containing subviral particles that have been obtained, for example, by quantitative biochemical dissociation of the virion (Vitale, Roine, Bamford, & Corcelli, 2013). The first indication of the presence of lipids in the virion is the sensitivity to organic solvents such as chloroform or the low buoyant density of the virion. Sudan Black staining of the highly purified virions analyzed in sodium dodecyl sulfate–polyacrylamide gel electrophoresis has also been used as the preliminary indication of membrane lipids or lipid-modified proteins (Pietila¨ et al., 2012; Prat, Lamy, & Weill, 1969). Further analyses of viral lipids have traditionally been conducted using organic solvent extraction followed by thin-layer chromatography (TLC), mass spectrometry (MS; e.g., electrospray ionization, ESI-MS), and/or nuclear magnetic resonance spectroscopy (NMR; Corcelli & Lobasso, 2006; London & Feigenson, 1979). Recently, a new method based on the direct detection of viral lipids by matrix-assisted laser desorption/ionization–time-offlight/mass spectrometer analysis using 9-aminoacridine as the matrix has been developed (Murphy & Gaskell, 2011; Vitale et al., 2013). This method avoids several drawbacks encountered in the traditional methods such as quantitative and systematic biases in the solvent extraction. The amount of viral material required for the analysis is miniscule compared to the traditional methods, and reliable detection of the minor components of viral lipids is possible (Vitale et al., 2013).

2. FUNCTION AND SIGNIFICANCE OF LIPIDS IN PROKARYOTIC VIRUS LIFE CYCLE Lipids have essential roles in different stages of the viral life cycle (entry, assembly, and exit) if the virus contains lipids as structural components of the virion. Although the exact mechanisms of bacteriophage lipid acquisition still require further research, the role of lipids in the life cycle is well established for PM2, PRD1, and ϕ6, the type species of the bestcharacterized families of lipid-containing bacteriophages, Corticoviridae, Tectiviridae (Section 3.1; Fig. 3), and Cystoviridae (Section 3.2; Fig. 3), respectively (Oksanen, Poranen, & Bamford, 2010). Compared to bacteria, archaea seem to have relatively more lipid-containing viruses, although to date, only approximately 100 archaeal viruses are known (Atanasova

Prokaryotic Viruses with Lipids

19

Figure 3 Sequential steps of the utilization of the viral membranes during the entry and assembly of dsDNA bacteriophage PRD1 with an internal membrane (A–G) and dsRNA phage ϕ6 with an external one (H–O). The host outer membrane (OM), peptidoglycan layer (PG), and cytoplasma membrane (CM) are colored with yellow, gray, and orange, respectively. The phage-specific membranes are in red. (A) PRD1 recognizes the cell envelope-associated DNA transfer complex via the receptor recognition protein P2 at the virion vertices. (B) After the binding, the particle is reorientated on the cell surface triggering the release of the vertex complexes (proteins P2, P5, P31, P16, and peripentonal P3) leading to the transformation of the inner membrane to a membraneous tail tube, which penetrates the cell envelope. The membrane-associated lytic transglycosylase protein P7 digests locally an opening to the peptidoglycan layer. The linear dsDNA genome with terminal proteins is transferred to the cytoplasm through the tail tube. Other proteins involved in the cell envelope penetration are at least P11, P14, P18, and P32. (C) Upon viral protein translation, the membrane-associated phage proteins are addressed to the host cytoplasmic membrane. (D) During the virion assembly, the phage-specific membrane batch at the CM is (Continued)

20

Nina S. Atanasova et al.

et al., 2012; Pietila¨ et al., 2014; Pina, Bize, Forterre, & Prangishvili, 2011). Consequently, detailed information about archaeal virus lipids is scarce, but represents an interesting field of research, as archaeal lipids are structurally very different from those of bacteria and eukaryotes (Section 1.1; Roine & Bamford, 2012; Sprott, 2011). In general, prokaryotic lipidcontaining viruses can be classified into three categories: those that contain an external or internal lipid bilayer, and those that have lipids as protein modifications. Entry is the first step in the virus life cycle in which viral membranes have an essential role. Viruses with an external membrane enter the cell by fusion of the viral membrane with that of the host (Cann, 2005). In the case of dsRNA bacteriophages with segmented genomes and an external membrane, the host membrane participating in the fusion is the outer membrane of the Gram-negative host bacterium (Poranen & Bamford, 2008). Membrane fusion with the cytoplasmic membrane of the host has been suggested for the pleomorphic mycoplasmaviruses (Section 3.3) as mycoplasmas lack the cell wall and the outer membrane (Putzrath & Maniloff, 1977). The entry mechanism is not known for any archaeal virus, although at least pleolipoviruses (Section 3.3) most probably use by membrane fusion (Pietila¨ et al., 2009; Roine & Oksanen, 2011). Some of the inner Figure 3—Cont'd pinched off with the help of phage-encoded nonstructural membrane-bound scaffolding protein P10. The particle assembly is also assisted by the host GroEL–GroES chaperonin complex and phage-encoded soluble assembly factors P17 and P33. (E) The virus capsid-associated proteins are added on the surface of the internal phage membrane enabling the formation of the PRD1 procapsid. (F) The phage genome is packaged by the packaging ATPase P9 through the membrane-bound unique vertex. (G) Mature virions are released upon host cell lysis. (H) The receptorbinding spike protein P3 of phage ϕ6 binds to the pilus receptor that retracts bringing the virion to face the outer membrane. (I) Translocation of P3 exposes the fusogenic protein P6 leading to the fusion of the viral membrane with the host outer membrane. Nucleocapsid (NC) surface protein P5 degrades a local opening to the peptidoglycan layer allowing the virus particle to face the cytoplasmic membrane of the host. (J) The NC surface protein P8 aids the acquisition of the cytoplasma membrane vesicle around the NC releasing the particle to the cell interior. (K) The internal polymerase complex (PC) particle is released from the membrane vesicle and starts transcription. (L) The ϕ6 particle assembly starts when the virus-specific membrane proteins are inserted to the cytoplasma membrane excluding locally the host proteins. (M) The NC associates with the virusspecific membrane, and the particle is release to the host cytoplasm with the aid of nonstructural assembly factor protein P12. (N) The P3 spike proteins are added to the virion surface anchored to protein P6 residing in the viral membrane. (O) Mature virions are released by cell lysis and ready to initiate the next life cycle.

Prokaryotic Viruses with Lipids

21

membrane-containing bacteriophages with linear dsDNA genomes use their membrane to form a tubular structure which serves as a DNA injection apparatus (Bamford & Mindich, 1982; Peralta et al., 2013). This has been well documented for the phages PRD1 and Bam35 which infect Gramnegative and Gram-positive hosts, respectively (Gaidelyte, CvirkaiteKrupovicˇ, Daugelavicˇius, Bamford, & Bamford, 2006; Peralta et al., 2013). Entry mechanisms of prokaryotic viruses with circular genomes are mostly unknown. However, it is considered that the entry of PM2, which has a circular supercoiled dsDNA genome, involves the fusion of the viral membrane with the host outer membrane (Cvirkaite-Krupovicˇ, Krupovicˇ, Daugelavicˇius, & Bamford, 2010; Kivela¨, Daugelavicˇius, Hankkio, Bamford, & Bamford, 2004). Inner membrane-containing viruses with either linear or circular genomes are known to infect archaea, but since the archaeal cell wall structure differs significantly from the bacterial one (Kandler & Konig, 1998), it remains to be seen how archaeal inner membrane-containing viruses enter the cell. Following entry and replication, lipids are incorporated into the new virions by yet mostly unrevealed ways during a complex assembly process or during egress if the virus exits the cell without lysis. This is introduced briefly below followed by more detailed descriptions about the different lipid-containing prokaryotic viruses.

2.1. How prokaryotic viruses acquire their lipids Prokaryotic viruses have no genes for lipid synthesis of their membranes. Thus, lipids are obtained from the host cytoplasmic membranes. This acquisition can result into a somewhat direct copy of the host lipid composition or be more or less selective. Although this selectivity depends on the particular virus, the localization of the lipid bilayer in the virion architecture might give insights into the mode of lipid acquisition. 2.1.1 Viruses with an external membrane Most of the known viruses with an external lipid bilayer (envelope) acquire their lipids as they bud out from the host cell. Thus, their lipid composition greatly resembles that of the host (Kuhn & Rossmann, 2005). Budding is a complex process, which is unique for different viruses, but always includes interactions between the viral integral membrane proteins and the host lipid bilayer proteins (Garoff, Hewson, & Opstelten, 1998). The maturation and release of this type of viruses is often a continuous process, and the viruses are

22

Nina S. Atanasova et al.

able to infect their host cells persistently (Cann, 2005). Among the known prokaryotic viruses, budding is suggested as the mode of exit for pleomorphic archaeal and bacterial viruses (pleolipoviruses, plasmaviruses, and the mycoplasma phage L172; Section 3.3; Dybvig et al., 1985; Pietila¨ et al., 2009; Putzrath & Maniloff, 1977). Because no established/confirmed models of prokaryotic virus budding are available at the moment, practically all the current knowledge considering this exit mechanism relies on eukaryotic virus research and is not discussed here. Another type of prokaryotic virus architecture involving an external lipid membrane is portrayed by ϕ6 and other viruses in the family Cystoviridae (Table 1; Section 3.2; Sarin et al., 2012). This virus morphology is characterized by an enveloped icosahedral nucleocapsid (NC) enclosing the dsRNA genome. So far, such viruses (or any viruses with an RNA genome) have not been observed for archaea. ϕ6-like viruses acquire their lipids after the assembly of the NC. The lipids forming the envelope are selected from the host cytoplasmic membrane (see Section 3.2 for details). 2.1.2 Viruses with a membrane underneath the icosahedral capsid Viruses with a membrane inside the protein capsid are common in the environment. Such bacteriophages infecting Gram-positive bacteria have temperate life styles, while the phages of the Gram-negative bacterial hosts are virulent (Oksanen & Bamford, 2012a). Archaeal inner membranecontaining viruses infect hosts belonging to the two major phyla Euryarchaeota or Crenarchaeaota. Both temperate and virulent life styles have been observed for these viruses (Pina et al., 2011). Virus life cycle has been studied in detail for PRD1. The membrane is derived from the host cytoplasmic membrane in a process where the major capsid protein (MCP) acquires the virus-specific lipid vesicle from the plasma membrane (Mindich, Bamford, McGraw, & Mackenzie, 1982; Rydman, Bamford, & Bamford, 2001). The virus has a unique vertex which is a conduit for the genome translocation into the empty procapsid (Str€ omsten, Bamford, & Bamford, 2003). Viruses are released by lysis of the host cell. The entry and lipid acquisition of PRD1-like viruses infecting Gram-positive bacteria are suggested to occur in a fashion similar to PRD1 (Ravantti et al., 2003). Lipid acquisition is thought to follow similar pathways also for the described archaeal inner membrane-containing viruses (Bamford et al., 2005; Brumfield et al., 2009; Maaty et al., 2006). The lipids of Sulfolobus turreted icosahedral virus (STIV) have been studied in detail and

Prokaryotic Viruses with Lipids

23

are known to be acidic species of crenarchaeal isoprenoid glycerol dialkyl tetraether lipids that are selectively acquired from the host (Maaty et al., 2006). Interestingly, the release of STIV, which infects a crenarchaeal host, involves the formation of the seven-sided pyramidal protrusions into the host cytoplasmic membrane causing cellular lysis (Brumfield et al., 2009). The lipid acquisition of inner membrane-containing viruses is usually selective, which might be due to features such as membrane curvature induced by the icosahedral capsid and/or specific lipid–protein interactions (see Section 3.1 for details; Bamford et al., 2005; Laurinavicˇius, Bamford, & Somerharju, 2007; Laurinma¨ki, Huiskonen, Bamford, & Butcher, 2005). 2.1.3 Viruses with lipids as structural protein modifications In some cases, lipids can be found in viruses only as structural protein modifications with no bilayer structure. Such viruses are rare, and these modifications are most probably incorporated into the viral proteins by host enzymes. These lipid modifications are considered to enable protein– protein interactions facilitating virion assembly (Hruby & Franke, 1993). The most commonly observed lipid modifications in the proteins of animal viruses are myristoylation and palmitoylation, which involve the co- or posttranslational additions of myristate or palmitate fatty acids to the target proteins (Hruby & Franke, 1993). Poliovirus is an example of a eukaryotic nonenveloped icosahedral virus, which contains lipids only as myristoylation of the internal capsid protein (Paul, Schultz, Pincus, Oroszlan, & Wimmer, 1987). In animal viruses, myristoylation has been suggested to be essential in the encapsidation of the viral genome and/or in assembly and is thought to be catalyzed by the host N-myristoyltransferase (Moscufo, Simons, & Chow, 1991). To our knowledge, virions that contain lipids only as modifications of structural proteins have not previously been described for bacteriophages. Archaeal lemon-shaped virus His1 is the only known prokaryotic virus, which lacks a bilayer but contains lipids as modification of the single major structural protein (Fig. 1; Section 3.5; Pietila¨ et al., 2013). However, the nature of the lipid modification is currently unknown (Pietila¨ et al., 2013). It is possible that the crenarchaeal fuselloviruses have similar lipid modifications and lack a bilayer, but it remains to be determined. In addition, lipid-modified spike proteins have been observed for two pleolipoviruses, HGPV-1 and His2, in which the virion is a membrane vesicle (Pietila¨ et al., 2012).

24

Nina S. Atanasova et al.

3. CURRENTLY KNOWN LIPID-CONTAINING BACTERIAL AND ARCHAEAL VIRUSES To date, the number of described bacteriophage isolates is above 6000, whereas there are only some 100 described archaeal viruses (Ackermann & Prangishvili, 2012; Pietila¨ et al., 2014). In spite of this, the observed diversity of virion morphotypes is higher for archaea predicting that novel viral architectures will be discovered in the future. Surprisingly, when taking into account the overall estimated abundance of viruses in the biosphere (above 1031; Suttle, 2005), we have only observed half-a-dozen morphotypes for bacteriophages and about a dozen for the archaeal ones (Fig. 1). The reasonably late discovery of lipids particularly in archaeal viruses has brought surprises and points toward additional diversity. In this chapter, we summarize the information on the lipids in bacterial and archaeal viruses.

3.1. Icosahedral viruses with an inner membrane Before the year 1968, there were no reports on bacteriophages having lipids as their structural component, and archaea were not even recognized as a distinct domain (Woese & Fox, 1977). Bacteriophage PM2 was the first prokaryotic virus for which lipids were observed as structural components of the virion (Camerini-Otero & Franklin, 1972; Espejo & Canelo, 1968b). These phage-infecting Pseudoalteromonas cells were fished out from sea water samples taken from the Pacific Ocean about 1 mile from Vina del Mar, Chile (Espejo & Canelo, 1968a). PM2 is an icosahedral virus with an internal lipid o, Kalkkinen, & Bamford, 1999). bilayer (Kivela¨ et al., 2004; Kivela¨, Ma¨nnist€ Later architecturally similar bacteriophages have been described. To date, it is known that archaeal cells are also infected by viruses that are structurally related to PM2. Currently, there are 23 known icosahedral viruses with an internal membrane of which 17 and 6 infect bacteria and archaea, respectively (Table 1). A common feature of these viruses is that their lipid composition differs from that of their host cell cytoplasmic membrane. All these viruses have a dsDNA genome, which is either linear or circular. The majority of inner membrane-containing viruses with a bacterial host has been classified into the family Tectiviridae, with PRD1 as the type species (Oksanen & Bamford, 2012a). These viruses infect either Gram-negative or Gram-positive bacteria (Table 1). In addition to bacteriophage PM2, which is still the only recognized member of the Corticoviridae family, a thermophilic bacteriophage P23-77 with similar morphology to PRD1 has been

Prokaryotic Viruses with Lipids

25

described for a Thermus host ( Jaatinen et al., 2008; Rissanen et al., 2012). Recently, a highly halophilic bacteriophage, Salisaeta icosahedral phage 1 (SSIP-1), was found from an experimental pond in Israel (unassigned family; Aalto et al., 2012). All these phages are virulent, except those infecting Gram-positive bacteria. The first described archaeal icosahedral virus with an internal membrane was STIV, isolated from an acidic high-temperature hot spring (Rice et al., 2004). A few years later, another crenarchaeal virus, STIV-2 was described (Happonen et al., 2010). STIV-2 is related to STIV and also infects Sulfolobus cells. To date, four viruses, SH1, HHIV-2, PH1, and SNJ1, similar to STIV and STIV-2 are known to infect euryarchaeal hosts ( Jaakkola et al., 2012; Porter et al., 2005, 2013; Zhang et al., 2012). These viruses infect either halophilic Haloarcula or Natrinema strains (Table 1). All the archaeal icosahedral viruses with an internal membrane are virulent, except SNJ1, which is temperate. The classification of these viruses is under discussion (Pawlowski et al., 2014). There is atomic-level information about two inner membranecontaining prokaryotic viruses. The best described is PRD1, isolated 40 years ago from sewage sample taken from Kalamazoo, MI, USA (Olsen et al., 1974). In addition to this, studies on bacteriophage PM2 have provided insights into the assembly of icosahedral viruses with an internal membrane. PRD1 and PM2 virus structures have been solved by X-ray ˚ resolution, respectively, allowing the detailed crystallography at 4 and 7 A analysis of the membranes and the membrane proteins (Abrescia et al., 2004, 2008; Cockburn et al., 2004). The difference in the genome type (linear vs. circular) dictates that the two viruses are bound to have different mechanisms for genome delivery, virus assembly, and genome packaging.

3.1.1 Lipids in the corticovirus PM2 with a circular supercoiled dsDNA genome PM2 is a virulent virus infecting two Gram-negative Pseudoalteromonas species (Kivela¨ et al., 1999; Oksanen & Bamford, 2012b). A significant portion of the available aquatic bacterial genomes contains PM2-like elements indicating that corticoviruses are wide-spread in marine environments (Krupovicˇ & Bamford, 2007). The genome is a highly negatively supercoiled, circular dsDNA molecule of 10,079 bp in length (Espejo, Canelo, & Sinsheimer, 1969; Ma¨nnist€ o et al., 1999). Other icosahedral viruses with an internal membrane and a circular dsDNA genome are STIV,

26

Nina S. Atanasova et al.

STIV-2, SNJ1, P23-77, and SSIP-1 (Table 1). Obviously, unique entry and assembly mechanisms for delivery and packaging of circular molecules have evolved. The viral membrane is most probably involved also in these processes which are rather poorly understood. The diameter of PM2 virion is 57 nm, and its mass is 47 MDa, which is divided to lipid (14%), nucleic acid (14%), and protein (72%; Franklin, Hinnen, Scha¨fer, & Tsukagoshi, 1976). The studies on PM2 virion structure have revealed that the membrane is well ordered following the icosahedral shape of the capsid (Abrescia et al., 2008; Huiskonen, Kivela¨, Bamford, & ˚ , which is Butcher, 2004). The thickness of the PM2 membrane is 29 A ˚ 3 A less than that of PRD1. This might be a consequence of the higher density of ordered transmembrane helices in PM2 (Abrescia et al., 2008; Cockburn et al., 2004). Most of the PM2 structural proteins are associated with the viral membrane (Kivela¨ et al., 2004). Only two proteins, the receptor-binding protein P1 and the MCP P2 form the capsid shell (Abrescia et al., 2008; Huiskonen et al., 2004; Kivela¨ et al., 2004). The other eight structural protein species (P3–P10) occupy about one-third of the viral membrane volume (Kivela¨, Kalkkinen, & Bamford, 2002; Laurinavicˇius et al., 2007; Scha¨fer, Hinnen, & Franklin, 1974). The virion lipid composition (64% PG, 36% PE, and traces of acyl-PG) deviates drastically from that of the host’s cytoplasmic membrane (25% PG, 75% PE; Laurinavicˇius et al., 2007). In addition, the different species of phospholipids are asymmetrically distributed in the membrane leaflets (Laurinavicˇius et al., 2007). Two transmembrane proteins P3 and P6, whose copy numbers are the highest among the PM2 membrane-associated proteins, form a well ordered, planar protein complex lying tangentially to the membrane under the capsid shell (Abrescia et al., 2008). This lattice of these membrane proteins mediates the interaction between the membrane and the icosahedral capsid shell. Similar to all membrane-containing viruses, also PM2 uses its membrane in genome delivery, but in a different way than PRD1 (Section 3.1.4), as the internal membrane of PM2 does not transform into a tail tube. After PM2 has recognized the cell surface receptor via the distal tip of the receptorbinding protein P1, the viral protein shell seemingly dissociates exposing the inner membrane surface (Abrescia et al., 2008; Kivela¨ et al., 2004). It has been proposed that the internal membrane mediates the translocation of the supercoiled genome across the host cell envelope via fusion of the viral membrane with host outer membrane (Kivela¨ et al., 2004). During this entry step, the integral membrane protein P10 most probably interacts with the

Prokaryotic Viruses with Lipids

27

host membrane (Kivela¨, Madonna, Krupovicˇ, Tutino, & Bamford, 2008). The genome penetration through the cytoplasmic membrane is dependent on calcium ions (Cvirkaite-Krupovicˇ, Krupovicˇ, et al., 2010). How is the supercoiled DNA incorporated into the membrane vesicle covered by the protein capsid? It is known that PM2 maturation does not proceed through empty membrane-containing procapsids as is the case for PRD1 (Section 3.1.5). It is established that PM2 DNA is replicated via the rolling circle mechanism in proximity to the host cytoplasmic membrane. However, it is not known, whether the replication is coupled to the o et al., 1999). The majority of DNA encapsidation (Brewer, 1978; Ma¨nnist€ the synthesized virus-specific proteins are associated with the host cell membrane (Datta, Braunstein, & Franklin, 1971). Results from transmission electron microscopy (TEM) experiments have proposed that during PM2 infection the maturation might occur via virus-sized empty vesicles (Dahlberg & Franklin, 1970). Empty vesicles have also been seen in cells infected with a temperature-sensitive PM2 mutant, in which the shift to permissive temperature induces the virion maturation process (Brewer, 1978, 1980) suggesting that empty vesicles are packaged. However, the virion architecture and the interactions seen in the virion X-ray structure (Abrescia et al., 2008) illustrate possible events leading to PM2 virion formation. The membrane proteins might act together with the condensed supercoiled genome nucleating virus assembly by initiating the bending of the membrane vesicle. This eventually would lead to the pinching off of DNA-filled vesicles resulting in coupled membrane morphogenesis, genome encapsidation, and capsid assembly. The highly organized transmembrane proteins (the lattice formed of P3 and P6) on the membrane surface can act as a scaffold for capsid assembly. The new virus particles always appear adjacent to or in association with the host cytoplasmic membrane, which apparently is the place for the virus assembly (Cota-Robles, Espejo, & Haywood, 1968). 3.1.2 Lipids in PRD1 and related viruses Viruses of the Tectiviridae family have a linear dsDNA genome and a proteinrich internal membrane enclosed in an icosahedral capsid shell. The genomes have inverted terminal repeats and a covalently linked 50 -terminal proteins used in the replication and packaging (Bamford, McGraw, MacKenzie, & Mindich, 1983; Ziedaite, Kivela¨, Bamford, & Bamford, 2009). Tectiviruses can be divided into two groups: viruses infecting Gram-negative bacteria and Gram-positive bacteria (Oksanen & Bamford, 2012a). The phages

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infecting Gram-negative bacteria include PRD1, PR3, PR4, PR5, PR722, and L17. They are very closely related having genome sequence identity between 91.9% and 99.8% (Saren et al., 2005; Table 1). These viruses have been isolated from distant locations all over the world, and the most different structural viral proteins are those responsible for host recognition (Saren et al., 2005). In addition, three phages, P37-14, P23-77, and ϕIN93, infecting Thermus strains have been reported, of which P23-77 is rather well described at genomic and structural levels ( Jaatinen et al., 2008; Jalasvuori et al., 2009; Rissanen et al., 2012, 2013). P23-77 and ϕIN93 share some sequence similarity, their genomes are colinear, and the virion proteins of P23-77 and ϕIN93 are 75% similar ( Jalasvuori et al., 2009). The recently isolated phage, KHP30, infecting Helicobacter pylori might also be a virus with an internal membrane (Uchiyama et al., 2013). The group of tectiviruses with Gram-positive Bacillus hosts currently includes Bam35, AP50, Wip1, GILO1, GIL16, and ϕNS11 (Table 1). These phages have also similarity at the nucleotide sequence level (Kan et al., 2013). The virion morphologies of PRD1 and Bam35 are indistinguishable, and the gene order is conserved in their genomes. In addition, their nucleotide identity across the genomes is 42% which, however, is very patchy (Ravantti et al., 2003). The placement of P23-77 (and maybe P37-14) in the family Tectiviridae is controversial, since the genome is circular, while the other tectiviruses have linear genomes (Oksanen & Bamford, 2012a). 3.1.3 Lipids of PRD1 form an icosahedrally ordered membrane The host range of PRD1 is broad including various Gram-negative bacteria such as Salmonella enterica, Escherichia coli, and Pseudomonas aeruginosa. However, the phage infects only strains which harbor an IncP-, IncW-, or IncNincompatibility group conjugative plasmid (Olsen et al., 1974). The genetic system available for PRD1 with a number of suppressor-sensitive virus mutants (Mindich, Cohen, & Weisburd, 1976) has been the key for success making PRD1 one of the best-described virus systems. The extensive genetic, biochemical, and structural approaches to study PRD1 have yielded invaluable knowledge about DNA delivery, virion assembly, and genome packaging. The structure of the PRD1 virus with a diameter of about 65 nm and a mass of about 66 MDa has been determined by X-ray crystallography at ˚ resolution (Abrescia et al., 2004; Cockburn et al., 2004). Fifteen per4 A cent of the virion mass is lipid, the rest being protein (70%) and DNA (15%; Bamford, Caldentey, & Bamford, 1995). The membrane constitutes

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of both protein and lipid in an approximate ratio of 1:1 (Davis, Muller, & Cronan, 1982). It is icosahedral and resides underneath the capsid shell enveloping the dsDNA genome. Half of the 18 PRD1 structural protein species are associated with the membrane. The vertex-stabilizing protein P16 at the fivefold vertices is the only icosahedrally ordered membrane protein which is visible in the electron density map (Abrescia et al., 2004). Laser Raman spectroscopy of PRD1 has shown that the lipids are in the liquid crystalline phase, and the membrane proteins are mainly α-helical (Bamford, Bamford, Towse, & Thomas, 1990; Tuma, Bamford, Bamford, Russell, & Thomas, 1996). The PRD1 membrane is well ordered, and it follows the icosahedral shape of the capsid (Cockburn et al., 2004). It is composed of 52% PE, 43% PG, and 5% CL (Laurinavicˇius, Ka¨kela¨, Somerharju, et al., 2004). The relative lipid composition in the PRD1 membrane is different from the outer or cytoplasmic membranes of the host bacterium S. enterica (for the host: 80% PE, 12% PG, and 8% CL; Laurinavicˇius, Ka¨kela¨, Somerharju, et al., 2004) showing that the virus acquires its lipids selectively upon assembly. The viral membrane comprises 26,000 lipid molecules asymmetrically distributed between the membrane leaflets (Cockburn et al., 2004; Laurinavicˇius et al., 2007). PG and CL are enriched in the outer membrane leaflet, whereas the zwitterionic PE is mainly found in the inner leaflet, facilitating close interactions with the genome (Cockburn et al., 2004; Laurinavicˇius et al., 2007). The enrichment of PG has also been observed in phage PM2 (Section 3.1.1; Laurinavicˇius et al., 2007). The asymmetric distribution may be a consequence of the shapes and charges of different phospholipid molecules and their interactions with the surrounding membrane proteins. In addition, the membrane proteins may affect the lipid composition locally driving the selective uptake of the lipids from the host pool. 3.1.4 PRD1 genome delivery occurs through a membranous tunneling nanotube The internal membrane of PRD1 is capable of forming a tubular structure, which has a central role in the delivery of DNA during infection (Bamford & Mindich, 1982; Grahn, Daugelavicˇius, & Bamford, 2002a, 2002b; Lundstr€ om, Bamford, Palva, & Lounatmaa, 1979; Peralta et al., 2013; Fig. 3A and B). The process is initiated by specific recognition of the cellular receptor (IncP plasmid-encoded DNA transfer complex) by the receptor recognizing fivefold spike protein P2 (Grahn, Caldentey, Bamford, &

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Bamford, 1999; Mindich et al., 1982). The irreversible binding to the cell surface receptor, which is possibly guided by the other fivefold spike protein P5, positions the virus particle to the cell surface and triggers structural changes in the virion. This leads to dissociation of the receptor recognizing fivefold vertex complexes composed of the receptor-binding protein P2, spike protein P5, penton protein P31, and vertex-stabilizing membrane protein P16, as well as the surrounding peripentonal capsid protein P3 trimers from the virion (Peralta et al., 2013). The openings formed at the vertices allow the capsid–membrane interactions to be destabilized, and the membrane vesicle is transformed into a tube-like structure that crosses the cell envelope (Peralta et al., 2013). The tube, which is 4.8 nm thick and 50 nm in length, is capable of penetrating the host envelope (the envelope of Salmonella is 15 nm thick) and protects the genome upon delivery to the cytoplasm. It has been proposed that the PRD1 membrane tube passes through the unique vertex also used for DNA packaging (Hong et al., 2014; Peralta et al., 2013; Str€ omsten, Bamford, et al., 2003). In the packaged virion, the putative interaction between the packaging ATPase P9 and the viral genome via the terminal protein P8 might serve as the nucleation point and facilitate tail tube formation through the vertex (Karhu, Ziedaite, Bamford, & Bamford, 2007; Peralta et al., 2013). However, the packaged genome is not a prerequisite for the tube formation, since the PRD1 procapsids devoid of the genome are still capable of forming the tube (Bamford & Mindich, 1982; Peralta et al., 2013). Cellular tomography of PRD1-infected cells allowed visualization of the entire tail tube penetrating the cell envelope. In most cases, the tail tube entered the cell surface almost orthogonally (Peralta et al., 2013). Several integral viral membrane proteins (P7, P14, P18, and P32) have been shown to be crucial for the formation of the membrane tube (Bamford & Mindich, 1982; Grahn et al., 2002a). In addition, the major membrane protein P11 (the adhesive factor of the membrane) most probably operates after receptor binding and interacts with the outer membrane of the host bacterium (Grahn et al., 2002a). For cell wall digestion, PRD1 uses a membraneassociated peptidoglycan degrading enzyme P7 which is a transglycosylase located in the viral membrane (Rydman & Bamford, 2000, 2002). The visualization of the tube by cryoelectron microscopy revealed different regions with either high or low densities indicating that the membrane proteins might act as a scaffold for the tube (Peralta et al., 2013).

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Once the tail tube has reached the cytoplasm, the viral genome is released most probably through the opening of the tip of the tail tube. The internal diameter of the tail tube is 4.5 nm, which allows one viral dsDNA chain to be translocated into the host cytoplasm (Peralta et al., 2013). The DNA release might be fuelled initially by the energy stored in the pressurized capsid and/or a change in the osmotic pressure (Cockburn et al., 2004; Peralta et al., 2013). 3.1.5 Assembly and packaging of internal membrane-containing bacteriophage PRD1 During PRD1 infection, no transcriptional activation of the host bacterium genes involved in the phospholipid biosynthesis has been recorded (Poranen et al., 2006). The overall changes in the host bacterium during virus infection are almost nonexistent and PRD1 utilizes only a small fraction (5–15%) of the synthesizing capacity of the host cell (Poranen et al., 2006). Fifteen minutes postinfection, a number of virus capsid-associated proteins, such as the MCP P3 trimers, receptor-binding protein P2 monomers, spike protein P5 trimers, and penton protein P31 pentamers are found soluble in the cytoplasm. The virus-encoded membrane proteins such as P7, P14, P11, and P18, on the other hand, are addressed to the host cytoplasmic membrane (Mindich et al., 1982; Fig. 3C). The correct folding of the viral proteins including several membrane proteins is mediated by the host GroEL/GroES chaperonins (Ha¨nninen et al., 1997). Most probably, the membrane proteins are clustered together in the host cytoplasmic membrane, where the lipid–protein interactions might serve as the nucleation point for procapsid assembly (Fig. 3D). The membrane-bound nonstructural scaffolding protein P10 guides the particle assembly, in which the virus-specific membrane area derived from the host cytoplasmic membrane is assembled together with the capsid-associated proteins (Rydman et al., 2001). Protein P30 is located on the surface of the viral membrane and is essential for particle assembly as it defines the size of the icosahedral particle by acting as a molecular tape measure protein (Abrescia et al., 2004; Rydman et al., 2001). In addition, phage-encoded nonstructural assembly factors P10, P17, and most probably P33 have roles in the procapsid formation (Mindich et al., 1982). It has been shown that protein P17, which is a soluble tetramer, is capable of binding to the positively charged lipid membranes (Caldentey et al., 1999; Holopainen, Saily, Caldentey, & Kinnunen, 2000).

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The empty membrane-containing procapsids, which are visible 40 min after infection, contain all structural proteins, except the packaging ATPase P9 and the genome terminal protein P8 (Fig. 3E; Mindich et al., 1982; Str€ omsten, Bamford, et al., 2003). PRD1 has a unique vertex through which the virus genome is packaged powered by the packaging ATPase P9 (Fig. 3F; Gowen, Bamford, Bamford, & Fuller, 2003; Karhu et al., 2007; Str€ omsten, Bamford, et al., 2003; Str€ omsten, Bamford, & Bamford, 2005; Ziedaite et al., 2009; Hong et al., 2014). Other unique vertex proteins are the packaging efficiency factor P6 and two small integral membrane proteins P20 and P22, which together form a conduit mediating the translocation of the viral DNA into the internal membrane vesicle of the procapsid (Hong et al., 2014). During packaging, the internal membrane slightly expands, and the interactions between the capsid and the underlying membrane increase (Butcher, Bamford, & Fuller, 1995; San Martin et al., 2002). In the virion, the curvature of the membrane is the highest underneath the fivefold vertices and at the edges of the facets (Cockburn et al., 2004). At these sites, the membrane is connected to the capsid shell either by the vertex-stabilizing membrane protein P16 at the fivefold vertices or by a series of interactions between the N-termini of the MCP P3 on the facet edges (Abrescia et al., 2004; Cockburn et al., 2004; Jaatinen et al., 2008). In addition to the membrane-anchored unique packaging vertex structure of PRD1, the only available structural information considering a packaging ATPase of an internal membrane-containing icosahedral virus is derived from studies of the archaeal virus STIV-2. The X-ray crystallographic structure of STIV-2 packaging ATPase B204 revealed that it belongs to the FtsK-HerA superfamily of P-loop ATPases, whose cellular and viral members have been suggested to have the same mechanism for DNA translocation (Happonen et al., 2010, 2013).

3.2. Enveloped icosahedral viruses: Phage ϕ6 and its relatives A relatively common bacterial virus morphotype is the one with an internal icosahedrally organized NC surrounded by a lipid envelope. There are some 10 isolates described so far (Table 1; Fig. 1). All this type of viruses have a segmented dsRNA genome (three segments), which resides inside the polymerase complex (PC). These viruses are classified into the family Cystoviridae. The PC is the innermost icosahedral structural element that is matured to a NC upon addition of a protein shell made of protein P8 around the PC. Due to the dsRNA genome, these viruses have to deliver

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their PC into the host cytoplasm as the host bacteria are not capable of replicating dsRNA. Within the cell, the PC particle translocates the transcribed ssRNA genomic segments from the interior of the PC to the cytoplasm where they are either used to guide protein synthesis or, later in the infection cycle, packaged to the newly formed PC particles where minus strand synthesis (replication) takes place. The hosts are Gram-negative bacteria with an outer and cytoplasmic membranes and a rigid peptidoglycan layer in between. The vast majority of information obtained for these viruses comes from the original isolate designated as ϕ6, which infects a plant pathogenic Pseudomonas syringae bacterium (Vidaver et al., 1973). The entire NC must be internalized for progeny production in a process where the viral and host membranes are crucial for penetrating the cell envelope. In this chapter, we follow the ϕ6 membrane throughout the infection cycle (Fig. 3H–O), the topic of this review. 3.2.1 Involvement of the membranes in ϕ6 entry There are receptor-binding spikes on the ϕ6 virion surface that are made of protein P3. These spikes are anchored to the virion through the integral membrane protein P6. For ϕ6, the receptor is the side of a type IV pilus (host pathogenesis factor) extending from the host cell surface (Fig. 3H; Mindich, Sinclair, & Cohen, 1976; Roine, Nunn, Paulin, & Romantschuk, 1996; Roine, Raineri, Romantschuk, Wilson, & Nunn, 1998; Romantschuk & Bamford, 1985, 1986). Other ϕ6-like viruses may use different cell surface structures for attachment. The retraction of the pilus filament brings the virion into contact with the cell surface (Romantschuk & Bamford, 1985; Romantschuk, Olkkonen, & Bamford, 1988). In this process, the P3-spike protein dislocates to expose protein P6 (Fig. 3I). Protein P6 has been shown to have fusogenic activity leading to the fusion of the viral membrane with the host outer membrane (Bamford, Palva, & Lounatmaa, 1976; Bamford, Romantschuk, & Somerharju, 1987; Lounatmaa & Bamford, 1978). This process locates the NC to face the peptidoglycan layer. There is a lytic enzyme, protein P5, associated with the NC surface digesting locally the peptidoglycan layer (Caldentey & Bamford, 1992; Hantula & Bamford, 1988; Kakitani, Iba, & Okada, 1980; Mindich, Lehman, & Huang, 1979; Romantschuk & Bamford, 1981). This positions the NC to face the host plasma membrane. The NC surface protein P8 is membrane active pushing the NC to acquire a plasma membranedriven envelope enclosing the NC in an endocytotic-like event (Fig. 3J; Ojala, Romantschuk, & Bamford, 1990; Olkkonen, Ojala, & Bamford,

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1991; Poranen, Daugelavicˇius, Ojala, Hess, & Bamford, 1999; Romantschuk et al., 1988). How the NC dissociates to release the P8 shell and how the PC escapes from the membrane vesicle and starts transcription are currently less understood than the early stages of entry. It is considered that the membrane-active nature of P8 and pH may be crucial factors in this process (Bamford, Bamford, Li, & Thomas, 1993; Cvirkaite-Krupovicˇ, Poranen, & Bamford, 2010; Huiskonen et al., 2006; Poranen et al., 1999; Tuma, Bamford, Bamford, & Thomas, 1999). 3.2.2 How ϕ6 acquires its membrane envelope The entering PC secretes the first transcripts into the cytoplasm-producing proteins needed for the empty PC assembly (Fig. 3K). Later in the life cycle, there is an increase of synthesis of envelope constituents and protein P8 assuring the maturation of the NC and formation of the viral envelope (Fig. 3L; Poranen, Tuma, & Bamford, 2008). The most crucial components for the membrane assembly are the integral membrane protein P9 that is inserted to the host plasma membrane and an assembly factor protein P12. P9 synthesis defines a virus-specific membrane region from where the host cytoplasmic membrane proteins are expelled (Fig. 3M). With the aid of nonstructural protein P12, a virus-specific vesicle that encloses the NC, is formed. These enveloped viral particles are released to the cell interior where the receptor-binding protein P3 is added to the particle finalizing the virion assembly process (Fig. 3N; Bamford et al., 1976; Ellis & Schlegel, 1974; Gonzalez, Langenberg, Van Etten, & Vidaver, 1977; Johnson & Mindich, 1994; Mindich et al., 1979; Stitt & Mindich, 1983a, 1983b). The acquirement of the membrane from the plasma membrane is an intriguing event due to the release of the virions to the cell interior instead of budding through the plasma membrane. The reason for such an assembly pathway obviously is the presence of the rigid peptidoglycan layer and the outer membrane of the host. The topology of the viral membrane in respect to the NC is particularly intriguing (Stitt & Mindich, 1983b). Either the NC (carrying the external P8 shell) recognizes the P9 batch in the cytoplasmic membrane and inverts to be released to the cell interior or a virus-specific vesicle with P9 and P8 is formed possibly attached to the host cytoplasmic membrane. P8 is in the context of the NC or alternatively, associated with the P9 in the virus-specific vesicle (Sarin et al., 2012). Finally, the NC is internalized to these virus-specific vesicles that are pinched off to release the viral particle to the cell interior. There is morphological electron microscopic support using mutant viruses for the presence of cytoplasmic

Prokaryotic Viruses with Lipids

35

membrane bags with internal NCs (Bamford, 1980). At the end of the life cycle, the virions are released by the host cell lysis (Kakitani et al., 1980; Mindich & Lehman, 1979; Romantschuk & Bamford, 1981). 3.2.3 Lipids of the ϕ6 virion and its host The phospholipid class and molecular species compositions of bacteriophage ϕ6 and its P. syringae host vary considerably pointing to specific mechanism to acquire the lipid (and protein) constituents for the virus. The phage contains significantly more PG and less PE than the cytoplasmic or outer membranes of its host. In addition, the phospholipid molecular species composition of the viral membrane also differs from those of the host membranes. However, it resembles more that of the cytoplasmic membrane than the outer membrane, which is in line with the observation that ϕ6 derives its phospholipids from the host cytoplasmic membrane (Laurinavicˇius, Ka¨kela¨, Bamford, & Somerharju, 2004). The outer leaflet contains approximately equal amounts of PE and PG. In the inner leaflet, PE is considerably enriched (65% PE and 31% PG; Laurinavicˇius et al., 2007). The phospholipid shape and charge as well as the interactions with the membrane proteins are the major selective phenomena. It should be noted that the membrane of ϕ6 is rich in proteins and the curvature and surface areas of the different leaflets are considerably different with some 40% more surface area in the outer leaflet (Laurinavicˇius, Ka¨kela¨, Bamford, et al., 2004). Obtaining and combining additional structural and biochemical data will allow to further refine the biogenesis of the ϕ6 membrane.

3.3. Vesicular pleomorphic viruses The word pleomorphic is commonly used to describe the morphology of enveloped viruses with asymmetric or variable virion architecture. Among the known prokaryotic viruses, L2 and L172 infecting mycoplasmas and pleolipoviruses infecting halophilic archaea, have a spherical, pleomorphic virion architecture (Table 1 and Fig. 1). Plasmavirus L2 has been classified into the family Plasmaviridae, but the other viruses are currently unclassified (Maniloff, 2012). The proposed viral family Pleolipoviridae includes the seven known pleolipoviruses (Pietila¨ et al., 2012) representing related viruses with different genome types (circular ssDNA and circular or linear dsDNA; Table 1 and Fig. 4; Sencˇilo et al., 2012). Interestingly, the genomes of the mycoplasma viruses L2 and L172 are ds and ssDNA, respectively, and the sizes correspond to those described for the genomes of pleolipoviruses (see below; Dybvig et al., 1985; Sencˇilo et al., 2012).

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Figure 4 Archaeal pleolipoviruses. The width of the transmission electron micrographs of the negatively stained virions represents 100 nm. The protein profile of the purified virions analyzed by SDS-PAGE with Coomassie blue and Sudan Black staining is shown on the right. The hypothetical positions of HGPV-1 VP2 and His2 VP32 proteins are marked with rectangles as they do not bind Coomassie Blue. The lipid-modified spike proteins of HGPV-2 and His2 are indicated by an asterisk. Molecular mass standards are shown on the far right.

Transmission electron micrographs of the prokaryotic pleomorphic viruses resemble each other, and the different viruses have roughly similar virion diameters. While pleolipoviruses are known to be decorated by spikes, so far such structural elements have not been observed for the mycoplasma viruses by the negative stain TEM (King et al., 2012; Maniloff, 2012; Pietila¨ et al., 2009, 2012; Roine et al., 2010). However, visualization of such virion surface structures may require additional methods and a higher resolution. The life cycles of all the prokaryotic pleomorphic viruses are nonlytic and persistent (Pietila¨ et al., 2012; Roine & Oksanen, 2011). The only significant difference in the life style is that L2 and L172 integrate in the host chromosome while pleolipoviruses have not been found to have integrases (Putzrath & Maniloff, 1977, 1978; Sencˇilo et al., 2012). However, several proviral pleolipovirus-like elements (plasmids and proviruses) in the genomes of some halophilic archaeal strains have been described (Sencˇilo et al., 2012), indicating that integration may be part of the life style of these viruses. Although the genomes of L172 and L2 are not related, the major structural proteins of both viruses are around 15–20 and 60–70 kDa (Dybvig et al., 1985), similar to those observed for pleolipoviruses. The genomes of pleolipoviruses are not related to plasmavirus L2 (E. Roine,

Prokaryotic Viruses with Lipids

37

personal communication), and no sequence data are currently available for L172. This, however, does not exclude relatedness at the level of virion architecture, which is considered to indicate common ancestry (Abrescia, Bamford, Grimes, & Stuart, 2012; Bamford, 2003). 3.3.1 Asymmetric lipid vesicles as viruses What makes pleolipoviruses special compared to other spherical pleomorphic viruses is the absence of a NC underneath the lipid membrane (Pietila¨ et al., 2009, 2010). In addition, the membrane bilayer of these viruses is covered with spike proteins, which are considered to partly contribute to the pleomorphic nature of the virions. These properties result in the characteristic “floppy,” membrane vesicle-like appearance. Like most of the known haloarchaeal viruses, pleolipoviruses require high NaCl concentrations for infectivity (Pietila¨ et al., 2010, 2012). The described pleolipoviruses (Fig. 4) infect halophilic archaea from the genera Halorubrum, Haloarcula, and Halogeometricum. Like their hosts, these viruses have been isolated from spatially distant hypersaline environments indicating worldwide distribution (Atanasova et al., 2012; Bath et al., 2006; Pietila¨ et al., 2009, 2012; Roine et al., 2010). The first and the best-described pleolipovirus, Halorubrum pleomorphic virus 1 (HRPV-1), was described in 2009 as the first archaeal virus with an ssDNA genome (circular 7048 nt; Pietila¨ et al., 2009). This virus and its host Halorubrum sp. were isolated from an Italian saltern in Trapani, Sicily. The major structural proteins of HRPV-1 are VP4 (spike protein, 53 kDa) and VP3 (integral membrane protein, 14 kDa; Pietila¨ et al., 2009). In addition, the putative ATPase (VP8) was detected as a minor protein component of the virion. The randomly distributed spikes are glycosylated and considered to serve as receptor binding and fusion proteins (Kandiba et al., 2012; Pietila¨ et al., 2010). TLC analysis of viral and host lipids showed that the virus acquires its lipids unselectively from the host (Pietila¨ et al., 2010). The viral lipids are composed of CL, PG, PG-Me, and PGS by MS. The absence of a NC was shown by quantitative biochemical dissociation studies (Pietila¨ et al., 2010). HRPV-1 is a nonlytic virus, which persists in the host cells allowing continuous virus production yielding high titers (Pietila¨ et al., 2009). In 2010, another virus, Haloarcula hispanica pleomorphic virus 1 (HHPV-1), with morphology similar to HRPV-1, was reported. Interestingly, the genome of this virus consisted of dsDNA (circular 8082 bp; Roine et al., 2010). Despite the different genome types, these two viruses were found to have gene synteny and sequence similarity proving their

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relatedness. In addition, HRPV-1 and HHPV-1 were found to be related to pHK2 plasmid of Haloferax sp. and to a proviral element in the genome of Haloferax volcanii (Roine & Oksanen, 2011). It is suggested that pHK2 might be a temperate virus resembling HRPV-1 and HHPV-1 (Roine et al., 2010). The viral lipids of HHPV-1 are similar to those observed for HRPV-1, and the virus acquires them unselectively from the host. The life cycle, protein profile, virion size, and morphology are similar to HRPV-1, but the spike of HHPV-1 is not glycosylated (Roine et al., 2010). A global survey of nine different hypersaline environments resulted in the isolation of four new pleomorphic viruses similar to HRPV-1 and HHPV-1 (Fig. 4; Atanasova et al., 2012). These six viruses as well as His2, which had previously been described as a spindle-shaped virus (Bath et al., 2006) but shown to have sequence similarity and morphological resemblance to HRPV-1 and HHPV-1, were included in comparative studies of virion architecture, life cycles, and genomics (Pietila¨ et al., 2012; Sencˇilo et al., 2012). Virion morphology was studied by cryoelectron microscopy and tomography revealing that all the viruses were roughly spherical and decorated with spikes having virion diameters from 40 to 70 nm (Fig. 4). Virion protein profiles of the new isolates were similar to the previously characterized ones, except that Halogeometricum pleomorphic virus 1 (HGPV-1) contained two membrane proteins and His2 contained two spike proteins. In addition, the spikes of these two viruses were found to be lipid modified (Pietila¨ et al., 2012). The lipid composition of the seven pleomorphic viruses resembles that of their hosts indicating unselective lipid acquisition. All viruses except HGPV-1 contained similar lipids than HRPV-1 and HHPV-1 (see above). In HGPV-1, as well as its host Halogeometricum sp. CG-9, the band corresponding to PGS in TLC was absent. Based on the presence of lipids and the nonlytic life style, it has been suggested that pleolipoviruses acquire their lipids and exit their host cells by budding (Pietila¨ et al., 2012). All the seven pleolipoviruses have a conserved cluster of five genes (including those encoding for the spike and internal membrane proteins) that are related to each other based on synteny and amino acid sequence similarity (Sencˇilo et al., 2012). In addition, it seems that pleomorphic viruses with circular genomes replicate via rolling circle replication, while protein-primed replication is suggested for His2 (Bath et al., 2006). The genomes of HRPV-3 and HGPV-1 have unusual, short single-stranded regions in their genomes suggesting that they might use a different replication strategy (Sencˇilo et al., 2012).

Prokaryotic Viruses with Lipids

39

3.3.2 Bacterial vesicular viruses Acholeplasma laidlawii virus L2 is the type species, and the only approved member, of the genus Plasmavirus in the family Plasmaviridae. Other candidates in the genus are the Acholeplasma phages M1, O1, v1, v2, v4, v5, and v7, but to date, these viruses remain unclassified (Maniloff, 2012). L2 virions are pleomorphic and approximately 80 nm in diameter. The viral genome is 11,965-bp circular dsDNA and contains a putative integrase. The virus is able to establish lysogeny with the host, A. laidlawii (strain Ja1; Dybvig & Maniloff, 1983). The productive virus life cycle is nonlytic and persistent retarding the host growth rate. After 6 h postinfection, the virus is able to persist in the host cells, which in turn can give rise to new carrier cell clones or cells that are resistant to the virus (Putzrath & Maniloff, 1977). In the lysogenic state, L2 integrates into a unique site in the host genome (Dybvig & Maniloff, 1983). L2 was isolated in England by washing the host cultures with phosphatebuffered saline (Gourlay, 1971). Virus production of infected cells is increased almost threefold by ultra violet radiation or mitomycin C (Putzrath & Maniloff, 1978). L2 acquires its lipids, which form the outer membrane of the virion, unselectively from A. laidlawii. It is considered that L2 obtains its lipid envelope during budding from the host cell (Al-Shammari & Smith, 1981). The L2 virion contains major structural proteins of 19, 58, 61, and 64 kDa, but additional minor protein species have been observed (Maniloff, 2012). L172 is the first described enveloped phage with an ssDNA genome (Dybvig et al., 1985). The virus was isolated from A. laidlawii strain S2 in the former Czechoslovakia (Liska, 1972). The presence of an envelope was first suggested based on sensitivity to detergents and organic solvents (Gourlay, Wyld, Garwes, & Pocock, 1979). Furthermore, the lipids were studied by TLC and confirmed to be unselectively acquired from the host (Al-Shammari & Smith, 1981). L172 viral lipids consist of phospho-, glyco-, and phosphoglycolipids and the fatty acid composition resembles that of the host (Al-Shammari & Smith, 1981). The virus is temperate and able to establish lysogeny. The productive infection cycle is nonlytic, and the virus exits the host presumably by budding. The 14.0-kbp circular ssDNA genome of L172 shows no homology to the dsDNA genome of L2 (Dybvig et al., 1985). However, the methods used at that time (hybridization) do not allow detailed comparison of sequence data. Virion size (60–80 nm), morphology, nonlytic life style, and structural protein profiles indicate that the two viruses may be related.

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3.4. Prokaryotic viruses with helical symmetry: With or without a membrane Viruses exhibiting helical virion symmetry are infecting both bacterial and archaeal hosts (Fig. 1 and Table 1). Bacterial helical viruses belong to the family Inoviridae, whereas archaeal ones are classified into two families: Lipothrixviridae and Rudiviridae, which are grouped into a single-order Ligamenvirales (Day, 2012; Prangishvili, 2012a, 2012b; Prangishvili & Krupovicˇ, 2012). Viruses belonging to these families have either filamentous or rod-shaped virions essentially containing a proteinaceous core and an ss or dsDNA genome (Day, 2012; Prangishvili, 2012a, 2012b). Of the three families, only the members of Lipothrixviridae have lipid constituents in the virion (Prangishvili, 2012b). 3.4.1 Lipothrixviruses: Helical viruses with a membrane envelope Up to date, a total of 12 lipothrixviruses infecting hyperthermophiles belonging to the genera Sulfolobus, Acidianus, and Thermoproteus have been reported (Bize et al., 2008; Ha¨ring, Vestergaard, Brugger, et al., 2005; Prangishvili, 2012b; Vestergaard et al., 2008; Table 1). Similarly to the majority of other hyperthermophilic archaeal viruses, lipothrixviruses of archaea in the genera Sulfolobus and Acidianus do not cause host cell lysis and are thought to establish a persistent infection (Arnold et al., 2000; Bettstetter et al., 2003; Bize et al., 2008; Ha¨ring, Vestergaard, Brugger, et al., 2005; Vestergaard et al., 2008). On the contrary, viruses infecting hosts belonging to the genus Thermoproteus have been suggested to be temperate (Zillig et al., 1988). Lipothrixvirus particles are flexible filaments minimally consisting of genomic DNA covered by a proteinaceous inner core and surrounded by an outer lipid membrane (Prangishvili, 2012b). Based on the details in virion organization, lipothrixviruses are divided into three established and one proposed genera: Alpha-, Beta-, Gamma-, and “Deltalipothrixviruses” (Prangishvili, 2012b). The genomes of lipothrixviruses are linear dsDNA molecules ranging from approximately 14 kb (TTV1) to 56 kb (DAFV; Neumann et al., 1989; Zillig et al., 1994). The nature of the genome termini is not clear, but it has been hypothesized that the termini may have an unknown chemical modification or may be associated with proteins (Arnold et al., 2000; Bettstetter et al., 2003; Bize et al., 2008; Ha¨ring, Vestergaard, Brugger, et al., 2005; Janekovic et al., 1983; Vestergaard et al., 2008). Lipothrixvirus

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genomes have modular organization shaped by horizontal gene transfer (HGT) between viruses within and outside the Lipothrixviridae family. Despite HGT obscuring the border between families and genera, a phylogenetic tree built based on the conserved hypothetical protein shared among all sequenced lipothrixviruses except for TTV1 verified the designation of lipothrixviruses into the existing genera (Vestergaard et al., 2008). 3.4.1.1 Alphalipothrixviruses

Alphalipothrixvirus Thermoproteus tenax virus 1 (TTV1) is one of the earliest and the best-characterized archaeal viruses ( Janekovic et al., 1983; Table 1). TTV1 particles are approximately 400 nm long and 40 nm wide filaments. Virion inner core is composed of genomic DNA superhelix wound around a central cavity and covered by DNA-binding proteins P1 and P2. Unlike other lipothrixviruses, TTV1 has an additional layer between the inner core and the outer lipid membrane. This layer consists of the helically arranged protein P3 delimited by the cap structures made of the protein P4. It has been suggested that this proteinaceous layer causes greater TTV1 virion rigidity compared to other lipothrixviruses (Arnold et al., 2000; Bize et al., 2008; Janekovic et al., 1983; Vestergaard et al., 2008; Zillig et al., 1988). TTV1 outer membrane was shown to have qualitatively the same, but quantitatively different lipid species than its host (Zillig et al., 1988). 3.4.1.2 Betalipothrixviruses

Betalipothrixvirus genus has eight members: T. tenax viruses 2 and 3 (TTV2 and TTV3), Desulforolobus ambivalens filamentous virus (DAFV), Acidianus filamentous viruses 3, 6, 7, 8, and 9 (AFV3, AFV6, AFV7, AFV8, and AFV9), and the type species Sulfolobus islandicus filamentous virus 1 (SIFV; Table 1). Virions are on average 2000 nm long and 25 nm wide filaments typically having tapered ends terminating with different number of fibers (Arnold et al., 2000; Bize et al., 2008; Janekovic et al., 1983; Vestergaard et al., 2008; Zillig et al., 1994). Particles of SIFV and AFV3 viruses have been studied in more detail, and the proposed models for their structures are in good agreement (Arnold et al., 2000; Vestergaard et al., 2008). According to the models, virion inner core has a nucleosome-like structure composed of two major structural proteins forming doughnutshaped subunits arranged in a zipper-like array with the genomic DNA wrapped around them. This model is fundamentally different from the suggested structure of TTV1 virion inner core. Both SIFV and AFV3 virions have fairly similar lipid constituents as their hosts (Arnold et al., 2000;

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Vestergaard et al., 2008). However, one lipid species found in S. islandicus is more abundant in the virus than in the host. In addition, there seems to be one modified host lipid species in the SIFV envelope, which is suggested to result from glycosylation of host lipids by virus-encoded glycosyl transferases (Arnold et al., 2000). 3.4.1.3 Gammalipothrixviruses and deltalipothrixviruses

Gammalipothrixvirus genus is represented by a single virus—Acidianus filamentous virus 1 (AFV1). AFV1 virions are 900 nm long and 24 nm wide filaments with claw-like structures at both termini (Table 1 and Fig. 5). The claw-like structures are connected to the main viral body (inner core and outer envelope) via narrow appendages with collars. As determined by TLC, one of the host lipids was absent from AFV1 particles, and the majority of the other lipids had different mobilities arguing that the viruses might modify the acquired host lipids (Bettstetter et al., 2003). One more proposed genus belonging to Lipothrixviridae is “Deltalipothrixvirus,” which includes a single isolate—Acidianus filamentous virus 2 (AFV2). Although AFV2 shares morphological similarity and

Figure 5 Lipothrixviruses. Electron micrographs of negatively stained virions of Acidianus filamentous virus 1 (AFV1) from the genus Gammalipothrixvirus. The bars represent 100 nm. Used with the permission from David Prangishvili and the International Committee on Taxonomy of Viruses (ICTV).

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a number of homologues with other lipothrixviruses, no lipids have been detected in AFV2 particles (Ha¨ring, Vestergaard, Brugger, et al., 2005).

3.5. Lemon-shaped viruses are specific for archaea Virus-like particles (VLPs) resembling lemons are abundant in both hyperthermic and hypersaline environments, where archaea typically dominate. Lemon- or spindle-shaped VLPs have also been observed in samples from freshwater sediments and an Antarctic lake (Borrel et al., 2012; Lo´pezBueno et al., 2009; Rachel et al., 2002; Santos et al., 2007; Sime-Ngando et al., 2011). So far, such viruses are archaea specific as no spindle-shaped virus has been described to infect bacteria or eukaryotes (King et al., 2012). Although spindle-shaped viruses share the lemon-shaped core, they can be divided into three groups based on the tail structure: (i) one short, (ii) long tail attached to one of the pointed ends, or (iii) one long tail attached to each end (King et al., 2012). 3.5.1 Viruses with one short tail His1 is so far the only spindle-shaped virus isolated from hypersaline environments (Bath & Dyall-Smith, 1998). His1 virions have a very short tail at one end, and the particles are flexible and their size may vary (Fig. 6A; Bath & Dyall-Smith, 1998; Pietila¨ et al., 2013). Although this flexibility may indicate the presence of a membrane, chloroform–methanol extraction and TLC have shown that the virions contain no free lipids. However, the MCP of His1, VP21, exists in two forms in the virion and one of them seems to be lipid modified (Fig. 6B). Thus, this modification may contribute to the plasticity of the virion (Pietila¨ et al., 2013). In addition to the MCP, a few minor structural proteins of His1 virions have been identified (Pietila¨ et al., 2013). Recently, the first DNA ejection study was performed for an archaeal virus using His1 as a model. It was concluded that the genome ejection from His1 virions to host cells is dependent on host factors as the ejection triggered by detergents was randomly paused and incomplete (Hanhija¨rvi, Ziedaite, Pietila¨, Haeggstr€ om, & Bamford, 2013). His1 resembles morphologically other short-tailed spindle-shaped viruses, but it is the only one described to have a linear dsDNA genome and to encode a putative, type-B DNA polymerase for protein-primed replication (Bath et al., 2006). Consequently, His1 is currently classified into a floating genus, Salterprovirus (King et al., 2012). The majority of short-tailed spindle-shaped viruses infect hyperthermophilic crenarchaea and are classified into the viral family Fuselloviridae, whose type species is Sulfolobus

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Figure 6 Morphology and protein composition of spindle-shaped virus His1. (A) Transmission electron micrographs of purified His1 virions which have been negatively stained with uranyl acetate (upper panel) and ammonium molybdate (lower panel). Arrows indicate large-sized particles. Ammonium molybdate results in highly elongated particles. Scale bar represents 100 nm. (B) Protein profiles of purified His1 virions in a tricine-SDS-polyacrylamide gel stained with Coomassie blue for proteins (left panel), with Sudan Black for lipids and lipoproteins (middle panel), or first with Sudan Black and then with Coomassie blue (right panel). St indicates molecular mass markers (VP, for virion protein).

spindle-shaped virus 1 (SSV1; King et al., 2012). Fuselloviruses have circular dsDNA genomes of similar size and they share gene synteny as well as significant nucleotide sequence similarity (Redder et al., 2009). SSV1 virions contain one major structural protein, VP1, and three minor ones (Menon et al., 2008; Reiter, 1987). SSV1 VP1 and His1 VP21 are similar at the amino acid level indicating that His1 may be related to fuselloviruses despite their different genome types (Pietila¨ et al., 2013). Interestingly, SSV1 VP1 has been reported to form two bands in a gel similar to His1 VP21 (Pietila¨ et al., 2013; Reiter, 1987). Like His1 virions, fuselloviral virions are rather flexible, but the presence of a lipid membrane is controversial (Martin et al., 1984; Redder et al., 2009). However, as is the case in His1 virions, the MCP of SSV1 may be lipid modified, explaining the two bands in the gel and the virion flexibility. A lemon-shaped VLP isolated from a hyperthermophilic euryarchaeon, Pyrococcus abyssi virus 1 (PAV1), resembles morphologically fuselloviruses

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and shares their genome type (Geslin et al., 2003, 2007; Gorlas et al., 2012). However, they have no significant sequence similarity, and thus PAV1 remains unclassified (Geslin et al., 2007; Gorlas et al., 2012). SSV1 and His1 virions and PAV1 particles are composed of one MCP with two hydrophobic domains (Geslin et al., 2003, 2007; Pietila¨ et al., 2013; Reiter, 1987). Thus, all these MCPs may have the same fold derived from a common ancestor and be lipid modified. 3.5.2 Viruses with one or two long tails In addition to spindle-shaped viruses with a short tail, virus isolates with either one or two long tails have been described. These viruses infect hyperthermophilic crenarchaea (Mochizuki, Sako, & Prangishvili, 2011; Prangishvili et al., 2006; Xiang et al., 2005). Sulfolobus tengchongensis spindle-shaped virus 1 (STSV1) has one long tail and is the largest spindle-shaped virus described so far (Xiang et al., 2005). However, the tail length varies and also two-tailed forms have occasionally been observed. TLC analysis indicates that the virion contains lipids which are selectively acquired from the host cell membrane (Xiang et al., 2005). Very recently, STSV2 was described, with 80% protein sequence identity to STSV1. Similarly to STSV1, the virus contains selectively acquired lipids, and the presence of a membrane has been suggested (Erdmann et al., 2014). So far, only one spindle-shaped virus with two long tails has been isolated. Acidianus two-tailed virus (ATV) is classified into the viral family Bicaudaviridae (King et al., 2012). The extraordinary feature of this virus is the capability to form the tails outside the host cells (Ha¨ring, Vestergaard, Rachel, et al., 2005; Prangishvili et al., 2006). It has been proposed that STSV1 may belong to the same viral family as ATV due to their similar appearance (Pina et al., 2011). Furthermore, ATV and STSV1 share nine homologous genes (Krupovicˇ, White, Forterre, & Prangishvili, 2012). In addition, the MCP of ATV with a unique four-helix bundle fold shares significant sequence similarity with STSV1 MCP for which no structure is available (Goulet et al., 2010; Krupovicˇ et al., 2012; Prangishvili et al., 2006; Xiang et al., 2005). However, ATV has several major structural proteins while STSV1 has only one, and no lipids were obtained from ATV virions after chloroform–methanol extraction (Prangishvili et al., 2006; Xiang et al., 2005). Like spindle-shaped viruses with a short tail, long-tailed isolates are nonlytic (Xiang et al., 2005). However, compared to short-tailed viruses,

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long-tailed ones have clearly larger genomes and virions (Mochizuki et al., 2011; Prangishvili et al., 2006; Xiang et al., 2005). Thus, both viruses seem to form their own groups.

3.6. Archaeal spherical viruses with helical NCs have an envelope In the beginning of the new millennium, several viruses with most intriguing, previously unknown morphologies were described infecting crenarchaeal hyperthermophiles. Not only did these viruses look unusual, but they also had gene sequences giving no matches in the public databases (Prangishvili & Garrett, 2004; Rachel et al., 2002). While some of these unique viral morphotypes (such as droplet- or bottle-shaped viruses) were never subjected to lipid analyses, Pyrobaculum spherical virus (PSV), the type species of the genus Globulovirus in the family Globuloviridae, is known to be surrounded by an outer lipid membrane which consists of selectively acquired host lipids (Ha¨ring et al., 2004). In addition to PSV, another globulovirus, Thermoproteus tenax spherical virus 1 (TTSV1) which morphologically resembles PSV, has been described (Ahn et al., 2006). The membrane envelope of globuloviruses encloses a helical nucleoprotein particle with linear dsDNA. The enveloped virions of PSV are 100 nm in diameter and contain variable numbers of spherical protrusions of about 15 nm in length. The buoyant density in CsCl is around 1.3 g ml1 which is typical for lipid-containing viruses. Interestingly, during isopycnic centrifugation, part of the PSV virions are disrupted which causes the release of the helical nucleoprotein core. The genome of PSV is a 28,337-bp long linear dsDNA molecule with inverted terminal repeats. The majority of the putative genes reside on one DNA strand (Ha¨ring et al., 2004). The major structural protein of the virus is 33 kDa in size, and the two minor ones have a mass of 16 and 20 kDa (King et al., 2012). The virion proteins show no matches to public data bases but have some degree of similarity to those of TTSV1. The infection of PSV is nonlytic and persistent. Viral lipids have been extracted by chloroform– methanol and analyzed by TCL. Two of the lipids have identical mobilities to those of the host and two are different indicating selective acquisition and modification of host lipids. PSV and its host Pyrobaculum sp. D11 were isolated from a neutral hot spring (85 °C) in Yellow Stone National Park (Ha¨ring et al., 2004). DNA metagenomic sequences related to PSV were detected in the same environment a few years later (Schoenfeld et al., 2008). In addition to Pyrobaculum sp., PSV infects a T. tenax strain, which

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is closely related to the original host. Regardless of the strict anaerobic nature of the hosts, the virions are stable when stored in the normal atmosphere (Ha¨ring et al., 2004). TTSV1 (Ahn et al., 2006) was isolated from the enrichment culture of the host T. tenax strain YS44 which was obtained from Indonesian hot springs in Siteri. No lipid analysis has been performed for TTSV1, but the morphology, which resembles that of PSV, indicates that the 70-nm virion is surrounded by a lipid envelope. Transmission electron micrographs of TTSV1 include virus particles which are aggregated as chains. The density of the virion is 1.29 g ml1 indicating presence of lipids. The genome of the virus is a linear dsDNA molecule of 21.6 kbp with no covalently attached proteins (Ahn et al., 2006). The viral life cycle is persistent and nonlytic, and the virus does not integrate into the host cell genome. The virus has one major structural protein of 10 kDa and two minor ones of 20 and 25 kDa in mass. The putative coding DNA sequences of TTSV1 are somewhat similar to those of PSV but have no matches in public databases.

4. CONCLUSIONS Prokaryotic viral lipids are derived from the cytoplasmic membranes of their host organism. Consequently, bacteriophage lipids are significantly different from those found in archaeal viruses. The main differences lay in the structure of the core lipids, which for bacteria (and eukaryotes) are based on fatty acid chains ester linked to glycerol and for archaea isoprenoid side chains ether linked to the glycerol backbone. Depending on the virion structure, derivatives of these core lipids are either incorporated into the virion during assembly of the icosahedral procapsids (internal membranecontaining viruses) or translocated over a preassembled icosahedrally symmetric particle (enveloped bacterial dsRNA viruses) as well as budding through the plasma membrane (enveloped bacterial and archaeal DNA viruses). Head-tailed icosahedral, icosahedral inner membrane-containing, and pleomorphic viral morphologies, of which the latter two contain lipids, are shared among bacterial and archaeal viruses (Fig. 1). Interestingly, the selectivity of lipid acquisition from the host seems to be dependent on virus architecture. All known icosahedral inner membrane-containing and enveloped dsRNA prokaryotic viruses take up their lipids selectively, while the pleomorphic viruses have close or almost identical lipid composition when compared to that of their host cytoplasmic membrane.

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Although lipid-containing bacteriophages have been studied already for several decades, the exact mechanism for lipid acquisition remains a challenge. The best-characterized model viruses are PM2, PRD1, and ϕ6. The viral membrane acquisition of these viruses has opposite end points (Fig. 3). The PM2 and PRD1 membranes lie underneath the protein capsid while in ϕ6 the membrane surrounds the NC (Fig. 3). This implies that the mechanisms of membrane acquirements must also be very different. In the case of these three viruses, the membrane is used as the genome delivery device. In PM2 (circular supercoiled dsDNA genome), the protein capsid dissociates and the membrane is suggested to fuse with the host outer membrane. In PRD1, which has a linear dsDNA genome, the membrane is converted into a tail tube that has all the activities needed to penetrate the host cell envelope. ϕ6 (segmented dsRNA genome inside the NC), on the other hand, uses a unique mechanism where all three membranes (viral membrane, OM, and CM) are involved. All these viruses translocate their progeny virions into the host cell cytoplasm prior to lysis. Due to the short history of archaeal virus research, no such models as above are available, and so far, a detailed comparison is not possible. In addition to the previously mentioned morphotypes that are shared by bacterial and archaeal viruses, archaea are infected by viruses with unique morphotypes, of which many contain lipids. However, for some of these viruses, the lipid status is unknown. An abundant group of archaeal viruses is the lemon-shaped ones with variable tail structures (exemplified by His1 with one short tail, STSV1 with one long tail, and ATV with two long tails; Fig. 1). Fuselloviruses and His1 seem to group together, whereas the long-tailed lemon-shaped viruses might form their own cluster. Currently, STSV1 is the only lemon-shaped virus for which lipids have been detected by TLC. ATV has been considered to be related to STSV1, but does not seem to contain lipids. His1 on the other hand contains lipids only as protein modifications. Similar observations have been made among the helical archaeal viruses. The lipid-containing lipothrixviruses are suggested to have a common origin with rudiviruses, for which no lipids have been detected. This is reminiscent to PRD1-type of viruses (double-beta barrel MCP fold), which either do or do not contain a lipid membrane. In the case of pleolipoviruses and plasmaviruses, for which no rigid protein capsid exists, the membrane also serves to protect the naked viral genome. This type of viruses thus depends on the presence of the membrane establishing the virion.

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Viruses can be classified into structure-based viral lineages according to the conserved MCP fold and the architectural principles of the virion. The core elements related to virion structure and assembly are conserved and thus can be applied to study virus evolution, while functions related to interactions with the host are in constant change. To date, four structure-based viral lineages have been established and several others are being proposed. These lineages contain structurally related viruses that infect organisms from all three domains of life and are considered to have a common origin. The presence of the membrane is not coupled to the virion architecture and viruses in the same structure-based lineage may or may not have lipids. However, viruses with the membrane vesicle architecture (pleolipoviruses and plasmaviruses) always contain lipids as they lack the proteinaceous capsid.

ACKNOWLEDGMENTS We thank Professor Angela Corcelli for critically reading the manuscript. This work was supported by Academy Professor (Academy of Finland) funding grants 255342 and 256518 (D. H. B.). We thank Academy of Finland (grants 271413 and 272853) and University of Helsinki for the support to EU ESFRI Instruct Centre for Virus Production. A. S. is a University of Helsinki Fellow in the Doctoral Program in Microbiology and Biotechnology.

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INDEX Note: Page numbers followed by “f ” indicate figures and “t ” indicate tables.

A Acholeplasma laidlawii virus L2, 39 Acidianus, 40 Acidianus filamentous virus (AFV) type 1, 4t, 42, 42f type 2, 4t, 42–43 type 3, 4t type 6, 4t type 7, 4t type 8, 4t type 9, 4t Acidianus spindle-shaped virus 1 (ASV1), 4t Acidianus two-tailed virus (ATV), 45 Agrobacterium-mediated transient expression, 130–131 AIM2, 77–78 Alphaherpesvirus DNA sensors AIM2, 77–78 cGAS, 81–83 DAI, 75–77 double-stranded, 64–65 IFI16, 78–81 immunity to, 66–67 innate immune system, 67–70 intracellular, 75 model of, 69f RIG-I, 83–84 RNA Pol III, 83–84 structure and entry, 64f TLR9, 70–74 vaccine, 88–90 viral DNA to, 84–85 Alphalipothrixviruses, 41 Anti-MCV antibodies, 209–210 Anti-TNF therapy, 210 AP50, 4t Aphids, potyvirus transmission, 138–142 Apoptosis cFLIP and, 224–226 gammaherpesvirus FLIPs, 223–224 MC159 and, 220–222 MC160 and, 222–223

Arabidopsis thaliana, 137 TuMV infection on, 161–163, 164 Archaea core structures, 3–17 lemon-shaped viruses for, 43–46 morphotypes, 3f pleolipoviruses, 36f spherical viruses, 46–47 Archaeol, 3–17 Asymmetric lipid vesicles as viruses, 37–38 Aujeszky’s disease, 66

B Bacterial vesicular viruses, 39 Bacteriophages, 2 infecting gram-positive bacteria, 22 morphotypes, 3f PM2, 24–25 Bam35, 4t Bean yellow mosaic virus (BYMV), 148–149 Betalipothrixviruses, 41–42 Bimolecular fluorescence complementation (BiFC), 122–123 Budding, 21–22

C Caldarchaeol, 3–17 Cantharidin, 211–212 Capsicum annuum, 145–146 Capsid, 64f Capsid-only strategy, 138–139 Cardiolipins (CLs), 17 Cauliflower mosaic virus (CaMV), 140–141 CD95, 218 Cell surface receptor, 29–30 Cell-to-cell movements, 128–132 Cellular FLIP (cFLIP), 216, 218–219 and apoptosis, 224–226 and IRF3 inhibition, 234–235 and NF-κB activation, 231–232 Cellular tomography, PRD1, 30 cFLIP. See Cellular FLIP (cFLIP)

253

254

Index

cGAS, 81–83 CI/P3N-PIPO complex, 131 CI protein, 108t, 115 mutants, 130–131 Cloning capacity, potyviruses, 165 Clover yellow vein virus (ClYVV), 148–150 Corticovirus PM2, 25–27 CP-interacting protein (CPIP), 125–126 CP-Nter domain, 134 CP protein, 108t, 113, 118–119 N-terminal region of, 118 of PVA, 118–119 in RNA replication, 125–126 in virus–vector interaction, 138–139 Cryotherapy, 211–212 C-terminal CP (CP-Cter) domain, 134 C-terminal domain, 113, 138–139 Curettage, 211–212 CXCL9, 72 CXCL10, 72 Cyclic GMP–AMP (cGAMP), 81

Epidermal growth factor (EGF), 206–207 Epidermal growth factor receptor (EGFR), 206–207 ER exit sites (ERES), 120 Eukaryotic initiation factor (eIF) eIF4E, 125, 132, 147–149 eIF4G, 125

D

G

DAG motif, 139 DAI. See DNA-dependent activator of IFNregulatory factors (DAI) DBP1, 124 Death effector domain (DED) motif, 218, 219f Death receptor (DR) family, 216 Deltalipothrixvirus, 42–43 Dendritic cells (DCs), 67 Dermoscopy, 211 Desulforolobus ambivalens filamentous virus (DAFV), 4t Diacylglycerol, 17f DNA-dependent activator of IFNregulatory factors (DAI) DNA sensors, 75–77 overexpression, 76 DNA-induced signaling, 85–88 DOCK8 gene, 210 dsDNA genome, 25–27

Gammaherpesvirus FLIPs, 223–224 Gammalipothrixviruses, 42–43 Genome replication, 212 Genome structures, potyvirus in, 103–105 GIL01, 4t GIL16, 4t Globuloviridae, 46 Glutathione peroxidase homolog, 237 Gram-positive Bacillus, 28 Gram-positive bacteria, 22 Green fluorescent protein (GFP), 120–121

E eEF1A, 123 Empty membrane-containing procapsids, 32 Endoplasmic reticulum (ER), 114

F FADD, 218 FLICE-inhibitory proteins (FLIPs) apoptosis, 220–223 gammaherpesvirus, 223–224 IRF3 activation and, 232–235 and NF-κB activation, 226–232 procaspase-8, 218–219 signaling events, 216–226 Fluorescence resonance energy transfer (FRET), 75–76 FRET-based real-time PCR, 211 Fuselloviridae, 43–44

H Haloarcula hispanica icosahedral virus 2 (HHIV-2), 4t Haloarcula hispanica pleomorphic virus 1 (HHPV-1), 4t, 37–38 Halogeometricum pleomorphic virus 1 (HGPV-1), 4t, 23, 38 Halorubrum pleomorphic virus (HRPV) type 1, 4t, 37, 38 type 2, 4t type 3, 4t, 38 type 6, 4t

255

Index

HCPro protein, 103–105, 106–113, 108t aphid proteins interacted with, 140 heterologous potyvirus particles, 139–140 N-terminal domain, 138–139 in potyvirus multiplication, 121–122 PVA, 131 receptor of, 140 and RNA silencing, 152–155 structure modeling, 151–152 symptomatology, 156 Helical symmetry, prokaryotic viruses, 40–43 Helical viruses, membrane envelope, 40–43 Helper Component (HC), 107, 138–139 Herpes simplex virus (HSV), 65, 66 TLR9 and, 71–72, 73–74 type 1, 88–89 type 2, 88–89 Herpesviridae family, 64–65 His virus 1 (His1), 4t, 23, 43–45, 44f His virus 2 (His2), 4t, 23 Homologous proteins, 103–105 Host cell response, to MCV, 207–208 Host virus, pathosystems, 161–163 Human foreskin fibroblasts (HFFs), 79

I Icosahedral viruses with inner membrane, 24–32 phage φ6, 32–35 IL-18-binding protein, 235–236 I-MC. See Inflammatory MC (I-MC) Imiquimod application, 211–212 Immune evasion mechanisms, MCV, 214–216, 215t Immune responses, MCV, 208–209 Immunity, alphaherpesviruses, 66–67 Immunohistochemical staining, 206–207 Infected cell protein (ICP) ICP0, 86–87 ICP27, 87–88 ICP34.5, 87 Inflammatory MC (I-MC), 208–209 Innate DNA sensing, 67–70 Inner membrane, icosahedral viruses, 24–32

Interferon-inducible protein 16 (IFI16) DNA sensors, 78–81 RNA interference, 80 Interferon regulatory factor 3 (IRF3) inhibition, 75 cFLIP and, 234–235 in fibroblasts, 76 and FLIPs, 232–235 MC159 and, 233–234 MC160 and, 234 Interferons (IFNs), 66–67 IFNβ, 207–208 type I, 207–208 Intracellular DNA sensors, 75 Intracellular movements, 128–132 Ipomoviruses, 103–105

J JPN1, 163–164

K 6K1, 114–115 6K2, 116–117, 120–121 Kaposi’s sarcoma herpesvirus (KSHV), 218–219, 223–224 Keratin13, 223–224, 229–231 Keratin14, 206–207 Keratin16, 206–207 Keratinocyte differentiation, 205–206 KHP30, 4t Knockout (KO) mutant, 124

L L2, 4t L17, 4t L172, 4t, 127 Lemon-shaped viruses, 43–46 Lettuce mosaic virus (LMV), 148–149 Linear ubiquitin chain assembly complex (LUBAC), 216–218 Lipid acquisition, 22–23 Lipid-containing archaeal viruses, 24–47 Lipid-containing bacterial viruses, 24–47 Lipid-containing phages, 2 Lipid-containing prokaryotic viruses, 2, 3–18, 4t and dsDNA, 19f, 20–21 and dsRNA, 19f, 20–21

256 Lipid-containing prokaryotic viruses (Continued ) with external membrane, 20–22 function and significance, 18–23 and icosahedral capsid, 22–23 NC surface protein, 19f origin and detection, 3–18 structural protein modifications, 23 Lipothrixviridae, 40, 42–43 Lipothrixviruses, 40–43, 42f LUBAC. See Linear ubiquitin chain assembly complex (LUBAC)

M Macrosiphum euphorbiae, 141–142 Major capsid protein (MCP), 22–23 MATH domain-containing proteins, 137 MBs. See Molluscum bodies (MBs) MC. See Molluscum contagiosum (MC) MC159 and apoptosis, 220–222 and IRF3 inhibition, 233–234 NF-κB activation and, 226–228 MC160 and apoptosis, 222–223 and IRF3 inhibition, 234 NF-κB activation and, 228–229 mCherry, 120–121 MCV. See Molluscum contagiosum virus (MCV) MEFs. See Murine embryo fibroblasts (MEFs) Membrane-anchored unique packaging vertex, 32 Membrane-containing procapsids, 32 Membrane envelope, 40–43 Membranous tunneling nanotube, 29–31 Methyl ester of phosphatidylglycerol (PGPMe), 17 Microtubule organizing center (MTOC), 64f Molluscipoxvirus genus, 202–203, 204 Molluscum bodies (MBs), 205–206 Molluscum contagiosum (MC) MC007, 237 MC54, 235–236 MC66, 237 MC148, 236–237

Index

Molluscum contagiosum virus (MCV), 202–203 acquired immunity, 209 characteristics, 203–204 diagnosis and treatment, 211–212 electron microscopy, 205f epidemiology, 209–210 host cell response, 207–208 host range, 214 immune evasion mechanisms, 214–216, 215t immune responses, 208–209 incidence tracking, 210 infection rates, 210 inhibition of apoptosis, 220–222 initial stages, 206–207 lesion characterization, 206–208 lesion development, 204–206 maturation, 205–206 MC007, 237 MC54, 235–236 MC66, 237 MC148, 236–237 MC159, 216, 220–222 MC160, 216, 222–223 morphogenesis, 205–206, 213–214 and PCR amplification, 212–213 replication, 203–204, 212–213 subtypes, 203 in tissue culture systems, 212–214 transmission, 205 VACV and, 203, 204 Monkeypox virus (MPX), 202 Murine cDCs, 72 Murine embryo fibroblasts (MEFs), 207–208 Myristoylation, 23 Myzus persicae, 141–142

N Natural killer (NK) cells, 67 NBS-LRR genes, 152 NF-κB activation cFLIPs and, 231–232 FLIPs and, 226–232 K13vFLIP, 229–231 and MC159, 226–228

Index

and MC160, 228–229 TLR9 signaling, 73–74 TNFR1, 216–218 NIa, 116–117 NIaPro, 108t, 117, 165–166 NIb, 108t, 117–118 Nicotiana benthamiana, 114–115, 153 gene expression profiles, 164 and TuMV, 141–142 Nicotiana glutinosa, 153 Nicotiana tabacum, 145–146 NI-MC. See Noninflammatory MC (NI-MC) NOD-like receptor (NLR), 77 50 -noncoding region (NCR), 103–105 Noncolonizer aphid species, 141–142 Noninflammatory MC (NI-MC), 208–209 N-terminal domain, 113 of CP, 118 of HCPro, 138–139 N-terminal sequence of 6K2 (6K2-N), 135–136 Nucleocapsid (NC) surface protein, 19f helical, 46–47 lipid envelope, 32–33 P8, 33–34 P9, 34–35

O Open reading frame (ORF), 103–105 ORF71, 224 PISPO, 107 virus, 204 Orthopoxvirus genus, 202–203

P Palmitoylation, 23 Papaya ringspot virus (PRSV), 146 Pathogen-associated molecular pattern (PAMP), 67–68 Pathosystems assayed, 163–164 host/potyvirus, 161–163 plant/potyvirus interactions in, 144–164

257 Pattern recognition receptors (PRRs), 67–68 PCaP1 functional relevance of, 130–131 PIPO domain with, 131 PCR amplification, 211, 212–213 Pea seed-borne mosaic virus (PSbMV), 132 PH1, 4t φNS11, 4t φ6, 4t host, 35 and icosahedral viruses, 32–35 involvement, 33–34 and membrane envelope, 34–35 φ2954, 4t φ7-φ14, 4t Pinellia ternata, 114–115 PIPO, 114–115 PISPO, 107 Pisum sativum, 146–147 Plant–potyvirus confrontation, 152 Plant virus, 144–164 Plasmacytoid DCs (pDCs), 71–72, 84 Pleolipoviruses, 36f Pleomorphic viruses prokaryotic, 36–37 vesicular, 35–39 Plum pox virus (PPV), 114–115 Pisum sativum, 146–147 Prunus, 137 PM2, 4t, 24–25 DNA, 27 and dsDNA genome, 25–27 maturation, 27 P3N-PIPO, 108t Poliovirus, 23 Poly(A)-binding protein 2 (PABP2), 122–123 Polymerase complex (PC), 32–33 Potato virus A (PVA), 118–119, 131 HCPro, 131 systemic infection, 134–135 Potato virus Y (PVY), 131, 151–152 Potyviral long-distance movement host factors involved in, 136–138 in phloem cells, 132–133 in vascular system, 133–134 viral determinants involved in, 134–136

258 Potyvirus, 102 biological and biochemical features, 105–119, 106f biotechnological applications, 103, 165–166 CI, 108t cloning capacity, 165 evolutionary abilities of, 145–152 in genome structures, 103–105 genomic maps, 104f genomic RNAs, 103–105 interactions in pathosystems, 144–164 P1, 103–105, 106–107, 108t P3, 108t pathosystems, 161–163 in planta interactions, 102 P3N-PIPO, 108t production, 102 proteins, 108t in situ hybridization, 161–163 symptomatology, 155–164 terminal protein, 104f VPg, 108t, 134–135 Potyvirus movement cell-to-cell movements, 128–132 intracellular movements, 128–132 long-distance (see Potyviral long-distance movement) Potyvirus multiplication HCPro in, 121–122 P1 protein for, 122 putative functions of, 125–128 subcellular localization of, 119–121 viral and plant factors in, 121–125 Potyvirus transmission acquisition, 141 by aphids, 138–142 seed transmission, 142–144 Potyvirus VPg-interacting protein (PVIP), 132 Poxviridae family, 202 Poxvirus DNA replication, 212–213 PR3, 4t PR4, 4t PR5, 4t PR722, 4t pRb-binding protein, 237

Index

PRD1, 2, 4t assembly and packaging, 31–32 cellular tomography, 30 form icosahedrally ordered membrane, 28–29 lipids in, 27–28 and membranous tunneling nanotube, 29–31 Procaspase-8, 218–219 Prokaryotic pleomorphic viruses, 36–37 Prokaryotic viruses, 2 with helical symmetry, 40–43 lipid membranes of, 2 lipids in (see Lipid-containing prokaryotic viruses) morphotypes of, 3f Protein P1, 103–105, 106–107, 108t, 122 P3, 108t, 114–115, 150–151 P10, 31 P16, 28–29 P17, 31 P23-77, 4t P30, 31 P37-14, 4t Prunus, 137, 146–147 PRV, 66 PSbMV, seed transmission, 142–144 PSI-K expression, 152 PTK motif, 139 pUS3 kinase, 87 Putative functions, potyvirus multiplication, 125–128 PYHIN, 77 Pyrobaculum spherical virus (PSV), 46 type1, 4t Pyrococcus abyssi virus 1 (PAV1), 4t, 44–45

R Reflectance confocal microscopy, 211 Restriction to TEV Movement (RTM), 136–137 type 1, 136–137 type 2, 136–137 type 3, 136–137 Rhopalosiphum maidis, 141–142 Ribonucleoprotein (RNP) complex, 125

259

Index

RIG-I, 83–84 RIP1, 216–218 RNA Pol III, 83–84 RNA silencing DCL3 and RDR6, 143–144 HCPro and, 152–155 suppression, 106–107 RNA synthesis, 125–126 Rudiviridae, 40

S Salisaeta icosahedral phage 1 (SSIP-1), 4t Salterprovirus, 43–44 Seed transmission of plant viruses, 142–144 in reciprocal cross-pollination, 143 of SMV, 143–144 SH1, 4t SHA3, 137 Signaling events, TNFR1, 216–226, 217f Size-exclusion limit (SEL), 129 Small RNA binding, 155 Sn-glycerol-1-phosphate (G1P), 3–17 Sn-glycerol-3-phosphate (G3P), 3–17 SNJ1, 4t Soybean mosaic virus (SMV), 114–115, 143–144 Spindle-shaped virus His1, 44f Stimulator of interferon genes (STING), 68–70, 82 Subcellular localization, 119–121 Sulfolobus, 40 Sulfolobus islandicus filamentous virus 1 (SIFV), 4t, 41–42 Sulfolobus spindle-shaped virus (SSV), 4t, 43–44 Sulfolobus spindle-shaped virus Kamchatka1 (SSVk1), 4t Sulfolobus spindle-shaped virus Ragged hills (SSVrh), 4t Sulfolobus tengchongensis spindle-shaped virus (STSV) type 1, 4t, 45 type 2, 4t, 45 Sulfolobus turreted icosahedral virus (STIV), 4t, 22–23, 25 type 2, 4t, 25

Sunflower chlorotic mottle virus (SuCMoV), 159–160 Symptomatology, 155–164 Syp71, 124

T TANK-binding kinase 1 (TBK1), 88 Tectiviruses, 27–28 TEV-At17, 161–163 Thermococcus prieurii virus 1 (TPV1), 4t Thermoproteus, 40 Thermoproteus tenax spherical virus 1 (TTSV1), 4t, 46–47 Thermoproteus tenax virus (TTV) type 1, 4t, 41 type 2, 4t, 41–42 type 3, 4t, 41–42 Tissue culture systems, 212–214 TNF. See Tumor necrosis factor (TNF) TNFR1. See Tumor necrosis factor receptor 1 (TNFR1) Tobacco etch virus (TEV), 120, 145–146 Tobacco vein mottling virus (TVMV), 141 Toll-like receptor 9 (TLR9), 68–70 deficiency in mice, 73–74 DNA sensors, 70–74 and HSV, 71–72, 73–74 inflammatory cytokines, 72 pDCs, 84 Toll-like receptors (TLRs), 67–68, 207–208 TRAF6, 87 Transmission electron microscopy (TEM), 27 Tumor necrosis factor (TNF), 207–208 Tumor necrosis factor receptor 1 (TNFR1), 216–226, 217f Turnip mosaic virus (TuMV), 113, 124 Arabidopsis thaliana, 161–163, 164 host adaptation, 146 Nicotiana benthamiana, 141–142 P3N-PIPO, 130–131 VPg, 126–127

U UK1, 163–164

260

V Vaccine design, 88–90 Vaccinia virus (VACV), 202 activating EGFR, 206–207 in cytoplasm, 212 and MCV, 203, 204 in nucleotide biosynthesis, 213–214 Vaccinia virus growth factor (VGF), 206–207 VACV. See Vaccinia virus (VACV) Vap27-1, 123 Varicella zoster virus (VZV), 65 Variola virus (VAR), 202 Vesicular pleomorphic viruses, 35–39 vFLIP. See Viral FLIP (vFLIP) VGF. See Vaccinia virus growth factor (VGF) viral cDNA, 102 Viral chemokine, 236–237 Viral DNA, 84–85 Viral FLIP (vFLIP), 216, 229–231 Viral replication, 71–72

Index

Viral RNA, 127, 132–133 Virion morphology, 38 Viruses adaptation to host, 147 asymmetric lipid vesicles as, 37–38 with one short tail, 43–45 with one/two long tails, 45–46 Virus-like particles (VLPs), 43 VPg, 108t, 116 interaction, 147–149 in long-distance movement, 134–135 mutations, 127 of potyviruses, 134–135 in viral translation, 126–127 in vivo evidence, 127

W Wheat streak mosaic virus (WSMV), 115 Wip1, 4t

Z Zucchini yellow mosaic virus (ZYMV), 140