Genes, Genetics and Transgenics for Virus Resistance in Plants [1 ed.] 9781910190821, 9781910190814

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Genes, Genetics and Transgenics for Virus Resistance in Plants

Edited by

Basavaprabhu L. Patil

Caister Academic Press

Genes, Genetics and Transgenics for Virus Resistance in Plants https://doi.org/10.21775/9781910190814

Edited by Basavaprabhu L. Patil ICAR-National Research Centre on Plant Biotechnology (NRCPB) LBS centre, IARI, Pusa campus New Delhi, India

Caister Academic Press

Copyright © 2018 Caister Academic Press Norfolk, UK www.caister.com British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN: 978-1-910190-81-4 (paperback) ISBN: 978-1-910190-82-1 (ebook) Description or mention of instrumentation, software, or other products in this book does not imply endorsement by the author or publisher. The author and publisher do not assume responsibility for the validity of any products or procedures mentioned or described in this book or for the consequences of their use. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without the prior permission of the publisher. No claim to original U.S. Government works. Cover design adapted from images provided by Basavaprabhu L. Patil. Ebooks Ebooks supplied to individuals are single-user only and must not be reproduced, copied, stored in a retrieval system, or distributed by any means, electronic, mechanical, photocopying, email, internet or otherwise. Ebooks supplied to academic libraries, corporations, government organizations, public libraries, and school libraries are subject to the terms and conditions specified by the supplier.

Contents

Forewordv Prefacevii 1

Mechanisms of Virus Resistance in Plants

2

Role of Host Transcription Factors in Modulating Defense Response During Plant–Virus Interaction

25

3

Surfacing the Role of Epigenetics in Host–Virus Interaction

55

4

Molecular Markers as Tools for Identification and Introgression of Virus-Resistant Genes

87

M. E. Chrissie Rey and Vincent N. Fondong

Saurabh Pandey, Pranav P. Sahu, Ritika Kulshreshtha and Manoj Prasad

Namisha Sharma, Pranav P. Sahu, Ritika Kulshreshtha and Manoj Prasad

Mamta Sharma, Avijit Tarafdar, U. S. Sharath Chandran, Devashish R. Chobe and Raju Ghosh

1

5

Genetic Engineering for Virus Resistance in Plants: Principles and Methods

103

6

Tools and Techniques for Production of Double-stranded RNA and its Application for Management of Plant Viral Diseases

119

7

Transgenic Virus-Resistant Papaya: Current Status and Future Trends

141

8

Development and Delivery of Transgenic Virus-resistant Cassava in East Africa

159

9

Viruses Infecting Rice and their Transgenic Control

177

Whitefly-transmitted Begomoviruses and Advances in the Control of their Vectors

201

Basavaprabhu L. Patil

Andreas E. Voloudakis, Maria C. Holeva, Athanasios Kaldis and Dongho Kim

10

Gustavo Fermin, Paula Tennant and Sudeshna Mazumdar-Leighton Henry Wagaba, Andrew Kiggundu and Nigel Taylor

Gaurav Kumar, Shweta Sharma and Indranil Dasgupta

Surapathrudu Kanakala and Murad Ghanim

iv  | Contents

11

Virus-resistant Transgenic Tomato: Current Status and Future Prospects

221

12

Management of Geminiviruses Focusing on Small RNAs in Tomato

235

13

Viruses Infecting Banana and their Transgenic Management

255

14

Virus-induced Gene Silencing (VIGS) and its Applications

277

15

Possible Strategies for Establishment of VIGS Protocol in Chickpea

329

S.V. Ramesh and Shelly Praveen

Archana Singh and Sunil Kumar Mukherjee

Ramasamy Selvarajan, Chelliah Anuradha, Velusamy Balasubramanian, Sivalingam Elayabalan and Kanicheluam Prasanya Selvam Deep Ratan Kumar, Tejbhan Saini and Radhamani Anandalakshmi Ranjita Sinha and Muthappa Senthil-Kumar

Index345

Foreword

In 1986, the first report on the use of genetic engineering to control Tobacco mosaic virus in tobacco plants was published and it opened the gate to a flood of publications for the possible control of many viruses in many hosts. At that time, this concept of engineering virus resistance was a breakthrough. Controlling plant viruses has always been a big challenge to breeders, and suddenly it was possible to control almost any virus in any crop through genetic engineering! Evidently, over the years many natural sources of resistance for numerous plant viruses had been identified; however, combining these resistance sources with other traits was always a challenge to breeders. Therefore, genetic engineering appeared as the solution to control plant viruses! But three decades later, we have to acknowledge that we have not seen the expected revolution in farmers’ fields. On the contrary, we have seen the emergence and outbreak of many new and known plant viral diseases, threatening food security. Even though some plants have been engineered with multiple virus resistance, they have never been commercialized. The engineered papaya with immunity to Papaya ringspot virus remains the most successful example of commercialization of virus-resistant transgenics. The failure to commercialize virus-resistant transgenics is the result not of technical or scientific problems or any sort of biological barrier, but mostly of political pressures from so-called ecological groups. In the meantime, improved technologies were developed and transferred to new crops and novel viruses, and in some instances made real scientific breakthroughs. However, the vested interests of these groups of fanatics, with a false claim of saving the earth’s ecology, raised biosafety standards to the point where only large

multinationals could afford them, which de facto prevented the application of these technologies to many important food crops in the world. In the beginning of the twenty-first century, the genomic revolution brought new hopes to control plant viruses by harnessing natural genes of resistance. Whole-genome sequencing of many plants, along with scores of novel DNA technologies, facilitated the use of modern tools in gene discovery for virus resistance. However, the introgression of these resistant loci was restricted due to their multigenic or recessive nature, making it difficult to transfer them to a suitable genetic background without using genetic engineering technologies. Recently, the discovery and use of gene-editing technologies such as CRISPR/Cas9 and TALENs may now allow plant virologists, genomics experts and breeders to work together for a breakthrough in controlling plant viruses. This can be a reality only if these technologies are not considered to be GM- technologies by policy makers. This book, Genes, Genetics and Transgenics for Virus Resistance in Plants, provides a very nice update on the status of current knowledge on the use of genetic engineering and other biotechnological strategies for the control of plant viruses. It is hoped that this information will be used in conjunction with the latest gene technologies to achieve the urgently needed scientific breakthroughs for the successful control of plant viruses, ultimately for the benefit of humankind. Claude M. Fauquet Director, Global Cassava Partnership for the 21st Century (GCP-21) International Center for Tropical Agriculture (CIAT), Cali, Colombia

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Preface

Viral diseases of crop plants cause significant yield losses, which is a major threat to global food security. Unlike other pests and pathogens, the only remedy available for control of plant viral diseases is through introgression of resistance trait, either through conventional breeding or through genetic engineering. Availability of few natural sources of virus resistance has hampered development of virus-resistant crop plants through conventional crop improvement methods. Thus genetic engineering for virus resistance is the sole option available for effective management of viral diseases. Since the first report on transgenic virus resistance in tobacco in 1986, huge progress has been made in our understanding of the molecular basis of virus resistance, complimented by the significant improvement in the tools and techniques used for genetic engineering. Despite major advancements in plant genomics and transgenics, there has been no commercialization of virus-resistant transgenic crops, except transgenic papaya. Thus, to provide an up-to-date reference book on genes, genetics and transgenics for virus resistance in plants for students, faculties and researchers, here we have compiled 15 diverse chapters. In the first chapter the

current knowledge on mechanisms of virus resistance in plants is discussed, followed by a chapter on the role of host transcription factors in modulating defence response during plant-virus interaction, and a chapter on the role of epigenetics in host-virus interactions. There is a chapter on how molecular markers could be employed as tools for identification and introgression of virus-resistant genes. This book also thoroughly discusses the principles and methods involved in the genetic engineering for virus resistance in plants. The book also elaborates on topical application of double-stranded RNA for control of plant viral diseases, without having to develop transgenic plants. Further, the book deals with individual crops such as papaya, cassava, rice, tomato, and banana, for which virus resistance has been accomplished by employing different transgenic technologies. The management of whitefly-transmitted begomoviruses and advances in the control of their vectors is also covered as an independent chapter. Virus-induced gene silencing (VIGS), another frontier area of research in which virus-derived silencing vectors are extensively used in gene function studies and functional genomics, is also discussed elaborately.

Mechanisms of Virus Resistance in Plants M. E. Chrissie Rey1* and Vincent N. Fondong2

1

1School of Molecular and Cell Biology, University of the Witwatersrand, Johannesburg, South Africa. 2Department of Biological Sciences, Delaware State University, Dover, DE, USA.

*Correspondence: [email protected] https://doi.org/10.21775/9781910190814.01

Abstract Considerable advances have been made in the past decade in understanding the mechanisms of virus resistance in plants. In particular, RNA silencing, resistance (R) protein-associated effector-triggered immunity (ETI), and recessive resistance (impaired susceptibility), have been at the forefront. While conserved pathogen associated molecular patterns (PAMPs) have generally not been identified in viruses, it is highly likely that virus-encoded glycoproteins or proteins such as the coat protein, and ribonucleoprotein complexes are analogous to PAMPs and can trigger pathogen triggered immunity (PTI)-like responses. More recently, the role of the chloroplast, and its signalling with the nucleus and mitochondrion, has attracted attention in virus resistance studies. Developments in highthroughput DNA and RNA sequencing, as well as protein and metabolite technologies, have enabled unravelling of defence responses to virus at a global level, leading to the current belief that resistance mechanisms form a complex of interacting networks. Unravelling these networks will provide opportunities for the next generation of new strategies for plant virus resistance engineering. Introduction Plant viruses cause physiological disorders that result in plant diseases that may be of significant economic and agronomic significance. These viral diseases reduce both quality and yield, with

annual losses estimated to be more than $30 billion worldwide (Sastry and Zitter, 2014). Cassava mosaic geminiviruses (CMGs) alone, for example, cause more than 25 million tons of losses annually in the cassava crop (Manihot esculenta Crantz) in Africa, India and Sri Lanka (Thresh and Cooter, 2005). Climate change, as well as increased agricultural intensification, has played a role in the increased spread of plant viral pathogens (Nicaise, 2014). Several natural resistance mechanisms that offer protection against viral pathogens exist in plants (Maule et al., 2009; Mandidi and Scholthof, 2013). In plants, the first line of defence against pathogens is pathogen-triggered immunity (PTI), which is mediated by host transmembrane pattern recognition receptors that detect a broad range of pathogen-associated molecular patterns (PAMPs) (Wang et al., 2012; Zvereva et al., 2012; Seo et al., 2013). To date, no conserved PAMPs have been identified in virus infections, and primary basal defence is largely based on endogenous RNA silencing. RNA silencing is an evolutionarily conserved mechanism where the plant host targets invading nucleic acids, including viruses. Defence responses in plants are complex and include many levels, notably immunity, partial resistance, quantitative resistance, generalized resistance, mature-plant resistance, durable resistance, and tolerance. Plants exhibiting immunity are unable to support virus replication and lack virus symptoms (Lecoq et al., 2004), while resistant plants limit virus multiplication by interfering

2  | Rey and Fondong

with the replication cycles, thus displaying different levels of resistance (Fraile and Garcia-Arenal, 2010). Plants may exhibit tolerance, where they sustain virus replication, but with no apparent symptoms. Plants displaying virus tolerance may exhibit low levels of symptoms throughout the lifespan of the plant, compared with susceptible plants (Bengyella et al., 2015). Resistance to viruses is not always genetically predetermined and can be highly adaptive in nature. This is the case with resistance based on RNA silencing, which appears to play key roles in both induced and basal resistance to viruses. The last decades have seen substantial advances in the molecular characterization of interactions between viral pathogens and their plant hosts, and this has resulted in the discovery of novel strategies that are being exploited in crop improvement efforts. Thus, a good understanding of plant defence mechanisms could lead to the development of effective ways to generate virus-resistant plants. In this chapter, we explore current understanding of mechanisms governing plant virus resistance. RNA silencing: a natural virus resistance mechanism in plants Antiviral post-transcriptional gene silencing In plants, as in other eukaryotic organisms, RNA silencing plays key regulatory roles in various biological processes, including plant growth and development, and host resistance to pathogen attack. Plant antiviral RNA silencing is triggered by highly structured or double-stranded (ds) viral RNAs that are recognized and processed by host Dicer-Like (DCL) proteins into primary 21–24 nt virus-derived small interfering RNAs (vsiRNAs), which associate with ARGONAUTE (AGO) effector proteins to target viral complementary RNA or DNA. The critical step in RNA silencing antiviral response is thus production of vsiRNAs (Zhang et al., 2015). In RNA viruses, vsiRNAs have been suggested to be processed from highly structured genomic RNA (Molnár et al., 2005; Koukiekolo et al., 2009; Zhang et al., 2015) and/or from replication intermediates (RIs). Interestingly, as illustrated in Cymbidium ring spot virus, vsiRNAs derived from the positive strand of the viral dsRNA far outnumber those derived from the negative strand,

suggesting that for this virus and likely many others (Molnár et al., 2005; Ho et al., 2006; Donaire et al., 2009), vsiRNAs originate mainly from structured RNAs than from dsRNAs. Indeed, several recent studies have revealed that there are hotspots in viral genomes that form regulatory stem–loop structures for vsiRNAs production (Xu et al., 2012; Vives et al., 2013; Visser et al., 2014). It has also been shown that host-encoded RNA-dependent RNA polymerases (RDRs) are required for the production of vsiRNAs in some RNA viruses, suggesting that vsiRNAs biogenesis is likely similar to biogenesis of endogenous siRNA (Qi et al., 2009; Garcia-Ruiz et al., 2010; Wang et al., 2010, 2012). Viral dsRNAs structures have been designated as virus-associated molecular pattern (VAMP; a form of PAMP) (Ruiz-Ferrer and Voinnet, 2009) and are processed by DCL proteins into vsiRNAs of different lengths (DCL4: 21 nt, DCL2: 22 nt, DCL3: 24 nt) (Deleris et al., 2006). Although both DCL4 and DCL2 play essential roles in plant defence against viruses, DCL4 has a more dominant role since 21 nt siRNAs are usually more abundant than 22 nt siRNAs following RNA virus infection, and loss-offunction mutations in DCL4 have a stronger impact on viral resistance than mutations in DCL2 (Wang et al., 2012). It is however important to note that the functional predominance of DCL4 over DCL2 could be partly due to the more potent antiviral silencing effect of 21 nt siRNAs, compared with the 22 nt species (Wang et al., 2011). In DNA viruses, vsiRNAs formation has been suggested to originate from complementarity of overlapping transcripts synthesized by bidirectional transcription that creates regions of dsRNAs that serve as templates for vsiRNAs production (Chellappan et al., 2004; Blevins et al., 2011; Aregger et al., 2012). Also, similar to RNA virus vsiRNA biogenesis, DNA virus vsiRNA production could be through folding of viral RNA transcripts into stem–loop structures (Vanitharani et al., 2005; Wang et al., 2012). All four DCLs have been found to contribute to the production of 21–24 nt vsiRNAs in the interactions between A. thaliana and cauliflower mosaic virus (CaMV), a dsDNA virus or cabbage leaf curl virus (CaLCuV), a ssDNA virus (Moissiard and Voinnet, 2006; Blevins et al., 2011; Aregger et al., 2012). The next step in vsiRNA processing involves AGO proteins, which are essential in antiviral defence against both RNA and DNA viruses.

Mechanisms of Virus Resistance in Plants |  3

Indeed, AGO1, AGO2, AGO4, AGO5, AGO7 and AGO10 are known to display antiviral activity in Arabidopsis, and AGO1 and AGO18 play antiviral defence roles in rice (reviewed in Carbonell and Carrington, 2015; Calil and Fontes, 2017). Only one strand of the vsiRNA duplex, processed by DCL proteins, associates with a member of the AGO protein family, the core component of the RISC (reviewed in Mallory and Vaucheret, 2010). Once loaded into the RISC, vsiRNA directs the complex to target and silence cognate viral RNAs. It has been shown that AGO1 is the dominant AGO in vsiRNA-directed antiviral defence even though AGO7 and AGO2 have also been found to play a role (Harvey et al., 2011; Qu et al., 2008; Jaubert et al., 2011; Wang et al., 2011). Recent evidence has demonstrated that, alternatively to the cleavage of viral RNA targets, host cells can suppress viral protein translation to silence viral RNA. The next level of vsiRNA processing involves RDRs, which synthesize cleaved RNA transcripts into dsRNAs for secondary vsiRNA processing. For example, biogenesis of tobacco rattle virus (TRV) vsiRNAs has been found to combine the activity of RDR1, RDR2 and RDR6 (Donaire et al., 2008). Correspondingly, knockdown of RDR6 rendered rice hypersusceptible to Rice stripe virus (RSV) with increased accumulation of RSV genomic RNA and reduced RSV vsiRNA accumulation ( Jiang et al., 2012). Similar observations were made in rice infected by rice dwarf virus (RDV) (Hong et al., 2015). Antiviral transcriptional gene silencing It has been shown that DNA viruses induce transcriptional gene silencing (TGS), leading to suppression of viral gene expression at the level of transcription in the nucleus (Kanazawa et al., 2011). This process is due to viral DNA methylation, which is mediated by vsiRNAs (Vanitharani et al., 2005; Moissiard and Voinnet, 2006). DNA methylation at gene promoters can be triggered by dsRNAs through the RNA-directed DNA methylation (RdDM) pathway. RdDM requires DCL3 protein, which processes viral dsRNAs into 24 nt vsiRNAs, the latter then guide AGO4 for de novo methylation of viral DNA (Chan et al., 2004; Xie et al., 2004; Henderson et al., 2006). It was reported, for example, that Arabidopsis plants defective in

methyltransferase or related cofactor activity are hypersensitive to geminivirus infection, suggesting that the viral genome is targeted by RdDM (Raja et al., 2008). Evidence that DLC3-dependent 24 nt vsiRNAs accumulate in plants infected with caulimoviruses and geminiviruses (Moissiard and Voinnet, 2006; Blevins et al., 2011) shows the important role played by the RdDM pathway in silencing DNA viruses. Indeed, DRB3, AGO4, Pol IV, Pol V, DRM1 and DRM2, all hallmarks of DNA methylation, have been implicated in the antiviral mechanism against geminiviruses CaLCuV and beet curly top virus (BCTV) (Raja et al., 2008; Raja et al., 2014). However, a separate study found no significant effect of AGO4, Pol IV or Pol V on the accumulation of CalCuV or CaMV (Blevins et al., 2011). Thus, our understanding of the specific requirements for antiviral RdDM is still incomplete. Viral suppressors of RNA silencing Many plant viruses are now known to encode proteins that counter RNA silencing – the so-called viral suppressors of RNA silencing (VSR). A comprehensive review on the current knowledge of the diverse VSRs and their strategies to suppress vsiRNAs biogenesis is available in Csorba et al. (2015). These VSRs target different steps in the RNA silencing pathway. For example, Rice yellow mottle virus P protein inhibits vsiRNA processing by blocking DCL function (Lacombe et al., 2010) and CaMV transactivator protein P6 interacts directly with DCL4 cofactor, DRB4, to block DCL cleavage of dsRNA (Haas et al., 2015). Also, rice yellow stunt virus (RYSV) protein P6 interacts directly with RDR6 to block secondary vsiRNA synthesis (Guo et al., 2013) and the V2 protein of the ssDNA virus, tomato yellow leaf curl virus (TYLCV) interacts with SGS3, a cofactor of RDR6, to inhibit synthesis of dsRNA substrate (Glick et al., 2008). The potyvirus helper component proteinase (HcPro), which is one of the best-characterized VSR, has been shown to interact with RAV2 (the ethylene-inducible transcription factor) to downregulate RDR6 (Zhang et al., 2015). Some VSRs bind to long dsRNA or siRNA duplexes to inhibit siRNA biogenesis or RISC formation as found in Tomato chlorosis virus p22, which binds to long dsRNAs to inhibit cleavage by RISC (Landeo-Rios et al., 2016). Also, the tombusvirus p19 protein binds to vsiRNA duplexes, thus preventing vsiRNA

4  | Rey and Fondong

loading into the AGO effector proteins and assembly of RISC (Silhavy et al., 2002). Correspondingly, tobacco mosaic virus (TMV) P126 and potyvirus HcPro were found to destabilize vsiRNAs by interacting and inhibiting HEN1-mediated methylation of vsiRNAs (Vogler et al., 2007). In a very direct strategy, Sweet potato chlorotic stunt virus RNA endoribonuclease III (RNase III) cleaves siRNA duplexes into approximately 14-bp products that are inactive in mediating silencing (Cuellar et al., 2009). Many VSRs target proteins involved in the effector steps of RNA silencing. For instance, it was shown that cucumber mosaic virus (CMV) Fny strain 2b protein interacts with AGO1 and inhibits its activity in RISC (Zhang et al., 2006) and Sugarcane yellow leaf virus P0 protein mediates the degradation of AGO1 (Pazhouhandeh et al., 2006; Baumberger et al., 2007). Furthermore, Turnip crinkle virus P38, has been reported to use its encoded glycine/tryptophan (GW) motifs to mimic GW motif-containing cellular proteins that are known to be important partners of AGOs, thereby attracting and disarming host AGO proteins and the formation of RISC ( Jin and Zhu, 2010). Accordingly, the versatile HcPro plays multiple roles in vsiRNA biogenesis, such as vsiRNA duplex binding and interacting with HEN1 to block HEN1 methylation of vsiRNA duplex, both of which inhibit vsiRNA loading into RISC (Zhang et al., 2015). It has also been suggested that HcPro attenuates viral RNA translational repression through association with AGO1 and ribosomes (Ivanov et al., 2016). VSRs are also known to inhibit RNA silencing indirectly through interaction with miRNAs that regulate enzymes of the RNA silencing pathway; such is the case of the tombusvirus P19, which has been found to induce the expression of miR168 that targets AGO1 mRNA (Várallyay et al., 2010). This action results in reduced accumulation of AGO1 protein. Furthermore, miR168, unlike other endogenous miRNAs, is not efficiently bound by P19, resulting in the selective increased loading of miR168 into AGO1 and the related AGO10 (Várallyay et al., 2010). Because miR168 directly down-regulates AGO1 mRNA stability and translation as part of a regulatory feedback loop, the net effect of this selective binding process is a sharp reduction in cellular AGO1 levels in addition to the direct vsiRNA sequestration by P19 (Pumplin

and Voinnet, 2013). It has also been reported that plants that are infected by a tombusvirus exhibit increased transcription of MIR168, thus further increasing the levels of miR168 available for AGO1 down-regulation (Várallyay et al., 2010; Pumplin and Voinnet, 2013). It has been shown that some VSRs, especially those of DNA viruses, suppress TGS in plants, by reducing DNA methylation (Buchmann et al., 2009). For example, Tomato yellow leaf curl China virus βC1 protein was reported to interact with and block the activity of S-adenosyl homocysteine hydrolase, a methyl cycle enzyme required for cytosine methylation (Yang et al., 2011). Correspondingly, BCTV C2 protein attenuates the 26S proteasome-mediated degradation of S-adenosyl-methionine decarboxylase through physical interaction, resulting in inhibition of DNA methylation (Zhang et al., 2011). These studies provide evidence supporting the view that the RdDM plays an important role in plant defence against DNA viruses (Wang et al., 2012). Remarkably, the 2b protein of CMV, an RNA virus, also interferes with the RdDM pathway (Hamera et al., 2016). Together, these diverse modes of action by VSRs from different viruses suggest that they have evolved separately to suit specific virus–host interactions. Cross-protection Prior infection of a host by a protective mild strain of a virus can prevent subsequent infection of the host by a closely related strain or homologous virus (McKinney, 1929). This phenomenon is referred to as cross-protection and has been used to control especially diseases caused by RNA viruses such as Citrus tristeza virus (Muller and Cota, 1977), Papaya ringspot virus (Yeh and Gonsalves, 1984), Zucchini yellow mosaic virus (Lecoq et al., 1991), and TMV (Lu et al., 1998). The molecular mechanism underpinning cross-protection was shown to be protein- and/or RNA-based (Lin et al., 2007a). Thus, Pepper mild mottle virus (PMMoV) containing a mutated 126-kDa replicase, for example, was shown to display attenuated symptoms and could cross-protect pepper plants from subsequent PMMoV infection (Yoon et al., 2006). Correspondingly, the coat protein (CP) of TMV has been shown to cross-protect infection by a homologous virus, due presumably to the ability of the CP to inhibit

Mechanisms of Virus Resistance in Plants |  5

uncoating of the second strain (Beachy, 1999; Lin et al., 2007a). It is also possible that peptides of related viruses can interact with high affinity to the essential domains of host proteins to inhibit infection by related viruses, thereby providing cross-protection (Rudolph et al., 2003). For instance, the GAC mild strain of ZYMV that has two mutations in the conserved motifs of HcPro has been observed to cross-protect courgette squash against infection by the virulent strain (Lin et al., 2007b). Effector-triggered immunity and resistance genes The fundamental mechanisms of natural resistance in plants are well established. Essentially, upon attack, the plant recognizes pathogens through cell surface pattern recognition receptors (PRRs), which recognize pathogen-associated molecular patterns (PAMPs) and activate PAMP-triggered immunity (PTI), which can be suppressed by the pathogen (Bigeard et al., 2015). In turn, plants respond to PTI suppression by inducing effectortriggered immunity (ETI), in which dedicated plant resistance (R) proteins recognize highly specific effector or avirulence factors produced by microbes ( Jones and Dangl, 2006). Virus pathogens in turn produce virulence factors, called effectors, which suppress PTI (Shan et al., 2008). This host defence and pathogen counter-defence cycle is known as the co‑evolutionary arms race. A majority of the known R proteins belong either to the coiled-coil (CC)- or Toll/interleukin-1 receptor (TIR)- nucleotide-binding leucine-rich repeat (NLR) protein families (see review in Gururani et al., 2012; Calil and Fontes, 2017). Both the N and LRR terminus domains (CC or TIR) are critical for resistance responses. These domains often function indirectly through interactions with host accessory proteins to mediate recognition of virus effectors (Moffett, 2009). Generally, in plant–pathogen interactions, the immune responses downstream of R protein activation are associated with a range of signalling responses such as phytohormone induction, in particular salicylic acid (SA), reactive oxygen species (ROS) production, calcium ion influx, and mitogen-activated protein kinases (MAPK) activation. Massive global transcriptional and translational reprogramming occurs, including the induction of genes associated with defence

responses (see review in Calil and Fontes, 2017). Frequently, R protein activation leads to the induction of the hypersensitive response (HR) where infected adjacent cells undergo programmed cell death (PCD), confining the pathogen to the local site of infection. Concomitantly, with the induction of the local defence response, R protein activation also activates defence signalling at distal tissues of infection, referred to as systemic acquired resistance (SAR). Strikingly, as shown in Fig. 1.1, plants defend themselves against viruses through similar PTI pathways employed against non-viral pathogens. Thus, DCL perception and cleavage of viral dsRNA structures or virus-associated molecular patterns (VAMPs) is analogous to PRR recognition of PAMP, and it triggers the PTI pathways against the virus. In turn, successful viruses encode specialized effectors that are able to suppress PTI. To counter, specific R gene products interact with these effectors (playing the role of avirulence factors), leading to ETI (Fig. 1.2). This results in viral RNA degradation, a process that can be suppressed by VSRs; VRSs are thus the second class of virus-encoded effectors. It is important to note that the tobacco N gene that confers resistance to TMV was the first R gene to be identified (Whitham et al., 1994), and the Avr gene of TMV was found to be the replicase gene (Wang et al., 2012); many other R genes involved in antiviral resistance in plants have been identified (see review in Gouveia et al., 2017). Intriguingly, antiviral immune responses in plants have revealed roles for sRNAs in the regulation of both PTI and ETI signalling (Fei et al., 2013; Li et al., 2014). In dicots, for instance, most phased secondary siRNAs (phasiRNAs), whose biogenesis is triggered by specific miRNAs, are from families of disease resistance genes belonging to NLR family of proteins (Zhai et al., 2011; Fei et al., 2013; Arikit et al., 2014) (Fig. 1.1). A good illustration of the important role of miRNAs in NLR is the action of miR482/2118 superfamily, which targets the P-loop motif of NLR genes in tomato (Shivaprasad et al., 2012), cotton (Zhu et al., 2013), and cassava (Allie et al., 2014; Khatabi et al., 2016). Furthermore, two tobacco miRNAs, miR6019 and miR6020, guide sequence–specific cleavage of transcripts encoding the TIR-NLR immune receptor N, which confers resistance to TMV (Li et al., 2012). Also, several miRNA families that target NLRs in

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Figure 1.1  Effector-triggered immunity in virus–plant interactions based on the plant immune response models proposed by Jones and Dangl (2006) and Fei et al. (2016). Following virus entry into a plant cell, highly structured or double-stranded (ds) viral RNAs are recognized as a virus-associated molecular pattern (VAMP; a form of PAMP) by host DCLs, which process the dsRNAs into vsiRNAs; this constitutes PAMP-triggered immunity (PTI). Some viruses encode suppressors of RNA silencing (VSRs) or virulence factors, which block DCL activity, thus ensuring successful establishment of the virus. Other virus encoded effectors (e.g. effector1 and effector2) are produced and in turn are recognized by specific R gene products, which interact with these effectors (thus playing the role of avirulence factors), leading to effector triggered immunity (ETI), an amplified version of PTI that often passes a threshold for induction of HR. Continuous auxin immune signalling can be inhibited by miRNA-mediated processing of genes involved in the signalling pathway into phasiRNAs. Also, some miRNAs, including especially members of the miR482/miR2118 superfamily, can trigger phasiRNA biogenesis from R genes, and these phasiRNAs may function synergistically with miRNAs either in cis or trans to suppress R-gene transcript levels.

tomato infected by the geminivirus Tomato yellow leaf curl Sardinia virus have been reported (Miozzi et al., 2014). Currently, it is not clear whether phasiRNA targeting of NLRs is to inhibit the ability of NLR proteins to mount ETI responses or whether it is to prevent overaccumulation of NLRs that could result in constitutive defence pathway activation in the absence of the pathogen. Immune signalling molecules and signalling responses One of the earliest downstream signalling events following virus and other pathogen perceptions is calcium influx and reactive oxygen (ROS) or

reactive nitrogen (RNS) species production (Fig. 1.2), which act as secondary messengers (Boller and Felix, 2009; Sewelam et al., 2016). Subsequently, activation of mitogen-activated protein kinase (MAPK) pathways acts as a central hub of a complex network of defence responses, involving phytohormones, nitric oxide, and sugars (Meng and Zhang, 2013). Protein kinase (PK) and MAPK signalling networks serve specific and overlapping roles in controlling enzymes, hormones and transcription factors (TFs) (reviewed in Tena et al., 2011). It was shown, for example, that following SA-dependent defence, at least 49 WRKY genes were differentially expressed in Arabidopsis (Dong et al., 2003). It appears that certain TF families are

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Figure 1.2  A non-exhaustive overview of plant defence mechanisms induced by viruses. Viruses enter plant cells through wounding or insect-mediated transmission and yet to be identified extracellular host cell membrane pathogen recognition receptors (PRRs) likely recognize the virion or viral ribonucleoprotein (vRNP). Once in the cell, viral proteins, including the coat protein (CP), movement protein (MP) are synthesized. Additionally, viral RNA or mRNA can trigger post-transcriptional gene silencing (PTGS) whereby siRNAs target homologous RNA for degradation, or interfere with viral protein translation. DNA viruses that replicate in the nucleus may also be targeted by TGS. Viral proteins act as avirulence (avr) factors and are recognized by cytosolic resistance (R) proteins, such as nucleotide binding-leucine rich repeat (NLR) proteins, which trigger effector triggered immunity (ETI). Non-R-receptors may also trigger ETI. ETI is usually, but not exclusively, involved in localized defence [hypersensitive response (HR)] which is associated with programmed cell death and containment of the virus. ETI may lead to salicylic acid (SA)-associated systemic acquired resistance (SAR) throughout the plant. Defence responses are initiated by global transcriptome, proteome and metabolome re-programming, induced by receptor-virus component mediated secondary signalling, involving MAPK, hormone and reactive oxygen species and reactive nitrogen species (ROS/RNS) pathways. Re-programming is controlled by transcription factors (TFs) and miRNA-post-transcriptional regulation. Signal transduction to the nucleus and chloroplast may be regulated by NPR1 (non-expressor of pathogenesis-related protein), which sends signals between the chloroplast and the nucleus through for example stromules. ROS and RNS regulate immune responses in several ways, including modification of DNA by methylation, or TGS of DNA viruses. NO is also involved in defence regulation through S-nitrosylation of proteins, and crosstalk with jasmonic acid and ethylene (JA and ET) signalling pathways, as well as a key regulator of SAR. ABA, abscisic acid; AVR, avirulence factor; CP, coat protein; ET, ethylene; ETI, effector triggered immunity; GD, endogenous guardee or decoy protein; HR, hypersensitive response; MAPK, mitogen activated protein kinase; MP, movement protein; NO, nitic oxide; PAMPs, pattern associated molecular patterns; PTI, PAMP-triggered immunity; PTGS, post-transcriptional gene silencing; R, resistance; siRNAs, small interfering RNAs; ROS, reactive oxygen species; RNS, reactive nitrogen species; SA, salicylic acid; SAR, systemic acquired resistance; SN, S-nitrosylation; TFs, transcription factors; TR, transcription regulation; PTR, post-transcriptional regulation; vRNP, viral nucleoprotein.

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particularly dedicated to regulating host immune responses, and although not exclusive, these comprise members of the AP2/ERF, bHLH, bZIP, MYB, NAC and WRKY TF families (Tsuda and Sommsich, 2015). Moreover, the WKRY TFs play a broad and pivotal role in regulating defence responses (reviewed in Eulgem and Somssich, 2007). Thus, the pepper WRKY gene, CaWRKYa, encoding a group I WRKY member is induced during HR in hot pepper (Capsicum annuum) upon TMV infection (Huh et al., 2015). CaWRKYa acts as a positive regulator of transcriptional reprogramming of pathogenesis-related (PR) gene expression and HR. A more recent study in tomato revealed WRKY Group III TFs to be differentially expressed in response to TYLCV infection (Huang et al., 2016). These, and other studies, highlight the central role that MAPK and TFs play in network signalling during plant virus defences. Central to this disease signalling are important molecules, including phytohormones, ROS and nitric oxide (NO), as discussed below. Phytohormones Phytohormones function as signalling molecules for stimulation of plant innate immunity and subsequent activation of defence responses to pathogens (reviewed in Pieterse et al., 2009; Alazem and Lin, 2015). While salicylic acid (SA), jasmonic acid ( JA) and ethylene (ET) are primarily involved in virus resistance (Derksen et al., 2013), other plant development associated hormones such as auxins (Aux), gibberellins (GA), brassinosteroids (BR) and abscisic acid (ABA) contribute to host defence (Durbak et al., 2012). ABA, for example, has been shown to limit virus spread through inhibition of β-1,3-glucanase, which degrades callose (Flors et al., 2005), and JA plays a positive defence role in compatible plant–virus interactions, while disruption of the SA pathway compromises plant resistance to viruses (Alazem and Lin, 2015). Correspondingly, SA is a key factor in basal defence and R gene induced systemic acquired resistance (SAR) (Vlot et al., 2009). Moreover, activation of SA biosynthesis and signalling leads to broad-spectrum defence, characterized by accumulation of ROS, pathogenesis-related proteins (PR), callose deposition and HR (Collum and Culver, 2016). Increasing evidence indicates that phytohormones also control components of the RNA

silencing system at both transcriptional and posttranscriptional levels, including the siRNA- and miRNA-mediated antiviral pathways (Alazem and Lin, 2015). For example, expression of RDR1, an important component of antiviral RNA silencing, has been known to increase upon exogenous application of phytohormones (Hunter et al., 2013). Also, mutation of components of sRNA pathways, including DCL1 and AGO1, caused ABA hypersensitivity and modification of sRNA pathways, enhanced the ABA response (Zhang et al., 2008). It was observed that plants expressing the bacterial salicylate hydroxylase (NahG), an enzyme that degrades SA, produced lower levels of siRNAs upon infection by Plum pox virus (Alamillo et al., 2006), thus further implicating SA in siRNA production. It was recently found that SA can directly inhibit replication of Tomato bushy stunt virus by competitively binding cytosolic glyceraldehyde 3-phosphate dehydrogenase (Tian et al., 2015). It was also reported that SA depletion can negate resistance conferred by potato Ny-1 R gene against potato virus Y (PVY) (Baebler et al., 2014). In addition to SA, JA and ET are known to play a role in signalling of plant virus resistance (Fig. 1.2) and all three hormones are generally antagonistic even though synergism between these pathways has been reported (Mur et al., 2006). JA-SA antagonism has been shown to activate RCY1 resistance to CMV, where a mutant allele of the JA receptor, CO11, restored resistance (Takahashi et al., 2004). Cytokinins are also implicated in resistance induction against several viruses (Sano et al., 1996) and Pogany et al. (2004) showed that induction of cytokinins was associated with a decrease in susceptibility to Tobacco necrosis virus. To counter host defence, viruses are known to manipulate plant hormones to disarm defence responses (Collum and Culver, 2016). For example, TMV replicase protein disrupts the localization and function of interacting auxin/indole acetic acid (IAA) proteins (Padmanabhan et al., 2006), and CaMV P6 protein can inhibit SA-dependent defences by altering the expression and localization of the SA receptor non-expressor of pathogenesisrelated proteins 1 (NPR1) (Love et al., 2012). There is indeed increasing evidence that cross-talk between phytohormone pathways is essential in the regulation of virus resistance (Collum and Culver, 2016).

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Reactive oxygen species Signalling cascades that trigger activation of defence-responsive genes characterize events that occur downstream of PTI and ETI. At the heart of these cascades are reactive oxygen species (ROS) and RNS, including nitric oxide (NO), nitrosium (NO+) ions and peroxynitrite (ONOO–). These signalling molecules are ubiquitous and are involved in various networking processes in plants responses to biotic and abiotic stresses (reviewed in Torres et al., 2006; Domingos et al., 2015; Hernandez et al., 2016). Production of ROS is typically biphasic, with a transitory first phase occurring shortly after interaction between pathogen and host cell membrane receptors, and a second sustained phase associated with signalling of several defence mechanisms and HR (Torres, 2010). During virus-resistance associated HR, plants accumulate ROS that is probably generated by an NADPH oxidase (Moeder et al., 2005). A small GTPase Rac has been shown to be involved in regulating NADPH; indeed, suppression of TMV-induced hypersensitive-type necrosis in tobacco at high temperature, is associated with down-regulation of NADPH oxidase, and stimulation of dehydroascorbate reductase (Moeder et al., 2005; Király et al., 2008). In addition to its role in HR, ROS plays several other roles in defence responses to virus infection, including crosslinking of cell wall glycoproteins and induction of PR proteins (Torres, 2010); ROS also acts through redox control of transcription factors and interaction with other signalling components such as MAPK (Mou et al., 2003) and phytohormones (Torres, 2010). Although its elevation is generally associated with resistance responses, it has been shown that high levels of ROS and ROS compounds accumulate in TYLCV-susceptible tomato compared with resistant ones (Moshe et al., 2012). This may suggest that resistant tomato exhibits greater tolerance to stress than susceptible tomato. Nitric oxide Nitric oxide (NO), as a multifunctional messenger molecule, mediates biological functions through chemical reactions of RNS (Bellin et al., 2013). Indeed, NO has emerged as a major player in plant resistance responses to virus infections (Mur et al., 2006). For example, it was found that in susceptible tomato, TMV infection results in NO production, leading to the inductions of both mitochondrial

alternative electron transport and basal defence (Fu et al., 2010). There is evidence that upon infection by TMV, resistant tobacco displays high levels of NO compared with susceptible tobacco (Durner et al., 1998). Correspondingly, tobacco that is treated with NO donors expresses defence genes (Song and Goodman, 2001) and limits the spread of TMV (Durner et al., 1998) and potato virus X (Li et al., 2014). In addition, infection of Hibiscus cannabinus by the geminivirus mesta yellow vein mosaic virus (MYVMV) results in an increase in the production of NO and tyrosine-nitrated proteins (Sarkar et al., 2010). Overall, NO has been shown to be involved in modulating the activity of many defence-related proteins through S-nitrosylation (Fig. 1.2) (Tada et al., 2008; Kovacs et al., 2015) and in the function of the SAR signalling pathway (Song and Goodman, 2001). Mechanistically, NO exerts its influence through cooperation with H2O2 or regulation of ROS in activating HR, and indirectly through modulation of the major SA, JA, ET and cytokinin pathways (reviewed in Mur et al., 2013). For example, NO and ROS play central roles in SA expression through regulation of NPR1, which interacts with a range of TGA-class transcription factors that bind to TGACG motifs encoded within the promoters of SA-induced defence genes (Zhang et al., 1999; Peleg-Grossman et al., 2010). Hypersensitive response, systemic necrosis and systemic acquired resistance ETI is often accompanied by rapid localized cell death in and around the pathogen infection site. This form of programmed cell death (PCD) is known as the hypersensitive response (HR) (reviewed in Lam et al., 2001). It is also accompanied by induction of broad plant defence responses, including production of defence hormones, NO, and ROS (Mandadi and Scholthof, 2013). HR is usually associated with several morphological and physiological changes in the cell, including effects on the chloroplast and mitochondrion, which plays an important role in HR and PCD responses in plants; for instance, ROS is derived from electrontransfer intermediates in the inner mitochondrial membrane (Lam et al., 2001). HR affects calcium ion homeostasis, leading to altered membrane permeability, and during HR several vacuolar caspase-like proteinases are activated and contribute to

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cell death or necrosis (Mur et al., 2008). Although tissue necrosis is largely associated with R gene triggered HR, it can be uncoupled from the resistance response. For example, the R gene Rx1 from potato recognizes the PVX CP and inhibits PVX, independent of HR-associated necrosis (Bendahmane et al., 1999). As expected, factors that mediate HR against diverse viral pathogens are conserved across plant genera (reviewed in Mandadi and Scholthof, 2013). While systemic necrosis (SN) is found in compatible virus–host interactions, HR-resistance associated necrosis and SN share similarities at the molecular and biochemical levels (Mandadi and Scholthof, 2013). For example, both HR and SN depend on a functional SGT1/RAR1 complex and both require MAPK kinase kinase (MAPKKK) and MEK2 signalling (Komatsu et al., 2010), suggesting that SN may promote antiviral immunity during a compatible virus–host interaction. SAR is triggered during an incompatible interaction involving viral Avr and R proteins, and resistance is transduced to non-infected distal tissues, resulting in elevated levels of phytohormones such as JA and SA (reviewed in Vlot et al., 2008). It is thought that SAR is long-lasting due to accompanying epigenetic modifications, such as chromatin remodelling and DNA methylation, which leads to SAR signalling (Spoel and Dong, 2012). While several metabolites have been proposed as putative signals in SAR, it is likely that SAR involves interaction between multiple signals such as glycerolipids, methyl salicylate (MeSA) and lipid-transfer proteins (Liu et al., 2011). Ubiquitin proteasome system in virus defence The ubiquitin proteasome system (UPS) regulates cellular activities including the cell cycle, transcription and signal transduction, and is also involved at all stages of viral defence (Citovsky et al., 2009). For instance, UPS has been shown to be involved in N gene-mediated HR and resistance response in TMV-infected tobacco and the MPs of TMV and TYMV are specifically targeted for degradation by the host UPS (Drugeon and Jupin, 2002). Several viruses also target multiple UPS components to promote virulence, thus stressing the important role of UPS in the balance between resistance and susceptibility.

Chloroplasts and plant immunity Growing evidence over the past decade has shown that the chloroplast plays a central role in pathogen immunity (reviewed in Zhao et al., 2016; Bhattacharyya and Chakraborty, 2017). For instance, the chloroplast is a major generator of ROS and H2O2 and a burst of chloroplast-associated ROS can be detected during virus infection in incompatible interactions (Hakmaori et al., 2012) and is essential for HR (Torres et al., 2006). Although ubiquitous in plants, there is no consensus on the central source of NO (Domingos et al. 2015), and no nitric oxide synthase (NOS) found in animals has been reported in plants. Arabidopsis nitric oxide-associated protein 1 (AtNOA1), previously linked with NO production, is indeed a nuclear encoded, chloroplast translation factor (Flores et al., 2008; Moreau et al., 2008), and a member of the conserved circularly permutated GTPase (cGTPase) of the YlqF/YawG family (Moreau et al., 2008, 2010; Sudhamsu et al., 2008). The nature of NOA1/cGTPase homologues involvement in virus defence responses is yet unclear. It is possible that NOA1 participation in disease response could be through its association with the chloroplast (Bhat et al., 2013; Liu et al., 2014). The chloroplast is also the main sites of production of SA and JA (Padmanabhan and Dinesh-Kumar, 2010; Caplan et al., 2015; Serrano et al., 2016) and is a rich source of ROS in response to pathogen attack. Moreover, a large pool of Ca2+, whose levels change in response to virus attack, is stored in the chloroplast (Mur et al., 2008), and it has been shown that a chloroplast-localized calciumsensing receptor is involved in basal resistance and R gene-mediated defence (Nomura et al., 2012). Furthermore, the N receptor interacting protein (NPRI) normally accumulates in the chloroplast, however, in N gene-containing tobacco infected by TMV, it relocalises to the cytoplasm and nucleus and plays a role in defence (Caplan et al., 2008). Consistent with the important role of the chloroplast during pathogen attack, tubular extensions, or stromules, also relocate from the chloroplast to nucleus during innate immunity, and correlate with accumulation of both NRIP1 defence protein and H2O2 in the nucleus (Caplan et al., 2015). It has for example been shown that upon Abutilon mosaic virus infection, stromules are induced and relocate to the nucleus (Krenz et al., 2012).

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Chloroplast components and proteins, such as thylakoids and RuBisCO, are also involved in virus resistance (reviewed in Bhattacharyya and Chakraborty, 2017). Thus, silencing of N. benthamiana photosystem I PSI-K, a chloroplast thylakoid protein that interacts in vitro with CI protein of PPV, enhanced PPV accumulation ( Jimenez et al., 2006). In RuBisCO small subunit (RbCS) silenced N. benthamiana, expression of PR1 gene was suppressed and Tomato mosaic virus was capable of inducing local necrosis (Zhao et al., 2013). It is now known that chloroplast-mediated defence response is largely light regulated (Kangasjärvi et al., 2012), thus it has been suggested that signals from the chloroplast mediate crosstalk between light acclimation and disease resistance in Arabidopsis (Sewelam et al., 2016). Chloroplast mediated defence response can also be through virus-R gene recognition associated ETI. Hence, several TIR-NLR proteins from different families have been shown to contain putative chloroplast localization signals, suggesting an extensive network of chloroplast-based pathogen recognition systems (Bhattacharyya and Chakraborty, 2017). Primary and secondary metabolism and virus resistance Following plant virus infection, massive reprogramming of metabolome, proteome and transcriptome occurs, and several studies have demonstrated that major differences can be observed between resistant/tolerant and susceptible cultivars (Allie et al., 2014; Maruthi et al., 2014; Sade et al., 2014). One of the most frequent plant responses to virus infection is a down-regulation of photosynthesis and other primary plant metabolic pathways (Rojas et al., 2014; Llave, 2016). It has been suggested that the role of primary metabolism during plant–pathogen interactions is to support cellular energy requirements for plant defence (Kangasjarvi et al., 2012). Chloroplasts are a common target of many plant viruses (Li et al., 2016), indeed general proteins or transcripts encoding proteins for chloroplast functioning and photosynthesis are repressed (Mochizuki et al., 2014). Significant differences in the levels of some metabolites (polyamines, amino acids, phenolic, indolic metabolites) have been observed between TYLCV-infected susceptible tomato and tomato line 902 line, which

carries the Ty-5 resistance trait from wild tomato (Solanum peruvianum) (Sade et al., 2014). This suggests a possible role for the phenylpropanoid, tryptophan and urea/polyamine pathways in resistance to geminiviruses. A two stage response was associated with resistance, namely, a coordinated response of primary metabolites such as amino acids, sugars and tricarboxylic acid (TCA) cycle intermediates, and secondly, a less tightly regulated metabolite response. This two-stage response has also been reported in metabolomics profiling of TMV infected plants (Bazzini et al., 2011). The defence cascade mediated by metabolism of carbohydrates, amino acids, lipids, and photorespiration is also linked to the defence signalling cascade (reviewed in Rojas et al., 2014). Glucose, fructose and sucrose can induce pathogenesis-related (PR) genes and photo-assimilate responding gene 1 (PAR1) (Herbers et al., 1996). These sugars are recognized as signalling molecules in plants, and the coordination of sugar and hormonal pathways, in addition to their roles as carbon and energy sources, likely plays a role in virus innate immunity (Bolouri Mogbaddam et al., 2012). Indeed, virus infection often leads to increased sugar levels (Shalitin and Wolf, 2000) which are thus signal intensification of immune responses (Gomez-Ariza et al., 2007). The cellular sucrose–hexose ratio and altered sucrose to starch ratios are important factors in cellular responses (Xiang et al., 2011). Hence, TYLCV infection of resistant tomato was reported to lead to silencing of the hexose transport gene (Vidavsky and Czosnek, 1998; Sade et al., 2013); in sucrose transporter gene (LeHT1)-silenced plants, sugar profiles were similar to those in susceptible plants, thus implicating LeHT1 and sugars in TYLCV resistance. Trehalose is another non-reducing sugar that can induce resistance to pathogens and was suggested to be an essential signal in defence (Singh et al., 2011). Accordingly, trehalose-6-phosphate synthase/phosphatase 11 (TPS11) expression was up-regulated in Arabidopsis infected by TMV (Golem and Culver, 2003). Although more information on the role of sugars in virus resistance responses is still to be determined, it is likely that ROS or RNS may play a role in a complex crosstalk between sugar and hormone signalling pathways. Thus, sugar signals may contribute to immune response priming and biodegradable sugar compounds could be further

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explored for use as alternatives to toxic agrochemicals (Bolouri-Mogbaddam et al., 2012). Phenotypes of virus resistance: tolerance and symptom recovery The type and mechanism of resistance deployed by plants against infecting viruses determines the nature of phenotype displayed by plants. Resistance manifestations range from immunity to tolerance. Plants with immunity to a virus display complete resistance due to the inability of the virus to replicate may be due to cell death and thus containment of the virus at the infection site. In contrast, resistant plants limit virus multiplication by interfering with the replication cycle, resulting in mild symptoms. Thus, there are different levels of resistance. Some plants display resistance in the form of tolerance, in which symptoms are reduced compared with susceptible plants but virus replication persists throughout the life of the plant (Bengyella et al., 2015). Tolerance has been associated in some instances with accumulation of vsiRNAs and an increase in defence-associated host gene expression (Sahu et al., 2010). Recovery from virus infection is characterized by initial severe systemic symptoms, which progressively decrease, resulting in some cases in complete disappearance of symptoms in newly developed leaves (Ghoshal and Sanfaçon, 2014; Ma et al., 2015; Nie and Molen, 2015; Patil and Fauquet, 2015). In this case, the recovered leaves display lower levels of virus replication (Bengyella et al., 2015). Recovery may also be due to an adaptive response involving vsiRNAs (Hagen et al., 2008) and dependent on temperature (Ghoshal and Sanfaçon, 2014). Recovery, which was first reported in tobacco plants infected by Tobacco ringspot virus (Wingard, 1928), occurs in both RNA and DNA viruses. Several lines of evidence from N. tabacum (Loebenstein et al., 1977) and N. benthamiana (Lu et al., 2012) infected with the M- strain of CMV(MCMV) have shown that disease development can involve initial and secondary pathogenicity processes that are interrupted by a transient recovery phase. Recovery from geminivirus infection has been observed in several plant species, including Cucumis melo (cantaloupe) and Citrullus lanatus (watermelon) infected by Curcubit leaf crumple virus

(Hagen et al., 2008), pepper plants infected by Pepper golden mosaic virus (Rodríguez-Negrete et al., 2009) and tomato infected with Tomato leaf curl New Delhi virus (Sahu et al., 2010). In cassava, recovery has been observed in plants infected by African cassava mosaic virus (Fondong et al., 2000), Sri Lankan cassava mosaic virus (Chellappan et al., 2004) and South African cassava mosaic virus (SACMV) (Allie et al., 2014). While tolerance and recovery phenotypes are often associated with RNA silencing (Ghoshal and Sanfaçon, 2014), it is possible that R genes play key roles in both phenomena. Interestingly, in SACMVinfected susceptible cassava T200, expression of a significant number of R gene analogues (RGAs) was down-regulated at 32 and 67 dpi, however, this was not observed in SACMV-infected tolerant TME3 (Allie et al., 2014). Putative R protein analogues with amide-like indole-3-accetic acid–Ile-Leu-Arg (IAA-ILR) and LRR-kinase conserved domains were observed in tolerant TME3 but not in the susceptible genotype (Louis and Rey, 2015). Phylogenetic and expression analyses demonstrated that diverse R genes unique to TME3 are differentially expressed during tolerance and recovery. It is therefore clear that tolerance and recovery in plant virus infections involve multiple mechanisms (Louis and Rey, 2015). Network modelling: understanding plant immunity Viral pathogens have been shown to induce multiple integrative networks of resistance mechanisms (Elena et al., 2011) (overview depicted in Fig. 1.2). Early studies in the analysis of virus-regulated genes (VRGs) have shown that viruses alter the transcription of master transcription factors and hub proteins, as determined from protein–protein interaction and transcriptional regulatory networks (TRNs), and from high-throughput transcriptome and proteome data (Geisler-Lee et al., 2007). A major component of plant defence is transcriptional and translational modulation of the genome. A plant-pathogen immune network, PPIN-1, including 552 host immune proteins in Arabidopsis infected by Pseudomonas syringae was recently established (Windram et al., 2014). A key observation from this network was that the majority of pathogens effectors were indirectly connected to

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immune receptors. Accordingly, deciphering largescale protein–protein networks that underpin plant immunity to virus infection will prove invaluable in the development of virus-resistant crops. Engineering for virus resistance Since the mid-1980s when it was first observed that plants transformed with a viral gene (i.e. genetic engineering) were resistant to the virus (Abel et al., 1986), genetic engineering has become an important tool in plant virus control efforts (see reviews in Cillo and Palukaitis, 2014; Galvez et al., 2014). This type of resistance was thought to be due to the ability of the expressed transgene to disrupt the assembly of the infecting viral protein in the plant, thereby conferring resistance. Thus, the resistance exhibited by tobacco plants containing the CP of TMV was suggested to be due to the ability of the transgenic CP to disassemble infecting TMV particles (Beachy, 1999). Correspondingly, the resistance displayed by transgenic tobacco plants expressing a defective TMV movement protein (dMP) was thought to be due to competition for the binding sites in the plasmodesmata between the dMP and the wild-type virus encoded MP (Lapidot et al., 1993). Unlike CP- and MP-mediated resistance, the ability of viral replicase protein to confer resistance was suggested to be due largely to RNA-mediated resistance (reviewed in Cillo and Palukaitis, 2014). It subsequently became clear that noncoding regions of a viral genome, viral sequences in sense or antisense orientation or in double-stranded forms, confer resistance by targeting viral RNA through the RNA silencing mechanism (Baulcombe, 1996; Cillo and Palukaitis, 2014; Galvez et al., 2014; Waterhouse et al., 1998). Thus, viral RNAs are processed into siRNAs, which guide the silencing complex to target viral mRNA for degradation or translation inhibition at the post-transcriptional level; or to cause an epigenetic modification at the transcriptional level, dependent on RNA-directed DNA methylation (reviewed in Castel and Martienssen, 2013; Vaucheret, 2006). Because sRNAs determine the specificity of the RNA silencing through base-pairing recognition of their complementary target RNAs (Ding and Lu, 2011), artificial microRNAs (amiRNAs) and trans-acting short interfering RNAs (tasiRNAs), have also been shown to confer resistance to RNA

and DNA plant viruses (Cillo and Palukaitis, 2014; Galvez et al., 2014; Teotia et al., 2016). The ease of designing specific and targeted technologies based on RNA silencing has provided new tools in virus control efforts. Advances made in this post-genomic era have yet led to the development of a reverse genetics approach, designated Targeting Induced Local Lesions IN Genomes (TILLING), for the control of viruses in crops (Gauffier et al., 2016; Nieto et al., 2007; Piron et al., 2010). TILLING is used to detect unknown single nucleotide polymorphisms (SNPs) in genes of interest using enzymatic digestion and deep sequencing to identify the mutations (Henikoff et al., 2004; Kurowska et al., 2011; McCallum et al., 2000). Plant virus control strategies have employed mainly a variant of TILLING, known as EcoTILLING, which examines natural genetic variation in populations (Comai et al., 2004). In EcoTILLING, virus resistance alleles are quarried for defective alleles whose gene products interfere with the virus infection cycle (Diaz-Pendon et al., 2004; Gauffier et al., 2016; Nieto et al., 2007; Piron et al., 2010; Robaglia and Caranta, 2006; Truniger and Aranda, 2009). The EcoTILLING approach was used to identify a non-silent allelic variant of eIF4E from 113 EcoTILLING accessions of melon and the variant was found to confer resistance to melon necrotic spot virus (MNSV) (Nieto et al., 2007). Accordingly, five new resistance alleles of eIF4E that conferred resistance to PVY in Capsicum were reported by Ibiza et al. (2010) and a splicing mutant of eIF4E from a mutagenized population was found to confer immunity to PVY and Pepper mottle virus (Menda et al., 2004). Because EcoTILLING identifies natural resistance, it will likely continue to be adopted as a method of choice in virus control efforts. The plant virus control toolbox has in the last few years been supplemented with a clustered regulatory interspaced short palindromic repeats (CRISPR) and CRISPR-associated nuclease 9 (Cas9) (CRISPR-Cas9) system (Bhaya et al., 2011), a highly efficient and specific gene editing method, used to target plant viruses. The Streptococcus pyogenes endonuclease Cas9 (SpCas9) CRISPR-Cas9 system has been harnessed for efficient genome editing and gene regulation in plants (see reviews by Baltes and Voytas, 2015; Belhaj et al., 2015; Liu et al., 2016) and other eukaryotes

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(Cong et al., 2013; Haurwitz et al., 2010; Jinek et al., 2013). The CRISPR-Cas9 system has been shown in many instances to be effective in controlling plant DNA viruses (see review in Fondong et al., 2016). Importantly, a robust multiplexing strategy for this system, whereby the guide RNA is engineered as a single polycistronic gene based on endogenous tRNA-processing system, has been developed (Xie et al., 2015). In addition to directly targeting the viral genome using the CRISPR-Cas9 system, several other CRISPR-Cas9 platforms have been described, including non-transgenic delivery of CRISPR-Cas9 cassette in plants, as well as CRISPR-Cas9 gene drive modification of insect virus vector populations (see review in Fondong et al., 2016). The early applications of CRISPR-Cas9 have been to target DNA genome viruses since Cas9 is a DNA endonuclease. However, recently, a RNA Cas9 endonuclease from Francisella novicida was successfully used to inhibit hepatitis C virus, a positive sense single-stranded RNA virus, in eukaryotic cells (Price et al., 2015), thereby expanding the scope of this approach in virus control, especially given that most plant viruses have an RNA genome. Future trends Despite advances in the understanding of molecular mechanisms involved in plant virus resistance over the past two decades, the ‘magic switch’ to trigger immunity remains elusive. Thanks to advances in deep sequencing, most studies have focused only on transcriptional changes occurring during growthrelated activities in response to an attack (Huot et al., 2014; Pajerowska-Mukhtar et al., 2012). In fact, thus far, there has been a near dearth of information on mRNA translational changes occurring during immune induction. Recently, however, a novel strategy of plant antiviral defence based on suppression of host global translation, mediated by the transmembrane immune receptor NIK1, (nuclear shuttle protein (NSP)-Interacting Kinase1), has been demonstrated in begomoviruses (Machado et al., 2017). Global translational reprogramming has been shown to be fundamental in immune responses (Xu et al., 2017a) and mRNA translation efficiency during PTI was linked to a highly enriched mRNA consensus sequence, R-motif, which consists mostly of purines (Xu et al., 2017a). This mRNA R-motif regulates protein translation

in response to PTI induction through interactions with poly(A)-binding proteins. Furthermore, evidence that translation of key immune regulatorsis rapidly and transiently induced upon pathogen challenge (Xu et al., 2017a) has led to the identification of two pathogen-responsive upstream open reading frames (uORFs) that mediate translational control of immune response transcription factors. It has therefore been suggested that uORFs can be used as a broad-spectrum pathogen control strategy, where it rapidly de-represses translation of pre-existing mRNAs immune transcription factors upon pathogen attack (Xu et al., 2017b). In such a scenario, the challenge in future would be to identify master immune regulators so as to apply uORFs in engineering durable broad-spectrum virus resistance, without compromising plant fitness. References Abel, P.P., Nelson, R.S., De, B., Hoffmann, N., Rogers, S.G., Fraley, R.T., and Beachy, R.N. (1986). Delay of disease development in transgenic plants that express the Tobacco mosaic virus coat protein gene. Science 232, 738–743. Alamillo, J.M., Saénz, P., and García, J.A. (2006). Salicylic acid-mediated and RNA-silencing defense mechanisms cooperate in the restriction of systemic spread of Plum pox virus in tobacco. Plant J. 48, 217–227. Depuydt, S., and Hardtke, C.S. (2011). Hormone signalling crosstalk in plant growth regulation. Curr. Biol. 21, R365–73. https://doi.org/10.1016/j.cub.2011.03.013 Alazem, M., and Lin, N.S. (2015). Roles of plant hormones in the regulation of host–virus interactions. Mol. Plant Pathol. 16, 529–540. https://doi.org/10.1111/ mpp.12204 Albar, L., Bangratz-Reyser, M., Hébrard, E., Ndjiondjop, M.N., Jones, M., and Ghesquière, A. (2006). Mutations in the eIF(iso)4G translation initiation factor confer high resistance of rice to Rice yellow mottle virus. Plant J. 47, 417–426. Allie, F., Pierce, E.J., Okoniewski, M.J., and Rey, C. (2014). Transcriptional analysis of South African Cassava mosaic virus-infected susceptible and tolerant landraces of cassava highlights differences in resistance, basal defense and cell wall associated genes during infection. BMC Genomics 15, 1006. https://doi.org/10.1186/14712164-15-1006 Aregger, M., Borah, B.K., Seguin, J., Rajeswaran, R., Gubaeva, E.G., Zvereva, A.S., Windels, D., Vazquez, F., Blevins, T., Farinelli, L., et al. (2012). Primary and secondary siRNAs in geminivirus-induced gene silencing. PLOS Pathog. 8, e1002941. https://doi.org/10.1371/journal. ppat.1002941 Arikit, S., Xia, R., Kakrana, A., Huang, K., Zhai, J., Yan, Z., Valdés-López, O., Prince, S., Musket, T.A., Nguyen, H.T., et al. (2014). An atlas of soybean small RNAs identifies phased siRNAs from hundreds of coding genes.

Mechanisms of Virus Resistance in Plants |  15

Plant Cell 26, 4584–4601. https://doi.org/10.1105/ tpc.114.131847 Baebler, Š., Witek, K., Petek, M., Stare, K., Tušek-Žnidarič, M., Pompe-Novak, M., Renaut, J., Szajko, K., StrzelczykŻyta, D., Marczewski, W., et al. (2014). Salicylic acid is an indispensable component of the Ny-1 resistancegene-mediated response against Potato virus Y infection in potato. J. Exp. Bot. 65, 1095–1109. https://doi. org/10.1093/jxb/ert447 Baltes, N.J., and Voytas, D.F. (2015). Enabling plant synthetic biology through genome engineering. Trends Biotechnol. 33, 120–131. https://doi.org/10.1016/j. tibtech.2014.11.008 Baulcombe, D.C. (1996). Mechanisms of pathogen-derived resistance to viruses in transgenic plants. Plant Cell 8, 1833–1844. https://doi.org/10.1105/tpc.8.10.1833 Baumberger, N., Tsai, C.H., Lie, M., Havecker, E., and Baulcombe, D.C. (2007). The Polerovirus silencing suppressor P0 targets ARGONAUTE proteins for degradation. Curr. Biol. 17, 1609–1614. Bazzini, A.A., Manacorda, C.A., Tohge, T., Conti, G., Rodriguez, M.C., Nunes-Nesi, A., Villanueva, S., Fernie, A.R., Carrari, F., and Asurmendi, S. (2011). Metabolic and miRNA profiling of TMV infected plants reveals biphasic temporal changes. PLOS ONE 6, e28466. https://doi.org/10.1371/journal.pone.0028466 Beachy, R.N. (1999). Coat-protein-mediated resistance to Tobacco mosaic virus: discovery mechanisms and exploitation. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 354, 659–664. https://doi.org/10.1098/rstb.1999.0418 Bednarek, P. (2012). Chemical warfare or modulators of defence responses – the function of secondary metabolites in plant immunity. Curr. Opin. Plant Biol. 15, 407–414. Belhaj, K., Chaparro-Garcia, A., Kamoun, S., Patron, N.J., and Nekrasov, V. (2015). Editing plant genomes with CRISPR/Cas9. Curr. Opin. Biotechnol. 32, 76–84. https://doi.org/10.1016/j.copbio.2014.11.007 Bellin, D., Asai, S., Delledonne, M., and Yoshioka, H. (2013). Nitric oxide as a mediator for defense responses. Mol. Plant Microbe Interact. 26, 271–277. https://doi. org/10.1094/MPMI-09-12-0214-CR Bendahmane, A., Kanyuka, K., and Baulcombe, D.C. (1999). The Rx gene from potato controls separate virus resistance and cell death responses. Plant Cell 11, 781–792. Bengyella, L., Sayanika, D., Waikhom, D., Allie, F., and Rey, C. (2015). Virus tolerance and recovery from viral induced symptoms in plants are associated with transcriptome reprogramming. Plant Mol. Biol. 89, 243–252. Bhat, S., Folimonova, S.Y., Cole, A.B., Ballard, K.D., Lei, Z., Watson, B.S., Sumner, L.W., and Nelson, R.S. (2013). Influence of host chloroplast proteins on Tobacco mosaic virus accumulation and intercellular movement. Plant Physiol. 161, 134–147. https://doi.org/10.1104/ pp.112.207860 Bhattacharyya, D., and Chakraborty, S. (2017). Chloroplast: the Trojan horse in plant-virus interaction. Mol. Plant Pathol. 13, 385–392. Bhaya, D., Davison, M., and Barrangou, R. (2011). CRISPRCas systems in bacteria and archaea: versatile small RNAs for adaptive defense and regulation. Annu. Rev.

Genet. 45, 273–297. https://doi.org/10.1146/annurevgenet-110410-132430 Bigeard, J., Colcombet, J., and Hirt, H. (2015). Signaling mechanisms in pattern-triggered immunity (PTI). Mol Plant 8, 521–539. https://doi.org/10.1016/j. molp.2014.12.022 Blevins, T., Rajeswaran, R., Aregger, M., Borah, B.K., Schepetilnikov, M., Baerlocher, L., Farinelli, L., Meins, F., Hohn, T., and Pooggin, M.M. (2011). Massive production of small RNAs from a non-coding region of Cauliflower mosaic virus in plant defense and viral counter-defense. Nucleic Acids Res. 39, 5003–5014. https://doi.org/10.1093/nar/gkr119 Boller, T., and Felix, G. (2009). A renaissance of elicitors: perception of microbe-associated molecular patterns and danger signals by pattern-recognition receptors. Annu. Rev. Plant Biol. 60, 379–406. https://doi.org/10.1146/ annurev.arplant.57.032905.105346 Bolouri Moghaddam, M.R., and Van den Ende, W. (2012). Sugars and plant innate immunity. J. Exp. Bot. 63, 3989– 3998. https://doi.org/10.1093/jxb/ers129 Brutus, A., and He, S.Y. (2010). Broad-spectrum defense against plant pathogens. Nat. Biotechnol. 28, 330–331. https://doi.org/10.1038/nbt0410-330 Calil, I.P., and Fontes, E.P.B. (2017). Plant immunity against viruses: antiviral immune receptors in focus. Ann. Bot. 119, 711–723. https://doi.org/10.1093/aob/mcw200 Caplan, J.L., Mamillapalli, P., Burch-Smith, T.M., Czymmek, K., and Dinesh-Kumar, S.P. (2008). Chloroplastic protein NRIP1 mediates innate immune receptor recognition of a viral effector. Cell 132, 449–462. https://doi.org/10.1016/j.cell.2007.12.031 Caplan, J.L., Kumar, A.S., Park, E., Padmanabhan, M.S., Hoban, K., Modla, S., Czymmek, K., and DineshKumar, S.P. (2015). Chloroplast Stromules Function during Innate Immunity. Dev. Cell 34, 45–57. https:// doi.org/10.1016/j.devcel.2015.05.011 Carbonell, A., and Carrington, J.C. (2015). Antiviral roles of plant ARGONAUTES. Curr. Opin. Plant Biol. 27, 111–117. https://doi.org/10.1016/j.pbi.2015.06.013 Castel, S.E., and Martienssen, R.A. (2013). RNA interference in the nucleus: roles for small RNAs in transcription, epigenetics and beyond. Nat. Rev. Genet. 14, 100–112. https://doi.org/10.1038/nrg3355 Chan, S.W., Zilberman, D., Xie, Z., Johansen, L.K., Carrington, J.C., and Jacobsen, S.E. (2004). RNA silencing genes control de novo DNA methylation. Science 303, 1336. https://doi.org/10.1126/ science.1095989 Chellappan, P., Vanitharani, R., Pita, J., and Fauquet, C.M. (2004). Short interfering RNA accumulation correlates with host recovery in DNA virus-infected hosts, and gene silencing targets specific viral sequences. J. Virol. 78, 7465–7477. https://doi.org/10.1128/JVI.78.14.74657477.2004 Cillo, F., and Palukaitis, P. (2014). Transgenic resistance. Adv. Virus Res. 90, 35–146. Citovsky, V., Zaltsman, A., Kozlovsky, S.V., Gafni, Y., and Krichevsky, A. (2009). Proteasomal degradation in plant-pathogen interactions. Semin. Cell Dev. Biol. 20, 1048–1054. https://doi.org/10.1016/j. semcdb.2009.05.012 Collum, T.D., and Culver, J.N. (2016). The impact of phytohormones on virus infection and disease. Curr

16  | Rey and Fondong

Opin Virol 17, 25–31. https://doi.org/10.1016/j. coviro.2015.11.003 Comai, L., Young, K., Till, B.J., Reynolds, S.H., Greene, E.A., Codomo, C.A., Enns, L.C., Johnson, J.E., Burtner, C., Odden, A.R., et al. (2004). Efficient discovery of DNA polymorphisms in natural populations by Ecotilling. Plant J. 37, 778–786. Cong, L., Ran, F.A., Cox, D., Lin, S., Barretto, R., Habib, N., Hsu, P.D., Wu, X., Jiang, W., Marraffini, L.A., et al. (2013). Multiplex genome engineering using CRISPR/ Cas systems. Science 339, 819–823. https://doi. org/10.1126/science.1231143 Csorba, T., Kontra, L., and Burgyán, J. (2015). Viral silencing suppressors: Tools forged to fine-tune host-pathogen coexistence. Virology 479-480, 85–103. https://doi. org/10.1016/j.virol.2015.02.028 Cuellar, W.J., Kreuze, J.F., Rajamäki, M.L., Cruzado, K.R., Untiveros, M., and Valkonen, J.P. (2009). Elimination of antiviral defense by viral RNase III. Proc. Natl. Acad. Sci. U.S.A. 106, 10354–10358. https://doi.org/10.1073/ pnas.0806042106 Deleris, A., Gallego-Bartolome, J., Bao, J., Kasschau, K.D., Carrington, J.C., and Voinnet, O. (2006). Hierarchical action and inhibition of plant Dicer-like proteins in antiviral defense. Science 313, 68–71. Diaz-Pendon, J.A., Truniger, V., Nieto, C., Garcia-Mas, J., Bendahmane, A., and Aranda, M.A. (2004). Advances in understanding recessive resistance to plant viruses. Mol. Plant Pathol. 5, 223–233. https://doi.org/10.1111/ j.1364-3703.2004.00223.x Ding, S.W., and Lu, R. (2011). Virus-derived siRNAs and piRNAs in immunity and pathogenesis. Curr. Opin. Virol. 1, 533–544. https://doi.org/10.1016/j. coviro.2011.10.028 Derksen, H., Rampitsch, C., and Daayf, F. (2013). Signaling cross-talk in plant disease resistance. Plant Sci. 207, 79–87. https://doi.org/10.1016/j.plantsci.2013.03.004 Domingos, P., Prado, A.M., Wong, A., Gehring, C., and Feijo, J.A. (2015). Nitric oxide: a multitasked signaling gas in plants. Mol. Plant 8, 506–520. https://doi. org/10.1016/j.molp.2014.12.010 Donaire, L., Barajas, D., Martínez-García, B., MartínezPriego, L., Pagán, I., and Llave, C. (2008). Structural and genetic requirements for the biogenesis of Tobacco rattle virus-derived small interfering RNAs. J. Virol. 82, 5167–5177. https://doi.org/10.1128/JVI.00272-08 Donaire, L., Wang, Y., Gonzalez-Ibeas, D., Mayer, K.F., Aranda, M.A., and Llave, C. (2009). Deep-sequencing of plant viral small RNAs reveals effective and widespread targeting of viral genomes. Virology 392, 203–214. https://doi.org/10.1016/j.virol.2009.07.005 Dong, J., Chen, C., and Chen, Z. (2003). Expression profiles of the Arabidopsis WKRY group III transcription factors are part of different plant defense response. Plant Mol. Biol. 51, 21–37. Drugeon, G., and Jupin, I. (2002). Stability in vitro of the 69K movement protein of Turnip yellow mosaic virus is regulated by the ubiquitin-mediated proteasome pathway. J. Gen. Virol. 83, 3187–3197. Durbak, A., Yao, H., and McSteen, P. (2012). Hormone signaling in plant development. Curr. Opin. Plant Biol. 15, 92–96. https://doi.org/10.1016/j.pbi.2011.12.004

Durner, J., Wendehenne, D., and Klessig, D.F. (1998). Defense gene induction in tobacco by nitric oxide, cyclic GMP, and cyclic ADP-ribose. Proc. Natl. Acad. Sci. U.S.A. 95, 10328–10333. Elena, S.F., Carrera, J., and Rodrigo, G. (2011). A systems biology approach to the evolution of plant-virus interactions. Curr. Opin. Plant Biol. 14, 372–377. https://doi.org/10.1016/j.pbi.2011.03.013 Eulgem, T., and Somssich, I.E. (2007). Networks of WRKY transcription factors in defense signaling. Curr. Opin. Plant Biol. 10, 366–371. Fan, J., and Doerner, P. (2012). Genetic and molecular basis of nonhost disease resistance: complex, yes; silver bullet, no. Curr. Opin. Plant Biol. 15, 400–406. https://doi. org/10.1016/j.pbi.2012.03.001 Fei, Q., Xia, R., and Meyers, B.C. (2013). Phased, secondary, small interfering RNAs in posttranscriptional regulatory networks. Plant Cell 25, 2400–2415. https://doi. org/10.1105/tpc.113.114652 Fei, Q., Zhang, Y., Xia, R., and Meyers, B.C. (2016). Small RNAs add zing to the zig-zag-zig model of plant defenses. Mol. Plant Microbe Interact. 29, 165–169. https://doi. org/10.1094/MPMI-09-15-0212-FI Flores-Pérez, U., Sauret-Güeto, S., Gas, E., Jarvis, P., and Rodríguez-Concepción, M. (2008). A mutant impaired in the production of plastome-encoded proteins uncovers a mechanism for the homeostasis of isoprenoid biosynthetic enzymes in Arabidopsis plastids. Plant Cell 20, 1303–1315. https://doi.org/10.1105/ tpc.108.058768 Flors, V., Ton, J., Jakab, G., and Mauch-Mani, B. (2005). Abscisic acid and callose: team players in defence against pathogens? J. Phytopathol. 153, 377–383. Fondong, V.N., Thresh, J.M., and Fauquet, C.M. (2000). Field experiments in Cameroon on Cassava mosaic virus disease and the reversion phenomenon in susceptible and resistant cassava cultivars. Int. J. Pest Manag. 46, 211–217. Fondong, V.N., Nagalakshmi, U., and Dinesh-Kumar, S.P. (2016). Novel Functional Genomics Approaches: A Promising Future in the Combat Against Plant Viruses. Phytopathology 106, 1231–1239. https://doi. org/10.1094/PHYTO-03-16-0145-FI Fraile, A., and García-Arenal, F. (2010). The coevolution of plants and viruses: resistance and pathogenicity. Adv. Virus Res. 76, 1–32. https://doi.org/10.1016/S00653527(10)76001-2 Fu, L.J., Shi, K., Gu, M., Zhou, Y.H., Dong, D.K., Liang, W.S., Song, F.M., and Yu, J.Q. (2010). Systemic induction and role of mitochondrial alternative oxidase and nitric oxide in a compatible tomato-Tobacco mosaic virus interaction. Mol. Plant Microbe Interact. 23, 39–48. https://doi. org/10.1094/MPMI-23-1-0039 Galvez, L.C., Banerjee, J., Pinar, H., and Mitra, A. (2014). Engineered plant virus resistance. Plant Sci. 228, 11–25. https://doi.org/10.1016/j.plantsci.2014.07.006 Garcia-Ruiz, H., Takeda, A., Chapman, E.J., Sullivan, C.M., Fahlgren, N., Brempelis, K.J., and Carrington, J.C. (2010). Arabidopsis RNA-dependent RNA polymerases and dicer-like proteins in antiviral defense and small interfering RNA biogenesis during Turnip Mosaic Virus infection. Plant Cell 22, 481–496. https://doi. org/10.1105/tpc.109.073056

Mechanisms of Virus Resistance in Plants |  17

Gauffier, C., Lebaron, C., Moretti, A., Constant, C., Moquet, F., Bonnet, G., Caranta, C., and Gallois, J.L. (2016). A TILLING approach to generate broad-spectrum resistance to potyviruses in tomato is hampered by eIF4E gene redundancy. Plant J. 85, 717–729. https:// doi.org/10.1111/tpj.13136 German-Retana, S., Walter, J., Doublet, B., Roudet-Tavert, G., Nicaise, V., Lecampion, C., Houvenaghel, M.C., Robaglia, C., Michon, T., and Le Gall, O. (2008). Mutational analysis of plant cap-binding protein eIF4E reveals key amino acids involved in biochemical functions and potyvirus infection. J. Virol. 82, 7601– 7612. https://doi.org/10.1128/JVI.00209-08 Ghoshal, B., and Sanfaçon, H. (2015). Symptom recovery in virus-infected plants: Revisiting the role of RNA silencing mechanisms. Virology 479-480, 167–179. https://doi.org/10.1016/j.virol.2015.01.008 Geisler-Lee, J., O’Toole, N., Ammar, R., Provart, N.J., Millar, A.H., and Geisler, M. (2007). A predicted interactome for Arabidopsis. Plant Physiol. 145, 317–329. Glick, E., Zrachya, A., Levy, Y., Mett, A., Gidoni, D., Belausov, E., Citovsky, V., and Gafni, Y. (2008). Interaction with host SGS3 is required for suppression of RNA silencing by Tomato yellow leaf curl virus V2 protein. Proc. Natl. Acad. Sci. U.S.A. 105, 157–161. https://doi.org/10.1073/pnas.0709036105 Golem, S., and Culver, J.N. (2003). Tobacco mosaic virus induced alterations in the gene expression profile of Arabidopsis thaliana. Mol. Plant Microbe Interact. 16, 681–688. https://doi.org/10.1094/ MPMI.2003.16.8.681 Gomez-Ariza, J., Campo, S., Rufat, M., Estopa, M., Messeguer, J., San Segundo, B., and Coca, M. (2007). Sucrose-mediated priming of plant defence responses and broad spectrum disease resistance by overexpression of the maize pathogenesis related PRms protein in rice plants. Mol. Plant Microbe Interact. 20, 832–842. Gouveia, B.C., Calil, I.P., Machado, J.P., Santos, A.A., and Fontes, E.P. (2016). Immune receptors and co-receptors in antiviral innate immunity in plants. Front Microbiol 7, 2139. https://doi.org/10.3389/fmicb.2016.02139 Guo, H., Song, X., Xie, C., Huo, Y., Zhang, F., Chen, X., Geng, Y., and Fang, R. (2013). Rice yellow stunt rhabdovirus protein 6 suppresses systemic RNA silencing by blocking RDR6-mediated secondary siRNA synthesis. Mol. Plant Microbe Interact. 26, 927–936. https://doi. org/10.1094/MPMI-02-13-0040-R Gururani, M.A., Venkatesh, J., Upadhyaya, C.P., Nookaraju, A., Pandey, S.K., and Park, S.W.(2012). Plant disease resistance genes: current status and future directions. Physiol. Mol. Plant Pathol. 78, 51–65. Haas, G., Azevedo, J., Moissiard, G., Geldreich, A., Himber, C., Bureau, M., Fukuhara, T., Keller, M., and Voinnet, O. (2015). Nuclear import of CaMV P6 is required for infection and suppression of the RNA silencing factor DRB4. EMBO J. 34, 2591–2592. https://doi. org/10.15252/embj.201570060 Hagen, C., Rojas, M.R., Kon, T., and Gilbertson, R.L. (2008). Recovery from Cucurbit leaf crumple virus (family Geminiviridae, genus Begomovirus) infection is an adaptive antiviral response associated with changes in viral small RNAs. Phytopathology 98, 1029–1037. https://doi.org/10.1094/PHYTO-98-9-1029

Hamera, S., Yan, Y., Song, X., Chaudhary, S.U., Murtaza, I., Su, L., Tariq, M., Chen, X., and Fang, R. (2016). Expression of Cucumber mosaic virus suppressor 2b alters FWA methylation and its siRNA accumulation in Arabidopsis thaliana. Biol. Open 5, 1727–1734. https:// doi.org/10.1242/bio.017244 Harvey, J.J., Lewsey, M.G., Patel, K., Westwood, J., Heimstädt, S., Carr, J.P., and Baulcombe, D.C. (2011). An antiviral defense role of AGO2 in plants. PLOS ONE 6, e14639. https://doi.org/10.1371/journal.pone.0014639 Haurwitz, R.E., Jinek, M., Wiedenheft, B., Zhou, K., and Doudna, J.A. (2010). Sequence- and structurespecific RNA processing by a CRISPR endonuclease. Science 329, 1355–1358. https://doi.org/10.1126/ science.1192272 Henderson, I.R., Zhang, X., Lu, C., Johnson, L., Meyers, B.C., Green, P.J., and Jacobsen, S.E. (2006). Dissecting Arabidopsis thaliana DICER function in small RNA processing, gene silencing and DNA methylation patterning. Nat. Genet. 38, 721–725. Henikoff, S., Till, B.J., and Comai, L. (2004). TILLING. Traditional mutagenesis meets functional genomics. Plant Physiol. 135, 630–636. https://doi.org/10.1104/ pp.104.041061 Herbers, K., Meuwly, P., Frommer, W.B., Metraux, J.P., and Sonnewald, U. (1996). Systemic Acquired Resistance Mediated by the Ectopic Expression of Invertase: Possible Hexose Sensing in the Secretory Pathway. Plant Cell 8, 793–803. https://doi.org/10.1105/tpc.8.5.793 Hernández, J.A., Gullner, G., Clemente-Moreno, M.J., Künstler, A., Juhász, C., Díaz-Vivancos, P., and Király, L. (2016). Oxidative stress and antioxidative responses in plant–virus interactions. Physiol. Mol. Plant Pathol. 94, 134–148. Ho, T., Pallett, D., Rusholme, R., Dalmay, T., and Wang, H. (2006). A simplified method for cloning of short interfering RNAs from Brassica juncea infected with Turnip mosaic potyvirus and Turnip crinkle carmovirus. J. Virol. Methods 136, 217–223. Hong, W., Qian, D., Sun, R., Jiang, L., Wang, Y., Wei, C., Zhang, Z., and Li, Y. (2015). OsRDR6 plays role in host defense against double-stranded RNA virus, Rice Dwarf Phytoreovirus. Sci Rep 5, 11324. https://doi. org/10.1038/srep11324 Huang, Y., Li, M.-Y., Wu, P., Xu, Z.-S., Que, F., Wang, F., and Xiong, A.-S. (2016). Members of WRKY Group III transcription factors are important in TYLCV defense signalling pathway in tomato (Solanum lycopersicum). BMC Genomics 17, 788. https://doi.org/10.1186/ s12864-016-3123-2 Huh, S.U., Lee, G.J., Jung, J.H., Kim, Y., Kim, Y.J., and Paek, K.H. (2015). Capsicum annuum transcription factor WRKYa positively regulates defense response upon TMV infection and is a substrate of CaMK1 and CaMK2. Sci Rep 5, 7981. https://doi.org/10.1038/ srep07981 Hunter, L.J., Westwood, J.H., Heath, G., Macaulay, K., Smith, A.G., Macfarlane, S.A., Palukaitis, P., and Carr, J.P. (2013). Regulation of RNA-dependent RNA polymerase 1 and isochorismate synthase gene expression in Arabidopsis. PLOS ONE 8, e66530. https://doi.org/10.1371/journal.pone.0066530

18  | Rey and Fondong

Huot, B., Yao, J., Montgomery, B.L., and He, S.Y. (2014). Growth-defense tradeoffs in plants: a balancing act to optimize fitness. Mol Plant 7, 1267–1287. Ibiza, V.P., Cañizares, J., and Nuez, F. (2010). EcoTILLING in Capsicum species: searching for new virus resistances. BMC Genomics 11, 631. https://doi. org/10.1186/1471-2164-11-631 Ivanov, K.I., Eskelin, K., Bašić, M., De, S., Lõhmus, A., Varjosalo, M., and Mäkinen, K. (2016). Molecular insights into the function of the viral RNA silencing suppressor HCPro. Plant J. 85, 30–45. https://doi. org/10.1111/tpj.13088 Jaubert, M., Bhattacharjee, S., Mello, A.F., Perry, K.L., and Moffett, P. (2011). ARGONAUTE2 mediates RNAsilencing antiviral defenses against Potato virus X in Arabidopsis. Plant Physiol. 156, 1556–1564. https://doi. org/10.1104/pp.111.178012 Jiang, L., Qian, D., Zheng, H., Meng, L.Y., Chen, J., Le, W.J., Zhou, T., Zhou, Y.J., Wei, C.H., and Li, Y. (2012). RNAdependent RNA polymerase 6 of rice (Oryza sativa) plays role in host defense against negative-strand RNA virus, Rice stripe virus. Virus Res. 163, 512–519. https:// doi.org/10.1016/j.virusres.2011.11.016 Jiménez, I., López, L., Alamillo, J.M., Valli, A., and García, J.A. (2006). Identification of a Plum pox virus CI-interacting protein from chloroplast that has a negative effect in virus infection. Mol. Plant Microbe Interact. 19, 350– 358. https://doi.org/10.1094/MPMI-19-0350 Jin, H., and Zhu, J.K. (2010). A viral suppressor protein inhibits host RNA silencing by hooking up with Argonautes. Genes Dev. 24, 853–856. https://doi. org/10.1101/gad.1927310 Jinek, M., East, A., Cheng, A., Lin, S., Ma, E., and Doudna, J. (2013). RNA-programmed genome editing in human cells. Elife 2, e00471. https://doi.org/10.7554/ eLife.00471 Jones, J.D., and Dangl, J.L. (2006). The plant immune system. Nature 444, 323–329. https://doi.org/10.1038/ nature05286 Kanazawa, A., Inaba, J.I., Shimura, H., Otagaki, S., Tsukahara, S., Matsuzawa, A., Kim, B.M., Goto, K., and Masuta, C. (2011). Virus-mediated efficient induction of epigenetic modifications of endogenous genes with phenotypic changes in plants. Plant J. 65, 156–168. https://doi. org/10.1111/j.1365-313X.2010.04401.x Kangasjärvi, S., Neukermans, J., Li, S., Aro, E.M., and Noctor, G. (2012). Photosynthesis, photorespiration, and light signalling in defence responses. J. Exp. Bot. 63, 1619–1636. https://doi.org/10.1093/jxb/err402 Khatabi, B., Arikit, S., Xia, R., Winter, S., Oumar, D., Mongomake, K., Meyers, B.C., and Fondong, V.N. (2016). High-resolution identification and abundance profiling of cassava (Manihot esculenta Crantz) microRNAs. BMC Genomics 17, 85. https://doi. org/10.1186/s12864-016-2391-1 Király, L., Hafez, Y.M., Fodor, J., and Király, Z. (2008). Suppression of Tobacco mosaic virus-induced hypersensitive-type necrotization in tobacco at high temperature is associated with down-regulation of NADPH oxidase and superoxide and stimulation of dehydroascorbate reductase. J. Gen. Virol. 89,799–808. Komatsu, K., Hashimoto, M., Oseki, J., Yamaji, Y., Maejima, K., Senshu, H., Himeno, M., Kagiwada, S., and Namba,

S. (2010). Virus-induced systemic necrosis in plants involve both programmed cell death and the inhibition of viral multiplication which are regulated by independent pathways. Mol. Plant Microbe Interact. 23, 283–293. Koukiekolo, R., Jakubek, Z.J., Cheng, J., Sagan, S.M., and Pezacki, J.P. (2009). Studies of a viral suppressor of RNA silencing p19-CFP fusion protein: a FRET-based probe for sensing double-stranded fluorophore tagged small RNAs. Biophys. Chem. 143, 166–169. https:// doi.org/10.1016/j.bpc.2009.05.001 Kovacs, I., Durner, J., and Lindermayr, C. (2015). Crosstalk between nitric oxide and glutathione is required for NONEXPRESSOR OF PATHOGENESIS-RELATED GENES 1 (NPR1)-dependent defense signalling in Arabidopsis thaliana. New Phytol. 208, 860–872. Kurowska, M., Daszkowska-Golec, A., Gruszka, D., Marzec, M., Szurman, M., Szarejko, I., and Maluszynski, M. (2011). TILLING: a shortcut in functional genomics. J. Appl. Genet. 52, 371–390. https://doi.org/10.1007/ s13353-011-0061-1 Kliebenstein, D.J., and Osbourn, A. (2012). Making new molecules - evolution of pathways for novel metabolites in plants. Curr. Opin. Plant Biol. 15, 415–423. https:// doi.org/10.1016/j.pbi.2012.05.005 Krenz, B., Jeske, H., and Kleinow, T. (2012). The induction of stromule formation by a plant DNA-virus in epidermal leaf tissues suggests a novel intra- and intercellular macromolecular trafficking route. Frontiers in Plant Sci. 3, 1–12. https://doi.org/10.3389/fpls.2012.00291 Lam, E., Kato, N., and Lawton, M. (2001). Programmed cell death, mitochondria and the plant hypersensitive response. Nature 411, 848–853. https://doi. org/10.1038/35081184 Landeo-Ríos, Y., Navas-Castillo, J., Moriones, E., and Cañizares, M.C. (2016). The p22 RNA silencing suppressor of the crinivirus Tomato chlorosis virus preferentially binds long dsRNAs preventing them from cleavage. Virology 488, 129–136. https://doi. org/10.1016/j.virol.2015.11.008 Lapidot, M., Gafny, R., Ding, B., Wolf, S., Lucas, W.J., and Beachy, R.N. (1993). A dysfunctional movement protein of Tobacco mosaic virus that partially modifies the plasmodesmata and limits virus spread in transgenic plants. Plant J. 4, 959–970. Lecoq, H.B., Lemaire, J.M., and Wipf-Scheibel, C. (1991). Control of zucchini mosaic virus in squash by crossprotection. Plant Dis. 75, 208–211. Lecoq, H., Moury, B., Desbiez, C., Palloix, A., and Pitrat, M. (2004). Durable virus resistance in plants through conventional approaches: a challenge. Virus Res. 100, 31–39. https://doi.org/10.1016/j.virusres.2003.12.012 Lee, H.A., Lee, H.Y., Seo, E., Lee, J., Kim, S.B., Oh, S., Choi, E., Choi, E., Lee, S.E., and Choi, D. (2017). Current understandings of plant nonhost resistance. Mol. Plant Microbe Interact. 30, 5–15. https://doi.org/10.1094/ MPMI-10-16-0213-CR Li, Z., Liang, W.-S., and Carr, J.P. (2014). Effects of modifying alternative respiration on nitric oxideinduced virus resistance and PR1 protein accumulation. J. Gen. Virol. 95, 2075–2081. https://doi.org/10.1099/ vir.0.066662-0 Li, Y., Lu, Y.G., Shi, Y., Wu, L., Xu, Y.J., Huang, F., Guo, X.Y., Zhang, Y., Fan, J., Zhao, J.Q., et al. (2014). Multiple

Mechanisms of Virus Resistance in Plants |  19

rice microRNAs are involved in immunity against the blast fungus Magnaporthe oryzae. Plant Physiol. 164, 1077–1092. https://doi.org/10.1104/pp.113.230052 Li, F., Pignatta, D., Bendix, C., Brunkard, J.O., Cohn, M.M., Tung, J., Sun, H., Kumar, P., and Baker, B. (2012). MicroRNA regulation of plant innate immune receptors. Proc. Natl. Acad. Sci. U.S.A. 109, 1790–1795. https:// doi.org/10.1073/pnas.1118282109 Lin, S.-S., Henriques, R., Wu, H.-W., Niu, Q.-W., Yeh, S.-D., and Chua, N.-H. (2007a). Strategies and mechanisms of plant virus resistance. Plant Biotechnol. Rep. 1, 125–134. Lin, S.-S., Wu, H.W., Jan, F.J., Hou, R.F., and Yeh, S.D. (2007b). Modifications of the HC-Pro of zucchini yellow mosaic potyvirus for generation of attenuated mutants for cross-protection against severe infection. Phytopathol. 97, 287–296. https://doi.org/10.1094/ PHYTO-97-3-0287 Liu, P.-P., von Dahl, C.C., Park, S.-W., and Klessig, D.F. (2011). Interconnection between methyl salicylate and lipid-based long distance signalling during the development of systemic acquired resistance in Arabidopsis and tobacco. Plant Physiol. 155, 1762–1768. https://doi.org/10.1104/pp.110.171694 Liu, J., Yang, J., Bi, H., and Zhang, P. (2014). Why mosaic? Gene expression profiling of African Cassava mosaic virus-infected cassava reveals the effect of chlorophyll degradation on symptom development. J. Integr. Plant Biol. 56, 122–132. https://doi.org/10.1111/jipb.12133 Liu, D., Hu, R., Palla, K.J., Tuskan, G.A., and Yang, X. (2016). Advances and perspectives on the use of CRISPR/Cas9 systems in plant genomics research. Curr. Opin. Plant Biol. 30, 70–77. https://doi.org/10.1016/j. pbi.2016.01.007 Llave, C. (2016). Dynamic cross-talk between host primary metabolism and viruses during infections in plants. Curr. Opin. Virol. 19, 50–55. https://doi.org/10.1016/j. coviro.2016.06.013 Loebenstein, G., Cohen, J., Shabtai, S., Coutts, R.H., and Wood, K.R. (1977). Distribution of Cucumber mosaic virus in systemically infected tobacco leaves. Virology 81, 117–125. Love, A.J., Geri, C., Lird, J., Carr, C., Yun, B.-W., Loake, G.J., Tada, Y., Sadanandom, A., and Milner, J.J. (2012). Cauliflower mosaic virus protein P6 inhibits signalling responses to salicylic acid and regulates innate immunity. PLOS ONE 7:e47535. https://doi.org/10.1371/ journal.pone.0047535 Louis, B., and Rey, C. (2015). Resistance gene analogs involved in tolerant cassava–geminivirus interaction that shows a recovery phenotype. Virus Genes 51, 393–407. https://doi.org/10.1007/s11262-015-1246-1 Lu, B., Stubbs, G., and Culver, J.N. (1998). Coat protein interactions involved in tobacco mosaic tobamovirus cross-protection. Virology 248, 188–198. Lu, J., Du, Z.X., Kong, J., Chen, L.N., Qiu, Y.H., Li, G.F., Meng, X.H., and Zhu, S.F. (2012). Transcriptome analysis of Nicotiana tabacum infected by Cucumber mosaic virus during systemic symptom development. PLOS ONE 7, e43447. https://doi.org/10.1371/ journal.pone.0043447 Ma, X., Nicole, M.C., Meteignier, L.V., Hong, N., Wang, G., and Moffett, P. (2015). Different roles for RNA silencing and RNA processing components in virus recovery and

virus-induced gene silencing in plants. J. Exp. Bot. 66, 919–932. https://doi.org/10.1093/jxb/eru447 Machado, J.P.B., Calil, I.P., Santos, A.A., and Fontes, E.P.B. (2017). Translational control in plant antiviral immunity. Genet. Mol. Biol. 40 (Suppl. 1), 292–304. Maiti, S., Paul, S., and Pal, A. (2011). Non-TIR-NB-LRR in MYMIV resistance. Mol. Biotechnol. 52, 217–233. Mallory, A., and Vaucheret, H. (2010). Form, function, and regulation of ARGONAUTE proteins. Plant Cell 22, 3879–3889. https://doi.org/10.1105/tpc.110.080671 Mandadi, K.K., and Scholthof, K.B.G. (2013). Plant immune responses against viruses: how does a virus cause disease? Plant Cell 25, 1489–1505. Maruthi, M.N., Bouvaine, S., Tufan, H.A., Mohammed, I.U., and Hillocks, R.J. (2014). Transcriptional response of virus-infected cassava and identification of putative sources of resistance for cassava brown streak disease. PLOS ONE 9, e96642. https://doi.org/10.1371/ journal.pone.0096642 Maule, A.J., Caranta, C., and Boulton, M.I. (2007). Sources of natural resistance to plant viruses: status and prospects. Mol. Plant Pathol. 8, 223–231. https://doi. org/10.1111/j.1364-3703.2007.00386.x McCallum, C.M., Comai, L., Greene, E.A., and Henikoff, S. (2000). Targeting induced local lesions IN genomes (TILLING) for plant functional genomics. Plant Physiol. 123, 439–442. McKinney, H.H. (1929). Mosaic diseases in Canary Islands, West Africa, and Gibraltar. J. Agric. Res. 39, 557–578. Menda, N., Semel, Y., Peled, D., Eshed, Y., and Zamir, D. (2004). In silico screening of a saturated mutation library of tomato. Plant J. 38, 861–872. https://doi. org/10.1111/j.1365-313X.2004.02088.x Meng, X., and Zhang, S. (2013). MAPK cascades in plant disease resistance signaling. Annu. Rev. Phytopathol. 51, 245–266. https://doi.org/10.1146/annurevphyto-082712-102314 Miozzi, L., Napoli, C., Sardo, L., and Accotto, G.P. (2014). Transcriptomics of the interaction between the monopartite phloem-limited geminivirus Tomato yellow leaf curl Sardinia virus and Solanum lycopersicum highlights a role for plant hormones, autophagy and plant immune system fine tuning during infection. PLOS ONE 9, e89951. https://doi.org/10.1371/ journal.pone.0089951 Mochizuki, T., Ogata, Y., Hirata, Y., and Ohki, S.T. (2014). Quantitative transcriptional changes associated with chlorosis severity in mosaic leaves of tobacco plants infected with Cucumber mosaic virus. Mol. Plant Pathol. 15, 242–254. Moeder, W., Yoshioka, K., and Klessig, D.F. (2005). Involvement of the small GTPase Rac in the defense responses of tobacco to pathogens. Mol. Plant Microbe Interact. 18, 116–124. https://doi.org/10.1094/ MPMI-18-0116 Moffett, P. (2009). Mechanisms of recognition in dominant R gene mediated resistance. Adv. Virus Res. 75, 1–33. https://doi.org/10.1016/S0065-3527(09)07501-0 Moissiard, G., and Voinnet, O. (2006). RNA silencing of host transcripts by cauliflower mosaic virus requires coordinated action of the four Arabidopsis Dicer-like proteins. Proc. Natl. Acad. Sci. U.S.A. 103, 19593–19598.

20  | Rey and Fondong

Molnár, A., Csorba, T., Lakatos, L., Várallyay, E., Lacomme, C., and Burgyán, J. (2005). Plant virus-derived small interfering RNAs originate predominantly from highly structured single-stranded viral RNAs. J. Virol. 79, 7812–7818. Moshe, A., Pfannstiel, J., Yariv, B., Kolot, M., Sobol, I., Csoznek, H., and Gorovits., R. (2012). Stress responses to Tomato yellow leaf curl virus (TYLCV) infection of resistant and susceptible tomato plants are different. Metabolomics S1:006. https://doi: 10.4172/21530769.S1-006 Mou, Z., Fan, W., and Dong, X. (2003). Inducers of plant systemic acquired resistance regulate NPR1 function through redox changes. Cell 113, 935–944. Moreau, M., Lee, G.I., Wang, Y., Crane, B.R., and Klessig, D.F. (2008). AtNOS/AtNOA1 is a functional Arabidopsis thaliana cGTPase and not a nitric-oxide synthase. J. Biol. Chem. 283, 32957–32967. https://doi. org/10.1074/jbc.M804838200 Moreau, M., Lindermayr, C., Durner, J., and Klessig, D.F. (2010). NO synthesis and signaling in plants – where do we stand? Physiol Plant 138, 372–383. https://doi. org/10.1111/j.1399-3054.2009.01308.x Moury, B., Janzac, B., Ruellan, Y., Simon, V., Ben Khalifa, M., Fakhfakh, H., Fabre, F., and Palloix, A. (2014). Interaction patterns between Potato virus Y and eIF4Emediated recessive resistance in the Solanaceae. J. Virol. 88, 9799–9807. https://doi.org/10.1128/JVI.00930-14 Muller, G.W., and Costa, A.S. (1977). Tristeza control in Brazil by preimmunization with mild strains. Proc. Int. Soc. Citric 3, 868–872. Mur, L.A., Carver, T.L., and Prats, E. (2006). NO way to live; the various roles of nitric oxide in plant-pathogen interactions. J. Exp. Bot. 57, 489–505. https://doi: 10.1093/jxb/erj052 Mur, L.A., Kenton, P., Lloyd, A.J., Ougham, H., and Prats, E. (2008). The hypersensitive response: the centenary is upon us but how much do we know? J. Exp. Bot. 59, 501–520. https://doi: 10.1093/jxb/erm239 Mur, L.A., Prats, E., Pierre, S., Hall, M.A., and Hebelstrup, K.H. (2013). Integrating nitric oxide into salicylic acid and jasmonic acid/ ethylene plant defense pathways. Front. Plant Sci. 4, 215. https://doi.org/10.3389/ fpls.2013.00215 Nicaise, V., German-Retana, S., Sanjuán, R., Dubrana, M.P., Mazier, M., Maisonneuve, B., Candresse, T., Caranta, C., and LeGall, O. (2003). The eukaryotic translation initiation factor 4E controls lettuce susceptibility to the Potyvirus Lettuce mosaic virus. Plant Physiol. 132, 1272–1282. Nicaise, V. (2014). Crop immunity against viruses: outcomes and future challenges. Front. Plant Sci. 5, 660. https://doi.org/10.3389/fpls.2014.00660 Nie, X., and Molen, T.A. (2015). Host recovery and reduced virus level in the upper leaves after Potato virus Y infection occur in tobacco and tomato but not in potato plants. Viruses 7, 680–698. https://doi.org/10.3390/ v7020680 Nieto, C., Piron, F., Dalmais, M., Marco, C.F., Moriones, E., Gómez-Guillamón, M.L., Truniger, V., Gómez, P., Garcia-Mas, J., Aranda, M.A., et al. (2007). EcoTILLING for the identification of allelic variants of melon eIF4E, a

factor that controls virus susceptibility. BMC Plant Biol. 7, 34. Nomura, H., Komori, T., Uemura, S., Kanda, Y., Shimotani, K., Nakai, K., Furuichi, T., Takebayashi, K., Sugimoto, T., Sano, S., et al. (2012). Chloroplast-mediated activation of plant immune signalling in Arabidopsis. Nat. Commun. 3, 926. https://doi.org/10.1038/ncomms1926 Padmanabhan, M.S., Shiferaw, H., and Culver, J.N. (2006). The Tobacco mosaic virus replicase protein disrupts the localization and function of interacting Aux/IAA proteins. Mol. Plant Microbe Interact. 19, 864–873. https://doi.org/10.1094/MPMI-19-0864 Padmanabhan, M.S., and Dinesh-Kumar, S.P. (2010). All hands on deck—the role of chloroplasts, endoplasmic reticulum, and the nucleus in driving plant innate immunity. Mol. Plant Microbe Interact. 23, 1368–1380. https://doi.org/10.1094/MPMI-05-10-0113 Pajerowska-Mukhtar, K.M., Wang, W., Tada, Y., Oka, N., Tucker, C.L., Fonseca, J.P., and Dong, X. (2012). The HSF-like transcription factor TBF1 is a major molecular switch for plant growth-to-defense transition. Curr. Biol. 22, 103–112. https://doi.org/10.1016/j. cub.2011.12.015 Patil, B.L., and Fauquet, C.M. (2015). Studies on differential behavior of cassava mosaic geminivirus DNA components, symptom recovery patterns, and their siRNA profiles. Virus Genes 50, 474–486. https://doi. org/10.1007/s11262-015-1184-y Pazhouhandeh, M., Dieterle, M., Marrocco, K., Lechner, E., Berry, B., Brault, V., Hemmer, O., Kretsch, T., Richards, K.E., Genschik, P., et al. (2006). F-box-like domain in the polerovirus protein P0 is required for silencing suppressor function. Proc. Natl. Acad. Sci. U.S.A. 103, 1994–1999. Peleg-Grossman, S., Melamed-Book, N., Cohen, G., and Levine, A. (2010). Cytoplasmic H2O2 prevents translocation of NPR1 to the nucleus and inhibits the induction of PR genes in Arabidopsis. Plant Signal Behav. 5, 1401–1406. Pieterse, C.M., Leon-Reyes, A., Van der Ent, S., and Van Wees, S.C. (2009). Networking by small-molecule hormones in plant immunity. Nat. Chem. Biol. 5, 308– 316. https://doi.org/10.1038/nchembio.164 Piron, F., Nicolaï, M., Minoïa, S., Piednoir, E., Moretti, A., Salgues, A., Zamir, D., Caranta, C., and Bendahmane, A. (2010). An induced mutation in tomato eIF4E leads to immunity to two potyviruses. PLOS ONE 5, e11313. https://doi.org/10.1371/journal.pone.0011313 Pogany, M., Koehl, J., Heiser, I., Elstner, E., and Barna, B. (2004). Juvenility of tobacco induced by cytokinin gene introduction decreases susceptibility to Tobacco necrosis virus and confers tolerance to oxidative stress. Physiol. Mol. Plant Pathol. 65, 39–47. Price, A.A., Sampson, T.R., Ratner, H.K., Grakoui, A., and Weiss, D.S. (2015). Cas9-mediated targeting of viral RNA in eukaryotic cells. Proc. Natl. Acad. Sci. U.S.A. 112, 6164–6169. https://doi.org/10.1073/ pnas.1422340112 Pumplin, N., and Voinnet, O. (2013). RNA silencing suppression by plant pathogens: defence, counterdefence and counter-counter-defence. Nat. Rev. Microbiol. 11, 745–760. https://doi.org/10.1038/ nrmicro3120

Mechanisms of Virus Resistance in Plants |  21

Qi, X., Bao, F.S., and Xie, Z. (2009). Small RNA deep sequencing reveals role for Arabidopsis thaliana RNAdependent RNA polymerases in viral siRNA biogenesis. PLOS ONE 4, e4971. https://doi.org/10.1371/journal. pone.0004971 Qu, F., Ye, X., and Morris, T.J. (2008). Arabidopsis DRB4, AGO1, AGO7, and RDR6 participate in a DCL4-initiated antiviral RNA silencing pathway negatively regulated by DCL1. Proc. Natl. Acad. Sci. U.S.A. 105, 14732–14737. https://doi.org/10.1073/pnas.0805760105 Raja, P., Jackel, J.N., Li, S., Heard, I.M., and Bisaro, D.M. (2014). Arabidopsis double-stranded RNA binding protein DRB3 participates in methylation-mediated defense against geminiviruses. J. Virol. 88, 2611–2622. https://doi.org/10.1128/JVI.02305-13 Raja, P., Sanville, B.C., Buchmann, R.C., and Bisaro, D.M. (2008). Viral genome methylation as an epigenetic defense against geminiviruses. J. Virol. 82, 8997–9007. https://doi.org/10.1128/JVI.00719-08 Robaglia, C., and Caranta, C. (2006). Translation initiation factors: a weak link in plant RNA virus infection. Trends Plant Sci. 11, 40–45. Rodríguez-Negrete, E.A., Carrillo-Tripp, J., and Rivera-Bustamante, R.F. (2009). RNA silencing against geminivirus: complementary action of posttranscriptional gene silencing and transcriptional gene silencing in host recovery. J. Virol. 83, 1332–1340. https://doi.org/10.1128/JVI.01474-08 Rojas, C.M., Senthil-Kumar, M., Tzin, V., and Mysore, K.S. (2014). Regulation of primary plant metabolism during plant-pathogen interactions and its contribution to plant defense. Front. Plant Sci. 5, 17. https://doi. org/10.3389/fpls.2014.00017 Ronald, P.C., and Shirasu, K. (2012). Front-runners in plant-microbe interactions. Curr. Opin. Plant Biol. 15, 345–348. https://doi.org/10.1016/j.pbi.2012.06.001 Rudolph, C., Schreier, P.H., and Uhrig, J.F. (2003). Peptide-mediated broad-spectrum plant resistance to tospoviruses. Proc. Natl. Acad. Sci. U.S.A. 100, 4429– 4434. https://doi.org/10.1073/pnas.0730832100 Ruiz-Ferrer, V., and Voinnet, O. (2009). Roles of plant small RNAs in biotic stress responses. Annu. Rev. Plant Biol. 60, 485–510. https://doi.org/10.1146/annurev. arplant.043008.092111 Sade, D., Brotman, Y., Eybishtz, A., Cuadros-Inostroza, A., Fernie, A.R., Willmitzer, L., and Czosnek, H. (2013). Involvement of the hexose transporter gene LeHT1 and of sugars in resistance of tomato to Tomato yellow leaf curl virus. Mol. Plant 6, 1707–1710. https://doi. org/10.1093/mp/sst036 Sade, D., Shriki, O., Cuadros-Inostroza, A., Tohge, T., Semel, Y., Haviv, Y., Willmitzer, L., Fernie, A.R., Czosnek, H., and Brotman, Y. (2014). Comparative metabolomics and transcriptomics of plant response to Tomato yellow leaf curl virus infection in resistant and susceptible cultivars. Metabolomics 11, 81–97. https://doi:10.1007/s11306014-0670-x Sahu, P.P., Rai, N.K., Chakraborty, S., Singh, M., Chandrappa, P.H., Ramesh, B., Chattopadhyay, D., and Prasad, M. (2010). Tomato cultivar tolerant to Tomato leaf curl New Delhi virus infection induces virus-specific short interfering RNA accumulation and defence-associated

host gene expression. Mol. Plant Pathol. 11, 531–544. https://doi.org/10.1111/j.1364-3703.2010.00630.x Sarkar, T.S., Majumdar, U., Roy, A., Maiti,D., Goswamy, M.G., Bhattacharjee, A., Ghosh, S.K., and Sanjay Ghosh, S. (2010). Production of Nitric Oxide in host-virus interaction. A case study with a compatible begomovirus-kenaf host-pathosystem. Plant Signal. Behav. 5, 668–676. Sano, H., Seo, S., Koizumi, N., Niki, T., Iwamura, H., and Ohashi, Y. (1996). Regulation by cytokinins of endogenous levels of jasmonic and salicylic acids in mechanically wounded tobacco plants. Plant Cell Physiol. 37,762–769. Sastry, S.K., and Zitter, T.A. (2014). Management of virus and viroid diseases of crops in the tropics. In Plant Virus and Viroid Diseases in the Tropics, Vol.2, Epidemiology and Management (Dordrecht: Springer), pp. 149–480. Seo, S., Okamoto, M., Iwai, T., Iwano, M., Fukui, K., Nakajima, N., and Ohashi, Y. (2000). Reduced levels of chloroplast FtsH protein in Tobacco mosaic virus infected leaves accelerate the hypersensitive response. Plant Cell 12, 917–932. Seo, J.K., Wu, J., Lii, Y., Li, Y., and Jin, H. (2013). Contribution of small RNA pathway components in plant immunity. Mol. Plant Microbe Interact. 26, 617–625. https://doi. org/10.1094/MPMI-10-12-0255-IA Serrano, I., Audran, C., and Rivas, S. (2016). Chloroplasts at work during plant innate immunity. J. Exp. Bot. 67, 3845–3854. https://doi.org/10.1093/jxb/erw088 Sewelam, N., Kazan, K., and Schenk, P.M. (2016). Global plant stress signaling: reactive oxygen species at the cross-road. Front. Plant Sci. 7, 187. https://doi. org/10.3389/fpls.2016.00187 Shalitin, D., and Wolf, S. (2000). Cucumber mosaic virus infection affects sugar transport in melon plants. Plant Physiol. 123, 597–604. Shan, L., He, P., Li, J., Heese, A., Peck, S.C., Nürnberger, T., Martin, G.B., and Sheen, J. (2008). Bacterial effectors target the common signalling partner BAK1 to disrupt multiple MAMP receptor-signalling complexes and impede plant immunity. Cell Host Microbe 4, 17–27. Shivaprasad, P.V., Chen, H.M., Patel, K., Bond, D.M., Santos, B.A., and Baulcombe, D.C. (2012). A microRNA superfamily regulates nucleotide binding site-leucinerich repeats and other mRNAs. Plant Cell 24, 859–874. https://doi.org/10.1105/tpc.111.095380 Silhavy, D., Molnár, A., Lucioli, A., Szittya, G., Hornyik, C., Tavazza, M., and Burgyán, J. (2002). A viral protein suppresses RNA silencing and binds silencinggenerated, 21- to 25-nucleotide double-stranded RNAs. EMBO J. 21, 3070–3080. https://doi.org/10.1093/ emboj/cdf312 Singh, V., Louis, J., Ayre, B.G., Reese, J.C., Pegadaraju, V., and Shah, J. (2011). Trehalose phosphate synthase 11-dependent trehalose metabolism promotes Arabidopsis thaliana defense against the phloem-feeding insect Myzus persicae. Plant J. 67, 94–104. https://doi. org/10.1111/j.1365-313X.2011.04583.x Song, F., and Goodman, R.M. (2001). Activity of nitric oxide is dependent on, but is partially required for function of, salicylic acid in the signalling pathway in tobacco systemic acquired resistance. Mol. Plant Microbe Interact. 14, 1458–1462.

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Spoel, S.H., and Dong, X. (2012). How do plants achieve immunity? Defence without specialized immune cells. Nat. Rev. Immunol. 12, 89–100. https://doi. org/10.1038/nri3141 Sudhamsu, J., Lee, G.I., Klessig, D.F., and Crane, B.R. (2008). The structure of YqeH. An AtNOS1/AtNOA1 ortholog that couples GTP hydrolysis to molecular recognition. J. Biol. Chem. 283, 32968–32976. https:// doi.org/10.1074/jbc.M804837200 Svitkin, Y.V., Herdy, B., Costa-Mattioli, M., Gingras, A.C., Raught, B., and Sonenberg, N. (2005). Eukaryotic translation initiation factor 4E availability controls the switch between cap-dependent and internal ribosomal entry site-mediated translation. Mol. Cell. Biol. 25, 10556–10565. Tada, Y., Spoel, S.H., Pajerowska-Mukhtar, K., Mou. Z., Song, J., Wang, C., Zuo, J., and Dong, X. (2008). Plant immunity requires conformational charges of NPR1 via s-nitrosylation and thioredoxins. Science 321, 952–956. Takahashi, H., Kanayama, Y., Zhang, M.S., Kusano, T., Hase, S., Ikegami, M., and Shah, J. (2004). Antagonistic interactions between the SA and JA signalling pathways in Arabidopsis modulate expression of defense genes and gene-for-gene resistance to Cucumber mosaic virus. Plant Cell Physiol. 45, 803–809. Tena, G., Boudsocq, M., and Sheen, J. (2011). Protein kinase signaling networks in plant innate immunity. Curr. Opin. Plant Biol. 14, 519–529. https://doi.org/10.1016/j. pbi.2011.05.006 Teotia, S., Singh, D., Tang, X., and Tang, G. (2016). Essential RNA-based technologies and their applications in plant functional genomics. Trends Biotechnol. 34, 106–123. https://doi.org/10.1016/j.tibtech.2015.12.001 Thresh, J.M., and Cooter, R.J. (2005). Strategies for controlling Cassava mosaic virus disease in Africa. Plant Pathol. 54, 587–614. https://doi: 10.1111/j.13653059.2005.01282.x Tian, M., Sasvari, Z., Gonzalez, P.A., Friso, G., Rowland, E., Liu, X.M., van Wijk, K.J., Nagy, P.D., and Klessig, D.F. (2015). Salicylic acid inhibits the replication of tomato bushy stunt virus by directly targeting a host component in the replication complex. Mol. Plant Microbe Interact. 28, 379–386. https://doi.org/10.1094/MPMI-09-140259-R Torres, M.A., Jones, J.D., and Dangl, J.L. (2006). Reactive oxygen species signaling in response to pathogens. Plant Physiol. 141, 373–378. Torres, M.A. (2010). ROS in biotic interactions. Physiol Plant 138, 414–429. https://doi.org/10.1111/j.13993054.2009.01326.x Truniger, V., and Aranda, M.A. (2009). Recessive resistance to plant viruses. Adv. Virus Res. 75, 119–159. https:// doi.org/10.1016/S0065-3527(09)07504-6 Tsuda, K., and Somssich, I.E. (2015). Transcriptional networks in plant immunity. New Phytol. 206, 932–947. https://doi.org/10.1111/nph.13286 Vanitharani, R., Chellappan, P., and Fauquet, C.M. (2005). Geminiviruses and RNA silencing. Trends Plant Sci. 10, 144–151. Várallyay, E., Válóczi, A., Agyi, A., Burgyán, J., and Havelda, Z. (2010). Plant virus-mediated induction of miR168 is associated with repression of ARGONAUTE1

accumulation. EMBO J. 29, 3507–3519. https://doi. org/10.1038/emboj.2010.215 Vaucheret, H. (2008). Plant Argonautes. Trends Plant Sci. 13, 350–358. Vidavsky, F., and Czosnek, H. (1998). Tomato breeding lines resistant and tolerant to Tomato yellow leaf curl virus issued from Lycopersicon hirsutum. Phytopathology 88, 910– 914. https://doi.org/10.1094/PHYTO.1998.88.9.910 Visser, M., Maree, H.J., Rees, D.J., and Burger, J.T. (2014). High-throughput sequencing reveals small RNAs involved in ASGV infection. BMC Genomics 15, 568. https://doi.org/10.1186/1471-2164-15-568 Vives, M.C., Velázquez, K., Pina, J.A., Moreno, P., Guerri, J., and Navarro, L. (2013). Identification of a new enamovirus associated with citrus vein enation disease by deep sequencing of small RNAs. Phytopathology 103, 1077–1086. https://doi.org/10.1094/PHYTO03-13-0068-R Vlot, A.C., Klessig, D.F., and Park, S.W. (2008). Systemic acquired resistance: the elusive signal(s). Curr. Opin. Plant Biol. 11, 436–442. https://doi.org/10.1016/j. pbi.2008.05.003 Vlot, A.C., Dempsey, D.A., and Klessig, D.F. (2009). Salicylic Acid, a multifaceted hormone to combat disease. Annu. Rev. Phytopathol. 47, 177–206. https:// doi.org/10.1146/annurev.phyto.050908.135202 Vogler, H., Akbergenov, R., Shivaprasad, P.V., Dang, V., Fasler, M., Kwon, M.O., Zhanybekova, S., Hohn, T., and Heinlein, M. (2007). Modification of small RNAs associated with suppression of RNA silencing by tobamovirus replicase protein. J. Virol. 81, 10379– 10388. Wang, X.B., Wu, Q., Ito, T., Cillo, F., Li, W.X., Chen, X., Yu, J.L., and Ding, S.W. (2010). RNAi-mediated viral immunity requires amplification of virus-derived siRNAs in Arabidopsis thaliana. Proc. Natl. Acad. Sci. U.S.A. 107, 484–489. https://doi.org/10.1073/ pnas.0904086107 Wang, X.B., Jovel, J., Udomporn, P., Wang, Y., Wu, Q., Li, W.X., Gasciolli, V., Vaucheret, H., and Ding, S.W.(2011). The 21-nucleotide, but not 22-nucleotide, viral secondary small interfering RNAs direct potent antiviral defense by two cooperative argonautes in Arabidopsis thaliana. Plant Cell 23, 1625–1638. Wang, M.B., Masuta, C., Smith, N.A., and Shimura, H. (2012). RNA silencing and plant viral diseases. Mol. Plant Microbe Interact. 25, 1275–1285. https://doi. org/10.1094/MPMI-04-12-0093-CR Waterhouse, P.M., Graham, M.W., and Wang, M.B. (1998). Virus resistance and gene silencing in plants can be induced by simultaneous expression of sense and antisense RNA. Proc. Natl. Acad. Sci. U.S.A. 95, 13959–13964. Windram, O., Penfold, C.A., and Denby, K.J. (2014). Network modeling to understand plant immunity. Annu. Rev. Phytopathol. 52, 93–111. https://doi. org/10.1146/annurev-phyto-102313-050103 Wingard, S.A. (1928).Hosts and symptoms of ring spot, a virus disease of plants. J. Agric. Res. 37, 127–153. Xiang, L., Le Roy, K., Bolouri-Moghaddam, M.R., Vanhaecke, M., Lammens, W., Rolland, F., and Van den Ende, W. (2011). Exploring the neutral invertaseoxidative stress defence connection in Arabidopsis

Mechanisms of Virus Resistance in Plants |  23

thaliana. J. Exp. Bot. 62, 3849–3862. https://doi. org/10.1093/jxb/err069 Xie, Z., Johansen, L.K., Gustafson, A.M., Kasschau, K.D., Lellis, A.D., Zilberman, D., Jacobsen, S.E., and Carrington, J.C. (2004). Genetic and functional diversification of small RNA pathways in plants. PLOS Biol. 2, E104. https://doi.org/10.1371/journal. pbio.0020104 Xie, K., Minkenberg, B., and Yang, Y. (2015). Boosting CRISPR/Cas9 multiplex editing capability with the endogenous tRNA-processing system. Proc. Natl. Acad. Sci. U.S.A. 112, 3570–3575. https://doi.org/10.1073/ pnas.1420294112 Xu, Y., Huang, L., Fu, S., Wu, J., and Zhou, X. (2012). Population diversity of Rice stripe virus-derived siRNAs in three different hosts and RNAi-based antiviral immunity in Laodelphgax striatellus. PLOS ONE 7, e46238. https://doi.org/10.1371/journal.pone.0046238 Xu, G., Greene, G.H., Yoo, H., Liu, L., Marqués, J., Motley, J., and Dong, X. (2017a). Global translational reprogramming is a fundamental layer of immune regulation in plants. Nature 545, 487–490. https://doi. org/10.1038/nature22371 Xu, G., Yuan, M., Ai, C., Liu, L., Zhuang, E., Karapetyan, S., Wang, S., and Dong, X. (2017b). uORF-mediated translation allows engineered plant disease resistance without fitness costs. Nature 545, 491–494. https://doi. org/10.1038/nature22372 Yang, X., Xie, Y., Raja, P., Li, S., Wolf, J.N., Shen, Q., Bisaro, D.M., and Zhou, X. (2011). Suppression of methylationmediated transcriptional gene silencing by βC1-SAHH protein interaction during geminivirus-betasatellite infection. PLOS Pathog. 7, e1002329. https://doi. org/10.1371/journal.ppat.1002329 Yeh, S.D., and Gonsalves, D. (1984). Evaluation of induced mutants of Papaya ringspot virus for control by crossprotection. Phytopathology 74, 1086–1091. Yoon, J.Y., Ahn, H.I., Kim, M., Tsuda, S., and Ryu, K.H. (2006). Pepper mild mottle virus pathogenicity determinants and cross-protection effect of attenuated mutants in pepper. Virus Res. 118, 23–30. https://doi. org/10.1016/j.virusres.2005.11.004

Zhai, J., Jeong, D.H., De Paoli, E., Park, S., Rosen, B.D., Li, Y., González, A.J., Yan, Z., Kitto, S.L., and Grusak, M.A. et al. (2011). MicroRNAs as master regulators of the plant NB-LRR defense gene family via the production of phased, transacting siRNAs. Genes Dev. 25, 2540–2553. https://doi.org/10.1101/gad.177527.111 Zhao, J., Zhang, X., Hong, Y., and Liu, Y. (2016). Chloroplast in plant–virus interaction. Front. Microbiol. 7, 1565. https://doi.org/10.3389/fmicb.2016.01565 Zhang, J.F., Yuan, L.J., Shao, Y., Du, W., Yan, D.W., and Lu, Y.T. (2008). The disturbance of small RNA pathways enhanced abscisic acid response and multiple stress responses in Arabidopsis. Plant Cell Environ. 31, 562–574. https://doi.org/10.1111/j.13653040.2008.01786.x Zhang, Y., Fan, W., Kinkema, M., Li, X., and Dong, X. (1999). Interaction of NPR1 with basic leucine zipper protein transcription factors that bind sequences required for salicylic acid induction of the PR-1 gene. Proc. Natl. Acad. Sci. U.S.A. 96, 6523–6528. Zhang, C., Wu, Z., Li, Y., and Wu, J. (2015). Biogenesis, function, and applications of virus-derived small RNAs in plants. Front. Microbiol. 6, 1237. https://doi. org/10.3389/fmicb.2015.01237 Zhang, X., Yuan, Y.R., Pei, Y., Lin, S.S., Tuschl, T., Patel, D.J., and Chua, N.H. (2006). Cucumber mosaic virus-encoded 2b suppressor inhibits Arabidopsis Argonaute1 cleavage activity to counter plant defense. Genes Dev. 20, 3255– 3268. https://doi.org/10.1101/gad.1495506 Zhang, Z., Chen, H., Huang, X., Xia, R., Zhao, Q., Lai, J., Teng, K., Li, Y., Liang, L., Du, Q., et al. (2011). BSCTV C2 attenuates the degradation of SAMDC1 to suppress DNA methylation-mediated gene silencing in Arabidopsis. Plant Cell 23, 273–288. https://doi. org/10.1105/tpc.110.081695 Zvereva, A.S., and Pooggin, M.M. (2012). Silencing and innate immunity in plant defense against viral and nonviral pathogens. Viruses 4, 2578–2597. https://doi. org/10.3390/v4112578

Role of Host Transcription Factors in Modulating Defense Response During Plant–Virus Interaction

2

Saurabh Pandey1, Pranav P. Sahu2, Ritika Kulshreshtha1 and Manoj Prasad1*

1National Institute of Plant Genome Research, New Delhi, India.

2Max Planck Institute for Plant Breeding Research, Cologne, Germany.

*Correspondence: [email protected] https://doi.org/10.21775/9781910190814.02

Abstract Plants are vulnerable to several environmental stresses either biotic or abiotic due to their sessile nature. Consequently, they assimilate various responses to counter and acclimate in ever-changing environments. One of the fascinating response is transcriptional reprogramming of cell leading to defence or stress adaptation. It is imperative to recognize transcription factors which are associated with plant defence responses against biotic agents such as viruses. Members of families belonging to WRKY family of transcription factor, myeloblastosis-related proteins (MYB), basic leucine zipper (bZIP), Apetela2/ethylene-responsive element binding (AP2/ERF), and NAC transcription factors have been shown to be associated with innumerable defence response against plant virus. These TF family members interact directly or indirectly to modulate defence response by activation or repression of downstream signalling pathways. Hence the gaining insight between plant virus and TFs interplay and deciphering the alterations in defence pathway are the prerequisite to engineer crops for tolerance to biotic stresses. In this chapter, we have attempted to summarize the explicit role of TFs in modulating the expression of defence genes during plant–virus interaction.

Introduction Plants are open to several environmental stresses either biotic or abiotic due to their sessile nature. Different viruses, bacteria, fungi and nematodes are the major source of the biotic stress (Van Verk et al., 2009). Plants possess different mechanism to deal with and defend itself against all these stresses. These include specific pathways of defence for necrotrophic and biotrophic pathogen. Defence against necrotrophic pathogens is mainly mediated by jasmonic acid ( JA) pathway, whereas salicylic acid (SA) pathway plays an imperative part against biotrophic pathogens (Dong, 1998; Howe, 2004). Apart from these hormone based signalling network, transcription is the foremost step in gene expression and this process is regulated by transcription factors (TFs) results in either activation or repression of defence gene expression (Fig. 2.1). It is well established that plant transcription factors play a significant part in the transcriptome regulation of various pathways to combat these stresses. In Arabidopsis thaliana more than 1500 TFs have been reported, till date. The TFs contain a DNA binding domain (DBD) which precisely identifies the target DNA sequence establishing a transcriptional complex and thus modulate gene expression (Ikeda and Ohme-Takagi, 2009).

26  | Pandey et al.

Biotic stress Virus

Fungi

Bacteria

Nematodes, insects

Signal perception SA mediated pathway (Biotrophic pathogen)

JA mediated pathway (Necrotrophic pathogen) Activation of Defence related TFs

AP2/ERF

MYB

bZIP

MYC

NAC

WRKY

Activation/repression of target induced/suppressed genes (PR genes) Stress tolerance

Figure 2.1 Schematic representation of transcriptional regulatory cis elements involved in biotic stress tolerance.

Diverse families of transcription factor are regulating plant defence against biotic stress (Table 2.1). Among them important families are WRKY family of transcription factor, myeloblastosis related TF family (MYB), basic leucine zipper family (bZIP), Apetela2/ethylene-responsive element binding family, NAC-TF family having No apical meristem (NAM), Arabidopsis transcription activator factor (ATAF1/2) and cup-shaped cotyledon (CUC). These domains define the different TF families, and act by binding to the DNA cis-elements associated with stress responses (Van verk et al., 2009; Singh et al., 2002). Regulation of different function of TF is an important process for triggering or suppressing signal transduction pathway by their interaction through regulatory molecules. Processes involved in plant immune responses such as pathogenassociated molecular pattern (PAMP)-triggered immunity (PTI) and effector-triggered immunity (ETI) are modulated by TF by interacting with different recognition or signalling proteins (Kaltdorf and Nassem, 2013; Pajerowska-mukhtar et al., 2013; Fu et al., 2012). These transcription factors having different interacting partners which regulate their activity like MAP kinase, which is regulating the WRKY TF activity (Asai et al., 2002).

In the subsequent section, we discuss the role of different transcription factors involved in plant defence response and there mode of interaction with different proteins to regulate the associated process. WRKY transcription factor family protein Structural constituent of WRKY The most important families of transcriptional regulators in plants is WRKY transcription factors which involves in various signalling cascades that regulate numerous plant responses and processes (Rushton et al., 2010). It has DNA binding domain of 60 residues in length denoted as WRKY domain in N-terminal and at C-terminal having zinc-finger structure (Rushton et al., 1996). WRKY transcription factors are classified into three groups on the basis of WRKY domain numbers (Group I, with two domains, Groups II and III, with a single domain) and their Zn fingers arrangement (C2H2 in group I and II while C2HC in Group III). WRKY transcription factors shows high affinity of binding to DNA sequence (C/T)TGAC(T/C) termed as W-box (Eulgem et al., 2000).

Table 2.1  Different transcription factors and their role in plant defence Transcription factor

Members

Plant

Functions

Reference

MYC

NaMYC2

Nicotiana attenuata

Plant resistance

Woldemariam et al. (2013)

AtMYC2

Arabidopsis

JA-induced with disease resistance

Boter et al. (2004)

AtMYC2

Arabidopsis

Induced by JA and negatively and positively regulate pathogen and wound responsive genes. JA-mediated tolerance to herbivore Helicoverpa armigera

Lorenzo et al. (2004)

NaMYC2

Nicotiana attenuata

Resistance to herbivores

Woldemariam et al. (2013)

MYC2-like (JAM) 1, 2 and 3

Nicotiana attenuata

As targets of MYC2 and negatively regulates defence against herbivores

Sasaki-Sekimoto et al. (2013)

RAP2.3 interaction with TGA4 in defence pathway

Arabidopsis

Plant defence

Büttner and Singh (1997)

RAP2.2

Arabidopsis

Resistance to Botrytis cinerea

Zhao et al. (2012)

OsERF922

Oryza sativa

Repressor of resistance to Magnaporthe oryzae

Liu et al. (2012)

Tsil1

Tobacco

Biotic stress and osmotic stress tolerance

Park et al. (2001)

RAV

Tomato

Resistance to R. solanacearum

Li et al. (2011)

SlRAV2

Tomato

Induces SlERF5 expression, increasing tolerance to bacterial wilt

Zhao et al. (2012), Moffat et al. (2012)

ERF5,6 and RAp2.2

Arabidopsis

Positive regulator in resistance to B. cinerea

Zhao et al. (2012), Moffat et al. (2012)

OsERF3

Oryza sativa

Resistance to Chilo suppressalis linked with MAPK

Lu et al. (2011)

OsERF922

Oryza sativa

Negative regulators-RNA i plants resistance to M. oryzae PR gene expression phenyalanine ammonia lyase

Liu et al. (2012)

PDF1.1 and 1.2

Arabidopsis

Activation upon phosphorylation of ERF6 by MPK3/MPK6 Meng et al. (2013) giving resistance to B. cinerea

AaORA

Artemisia annua

Increase in expression of defence related genes

Lu et al. (2013)

XopDtypeIII

Xanthomonas euvesicatoria

Targets and desumolytes SlERF4 preventing ethylene production and ethylene related immunity

Kim et al. (2013)

OPBP1

Oryza sativa

Resistance to Pseudomonas syringae and Phytophthora parasitica

Guo et al. (2004)

AP2/ERF

Ap2/EREBP

Table 2.1 Continued Transcription factor

Members

Plant

Functions

Reference

MYB

AtMYB030

Arabidopsis

Hypocotyl elongation, brassinosteroid pathway

Li et al. (2009), Segarra et al. (2009)

AtMYB44

Arabidopsis

bZIP

Plant defence response against aphid

Liu et al. (2010)

AtMYB060/AtMYB094/AtMYB096 Arabidopsis

Biotic stress response

Cominelli et al. (2005), Seo and Park (2010)

MYB 30

Positive regulator of the HR and resistance to bacteria

Froidure et al. (2010)

AtMYB12/PFG1

Arabidopsis thaliana

Overexpression leads to insect resistance

Misra et al. (2010), Mitsunami et al. (2014), Schenke et al. (2011)

AtMYB96

Arabidopsis thaliana

Arabidopsis activation tagged overexpression leads to pathogen resistance

Seo et al. (2010)

AtBOS1/MYB108

Arabidopsis thaliana

Knock out provides resistance to Alternaria brassicicola, Pseudomonas syringae pv. tomato and Peronospora parasitica

Mengiste et al. (2003)

AtMYB15

Arabidopsis thaliana

Resistance to the green peach aphid

Liu et al. (2010)

OsMYB4

Oryza sativa

Resistance to pathogen attack

Vannini et al. (2006)

TaPIMP1

Triticum aestivum

Overexpression induce resistance to Ralstonia solanacearum and Bipolaris sorokiniana

Liu et al. (2011), Zang et al. (2012)

SpMYB

Solanum pimpinellifolium

Overexpression leads to resistance to Alternaria alternata

Li et al. (2014)

CabZIP1

Capsicum annum

After pathogen infection, decrease in plant growth

Kuhlmann et al. (2003)

OBF protein

Arabidopsis

Induction of PR gene expression by SA

Sato et al. (1996)

AtbZIP10

Arabidopsis

Cell death and basal defence response

Kaminaka et al. (2006)

TGA members

Arabidopsis

Induction of PR gene expression

Despres et al. (2000)

rTGA2.1, rTGA2.2, rTGA2.3, rLG2, TGAL2, TGAL4

Oryza sativa

Induction of SA-responsive gene expression

Chern et al. (2014)

VvbZIP23

Vitisvinifera

Regulation of biotic and abiotic stress responses

Tak et al. (2013)

PPI1

Capsicum chinense

Regulation of defence gene expression

Lee et al. (2002)

OsTGAP1

Oryza sativa

Regulation of genes involved in the biosynthesis of diterpenoids and plant defence

Okada et al. (2009)

SlAREB1

Solanum lycopersicum

Pathogens response

Orellana et al. (2010)

Table 2.1 Continued Transcription factor

Members

Plant

Functions

Reference

WRKY

HvWRKY1 and HvWRKY2

H. vulgare

Negative regulators of PTI

Shen et al. (2007)

OsWRKY1

Oryza sativa subsp. japonica

Resistance to Magnaporthe grisea

Tao et al. (2009)

OsWRKY2

O. sativa subsp. indica

Resistance to Magnaporthe grisea

Tao et al. (2009)

NaWRKY3 and NaWRKY6

Nicotiana attenuate

Responses to herbivory

Skibbe et al. (2008)

AtWRKY23

Arabidopsis

Nematode resistance

Grunewald et al. (2008)

wrky 33

Arabidopsis

Resistance to Botrytis cinerea and Alternaria brassicicola

Li et al. (2012)

NbWRKY8

Nicotianabenthamiana

Resistance to Phytophthora infestans and Colletotrichumorbiculare

Ishihama et al. (2011)

HvWRKY1,2

Hordeum vulgare

Resistance to Blumeria graminis

Shen et al. (2007)

OsWRKY45

Oryza sativa

Resistance to Magnaporthe oryzae

Inoue et al. (2013)

WRKY52/RRS1

Arabidopsis

Resistance to Ralstonia solanacearum

Deslandes et al. (2002)

WRKY8

Arabidopsis

Negative and positive regulator of basal defence to Pseudomonas syringae and B. cinerea, respectively

Chen et al. (2010)

WRKY11

Arabidopsis

The loss of function conferred resistance to both avirulent Journot-Catalino et al. and virulent P. syringae pv. tomato strains (2006)

CaWRKY1

Capsicum annuum

Silencing disease symptoms caused Xanthomonas axonopodis pv. vesicatoria race 1

Oh et al. (2008)

CaWRKY58

Capsicum annuum

Silencing disease symptoms caused R. solanacearum

Wang et al. (2013)

OsWRKY62.1

Rice

interaction with Xa21, negatively regulates defence as well as in basal defence against Xanthomonas oryzae pv. oryzae

Peng et al. (2008)

OsWRKY45–1, 45–2

Rice

Loss of function gives resistance to X. oryzaepv. oryzae and X. oryzaepv. oryzicola

Tao et al. (2009)

WRKY

Arabidopsis

Induced within 18 h after infection with B. cinerea

Windram et al. (2012)

Arabidopsis

Target of NPR1 (microarray expression data)

Wang et al. (2006)

Table 2.1 Continued Transcription factor

Members

Plant

Functions

Reference

Grape

Expression was altered during powdery mildew Erysiphenecator infection

Guo et al. (2014)

AtWRKY38/48/62

Arabidopsis

Negatively regulate the defence against Pseudomonas syringae

Xing et al. (2008), Kim et al. (2008)

AtWRKY22/29

Arabidopsis

Enhanced resistance by involved in MAPK signalling against P. syringae

Asai et al. (2002)

BdWRKY8/34/50/69/70

Brachpodiumdistachyon

U-pregulated in F. graminearum infection

Wen et al. (2014)

CaWRKY27

Pepper

Upon R. solanacearum infection, positive regulation of SA/JA/ET signalling

Dang et al. (2014)

GhWRKY39–1

Cotton

Overexpression regulates its Resistance to R. solanacearum

Shi et al. (2014)

OsWRKY22

Rice

Resistance to Pyriculariaoryzae Cav.

Cheng et al. (2014)

HvNAC6

Hordeum vulgare

HvNAC6 positively regulates penetration. Resistant towards Blumeriagraminis f. sp. hordei (Bgh) attack

Jensen et al. (2007)

ataf1–1

Arabidopsis thaliana

Loss-of-function mutants have attenuated penetration resistance towards Bgh attack

Jensen et al. (2008)

ATAF1, PR1

A. thaliana

ATAF1 negatively regulates resistance to Botrytis cinerea

Wu et al. (2009)

ATAF1, PR-1, PR-5, NPR1, PDF1.2

A. thaliana

ATAF1 negatively regulates resistance to Pseudomonas syringae, B. cinerea, Alternaria brassicicola

Wang et al. (2009a)

ATAF2, PR1, PR2, PR4, PR5, PDF1.1, PDF1.2

A. thaliana

ATAF2 negatively regulates resistance to Fusarium oxysporum, represses pathogenesis-related proteins

Delessert et al. (2005)

ATAF2, PR1, PR2, PDF1.2

A. thaliana

Reduced Tobacco mosaic virus accumulation, increased pathogenesis-related genes

Wang et al. (2009b)

ATAF2, NIT2

A. thaliana

Defence hormones, pathogen infection

Huh et al. (2012)

WRKY

NAC

ANAC019, ANAC055

A. thaliana

Defence disease, JA pathway

Bu et al. (2008)

NTL6, PR1, PR2, PR5

A. thaliana

Positive regulator of pathogen resistance to P. syringae

Seo et al. (2010)

ANAC042, P450

A. thaliana

Regulation of camalexin biosynthesis, pathogen infection

Saga et al. (2012)

SlNAC1

Nicotiana benthamiana

Increased Tomato leaf curl virus (TLCV) DNA accumulation

Selth et al. (2005)

Table 2.1 Continued Transcription factor

Members

Plant

Functions

Reference

OsNAC4

Oryza sativa

Inducer of HR cell death upon Acidovoraxavenae infection, loss of plasma membrane integrity, nuclear DNA fragmentation

Kaneda et al. (2009)

OsNAC6, PR protein 1,

O. sativa

Slightly increased tolerance to rice blast Disease

Nakashima et al. (2007)

rim1–1

O. sativa

Resistance to Rice dwarf virus (RDV), susceptible to Rice transitory yellowing virus (RTYV) and RSV

Yoshii et al. (2009)

Os02g34970, Os02g38130, Os11g03310, Os11g03370, Os11g05614, Os12g03050

O. sativa

Rice grassy stunt virus, Rice ragged stunt virus, Rice dwarf virus, Rice black-streaked dwarf virus, Rice transitory yellowing virus infections

Nuruzzaman et al. (2010)

OsNAC19

O. sativa

Disease resistance

Lin et al. (2007)

GRAB1, GRAB2

Triticum monococcum

Inhibited Wheat dwarf virus replication

Xie et al. (1999)

ATAF2

Tobacco

Tobacco mosaic virus

Wang et al. (2009b)

ONAC122 and ONAC131 Brome mosaic virus (BMV)

O. sativa

Defence responses against Magnaporthe grisea

Sun et al. (2013)

SlNAC1

Solanum lycopersicum

Upregulated during Pseudomonas Infection

Huang et al. (2012)

CaNAC1

Cicer arietinum

Defence responses against pathogen

Oh et al. (2005)

GmNAC6

Glycine max

Responses to biotic signals, osmotic stress-induced

Faria et al. (2011)

TLCV, SlNAC1

Solanum lycopersicum

Enhances viral replication

Selth et al. (2005)

Stprx2, StNAC

Solanum tuberosum

Wounding and pathogen response

Collinge and Boller (2001)

NAC TF -NTL6

A. thaliana

Binds to promoter genes of PR genes

Seo et al. (2010)

ATAF1 OE

A. thaliana

Imparts tolerance to drought and susceptibility to B. cinerea

Wu et al. (2009)

JA2, JA2-like

Solanum lycopersicum

Pathogen-induced stomatal closure and reopening

Du et al. (2014)

StNTP1, StNTP2

Solanum tuberosum

Interact with Rx effector and suppresses defence responses against P. infestans

McLellan et al. (2013)

TMV replicase

Interact with ATAF2 suppressing basal host defence

Wang et al. (2009)

NTL9

Interact with HopD1 a type III effector from P. syringae suppressing ETI responses

Block et al. (2014)

SPL

NbSPL6 N

N. benthamiana

N-mediated resistance to TMV

Padmanabhan et al. (2013)

CCCHtype zinc-finger protein

C3H12

Rice

Resistance to X. oryzae pv. oryzae

Deng et al. (2012)

32  | Pandey et al.

Functions of WRKY-TF in plant defence WRKY-TF activates or represses transcription based on their promoter context. For example in yeast, Arabidopsis WRKY53 (AtWRKY53) has been reported to activate or repress the reporter gene related to promoter positions (Miao et al., 2004). The results were confirmed in Arabidopsis protoplasts by co-transforming the AtWRKY53 with the reporter constructs, where it negatively regulates its promoter but activates other promoters by acting as activators (Miao et al., 2004). Similarly, OsWRKY72 and OsWRKY77 have been shown by transient expression studies to activate the ABA induction and repress the GA signalling in aleurone layers (Xie et al., 2006). The gene activation role of WRKY transcription factor have been shown in parsley, where PcWRKY1 transcription factor activates the PcPR10 gene (Ulker and Sommisch, 2004). This activation is explained by a model of WRKY and W-box intermediated transcriptional control. The aforementioned, receptor-moderated identification of a pathogen triggers the stimulation of mitogen activated protein kinase (MAPK) pathway, and that eventually results in movement of the protein kinase (MPK) towards the nucleus. In the nucleus it can alter the attached WRKY factors precisely. This contact results into the freeing of WRKY proteins in cytosol and they are replaced by other WRKY transcription factors binds to their respective W-box element and finally activates PcWRKY1 and Pathogen related gene-10 (Fig. 2.2). HvWRKY1 and HvWRKY2 are the examples of the WRKY-TF that act as repressor of genes related to basal defence (Shen et al., 2007). Moreover, MAP kinase pathways are the control point for the WRKY-TF activity, as revealed in various studies (Asai et al., 2002; Miao et al., 2007; Rasmussen et al., 2012). Interestingly, histone modification also assist and act as another control point which may regulate activation and repression of the WRKY transcription factors. AtWRKY38, AtWRKY53, AtWRKY62, AtWRKY 70 have been shown to be regulated by histone modification pattern (Fig. 2.2) such as H3K4me2, H3K4me3, H3K27me2 or H3K27me3 (Alvarez-venegez et al., 2007; Kim et al., 2008; Ay et al., 2009). Hence, WRKY-TF may have an impact on development and defence related genes. Another way of regulation of WRKY is through the small RNAs. For example,

AtWRKY18 have a crucial function in the modulation of defence response in case of powdery mildew (Pandey and Sommisch, 2009). WRKY genes promoters are enriched in W-boxes and provide another way of regulation of WRKY factors that is auto-regulation and cross-regulation (Pandey and Sommisch, 2009). Members of WRKY TFs have been described to be linked with plant defence system through the modulation of transcription of defence contributing genes (Eulgem and Sommisch, 2007). Hence, these are the key control element of plant immune responses and regulate its component ETI, PTI/MTI, basal defence and systemic acquired resistance. Many overexpression and knockdown studies have shown their involvement in plant defence and provide the impetus to decipher their role in the defence signalling networks (Sun et al., 2003, Van verk, 2008, Pandey and Sommisch, 2009). Within these networks MAP kinases targets WRKY proteins to subsequent pathways (Popescu et al., 2009). In barley, resistance gene Mildew resistance locus A (MLA) in cytoplasm requires avirulence factor AVR10 from fungus for recognition and eliciting the ETI response for powdery mildew (Blumeria graminis f. sp. Hordei). MLA was also associated with HvWRKY1 and HvWRKY2 specific functions in the nucleus as shown in (Fig. 2.2) (Shen et al., 2007). Both, HvWRKY1 and HvWRKY2 work as negative controllers of PTI, consequently providing the link between the two important defence signalling pathways: activation of salicylic acidand jasmonic acid/ethylene-mediated signalling pathways regulated via WRKY11 and WRKY 70, respectively, as shown by Jiang et al. (2016) in Arabidopsis. Which modulate the induced systemic resistance in an NPR1-dependent manner triggered by Bacillus cereus AR156 provided evidence for wide-ranging resistance towards Pseudomonas syringae pv. Tomato DC3000 by non-pathogenic rhizobacteria ( Jiang et al., 2016). AtWRKY52/RRS1 have been displayed to convene resistance to bacterial pathogen Ralstonia solanacearum; however, the encoded protein is rather unique then seems to function as an R protein, providing direct involvement of the WRKY factor in defence as shown in Fig. 2.2 (Deslandes et al., 2002). Further, additional components of these pathways identified, which links the WRKY factors with the end stages of the pathway.

Host Transcription Factors in Modulating Defence Response |  33

Tobacco mosaic virus

Pathogen attack

W-box

l tro Con ts n poi

WRKY

MAP kinase Histone modification Small RNAs

AVR10 MAP kinase cascades

Autoregulation and cross-regulation

Interaction with MLA MAPK accumulation PcWRKY1

MLA

activation PcPR10 HvWRKY1

HvWRKY2 ETI responses

WRKY 1-3 N genes

JA Signalling (COL1) Nucleus

Production of PR genes product

Resistance

Figure 2.2  Illustrative representation of mode of action of WRKY TFs. Pathogen interacts with W-box of WRKY TF which activates the signalling cascades of MAP kinase. This induction leads to MAPK accumulation inside the nucleus, where it alters the binding of WRKY to their W-Box elements and finally activates PcWRKY1 and PcPR10 genes and provide resistance response. In barley, avirulence factor AVR10 interacts with Mildew resistance locus A which subsequently act with HvWRKY1 and HvWRKY2 to stimulate the effector-triggered immunity (ETI) responses to provide resistance to powdery mildew. Similarly, Tobacco mosaic virus acts through MAP kinase pathway components, which in turn act with WRKY1–3–N-gene module in tobacco with JA signalling component COL1 to provide resistance to the virus. Control of WRKY-mediated resistance through signalling cascade (MAP Kinase), Histone modification, small RNAs and its autoregulation and cross regulation through salicylic acid/jasmonic acid pathways.

In rice OsWRKY45-1 (allele in japonica) and OsWRKY45–2 (allele in indica), encoding two proteins that differ in 10 amino acids, shows dissimilar functions during rice–bacteria interactions (Tao et al., 2009). They act as positive regulators and shows resistance to Magnaporthe grisea a rice fungal pathogen in overexpression studies (Tao et al., 200; Shimono et al., 2007), but were differentially regulated in response to Xanthomonas oryzae (bacterial rice leaf blight). OsWRKY45-1 act as negative regulator whereas OsWRKY45–2 as positive regulator, in providing resistance. Both alleles work differentially because of their pathway specific function. For example, OsWRKY45-1 controls both SA and JA level; however, OsWRKY45-2 appears to considerably modulate only JA levels. WRKY TF also plays role in modulating insect feeding behaviour. Kloth et al. (2016) reported

WRKY 22 induced by green peach aphid Myzuspersicae through genome-wide association mapping. They have shown Salicylic acid and jasmonic acid interplay by overexpression and knockout plant study, thus making wrky22 a potential target in manipulating aphid infection through host plant defence. Herbivory response also involves WRKY factors, for example in Nicotiana attenuate (a native tobacco) the WRKY gene, NaWRKY3 was essential for NaWRKY6 switching mediated through fatty acid–amino conjugates in oral secretion of Manduca sexta larva. If we silence either or both genes, plants became very susceptible to herbivores. Therefore, these WRKY genes might assist plants to distinguish between mechanical wounding and herbivore attack (Skibbe et al., 2008). Nematode resistance is another dimension for WRKY TF roles

34  | Pandey et al.

in plant defence. AtWRKY23 gene from Arabidopsis is up-regulated at the time of nematode infestation and their knockdown demonstrate improved resistance towards Heterodera schachtii the cyst nematode (Grunewald et al., 2008). The role of WRKY-TF has also been evident during plant–virus interactions. This transcription factor works at a different level of defence pathway, and its interrelation with defence-associated genes has been shown in tobacco plant for N-mediated resistance towards Tobacco mosaic virus. Here silencing of WRKY 1–3 affected the N-mediated resistance along with MYB down-regulation (Fig. 2.2) (Liu et al., 2004). Metabolic pathways manipulation is another way to impart resistance to biotic agents. In potato, the StWRKY1 transcription factor modulates the potato–Phytophthora infestans interface by altering phenyl propanoid pathway genes. Further promoter region analysis revealed that through StWRKY1 binding to promoters of hydroxy cinnamic acid amide (HCAA) biosynthetic genes which are responsible for secondary cell wall synthesis encodes 4-coumarate:CoA ligase and tyramine hydroxycinnamoyl transferase, reinforces the resistance to fungal pathogens (Yogendra et al., 2015). Studying these pathways might provide ways to manipulate these TF for developing plant resistance to viruses. MYB transcription factor family protein Structural constituent of MYB-TF The myeloblastosis-related (MYB) family of proteins is vast, having divergent functions and characterized exclusively in all eukaryotes. Maximum MYB TFs family proteins contain different number of MYB domain repeats which confer capacity to attach to DNA and work as transcription factors. Broad functional classification of MYB in Arabidopsis thaliana (Dubos et al., 2010) showed their involvement in different plant processes. The first MYB gene to be recognized was C1 from Zea mays (Paz-Ares et al., 1987). The MYB proteins possess two dissimilar regions, a preserved MYB DNA-binding domain at N-terminal as well as diverse modulator region at C-terminal accountable for controlling action of the protein. The essential

feature of MYB transcription factor is MYB DNAbinding domain, and the MYB family could be separated into four different classes, based upon number of DNA-binding domain as 1R-, R2R3-, 3R- and 4R-MYB classes of proteins (Dubos et al., 2010, Stracke et al., 2001). MYB domain comprises four imperfect amino acid sequence repeats (R), which are having 52 amino acids of each repeats with three α-helices. Also helix–turn–helix (HTH) structure form by the second and third helices of the repeats with three regularly separate tryptophan (or hydrophobic) residues, which forms a hydrophobic core part in the three-dimensional HTH structure (Ogata et al., 1996). Typical MYB protein c-Myb having three repeats, denoted by means of R1, R2 and R3, in addition other MYB protein repeats are named conferring to their similarity to R1, R2 or R3 of c-MYB (Starcke et al., 2001). Functions of MYB-TF in plant defence R2R3-MYB protein class are plant specific as well as most prolific type in plants, having additionally 100 members of R2R3-MYB in the dicot and monocot plant genomes ( Jiang et al., 2004; Wilkins et al., 2009). But MYB gene family is symbolized by merely five 3R-MYB genes, out of the 190 genes coded for R2R3-MYB proteins in Arabidopsis (Stracke et al., 2001; Yanhui et al., 2006). A genomewide survey by Du et al. (2012a) for R2R3-MYB family of genes in maize genome shows presumed full set of R2R3-MYBs, and included total 157 characteristic R2R3-MYB encrypting genes. Also in Populus genome there are five 3R-MYB genes out of 192 R2R3-MYB genes (Wilkins et al., 2009). In soybean genome 252 MYB genes were reported and classified as R2R-MYB (2R-MYB) genes (244 in numbers), R1R2R3-MYB (3R-MYB) genes (6 in numbers), and R0R1R2R3- MYB (4R-MYB) genes (Du et al., 2012b). MYB TF act by means of positive and negative regulation of the gene expression. MYB30 is an important and well-characterized member of MYB transcription factor family. AtMYB30 expression was enhanced in response to pathogen attack, where it has been recognized as a positive controller of hypersensitive response (HR) performing by modulation of very-long-chain fatty acids (VLCFA) synthesis (Raffaele et al., 2008). Similarly, AtMYB30 has also been shown to

Host Transcription Factors in Modulating Defence Response |  35

via reactive oxygen intermediates after experiencing either biotic or abiotic stress (Mengiste et al., 2003). Moreover, in rice the OsLTR1 has been shown to regulate JA-dependent resistance response; however, AtMYB15, AtMYB34, AtMYB51 and AtMYB75 were related through wound response or defence meant for insect herbivory (Cheong et al., 2002; Johnson and Dowd, 2004). AtMYB102 has been described to be activated to provide defence facing the Pierisrapae (De Vos et al., 2006). However, AtMYB62 has been shown to be induced in feedback to phosphate deprivation (Devaiah et al., 2009), although AtMYB108 induces against biotic as well as abiotic stress responses (Mengiste et al., 2003). One of the members of MYB family AtMYB44 has been found to act in plant resistance response facing the aphid (Liu et al., 2010). Beneficial strains of Pseudomonas spp., Trichoderma, and Rhizobacteria triggered the defence pathways of similar kind where root specific MYB72 TF function as initial convergence point in the signalling pathways and provide resistance in Arabidopsis (Van der Ent et al., 2008; Segarra et al., 2009). In another report, AtMYB96 has been identified as a factor responsible for enhanced pathogen resistance by mediating ABA signals to induce SA-biosynthesis. Hence

interact with other transcription factors to regulate brassinosteroid pathway genes expression which directs hypocotyls cell elongation (Li et al., 2009). The regulation of MYB30 is mediated by MIEL1 (MYB30 interacting E3 Ligase 1) an Arabidopsis ring type E3 ubiquitin ligase interacting at nucleus which expedite the proteasomal degradation and thus its down-regulation. In non-infected plant MIEL1 leads to defence and protect the cell death by degrading MYB30 (Fig. 2.3). However, after bacterial inoculation MIEL1 suppressed and hence MYB30 was expressed and accumulate at inoculation site, thus providing HR and defence against pathogen (Marino et al., 2013). Bacteria evolve to infect plants by targeting some of the transcription factors. XopD, a type III effectors (T3E) from Xanthomonas compestris pv. Vesicatoria targets the MYB30 and inhibit transcriptional activation of MYB30 VLFCA-related targeted genes and suppression of Arabidopsis defence also shown in Fig. 2.3 (Cannonea et al., 2011). The R2R3-MYB-like members of MYB family has linked with JA reliant defence responses. For instance the botrytis susceptible 1 (BOS1) was found to elicit stress response in case of both biotic and abiotic stress signals in Arabidopsis by involving JA signalling pathway and impart reaction to signals

Pathogen

Viral attack

XopD Xcv MYB30 accumulation

MYB30 Suppression

MYB30

NIK1 Suppress the MYB accumulation

VLFCA Genes

NIK1 LIMYB

Tolerance to begomovirus

MIEL1

Cuticular WAX accumulation

Ubiquitination AtMYB30 HR Response Nucleus

Downregulation

Proteasome

Figure 2.3  MYB TF network in biotic stress. Upon pathogen infection MYB 30 is up-regulated and modulate very-long-chain fatty acids (VLFCA) gene responsible for cuticular wax accumulation also induces HR response. MYB30 interacting E3 Ligase 1 (MIEL1) Binds to AtMYB30 and ubiquitination of it leads to degradation followed by suppression of defence response. Same strategy to target TF followed by Xanthomonas compestris pv. Vesicatoria by secretion of type III effectors (T3E) to circumvent the plant defence response. Immune receptor NIK1 interact with LIMYB after viral infection in the nucleus and providing tolerance to begomoviruses.

36  | Pandey et al.

MYB96 links the ABA and SA pathways at the molecular level (Seo and Park, 2010). Involvement of MYB TF in disease resistance was shown by overexpression of Thinopyrum intermedium MYB TF (TiMYB2R1) in wheat. Transgenic plant demonstrates the sequential elevation of the defence-related genes (Liu et al., 2013). MYB TF functions as a negative controller to reduce the viral load and provides viral immunity in A. thaliana (Zorzatto et al., 2015). Herein immune receptor NIK1 which is a leucine-rich repeat receptor-like kinase (LRR-RLK) interact with the LIMYB in the nucleus and leads to the downstream suppression of translation as schematically shown in Fig. 2.3. By overexpression study it was observed that LIMYB can repress the ribosomal genes transcription, which may lead to the protein translation and reduce viral protein association with ribosome, thus providing tolerance to Begomovirus (Zorzatto et al., 2015).

AP2/ERF transcription factor family protein Structural element of AP2/ERF-TF These TFs are members of ERF subfamily of AP2 transcription factor family. The characteristic feature of AP2/ERF TF family proteins is a minimum of one DNA-binding domain (Fig. 2.4), termed the AP2 domain, is further classified into the ERF, AP2 and RAV families of proteins. The proteins having a particular AP2 domain (single domain) whose genomic sequence with minor extent of introns are classified in ERF TF family (Nakano et al., 2006). The characteristic of the AP2/ERF superfamily is conserved DNA binding domain composed of 60 amino acid residues that convene a distinctive 3-D folding and structured into a three antiparallel beta-sheets making a sheet along with a parallel alpha-helix (Allen et al., 1998). AP2/ERF DNA binding domain has been alienated amongst

A)

Upregulation Pathogen attack

Chitinase

AP2/ERF

Osmotin

Botrytis cinerea

PR genes

ET responses

Defense responses

Defence related genes

JA/SA/Ethylene pathway modulation

MAPK

RAP2.2 AP2

TPR/TP L B3

Activation

GCC box elements

GmERF3

Defence to TMV

Target genes

B) AP2

Ub

AP2

AP2 AP2

Proteasome B3

Nucleus

Figure 2.4  Illustration of AP2/ERF TF in plant defence. (A) After the pathogen attacks AP2/ERF transcription factors are induced and upregulate various genes such as defence-related genes, chitinase, osmotin, PR genes to support plant defence. AP2/ERF can act as transcriptional repressor when it interact with co-repressor Topless-related (TPR) and Topless (TPL) with B3 repression domain to further repress the target genes. By upregulation of MAPK and JA/SA/Ethylene pathway It will attaches to GCC box elements and directly activate the resistance genes as GmERF3 activation leads to resistance to Tobacco mosaic virus. Also Botrytis cinerea infection activates the RAP2.2 which induces defence responses against it. AP2/ERF is regulated by its ubiquitination through 26s proteosomal pathway, which leads to its degradation. B) Domain structure of AP2, ERF and RAV members of AP2/ERF family.

Host Transcription Factors in Modulating Defence Response |  37

DREB and ERF domains contained by the ERF family depending on the uniqueness of residues at precise sites (Sakuma et al., 2002). Alterations in DNA affinity as well as specificity of these two subfamilies attributed to their amino acid sequence difference. Further it has been shown that various DREB proteins interact with A/GCCGAC element, frequently related with ABA, drought and cold receptive genes (Stockinger et al., 1997). On other hand, the ERF subfamily members precisely interact in vitro with the AGCCGCC element, known as the GCC-box, that is frequently present upstream of genes that react with ethylene, different pathogens and also in case of wounding (OhmeTakagi and Shinshi, 1995). Functions of AP2/ERF-TF in plant defence The AP2 transcription factor family have genes that encode proteins involved in the regulation of disease resistance pathways. AP2/ERF-TFs can act as either activators or repressors subject to whether they will activate or repress transcription of their target gene. Moreover, transcriptional repressors can be defined as active repressors, which comprise repression domain (RD) with repression action to DNA binding domain, or passive repressors, which suppress the transcription by competing for binding of target sequence with the activators (Licausi et al., 2013). Activation domains observed in AP2-TF are usually acidic; for example, in tobacco protoplast the tobacco ERF2 and ERF4 N-terminal and/or C-terminal acidic parts turn as activation domains (Ohta et al., 2000). Repression domains (RDs) are distinct and have three plant-specific RDs, namely the TLLLFR motif (Matsui et al., 2008), ERF-associated amphiphilic repression motif (EAR: LxLxLx/DLNxxP) (Ohta et al., 2001, Hiratsu et al., 2003), and the B3 repression domain (BRD; RLFGV) (Ikeda and Ohme-Takagi, 2009). ERFs proteins that comprise an EAR/BRD-motif act as transcriptional repressors and by acting together through the transcriptional corepressors Topless-related (TPR) and Topless (TPL), suppress the target genes transcription (Causier et al., 2012). Also, different Arabidopsis proteins have EAR-like motifs for example AP2–8 (ANT-LIKE 6, AIL6) AP2–12 (ANT-LIKE7, AIL7), AP2–13 (AP2) and AP2–14 (TARGET OF EAT 1, TOE1). While AP2–13 positively regulates the floral development,

it works as negative controller in case of AGAMOUS gene (Drews et al., 1991). Modulation at the post-transcriptional level is another way of control over the AP2 TFs. It involves alternate splicing, protein level control and protein stability control by the 26S proteasomal pathways. Alternate splicing has been reported as one of the major cause in the rapid build-up of DREB2-like sequences in different grass family members (Xue and Loveridge, 2004; Egawa et al., 2006; Qin et al., 2007). Moreover, for members of the ERF-VII group in Arabidopsis as well as in tomato (Solanum lycopersicum) alternative splicing has been reported (Pirrello et al., 2006; Licausi et al., 2010). Protein stability is another method of AP2 TFs regulation; for example, it has been proposed tht the ERF-VI family of cytokinin-responsive factors (CRFs) is involved in phosphorylation which is facilitated through histidine kinases and histidine-containing phosphotransfer proteins and their nuclear transfer from the cytosol (Rashotte et al., 2006). It has been reported that ERF-IXb group member AtERF104 works as a positive regulator in response to pathogen in Arabidopsis, and steadied by its combination with MPK6 (Bethke et al., 2009). Phosphorylation negatively regulates the transcription of different homologues of DREB2A (Sakuma et al., 2006; Agarwal et al., 2007). 26S Proteasome also regulates AP2/ERF proteins and this kind of regulation has been reported in DREB family. Indeed, the richness of the Arabidopsis DREB2A was moderated by the DRIP1 and DRIP2 the RING-E3 ligases in non-stress environments (Qin et al., 2007). Related mechanism triggers for proteosomal degradation of drought-responsive Arabidopsis ERF53 upon ubiquitination by RGL1 and RGL2 (Cheng et al., 2012). Disease-associated stimuli such as salicylic acid, ethylene (ET), and jasmonic acid, and infection by virulent or avirulent pathogens (Fig. 2.4) play a part in regulation and have been shown for a number of ERF genes (De vos et al., 2005). Several ERF genes show resistance/tolerance to different biotic stresses upon ectopic expression in discrete plants. For example, a number of ERFs stimulate the transcription of different kind of defence-related genes, chitinase, osmotin, pathogenesis-related (PR) genes and β-1,3-glucanase (Gutterson and Reuber, 2004) (Fig. 2.4). Study of ectopic expression of GmERF3 gene which is an AP2/ERF factor in transgenic tobacco plants

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induces the PR genes expression and higher resistance to different pathogens such as Tobacco mosaic virus, Ralstonia solanacearum and Alternaria alternate (Zhang et al., 2009). Although the target gene core which is controlled by each ERF has not been entirely elucidated, many ERF genes, such as ERF1, as well as its different homologues from ERF-IX group has been comprehensively characterized as probable candidate involved in pathogen responses (Lorenzo et al., 2003; Zarei et al., 2011; Moffat et al., 2012). Furthermore, ethylene (ET) biosynthesis regulated by various ERF proteins as reported by Zhang et al. (2009) and Li et al. (2011). OPBP1 an ERF protein from tobacco improves resistance to pathogens after their ectopic expression in rice (Chen and Guo, 2008). Moreover, RAP2.2 an Arabidopsis gene provide resistance to B. cinerea and ET responses represented in Fig. 2.4 (Zhao et al., 2012) and rice OsERF922 work as negative regulator (Fig. 2.4) for providing resistance to Magnaporthe oryzae by decreasing the level of genes of phytoalexin biosynthetic pathway (Liu et al., 2012). Similarly, tomato Pti5 transcriptional up-regulation was shown against the potato aphid Macrosiphum euphorbiae (Thomas) invasion, and provide augmented plant defence response to control the population growth of the insect (Wu et al., 2015). In conclusion, condition-specific expression of AP2/ERF transcription factor offers a way to impart stress tolerance against plant viruses through modulation of ethylene, jasmonic and salicylic acid pathways. bZIP transcription factor family Structural element of bZIP-TF The bZIP domain containing transcription factor family is one of biggest families of transcriptional factors in eukaryotes. bZIP transcription factors involved in the regulation of various aspects like floral development, seed maturation, abiotic stress and mediating defence responses to counter pathogens in plants ( Jakoby et al., 2002). Members of bZIP TF family proteins in Arabidopsis (AtbZIPs) divided into 10 individual groups: A, B, C, D, E, F, G, H, I and S ( Jakoby et al., 2002) each group having similar sequences at basic region with common features like leucine zipper domain size and position in the protein.

Functions of bZIP-TF in plant defence Proteins of bZIP-TF family interact specifically with other proteins for the regulation of its activity, cellular compartment localization and role in the course of resistance responses against biotic agents have been described ( Jakoby et al., 2002; Alves et al., 2013). The cis-element-binding TGA (TGACGTCA) proteins, is a Group D member of bZIP proteins in Arabidopsis, which act in the SA mediated signalling and associated with responses against biotic stress ( Jakoby et al., 2002). During pathogen responses TGA members act together with the ankyrin repeat protein family members, precisely NPR1 (non-expresser of PR-1), which are the crucial constituents in the defence network intermediated by SA signalling (Fu et al., 2012; Kaltdorf et al., 2013; Pajerowska-mukhtar et al., 2013). Moreover during SA signalling, translocation of NPR1 to the nucleus is the key regulatory step. In normal conditions nearly all NPR1 is maintained as an oligomer in the cytoplasm by the formation of intermolecular disulfide bonds (Fu et al., 2012; Pieterse et al., 2012). When pathogen attacks, synthesis of SA trigger modifications in the cellular redox state via ROS formation (Fu et al., 2012; Pieterse et al., 2012), which bring on monomerization of NPR1 by action of H3 and H5 thioredoxins (TRXH3/H5). However, in cells induced with SA, NPR1 in monomeric form transfer into nucleus by nuclear pore complex (Fig. 2.5) (Fu et al., 2012; Kaltdorf et al., 2013; Pajerowska-mukhtar et al., 2013; Pieterse et al., 2012). Moreover, NPR1 monomers attaches to promoter region of SA-responsive genes by the help of interaction with TGA family members (bZIP). During the translocation and interaction, NPR1 gets phosphorylated and after that E3 ubiquitin ligase ubiquitinate it because it has a great affinity for phosphorylated form of NPR1. Ubiquitination of NPR1 leads to its degradation mediated through proteasome complex. Protein homologues of NPR1, (NPR3 and NPR4) work like SA receptor in degradation process. Here NPR3/NPR4 work as Cullin 3 E3 ubiquitin ligase adapter, that enable ubiquitination followed by degradation of NPR1 and are modulated by SA (Fig. 2.5) (Fu et al., 2012; Kaltdorf et al., 2013; Pajerowska-Mukhtar et al., 2013; Pieterse et al., 2012). Mutant studies in Arabidopsis provided knowledge about new interacting partners for TGA.

Host Transcription Factors in Modulating Defence Response |  39

Redox potential change ROS Formation NPR1

Pathogen Salicylic acid

NPR1 TRX-H3/H5 Monomerization

P NPR1 bZIP

Ub

Tolerance to viruses

NPR3/4

SA Responsive genes Nucleus

Proteasome

Figure 2.5  Simplified representation of bZIP TF family in plant defence. Upon pathogen infection salicylic acid synthesis occur and it changes the redox potential of cell via ROS formation leading to monomerization of nonexpressor of PR-1 (NPR1) by action of H3 and H5 thioredoxins (TRX-H3/H5). This monomer NPR1 is transferred through nuclear pore complex into nucleus where it attaches to promoter region of SA-Responsive genes with the help of interaction with bZIP TFs. NPR1 after interaction gets phosphorylated and with the help of NPR3/4 it gets ubiquitinated and degraded by Proteasome.

Double mutants, npr3npr4, in Arabidopsis mount up great ranks of NPR1 as well as showing insensitiveness towards stimulation of systemic acquired resistance (Fu et al., 2012). Some reports also established the interface among 17 CC-type glutaredoxins and TGA2 bZIP transcription factors (Zander et al., 2012). Among CC-type glutaredoxins and TGA proteins proposed to facilitate defence responses against pathogens and it is involved in different processes of plant development (Zander et al., 2012). Other transcription factors also interact with TGA as interactions of WRKY TF have been shown in tobacco where the NtWRKY12 and TGA protein act together in vitro and in vivo (Van verk et al., 2011). Furthermore, RF2a and RF2b overexpression in rice providing resistance to Rice tungro disease caused by Rice tungro bacillform virus (Daia et al., 2008). HD-Zip family member also negatively regulate the defence responses. In Arabidopsis HD-Zip family member HAT1- HAT1OX and hat1, hat1hat3, hat1hat2hat3 mutants upon challenging with Cucumber mosaic virus showed susceptibility and tolerance against the virus. Anti-CMV defence

response in above mentioned plants is associated with antioxidant and defence related genes that are down-regulated in HAT1OX but upregulated in mutant ones. Further salicylic acid level also follows the same pattern in the overexpressed and mutant plants (Zou et al., 2016). Together, these stories established the connection of bZIP TFs in SA signalling, which could be coupled for evolving improved genotypes capable of virus attack. Myelocytomatosis-related transcription factor family proteins Structural organization of MYC-TF The myelocytomatosis-related family (MYC) TFs subfamily categorized by the occurrence of a basic helix–loop–helix (bHLH) domain and established in eukaryotes. This bHLH domain at their N-terminal containing a basic DNA-binding region consist of nearly 15 amino acids by way of a large number of basic residues, and at C-terminal contains hydrophobic residues which work as dimerization

40  | Pandey et al.

domain, and forms two alpha-helices separated by a loop (Toledo-Oritz et al., 2003). Functions of MYC-TF in plant defence MYC-TFs are key transcriptional regulators and function as a positive controller for JA-responsive genes and wound resistance genes expression. However, it also acts as a negative controller in the course of the pathogen defence genes expression (Van verk, 2009; Lorenzo et al., 2004). During the pathogen attacks, plant produces a bioactive form of JA ( JA conjugated with isoleucine JA-Ileu), that binds to its receptor CORONATINE INSENSITIVE-1 (COI1). This COI1 is an F-box protein which partners by way of interaction with various proteins like cullin, SKP1 and RBX1 proteins, to form the SCFCOI1 complex. After that jasmonatezim-domain ( JAZ) proteins through different interacting partners bring about JAZ unbinding arising out of MYC. Moreover, JAZ interacts, with the SCFCOI1 complex with the help of its

A)

Jas domain and further ubiquitination takes place by the complex followed by its degradation by 26S proteasome machinery. This JA-Ileu with the adjacent sequence permits the binding of protein to COI1, which in turn leads to alteration in the jasmonate-zinc-finger protein expression in inflorescence meristem (Lorenzo et al., 2004; Chini et al., 2007, 2009; Cheng et al., 2011). So, JAZ proteolysis promotes the release and activation of MYC in the presence of JA-Ileu. Moreover, activation of MYC TF triggers the expression of additional transcription factors, like MYBs and WRKYs, which plays crucial role in plant defence (Fig. 2.6) (Yan et al., 2013). Furthermore, MYC stimulates the transcription of the JAZ protein, and restores JA at their basal level (Yan et al., 2013). Structurally JAZ proteins family consisting of 12 different proteins that comprise a centrally located ZIM domain having 28 amino acids of Jas domain at C-terminal as well as carboxy-terminal domain in the N-terminal region (Fig. 2.6). Commonly JAZ proteins for example JAZ3 or JAZ10.1 Tolerance to viruses

Pathogen attack

Jasmonic acid

JA-Ileu

SKP1

MYC2

RBX1

SCFCOI1 complex

B)

WRKY

JAZ

COI 1 Cullin

Increased transcription

MYB

JAZ3

Ub

Nucleus JAZ ZIM

CT

Proteasome

Figure 2.6 Schematic description of MYC TF family in plant defence. (A) During pathogen attacks plant produces jasmonic acid Bioactive form JA-Ileu which binds to its receptor CORONATINE INSENSITIVE-1 (COI1) and forms SCFCOI1 complex by interaction with Cullin, SKP1 and RBX1. This SCFCOI1 complex interact with JAZ (JAZ3) domain by the help of its Jas domain and this complex further ubiquitinated and degraded by 26s Proteasomal machinery. This proteolysis promotes the release and activation of MYC (MYC2) TF, which in turn triggers MYB and WRKY and regulate defence responses. MYC also stimulate the transcription of JAZ protein which restores Jasmonic acid (JA) to its basal level and provide tolerance to viruses. (B) Structure of JAZ proteins containing ZIM and carboxyterminal (CT) domains.

Host Transcription Factors in Modulating Defence Response |  41

act together by means of MYC and have the ability to negatively control its action when JA-Ileu is absent. Also MYC2 attaches to G-box motifs next to genes regulated by JA where JAZ proteins inhibit MYC2 action by a straight interaction among JAZ (C-terminus) and MYC2 (N-terminus) (Chini et al., 2007). Truncation of Jas domain do not affect the binding of JAZ proteins with MYC2, which shows it will act through its N-terminal portion, as a dominantnegative repressor (Memelink, 2009). However, present model for communication and control of MYC is no more factual to all JAZ proteins since other JAZ proteins (like JAZ3) interact with MYC2 by an altered pathways. Also deletion of Jas domain in JAZ3 contribute to make this protein incapable of interacting with MYC2, here the Jas domain is adequate for interaction among JAZ3 with MYC2 (Chini et al., 2009). Thus, it may be suggested that, JAZ3 act together by means of dimer and binds through Jas domain to MYC2, defeating its act. Furthermore, MYC2 is permanently disarmed by the truncation of protein resultant from a deletion at C-terminal region of JAZ3. Further as shown in studies this interface arises via heterodimerization through additional JAZ protein by its N-terminal domain, consecutively, attaches irreversibly to MYC2, therefore performing like dominant-negative repressor (Memelink, 2009). As shown in Arabidopsis, MYC proteins differentially interact with JAZ proteins as MYC2 is probably interacting with all 12 of the JAZ proteins; however, MYC3 establishes a robust interaction with eight of these proteins ( JAZ1, JAZ2, JAZ5, JAZ6, JAZ8, JAZ9, JAZ10 and JAZ11) (Cheng et al., 2011) and MYC4 work together with JAZ1, JAZ3 and JAZ9 (Van verk, 2009). All of these interactions follow the similar mechanism as for MYC2 (Van verk 2009; Cheng et al., 2011). Jasmonic acid ( JA)-signalling operates as a key component in facilitating defence responses against Fusarium oxysporum. JAZ7 relates with transcription factors working as activating (MYC3, MYC4) or repressing ( JAM1) agent of JA-signalling and comprises an efficient EAR repressor motif facilitating transcriptional suppression through corepressor TOPLESS (TPL). It directly recruits TPL, in wild type plants JAZ7 act as a repressor inside JA–response complex. But in jaz7-1D plants, misregulated ectopic JAZ7 showing hyper-activation

of JA-signalling via unsettling COI1–JAZ-TPL-TF complexes (Thatcher et al., 2016) Jasmonic acid signalling in turn induces JAresponsive genes which ultimately modulates resistance genes for providing resistance. It is reported that OsJAZ8C act as negative regulator of JA-induced resistance to Xanthomonas oryzae pv. oryzae (Xoo) in rice (Yamada et al., 2012). As OsMYC2 by interaction with JAZ interacting domain of OsJAZ proteins act as positive regulator of jasmonic acid signalling provides resistance to Xanthomonas oryzae pv. oryzae (Xoo) (Uji et al., 2016). Based on these data, in conclusion JA playing a vital role in resistance to Xoo and OsJAZ8 turns as a suppressor of JA signalling in rice plant (Yamada et al., 2012). NAC transcription factor family protein Structure and function of NAC-TF NAC is one of the emerging transcription factor family works in modulation of defence signalling pathways in plants. The distinctive feature of this transcription factors family is the presence of NAC domain. The NAC TFs superfamily can be classified into seven subfamilies and each subfamily defines the functions of corresponding NAC genes (Pei et al., 2013). NAM, ATAF and CUC are the important subfamilies of NAC genes involved in plant response to abiotic stresses in addition to biotic stresses. Pathogens produce different proteins to impede with the role of NAC TFs and thus disarming the plant defence and leads to colonization as suggested by recent studies. Phytophthora infestans produces LxLR effector (Pi03192), which in turn combine with NAC TFs designated as NAC targeted by phytophthora 1 and 2 (NTP1 and NTP2). Then this interface follows in the endoplasmic reticulum and ward off NTP1 localization towards the nucleus (Fig. 2.7). Augmented susceptibility to infection by P. infestans has been revealed by virus-induced gene silencing (VIGS) of genes encrypting these two NAC factors, signifying their role in plant defence (McLellan et al., 2013). Viral proteins interaction with NAC transcription factors has also shown by different studies. NAC TF expressed in rice plant after different virus

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Figure 2.7 Schematic description of NAC TF family in plant defence. Pathogens during attack produces effector molecules LxLR by Phytophthora infestans, which in turn combine with NAC targeted by phytophthora 1 and 2 (NTP1 and NTP2) in endoplasmic reticulum. This interaction retard the NTP1 localization inside the nucleus and thus increased the susceptibility to infection. During the viral attack Turnip crinkle virus-interacting protein (TIP) interact with TCV coat protein (CP). Then TIP-CP module activates salicylic acid pathway genes by unknown intermediate and reduce the plant basal defence against TCV. But hypersensitive response protein (HRT) protect the TIP by perceiving alteration in TIP triggered by TIP–CP interaction and provide tolerance to virus. Moreover Tobacco mosaic virus (TMV) induces ATAF2 and diminishes the basal defence pathway by repressing the salicylic acid pathway genes. In the absence of pathogen suppressors of non-expresser of PR genes inducible 1 (SNI1) interacts with calmodulin–regulated NAC transcriptional repressor (CBNAC) in Arabidopsis and repress the PR genes expression. When inducer is present during pathogen attack NPR1 translocate towards nucleus and interact through TGA thus induces PR gene expression and provides tolerance to pathogens.

infection (Nuruzzaman et al., 2015). Upon infection to various viruses like Rice black-streaked dwarf virus (RBSDV), Rice dwarf virus (RDV), Rice grassy stunt virus (RGSV), Rice transitory yellowing virus (RTYV) and Rice ragged stunt virus (RRSV) members of NAC family NAC22, ONAC2, ONAC3, SND and ANAC34 all were down-regulated. Which suggest the role of NAC TF in plant viral infections. One of the Arabidopsis NAC protein, designated as Turnip crinkle virus-interacting protein (TIP), precisely interacts with the TCV coat protein (CP). TIP act as transcriptional activator to stimulate defence in plant, which in turn provide basal level of resistance to the virus. Also coat protein of Turnip crinkle virus interacts with TIP and restricts the basal resistance to the virus. Corresponding resistant plant upon infection with mutated TCV (R6A) shows the breakdown of resistance which is hypothesized

due to TIP–CP interaction. Moreover, these TCV mutants are similar in accumulation rates but defective in salicylic acid signalling suggesting TIP role through these pathways (Donze et al., 2014). After infection cells made viral CP, which bind to TIP to decrease basal resistance and stimulate the prompt infection. The CP works as a virulence factor here to provide resistance. A resilient HR-mediated resistance response in resistant plants can be seen, where these plants express a hypersensitive response protein (termed HRT) that can possibly protect the TIP protein by perceiving an alteration in TIP triggered by TIP–CP interface (Ren et al., 2000). Further, yeast two hybrid and in planta immunoprecipitation assays revealed an interface among the TMV 126-/183-kDa replicase protein(s) helicase domain and ATAF2 which is an Arabidopsis NAC domain containing TFs (Wang et al., 2009a).

Host Transcription Factors in Modulating Defence Response |  43

Here, ATAF2 transcriptionally induced by the TMV infection. Thus, by targeting ATAF2, TMV subdues the basal defence pathways in the course of the compatible virus–host interaction (Ren et al., 2000). This assumption is confirmed by the diminished capacity of SA to transcriptionally stimulate genes related with plant defence in TMV-infested tissues (Wang et al., 2009b). NAC TFs moreover interact with different protein suppressors involved in plant defence. When pathogen is absent, protein SNI1-Suppressor of Non-expresser of PR Genes Inducible 1, interacts with a calmodulin-regulated NAC transcriptional repressor CBNAC in Arabidopsis (Kim et al., 2012). CBNAC then attached to the PR1 promoter at E0-1-1 element, meanwhile SNI1 augments the DNA-binding activity of CBNAC, leads to improved repression of the PR1 gene by SNI1 (Kim et al., 2012). Inducer existence in the course of pathogen attack, leads to translocation of more quantity of active NPR1 towards nucleus where it interacts through TGA and inducing the PR1 gene expression (Kim et al., 2012). This SNI1 and CBNAC protein complex perhaps disassembled by NPR1, calmodulin or additional unidentified pathways (Fig. 2.7). Thus, particular influence of NACs with suitable promoter suggests a promising way to considerably alter disease stress response in plants. Other transcription factors Viral component interaction with transcription factors can cause the transcriptional conditioning of a number of pathways. Alfalfa mosaic virus interacts with bHLH family member TF ILR3 through its coat protein and brings the translocation of fraction of ILR3 from nucleus to nucleolus. Loss of function mutants of ILR3 shows higher level of ROS, PR1 mRNAs, salicylic acid, and jasmonic acid contents (Paricio and Pallas, 2017). Our current knowledge about these target genes stems mainly from the studies performed using large-scale transcriptome analyses in plants that overexpress TFs. Deeper analyses can better elucidate their roles under biotic stress response. Future perspectives Advent of genomics and sequencing technology has provided immense opportunity in the field of

TF–virus interaction module. Combining bioinformatics and experimental approaches we can explore this field. In this chapter, we have illustrated the eminent role of TFs as modern tools for genomic studies to enhance plant tolerance against several biotic stresses. Substantial advancements have been actualized in elucidating various defensive roles of TFs to biotic stress, for instance virus and bacteria, while a considerable number of candidate TF genes were previously validated. Tracking down the fundamental TFs from their huge family pool and sequentially comprehending their extensive role still needs to be decoded. The research facilities in providing ambient conditions to establish transgenic lines with an elevated levels of TFs, face a huge set back. Similarly, the existent redundant data need to be sorted out in order to characterize and categorize the TFs. Genome wide identification of different transcription factors upon virus infections as mentioned in the text are the examples of such combinations. We can identify different transcription factors upon viral infections through bioinformatics and validate them through experiments in wet lab and further explore their role in defence pathways. Moreover validated transcription factor-virus defence module can be utilized in transgenic generation (Fig. 2.8) or can be incorporated in breeding programmes. Understanding the role of TF in the regulatory gene network responsive for biotic stress tolerance can help researchers and breeders to improve the crop for these stresses and thus improve the crop productivity. Plant TF play major role for providing biotic stresses. The major TF families for biotic stress tolerance are WRKY, MYB, AP2/ERF, bZIP, MYC and NAC as identified in Arabidopsis and many other plants. These TFs are the useful targets of biotechnological manipulation through their overexpression and silencing, as shown by the examples in quoted in this chapter. TF may regulate a number of genes related to defence pathways in plants so by modulating these transcription factors we can increase the biotic stress resistance as we have seen from the examples of different TF families. Comprehensive crosstalk amongst the mentioned diverse signal transduction pathways by means of TF permits the plant to adjust its defences facing diverse sorts of insect and pathogen attackers. Transcription factors are the proteins and the genes of transcription factors can be modified

Identification of Plant transcription factors Bioinformatics approach -Genome level

Validation by experiments -Gain of function

Validation of TF in defense mechanism

1. Transient overexpression

-Target gene identification

1. TF identification

2. Transgenic plants

-Transcriptional network analysis

2. Phylogenetic relationship

-Loss of function

-Hormonal signaling pathways and crosstalk

3. Evolutionary relationship 4. Cis-element scanning

1. RNAi 2. VIGS 3. T-DNA mutagenesis

5. Comparative genomics

-TF –DNA interaction

-Transcriptome level

1. In-vitro methods

1.Differential expression 2.Co-expression

*DIP-ChIP * EMSA 2. In-vivo methods

-Proteome level

* ChIP-chip

1. TFs interaction

* ChIP-seq

2. Post-translational modification

-Post-translational modification -Metabolite synthesis for defense

Purpose of Plant TF study in Plant defense responses -Elucidation of plant TF in plant defense responses -Improved resistant/tolerant crop cultivars through genetic engineering

* DamID * DNaseI seq

Figure 2.8  Schematic representation of utilization of TF-virus module for crop improvement.

Host Transcription Factors in Modulating Defence Response |  45

in the way we generate the genetically modified crops. Moreover, these TF can carry nonspecific constitutive promoter or specific promoter such as tissue, time or condition specific. Transcriptional reprogramming and crosstalk among signalling pathways between different TFs under bacteria, fungus, insects and viruses infection pave the way for understanding intricate mechanism of plant defence response. Multiple stress-responsive TF genes are the good candidate for developing comprehensive disease resistant crops, therefore, needs significant attention of modern scientific studies (Prasch and Sonnewald, 2015). Modifying these candidate genes through genetic manipulation in crop plants might be a powerful resource contributing to combating biotic stress. Thus, identification of transcription factor which are associated with the biotic stress tolerance could create ample opportunities for crop improvement by generating these modified GM crops. References

Agarwal, P., Agarwal, P.K., Nair, S., Sopory, S.K., and Reddy, M.K. (2007). Stress-inducible DREB2A transcription factor from Pennisetum glaucum is a phosphoprotein and its phosphorylation negatively regulates its DNAbinding activity. Mol. Genet. Genomics 277, 189–198. https://doi.org/10.1007/s00438-006-0183-z Allen, M.D., Yamasaki, K., Ohme-Takagi, M., Tateno, M., and Suzuki, M. (1998). A novel mode of DNA recognition by a beta-sheet revealed by the solution structure of the GCC-box binding domain in complex with DNA. EMBO J. 17, 5484–5496. https://doi.org/10.1093/ emboj/17.18.5484 Alvarez-Venegas, R., Abdallat, A.A., Guo, M., Alfano, J.R., and Avramova, Z. (2007). Epigenetic control of a transcription factor at the cross section of two antagonistic pathways. Epigenetics 2, 106–113. Alves, M.S., Dadalto, S.P., Gonçalves, A.B., De Souza, G.B., Barros, V.A., and Fietto, L.G. (2013). Plant bZIP transcription factors responsive to pathogens: a review. Int. J. Mol. Sci. 14, 7815–7828. https://doi. org/10.3390/ijms14047815 Asai, T., Tena, G., Plotnikova, J., Willmann, M.R., Chiu, W.L., Gomez-Gomez, L., Boller, T., Ausubel, F.M., and Sheen, J. (2002). MAP kinase signalling cascade in Arabidopsis innate immunity. Nature 415, 977–983. https://doi.org/10.1038/415977a Asai, T., Tena, G., Plotnikova, J., Willmann, M.R., Chiu, W.L., Gomez-Gomez, L., Boller, T., Ausubel, F.M., and Sheen, J. (2002). MAP kinase signalling cascade in Arabidopsis innate immunity. Nature 415, 977–983. https://doi.org/10.1038/415977a Ay, N., Irmler, K., Fischer, A., Uhlemann, R., Reuter, G., and Humbeck, K. (2009). Epigenetic programming via histone methylation at WRKY53 controls leaf senescence in Arabidopsis thaliana. Plant J. 58, 333–346. https://doi.org/10.1111/j.1365-313X.2008.03782.x

Bethke, G., Unthan, T., Uhrig, J.F., Pöschl, Y., Gust, A.A., Scheel, D., and Lee, J. (2009). Flg22 regulates the release of an ethylene response factor substrate from MAP kinase 6 in Arabidopsis thaliana via ethylene signaling. Proc. Natl. Acad. Sci. U.S.A. 106, 8067–8072. https:// doi.org/10.1073/pnas.0810206106 Block, A., Toruño, T.Y., Elowsky, C.G., Zhang, C., Steinbrenner, J., Beynon, J., and Alfano, J.R. (2014). The Pseudomonas syringae type III effector HopD1 suppresses effector-triggered immunity, localizes to the endoplasmic reticulum, and targets the Arabidopsis transcription factor NTL9. New Phytol. 201, 1358– 1370. https://doi.org/10.1111/nph.12626 Boter, M., Ruíz-Rivero, O., Abdeen, A., and Prat, S. (2004). Conserved MYC transcription factors play a key role in jasmonate signaling both in tomato and Arabidopsis. Genes Dev. 18, 1577–1591. https://doi.org/10.1101/ gad.297704 Bu, Q., Jiang, H., Li, C.B., Zhai, Q., Zhang, J., Wu, X., Sun, J., Xie, Q., and Li, C. (2008). Role of the Arabidopsis thaliana NAC transcription factors ANAC019 and ANAC055 in regulating jasmonic acid-signaled defense responses. Cell Res. 18, 756–767. https://doi. org/10.1038/cr.2008.53 Büttner, M., and Singh, K.B. (1997). Arabidopsis thaliana ethylene-responsive element binding protein (AtEBP), an ethylene-inducible, GCC box DNA-binding protein interacts with an ocs element binding protein. Proc. Natl. Acad. Sci. U.S.A. 94, 5961–5966. Canonne, J., Marino, D., Jauneau, A., Pouzet, C., Brière, C., Roby, D., and Rivas, S. (2011). The Xanthomonas type III effector XopD targets the Arabidopsis transcription factor MYB30 to suppress plant defense. Plant Cell 23, 3498–3511. https://doi.org/10.1105/tpc.111.088815 Causier, B., Ashworth, M., Guo, W., and Davies, B. (2012). The TOPLESS interactome: a framework for gene repression in Arabidopsis. Plant Physiol. 158, 423–438. https://doi.org/10.1104/pp.111.186999 Chen, L., Zhang, L., and Yu, D. (2010). Wounding-induced WRKY8 is involved in basal defense in Arabidopsis. Mol. Plant Microbe Interact. 23, 558–565. https://doi. org/10.1094/MPMI-23-5-0558 Chen, X., and Guo, Z. (2008). Tobacco OPBP1 enhances salt tolerance and disease resistance of transgenic rice. Int J Mol Sci 9, 2601–2613. https://doi.org/10.3390/ ijms9122601 Cheng, H., and Wang, S. (2014). WRKY-Type transcription factors: A significant factor in rice–pathogen interactions. Scientia Sinica Vitae 44, 784. Cheng, M.C., Hsieh, E.J., Chen, J.H., Chen, H.Y., and Lin, T.P. (2012). Arabidopsis RGLG2, functioning as a RING E3 ligase, interacts with AtERF53 and negatively regulates the plant drought stress response. Plant Physiol. 158, 363–375. https://doi.org/10.1104/pp.111.189738 Cheng, Z., Sun, L., Qi, T., Zhang, B., Peng, W., Liu, Y., and Xie, D. (2011). The bHLH transcription factor MYC3 interacts with the Jasmonate ZIM-domain proteins to mediate jasmonate response in Arabidopsis. Mol. Plant 4, 279–288. https://doi.org/10.1093/mp/ssq073 Cheong, Y.H., Chang, H.S., Gupta, R., Wang, X., Zhu, T., and Luan, S. (2002). Transcriptional profiling reveals novel interactions between wounding, pathogen, abiotic stress, and hormonal responses in Arabidopsis.

46  | Pandey et al.

Plant Physiol. 129, 661–677. https://doi.org/10.1104/ pp.002857 Chern, M., Bai, W., Ruan, D., Oh, T., Chen, X., and Ronald, P.C. (2014). Interaction specificity and coexpression of rice NPR1 homologs 1 and 3 (NH1 and NH3), TGA transcription factors and Negative Regulator of Resistance (NRR) proteins. BMC Genomics 15, 461. https://doi.org/10.1186/1471-2164-15-461 Chini, A., Fonseca, S., Chico, J.M., Fernández-Calvo, P., and Solano, R. (2009). The ZIM domain mediates homoand heteromeric interactions between Arabidopsis JAZ proteins. Plant J. 59, 77–87. https://doi.org/10.1111/ j.1365-313X.2009.03852.x Chini, A., Fonseca, S., Fernández, G., Adie, B., Chico, J.M., Lorenzo, O., García-Casado, G., López-Vidriero, I., Lozano, F.M., Ponce, M.R., et al. (2007). The JAZ family of repressors is the missing link in jasmonate signalling. Nature 448, 666–671. Collinge, M., and Boller, T. (2001). Differential induction of two potato genes, Stprx2 and StNAC, in response to infection by Phytophthora infestans and to wounding. Plant Mol. Biol. 46, 521–529. Cominelli, E., Galbiati, M., Vavasseur, A., Conti, L., Sala, T., Vuylsteke, M., Leonhardt, N., Dellaporta, S.L., and Tonelli, C. (2005). A guard-cell-specific MYB transcription factor regulates stomatal movements and plant drought tolerance. Curr. Biol. 15, 1196–1200. Dai, S., Wei, X., Alfonso, A.A., Pei, L., Duque, U.G., Zhang, Z., Babb, G.M., and Beachy, R.N. (2008). Transgenic rice plants that overexpress transcription factors RF2a and RF2b are tolerant to rice tungro virus replication and disease. Proc. Natl. Acad. Sci. U.S.A. 105, 21012–21016. https://doi.org/10.1073/pnas.0810303105 Dang, F., Wang, Y., She, J., Lei, Y., Liu, Z., Eulgem, T., Lai, Y., Lin, J., Yu, L., Lei, D., et al. (2014). Overexpression of CaWRKY27, a subgroup IIe WRKY transcription factor of Capsicum annuum, positively regulates tobacco resistance to Ralstonia solanacearum infection. Physiol. Plant 150, 397–411. https://doi.org/10.1111/ ppl.12093 De Vos, M., Denekamp, M., Dicke, M., Vuylsteke, M., Van Loon, L., Smeekens, S.C., and Pieterse, C.M. (2006). The Arabidopsis thaliana transcription factor AtMYB102 functions in defense against the insect herbivore Pieris rapae. Plant Signal Behav 1, 305–311. De Vos, M., Van Oosten, V.R., Van Poecke, R.M., Van Pelt, J.A., Pozo, M.J., Mueller, M.J., Buchala, A.J., Métraux, J.P., Van Loon, L.C., Dicke, M., et al. (2005). Signal signature and transcriptome changes of Arabidopsis during pathogen and insect attack. Mol. Plant Microbe Interact. 18, 923–937. https://doi.org/10.1094/MPMI-18-0923 Delessert, C., Kazan, K., Wilson, I.W., Van Der Straeten, D., Manners, J., Dennis, E.S., and Dolferus, R. (2005). The transcription factor ATAF2 represses the expression of pathogenesis-related genes in Arabidopsis. Plant J. 43, 745–757. Deng, H., Liu, H., Li, X., Xiao, J., and Wang, S. (2012). A CCCH-type zinc finger nucleic acid-binding protein quantitatively confers resistance to rice bacterial blight disease. Plant Physiol. 158, 876–889. https://doi. org/10.1104/pp.111.191379 Deslandes, L., Olivier, J., Theulieres, F., Hirsch, J., Feng, D.X., Bittner-Eddy, P., Beynon, J., and Marco, Y. (2002).

Resistance to Ralstonia solanacearum in Arabidopsis thaliana is conferred by the recessive RRS1-R gene, a member of a novel family of resistance genes. Proc. Natl. Acad. Sci. U.S.A. 99, 2404–2409. https://doi. org/10.1073/pnas.032485099 Després, C., DeLong, C., Glaze, S., Liu, E., and Fobert, P.R. (2000). The Arabidopsis NPR1/NIM1 protein enhances the DNA binding activity of a subgroup of the TGA family of bZIP transcription factors. Plant Cell 12, 279–290. Devaiah, B.N., Madhuvanthi, R., Karthikeyan, A.S., and Raghothama, K.G. (2009). Phosphate starvation responses and gibberellic acid biosynthesis are regulated by the MYB62 transcription factor in Arabidopsis. Mol Plant 2, 43–58. https://doi.org/10.1093/mp/ssn081 Dong, X. (1998). SA, JA, ethylene, and disease resistance in plants. Curr. Opin. Plant Biol. 1, 316–323. Donze, T., Qu, F., Twigg, P., and Morris, T.J. (2014). Turnip crinkle virus coat protein inhibits the basal immune response to virus invasion in Arabidopsis by binding to the NAC transcription factor TIP. Virology 449, 207–214. https://doi.org/10.1016/j.virol.2013.11.018 Drews, G.N., Bowman, J.L., and Meyerowitz, E.M. (1991). Negative regulation of the Arabidopsis homeotic gene AGAMOUS by the APETALA2 product. Cell 65, 991–1002. Dröge-Laser, W., Kaiser, A., Lindsay, W.P., Halkier, B.A., Loake, G.J., Doerner, P., Dixon, R.A., and Lamb, C. (1997). Rapid stimulation of a soybean protein-serine kinase that phosphorylates a novel bZIP DNA-binding protein, G/HBF-1, during the induction of early transcription-dependent defenses. EMBO J. 16, 726– 738. https://doi.org/10.1093/emboj/16.4.726 Du, H., Feng, B.R., Yang, S.S., Huang, Y.B., and Tang, Y.X. (2012a). The R2R3-MYB transcription factor gene family in maize. PLOS ONE 7, e37463. https://doi. org/10.1371/journal.pone.0037463 Du, H., Yang, S.S., Liang, Z., Feng, B.R., Liu, L., Huang, Y.B., and Tang, Y.X. (2012b). Genome-wide analysis of the MYB transcription factor superfamily in soybean. BMC Plant Biol. 12, 106. https://doi.org/10.1186/14712229-12-106 Du, M., Zhai, Q., Deng, L., Li, S., Li, H., Yan, L., Huang, Z., Wang, B., Jiang, H., Huang, T., et al. (2014). Closely related NAC transcription factors of tomato differentially regulate stomatal closure and reopening during pathogen attack. Plant Cell 26, 3167–3184. https://doi.org/10.1105/tpc.114.128272 Dubos, C., Stracke, R., Grotewold, E., Weisshaar, B., Martin, C., and Lepiniec, L. (2010). MYB transcription factors in Arabidopsis. Trends Plant Sci. 15, 573–581. https:// doi.org/10.1016/j.tplants.2010.06.005 Egawa, C., Kobayashi, F., Ishibashi, M., Nakamura, T., Nakamura, C., and Takumi, S. (2006). Differential regulation of transcript accumulation and alternative splicing of a DREB2 homolog under abiotic stress conditions in common wheat. Genes Genet. Syst. 81, 77–91. Encinas-Villarejo, S., Maldonado, A.M., Amil-Ruiz, F., de los Santos, B., Romero, F., Pliego-Alfaro, F., Muñoz-Blanco, J., and Caballero, J.L. (2009). Evidence for a positive regulatory role of strawberry (Fragariaananassa) Fa

Host Transcription Factors in Modulating Defence Response |  47

WRKY1 and Arabidopsis AtWRKY75 proteins in resistance. J. Exp. Bot. 60, 3043–3065. Eulgem, T., and Somssich, I.E. (2007). Networks of WRKY transcription factors in defense signaling. Curr. Opin. Plant Biol. 10, 366–371. Eulgem, T., Rushton, P.J., Robatzek, S., and Somssich, I.E. (2000). The WRKY superfamily of plant transcription factors. Trends Plant Sci. 5, 199–206. Faria, J.A., Reis, P.A., Reis, M.T., Rosado, G.L., Pinheiro, G.L., Mendes, G.C., and Fontes, E.P. (2011). The NAC domain-containing protein, GmNAC6, is a downstream component of the ER stress- and osmotic stress-induced NRP-mediated cell-death signaling pathway. BMC Plant Biol. 11, 129. https://doi.org/10.1186/1471-2229-11129 Froidure, S., Canonne, J., Daniel, X., Jauneau, A., Brière, C., Roby, D., and Rivas, S. (2010). AtsPLA2-alpha nuclear relocalization by the Arabidopsis transcription factor AtMYB30 leads to repression of the plant defense response. Proc. Natl. Acad. Sci. U.S.A. 107, 15281– 15286. https://doi.org/10.1073/pnas.1009056107 Fu, Z.Q., Yan, S., Saleh, A., Wang, W., Ruble, J., Oka, N., Mohan, R., Spoel, S.H., Tada, Y., Zheng, N., et al. (2012). NPR3 and NPR4 are receptors for the immune signal salicylic acid in plants. Nature 486, 228–232. https:// doi.org/10.1038/nature11162 Grunewald, W., Karimi, M., Wieczorek, K., Van de Cappelle, E., Wischnitzki, E., Grundler, F., Inzé, D., Beeckman, T., and Gheysen, G. (2008). A role for AtWRKY23 in feeding site establishment of plant-parasitic nematodes. Plant Physiol. 148, 358–368. https://doi.org/10.1104/ pp.108.119131 Guo, C., Guo, R., Xu, X., Gao, M., Li, X., Song, J., Zheng, Y., and Wang, X. (2014). Evolution and expression analysis of the grape (Vitisvinifera L.) WRKY gene family. J. Exp. Bot. 65, 1513–1528. Guo, Z.J., Chen, X.J., Wu, X.L., Ling, J.Q., and Xu, P. (2004). Overexpression of the AP2/EREBP transcription factor OPBP1 enhances disease resistance and salt tolerance in tobacco. Plant Mol. Biol. 55, 607–618. Gutterson, N., and Reuber, T.L. (2004). Regulation of disease resistance pathways by AP2/ERF transcription factors. Curr. Opin. Plant Biol. 7, 465–471. https://doi. org/10.1016/j.pbi.2004.04.007 Hiratsu, K., Matsui, K., Koyama, T., and Ohme-Takagi, M. (2003). Dominant repression of target genes by chimeric repressors that include the EAR motif, a repression domain, in Arabidopsis. Plant J. 34, 733–739. Howe, G. (2004).Jasmonates as signals in the wound response. J. Plant Growth Regul. 23, 223–227. Huang, H., Wang, Y., Wang, S., Wu, X., Yang, K., Niu, Y., and Dai, S. (2012). Transcriptome-wide survey and expression analysis of stress-responsive NAC genes in Chrysanthemum lavandulifolium. Plant Sci. 193-194, 18–27. https://doi.org/10.1016/j.plantsci.2012.05.004 Huh, S.U., Lee, S.B., Kim, H.H., and Paek, K.H. (2012). ATAF2, a NAC transcription factor, binds to the promoter and regulates NIT2 gene expression involved in auxin biosynthesis. Mol. Cells 34, 305–313. https:// doi.org/10.1007/s10059-012-0122-2 Ikeda, M., and Ohme-Takagi, M. (2009). A novel group of transcriptional repressors in Arabidopsis. Plant Cell

Physiol. 50, 970–975. https://doi.org/10.1093/pcp/ pcp048 Inoue, H., Hayashi, N., Matsushita, A., Xinqiong, L., Nakayama, A., Sugano, S., Jiang, C.J., and Takatsuji, H. (2013). Blast resistance of CC-NB-LRR protein Pb1 is mediated by WRKY45 through protein-protein interaction. Proc. Natl. Acad. Sci. U.S.A. 110, 9577– 9582. https://doi.org/10.1073/pnas.1222155110 Ishihama, N., Yamada, R., Yoshioka, M., Katou, S., and Yoshioka, H. (2011). Phosphorylation of the Nicotiana benthamiana WRKY8 transcription factor by MAPK functions in the defense response. Plant Cell 23, 1153– 1170. https://doi.org/10.1105/tpc.110.081794 Jakoby, M., Weisshaar, B., Dröge-Laser, W., VicenteCarbajosa, J., Tiedemann, J., Kroj, T., and Parcy, F. (2002). bZIP transcription factors in Arabidopsis. Trends Plant Sci. 7, 106–111. Jensen, M., Hagedorn, P., de Torres-Zabala, M., Grant, M., Rung, J., and Collinge, D. (2008). Transcriptional regulation by an NAC (NAMATAF1,2-CUC2) transcription factor attenuates ABA signaling for efficient basal defence towards Blumeriagraminis f sphordei in Arabidopsis. Plant J. 56, 867–880. Jensen, M.K., Rung, J.H., Gregersen, P.L., Gjetting, T., Fuglsang, A.T., Hansen, M., Joehnk, N., Lyngkjaer, M.F., and Collinge, D.B. (2007). The HvNAC6 transcription factor: a positive regulator of penetration resistance in barley and Arabidopsis. Plant Mol. Biol. 65, 137–150. https://doi.org/10.1007/s11103-007-9204-5 Jiang, C.H., Huang, Z.Y., Xie, P., Gu, C., Li, K., Wang, D.C., Yu, Y.Y., Fan, Z.H., Wang, C.J., Wang, Y.P., et al. (2016). Transcription factors WRKY70 and WRKY11 served as regulators in rhizobacterium Bacillus cereus AR156induced systemic resistance to Pseudomonas syringae pv. tomato DC3000 in Arabidopsis. J. Exp. Bot. 67, 157–174. https://doi.org/10.1093/jxb/erv445 Jiang, C., Gu, J., Chopra, S., Gu, X., and Peterson, T. (2004). Ordered origin of the typical two- and three-repeat Myb genes. Gene 326, 13–22. Johnson, E.T., and Dowd, P.F. (2004). Differentially enhanced insect resistance, at a cost, in Arabidopsis thaliana constitutively expressing a transcription factor of defensive metabolites. J. Agric. Food Chem. 52, 5135–5138. https://doi.org/10.1021/jf0308049 Journot-Catalino, N., Somssich, I.E., Roby, D., and Kroj, T. (2006). The transcription factors WRKY11 and WRKY17 act as negative regulators of basal resistance in Arabidopsis thaliana. Plant Cell 18, 3289–3302. Kaltdorf, M., and Naseem, M. (2013). How many salicylic acid receptors does a plant cell need? Sci Signal 6, jc3. https://doi.org/10.1126/scisignal.2003944 Kaminaka, H., Näke, C., Epple, P., Dittgen, J., Schütze, K., Chaban, C., Holt, B.F., Merkle, T., Schäfer, E., Harter, K., et al. (2006). bZIP10-LSD1 antagonism modulates basal defense and cell death in Arabidopsis following infection. EMBO J. 25, 4400–4411. Kaneda, T., Taga, Y., Takai, R., Iwano, M., Matsui, H., Takayama, S., Isogai, A., and Che, F.S. (2009). The transcription factor OsNAC4 is a key positive regulator of plant hypersensitive cell death. EMBO J. 28, 926–936. https://doi.org/10.1038/emboj.2009.39 Kim, H.S., Park, H.C., Kim, K.E., Jung, M.S., Han, H.J., Kim, S.H., Kwon, Y.S., Bahk, S., An, J., Bae, D.W., et al. (2012).

48  | Pandey et al.

A NAC transcription factor and SNI1 cooperatively suppress basal pathogen resistance in Arabidopsis thaliana. Nucleic Acids Res. 40, 9182–9192. https:// doi.org/10.1093/nar/gks683 Kim, J.G., Stork, W., and Mudgett, M.B. (2013). Xanthomonas type III effector XopD desumoylates tomato transcription factor SlERF4 to suppress ethylene responses and promote pathogen growth. Cell Host Microbe 13, 143–154. https://doi.org/10.1016/j. chom.2013.01.006 Kim, K.C., Lai, Z., Fan, B., and Chen, Z. (2008). Arabidopsis WRKY38 and WRKY62 transcription factors interact with histone deacetylase 19 in basal defense. Plant Cell 20, 2357–2371. https://doi.org/10.1105/ tpc.107.055566 Kim, K.C., Lai, Z., Fan, B., and Chen, Z. (2008). Arabidopsis WRKY38 and WRKY62 transcription factors interact with histone deacetylase 19 in basal defense. Plant Cell 20, 2357–2371. https://doi.org/10.1105/ tpc.107.055566 Kloth, K.J., Wiegers, G.L., Busscher-Lange, J., van Haarst, J.C., Kruijer, W., Bouwmeester, H.J., Dicke, M., and Jongsma, M.A. (2016). AtWRKY22 promotes susceptibility to aphids and modulates salicylic acid and jasmonic acid signalling. J. Exp. Bot. 67, 3383–3396. https://doi.org/10.1093/jxb/erw159 Kuhlmann, M., Horvay, K., Stathmann, A., Heinekamp, T., Fischer, U., Böttner, S., and Dröge-Laser, W. (2003). The alpha-helical D1 domain of the bZIP transcription factor BZI-1 interacts with the ankyrin-repeat protein ANK1, and is essential for BZI-1 function, both in auxin signaling and pathogen response. J. Biol. Chem. 278, 8786–8794. Lee, S.J., Lee, M.Y., Yi, S.Y., Oh, S.K., Choi, S.H., Her, N.H., Choi, D., Min, B.W., Yang, S.G., and Harn, C.H. (2002). PPI1: a novel pathogen-induced basic region-leucine zipper (bZIP) transcription factor from pepper. Mol. Plant Microbe Interact. 15, 540–548. https://doi. org/10.1094/MPMI.2002.15.6.540 Li, C.W., Su, R.C., Cheng, C.P., Sanjaya, You, S.J., Hsieh, T.H., Chao, T.C., and Chan, M.T. (2011). Tomato RAV transcription factor is a pivotal modulator involved in the AP2/EREBPmediated defense pathway. Plant Physiol. 156, 213–227. Li, G., Meng, X., Wang, R., Mao, G., Han, L., Liu, Y., and Zhang, S. (2012). Dual-level regulation of ACC synthase activity by MPK3/MPK6 cascade and its downstream WRKY transcription factor during ethylene induction in Arabidopsis. PLOS Genet. 8, e1002767. https://doi. org/10.1371/journal.pgen.1002767 Li, J.B., Luan, Y.S., and Yin, Y.L. (2014). SpMYB overexpression in tobacco plants leads to altered abiotic and biotic stress responses. Gene 547, 145–151. https:// doi.org/10.1016/j.gene.2014.06.049 Li, L., Yu, X., Thompson, A., Guo, M., Yoshida, S., Asami, T., Chory, J., and Yin, Y. (2009). Arabidopsis MYB30 is a direct target of BES1 and cooperates with BES1 to regulate brassinosteroid-induced gene expression. Plant J. 58, 275–286. https://doi.org/10.1111/j.1365313X.2008.03778.x Li, Z., Zhang, L., Yu, Y., Quan, R., Zhang, Z., Zhang, H., and Huang, R. (2011). The ethylene response factor AtERF11 that is transcriptionally modulated by the

bZIP transcription factor HY5 is a crucial repressor for ethylene biosynthesis in Arabidopsis. Plant J. 68, 88–99. https://doi.org/10.1111/j.1365-313X.2011.04670.x Licausi, F., Ohme-Takagi, M., and Perata, P. (2013). APETALA2/ethylene responsive factor (AP2/ERF) transcription factors: mediators of stress responses and developmental programs. New Phytol. 199, 639–649. Licausi, F., van Dongen, J.T., Giuntoli, B., Novi, G., Santaniello, A., Geigenberger, P., and Perata, P. (2010). HRE1 and HRE2, two hypoxia-inducible ethylene response factors, affect anaerobic responses in Arabidopsis thaliana. Plant J. 62, 302–315. https://doi. org/10.1111/j.1365-313X.2010.04149.x Lin, R., Zhaom, W., Mengm, X., Wang, M., and Peng, Y. (2007). Rice gene OsNAC19 encodes a novel NACdomain transcription factor and responds to infection by Magnaporthegrisea. Plant Sci. 172, 120–130. Liu, D., Chen, X., Liu, J., Ye, J., and Guo, Z. (2012). The rice ERF transcription factor OsERF922 negatively regulates resistance to Magnaporthe oryzae and salt tolerance. J. Exp. Bot. 63, 3899–3911. https://doi.org/10.1093/ jxb/ers079 Liu, D., Chen, X., Liu, J., Ye, J., and Guo, Z. (2012). The rice ERF transcription factor OsERF922 negatively regulates resistance to Magnaporthe oryzae and salt tolerance. J. Exp. Bot. 63, 3899–3911. https://doi.org/10.1093/ jxb/ers079 Liu, H., Zhou, X., Dong, N., Liu, X., Zhang, H., and Zhang, Z. (2011). Expression of a wheat MYB gene in transgenic tobacco enhances resistance to Ralstonia solanacearum, and to drought and salt stresses. Funct. Integr. Genomics 11, 431–443. https://doi.org/10.1007/s10142-0110228-1 Liu, R., Lü, B., Wang, X., Zhang, C., Zhang, S., Qian, J., Chen, L., Shi, H., and Dong, H. (2010). Thirty-seven transcription factor genes differentially respond to a hairpin protein and affect resistance to the green peach aphid in Arabidopsis. J. Biosci. 35, 435-450. Liu, X., Yang, L., Zhou, X., Zhou, M., Lu, Y., Ma, L., Ma, H., and Zhang, Z. (2013). Transgenic wheat expressing Thinopyrum intermedium MYB transcription factor TiMYB2R-1 shows enhanced resistance to the takeall disease. J. Exp. Bot. 64, 2243–2253. https://doi. org/10.1093/jxb/ert084 Liu, Y., Schiff, M., and Dinesh-Kumar, S.P. (2004). Involvement of MEK1 MAPKK, NTF6 MAPK, WRKY/MYB transcription factors, COI1 and CTR1 in N-mediated resistance to Tobacco mosaic virus. Plant J. 38, 800–809. https://doi.org/10.1111/j.1365313X.2004.02085.x Lorenzo, O., Chico, J.M., Sánchez-Serrano, J.J., and Solano, R. (2004). Jasmonate-insensitive 1 encodes a MYC transcription factor essential to discriminate between different jasmonate-regulated defense responses in Arabidopsis. Plant Cell 16, 1938–1950. https://doi. org/10.1105/tpc.022319 Lorenzo, O., Chico, J.M., Sánchez-Serrano, J.J., and Solano, R. (2004). Jasmonate-insensitive 1 encodes a MYC transcription factor essential to discriminate between different jasmonate-regulated defense responses in Arabidopsis. Plant Cell 16, 1938–1950. https://doi. org/10.1105/tpc.022319

Host Transcription Factors in Modulating Defence Response |  49

Lorenzo, O., Piqueras, R., Sánchez-Serrano, J.J., and Solano, R. (2003). Ethylene response factor 1 integrates signals from ethylene and jasmonate pathways in plant defense. Plant Cell 15, 165–178. Lu, J., Ju, H., Zhou, G., Zhu, C., Erb, M., Wang, X., Wang, P., and Lou, Y. (2011). An EAR-motif-containing ERF transcription factor affects herbivore-induced signaling, defense and resistance in rice. Plant J. 68, 583–596. https://doi.org/10.1111/j.1365-313X.2011.04709.x Lu, X., Zhang, L., Zhang, F., Jiang, W., Shen, Q., Zhang, L., Lv, Z., Wang, G., and Tang, K. (2013). AaORA, a trichome-specific AP2/ERF transcription factor of Artemisia annua, is a positive regulator in the artemisinin biosynthetic pathway and in disease resistance to Botrytis cinerea. New Phytol. 198, 1191–1202. https:// doi.org/10.1111/nph.12207 Marino, D., Froidure, S., Canonne, J., Ben Khaled, S., Khafif, M., Pouzet, C., Jauneau, A., Roby, D., and Rivas, S. (2013). Arabidopsis ubiquitin ligase MIEL1 mediates degradation of the transcription factor MYB30 weakening plant defence. Nat. Commun. 4, 1476. https://doi.org/10.1038/ncomms2479 Mateo, A., Mühlenbock, P., Rustérucci, C., Chang, C.C., Miszalski, Z., Karpinska, B., Parker, J.E., Mullineaux, P.M., and Karpinski, S. (2004). LESION SIMULATING DISEASE 1 is required for acclimation to conditions that promote excess excitation energy. Plant Physiol. 136, 2818–2830. https://doi.org/10.1104/pp.104.043646 Matsui, K., Umemura, Y., and Ohme-Takagi, M. (2008). AtMYBL2, a protein with a single MYB domain, acts as a negative regulator of anthocyanin biosynthesis in Arabidopsis. Plant J. 55, 954–967. https://doi. org/10.1111/j.1365-313X.2008.03565.x McLellan, H., Boevink, P.C., Armstrong, M.R., Pritchard, L., Gomez, S., Morales, J., Whisson, S.C., Beynon, J.L., and Birch, P.R. (2013). An RxLR effector from Phytophthora infestans prevents re-localisation of two plant NAC transcription factors from the endoplasmic reticulum to the nucleus. PLOS Pathog. 9, e1003670. https://doi. org/10.1371/journal.ppat.1003670 Memelink, J. (2009). Regulation of gene expression by jasmonatehomones. Phytochemistry 70, 1560-1570. Meng, X., Xu, J., He, Y., Yang, K.Y., Mordorski, B., Liu, Y., and Zhang, S. (2013). Phosphorylation of an ERF transcription factor by Arabidopsis MPK3/MPK6 regulates plant defense gene induction and fungal resistance. Plant Cell 25, 1126–1142. https://doi. org/10.1105/tpc.112.109074 Mengiste, T., Chen, X., Salmeron, J., and Dietrich, R. (2003). The BOTRYTIS SUSCEPTIBLE1 gene encodes an R2R3MYB transcription factor protein that is required for biotic and abiotic stress responses in Arabidopsis. Plant Cell 15, 2551–2565. https://doi.org/10.1105/ tpc.014167 Miao, Y., Laun, T.M., Smykowski, A., and Zentgraf, U. (2007). Arabidopsis MEKK1 can take a short cut: it can directly interact with senescence-related WRKY53 transcription factor on the protein level and can bind to its promoter. Plant Mol. Biol. 65, 63–76. https://doi. org/10.1007/s11103-007-9198-z Miao, Y., Laun, T., Zimmermann, P., and Zentgraf, U. (2004). Targets of the WRKY53 transcription factor and

its role during leaf senescence in Arabidopsis. Plant Mol. Biol. 55, 853–867. Misra, P., Pandey, A., Tiwari, M., Chandrashekar, K., Sidhu, O.P., Asif, M.H., Chakrabarty, D., Singh, P.K., Trivedi, P.K., Nath, P., et al. (2010). Modulation of transcriptome and metabolome of tobacco by Arabidopsis transcription factor, AtMYB12, leads to insect resistance. Plant Physiol. 152, 2258–2268. https://doi.org/10.1104/ pp.109.150979 Mitsunami, T., Nishihara, M., Galis, I., Alamgir, K.M., Hojo, Y., Fujita, K., Sasaki, N., Nemoto, K., Sawasaki, T., and Arimura, G. (2014). Overexpression of the PAP1 transcription factor reveals a complex regulation of flavonoid and phenylpropanoid metabolism in Nicotiana tabacum plants attacked by Spodoptera litura. PLOS ONE 9, e108849. https://doi.org/10.1371/ journal.pone.0108849 Moffat, C.S., Ingle, R.A., Wathugala, D.L., Saunders, N.J., Knight, H., and Knight, M.R. (2012). ERF5 and ERF6 play redundant roles as positive regulators of JA/Et-mediated defense against Botrytis cinerea in Arabidopsis. PLOS ONE 7, e35995. https://doi. org/10.1371/journal.pone.0035995 Nakano, T., Suzuki, K., Fujimura, T., and Shinshi, H. (2006). Genome-wide analysis of the ERF gene family in Arabidopsis and rice. Plant Physiol. 140, 411–432. Nakashima, K., Tran, L.S., Van Nguyen, D., Fujita, M., Maruyama, K., Todaka, D., Ito, Y., Hayashi, N., Shinozaki, K., and Yamaguchi-Shinozaki, K. (2007). Functional analysis of a NAC-type transcription factor OsNAC6 involved in abiotic and biotic stress-responsive gene expression in rice. Plant J. 51, 617–630. Nuruzzaman, M., Sharoni, A.M., Satoh, K., Karim, M.R., Harikrishna, J.A., Shimizu, T., Sasaya, T., Omura, T., Haque, M.A., Hasan, S.M., et al. (2015). NAC transcription factor family genes are differentially expressed in rice during infections with Rice dwarf virus, Rice black-streaked dwarf virus, Rice grassy stunt virus, Rice ragged stunt virus, and Rice transitory yellowing virus. Front. Plant. Sci. 6, 676. https://doi.org/10.3389/ fpls.2015.00676 Ogata, K., Kanei-Ishii, C., Sasaki, M., Hatanaka, H., Nagadoi, A., Enari, M., Nakamura, H., Nishimura, Y., Ishii, S., and Sarai, A. (1996). The cavity in the hydrophobic core of Myb DNA-binding domain is reserved for DNA recognition and trans-activation. Nat. Struct. Biol. 3, 178–187. Oh, S.K., Lee, S., Yu, S.H., and Choi, D. (2005). Expression of a novel NAC domain-containing transcription factor (CaNAC1) is preferentially associated with incompatible interactions between chili pepper and pathogens. Planta 222, 876–887. https://doi.org/10.1007/s00425-0050030-1 Oh, S.K., Baek, K.H., Park, J.M., Yi, S.Y., Yu, S.H., Kamoun, S., and Choi, D. (2008). Capsicum annuum WRKY protein CaWRKY1 is a negative regulator of pathogen defense. New Phytol. 177, 977–989. https://doi. org/10.1111/j.1469-8137.2007.02310.x Ohme-Takagi, M., and Shinshi, H. (1995). Ethyleneinducible DNA binding proteins that interact with an ethylene-responsive element. Plant Cell 7, 173–182. https://doi.org/10.1105/tpc.7.2.173

50  | Pandey et al.

Ohta, M., Matsui, K., Hiratsu, K., Shinshi, H., and OhmeTakagi, M. (2001). Repression domains of class II ERF transcriptional repressors share an essential motif for active repression. Plant Cell 13, 1959–1968. Ohta, M., Ohme-Takagi, M., and Shinshi, H. (2000). Three ethylene-responsive transcription factors in tobacco with distinct transactivation functions. Plant J. 22, 29–38. Okada, A., Okada, K., Miyamoto, K., Koga, J., Shibuya, N., Nojiri, H., and Yamane, H. (2009). OsTGAP1, a bZIP transcription factor, coordinately regulates the inductive production of diterpenoid phytoalexins in rice. J. Biol. Chem. 284, 26510–26518. https://doi.org/10.1074/ jbc.M109.036871 Orellana, S., Yañez, M., Espinoza, A., Verdugo, I., González, E., Ruiz-Lara, S., and Casaretto, J.A. (2010). The transcription factor SlAREB1 confers drought, salt stress tolerance and regulates biotic and abiotic stress-related genes in tomato. Plant Cell Environ. 33, 2191–2208. https://doi.org/10.1111/j.1365-3040.2010.02220.x Padmanabhan, M.S., Ma, S., Burch-Smith, T.M., Czymmek, K., Huijser, P., and Dinesh-Kumar, S.P. (2013). Novel positive regulatory role for the SPL6 transcription factor in the N TIR-NB-LRR receptor-mediated plant innate immunity. PLOS Pathog. 9, e1003235. https://doi. org/10.1371/journal.ppat.1003235 Pajerowska-Mukhtar, K.M., Emerine, D.K., and Mukhtar, M.S. (2013). Tell me more: roles of NPRs in plant immunity. Trends Plant Sci. 18, 402–411. https://doi. org/10.1016/j.tplants.2013.04.004 Pandey, S.P., and Somssich, I.E. (2009). The role of WRKY transcription factors in plant immunity. Plant Physiol. 150, 1648–1655. https://doi.org/10.1104/ pp.109.138990 Paricio, F., and Pallas, V. (2017). The coat protein of Alfalfa mosaic virus interacts and interferes with the transcriptional activity of the bHLH transcription factorILR3 promoting salicylic acid-dependent defence signaling response. Mol. Plant Pathol. 18, 173–186. Park, J.M., Park, C.J., Lee, S.B., Ham, B.K., Shin, R., and Paek, K.H. (2001). Overexpression of the tobacco Tsi1 gene encoding an EREBP/AP2-type transcription factor enhances resistance to pathogen attack and osmotic stress in tobacco. Plant Cell 13, 1035–1046. Paz-Ares, J., Ghosal, D., Wienand, U., Peterson, P.A., and Saedler, H. (1987). The regulatory c1 locus of Zea mays encodes a protein with homology to myb protooncogene products and with structural similarities to transcriptional activators. EMBO J. 6, 3553–3558. Pei, H., Ma, N., Tian, J., Luo, J., Chen, J., Li, J., Zheng, Y., Chen, X., Fei, Z., and Gao, J. (2013). An NAC transcription factor controls ethylene-regulated cell expansion in flower petals. Plant Physiol. 163, 775–791. https://doi.org/10.1104/pp.113.223388 Peng, Y., Bartley, L.E., Chen, X., Dardick, C., Chern, M., Ruan, R., Canlas, P.E., and Ronald, P.C. (2008). OsWRKY62 is a negative regulator of basal and Xa21mediated defense against Xanthomonas oryzae pv. oryzae in rice. Mol. Plant 1, 446–458. https://doi.org/10.1093/ mp/ssn024 Pieterse, C.M., Van der Does, D., Zamioudis, C., LeonReyes, A., and Van Wees, S.C. (2012). Hormonal modulation of plant immunity. Annu. Rev. Cell Dev.

Biol. 28, 489–521. https://doi.org/10.1146/annurevcellbio-092910-154055 Pirrello, J., Jaimes-Miranda, F., Sanchez-Ballesta, M.T., Tournier, B., Khalil-Ahmad, Q., Regad, F., Latché, A., Pech, J.C., and Bouzayen, M. (2006). Sl-ERF2, a tomato ethylene response factor involved in ethylene response and seed germination. Plant Cell Physiol. 47, 1195–1205. Popescu, S.C., Popescu, G.V., Bachan, S., Zhang, Z., Gerstein, M., Snyder, M., and Dinesh-Kumar, S.P. (2009). MAPK target networks in Arabidopsis thaliana revealed using functional protein microarrays. Genes Dev. 23, 80–92. https://doi.org/10.1101/gad.1740009 Prasch, C.M., and Sonnewald, U. (2015). Signaling events in plants: stress factors in combination change the picture. Environ. Expt. Bot. 114, 4–14. Qin, F., Kakimoto, M., Sakuma, Y., Maruyama, K., Osakabe, Y., Tran, L.S., Shinozaki, K., and Yamaguchi-Shinozaki, K. (2007). Regulation and functional analysis of ZmDREB2A in response to drought and heat stresses in Zea mays L. Plant J. 50, 54–69. Raffaele, S., Vailleau, F., Léger, A., Joubès, J., Miersch, O., Huard, C., Blée, E., Mongrand, S., Domergue, F., and Roby, D. (2008). A MYB transcription factor regulates very-long-chain fatty acid biosynthesis for activation of the hypersensitive cell death response in Arabidopsis. Plant Cell 20, 752–767. https://doi.org/10.1105/ tpc.107.054858 Rashotte, A.M., Mason, M.G., Hutchison, C.E., Ferreira, F.J., Schaller, G.E., and Kieber, J.J. (2006). A subset of Arabidopsis AP2 transcription factors mediates cytokinin responses in concert with a two-component pathway. Proc. Natl. Acad. Sci. U.S.A. 103, 11081–11085. Rasmussen, M.W., Roux, M., Petersen, M., and Mundy, J. (2012). MAP Kinase Cascades in Arabidopsis Innate Immunity. Front. Plant Sci. 3, 169. https://doi. org/10.3389/fpls.2012.00169 Ren, T., Qu, F., and Morris, T.J. (2000). HRT gene function requires interaction between a NAC protein and viral capsid protein to confer resistance to Turnip crinkle virus. Plant Cell 12, 1917–1926. Rushton, P., Somssich, I., Ringler, P., and Shen, Q. (2010). WRKY transcription factors. Trends Plant Sci. 15, 247–258. Rushton, P.J., Torres, J.T., Parniske, M., Wernert, P., Hahlbrock, K., and Somssich, I.E. (1996). Interaction of elicitor-induced DNA-binding proteins with elicitor response elements in the promoters of parsley PR1 genes. EMBO J. 15, 5690–5700. Saga, H., Ogawa, T., Kai, K., Suzuki, H., Ogata, Y., Sakurai, N., Shibata, D., and Ohta, D. (2012). Identification and characterization of ANAC042, a transcription factor family gene involved in the regulation of camalexin biosynthesis in Arabidopsis. Mol. Plant Microbe Interact. 25, 684–696. https://doi.org/10.1094/MPMI-09-110244 Sakuma, Y., Liu, Q., Dubouzet, J.G., Abe, H., Shinozaki, K., and Yamaguchi-Shinozaki, K. (2002). DNA-binding specificity of the ERF/AP2 domain of Arabidopsis DREBs, transcription factors involved in dehydrationand cold-inducible gene expression. Biochem. Biophys. Res. Commun. 290, 998–1009. https://doi. org/10.1006/bbrc.2001.6299

Host Transcription Factors in Modulating Defence Response |  51

Sakuma, Y., Maruyama, K., Osakabe, Y., Qin, F., Seki, M., Shinozaki, K., and Yamaguchi-Shinozaki, K. (2006). Functional analysis of an Arabidopsis transcription factor, DREB2A, involved in drought-responsive gene expression. Plant Cell 18, 1292–1309. Sasaki-Sekimoto, Y., Jikumaru, Y., Obayashi, T., Saito, H., Masuda, S., Kamiya, Y., Ohta, H., and Shirasu, K. (2013). Basic helix-loop-helix transcription factors JASMONATE-ASSOCIATED MYC2-LIKE1 ( JAM1), JAM2, and JAM3 are negative regulators of jasmonate responses in Arabidopsis. Plant Physiol. 163, 291–304. https://doi.org/10.1104/pp.113.220129 Sato, F., Kitajima, S., Koyama, T., and Yamada, Y. (1996). Ethylene-induced gene expression of osmotin-like protein, a neutral isoform of tobacco PR-5, is mediated by the AGCCGCC cis-sequence. Plant Cell Physiol. 37, 249–255. Schenke, D., Böttcher, C., and Scheel, D. (2011). Crosstalk between abiotic ultraviolet-B stress and biotic (flg22) stress signalling in Arabidopsis prevents flavonol accumulation in favor of pathogen defence compound production. Plant Cell Environ. 34, 1849–1864. https:// doi.org/10.1111/j.1365-3040.2011.02381.x Segarra, G., Van der Ent, S., Trillas, I., and Pieterse, C.M. (2009). MYB72, a node of convergence in induced systemic resistance triggered by a fungal and a bacterial beneficial microbe. Plant Biol 11, 90–96. https://doi. org/10.1111/j.1438-8677.2008.00162.x Selth, L.A., Dogra, S.C., Rasheed, M.S., Healy, H., Randles, J.W., and Rezaian, M.A. (2005). A NAC domain protein interacts with Tomato leaf curl virus replication accessory protein and enhances viral replication. Plant Cell 17, 311–325. Seo, J.S., Sohn, H.B., Noh, K., Jung, C., An, J.H., Donovan, C.M., Somers, D.A., Kim, D.I., Jeong, S., Kim, C., Kim, H.M., Lee, S., Choi, Y.D., Moon, T.W., Kim, C.H., and Cheon, J. (2012). Expression of the Arabidopsis AtMYB44 gene confers drought/salt-stress tolerance in transgenic soybean. Mol. Breed. 29, 601–608. Seo, P.J., and Park, C.M. (2010). MYB96-mediated abscisic acid signals induce pathogen resistance response by promoting salicylic acid biosynthesis in Arabidopsis. New Phytol. 186, 471–483. https://doi.org/10.1111/ j.1469-8137.2010.03183.x Seo, P.J., Kim, M.J., Park, J.Y., Kim, S.Y., Jeon, J., Lee, Y.H., Kim, J., and Park, C.M. (2010). Cold activation of a plasma membrane-tethered NAC transcription factor induces a pathogen resistance response in Arabidopsis. Plant J. 61, 661–671. https://doi.org/10.1111/j.1365313X.2009.04091.x Shen, Q.H., Saijo, Y., Mauch, S., Biskup, C., Bieri, S., Keller, B., Seki, H., Ulker, B., Somssich, I.E., and Schulze-Lefert, P. (2007). Nuclear activity of MLA immune receptors links isolate-specific and basal disease-resistance responses. Science 315, 1098–1103. Shi, W., Hao, L., Li, J., Liu, D., Guo, X., and Li, H. (2014). The Gossypium hirsutum WRKY gene GhWRKY39-1 promotes pathogen infection defense responses and mediates salt stress tolerance in transgenic Nicotiana benthamiana. Plant Cell Rep. 33, 483-498. Shimono, M., Sugano, S., Nakayama, A., Jiang, C.J., Ono, K., Toki, S., and Takatsuji, H. (2007). Rice WRKY45

plays a crucial role in benzothiadiazole-inducible blast resistance. Plant Cell 19, 2064–2076. Singh, K., Foley, R.C., and Oñate-Sánchez, L. (2002). Transcription factors in plant defense and stress responses. Curr. Opin. Plant Biol. 5, 430–436. Skibbe, M., Qu, N., Galis, I., and Baldwin, I.T. (2008). Induced plant defenses in the natural environment: Nicotiana attenuata WRKY3 and WRKY6 coordinate responses to herbivory. Plant Cell 20, 1984–2000. https://doi.org/10.1105/tpc.108.058594 Stockinger, E.J., Gilmour, S.J., and Thomashow, M.F. (1997). Arabidopsis thaliana CBF1 encodes an AP2 domain-containing transcriptional activator that binds to the C-repeat/DRE, a cis-acting DNA regulatory element that stimulates transcription in response to low temperature and water deficit. Proc. Natl. Acad. Sci. U.S.A. 94, 1035–1040. Stracke, R., Werber, M., and Weisshaar, B. (2001). The R2R3-MYB gene family in Arabidopsis thaliana. Curr. Opin. Plant Biol. 4, 447–456. Sun, C., Palmqvist, S., Olsson, H., Borén, M., Ahlandsberg, S., and Jansson, C. (2003). A novel WRKY transcription factor, SUSIBA2, participates in sugar signaling in barley by binding to the sugar-responsive elements of the iso1 promoter. Plant Cell 15, 2076–2092. Sun, L., Zhang, H., Li, D., Huang, L., Hong, Y., Ding, X.S., Nelson, R.S., Zhou, X., and Song, F. (2013). Functions of rice NAC transcriptional factors, ONAC122 and ONAC131, in defense responses against Magnaporthe grisea. Plant Mol. Biol. 81, 41–56. https://doi. org/10.1007/s11103-012-9981-3 Tak, H., and Mhatre, M. (2013). Cloning and molecular characterization of a putative bZIP transcription factor VvbZIP23 from Vitis vinifera. Protoplasma 250, 333–345. https://doi.org/10.1007/s00709-012-0417-3 Tao, Z., Liu, H., Qiu, D., Zhou, Y., Li, X., Xu, C., and Wang, S. (2009). A pair of allelic WRKY genes play opposite roles in rice-bacteria interactions. Plant Physiol. 151, 936–948. https://doi.org/10.1104/pp.109.145623 Thatcher, L.F., Cevik, V., Grant, M., Zhai, B., Jones, J.D., Manners, J.M., and Kazan, K. (2016). Characterization of a JAZ7 activation-tagged Arabidopsis mutant with increased susceptibility to the fungal pathogen Fusarium oxysporum. J. Exp. Bot. 67, 2367–2386. https://doi. org/10.1093/jxb/erw040 Toledo-Ortiz, G., Huq, E., and Quail, P.H. (2003). The Arabidopsis basic/helix-loop-helix transcription factor family. Plant Cell 15, 1749–1770. Uji, Y., Taniguchi, S., Tamaoki, D., Shishido, H., Akimitsu, K., and Gomi, K. (2016). Overexpression of OsMYC2 results in the up-regulation of early JA-rresponsive genes and bacterial blight resistance in Rice. Plant Cell Physiol. 57, 1814–1827. https://doi.org/10.1093/pcp/pcw101 Ulker, B., and Somssich, I.E. (2004). WRKY transcription factors: from DNA binding towards biological function. Curr. Opin. Plant Biol. 7, 491–498. https://doi. org/10.1016/j.pbi.2004.07.012 Van der Ent, S., Verhagen, B.W., Van Doorn, R., Bakker, D., Verlaan, M.G., Pel, M.J., Joosten, R.G., Proveniers, M.C., Van Loon, L.C., Ton, J., et al. (2008). MYB72 is required in early signaling steps of rhizobacteria-induced systemic resistance in Arabidopsis. Plant Physiol. 146, 1293–1304. https://doi.org/10.1104/pp.107.113829

52  | Pandey et al.

Van Verk, M., Gatz, C., and Linthorst, H. (2009). Transcriptional regulation of plant defense responses. Adv. Bot. Res. 51, 397-438. van Verk, M.C., Neeleman, L., Bol, J.F., and Linthorst, H.J. (2011). Tobacco transcription factor NtWRKY12 interacts with TGA2.2 in vitro and in vivo. Front. Plant Sci. 2, 32. https://doi.org/10.3389/fpls.2011.00032 van Verk, M.C., Pappaioannou, D., Neeleman, L., Bol, J.F., and Linthorst, H.J. (2008). A Novel WRKY transcription factor is required for induction of PR-1a gene expression by salicylic acid and bacterial elicitors. Plant Physiol. 146, 1983–1995. https://doi.org/10.1104/pp.107.112789 Vannini, C., Iriti, M., Bracale, M., Locatelli, F., Faoro, F., Croce, P., Pirona, R., di Maro, A., Coraggio, I., and Genga, A. (2006). The ectopic expression of the rice OsMYB4 gene in Arabidopsis increases tolerance to abiotic, environmental and biotic stresses. Physiol. Mol. Plant Pathol. 69, 26-42. Wang, D., Amornsiripanitch, N., and Dong, X. (2006). A genomic approach to identify regulatory nodes in the transcriptional network of systemic acquired resistance in plants. PLOS Pathog. 2, e123. Wang, X., Basnayake, B.M., Zhang, H., Li, G., Li, W., Virk, N., Mengiste, T., and Song, F. (2009a). The Arabidopsis ATAF1, a NAC transcription factor, is a negative regulator of defense responses against necrotrophic fungal and bacterial pathogens. Mol. Plant Microbe Interact. 22, 1227–1238. https://doi.org/10.1094/ MPMI-22-10-1227 Wang, X., Goregaoker, S.P., and Culver, J.N. (2009b). Interaction of the Tobacco mosaic virus replicase protein with a NAC domain transcription factor is associated with the suppression of systemic host defenses. J. Virol. 83, 9720–9730. https://doi.org/10.1128/JVI.00941-09 Wang, X., Goregaoker, S.P., and Culver, J.N. (2009). Interaction of the Tobacco mosaic virus replicase protein with a NAC domain transcription factor is associated with the suppression of systemic host defenses. J. Virol. 83, 9720–9730. https://doi.org/10.1128/JVI.00941-09 Wang, Y., Dang, F., Liu, Z., Wang, X., Eulgem, T., Lai, Y., Yu, L., She, J., Shi, Y., Lin, J., et al. (2013). CaWRKY58, encoding a group I WRKY transcription factor of Capsicum annuum, negatively regulates resistance to Ralstonia solanacearum infection. Mol. Plant Pathol. 14, 131–144. https://doi.org/10.1111/j.13643703.2012.00836.x Wen, F., Zhu, H., Li, P., Jiang, M., Mao, W., Ong, C., and Chu, Z. (2014). Genome-wide evolutionary characterization and expression analyses of WRKY family genes in Brachypodium distachyon. DNA Res. 21, 327–339. https://doi.org/10.1093/dnares/dst060 Wilkins, O., Nahal, H., Foong, J., Provart, N.J., and Campbell, M.M. (2009). Expansion and diversification of the Populus R2R3-MYB family of transcription factors. Plant Physiol. 149, 981–993. https://doi.org/10.1104/ pp.108.132795 Windram, O., Madhou, P., McHattie, S., Hill, C., Hickman, R., Cooke, E., Jenkins, D.J., Penfold, C.A., Baxter, L., Breeze, E., et al. (2012). Arabidopsis defense against Botrytis cinerea: chronology and regulation deciphered by high-resolution temporal transcriptomic analysis. Plant Cell 24, 3530–3557. https://doi.org/10.1105/ tpc.112.102046

Woldemariam, M.G., Dinh, S.T., Oh, Y., Gaquerel, E., Baldwin, I.T., and Galis, I. (2013). NaMYC2 transcription factor regulates a subset of plant defense responses in Nicotiana attenuata. BMC Plant Biol. 13, 73. https://doi.org/10.1186/1471-2229-13-73 Wu, C., Avila, C.A., and Goggin, F.L. (2015). The ethylene response factor Pti5 contributes to potato aphid resistance in tomato independent of ethylene signalling. J. Exp. Bot. 66, 559–570. https://doi.org/10.1093/jxb/ eru472 Wu, Y., Deng, Z., Lai, J., Zhang, Y., Yang, C., Yin, B., Zhao, Q., Zhang, L., Li, Y., Yang, C., et al. (2009). Dual function of Arabidopsis ATAF1 in abiotic and biotic stress responses. Cell Res. 19, 1279–1290. https://doi.org/10.1038/ cr.2009.108 Xie, Q., Sanz-Burgos, A.P., Guo, H., García, J.A., and Gutiérrez, C. (1999). GRAB proteins, novel members of the NAC domain family, isolated by their interaction with a geminivirus protein. Plant Mol. Biol. 39, 647–656. Xie, Z., Zhang, Z.L., Zou, X., Yang, G., Komatsu, S., and Shen, Q.J. (2006). Interactions of two abscisic-acid induced WRKY genes in repressing gibberellin signaling in aleurone cells. Plant J. 46, 231–242. Xing, D.H., Lai, Z.B., Zheng, Z.Y., Vinod, K.M., Fan, B.F., and Chen, Z.X. (2008). Stress- and pathogen-induced Arabidopsis WRKY48 is a transcriptional activator that represses plant basal defense. Mol. Plant 1, 459–470. https://doi.org/10.1093/mp/ssn020 Xue, G.P., and Loveridge, C.W. (2004). HvDRF1 is involved in abscisic acid-mediated gene regulation in barley and produces two forms of AP2 transcriptional activators, interacting preferably with a CT-rich element. Plant J. 37, 326–339. Yamada, S., Kano, A., Tamaoki, D., Miyamoto, A., Shishido, H., Miyoshi, S., Taniguchi, S., Akimitsu, K., and Gomi, K. (2012). Involvement of OsJAZ8 in jasmonate-induced resistance to bacterial blight in rice. Plant Cell Physiol. 53, 2060–2072. https://doi.org/10.1093/pcp/pcs145 Yan, Y., Borrego, E., and Kolomiets, M. (2013). Lipid Metabolism; Baez, R., Ed. InTech: Rijeka, Croatia: Chapter 16, pp. 393-442. Yanhui, C., Xiaoyuan, Y., Kun, H., Meihua, L., Jigang, L., Zhaofeng, G., Zhiqiang, L., Yunfei, Z., Xiaoxiao, W., Xiaoming, Q., et al. (2006). The MYB transcription factor superfamily of Arabidopsis: expression analysis and phylogenetic comparison with the rice MYB family. Plant Mol. Biol. 60, 107–124. https://doi.org/10.1007/ s11103-005-2910-y Yogendra, K.N., Kumar, A., Sarkar, K., Li, Y., Pushpa, D., Mosa, K.A., Duggavathi, R., and Kushalappa, A.C. (2015). Transcription factor StWRKY1 regulates phenylpropanoid metabolites conferring late blight resistance in potato. J. Exp. Bot. 66, 7377–7389. https:// doi.org/10.1093/jxb/erv434 Yoshii, M., Shimizu, T., Yamazaki, M., Higashi, T., Miyao, A., Hirochika, H., and Omura, T. (2009). Disruption of a novel gene for a NAC-domain protein in rice confers resistance to Rice dwarf virus. Plant J. 57, 615–625. https://doi.org/10.1111/j.1365-313X.2008.03712.x Zander, M., Chen, S., Imkampe, J., Thurow, C., and Gatz, C. (2012). Repression of the Arabidopsis thaliana jasmonic acid/ethylene-induced defense pathway by TGAinteracting glutaredoxins depends on their C-terminal

Host Transcription Factors in Modulating Defence Response |  53

ALWL motif. Mol Plant 5, 831–840. https://doi. org/10.1093/mp/ssr113 Zarei, A., Körbes, A.P., Younessi, P., Montiel, G., Champion, A., and Memelink, J. (2011). Two GCC boxes and AP2/ ERF-domain transcription factor ORA59 in jasmonate/ ethylene-mediated activation of the PDF1.2 promoter in Arabidopsis. Plant Mol. Biol. 75, 321–331. https://doi. org/10.1007/s11103-010-9728-y Zhang, Z., Liu, X., Wang, X., Zhou, M., Zhou, X., Ye, X., and Wei, X. (2012). An R2R3 MYB transcription factor in wheat, TaPIMP1, mediates host resistance to Bipolaris sorokiniana and drought stresses through regulation of defense- and stress-related genes. New Phytol. 196, 1155–1170. https://doi.org/10.1111/j.14698137.2012.04353.x Zhang, Z., Zhang, H., Quan, R., Wang, X.C., and Huang, R. (2009). Transcriptional regulation of the ethylene response factor LeERF2 in the expression of ethylene biosynthesis genes controls ethylene production in tomato and tobacco. Plant Physiol. 150, 365–377. https://doi.org/10.1104/pp.109.135830

Zhao, Y., Wei, T., Yin, K.Q., Chen, Z., Gu, H., Qu, L.J., and Qin, G. (2012). Arabidopsis RAP2.2 plays an important role in plant resistance to Botrytis cinerea and ethylene responses. New Phytol. 195, 450–460. https://doi. org/10.1111/j.1469-8137.2012.04160.x Zorzatto, C., Machado, J.P., Lopes, K.V., Nascimento, K.J., Pereira, W.A., Brustolini, O.J., Reis, P.A., Calil, I.P., Deguchi, M., Sachetto-Martins, G., et al. (2015). NIK1mediated translation suppression functions as a plant antiviral immunity mechanism. Nature 520, 679–682. https://doi.org/10.1038/nature14171 Zou, L.J., Deng, X.G., Han, X.Y., Tan, W.R., Zhu, L.J., Xi, D.H., Zhang, D.W., and Lin, H.H. (2016). Role of Transcription Factor HAT1 in Modulating Arabidopsis thaliana Response to Cucumber mosaic virus. Plant Cell Physiol. 57, 1879–1889. https://doi.org/10.1093/pcp/ pcw109

Surfacing the Role of Epigenetics in Host–Virus Interaction Namisha Sharma1, Pranav P. Sahu2, Ritika Kulshreshtha1 and Manoj Prasad1*

3

1National Institute of Plant Genome Research, New Delhi, India.

2Max Planck Institute for Plant Breeding Research, Cologne, Germany.

*Correspondence: [email protected] https://doi.org/10.21775/9781910190814.03

Abstract Epigenetics is a mechanism which determines the phenotype of an organism by causing heritable (during cell division) but simultaneously reversible alterations/variations in gene expression. It is not related to alterations in the DNA sequence of the genotype. Geminiviruses are the most devastating plant viruses since they cause significant yield losses in world agriculture. The plant defence initiated against these DNA viruses is of special interest, specifically in regard to the role of epigenetic mechanism played in control of virus spread. These heritable and covalent modifications of DNA and histone in virus genome are mainly related to suppression of gene transcription, despite the differences between viruses, the role of epigenetics seems to be reasonably comparable. However, several key questions remain unanswered concerning the basic mechanism behind the epigenetic regulation of viruses via plant defence system. This book chapter specifically summarizes the recent advances on role of epigenetics in virus genome modification leading to silencing of viral genes and plant tolerance/resistance. Introduction Plants are constantly exposed to several stresses, which involve different types of abiotic and biotic stress factors. Biotic stress in plants is caused by various living organisms such as plant pathogens, for

example bacteria, viruses, fungi and parasites. Ever since the early expansion in agriculture, diseases due to these pathogens, have persisted on plants; thus, adversely affecting the plant growth and development, eventually leading to loss in agricultural yield. The initial approach to understand the nature of these diseases in plants has been reported in writings of Theophrastus (372–287 BC). Latest developments in plant biology unveiled that plants have specific defence mechanism to minimize the damage caused by pathogen infection. Several reports have highlighted the role of RNA silencing pathways in plant defence against viruses (Chellappan et al., 2004; Molnár et al., 2005; Akbergenov et al., 2006; Ruiz-Ferrer and Voinnet, 2009; PérezQuintero et al., 2010; Dalakouras et al., 2013; Raja et al., 2014; Andika et al., 2015). A significant association between the gene expression and chromatin modifications has been illustrated in plants during stress imposition (Pooggin et al., 2013; Sharma et al., 2013; Carbonell and Carrington, 2015; Zhang et al., 2015; Fortes and Gallusci, 2017). In case of DNA viruses the chromatin is subjected to siRNA mediated DNA methylation leading to epigenetic modifications against geminiviruses (Raja et al., 2010). The term ‘epigenetic’ was coined by Conrad H. Waddington (1956). Epigenetic is a crucial adaptive mechanism which determines the phenotype of a system. It is related to interaction of genes and their products causing heritable but simultaneously

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reversible alterations in the expression of genes, independent of DNA sequence variation (Berger, 2007). In eukaryotes, epigenetic modification persists in the course of mitotic divisions and can be inherited through meiotic divisions. It is associated with the reversible chemical modifications in DNA and histones and thus contributes notably to cell diversity by regulating the gene expression. It is an important mechanism which can influence the gene transcription, and regulates gene expression in response to different stress conditions and during development. In eukaryotic cell, epigenetic mechanisms can reconfigure the chromatin structure by some of the chemical modulations at DNA level through methylation of cytosine residues or by post-translational modifications of histones. Thus, it determines the binding of transcription regulators to DNA and level of mRNA aggregation at the transcriptional level (Sahu et al., 2013; Espinas et al., 2016). DNA methylation in plants is tissue-, species-, age-, organelle-, sequence-specific, as well as a dynamic and heritable phenomenon. DNA methylation plays a vital role in controlling gene expression and DNA replication. The observation from transgenic tobacco plants indicates that overexpression of a transgene induces DNA hypermethylation and transcriptional silencing of those loci containing sequences homologous to transgenes (Matzke et al., 1989). Further, this silencing was the result of small non-coding RNAs directing epigenetic changes termed as RNA-directed DNA methylation (RdDM). These heterochromatic siRNAs are 23–24 nt long. In plants, dsRNAs or siRNAs which contain homologous sequences to promoter regions of a gene can elicit promoter methylation and transcriptional gene silencing (TGS; Mette et al., 2000; Melquist and Bender, 2003; Sharma et al., 2013; Wendte and Pikaard, 2017). TGS could also be the cause of structural changes similar to heterochromatinization in genome. Apart from regulating various biological responses such as developmental processes, time of flowering, parent of origin imprinting, paramutation, suppression of expression of transposable elements (TEs), and repeat sequences, the epigenetic alterations have been reported to play a vital role in the plant immune system. Eradicating the role of epigenetics in providing resistance/tolerance against the viral stimuli in plants is important challenge to

understand the molecular and cellular processes which are controlling virus spread in plants. In this chapter, we examine the strategies employed by plants to initiate transcriptional gene silencing against virus via DNA methylation and the mechanism evolved by viruses as counteract against the plant defence system. Epigenetic modifications in plants Plants are an important system for studying the epigenetic regulations. Epigenetic mechanisms can induce metastable variations in gene expression patterns, facilitating the plants to survive in unfavourable environments. In plants the structure and stable repression or activation of chromatin is regulated by epigenetic modifications, comprising DNA methylation and histone modifications. DNA methylation involves two opposing processes of adding and removing a methyl group in DNA which is directed by small non-coding RNAs. Further, histone proteins are also exposed to several modifications such as acetylation, methylation, phosphorylation, sumoylation and ubiquitination. These histone modifications influence epigenetic states in plants and regulate the defence responsive genes during pathogen attacks. The reciprocation within DNA methylation, non-coding small RNAs and histone modification facilitates plants with diverse and robust epigenetic regulation. In the subsequent section, we have summarized the different epigenetic mechanisms and the key players in the epigenetic regulation in plants. Histone modifications and its role in epigenetic regulation In eukaryotes, the chromatin consists of the genomic DNA packaged as a complex with the histone proteins. These histone proteins form an octamer around which the genomic DNA is wrapped. The core histones H2A, H2B, H3 and H4 are characterized by two regions, a histonefold domain, the conserved region and N-terminal histone tail, variable and unstructured (Andrews and Luger, 2011). The epigenetic modification associated with regulation of gene expression is covalent post-translational modification of N-terminal tail of core histone protein at certain amino acid residues like lysine, arginine, serine and

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threonine. The modifications either lead to activation or repression of transcription depending on the type of modification. Histone acetyltransferases (HATs) catalysed acetylation of H3 and H4 lysine (K) at positions 4, 9, 27, 36 and 79 is important for positive regulation of transcription whereas deacetylation catalysed by histone deacetylase (HDAC) leads to negative regulation. The other form of modification, i.e. methylation, is the most abundant and affects transcription in a way depending on position and degree of methylation (Fig. 3.1). It is associated with the methylation of lysine (K) and arginine (R) residues catalysed by histone lysine methyltransferases (HKMTs) consisting of evolutionarily conserved Su(var)3–9, Enhancer-ofzeste and Trithorax (SET) domain (Thorstensen et al., 2011) and protein arginine methyltransferases (PRMTs) (Di and Bedford, 2011) respectively. Amongst these histone modifications, H3K9me2 is carried out by Arabidopsis Kryptonite (KYP) (Du et al., 2014) whereas H3K27 methylation is performed by the Polycomb repressive complex 2 (PRC2) (Liu et al., 2010; Zheng and Chen, 2011). Further, these modifications can be mono-, di- and trimethylation of lysine and mono- and dimethylation of arginine residue. Unlike histone acetylation, histone methylation is related to both activation and repression of gene expression. Active transcription is associated with H3K36me3, H3K48, H3K79

and dimethylation and trimethylation of histone H3 at lysine 4 (H3K4me2, H3K4me3), in contrast methylation at H3K9me2, H3K27, H4K20me1, H4R3me2 and H3R2me2 are responsible for chromatin condensation and hence transcription repression. Transcriptional activation is also linked to H3 phosphorylation at serine and threonine residues (Li et al., 2001). Conversely, methyl group removal from histone proteins is catalysed by lysine-specific demethylase1 (LSD1) and Jumonji C ( JmjC) demethylases which are evolutionarily conserved histone demethylases. Understanding the role of histone modification in developmental reprogramming and in response to various environmental stresses in plants has been progressed in recent years. Histone modifications are important players in vernalization and photomorphogenesis. H3K4me3 and histone acetylation is responsible for active expression of Flowering Locus C (FLC) resulting in late flowering whereas H3K9me2, H3K27me2 and histone deacetylation reverse this effect by repression of FLC in Arabidopsis thaliana (He, 2012). In addition to flowering time regulation, histone modification particularly H3K9ac contribute to light-induced activation of Long Hypocotyl 5 (HY5) and HY5 Homologue (HYH) and their downstream effectors like photosynthesis-related genes such as Photosystem I subunit F (PsaF) genes (Charron et al., 2009).

K5,K27 M H4 H2B H2A

H2B

H3

H4

H3

M

K12, K20 M

K4, K9, K14, K23, K27, K36, K48, K79 Figure 3.1 Histone modifications: nucleosome consists of four core histone proteins, H2A, H2B, H3 and H4. In response to unfavourable conditions post-translational modifications of histone including methylation, ubiquitinylation and acetylation, causes chromatin modification and alters the gene expression in plants. These modifications can activate/inactivate transcription of a gene. Acetylation at lysine residue of histone proteins is associated with gene activation whereas methylation is associated with both activation and inactivation of gene. K, lysine; red star, inactivation; blue star, activation; M, methylation.

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Promoter and coding region of another photosynthetic gene; Phosphoenolpyruvate carboxylase (Pepc) in maize showed light induced acetylation of H4K5 and H3K9 (Offermann et al., 2008). Moreover, regulation of gibberellin metabolism genes is also associated with light-induced acetylation of H3K27ac and trimethylation of H3K27me (Charron et al., 2009). Histone modifications are also important component of regulatory mechanism involved in biotic stress response. H3K9 is a negative regulator of viral gene expression and it is well illustrated that Arabidopsis harbouring mutation in genes encoding for histone H3K9 methyltransferases (associated with repressed genes) were observed to be hypersensitive to the infection by Beet curly top virus (Raja et al., 2008). In Nicotiana plants transformed with promoter region of African cassava mosaic geminivirus (ACMV), an induction in the level of H3K9 methylation was observed at intergenic region of ACMV promoter, supporting the hypothesis that plants utilize the histone modification mechanism in order to defend themselves from DNA viruses (Dogar, 2006). In a study on endogenous Petunia vein clearing virus (ePVCV), two lines of Petunia hybrid (RdC; rose du ciel and W138) varying in their resistance to the PVCV were used and in the cultivar (W138) with a reduced resistance to PVCV, the equivalent amount of H3mK9me2 and H3mK4 was observed. On the contrary, the level of H3mK9me2 was significantly increased in the stronger silencing strain (RdC) (Noreen et al., 2007), thus elucidating the role of histone methylation in virus resistance. Understanding the mechanism behind tolerance against viruses is an important strategy for developing virus-resistant plants. In this context dynamics of H3K9 histone methyltransferase, Kryptonite (NbKYP) was correlated with transcriptional regulation of viral genes in Nicotiana benthamiana. Further, it was illustrated that transcription activator protein (AC2) encoded by geminiviruses activates the expression of transcription repressor RAV2 (RELATED TO ABI3 and VP1) which regulates the expression of KYP helping in virus spread (Sun et al., 2015). In a recent study, importance of histone methylation in imparting tolerance against viral infection was evaluated in the pepper plants. In pepper plants infected with Pepper golden mosaic virus (PepGMV), the viral chromatin isolated from the symptomatic tissue was found to be associated

with H3K4me3 representing active transcription of viral genes. Contrastingly, repressive chromatin mark H3K9me2 was associated with the viral chromatin isolated from recovered tissue leading to repressed transcription of viral genes (CenicerosOjeda et al., 2016). These studies demonstrate the pivotal role histone modification in gene regulation during virus infection and illustrate that histone methylation might function in the anti-viral system. Plant DNA methylation DNA methylation mechanism is considered as an important factor of immune system during virus infection in plant cells (Blevins et al., 2006; Raja et al., 2008). DNA methylation controls the genome access to transcription factors and function of the genes and genome structure and integrity. TGS leads to suppression of transcription mainly within the transposons, chromosomal repeats and transgenic inserts; simultaneously, it also regulates the expression of coding genes. Within plants this gene body methylation is associated with CG methylation of the transcribed DNA regions. The function of body methylation is still unidentified, but it has been proposed that it might either avert transcription from intragenic promoters (specifically in case of viruses) or increase the splicing accuracy. During DNA replication, DNA methylation on the template strand can be inherited to guide thus maintaining the ‘epigenetic mark’. This DNA methylation can be transmitted through mitosis and meiotic divisions and can be reorganized during cellular reprogramming (Law and Jacobsen, 2010; Birnbaum and Roudier, 2017). DNA methylation involves two opposing processes of adding and removing a methyl group in DNA. In eukaryotes, DNA methylation is initiated by methylation at C5 (fifth carbon) of pyrimidine ring of cytosine [5-methylcytosine (5-meC)] or the sixth nitrogen atom of adenine nucleotides that confers gene silencing and plays crucial role in plant development and defence against viruses, transposons, and transgenes (Lister et al., 2008; Sahu et al., 2013). In plant genomes, methylation occurs in the target regions at both symmetric cytosine sites (CpG, and CpHpG, where H is A, T, or C) and at asymmetric sites (CpHpH, where H is A, T, or C) (Cokus et al., 2008). It has been observed that approximately 50% of plant genome to be methylated consisting of centromeric region and repetitive region (Matzke

Surfacing the Role of Epigenetics in Host–Virus Interaction |  59

and Mosher, 2014). Plants produce dsRNA specifically siRNAs during DNA virus infections ( Jones et al., 1999) or against inverted repeat transgenes (Aufsatz et al., 2002; Melquist and Bender, 2003) triggering DNA methylation of the complementary (identical) sequences. RNA directed DNA methylation (RdDM) is highly sequence specific and does not spread beyond the boundary of the trigger RNA (Pélissier et al., 1999). RdDM causes RNA-guided epigenetic modification of the genome. RdDM was first detected in viroid-infected tobacco plants (Wassenegger et al., 1994). RdDM is triggered by dsRNA that is processed into small RNAs of 21–24 nt, reinforcing a link with post-transcriptional gene silencing (PTGS) (Mette et al., 2000). In plants, dsRNAs trigger promoter methylation and TGS based on the homologous to promoter regions (Mette et al., 2000; Jones et al., 2001; Melquist and Bender, 2003). RNA directed DNA methylation The term ‘RNA directed DNA methylation’ was coined by Sanger and colleagues (1994). It describes the TGS phenomenon that requires small RNAs and it is a conserved event in plants, animals and fungi (Castel and Martienssen, 2013; Sharma et al., 2013). Cytosine methylation is initiated by the enzyme catalysed (adenosine kinase, ADK) translocation of methyl group from S-adenosyl methionine (SAM) to C5 of cytosine. In plant species, the level of 5mC ranges from 6% to 25% of total cytosines (Steward et al., 2002). During replication, methylation imprints at CpG and CpHpG sites can be re-established on newly synthesized sister strand by using hemi-methylated sites as the template. Contrastingly, methylation at CpHpH sequences cannot be conserved as after DNA replication one of the strands of DNA does not have a methylated cytosine at the respective position (Chan et al., 2005; Xie and Yu, 2015). The smallest DNA-target size for RdDM is 30 bp (Pélissier and Wassenegger, 2000). Through next-generation sequencing in different plant varieties and wild-type and mutant plants, the genome-wide patterns of DNA modification (especially methylation) at single-nucleotide resolution has been identified. Table 3.1 summarizes the components involved in RdDM.

Enzymes involved in chromatin modifications Methyltransferase Two types of DNA methyltransferases are recognized in plants, for methylation maintenance and establishment of methylation. The enzymes participating in RdDM are Domains Rearranged Methyltransferase (DRM; de novo methylation), Chromomethylase (CMT; methylation maintenance), Methyltransferase (MET; maintenance of methylation), Kryptonite (KYP; methylation maintenance and H3K9 methylation). These enzymes have diverse role in the de novo and maintenance of DNA methylation. DNA methyltransferase 1 (MET1), the conserved Dnmt1-type enzyme [DNA (cytosine-5-)-methyltransferase 1], sustains methylation at symmetrical dinucleotide (CpG) sites (Wendte and Pikaard, 2017). The met1 mutant plants have reduced CpG methylation (Vongs et al., 1993). Chromomethylase 3 (CMT3) is plant-specific enzyme and it catalysis methylation at CpHpG sequence predominantly at centromeric repeats and transposons (Lindroth et al., 2001; Tompa et al., 2002). Additionally, CMT3 along with Kryptonite9/SUVH4 (KYP9) mediates CpHpG DNA methylation at sites containing methylation at 9th residue; lysine of histone H3 (H3K9), a mark of repressed chromatin (Du et al., 2014). This pathway was discovered as the first direct link between DNA methylation and histone methylation. Recently, another chromomethylase gene, CMT2, was found to recognize H3K9me2 peptides and stimulate the methylation at CpHpH sites in pericentromeric heterochromatin (Stroud et al., 2014). Further, de novo methylation is initiated by domains rearranged methyltransferases, which includes two DNA methyltransferases, DRM1 and DRM2 in form of homodimer. DRM2, an orthologue of conserved Dnmt3 family (Cao et al., 2002; Chan et al., 2005), catalyses methylation at asymmetrical cytosine (CpHpH) sites and is recruited by heterochromatic siRNAs. Concurrently, a third gene family member, DRM3, a homologue of Dnmt3L, was determined to play an important role in establishing RdDM by stimulating the activity of DRM2 (Henderson et al., 2010). It has been proposed that DRM2 interacts with AGO4- siRNA complex and initiates methylation of cytosine on the template DNA strand. This

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Table 3.1 Enzymes involved in chromatin modifications Protein

Protein class

HEN1

Biochemical activity

Pathway

References

sRNA methyltransferase Methylates last nucleotide of sRNA at 2ʹ-OH

All sRNA pathways

Yu et al. (2005)

DCL3

RNase III

24 nt siRNA synthesis

Chromatin modification

Qi et al. (2005)

RDR2

RNA dependent RNA Polymerase

rasiRNA synthesis

Chromatin modification

Xie et al. (2004)

AGO4

RNA slicer

RdDM

Chromatin modification

Zilberman et al. (2003)

AGO6

RNA slicer

RdDM

Chromatin modification

Zilberman et al. (2003)

AGO9

RNA slicer

RdDM

Chromatin modification during female gametophyte formation

Havecker et al. (2010)

HDA6

Histone deacetylase

Remove acetyl group

Chromatin modification

Aufsatz et al. (2002)

DRD1

SNF2-like chromatin remodelling factor

RdDM

Chromatin modification

Kanno et al., (2004)

CMT3

DNA methyl-transferase

Methylation cytosine

Chromatin modification

Chan et al. (2005)

DRM1/2 DNA methyl-transferase

Methylation of cytosine

Chromatin modification

Cao and Jacobsen (2002)

MET1

DNA methyl-transferase

Methylation of Cytosine

Chromatin modification

Aufsatz et al. (2004)

KYP

DNA methyl-transferase

H3 K9 methyl-transferase

Chromatin modification

Du et al. (2014)

DDM1

DNA methyl-transferase

Maintenance of CG methylation

Chromatin modification

Matzke et al. (2005)

SGS3

Coiled-coil protein

siRNA amplification

siRNA

Mourrain et al. (2000)

DRB4

dsRBP

Interacts with DCL4

tasiRNA

Qu et al. (2008)

RNA PolII

DNA dependent RNA Polymerase

Transcribes the target gene

siRNA miRNA

Bartel (2004)

RNA PolIV

DNA dependent RNA Polymerase

ssRNA production for synthesis of 24 nt siRNA

Chromatin modification

Smith et al. (2007)

RNA PolV

DNA dependent RNA Polymerase

Synthesis of scaffold ssRNA which interacts with the siRNA

Chromatin modification

Haag and Pikaard (2011)

Classy 1 Chromatin remodeller

Synthesis of dsRNA

Chromatin modification

Smith et al. (2007)

ROS1

Repressor of silencing

Demethylation of DNA

Chromatin modification

Gehring et al. (2009)

DML2/3

Demeter-like2/3

Demethylation of DNA

Chromatin modification

Gehring et al. (2009)

RDM1

RNA-directed DNA methylation

RNA Pol V transcription

Chromatin modification

Gao et al. (2010)

SHH1

Sawadee Homeodomain Interacts with H3K9me and Homologue 1 recruits PolIV to the loci

Chromatin modification

Borges and Martienssen (2015)

KTF1

Kow domain-containing transcription factor 1

recruits AGO4 onto the PolV

Chromatin modification

He et al. (2009)

MORC6

Microrchidia 6

interact with PolV

Chromatin modification

Brabbs et al. (2013)

Surfacing the Role of Epigenetics in Host–Virus Interaction |  61

binding of DRM2 on the target region is regulated by base pairing between siRNAs and transcript of Pol V (Zhong et al., 2014). Contrastingly, in mammals, dual role of DNMTs has been proposed, having role in CpG methylation and active demethylation of methylated CpG islands via deamination (Métivier et al., 2008). Apart from this repressor of silencing1 (ROS1), demeter-like2 (DML2), and DML3 are DNA glycosylases that actively mediate DNA demethylation via a base excision repair process (Gehring et al., 2009). Dicer-like enzymes RdDM involves synthesis of 24 nt siRNAs which elicits de novo cytosine methylation in a sequencespecific manner. The biogenesis of these sRNAs involves core proteins of RNAi machinery: Dicerlike proteins (DCL), RNA-dependent RNA polymerase (RDR) and Argonaute (AGO). Dicer is an evolutionarily conserved large 1 (RNase III) type enzyme that cleaves dsRNA to produce 21–24 nt small RNAs (sRNA). Dicer proteins have two dsRNA binding domains, PAZ (Piwi/ Argonaute/Zwille) domain at one end and dual RNase III domains at another end and other domains such as DEAD box and helicase-C. It associates with dsRNA via its PAZ domain and then dices it with RNase III domains to produce 21–26 nt long siRNAs (Bernstein et al., 2001). Four DCL proteins are encoded in Arabidopsis genome (DCL1-DCL4) and each one has precise key functions in specific silencing pathways. Amongst these DCL1 is predominantly involved in miRNA synthesis, while DCL3 in 24 nt heterochromatic siRNA biogenesis (Ramachandran and Chen, 2008b). DCL2 and DCL4 are destined for virusderived siRNAs (vsiRNAs) biogenesis (Bologna and Voinnet, 2014). DCL4 initiates the generation of 21 nt siRNAs and is the principal component of antiviral defence. DCL2 acts as substitute required for synthesis of 22 nt siRNAs and are activated specifically when DCL4 is inactive/absent in Arabidopsis (Wang et al., 2011). Deep sequencing of virus-derived siRNAs revealed that 21 nt siRNA are more profuse in comparison to the 22 nt siRNA, thus DCL4 serve as the dominant viral sensor (Raja et al., 2010). It was observed that in plants infected with DNA viruses, the 24 nt vsiRNAs produced by DCL3 are predominant than 21- and 22 nt siRNAs (Blevins et al., 2006). DCL3 works in association

with RNA-binding proteins such as Tough (TGH), MOS4 and Dawdle (DDL) required for efficient processing of dsRNA (Yu et al., 2008; Ren et al., 2012). It also interacts with CELL DIVISION CYCLE 5 (CDC5), a MYB-related DNA binding protein, and PRL1, conserved WD-40 containing protein, for optimal activities and synthesis of siRNAs (Zhang et al., 2013, 2014). Simultaneously, several studies demonstrated that DCL1 promotes DCL3- and DCL4- derived siRNA accumulation upon dsDNA geminiviruses (CaMV or CaLCuV) infection (Moissiard and Voinnet, 2006). Contrastingly, study on dcl mutants revealed the antagonistic role of DCL1 in viral resistance. In Turnip crinkle virus (TCV) infected plants; DCL4 and DCL3 were negatively regulated by DCL1 and down-regulation of DCL1 led to their higher expression (Qu et al., 2008). Thus, the role of DCLs are not yet clearly understood and it needs to be further validated using quadruple dcl mutant plants. The sRNAs produced from DCL-mediated processing are methylated at 2′-OH of 3′ terminal nucleotides by HEN1 methyltransferase (Yu et al., 2005). This methylation protects sRNA molecules from uridylation and degradation by exoribonucleases like sRNA degrading nucleases (SDN1–3) (Li et al., 2005; Ramachandran and Chen, 2008a). Argonaute The processed and methylated siRNAs duplex is then incorporated into the effector complex termed as RNA induced silencing complex (RISC). It is a ribonucleoprotein complex, functionally reorganized with endonucleases called Argonaute (AGO) proteins and other uncharacterized components. These are the effector proteins and are conserved from Archaea to humans (Tolia and Joshua-Tor, 2007). These proteins have a bilobed structure, constituting of four regions: N-terminal domain, PAZ, MID and PIWI domain. The PAZ domain binds the two nucleotide 3′-overhang of sRNA whereas PIWI domain adapt RNaseH activity and it hydrolyses the target mRNA (Song et al., 2004; Cenik and Zamora, 2011). MID domain anchors 5′ end of guide strand sRNA (Vaucheret, 2008). Arabidopsis genome encodes for ten AGOs, but precise role of several AGOs in plant immune response needs to be explored. AGO1, AGO2, AGO5 and AGO7 were identified as a prerequisite for the protection against virulent viruses. AGO1 is the

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main slicer endonuclease that cleaves the targets of siRNA and miRNA whereas AGO7 is the subordinate slicer (Qu et al., 2008). AGO2 was reported to function during Cucumber mosaic virus (CMV) infection (Harvey et al., 2011), AGO10 has analogous functions as AGO1 (Baulcombe, 2004) and AGO4, AGO6 and AGO9 are essential in the TGS mechanism (Zilberman et al., 2003; Havecker et al., 2010). It has been reported that AGO4 and AGO6 varied in their co-localization with DNA dependent RNA polymerases, AGO6 associates with Pol V in the nucleoplasm whereas AGO4 localizes with Pol V in perinucleolar and Pol II in nucleoplasm (Duan et al., 2015). Amongst these, AGO4 is the key slicer in RdDM that initiates the repression of chromatin in sequence specific by channelizing the interaction between RNA polymerase V and DRM2 (Zhong et al., 2014). AGO4 interacts with large subunit of Pol V and initiates base pairing between 24 nt siRNA and Pol V dependent transcript. Kow domain-containing transcription factor 1 (KTF1)/ SPT5L act as a scaffold protein and regulates the interaction between AGO4 and Pol V (Rowley et al., 2011). Recently, it was observed that AGO4 and siRNA conjugates are also present in the cytoplasm. This study revealed that AGO4 might bind to 24 nt siRNA in the cytoplasm and this complex then moves to the nucleus where it initiates the chromatin modification (Ye et al., 2012). Further, AGO9 was also found to play an important role during RdDM via Pol IV-dependent mechanism (Olmedo-Monfil et al., 2010). It has been found to function in the female gametophyte and silence the transposon elements. Further, phenotypically, the ago9 mutant resembled the double mutant of Pol IV and Pol V thus signifying the role of AGO9 in RdDM (Olmedo-Monfil et al., 2010). Subsequently, it was observed that methylation of LTR is targeted by AGO6-associated with 24 nt siRNA and Pol V. Thus, it was proposed that AGO6 is the main component in Arabidopsis RdDM (Haag and Pikaard, 2011). On the other hand, recently it was elucidated that AGO6 function in RDR6-RdDM in which 21–22 nt siRNAs guide AGO6 to the target region on chromatin. It was hypothesized that Pol IV-RdDM originated from AGO6-RdDM through the course of evolution in Arabidopsis (McCue et al., 2015). Similarly, AGO2 was also found to induce DNA methylation in

association with 21–22 nt siRNA via non-canonical mechanism (Pontier et al., 2012). DNA dependent RNA polymerase IV and V Two types of plant-specific RNA polymerases (Pol) are prerequisite for the RdDM pathway: Pol IV and Pol V, which are evolved from Pol II. Pol IV and Pol V consist of 12 subunits and the largest subunits are NRPD1 and NRPE1, respectively (Haag and Pikaard, 2011). The role of Pol IV and Pol V in RdDM was revealed by studying the nrpd1 and nrpe1 mutants and it was observed that these polymerases have a non-redundant function in RdDM (Pontier et al., 2005). Apart from transcriptional gene silencing, Pol IV and V also function in RNA primed-DNA repair mechanism (Haag et al., 2012). Pol IV is required for synthesis of 24 nt siRNAs in association with RNA-dependent RNA polymerase 2 (RDR2). Pol IV acts upstream of RDR2 and Pol IV-dependent single-stranded (ss) transcripts and converted to double-stranded (ds) transcripts by RDR2 which are cleaved by DCL3 to generate 24 nt siRNAs. These 24 nt siRNAs then loaded into AGO4 and this complex initiates the DNA methylation by binding onto the Pol V-dependent transcript. As discussed earlier, AGO4 interacts with Pol V and alleviate the complex (Rowley et al., 2011). Further, it was shown that Defective in RNA-directed DNA Methylation (DRD1), a putative SWI2/SNF2-like chromatin remodelling ATPase (Huettel et al., 2007); Defective in Meristem Silencing 3 (DMS3), consisting of the hinge domains of cohesins and condensins (Wierzbicki et al., 2009); DMS11, consisting of ATPase domain (Lorković et al., 2012) and RNA-directed DNA Methylation1 (RDM1), a single-stranded DNA binding protein (Gao et al., 2010) are regulators of RdDM in association with Pol V. These proteins form a complex termed DDR complex. DRD1 and DMS3 are prerequisite for synthesis of Pol V-dependent transcripts (Law et al., 2010; Matzke and Mosher, 2014) whereas RDM1 serves as a scaffold protein between AGO4-siRNA-Pol V complex and DRM2. Besides these proteins, SU(VAR)3–9 HOMOLOGUE 2 (SUVH2) and SUVH9, methyl-DNA binding proteins are essential for link between Pol V and chromatin (Liu et al., 2014). The transcripts of Pol V are 200 nt in length and

Surfacing the Role of Epigenetics in Host–Virus Interaction |  63

lack poly-A tails and 5′ ends consists of capped/5′ triphosphates and 5′ monophosphates (Wierzbicki et al., 2008) whereas Pol IV transcripts also lack poly-A tails along with the 5′ cap (Li et al., 2015). RNA-dependent RNA polymerase Arabidopsis encodes six RNA dependent RNA polymerases (RDRs) which play a role in vsiRNA generation and confer resistance to geminiviruses (Butterbach et al., 2014). RDRs convert the ssRNA molecule to dsRNA molecule. RDR6 contributes to systemic signalling by synthesis of dsRNA using target mRNA as the template and vsiRNA as the primer leading to de novo production of complementary RNA strand in association with SGS3. Further, RDR6 contributes to the synthesis of distinct classes of siRNAs produced by cleaving activity of DCL1, DCL2 or DCL4 (Mlotshwa et al., 2002). In contrast, RDR2 has been revealed to direct chromatin modification by aiding DCL3 in the biogenesis of 24 nt siRNAs (Xie and Yu, 2015). It has been reported to interact with Pol IV larger subunit NRPE1, DCL3 and AGO4 (Li et al., 2006). Mechanism of RdDM The sRNA sequencing experiments predicted that in plants, 24 nt small RNAs are most profuse sRNAs species. However, the individual heterochromatic siRNA loci are not sustained even in the closely related species (Ma et al., 2010). In canonical model (Fig. 3.2), binding between 24 nt siRNAs and complementary (scaffold) RNAs instigate the DNA methylation mechanism (Matzke and Mosher, 2014). The biogenesis of heterochromatic siRNA initiates with transcription by DNA-dependent RNA polymerase (Pol IV) to produce ssRNAs at its target loci. RDR2 catalyse the conversion of these transcribed sRNAs to dsRNA in association with the chromatin remodeller, Classy 1 [CLSY1] (Borges and Martienssen, 2015). The binding of Pol IV onto the target region on the chromatin is regulated by Sawadee Homeodomain Homologue 1 (SHH1) which interacts with H3K9me and recruits Pol IV to the loci. Further, these dsRNAs are cleaved by DCL3 to produce 24 nt siRNAs, and are stabilized by methylation at their 3′-OH groups by HUA Enhancer 1 (HEN1) ( Ji and Chen, 2012). The guide strand incorporates into AGO4/AGO6 or AGO9 (expressed specifically in reproductive tissue) containing RITS complex, which then

enters the RNA Pol V-mediated pathway of de novo DNA methylation. KTF1 acts as the bridge and recruits AGO4 onto the Pol V largest subunit. Pol V transcripts activate the heterochromatin formation. It interacts with the methyl-DNA-binding proteins, SET domain containing SU (VAR) 3–9 homologue 2 (SUVH2) and SUVH9 and transcribes the target locus. Pol V transcripts function as a scaffold RNA which interacts with the siRNA associated with AGO4/6. Heterochromatic siRNAs bound AGO4/6 interacts with chromatin modifiers such as DRM2 which catalyses de novo methylation at the siRNA-targeted site (Haag and Pikaard, 2011). RNA-directed DNA Methylation 1 (RDM1) interacts with AGO4 and DRM2, thus plays a crucial role in recruitment of DRM2 onto the target sequence. Concurrently, RDM1 also interacts with Defective in RNA-directed DNA Methylation 1 (DRD1) to open the DNA duplex and regulates Pol V-mediated transcription (Gao et al., 2010). Subsequently, the putative cohesion-like proteins DMS3 and MICRORCHIDIA 6 (MORC6) interact with Pol V and assist in creating and maintaining the unwound state of target region. Interestingly, in A. thaliana, many endogenous sRNAs are derived from intergenic regions and repeats, which might potentially target chromatin modifications to enhancer elements or promoter regions and transposons (Xie et al., 2004). Along with Pol IV and Pol V, Pol II might synthesize 24 nt siRNAs and scaffold RNAs (Zheng et al., 2009) and similar to Pol V, Pol II also consists the AG hook domain required for interaction with AGO4/6 (Zheng et al., 2009). Simultaneously, several alternative pathway ‘non-canonical RdDM’ that requires components of PTGS have been reported recently. These pathways include: RdDM induced by miRNA and tasiRNA Besides PTGS, mature microRNA (miRNAs) and trans-acting siRNAs (tasiRNAs) might direct the chromatin remodelling based on the type of AGO protein and the size of the loaded miRNA (Fig. 3.3). In plants, canonical biogenesis of 21–22 nt miRNAs involves DCL1 and AGO1 proteins that target and cleave the complementary messenger RNAs (mRNAs). However, in studies on rice, moss and Arabidopsis 24 nt miRNA bound to AGO4 were identified to initiate DNA methylation at the target loci (Chellappan et al., 2010; Khraiwesh et

64  | Sharma et al. Hcy

ADK

Methionine

SAH SAM

ssDNA

PolIV trasncript

CLSY1

M M M

PolV

Pol IV

CMT3 MET1 KYP2

DRM2 AGO4

R DRM2 D M 1

RDR2

dsRNA DCL3

24 nt siRNA duplex

HEN1

AGO4

Methylated siRNA duplex

AGO-RITS

Figure 3.2 RNA dependent DNA methylation canonical pathway: RdDM is initiated by Pol IV- dependent transcription. Further, 24 nt siRNAs are synthesized by the activity of RDR2 which convert the singlestranded transcript of Pol IV to dsRNAs. The dsRNAs are then cleaved by DCL3 to produce 24 nt siRNAs. These siRNAs are incorporated into RITS complex consisting of Ago4/6. This siRNA–Ago complex then interacts with Pol V transcript and initiates methylation of the complementary DNA sequence by recruiting Domains rearranged methyltransferase 2 (DRM2). This methylation is maintained by Chromomethylase (CMT; methylation maintenance), Methyltransferase (MET; maintenance of methylation), Kryptonite (KYP; methylation maintenance and H3K9 methylation). DNA methylation subsequently aids in histone modifications leading to the heterochromatin formation.

al., 2010; Wu et al., 2010). The hairpin molecule is cleaved by DCL3 to produce 24 nt miRNAs which loads onto AGO4 which induces chromatin modifications. Apart from miRNAs, tasiRNAs are also involved in TGS pathway. Their biogenesis is induced by binding of miRNA onto the TAS RNA precursor and its cleavage by DCL4. These 21 nt tasiRNAs form complex with AGO1 and initiate PTGS. However, some tasiRNAs are processed by DCL1 to produce 21 nt tasiRNAs which are incorporated in AGO4/6 that can participate in RdDM

by interacting with its target region within the Pol V transcript (Nuthikattu et al., 2013). RDR6- dependent DNA methylation Several findings report that canonical pathway alone does not regulate the virus infection in plants (Herr et al., 2005; Raja et al., 2008). RDR6, a component of PTGS pathway which produces dsRNA from ssRNA as template, was identified to play an important role in RdDM against viruses (Stroud et al., 2013). The transcripts of Pol II are converted to

CMT3 MET1 KYP2

miRNA gene

ssDNA

Pol II

PolIV trasncript

Pri-miRNA DCL3

CLSY1

Pre-miRNA

M M M

PolV

Pol IV

DRM2 AGO4

R DRM2 D M 1

RDR2

dsRNA DCL3 24 nt miRNA duplex

HEN1

AGO4

Methylated siRNA duplex

AGO-RITS

Figure 3.3  MiRNA and tasiRNA induced RdDM: apart from siRNAs, hairpin-loop transcripts of miRNA and tasiRNAs produced by miRNA–mediated cleavage can be cleaved by DCl3 to produce 24 nt miRNAs or siRNAs. These mature miRNAs and siRNAs are incorporated in AGO4/6 that can participate in RdDM and initiate DNA methylation.

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dsRNA by RDR6 and cleaved by DCL2/4 to produce 21–22 nt siRNAs which are then loaded onto AGO2 (Fig. 3.4). This AGO2-siRNA (21–22 nt) complex interacts with Pol V transcripts and activates methylation by recruiting DRM2. This methylation further triggers the canonical RdDM mechanism via Pol IV, RDR2 and DCL3. Since, DCL2/DCL4 is occupied for synthesis of 21–22 nt siRNAs, DLC3 is activated for generating 24 nt siRNAs and initiate RdDM (Marí-Ordóñez et al., 2013). This de novo methylation is then maintained by MET1, CMT3 and DDM1 (Panda and Slotkin, 2013). Role of RdDM in plant defence against geminiviruses Geminiviridae In 2015, International Committee on Taxonomy of Viruses (ICTV) has distinguished seven orders, 111 families, 27 subfamilies, 587 genera, and 3704 species of viruses (ICTV Virus Taxonomy 2015; www.ictvonline.org/virusTaxonomy). Amongst 111 families, 82 families are not yet assigned to any order. One such family of plant viruses is Geminiviridae which is one of the diverse and most important families of plant viruses that infect a wide range of plant species. Geminiviruses are characterized by their geminate shaped, twinned icosahedral particles with quasi-isomeric morphology with 110 copies of Coat protein (CP) fitted in 22 pentameric capsomeres. Its genome is ≈ 2.5 kb to 3.0 kb in size, organized as covalently closed and circular ssDNA (Rojas et al., 2005). Geminiviruses replicate via dsDNA replicative intermediates by rolling-circle replication (RCR) mechanism within the host nucleus. Viral dsDNAs associate with cellular histones and organize as minichromosomes (Pilartz and Jeske, 1992). As geminiviruses encode few proteins and depend on host machinery for replication and transcription, it emphasis their use as models for fundamental host processes; DNA replication, transcription, cell cycle and epigenetic regulation (Hanley-Bowdoin et al., 2004). They are transmitted through insects and could be bipartite or monopartite. Monopartite viruses contain single virus genome, i.e. DNA-A, while bipartite viruses comprise two genomic components, i.e. DNA-A and DNA-B. Geminiviridae is

grouped into nine genera and every genus has a CP which forms the viral capsid and replicationassociated protein (Rep) which is required for production of dsDNA intermediate via RCR mechanism. Geminiviruses have bi-directional promoter for the transcription of either the virion (V) or complementary (C) sense DNA strand-specific transcripts. It has a common intergenic region (IR) of about 300 bp. IR contains a conserved nonanucleotide sequence 5′-TAATATTAC-3′ which forms a stem loop structure from where synthesis of ssDNA begins for virus replication (Bisaro, 1996). Apart from mono- or bipartite genome, presence of an additional molecule, i.e. satellite DNA has also been reported (Dry et al., 1997; Fiallo-Olivé et al., 2012). Satellite DNAs (DNA-β or DNA-α) of monopartite viruses are smaller in size (≈ 1.4 kb), single functional protein encoding, and are required for characteristic symptom development of the disease. Classification of Geminiviridae The number of genomic molecules, its organization, type of insect vector, and host plants infected act as the criteria to classify Geminiviridae family into nine genera: Becurtovirus, Begomovirus, Capulavirus, Curtovirus, Eragrovirus, Grablovirus, Mastrevirus, Topocuvirus and Turncurtovirus (Sahu et al., 2014a; Zerbini et al., 2017). The genus Curtovirus of the Geminiviridae family has a monopartite genome which encodes seven proteins. Leafhoppers or treehoppers are the main vector for transmission of the viruses belonging to this group. Genus Becurtovirus has two major species, i.e. Beet curly top Iran virus and Spinach curly top Arizona virus. They are biologically similar to genus Curtovirus; however, genome organization (monopartite genome encoding five proteins) resembles the genus Mastrevirus. Viruses of this genus are transmitted through leafhoppers and cause curly top disease in various dicot species. Eragroviruses (Eragrostis curvula streak virus) are genomically monopartite in nature, encoding four proteins, but their mode of spreading is still unknown. The genus containing the second largest number of species, Mastrevirus, has a monopartite genome consisting of four ORFs, two in a positive sense and other two in a negative sense. The members of this genus infect monocotyledonous plants and are transmitted by leafhoppers. Topocuvirus

CMT3 MET1 KYP2

miRNA gene

ssDNA

Pol IV

Pol II PolIV trasncript

RDR6

DCL2/4

CLSY1

M M M

PolV

DRM2 AGO4

R DRM2 D M 1

RDR2

dsRNA DCL3

HEN1

24 nt miRNA duplex

HEN1

AGO4/6

Methylated siRNA duplex

AGO-RITS

Figure 3.4  RDR6-dependent non-canonical pathway of RNA dependent DNA methylation: plants have also developed non-canonical pathway as a defence mechanism against lately integrated transposons. It involves transcripts of transposon via Pol II. These transcripts are converted into dsRNAs by RDR6 which are then diced by DCl2/4 to generate 21–22 nt siRNAs. As a deviation from PTGS pathway, these 21–22 nt siRNAs are recruited on to AGO2 which interacts with Pol V and DRM2 initiating de novo methylation.

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(Tomato pseudo-curly top virus) have monopartite genome enclosed of six ORFs. Topocuvirus mainly infects dicotyledons and is known to be transmitted by tree hoppers. Turncurtovirus comprises a single member (Turnip curly top virus). It has monopartite genome which comprises six ORFs encoding various proteins which are necessary for virus replication and systemic movement upon infection. Similar to eragroviruses, mode of Turncurtovirus spread is still unidentified. In the subsequent section, a detailed description of begomoviruses is provided. Begomovirus Begomoviruses are the causative agent of numerous diseases in economically important dicotyledonous plants. The genus name was assigned on the name of its species: Bean golden mosaic virus (BGMV). Viruses of this genus hinder the agricultural production in tropical and subtropical regions of the world (Rojas and Gilbertson, 2008). Begomoviruses redirect host machineries by initiating re-replication, interfering cell signalling and transcriptional controls and inhibiting defence pathways (HanleyBowdoin et al., 2013). Based on phylogenetic studies, begomoviruses can be divided into two groups, Old World (OW) and New World (NW) begomoviruses and they are genetically distinct (Briddon et al., 2010). The genome of NW viruses is bipartite, whereas the OW begomoviruses have either a monopartite genome along with satellite

DNAs or a bipartite genome (Fauquet et al., 2008). OW includes areas of eastern hemisphere, Asia, Africa, Europe and NW includes western hemisphere, America (Padidam et al., 1999). Besides, OW viruses are genetically more diverse and additionally have a conserved gene, AV2/V2 (precoat protein) in DNA-A which is not encoded by the NW begomoviruses (Rybicki, 1994). As it has been reported that begomoviruses have higher tendency for recombination and attaining new DNA components; thus, they are extremely vulnerable to evolutionary process, leading to emergence of new virulent strains. Owing to their association with severe diseases in economically and socially relevant crops, researchers have made tremendous progress to illustrate and understand the biology of these viruses. Genome organization of Begomovirus Begomoviruses can be bipartite (two genomic components) or monopartite (single component). Bipartite begomoviruses consist of two genome components DNA-A and DNA-B ≈ 2.6 kb (Fig. 3.5). DNA-A consists of six ORFs which help in virus replication and transcription whereas DNA-B encodes two ORFs responsible for virus movement. DNA-A virion/positive sense strand encodes only two proteins, AV/CP and pre-coat protein (AV2) required for virus enveloping, movement in plant and suppression of host defence (Rojas et al., 2005). The complementary/negative sense-strand

IR

IR

DNA A (≈2.7 Kb)

DNA B (≈2.6 Kb)

N SP

CP

AC

4

AV 2

MP

p

Re TrAP

RE

n

Figure 3.5  Organization of geminivirus genome: orientations of corresponding open reading frames (ORFs) are depicted with arrows. DNA-A encoded proteins, i.e. AC1/C1 (replication initiation protein); AC2/C2 (transcription activator); AC3/C3 (replication enhancer); AV/V1 (coat protein) are depicted in arrows according to their orientation in the genome. DNA-B specific ORFs encoding BV1/NSP (Nuclear shuttle protein) and BC1/ MP (Movement protein) are shown.

Surfacing the Role of Epigenetics in Host–Virus Interaction |  69

encodes for AC1/Rep, transcription activator protein (AC2/TrAP), replication enhancer (AC3/ REn), and pathogenesis related protein (AC4). DNA-B complementary strand has ORF for movement protein (BC1/MP) and the virion strand encodes for nuclear shuttle protein (BV1/NSP) (Fauquet et al., 2008). The ORFs on DNA-A and DNA-B are transcribed in bi-directional manner and are separated by a common region (CR) of 180–200 nt. This noncoding CR or IR in geminivirus genomes contains origin of replication required for RCR and regions for initiating transcription of viral genes. Within IR there are promoter elements such as TATA boxes and sequences recognized by various transcription factors including ‘AG’ motif and G-box (Hanley-Bowdoin et al., 2000; Orozco et al., 1997). IR contains a conserved GC rich inverted repeat called as the hairpin motif that forms a cruciform or stem–loop structure. This GC-rich stem encloses a consensus AT-rich nonanucleotide sequence, 5′-TAATATTAC-3’, in the loop conserved among geminivirus genomes where Rep-mediated nicking initiates RCR. IR consists of repeated upstream iterative sequence motifs known as ‘iterons’ for binding of Rep proteins. The region between repeats of iteron and stem loop is called as origin of replication or ori. Functions of Begomovirus-encoded proteins Coat protein (AV1) is a protein of ≈ 28 kDa that is implicated in the encapsidation of the new viral genome (Briddon et al., 1990; Boulton, 2002). It has the ability to interact with both ssDNA and dsDNA (Liu et al., 1997; Kunik et al., 1998; Palanichelvam et al., 1998). CP has been found to be nuclear localized and its interaction with α-importin is essential for cytoplasmic trafficking of the host during virus infection (Liu et al., 1999). In monopartite viruses, it facilitates cell to cell and long-distance movement of viral genome (Boulton et al., 1989; Liu et al., 1997; Kotlizky et al., 2000). It is also recognized as a determinant of vector specificity (Noris et al., 1998; Höhnle et al., 2001). In the virus-infected cells, pre-coat protein (AV2) is found to be localized in the cytoplasm as well as cell periphery (Chowda-Reddy et al., 2008). AV2 acts as a suppressor of plant RNA interference (RNAi) machinery (Chowda-Reddy et al.,

2008). Mutation in pre-coat protein of bipartite Begomovirus ToLCNDV (Tomato leaf curl New Delhi virus) exhibited reduction in the viral DNA accumulation in infected plants (Padidam et al., 1996). It also facilitates the intracellular, intercellular and systemic movement of the monopartite begomoviruses (Stanley et al., 1992; Hormuzdi and Bisaro, 1993). AC1 (Rep) encodes for ≈ 40 kDa multifunctional protein which is most essential factor for initiation and termination of viral genome replication (Fontes et al., 1994; Hanley-Bowdoin et al., 2000). It can exclusively bind to a repeated consensus sequence present in IR of dsDNA viral genome. It cleaves and ligates DNA at conserved nonanucleotide sequence TAATATT↓AC within a hairpin loop of the plus-strand origin (Lazarowitz et al., 1992; Orozco et al., 1997). Rep is a key component in the interaction with plant cell cycle regulatory elements (Kong et al., 2000; Kong and Hanley-Bowdoin, 2002), cellular proteins (Morilla et al., 2006) along with the viral REn protein (Settlage et al., 2005). Recent information suggests the role of Rep in transcriptional gene silencing (Rodríguez-Negrete et al., 2013). During replication of plus-strand, it acts as a DNA helicase to unwind viral DNA (Bisaro, 1996). Apart from its role in viral genome replication, Rep interacts with several host factors involved in DNA replication. Rep plays an essential role in activating the host replication machinery by interacting with the plant cell cycle regulatory elements. By its cyclin interaction motif (RXL motif) Rep interacts with retinoblastomarelated protein (RBR) (Gutierrez, 2000). This interaction activates E2F-family transcription factors which in turn activates transcription of S-phase-specific genes, thus removing the cell cycle arrest. It also plays an essential role in activating the replication of viral as well as plant DNA. It binds to subunits of host DNA polymerase complexes (Castillo et al., 2003; Bagewadi et al., 2004; Settlage et al., 2005; Bruce et al., 2011), ssDNA binding proteins (Singh et al., 2007; Lozano-Durán et al., 2011), and recombination/repair process associated proteins to impede host DNA replication event. These critical roles in the genome replication and interactions with multiple proteins make Rep an exceptional target for antiviral resistance strategy by the expression of mutant proteins. Host genes play an important role during virus

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infection and act as a counter-defence mechanism. On the other hand, viruses are also armed with suppressors of host gene activation and gene silencing. In this regard, AC2 from bipartite and monopartite begomoviruses serves as a factor for virus pathogenicity and suppression of gene silencing (Voinnet et al., 1999; Trinks et al., 2005; Vanitharani et al., 2005; Chowda-Reddy et al., 2008). It was observed that expression of TrAP proteins from geminiviruses decreased the resistance in N. benthamiana and tobacco plants (Sunter et al., 2001; Hao et al., 2003). Expression of AC3 encoding the replication enhancer protein is determined by the unidirectional left promoter (Shivaprasad et al., 2005). Interaction of this protein with Rep and PCNA is one of the perquisites of viral DNA replication process (Castillo et al., 2003; Settlage et al., 2005). REn of Tomato leaf curl Kerala virus has been identified to interact with Rep, which assists in enhancement of ATPase activity of Rep (Pasumarthy et al., 2011). Disruption of AC4 resulted in reduced viral DNA accumulation and symptom development ( Jupin et al., 1994; Teng et al., 2010). It has also been evidenced that AC4 may suppress the post-transcriptional gene silencing (PTGS) mechanism of host plant (Vanitharani et al., 2005; Fondong et al., 2007). Nuclear shuttle protein (BV1/NSP) is requisite for shuttling of newly synthesized viral DNA from nucleus to cytoplasm while MP actively binds with NSP-viral DNA complex and participates in cellto-cell movement of virus through plasmodesmata (Gafni and Epel, 2002). The MPs are also pathogenicity determinant of bipartite begomoviruses and the mutation at 3′ region has been linked with symptom development (Gafni and Epel, 2002). Mechanism of replication, transcription and movement of geminiviruses Geminivirus DNA replicates through RCR mechanism and requires host cell cycle machinery ( Jeske et al., 2001). When viral ssDNA (minus strand genome) is released from virions it is converted into an intermediate dsDNA replicative form (RF) of viral genome which acts as a template for replication and transcription (Gutierrez, 2000). In contrast to mastreviruses which have ≈ 80 bp primer complementary to the small region of IR,

begomoviruses depend on host proteins such as primase to produce a primer that initiate complementary strand synthesis (Hayes et al., 1988). This RF assembles into minichromosomes and is transcribed by host DNA-dependent RNA polymerase II (Pol II) into the viral transcripts. The interaction between Rep, REn and RBR interrupts the RBRE2F binding and leads to expression of host genes required for replication. Replication of geminivirus genome is initiated by nicking activity of Rep at nonanucleotide ‘TAATATTAC’ located in stem–loop structure of IR region. This causes cleavage of the phosphodiester bond and Rep gets covalently linked to the 5′ end of the cleaved DNA via a phosphotyrosine linkage (Laufs et al., 1995). The 3′ terminus is extended by host DNA polymerase using minus strand as template which finally relocates the parental plusstrand from the intact minus-strand template (Fig. 3.6). After synthesis of viral strand, the nonanucleotide is again cleaved followed by ligation due to nicking and closing activity of Rep, thereby leading to generation of the new viral DNA unit ( Jeske et al., 2001). Using dsDNA as template, geminivirus genome is transcribed in a bi-directional mode by host Pol II. The transcription of geminivirus generates overlapping RNA fragments, which are polycistronic in nature. In the later stages of infection Rep binds to iterons in the ori and inhibits its own transcription, thus enhancing the promoter strength for downstream ORFs. This initiates the production of TrAP which in turn activates the transcription of late viral genes, CP, NSP and MP (Vanitharani et al., 2005). NSP binds to the viral DNA and shuttles it across the nuclear membrane into the cytoplasm where it interacts with MP which traffics it across the plasmodesmata. The final stage of replication cycle involves encapsidation of virus genome by CP into virions which is then taken up by insect vector. Further, monopartite begomoviruses are restricted to phloem and are not transmissible through sap, in contrast to several bipartite begomoviruses which are sap transmissible and enter both phloem and non-phloem tissues (Fig. 3.6). Another mode of replication, recombinationdependent replication (RDR), has been reported in several geminiviruses such as ACMV, ToLCV, Abutilon mosaic virus (AbMV), Tomato golden mosaic virus (TGMV), BCTV and TYLCV. The

Surfacing the Role of Epigenetics in Host–Virus Interaction |  71

Primer synthesis Binding of NSP on virion sense strand

Complementary strand synthesis

dsDNA

Cell to cell movement of viral genome via MP

Nucleus

Transcription Strand Displacement Nicking by Rep

Translation Rep TrAP CP REn NSP MP

Nucleus

Cytoplasm

Figure 3.6  Schematic representation for geminivirus replication and movement in plants: geminivirus enters the cell and after uncoating, single-stranded circular viral DNA enters the nucleus of plant cell. The viral replication cycle is initiated by RNA primer synthesis and complementary strand synthesis by host DNA polymerase leading to the synthesis of dsDNA. This dsDNA serves as the template for transcription and viral proteins are synthesized in the cytoplasm. Rep is transported into the nucleus and initiates the rolling circle replication. With the help of nuclear shuttle protein, the newly synthesized circular single-stranded DNAs move from nucleus to cytoplasm. Movement proteins aid in movement of viral genome to other cells and various parts of host. Rep (Replication associated protein), TrAP (Transcription activator protein), CP (Coat protein), REn (Replication enhancer protein), NSP (Nuclear shuttle protein) and MP (Movement protein).

interaction between host recombination protein, RAD54 and Rep might play an important role in recombination. RDR is initiated by binding between single-stranded region of the partially replicated dsDNA and a closed, circular dsDNA at homologous site followed by homologous recombination and elongation of ssDNA. The recombinant dsDNA produced acts as a template for both ssDNA and dsDNA synthesis ( Jeske et al., 2001; Preiss and Jeske, 2003). TGS-mediated resistance to geminiviruses Plants perceive various endogenous and environmental stimuli including attack by a spectrum of pathogens, leading to changes in gene expression.

Plants have developed multifaceted defence mechanism to protect themselves from potentially harmful effects of various biotic factors (Muthamilarasan and Prasad, 2013). It is noteworthy that most studies on plant defence response during virus infection involve host-encoded sRNAs and associated silencing pathways (Padmanabhan et al., 2009; Zvereva and Pooggin, 2012), instead of hypersensitive response (HR). These sRNAs include miRNA (21–22 nt) that are endogenously encoded non-coding parts of the genome, and siRNAs (21–24 nt) which are generated by dicing of exogenous/endogenous dsRNA. The exogenous siRNAs originating from the molecules of infecting virus are termed as vsiRNAs and they aid in initiating the RNA silencing-mediated antiviral immunity.

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The vsiRNAs are generated from long dsRNA which are produced from exogenous molecules and are processed in DCL-dependent manner. These precursor dsRNAs are derived (a) from replicative intermediate dsRNA, produced during genomic replication of RNA viruses by virus-encoded RNA polymerases, or through bi-directional transcription in the case of DNA viruses, leading to overlapping sense and antisense transcripts; or (b) by convergent transcription which requires hybridization of unrelated RNA molecules with sequence complementarity. They are also produced from (c) structure-associated siRNA, synthesized from inverted-repeat viral transcripts with imperfectly base paired secondary structure (Fig. 3.7; Sharma et al., 2013). The sequencing and experimental data of siRNAs propose that these single-stranded viral transcripts with stem–loop structure are the hot spots for generation of vsiRNA initially and act as substrate for plant DCLs (Molnár et al., 2005). Simultaneously, sequencing-based approach also revealed that vsiRNAs are not randomly distributed along the viral genome, and majority of viral siRNAs map to the positive strand of viral genome (Molnár et al., 2005; Ho et al., 2007; Donaire et al., 2009; Qi et al., 2009). In plants, RdDM involves de

CV

novo methylation at cytosine-residue, irrespective of sequence context (Mette et al., 2000; Lewsey et al., 2016; Xie and Yu, 2015), in coding and promoter regions of silenced genes. It was observed that pathogen stress might shape the plant genome. In plants, epigenetic mechanism contributes in adapting towards specific environmental conditions. There are reports in which virus infection of tomato plants triggered changes in DNA methylation of plant genome (Mason et al., 2008). In tobacco plants infected with TMV, significant hypermethylation at CpG and CpHpG sites was observed. Antagonistically, loci-specific bisulfite sequencing analysis revealed hypomethylation in N gene-like R genes loci. This epigenetic response induced by pathogen attack might be a signal of an adaptive response by plants to be attacked by new pathogens (Kovalchuk et al., 2003; Boyko et al., 2007). Apart from regulating the plant growth and development, DNA methylation acts against exogenous DNA viruses. The dsRNA generated during viral infection can initiate sRNAs directed de novo DNA methylation via RdDM pathway resulting in transcriptional silencing of viruses (Chan et al., 2004; Raja et al., 2010; Yadav and Chattopadhyay,

Convergent replication

Bi-directional Transcription

Secondary Structure

dsRNA DCL

siRNA duplex

Figure 3.7  Schematic representation of different types of precursor dsRNAs: the dsRNA precursors of siRNA are derived from convergent transcription and via bi-directional transcription. The secondary structures within the viral genome or mRNA transcript are also a key source of dsRNA for synthesis of siRNA. DCL, Dicer like protein; dsRNA, double-stranded RNA; siRNA, small interfering RNA.

Surfacing the Role of Epigenetics in Host–Virus Interaction |  73

2011; Ceniceros-Ojeda et al., 2016; Deuschle et al., 2016; Jackel et al., 2016). It causes virus genome modification by initiating methylation at cytosine residues in virus DNA and associated histones and suppresses the transcription and/or replication of virus minichromosome (Raja et al., 2010, 2014; Paprotka et al., 2011; Ceniceros-Ojeda et al., 2016; Deuschle et al., 2016; Jackel et al., 2016). Geminiviruses are DNA viruses that have ssDNA genomes and replicate in the host nucleus by rolling-circle amplification producing dsDNA intermediates. These dsDNA intermediates act as template for both virus replication and transcription, and form minichromosomes in association with plant cellular histone proteins (Pilartz and Jeske, 2003, Vanitharani et al., 2005; Bisaro, 2006,). In plant host system, geminiviruses are targeted by RNA silencing and transcripts of virus genome are regulated by PTGS (Waterhouse et al., 2001; Raja et al., 2010; Patil and Fauquet, 2015). Simultaneously, several reports supported the theory that plants utilize DNA methylation to initiate defence against geminiviruses. SiRNAs generated against geminiviruses are derived from dsRNAs produced by hybridi­zation of sense and antisense transcripts (Akbergenov et al., 2006; Blevins et al., 2011; Ding and Voinnet, 2007). In case of geminivirus, these dsRNAs are produced during virus replication by the action of host RDRs, or by transcription of inverted repeats or converged promoters (Xie et al., 2004; Llave, 2010; Raja et al., 2010; Aregger et al., 2012, Szittya and Burgyán, 2013). These siRNAs can be of two types based on their size; the first is the long (24–26 nt) siRNA class, which correlates with sequence-specific methylation of target DNA for transcriptional suppression (TGS) and chromatin modulation; and the second is the short (21–22 nt) siRNA class, which is related to mRNA degradation (PTGS) (Hamilton et al., 2002; Pantaleo, 2011; Pooggin, 2013). Virus-derived siRNAs have been extensively studied during various plant– virus interactions (Chellappan et al., 2004; Molnár et al., 2005; Akbergenov et al., 2006; Ruiz-Ferrer and Voinnet, 2009; Dalakouras et al., 2013; Raja et al., 2014). The siRNA-mediated gene silencing has been shown to be involved in the recovery from the specific virus infection in plants (RodríguezNegrete et al., 2009; Sahu et al., 2010; Rajeswaran et al., 2014). For example, in Tomato golden mosaic virus (TGMV) and ACMV, in vitro methylation of

virus DNA retarded the virus replication in Nicotiana protoplasts (Brough et al., 1992; Ermak et al., 1993). Moreover, plants with mutations in genes for cytosine or histone methyltransferase, methyl cycle enzymes, and DCL proteins and components of RdDM pathways were hypersensitive to geminivirus infection as compare to wild-type and this susceptibility was due to reduced DNA methylation (Raja et al., 2008). Further, virus genome in the recovered host tissue was hypermethylated and a positive association was demonstrated between increased DNA methylation of virus genome and recovery of host plants (Hagen et al., 2008; Raja et al., 2008; Rodríguez-Nergete et al., 2009; Paprotka et al., 2011; Yadav and Chattopadhyay, 2011). In another study, it was demonstrated that the virus minichromosomes from symptomatic tissue of PepGMV-infected pepper had relaxed conformation and reduced level of DNA methylation. On the other hand, minichromosome isolated from recovered tissue, was compact and hypermethylated (Ceniceros-Ojeda et al., 2016). Recently, a study on Musa acuminata-infecting Banana streak pararetroviruses (BSV) revealed that 21–22  nt siRNAs targeted the virus coding regions whereas 24 nt siRNAs were found to be evenly targeting the whole virus genome (Rajeswaran et al., 2014). Moreover, antagonism of viroid on the geminivirus infection was also linked to DNA methylation (Torchetti et al., 2016). It was observed that in tomato co-infected with Tomato yellow leaf curl Sardinia virus (TYLCSV) and Potato spindle tuber viroid (PSTVd), the genome of TYLCSV was hypermethylated. It was a consequence of significant up-regulation of key genes involved in DNA methylation (MET1, CMT3, and DDM1) via PSTVd. Symptom development and disease severity differs from plant to plant and even between the genotypes of a particular host species. In our previous study, a naturally tolerant cultivar of tomato, namely H-88-78-1, was identified which has reduced infectivity and virus titre at 21 dpi in contrast to a susceptible cv. Punjab Chhuhara (Sahu et al., 2010). A crucial role of antivirus activity of siRNA was established, as in cv. H-88-78-1 lower abundance of virus genome was correlated with a relatively higher level of vsiRNAs production (Sahu et al., 2010). In order to understand the molecular mechanism of differential natural tolerance in tomato against ToLCNDV, susceptible

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and tolerant tomato cultivars were studied for the siRNA accumulation and pattern of distribution of siRNA along the virus genome after 21 dpi. It was observed that three regions (A1, A7 and A8) in DNA-A corresponding to IR and AC1 were the preferential targets of siRNA accumulation in the tolerant cultivar. In the susceptible cultivar, no such differential pattern was observed. Evaluation of siRNA accumulation demonstrated that tolerant cultivar synthesized more IR-specific siRNAs, amongst which 24 nt siRNAs were more abundant, and AC1-specific siRNAs which were mainly 21–22 nt long (Sahu et al., 2014b). Thus, based on these observations it can be proposed that PTGS targets the coding regions of virus genome leading to the degradation of virus transcripts while noncoding/intergenic regions are targeted by the TGS system. Further, it was inferred that both siRNA mediated TGS and PTGS phenomenon play crucial role in governing the tolerance in cv. H-88–78–1. These mechanisms alter the transcription rates of virus genes by initiating RNA degradation which may cause reduced infection. It was observed that transcription of a gene is dependent on degree of methylation of the virus promoter region and methylation within the gene. These results validate that geminivirus are the models for studying the mechanism of genome methylation and pathways

regulating cytosine methylation in plants (Sahu et al., 2014b). Suppressor of TGS encoded by geminiviruses It has been observed that plants synthesize 24 nt vsiRNAs during DNA virus infection. These 24 nt siRNAs can initiate DNA methylation of viral genome, ultimately leading to transcriptional silencing restricting the virus infection. Contrastingly, there is evidence demonstrating the presence of unmethylated viral dsDNA in the nucleus, producing Pol II-mediated transcripts of viral genes. Hence, DNA viruses might escape or repress RdDM. This inhibition of methylation cycle additionally provides evidence that the genome of virus is the main target for small-RNA-directed methylation. To counter this defence response, the genome of virus encodes suppressor proteins which suppress silencing mechanisms by interacting and inhibiting host proteins involved in silencing process. Table 3.2 summarizes the viral suppressors encoded by geminiviruses. Diverse viral suppressors have been identified which evolved independently and differ in their structure and function. Studies demonstrated that TrAP (AC2/C2/AL2/L2) is one of the most important suppressors of plant defence. AL2

Table 3.2  Virus-encoded suppressors of gene silencing Virus

Suppressor

Suppression Mechanism

Reference

Mungbean yellow mosaic AC2 virus

Interact with ADK required for S-adenosylmethionine (SAM) synthesis

Trinks et al. (2005)

Cabbage leaf curl virus (CaLCuV)

AC2

Interact with ADK required for S-adenosylmethionine (SAM) synthesis

Liu et al. (2014)

Tomato golden mosaic virus

AC2

Interact with ADK required for S-adenosylmethionine (SAM) synthesis

Wang et al. (2003)

Beet severe curly top virus

C2

Attenuates the degradation of SAMDC1 and represses the DNA methylation

Zhang et al. (2011)

ICMV

AC2

Inhibits KYP, histone methyltransferase

Sun et al. (2015); Castillo-Ganzalez et al. (2015)

Tomato yellow leaf curl China virus

βC1

Inhibits the activity of S-adenosyl Homocysteine Hydrolase (SAHH), required for synthesis of SAM

Yang et al. (2011)

Tomato yellow leaf curl virus

V2

Not known

Wang et al. (2014)

Tomato yellow leaf curl Sardinia virus (TYLCSV)

Rep/AC1

Reduces the transcript level of CMT3 and MET1

Rodríguez-Negrete et al. (2013)

ACMV

AC4

ss-sRNA binding

Chellappan et al. (2005)

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of Tomato golden mosaic virus (TGMV) and L2 of Beet curly top virus (BCTV) proteins act as the transcription activator and silencing suppressor. AL2/ L2 suppress host kinase proteins such as Adenosine kinase (ADK), which is prerequisite for translocation of methyl group from S-adenosyl-methionine (SAM) synthesis; a cofactor for methyltransferase, to C5 of cytosine (Wang et al., 2003; Buchmann et al., 2009; Yang et al., 2013). Recently, a study in Nicotiana benthamiana revealed that suppression of silencing by AL2 and L2 depends on the plant developmental stage ( Jamie et al., 2015). Both AL2 and L2 were able to inhibit silencing during vegetative phase whereas AL2 was found to reverse TGS in the reproductive stage also, proposing variation in the silencing mechanism during development stages in response to virus infection. Further, C2 regulates the proteasome-mediated degradation of S-adenosyl-methionine decarboxilase 1 (SAMDC1), which catalysis decarboxylation of SAM (dcSAM; competitive inhibitor of SAM). Increased levels of dcSAM further leads to reduced viral DNA methylation (Zhang et al., 2011). Apart from ADK and SAMDC1-based suppression of plant defence system, AC2 was also found to interact with RDR6 and AGO1 (Kumar et al., 2015) and histone methyltransferase, SUVH4/KYP (Castillo-González et al., 2015) thus inhibit the RNA silencing mechanism. Another viral protein essential for virus replication, Rep protein, exhibits TGS inhibition activity by reducing the expression of MET1 and CMT3 plant proteins and regulating the CG methylation level. Nicotiana benthamiana plants overexpressing TYLCSV Rep showed reduced level of MET1 and CMT3 transcripts and viral DNA methylation (Rodríguez-Negrete et al., 2013). Study on Tomato yellow leaf curl China virus (TYLCCNV) showed that the DNA betasatellite (TYLCCNB) required for the virulence of virus interacts with S-adenosyl homocysteine hydrolase (SAHH), a component of TGS (Yang et al., 2011). Thus, it can be hypothesized that viruses target the components of TGS machinery to suppress the plant defence system. These suppressor proteins can be targeted by methods such as RNA interference-based silencing through hairpin-based silencing, artificial miRNA and artificially targeted DNA methylation to enhance the virus tolerance.

Conclusion In recent years various studies have reported the roles of sRNAs in plants including regulation of gene expression at different regulatory levels, plant development and response against different stresses (biotic and abiotic). These sRNAs control the transcription of protein encoding genes by chromatin remodelling and initiating the DNA methylation. Epigenetic modifications manipulate the access of DNA for binding of transcription factor and enhancer proteins, thus regulate the degree of expression of gene. Further, siRNA-mediated TGS phenomenon plays crucial role in governing the tolerance against geminiviruses. The siRNA-mediated epigenetic modifications alter the transcription rates of virus genes by initiating DNA methylation which may cause reduced infection. Plants have developed siRNA-mediated de novo methylation and its maintenance against cytosines within the viral genome at both asymmetrical and symmetrical sites and repress the virus transcription. Apart from genes required for its replication and transcription, geminiviruses have been reported to encode suppressor proteins which repress the silencing mechanism and evade the plant defence system. The repression of methylation cycle by viral proteins provides support to the idea that genome of virus is the main target for small-RNA-directed methylation (Fig. 3.8). There is evidence that virus infection initiates epigenetic modifications within the host plant genome (Kovalchuk et al., 2003; Boyko et al., 2007; Mason et al., 2008). It can be hypothesized that this virus-induced epigenetic modifications can be transmitted to progeny by meiosis through meristem, thus inheriting the adaptive defence state in progeny. Future prospective Most of knowledge regarding role of epigenetics against viral pathogen is based on the evaluation of the mutant plants. The natural variation amongst different genotypes varying in their tolerance/resistance to viruses needs to be explored to understand the role of epigenetic adaptations in plant survival during virus infection. Several epigenetic techniques such as Chromatin immunoprecipitation

Geminivirus

ADK

Hcy

AC2

ssDNA Methionine

SAHH

Viral chromatin (dsDNA)

SAM

Pol IV PolIV trasncript

AC4

CLSY1

CMT3 MET1 KYP2 DRM2

PolV AGO4

RDR2

dsRNA

SAH

SAMDC1

AC1 AC2

βC1 dcSAM

Proteasomemediated degradation

R DRM2 D M 1

DCL3 24 nt siRNA duplex

AGO4

Methylated siRNA duplex

AGO-RITS

NUCLEUS

HEN1

Figure 3.8  Schematic illustration of antiviral transcriptional silencing machinery in plants and counter strategy by virus encoded silencing suppressors proteins: Viral DNA methylation is initiated by synthesis of 24 nt siRNAs which are recruited onto AGO4 leading viral genome methylation via DRM2. Different virus suppressors interrupt the methylation pathway, thus inhibiting the plant defence system. The viral suppressors are indicated in red colour within the figure.

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(ChIP), deep sequencing, single-cell restriction analysis of methylation (SCRAM), Bisulfite sequencing, RNA-antisense purification with mass spectrometry (RAP-MS), Methylated DNA immunoprecipitation (MeDIP) can be utilized to identify the histone modifications, and DNA methylation associated with the virus genome. Recently, genome editing approaches specifically clustered regularly interspaced short palindromic repeats (CRISPR)-Cas9 system have been used to generate resistance to geminiviruses. This system has been used to target different genomic regions of geminiviruses such as Bean yellow dwarf virus, Beet curly top virus, Merremia mosaic virus and Tomato yellow leaf curl virus to provide resistance in N. benthamiana and Arabidopsis plants. The methylated genomic regions of virus identified through different epigenetic techniques can be targeted for genome editing approaches during preparation of transgenic plants to confer resistance to viruses. In view of the fact that the exact mechanism of epigenetics is still not completely understood, thus determining the genome methylation pattern of geminiviruses, substrate/chromatin marks required for epigenetic modification and the mechanism that exists for viral genome methylation would help in understanding the mechanism and functions of epigenetics. Investigations are required to unveil the complex mechanism of epigenetics and the regulatory mechanism. Additionally, we need to identify the chromatin modifications regulating the plant immunity against the pathogens and their inheritance. References

Akbergenov, R., Si-Ammour, A., Blevins, T., Amin, I., Kutter, C., Vanderschuren, H., Zhang, P., Gruissem, W., Meins, F., Hohn, T., et al. (2006). Molecular characterization of geminivirus-derived small RNAs in different plant species. Nucleic Acids Res. 34, 462–471. Andika, I.B., Maruyama, K., Sun, L., Kondo, H., Tamada, T., and Suzuki, N. (2015). Different Dicer-like protein components required for intracellular and systemic antiviral silencing in Arabidopsis thaliana. Plant Signal Behav. 10, e1039214. https://doi.org/10.1080/155923 24.2015.1039214 Andrews, A.J., and Luger, K. (2011). Nucleosome structure(s) and stability: variations on a theme. Annu. Rev. Biophys. 40, 99–117. https://doi.org/10.1146/ annurev-biophys-042910-155329 Aregger, M., Borah, B.K., Seguin, J., Rajeswaran, R., Gubaeva, E.G., Zvereva, A.S., Windels, D., Vazquez, F., Blevins, T., Farinelli, L., et al. (2012). Primary and secondary siRNAs in geminivirus-induced gene silencing. PLOS

Pathog. 8, e1002941. https://doi.org/10.1371/journal. ppat.1002941 Aufsatz, W., Mette, M.F., van der Winden, J., Matzke, M., and Matzke, A.J. (2002). HDA6, a putative histone deacetylase needed to enhance DNA methylation induced by double-stranded RNA. EMBO J. 21, 6832–6841. Bagewadi, B., Chen, S., Lal, S.K., Choudhury, N.R., and Mukherjee, S.K. (2004). PCNA interacts with Indian mung bean yellow mosaic virus rep and down-regulates Rep activity. J. Virol. 78, 11890–11903. Baulcombe, D. (2004). RNA silencing in plants. Nature. 43, 356-363. Berger, S.L. (2007). The complex language of chromatin regulation during transcription. Nature 447, 407–412. Bernstein, E., Caudy, A.A., Hammond, S.M., and Hannon, G.J. (2001). Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409, 363– 366. https://doi.org/10.1038/35053110 Birnbaum, K.D., and Roudier, F. (2017). Epigenetic memory and cell fate reprogramming in plants. Regeneration 4, 15–20. https://doi.org/10.1002/reg2.73 Bisaro, D.M. (1996). Geminivirus DNA Replication. Cold Spring Harbor Laboratory Press. 9, 833-854. Bisaro, D.M. (2006). Silencing suppression by geminivirus proteins. Virology 344, 158–168. Blevins, T., Rajeswaran, R., Aregger, M., Borah, B.K., Schepetilnikov, M., Baerlocher, L., Farinelli, L., Meins, F., Hohn, T., and Pooggin, M.M. (2011). Massive production of small RNAs from a non-coding region of Cauliflower mosaic virus in plant defense and viral counter-defense. Nucleic Acids Res. 39, 5003–5014. https://doi.org/10.1093/nar/gkr119 Blevins, T., Rajeswaran, R., Shivaprasad, P.V., Beknazariants, D., Si-Ammour, A., Park, H.S., Vazquez, F., Robertson, D., Meins, F., Hohn, T., et al. (2006). Four plant Dicers mediate viral small RNA biogenesis and DNA virus induced silencing. Nucleic Acids Res. 34, 6233–6246. Bologna, N.G., and Voinnet, O. (2014). The diversity, biogenesis, and activities of endogenous silencing small RNAs in Arabidopsis. Annu Rev Plant Biol 65, 473–503. https://doi.org/10.1146/annurevarplant-050213-035728 Borges, F., and Martienssen, R.A. (2015). The expanding world of small RNAs in plants. Nat. Rev. Mol. Cell Biol. 16, 727–741. https://doi.org/10.1038/nrm4085 Boulton, M.I. (2002). Functions and interactions of Mastrevirus gene products. Physiol. Mol. Plant Pathol. 60, 243-255. Boulton, M.I., Steinkellner, H., Donson, J., Markham, P.G., King, D.I., and Davies, J.W. (1989). Mutational analysis of the virion-sense genes of maize streak virus. J. Gen. Virol. 70, 2309–2323. https://doi.org/10.1099/00221317-70-9-2309 Boyko, A., Kathiria, P., Zemp, F.J., Yao, Y., Pogribny, I., and Kovalchuk, I. (2007). Transgenerational changes in the genome stability and methylation in pathogen-infected plants: (virus-induced plant genome instability). Nucleic Acids Res. 35, 1714–1725. Briddon, R.W., Pinner, M.S., Stanley, J., and Markham, P.G. (1990). Geminivirus coat protein gene replacement alters insect specificity. Virology 177, 85–94.

78  | Sharma et al.

Briddon, R.W., Patil, B.L., Bagewadi, B., Nawaz-ul-Rehman, M.S., and Fauquet, C.M. (2010). Distinct evolutionary histories of the DNA-A and DNA-B components of bipartite begomoviruses. BMC Evol Biol. 10, 97. Brough, C.L., Gardiner, W.E., Inamdar, N.M., Zhang, X.Y., Ehrlich, M., and Bisaro, D.M. (1992). DNA methylation inhibits propagation of Tomato golden mosaic virus DNA in transfected protoplasts. Plant Mol. Biol. 18, 703–712. Bruce, G., Gu, M., Shi, N., Liu, Y., and Hong, Y. (2011). Influence of retinoblastoma-related gene silencing on the initiation of DNA replication by African Cassava mosaic virus Rep in cells of mature leaves in Nicotiana benthamiana plants. Virol. J. 8, 561. https://doi. org/10.1186/1743-422X-8-561 Buchmann, R.C., Asad, S., Wolf, J.N., Mohannath, G., and Bisaro, D.M. (2009). Geminivirus AL2 and L2 proteins suppress transcriptional gene silencing and cause genome-wide reductions in cytosine methylation. J. Virol. 83, 5005–5013. https://doi.org/10.1128/ JVI.01771-08 Butterbach, P., Verlaan, M.G., Dullemans, A., Lohuis, D., Visser, R.G., Bai, Y., and Kormelink, R. (2014). Tomato yellow leaf curl virus resistance by Ty-1 involves increased cytosine methylation of viral genomes and is compromised by Cucumber mosaic virus infection. Proc. Natl. Acad. Sci. U.S.A. 111, 12942–12947. https://doi. org/10.1073/pnas.1400894111 Cao, X., and Jacobsen, S.E. (2002). Role of the Arabidopsis DRM methyltransferases in de novo DNA methylation and gene silencing. Curr. Biol. 12, 1138–1144. Carbonell, A., and Carrington, J.C. (2015). Antiviral roles of plant ARGONAUTES. Curr. Opin. Plant Biol. 27, 111–117. https://doi.org/10.1016/j.pbi.2015.06.013 Castel, S.E., and Martienssen, R.A. (2013). RNA interference in the nucleus: roles for small RNAs in transcription, epigenetics and beyond. Nat. Rev. Genet. 14, 100–112. https://doi.org/10.1038/nrg3355 Castillo, A.G., Collinet, D., Deret, S., Kashoggi, A., and Bejarano, E.R. (2003). Dual interaction of plant PCNA with geminivirus replication accessory protein (Ren) and viral replication protein (Rep). Virology 312, 381–394. Castillo-González, C., Liu, X., Huang, C., Zhao, C., Ma, Z., Hu, T., Sun, F., Zhou, Y., Zhou, X., Wang, X.J., et al. (2015). Geminivirus-encoded TrAP suppressor inhibits the histone methyltransferase SUVH4/KYP to counter host defense. Elife 4, e06671. https://doi.org/10.7554/ eLife.06671 Ceniceros-Ojeda, E.A., Rodríguez-Negrete, E.A., and Rivera-Bustamante, R.F. (2016). Two Populations of Viral Minichromosomes Are Present in a GeminivirusInfected Plant Showing Symptom Remission (Recovery). J. Virol. 90, 3828–3838. https://doi. org/10.1128/JVI.02385-15 Cenik, E.S., and Zamore, P.D. (2011). Argonaute proteins. Curr. Biol. 21, R446–9. https://doi.org/10.1016/j. cub.2011.05.020 Chan, S.W., Henderson, I.R., and Jacobsen, S.E. (2005). Gardening the genome: DNA methylation in Arabidopsis thaliana. Nat. Rev. Genet. 6, 351–360. Chan, S.W., Zilberman, D., Xie, Z., Johansen, L.K., Carrington, J.C., and Jacobsen, S.E. (2004). RNA silencing genes control de novo DNA methylation.

Science 303, 1336. https://doi.org/10.1126/ science.1095989 Charron, J.B., He, H., Elling, A.A., and Deng, X.W. (2009). Dynamic landscapes of four histone modifications during deetiolation in Arabidopsis. Plant Cell 21, 3732– 3748. https://doi.org/10.1105/tpc.109.066845 Chellappan, P., Vanitharani, R., and Fauquet, C.M. (2005). MicroRNA-binding viral protein interferes with Arabidopsis development. Proc. Natl. Acad. Sci. U.S.A. 102, 10381–10386. Chellappan, P., Vanitharani, R., Pita, J., and Fauquet, C.M. (2004). Short interfering RNA accumulation correlates with host recovery in DNA virus-infected hosts, and gene silencing targets specific viral sequences. J. Virol. 78, 7465–7477. https://doi.org/10.1128/JVI.78.14.74657477.2004 Chellappan, P., Xia, J., Zhou, X., Gao, S., Zhang, X., Coutino, G., Vazquez, F., Zhang, W., and Jin, H. (2010). siRNAs from miRNA sites mediate DNA methylation of target genes. Nucleic Acids Res. 38, 6883–6894. https://doi. org/10.1093/nar/gkq590 Chowda-Reddy, R.V., Achenjang, F., Felton, C., Etarock, M.T., Anangfac, M.T., Nugent, P., and Fondong, V.N. (2008). Role of a geminivirus AV2 protein putative protein kinase C motif on subcellular localization and pathogenicity. Virus Res. 135, 115–124. https://doi. org/10.1016/j.virusres.2008.02.014 Cokus, S.J., Feng, S., Zhang, X., Chen, Z., Merriman, B., Haudenschild, C.D., Pradhan, S., Nelson, S.F., Pellegrini, M., and Jacobsen, S.E. (2008). Shotgun bisulphite sequencing of the Arabidopsis genome reveals DNA methylation patterning. Nature 452, 215–219. https:// doi.org/10.1038/nature06745 Dalakouras, A., Dadami, E., and Wassenegger, M. (2013). Viroid-induced DNA methylation in plants. Biomol Concepts 4, 557–565. https://doi.org/10.1515/bmc2013-0030 Deuschle, K., Kepp, G., and Jeske, H. (2016). Differential methylation of the circular DNA in geminiviral minichromosomes. Virology 499, 243–258. Di-Lorenzo, A., and Bedford, M.T. (2011). Histone arginine methylation. FEBS Lett. 585, 2024-2031. Ding, S.W., and Voinnet, O. (2007). Antiviral immunity directed by small RNAs. Cell 130, 413–426. Dogar, A.M. (2006). RNAi dependent epigenetic marks on a geminivirus promoter. Virol. J. 3, 5. Donaire, L., Wang, Y., Gonzalez-Ibeas, D., Mayer, K.F., Aranda, M.A., and Llave, C. (2009). Deep-sequencing of plant viral small RNAs reveals effective and widespread targeting of viral genomes. Virology 392, 203–214. https://doi.org/10.1016/j.virol.2009.07.005 Dry, I.B., Krake, L.R., Rigden, J.E., and Rezaian, M.A. (1997). A novel subviral agent associated with a geminivirus: the first report of a DNA satellite. Proc. Natl. Acad. Sci. U.S.A. 94, 7088–7093. Du, J., Johnson, L.M., Groth, M., Feng, S., Hale, C.J., Li, S., Vashisht, A.A., Wohlschlegel, J.A., Patel, D.J., and Jacobsen, S.E. (2014). Mechanism of DNA methylationdirected histone methylation by Kryptonite. Mol. Cell 55, 495–504. https://doi.org/10.1016/j. molcel.2014.06.009 Duan, C.G., Zhang, H., Tang, K., Zhu, X., Qian, W., Hou, Y.J., Wang, B., Lang, Z., Zhao, Y., Wang, X., et al. (2015).

Surfacing the Role of Epigenetics in Host–Virus Interaction |  79

Specific but interdependent functions for Arabidopsis AGO4 and AGO6 in RNA-directed DNA methylation. EMBO J. 34, 581–592. https://doi.org/10.15252/ embj.201489453 Ermak, G., Paszkowski, U., Wohlmuth, M., Mittelsten Scheid, O., and Paszkowski, J. (1993). Cytosine methylation inhibits replication of African Cassava mosaic virus by two distinct mechanisms. Nucleic Acids Res. 21, 3445–3450. Espinas, N.A., Saze, H., and Saijo, Y. (2016). Epigenetic control of defense signaling and priming in plants. Front. Plant Sci. 7, 1201. https://doi.org/10.3389/ fpls.2016.01201 Fauquet, C.M., Briddon, R.W., Brown, J.K., Moriones, E., Stanley, J., Zerbini, M., and Zhou, X. (2008). Geminivirus strain demarcation and nomenclature. Arch. Virol. 153, 783–821. https://doi.org/10.1007/ s00705-008-0037-6 Fiallo-Olivé, E., Martínez-Zubiaur, Y., Moriones, E., and Navas-Castillo, J. (2012). A novel class of DNA satellites associated with New World begomoviruses. Virology 426, 1–6. https://doi.org/10.1016/j.virol.2012.01.024 Fondong, V.N., Reddy, R.V., Lu, C., Hankoua, B., Felton, C., Czymmek, K., and Achenjang, F. (2007). The consensus N-myristoylation motif of a geminivirus AC4 protein is required for membrane binding and pathogenicity. Mol. Plant Microbe Interact. 20, 380–391. https://doi. org/10.1094/MPMI-20-4-0380 Fontes, E.P., Eagle, P.A., Sipe, P.S., Luckow, V.A., and HanleyBowdoin, L. (1994). Interaction between a geminivirus replication protein and origin DNA is essential for viral replication. J. Biol. Chem. 269, 8459–8465. Fortes, A.M., and Gallusci, P. (2017). Plant stress responses and phenotypic plasticity in the epigenomics era: perspectives on the grapevine scenario, a model for perennial crop plants. front. Plant Sci. 8, 82. https://doi. org/10.3389/fpls.2017.00082 Gafni, Y., and Epel, B.L. (2002). The role of host and viral proteins in intra- and inter-cellular trafficking of geminiviruses. Physiol. Mol. Plant Path. 60, 231-241. Gao, Z., Liu, H.L., Daxinger, L., Pontes, O., He, X., Qian, W., Lin, H., Xie, M., Lorkovic, Z.J., Zhang, S., et al. (2010). An RNA polymerase II- and AGO4-associated protein acts in RNA-directed DNA methylation. Nature 465, 106–109. https://doi.org/10.1038/nature09025 Gehring, M., Reik, M., and Henikoff, S. (2009). DNA demethylation by DNA repair. Trends Genet. 25, 82–90. Gutierrez, C. (2000). DNA replication and cell cycle in plants: learning from geminiviruses. EMBO J. 19, 792–799. https://doi.org/10.1093/emboj/19.5.792 Haag, J.R., and Pikaard, C.S. (2011). Multisubunit RNA polymerases IV and V: purveyors of non-coding RNA for plant gene silencing. Nat. Rev. Mol. Cell Biol. 12, 483–492. https://doi.org/10.1038/nrm3152 Haag, J.R., Ream, T.S., Marasco, M., Nicora, C.D., Norbeck, A.D., Pasa-Tolic, L., and Pikaard, C.S. (2012). In vitro transcription activities of Pol IV, Pol V, and RDR2 reveal coupling of Pol IV and RDR2 for dsRNA synthesis in plant RNA silencing. Mol. Cell 48, 811–818. https:// doi.org/10.1016/j.molcel.2012.09.027 Hagen, C., Rojas, M.R., Kon, T., and Gilbertson, R.L. (2008). Recovery from Cucurbit leaf crumple virus (family Geminiviridae, genus Begomovirus) infection is

an adaptive antiviral response associated with changes in viral small RNAs. Phytopathology 98, 1029–1037. https://doi.org/10.1094/PHYTO-98-9-1029 Hamilton, A., Voinnet, O., Chappell, L., and Baulcombe, D. (2002). Two classes of short interfering RNA in RNA silencing. EMBO J. 21, 4671–4679. Hanley-Bowdoin, L., Bejarano, E.R., Robertson, D., and Mansoor, S. (2013). Geminiviruses: masters at redirecting and reprogramming plant processes. Nat. Rev. Microbiol. 11, 777–788. https://doi.org/10.1038/ nrmicro3117 Hanley-Bowdoin, L., Settlage, S.B., and Robertson, D. (2004). Reprogramming plant gene expression: a prerequisite to geminivirus DNA replication. Mol. Plant Pathol. 5, 149–156. https://doi.org/10.1111/j.13643703.2004.00214.x Hanley-Bowdoin, L., Settlage, S.B., Orozco, B.M., Nagar, S., and Robertson, D. (2000). Geminiviruses: models for plant DNA replication, transcription, and cell cycle regulation. Crit. Rev. Biochem. Mol. Biol. 35, 105–140. Hao, L., Wang, H., Sunter, G., and Bisaro, D.M. (2003). Geminivirus AL2 and L2 proteins interact with and inactivate SNF1 kinase. Plant Cell 15, 1034–1048. Harvey, J.J., Lewsey, M.G., Patel, K., Westwood, J., Heimstädt, S., Carr, J.P., and Baulcombe, D.C. (2011). An antiviral defense role of AGO2 in plants. PLOS ONE 6, e14639. https://doi.org/10.1371/journal.pone.0014639 Havecker, E.R., Wallbridge, L.M., Hardcastle, T.J., Bush, M.S., Kelly, K.A., Dunn, R.M., Schwach, F., Doonan, J.H., and Baulcombe, D.C. (2010). The Arabidopsis RNAdirected DNA methylation argonautes functionally diverge based on their expression and interaction with target loci. Plant Cell 22, 321–334. https://doi. org/10.1105/tpc.109.072199 Hayes, R., MacDonald, H., Coutts, R., and Buck, K. (1988). Priming of complementary DNA synthesis in vitro by small DNA molecules tightly bound to virion DNA of Wheat dwarf virus. J. Gen.Virol. 69, 1345-1350. He, Y. (2012). Chromatin regulation of flowering. Trends Plant Sci. 17, 556–562. https://doi.org/10.1016/j. tplants.2012.05.001 Henderson, I.R., Deleris, A., Wong, W., Zhong, X., Chin, H.G., Horwitz, G.A., Kelly, K.A., Pradhan, S., and Jacobsen, S.E. (2010). The de novo cytosine methyltransferase DRM2 requires intact UBA domains and a catalytically mutated paralog DRM3 during RNAdirected DNA methylation in Arabidopsis thaliana. PLOS Genet. 6, e1001182. https://doi.org/10.1371/ journal.pgen.1001182 Herr, A.J., Jensen, M.B., Dalmay, T., and Baulcombe, D.C. (2005). RNA polymerase IV directs silencing of endogenous DNA. Science 308, 118–120. Ho, T., Wang, H., Pallett, D., and Dalmay, T. (2007). Evidence for targeting common siRNA hotspots and GC preference by plant Dicer-like proteins. FEBS Lett. 581, 3267–3272. Höhnle, M., Höfer, P., Bedford, I.D., Briddon, R.W., Markham, P.G., and Frischmuth, T. (2001). Exchange of three amino acids in the coat protein results in efficient whitefly transmission of a nontransmissible Abutilon mosaic virus isolate. Virology 290, 164–171. Hormuzdi, S.G., and Bisaro, D.M. (1993). Genetic analysis of Beet curly top virus: evidence for three virion sense

80  | Sharma et al.

genes involved in movement and regulation of singleand double-stranded DNA levels. Virology 193, 900–909. Huettel, B., Kanno, T., Daxinger, L., Bucher, E., van der Winden, J., Matzke, A.J., and Matzke, M. (2007). RNA-directed DNA methylation mediated by DRD1 and Pol IVb: a versatile pathway for transcriptional gene silencing in plants. Biochim. Biophys. Acta 1769, 358–374. ICTV Virus Taxonomy. (2014). Virus Taxonomy: 2014 Release. EC 46, Montreal, Canada. (www.ictvonline. org/virustaxonomy.asp). Jackel, J.N., Buchmann, R.C., Singhal, U., and Bisaro, D.M. (2015). Analysis of geminivirus AL2 and L2 proteins reveals a novel AL2 silencing suppressor activity. J. Virol. 89, 3176–3187. https://doi.org/10.1128/JVI.02625-14 Jackel, J.N., Storer, J.M., Coursey, T., and Bisaro, D.M. (2016). Arabidopsis RNA Polymerases IV and V Are Required To Establish H3K9 Methylation, but Not Cytosine Methylation, on Geminivirus Chromatin. J. Virol. 90, 7529–7540. https://doi.org/10.1128/ JVI.00656-16 Jeske, H., Lütgemeier, M., and Preiss, W. (2001). DNA forms indicate rolling circle and recombination-dependent replication of Abutilon mosaic virus. EMBO J. 20, 6158– 6167. https://doi.org/10.1093/emboj/20.21.6158 Ji, L., and Chen, X. (2012). Regulation of small RNA stability: methylation and beyond. Cell Res. 22, 624– 636. https://doi.org/10.1038/cr.2012.36 Jones, L., Ratcliff, F., and Baulcombe, D.C. (2001). RNAdirected transcriptional gene silencing in plants can be inherited independently of the RNA trigger and requires Met1 for maintenance. Curr. Biol. 11, 747–757. Jupin, I., De Kouchkovsky, F., Jouanneau, F., and Gronenborn, B. (1994). Movement of tomato yellow leaf curl geminivirus (TYLCV): involvement of the protein encoded by ORF C4. Virology 204, 82–90. Khraiwesh, B., Arif, M.A., Seumel, G.I., Ossowski, S., Weigel, D., Reski, R., and Frank, W. (2010). Transcriptional control of gene expression by microRNAs. Cell 140, 111–122. https://doi.org/10.1016/j.cell.2009.12.023 Kong, L.J., and Hanley-Bowdoin, L. (2002). A geminivirus replication protein interacts with a protein kinase and a motor protein that display different expression patterns during plant development and infection. Plant Cell 14, 1817–1832. Kong, L.J., Orozco, B.M., Roe, J.L., Nagar, S., Ou, S., Feiler, H.S., Durfee, T., Miller, A.B., Gruissem, W., Robertson, D., et al. (2000). A geminivirus replication protein interacts with the retinoblastoma protein through a novel domain to determine symptoms and tissue specificity of infection in plants. EMBO J. 19, 3485–3495. https:// doi.org/10.1093/emboj/19.13.3485 Kotlizky, G., Boulton, M.I., Pitaksutheepong, C., Davies, J.W., and Epel, B.L. (2000). Intracellular and intercellular movement of maize streak geminivirus V1 and V2 proteins transiently expressed as green fluorescent protein fusions. Virology 274, 32–38. https://doi. org/10.1006/viro.2000.0415 Kovalchuk, I., Kovalchuk, O., Kalck, V., Boyko, V., Filkowski, J., Heinlein, M., and Hohn, B. (2003). Pathogen-induced systemic plant signal triggers DNA rearrangements.

Nature 423, 760–762. https://doi.org/10.1038/ nature01683 Kumar, V., Mishra, S.K., Rahman, J., Taneja, J., Sundaresan, G., Mishra, N.S., and Mukherjee, S.K. (2015). Mungbean yellow mosaic Indian virus encoded AC2 protein suppresses RNA silencing by inhibiting Arabidopsis RDR6 and AGO1 activities. Virology 486, 158–172. https://doi.org/10.1016/j.virol.2015.08.015 Kunik, T., Palanichelvam, K., Czosnek, H., Citovsky, V., and Gafni, Y. (1998). Nuclear import of the capsid protein of Tomato yellow leaf curl virus (TYLCV) in plant and insect cells. Plant J. 13, 393–399. Laufs, J., Schumacher, S., Geisler, N., Jupin, I., and Gronenborn, B. (1995). Identification of the nicking tyrosine of geminivirus Rep protein. FEBS Lett. 377, 258–262. Law, J.A., and Jacobsen, S.E. (2010). Establishing, maintaining and modifying DNA methylation patterns in plants and animals. Nat. Rev. Genet. 11, 204–220. https://doi.org/10.1038/nrg2719 Law, J.A., Ausin, I., Johnson, L.M., Vashisht, A.A., Zhu, J.K., Wohlschlegel, J.A., and Jacobsen, S.E. (2010). A protein complex required for polymerase V transcripts and RNA- directed DNA methylation in Arabidopsis. Curr. Biol. 20, 951–956. https://doi.org/10.1016/j. cub.2010.03.062 Lazarowitz, S.G., Wu, L.C., Rogers, S.G., and Elmer, J.S. (1992). Sequence-specific interaction with the viral AL1 protein identifies a geminivirus DNA replication origin. Plant Cell 4, 799–809. https://doi.org/10.1105/ tpc.4.7.799 Lewsey, M.G., Hardcastle, T.J., Melnyk, C.W., Molnar, A., Valli, A., Urich, M.A., Nery, J.R., Baulcombe, D.C., and Ecker, J.R. (2016). Mobile small RNAs regulate genome-wide DNA methylation. Proc. Natl. Acad. Sci. U.S.A. 113, E801–10. https://doi.org/10.1073/ pnas.1515072113 Li, C.F., Pontes, O., El-Shami, M., Henderson, I.R., Bernatavichute, Y.V., Chan, S.W., Lagrange, T., Pikaard, C.S., and Jacobsen, S.E. (2006). An ARGONAUTE4containing nuclear processing center colocalized with Cajal bodies in Arabidopsis thaliana. Cell 126, 93–106. Li, J., Gorospe, M., Hutter, D., Barnes, J., Keyse, S.M., and Liu, Y. (2001). Transcriptional induction of MKP-1 in response to stress is associated with histone H3 phosphorylation-acetylation. Mol. Cell. Biol. 21, 8213–8224. https://doi.org/10.1128/ MCB.21.23.8213-8224.2001 Li, J., Yang, Z., Yu, B., Liu, J., and Chen, X. (2005). Methylation protects miRNAs and siRNAs from a 3′-end uridylation activity in Arabidopsis. Curr. Biol. 15, 1501–1507. Li, S., Vandivier, L.E., Tu, B., Gao, L., Won, S.Y., Li, S., Zheng, B., Gregory, B.D., and Chen, X. (2015). Detection of Pol IV/RDR2-dependent transcripts at the genomic scale in Arabidopsis reveals features and regulation of siRNA biogenesis. Genome Res. 25, 235–245. https://doi. org/10.1101/gr.182238.114 Lindroth, A.M., Cao, X., Jackson, J.P., Zilberman, D., McCallum, C.M., Henikoff, S., and Jacobsen, S.E. (2001). Requirement of CHROMOMETHYLASE3 for maintenance of CpXpG methylation. Science 292, 2077–2080. https://doi.org/10.1126/science.1059745

Surfacing the Role of Epigenetics in Host–Virus Interaction |  81

Lister, R., O’Malley, R.C., Tonti-Filippini, J., Gregory, B.D., Berry, C.C., Millar, A.H., and Ecker, J.R. (2008). Highly integrated single-base resolution maps of the epigenome in Arabidopsis. Cell 133, 523–536. https:// doi.org/10.1016/j.cell.2008.03.029 Liu, C., Lu, F., Cui, X., and Cao, X. (2010). Histone methylation in higher plants. Annu. Rev. Plant Biol. 61, 395–420. https://doi.org/10.1146/annurev. arplant.043008.091939 Liu, H., Boulton, M.I., and Davies, J.W. (1997). Maize streak virus coat protein binds single- and double-stranded DNA in vitro. J. Gen. Virol. 78, 1265–1270. https://doi. org/10.1099/0022-1317-78-6-1265 Liu, H., Boulton, M.I., Thomas, C.L., Prior, D.A., Oparka, K.J., and Davies, J.W. (1999). Maize streak virus coat protein is karyophyllic and facilitates nuclear transport of viral DNA. Mol. Plant Microbe Interact. 12, 894–900. https://doi.org/10.1094/MPMI.1999.12.10.894 Liu, L., Chung, H.Y., Lacatus, G., Baliji, S., Ruan, J., and Sunter, G. (2014). Altered expression of Arabidopsis genes in response to a multifunctional geminivirus pathogenicity protein. BMC Plant Biol. 14, 302. https:// doi.org/10.1186/s12870-014-0302-7 Liu, Z.W., Shao, C.R., Zhang, C.J., Zhou, J.X., Zhang, S.W., Li, L., Chen, S., Huang, H.W., Cai, T., and He, X.J. (2014). The SET domain proteins SUVH2 and SUVH9 are required for Pol V occupancy at RNA-directed DNA methylation loci. PLOS Genet. 10, e1003948. https:// doi.org/10.1371/journal.pgen.1003948 Llave, C. (2010). Virus-derived small interfering RNAs at the core of plant-virus interactions. Trends Plant Sci. 15, 701–707. https://doi.org/10.1016/j. tplants.2010.09.001 Lorković, Z.J., Naumann, U., Matzke, A.J., and Matzke, M. (2012). Involvement of a GHKL ATPase in RNAdirected DNA methylation in Arabidopsis thaliana. Curr. Biol. 22, 933–938. https://doi.org/10.1016/j. cub.2012.03.061 Lozano-Durán, R., Rosas-Díaz, T., Luna, A.P., and Bejarano, E.R. (2011). Identification of host genes involved in geminivirus infection using a reverse genetics approach. PLOS ONE 6, e22383. https://doi.org/10.1371/ journal.pone.0022383 Ma, Z., Coruh, C., and Axtell, M.J. (2010). Arabidopsis lyrata small RNAs: transient MIRNA and small interfering RNA loci within the Arabidopsis genus. Plant Cell 22, 1090–1103. https://doi.org/10.1105/tpc.110.073882 Marí-Ordóñez, A., Marchais, A., Etcheverry, M., Martin, A., Colot, V., and Voinnet, O. (2013). Reconstructing de novo silencing of an active plant retrotransposon. Nat. Genet. 45, 1029–1039. https://doi.org/10.1038/ ng.2703 Mason, G., Noris, E., Lanteri, S., Acquadro, A., Accotto, G.P., and Portis, E. (2008). Potentiality of methylationsensitive amplification polymorphism (MSAP) in identifying genes involved in tomato response to Tomato yellow leaf curl Sardinia virus. Plant Mol. Biol. Rep. 26, 156-173. Matzke, M.A., and Mosher, R.A. (2014). RNA-directed DNA methylation: an epigenetic pathway of increasing complexity. Nat. Rev. Genet. 15, 394–408. https://doi. org/10.1038/nrg3683

Matzke, M.A., Primig, M., Trnovsky, J., and Matzke, A.J. (1989). Reversible methylation and inactivation of marker genes in sequentially transformed tobacco plants. EMBO J. 8, 643–649. McCue, A.D., Panda, K., Nuthikattu, S., Choudury, S.G., Thomas, E.N., and Slotkin, R.K. (2015). ARGONAUTE 6 bridges transposable element mRNA-derived siRNAs to the establishment of DNA methylation. EMBO J. 34, 20–35. https://doi.org/10.15252/embj.201489499 Melquist, S., and Bender, J. (2003). Transcription from an upstream promoter controls methylation signalling from an inverted repeat of endogenous genes in Arabidopsis. Genes Dev. 17, 2036-2047. Métivier, R., Gallais, R., Tiffoche, C., Le Péron, C., Jurkowska, R.Z., Carmouche, R.P., Ibberson, D., Barath, P., Demay, F., Reid, G., et al. (2008). Cyclical DNA methylation of a transcriptionally active promoter. Nature 452, 45–50. https://doi.org/10.1038/nature06544 Mette, M.F., Aufsatz, W., van der Winden, J., Matzke, M.A., and Matzke, A.J. (2000). Transcriptional silencing and promoter methylation triggered by doublestranded RNA. EMBO J. 19, 5194–5201. https://doi. org/10.1093/emboj/19.19.5194 Mlotshwa, S., Voinnet, O., Mette, M.F., Matzke, M., Vaucheret, H., Ding, S.W., Pruss, G., and Vance, V.B. (2002). RNA silencing and the mobile silencing signal. Plant Cell 14, S289–301. Moissiard, G., and Voinnet, O. (2006). RNA silencing of host transcripts by cauliflower mosaic virus requires coordinated action of the four Arabidopsis Dicer-like proteins. Proc. Natl. Acad. Sci. U.S.A. 103, 19593–19598. Molnár, A., Csorba, T., Lakatos, L., Várallyay, E., Lacomme, C., and Burgyán, J. (2005). Plant virus-derived small interfering RNAs originate predominantly from highly structured single-stranded viral RNAs. J. Virol. 79, 7812–7818. Morilla, G., Castillo, A.G., Preiss, W., Jeske, H., and Bejarano, E.R. (2006). A versatile transreplication-based system to identify cellular proteins involved in geminivirus replication. J. Virol. 80, 3624–3633. Muthamilarasan, M., and Prasad, M. (2013). Plant innate immunity: an updated insight into defense mechanism. J. Biosci. 38, 433–449. Noreen, F., Akbergenov, R., Hohn, T., and Richert-Pöggeler, K.R. (2007). Distinct expression of endogenous Petunia vein clearing virus and the DNA transposon dTph1 in two Petunia hybrida lines is correlated with differences in histone modification and siRNA production. Plant J. 50, 219–229. Noris, E., Vaira, A.M., Caciagli, P., Masenga, V., Gronenborn, B., and Accotto, G.P. (1998). Amino acids in the capsid protein of Tomato yellow leaf curl virus that are crucial for systemic infection, particle formation, and insect transmission. J. Virol. 72, 10050–10057. Nuthikattu, S., McCue, A.D., Panda, K., Fultz, D., DeFraia, C., Thomas, E.N., and Slotkin, R.K. (2013). The initiation of epigenetic silencing of active transposable elements is triggered by RDR6 and 21-22 nucleotide small interfering RNAs. Plant Physiol. 162, 116–131. https://doi.org/10.1104/pp.113.216481 Offermann, S., Dreesen, B., Horst, I., Danker, T., Jaskiewicz, M., and Peterhansel, C. (2008). Developmental and environmental signals induce distinct histone acetylation

82  | Sharma et al.

profiles on distal and proximal promoter elements of the C4-Pepc gene in maize. Genetics 179, 1891–1901. https://doi.org/10.1534/genetics.108.087411 Olmedo-Monfil, V., Durán-Figueroa, N., Arteaga-Vázquez, M., Demesa-Arévalo, E., Autran, D., Grimanelli, D., Slotkin, R.K., Martienssen, R.A., and Vielle-Calzada, J.P. (2010). Control of female gamete formation by a small RNA pathway in Arabidopsis. Nature 464, 628–632. https://doi.org/10.1038/nature08828 Orozco, B.M., Miller, A.B., Settlage, S.B., and HanleyBowdoin, L. (1997). Functional domains of a geminivirus replication protein. J. Biol. Chem. 272, 9840–9846. Padidam, M., Beachy, R.N., and Fauquet, C.M. (1996). The role of AV2 (‘precoat’) and coat protein in viral replication and movement in tomato leaf curl geminivirus. Virology 224, 390–404. Padidam, M., Sawyer, S., and Fauquet, C.M. (1999). Possible emergence of new geminiviruses by frequent recombination. Virology 265, 218–225. https://doi. org/10.1006/viro.1999.0056 Padmanabhan, C., Zhang, X., and Jin, H. (2009). Host small RNAs are big contributors to plant innate immunity. Curr. Opin. Plant Biol. 12, 465–472. https://doi. org/10.1016/j.pbi.2009.06.005 Palanichelvam, K., Kunik, T., Citovsky, V., and Gafni, Y. (1998). The capsid protein of Tomato yellow leaf curl virus binds cooperatively to single-stranded DNA. J. Gen. Virol. 79, 2829–2833. https://doi.org/10.1099/00221317-79-11-2829 Panda, K., and Slotkin, R.K. (2013). Proposed mechanism for the initiation of transposable element silencing by the RDR6-directed DNA methylation pathway. Plant Signal Behav 8, e25206. https://doi.org/10.4161/psb.25206 Pantaleo, V. (2011). Plant RNA silencing in viral defence. Adv. Exp. Med. Biol. 722, 39–58. https://doi. org/10.1007/978-1-4614-0332-6_3 Paprotka, T., Deuschle, K., Metzler, V., and Jeske, H. (2011). Conformation-selective methylation of geminivirus DNA. J. Virol. 85, 12001–12012. https://doi. org/10.1128/JVI.05567-11 Pasumarthy, K.K., Mukherjee, S.K., and Choudhury, N.R. (2011). The presence of Tomato leaf curl Kerala virus AC3 protein enhances viral DNA replication and modulates virus induced gene-silencing mechanism in tomato plants. Virol. J. 8, 178. https://doi.org/10.1186/1743422X-8-178 Patil, B.L., and Fauquet, C.M. (2015). Studies on differential behavior of cassava mosaic geminivirus DNA components, symptom recovery patterns, and their siRNA profiles. Virus Genes 50, 474–486. https://doi. org/10.1007/s11262-015-1184-y Pélissier, T., and Wassenegger, M. (2000). A DNA target of 30 bp is sufficient for RNA-directed DNA methylation. RNA 6, 55–65. Pélissier, T., Thalmeir, S., Kempe, D., Sänger, H.L., and Wassenegger, M. (1999). Heavy de novo methylation at symmetrical and non-symmetrical sites is a hallmark of RNA-directed DNA methylation. Nucleic Acids Res. 27, 1625–1634. Pérez-Quintero, A.L., Neme, R., Zapata, A., and López, C. (2010). Plant microRNAs and their role in defense against viruses: a bioinformatics approach. BMC Plant

Biol. 10, 138. https://doi.org/10.1186/1471-2229-10138 Pilartz, M., and Jeske, H. (1992). Abutilon mosaic geminivirus double-stranded DNA is packed into minichromosomes. Virology 189, 800–802. Pilartz, M., and Jeske, H. (2003). Mapping of abutilon mosaic geminivirus minichromosomes. J. Virol. 77, 10808–10818. Pontier, D., Picart, C., Roudier, F., Garcia, D., Lahmy, S., Azevedo, J., Alart, E., Laudié, M., Karlowski, W.M., Cooke, R., et al. (2012). NERD, a plant-specific GW protein, defines an additional RNAi-dependent chromatin-based pathway in Arabidopsis. Mol. Cell 48, 121–132. https:// doi.org/10.1016/j.molcel.2012.07.027 Pontier, D., Yahubyan, G., Vega, D., Bulski, A., SaezVasquez, J., Hakimi, M.A., Lerbs-Mache, S., Colot, V., and Lagrange, T. (2005). Reinforcement of silencing at transposons and highly repeated sequences requires the concerted action of two distinct RNA polymerases IV in Arabidopsis. Genes Dev. 19, 2030–2040. Pooggin, M.M. (2013). How can plant DNA viruses evade siRNA-directed DNA methylation and silencing? Int. J. Mol. Sci. 14, 15233–15259. https://doi.org/10.3390/ ijms140815233 Preiss, W., and Jeske, H. (2003). Multitasking in replication is common among geminiviruses. J. Virol. 77, 2972– 2980. Qi, X., Bao, F.S., and Xie, Z. (2009). Small RNA deep sequencing reveals role for Arabidopsis thaliana RNAdependent RNA polymerases in viral siRNA biogenesis. PLOS ONE 4, e4971. https://doi.org/10.1371/journal. pone.0004971 Qu, F., Ye, X., and Morris, T.J. (2008). Arabidopsis DRB4, AGO1, AGO7, and RDR6 participate in a DCL4-initiated antiviral RNA silencing pathway negatively regulated by DCL1. Proc. Natl. Acad. Sci. U.S.A. 105, 14732–14737. https://doi.org/10.1073/pnas.0805760105 Raja, P., Jackel, J.N., Li, S., Heard, I.M., and Bisaro, D.M. (2014). Arabidopsis double-stranded RNA binding protein DRB3 participates in methylation-mediated defense against geminiviruses. J. Virol. 88, 2611–2622. https://doi.org/10.1128/JVI.02305-13 Raja, P., Sanville, B.C., Buchmann, R.C., and Bisaro, D.M. (2008). Viral genome methylation as an epigenetic defense against geminiviruses. J. Virol. 82, 8997–9007. https://doi.org/10.1128/JVI.00719-08 Raja, P., Wolf, J.N., and Bisaro, D.M. (2010). RNA silencing directed against geminiviruses: post-transcriptional and epigenetic components. Biochim. Biophys. Acta 1799, 337–351. https://doi.org/10.1016/j. bbagrm.2010.01.004 Rajeswaran, R., Seguin, J., Chabannes, M., Duroy, P.O., Laboureau, N., Farinelli, L., Iskra-Caruana, M.L., and Pooggin, M.M. (2014). Evasion of short interfering RNA-directed antiviral silencing in Musa acuminata persistently infected with six distinct Banana streak pararetroviruses. J. Virol. 88, 11516–11528. https://doi. org/10.1128/JVI.01496-14 Ramachandran, V., and Chen, X. (2008a). Degradation of microRNAs by a family of exoribonucleases in Arabidopsis. Science 321, 1490–1492. https://doi. org/10.1126/science.1163728

Surfacing the Role of Epigenetics in Host–Virus Interaction |  83

Ramachandran, V., and Chen, X. (2008b). Small RNA metabolism in Arabidopsis. Trends Plant Sci. 13, 368– 374. https://doi.org/10.1016/j.tplants.2008.03.008 Ren, G., Xie, M., Dou, Y., Zhang, S., Zhang, C., and Yu, B. (2012). Regulation of miRNA abundance by RNA binding protein TOUGH in Arabidopsis. Proc. Natl. Acad. Sci. U.S.A. 109, 12817–12821. https://doi. org/10.1073/pnas.1204915109 Rodríguez-Negrete, E., Lozano-Durán, R., Piedra-Aguilera, A., Cruzado, L., Bejarano, E.R., and Castillo, A.G. (2013). Geminivirus Rep protein interferes with the plant DNA methylation machinery and suppresses transcriptional gene silencing. New Phytol. 199, 464–475. https://doi. org/10.1111/nph.12286 Rodríguez-Negrete, E.A., Carrillo-Tripp, J., and Rivera-Bustamante, R.F. (2009). RNA silencing against geminivirus: complementary action of posttranscriptional gene silencing and transcriptional gene silencing in host recovery. J. Virol. 83, 1332–1340. https://doi.org/10.1128/JVI.01474-08 Rojas, M.R., and Gilbertson, R.L. (2008). Emerging plant viruses: a diversity of mechanisms and opportunities. In: Roossinck MJ, eds. Plant Virus Evolution. SpringerVerlag, Berlin, pp. 27–51. Rojas, M.R., Hagen, C., Lucas, W.J., and Gilbertson, R.L. (2005). Exploiting chinks in the plant’s armor: evolution and emergence of geminiviruses. Annu. Rev. Phytopathol. 43, 361–394. https://doi.org/10.1146/ annurev.phyto.43.040204.135939 Rowley, M.J., Avrutsky, M.I., Sifuentes, C.J., Pereira, L., and Wierzbicki, A.T. (2011). Independent chromatin binding of ARGONAUTE4 and SPT5L/KTF1 mediates transcriptional gene silencing. PLOS Genet. 7, e1002120. https://doi.org/10.1371/journal. pgen.1002120 Ruiz-Ferrer, V., and Voinnet, O. (2009). Roles of plant small RNAs in biotic stress responses. Annu. Rev. Plant Biol. 60, 485–510. https://doi.org/10.1146/annurev. arplant.043008.092111 Rybicki, E.P. (1994). A phylogenetic and evolutionary justification for three genera of Geminiviridae. Arch. Virol. 139, 49–77. Sahu, P.P., Pandey, G., Sharma, N., Puranik, S., Muthamilarasan, M., and Prasad, M. (2013). Epigenetic mechanisms of plant stress responses and adaptation. Plant Cell Rep. 32, 1151–1159. https://doi. org/10.1007/s00299-013-1462-x Sahu, P.P., Rai, N.K., Chakraborty, S., Singh, M., Chandrappa, P.H., Ramesh, B., Chattopadhyay, D., and Prasad, M. (2010). Tomato cultivar tolerant to Tomato leaf curl New Delhi virus infection induces virus-specific short interfering RNA accumulation and defence-associated host gene expression. Mol. Plant Pathol. 11, 531–544. https://doi.org/10.1111/j.1364-3703.2010.00630.x Sahu, P.P., Sharma, N., Puranik, S., and Prasad, M. (2014b). Post-transcriptional and Epigenetic Arms of RNA Silencing: A Defense Machinery of Naturally Tolerant Tomato Plant Against Tomato leaf curl New Delhi virus. Plant Mol. Biol. Rep. 32, 1015. Sahu, P.P., Sharma, N., Puranik, S., Muthamilarasan, M., and Prasad, M. (2014a). Involvement of host regulatory pathways during geminivirus infection: a novel platform for generating durable resistance. Funct. Integr.

Genomics 14, 47–58. https://doi.org/10.1007/s10142013-0346-z Settlage, S.B., See, R.G., and Hanley-Bowdoin, L. (2005). Geminivirus C3 protein: replication enhancement and protein interactions. J. Virol. 79, 9885–9895. Sharma, N., Sahu, P.P., Puranik, S., and Prasad, M. (2013). Recent advances in plant-virus interaction with emphasis on small interfering RNAs (siRNAs). Mol. Biotechnol. 55, 63–77. https://doi.org/10.1007/ s12033-012-9615-7 Shivaprasad, P.V., Akbergenov, R., Trinks, D., Rajeswaran, R., Veluthambi, K., Hohn, T., and Pooggin, M.M. (2005). Promoters, transcripts, and regulatory proteins of Mungbean yellow mosaic geminivirus. J. Virol. 79, 8149–8163. Singh, D.K., Islam, M.N., Choudhury, N.R., Karjee, S., and Mukherjee, S.K. (2007). The 32 kDa subunit of replication protein A (RPA) participates in the DNA replication of Mung bean yellow mosaic India virus (MYMIV) by interacting with the viral Rep protein. Nucleic Acids Res. 35, 755–770. Song, J.J., Smith, S.K., Hannon, G.J., and Joshua-Tor, L. (2004). Crystal structure of Argonaute and its implications for RISC slicer activity. Science 305, 1434–1437. https://doi.org/10.1126/science.1102514 Stanley, J., Latham, J.R., Pinner, M.S., Bedford, I., and Markham, P.G. (1992). Mutational analysis of the monopartite geminivirus Beet curly top virus. Virology 191, 396–405. Steward, N., Ito, M., Yamaguchi, Y., Koizumi, N., and Sano, H. (2002). Periodic DNA methylation in maize nucleosomes and demethylation by environmental stress. J. Biol. Chem. 277, 37741–37746. https://doi. org/10.1074/jbc.M204050200 Stroud, H., Do, T., Du, J., Zhong, X., Feng, S., Johnson, L., Patel, D.J., and Jacobsen, S.E. (2014). Non-CG methylation patterns shape the epigenetic landscape in Arabidopsis. Nat. Struct. Mol. Biol. 21, 64–72. https:// doi.org/10.1038/nsmb.2735 Stroud, H., Greenberg, M.V., Feng, S., Bernatavichute, Y.V., and Jacobsen, S.E. (2013). Comprehensive analysis of silencing mutants reveals complex regulation of the Arabidopsis methylome. Cell 152, 352–364. https://doi. org/10.1016/j.cell.2012.10.054 Sun, Y.W., Tee, C.S., Ma, Y.H., Wang, G., Yao, X.M., and Ye, J. (2015). Attenuation of Histone Methyltransferase KRYPTONITE-mediated transcriptional gene silencing by Geminivirus. Sci Rep 5, 16476. https://doi. org/10.1038/srep16476 Sunter, G., Sunter, J.L., and Bisaro, D.M. (2001). Plants expressing Tomato golden mosaic virus AL2 or Beet curly top virus L2 transgenes show enhanced susceptibility to infection by DNA and RNA viruses. Virology 285, 59–70. https://doi.org/10.1006/viro.2001.0950 Szittya, G., and Burgyán, J. (2013). RNA interferencemediated intrinsic antiviral immunity in plants. Curr. Top. Microbiol. Immunol. 371, 153–181. https://doi. org/10.1007/978-3-642-37765-5_6 Teng, K., Chen, H., Lai, J., Zhang, Z., Fang, Y., Xia, R., Zhou, X., Guo, H., and Xie, Q. (2010). Involvement of C4 protein of Beet severe curly top virus (family Geminiviridae) in virus movement. PLOS ONE 5, e11280. https://doi. org/10.1371/journal.pone.0011280

84  | Sharma et al.

Thorstensen, T., Grini, P.E., and Aalen, R.B. (2011). SET domain proteins in plant development. Biochim. Biophys. Acta 1809, 407–420. https://doi. org/10.1016/j.bbagrm.2011.05.008 Tolia, N.H., and Joshua-Tor, L. (2007). Slicer and the argonautes. Nat. Chem. Biol. 3, 36–43. Tompa, R., McCallum, C.M., Delrow, J., Henikoff, J.G., van Steensel, B., and Henikoff, S. (2002). Genome-wide profiling of DNA methylation reveals transposon targets of CHROMOMETHYLASE3. Curr. Biol. 12, 65–68. Torchetti, E.M., Pegoraro, M., Navarro, B., Catoni, M., Di Serio, F., and Noris, E. (2016). A nuclear-replicating viroid antagonizes infectivity and accumulation of a geminivirus by upregulating methylation-related genes and inducing hypermethylation of viral DNA. Sci Rep 6, 35101. https://doi.org/10.1038/srep35101 Trinks, D., Rajeswaran, R., Shivaprasad, P.V., Akbergenov, R., Oakeley, E.J., Veluthambi, K., Hohn, T., and Pooggin, M.M. (2005). Suppression of RNA silencing by a geminivirus nuclear protein, AC2, correlates with transactivation of host genes. J. Virol. 79, 2517–2527. Vanitharani, R., Chellappan, P., and Fauquet, C.M. (2005). Geminiviruses and RNA silencing. Trends Plant Sci. 10, 144–151. Vaucheret, H. (2008). Plant ARGONAUTES. Trends Plant Sci. 13, 350–358. Voinnet, O., Pinto, Y.M., and Baulcombe, D.C. (1999). Suppression of gene silencing: a general strategy used by diverse DNA and RNA viruses of plants. Proc. Natl. Acad. Sci. U.S.A. 96, 14147–14152. Vongs, A., Kakutani, T., Martienssen, R.A., and Richards, E.J. (1993). Arabidopsis thaliana DNA methylation mutants. Science 260, 1926–1928. Waddington, C.H. (1956). The genetic assimilation of the bithorax phenotype. Evolution. 10, 1–13. Wang, B., Li, F., Huang, C., Yang, X., Qian, Y., Xie, Y., and Zhou, X. (2014). V2 of Tomato yellow leaf curl virus can suppress methylation-mediated transcriptional gene silencing in plants. J. Gen. Virol. 95, 225-230. Wang, H., Hao, L., Shung, C.Y., Sunter, G., and Bisaro, D.M. (2003). Adenosine kinase is inactivated by geminivirus AL2 and L2 proteins. Plant Cell 15, 3020–3032. https:// doi.org/10.1105/tpc.015180 Wang, X.B., Jovel, J., Udomporn, P., Wang, Y., Wu, Q., Li, W.X., Gasciolli, V., Vaucheret, H., and Ding, S.W. (2011). The 21-nucleotide, but not 22-nucleotide, viral secondary small interfering RNAs direct potent antiviral defense by two cooperative argonautes in Arabidopsis thaliana. Plant Cell. 23, 1625–1638. Wassenegger, M., Heimes, S., Riedel, L., and Sänger, H.L. (1994). RNA-directed de novo methylation of genomic sequences in plants. Cell 76, 567–576. Waterhouse, P.M., Wang, M.B., and Lough, T. (2001). Gene silencing as an adaptive defence against viruses. Nature 411, 834–842. https://doi.org/10.1038/35081168 Wendte, J.M., and Pikaard, C.S. (2017). The RNAs of RNAdirected DNA methylation. Biochim. Biophys. Acta 1860, 140–148. Wierzbicki, A.T., Haag, J.R., and Pikaard, C.S. (2008). Noncoding transcription by RNA polymerase Pol IVb/ Pol V mediates transcriptional silencing of overlapping and adjacent genes. Cell 135, 635–648. https://doi. org/10.1016/j.cell.2008.09.035

Wierzbicki, A.T., Ream, T.S., Haag, J.R., and Pikaard, C.S. (2009). RNA polymerase V transcription guides ARGONAUTE4 to chromatin. Nat. Genet. 41, 630– 634. https://doi.org/10.1038/ng.365 Wu, L., Zhou, H., Zhang, Q., Zhang, J., Ni, F., Liu, C., and Qi, Y. (2010). DNA methylation mediated by a microRNA pathway. Mol. Cell 38, 465–475. https://doi. org/10.1016/j.molcel.2010.03.008 Xie, M., and Yu, B. (2015). siRNA-directed DNA Methylation in Plants. Curr. Genomics 16, 23–31. https://doi.org/10.2174/1389202915666141128002 211 Xie, Z., Johansen, L.K., Gustafson, A.M., Kasschau, K.D., Lellis, A.D., Zilberman, D., Jacobsen, S.E., and Carrington, J.C. (2004). Genetic and functional diversification of small RNA pathways in plants. PLOS Biol. 2, E104. https://doi.org/10.1371/journal. pbio.0020104 Yadav, R.K., and Chattopadhyay, D. (2011). Enhanced viral intergenic region specific siRNA accumulation and DNA methylation correlates with resistance to a geminivirus. Mol. Plant Microbe Interact. 24, 1189-1197. Yang, L.P., Fang, Y.Y., An, C.P., Dong, L., Zhang, Z.H., Chen, H., Xie, Q., and Guo, H.S. (2013). C2-mediated decrease in DNA methylation, accumulation of siRNAs, and increase in expression for genes involved in defense pathways in plants infected with Beet severe curly top virus. Plant J. 73, 910–917. https://doi.org/10.1111/ tpj.12081 Yang, X., Xie, Y., Raja, P., Li, S., Wolf, J.N., Shen, Q., Bisaro, D.M., and Zhou, X. (2011). Suppression of methylationmediated transcriptional gene silencing by βC1-SAHH protein interaction during geminivirus-betasatellite infection. PLOS Pathog. 7, e1002329. https://doi. org/10.1371/journal.ppat.1002329 Ye, R., Wang, W., Iki, T., Liu, C., Wu, Y., Ishikawa, M., Zhou, X., and Qi, Y. (2012). Cytoplasmic assembly and selective nuclear import of Arabidopsis Argonaute4/ siRNA complexes. Mol. Cell 46, 859–870. https://doi. org/10.1016/j.molcel.2012.04.013 Yu, B., Bi, L., Zheng, B., Ji, L., Chevalier, D., Agarwal, M., Ramachandran, V., Li, W., Lagrange, T., Walker, J.C., et al. (2008). The FHA domain proteins DAWDLE in Arabidopsis and SNIP1 in humans act in small RNA biogenesis. Proc. Natl. Acad. Sci. U.S.A. 105, 10073– 10078. https://doi.org/10.1073/pnas.0804218105 Yu, B., Yang, Z., Li, J., Minakhina, S., Yang, M., Padgett, R.W., Steward, R., and Chen, X. (2005). Methylation as a crucial step in plant microRNA biogenesis. Science 307, 932–935. Zhang, C., Wu, Z., Li, Y., and Wu, J. (2015). Biogenesis, Function, and Applications of Virus-Derived Small RNAs in Plants. Front. Microbiol 6, 1237. https://doi. org/10.3389/fmicb.2015.01237 Zhang, S., Liu, Y., and Yu, B. (2014). PRL1, an RNAbinding protein, positively regulates the accumulation of miRNAs and siRNAs in Arabidopsis. PLOS Genet. 10, e1004841. https://doi.org/10.1371/journal. pgen.1004841 Zhang, S., Xie, M., Ren, G., and Yu, B. (2013). CDC5, a DNA binding protein, positively regulates posttranscriptional processing and/or transcription of primary microRNA transcripts. Proc. Natl. Acad. Sci.

Surfacing the Role of Epigenetics in Host–Virus Interaction |  85

U.S.A. 110, 17588–17593. https://doi.org/10.1073/ pnas.1310644110 Zhang, Z., Chen, H., Huang, X., Xia, R., Zhao, Q., Lai, J., Teng, K., Li, Y., Liang, L., Du, Q., et al. (2011). BSCTV C2 attenuates the degradation of SAMDC1 to suppress DNA methylation-mediated gene silencing in Arabidopsis. Plant Cell 23, 273–288. https://doi. org/10.1105/tpc.110.081695 Zheng, B., and Chen, X. (2011). Dynamics of histone H3 lysine 27 trimethylation in plant development. Curr. Opin. Plant Biol. 14, 123–129. https://doi. org/10.1016/j.pbi.2011.01.001 Zheng, B., Wang, Z., Li, S., Yu, B., Liu, J.Y., and Chen, X. (2009). Intergenic transcription by RNA polymerase II coordinates Pol IV and Pol V in siRNA-directed transcriptional gene silencing in Arabidopsis. Genes Dev. 23, 2850–2860. https://doi.org/10.1101/gad.1868009

Zhong, X., Du, J., Hale, C.J., Gallego-Bartolome, J., Feng, S., Vashisht, A.A., Chory, J., Wohlschlegel, J.A., Patel, D.J., and Jacobsen, S.E. (2014). Molecular mechanism of action of plant DRM de novo DNA methyltransferases. Cell 157, 1050–1060. https://doi.org/10.1016/j. cell.2014.03.056 Zilberman, D., Cao, X., and Jacobsen, S.E. (2003). ARGONAUTE4 control of locus-specific siRNA accumulation and DNA and histone methylation. Science 299, 716–719. https://doi.org/10.1126/ science.1079695 Zvereva, A.S., and Pooggin, M.M. (2012). Silencing and innate immunity in plant defense against viral and nonviral pathogens. Viruses 4, 2578–2597. https://doi. org/10.3390/v4112578

Molecular Markers as Tools for Identification and Introgression of VirusResistant Genes

4

Mamta Sharma*, Avijit Tarafdar, U. S. Sharath Chandran, Devashish R. Chobe and Raju Ghosh

International Crops Research Institute for the Semi-Arid Tropics (ICRISAT), Hyderabad, Telengana, India. *Correspondence: [email protected] https://doi.org/10.21775/9781910190814.04

Abstract A majority of the plant viral diseases are spread by insect vectors. Control of vectors using chemicals is a common practice for management of viral diseases. Although, to prevail such situations is still impractical as high recurring costs of the pesticides. Emergence of insecticide resistant insect populations, human anxiety regarding pesticide residue and its side effects also are other concern. Hence development of virus-resistant crop plants is the need of future. In order to accelerate the current scenario of virus resistance breeding, molecular markers could function as a key tool to help and skip several generations of crossing and analysis altogether, thereby saving precious time. The researchers associated with the science of plant breeding and plant pathology has discovered reliable and rapid diagnosis techniques for many viral diseases using different molecular markers. Screening of viral disease resistance lines by marker-assisted selection (MAS) is most common technique, because phenotypic selection of virus-resistant lines is always not convenient. It has a immense importance to come across the mandate of resistance breeding, hence markers like SCAR, RFLP, RAPD, SSR, AFLP, TRAP and CAPS known for their powerful genetic association could be used to supply complementary information to the classical genetic analyses. In resistance breeding programme gene tagging of particular

trait is a necessary object for map based gene cloning and MAS. But mapping of genetic linkage is a time consuming procedure. In recent years, it is possible to identify markers directly without drawing any genetic linkage map. Markers based on nucleotide-binding and leucine-rich repeat domain (NB-LRR) is an advanced tool for identification of disease resistant genes. Markers based on molecular genetics can be used to determine the monogenic or polygenic disease resistance in crops which could provide durable resistance to wide range of pathogens. The gene pyramiding technique in crops is the best way for developing durable resistance (multigenic resistance) against multiple diseases. Based on molecular markers, identified segregating population in crops with viral disease resistance is needed to be confirmed against challenged viral inoculums either in controlled environment or in natural field conditions. Introduction Developing diagnostic tests for viral pathogens often pose great difficulty and challenges in mass multiplication of inoculum and its preservation. Identification of several races and pathotypes of viral infectious agents of plant is still more complex. On the other hand, resistance selection for disease in adult plant is difficult as well as very expensive also. This is where the use of markers comes into

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play. Since last three decades, for genetic analysis molecular markers are used as an essential tool and being used to decode genetic information. To understand the genetic assortment, markers are categorized into three groups’, namely molecular (genotypic), biochemical and morphological (phenotypic), where the first one is based on DNA polymorphism. Molecular markers are generally of two types, i.e. PCR based, such as amplified fragment length polymorphism (AFLP) (Vos et al., 1995), sequence characterized amplified regions (SCARs) (Michelmore et al., 1991) and random amplified polymorphic DNA (RAPD) (Williams et al., 1990) etc. and nucleotide (DNA and RNA) hybridization based such as restriction fragment length polymorphism (RFLP) (Botstein et al., 1980). Genetic markers are now employed for various resistance breeding programmes. The basic concept behind marker-assisted selection (MAS) is the possibility to locate the gene associated with the marker. The probable chances of detecting a disease resistance gene are inversely proportional to the genetic distance between the gene and the marker. A larger population from several crosses is required for better estimation of the genetic distance between the gene and the marker, as the genetic distance varies greatly between different crosses (Messeguer et al., 1991). Over the years, many quick and reliable detection techniques have been developed for many viral diseases by suitably making use of molecular markers. For example, linkage between molecular markers and major viral disease resistance in tomato such as Tomato mosaic virus (Sobir et al., 2000). Breeding for resistance to viral disease has greatly contributed to the improved yield and quality of many crops and, evidently, has led to the identification disease resistance genes. Even though in vivo screening is a prerequisite for confirmation of MAS, utilization of molecular markers can reduce the undesired population size and further improve the pace of selection programmes. Why use of molecular markers in plant virus resistance? Majority of plant viral diseases are spread by insect vectors such as aphids, white flies, mealybugs and thrips and a few by nematodes. Potato virus Y

(PVY) is known to be transmitted in many plants (especially in Solanaceae) by more than 40 aphid species in a non-persistent manner (Scholthof et al., 2011). Even though control of vectors using chemicals is a common practice in controlling the spread of the virus, it is still impractical due to the prevailing situations such as high recurring costs of the pesticides, emergence of insecticide resistant insect populations, human concerns regarding pesticide residue and its side effects. Hence developing virus-resistant crop plants yield more potential benefits, as it is more economical and eco-friendly way of managing the viral disease. In order to accelerate the current scenario of breeding virus-resistant crops, molecular markers could function as a key tool to help skip several generations of crossing and analysis altogether, thereby saving precious time. Molecular markers function as tags that could be used for gene identification and locate them with respect to other specific genes. For better understanding of the molecular basis of resistance and further advancement of the resistance breeding programmes, genetic mapping of disease resistance genes is an important tool. However, its success is also dependent on the genetic polymorphic markers of a segregating population as well as, various pathological diagnostic tests. In most of the backcross breeding programmes molecular markers gain their importance as an indispensable tool for gene pyramiding while tracing the resistance genes (Narasimhulu et al., 2013). A tight linkage amongst disease resistance genes and molecular markers is of pronounced value in breeding programmes involving disease resistance, which allows breeders to keep track of the DNA markers over the generations and not hold on till the phenotypic expression of resistance genes. For example, molecular markers aided selection of plants, having both Ty-3 and Ty-2 genes versus plants with Ty-3 alone against tomato yellow leaf curl disease, while this probable difference could not have been made out solely based on symptoms alone. If a resistance gene is linked to co-dominant markers in a heterozygous plant species, it could be identified easily and can be introgressed in a short number of generations (Tanksley, 1983; Young and Tanksley, 1989) for developing desired homozygous resistant population through breeding programme. For example, Jefferies et al. (2003) developed Barley yellow dwarf virus (BYDV)

Molecular Markers for Virus-Resistant Genes |  89

resistance breeding lines using a co-dominant marker YLM linked to Yd2 gene. Markers could be helpful in analysis of polygenic resistance as well as in multi-genic resistance (durable resistance) where several resistance genes need to be cumulated in a single genotype (Melchinger, 1990). For molecular characterization of disease resistance gene cluster and to determine their mode of action the molecular markers could also be used as a starting point for cloning the particular locus. Source of resistance One of the major aspects one should deal with while developing a disease resistant variety is the availability of a source of resistance. These may be genes innate to a particular crop species that can either be dominant (majority of cases) or recessive in nature. For example, in barley, Ordon et al. (2003) mapped several recessive resistance genes (e.g. rym4, rym5, rym9, rym11 and rym13) for yellow mosaic disease and resistance lines to disease identified through screening programmes. So many resistance genes have been identified in different crops that exist in various collections around the world (Table 4.1), while several more novel resistance genes are still being discovered from wild spp. of different crops (Barker, 1996). For example, various resistance genes are identified from various crops for monogenic dominant resistance to different viral diseases. Non-host resistance is another newly ventured source for resistance but yet needs to be exploited. Let’s consider the case of potato (Solanum tuberosum), where viruses pose a severe threat to its cultivation, due to its primary infection in vegetative stage as well as subsequent infection in its tuberous stage. Potato virus Y (PVY) is the fifth most important plant virus, worldwide. In some existing cultivars and improved breeding lines of potato, multiple copies of virus resistance genes are present naturally, while in others, different genes conferring virus resistances have been combined (Solomon-Blackburn, 1998). In wild relatives, two types of resistance genes, Ry and Ny are identified against PVY (Cockerham, 1970; Valkonen et al., 1996). The Ry gene, confers symptomless extreme resistance (ER), while the second, Ny gene prevent spread of virus within the plant tissues under specific environmental circumstances through hypersensitive reaction (HR).

Cultivated tomato, which is another solanaceous crop, is highly susceptible to Tomato yellow leaf crinkle virus (TYLCV), hence relying on the breeding for transfer of resistance genes from wild relatives of cultivated tomato. Five resistance gene loci, four dominant genes (Ty-1, Ty-2, Ty-3, Ty-4) and one recessive gene (ty-5) against TYLCV (Anbinder et al., 2009) are reported to be present in some Solanum spp. (S. peruvianum, S. pimpinellifolium, S. chilense, S. cheesmaniae and S. habrochaites) (Vidavski, 2007). Types of resistance As like the resistance provided by the plants against many fungal and bacterial patho-systems, viral disease resistance can also be broadly categorized into two types of resistance monogenic and polygenic. Genetic linkages between monogenic disease resistance genes and molecular markers are very common, while majority of the complex disease resistances are oligogenic or polygenic in nature, i.e. controlled by cluster of genes or multiple genes (Mather and Jink, 1971). For example, the dominant gene Rsc-7 is responsible in controlling Soybean mosaic virus (SMV) disease (Fu et al., 2005). Extreme resistance (ER) and hypersensitive response (HR) are well recognized in plants after viral infection and it was proposed by Solomon Blackburn and Barker (2001). Multiplication of virus during the primitive stage of infection in host plant is prevented by genes responsible for ER and does not involve cell death (Gilbert et al., 1998), while HR provides rapid defence response which results in necrosis thereby preventing any further spread of infection (Dixon et al., 1994). Though plants with resistance to virus are prone to infection, the virus accumulation/aggregation only reaches comparatively low concentration within the plant due to lower multiplication rate, thereby delaying the disease establishment and lowering the disease severity. This type of resistance can be termed as ‘moderate resistance’ (Valkonen et al., 1996). Limitation to virus movement is another form of resistance offered by the plants. For example, in case of Potato leaf roll virus (PLRV) it has been long known that some potato clones could give rise to a lesser percentage of virus-infected daughter tubers from the diseased mother plant (Hutton and Brock, 1953; Barker, 1987).

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Table 4.1  List of mapped based genes conferring virus resistance using various kinds of molecular markers in some major crops No. Host

Viral pathogen

Resistance gene

Marker

1

Potato virus Y

Ry-fsto

CAPS

Solanum stoloniferum

Flis et al. (2005)

Potato virus X

Rx

GP 34

Cultivar Cara(Rx genotype)

Bendhamane et al. (1997)

RFLP

Lycopersicon chilense

Zamir et al. (1994)

2

3

4

5

Potato

Tomato

Soybean

Rice

Barley

Tomato yellow leaf Ty-1 curl virus

Source of resistance

Reference

Tomato spotted wilt virus

Sw-5

SNP

–

Shi et al. (2011)

Soybean mosaic virus

Rsv-4

SSR

LR2 (Resistant line)

Hayes et al. (2000)

Rsc-7

SSR

Kefeng No. 1

Fu et al. (2006)

Rice tungro spherical virus

RTSV resistant RFLP gene

ARC11554 (IRGC)

Sebastian et al. (1996)

Rice stripe virus

RSV1

SSR

–

Zhao et al. (2010)

Barley yellow mosaic virus

rym-1, rym-5

RFLP and Donor line ‘Y4’ CAPS

Okada et al. (2003)

Barley yellow dwarf virus

Yd2

AFLP

–

Paltridge et al. (1998)

6

Pigeonpea

Pigeonpea sterility SMD mosaic virus

AFLP

BRG 3

Ganapthyet al. (2009)

7

Beet

Beet necrotic yellow vein virus

–

RAPD and STS

Holly1–4, R104, R128

Scholten et al. (1997)

8

Bean

Bean golden mosaic virus

bgm-1

RAPD

Garrapato

Urrea et al. (1996)

9

Citrus

Citrus tristeza virus CTV gene

SSR

Ponciru strifoliata

Christofani and Machado (2000)

10

Blackgram

Mungbean mosaic MYMV resistance virus gene

ISSR and SCAR

TU 94–2

Souframanien and Gopalakrishna (2006)

11

Cowpea

Cowpea Yellow Mosaic Virus

CYMV resistance gene

SSR

GC-3

Gioi et al. (2012)

12

Wheat

Streak mosaic virus

Wsm1

STS

–

Talbert et al. (1996)

Barley yellow dwarf virus

Bdv2

SCAR

TAF46

Stoutjesdijk et al. (2001)

Turnip mosaic virus

TuRBO1

RFLP

–

Walsh et al. (1999)

13

Rapeseed

Detection of quantitative trait loci The association between markers and quantitative trait loci (QTLs) play vital role in resistance breeding of important crops. Using various statistical tools the mapping of QTLs can be performed very easily. With the help of molecular markers, polygenic disease resistance can be subdivided,

and its individual genes contributing for resistance could be studied. In many cases degree phenotypic traits are influenced by various environmental factors. Therefore the association between genotypic marker and the phenotypic traits can be sorted out using statistical approaches like ANOVA. To use this kind of method, it requires the markers for disease resistance genes to have been mapped. Hence, the phenotypic values are the dependent variables and

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the genotypic markers correspond to the treatment. Analysis of variance models having increasing complexity provides precise information regarding the genetic basis of resistance in crop plants (Lefebvre, 1993). The interval in mapping approach helps to provide association with molecular markers. Using the maximum likelihood equation, the method provides an estimate also, which can be expressed as limit of detection (LOD) score of the likelihood of the presence of a QTL for regular intervals throughout the genome based on flanking marker information. This is useful for genomic study and in the sequencing of the genome of an organism. The LOD score or value mainly depends on the QTL localization with respect to flanking markers and the degree of its extent; and also it depends on the likelihood that there is presence QTL in chromosome. While examining the curves signifying LOD, this technique would be suitable because it takes into account the recombination rates of different markers. Researchers have also been testing putative QTLs with non-parametric statistical tests because the disease resistance needs to be assessed with ordinal scales while data always do not show a normal distribution, (Young and Tanksley, 1989). Identification of disease resistant lines using genetic markers The crop may be affected by viral, bacterial or fungal pathogens but the strategy of cloning resistancegene homologue (RGH) has been recognized as an effective tool for finding markers and R-genes associated with diseases. Physical association with RGH clones on large-insert DNA clones such as bacterial artificial chromosomes (BACs) can be used for identifying microsatellite or SSR markers. Recent trends in biotechnological approaches open new pathways towards molecular resistance breeding research by utilizing various molecular markers, thereby enhancing and hastening the conventional breeding programme. In view of this, MAS has a great potential to fulfil this demand of resistance breeding, hence the markers like SCAR, RFLP, RAPD, SSR, AFLP, TRAP and CAPS could be used to supply complementary information to the classical genetic analyses. Some of them are given below for their contribution to virus resistance breeding.

RAPD Random amplified polymorphic DNA (RAPD) is a type of molecular marker based on PCR amplification reaction using genomic DNA. By using RAPD we can identify the resistance traits bearing plants from a mixed population. In RAPD segments of amplified DNA are random. For performing RAPD, small oligos need to be synthesized. The arbitrary oligos usually used in RAPD comprises of 8–12 nucleotides. It is simple, rapid and inexpensive technique. By resolving the resulted PCR-products in gel electrophoresis, a semi-unique profile pattern is observed. Using this pattern we can identify the populations bearing the desired resistance traits. RAPD along with bulk segregant analysis (BSA) can successfully be combined together to identify markers closely associated with any traits of economic importance (Michelmore et al., 1991) in this case disease/virus resistance. RAPD marker analyses a very small change in genome of an organism. Using of RAPD markers selection of variant in plant is very simple and fast. The marker can easily distinguish between normal plants and variant plants. For development of Tobacco mosaic virus (TMV) resistant lines in tomato, RAPD has been used to identify the resistant tomato inbred lines within the heterozygous F1 hybrids carrying Tm-2a gene. RFLP Restriction fragment length polymorphism (RFLP) was developed in early 1980s and known to be the first molecular marker. DNA sequence is the source of genetic information. Genetic diversity within species is dependent on the variation in DNA sequence present within the chromosome (Farooq and Azam, 2002). Plants are able to replicate their DNA with high precision and rapidity, but many mechanisms are present which are still responsible for alterations in DNA. The loss or gain of a recognition sites due to simple or large base pair alterations as a result of processes like inversion, translocation, transpositions or deletion lead to formation of restriction fragments of different lengths. RFLP marker provides a method to directly pursue chromosome segments during recombination as they obey Mendelian rules and is of aid while constructing genetic maps. During meiosis when plants produce gametes, recombination occurs in chromosomes by crossing over and this recombination plays an important role in conventional genetic

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mapping. Use of RFLP to assist the improvement of yield and host plant resistance has been reported in two maize viral pathogens, Maize dwarf mosaic virus (MDMV) and Maize chlorotic dwarf virus (MCDV). In maize, a tightly bracketed major resistance gene against MDMV was found to be located in the short arm of chromosome 6 while mapping. However, it is always not essential to use RFLPs for the resistant selection, as artificial inoculation techniques are highly effective with largely dominant resistance and modifiers apparently controlling the expression of resistance in several susceptible genetic backgrounds. On the other hands, there was no resistance factors identified for MCDV. As the labour and cost involved in conventional pathological screening technique for identification of MCDV resistance germplasm is too high, use of RFLP markers tightly linked to loci associated with the genes conferring MCDV resistance used for developing resistant lines. AFLP Amplified fragment length polymorphism (AFLP) is a novel PCR based molecular technique for finger printing of DNA obtained from any source, or, of any complexity. In the process of AFLP, total genomic DNA is fragmented by using two restriction enzymes that cuts the DNA and forms blunt ends. The resultant DNA strands are then ligated with double-stranded nucleotide adapters to serve as primer binding sites during PCR amplification. Primers having restriction site sequence and additional nucleotides at the 3′-end are used as selective complementary agents to the adapter, in order to amplify a subset of ligated fragments. Polymorphisms are identified after gel electrophoresis (polyacrylamide gels) by observing the presence or absence of DNA fragments following analysis. AFLP is extensively used by molecular breeders working on plant virus resistance for the development of genetic maps having high resolution and positional cloning of desired genes. It is a powerful tool when we develop a resistance population of plants without having prior genomic sequence information as it has ability to rapidly produce a large number of marker fragments from its genomic DNA. This method has an added advantage that it requires very little amounts of DNA template as compared to other fingerprinting methods such as

RAPD and inter-simple sequence repeats (ISSR). Despite all these fact AFLP technique is still a relatively labour-intensive method and can be easily multiplexed. AFLP patterns can be evaluated fully automated for phylogenetic analyses, but manual refinement is needed for comparisons of similar patterns. For example, high-resolution genetically and physically modified mapping of the Rx gene using AFLP for high level of resistance to Potato virus X in tetraploid potato has been reported (Bendahmane et al., 1997). Also in tomato, the resistance genotype selection against Tomato yellow leaf curl virus (TYLCV) was conferred by AFLP in two genes Ty-1 and Ty-3. SSR Simple sequence repeat (SSRs) or short tandem repeats are known to be an ideal genetic marker, widely used in all higher organisms for identifying genetic differences between or within the closely related species (Farooq and Azam, 2002). To amplify the markers, several types of oligomers are generally designed, which consist of tandemly repeated oligomers having 2–7 base pair units like mono-, di-, tri-, tetr-a and penta-nucleotides (A, T, AT, GA, AGG, AAAG etc.) in a usual recurring manner and it can be arranged in such a way that it forms different lengths of repeated motifs. These recurrences are extensively dispersed all over the plant genome and tend to be displayed as a high level of genetic variation. The variations present within the number of tandemly repetitive units confer a high polymorphic banding pattern (Farooq and Azam, 2002) which can be identified through PCR, using locus specific flanking primers. SSR provides many desirable marker properties which incorporate high levels of polymorphism, unambiguous designation of alleles, information content and even high reproducibility, dispersal, selective neutrality, co-dominance, rapid and simple genotyping assays. In genetic mapping, genome analysis, and also for the marker-assisted resistance breeding, microsatellites (SSR) markers have become a suitable choice to manage the extensive range of applications (Ayres et al., 1997; Weising et al., 1998). Between cowpea (Vigna unguiculata) genotypes GC-3 (standard resistant) and Chirodi (susceptible), SSR primer pairs were employed to detect various polymorphisms. These

Molecular Markers for Virus-Resistant Genes |  93

markers were then used to carry out PCR in all the individual cowpea lines to identify the resistant genes. The linkage of SSR markers to Cowpea yellow mosaic virus (CYMV) resistance was evaluated using 20 CYMV resistance germplasm lines and 20 susceptible germplasm lines of Vigna unguiculata. This proved the highly heritable nature of the QTL’s for CYMV resistance trait. Thus identified markers provide information to select Vigna unguiculata germplasm lines resistance to CYMV would be very useful in resistance breeding programmes (Gioi et al., 2012). SCAR Sequence-characterized amplified region (SCAR) markers are RAPD-derived stable molecular markers. The basic concept behind SCAR is to generate co-dominant markers from dominant markers. As the process of RAPD is very tedious, SCAR markers developed through molecular cloning is the simplest way to identify the unique polymorphisms present within the population (Rajesh et al., 2013). SCAR markers owing to the higher annealing temperatures and long primers with sequence specificity, contributes to extreme levels of polymorphism (Kumla et al., 2012). SR 2, a co-dominant SCAR marker was derived from dominant RAPD marker for the identification of bgm-1 gene in common bean (Urrea et al., 1996). From mixture of different genetic background in common bean, the useful population resistant against Bean golden yellow mosaic virus carrying bgm-1 gene was observed only in another population of common bean except G2402 (or Garrapato) and the population derived from it. Thus the polymorphisms generated using SCAR markers are very useful in recognizing the gene from various genetic backgrounds. Thus, the uniqueness of marker polymorphism and its consistency over different laboratories, seasons and diverse genetic backgrounds have aided in its wide use. Lately, a second SCAR marker was advanced from the W12.700 RAPD for identifying QTL having a cluster of genes located in the linkage group b04 (Miklas et al., 2000), and has also been included into the International Center for Tropical Agriculture (CIAT) breeding programme. Also the SCAR primer (SW 13) was derived from the decamer RAPD primer (OW13) which was associated with I-gene, which contributes to

the resistance to the Bean common mosaic virus (BCMV) in common bean. TRAP The target region amplified polymorphism (TRAP), which is a PCR-based marker that was designed to detect intragenic polymorphisms. These markers are generated with fixed sense primers about 18 nucleotides, designed from a gene or expressed sequence tag (EST) and an antisense arbitrary primer of the same approximate size, designed with the AT- or a GC- motif, in order to direct them to hybridize with introns or exons, respectively. These properties of TRAP markers make them an interesting tool to detect polymorphisms that can be useful for genetic improvement when analysing gene-rich regions of a genome. TRAP and AFLP technique was tested for 10 sugarcane genotypes and when they were compared, a very similar number of average bands were obtained (47 vs. 52). However, TRAP markers revealed a much higher polymorphism, 59.7%, than the AFLP markers, which showed 34.43%. These results indicated that TRAP markers are more effective than AFLP when analysing the sugarcane genome, as it is easier, faster and more economic than applying AFLP markers and more efficient as seen by the number of polymorphic loci obtained for each reaction. It must be highlighted that in most cases a combination of different marker techniques is recommended for genetic diversity studies, since they could reflect different aspects of the genome studied as well as providing more precise information. Nevertheless, TRAP markers are routinely applied as a first approximation in genetic diversity studies. TRAP technology has replaced AFLP analysis completely and is now routinely employed to genetically characterize propagated plant material. In regards of productivity, efficiency and safety, meristem micropropagated plants are quite advantageous as they are healthy high-yielding plants that in the short term will replace old and/or infected materials. The primers for the TRAP markers of the disease resistance gene designed from the sequenced expressed sequence tag (EST) which is already submitted in the Compositae genomics database or sequenced resistance gene analogue (RGA). The TRAP marker system has been found to have potential for mapping genomic regions of the common bean

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linked with disease resistance (Miklas et al., 2006). Eighty-five TRAP markers for R-genes against ashy stem blight, Bean golden yellow mosaic virus (BGYMV) and common bacterial blight have been identified from common bean, out of which 17 of TRAP markers were found in BAT 93/Jalo EEP558 core mapping population and six TRAP markers in Dorado/XAN 176 mapping population which is linked with well identified QTL. CAPS Another method in molecular biology, for quick analysis of genetic markers, i.e. PCR based RFLP is named as Cleaved Amplified Polymorphic Sequence or CAPS. In this method, PCR amplification is carried out at the altered restriction site. Resulting PCR products are digested with the restriction enzyme and fractionated using gel electrophoresis (agarose or acrylamide). The fractionated PCR products can be observed as clear distinguishable patterns. The gene closely associated with CAPS markers can easily be identified in MAS, which are extensively incorporated in resistance breeding of several crops such as wheat, barley, soybean, potato, tomato and other crops for the development of tolerance and resistance to pathogens. In several cereal crops, CAPS method was successfully employed for the genetic mapping of individual genes conferring tolerance to pathogens, as well as QTLs associated with important quantitative traits such as overall plant development and grain quality. CAPS can also be applied as a technique for the analysis of evolutionary aspects of both crops and the pathogen such as genetic polymorphism and phylogeny in closely related species. In barley, soil-borne viruses (fungal transmitted) like Barley mosaic virus complex (BaYMV, BaMMV, BaYMV-2) are known for their potential spread and yield losses and thereby of high importance. This leads to the concern of resistance breeding method having simple, economic, and superior selection methods. Targeted site mutations leading to Single Nucleotide Polymorphisms (SNPs) responsible for resistance can be identified directly by designing suitable CAPS marker. In barley, rym4/rym5 alleles of the Hv-eIF4E gene conferring resistance to Barley yellow mosaic virus complex were identified through this method (Sedláček et al., 2010).

Biochemical markers The biochemical markers are generally proteins which are produced by gene expression. By using of electrophoresis and staining, these proteins could easily be isolated, extracted and characterized. Isozymes are commonly used as biochemical markers which are resultant products of several alleles. They help in catalysing similar enzymatic reactions. The alleles are from either one or several genes. As the segregation analysis of monomeric and dimeric isozymes is very easy, these are most often used in plant breeding as biochemical marker. Although any biochemical marker related to viral disease resistant in plant is unavailable, isozyme for resistant to fungal diseases are reported. For example, Bournival et al. (1989) studied the isoezymatic activity of Got2 for its association to the tomato resistance to Fusarium oxysporum race 3. It is notable that within a cultivated species the polymorphism of isozyme markers is found to be fairly poor. Marker assisted selection for identification of viral resistance line For screening of viral disease resistance lines using molecular markers, marker assisted selection (MAS) is very useful, because phenotypic selection of viral resistant lines is always not convenient or sometimes it is very difficult to direct screening with challenged inoculums. On the other hand, the method is precise and fast (Watanabe, 1994) and convenient in backcross breeding. The importance of MAS is to reduce labour and land which are required for screening of large number of population in conventional resistance breeding programmes at an early in vitro level. Molecular markers can be used in marker-assisted backcross breeding and resistance gene introgression programmes from wild species across the population for inherited traits and can easily be selected against the unwanted characters (non-resistant traits) carried by wild relatives (Young and Tanksley, 1989) (Fig. 4.1). Only a minute amount of plant tissues is needed for MAS, so it can reduce a number of years from the actual time frame in selecting true disease resistant germplasm or breeding lines. Generally, the success of MAS depends on close association

Molecular Markers for Virus-Resistant Genes |  95

Figure 4.1  Schematic flowchart of introgression of resistance genes in plants using marker assisted backcross breeding.

of resistance genes and markers which are tightly linked to it, such as QTLs. In potato, the gene Ryadg was put into application in MAS for identification of virus resistance in breeding lines but, so far, not reported to be very effective and applicable. Over the past two decades, research on molecular markers enhanced the suitability and efficiency of MAS in resistance selection especially by the utilization of SSR markers. SSR marker have more profound advantage over the AFLP, as it can minimize the constraint regarding population specificity found with the later (Milbourne et al., 1998). The bgm-1 gene for the resistance of Bean golden yellow mosaic virus (BGYMV) and bc-3 gene for resistance to Bean common mosaic virus (BCMV) in bean were selected for MAS selection. Multiple flanking markers for selection of a dominant CMD2 gene related to resistance of Cassava mosaic virus (CMV) are also reported in cassava (Akano et al., 2002).

Gene tagging and strategies for targeted mapping Previously, in order to find the markers linked with disease resistance genes, mapping of genetic linkage was a prerequisite. But development of genetic linkage maps is a time consuming procedure. In recent times, it could have been possible to identify markers directly without drawing any genetic linkage map. Use of molecular markers for finding polygenic traits, is not suitable since it is basically restricted to monogenic traits, as it is marked to the particular genomic region coding for the desired trait. To identify the markers in chromosomes or chromosome arms of an aneuploid line for the genes responsible for disease resistance is the best example for this aspect (Lefebvre, 1993). However, gene tagging of a particular trait is a need of MAS and map-based gene cloning. In tomato TMV resistance Tm-2 locus was identified by targeted mapping.

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NB-LRR gene clusters analysis for viral disease resistant Nucleotide-binding and leucine-rich repeat domains (NB-LRRs) provide novel tools in modern era for identifying markers related to disease resistant genes ( Jupe, 2012) (Table 4.2). In potato, organization of NB-LRRs and phylogenetic relationship were analysed during whole genome sequencing of cv. Sarpo Mira, and PVY resistance gene Ny-Smira was identified in the long arm of chromosome IX. Potential R-gene locus of potato was identified by analysing of Ry364, TG328 and 38–510 markers against BLASTn search tools and positions of NB-LRR gene cluster was compared ( Jupe et al., 2013). The comparison of genomic map shows that the resistance Ny-Smira gene is located within the resistance gene hotspot that contains 46 NB-LRR genes. In other findings, the genes TNL and CNL which are related to the late blight resistance was found in the C64 gene cluster of potato which contains gene Rpi-vnt1, homologous to the Tm-2 gene in tomato. The Tm-2 gene is known to provide resistance to Tomato mosaic virus (Lanfermeijer et al., 2003). Gene pyramiding Durable and broad resistance in plants to a wide range of pathogens including viruses is desired but multiple disease resistance often doubtful in conventional breeding as the resistance is affected by multiple factors such as environments which sometimes favour the pathogen and disease, resistant genes which are commonly deployed in cultivars and other factors like pathogen populations and selection pressure, genetic mutation and recombination/gene flow, reproduction/replication, and persistence in environment. Breeding inbred lines with multiple disease resistance is more desirable but often difficult to achieve. Because selection of screening methods and management of segregating populations for desired traits are very difficult. Gene pyramiding in crops is the best way for developing of durable resistance (multigenic resistance) against multiple diseases. Genetic markers can be used for mapping genes to determine the monogenic or polygenic disease resistance in crops which will give durable resistance to wide range of pathogens. The World Vegetable Center (AVRDC) developed tomato lines with multiple

disease resistant, including viral, bacterial and fungal diseases. The lines are known to be resistant against begomoviruses such as tomato yellow leaf curl disease, Tobacco mosaic virus (TMV), Ralstonia solanacearum causing bacterial wilt, Phytophthora infestans causing late blight, Stemphyllium spp. causing grey leaf spot and Fusarium oxysporum f. sp. lycopersici race 2. The AVRDC developed commercial cultivars of tomato resistant to tomato yellow leaf curl disease by introgressing six genes, namely Ty-1/Ty-3, Ty-2, Ty-4, Ty-5 and Ty-6) (Hutton and Scott, 2012), also three resistance loci, namely Rsv1, Rsv3 and Rsv4, are identified against seven strains of Soybean mosaic virus (SMV) (Shi et al., 2007) (Fig. 4.2). Evaluation of viral disease resistant lines So many genes involved in disease resistance have been mapped in commercially important crops. These genes are linked with molecular markers and available for MAS. Based on molecular markers, identification of segregating population in crops with viral disease resistant needs to be confirmed against challenging viral inoculum, either in controlled environment or in natural field conditions. Effective selections of those segregating populations for particular viral disease resistance need to have standard screening methods, i.e. cost-effective and can permit rapid screening of thousands of plants under high disease pressure. In general, disease screening procedure employs natural disease pressure for field testing e.g. Pigeonpea sterility mosaic virus (Sharma et al., 2015) and Begomovirus in urdbean (Biswas et al., 2009), mungbean, (Biswas et al., 2015), and inoculation of specific viral strains in greenhouse/growth room e.g. Citrus tristeza virus (Biswas et al., 2009). Future prospects Over the few past decades, research has led to the development of virus-resistant crops through conventional breeding methods, but most of the research is still limited at greenhouse level. Among them, few findings are followed up to field level experiments to prove the resistance ability of the developed breeding lines. It is important to implement marker assisted resistance breeding lines in

Table 4.2.  The list of resistance genes against Avr- genes of plant viruses which are cloned and organized into the NB-LRRs and the non-NB-LRRs (courtesy of de Ronde et al., 2014) Family of host plant Host plant Brassicaceae

Arabidopsis thaliana

Resistance N terminal end/ type protein type Recognizes

Virus genus

Avr gene

Type of interaction

HRT

NB-LRR

Turnip crinkle virus

Carmovirus

CP

Hypersensitive

RCY1

NB-LRR

Cucumber mosaic virus

Cucumovirus CP

JAX1

Non NBLRR

Jacalin-like

Broad resistance against potexvirus

Potexvirus

Unknown

Jasmonic acidinducing SAR

RTM1

Non NBLRR

Jacalin-like

Tobacco etch virus

Potyvirus

CP

Jasmonic acidinducing SAR; restricts viral systemic movement

RTM2

Non NBLRR

HSPs domain

Plum pox virus

Potyvirus

CP

Not involved in heat tolerance; restricts viral systemic movement

RTM3

Non NBLRR

Meprin and Lettuce mosaic virus TRAF homology (MATH) domain

Potyvirus

CP

Restricts viral systemic movement

Common name R gene Mouse ear cress

Coiled coil (CC)

Field mustard

BcTuR3

NB-LRR

Toll and Interleukin-1 Receptor (TIR)

Turnip mosaic virus

Potyvirus

Unknown

Unknown

Cucurbitaceae

Cucumis melo

Muskmelon

Pvr1, Pvr2

NB-LRR

TIR

Papaya ringspot virus

Potyvirus

Viral genomelinked protein

Amino acid changes polymorphism

Fabaceae

Glycine max

Soybean

Rsv1

NB-LRR

CC

Soybean mosaic virus

Potyvirus

P3+ HCPro

Extreme resistance/ hyperresistance

Phaseolus vulgaris

Kidney bean

I (locus)

NB-LRR

CC

Bean common mosaic virus, Bean necrotic mosaic virus, Blackeye cowpea mosaic virus, Azuki mosaic virus, Cowpea aphid-borne mosaic virus, Passionfruit woodiness virus, Soybean mosaic virus, Thailand passiflora virus, Watermelon mosaic virus, Zucchini yellow mosaic virus

Potyvirus

Unknown

Extreme resistance/ hyperresistance/ phloem necrosis

Molecular Markers for Virus-Resistant Genes |  97

Brassica campestris

Family of host plant Host plant

Common name R gene

Resistance N terminal end/ type protein type Recognizes

Virus genus

Fabaceae

Vigna mungo

Black gram

CYR1

NB-LRR

CC

Mungbean yellow mosaic virus

Begomovirus CP

Incompatible response via a cognate CYR1 genemediated interaction

Rutaceae

Poncirus trifoliata

Trifoliate orange

Ctv (locus)

NB-LRR

CC

Citrus tristeza virus

Closterovirus Unknown

Unknown

Solanaceae

Capsicum annuum

Capsicum

L-locus: L1, L2, L3, L4

NB-LRR

CC

Tobacco mosaic virus, Tomato mosaic virus, Tobacco mild green mosaic virus, Bell pepper mottle virus, Paprika mild mottle virus, Obuda pepper virus, Pepper mild mottle virus

Tobamovirus

CP (all)

Unknown

Pvr1, Pvr2

NB-LRR

TIR

Broad-spectrum potyvirus resistance

Potyvirus

Viral genomelinked protein

Amino acid changes polymorphism

NRG1

NB-LRR

CC

Tobacco mosaic virus

Tobamovirus

p50 (Helicase)

Virus-induced gene silencing

N

NB-LRR

TIR

Ty-1, Ty-3

Non NBLRR

RNA– dependent RNA polymerase

Tomato yellow leaf curl virus

Begomovirus No

Solanum hirsutum

Tm-1

Non NBLRR

TIM-barrel-like domain protein

Tomato mosaic virus

Tobamovirus

Replicase, Hypersensitive Helicase domain

Solanum peruvianum

Sw5b

NB-LRR

CC

Tomato spotted wilt virus and other tospoviruses

Tospovirus

NSm

Hypersensitive

Tm-2, Tm-22

NB-LRR

CC

Tobacco mosaic virus, Tomato mosaic virus and other tobamoviruses

Tobamovirus

30 kD MP

Hypersensitive

Nicotiana benthamiana

Tobacco

Nicotiana glutinosa Solanum chilense

Wild tomato

Avr gene

Type of interaction

Transcriptional gene silencing; RNAi defense mechanism

Solanum stoloniferum

Wild potato

Ry-fasto

NB-LRR

Unknown

Potato virus Y

Potyvirus

Unknown

Extreme resistance

Solanum tuberosum

Potato

Rx1, Rx2

NB-LRR

CC

Potato virus X and other potex viruses

Potexvirus

CP

Extreme resistance/ hyperresistance

Potato virus Y

Potyvirus

Unknown

Unknown

Unknown

Hypersensitive

Y-1

NB-LRR

TIR

Ny-1

NB-LRR

Unknown

98  | Sharma et al.

Table 4.2.  Continued.

Molecular Markers for Virus-Resistant Genes |  99

Figure 4.2.  Procedure of pyramiding Rsv 1, Rsv 3 and Rsv 4 genes for SMV resistance (courtesy Shi et al., 2007)

fields as a tool for integrated disease management strategies in essential crops. Agro-ecosystem in a sustainable agriculture is important parameters, and determined by several factors that effects on disease progression and crop development. Conversely, most of the commercial production of the resistant cultivars is completely depending on few research institutes, and rarely followed by any private research and development programmes. Significant expansion of resistant cultivars is needed due to growing demand for organic food which is expected over the next decade. In parallel with success story of marker assisted resistance breeding, several information and key factors are also identified on the aspect linked to gaps of implementation between published research findings and production, and commercialization of resistant varieties.

Conclusion In our present century, due to climatic effects viral diseases are emerging along with increasing vector populations and causes substantial yield losses in crops. Utilization of genetic resistance in farming system could be a better substitute of chemical based management of viral diseases. To improve climate smart eco-friendly farming system, the researchers are trying to identify novel ‘R’ genes from host/non-host plants to reinforce the efficient control measure. With the changes in legislation at country level, development of new cultivars carrying virus resistance genes and its use in farming practices is of prime importance to reduce of chemical usage. Practically, at present use of resistant cultivars is not comparable with chemical management. Hence, extensive utilization of knowledge

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related to marker assisted breeding using advanced molecular technologies for developing virus resistance crops is needed. References

Akano, O., Dixon, O., Mba, C., Barrera, E., and Fregene, M. (2002). Genetic mapping of a dominant gene conferring resistance to cassava mosaic disease. Theor. Appl. Genet. 105, 521–525. https://doi.org/10.1007/s00122-0020891-7 Anbinder, I., Reuveni, M., Azari, R., Paran, I., Nahon, S., Shlomo, H., Chen, L., Lapidot, M., and Levin, I. (2009). Molecular dissection of Tomato leaf curl virus resistance in tomato line TY172 derived from Solanum peruvianum. Theor. Appl. Genet. 119, 519–530. https:// doi.org/10.1007/s00122-009-1060-z Ayres, N.M., McClung, A.M., Larkin, P.D., Bligh, H.F.J., Jones, C.A., and Park, W.D. (1997). Microsatellites and a single-nucleotide polymorphism differentiate apparent amylase classes in an extended pedigree of US rice germplasm. Theor. Appl. Genet. 7, 773–781. Barker, H. (1996). Inheritance of resistance to potato viruses Y and A in progeny obtained from potato cultivars containing gene Ry:evidence for a new gene for extreme resistance to PVY. Theor. Appl. Genet. 93, 710–716. https://doi.org/10.1007/BF00224066 Bendahmane, M., Fitchen, J.H., Zhang, G., and Beachy, R.N. (1997). Studies of coat protein-mediated resistance to tobacco mosaic tobamovirus: correlation between assembly of mutant coat proteins and resistance. J. Virol. 71, 7942–7950. Biswas, K.K., Tarafdar, A., Jayakumar, B.K., and Pun, K.B. (2009b). Strategy for the production of virus-free mandarin (Citrus reticulata) planting materials in the Darjeeling hills of India. Indian Phytopathol. 62, 376–380. Biswas, K.K., Biswas, K., and Tarafdar, A. (2015). Multiple and mixed infections with yellow mosaic, leaf crinkle and bud necrosis disease complex in mungbean: a threat to cultivation of mungbean in India. Legume Res. 38, 382–388. Biswas, K.K., Tarafdar, A., and Kumar.A. (2009a). Multiple infection in urdbean (Vigna mungo) in natural condition by begomovirus, tospovirus and urdbean leaf crinkle virus complex. Indian Phytopathol. 62, 75–82. Botstein, D., White, R.L., Skolnick, M., and Davis, R.W. (1980). Construction of a genetic linkage map in man using restriction fragment length polymorphisms. Am. J. Hum. Genet. 32, 314–331. Bournival, B.L., Scott, J.W., and Vallejos, C.E. (1989). An isozyme marker for resistance to race 3 of Fusarium oxysporum f. sp. lycopersici in tomato. Theor. Appl. Genet. 78, 489–494. https://doi.org/10.1007/BF00290832 Cockerham, G. (1970). Genetical studies on resistance to potato viruses X and Y. Heredity 254, 309–348. Cristofani, M., and Machado, M.A. (1998). Molecular markers linked to Citrus tristeza virus (CTV) resistance gene in cultivars and hybrids of Poncirus trifoliata. In proceeding of First International Citrus Biotechnology Symposium, pp 535.

de Ronde, D., Butterbach, P., and Kormelink, R. (2014). Dominant resistance to plant viruses. Front Plant Sci. 5, 307. https://doi.org/10.3389/fpls.2014.00307 Farooq, S., and Azam, F. (2002). Molecular markers in plant breeding: concepts and characterization. Pak. J. Biol. Sci.5, 1135–1140. Flis, B., Hennig, J., and Strzelczyk-Żyta, D. (2005). Mol. Breeding 15, 95. https://doi.org/10.1007/s11032-0042736-3. Fu, F.X., Zhang, Y., Bell, P.R.F., and Hutchins. D.A. (2005). Phosphate uptake and growth kinetics of Trichodesmium (Cyanobacteria) isolates from the North Atlantic Ocean and the Great Barrier Reef, Australia. J. Phycol. 41, 62–73. Fu, S., Zhan, Y., Zhi, H., Gai, J., and Yu, D. (2006). Mapping of SMV resistance gene Rsc-7 by SSR markers in soybean. Genetica 128, 63–69. https://doi.org/10.1007/s10709005-5535-9 Ganapathy, K.N. (2009). Identification of AFLP markers linked to sterility mosaic disease in pigeonpea Cajanus cajan (L.). Millsp. International Journal of Integrative Biology 73, 145. Gioi, T.D., K.S., Boora, and Chaudhary, K. (2012). Identification and characterization of SSR markers linked to yellow mosaic virus resistance gene(s) in cowpea. Int. J. Plant Res. 2, 1–8. Hayes, A.J., Ma, G.G., Buss, R., and Maroof, M.A.S. (2000). Molecular marker mapping of RSV4, a gene conferring resistance to all known strains of Soybean mosaic virus. Crop Sci. 40, 1434–1437. Hutton, S.F., Scott, J.W., and Schuster, D.J. (2012). Recessive resistance to Tomato yellow leaf curl virus from the tomato cultivar Tyking Is located in the same region as Ty-5 on Chromosome 4. Hort. Science 47, 324–327. Jefferies, S.P., King, B.J., Barr, A.R., Warner, P., Logue, S.J., and Langridge, P. (2003). Marker-assisted backcross introgression of the Yd2 gene conferring resistance to Barley yellow dwarf virus in barley. Plant Breeding 1, 225–256. Jupe, F., Pritchard, L., Etherington, G.J., Mackenzie, K., Cock, P.J., Wright, F., Sharma, S.K., Bolser, D., Bryan, G.J., Jones, J.D., et al. (2012). Identification and localisation of the NB-LRR gene family within the potato genome. BMC Genomics 13, 75. https://doi. org/10.1186/1471-2164-13-75 Jupe, F., Witek, K., Verweij, W., Sliwka, J., Pritchard, L., Etherington, G.J., Maclean, D., Cock, P.J., Leggett, R.M., Bryan, G.J., et al. (2013). Resistance gene enrichment sequencing (RenSeq) enables re-annotation of the NB-LRR gene family from sequenced plant genomes and rapid mapping of resistance loci in segregating populations. Plant J. 76, 530–544. https://doi. org/10.1111/tpj.12307 Kanyuka, B., and Baulcombe, D.C. (1997). High resolution and physical mapping of the Rx gene for extreme resistance to Potato virus Xin tetraploid potato. Theor. Appl. Genet. 95, 153–162. Kumla, S., Doolgindachbaporn, S., Sudmoon, R., and Sattayasai, N. (2012). Genetic variation, population structure and identification of yellow catfish, Mystusne murus (CandV) in Thailand using RAPD, ISSR and SCAR marker. Mol. Biol. Rep, 39, 5201–5210.

Molecular Markers for Virus-Resistant Genes |  101

Lanfermeijer, F.C., Dijkhuis, J., Sturre, M.J. G., and Hille, J. (2003). Cloning and characterizatíon of the durable Tomato mosaic virus resistance gene Tm-22 from Lycopersicon esculentum. Plant Mol. Biol. 52, 1037–1049. Lefebvre, V. (1993). Marquagemoléculaire et analyse de resistances polygéniques: interaction piment (Capsicum annuum L.) Phytophthora capsici Leon. Ph.D. thesis, Université de Paris XI-Orsay, France, pp 182. Mather, K., and Jiniks, J.L. (1971). Biometrical genetics (Chapman and Hall, London). Melchinger, A.E. (1990). Use of molecular markers in breeding for oligogenic disease resistance. Plant Breeding, 104, 1–19. https://doi.org/10.1111/j.1439-0523.1990. tb00396.x. Messeguer, R., Ganal, M., de Vicente, M.C., Young, N.D., Bolkan, H., and Tanksley, S.D. (1991). High resolution RFLP map around the root knot nematode resistance gene (Mi) in tomato. Theor. Appl. Genet. 82, 529–536. https://doi.org/10.1007/BF00226787 Michelmore, R.W., Paran, I., and Kesseli, R.V. (1991). Identification of markers linked to disease-resistance genes by bulked segregant analysis: a rapid method to detect markers in specific genomic regions by using segregating populations. Proc. Natl. Acad. Sci. U.S.A. 88, 9828–9832. Miklas, P.N., Kelly, J.D., Beebe, S.E., and Blair, M.W. (2006). Common bean breeding for resistance to biotic and abiotic stresses: from classical to MAS breeding. Euphytica.147, 105–131. Miklas, P.N., Stone, V., Daly, M.J., Stavely, J.R., Steadman, J.R., Bassett, M.J., Delorme, R., and Beaver, J.S. (2000). Bacterial, fungal, and viral disease resistance loci mapped in a recombinant inbred common bean population (‘Dorado’/XAN 176). J. Am. Soc. Hort. Sci. 125, 476–481. Milbourne, D., Meyer, R.C., Collins, A.J., Ramsay, L.D., Gebhardt, C., and Waugh, R. (1998). Isolation, characterization and mapping of simple sequence repeat loci in potato. Mol. Gen. Genet. 259, 233–245. Narasimhulu, R., Kenchanagoudar, P.V., Gowda, M.V.C., and Sekhar, L. (2013). Genetic variability and correlation studies for selection of multiple disease resistance lines in two crosses of peanut. Bioinfolet. 10, 183–186. Okada, Y., Kashiwazaki, S., Kanatani, R., Arai, S., and Ito, K. (2003). Effects of barley yellow mosaic disease resistant gene rym1 on the infection by strains of Barley yellow mosaic virus and Barley mild mosaic virus. Theor. Appl. Genet. 106, 181–189. https://doi.org/10.1007/ s00122-002-1019-9 Ordon, F., Friedt, W., Scheurer, K., Pellio, B., Werner, K., Weiskorn, C., Neuhaus, G., Huth, W., Habekuss, A., and Graner, A. (2004). Marker based strategies for resistance breeding to viruses (BaMMV, BaYMV, BaYMV-2, BYDV) in barley (H. vulgare L.). In Proceeding of Eucarpia Cereal Section, 21–25 November, Salsomaggiore, Italy. Paltridge, N.G., Collins, N.C., Bendahmane, A., and Symons, R.H. (1998). Development of YLM, a codominant PCR marker closely linked to the Yd2 gene for resistance to barley yellow dwarf disease. Theor. Appl. Genet. 96, 1170–1177. Rajesh, M.K., Jerard, B.A., Preethi, P., Thomas, R.J., Fayas, T.P., and Rachana, K.E. (2013). Development of a

RAPD-derived SCAR marker associated with tall-type palm trait in coconut. Sci. Hortic. 150, 312–316. Scholten, O.E., Klein-Lankhorst, R.M., Esselink, D.G., De Bock, T.S., and Lange, W. (1997). Identification and mapping of random amplified polymorphic DNA (RAPD) markers linked to resistance to beet necrotic yellow vein virus (BNYVV) in Beta accessions. Theor. Appl. Genet. 94, 123–130. https://doi.org/10.1007/ s001220050390 Scholthof, K.B., Adkins, S., Czosnek, H., Palukaitis, P., Jacquot, E., Hohn, T., Hohn, B., Saunders, K., Candresse, T., Ahlquist, P., et al. (2011). Top 10 plant viruses in molecular plant pathology. Mol. Plant Pathol. 12, 938–954. https://doi.org/10.1111/j.13643703.2011.00752.x Sebastian, L.S., Ikeda, R., Huang, N., Imbe, T., Coffman, W.R., and McCouch, S.R. (1996). Molecular mapping of resistance to Rice tungro spherical virus and green leafhopper. Phytopathology 86, 25–30. Sedláček, T., Mařík, P., and Chrpová, J. (2010). Development of CAPS marker for Identification of rym4 and rym5 Alleles conferring resistance to the Barley yellow mosaic virus complex in barley. Czech J. Genet. Plant Breed. 46, 159–163. Sharma, M., Telangre, R., Ghosh, R., and Pandey, S. (2015). Multi-environment field testing to identify broad, stable resistant to sterility mosaic disease of pigeonpea. J. Gen. Plant Pathol. https://doi.org/10.1007/s10327-0150585-z. Shi, A., Chen, P., Li, D., Zheng, C., Zhang, B., and Hou, A. (2007). Pyramiding multiple genes for resistance to Soybean mosaic virus in soybean using molecular markers. Mol. Breeding. https://doi.org/10.1007/ s11032-008-9219-x. Shi, A., Vierling, R., Grazzini, R., Chen, P., Caton, H., and Panthee, D. (2011). Identification of molecular markers for Sw-5 gene of Tomato spotted wilt virus resistance. Am. J. Biotech. Mol. Sci., 1, 8–16. Sobir, Ohmori, T., Murata, M., and Motoyoshi, F. (2000). Molecular characterization of the SCAR markers tightly linked to the Tm-2 locus of the genus Lycopersicon. Theor. Appl. Genet. 101, 64–69. Solomon-Blackburn, R. (1998). Progress in breeding potatoes for resistance to virus diseases. Aspects Appl. Biol. 52, 299–304. Solomon-Blackburn, R.M., and Barker, H. (2001). A review of host major-gene resistance to potato viruses X, Y, A and V in potato, genes, genetics and mapped locations. Heredity 86, 8–16. Souframanien, J., and Gopalakrishna, T. (2006). ISSR and SCAR markers linked to the mungbean yellow mosaic virus (MYMV) resistance gene in blackgram [Vignamungo (L.) Hepper]. Plant Breeding 125, 619–622. Stoutjesdijk, P., Kammholz, S.J., Kleven, S., Matsay, S., Banks, P.M., and Larkin, P.J. (2001). PCR-based molecular marker for the Bdv2 Thinopyrum intermedium source of Barley yellow dwarf virus resistance in wheat. Crop and Pasture Science 52, 1383–1388. Talbert, L.E., Bruckner, P.L., and Smith, L.Y. (1996). Development of PCR markers linked to resistance to wheat streak mosaic virus in wheat. Theoret. Appl. Genet. 93, 463–7.

102  | Sharma et al.

Tanksley, S.D., and Orton, T.J. (1983). Isozymes. In Plant Breeding and Genetics, Tanksley, S.D, and Orton, T.J., eds. (Elsevier, Amsterdam). Urrea, C.A., Miklas, P.N., Beaver, J.S., and Riley, R.H. (1996). A co-dominant RAPD marker useful for indirect selection of BGMV resistance in common bean. J. Am. Soc. Hort. Sci. 121, 1035–1039. Valkonen, J.P.T., Jones, R.A.C., Slack, S.A., and Watanabe, K.N. (1996). Resistance specificities to virus in potato, standardization of nomenclature. Plant Breeding 115, 433–438. Vidavski, F.S. (2007). Exploitation of resistance genes found in wild tomato species to produce resistant cultivars; pile up of resistant genes. In Tomato yellow leaf curl virus disease: management, molecular biology, breeding for resistance, Czosnek H, ed. pp. 363–372. Vos, P., Hogers, R., Bleeker, M., Reijans, M., van de Lee, T., Hornes, M., Frijters, A., Pot, J., Peleman, J., Kuiper, M., and Zabeau, M. (1995). AFLP: a new technique for DNA fingerprinting. Nucleic Acids Res. 11, 4407–4441. Walsh, J., Sharpe, A., and Jenner, C. (1999). Characterisation of resistance to turnip mosaic virus in oilseed rape (Brassica napus) and genetic mapping of TuRB01. Theor. Appl. Genet. 99, 1149–1154. Watanabe, M., Takasaki, T., Toriyama, K., Yamakawa, S., Isogai, A., Suzuki, A., and Hinata, K. (1994). A high

degree of homology exists between the protein encoded by SLG and the S receptor domain encoded by SRK in self-incompatible Brassica campestris L. Plant Cell Physiol. 35, 1221–1229. Weising, K., Winter, P., Huttel, B., and Kahl, G. (1998). Microsatellite markers for molecular breeding. J. Crop Prod. 1, 113–143. Williams, J.G., Kubelik, A.R., Livak, K.J., Rafalski, J.A., and Tingey, S.V. (1990). DNA polymorphisms amplified by arbitrary primers are useful as genetic markers. Nucleic Acids Res. 18, 6531–6535. Young, N.D., and Tanksley, S.D. (1989). Restriction fragment length polymorphism maps and the concept of graphical genotypes. Theor. Appl. Genet. 77, 95–101. https://doi.org/10.1007/BF00292322 Zamir, D., Ekstein-Michelson, I., Zakay, Y., Navot, N., Zeidan, M., Sarfatti, M., Eshed, Y., Harel, E., Pleban, T., van-Oss, H., et al. (1994). Mapping and introgression of a Tomato yellow leaf curl virus tolerance gene, TY-1. Theor. Appl. Genet. 88, 141–146. https://doi.org/10.1007/ BF00225889 Zhao, F., Cai, Z., Hu, T., Yao, H., Wang, L., Dong, N., Wang, B., Ru, Z., and Zhai, W. (2010). Genetic analysis and molecular mapping of a novel gene conferring resistance to Rice stripe virus. Plant Mol. Biol. Rep. 28, 512–518.

Genetic Engineering for Virus Resistance in Plants: Principles and Methods

5

Basavaprabhu L. Patil

ICAR-National Research Centre on Plant Biotechnology, New Delhi, India. *Correspondence: [email protected], [email protected] https://doi.org/10.21775/9781910190814.05

Abstract Viral diseases are a major menace for cultivation of crop plants across the globe. Non-availability of virucides and sources of host-plant resistance for their introgression in farmers preferred cultivars has put up major challenge for control of plant viral diseases. Hence genetic engineering for development of virus-resistant transgenic crop plants is a promising option when compared to other conventional tools and techniques. Since the first report of virus-resistant transgenic crops in 1986, significant progress has been made in understanding the molecular basis of virus resistance and also in the tools and techniques used for plant genetic engineering. Despite major advancement in the area of plant biotechnology, in the last 30 years, there has been no significant increase in the deployment of virus-resistant transgenic crops, except for few examples such as papaya. Thus, to help the plant biologists, here in this chapter we have compiled and briefly discuss all the tools and techniques available for engineering virus resistance in crop plants. Of all the strategies currently available for engineering virus resistance, RNA-interferencebased technology has shown greater success. The recently developed genome editing technology has also given new hope for virus-resistant transgenic crops. Without compromising the safety aspects of these transgenic crops the regulatory procedures followed in the approval of virus-resistant transgenic crops for farmers’ cultivation have to

be refined to ensure that the benefit reaches to the farmers without much delay. Introduction Plant viruses infect almost all the plant species and across the world plant viral diseases are known to cause severe economic losses in cultivated crop plants (Hull, 2014). Plant viruses show humungous diversity in their composition and structure and come in variable shapes and sizes. Viruses of different taxa have different genome composition and organization, the genetic material can be made of DNA or RNA, it may be present as single or double-stranded, linear or circular, monomer or multi-segmented, and the RNA genome can occur as plus sense, negative sense or ambisense (Fauquet et al., 2005). Irrespective of their structural and genome organization, all viruses are obligate, intracellular parasites, which hijack host’s biological machinery to encode their proteins and to proliferate within the plant. Some of the conventional ways to manage plant viral diseases are phytosanitation, change in cropping/cultural practices, employing insecticides to control insect vectors that transmit the viruses, and mild strain protection. However, these conventional methods have not been completely successful in controlling the plant viral diseases and the menace caused by plant viral diseases continues. Of all these virus control strategies, phytosanitation and

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cultivation of virus-resistant crop varieties are the most pragmatic approaches for economic management of plant viral diseases (Kyle, 1993). The virus resistance trait can be governed by genes which are dominant, co-dominant, recessive, or additive in nature (Whitham and Hajimorad, 2016). However evolution of resistant breaking virus strains has already broken down the limited sources of virus resistance that have been introgressed into cultivated crops. Additionally, the virus resistance genes which are recessive in nature or in those cases where resistance is governed by multiple genes are difficult to introgress in cultivated crop varieties through plant breeding technologies. Unlike the fungicides and bactericides that are commercially available for the control of plant pathogens such as fungus and bacteria, there are no viricides developed for the control of viral diseases. The major reason for the absence of a viricide is viruses do not have their own distinct biochemical pathway from their hosts unlike other plant pathogens such as fungi and bacteria. Since viruses hijack host biochemical pathways for its proliferation, any chemical that may be applied to check the viruses may potentially have a negative impact on the host or its biochemical pathways (Lindbo and Falk, 2017). Hence chemical control of insect vectors such as aphids, whiteflies, thrips and leafhoppers which are involved in transmission of plant viruses, through application of pesticides is the most widely followed approach by the farming community (Whitefield et al., 2015). However, these chemicals are not very effective on vectors such as aphids which transmit the viruses in a non-circulative and non-persistent manner, wherein the aphids acquire and transmit viruses in a fraction of time, before the pesticide starts acting on it. Further, many insect vectors have become immune to chemical pesticides and these are no longer effective. Additionally, these carcinogenic pesticides pollute our environment; mainly the water bodies and is the major cause of increase in number of cancer cases. Cross-protection is yet another important virus management strategy which has been successful in selected cases. It is a type of induced resistance which is based on the principal that prior infection with one virus gives protection against a closely related virus. This was first demonstrated for Tobacco mosaic virus (TMV) (McKinney, 1929), later it was commercially employed for management of Citrus

tristeza virus in citrus orchards and was also demonstrated to work against potyviruses such as Papaya ring spot virus (PRSV) and Zucchini yellow mosaic virus (ZYMY) (Ziebell and Carr, 2010). In the last few decades significant advances are made in understanding of the plant viruses, their biology, interaction with host plant and the insect vector, viral pathogenesis and host resistance mechanisms (Cillo and Palukaitis, 2014). Genetic engineering for virus resistance is a highly promising strategy to develop virus-resistant transgenic crops, wherein all the agronomic traits are preserved despite the gain in viral resistance, in contrast to the crop improvement through breeding. The transgenic technologies that are used for engineering virus resistance in crop plants can be broadly classified into pathogen-derived virus resistance (PDR) and plant-derived virus resistance. Pathogen-derived virus resistance is achieved by primarily transferring the virus-derived genes, including viral coat protein, replicase, movement protein, defective interfering RNA, non-coding RNA sequences, and protease, into susceptible plants. The plant derived or non-viral genes, such as R genes, ribosome-inactivating proteins (RIP), protease inhibitors, dsRNAse, RNA modifying enzymes, and scFvs, have also been used successfully to engineer virus resistance in plants (Cillo and Palukaitis, 2014). Technological advances in RNA-interference (RNAi) and Genome editing has ushered a new hope for the management of plant viral diseases. In this chapter we will discuss the principals involved in each of these transgenic strategies and also briefly discuss about the successful application of these tools and techniques to accomplish virus-resistant crop plants. Coat protein-mediated virus resistance The concept of pathogen (or parasite)-derived resistance was first given by Sanford and Johnston in 1985 (Sanford and Johnston, 1985) with the assumption that the coat protein (CP) was the likely determinant of cross-protection. Subsequently work from Dr Roger Beachy’s lab at Washington University (St. Louis, MO, USA) demonstrated that transgenic expression of TMV CP gene in tobacco plants gave significant resistance to TMV infection (Abel et al., 1986). Although none of the

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transgenic tobacco plant lines were immune to TMV infection, many of the lines showed reduced infection and some of them delayed development of symptoms. Further with the increase in the levels of viral inoculum the resistance level was reduced. Additionally the TMV challenge experiments revealed that the resistance was displayed only against the TMV virions, but not against the purified viral RNA or to the virus particles that were partially uncoated (Nelson et al., 1987; Register and Beachy, 1988). This implicated that one of the very early step in virus infection process, such as the uncoating of the virion RNA was affected by overexpression of CP in the host cells (Osbourn et al., 1989; Wisniewski et al., 1990). Polysome-mediated virion disassembly is assumed to be the primary first step in the establishment of TMV infection in the plant cells. Further molecular dissection indicated that the resistance was primarily to virus particles, which likely interfered in the initial events during the viral infection process. Later studies using in vitro encapsidated TMV genomic RNA carrying reporter genes also suggested that virion disassembly was mainly reduced in the genetically engineered plants. Transgenic expression of mutated TMV CP variants, revealed that some of the CP mutants enhanced the CP–CP interactions resulting in increased aggregation of CPs, accompanied by enhanced resistance levels compared to the transgenic tobacco expressing wild type CP (Asurmendi et al., 2004, 2007; Bendahmane et al., 1997; Lu et al., 1998). When these studies were replicated using the CP-transgenic protoplasts there was a drastic reduction in the expression of TMV movement protein, as well as the formation of replication complexes and this had a strong correlation with delay in the appearance of viral symptoms (Asurmendi et al., 2004; Bendahmane et al., 2007). However, it is also possible that other events of the virus infection process could also be affected and this phenomenon was termed ‘Coat Proteinmediated resistance (CPMR)’ (Beachy, 1999). This breakthrough ushered development of virus-resistant transgenics in other crop plants using the CPMR approach. In a span of three years CPMR strategy was evaluated on viruses representing at least five different taxa and varying resistance levels were observed (Beachy et al., 1990). Transgenic studies against Cucumber mosaic virus (CMV) and Potato virus Y and Potato leaf roll virus (PLRV)

showed that the levels of resistance was not directly correlated with the expression levels of corresponding viral CPs (Cuozzo et al., 1988; Quemada et al., 1991; Lawson et al., 1990; Kawchuk et al., 1990). The first virus-resistant transgenic crop to be deregulated and commercialized is squash (Cucurbita pepo) in United States in the 1990s (Tricoli et al., 1995). Squash is severely affected by CMV and at least two other potyviruses, namely Zucchini yellow mosaic virus (ZYMV) and Watermelon mosaic virus (WMV) and multiple virus-resistant squash could not be obtained through conventional breeding efforts (Arce-Ochoa et al., 1995). Transgenic squash was developed through Agrobacterium mediated transformation of a construct with a promoter driving the CP genes from ZYMV, WMV and CMV and the transgenic plants obtained were resistant to all the three viruses (Tricoli et al., 1995). Field trials of this virus-resistant transgenic squash resulted in 50-fold higher yields than the corresponding non-transgenic squash (Fuchs et al., 1998). Virus-resistant transgenic squash accounted for 12% of the total squash produced in the USA, in 2006, there was an estimated profit of US$24 million ( Johnson et al., 2007). Until now this transgenic squash cultivated in the US and still shows significant multi-virus resistance (Lindbo and Falk, 2017). Similarly transgenic potato overexpressing PVY CP showed high level of resistance to PVY (Kaniewski et al., 1990). However potato plants transgenic for CP of Potato leaf roll virus (PLRV, genus Polerovirus) failed to show virus resistance in the field trials, but resistance was accomplished when the replicase gene was overexpressed (Kaniewski and Thomas, 2004). Development of virus-resistant transgenic papaya is the most successful example which used CPMR strategy and this is discussed in a dedicated chapter on virusresistant transgenic papaya of this book and more details can also be found in Fitch (2016), Fuchs and Gonsalves (2007, 2015), and Kumari et al. (2015). Plum is another important fruit tree cultivated in temperate countries of Americas and Europe which is severely affected by Sharka disease, caused by Plum pox virus (PPV, genus: Potyvirus), which is transmitted by the aphids (García et al., 2014). After the first report of Sharka disease from Bulgaria in 1900s, epidemics of this disease have been reported from Spain, USA and Canada, where in

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rouging of millions of infected plum trees has been done to eradicate PPV (Cambra et al., 2006; Scorza et al., 2013). The US Department of AgricultureAgricultural Research Service (USDA-ARS) has developed a transgenic plum variety ‘Honeysweet’, which was deregulated after 16 years of greenhouse screening and field evaluation (Scorza et al., 2013). However, until now the virus-resistant transgenic plum is not cultivated anywhere in the world (Lindbo and Falk, 2017). Other than the United States, China and Brazil are the only countries that have allowed cultivation of virus-resistant transgenic plants ( James, 2009; Tollefson, 2011). Interference from heterologous viral genes Replicase mediated virus resistance After the success obtained in achieving virus resistance by transgenically expressing the viral CP, researchers pondered if other viral genes would be equally or more effective than the CP gene, thus contributing for the pathogen-derived resistance. In one of the first such experiment, when the 54-kDa RNA-dependent RNA polymerase (RdRp or Replicase) of TMV was transformed into tobacco, a very high level of resistance was obtained, although the expressed Rep protein remained undetected (Golemboski et al., 1990). Soon there was a rush to achieve ‘replicase-mediated resistance’ against viruses of other taxa (Anderson et al., 1992). Significant resistance was accomplished when the replicase of Pea early browning virus, PVY and CMV were transgenically expressed. Different versions of the replicase genes, such as full length, truncated and mutants were screened for transgenic resistance to corresponding viruses. Resistance was achieved in all the versions of replicase, implying that the replicase-based resistance did not require expression of a functional protein or the protein as such. This led to a hypothesis that the observed Rep-based resistance could be RNA-mediated, which is most likely to be post-transcriptional gene silencing (PTGS), discussed in later sections of this chapter (Marano and Baulcombe, 1998; Mueller et al., 1995). The Rep mediated resistance was confined to a narrow spectrum of plant viruses and was not as effective as CPMR. However, the plant infecting

single-stranded DNA viruses, such as the geminiviruses, which do not encode polymerases, but their replication-associated protein (Rep or AC1) interacts with host encoded polymerases to carry out replication of the viral genome. Transgenic expression of Rep gene of geminiviruses, such as African Cassava mosaic virus (ACMV), inhibited the replication and proliferation of viral DNA and there was a positive correlation between the amount of Rep transcript and the level of viral resistance (Hong and Stanley, 1995). The truncated 210 amino acid long Rep protein strongly inhibited the virus replication in protoplasts and triggered resistance when expressed at high levels, which was in contrast to the expression of a dominant negative mutant of Tomato yellow leaf curl Sardinia virus (TYLCSV) (Noris et al., 1996; Brunetti et al., 1997). This dominant negative mutant of Rep, which lacks a conserved NTP-binding domain, works by inhibiting the expression of the viral Rep protein and by forming non–functional complexes with the native Rep protein encoded by the virus (Lucioli et al., 2003). Similarly Rep proteins mutated in the orior NTP-binding sites were engineered to produce transgenic plants resistant to other geminiviruses, such as Bean golden mosaic virus (BGMV; Hanson and Maxwell, 1999). Movement protein mediated virus resistance Plant viruses encode specialized proteins termed ‘movement proteins’ (MPs) that help cell-to-cell movement of the virions or its ribonucleoprotein complex. The successful interaction between the movement proteins and the plasmodesmata proteins is crucial for the formation of tubules to allow intercellular trafficking of virions and/or ribo–nucleoprotein complexes comprising viral RNA along with one or more of the virus-encoded proteins. The movement proteins bind to the viral genome (DNA or RNA) through their nucleic acid binding domains and they also have a different domain for modification of the host plant plasmodesmata. Mutations in the DNA/RNA binding domain of MPs allow their interaction with the plasmodesmata; however, they interfere with native MP-mediated plasmodesmatal trafficking of viral DNA/RNA. Application of such strategies can help in achieving broad spectrum resistance to diverse range of plant viruses that rely upon

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similar plasmodesmata components to establish infections. Transgenic expression of 30-kDa defective movement protein of TMV could not only show resistance to the homologous virus, but also to other tobacco infecting viruses such as Alfalfa mosaic virus (Family: Bromoviridae), Peanut chlorotic streak virus (Family: Caulimoviridae), Tobacco rattle virus, Tobacco ringspot virus (Family: Comoviridae), and CMV (Cooper et al., 1995; Lapidot et al., 1993; Malyshenko et al., 1993). Similarly transgenic expression of 12-kDa MP of PVX in potato (Seppanen et al., 1997), and 13-kDa MP of White clover mosaic virus (WClMV) in the model host Nicotiana benthamiana, resulted in the resistance to the corresponding viruses, as well as resistance to viruses of other genera and/or families in certain cases (Beck et al., 1994; Cooper et al., 1995). However overexpression of the 12 kDa MP of PVX in tobacco resulted in resistance to PVX alone and not to other viruses such as TMV and PVY (Kobayashi et al., 2001). In contrast to the plasmodesmata-modifying movement proteins, the viruses belonging to Caulimoviridae and Comoviridae families code for a protein that forms tubules which help in the trafficking of virions from virus-infected cells to the neighbouring cells. Transgenic expression of wild-type MP of Cowpea mosaic virus (CPMV) in N. benthamiana conferred a very specific resistance to different strains of CPMV, but not to other members of Comovirus genus (Sijen et al., 1995), while expression of wild-type MP of Potato leaf roll virus (PLRV) in potato delivered broad-spectrum resistance to PLRV, PVY, and PVX (Tacke et al., 1996). Overexpression of truncated MPs of plantinfecting begomoviruses (Family: Geminiviridae) with single-stranded circular DNA as their genome resulted in significant resistance to both homologous and heterologous viruses (Duan et al., 1997; Hou et al., 2000). However, their resistance in the real field situation is yet to be confirmed and any undesirable effects on the plant due to the interference by MPs in plant communication is not ruled out. RNA-mediated virus resistance While a large number of labs were working for protein-mediated virus resistance, there were parallel efforts being made to evaluate the efficacy of RNA mediated resistance to plant viruses. To understand

the basis of resistance the native or modified forms of viral sequences were transgenically expressed as RNA, either in sense or antisense orientation, without getting translated into a protein. The very first evidence on RNA mediated resistance was shown by Lindbo and Dougherty (1992a,b), when they overexpressed non-translatable TEV CP sequence. When challenged by TEV about 30% of the transgenic plant lines showed a TEV-immune phenotype (Lindbo et al., 1993). Molecular analysis revealed that there was very high expression of the transgene RNAs and the TEV RNAs were not capable of replicating in the cells of these resistant transgenic plants and further the resistance could not be overcome even by very high levels of viral inoculum (Dougherty et al., 1994; Lindbo et al., 1993). This resistance was hypothesized to be through post-transcriptional gene silencing (PTGS) and a model for PTGS was proposed by Lindbo et al. (1993). This generated significant curiosity among the biologists and further lead to the prediction that the small RNAs generated from the aberrant and overexpressed transgenes were part of the sequence-specific RNA targeting and degradation (Dougherty and Parks, 1995). Subsequently research in the area of PTGS in both plant and animal systems helped in thorough understanding and fine tuning of the PTGS phenomenon. Additionally the expression of natural or artificial Defective Interfering (DI) RNAs or DNAs (Frischmuth and Stanley, 1994; Stenger, 1994; Patil and Dasgupta, 2006), and the use of satellite RNAs (satRNAs) were also explored to achieve virus resistance in plants (Kim et al., 1997; Kim et al., 1995). As the understanding of the RNA-interference (RNAi) phenomenon improved, RNAi was employed with various modifications and improvements through production of different versions of small RNAs, such as siRNAs (small interfering RNAs), amiRNAs (artificial microRNAs), shRNAs (short hairpin RNAs), and tasiRNAs (trans-acting siRNAs), to accomplish virus resistance through homology dependent gene silencing or post-transcriptional gene silencing (PTGS). Readers are requested to refer to Lindbo and Dougherty (2005) for a more detailed review on the history of PTGS, RNA silencing of transgenes and virus resistance. Our current understanding of RNAi provides new perspective on the old experimental results. The immunity of the

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plants expressing non-translatable TEV CP can be attributed to RNAi which probably is an outcome of dsRNA formation (Lindbo et al., 1993). These dsRNA molecules are processed by series of enzymes such as DCLs and the AGO proteins of the RISC complex (Bernstein et al., 2001; Hammond et al., 2000; Liu et al., 2004). Re-investigation of ‘CPMR’ in the deregulated virus-resistant transgenic papaya and plum revealed that the part of the resistance could be an RNA-mediated resistance (Scorza et al., 2001; Tennant et al., 2001). Additionally, some of the ‘replicase-mediated resistance’ have also been attributed to RNAi (Marano and Baulcombe, 1998; Vazquez et al., 2001), although some of the replicase-mediated resistance is still protein mediated, perhaps this is in addition to RNAi (Donson et al., 1993; Prins et al., 2008). Later, there have been significant numbers of RNAi-based transgenics developed by transgenically expressing hairpin RNA (Helliwell and Waterhouse, 2005; Mlotshwa et al., 2008) or double-stranded RNA (hpRNA or dsRNA) (Tenllado et al., 2004; Waterhouse and Fusaro, 2006). RNAi can be induced by expression of an antisense RNA (asRNA) or a double-stranded RNA (dsRNA) or hairpin RNA (hpRNA). The host gene silencing machinery checks both RNA and DNA viruses as a function of plant’s innate immunity. However, the viruses have evolved a special class of proteins that function as silencing suppressors to counteract the induced RNA silencing in their host plants (Vanitharani et al., 2005; Bisaro, 2006). Several studies have revealed that the expression of dsRNA is significantly more efficient trigger of RNAi than the use of asRNA. This is mainly because of instantaneous production of siRNA by the dsRNAs, which trigger the RNAi system even in the absence of a virus, the transcripts of which are substrates for the Dicer-like (DCL) proteins (Waterhouse et al., 1998; Fusaro et al., 2006). Further through transient agroinfiltration studies of RNAi constructs targeting different reporter genes, it has been shown that both the light intensity and the temperature affect the systemic spread of silencing signal across the plant (Patil and Fauquet, 2015). RNAi has been used to develop transgenic resistance to Cucumber mosaic virus, Zucchini yellow mosaic virus and Watermelon mosaic virus 2 (Klas et al., 2006; Tricoli et al., 1995), Potato leaf roll virus, Potato virus Y and Potato virus X (Thomas et al., 2000), Papaya ring spot virus

(Krubphachaya et al., 2007), Plum pox virus (Hily et al., 2004; Kundu et al., 2008), Cassava mosaic and brown streak viruses (Patil et al., 2011, 2016; Yadav et al., 2011; Ogwok et al., 2012) and geminiviruses infecting legumes (Pooggin et al., 2003; Kumar et al., 2017). Some of these virus-resistant transgenic works are discussed in greater detail in various chapters of this book. Exogenous application of dsRNA by coating it on nano-sheets is also getting major attention as a novel strategy for plant protection and this is discussed elaborately in a dedicated chapter of this book (Voloudakis et al., 2015; Mitter et al., 2017a,b). After successful implementation of RNAi to accomplish resistance to plant viruses, further medications/improvements were made in this technology to achieve resistance by expression of artificial microRNAs (amiRNAs) (Niu et al., 2006). Since the first demonstration of amiRNA-based virus resistance, this strategy has been successfully employed against cucumoviruses, potex viruses, ipomoviruses, begomoviruses, and tospoviruses (Ai et al., 2011, 2013; Duan et al., 2008; Mitter et al., 2016; Qu et al., 2007; Wagaba et al., 2016; Zhang et al., 2011). This book consists of a dedicated chapter on ‘Management of geminiviruses focusing on small RNAs in tomato’ which discusses on the use of amiRNA and tasiRNA-based resistance strategies for the transgenic control of geminiviruses. Our lab in collaboration with others, with funding from Dept. of Biotechnology (DBT, Govt. of India) is currently working on a novel smallRNA-based technology called as ‘miRNA-Induced Gene Silencing’ (MIGS), which is primarily based on the ability of miR173 to trigger the generation of secondary siRNAs (tasiRNAs) from its target sequences (Felippes et al., 2012). MIGS involves targeting the miR173 to a viral sequence, for which silencing is desired and this is achieved by addition of the miRNA target site in the immediate upstream of the viral/gene sequence of interest (Felippes, 2013). Genome editing as a tool for engineering virus resistance The recent advent of genome editing technologies has revolutionized crop improvement through genetic engineering. Through genome editing one can directly edit or introduce the desired alleles, by inducing desired mutations in the plant’s genome

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through different molecular tools and techniques (Hsu et al., 2014). This involves nuclease mediated double-stranded DNA break in the targeted genome sequence of an organism followed by DNA repair, either through homology-directed repair (HDR) or by non-homologous end joining (NHEJ), thus introducing modification at a target site (Gorbunova and Levy, 1997; Puchta et al., 1996). Some of the popular genome editing tools used until recently are, zinc finger nucleases (ZFNs) technology and transcription activator-like effector nucleases (TALENs) which mediate target sequence editing through DNA-protein based interaction (Christian et al., 2010; Kim et al., 1996). However, due to less specificity of recognition between the protein residues and the target DNA sequence, ZFNs and TALENs are not widely used (Belhaj et al., 2013). Subsequently a new method based on clustered regularly interspaced short palindromic repeats/CRISPR-associated (CRISPR/ Cas) has been developed which is guided by short RNAs to accurately carry out targeted changes in the genome of an organism (Cong et al., 2013; Mali et al., 2013). From the time of its discovery in 2013, the RNA-guided gene/genome editing using the CRISPR-Cas9 derived from the bacterium Streptococcus pyogenes has revolutionized the genome editing technology in crop plants. The native function of the CRISPR-Cas9 in a bacterium is to provide immunity against the invading phages and conjugative plasmids. The CRISPR-Cas9 creates Non-Homologous End Joining repair of the double-strand breaks in DNA, resulting in insertions or deletions of few base pairs in the target sequence. CRISPR-Cas9 based genome editing has been demonstrated in various plant families such as Solanaceae, Cruciferae, Poaceae and Fabaceae and it is also being explored to develop personalized therapies for individual patients with HIV-1 variants (Hu et al., 2014). Majority of plant virus resistance genes are recessive in nature, including the eukaryotic translation initiation factors eIF4E (Kang et al., 2005; Maule et al., 2007; Palukaitis and Carr, 2008). A complex of such initiation factors and other host factors bind to the viral 5′ cap structure and the 3′ poly-A tail of mRNA for translation. The copy numbers of the eIF4E gene and its isoforms vary among plant species. Potyviruses in particular, associate with these proteins, through the viral-encoded protein (VPg)

bound to the 5′ end of the viral RNA (Robaglia and Caranta, 2006). Broad spectrum virus resistance has been displayed by silencing the eIF4E gene in tomato and melon (Chandrasekaran et al., 2016). A broad spectrum resistance was obtained when the T3 generation homozygous plants with deletions in the eIF4E gene were challenged with diverse viruses of the Potyviridae family such as Cucumber vein yellowing virus, Zucchini yellow mosaic virus and Papaya ring spot mosaic virus-W (Chandrasekaran et al., 2016). In addition to the control of plant infecting RNA viruses, the plant DNA viruses, mainly the geminiviruses such as Tomato yellow leaf curl virus (TYLCV), Cabbage leaf curl virus (CaLCuV), Cotton leaf curl Kokhran virus (CLCuKV) and Beet severe curly top virus (BSCuTV) have also been controlled by employing CRISPR/Cas9 technology (Ali et al., 2015, 2016; Yin et al., 2015; Ji et al., 2015). However in this case the sgRNAs had sequence identity with the geminiviral genome sequence, in contrast to the host plant genes in the case of potyviruses. Thus by using an array of sgRNAs, with homology to diverse viral sequences, the CRISPR/ Cas9 technology can be used for control of multiple and diverse viruses infecting crop plants. Virus resistance through nonviral sources Employing host genes for virus resistance Plants respond to an attack by a pathogen through a range of molecular mechanisms, some of these mechanisms are the first basal defence response, while others are more specific in nature (Dangl and Jones, 2001). Several genes regulating multiple pathways have been shown to have role in manifestation of these defence mechanisms. Plants naturally encode for resistance gene/s in response to an invasion by a pathogen. An incompatible reaction in a specific plant resistance gene and viral avirulence gene combinations result in a hypersensitive reaction (HR) in which case the adjacent cells undergo programmed cell death, ultimately resulting in isolation of viral infections. These genes are broadly classified into three different categories: dominant resistance (R) genes, recessive resistance genes, and transcription factors (TFs) (Maule et al., 2007; Palukaitis and Carr, 2008). The translational

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products of R genes recognize specific effectors of the pathogens termed as avirulence proteins encoded by avr genes, complying the gene-for-gene hypothesis. Majority of the R genes have nucleotide-binding leucine rich repeat (NB-LRR) and extracellular LRR resistance proteins. The members of NB-LRR can possess either an amino-terminal coiled-coil (CC) or Toll/interleukin-1 receptor (TIR) domains. The structural and functional characteristics of the R-genes are extensively reviewed in past publications (Belkhadir et al., 2004; Chisholm et al., 2006; Dangl and Jones, 2001). The first R gene to be identified and characterized was in Nicotiana glutinosa against TMV which was termed as ‘N’ gene, coding for TIR-NB-LRR class protein. The Hypersensitive reaction (HR) was the outcome of resistant phenotype resulting in necrotic spots on the inoculated leaves. Transgenic expression of the N gene in tomato gave HR phenotype, thus restricting the infection of TMV and ToMV to the initial sites of infection and there was no systemic spread of the virus. Two CC-NBS-LRR class R genes, Tm22 and Sw5, are extensively introgressed in commercial tomato varieties for resistance to ToMV and TSWV, respectively. When these R genes were transgenically expressed in tobacco, both the genes resulted in a resistant phenotype similar to that observed in the source host (Lanfermeijer et al., 2004; Spassova et al., 2001). In at least one case, transgenic expression of the non-viral R gene Prf R gene in tomato against Pseudomonas syringae pathovar tomato resulted in tolerance to a range of bacterial pathogens, as well as to TMV, through activation of systemic acquired resistance (Oldroyd and Staskawicz, 1998). Recessive resistance genes are actually the mutant alleles of the susceptibility genes that encode for proteins that are indispensable for viral attack. The plants carrying these alleles have to be homozygous to exhibit the resistance phenotype. The recessive R genes exhibiting resistance to plant viruses are mostly the Eukaryotic translation initiation factors (eIFs), and particularly the eIF4E and eIF4G protein families (Kang et al., 2005; Maule et al., 2007; Palukaitis and Carr, 2008). Most of the recessive R genes identified are against the potyviruses, which rely upon the host eIFs for different functions in their replication and translation (Robaglia and Caranta, 2006). Besides the dominant and recessive resistance genes, a third category of host/plant genes that have

been used for generating virus resistance plants are the Transcription Factors (TFs), which are the master regulators for the gene expression of single or a cluster of genes (Dai et al., 2008). The major function of TFs is to fine-tune the response of the host plants against different environmental stresses. In this chapter it will not be discussed in greater detail, since Chapter 2 is dedicated to the role of host transcription factors in modulating defence response during plant–virus interaction. Resistance to infections of tobamoviruses such as TMV and ToMV in tobacco and PMMoV in hot pepper has been achieved by overexpression of different members of ethylene response factors (ERFs), which are mostly associated with defence response pathways through ethylene production (Cao et al., 2005; Fischer and Dröge-Laser, 2004; Shin et al., 2002; Zhang et al., 2009). In certain cases, in addition to broad-spectrum resistance to diverse pathogens, tolerance to some abiotic stresses has also been reported by overexpression of TFs of ERF family (Cao et al., 2005). Broad-spectrum virus resistance has been accomplished by antisense inhibition of a host gene coding for S-adenosylhomocysteine hydrolase (SAHH), which is critical for a number of S-adenosylmethionine-dependent reactions, including the capping of the 5′-end of viral RNAs during replication (Masuta et al., 1995). However, the majority of the SAHH-expressing transgenic plants showed abnormal growth, as a result of enhanced cytokinin activity, in the absence of virus infection. Lectins, proteins that reversibly bind carbohydrates, have also been reported to exhibit virus resistance when transgenically expressed (Yamaji et al., 2012). Lectins exist in most living organisms; however their biological significance is not well understood. A large number of plant lectins are induced under different biotic and abiotic stresses and show antibacterial, antifungal, and anti-insect activities, implying that plant lectins have defensive roles (Van Damme et al., 2004). There are at least two lectins RTM1 and JAX1 (jacalin-type lectin), that are known to show resistance to potyviruses and potexviruses when transgenically expressed in plants (Chisholm et al., 2000; Yamaji et al., 2012). Ribosome-inactivating proteins Ribosome-inactivating proteins (RIPs) are primarily the RNA N-glycosidases that specifically remove

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the purine base at A4324 of 28S rRNA, resulting in detachment of the 3′-end of the substrate RNA, and hence making rRNA a non-functional entity in the protein translation. RIPs accumulate mainly in the seeds and various other parts of the plants. Transgenic expression of several RIPs, such as the pokeweed (Phytolacca americana) antiviral protein (PAP), have been studied for their ability to check virus infection. Overexpression of one of the three PAPs from P. americana, showed broad spectrum resistance to PVX in tobacco and potato, and to PVY in tobacco, potato, and N. benthamiana (Lodge et al., 1993). Transgenic overexpression of PAP in Brassica napus provided resistance to TuMV (Zhang et al., 1999). As the wild-type PAP affects the transformation efficiency and the plant development, several mutants of PAP were used in most transgenic events to minimize side effects on the host plant (Lodge et al., 1993; Tumer et al., 1997). Similarly, transgenic expression of a RIP from Phytolacca insularis in potato showed certain level of protection against PVX, PVY, and PLRV (Moon et al., 1997). Transgenic expression of dianthin, a RIP from Dianthus caryophyllus, in N. benthamiana using a promoter from ACMV AV1 (which is transactivated by the AC2), showed tolerance to ACMV, but not to other geminiviruses (Hong et al., 1996). However, the mechanism of action of RIPs is not properly understood, although they are presumed to have both direct effects on the viral RNAs, as well as induction of host defence response (Wang and Tumer, 2000). Further, the different levels of resistance observed by use of different RIPs in different plant species against different viruses was not consistent and hence was not promising for their large-scale application and commercialization. Virus resistance through protease inhibitors from plants Anti-viral plantibodies One of the novel approaches used for transgenic control of plant viruses is by expression of antibodies, popularly used for recognition of animal pathogens. Unlike the immune system of animals, the plants do not possess these antibodies; however the affinities of selected antibodies can be good enough to disrupt essential functions of a viral protein in plants. Although the production of

single-chain variable fragment (scFv) antibodies is technically difficult, the significant advancements in the phage display approach and generation of synthetic scFv libraries have made this strategy more promising (Ziegler and Torrance, 2002). Initially it was shown that both polyclonal antibodies (Düring et al., 1990; Hiatt et al., 1989) and scFv antibodies (Firek et al., 1993; Owen et al., 1992) could be expressed stably in transgenic tobacco plants. Subsequently the first successful demonstration of an antibody-mediated resistance to a plant virus was made by transgenic expression of a scFv against Artichoke mottled crinkle virus (AMCV) in N. benthamiana plants (Tavladoraki et al., 1993). However the levels of resistance obtained were not significant, the resistant phenotype was mostly delay in symptom expression or reduced levels of viral infection. Subsequent to this not much progress was made, as a number of questions remained unanswered, such as the choice of the appropriate target in the virus, the levels of scFv protein expression required, and stability of the expressed proteins (Nölke et al., 2004). With the increase in the understanding of the structure of antibodies, their stability was improved and new protein scaffolds were designed that could be employed as protein-binding alternatives to antibodies, to overcome the deficiencies of scFvs. Later the level of resistance was enhanced, through improvements in plantibody stability and their targeting, and also plantibodies were generated against non-structural viral proteins, such as replication-associated/protease proteins (Gargouri-Bouzid et al., 2006; Nickel et al., 2008) and viral replicases (Boonrod et al., 2004; Gil et al., 2011). Systemic acquired resistance In response to the viral infections plant show hypersensitive reaction (HR), which in the beginning is restricted to the sites of viral infection and later it spreads systemically. This resistance reaction is referred as systemic acquired resistance (SAR), which is characterized by coordinated activation of cluster of genes in the distal parts of the plant. Once SAR is triggered, there is accumulation of salicylic acid (SA), resulting in increased expression of pathogenesis related (PR) proteins and the activation of phenylpropanoid pathway which leads to accumulation of phenolic compounds, enhanced production of active oxygen species and deposition

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of lignin and suberin is reinforced in the cell walls. The role of SA in imparting resistance to TMV has been proved by transgenically overexpressing bacterial salicylate hydroxylase (NahG) gene in tobacco, thus decreasing its endogenous salicylic acid and hence resulting in its susceptibility to TMV (Delaney, 1994). SA binds to the catalase enzyme, mainly involved in degradation of hydrogen peroxide (H2O2), which subsequently leads to enhanced accumulation of H2O2. Severe reduction in catalase activity of approx. 90% was achieved by transgenic antisense expression of catalase-1 or catalase-2 genes, which further resulted in chlorosis or necrosis in the lower leaves (Takahashi et al., 1997). These transgenic tobacco plants displayed enhanced accumulation SA and PR, ultimately leading to increased resistance to TMV. Conclusion In contrast to the chemical control as a major option for control of pests and pathogens, the only option available for management of plant viral diseases is genetic improvement. Although crop improvement through plant breeding is an important option for introgressing the virus resistance trait in the target plant genotype, it has been a hard task to identify appropriate sources of virus resistance that could be used in breeding programme without compromising with other agronomic/yield/quality traits. Hence genetic engineering remains as the most promising option for the management of plant viral diseases. Of all the genetic engineering technologies described in this chapter, RNA-interference technology is the most promising tool to accomplish virus resistance. Majority of the published reports of transgenic virus resistance have used model hosts plants, such as Arabidopsis and N. benthamiana. With world population projected to reach 9.8 billion by the year 2050, the scientific community is hard pressed to translate their research findings to field applications, for the benefit of the farmers and the humankind at large. Hence it is high time to employ the proven and promising genetic engineering strategies in the field crops. One of the major hurdles in developing virus-resistant transgenic field crops is non-availability of established and reproducible tissue culture and transformation technology. Although some progress has been made in development of virus-resistant transgenic

crops, screening of these transgenic plant lines has either ended up in the lab or the greenhouses, without reaching the farmers’ fields. Hence intensive and aggressive efforts are required to generate large number of transgenic plant lines, so that the best one can be identified for better performance in the field scenario. It is also important to target multiple viruses to have broad-spectrum and durable field resistance. This can only be achieved if one can have sound data on the diversity of plant viruses infecting a crop species in a particular geographical region. Other factors that are limiting to successful introduction and cultivation of transgenic field or horticultural crops are intellectual property rights (IPR), poor bio-safety regulations, high cost of transgenic crop development and negative campaigns by some of the activists. Development of novel, acceptable and cost effective tools and techniques should be the priority, along with partnership between industries and academic institutes to access patented technologies at a reasonable price. It is also important to enhance the financial support for production and deregulation of transgenic crops, which includes the creation of appropriate infrastructure for implementation of biosafety protocols in order to enable the farmers to obtain a major share of the gains from these transgenic crops developed by industries. Several virus-resistant transgenic crops are in the pipeline for deregulation and their early release will facilitate affordable and secure desperately required food supply in the developing and underdeveloped regions of the globe. Acknowledgements BLP would like to acknowledge Donald Danforth Plant Science Center, St. Louis (USA), Indian Council of Agricultural Research and Department of Biotechnology (India) for the opportunity and funding to carry out research work on virusresistant transgenic crops during his tenure as Post-Doctoral Fellow and Senior Scientist. References Abel, P.P., Nelson, R.S., De, B., Hoffmann, N., Rogers, S.G., Fraley, R.T., and Beachy, R.N. (1986). Delay of disease development in transgenic plants that express the Tobacco mosaic virus coat protein gene. Science 232, 738–743. Ai, T., Zhang, L., Gao, Z., Zhu, C.X., and Guo, X. (2011). Highly efficient virus resistance mediated by artificial

Genetic Engineering for Virus Resistance in Plants |  113

microRNAs that target the suppressor of PVX and PVY in plants. Plant Biol. 13, 304–316. https://doi. org/10.1111/j.1438-8677.2010.00374.x Ali, I., Amin, I., Briddon, R.W., and Mansoor, S. (2013). Artificial microRNA-mediated resistance to the monopartite begomovirus Cotton leaf curl Burewala virus. Virol. J. 10, 231. https://doi.org/10.1186/1743422X-10-231 Ali, Z., Abulfaraj, A., Idris, A., Ali, S., Tashkandi, M., and Mahfouz, M.M. (2015). CRISPR/Cas9-mediated viral interference in plants. Genome Biol. 16, 238. https:// doi.org/10.1186/s13059-015-0799-6 Ali,Z.,Ali,S., Tashkandi, M., Zaidi, S.S., and Mahfouz, M.M. (2016). CRISPR/Cas9-Mediated Immunity to Geminiviruses: Differential Interference and Evasion. Sci. Rep.6, 26912. https://doi.org/10.1038/srep26912 Anderson, J.M., Palukaitis, P., and Zaitlin, M. (1992). A defective replicase gene induces resistance to Cucumber mosaic virus in transgenic tobacco plants. Proc. Natl. Acad. Sci. U.S.A. 89, 8759–8763. Arce-Ochoa, J.P., Dainello, F., Pike, L.M., and Drews, D. (1995). Transgenic summer squash hybrids and their parental hybrid line. Hort. Science 30, 492–493. Asurmendi, S., Berg, R.H., Koo, J.C., and Beachy, R.N. (2004). Coat protein regulates formation of replication complexes during Tobacco mosaic virus infection. Proc. Natl. Acad. Sci. U.S.A. 101, 1415–1420. https://doi. org/10.1073/pnas.0307778101 Beachy, R.N. (1999). Coat-protein-mediated resistance to Tobacco mosaic virus: discovery mechanisms and exploitation. Philos. Trans. R. Soc. Lond., B, Biol. Sci. 354, 659–664. https://doi.org/10.1098/rstb.1999.0418 Beachy, R.N., Loesch-Fries, S., and Tumer, N. (1990). Coat protein-mediated resistance to virus infection. Annu. Rev. Phytopathol. 28, 451–474. Beck, D.L., Van Dolleweerd, C.J., Lough, T.J., Balmori, E., Voot, D.M., Andersen, M.T., O’Brien, I.E., and Forster, R.L. (1994). Disruption of virus movement confers broad-spectrum resistance to systemic infection by plant viruses with a triple gene block. Proc. Natl. Acad. Sci. U.S.A. 91, 10310–10314. Belhaj, K., Chaparro-Garcia, A., Kamoun, S., and Nekrasov, V. (2013). Plant genome editing made easy: targeted mutagenesis in model and crop plants using the CRISPR/Cas system. Plant Methods 9, 39. https://doi. org/10.1186/1746-4811-9-39 Belkhadir, Y., Subramaniam, R., and Dangl, J.L. (2004). Plant disease resistance protein signaling: NBS-LRR proteins and their partners. Curr. Opin. Plant Biol. 7, 391–399. https://doi.org/10.1016/j.pbi.2004.05.009 Bendahmane, M., Chen, I., Asurmendi, S., Bazzini, A.A., Szecsi, J., and Beachy, R.N. (2007). Coat proteinmediated resistance to TMV infection of Nicotiana tabacum involves multiple modes of interference by coat protein. Virology 366, 107–116. Bernstein, E., Caudy, A.A., Hammond, S.M., and Hannon, G.J. (2001). Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409, 363– 366. https://doi.org/10.1038/35053110 Bisaro, D.M. (2006). Silencing suppression by geminivirus proteins. Virology 344, 158–168. Boonrod, K., Galetzka, D., Nagy, P.D., Conrad, U., and Krczal, G. (2004). Single-chain antibodies against a

plant viral RNA-dependent RNA polymerase confer virus resistance. Nat. Biotechnol. 22, 856–862. https:// doi.org/10.1038/nbt983 Brunetti, A., Tavazza, M., Noris, E., Tavazza, R., Caciagli, P., and Ancora, G., et al. (1997). High expression of truncated viral Rep protein confers resistance to Tomato yellow leaf curl virus in transgenic tomato plants. Mol. Plant-Microbe Interact. 10, 571–579. Cambra, M., Capote, N., Myrta, A., and Llacer, G. (2006). Plum pox virus and the estimated costs associated with sharka disease. EPPO Bull. 36, 202–204. Cao, Y., Wu, Y., Zheng, Z., and Song, F. (2005). Overexpression of the rice EREBP-like gene OsBIERF3 enhances disease resistance and salt tolerance in transgenic tobacco. Physiol. Mol. Plant Pathol. 67, 202–211. Chandrasekaran, J., Brumin, M., Wolf, D., Leibman, D., Klap, C., Pearlsman, M., Sherman, A., Arazi, T., and GalOn, A. (2016). Development of broad virus resistance in non-transgenic cucumber using CRISPR/Cas9 technology. Mol. Plant Pathol. 17, 1140–1153. https:// doi.org/10.1111/mpp.12375 Chisholm, S.T., Coaker, G., Day, B., and Staskawicz, B.J. (2006). Host-microbe interactions: shaping the evolution of the plant immune response. Cell 124, 803–814. Chisholm, S.T., Mahajan, S.K., Whitham, S.A., Yamamoto, M.L., and Carrington, J.C. (2000). Cloning of the Arabidopsis RTM1 gene, which controls restriction of long-distance movement of tobacco etch virus. Proc. Natl. Acad. Sci. U.S.A. 97, 489–494. Christian, M., Cermak, T., Doyle, E.L., Schmidt, C., Zhang, F., Hummel, A., Bogdanove, A.J., and Voytas, D.F. (2010). Targeting DNA double-strand breaks with TAL effector nucleases. Genetics 186, 757–761. https://doi. org/10.1534/genetics.110.120717 Cillo,F., and Palukaitis, P. (2014). Transgenic resistance. Adv. Virus Res. 90, 35–146. https://doi.org/10.1016/ B978-0-12-801246-8.00002-0 Cong, L., Ran, F.A., Cox, D., Lin, S., Barretto, R., Habib, N., Hsu, P.D., Wu, X., Jiang, W., Marraffini, L.A., et al. (2013). Multiplex genome engineering using CRISPR/ Cas systems. Science 339, 819–823. https://doi. org/10.1126/science.1231143 Cooper, B., Lapidot, M., Heick, J.A., Dodds, J.A., and Beachy, R.N. (1995). A defective movement protein of TMV in transgenic plants confers resistance to multiple viruses whereas the functional analog increases susceptibility. Virology 206, 307–313. Cuozzo, M., O’Connell, K.M., Kaniewski, W., Fang, R., Chua, N.H., and Tumer, N. (1988). Viral protection in transgenic tobacco plants expressing the Cucumber mosaic virus coat protein or its antisense RNA. Nat. Biotechnol. 6, 549–557. Dai, S., Wei, X., Alfonso, A.A., Pei, L., Duque, U.G., Zhang, Z., Babb, G.M., and Beachy, R.N. (2008). Transgenic rice plants that overexpress transcription factors RF2a and RF2b are tolerant to rice tungro virus replication and disease. Proc. Natl. Acad. Sci. U.S.A. 105, 21012–21016. https://doi.org/10.1073/pnas.0810303105 Dangl, J.L., and Jones, J.D. (2001). Plant pathogens and integrated defence responses to infection. Nature 411, 826–833. https://doi.org/10.1038/35081161

114  | Patil

de Felippes, F.F. (2013). Down-regulation of plant genes with miRNA-induced gene silencing. Methods Mol. Biol. 942, 379–387. https://doi.org/10.1007/978-162703-119-6_20 de Felippes, F.F., Wang, J.W., and Weigel, D. (2012). MIGS: miRNA-induced gene silencing. Plant J. 70, 541–547. https://doi.org/10.1111/j.1365-313X.2011.04896.x Delaney,T.P., Uknes, S., Vernooij, B., Friedrich, L., Weymann, K., Negrotto, D., Gaffney, T., Gut-Rella, M., Kessmann, H., Ward, E., et al. (1994). A central role of salicylic Acid inplant diseaseresistance. Science 266,1247–1250. Donson, J., Kearney, C.M., Turpen, T.H., Khan, I.A., Kurath, G., Turpen, A.M., Jones, G.E., Dawson, W.O., and Lewandowski, D.J. (1993). Broad resistance to tobamoviruses is mediated by a modified Tobacco mosaic virus replicase transgene. Mol. Plant Microbe Interact. 6, 635–642. Dougherty, W.G., and Parks, T.D. (1995). Transgenes and gene suppression: telling us something new? Curr. Opin. Cell Biol. 7, 399–405. Duan, C.G., Wang, C.H., Fang, R.X., and Guo, H.S. (2008). Artificial MicroRNAs highly accessible to targets confer efficient virus resistance in plants. J. Virol. 82, 11084– 11095. https://doi.org/10.1128/JVI.01377-08 Duan, Y.P., Powell, C.A., Webb, S.E., Purcifull, D.E., and Hiebert, E. (1997). Geminivirus resistance in transgenic tobacco expressing mutated BC1 protein. Mol. PlantMicrobe Interact. 10, 617–623. Düring, K., Hippe, S., Kreuzaler, F., and Schell, J. (1990). Synthesis and self-assembly of a functional monoclonal antibody in transgenic Nicotiana tabacum. Plant Mol. Biol. 15, 281–293. Fauquet, C.M., Mayo, M.A., Maniloff, J., Desselberger, U., and Ball, L.A. (2005). Virus taxonomy: eighth report of the International Committee on Taxonomy of Viruses (Elsevier Academic Press, London). Firek, S., Draper, J., Owen, M.R., Gandecha, A., Cockburn, B., and Whitelam, G.C. (1993). Secretion of a functional single-chain Fv protein in transgenic tobacco plants and cell suspension cultures. Plant Mol. Biol. 23, 861–870. Fischer, U., and Dröge-Laser, W. (2004). Overexpression of NtERF5, a new member of the tobacco ethylene response transcription factor family enhances resistance to Tobacco mosaic virus. Mol. Plant Microbe Interact. 17, 1162–1171. https://doi.org/10.1094/ MPMI.2004.17.10.1162 Fitch, M.M.M. (2016). Update on gene transfer biotechnology of papaya. Acta Hortic. 1111, 7–18. Frischmuth, T., and Stanley, J. (1994). Beet curly top virus symptom amelioration in Nicotiana benthamiana transformed with a naturally occurring viral subgenomic DNA. Virology 200, 826–830. Fuchs, M., and Gonsalves, D. (2007). Safety of virusresistant transgenic plants two decades after their introduction: lessons from realistic field risk assessment studies. Annu Rev Phytopathol 45, 173–202. https:// doi.org/10.1146/annurev.phyto.45.062806.094434 Fuchs, M., Tricoli, D.M., Carney, K.J., Schesser, M., McFerson, J.R., and Gonsalves, D. (1998). Comparative virus resistance and fruit yield of transgenic squash with single and multiple coat protein genes. Plant Dis. 82, 1350–1356.

García, J.A., Glasa, M., Cambra, M., and Candresse, T. (2014). Plum pox virus and sharka: a model potyvirus and a major disease. Mol. Plant Pathol. 15, 226–241. https://doi.org/10.1111/mpp.12083 Gargouri-Bouzid, R., Jaoua, L., Rouis, S., Saïdi, M.N., Bouaziz, D., and Ellouz, R. (2006). PVY-resistant transgenic potato plants expressing an anti-NIa protein scFv antibody. Mol. Biotechnol. 33, 133–140. Gil, M., Esteban, O., Garcia, J.A., Pena, L., and Cambra, M. (2011). Resistance to Plum pox virus in plants expressing cytosolic and nuclear single-chain antibodies against viral NIb replicase. Plant Pathol. 60, 967–976. Golemboski, D.B., Lomonossoff, G.P., and Zaitlin, M. (1990). Plants transformed with a Tobacco mosaic virus nonstructural gene sequence are resistant to the virus. Proc. Natl. Acad. Sci. U.S.A. 87, 6311–6315. Gorbunova, V., and Levy, A.A. (1997). Non-homologous DNA end joining in plant cells is associated with deletions and filler DNA insertions. Nucleic Acids Res. 25, 4650–4657. Hammond, S.M., Bernstein, E., Beach, D., and Hannon, G.J. (2000). An RNA-directed nuclease mediates posttranscriptional gene silencing in Drosophila cells. Nature 404, 293–296. https://doi.org/10.1038/35005107 Hanson, S.F.,and Maxwell, D.P. (1999). trans-Dominant Inhibition of Geminiviral DNA Replication by Bean Golden Mosaic Geminivirus rep Gene Mutants. Phytopathology 89, 480–486. https://doi.org/10.1094/ PHYTO.1999.89.6.480 Helliwell, C.A., and Waterhouse, P.M. (2005). Constructs and methods for hairpin RNA-mediated gene silencing in plants. Meth. Enzymol. 392, 24–35. Hiatt, A., Cafferkey, R., and Bowdish, K. (1989). Production of antibodies in transgenic plants. Nature 342, 76–78. https://doi.org/10.1038/342076a0 Hily,J.M., Scorza, R., Malinowski, T., Zawadzka, B., and Ravelonandro, M. (2004). Stability of gene silencingbased resistance to Plum pox virus in transgenic plum (Prunus domestica L.) under field conditions. Transgenic Res.13, 427–436. Hong, Y., and Stanley, J. (1996). Virus resistance in Nicotiana benthamiana conferred by African Cassava mosaic virus replication-associated protein (AC1) transgene. Mol. Plant-Microbe Interact. 9, 219–225. Hong, Y., Saunders, K., Hartley, M.R., and Stanley, J. (1996). Resistance to geminivirus infection by virus-induced expression of dianthin in transgenic plants. Virology 220, 119–127. Hou, Y.M., Sanders, R., Ursin, V.M., and Gilbertson, R.L. (2000). Transgenic plants expressing geminivirus movement proteins: abnormal phenotypes and delayed infection by Tomato mottle virus in transgenic tomatoes expressing the Bean dwarf mosaic virus BV1 or BC1 proteins. Mol. Plant Microbe Interact. 13, 297–308. https://doi.org/10.1094/MPMI.2000.13.3.297 Hsu, P.D., Lander, E.S., and Zhang, F. (2014). Development and applications of CRISPR-Cas9 for genome engineering. Cell 157, 1262–1278. https://doi. org/10.1016/j.cell.2014.05.010 Hu, W., Kaminski, R., Yang, F., Zhang, Y., Cosentino, L., Li, F., Luo, B., Alvarez-Carbonell, D., Garcia-Mesa, Y., Karn, J., et al. (2014). RNA-directed gene editing specifically eradicates latent and prevents newHIV-1 infection. Proc.

Genetic Engineering for Virus Resistance in Plants |  115

Natl. Acad. Sci. U.S.A. 111, 11461–11466. https://doi. org/10.1073/pnas.1405186111 Hull, R. (2014). Matthews’ Plant Virology (Academic Press, San Diego.) James, C. (2009). Global status of commercialized biotech. GM Crops 2008 (ISAAA, New York). Ji, X., Zhang, H., Zhang, Y., Wang, Y., and Gao, C. (2015). Establishing a CRISPR-Cas-like immune system conferring DNA virus resistance in plants. Nat Plants 1, 15144. https://doi.org/10.1038/nplants.2015.144 Johnson, S.R., Strom, S., and Grillo, K. (2007). Quantification of the impacts on US agriculture of biotechnology-derived crops planted (2006). (National Center for Food and Agriculture Policy, Washington). Kang, B.C., Yeam, I., and Jahn, M.M. (2005). Genetics of plant virus resistance. Annu Rev Phytopathol 43, 581–621. https://doi.org/10.1146/annurev. phyto.43.011205.141140 Kaniewski, W., Lawson, C., Sammons, B., Haley, L., Hart, J., Delannay, X., and Tumer, N. (1990). Field resistance of transgenic Russet Burbank potato to effects of infection by Potato virus X and Potato virus Y. Nat. Biotechnol. 8, 750–754. Kaniewski, W.K., and Thomas, P.E. (2004). The potato story. AgBioForum 7, 41–46. Kawchuk, L.M., Martin, R.R., and McPherson, J. (1990). Resistance in transgenic potato expressing the Potato leaf roll virus coat protein gene. Mol. Plant-Microbe Interact. 3, 301–307. Kim, S.J., Lee, S.J., Kim, B.D., and Paek, K.H. (1997). Satellite-RNA-mediated resistance to Cucumber mosaic virus in transgenic plants of hot pepper (Capsicum annuum cv. Golden Tower). Plant Cell Reports 16, 825–830. Kim, S.J., Paek, K.H., and Kim, B.D. (1995). Delay of disease development in transgenic Petunia plants expressing cucumber mosaic-virus I 17N-satellite RNA. J. Amer. Soc. Hort. Sci. 120, 353–359. Kim, Y.G., Cha, J., and Chandrasegaran, S. (1996). Hybrid restriction enzymes: zinc finger fusions to Fok I cleavage domain. Proc. Natl. Acad. Sci. U.S.A. 93, 1156–1160. Klas, F.E., Fuchs, M., and Gonsalves, D. (2006). Comparative spatial spread overtime of Zucchini Yellow Mosaic Virus (ZYMV) and Watermelon mosaic virus (WMV) in fields of transgenic squash expressing the coat protein genes of ZYMV and WMV, and in fields of non transgenic squash. Transgenic Res. 15, 527–541. Kobayashi, K., Cabral, S., Calamante, G., Maldonado, S., and Mentaberry, A. (2001). Transgenic tobacco plants expressing the potato virus X open reading frame 3 gene develop specific resistance and necrotic ring symptoms after infection with the homologous virus. Mol. PlantMicrobe Interact. 14, 1274–1285. https:// doi.org/10.1094/MPMI.2001.14.11.1274 Krubphachaya, P., Jurícek, M., and Kertbundit, S. (2007). Induction of RNA-mediated resistance to Papaya ringspot virus type W. J. Biochem. Mol. Biol. 40, 404–411. Kumar, S., Tanti, B., Patil, B.L., Mukherjee, S.K., and Sahoo, L. (2017). RNAi-derived transgenic resistance to Mungbean yellow mosaic India virus in cowpea. PLOS ONE. 12: e0186786. Kumari, S., Trivedi, M., and Mishra, M. (2015). PRSV resistance in papaya (Carica papaya L.) through genetic engineering: A review. J. Appl. Hortic. 17, 243–248.

Kundu,J.K., Briard, P., Hily, J.M., Ravelonandro, M., and Scorza, R. (2008). Role of the 25-26 nt siRNA in the resistance of transgenic Prunus domestica graft inoculated with plum poxvirus. VirusGenes36,215–220. https://doi.org/10.1007/s11262-007-0176-y Kyle, M. (1993). Resistance to Viral Diseases of Vegetables. Timber Press, Portland, OR. Lanfermeijer, F.C., Jiang, G., Ferwerda, M.A., Dijkhuis, J., de Haan, P., Yang, R., etal. (2004). The durable resistance gene Tm-22 from tomato confers resistance to ToMV in tobacco and preserves its viral specificity. Plant Science 167, 687–692. Lapidot, M., Gafny, R., Ding, B., Wolf, S., Lucas, W.J., and Beachy, R.N. (1993). A dysfunctional movement protein of Tobacco mosaic virus that partially modifies the plasmodesmata and limits virus spread in transgenic plants. Plant J. 4, 959–970. Lawson, C., Kaniewski, W., Haley, L., Rozman, R., Newell, C., Sanders, P., and Tumer, N.E. (1990). Engineering resistance to mixed virus infection in a commercial potato cultivar: resistance to potato virus X and Potato virus Y in transgenic Russet Burbank. Biotechnology 8, 127–134. Lindbo, J.A., and Dougherty, W.G. (1992a). Pathogenderived resistance to a potyvirus: immune and resistant phenotypes in transgenic tobacco expressing altered forms of a potyvirus coat protein nucleotide sequence. Mol. Plant Microbe Interact. 5, 144–153. Lindbo, J.A., and Dougherty, W.G. (1992b). Untranslatable transcripts of the tobacco etch virus coat protein gene sequence can interfere with tobacco etch virus replication in transgenic plants and protoplasts. Virology 189, 725–733. Lindbo, J.A., and Dougherty, W.G. (2005). Plant pathology and RNAi: a brief history. Annu. Rev. Phytopathol. 43, 191–204. https://doi.org/10.1146/annurev. phyto.43.040204.140228 Lindbo, J.A., and Falk, B.W. (2017). The Impact of ‘Coat Protein-Mediated Virus Resistance’ in Applied Plant Pathology and Basic Research. Phytopathology 107, 624–634. https://doi.org/10.1094/PHYTO-12-160442-RVW Lindbo, J.A., Silva-Rosales, L., Proebsting, W.M., and Dougherty, W.G. (1993). Induction of a Highly Specific Antiviral State in Transgenic Plants: Implications for Regulation of Gene Expression and Virus Resistance. Plant Cell 5, 1749–1759. https://doi.org/10.1105/ tpc.5.12.1749 Liu, J., Carmell, M.A., Rivas, F.V., Marsden, C.G., Thomson, J.M., Song, J.J., Hammond, S.M., Joshua-Tor, L., and Hannon, G.J. (2004). Argonaute2 is the catalytic engine of mammalian RNAi. Science 305, 1437–1441. https:// doi.org/10.1126/science.1102513 Lodge, J.K., Kaniewski, W.K., and Tumer, N.E. (1993). Broad-spectrum virus resistance in transgenic plants expressing pokeweed antiviral protein. Proc. Natl. Acad. Sci. U.S.A. 90, 7089–7093. Lucioli, A., Noris, E., Brunetti, A., Tavazza, R., Ruzza, V., Castillo, A.G., Bejarano, E.R., Accotto, G.P., and Tavazza, M. (2003). Tomato yellow leaf curl Sardinia virus rep-derived resistance to homologous and heterologous geminiviruses occurs by different mechanisms and is overcome if virus-mediated transgene silencing is activated. J. Virol. 77, 6785–6798.

116  | Patil

Mali, P., Yang, L., Esvelt, K.M., Aach, J., Guell, M., DiCarlo, J.E., Norville, J.E., and Church, G.M. (2013). RNA-guided human genome engineering via Cas9. Science 339, 823–826. https://doi.org/10.1126/ science.1232033 Malyshenko, S.I., Kondakova, O.A., Nazarova, J.V., Kaplan, I.B., Taliansky, M.E., and Atabekov, J.G. (1993). Reduction of Tobacco mosaic virus accumulation in transgenic plants producing nonfunctional viral transport proteins. J. Gen. Virol. 74, 1149–1156. Marano, M.R., and Baulcombe, D. (1998). Pathogenderived resistance targeted against the negative-strand RNA of Tobacco mosaic virus: RNA strand-specific gene silencing. Plant J. 13, 537–546. Masuta, C., Tanaka, H., Uehara, K., Kuwata, S., Koiwai, A., and Noma, M. (1995). Broad resistance to plant viruses in transgenic plants conferred by antisense inhibition of a host gene essential in S-adenosylmethionine-dependent transmethylation reactions. Proc. Natl. Acad. Sci. U.S.A. 92, 6117–6121. Maule, A.J., Caranta, C., and Boulton, M.I. (2007). Sources of natural resistance to plant viruses: status and prospects. Mol. Plant Pathol. 8, 223–231. https://doi. org/10.1111/j.1364-3703.2007.00386.x McKinney, H.H. (1929). Mosaic disease in the Canary Islands, West Africa, and Gibralter. J. Agric. Res. 39, 557–578. Mitter, N., Worrall, E.A., Robinson, K.E., Li, P., Jain, R.G., Taochy, C., Fletcher, S.J., Carroll, B.J., Lu, G.Q., and Xu, Z.P. (2017a). Clay nanosheets for topical delivery of RNAi for sustained protection against plant viruses. Nat Plants 3, 16207. https://doi.org/10.1038/ nplants.2016.207 Mitter, N., Worrall, E.A., Robinson, K.E., Xu, Z.P., and Carroll, B.J. (2017b). Induction of virus resistance by exogenous application of double-stranded RNA. Curr Opin Virol 26, 49–55. Mlotshwa, S., Pruss, G.J., and Vance, V. (2008). Small RNAs in viral infection and host defense. Trends Plant Sci. 13, 375–382. https://doi.org/10.1016/j. tplants.2008.04.009 Moon, Y.H., Song, S.K., Choi, K.W., and Lee, J.S. (1997). Expression of a cDNA encoding Phytolacca insularis antiviral protein confers virus resistance on transgenic potato plants. Mol. Cells 7, 807–815. Mueller, E., Gilbert, J., Davenport, G., Brigneti, G., and Baulcombe, D.C. (1995). Homology-dependent resistance: transgenic resistance in plants related to homology-dependent gene silencing. Plant J. 7, 1001– 1013. Nelson, R.S., Abel, P.P., and Beachy, R.N. (1987). Lesions and virus accumulation in inoculated transgenic tobacco plants expressing the coat protein gene of Tobacco mosaic virus. Virology 158, 126–132. Nickel, H., Kawchuk, L., Twyman, R.M., Zimmermann, S., Junghans, H., Winter, S., Fischer, R., and Prüfer, D. (2008). Plantibody-mediated inhibition of the Potato leafroll virus P1 protein reduces virus accumulation. Virus Res. 136, 140–145. https://doi.org/10.1016/j. virusres.2008.05.001 Niu, Q.W., Lin, S.S., Reyes, J.L., Chen, K.C., Wu, H.W., Yeh, S.D., and Chua, N.H. (2006). Expression of artificial

microRNAs in transgenic Arabidopsis thaliana confers virus resistance. Nat. Biotechnol. 24, 1420–1428. Nölke, G., Fischer, R., and Schillberg, S. (2004). Antibodybased resistance in plants. J. Plant Pathol. 86, 5–17. Noris, E., Accotto, G.P., Tavazza, R., Brunetti, A., Crespi, S., and Tavazza, M. (1996). Resistance to tomato yellow leaf curl geminivirus in Nicotiana benthamiana plants transformed with a truncated viral C1 gene. Virology 224, 130–138. Ogwok, E., Odipio, J., Halsey, M., Gaitán-Solís, E., Bua, A., Taylor, N.J., Fauquet, C.M., and Alicai, T. (2012). Transgenic RNA interference (RNAi)-derived field resistance to cassava brown streak disease. Mol. Plant Pathol. 13, 1019–1031. https://doi.org/10.1111/ j.1364-3703.2012.00812.x Oldroyd, G.E., and Staskawicz, B.J. (1998). Genetically engineered broad-spectrum disease resistance in tomato. Proc. Natl. Acad. Sci. U.S.A. 95, 10300–10305. Osbourn, J.K., Watts, J.W., Beachy, R.N., and Wilson, T.M. (1989). Evidence that nucleocapsid disassembly and a later step in virus replication are inhibited in transgenic tobacco protoplasts expressing TMV coat protein. Virology 172, 370–373. Owen, M., Gandecha, A., Cockburn, B., and Whitelam, G. (1992). Synthesis of a functional anti-phytochrome single-chain Fv protein in transgenic tobacco. Biotechnology 10, 790–794. Palukaitis, P., and Carr, J.P. (2008). Plant resistance responses to viruses. J. of Plant Pathol. 90, 153–171. Patil, B.L., and Dasgupta, I. (2006). Defective-interfering DNAs of plant viruses. CRC Crit. Rev. Plant Sci. 25, 47–64. Patil, B.L., and Fauquet, C. (2015). Light intensity and temperature affect systemic spread of silencing signal in transient agro-infiltration studies. Mol. Plant Pathol. 16, 484–494. Patil, B.L., Bagewadi, B., Yadav, J.S., and Fauquet, C.M. (2016). Mapping and identification of cassava mosaic geminivirus DNA-A and DNA-B genome sequences for efficient siRNA expression and RNAi based virus resistance by transient agro-infiltration studies. Virus Res. 213, 109–115. Patil, B.L., Ogwok, E., Wagaba, H., Mohammed, I.U., Yadav, J.S., Bagewadi, B., Taylor,N.J., Kreuze, J.F., Maruthi, M.N., Alicai, T., et al. (2011). RNAi-mediated resistance to diverse isolates belonging to two virus species involved in Cassava brown streak disease. Mol. Plant Pathol. 12, 31–41. https://doi.org/10.1111/j.13643703.2010.00650.x Pooggin, M., Shivaprasad, P.V., Veluthambi, K., and Hohn, T. (2003). RNAi targeting of DNA virus in plants. Nat. Biotechnol. 21, 131–132. https://doi.org/10.1038/ nbt0203-131b Prins, M., Laimer, M., Noris, E., Schubert, J., Wassenegger, M., and Tepfer, M. (2008). Strategies for antiviral resistance in transgenic plants. Mol. Plant Pathol. 9, 73–83. https://doi.org/10.1111/j.13643703.2007.00447.x Puchta, H., Dujon, B., and Hohn, B. (1996). Two different but related mechanisms are used in plants for the repair of genomic double-strand breaks by homologous recombination. Proc. Natl. Acad. Sci. U.S.A. 93, 5055– 5060.

Genetic Engineering for Virus Resistance in Plants |  117

Qu, J., Ye, J., and Fang, R. (2007). Artificial microRNAmediated virus resistance in plants. J. Virol. 81, 6690–6699. Quemada, H.D., Gonsalves, D., and Slightom, J. (1991). Expression of coat protein gene from Cucumber mosaic virus strain C in tobacco: Protection against infections by CMV strains transmitted mechanically or by aphids. Phytopathol. 81, 794–802. Register, J.C., and Beachy, R.N. (1988). Resistance to TMV in transgenic plants results from interference with an early event in infection. Virology 166, 524–532. Robaglia, C., and Caranta, C. (2006). Translation initiation factors: a weak link in plant RNA virus infection. Trends Plant Sci. 11, 40–45. Sanford, J.C., and Johnston, S.A. 1985. The concept of parasite-derived resistance-deriving resistance genes from the parasite’s own genome. J. Theor. Biol. 113, 395–405. Scorza, R., Callahan, A., Dardick, C., Ravelonandro, M., Polak, J., Malinowski, T., Zagrai, I., Cambra, M., and Kamenova, I. (2013). Genetic engineering of Plum pox virus resistance: ‘HoneySweet’ plum—from concept to product. Plant Cell Tissue Organ Cult. (PCTOC) 115, 1–12. Scorza, R., Callahan, A., Levy, L., Damsteegt, V., Webb, K., and Ravelonandro, M. (2001). Post-transcriptional gene silencing in Plum pox virus resistant transgenic European plum containing the plum pox potyvirus coat protein gene. Transgenic Res. 10, 201–209. Seppanen, P., Puska, R., Honkanen, J., Tyulinka, L.G., Fedorkin, O., and Morozov, S.Yu., et al. (1997). Movement protein-derived resistance to triple gene block-containing viruses. J. Gen. Virol. 78, 1241–1246. Shin, R., Park, J.M., An, J.M., and Paek, K.H. (2002). Ectopic expression of Tsi1 in transgenic hot pepper plants enhances host resistance to viral, bacterial, and oomycete pathogens. Mol. Plant Microbe Interact. 15, 983–989. https://doi.org/10.1094/MPMI.2002.15.10.983 Sijen, T., Wellink, J., Hendriks, J., Verver, J., and van Kammen, A. (1995). Replication of cowpea mosaic virus RNA1 and RNA2 is specifically blocked in transgenic Nicotiana benthamiana plants expressing the full-length replicase or movement protein genes. Mol. Plant-Microbe Interact. 8, 340–347. Spassova, M.I., Prins, T.W., Folkertsma, R.T., KleinLankhorst, R.M., Hille, J., and Goldbach, R.W., et al. (2001). The tomato gene Sw5 is a member of the coiled coil, nucleotide binding, leucine-rich repeat class of plant resistance genes and confers resistance to TSWV in tobacco. Mol. Breeding 7, 151–161. Stenger, D.C. (1994). Strain-specific mobilization and amplification of a transgenic defective-interfering DNA of the geminivirus Beet curly top virus. Virology 203, 397–402. Tacke, E., Salamini, F., and Rohde, W. (1996). Genetic engineering of potato for broadspectrum protection against virus infection. Nat. Biotechnology 14, 1597– 1601. Takahashi,H., Chen, Z., Du, H., Liu, Y., and Klessig, D.F. (1997). Development of necrosis and activation of disease resistance in transgenic tobacco plants with severely reducedcatalaselevels. Plant J. 11,993–1005.

Tavladoraki, P., Benvenuto, E., Trinca, S., De Martinis, D., Cattaneo, A., and Galeffi, P. (1993). Transgenic plants expressing a functional single-chain Fv antibody are specifically protected from virus attack. Nature 366, 469–472. https://doi.org/10.1038/366469a0 Tenllado, F., Llave, C., and Díaz-Ruíz, J.R. (2004). RNA interference as a new biotechnological tool for the control of virus diseases in plants. Virus Res. 102, 85–96. https://doi.org/10.1016/j.virusres.2004.01.019 Tennant, P., Fermin, G., Fitch, M.M., Manshardt, R.M., Slightom, J., and Gonsalves, D. (2001). Papaya ringspot virus resistance of transgenic rainbow and SunUp is affected by gene dosage, plant development, and coat protein homology. Eur. J. Plant Pathol. 107, 645–653. Thomas, P.E., Lawson, E.C., Zalewski, J.C., Reed, G.L., and Kaniewski, W.K. (2000). Extreme resistance to Potato leafroll virus in potato cv. Russet Burbank mediated by the viral replicase gene. Virus Res. 71, 49–62. Tollefson, J. (2011). Brazil cooks up transgenic bean. Nature 478, 168. https://doi.org/10.1038/478168a Tricoli, D., Carney, K.J., Russell, P.F., McMaster, J.R., Groff, D.W., Hadden, K.C., Himmel, P.T., Hubbard, J.P., Boeshore, M.L., and Quemada, H.D. (1995). Field evaluation of transgenic squash containing single or multiple virus coat protein gene constructs for resistance to Cucumber mosaic virus, Watermelon mosaic virus 2 and zucchini yellow mosaic virus. BioTechnology 13, 1458–1465. Tricoli, D.M., Carney, K.J., Russell, P.F., McMaster, J.R., Groff, D.W., Hadden, K.C., Himmel, P.T., Hubbard, J.P., Boeshore, M.L., and Quemada, H.D. (1995). Single or multiple virus coat protein gene constructs for resistance to Cucumber mosaic virus, Watermelon mosaic virus 2, and Zucchini yellow mosaic virus. Nat. Biotechnol. 13, 1458–1465. Tumer, N.E., Hwang, D.J., and Bonness, M. (1997). C-terminal deletion mutant of pokeweed antiviral protein inhibits viral infection but does not depurinate host ribosomes. Proc. Natl. Acad. Sci. USA 94, 3866– 3871. Van Damme, E.J.M., Peumans, W.J., Barre, A., and Rouge, P. (1998). Plant lectins: A composite of several distinct families of structurally and evolutionary related proteins with diverse biological roles. Crit. Rev. Plant Sci. 17, 575–692. Vanitharani, R., Chellappan, P., and Fauquet, C.M. (2005). Geminiviruses and RNA silencing. Trends Plant Sci. 10, 144–151. Vazquez Rovere, C., Asurmendi, S., and Hopp, H.E. (2001). Transgenic resistance in potato plants expressing Potato leaf roll virus (PLRV) replicase gene sequences is RNA-mediated and suggests the involvement of posttranscriptional gene silencing. Arch. Virol. 146, 1337–1353. Voloudakis, A.E., Holeva, M.C., Sarin, L.P., Bamford, D.H., Vargas, M., Poranen, M.M., and Tenllado, F. (2015). Efficient double-stranded RNA production methods for utilization in plant virus control. Methods Mol. Biol. 1236, 255–274. https://doi.org/10.1007/978-1-49391743-3_19 Wagaba, H., Patil, B.L., Mukasa, S., Alicai, T., Fauquet, C.M., and Taylor, N.J. (2016). Artificial micro RNA-derived

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resistance to Cassava brown streak disease. J Virol Methods 231, 38–43. Wang, P., and Tumer, N.E. (2000). Virus resistance mediated by ribosome inactivating proteins. Adv. Virus Res. 55, 325–355. Waterhouse, P.M., and Fusaro, A.F. (2006). Plant science. Viruses face a double defense by plant small RNAs. Science 313, 54–55. Waterhouse, P.M., Graham, M.W., and Wang, M.B. (1998). Virus resistance and gene silencing in plants can be induced by simultaneous expression of sense and antisense RNA. Proc. Natl. Acad. Sci. U.S.A. 95, 13959–13964. Whitfield, A.E., Falk, B.W., and Rotenberg, D. (2015). Insect vector-mediated transmission of plant viruses. Virology 479-480, 278–289. https://doi.org/10.1016/j. virol.2015.03.026 Whitham, S.A., and Hajimorad, M.R. (2016). Plant Genetic Resistance to Viruses. (Springer, AG Switzerland.) doi:10.1007/978-3-319-32919-2 4 Wisniewski, L.A., Powell, P.A., Nelson, R.S., and Beachy, R.N. (1990). Local and systemic spread of Tobacco mosaic virus in transgenic tobacco. Plant Cell 2, 559–567. https://doi.org/10.1105/tpc.2.6.559 Yadav, J.S., Ogwok, E., Wagaba, H., Patil, B.L., Bagewadi, B., Alicai, T., Gaitan-Solis, E., Taylor, N.J., and Fauquet, C.M. (2011). RNAi-mediated resistance to Cassava brown streak Uganda virus in transgenic cassava. Mol. Plant Pathol. 12, 677–687. https://doi.org/10.1111/ j.1364-3703.2010.00700.x Yamaji, Y., Kobayashi, T., Hamada, K., Sakurai, K., Yoshii, A., Suzuki, M., Namba, S., and Hibi, T. (2006). In vivo interaction between Tobacco mosaic virus RNAdependent RNA polymerase and host translation elongation factor 1A. Virology 347, 100–108.

Yin,K., Han, T., Liu, G., Chen, T., Wang, Y., Yu, A.Y., and Liu, Y. (2015). A geminivirus-based guide RNA delivery system for CRISPR/Cas9 mediated plant genome editing. Sci Rep5, 14926. https://doi.org/10.1038/ srep14926 Zaidi, S.S., Mansoor, S.,Ali,Z., Tashkandi, M., and Mahfouz, M.M. (2016). Engineering Plants for Geminivirus Resistance with CRISPR/Cas9 System. Trends Plant Sci. 21, 279–281. https://doi.org/10.1016/j. tplants.2016.01.023 Zhang, G., Chen, M., Li, L., Xu, Z., Chen, X., and Guo, J., et al. (2009). Over expression of the soybean GmERF3 gene, an AP2/ERF type transcription factor for increased tolerances to salt, drought, and diseases in transgenic tobacco. J of Experimental Botany 60, 3781–3796. Zhang, H., Tian, Y., Zhou, Y., Dang, B., Lan, H., and Song, G., et al. (1999). Introduction of pokeweed antiviral protein cDNA into Brassica napus and acquisition of transgenic plants resistant to viruses. Chinese Science Bulletin 44, 701–704. Zhang, X., Li, H., Zhang, J., Zhang, C., Gong, P., Ziaf, K., Xiao, F., and Ye, Z. (2011). Expression of artificial microRNAs in tomato confers efficient and stable virus resistance in a cell-autonomous manner. Transgenic Res. 20, 569–581. https://doi.org/10.1007/s11248-0109440-3 Ziebell, H., and Carr, J.P. (2010). Cross-protection: a century of mystery. Adv. Virus Res. 76, 211–264. https://doi.org/10.1016/S0065-3527(10)76006-1 Ziegler-Graff, V., Guilford, P.J., and Baulcombe, D.C. (1991). Tobacco rattle virus RNA-1 29K gene product potentiates viral movement and also affects symptom induction in tobacco. Virology 182, 145–155.

Tools and Techniques for Production of Double-stranded RNA and its Application for Management of Plant Viral Diseases

6

Andreas E. Voloudakis1*, Maria C. Holeva2, Athanasios Kaldis1 and Dongho Kim3

1Laboratory of Plant Breeding and Biometry, Department of Crop Science, Agricultural University of Athens, Athens,

Greece.

2Laboratory of Bacteriology, Benaki Phytopathological Institute, Kifissia, Greece. 3AgroRNA Genolution, Songpa-gu, Seoul, South Korea.

*Correspondence: [email protected] https://doi.org/10.21775/9781910190814.06

Abstract Due to the rapidly growing global population, food production and security is the major challenge of agriculture. Plant viruses are obligate parasites that in some instances could cause up to 100% losses in a crop (e.g. maize streak disease). Although difficult to accurately determine the global economic impact that plant viruses have on agriculture, it is estimated that US$60 billion loss in crop yields worldwide each year is due to plant viral diseases. RNA silencing (RNA interference, RNAi) is a conserved endogenous pathway of all higher eukaryotes, which controls gene expression. RNAi is induced by double-stranded RNA (dsRNA) and allows the cell to recognize aberrant genetic material in a highly sequence-specific manner ultimately leading to its degradation, thus protecting the cell from subcellular pathogens, such as viruses and transposons. DsRNA-mediated resistance has been exploited in transgenic plants to convey resistance to viruses and against insects, vectors of plant viruses, via host induced gene silencing (HIGS). A non-transgenic approach employing RNAi has been used where enzymatically synthesized specific dsRNA molecules, when applied directly onto

plant tissue, induce resistance to the cognate virus; as a result dsRNA molecules could be efficacious antiviral agents for crop protection. Next generation sequencing and bioinformatics analyses have provided a plethora of information and useful tools for the design and study of dsRNA application. In this chapter, the different methods for dsRNA production, both in vitro and in vivo, the means of direct application of the dsRNA molecules onto plants and several examples of non-transgenic dsRNA-mediated resistance are presented. Introduction FAO estimates that a 20–40% global crop yield reduction each year is due to the damage brought by plant pests and diseases, a percentage that threatens food security and agricultural sustainability. The information on the importance of viral diseases in crop production is incomplete, even to date, due to the wide range of crop/plants infected and the agro-ecologies studied, the evolution of pathogenic virus strains or strain combinations, the spread of viruses and virus vectors into new areas (control measures take time to implement) due to human

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activities and climate change, the variable intensity of relevant research worldwide, and the lack of data on losses plant viruses cause. As a result, the global picture of their ecological and economical impact is difficult to be accurately presented. However, it is accepted that plant viruses cause serious damage in agriculture worldwide, with the economic damage estimated to be US$60 billion loss in crop yields worldwide each year (https://en.wikipedia.org/ wiki/Plant_virus#cite_note-1). This is possibly an underestimated value since crop losses are also caused without obvious exhibition of viral symptoms or due to misinterpretation of viral symptoms for other effects (e.g. plant nutrient deficiencies). Mahy and Regenmortel (2009) estimated that reduction in global crop yields each year due to plant pathogenic viruses range from 10% to 15%. Plant viruses are obligate intracellular parasites, with the majority of them needing a vector for transmission to a new host plant thus ensuring their survival. Virus vectors include insects (aphids, whiteflies, thrips, leafhoppers, planthoppers), mites, nematodes and fungi. Vectors play a key role in determining the plant host range of the respective virus. The plant virus–vector interaction is genetically controlled and it depends on specific molecular interactions of components of viral as well as vector origin. It is well understood that elimination/control of the virus-vector is desirable in order to reduce viral epidemics. No antiviral agents for plants exist as opposed to chemicals available for insects, mites, nematodes, fungi and bacteria. Chemical control of virus vectors is the current strategy followed in order to reduce virus transmission on a crop. This control is accomplished by insecticide sprays, which in some cases are numerous and costly, or by integrated pest management (IPM). Insecticide spraying, which potentially has harmful environmental and health side effects, could induce the development of insect-resistant strains that will prevail over time rendering insecticide use ineffective. Thus the identification of novel, environment-friendly alternative means to control pests is needed; farmers facing the threat of insecticide resistance development will more easily accept nucleic acid-based insecticides. Viroids comprise a group of plant pathogens that possess a small (256–401 nt) circular ssRNA genome, which self-folds leading to a secondary structure, without encoding for any protein. Viroids

replicate either in the nucleus or chloroplast via a rolling-circle method and indirectly alter plant gene expression resulting in diseases. In the literature there are a couple of ‘Top Ten’ lists regarding plant viruses; the first one ranking plant viruses according to international community voting (Scholthof et al., 2011) and the second one according to the economic impact to humanity if the infected worldwide-important food crops are affected (Rybicki, 2015). The control of plant virus and their epidemics – apart from the chemical control of virus vectors – is accomplished mainly by employing one or a combination of the following means. Plant resistance to viruses Resistance of plants against viruses -and their vectors- could be achieved via utilization of resistance (R) genes (dominant or recessive) where such genetic material is available (for reviews see Soosaar et al., 2005; de Ronde et al., 2014; chapters in the present book); this is considered the most desirable plant resistance mechanism against viruses. Resistance to viruses in plants is usually established by confining the invading virus in a necrotic lesion (outcome of the hypersensitive response, HR) in the site of entry and the immediately adjacent cells, thus inhibiting virus spread in the neighbouring healthy cells and preventing further infection. RNA interference (RNAi) establishes a very strong defence against viruses at the post-transcriptional level, contributing to the innate immune plant response. RNAi will be discussed later in this chapter. RNA decay pathways compose a conserved mechanism that aids in the endogenous gene expression control via elimination of dysfunctional transcripts leading to the desired mRNA quality and abundance. With increase of our knowledge of mRNA decay and RNAi it becomes more apparent that there is a genetic overlap between the two pathways (Christie et al., 2011). RNA quality control (RQC) and RNAi pathways interplay, with the degradation of aberrant RNAs by RQC preventing their entry in the RNAi pathway and thus reducing the amounts of siRNAs produced from those RNAs. Recently, it was found that coat and movement proteins of Tobacco mosaic virus (TMV) play a key role in inducing RNA decay pathways in tobacco impairing RNAi, presumably due to the

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reduction of the viral siRNAs produced that would lead into miss-function of the siRNA amplification step of RNAi (Conti et al., 2017). Nonsense-mediated decay (NMD), a host RQC mechanism, was proposed to serve as a general viral restriction mechanism in plants (Garcia et al., 2014). Study of this plant resistance mechanism, that operates independent of the RNAi mechanism (for RNAi see below), is genetically based and poorly as yet characterized in plants. Viruses depend heavily on the host proteins (such as the translational machinery) to complete their life cycle. Host plants have evolved translational defence mechanisms that impair viral infection, with the host-mediated translational suppression described as an efficient means to specifically suppress viral mRNA translation. Plants have evolved recessive resistance genes (more durable), with mutations in genes encoding translation initiation factors (eIFs), which are normally required for viral mRNA translation and infection. Due to functional redundancy of the eIF isoforms, loss-of-function mutations of one isoform could provide virus resistance without compromising plant cell translation and thus growth. It is noteworthy mentioning that CRISPR/Cas9 was employed to develop novel genetic resistance to Turnip mosaic virus (TuMV) in Arabidopsis thaliana by deletion of eIF(iso)4E (Pyott et al.,2016) and against Cucumber vein yellowing virus, Zucchini yellow mosaic virus and Papaya ring spot mosaic virus-W in cucumber by disrupting eIF4E (Chandrasekaran et al., 2016). Along these lines, it was shown that activation of the trans-membrane immune receptor NIK1 [nuclear shuttle protein (NSP)-interacting kinase 1] promotes the down-regulation of translational machinery associated genes, culminating in the inhibition of viral and host mRNAs translation, which causes an increase in tolerance to begomoviruses (Zorzatto et al., 2015). Pathogen derived resistance to viruses Cross-protection in plants against viruses is a well known ‘immunization’ phenomenon where a mild virus isolate, upon inoculation onto its host, can protect the plant host against a severe isolate of the same virus (Gal-On and Shiboleth, 2006). The molecular mechanism of cross-protection still remains unclear, although several lines of evidence

imply that the resistance could be protein- and/or RNA-mediated. Cross-protection or host genetic resistance to a virus could be at risk if another virus infects the same host plant. There are several reports of plant virus synergism in the field (e.g. potyvirus-PVX, CMV-TYLCV, etc). In such a case one virus could suppress the resistance to another virus leading to disease epidemic and establishing a warning to plant breeders raising virus-resistant transgenic crops. Pathogen derived resistance (PDR) (Sanford and Johnston, 1985) is considered a scientific development of the cross-protection method. The first attempt to utilize the concept of pathogen derived resistance, namely the introduction of small or long fragments of the virus genome into the plant host genome, proposed by Sanford and Johnston, resulted in the pioneering work on coatprotein-mediated resistance to Tobacco mosaic virus by the Beachy lab (Abel et al., 1986). Since then multiple strategies have been developed to engineer resistance into transgenic plants. This includes expression of the target protein (protein-based resistance) or the corresponding RNA molecule (RNA-mediated resistance). RNAi-mediated resistance to viruses RNA interference (RNAi) or RNA silencing is an endogenous gene expression control mechanism that is present in all eukaryotes (Baulcombe, 2004; Csorba et al., 2009; Waterhouse et al., 2001). RNAi was first recognized in plants in early 1990s, where it was called post-transcriptional gene silencing (PTGS) (Lindbo, 2012). RNAi is an important defence mechanism against viruses that is triggered by viral double-stranded (ds) RNA molecules. These dsRNA precursors may arise from replicative viral intermediates, from RNA secondary structures of the viral genome, from the annealing of sense and anti-sense RNAs, or from the action of host RNAdependent RNA polymerases (RDRs) (Ding, 2010). The dsRNA molecules are detected by the Dicer-like (DCL) proteins -that possess RNase type III-like activity- and cleaved into short interfering (si) RNA duplexes (Aliyari and Ding, 2009), with the majority of the siRNAs having a length of 21–24 nucleotides (nt) (Bernstein et al., 2001; Blevins et al., 2006). SiRNAs are incorporated into RNA inducing silencing complexes (RISCs) by

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specifically interacting with a member of the Argonaute (AGO) family of proteins (Hutvagner and Simard, 2008). The one strand (passenger strand) is degraded by the innate endonuclease RNase H-like activity of AGO proteins. The remaining strand (guide strand) serves as the guide molecule that allows the RISC complex to come in close proximity with the viral target mRNAs and to cleave them in a sequence specific manner (Baumberger and Baulcombe, 2005). RDRs are enzymes that use single-stranded RNAs as templates to create the complementary strand, thus producing dsRNA. The importance of RDRs comes from the fact that by producing new long dsRNA molecules, that subsequently will be recognized by DCL proteins, they promote the generation of secondary siRNAs and as a result increase the pool of siRNAs that can exert an antiviral silencing effect (Garcia-Ruiz et al., 2010; Wang et al., 2010). RNAi plays a key role in plant defence against viruses (Wang et al., 2012) and has therefore been exploited in plant biotechnology to induce resistance via transgenesis (Duan et al., 2012). Genetic material introduced in the host genome includes dsRNA, siRNAs and artificial micro RNAs (amiRNAs). The selection of the genetic material was made easy due to the rich sequence information available for plant viruses as a result of genomics analyses and the use of next generation sequencing. It should be noted that viruses have evolved suppressors of RNAi (for a review see Csorba et al., 2015) in order to escape the RNAi mechanism. One could imagine that an evolutionary arms race between hosts and viruses exists with RNAi in the central part. The specificity of RNAi was initially described as high owing to the sequence homology of the siRNA with the target mRNA. However, this needs to be revisited since some reports have shown nonspecific effects (unintended gene silencing) that are designated as off-target effects (Senthil-Kumar and Mysore, 2011). Such unintended gene silencing could affect the outcome of functional gene analyses where RNAi is used as a reverse genetics tool, as well as affect gene expression in non-target organisms (e.g. human and animals) when dsRNA is applied. For the precise selection of sequences to be used in RNAi, bioinformatics tools (e.g. http:// plantgrn.noble.org/pssRNAit/) have been developed that perform scanning in genomes of host or

non-target organisms for off-target sequences enabling as a result the design of more specific RNAi constructs. A new transgene-based host-induced gene silencing (HIGS) strategy was developed in order to protect plants from vectors of plant viruses such as insects and nematodes (Hunter et al., 2012; Tamilarasan and Rajam, 2013; Koch and Kogel, 2014). In this method, the transgenic plant produces dsRNA or siRNAs of a key endogenous gene of the insect; uptake of these dsRNA/siRNAs will lead into silencing of the gene in the insect and as a result reduction of pest population, which is a desirable outcome that greatly affects virus disease epidemics. It was shown that host plants were protected from insects when they expressed dsRNAs (e.g. hairpin dsRNA) targeting vital insect genes (Baum et al., 2007; Mao et al., 2007; Gordon and Waterhouse, 2007). Furthermore, application of ingestible dsRNA that reduces the expression of detoxification insect genes, encoding proteins that detoxify plant toxic compounds (e.g. cotton gossypol), could become a promising strategy for controlling insect pests. It is anticipated that ingestible dsRNA (transgene-encoded or exogenously applied) could join the farmer’s armoire alongside the Bt transgenes that have shown very good success, replacing chemical insecticides for a range of crops, in IPM. Such a nucleotide specific gene silencing method could also be applicable for many important insect pests where Bt protection is not amenable to and where there is an imminent danger of developing Bt resistance. HIGS is limited by several factors, namely the availability of transformation protocols and difficulties in making stable transgene expression in many crop species, as well as the public’s concern for the production of genetically modified crops (GMOs). In order to circumvent the issues raised by GMO creation through transgenesis, nontransgenic methods -exploiting RNAi- have been developed to induce resistance to plant viruses. It is proposed that the exogenously applied dsRNA, upon introduction into the plant cell, mimics the viral dsRNA intermediate (aberrant or not), and triggers RNAi (Tenllado et al., 2004). This may lead to the production of siRNAs that are incorporated into the nuclease complex (RNA induced silencing complex, RISC) responsible for degradation of the cognate target RNA. The method of ‘dsRNA

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vaccination’ has been used with success against several RNA viruses as well as viroids (Table 6.1). Other plant resistance mechanisms against viruses RNA-mediated transcriptional gene silencing (TGS) is a conserved phenomenon that occurs in fungi, plants and animals. In plants, RNA-mediated TGS was reported first by Wassenegger et al. (1994), designated as RNA-directed DNA methylation (RdDM), a process essential for suppressing transposons, repairing DNA damage caused by stress, stabilizing the genome and maintaining cell identity, as well as defending against exogenous DNAs (e.g. transgenes). Plants produce 24-nt small interfering (si) RNAs and long non-coding RNAs (80 nt long) to direct de novo DNA methylation and thus TGS. RdDM is mediated by two plant-specific DNA-dependent RNA polymerases, Pol IV and Pol V; Pol IV functions to initiate siRNA biogenesis, while Pol V generates scaffold transcripts that recruit downstream RdDM factors (Pooggin et al., 2013). It was recently found that a class of Dicerindependent non-coding RNAs largely guides RdDM in plants, and that siRNAs are required to maintain DNA methylation at only a subset of loci (Yang et al., 2016). RdDM being a nuclear branch of the plant RNA silencing machinery regulates gene expression and defends against invasive nucleic acids such as transposons, transgenes and viruses. Transcriptional arrest of viral mini-chromosomes (Ghoshal and Sanfacon, 2015) could lead to reduction in infection. However, there is evidence indicating that DNA viruses (geminiviruses and pararetoviruses) most likely evade or suppress RdDM (Pooggin et al., 2013). Specialized mechanisms of replication and silencing evasion could rescue viral DNA mini-chromosomes from repressive methylation, established by the RdDM pathway, interfering thus with the desirable transcriptional and posttranscriptional silencing of viral genes. The usefulness of genomics and in particular the advent of next generation sequencing, at an affordable cost, should be appreciated in our days where we have in our possession a great amount of genetic information available. Recently, genome editing (CRISPR/Cas 9 and other methods) in plants has been proposed to generate virus resistance (Zaidi et al.,2016). This precision engineering could be

accomplished via transgenesis or by employing autonomously replicating virus-based vectors (DNA and RNA viruses) that deliver genomeediting reagents in plants (Zaidi and Mansoor, 2017). Genome editing against geminivirus has been achieved (Ali et al., 2016), increasing thus the number of approaches available for virus protection. Resistance to potyviruses was achieved through genome editing of eIF(iso)4E or eIF-4E (Pyott et al., 2016; Chandrasekaran et al., 2016), taking thus advantage of the translational control of plant immunity (Machado et al., 2017). The search of susceptibility genes in plant–virus interactions has attracted the interest of the scientific community especially with the advent of RNAi and genome editing technologies. Such searches in the genomics era, with the bioinformatics tools available, are relative easy since both transcriptomics and proteomics analyses of plant–virus interactions increase in number rapidly. DsRNA production Laboratory-level dsRNA production Two methodological approaches have been described for the production of dsRNA molecules for topical application in plants. Both approaches are based on an enzyme-mediated reaction transcribing a selected target DNA sequence to dsRNA. The first approach is performed in a test tube (‘in vitro’ approach), while the second one employs specially engineered bacterial strains producing the dsRNA molecules inside their cells (‘in vivo’ approach). These approaches, also reviewed by Robinson et al. (2014), Voloudakis et al. (2015) and Mitter et al. (2017), are briefly presented below. In vitro dsRNA production For the in vitro approach (Fig. 6.1), the targetedDNA viral sequence to be used as template for transcription, can be: i

Amplified by targeted-DNA specific primers (F1, R1) and cloned in a bidirectional transcription plasmid-vector with opposing T7 DNA-dependent RNA polymerase (DdRp) promoters flanking the polylinker sites, e.g. LITMUS (NEB, USA) or L4440 (Addgene, USA). Then, the cloned insert is PCR-amplified

Table 6.1 List of plant viruses and viroids against which dsRNA was exogenously applied on host plants for virus control Target virus/viroid

Family

Gene

Size (bp)

Host plant

Production method

Protection (%) Reference

Pepper mild mottle virus (PMMoV)

Virgaviridae

Replicase

977,596

N. benthamiana

In vitro

82

Tenllado and DíazRuíz (2001)

Tobacco etch virus (TEV)

Potyviridae

HC-Pro

1483

Tobacco

IN vitro

100

Tenllado and DíazRuíz (2001)

Alfalfa mosaic virus (AlMV)

Bromoviridae

RNA3

1124

N. benthamiana

In vitro

100

Tenllado and DíazRuíz (2001)

Plum pox virus (PPV)

Potyviridae

HC-Pro

1492

N. benthamiana

In vivo1

83

Tenllado et al. (2003)

CP

1081

73

Pepper mild mottle virus (PMMoV)

Virgaviridae

Replicase, IR 54

977

N. benthamiana

In vivo1

100

Tenllado et al. (2003)

Cucumber mosaic virus (CMV)

Bromoviridae

CP

657

Tobacco

IN vitro

40

Ηoleva et al. (2006)

50

Tomato CP

657

Tobacco

In vitro

40

In vivo

65

In vitro

38

1

2b

400

Ηoleva et al. (2007)

CP + 2b

657 + 400

Pospiviroidae

-

365

Potato spindle tuber viroid (PSTVd)

Pospiviroidae

-

353

Chrysanthemum chlorotic mottle viroid (CChMVd)

Avsunviroidae

-

356

Tobacco mosaic virus (TMV)

Virgaviridae

CP

480

p126

666

CP

480

In vivo

50

Yin et al. (2009)

In vivo

34–66

Sun et al. (2010)

Citrus exocortis viroid (CEVd)

various

In vitro

45

In vitro

50

Tomato

In vitro

delay

Chrysanthemum

In vitro

50

Tobacco

In vitro

50

Konakalla et al. (2016)

In vitro

65

Konakalla et al. (2016)

Gynura, Tomato

Carbonell et al. (2008)

Table 6.1 Continued Host plant

Production method

Protection (%) Reference

Tobacco

In vivo

46–72

Sun et al. (2010)

Maize

In vivo

70, 80

Gan et al. (2010)

≈ 500

Pea

in vivo

55

Safarova et al. (2014)

236

Orchid

in vivo

50

Lau et al. (2014) Mitter et al. (2017) Kaldis et al. (2018)

Target virus/viroid

Family

Gene

Size (bp)

Potato virus Y (PVY)

Potyviridae

various

Sugarcane mosaic virus (SCMV)

Potyviridae

CP

327,394

Pea seed-borne mosaic virus (PsbMV)

Potyviridae

CP

Cymbidium mosaic virus (CymMV)

Alphaflexiviridae

CP

Cucumber mosaic virus (CMV)

Bromoviridae

2b

330

Tobacco

In vivo

762

Zucchini yellow mosaic virus (ZYMV)

Potyviridae

HC-Pro

588

Cucumber, Watermelon

In vitro

82, 50

CP 1In 2In

498

vivo dsRNA production was made in Escherichia coli HT115 strain (RNAse II-deficient strain). this experiment the dsRNA was not naked but loaded onto ‘bioclay’ prior to spraying.

70, 43

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Figure 6.1  Schematic representation of five methods (i to v) for in vitro production of dsRNA. For each method, the main steps to generate the template for the in vitro transcription are indicated. In method (i) step 3 can be followed by steps 4a and 5a, or 4b and 5b. Diagrams are not drawn to scale.

ii

using a single T7 DdRp promoter-specific primer, which results in a PCR product with the target DNA sequence flanked by T7 DdRp promoters. In vitro transcription of this PCR product produces dsRNA. Alternatively, the plasmid containing the target sequence is linearized by restriction digestion downstream of the cloned insert, thus, generating a template for each RNA strand. The linearized templates are pooled for simultaneous transcription that synthesizes dsRNA (HiScribe RNAi Transcription Kit of BioLabs, USA; Holeva et al., 2006) (Fig. 6.1i). Amplified by targeted-DNA specific primers

(F1, R1) and cloned in a unidirectional transcription plasmid-vector with a single T7 DdRp promoter sequence present upstream of the polylinker site. In this case, two separate plasmid clones having the same target sequence in opposite orientations need to be constructed. Each clone should contain the T7 DdRp promoter at both ends of the target sequence. The two plasmid clones are linearized by restriction digestion downstream of the cloned insert, thus, generating a template for each RNA strand. The two complementary RNA strands synthesized are mixed and hybridized post-transcriptionally to form

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dsRNA (Tenllado and Diaz-Ruiz, 2001) (Fig. 6.1ii). iii Amplified by targeted-DNA specific primers in two PCR reactions, in which the T7 DdRp promoter sequence has been introduced at the 5΄ end of either of the amplification primers. Thus four primers and two PCR reactions are necessary for the production of the transcription templates. The resulting two PCR products are then used for the transcription and production of dsRNA (T7 RiboMAX™ Express RNAi System, Promega, USA) (Fig. 6.1iii). iv Amplified by targeted-DNA specific primers that carry a linker sequence at their 5′ ends. The PCR product is then re-amplified by a single primer with the T7 DdRp promoter sequence at its 5′ end and the linker sequence at its 3′ end. The resulting PCR product has at both ends the T7 DdRp promoter sequence. In vitro transcription of this PCR product produces dsRNA (Holeva et al., 2007; Konakalla et al., 2016; Kaldis et al., 2018) (Fig. 6.1iv). v Amplified with primers designed separately for each target sequence. Both primers contain 17–22 nucleotides of the target sequence at the 3′ end. In addition, the forward primer contains the T7 DdRp promoter sequence at its 5′ ends, and the reverse primer contains the phi 6 (φ6) RNA dependent RNA polymerase (RDR) promoter sequence at its 5′ end. The resulting PCR product is transcribed into ssRNA by T7 DdRp which is then replicated by φ6 RDR into dsRNA. The in vitro transcription and the replication reaction take place in a single incubation step (Aalto et al., 2007) (Fig. 6.1v). In vivo dsRNA production For the in vivo approach, three Escherichia coli strains and one Pseudomonas syringae strain have been reported in the literature as suitable for the production of dsRNA and these bacterial expression systems are briefly presented here. The most frequently used is the E. coli HT115 (DE3) strain (Timmons et al., 2001), which is an RNAse III-deficient strain expressing T7 DdRp from an isopropyl b-D-1 thiogalactopyranoside (IPTG)-inducible promoter. E. coli HT115 (DE3) is transformed with a plasmid construct (transcription vector) carrying the targeted-DNA sequence between T7 DdRp promoters (Fig. 6.2i). Upon

IPTG-induction for 2–4 hours at 37°C, dsRNA is produced and accumulated in the bacterial cells from where it is purified by standard phenol-chloroform treatment, or French press lysis or sonication (Tenllado et al., 2003; Gan et al., 2010) (Fig. 6.2ii). The E. coli M-JM109LacY is another RNAse IIIdeficient strain, which has been similarly used for dsRNA production upon IPTG induction and is reported to produce higher amounts of dsRNA in relation to E. coli HT115 (DE3) (Yin et al., 2009). More recently, an RNase III competent E. coli strain carrying a construct encoding p19 (a siRNA binding protein of the plant RNA virus tombusvirus) has been utilized to produce a pool of siRNAs for exogenous application. In this strain, the targeted-DNA sequence is transcribed by T7 DdRp. The dsRNA produced is digested by the bacterial RNase III into ≈ 21 nt duplexes which form complexes with p19 protein. After affinity capture of the complexes, a pool of siRNAs is eluted and is ready for application (Huang et al., 2013). Furthermore, Pseudomonas syringae LM2691 strain has been utilized for large-scale dsRNA production. This strain harbours three plasmids: (a) plasmid pLM1086, which constitutively expresss T7 DdRp; (b) pLM991, which contains a cDNA copy of φ6 genome segment L under T7 DdRp promoter and a kanamycin resistance gene; and (c) pSinsert, which carries the target DNA sequence between the RNA packaging and replication signals specific for the φ6 genome segment S, under the T7 DdRp promoter. The transcribed ssRNAs of the target sequence are packaged into φ6 procapsids where they are converted to dsRNA. These dsRNAs serve as template for transcription and lead to amplification of dsRNA-filled polymerase complexes (Aalto et al., 2007; Voloudakis et al., 2015). Industrial dsRNA production – a perspective The discovery that long dsRNA can be taken up and processed to trigger RNAi effect in plants, insects and possibly some invertebrates, such as shrimp, is an exciting one for industry. If a target-specific dsRNA is sprayed to a target organism, serving as a target-specific viricide, insecticide, herbicide etc will not raise significant environmental concerns over the non-selective toxic effect of the chemical alternatives, where present. Even though the

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Figure 6.2  In vivo approach for production of dsRNA in E. coli HT115 (DE3) cells. (i) Diagram of the bacterial cell carrying the plasmid construct for the production of dsRNA of the targeted sequence (Lac-Pro: Lactose-responsive promoter; T7 DdRp gene: T7 DNA dependent RNA polymerase gene; TetR gene: tetracycline resistance gene). (ii) Main steps for the production of dsRNA using the E. coli HT115 (DE3) strain carrying the plasmid construct described above.

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stability and efficacy of dsRNA is assured, we are facing the issue of dsRNA mass production as an agrochemical product and its associated cost. The two methods, described above, need to be compared at an industrial level in order to determine the method of choice. With a shallow understanding, it may be easy to conclude that RNA produced from bulky E. coli cultures may be the better choice due to its lower cost. In fact, several reports claim that the method may be useful in agricultural as well as aquacultural applications. However, the following issues should be extensively considered for large-scale dsRNA production. First, the quality and quantity of dsRNA from the two methods differs. For the in vivo method dsRNA quality and quantity greatly depend on the boundary sequences of the dsRNA sequence. Unless a T7 polymerase termination signal is introduced, the RNA transcription can extend further than the dsRNA region. When the bidirectional transcriptional reaction is performed, it is believed that aberrant forms of dsRNA -with a long and heterogeneous ssRNA tail- could be made. In contrast, when the PCR product containing two inverted promoters (see Fig. 6.1) and the target sequence is transcribed in vitro, the length of dsRNA is defined by dsRNA and additional short extended ssRNA tails. The short ssRNA tail can be trimmed off with an ssRNA specific nuclease. A comparison of the two methods is presented in Fig. 6.3. An identical

dsRNA of 310 bp was synthesized utilizing each method and its quality was compared. After the induction of T7 polymerase, the target RNA was produced (lanes 1 and 2). The derived dsRNA was heterogeneous even after dsRNA purification using a dsRNA purification kit (lanes 3 and 4). On the other hand, the dsRNA produced from the in vitro reaction shows homogeneity and was further enriched with the purification kit (lanes 5 and 6). Second, each dsRNA synthesis method requires its unique system. Although there is room for improvement in regards to cultivation conditions, the in vivo method must handle up to 600 to 800 l of culture volume in order to make 1 kg of dsRNA. The cost of media can be reduced through the development of cheaper alternative ingredients. Even when the high cost of the IPTG (inducer of expression molecule) is ignored, the system for continuous cultivation and centrifugation could be a big financial burden. In contrast, the in vitro method produces around 2–3 mg of purified RNA per ml of reaction. To make 1 kg of RNA, 400 l of transcriptional reaction is required. The reaction can be done in multiples of 10 l reaction tanks with a temperature control system. A PCR reaction system that can handle more than 1 l is essential for the in vitro transcriptional reaction. Additional requirement is the bulky supply of T7 RNA polymerase and pyrophosphatase. Third, the purification of dsRNA from each

Figure 6.3  Comparison of two RNA molecules produced by either the in vivo or in vitro method. A 310 bp dsRNA (see arrow) was cloned in a L4440 vector and transformed into the host bacterial strain E. coli HT115 (DE3). Four hours before or after the induction, the total nucleic acid was purified using the 5 min Cell/Virus DNA/RNA extraction kit (Biofactories, Inc. USA) (lanes 1 and 2). Both samples were further purified using the 5 min dsRNA Extraction Kit in order to enrich dsRNA (lanes 3 and 4). The same clone was used for PCR amplification and in vitro transcriptional reaction. The dsRNA produced by the reaction was analysed on 1% agarose gel before and after purification using the 5 min dsRNA Extraction Kit (lanes 5 and 6).

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system is the most essential factor to consider. Although unreported, the processing and purification of dsRNA from the in vivo bacterial culture have several critical factors. The purification of dsRNA after lysis of the bulk cell pellet requires technical breakthroughs in order to lower the cost of this step. Without a simple physical method to separate the dsRNA from other RNA, DNA and proteins, serial RNA extraction and sequential dsRNA enrichment steps must be included in order to achieve a pure product. However, the quality of dsRNA is often challenged by the presence of unwanted molecules, such as endotoxin and E. coli cells. Several attempts have been made to circumvent the rather complicated procedures. The bacteria in the culture could be inactivated by formamide without further purification. Regardless of its efficacy, the product may need a full safety evaluation for the presence of endotoxin and E. coli contamination before agricultural application. In contrast, RNA from in vitro transcription can be purified by simple physical methods: precipitation and washing with solutions of unique salt compositions and appropriate pH. With a couple of centrifugation steps, after enzymatic treatment, the desired dsRNA quality could be acquired. In addition to its homogeneity, the in vitro-produced dsRNA has a lesser chance of being contaminated with endotoxin or bacteria. If dsRNA is to be developed as sequence-specific agricultural product, the cost must be considered. The current cost of in vitro production is partitioned as follows: 35% for rNTP, 30% for DNA template synthesis, 5% for enzymes, 10% for purification and 20% for other chemicals in the transcription reaction. The cost of rNTP is the greatest, but can be reduced with increased market demand. The high cost of the DNA template can be reduced if the used DNA template is recycled after a physical separation from the reaction mixture. Although the cost of enzyme is relatively low, it can be further reduced using enzymes conjugated with an affinity tag. The used enzymes can be recycled after the reaction through affinity binding to a corresponding resin. When the demand for dsRNA in the global market is more than 100 kg/year as raw material, the price of 1 kg of dsRNA can be reduced to about $15,000. Additional field trials may be required to determine the precise unit price of dsRNA after determining the effective dose and appropriated formulations.

DsRNA application DsRNA delivery by mechanical application Studies have shown that dsRNA molecules, produced by either the in vitro or in vivo method, could be applied onto plant tissue by rubbing the dsRNA solution on plant leaf tissue where abrasive (e.g. carborundum, celite) has been previously applied to (Fig. 6.4A). This resembles the mechanical inoculation of plants with virus-containing plant sap for establishing infection. Alternatively, spraying the dsRNA solution (e.g. bacterially produced dsRNAs [Mitter et al., 2017 and references within]) onto plant could also induce resistance in the host. This means that the above-described dsRNA application could be useful only for laboratory use as it is a labour-intensive method; thus, it is not foreseen to be used in high-throughput dsRNA application in agriculture. DsRNA delivery by bombardment Various equipment, of varying costs, have been developed for the delivery of nucleic acids into plants in a high-throughput manner. Such equipment will facilitate, expedite and keep the cost of application for the dsRNA application step low. For example, the HandGun for particle bombardment (Fig.6.4B), the BIM-10 (Fig. 6.4C), the CONRAD air-brush gun AFC-250A (Fig. 6.4D), the Biojector-2000 (Fig. 6.4E) and a hand-held air-gun (Fig. 6.4F) have been used for delivery of dsRNAs or other nucleic acids into plants. Other means of dsRNA delivery onto plants The cost of dsRNA application in agriculture should be low and therefore cheap means of dsRNA delivery are needed. In this respect, means such as spraying, drenching (root soaking), delivery by symbionts or plant viruses, trunk injections are considered. Spray induced gene silencing (designated as SIGS) is a new innovative strategy for protecting crops from pathogen infection. SIGS is powerful, fast and will not increase the cost of treating plants if dsRNA is to be included in the spraying programme of a crop. The integration of nanotechnology in SIGS could improve significantly

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Figure 6.4 Means and devices for double-stranded RNA delivery into plants. (A) Mechanical delivery application method of dsRNA on plant leaf surfaces. (B) HandGun for particle bombardment (courtesy of V. Gaba, ARO, Volcani Center; IL, Gal-On et al., 1997). (C) BIM-10 in action delivering dsRNA onto plants (courtesy of GeORION Ltd., GR). The liquid bottle contains the dsRNA along with carborundum as an abrasive. (D) CONRAD air-brush gun AFC-250A (courtesy of A. Dalakouras and M. Wassenegger, RLP AgroScience, DE; Dalakouras et al., 2016). (E) Biojector 200 delivering nucleic acids in citrus stem (courtesy of I. Lavagi, T. Dang and G. Vidalakis, University of California, Riverside, USA). (F) Airgun bombardment device delivering infectious viral clone (courtesy of L. Predajna, Z. Subr and M. Glasa, Institute of Virology, SK; Predajna et al., 2010).

dsRNA application onto plant surfaces (e.g. nontoxic, degradable spray of dsRNAs; Mitter et al., 2017). Various strategies using chemically modified siRNAs, liposomes and nanoparticles have been considered and developments in delivery of silencing RNAs are expected in the near future. Drenching is also a possibility since it has been shown that plant roots absorb dsRNA (Li et al., 2015). It was shown that RNAi could be activated

by non-pathogenic bacteria engineered to manufacture and deliver silencing short hairpin RNA (shRNA) to target cells (Keates et al., 2008). Such trans-kingdom RNAi (tkRNAi) could provide a viable means to accomplish therapeutic RNAi in agriculture if the appropriate bacterium is selected, eliminate the cost of siRNA manufacture and could facilitate high-throughput in vivo functional genomics screening. Plant symbionts could be vehicles to deliver dsRNA in planta. Several developments are

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required before this approach becomes applicable. The symbiont needs to be culturable, amenable to transformation and to well-established molecular techniques. The issue of interaction with the host species needs to be considered. Plant viruses have been suggested to perform virus-induced gene silencing (VIGS). The generation of symptomless (mild symptoms may be accepted) virus-vectors or isolation of similar from nature is needed for the host species targeted. Such genetic material should be well characterized molecularly and biologically. There is a limitation of size that could be expressed in VIGS vectors and posses some restrictions in adopting this method. Trunk injection similar to that used for administration of chemical compounds against fungi, bacteria, phytoplasmas, insects or nutrient deficiency problems (Dal Maso et al., 2017; Huang et al., 2016; Gurr et al., 2016; Aćimović et al., 2015; Fernandez-Escobar et al., 1993) may also be useful for dsRNA application. This application method needs a special injection tool; however it uses the least amount of active compound in comparison to spraying the tree and limits environmental exposure. DsRNA application in agriculture – An industrial perspective With advances in genomic studies, including sequencing, dsRNA products may be the next generation of crop protection technology. With its target specificity but lack of environmental toxicity, the product is incredibly promising. Since dsRNA is not very stable in the field, the residual environmental issue can also be minimized. When industrial product development is considered, one must address the general principles for an agricultural product. It should have a potent efficacy, low cost, high safety, as well as consistent quality. Although there are multiple studies showing that dsRNA approaches are applicable to diverse target organisms, the actual product development may wait for large field application. DsRNA-mediated resistance Exogenous application of dsRNA molecules onto plants for the effective control of plant virus diseases and/or their insect-vectors has to consider two important features that could determine the

usefulness of the application in agriculture: a) the life-span of dsRNA and b) the capacity of dsRNA to spread systemically in planta. These features are discussed below. Life-span of dsRNA Several studies suggest that dsRNA molecules are much less prone to degradation relative to single-stranded RNA molecules. Semi-quantitative RT-PCR assays in watermelon and squash plants, upon topical application of dsRNA, revealed that the dsRNA remained remarkably stable over time at the leaves of topical application in watermelon and squash [at least 14 days post inoculation (dpi)] (Fig. 6.5). The gradual degradation from 6 to 14 dpi may be due to the processing of the exogenous dsRNA by the plant RNAi machinery for the production of siRNAs. By using a semi-quantitative stem–loop RT-PCR specific siRNAs, deriving from a long dsRNA fragment of the HC-Pro gene of Zucchini yellow mosaic virus (ZYMV), were detected suggesting that the applied dsRNA is processed in planta being a source of siRNAs with an antiviral activity that could last for a significant period of time (Kaldis et al., 2017). Results from different research groups, which utilize variable protocols for the exogenous application of dsRNA onto plants (carborundum, spraying, drenching), confirm that the stability of dsRNA molecules in planta for a relatively long time is a common situation for many plant species, both monocots and dicots. For example, by using real-time RT-PCR the exogenously applied dsRNA for p126 gene of TMV was detected in local leaves of tobacco plants for at least 9 dpi (Konakalla et al., 2016). Detection of exogenous dsRNA was achieved for several weeks after its application in tomato plants (Voloudakis’ lab). Another study in barley leaves showed that a dsRNA targeting the CYP51 genes of Fusarium graminearum was detected by Northern blotting at the site of application upon spraying, for at least 7 days (Koch et al., 2016). Wang and colleagues showed, in an indirect way, the durability of dsRNA application for protection purposes. They sprayed the surface of a variety of fruits and vegetables with dsRNA targeting the DCL1 and DCL2 genes of the harmful fungus Botrytis cinerea and found that the dsRNA confers protection for up to 8 days after spaying, suggesting that 8 days constitute the active life-span of dsRNA

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Figure 6.5  Detection of dsRNA ZYMV HC-Pro in local (treated) and systemic (non-treated) leaves of watermelon and squash plants. (A) Watermelon: dsRNA ZYMV HC-Pro was exogenously applied onto one leaf of each plant, via mechanical inoculation (carborundum). RNA extraction was performed from local and systemic tissue, at regular time points, from 1 to 14 dpi (days post inoculation). The presence of dsRNA ZYMV HC-Pro at each time point was determined by semi-quantitative RT-PCR. Different numbers of PCR cycles were used, to clearly visualize the degradation pattern of dsRNA over time, and the differences between local and systemic tissue. Results from non-treated plants (mock) are shown on the left. (B) Squash: dsRNA ZYMV HC-Pro dsHC application and subsequent analysis were performed as described above for watermelon plants. In both (A) and (B), a 100 bp DNA ladder was used (New England Biolabs, USA).

upon spraying and after this time point dsRNA falls below a critical threshold (Wang et al., 2016). San Miguel and Scott (2016); using a slightly different approach in potato, they concluded that the active life-span of dsRNA molecules could be even longer. They exogenously applied dsRNA targeting the actin gene of a potato pest on the surface of potato leaves, and left the leaves to dry. Their results showed that the dsRNA, once dried on the surface, is very resistant to leaf wash by water, and is able to confer protection against the respective insect pest for at least four weeks. The variation in the period of active life-span of dsRNA molecules presented above may be due to the variable modes of application or to the different plant species. This information will be very useful in experimental trials in field conditions, in which repeated treatments with dsRNA at regular time points could result in efficient protection against pathogens causing devastating plant diseases.

Adherence and stability of dsRNA molecules could extend its life span and effectiveness. For example, topical application of BioClay-loaded dsRNA was reported to significantly prolong and increase the systemic protection of tobacco against Cucumber mosaic virus (CMV) (Mitter et al., 2017). In planta movement of dsRNA Earlier studies in tobacco did not detect dsRNA accumulation in systemic tissues after its exogenous application in local leaves, suggesting that the applied dsRNA does not possess capacity for long transport (Tenllado and Díaz-Ruíz, 2001). However, these studies were based on Northern blotting, whose sensitivity may not be high enough if the concentration of dsRNA molecules in systemic leaves is below the detection limit. Indeed, as revealed by RT-PCR assays in watermelon and squash plants (Fig. 6.5), dsRNA amplification zones at the systemic (untreated) leaves clearly

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can be detected up to 14 dpi if the number of PCR cycles increase to 35. Interestingly, the degradation pattern of dsRNA at systemic leaves is similar to the one found for the local (treated) leaves, suggesting that similar mechanisms are involved, e.g. the processing of the dsRNA by the endogenous RNAi machinery. Similar conclusions can be drawn from other studies also. For example, application of dsRNA for p126 gene of Tobacco mosaic virus (TMV) in tobacco results in a rapid systemic movement from as early as 1 hpi (hour post inoculation) up to 9 dpi, as revealed by real-time RT-PCR (Konakalla et al., 2016). In tomato plants, detection of exogenously applied dsRNA at systemic tissue was achieved for several weeks after dsRNA application at local leaves (Voloudakis’ lab). Additional evidence for the capacity of dsRNA for long transport exists from studies that use drenching as the application method of dsRNA (Hunter et al., 2012; Li et al., 2015). These studies, which were performed in trees, grapevines, Arabidopsis, rice and maize, showed that dsRNA applied by irrigation can be absorbed by plant roots and transferred long distance. The most possible road for the dsRNA transport is through the plant vasculature. Indeed, Koch and colleagues in their elegant study provided the most direct proof for the capacity of dsRNA to move throughout the plant via its vasculature. The researchers produced fluorescently labelled dsRNA molecules targeting the CYP51 genes of Fusarium graminearum and topically applied them on a restricted area of barley leaves. Then, by using confocal laser scanning microscopy, they detected the labelled dsRNAs inside mesophyll cells, xylem vessels, phloem parenchyma cells and companion

cells, proving that the dsRNA from the leaf surface is absorbed by the mesophyll plant cells and then transferred to other areas through the vascular system. Interestingly, this transport has a bioactive potential, because it confers protection against Fusarium graminearum even when this fungus is applied to a distal part of the leaf, not subjected to dsRNA spaying (Koch et al., 2016). The above studies reinforce the concept of exogenous dsRNA application as a bioprotective method against several diseases, including virus diseases, because the movement of dsRNA means that there is spreading of the RNA silencing effect. The reasons for the lower amount of detected dsRNA in systemic versus treated leaves are not well understood, and further work is needed to clarify this point. One possibility is that the plant vasculature may have certain limitations and does not allow transport of high amounts of dsRNA. Another explanation could be that during transport there are significant losses due to unknown dsRNA degradation mechanisms. To improve the protective effect of the applied dsRNA, one possible way that needs further experimentation is to use chemical adjuvants that generally stabilize the bioactive compounds, in order to further increase the lifespan and the action of dsRNAs (Wang et al., 2016). Examples of topical dsRNA application against plant viruses DsRNA molecules have been produced in vitro and in vivo and applied in different pathosystems with very good results in viral protection (Fig. 6.6 and Table 6.1).

Figure 6.6  Results of the application of dsRNA at large scale. Photographs of cucurbit plants (25 days after dsRNA vaccination). (i) Plants have received only Zucchini mosaic virus (ZYMV). (ii) Plants have received ZYMV along with the dsRNA vaccine, namely the dsRNA for ZYMV_HC-Pro); plants having larger biomass (courtesy of GeORION Ltd., GR).

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Application of dsRNA (in agriculture/ aquaculture) beyond plant virus control DsRNA could be the future insecticide. It has been shown that spraying specific dsRNA molecules could control insects, especially beetles including the western maize rootworm and Colorado potato beetle (Bolognesi et al., 2012; Baum et al., 2007; Palli, 2014). Major agrochemical companies are in the process of developing sprayable dsRNAbased insecticides that target agricultural pests. Alternatively, artificial feed or bait containing a species-specific pheromone can be formulated along with specific dsRNA. This approach may be far more cost-effective with minimal environmental effect. Aiming at the control of insect-vectors, the exogenously applied onto tomato leaves dsRNA was detected in by mites, aphids and whiteflies supporting the notion that topical dsRNA application could be used for dsRNA delivery to plant pests (Gogoi et al., 2017). This provides an alternative, and most importantly, a non-transgenic approach to deliver dsRNAs into plant pests in order to target vital genes of the targeted pest. As a result this method could be added in the IPM armoire of crop pests. Honeybees are the main pollinators for flowering plants in nature, and thus they maintain and drive the biodiversity of ecosystems. Without pollination, it is estimated that one third of our dinner table food will disappear. Several studies have shown that orally consumed dsRNA in sugar solution suppresses viral infection in the honeybee (Maori et al., 2009). After eradicating the possibility of dsRNA (insect protector) as general immune stimulant in honeybees, applications may be extended to diverse types of viral diseases. Shrimp cultivation has a market value of more than $15 billion. A significant amount of shrimp cultivation has been damaged by the outbreak of viral diseases such as the White spot syndrome virus (WSSV). Studies have shown that the dsRNA in shrimp is processed into siRNAs for antiviral defence (Nilsen et al., 2017). An intramuscular injection of dsRNA into shrimp shows prominent protection against the WSSV. For practical field applications, the dsRNA should be coated with shrimp feed. With data from field trials indicating viral protection, feed formulated with dsRNA may

be one of the most valuable products utilizing the principle. Other uses of dsRNA, which are not related to viruses, have been proposed and proven that could control the targeted organism (e.g. fungi) upon exogenous application. Limitations of topical dsRNAmediated resistance There are several concerns in using the topical application of dsRNA. First, the RNAi off-target effects employing the dsRNA-mediated resistance either via transgenesis or transiently, as discussed in this chapter, as well as CRISPR/Cas9 target selection is of concern. The selection of the target sequence needs to be done wisely. Available bioinformatics tools that could screen DNA sequences for offtarget effects prove that computational analyses are heavily involved in decision making of the target sequence. In order for the dsRNA molecules to reach an industrial application, the environmental fate of dsRNA needs to be determined. It was found that dsRNA was degraded and its biological activity was undetectable within approximately 2 days after application to soil; dsRNA was undetectable as measured by QuantiGene (Dubelman et al., 2014). Thus, dsRNA is considered not to be a very stable molecule in the field. In contrast, dsRNA stability is desirable within plant. Current evidence indicates that dsRNA could be detected in planta by RT-PCR for at least two (Fig. 6.5) to more weeks (Voloudakis laboratory). The long-term effectiveness of the dsRNA vaccination approach against plant viruses depends on it’s in planta stability and needs to be determined for the different host species. Within plant, dsRNA amounts decline over time since dsRNA molecules do not amplify in planta; dsRNA could be detected for up to 14 days post application in watermelon and squash (Fig. 6.5). This could have positive consequences for keeping an active antiviral RNAi in the host if the concentration threshold is met. It was proposed that naked dsRNA could ‘survive’ in planta for about five days upon foliar application (Mitter et al., 2017). Application strategies can be improved by mixing the dsRNAs with stabilizing chemical reagents that will increase the length and induction of plant protection through RNAi. Such an invention was proposed by using

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‘bioclay’ with the dsRNA detection made for ≈ 30 days after topical application (Mitter et al., 2017). DsRNA vaccinated plants, as transgenic plants, could be at risk if another virus, except the one that is targeted, infects the dsRNA-treated plants. In such cases (e.g. CMV-TYLCV in tomato [Butterbach et al., 2014]) the dsRNA treatment will be ineffective. Although this is a serious warning to plant breeders, in case of dsRNA-mediated transgenic resistance, it should also be considered in the transient application of dsRNA. The use of hybrid dsRNA molecules, namely constructs that contain sequences for both viruses, could reduce the risk of such negative effects due to plant virus synergism. Conclusion and future prospects Transgenic RNAi has been used against plant viruses in the lab since 1990s. However, in areas of the world where transgenic plants are not permitted, the non-transgenic approaches described in this chapter could exploit the plant RNAi for the control of viruses infecting crops. Furthermore, the non-transgenic application of dsRNA could be employed for transformation of recalcitrant plant species and resistance induction could be developed rapidly against evolving viruses or emerging viral diseases. Advancements in delivery means of dsRNA will provide significant advantages to this transient RNA silencing method for field scale application. Viruses have a limited coding capacity and they depend on the host in order to complete their life cycle, such as the need of translation components. A large number of host proteins interact in viral infection with viruses exploiting the host protein synthesis machinery; such host proteins could be considered products of susceptibility genes. Topical application of dsRNA leading to silencing these susceptibility genes, if plant development is not compromised, could be a novel means to induce virus control. The use of transient silencing assays such as the dsRNA vaccination and the virus-induced gene silencing (VIGS) could be used to quickly screen for candidate susceptibility genes, especially for plant species that are difficult to transform. The environmental fate of double-stranded RNA in agricultural soils is high in the research agenda. DsRNA is unlikely to persist or accumulate

freely in the environment for long time (Dubelman et al., 2014) and as a result it could be considered an environmentally friendly molecule. Such studies will allow the exposure of regulatory agencies to dsRNA in agricultural systems and thus help advance risk assessment of RNAi crop protection technology prior to its implementation. Off-target effects could potentially compromise RNAi’s biosafety and thus the topical application of dsRNAs. Improved design of highly specific RNAi constructs (Pooggin, 2017) (and CRISPR/Cas 9 targets) is needed and this could be accomplished by computational analysis and subsequent validation of designed RNAi constructs contributing thus in a full safety product evaluation. It is well accepted that the risk for off-target effect increases with the length of the dsRNA sequence. The current data available indicate that dsRNA as small as ≈ 400 bp (see Table 6.1) could be used reducing the abovementioned risk. The use of RNAi to combat insects (e.g. vectors of plant viruses) is anticipated to become an integral component of crop IPM worldwide in the near future. For practical reasons there is a need to have ample amounts of dsRNA molecules in planta for enabling trans-kingdom RNAi leading to an effective pest control. The latter may be difficult to achieve since dsRNA will be degraded by plant RNAi and thus means of avoiding plant RNAi need to be improvised. The molecular aspects of RNAi in pests of plants (e.g. differences between plant and pest RNAi), the escape of dsRNA degradation by the pest and improvement in dsRNA import in the cytoplasm of pests need to be examined in detail in order to develop the future dsRNA-based insecticides. One should not underestimate the usefulness of exogenous application of dsRNA to screen for the most efficient RNAi constructs that could eventually get incorporated in the plant genome (transgenic RNAi). For example, the use of transient silencing assays such as the dsRNA vaccination and VIGS could be used to quickly screen for candidate plant susceptibility genes. Furthermore, this method could be used to determine the minimum length needed for an efficient RNAi construct having in mind that reduction of the risk for off-target effects is mandated. Other issues that need investigation in the future are the induction of RdDM as a potential epigenetic

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antiviral means and the effect of temperature in the topical application of dsRNA. Advances in the area of plant amplicon discovery could lead in the development of molecules/vehicles that could produce dsRNAs continuously in planta, thus contributing in long lasting effect. Circular molecules have been computationally identified in plants and this could pave the way of developing useful amplicons. Along these lines, the area of cross-protection could be revisited in the near future. Basic research in pathways that run prior to and/or in parallel, or interplay with the RNAi pathway will spark new ideas for virus control. As novel opportunities arise for plant antiviral means, the exogenous dsRNA application could find new targets for the regulation of plant genes involved in virus proliferation during infection. For example, specific activation of autophagy or proteasome degradation of plant virus proteins would enhance plant resistance to viruses. Peptide-nucleic acid interactions will continue to play a key role in developing antiviral biotechnological approaches. Nanotechnology advents could improve the length and efficiency of dsRNA as inducer of RNAi. Regarding the latter, research is under way in the medical sector and could possibly be transferred in agriculture field if economics allow it. Finally, the cost of dsRNA manufacturing and application as well as the regulatory framework will influence whether at the end of the day this methodology will see an industrial application. The assessment of safety aspects and the consumer perception of this non-transgenic technology need to be investigated so that the public will accept the future introduction of such means of control. Acknowledgements Some of the data presented in this review have been obtained in the frame of the projects: A ‘Pythagoras II’ funded by General Secretariat of Research and Technology of Greece B ‘sRNAvac’ funded by General Secretariat of Research and Technology of Greece C ‘COST FA0806’ funded by Cooperation in Science and Technology (COST), EU D ‘Erasmus Mundus Action BRAVE’ funded by Education, Audiovisual and Culture Executive Agency, EU.

Projects were conducted at the Laboratory of Plant Breeding and Biometry of the Agricultural University of Athens (coordinator of projects a, b, c and d: A.E. Voloudakis), as well as at the Laboratory of Bacteriology of the Benaki Phytopathological Institute (participant in projects a, b and c). References Aalto, A.P., Sarin, L.P., van Dijk, A.A., Saarma, M., Poranen, M.M., Arumäe, U., and Bamford, D.H. (2007). Largescale production of dsRNA and siRNA pools for RNA interference utilizing bacteriophage phi6 RNAdependent RNA polymerase. RNA 13, 422–429. Abel, P.P., Nelson, R.S., De, B., Hoffmann, N., Rogers, S.G., Fraley, R.T., and Beachy, R.N. (1986). Delay of disease development in transgenic plants that express the Tobacco mosaic virus coat protein gene. Science 232, 738–743. Aćimović, S.G., Zeng, Q., McGhee, G.C., Sundin, G.W., and Wise, J.C. (2015). Control of fire blight (Erwinia amylovora) on apple trees with trunk-injected plant resistance inducers and antibiotics and assessment of induction of pathogenesis-related protein genes. Front. Plant Sci. 6, 16. https://doi.org/10.3389/ fpls.2015.00016 Ali, Z., Ali, S., Tashkandi, M., Zaidi, S.S.-A., Mahfouz, M.M. (2016). CRISPR/Cas9-mediated immunity to Geminiviruses: Differential Interference and Evasion. Sci Rep 6, 26912. https://doi.org/10.1038/srep26912 Aliyari, R., and Ding, S.W. (2009). RNA-based viral immunity initiated by the Dicer family of host immune receptors. Immunol. Rev. 227, 176–188. https://doi. org/10.1111/j.1600-065X.2008.00722.x Baulcombe, D.C. (2004). RNA silencing in plants. Nature 431, 356-363. Baum, J.A., Bogaert, T., Clinton, W., Heck, G.R., Feldmann, P., Ilagan, O., Johnson, S., Plaetinck, G., Munyikwa, T., Pleau, M., et al. (2007). Control of coleopteran insect pests through RNA interference. Nat. Biotechnol. 25, 1322–1326. Baumberger, N., and Baulcombe, D.C. (2005). Arabidopsis ARGONAUTE1 is an RNA Slicer that selectively recruits microRNAs and short interfering RNAs. Proc. Natl. Acad. Sci. U.S.A. 102, 11928–11933. Bernstein, E., Caudy, A.A., Hammond, S.M., and Hannon, G.J. (2001). Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409, 363– 366. https://doi.org/10.1038/35053110 Blevins, T., Rajeswaran, R., Shivaprasad, P.V., Beknazariants, D., Si-Ammour, A., Park, H.S., Vazquez, F., Robertson, D., Meins, F., Hohn, T., et al. (2006). Four plant Dicers mediate viral small RNA biogenesis and DNA virus induced silencing. Nucleic Acids Res. 34, 6233–6246. Bolognesi, R., Ramaseshadri, P., Anderson, J., Bachman, P., Clinton, W., Flannagan, R., Ilagan, O., Lawrence, C., Levine, S., Moar, W., et al. (2012). Characterizing the mechanism of action of double-stranded RNA activity against western corn rootworm (Diabrotica virgifera virgifera LeConte). PLOS ONE 7, e47534. https://doi. org/10.1371/journal.pone.0047534

138  | Voloudakis et al.

Butterbach, P., Verlaan, M.G., Dullemans, A., Lohuis, D., Visser, R.G., Bai, Y., and Kormelink, R. (2014). Tomato yellow leaf curl virus resistance by Ty-1 involves increased cytosine methylation of viral genomes and is compromised by Cucumber mosaic virus infection. Proc. Natl. Acad. Sci. U.S.A. 111, 12942–12947. https://doi. org/10.1073/pnas.1400894111 Carbonell, A., Martínez de Alba, A.E., Flores, R., and Gago, S. (2008). Double-stranded RNA interferes in a sequence-specific manner with the infection of representative members of the two viroid families. Virology 371, 44–53. Chandrasekaran, J., Brumin, M., Wolf, D., Leibman, D., Klap, C., Pearlsman, M., Sherman, A., Arazi, T., and GalOn, A. (2016). Development of broad virus resistance in non-transgenic cucumber using CRISPR/Cas9 technology. Mol. Plant Pathol. 17, 1140–1153. https:// doi.org/10.1111/mpp.12375 Christie, M., Brosnan, C.A., Rothnagel, J.A., and Carroll, B.J. (2011). RNA decay and RNA silencing in plants: competition or collaboration? Front. Plant Sci. 2, 99. https://doi.org/10.3389/fpls.2011.00099 Conti, G., Zavallo, D., Venturuzzi, A.L., Rodriguez, M.C., Crespi, M., and Asurmendi, S. (2017). TMV induces RNA decay pathways to modulate gene silencing and disease symptoms. Plant J. 89, 73–84. https://doi. org/10.1111/tpj.13323 Csorba, T., Kontra, L., and Burgyán, J. (2015). Viral silencing suppressors: Tools forged to fine-tune host-pathogen coexistence. Virology 479-480, 85–103. https://doi. org/10.1016/j.virol.2015.02.028 Csorba, T., Pantaleo, V., and Burgyán, J. (2009). RNA silencing: an antiviral mechanism. Adv. Virus Res. 75, 35–71. https://doi.org/10.1016/S00653527(09)07502-2 Dal Maso, E., Cocking, J., and Montecchio, L. (2017). An enhanced trunk injection formulation of potassium phosphite against chestnut ink disease. Arboricult. J. 39, 125-141. Dalakouras, A., Wassenegger, M., McMillan, J.N., Cardoza, V., Maegele, I., Dadami, E., Runne, M., Krczal, G., and Wassenegger, M. (2016). Induction of silencing in plants by high-pressure spraying of in vitro-synthesized small RNAs. Front. Plant Sci. 7, 1327. https://doi. org/10.3389/fpls.2016.01327 de Ronde, D., Butterbach, P., and Kormelink, R. (2014). Dominant resistance to plant viruses. Front. Plant Sci. 5, 307. https://doi.org/10.3389/fpls.2014.00307 Ding, S.W. (2010). RNA-based antiviral immunity. Nat. Rev. Immunol. 10, 632–644. https://doi.org/10.1038/ nri2824 Duan, C.G., Wang, C.H., and Guo, H.S. (2012). Application of RNA silencing to plant disease resistance. Silence 3, 5. https://doi.org/10.1186/1758-907X-3-5 Dubelman, S., Fischer, J., Zapata, F., Huizinga, K., Jiang, C., Uffman, J., Levine, S., and Carson, D. (2014). Environmental fate of double-stranded RNA in agricultural soils. PLOS ONE 9, e93155. https://doi. org/10.1371/journal.pone.0093155 Fernandez-Escobar, R., Barranco, D., and Benlloch, M. (1993). Overcoming iron chlorosis in olive and peachtrees using a low-pressure trunk-injection method. HortSci. 28, 192-194.

Gal-On, A., Meiri, E., Elman, C., Gray, D.J., and Gaba, V. (1997). Simple handheld devices for the efficient infection of plants with viral encoding constructs by particle bombardment. J. Virol. Methods 64, 103-110. Gal-On A., and Shiboleth, Y.M. (2006). Cross-protection. In: Natural Resistance Mechanisms of Plants to Viruses. Loebenstein G., Carr, J.P. (eds). Springer, Dordrecht. Gan, D., Zhang, J., Jiang, H., Jiang, T., Zhu, S., and Cheng, B. (2010). Bacterially expressed dsRNA protects maize against SCMV infection. Plant Cell Rep. 29, 1261–1268. https://doi.org/10.1007/s00299-010-0911-z Garcia-Ruiz, H., Takeda, A., Chapman, E.J., Sullivan, C.M., Fahlgren, N., Brempelis, K.J., and Carrington, J.C. (2010). Arabidopsis RNA-dependent RNA polymerases and dicer-like proteins in antiviral defense and small interfering RNA biogenesis during Turnip mosaic virus infection. Plant Cell 22, 481–496. https://doi. org/10.1105/tpc.109.073056 Garcia, D., Garcia, S., and Voinnet, O. (2014). Nonsensemediated decay serves as a general viral restriction mechanism in plants. Cell Host Microbe 16, 391–402. https://doi.org/10.1016/j.chom.2014.08.001 Ghoshal, B., and Sanfaçon, H. (2015). Symptom recovery in virus-infected plants: Revisiting the role of RNA silencing mechanisms. Virology 479-480, 167–179. https://doi.org/10.1016/j.virol.2015.01.008 Gogoi, A., Sarmah, N., Kaldis, A., Perdikis, D., and Voloudakis, A. (2017). Plant insects and mites uptake double-stranded RNA upon its exogenous application on tomato leaves. Planta 246, 1233–41. Gordon, K.H., and Waterhouse, P.M. (2007). RNAi for insect-proof plants. Nat. Biotechnol. 25, 1231–1232. Gurr, G.M., Johnson, A.C., Ash, G.J., Wilson, B.A., Ero, M.M., Pilotti, C.A., Dewhurst, C.F., and You, M.S. (2016). Coconut lethal yellowing diseases: A Phytoplasma threat to palms of global economic and social significance. Front. Plant Sci. 7, 1521. https://doi. org/10.3389/fpls.2016.01521 Holeva, M., Sclavounos, A.P., Kyriakopoulou, P.E., and Voloudakis, A.E. (2006). In vitro produced dsRNA induces resistance to a sever Hellenic CMV isolate in tobacco and tomato. 8th International Congress of Plant Molecular Biology, 20-25 August 2006, Adelaide, Australia. Holeva, M.C., Sclavounos, A.P., Milla, S.P., Kyriakopoulou, P.E., and Voloudakis, A.E. (2007). External application of dsRNA of the capsid protein (CP) or 2b gene of CMV reduces the severity of CMV-infection in tobacco. ΧΙΙΙ International Congress on Molecular Plant-Microbe Interactions, 21-27 July 2007, Sorrento, Italy. Huang, L., Jin, J., Deighan, P., Kiner, E., McReynolds, L., and Lieberman, J. (2013). Efficient and specific gene knockdown by small interfering RNAs produced in bacteria. Nat. Biotechnol. 31, 350–356. https://doi. org/10.1038/nbt.2537 Huang, J., Zhang, J., Li, Y., Li, J., and Shi, X.H. (2016). Evaluation of the effectiveness of insecticide trunk injections for control of Latoia lepida (Cramer) in the sweet olive tree Osmanthus fragrans. PeerJ 4, e2480. https://doi.org/10.7717/peerj.2480 Hunter, W.B., Glick, E., Paldi, N., and Bextine, B.R. (2012). Advances in RNA interference: dsRNA treatment in

Production and Application of Double-Stranded RNA Against Plant Viruses |  139

trees and grapevines for insect pest suppression. SWest Entomol. 37, 85-87. Hutvagner, G., and Simard, M.J. (2008). Argonaute proteins: key players in RNA silencing. Nat. Rev. Mol. Cell Biol. 9, 22–32. Kaldis, A., Berbati, M., Melita, O., Reppa, C., Holeva, M., Otten, P., and Voloudakis, A. (2018). Exogenously applied dsRNA molecules deriving from the Zucchini yellow mosaic virus (ZYMV) genome move systemically and protect cucurbits against ZYMV. Mol. Plant Pathol. 19, 883–95. https://doi.org/10.1111/mpp.12572 Keates, A.C., Fruehauf, J., Xiang, S., and Li, C.J. (2008). TransKingdom RNA interference: a bacterial approach to challenges in RNAi therapy and delivery. Biotechnol. Genet. Eng. Rev. 25, 113–127. Koch, A., and Kogel, K.H. (2014). New wind in the sails: improving the agronomic value of crop plants through RNAi-mediated gene silencing. Plant Biotechnol. J. 12, 821–831. https://doi.org/10.1111/pbi.12226 Koch, A., Biedenkopf, D., Furch, A., Weber, L., Rossbach, O., Abdellatef, E., Linicus, L., Johannsmeier, J., Jelonek, L., Goesmann, A., et al. (2016). An RNAi-based control of Fusarium graminearum infections through spraying of long dsRNAs involves a plant passage and is ccontrolled by the fungal silencing machinery. PLOS Pathog. 12, e1005901. https://doi.org/10.1371/ journal.ppat.1005901 Konakalla, N.C., Kaldis, A., Berbati, M., Masarapu, H., and Voloudakis, A.E. (2016). Exogenous application of double-stranded RNA molecules from TMV p126 and CP genes confers resistance to TMV in tobacco. Planta 244, 961–969. https://doi.org/10.1007/s00425-0162567-6 Lau, S.E., Mazumdar, P., Hee, T.W., Song, A.L.A., Othman, R.Y., and Harikrishna, J.A. (2014). Crude extracts of bacterially-expressed dsRNA protect orchid plants against Cymbidium mosaic virus during transplantation from in vitro culture. J. Hort. Sci. Biotechnol. 89, 569-576 Li, H., Guan, R., Guo, H., and Miao, X. (2015). New insights into an RNAi approach for plant defence against piercing-sucking and stem-borer insect pests. Plant Cell Environ. 38, 2277–2285. https://doi.org/10.1111/ pce.12546 Lindbo, J.A. (2012). A historical overview of RNAi in plants. Methods Mol. Biol. 894, 1–16. https://doi. org/10.1007/978-1-61779-882-5_1 Machado, J.P.B., Calil, I.P., Santos, A.A., and Fontes, E.P.B. (2017). Translational control in plant antiviral immunity. Genet. Mol. Biol. 40 (Suppl. 1), 292–304. Mahy, B.W.J., and van Regenmortel, M.H.V. (2009). Desk Encyclopedia of Plant and Fungal Virology. Cambridge, MA: Academic Press. Mao, Y.B., Cai, W.J., Wang, J.W., Hong, G.J., Tao, X.Y., Wang, L.J., Huang, Y.P., and Chen, X.Y. (2007). Silencing a cotton bollworm P450 monooxygenase gene by plantmediated RNAi impairs larval tolerance of gossypol. Nat. Biotechnol. 25, 1307–1313. Maori, E., Paldi, N., Shafir, S., Kalev, H., Tsur, E., Glick, E., and Sela, I. (2009). IAPV, a bee-affecting virus associated with Colony Collapse Disorder can be silenced by dsRNA ingestion. Insect Mol. Biol. 18, 55–60. https:// doi.org/10.1111/j.1365-2583.2009.00847.x

Mitter, N., Worrall, E.A., Robinson, K.E., Li, P., Jain, R.G., Taochy, C., Fletcher, S.J., Carroll, B.J., Lu, G.Q., and Xu, Z.P. (2017a). Clay nanosheets for topical delivery of RNAi for sustained protection against plant viruses. Nat Plants 3, 16207. https://doi.org/10.1038/ nplants.2016.207 Mitter, N., Worrall, E.A., Robinson, K.E., Xu, Z.P., and Carroll, B.J. (2017b). Induction of virus resistance by exogenous application of double-stranded RNA. Curr Opin Virol 26, 49–55. Nilsen, P., Karlsen, M., Sritunyalucksana, K., and Thitamadee, S. (2017). White spot syndrome virus VP28 specific double-stranded RNA provides protection through a highly focused siRNA population. Sci Rep 7, 1028. https://doi.org/10.1038/s41598-017-01181-w Palli, S.R. (2014). RNA interference in Colorado potato beetle: steps toward development of dsRNA as a commercial insecticide. Curr. Opin. Insect Sci. 6, 1–8. https://doi.org/10.1016/j.cois.2014.09.011 Pooggin, M.M. (2013). How can plant DNA viruses evade siRNA-directed DNA methylation and silencing? Int. J. Mol. Sci. 14, 15233–15259. https://doi.org/10.3390/ ijms140815233 Pooggin, M.M. (2017). RNAi-mediated resistance to viruses: a critical assessment of methodologies. Curr. Opin. Virol. 26, 28–35. Predajňa, L., Nagyová, A., and Subr, Z. (2010). Simple and efficient biolistic procedure for the plant transfection with cDNA clones of RNA viruses. Acta Virol. 54, 303–306. Pyott, D.E., Sheehan, E., and Molnar, A. (2016). Engineering of CRISPR/Cas9-mediated potyvirus resistance in transgene-free Arabidopsis plants. Mol. Plant Pathol. 17, 1276–1288. https://doi.org/10.1111/mpp.12417 Robinson, K.E., Worrall, E.A., and Mitter, N. (2014). Double-stranded RNA expression and its topical application for non-transgenic resistance to plant viruses. J. Plant Biochem. Biotechnol. 23, 231–237. Rybicki, E.P. (2015). A Top Ten list for economically important plant viruses. Arch. Virol. 160, 17–20. https://doi.org/10.1007/s00705-014-2295-9 Safarova, D., Brasda, P., and Navratil, M. (2014). Effect of artificial dsRNA on infection of pea plants by Pea seed-borne mosic virus. Czsch J. Genet. Plant Breed. 50, 105-108. San Miguel, K., and Scott, J.G. (2016). The next generation of insecticides: dsRNA is stable as a foliar-applied insecticide. Pest Manag. Sci. 72, 801–809. https://doi. org/10.1002/ps.4056 Sanford, J.C., and Johnston, S.A. (1985). The concept of parasite-derived resistance-deriving resistance genes from the parasite’s own genome. J. Theor. Biol. 113, 395405. Scholthof, K.B., Adkins, S., Czosnek, H., Palukaitis, P., Jacquot, E., Hohn, T., Hohn, B., Saunders, K., Candresse, T., Ahlquist, P., et al. (2011). Top 10 plant viruses in molecular plant pathology. Mol. Plant Pathol. 12, 938–954. https://doi.org/10.1111/j.13643703.2011.00752.x Senthil-Kumar, M., and Mysore, K.S. (2011). Caveat of RNAi in plants: the off-target effect. Methods Mol. Biol. 744, 13–25. https://doi.org/10.1007/978-1-61779123-9_2

140  | Voloudakis et al.

Soosaar, J.L., Burch-Smith, T.M., and Dinesh-Kumar, S.P. (2005). Mechanisms of plant resistance to viruses. Nat. Rev. Microbiol. 3, 789–798. Tamilarasan, S., and Rajam, M.V. (2013). Engineering crop plants for nematode resistance through host-derived RNA interference. Cell Dev. Biol. 2, 114. Tenllado, F., and Díaz-Ruíz, J.R. (2001). Double-stranded RNA-mediated interference with plant virus infection. J. Virol. 75, 12288–12297. https://doi.org/10.1128/ JVI.75.24.12288-12297.2001 Tenllado, F., Llave, C., and Díaz-Ruíz, J.R. (2004). RNA interference as a new biotechnological tool for the control of virus diseases in plants. Virus Res. 102, 85–96. https://doi.org/10.1016/j.virusres.2004.01.019 Tenllado, F., Martínez-García, B., Vargas, M., and Díaz-Ruíz, J.R. (2003). Crude extracts of bacterially expressed dsRNA can be used to protect plants against virus infections. BMC Biotechnol. 3, 3. Timmons, L., Court, D.L., and Fire, A. (2001). Ingestion of bacterially expressed dsRNAs can produce specific and potent genetic interference in Caenorhabditis elegans. Gene 263, 103–112. Voloudakis, A.E., Holeva, M.C., Sarin, L.P., Bamford, D.H., Vargas, M., Poranen, M.M., and Tenllado, F. (2015). Efficient double-stranded RNA production methods for utilization in plant virus control. Methods Mol. Biol. 1236, 255–274. https://doi.org/10.1007/978-1-49391743-3_19 Wang, M., Weiberg, A., Lin, F.M., Thomma, B.P., Huang, H.D., and Jin, H. (2016). Bidirectional cross-kingdom RNAi and fungal uptake of external RNAs confer plant protection. Nat Plants 2, 16151. https://doi. org/10.1038/nplants.2016.151 Wang, M.B., Masuta, C., Smith, N.A., and Shimura, H. (2012). RNA silencing and plant viral diseases. Mol. Plant Microbe Interact. 25, 1275–1285. https://doi. org/10.1094/MPMI-04-12-0093-CR

Wang, X.B., Wu, Q., Ito, T., Cillo, F., Li, W.X., Chen, X., Yu, J.L., and Ding, S.W. (2010). RNAi-mediated viral immunity requires amplification of virus-derived siRNAs in Arabidopsis thaliana. Proc. Natl. Acad. Sci. U.S.A. 107, 484–489. https://doi.org/10.1073/ pnas.0904086107 Wassenegger, M., Heimes, S., Riedel, L., and Sänger, H.L. (1994). RNA-directed de novo methylation of genomic sequences in plants. Cell 76, 567–576. Waterhouse, P.M., Wang, M.B., and Lough, T. (2001). Gene silencing as an adaptive defence against viruses. Nature 411, 834–842. https://doi.org/10.1038/35081168 Yang, D.L., Zhang, G., Tang, K., Li, J., Yang, L., Huang, H., Zhang, H., and Zhu, J.K. (2016). Dicer-independent RNA-directed DNA methylation in Arabidopsis. Cell Res. 26, 1264. https://doi.org/10.1038/cr.2016.122 Yin, G., Sun, Z., Liu, N., Zhang, L., Song, Y., Zhu, C., and Wen, F. (2009). Production of double-stranded RNA for interference with TMV infection utilizing a bacterial prokaryotic expression system. Appl. Microbiol. Biotechnol. 84, 323–333. https://doi.org/10.1007/ s00253-009-1967-y Zaidi, S.S., and Mansoor, S. (2017). Viral Vectors for Plant Genome Engineering. Front. Plant Sci. 8, 539. https:// doi.org/10.3389/fpls.2017.00539 Zaidi, S.S., Tashkandi, M., Mansoor, S., and Mahfouz, M.M. (2016). Engineering Plant Immunity: Using CRISPR/ Cas9 to Generate Virus Resistance. Front. Plant Sci. 7, 1673. https://doi.org/10.3389/fpls.2016.01673 Zorzatto, C., Machado, J.P., Lopes, K.V., Nascimento, K.J., Pereira, W.A., Brustolini, O.J., Reis, P.A., Calil, I.P., Deguchi, M., Sachetto-Martins, G., et al. (2015). NIK1mediated translation suppression functions as a plant antiviral immunity mechanism. Nature 520, 679–682. https://doi.org/10.1038/nature14171

Transgenic Virus-Resistant Papaya: Current Status and Future Trends Gustavo Fermin1, Paula Tennant2 and Sudeshna Mazumdar-Leighton3*

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1Instituto Jardín Botánico de Mérida, Faculty of Sciences, Universidad de Los Andes, Mérida, Venezuela. 2Department of Life Sciences, The University of the West Indies, Mona, Jamaica.

3Plant Biotic Interactions Lab, Department of Botany, Delhi University, Delhi, India.

*Correspondence: [email protected] https://doi.org/10.21775/9781910190814.07

Abstract Viruses pose a major threat to worldwide production of papaya. Transgenic papaya varieties ‘Rainbow’ and ‘SunUp’ developed in the United States have provided durable resistance to local strains of Papaya ringspot virus (PRSV). Posttranscriptional gene silencing (PTGS) which exploits sequence homology between a transgene and the corresponding region of the invading viral genome has been used to successfully obtain PRSVresistant plants in South America, the Caribbean, Asia, and Australia. Many of these transgenic lines await government and/or public approval prior to commercialization. This review re-emphasizes the success of Hawaiian PRSV-resistant transgenic papayas for sustainable virus resistance in the field, along with the availability of a proven transgenic toolkit. However emergence of new PRSV strains and mixed infections with viruses like Papaya leaf distortion mosaic virus and Papaya mosaic virus pose new challenges for future adoption of transgenic virus-resistant plants. In addition to PRSV, geminiviruses causing papaya leaf curl disease in Asia, Papaya meleira virus (and Papaya virus Q) in Brazil and Mexico are important targets. There is need to monitor field-level diversity and evolution of viruses against the backdrop of transgenic technologies available for next generation virus-resistant papaya benefiting farmers worldwide.

Introduction Papaya (Carica papaya L.) is a fast-growing dicotyledonous species belonging to family Caricaceae. It is native to the plains of eastern Central America and is believed to have evolved into the economically useful gynodioecious form as a result of centuries of anthropogenic selection (Storey, 1976; OECD, 2005). Papaya grows abundantly in tropics and subtropics (Manshardt, 1992) and is of horticultural importance due to its high nutritional and medicinal value (Malabadi et al., 2011). Papaya is cultivated in home gardens as well as commercial plantations and regarded as a cash crop (Tennant et al., 2007; Bau et al., 2008). Ripe papaya fruit is a rich source of vitamins and dietary fibre while the unripe fruit is used as a vegetable and in salads (Manshardt, 1992; Aravind et al., 2013; Azad et al., 2014). Proteolytic enzymes like papain and chymopapain are obtained from papaya latex. These cysteine proteases are used for industrial applications such as beer clearing, production of chewing gum and cosmetics, degumming cocoon silks, leather tanning and meat tenderizing (OECD, 2005; Amri and Mamboya, 2012). Almost all parts of the papaya plant from its roots to the seeds are used in traditional medicine based on Indian ‘Ayurveda’ and Chinese herbal remedies (Ye and Li, 2010; Manohar, 2013). Various studies have investigated the role of papaya tissues and derivatives as anti-plasmodial, anti-cancer, anti-dengue,

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anti-septic, anti-parasitic, anti-inflammatory and anti-diabetic agents (Aravind et al., 2013). Global production of papaya According to estimates available from the Food and Agriculture Organization (FAO) of the United Nations, the total production of papaya in 2014 was about 11.6 million metric tons from a cultivated area of 433,057 hectares (ha). India is the world’s largest producer of papaya, producing 5.5m MT annually; followed by Brazil (1.6m MT), Indonesia (0.9m MT), Mexico (0.8m MT) and Nigeria (0.8m MT) (FAO, 2016). Various cultivars are grown for fruit and/or latex. Papaya production in Hawaii is said to be based on the ‘Solo’ cultivar where fruits are obtained from hermaphrodite trees of an inbred gynodioecious strain (Storey, 1976). In South Africa, ‘Hortus Gold’ is a popular variety grown for fruits borne by female plants of a dioecious strain (Storey, 1976). Dioecious cultivars ‘Bettina’ and ‘Petersen’ are cultivated in Australia. The Cuban variety ‘Maradol’ is cultivated extensively in Mexico (Noa-Carrazana et al., 2006). In India popular varieties include ‘Coorg Honey Dew’, ‘Pusa Dwarf ’, ‘Pusa Delicious’, and ‘Taiwan’ (Chadha, 2002). In Taiwan, the cultivar ‘Tainung No. 2’, derived from hybridization of the cultivars ‘Thailand’ and ‘Sunrise’, is very popular (Kung et al., 2009). Viruses: a key problem associated with papaya cultivation Common targets for breeding improved cultivars of papaya are virus resistance, hermaphroditic plants with a low frequency of sex change, dwarf varieties/tree size, delayed ripening and enhanced latex production (Storey, 1976; Ogata et al., 2016). Viruses pose serious threats to papaya cultivation worldwide. Table 7.1 provides a list of major papaya viruses for which genome sequence information is available. Generally, the type of virus and associated vector complex varies geographically and is influenced by varieties planted, cultivation practices and local climate. However, some papaya viruses that cause economic loss are global in nature and hence, important from perspectives of trade, industry and agriculture. Virus resistance continues to be a major goal for sustainable papaya cultivation. In addition to prophylaxis and disease management practices, mitigating economic loss incurred from viral diseases in cultivated papaya has involved

planting tolerant cultivars, cross-protection, and breeding for durable virus resistance by transgenic approaches. Transgenic papayas can provide reliable and broad-spectrum resistance to viruses specific to different agro-ecosystems. Development and adoption of transgenic virus-resistant papaya in Hawaii was a major breakthrough in the application of biotechnology to agriculture and will be discussed further. Global and emerging viruses of papaya Papaya ringspot virus Papaya ringspot virus (PRSV) is an important potyvirus infecting papaya in almost all regions where it is cultivated (Yeh and Gonsalves, 1984; Gonsalves, 1998). PRSV shares a long history with papaya production. It is the most widespread and destructive virus infecting papaya and can cause up to 100% yield loss. Subsequently it is perhaps the best-characterized virus infecting papaya. The severity and extent of viral infections in papaya tend to vary influencing fruit yield, tree vigour, or fruit quality (Tennant et al., 2007). PRSV infections of papaya cause reduction in acreage under production, employment crises, loss of foreign exchange and decrease in fruits for local market dispersal (Tennant et al., 2007). Various sap-sucking hemipteran aphids like Myzus persicae (Sulzer) and Aphis gossypii Glover transmit PRSV in a non-circulative, non-persistent manner (Purcifull et al., 1984; Kalleshwaraswamy and Kumar, 2008). PRSV is also sap-transmissible. Seed-transmission is rare (Gonsalves, 1998) but has been reported (Bayot et al., 1990). There are two biotypes based on host range: PRSV-p which naturally infects cucurbits and papaya, and PRSV-w which infects cucurbits but not papaya (Gonsalves, 1998). PRSV-p (henceforth referred to as PRSV) has been identified from Africa, tropical Asia, the Caribbean, Central and South America and the South Pacific (Fermin et al., 2015). Reports of potyviruses causing disease in papaya date back to 1929 in Jamaica, yet PRSV was first recognized as a major threat to papaya production in Hawaii in 1945 (Fermin et al., 2010). Reports of PRSV infecting papaya continue to appear from different parts of the world. International trade in fruit and seed,

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Table 7.1 A list of first reported complete/partial genome sequences of viruses from papaya Virus species; mode of transmission

Typical symptoms

Type of virus/ genome features Points of interest

References [RefSeq number]1

Family Potyviridae, Potyvirus Papaya ringspot virus, PRSV-p; Aphid, sap (and seed) transmitted

Yellowing, distortion, severe leaf mosaic, ‘ringspots’ on fruits, reduced fruit set, reduced fruit quality, poor growth

Papaya leaf distortion mosaic virus, PLDMV-p; Aphid transmitted

Resemble those caused Single-stranded by PRSV, distortion, leaf- positive, RNA, narrowing, stunting 10.1 kb

Single-stranded positive, RNA, 10.33 kb

First instance of commercial Yeh et al. (1992); transgenic papaya with Gonsalves et al. resistance to PRSV-Hawaii, (2010) [NC_001785] 1998; commercialized in China, 2006 Threat to PRSV-resistant transgenic papaya in China and Taiwan

Maoka and Hataya (2005); Kung et al. (2009); Tuo et al. (2013) [NC_005028]

Single-stranded positive, RNA, 6.67 kb

Low economic significance, mixed infection with PRSV in Mexico

Sit et al. (1989); Noa-Carrazana and Silva-Rosales (2001) [NC_001748]

Family Alphaflexiviridae, Potexvirus Papaya mosaic virus, PapMV; Aphid transmitted

Resemble those caused by PRSV, mosaics, stunting, distortion, spots on fruits

Family Geminiviridae, Begomovirus Papaya leaf curl virus, PaLCuV; Whitefly transmitted

Curled leaves, twisted petioles, stunting, vein enations

Single-stranded DNA mono/ bipartite, 2.75 kb

Broad host range, probably associated with satellites

Saxena et al. (1998) [NC_004147]

Papaya leaf curl Guandong virus, PaLCuGV; Whitefly transmitted

Downwards leaf curling, vein thickening, leaf chlorosis

Singlestranded DNA, monopartite, 2.74 kb

Not associated with alphaor betasatellite

Wang et al. (2004) [NC_005844]

Papaya leaf curl China virus, PaLCuCNV; Whitefly transmitted

Downwards leaf curling, vein thickening, leaf chlorosis

Singlestranded DNA, monopartite, 2.75 kb

Not associated with alphaor betasatellite

Wang et al. (2004) [NC_005321]

Papaya leaf crumple virus, PaLCrV; Whitefly transmitted

Severe curling of leaf Singlemargins, petioles twisted, stranded DNA, stunted, enations monopartite, 2.74 kb

Associated with Tomato leaf curl betasatellite, and Croton yellow vein mosaic betasatellite

Singh-Pant et al. (2012) [NC_014707]

Papaya leaf curl Faisalabad virus, PaLCuFV

Currently not available

Singlestranded DNA, monopartite, 2.75 kb

Currently not available

Unpublished [NC_033276]

Papaya lethal yellowing virus, PLYV, Saptransmitted, soil-borne, vector unidentified

Yellowing in young then mature leaves, wilting, greenish-brown circular spots on fruits

Single-stranded positive, RNA, 4.145 kb

Mixed infection with PRSV in natural and cross-protected papayas, currently classified as Sobemovirus

Pereira et al. (2012); Nascimento et al. (2010) [NC_018449]

Papaya meleira virus, PMeV, transmitted by infected latex, vector/ seed unproven

Spontaneous latex exudation, leaf tip burns, malformed fruits

Double-stranded Lowered proteolysis in latex Maciel-Zambolim RNA, 8.7 kb et al. (2003); Abreu et al. (2015b) [NC_028378]

Papaya virus Q, PpVQ, Not seed/aphid/graft transmitted

Currently not clear, disparate symptoms in mixed infections

RdRp gene

Unclassified

Mixed infections with PRSV (Ecuador) and PMeV (Brazil), assigned to Umbravirus

Quito-Avila et al. (2015); [KP165407]

*RefSeq refers to the reference genome sequence for each virus type available at www.ncbi.nlm.nih.gov.

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population expansions and dispersion of vectors may have contributed to the prevalence and spread of PRSV (Bateson et al., 2002). Papaya leaf distortion mosaic virus and Papaya mosaic virus Like PRSV, Papaya mosaic virus (PapMV) and Papaya leaf distortion mosaic virus (PLDMV) have a long history with papaya production and data on some genotypes are available but they are not as widespread and can be considered of local significance. As indicated in Table 7.1, papaya plants infected with PRSV, PLDMV, and PapMV exhibit similar symptoms viz. mosaic, distortion of leaves, yellow-green leaf discolorations, ring-spots on fruits, and water-soaked streaks on foliage (Tuo et al., 2013). Reliable methods based on EnzymeLinked Immunosorbent Assays (ELISAs) and Reverse-Transcriptase Polymerase Chain Reaction (RT PCR) have been developed that can routinely distinguish infections by PRSV and PLDMV potyviruses (Maoka and Hatoya, 2005; Bau et al., 2008; Shen et al., 2014). Mixed infections of PRSV with PLDMV in Taiwan, China and PRSV with PapMV in the Philippines have been reported (Bau et al., 2008; Cruz et al., 2009; Tuo et al., 2013). Another emerging virus in Brazil, and several countries of Central and South America is the potexvirus PapMV (Noa-Carrazana et al., 2006). Originally detected in the USA in 1962, PapMV occurs as single infection or as mixed infections with PRSV (Noa-Carrazana et al., 2006; Chavez-Calvillo et al., 2016). Interestingly the order in which either virus infects a susceptible host influences the severity of disease symptoms. Infection by PapMV before PRSV triggers antagonistic interactions, synthesis of pathogenesis-related (PR) plant defence proteins and alleviation of disease symptoms (ChavezCalvillo et al., 2016). Papaya meleira virus, Papaya lethal yellowing virus and leaf curl viruses of papaya In addition to the above-mentioned viruses of papaya, some recently described viruses below are attracting increasing attention and hold economic potential. Papaya sticky disease (or ‘meleira’ in Portuguese) was reported in 1980 from Brazil and recently from Mexico, where leaves and fruits of infected plants exude latex (Abreu et al., 2015).

Deep sequencing of latex-associated viruses indicated infection by Papaya meleira virus (PMeV) and Papaya virus Q (Sá Antunes et al., 2016). Another papaya virus reported from north-eastern Brazil is Papaya lethal yellowing virus (PLYV) belonging to genus Sobemovirus (Pereira et al., 2012). Begomoviruses belonging to Geminiviridae are another major widespread group of viruses in Asia (Varma and Malathi, 2003; Mishra et al., 2016). Disease in papaya caused by Papaya leaf curl virus disease was reported for the first time in 1939 from India (Nariani, 1956). In India, PRSV can cause 50–60% yield loss, while leaf curl disease is estimated to cause 70–80% yield loss (Chadha, 2002). Other begomoviruses (mostly monopartite with/without satellite molecules) that infect papaya include Papaya leaf curl virus (Saxena et al., 1998); Papaya leaf crumple virus or PaLCrV (Singh-Pant et al., 2012); Papaya leaf curl Guandong virus or PaLCuGuV and Papaya leaf curl China virus or PaLCuCNV (Wang et al., 2004). Infection of papaya by other begomovirus including Tomato leaf curl New Delhi virus (ToLCNDV) has also been reported (Maruthi et al., 2007; Singh-Pant et al., 2012). ToLCNDV is a new emerging whitefly transmitted begomovirus with global distribution (Moriones et al., 2017). Papaya plants co-infected with PRSV and begomoviruses were observed in the same field in North India (Fig. 7.1; Singh-Pant et al., 2012). Natural infection of papaya by three viruses including PRSV and the pandemic-causing begomovirus Tomato yellow leaf curl virus-IL (TYLCV-IL) was reported recently in Texas, USA (Alabi et al., 2017). These results suggest that papaya can serve as an alternate host and/or reservoir of disparate viruses. Hence, effective detection/diagnosis methods and control strategies that can reduce yield loss from one and/ or both viruses are essential. Genome and genetic diversity of papaya viruses PRSV is a positive, single-stranded RNA potyvirus encapsidated by a virus-encoded coat protein (CP). The viral RNA is poly-adenylated at the 3′ end while at the 5′ end it is covalently linked with the protein VPg (viral protein-genome linked). The genome is 10.324 kb long and translates into a single large polyprotein (3344 amino acids) which is subsequently cleaved into smaller proteins of varied functions. The arrangement of genes resembles

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Figure 7.1  A commercial field in Haryana, India with (A) healthy papaya plant and plants showing symptoms of (B) leaf curl disease, (C) mixed infection by begomovirus and PRSV, (D) PRSV. Typical symptoms of PRSV observed on (E) a fruit is shown.

other potyviruses. The cleaved proteins are P1 (63 kDa), helper component proteinase or HC-Pro (52 kDa), P3 (46 kDa), cylindrical inclusion protein or C1 (72 kDa), nuclear inclusion protein a, or NIa, (48 kDa), nuclear inclusion protein b or NIb (59 kDa), 35-kDa coat protein or CP (Yeh et al., 1992). An embedded ORF encoding a 25-kDa protein putative movement protein called PIPO (Pretty Interesting Potyviridae ORF) is also present (Chung et al., 2008). HC-Pro participates in aphid transmission, symptomatology, genome amplification, and suppression of viral gene silencing. PRSV is a highly variable virus species. The genetic variability of this virus is related to the geographical origin of the isolates (Bateson et al., 2002). Greatest diversity in the nucleotide sequences of CP and HC-Pro has been observed in isolates from India supporting an Asian origin of PRSV and India being the centre of origin (Olarte Castillo et al., 2011). In India, estimates of sequence variability of the cp gene of PRSV range from 11 to 18.5% ( Jain et al., 2004; Srinivasulu and Sai Gopal, 2011). The genome of the potyvirus, PLDMV is 10.153 kb in length. It encodes a polyprotein of 373.68 kDa is catalytically cleaved into the coat protein, P1, HC-Pro, P2, 6K1, C1, 6K2, Nla-VPg, Nla-Pro, and NIb (Maoka and Hataya, 2005; Shen et al., 2014). Another putative protein PIPO is encoded by an ORF located within the P2 gene. Different isolates of PLDMV that infect papaya share about 95–97% sequence similarity, while the biotype (PLDMV-p) that infects papaya differs from the biotype infecting cucurbits (PLDMV-c) at the 5′ region (Maoka and Hataya, 2005). PapMV is a positive, singlestranded RNA virus with a genome size of 6.66 kb. A cap of m7GpppN occurs at the 5′ end followed

by un-translated region. PLYV is also a positive, single-stranded RNA virus with a genome size of 4145 bases (Table 7.1). Four open reading frames (ORF1, ORF2a, ORF2b and ORF3) are present. The ORF1 encodes a protein of unknown function while ORF2a and ORF2b encode a poly-protein including a serine protease and a VPg. A frameshift mutation on the same strand as ORF2b produces the RdRp. ORF3 encodes the PLYV coat protein. Analysis of sequences encoding RNA-dependent RNA polymerase (RdRp) and coat protein from 21 isolates of PapMV isolates encoding from Brazil revealed low genetic variability and resembled homologues in Sobemovirus (Pereira et al., 2012). Leaf curl disease causing begomoviruses that infect papaya are typically monopartite with satellite molecules (Wang et al., 2004; Singh-Pant et al., 2012). These viruses have a circular single-stranded DNA A genome of approximately 2.4 kb associated with alphasatellite and/or betasatellite (Zhou, 2013). The DNA A genome codes for AV1 (coat protein or V1), AV2 (pre-coat protein or V2) on the sense strand, while the complementary strand contains genes for AC1 (replication associated protein or Rep), AC2 (transcriptional activator or TrAP), AC3 (replication enhancer or REn) and AC4 (pathogenicity determinant or C4) proteins that are essential for viral function (Patil and Fauquet, 2009). The DNA B genome (when present) encodes viral movement proteins BV1 (nuclear shuttle protein, NSP) and BC1 (movement protein, MP) (Rojas et al., 2005). Function of the DNA B genome is supplanted in monopartite begomoviruses by the satellite molecules. The β satellite and encoded βC1 protein is implicated in amelioration of disease symptoms, and host range (Zhou, 2013).

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Sequence diversity of begomovirus species (and satellite molecules) infecting papayas in India is high (Singh-Pant et al., 2012), supporting the contention of Indo-China being the centre of origin for begomoviruses (Nawaz-ul-Rehman and Fauquet, 2009). The 8.7 kb double-stranded RNA genome of Papaya meleira virus or PMeV contains two ORFs (Abreu et al., 2015). The first ORF has no known function and resembles uncharacterized homologues from mycoviruses. The ORF2 encodes a RdRp that contains domains resembling counterparts from different viral genera like Luteovirus, Rotavirus and Totivirus (Abreu et al., 2015). Papaya sticky disease in Mexico appears to be caused by two RNA species PMeV as well as an umbraviruslike ssRNA entity resembling Papaya virus Q or PpVQ (Sá Antunes et al., 2016). Partial genome sequence of PpVQ reported from Ecuador indicates presence of two ORFs of unassigned function (Quito-Avila et al., 2015). PpQV which lacks a coat protein may be associated with PMeV that plays the role of a helper virus role in causing papaya sticky disease (Antunes et al., 2016). Intriguingly, PMeV was detected in both symptomatic and asymptomatic papayas in Brazil (Sá Antunes et al., 2016), suggesting that it may be a persistent virus that affects disease manifestations in papaya especially during mixed infections. Control and management of viral diseases in papaya Traditional approaches for viral disease control The best documented strategies for control of papaya viruses target PRSV (Fermin et al., 2010). Disease management typically includes quarantine, eradication and avoidance by planting papaya in areas unaffected by the virus. However, these control measures can be expensive, laborious and not fool-proof (Gonsalves, 1998; Tripathi et al., 2008). Several countries in Latin America have successfully managed viral diseases in papaya using avoidance, roguing (even cucurbit elimination), weed eradication, land use change, crop rotation, intercropping, barrier plants, careful seed selection, and judicious planting schedules (reviewed in Fermin et al., 2010). Management of insect vectors

using insect traps, plastic mulch and aphicides to deter probing behaviour has also been demonstrated worldwide. In India where virus infestations of papaya are high, insecticide applications to control vector populations have been the primary method of disease control (Chadha, 2002). To date, not much has been published regarding control strategies for papaya sticky disease. However, it has been reported that roguing is essential. As disease symptoms appear after fruit setting, an apparently healthy -ooking plant may be actually be infected by PMeV (Abreu et al., 2015). Breeding for resistance and tolerant plants There is no known source of natural resistance to PRSV in C. papaya germplasm (Mekako and Nakasone, 1975; Conover and Litz, 1978). Resistance genes do exist naturally in distantly related Vasconcellea species (Tripathi et al., 2008). However, sexual incompatibility of wild species and cultivated papaya poses a challenge for development of PRSV-resistant varieties through traditional breeding methods (Manshardt, 1992; Gonsalves et al., 2006). Hence embryo rescue and wide hybridization methods including use of bridge species such as Vasconcellea parviflora have been attempted (Drew et al., 2005). A more phylogenetically distant plant, Cucumis metuliferus (Cucurbitaceae), also seems to harbour genes involved in resistance to PRSV and may potentially be useful in biotechnological projects aimed at engineering PRSV-resistant papayas (Lin et al., 2013). Development of tolerant varieties of papaya by introgression has also been attempted, but poor fruit quality and requirement of cumbersome back-crossing with commercial varieties has generally deterred development of this strategy for virus resistance (Fermin et al., 2010). In the case of PLDMV, natural resistance occurs in V. cundinamarcensis and intergeneric hybrids with C. papaya have shown promising results for virus resistance breeding (Tarora et al., 2016). Cross-protection against viral diseases Cross-protection for containment of PRSV has been mostly successful in Hawaii, Taiwan, Brazil and Venezuela (Yeh and Gonsalves, 1984; Fermin et al., 2010). In this field-based approach, inoculation of a plant with the attenuated strain of a particular

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virus confers protection against subsequent attack by a related strain of the same virus. Strains used for cross-protection are usually derived from singlelesion, field isolates that have been rendered mild by mutagenesis. Examples of mild strains developed for cross-protection in Hawaii are PRSV HA5–1 and PRSV HA6–1 (Yeh and Gonsalves, 1984). Current understanding of the molecular basis for cross-protection involves reduced suppression of gene silencing by attenuated strains. However, problems associated with availability, production and maintenance of the mild strains; logistics of field-level inoculations and absence of legislation have proven to be some major limitations of this approach (Tripathi et al., 2008, Fermin et al., 2010). Furthermore, cross-protection has proven to be only partially successful in containing PRSV in field studies (Azad et al., 2014). Thus, improved monitoring for virus outbreaks, developing diagnostic tools for early detection and characterization of the disease-causing agent(s) is urgently needed. Generally, developing resistance to viral diseases via traditional breeding methods has had limited success. Thus, other strategies must be adopted to combat yield losses in papaya crops due to viral diseases. Transgenic papayas for sustainable virus-resistance Improvements in agricultural biotechnology in the late 1980s, especially recombinant DNA technology and plant transformation protocols made it conceivable to introduce desired genes into papaya to combat viral diseases for crop improvement (Grumet, 1990). In 1998, two transgenic papaya varieties, ‘Rainbow’ and ‘SunUp’, became the first virus-resistant fruit crop to be commercialized in the USA (Gonsalves, 1998) and grown in Hawaii. The following section provides a brief account of PRSV-resistant transgenic papayas in Hawaii in the context of development, mechanism of resistance and ongoing efforts for durable resistance to papaya viruses. Pathogen-derived resistance: viral coat protein gene-based transgenic papaya Development of virus-resistant transgenic plants has typically been based on the concept of pathogen derived resistance (PDR), where any region of

the viral genome (translatable into protein and/or untranslatable) can be used to engineer resistance (Sanford and Johnston, 1985). PDR would control the invading virus by expression of virus-derived genes or fragments as transgenes in susceptible crop plants conferring resistance to same or closely related pathogens. Successful development of disease resistance phenotypes in transgenic tobacco, expressing the coat protein gene (cp) of Tobacco mosaic virus (TMV) provided early proof of the concept (Abel et al., 1986). Even though the exact mechanism was then unclear, cp gene mediated transgenic resistance was also demonstrated in the early 1990s against Cucumber mosaic virus, Potato virus Y, Potato leaf roll virus and Tobacco etch virus (Lindbo and Falk, 2017 and references therein). The earliest and most enduring practical applications of PDR against viruses came from using cp genes for the development of transgenic summer squash (Cucurbita pepo) against multiple viruses (Courgette yellow mosaic virus, Watermelon mosaic virus WMV-2 and Cucumber mosaic virus) and transgenic papaya against PRSV isolates from Hawaii (Tricoli et al., 1995; Lius et al., 1997; Gonsalves et al., 2004). These commercialized, transgenic lines currently account for significant areas under cultivation in the United States and have provided economic benefit to farmers over two decades (Lindbo and Falk, 2017). The role of RNA in mediating PDR by cellular degradation of RNA duplexes formed between transcripts from a transgene and the viral pathogen was established subsequently (Lindbo and Dougherty, 1992, 2005). The extent of sequence homology between a transgene and the corresponding region of viral genome prompts post-transcriptional gene silencing (PTGS). PTGS is a defence response to non-self RNA in plants and animals, where it is called RNA interference or RNAi (Lindbo and Dougherty, 2005). PTGS-mediated transgenic resistance depends on accumulation of 21–25 nt small-interfering RNAs (siRNAs) from degradation of target RNAs by a plant’s RNA surveillance system. This process comprises of RNA-induced silencing complex (RISC) with Dicer-like proteins (DCL) and specificity conferring Argonaute (AGO) guide proteins (Hamilton and Baulcombe, 1999; Brodersen and Voinnet, 2006). Though not elucidated until the early 1990s, PTGS is now known to be the predominant causal mechanism behind successful PDR-deploying

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transgenics against plant viruses (Lindbo and Falk, 2017). Transgenic virus-resistant papaya in Hawaii demonstrates an important practical application of the PDR concept with enduring impacts on agriculture as described below. Hawaiian success story Several reviews and articles describe the development of PRSV-resistant transgenic papaya for Hawaii, USA (Ferreira et al., 2002; Gonsalves et al., 2004; Fuchs and Gonsalves, 2007; Gonsalves, 2015). It is the first instance of a transgenic virusresistant edible fruit available commercially in the world that continues to show high level of resistance to Hawaiian strains of PRSV. It was developed in the late 1980s by a research team at Cornell University and the University of Hawaii comprising Dennis Gonsalves, Maureen Fitch, Richard Manshardt and Jerry Slightom (Gonsalves, 1998). The papaya cultivar Sunset was transformed with the cp gene of a mild, cross-protection strain PRSV HA 5–1. The transgene was introduced by particle gun bombardment of immature zygotic embryos into the host as a chimeric protein, with a leader sequence of the CMV cp gene fused in-frame at the N terminus of the full-length cp gene of PRSV HA 5–1 (Fitch et al., 1992; Fitch et al., 1993). Transgenic R1 plants of line 55-1 were highly resistant to Hawaiian PRSV strains but showed differential susceptibility to different geographical isolates of the virus, attesting to the importance of sequence homology between

the transgene and the targeted virus (Tennant et al., 2001). Experimental field trials showed similar results and provided convincing evidence that line 55-1 could control PRSV in Hawaii. Rainbow, a F1 hybrid of the yellow-fleshed Kapoho, the dominant cultivar in Puna and a homozygous selection of line 55-1 (SunUp) were produced. The transgenic plants showed full resistance to PRSV and grew prolifically in field trials conducted at Oahu, Hawaii (Lius et al., 1997). In1992, PRSV infection of papayas growing in the Puna district of Hawaii had assumed epidemic proportions. This caused significant reduction in yield (which decreased by almost 50%) in this major production centre of papaya. Attempts to contain the viral disease by management strategies using tolerant plants and cross-protection were in vain. Hence, the transgenic papayas were tested in field trials in Puna in 1995 and found to be successful in resisting PRSV infection (Fig. 7.2). Some field trials produced 125,000 pounds per acre per year, in contrast to non-transgenic fields that yielded only 5000 pounds per acre per year (Gonsalves et al., 2004). After successful field testing over several years, regulatory agencies like USDA-APHIS, FDA and EPA approved and deregulated transgenic papaya for its commercial utilization by 1997. Presently, transgenic papaya is widely sold in Hawaii and exported to markets in the US, Canada and Japan. It is estimated that 75% of the papaya grown in Hawaii is transgenic while the export market

Figure 7.2 (A) Transgenic papaya field trial in Puna, Hawaii from November 1996 showing non-transgenic plants (left) and transgenic plants (right). (B) An aerial view of field trial from May 1997 showing a solid block of PRSV-resistant transgenic papayas in the centre surrounded by non-transgenic papayas, and replicated blocks of transgenic, non-transgenic, tolerant and cross-protected papaya plants to the right. (Images used with permission of Professor Dennis Gonsalves.)

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went from a worth of $14 million dollars to $23 million dollars over a span of 3 years due to durability and reliability of the virus-resistance conferred by transgenic papaya (VIB 2014). Interestingly, continuous planting of transgenic virus-resistant papayas has lowered incidence of the PRSV itself, and so non-transgenic papayas can now be cultivated albeit with buffer zones of transgenic papayas (Fitch, 2016). However, the majority of papaya consumed globally is non-transgenic, susceptible to local viruses including PRSV and produced by traditional methods for disease management (Fermin et al., 2010). PRSV-resistant papayas (based on cp, NIb, and HC-pro transgenes) Table 7.2 highlights attributes of transgenic papaya developed by various research groups around the world. Most groups worked with regional papaya varieties and sequences of local PRSV strains for PDR-based transgenic virus resistance. A large number of constructs used for successful papaya transformation contained leader sequences from β-glucuronidase (uidA) gene or Cucumber mosaic virus along with Cauliflower mosaic virus 35S promoter (reviewed by Fermin et al., 2010). Untranslatable versions of transgenes were achieved by either adding a stop codon or causing a frame shift mutation in the cp gene sequence or using the 3’untranslated regions of viral genes. The use of untranslatable cp gene fragments had several advantages, especially minimization of toxicity and/or allergenicity likely associated with expression of a heterologous protein. PRSV resistance phenotypes obtained with these transgenes was based on PTGS (Tennant et al., 2001). The efficacy of viral resistance attained by cp gene-based PRSV resistance depends on (1) sequence homology between transgene and the viral genome, (2) evolution of virus strains with strong PTGS suppressing abilities, (3) developmental stage of the plant, (4) zygosity/genotype of the host plant and (5) environmental conditions (Pang et al., 1996; Tennant et al., 2001; Bau et al., 2003; Kung et al., 2015). Currently, these transgenic plants are at different stages of development, green house and field trials as well as processes of regulatory oversight. Except for transgenic PRSVresistant papaya that have been commercialized in China (Ye and Li, 2010), all the virus-resistant

lines await government approval for commercial cultivation. The following example demonstrates the adaptability of the transgenic approach to provide solutions to changing virus scenarios in the field. Transgenic papayas had been produced in Taiwan using untranslatable fragments from cp gene of a highly virulent strain, PRSV-YK. It was resistant to geographically diverse PRSV strains from Taiwan, Thailand, Hawaii, and Mexico (Bau et al., 2008). These plants showed resistance in field trails until emergence of a new pathotype of PLDMV in Taiwan (Bau et al., 2008) and Hainan province in southern China (Tuo et al., 2013). The PLDMV strain P-TW-WF from Taiwan is different in sequence and host range (infecting only papaya) from the Japanese strains which also infect cucurbits (Maoka and Hataya, 2005). The infection of transgenic PRSV-resistant papaya Tainung No. 2 in Taiwan by the relatively rare PLDMV strain was cause for much concern (Bau et al., 2008). In 2009, transgenic papaya derived from the cultivar Thailand containing truncated coat protein gene fragments from PLDMV-P-TW-WF and PRSV-YK isolates were obtained (Kung et al., 2009). Interestingly, the transgenic lines were resistant to both potyviruses in green house trials and referred to as ‘double resistant’. Resistance by PTGS and occurrence of siRNAs specific to the chimeric constructs were observed in the transgenic lines (Kung et al., 2009). Three transgenic lines (HR10-4, 14-1 and 14-3) were also resistant to PRSV strains from Hawaii, Thailand and Mexico (Kung et al., 2009). In the evolutionary arms race between viruses and host plants, emergence of new virulent strains is inevitable as exemplified by strain(s) PRSV-5-19 that could infect these chimeric double virus-resistant papayas (Kung et al., 2009). The Taiwanese PRSV strain 5-19 could break down resistance in transgenic papayas due to the presence of a suppressor encoded by HC-Pro in a sequence-independent manner (Yeh et al., 2010). Hence the gene encoding HC-Pro, best known as potyviral suppressor of gene silencing (Anandalakshmi et al., 1998) was used as untranslatable transgenes of various sizes to transform somatic embryos of variety Sunrise (Yeh et al., 2010). These plants showed accumulation of siRNA specific for the transgene, suggesting potential application as sources of resistance to new strains of PRSV.

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Table 7.2. Transformation events for virus resistance in papaya Method of Donor virus/isolate transformation Type

Research stages

References

Local variety

PRSV Bridgeman Downs

Biolistics

T cp gene

Field Testing

Lines et al. (2002)

Brazil

Sunrise solo, Sunset solo

PRSV Bahia

Biolistics

T and UT cp gene

Field Testing, Agronomic evaluation

Souza et al. (2005)

China

Huanong No. 1 PRSV

Agrobacterium UT rep gene

Commercial

Fitch (2010)

China

Sunset

PRSV, PLDMV

Agrobacterium T multi cp gene

Southern positive plants

Fitch (2010)

China

Sunrise, ZhongBai, Suizhonghong 48

PRSV

India

Pusa Delicious PRSV, PLCV

Country

Cultivar

Australia

India

Jia et al. (2017)

Screening Agrobacterium T cp gene, rep gene

Chandra et al. (2008)

PRSV

Biolistics

Chandrashekara et al. (2006)

T cp gene

Transient expression

Indonesia

Bangkok, Burung

PRSV Bogor; cp, Antisense ACC oxidase

Biolistics

Jamaica

Sunrise solo

PRSV Caymanas

Biolistics

T and UT cp gene

Field tested

Tennant et al. (2005)

Mexico

PRSV

Biolistics

T cp gene

Greenhouse

Fitch (2010)

Philippines Davao Solo

PRSV cp

Agrobacterium T cp gene

Field resistance

Lawas and Magdalita (2007)

Taiwan

Tainung no. 2

PRSV YK, PLDMV

Biolistics

T cp gene

Field resistance

Bau et al. (2003)

Thailand

KhakDum

PRSV Ratchaburi province

Biolistics

T cp gene

Field resistance

Kertbundit et al. (2007)

Thailand

Khaeknual

PRSV Chiang Mai

Biolistics

T cp gene

Field resistance

Phironrit et al. (2005)

US: Florida CV.F65

PRSV H1K

Agrobacterium T and UT cp gene

Passed FDA, Petition for deregulation

Davis and Ying (2004)

US: Hawaii Sunset solo

PRSV HA

Biolistics

Commercial

Gonsalves (1998); Tennant et al. (2005); Ferreira et al. (2002)

US: Virgin Islands

Washington, Yuen Nong

PRSV

Agrobacterium T cp gene

Field resistance

Zimmerman et al. (2005)

Venezuela

Tailandaroja

PRSV EV and LA strains

Agrobacterium T cp gene

Greenhouse

Fermin et al. (2004)

Damayanti et al. (2001); Fitch, (2010)

T cp gene

cp, coat protein gene; rep, replicase gene; T, translatable; UT; unstranslatable.

In China, transgenic virus-resistant papaya cultivar Huanong No. 1 containing the NIb gene from PRSV strain YS (Yellow Spot) was deregulated in

2006 (Ye and Li, 2010). Transgenic papaya developed in Guangdong province of China carrying the NIb gene fragment was resistant to PRSV isolates

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from Guangdong (Rao et al., 2012). However these transgenic plants were susceptible to PRSV strains from Hainan, highlighting high sequence variation in virus strains within China. In fact, PRSV isolates from Hainan belong to three clusters based upon multi-gene phylogeny and geographical distribution (Zhao et al., 2016). In 2015, an estimated 7000 hectares of transgenic virus-resistant papaya was cultivated in southern China also indicating high acceptance by papaya farmers ( James, 2015). In Hainan province and other regions of China where commercial PRSV-resistant papaya is cultivated (Guo et al., 2009), appearance of new strains of PRSV and susceptibility to PLDMV-DF (Tuo et al., 2013) are of serious concern. RNAi construct-based transgenic strategies for virus-resistance in papaya RNAi constructs that generate dsRNA capable of triggering gene silencing have been used successfully for virus resistance (Waterhouse et al., 1998; Patil et al., 2011; Kreuze and Valkonen, 2017). In this approach, short regions of viral genes (usually untranslatable) are present as inverted repeats or IR within each constructs, producing transcripts that form RNA hairpin loops and other secondary structures that are degraded via PTGS or RNAi pathways. An advantage of the approach is that short genomic fragments of diverse viruses can be used for silencing multiple and co-infectious viruses (Bucher et al., 2006). RNAi with cp gene constructs Recently, an RNAi approach was used to obtain resistance to virulent, sequence divergent strains of PRSV from Hainan, China which showed about 97–100% sequence variability at the genome level ( Jia et al., 2017). In this study, resistance to multiple strains from Hainan were observed in greenhouse experiments using RNAi hairpin constructs based on a conserved 544bp region of the cp gene separated by a short intron of 201bp. The transgene was delivered by particle gun bombardment to obtain a single insertion transgenic line-454. As expected siRNAs less than 50bp were detected in the transgenic papayas, along with suppression of disease symptoms in the green house experiments. These results suggest that this RNAi-based approach may be of practical application for PRSV resistance

in China. It may be noted that after deregulation in 2006, PRSV resistant transgenic papayas were already being grown in China ( James, 2015; Fermin et al., 2010). In India, recent success has been demonstrated in control of PRSV by foliar application of dsRNA synthesized in vitro (Voloudakis AE and Patil BL, personal communication). RNAi with rep gene constructs against begomoviruses The RNAi based approach using a rep gene (AC1) fragment from DNA-A genome of Bean golden mosaic virus (the eponymous begomovirus) has been used for generating resistance to golden mosaic disease in Phaseolus beans (cultivar Carioca) in Brazil (Bonfim et al., 2007). Transgenic virus-resistant line EMB-PV051–1; Embrapa 5.1 was commercialized as cultivar BRS FC401 RMD in 2011, an important step for future control strategies for single-stranded DNA begomoviruses that attack papayas. An important advantage of RNAiconstruct based transgenic strategies for crops is that the percentage of plant transformants that possess the desired virus resistance phenotype is high. The transgenic BGMV-resistant has shown successful field resistance in Brazil since 2007–2013 in conjunction with integrated pest management of the whitefly vector (Aragão, 2014). The transgenic produce is not yet available for export. amiRNA-mediated and CRISPR-Casbased virus resistance A recent approach for transgenic virus resistance involves microRNA (miRNA) and siRNAs, which together account for 21–24 nt RNA species in plants (Tomari and Zamore, 2005). These short RNAs differ in their biogenesis from siRNA that participate in PTGS. The former arise from pre-miRNA originating from single-stranded, endogenous plant transcripts that fold at selfcomplementary regions into loops and other secondary structures recognized by DCL. In case of siRNA, disparate molecules anneal to form dsRNA which are then cleaved by DCL from RISC, initiating PTGS. It has become possible to design artificial miRNA (amiRNA) with as little as 21 nt from a viral genome inserted within a plant miRNA gene to generate pathogen-specific small RNA and resistance phenotype (Niu et al., 2006). The success of amiRNA has been demonstrated against various

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viruses including begomoviruses (Ali et al., 2013; Wagaba et al., 2016). However no amiRNA-based virus-resistant transgenic plant has been commercialized yet. Another important process in the horizon of engineered disease resistance in plants is genome editing by CRISPR/Cas9 system (Bortesi and Fischer, 2015; Fondong et al., 2016; Noman et al., 2016) such that a small region of existing plant miRNA can be modified to target a virus gene for silencing. Transgenes based on Cas9/single guide RNA (sgRNA) can target expression of host translational factors (e.g. eIF4) that interact with potyviral VPg (Pyott, 2016). Recently, virus resistance has been displayed for PRSV-w, along with RNA viruses infecting cucumber by silencing the eIF4E gene in tomato and melon (Chandrasekaran et al., 2016). With rapid advances in this approach that can tap into natural resistance mechanisms of plants against viruses, results are awaited for next generation of transgenic virus-resistant papaya. Conclusion: challenges in adoption of transgenic, virusresistant papaya Development of transgenic papayas based on PDR and PTGS/RNAi strategies is a robust solution for debilitating viruses that cause economic loss. This approach has been proven with PRSV-resistance in Hawaii, USA. Large scale research, safety trials and publications addressing various concerns like gene flow, recombination of viral strains, hetero-encapsidation and allergenicity have been already been addressed with respect to the transgenic PRSVresistant papayas available commercially (Yeh and Gonsalves, 1994; Roberts et al., 2008; Fermin et al., 2011). Furthermore, a draft genome is available for ‘SunUp’ papaya (Ming et al., 2008), making it a very well-characterized system for understanding virus resistance in transgenic plants. Resistance in SunUp papaya to PRSV-HA has been associated with transcriptomic responses characteristic of plant defences against pathogens and no evidence of genes producing any known toxins or allergenic proteins (Fang et al., 2016). Transgenic PRSVresistant papaya remains one of the best examples of durable transgenic resistance in a horticultural crop. Unlike insect-resistant transgenic Bt crops grown around the world (Wan et al., 2012; Fabrick

et al., 2014), no breakdown in the PTGS-based transgenic virus-resistant plants has been observed for the cognate strain. In fact, the incidence of PRSV in Hawaii has been lowered (Fitch, 2016). Taken together, these results suggest that the transgenic PRSV-resistant Hawaiian papaya may be used as a template for developing resistance to other PRSV strains and viruses of regional importance. Even though use of transgenic papaya greatly improved papaya production in Hawaii, they have not been readily accepted in other parts of the world. Except for China, transgenic PRSV-resistant papayas developed elsewhere in the world await regulatory clearances, social acceptance and/or marketability (Tecson Mendoza et al., 2008; Fitch, 2010). Transgenic PRSV-resistant papaya has been commercialized in China and is now being cultivated (Ye and Li, 2010). It will be necessary to monitor their ability to improve papaya production as well as susceptibility to other viruses, pathogens and pests. Interestingly, contamination of native papaya plants and fruits in Hong Kong was recently demonstrated and found to originate from commercial/field-tested transgenic papayas from Hawaii, Taiwan and China. This prompted the authorities to remove labels for these papayas and treat them as non-transgenic ones (Agriculture, Fisheries and Conservation Department, 2017). Adoption of transgenic technology depends on local and region-specific factors including product demand and bio-safety considerations. Perceptions of environmental risks and food-safety concerns are major deterrents for social acceptance of transgenic technology. Furthermore, developing countries often lack means and infrastructure for enforcement of proper bio-safety laws. Awareness among farmers and consumers should be created with the help of various governmental and non-governmental organizations on the benefits of using genetically engineered crops and sustainable management practices. There is need for impetus to promote the science and strategies that will enable second and third generation transgenic virus-resistant papayas. Acknowledgements We thank Professor Dennis Gonsalves for permission to use images in Fig. 7.2. The research scholars (Parul Bhardwaj, Aashima Mehra and Tabasum Akhter) of Department of Botany at University of

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Delhi, India, are thanked for their input and feedback on drafts. The editor is thanked for his valuable suggestions on this chapter. References

Abel, P.P., Nelson, R.S., De, B., Hoffmann, N., Rogers, S.G., Fraley, R.T., and Beachy, R.N. (1986). Delay of disease development in transgenic plants that express the Tobacco mosaic virus coat protein gene. Science 232, 738–743. Abreu, P.M., Antunes, T.F., Magaña-Álvarez, A., Pérez-Brito, D., Tapia-Tussell, R., Ventura, J.A., Fernandes, A.A., and Fernandes, P.M. (2015a). A current overview of the Papaya meleira virus, an unusual plant virus.Viruses 7, 1853–1870. https://doi.org/10.3390/v7041853 Abreu, E.F., Daltro, C.B., Nogueira, E.O., Andrade, E.C., and Aragão, F.J. (2015b). Sequence and genome organization of papaya meleira virus infecting papaya in Brazil. Arch. Virol. 160, 3143–3147. https://doi.org/10.1007/ s00705-015-2605-x Agriculture, Fisheries and Conservation Department (AFCD) (2017). Genetically Modified Organisms (Control of Release) Ordinance Cap. 607. Expert Group. Report on the Survey of Genetically Modified Organisms in Hong Kong URL: www.afcd.gov.hk/ english/conservation/con_gmo/gmo_exp/files/ D i s c u s s i o n _ Pap er _ 0 2 _ G M O _ 2 0 1 7 _ G M Os _ Survey_2014_2016.pdf. Accessed 17 December 2017. Ali, I., Amin, I., Briddon, R.W., and Mansoor, S. (2013). Artificial microRNA-mediated resistance to the monopartite begomovirus Cotton leaf curl Burewala virus. Virol. J. 10, 231. https://doi.org/10.1186/1743422X-10-231 Amri, E., and Mamboya, F. (2012). Papain, a plant enzyme of biological importance: a review. Am. J. Biochem. Biotechnol. 8, 99–104. https://doi.org/10.3844/ ajbbsp.2012.99.104 Anandalakshmi, R., Pruss, G.J., Ge, X., Marathe, R., Mallory, A.C., Smith, T.H., and Vance, V.B. (1998). A viral suppressor of gene silencing in plants. Proc. Natl. Acad. Sci. U.S.A. 95, 13079–13084. Aragão, F.J.L. (2014). GM plants with RNAi-golden mosaic resistant bean. BMC Proceedings 8, O24. https://doi. org/10.1186/1753-6561-8-S4-O24 Aravind, G., Bhowmik, D., Duraivel, S., and Harish, G. (2013). Traditional and medicinal uses of Carica papaya. J. Med. Plants Stud. 1, 7–15. Azad, M.A., Amin, L., and Sidik, N.M. (2014). Gene technology for Papaya ringspot virus disease management. Scientific World Journal 2014, 768038. https://doi.org/10.1155/2014/768038 Bateson, M.F., Lines, R.E., Revill, P., Chaleeprom, W., Ha, C.V., Gibbs, A.J., and Dale, J.L. (2002). On the evolution and molecular epidemiology of the potyvirus Papaya ringspot virus. J. Gen. Virol. 83, 2575–2585. Bau, H.J., Kung, Y.J., Raja, J.A., Chan, S.J., Chen, K.C., Chen, Y.K., Wu, H.W., and Yeh, S.D. (2008). Potential threat of a new pathotype of Papaya leaf distortion mosaic virus infecting transgenic papaya resistant to Papaya ringspot virus. Phytopathology 98, 848–856. https://doi. org/10.1094/PHYTO-98-7-0848

Bayot, R.G., Villegas, V.N., Magdalita, P.M., Jovellana, M.D., Espino, T.M., and Exconde, S.B. (1990). Seed transmissibility of Papaya ringspot virus. Philipp. J. of Crop Sci. 15, 107–111. Bonfim, K., Faria, J.C., Nogueira, E.O., Mendes, E.A., and Aragão, F.J. (2007). RNAi-mediated resistance to Bean golden mosaic virus in genetically engineered common bean (Phaseolus vulgaris). Mol. Plant Microbe Interact. 20,7 17–726. https://doi.org/10.1094/MPMI-20-60717 Bortesi, L., and Fischer, R. (2015). The CRISPR/ Cas9 system for plant genome editing and beyond. Biotechnol. Adv. 33, 41–52. https://doi.org/10.1016/j. biotechadv.2014.12.006 Brodersen, P., and Voinnet, O. (2006). The diversity of RNA silencing pathways in plants. Trends Genet. 22, 268–280. Bucher, E., Lohuis, D., van Poppel, P.M., GeertsDimitriadou, C., Goldbach, R., and Prins, M. (2006). Multiple virus resistance at a high frequency using a single transgene construct. J. Gen.Virol. 87, 3697–3701. https://doi.org/10.1099/vir.0.82276-0 Chadha, K., Viswanath, C., and Dutt, S. (2002). Fruit Crops: Papaya. In Handbook of Horticulture (ICAR, New Delhi), pp. 255–259. Chandra, R., Mishra, M., Pati, R., Agarwal, S., and Jain, R. (2008). Shoot tip transformation in papaya (Carica papaya L.). Paper presented at II International Symposium on Papaya, Acta Hortic. 851, 219–226. https://doi.org/10.17660/Acta Hortic.2010.851.32 Chandrashekara, K.R., Kumar, G.A., Krishnamurthy, K.B., Jagadish, K., Rashmi, S., and Vani, A. (2006). Studies on transient expression of coat protein (CP) gene of Papaya ringspot virus (PRSV) towards developing transgenic resistance in papaya. Paper presented at Sixteenth Annual Convention and International Symposium of Indian Virological Society on Management of Vectorborne Viruses at ICRISAT (Hyderabad, India). Chandrasekaran, J., Brumin, M., Wolf, D., Leibman, D., Klap, C., Pearlsman, M., Sherman, A., Arazi, T., and GalOn, A. (2016). Development of broad virus resistance in non-transgenic cucumber using CRISPR/Cas9 technology. Mol. Plant Pathol. 17, 1140–1153. https:// doi.org/10.1111/mpp.12375 Chávez-Calvillo, G., Contreras-Paredes, C.A., MoraMacias, J., Noa-Carrazana, J.C., Serrano-Rubio, A.A., Dinkova, T.D., Tripp, M.C., and Silva-Rosales, L. (2016). Antagonism or synergism between Papaya ringspot virus and papaya mosaic virus in Carica papaya is determined by their order of infection. Virology 489, 179–191. https://doi.org/10.1016/j.virol.2015.11.026 Chung, B.Y., Miller, W.A., Atkins, J.F., and Firth, A.E. (2008). An overlapping essential gene in the Potyviridae. Proc. Natl. Acad. Sci. U.S.A. 105, 5897–5902. https://doi. org/10.1073/pnas.0800468105 Damayanti, D., Ambarwati, U.T., Mariska, I., Hutami, S., Purnamaningsih, R., and Herman, M. (2001). Transformation study of Carica papaya L. using CO-PRSV and antisense ACC oxidase genes via particle bombardment. Paper presented at UK Papaya Biotechnology Network Meeting (Vietnam). Davis, M.J., and Ying, Z. (2004). Development of papaya breeding lines with transgenic resistance to Papaya

154  | Fermin et al.

ringspot virus. Plant Dis. 88, 352–358. https://doi. org/10.1094/PDIS.2004.88.4.352 Drew, R., Siar, S., Dillon, S., Ramage, C., O’Brien, C., and Sajise, A. (2005). Intergeneric hybridisation between Carica papaya and wild Vasconcellea species and identification of a PRSV-P resistance gene. Paper presented at International Symposium on Biotechnology of Temperate Fruit Crops and Tropical Species 738. Fabrick, J.A., Ponnuraj, J., Singh, A., Tanwar, R.K., Unnithan, G.C., Yelich, A.J., Li, X., Carrière, Y., and Tabashnik, B.E. (2014). Alternative splicing and highly variable cadherin transcripts associated with field-evolved resistance of pink bollworm to bt cotton in India. PLOS ONE 9, e97900. https://doi.org/10.1371/journal. pone.0097900 Fermin, G., Inglessis, V., Garboza, C., Rangel, S., Dagert, M., and Gonsalves, D. (2004). Engineered resistance to Papaya ringspot virus in Venezuelan transgenic papayas. Plant Disease 88, 516–522. https://doi.org/10.1094/ PDIS.2004.88.5.516 Fermin, G.A., Castro, L.T., and Tennant, P.F. (2010). CP-transgenic and non-transgenic approaches for the control of Papaya ringspot: current situation and challenges. Transgenic Plant J. 4, 1–15. Fermín, G., Keith, R.C., Suzuki, J.Y., Ferreira, S.A., Gaskill, D.A., Pitz, K.Y., Manshardt, R.M.,Gonsalves, D., and Tripathi, S. (2011). Allergenicity assessment of the Papaya ringspot virus coat protein expressed in transgenic rainbow papaya. J. Agric. Food Chem. 59, 10006–10012. https://doi.org/10.1021/jf201194r Fermin, G., Randle, M., and Tennant, P. (2015). Papaya ringspot virus. In Diseases of Tropical and Subtropical Crops, Tennant, P., and Fermin, G., eds. (CAB International, Wallingford), pp. 144–161. Ferreira, S.A., Pitz, K.Y., Manshardt, R., Zee, F., Fitch, M., and Gonsalves, D. (2002). Virus coat protein transgenic papaya provides practical control of Papaya ringspot virus in Hawaii. Plant Dis. 86, 101–105. https://doi. org/10.1094/PDIS.2002.86.2.101 Fitch, M.M., Manshardt, R.M., Gonsalves, D., Slightom, J.L., and Sanford, J.C. (1992). Virus resistant papaya plants derived from tissues bombarded with the coat protein gene of Papaya ringspot virus. Nature Biotechnol. 101, 466–1472. https://doi.org/10.1038/nbt1192-1466 Fitch, M.M., Manshardt, R.M., Gonsalves, D., and Slightom, J.L. (1993). Transgenic papaya plants from Agrobacterium-mediated transformation of somatic embryos. Plant Cell Rep. 12, 245–249. https://doi. org/10.1007/BF00237128 Fitch, M.M. (2010). Papaya ringspot virus (PRSV) coat protein gene virus resistance in papaya: update on progress worldwide. Transgenic Plant J. 14, 16–28. Fitch, M. (2014). Update on gene transfer biotechnology of papaya. Paper presented at XXIX International Horticultural Congress on Horticulture: Sustaining Lives, Livelihoods and Landscapes (Brisbane, Australia). Food and Agriculture Organization (FAO) of the United Nations (2016). FAOSTAT. Available online: www. fao.org/faostat/en/#data/QC. Accessed 17 December 2017. Fondong, V.N., Nagalakshmi, U., and Dinesh-Kumar, S.P. (2016). Novel Functional Genomics Approaches: A Promising Future in the Combat Against Plant

Viruses. Phytopathology, 106, 1231–1239. https://doi. org/10.1094/PHYTO-03-16-0145-FI Fuchs, M., and Gonsalves, D. (2007). Safety of virusresistant transgenic plants two decades after their introduction: lessons from realistic field risk assessment studies. Annu. Rev. Phytopathol. 45, 173–202. https:// doi.org/10.1146/annurev.phyto.45.062806.094434 Gonsalves, D. (1998). Control of Papaya ringspot virus in papaya: a case study. Annu. Rev. Phytopathol. 36, 415– 437. https://doi.org/10.1146/annurev.phyto.36.1.415 Gonsalves, D. (2004). Transgenic papaya in Hawaii and beyond. Agbioforum 7, 36–40. Gonsalves, C., Lee, D.R., and Gonsalves, D. (2004). Transgenic virus resistant papaya: the Hawaiian ‘Rainbow’ was rapidly adopted by farmers and is of major importance in Hawaii today. APSnet Features. https://doi.org/10.1094/APSnetFeature-2004-0804 Gonsalves, D., Vegas, A., Prasartsee, V., Drew, R., Suzuki, J., and Tripathi, S. (2006). Developing papaya to control Papaya ringspot virus by transgenic resistance, intergeneric hybridization, and tolerance breeding. In Plant Breeding Reviews, Janick, J., ed. ( John Wiley and Sons, Hoboken), vol. 26, pp. 35–73. Gonsalves, D., Tripathi, S., Carr, J.B., and Suzuki, J.Y. (2010). Papaya ringspot virus. The Plant Health Instructor. https://doi.org/10.1094/PHI-I-2010-1004-01 Gonsalves, D. (2015). The wayward Hawaiian boy returns home. Annu. Rev. Phytopathol. 53, 1–17. https://doi. org/10.1146/annurev-phyto-080614-120314. Grumet, R. (1990). Genetically engineered plant virus resistance. Hort. Sci. 25, 508–513. Guo, J., Yang, L., Liu, X., Guan, X., Jiang, L., and Zhang, D. (2009). Characterization of the exogenous insert and development of event-specific PCR detection methods for genetically modified Huanong No. 1 papaya. J. Agric. Food Chem.57, 7205–7212. https://doi.org/10.1021/ jf901198x Hamilton, A.J., and Baulcombe, D.C. (1999). A species of small antisense RNA in posttranscriptional gene silencing in plants. Science 286, 950–952. https://doi. org/10.1126/science.286.5441.950 Jain, R.K., Sharma, J., Sivakumar, A.S., Sharma, P.K., Byadgi, A.S., Verma, A.K., and Varma, A. (2004). Variability in the coat protein gene of Papaya ringspot virus isolates from multiple locations in India. Arch. Virol. 149, 2435– 2442. https://doi.org/10.1007/s00705-004-0392-x James, C. (2015). 20th anniversary (1996 to 2015) of the global commercialization of biotech crops and biotech crop highlights in 2015. ISAAA Brief No. 51. (International Service for the Acquisition of Agribiotech Applications, New York). Jia, R., Zhao, H., Huang, J., Kong, H., Zhang, Y., Guo, J., Huang, Q., Guo, Y., Wei, Q., Zuo, J., et al. (2017). Use of RNAi technology to develop a PRSV-resistant transgenic papaya. Sci. Rep. 7, 12636. https://doi.org/10.1038/ s41598-017-13049-0 Kalleshwaraswamy, C.M., and Kumar, N.K. (2008). Transmission efficiency of Papaya ringspot virus by three aphid species. Phytopathology 98, 541–546. https:// doi.org/10.1094/PHYTO-98-5-0541 Kertbundit, S., Pongtanom, N., Ruanjan, P., Chantasingh, D., Tanwanchai, A., Panyim, S., and Juříček, M. (2007).

Papaya Virus and Transgenic Resistance |  155

Resistance of transgenic papaya plants to Papaya ringspot virus. Biol. Plantar. 51, 333–339. Kreuze, J.F., and Valkonen, J.P. (2017). Utilization of engineered resistance to viruses in crops of the developing world, with emphasis on sub-Saharan Africa. Curr. Opin. Virol. 26, 90–97. Kung, Y.J., Bau, H.J., Wu, Y.L., Huang, C.H., Chen, T.M., and Yeh, S.D. (2009). Generation of transgenic papaya with double resistance to Papaya ringspot virus and Papaya leaf-distortion mosaic virus. Phytopathology 99, 1312–1320. https://doi.org/10.1094/PHYTO-99-111312 Kung, Y.J., You, B.J., Raja, J.A., Chen, K.C., Huang, C.H., Bau, H.J., Yang, C.F., Huang, C.H., Chang, C.P., and Yeh, S.D. (2015). Nucleotide sequence-homologyindependent breakdown of transgenic resistance by more virulent virus strains and a potential solution. Sci. Rep. 5, 9804. https://doi.org/10.1038/srep09804 Lawas, T.P., and Magdalita, P. (2007). Biotech papaya resistant to PRSV now under confined trials. ABSPII Newslett 3, 1. Lin, Y.T., Jan, F.J., Lin, C.W., Chung, C.H., Chen, J.C., Yeh, S.D., and Ku, H.M. (2013). Differential gene expression in response to Papaya ringspot virus infection in Cucumis metuliferus using cDNA-amplified fragment length polymorphism analysis. PLOS ONE 8, e68749. https:// doi.org/10.1371/journal.pone.0068749 Lindbo, J.A., and Dougherty, W.G. (1992). Untranslatable transcripts of the tobacco etch virus coat protein gene sequence can interfere with tobacco etch virus replication in transgenic plants and protoplasts.Virology 189, 725–733. Lindbo, J.A., and Falk, B.W. (2017). The impact of ‘coat protein-mediated virus resistance’ in applied plant pathology and basic research. Phytopathology 107, 624–634. https://doi.org/10.1094/PHYTO-12-160442-RVW Lines, R.E., Persley, D., Dale, J.L., Drew, R., and Bateson, M.F. (2002). Genetically engineered immunity to Papaya ringspot virus in Australian papaya cultivars. Mol. Breed. 10, 119–129. Lius, S., Manshardt, R.M., Fitch, M.M.M., Slightom, J.L., Sanford, J.C., and Gonsalves, D. (1997). Pathogenderived resistance provides papaya with effective protection against Papaya ringspot virus. Mol. Breed. 3, 161-168. https://doi.org/10.1023/A:1009614508659 Maciel‐Zambolim, E., Kunieda‐Alonso, S., Matsuoka, K., De Carvalho, M.G., and Zerbini, F.M. (2003). Purification and some properties of papaya meleira virus, a novel virus infecting papayas in Brazil. Plant Pathol. 52, 389–394. https://doi.org/10.1046/j.13653059.2003.00855.x Malabadi, R.B., Kumar, S.V., Mulgund, G.S., and Nataraja, K. (2011). Induction of somatic embryogenesis in papaya (Carica papaya). Res. Biotechnol. 2, 40–55. Manshardt, R.M. (1992). Papaya. In Biotechnology of Perennial Fruit Crops, Hammerschlag, F.A., Litz, R.E., eds. (CAB International, Wallingford), pp. 489–511. Maoka, T., and Hataya, T. (2005). The Complete Nucleotide Sequence and Biotype Variability of Papaya leaf distortion mosaic virus. Phytopathology 95, 128–135. https://doi.org/10.1094/PHYTO-95-0128

Maruthi, M.N., Rekha, A.R., Mirza, S.H., Alam, S.N., and Colvin, J. (2007). PCR-based detection and partial genome sequencing indicate high genetic diversity in Bangladeshi begomoviruses and their whitefly vector, Bemisia tabaci. Virus Genes 34, 373–385. https://doi. org/10.1007/s11262-006-0027-2 Mekako, H.U., and Nakasone, H.Y. (1975). Floral development and compatibility studies of Carica species. J. Am. Soc. Hortic. Sci. 100, 145–148. Ming, R., Hou, S., Feng, Y., Yu, Q., Dionne-Laporte, A., Saw, J.H., Senin, P., Wang, W., Ly, B.V., Lewis, K.L., et al. (2008). The draft genome of the transgenic tropical fruit tree papaya (Carica papaya Linnaeus). Nature 452, 991–996. https://doi.org/10.1038/nature06856 Mishra, R., Gaur, R.K., and Patil, B.L. (2016). Current knowledge of viruses infecting papaya and their transgenic management. In Plant Viruses: Evolution and Management, Gaur, R., Petrov, N., Patil, B., Stoyanova, M., eds. (Springer, Singapore), pp. 189–203. https:// doi.org/10.1007/978-981-10-1406-211 Moriones, E., Praveen, S., and Chakraborty, S. (2017). Tomato leaf curl New Delhi virus: an emerging virus complex threatening vegetable and fiber crops.Viruses 9, E264. Nariani, T.K. (1956). Leaf curl of papaya. Indian Phytopathol. 9, 151–157. Nascimento, A.K.Q., Lima, J.A.A., Nascimento, A.L.L., Beserra Jr, E.A., and Purcifull, D.E. (2010). Biological, physical, and molecular properties of a papaya lethal yellowing virus isolate. Plant Dis. 94, 1206–1212. https://doi.org/10.1094/PDIS-11-09-0733 Nawaz-ul-Rehman, M.S., and Fauquet, C.M. (2009). Evolution of geminiviruses and their satellites.FEBS Lett. 583, 1825–1832. https://doi.org/10.1016/j. febslet.2009.05.045 Niu, Q.W., Lin, S.S., Reyes, J.L., Chen, K.C., Wu, H.W., Yeh, S.D., and Chua, N.H. (2006). Expression of artificial microRNAs in transgenic Arabidopsis thaliana confers virus resistance. Nat. Biotechnol. 24, 1420–1428. Niu, Q.W., Lin, S.S., Reyes, J.L., Chen, K.C., Wu, H.W., Yeh, S.D., and Chua, N.H. (2006). Expression of artificial microRNAs in transgenic Arabidopsis thaliana confers virus resistance. Nat. Biotechnol. 24, 1420–1428. https://doi.org/10.1038/nbt1255 Noa-Carrazana, J.C., and Silva-Rosales, L. (2001). First report of a Mexican isolate of papaya mosaic virus in papaya (Carica papaya) and pumpkin (Cucurbita pepo). Plant Dis. 85, 558–558. https://doi.org/10.1094/ PDIS.2001.85.5.558A Noa-Carrazana, J.C., González-de-León, D., RuizCastro, B.S., Piñero, D., and Silva-Rosales, L. (2006). Distribution of Papaya ringspot virus and papaya mosaic virus in papaya plants (Carica papaya) in Mexico. Plant Dis. 90, 1004–1011. https://doi.org/10.1094/PD-901004 Noman, A., Aqeel, M., and He, S. (2016). CRISPRCas9: tool for qualitative and quantitative plant genome editing. Front. Plant Sci. 7, 1740. https://doi. org/10.3389/fpls.2016.01740 Olarte Castillo, X.A., Fermin, G., Tabima, J., Rojas, Y., Tennant, P.F., Fuchs, M., Sierra, R., Bernal, A.J., and Restrepo, S. (2011). Phylogeography and molecular epidemiology of Papaya ringspot virus.

156  | Fermin et al.

Virus Res. 159, 132–140. https://doi.org/10.1016/j. virusres.2011.04.011 Ogata, T., Yamanaka, S., Shoda, M., Urasaki, N., and Yamamoto, T. (2016). Current status of tropical fruit breeding and genetics for three tropical fruit species cultivated in Japan: pineapple, mango, and papaya. Breed. Sci. 66, 69–81. https://doi.org/10.1270/jsbbs.66.69 Pang, S.Z., Ja, F.J., Carney, K., Stout, J., Tricoli, D.M., Quemada, H.D., and Gonsalves, D. (1996). Post‐ transcriptional transgene silencing and consequent tospovirus resistance in transgenic lettuce are affected by transgene dosage and plant development. Plant J. 9, 899–909. https://doi.org/10.1046/j.1365313X.1996.9060899.x Patil, B.L., and Fauquet, C.M. (2009). Cassava mosaic geminiviruses: actual knowledge and perspectives. Mol. Plant Pathol. 10, 685–701. https://doi.org/10.1111/ j.1364-3703.2009.00559.x Patil, B.L., Ogwok, E., Wagaba, H., Mohammed, I.U., Yadav, J.S., Bagewadi, B., Taylor,N.J., Kreuze, J.F., Maruthi, M.N., Alicai, T., et al. (2011). RNAi-mediated resistance to diverse isolates belonging to two virus species involved in Cassava brown streak disease. Mol. Plant Pathol. 12, 31–41. https://doi.org/10.1111/j.13643703.2010.00650.x Pereira, A.J., Alfenas-Zerbini, P., Cascardo, R.S., Andrade, E.C., and Murilo Zerbini, F. (2012). Analysis of the fulllength genome sequence of papaya lethal yellowing virus (PLYV), determined by deep sequencing, confirms its classification in the genus Sobemovirus. Arch. Virol. 157, 2009–2011. https://doi.org/10.1007/s00705-0121384-x Purcifull, D.E., Edwardson, J.R., Hiebert, E., and Gonsalves, D. (1984). Papaya ringspot virus. CMI/AAB Descr. Plant Viruses, 292, 8. Pyott, D.E., Sheehan, E., and Molnar, A. (2016). Engineering of CRISPR/Cas9-mediated potyvirus resistance in transgene-free Arabidopsis plants. Mol. Plant Pathol. 17, 1276–1288. https://doi.org/10.1111/mpp.12417 Quito-Avila, D.F., Alvarez, R.A., Ibarra, M.A., and Martin, R.R. (2015). Detection and partial genome sequence of a new umbra-like virus of papaya discovered in Ecuador. Eur. J. Plant Pathol. 143, 199–204. https://doi. org/10.1007/s10658-015-0675-y Rao, X.Q., Li, Y.Y., Ruan, X.L., Yan, C., Zhang, S.G., and Li, H.P. (2012). Variation of neomycin phosphotransferase II marker gene in transgenic papaya plants under the field cultivation. Food Biotechnol. 26, 293–306. https://doi. org/10.1080/08905436.2012.723605 Roberts, M., Minott, D.A., Tennant, P.F., and Jackson, J.C. (2008). Assessment of compositional changes during ripening of transgenic papaya modified for protection against Papaya ringspot virus. J. Sci. Food and Agriculture 88, 1911–1920. https://doi.org/10.1002/jsfa.3295 Rojas, M.R., Hagen, C., Lucas, W.J., and Gilbertson, R.L. (2005). Exploiting chinks in the plant’s armor: evolution and emergence of geminiviruses. Annu. Rev. Phytopathol. 43, 361–394. https://doi.org/10.1146/ annurev.phyto.43.040204.135939 Sá Antunes, T.F., Amaral, R.J., Ventura, J.A., Godinho, M.T., Amaral, J.G., Souza, F.O., Zerbini, P.A., Zerbini, F.M., and Fernandes, P.M. (2016). The dsRNA Virus Papaya Meleira Virus and an ssRNA Virus are associated with

Papaya Sticky Disease. PLOS ONE 11, e0155240. https://doi.org/10.1371/journal.pone.0155240 Sanford, J.C., and Johnston, S.A. (1985). The concept of parasite-derived resistance-deriving resistance genes from the parasite’s own genome. J. Theoretical Biol. 113, 395–405. https://doi.org/10.1016/S00225193(85)80234-4 Saxena, S., Hallan, V., Singh, B.P., and Sane, P.V. (1998). Leaf curl disease of Carica papaya from India may be caused by a bipartite geminivirus. Plant Dis. 82, 126. https:// doi.org/10.1094/PDIS.1998.82.1.126A Shen, W., Tuo, D., Yan, P., Li, X., and Zhou, P. (2014). Detection of Papaya leaf distortion mosaic virus by reverse-transcription loop-mediated isothermal amplification. J. Virol. Methods 195, 174–179. https:// doi.org/10.1016/j.jviromet.2013.09.011 Singh-Pant, P., Pant, P., Mukherjee, S.K., and MazumdarLeighton, S. (2012). Spatial and temporal diversity of begomoviral complexes in papayas with leaf curl disease. Arch. Virol. 157, 1217–1232. https://doi.org/10.1007/ s00705-012-1287-x Sit, T.L., Abouhaidar, M.G., and Holy, S. (1989). Nucleotide sequence of papaya mosaic virus RNA. J. Gen. Virol. 70, 2325–2331. https://doi.org/10.1099/0022-1317-70-92325 Souza Júnior, M.T., Nickel, O., and Gonsalves, D. (2005). Development of virus resistant transgenic papayas expressing the coat protein gene from a Brazilian isolate of Papaya ringspot virus. Fitopatol. Bras 30, 357–365. https://doi.org/10.1590/S0100-41582005000400004 Srinivasulu, M., and Gopal, D.S. (2012). Coat protein sequence comparison of south Indian isolates of Papaya ringspot virus with other Indian subcontinent isolates. Phytopathol. Mediterr. 50, 359–369. https://doi. org/10.14601/Phytopathol_Mediterr-9085 Storey, W.B. (1976). Papaya. In Evolution of Crop Plants, Simmonds, N.W., ed. (Longman, London), pp. 21–24. Tarora, K., Shudo, A., Kawano, S., Yasuda, K., Ueno, H., Matsumura, H., and Urasaki, N. (2016). Development of plants resistant to Papaya leaf distortion mosaic virus by intergeneric hybridization between Carica papaya and Vasconcellea cundinamarcensis. Breed. Sci. 66, 734–741. https://doi.org/10.1270/jsbbs.16107 Tecson Mendoza, E.M., Laurena, A.C., and Botella, J.R. (2008). Recent advances in the development of transgenic papaya technology. Biotechnol. Annu. Rev. 14, 423–462. https://doi.org/10.1016/S13872656(08)00019-7 Tennant, P., Fermin, G., Fitch, M.M., Manshardt, R.M., Slightom, J.L., and Gonsalves, D. (2001). Papaya ringspot virus resistance of transgenic Rainbow and SunUp is affected by gene dosage, plant development, and coat protein homology. Eur. J. Plant Pathol. 107, 645–653. https://doi.org/10.1023/A:1017936226557 Tennant, P., Ahmad, M.H., and Gonsalves, D. (2005). Field resistance of coat protein transgenic papaya to Papaya ringspot virus in Jamaica. Plant Dis. 89, 841–847. https://doi.org/10.1094/PD-89-0841 Tennant, P.F., Fermin, G.A., and Roye, M.E. (2007). Viruses infecting papaya (Carica papaya L.): etiology, pathogenesis, and molecular biology. Plant Viruses 1, 178–188.

Papaya Virus and Transgenic Resistance |  157

Tricoli, D.M., Carney, K.J., Russell, P.F., McMaster, J.R., Groff, D.W., Hadden, K.C., Himmel, P.T., Hubbard, J.P., Boeshore, M.L., and Quemada, H.D. (1995). Field evaluation of transgenic squash containing single or multiple virus coat protein gene constructs for resistance to Cucumber mosaic virus, Watermelon mosaic virus 2, and zucchini yellow mosaic virus. Nature Biotechnol. 13, 1458–1465. https://doi.org/10.1038/nbt1295-1458 Tripathi, S., Suzuki, J.Y., Ferreira, S.A., and Gonsalves, D. (2008). Papaya ringspot virus-P: characteristics, pathogenicity, sequence variability and control. Mol. Plant Pathol. 9, 269–280. https://doi.org/10.1111/ j.1364-3703.2008.00467.x Tuo, D., Shen, W., Yan, P., Li, C.h., Gao, L., Li, X., Li, H., and Zhou, P. (2013). Complete genome sequence of an isolate of papaya leaf distortion mosaic virus from commercialized PRSV-resistant transgenic papaya in China. Acta Virol. 57, 452–455. https://doi. org/10.419/av_2013_04_452 Varma, A., and Malathi, V.G. (2003). Emerging geminivirus problems: a serious threat to crop production. Ann. App. Biol. 142, 145–164. https://doi. org/10.1111/j.1744-7348.2003.tb00240.x VIB (2014). Virus resistant papaya in Hawaii: the local papaya industry’s life raft. In Facts Series: Virus resistant papaya in Hawaii (Gent, Belgium). Wagaba, H.,Patil, B.L., Mukasa, S., Alicai, T., Fauquet, C.M., and Taylor, N.J. (2016). Artificial microRNAderived resistance to Cassava brown streak disease. J. Virol. Methods 231, 38–43. https://doi.org/10.1016/j. jviromet.2016.02.004 Wan, P., Huang, Y., Wu, H., Huang, M., Cong, S., Tabashnik, B.E., and Wu, K. (2012). Increased frequency of pink bollworm resistance to Bt toxin Cry1Ac in China. PLOS ONE 7, e29975. https://doi.org/10.1371/journal. pone.0029975 Wang, X., Xie, Y., and Zhou, X. (2004). Molecular characterization of two distinct begomoviruses from Papaya in China.Virus Genes 29, 303–309.

Waterhouse, P.M., Graham, M.W., and Wang, M.B. (1998). Virus resistance and gene silencing in plants can be induced by simultaneous expression of sense and antisense RNA. Proc. Natl. Acad. Sci. U.S.A. 95, 13959– 13964. https://doi.org/10.1073/pnas.95.23.13959 Ye, C., and Li, H. (2010). 20 years of transgenic research in China for resistance to Papaya ringspot virus. Transgenic Plant J. 4, 58–63. Yeh, S.D., and Gonsalves, D. (1985). Translation of Papaya ringspot virus RNA in vitro: detection of a possible polyprotein that is processed for capsid protein, cylindrical-inclusion protein, and amorphousinclusion protein.Virology 143, 260–271.https://doi. org/10.1016/0042-6822(85)90113-8 Yeh, S.D., Jan, F.J., Chiang, C.H., Doong, T.J., Chen, M.C., Chung, P.H., and Bau, H.J. (1992). Complete nucleotide sequence and genetic organization of Papaya ringspot virus RNA. J. Gen. Virol. 73, 2531–2541. https://doi. org/10.1099/0022-1317-73-10-2531 Yeh, S.D., and Gonsalves, D. (1994). Practices and perspective of control of Papaya ringspot virus by crossprotection. In Advances in Disease Vector Research, Harris, K.F., ed. (Springer, New York), pp. 237–257. https://doi.org/10.1007/978-1-4612-2590-4_9 Yeh, S.D., Kung, Y.J., Bau, H.J., Yu, T.A., and Raja, J.A. (2010). Generation of a papaya hybrid variety with broad-spectrum transgenic resistance to Papaya ringspot virus and papaya leaf-distortion mosaic virus. Transgenic Plant J. 4, 37–44. Zhou, X. (2013). Advances in understanding begomovirus satellites. Annu. Rev. Phytopathol. l51, 357–381. https:// doi.org/10.1146/annurev-phyto-082712-102234 Zimmerman, T.W., Joseph, L., St. Brice, N., and Kowalski, J.A. (2007). Development and selection for homozygous transgenic papaya seedling. Acta Hortic. 740, 177–182. https://doi.org/10.17660/ActaHortic.2007.740.20

Development and Delivery of Transgenic Virus-resistant Cassava in East Africa Henry Wagaba,1 Andrew Kiggundu2 and Nigel Taylor2*

8

1National Crops Resources Research Institute, National Agricultural Research Organization (NARO), Kampala,

Uganda.

2Donald Danforth Plant Science Center, St. Louis, MO, USA.

*Correspondence: [email protected] https://doi.org/10.21775/9781910190814.08

Abstract Cassava (Manihot esculenta) is a major staple food crop in sub-Saharan Africa where it provides food security and income to rural communities. The two virus diseases Cassava mosaic disease (CMD) and Cassava brown streak disease (CBSD) constrain cassava production in these regions. RNA interference (RNAi) technology has been developed to address both diseases and has been shown to be effective in greenhouse and field studies. Field trials in Uganda and Kenya have demonstrated the potential of this technology to increase usable storage root yields greater than 50-fold under high CBSD pressure. This resistance is maintained across the cropping cycle and geographic locations from central Uganda to coastal Kenya, indicating its potential to help secure cassava production in East Africa. This review describes advances made in applying transgenic approaches to control CBSD and CMD and the challenges faced in delivering the improved planting materials to benefit cassava farmers in East Africa. Introduction Cassava (Manihot esculenta Crantz) is a tropical perennial shrub cultivated as a staple food crop by smallholder farmers across the tropics, where it is the most important source of dietary calories after rice and maize (FAO, 2015). Cassava tuberous roots are

high in carbohydrate and starch and can be cooked and processed in many ways for daily meals. Some communities also use the tender leaves as a green vegetable. Cassava generally performs well in places where soils are less fertile and rainfall patterns are less predictable. This makes cassava a crop resilient to stresses associated with climate change, ensuring that it will remain a central food security crop for the 21st century ( Jarvis et al., 2012). Additionally, cassava is an important industrial crop for animal feed (Morgan and Choct, 2016), starch (Marcon et al., 2007; Srinivas, 2007), food processing (Balagopalan, 2002), paper products (Aripin et al., 2013), textiles, cosmetics, pharmaceuticals (Nweke, 1996) biofuels and bio-plastics (Ademiluyi and Mepba, 2013; Nuwamanya et al., 2012; Tumwesigye et al., 2016a,b). World production of cassava exceeds 268 million metric tonnes (FAO, 2015), with more than 50% produced in sub-Saharan Africa. Despite Africa being the highest producer, most international trade is concentrated in Asia, with China and Thailand being the largest importer and exporter of raw cassava products, respectively (Curran and Cooke, 2008). Several African countries have recently earmarked cassava as an important industrial raw material and driver for economic growth through value addition, employment and income generation (Anyanwu et al., 2015). Despite these economic prospects, virus diseases remain a major threat to cassava production throughout

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sub-Saharan Africa and a constraint for its development as a commodity crop in these regions (Rey and Vanderschuren, 2017; Tomlinson et al., 2017). Importance of cassava mosaic disease and cassava brown streak disease Cassava mosaic disease (CMD) and cassava brown streak disease (CBSD) are the two most economically important diseases affecting cassava production in East and Central Africa. Both diseases are caused by viruses and result in significant losses in yield and quality of storage roots. Together CMD and CBSD contribute losses estimated at $1 billion per annum (IITA, 2014; Legg et al., 2006). CMD is caused by circular ssDNA cassava mosaic geminivirus (CMGs) belonging to the genus Begomovirus in the family Geminiviridae (Rey and Vanderschuren, 2017) and is endemic throughout sub-Saharan Africa. CMD was first reported in 1894 in the Usambara Mountain range in Northeast Tanzania, where infected plants were described as having symptoms of crippling/crinkling (Warburg, 1894). It was later found that the disease was transmitted by whiteflies (Chant, 1958), and caused by a virus (Harrison et al., 1977), later named the African Cassava mosaic virus (ACMV). Molecular characterization of ACMV isolates from CMD-affected plants in East Africa revealed that additional virus species existed, referred to as the East African Cassava mosaic virus (EACMV) (Frischmuth et al., 1997). To date, 11 distinct cassava mosaic species have been characterized worldwide from CMDinfected cassava, with nine present in sub-Saharan Africa (Brown et al., 2015; Rey and Vanderschuren, 2017; Rey et al., 2012). Crop losses due to CMD are greater when infection occurs at an early age, and when infected planting materials are used in a new cropping cycle. It is estimated that yield losses reaching 90% can occur in farmers’ fields depending on whether single or dual infections of viral species are present (Owor et al., 2004). CBSD causes estimated yield losses of up to $100 million per annum and presents a food security threat to families that rely on cassava as their main source of food (Manyong et al., 2012). In Tanzania alone, losses due to CBSD were estimated at more than $51 million (Ndyetabula et al., 2016). The disease was first described on field tests performed

in Amani coastal East Africa in the 1930s, when the name ‘brown streak’ was suggested (Storey, 1936). In 1950, Nichols (1950) described the disease’s geographical distribution to be limited to areas below 1000 metres above sea level. CBSD was first observed in Uganda in 1945 at Bukalasa experimental station within materials introduced earlier from Amani, Tanzania. Rouging of diseased materials led to eradication of the disease within the country until the early 2000s when CBSD re-emerged in Central Uganda (Alicai et al., 2016). The disease has since spread to the midhighlands of east Africa (Lake Victoria basin) across the Great Lakes region and into Central Africa, creating a pandemic. West Africa, the world’s largest cassava-growing region, is now under serious threat from CBSD. Early stem grafting experiments indicated that the causal agent of CBSD was a virus (Storey, 1936) with mechanical transmission of CBSD to various indicator hosts reported in the late 1950s (Lister, 1959). Another thirty-five years was required before virus particles were identified by electron microscopy analysis of CBSD-infected N. debneyi (Bock, 1994) and then cassava (Were et al., 2004). The Potyviridae sequence was finally confirmed seven years later through reverse transcription PCR (RT-PCR) on CBSD-infected N. benthamiana samples and identified as being closely related to Sweetpotato mild mottle virus (SPMMV, genus Ipomovirus, family Potyviridae) (Monger et al., 2001). For both CMD and CBSD, the biggest contributor to disease spread is movement and exchange of infected planting materials between cassava growers. Studies have shown that the CBSD pandemic is less associated with new virulent strains of the two causal viruses CBSV and UCBSV (Mbanzibwa et al., 2009a), but rather a result of the ‘super-abundance’ of whitefly populations (Legg et al., 2014b). These studies showed that a wave of high populations of whiteflies preceded waves of CBSD outbreaks. Other factors being investigated are whether cassava viruses manipulate plant hosts to attract more vectors, as has been reported elsewhere, and the role of the minimum temperature required for spread by the whitefly ( Jeremiah et al., 2015). Symptoms of CMD and CBSD Symptoms of CMD and the accompanying cellular modifications depend on whether one or more

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CMGs are present in the same ‘co-infected’ plant (Pita et al., 2001). CMD-infected plants show mosaic symptoms and distortion of leaves resulting in compromised photosynthetic capacity and consequent suppression of storage root yields (Legg et al., 2006) (Fig. 8.1). Reversion or recovery in cassava is also known to occur, attributed to incomplete systemic movement of the causal viruses such that whole branches or individual shoots become symptomless and free of detectable virus. When present in sufficient amount, such branches can provide uninfected cuttings for establishing the next cropping cycle (Mohammed et al., 2016). Symptoms of CMD can also be enhanced by recently discovered host plant DNA sequences referred to as sequence enhancing geminivirus symptoms 1 and 2 (SEGS-1 and SEGS-2). SEGS-1 and SEGS-2 are reported to be capable of breaking inherent CMD2-type resistance. The two SEGs differ in their mode of action. SEGS-1 is derived from the cassava genome and facilitates CMD infection, as either an integrated copy and/or an episome, while SEGS-2 is considered to have originated from the cassava genome but is now encapsulated into virions and

transmitted as an episome by the whitefly vectors (Ndunguru et al., 2016). Aerial display of CBSD symptoms varies depending on the cultivar and environmental conditions (Jeremiah et al., 2015). Typical CBSD symptoms include feathery chlorosis along leaf veins or circular chlorotic patches between the primary veins (Fig. 8.2A). Stems display brown necrotic streaks and in severe cases stem dieback (Fig. 8.2B). Symptoms in the tuberous roots consist of brown, corky necrosis of the starchy tissue and occasional radial constrictions (Fig. 8.2D and E). Investigations of the biochemical changes that occur in infected leaves indicate a reduction in chlorophyll a and b and carotenoids (Nuwamanya et al., 2017). The considerable variation in symptom types observed between cultivars, varieties and geographical area suggest differences in levels of viral infection, virulence and plant resistance (Hillocks and Maruthi, 2015). Due to variation in symptoms displayed in different cultivars and viral infections, recommendations to farmers as to how best identify and select clean plant material is challenging.

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Figure 8.1  Typical symptoms of cassava mosaic disease (CMD) on cassava. (A-C) Plants showing a distorted leaf with variations in mosaic and mottling symptoms; (D) asymptomatic healthy plant.

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Figure 8.2  Symptoms of cassava brown streak disease (CBSD). (A) Yellowing patches on the leaves and along leaf veins; (B) brown streaks on a young stem; (C) diseasefree cassava plant; (D) storage root deformation caused by CBSD; (E) brown necrosis in the storage roots; (F) healthy cassava roots.

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Aetiology of CMD and CBSD CMD is caused by a complex of single-stranded bipartite DNA viruses generally referred to as Cassava mosaic geminiviruses (CMGs) (Geminiviridae: Begomovirus) (Fauquet and Fargette, 1990). Nine distinct CMG species infect cassava in Africa including: African Cassava mosaic virus (ACMV), East African Cassava mosaic virus (EACMV), East African cassava mosaic Zanzibar virus (EACMZV), East African cassava mosaic Cameroon virus (EACMCV), East African cassava mosaic Malawi virus (EACMMV) and South African Cassava mosaic virus (SACMV). CMG genomes are composed of two circular single-stranded DNA (ssDNA) molecules, referred to as DNA A and DNA B of size 2.7–3.0 kb (Hanley-Bowdoin et al., 2013). DNA A encodes two overlapping virion-sense open reading frames (ORFs) involved in encapsidation (AV1) and silencing suppressor targeting post-transcriptional gene silencing (PTGS)(AV2 or V2) (Zhang et al., 2005), four partially overlapping complementary-sense ORFs necessary for replication (AC1, AC3) and for transcription and suppression of host-mediated gene silencing (AC2, AC4) (Böttcher et al., 2004). DNA B encodes two non-overlapping ORFs (BC1, BV1), each present on the complementary and virion strands that are involved in inter- and intra-cellular trafficking of the virus (Hanley-Bowdoin et al., 2013). The right- and left-ward transcriptional units of the DNA A and DNA B components, respectively, are separated by a homologous intergenic region (IR) of approximately 200 nucleotides (Hanley-Bowdoin et al., 2013). At least two closely related virus species are confirmed to cause CBSD: Cassava brown streak virus (CBSV) and Ugandan cassava brown streak virus (UCBSV) (Mbanzibwa et al., 2011a; Winter et al., 2010). Both species belong to the family Potyviridae, genus Ipomovirus, and possess positive sense single-stranded RNA (+ssRNA) genomes (Mbanzibwa et al., 2009a; Monger et al., 2001). Complete genome sequences of CBSV and UCBSV have shown ≈ 70% and ≈ 74% nucleotide and amino acid identity, respectively (Mbanzibwa et al., 2011b). A recent addition of 12 new genomes to the public database including seven CBSV and five of UCBSV, suggests up to four new species may be present (Ndunguru et al., 2015). CBSV-infected plants have been observed to produce more rapid

and severe CBSD symptoms than those on plants infected with UCBSV alone in field and laboratory experiments(Mohammed et al., 2012; Ogwok et al., 2016; Vanderschuren et al., 2012). The molecular mechanism behind this observation is not yet understood. Transmission of CMD and CBSD Both causal viruses of CMD and CBSD are transmitted by the whitefly vector species Bemisia tabaci, Gennadius (Maruthi et al., 2005). A recent report describes that spread of CBSD occurs systematically from a point source inoculum to a maximum of 17 m radius in the field (Maruthi et al., 2017). Increase in the number of the whitefly vector has raised fears that CBSD and new virulent forms of CMD will continue to spread into new territories in West Africa. This would likely result in increased food insecurity and impaired economic development. Climate change has most likely contributed to the spread of CMD and CBSD, as higher temperatures favour spread of the whitefly vector (Legg et al., 2014a). Further, B. tabaci seems to be evolving to adapt to warmer climates causing the diseases to spread into new geographical areas (Gilbertson et al., 2015). However, the majority of long distance transmissions of CMD and CBSD within farming communities occur via exchange of infected planting materials (Maruthi et al., 2017). Genetic diversity of CMD and CBSD viruses The complete genetic diversity of the Cassava mosaic virus complex was reviewed by Rey et al. (2012). This diversity includes: East African cassava mosaic (EACM) virus, the EACM-Kenya virus, the EACMMalawi virus, EACM-Zanzibar virus, Cassava mosaic Madagascar virus, and South African Cassava mosaic virus; African Cassava mosaic virus/East African Cassava mosaic virus recombinant variant East African Cassava mosaic virus (EACMV) Uganda; EACM-Cameroon virus; and the newest discovery, the African Cassava mosaic virus (ACMV)–Burkina Faso (Tiendrébéogo et al., 2012). On the Indian sub-continent, only two species are known to occur, Indian Cassava mosaic virus, and Sri Lankan Cassava mosaic virus (Saunders et al., 2002). The high diversity of Cassava mosaic virus species has implications

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on control measures employed or under development. In particular, breeding and transgenic approaches for resistance could be broken due to new variants caused in part to recombination as has been reported for EACMV, but also co-infections that worsen disease and productivity loss. The first diagnostic test for the virus causing CBSD was developed using the partial sequence of 1114 nucleotides from the coat protein (CP) gene of CBSV (Monger et al., 2001). The virus was subsequently named Cassava brown streak virus, and classified in the genus Ipomovirus, family Potyviridae. It was not until the full genome of the virus was available that it was compared to other Ipomoviridae (Mbanzibwa et al., 2009b) leading to the discovery that two distinct species cause CBSD (Winter et al., 2010). A recent review of the genome organization of the two virus species was reported by Patil et al. (2015). Additional genomic sequencing has brought 12 new genomes to the public GenBank, making a total of 26 total genomes to date (Ndunguru et al., 2015). Recombination patterns also identified eight and four recombination events amongst CBSV and UCBSV sequences respectively (Ndunguru et al., 2015). With this diversity, current diagnostic primers may not provide sufficiently accurate diagnosis of these viruses (Ndunguru et al., 2015). Phylogenetic analysis has also shown that CBSV evolves at least five times faster than UCBSV because it accumulates more non-synonymous mutations than the synonymous ones observed in UCBSV (Alicai et al., 2016). Co-infection of CBSV and UCBSV is common in field-grown cassava. However, highly susceptible cultivars accumulate high levels of viral RNAs of CBSV and UCBSV, while the less susceptible cultivars accumulate CBSV and negligible amounts of UCBSV (Ogwok et al., 2016). UCBSV is known to cause milder foliar symptoms than CBSV, indicating that CBSV is a more aggressive and virulent CBSD viral pathogen for reasons that remain largely unknown to date (Patil et al., 2011). Management of CBSD and CMD Management of CMD and CBSD is problematic and occurs through an integrated management approach involving phyto-sanitation, planting of CBSD and CMD-tolerant varieties, use of diseasefree (clean) planting materials and removal of

infected plants by rouging (Legg et al., 2014a). Control of the whitefly vector populations through spraying with chemical insecticides and biological methods can be effective in limiting the spread of disease. However, for the majority of cassava farmers in Africa who practise subsistence farming, the use of chemical sprays is not sustainable. Therefore, dissemination of resistant varieties remains the effective mechanism for securing yields in areas of high disease pressure (Horowitz et al., 2011). There are three major sources of host plant resistance to CMD currently exploited by breeders. These are referred to as CMD1, CMD2 and CMD3 (Beyene et al., 2016a). CMD1 is polygenic and recessive (Hahn et al., 1980; Fregene and PuontiKaerlas, 2002). CMD2 is dominant, monogenic and was discovered within related landraces collected from farmers’ fields in Nigeria and other West African countries (Akano et al., 2002; Okogbenin et al., 2012) CMD2-type landraces have been widely deployed in East, Central and West Africa (Legg et al., 2006; Rabbi et al., 2014). CMD3 exists as CMD2 plus a quantitative trait locus (QTL), identified in cultivar TMS 97/2205, and confers very high levels of resistance to CMD with little or no expression of disease on leaves (Okogbenin et al., 2012). To date, the gene(s) involved and the molecular mechanisms underlying CMD1, CMD2 and CMD3 resistance remain unknown. A role for transgenic cassava Cassava is a highly heterozygous outcrossing species. This coupled with long growing cycles and asynchronous flowering make genetic improvement using conventional means challenging (Fregene et al., 2003). Furthermore, the vegetative nature of cassava enables the accumulation of viruses and pests over successive cropping cycles. Advances in genomic sequencing (Bredeson et al., 2016) and molecular breeding methods have improved the precision and speed at which solutions can be obtained, but they alone cannot unlock the full potential of the crop. Genetic engineering can introduce genes for enhanced traits directly into farmer-preferred cultivars while maintaining farmer- and consumer-preferred traits. Genetic engineering also has the potential to introduce novel traits that do not exist or are lacking in the gene pool such as increased Fe and Zn concentrations

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and robust CBSD resistance. While a few CBSDresistant varieties such as ‘Namikonga’ exist, the trait is polygenic, making breeding for resistance to CMD and CBSD problematic. Transgenic, and more recently gene editing, technologies therefore offer attractive approaches for improving cassava varieties for use by farmers and breeders. Status of transgenic technologies in cassava Genetic engineering of cassava commenced in the 1990s (Chavarriaga-Aguirre et al., 2016). Current protocols for genetic transformation depend on the ability to form somatic embryos. (Chauhan et al., 2015; Taylor et al., 2012). Friable embryogenic callus (FEC) is the most frequently employed target tissue for transformation by co-culture with Agrobacterium tumefaciens. Previously, these processes were possible in a few model cassava varieties. Improvement in the tissue culture processes has expanded the range of transformable cultivars and increased efficiencies of transgenic plant recovery by up to 110-fold in some cultivars (Chauhan et al., 2015). Continued efforts are expanding the range of cultivars further (Chauhan et al., 2017) while development of alternative morphogenic systems present new potentials for application of transgenic and gene editing technologies (Chauhan and Taylor, 2017). Farmer-preferred African varieties with robust protocols in place for production of transgenic cassava include: TME 204, TME 419, TME 7, TME 3, TME 14, NASE 13, NASE 14, TMS 98/0505 and TMS 91/02324 (Bull et al., 2009; Chauhan et al., 2015). These have been transformed for different traits such as pro vitaminA and iron and zinc nutritional enhancement (Beyene et al., 2016a; Narayanan et al., 2015) and accelerated flowering (Bull et al., 2017), in addition to resistance to CMD and CBSD (Beyene et al., 2016a; Odipio et al., 2014; Ogwok et al., 2016; Vanderschuren et al., 2012; Wagaba et al., 2016a; Yadav et al., 2011). Transgenic strategies for CMD and CBSD resistance Initial studies on pathogen-derived resistance in N. benthamiana demonstrated that overexpression of the coat protein was not effective for controlling

ACMV (Frischmuth and Stanley, 1998). However, experiments in the early 2000s expressing double-stranded RNA from the AC1 gene (replication-associated protein gene) showed that resistance to ACMV could be achieved in transgenic cassava (Chellappan et al., 2004). Resistance to CMD was also obtained using a hairpin RNA construct targeting the viral promoter for post-transcriptional gene silencing (hp-TGS) (Vanderschuren et al., 2007). Targeting non-structural viral mRNAs of Rep (AC1), TrAP (AC2) and REn (AC3) using antisense mRNAs had also been reported to result in protection against ACMV (Zhang et al., 2005). In another study, a 155-nucleotide segment of ACMVKE AC1 expressed in the cassava cultivar 60444 as a hairpin double-stranded RNA (dsRNA) under the control of the constitutive CaMV 35S promoter showed strong resistance to the virus (Vanderschuren et al., 2009). Performance of transgenic cassava expressing the AC1 gene from EACMV has been demonstrated under field conditions in the experimental, CMD-susceptible variety 60444. Significant resistance to the EACMV was achieved over successive vegetative cropping cycles, while plants remained susceptible to infection with ACMV (Ogwok et al. unpublished). Unlike CBSD, significant success has been achieved in addressing CMD via conventional breeding, with breeders exploiting CMD1, CMD2 and CMD3-type sources of resistance (Okogbenin et al., 2012; Rabbi et al., 2014). Despite a lack of knowledge concerning the molecular mechanisms underlying CMD1, CMD2 and CMD3, farmerpreferred varieties carrying these sources of resistance have been selected and widely deployed across Africa (Dixon et al., 2010; Ntawuruhunga et al., 2013). Widespread reliance on the use of CMD2 is a cause for concern. Being monogenic/ monolocus and dominant in nature, it is highly favoured by breeders. However, two recent findings indicate that CMD2-type resistance can be broken in a rapid and predictable manner. Beyene et al. (2016b) report that CMD2-type cultivars lose resistance to CMD when they undergo somatic embryogenesis. In addition, the newly discovered phenomenon of SEGs can result in breakdown of CMD2-type resistance under laboratory and field conditions. Both reports increase concern over the durability of resistance to CMD in the many farmerpreferred cultivars deployed across large regions

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of East, Central and West Africa (Ndunguru et al., 2016). Interest in the use of transgenic approaches to CMD resistance therefore remains, and efforts to develop this further should be encouraged. Reports of CMD in Southeast Asia indeed show that there is need for continued efforts for resistance using these approaches (Karthikeyan et al., 2016; Wang et al., 2015). Greater successes have been achieved employing RNA interference (RNAi) technology against CBSD by expressing inverted repeat constructs generated from the coat protein (CP) gene of UCBSV and CBSV. Analysis of the CP sequence showed that the CP gene is the most conserved in both viruses (Mbanzibwa et al., 2011b). Targeting it therefore has potential to develop protection against both causal viruses. Successful control of CBSD was first demonstrated in Nicotiana benthamiana (Patil et al., 2011) and then in graft challenge experiments in transgenic cassava cv. 60444 (Yadav et al., 2011). Protection in the greenhouse was achieved against the UCBSV isolate from which the CP sequence was obtained. Performance in the field was superior and is described below. Vanderschuren et al. (2012) reported strong protection against CBSV and UCBSV by expressing an RNAi construct consisting of the CP sequence in the CMD2-type Nigerian landrace Okoiyawo (TME 7). Other virus genes have been targeted for resistance to CBSV and UCBSV using artificial microRNAs (amiRNAs) (Wagaba et al., 2016b). Some success was reported, but this did not match that achieved by expression of dsRNA from CP-derived hairpin constructs (Wagaba et al., 2013). Preliminary screens for resistance to CMD and CBSD Efficiency of vector transmission of CMGs and CBSVs varies greatly under different environmental conditions, and can be problematic to perform experimentally on a large scale. Therefore, before testing takes place in the field, various methods are utilized to evaluate cassava plants in screenhouse or greenhouse settings. These include graft inoculation from CMD- or CBSD-infected hosts to healthy plants (Vanderschuren et al., 2012; Wagaba et al., 2013; Yadav et al., 2011), direct delivery of DNA genomes as infectious clones via microparticle bombardment for assessment of CMD

(Beyene et al., 2016b; Chellappan et al., 2004), Agrobacterium-mediated inoculation of plants with cloned infectious DNA genomes and mechanical transmission of cloned viral DNA genomes by abrasion techniques (Ntui et al., 2015). These methods require up to 20 weeks for full assessment of CMD and/or CBSD symptom development after inoculation. Beyene et al. (2016a) recently described a method based on virus-induced gene silencing (VIGS) of the cassava SPINDLY (SPY) gene MeSPY1. Plants with robust resistance to CMD recover from this challenge, while susceptible lines undergo senescence of the apical meristem and subsequent death of whole plants. This VIGs-based system can be used to evaluate CMD resistance with data complete within 2–4 weeks of inoculation and can be applied as a high-throughput rapid screening system to assess transgenic lines and conventional germplasm under controlled greenhouse growth conditions. Additional methods using infectious clones are under development and are being tested on cassava for transmission of UCBSV and CBSVs. These promise to be important new tools for evaluating cassava germplasm in the greenhouse, helping to increase throughput for early selection of combined disease resistance before undertaking field trials. Developing transgenic virus-resistant cassava for deployment to African farmers VIRCA Plus, originally VIRCA (Taylor et al., 2012), is a collaborative project to develop cassava varieties with robust resistance to two viral plant diseases (CMD and CBSD) and deliver them to farmers, benefiting millions of households in East Africa and beyond. The project is field-testing transgenic cassava expressing RNAi technology that confers resistance to CBSD. It is pursuing strategies to combine this with resistance to CMD (which is endemic in the same areas as CBSD) and collecting data required by regulatory authorities in Uganda and Kenya for applications to release resistant cassava to smallholder farmers. VIRCA Plus is structured to include aspects of intellectual property, technology development and transfer, product development, biosafety, quality control, communication and stewardship required for genetically engineered crop product delivery

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(Taylor et al., 2012). The project is led by the Donald Danforth Plant Science Center (DDPSC), St. Louis, MO, USA, and conducts research and development activities together with the National Crops Resources Research Institute (NARO, NaCRRI), Uganda, the Kenya Agricultural and Livestock Research Organization (KALRO), Kenya, the National Root Crops Research Institute (NRCRI), Nigeria and the CGIAR system, through the International Institute for Tropical Agriculture (IITA), Nairobi, Kenya. Field-testing of transgenic cassava Many research activities culminate at the stage of greenhouse and laboratory testing with subsequent publication of results in the scientific literature. However, if the goal is to deliver benefits to farmers and consumers such as the case for the VIRCA Plus project, a significant programme of additional assessment is required. Before transgenic crops can be released for cultivation and consumption, they must undergo multiple rounds of testing and regulatory assessment, resulting in submission of comprehensive dossiers to regulatory authorities. Transgenic crops can only be made available to farmers after review and if approved by the designated national authorities. Confined field trials (CFTs) are an essential component of this process and are performed after proof-of-concept is shown under laboratory and greenhouse conditions. In CFTs, transgenic plants are grown within a field facility designed to ensure material and genetic confinement. Prior to implementation, all CFT sites are first approved by the national regulatory authority responsible in the respective countries. National regulatory authorities provide regulatory oversight before, during and after the completion of a CFT. In most cases, elite farmer-preferred varieties are the initial targets for genetic engineering with the intention of deploying the improved germplasm directly to farmers. As a vegetatively propagated crop, transgenic cassava plants are assessed in the hemizygous T0 state and in most cases will remain as T0 through field-testing, regulatory assessment and deployment. After regulatory approval, breeders may wish to access and incorporate the enhanced material into their breeding programmes.

CFTs are used to confirm the efficacy of the transgenic trait(s) under field conditions. With regards resistance to CBSD and CMD, it is important to perform these experiments under conditions of high disease pressure transmitted by the whitefly vector. Most commonly, initial field trials are composed of multiple independent transgenic events (between 10 and 30) that are assessed against each other and known controls for performance against the target disease and for presence of obvious phenotypic off-types. These ‘trait selection’ trials (TST) are followed by ‘yield selection’ trials (YST). In the latter, a smaller number of the best performing transgenic events (5–10) are tested, but in larger replications. This enables generation of statistically meaningful data for agronomic qualities such as yield, harvest index, plant architecture, and response to common pests and diseases beyond the target disease. YSTs are best carried out at multiple locations to take into account genetic by environmental interactions in performance of the transgenic events. Information generated within the YSTs is used to select lead events, usually the single best performing transgenic line and one backup, which enter multi-locational regulatory CFTs. Regulatory field trials (RFTs) focus on assessment of the lead lines for trait performance against the target disease and collection of data to demonstrate that unintentional changes have not taken place within the modified plant lines. Data for food and feed safety assessment are collected in accordance with Codex Alimentarius and guidelines published by OECD. Studies addressing environmental risk are also being carried out in order to meet requirements for compiling and submission of a regulatory dossier (Falck-Zepeda et al., 2012). Assessment of RNAi-mediated virus resistance to control CMD and CBSD requires that plants are tested in areas with high disease pressure. The Lake Victoria basin has a high incidence of CMD and CBSD. The Uganda National Crops Resources Research Institute (NaCRRI) at Namulonge, located in this basin, is therefore ideally placed to test plants in a research setting. The Ugandan National Biosafety Committee (NBC) has granted permission to perform CFTs at Namulonge and two additional sites in Uganda since 2009. Under these permissions, CFTs have been conducted to test plants expressing inverted repeats for EACMV AC1and CP sequences of UCBSV and CBSV.

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RNAi-mediated resistance to CMD AC1 is the only geminivirus gene indispensable for viral replication and has been the most successful for generating RNAi-derived resistance to CMD. First tested in N. benthamiana, Hong and Stanley (1996) generated transgenic plants expressing the AC1 gene from ACMV-KE. Resulting transgenic lines showed resistance to ACMV, characterized by reduced levels of viral DNA accumulation compared with control plants. Vanderschuren et al. (2009) described expression of a 155-nucleotide segment of ACMV-KE AC1 in the cassava cultivar 60444 as a hairpin double-stranded RNA (dsRNA) under the control of the CaMV 35S promoter. Consistent with the recent report of transient expression (Patil et al., 2016), transgenic cassava lines with high levels of AC1 siRNAs were observed to display very high resistance to ACMV, and levels of siRNAs were correlated with the observed resistance. Correspondingly, an inverted repeat RNAi construct containing the overlapping region between SLCMV AV1 and AV2 was recently used to transform cultivar KU50, which is cultivated extensively in Southeast Asia for fuel production (Ntui et al., 2015). Transgenic lines obtained displayed high levels of resistance to SLCMV compared with wild-type plants, and PCR analyses failed to detect viral DNA in systemic uninoculated leaves, suggesting immunity to SLCMV. The B-component of geminiviruses has also been shown to induce host resistance, although not commonly assessed. As described above, transgenic cassava plants of the CMD susceptible variety 60444 expressing a hairpin construct derived from the AC1 gene of EACMV displayed significant resistance to EACMV over successive vegetative cropping cycles at NaCRRI, Namulonge (Ogwok et al., unpublished). RNAi-mediated resistance to CBSD Initial understanding for efficacy of RNAi against CBSD was generated from three truncations of CP sequences derived from the UCBSV-[UG: Nam: 04] isolate. These consisted of the 894 nt near fulllength (FL), 397 nt N-terminal (NT) and 491 nt C-terminal (CT). The UCBSV-CP sequence was utilized as this was the only sequence available when these studies were initiated. During tests in the model host N. benthamiana, resistance was recorded against UCBSV (Patil et al., 2011) and

cross-protection shown against non-homologous CBSV isolates. The FL CP provided 100% protection in two of the highest siRNA-expressing lines with levels of resistance to the pathogens positively correlated with the level of siRNA accumulation detected by Northern blotting. Transgenic cassava plants of cultivar 60444 generated with the same RNAi constructs showed strong resistance to UCBSV when bud graft challenged under greenhouse conditions (Yadav et al., 2011). The seven best performing transgenic cassava plant lines derived from the FL-CP and NT-CP RNAi constructs were evaluated under CFT conditions at NaCRRI across a full growing season in 2009–2010. Similar to greenhouse results, all transgenic lines derived from FL-CP showed significant resistance to CBSD, while 90% of the non-transgenic control plants became heavily infected, and developed severe storage root necrosis. One line expressing the highest levels of CP-derived siRNAs showed very high resistance to CBSD and remained free of foliar and root necrotic symptoms across the 12-month cropping cycle (Ogwok et al., 2012). Performance of these resistant transgenic cassava lines was evaluated for durability of the RNAi-derived resistance in a subsequent vegetative generation by propagation of stem cuttings and establishment of a follow-up planting. Resistance to infection by CBSV and UCBSV was maintained, providing strong evidence for the potential of RNAi technology to generate stable, robust field-level resistance to CBSD (Odipio et al., 2014). Development of CBSD-resistant cassava for delivery to farmers in East Africa Knowledge gained from the field studies described above is being used to further develop a CBSD resistant cassava product with the intention to submit an application for commercial release to regulatory authorities in Uganda and Kenya. A pipeline was established within the VIRCA Plus project for high-throughput production and screening of transgenic plants that have low (1–2) copy T-DNA integrations, carry no plasmid vector backbone sequences, and that express CP-derived siRNAs at levels required to generate resistance to CBSD-causing viruses. A new RNAi construct was constructed

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Figure 8.3  Confined field trials of cassava cultivar TME 204 enhanced by RNAi for resistance to CBSD.

consisting of the near full-length CPs from both CBSV and UCBSV fused together to form an inverted repeat (designated p5001) (Beyene et al., 2016a). Expressing siRNAs from both CPs was considered the best way in which to generate robust and durable resistance to both causal pathogens. The enhanced construct was transformed into the Ugandan farmer-preferred, CMD-resistant cultivar TME 204 (Beyene et al., 2016a; Chauhan et al., 2015). Approximately 350 independent transgenic plant lines were required to identify 25 events that met the low T-DNA copy number and siRNA expression levels needed for advancement to field testing in Uganda and Kenya (Fig. 8.3). Prior to field testing, a subset of regenerated plants was assessed in the greenhouse by graft inoculation with the virulent CBSV strain Naliendele. Again, those lines

showing high levels of CP-derived siRNA accumulation demonstrated robust resistance to CBSD and remained asymptomatic in their leaves and storage roots (Beyene et al., 2016a). Twenty-five independent transgenic plant lines were established in CFTs in randomized replicated plots. After 12 months of exposure to whitefly-transmitted disease pressure at Namulonge, Uganda, foliar symptoms were observed on all non-transgenic TME 204 plants while sixteen out of twenty-five transgenic lines remained symptom-free. In all cases, the asymptomatic lines were also those accumulating high siRNA. Approximately 10,000 storage roots were assessed for presence of necrosis caused by CBSD. Between 80% and 100% of the non-transgenic TME 204 plants displayed CBSD within their storage roots,

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(A)

(B)

Figure 8.4  Cassava storage roots of cultivar TME 204 harvested after 12 months growth in a confined field trial with high CBSD pressure. (A) Slices through roots of non-transgenic plants showing severe brown necrotic CBSD symptoms; (B) slices through roots of plants modified by RNAi showing no CBSD symptoms.

with 96% – 100% of all roots from these plants carrying damage that would make them inedible and unmarketable (Wagaba et al., 2016a). In contrast, in the six best performing RNAi TME 204 lines, less than 2% of the storage roots had visible CBSD. For example, in the high siRNA-expressing line designated 5001-20, only one storage root out of the 275 assessed showed necrotic tissue. The remaining 274 were free of symptoms and would have been fit for consumption or sale. This represents at least a 5-fold increase in usable yields that could potentially be available to farmers. Within the same CFT, the local CBSD tolerant cultivars NASE 14 and NASE 3 both showed 88%–89% incidence of CBSD within their storage roots, indicating the efficacy of the RNAi in addressing the disease. As cassava is a vegetatively propagated crop, resistance to virus diseases must be maintained across stake-derived cropping cycles. This presents challenges compared to most seed-propagated crops, which start each cropping cycle with virus-free propagules. To address this question, stem cuttings were taken from CBSD-free plants described above and used to establish a second CFT at the same location. Over a second 12-month cycle, the RNAi TME 204 plants from 11 of the transgenic lines continued to perform well, with only one storage root showing indications of low-level infection with CBSD. Control plants of non-transgenic TME 204 established from diseasefree stakes acquired more than 95% incidence of

necrosis within their storage roots. A CFT established at Mtwapa (near Mombasa), Kenya with the same TME 204 transgenic lines also remained free of CBSD (Fig. 8.4). Across these locations, plants that remained asymptomatic for CBSD foliar and root symptoms tested negative for presence of CBSV and UCBSV by RT-PCR diagnostics (Wagaba et al., 2016a). Data confirming resistance to CBSD are important because they demonstrate that this RNAi technology has relevance across a wide geographical area when cassava is exposed to pathogens with differing genetic diversity (Wagaba et al., 2016a). This information will be critical for determining regions into which the final product can be deployed. Having achieved acceptable levels of resistance to UCBSV and CBSV, the project has taken two different pathways towards generating CBSD-resistant cassava materials for use by farmers and breeders. These are designed to generate an array of genetic backgrounds resistant to both CMD and CBSD. In the first approach, leading transgenic CBSDresistant TME 204 events are being crossed with CMD-resistant cultivars (using traditional breeding methods) to generate progeny that will also carry functional CMD2-, CMD3- and CMD1- type resistance. In the second strategy, the CMD-resistant Ugandan farmer-preferred cultivars NASE 13 and NASE 14 are being genetically transformed with the same p5001 inverted repeat construct that was successfully used to develop transgenic CBSDresistant TME 204 events.

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The regulatory environment in Uganda and Kenya In order to deliver crops that have been improved with transgenic technologies to farmers, the appropriate regulatory laws must be in place within a given country. Both Uganda and Kenya have recently made important progress in developing these legal structures and processes. The Ugandan Parliament passed the Biotechnology and Biosafety Act of 2017. Prior to this, Uganda had passed the Biotechnology and the Biosafety Policy in 2008, following the country’s ratification of the Cartagena Protocol on Biosafety in 2002. Under the Uganda National Council for Science and Technology (UNCST), the National Biosafety Committee (NBC) has a mandate to supervise transgenic research and development activities, including CFTs. Procedures for conducting laboratory/greenhouse genetic engineering activities (containment) and open field trials (confinement) were implemented by the Uganda National Council for Science and Technology (UNCST). UNCST developed guidelines for management of recombinant gene experimentation in the laboratory, greenhouse and Standard Operating Procedures (SOPs) for conducting and monitoring CFTs. Under this framework, CFTs have been approved and implemented for banana (black sigatoka, banana bacterial wilt, vitamin-A and parasitic nematode resistance), insect-resistant and herbicide-tolerant cotton, water efficient and insect-resistant maize, nitrogen use efficient rice, virus-resistant sweet potato, herbicide tolerant soybean, late blight resistant potato and virus-resistant cassava (Kwehangana and Kasule, 2017). With the recent passing of the Biotechnology and Biosafety Act by Parliament, UNCST will have the authority to decide whether to approve genetically engineered crop varieties and allow them to be available to Ugandan farmers for open cultivation. Similarly, Kenya signed the Cartagena Protocol on Biosafety in 2000 and ratified it in 2003. The National Biotechnology Development Policy was published in 2006. The Biosafety bill was signed into law in 2009 giving rise to Biosafety Act No. 2 of 2009 that sets up the National Biosafety Authority (NBA) as the national focal point of all biosafety matters in Kenya and makes provision for the establishment of a legal framework for the safe handling, use and transfer of genetically engineered organisms. The NBA exercises this general

supervision and control with a view to ensuring safety to human and animal health and protection of the environment. To date, the NBA has approved CFT sites in Alupe and Kakamega, Western Kenya; Kilifi–Mtwapa, Eastern Kenya; Muranga–ThikaKandara, Central Kenya and has approved more than 22 trials in total, and one limited/conditional environmental release for national performance trials (NPTs) in maize. Conclusion Over the last 5–8 years, important progress has been made towards delivering virus-resistant cassava planting materials to farmers in East Africa. Effective RNAi-mediated resistance to CMD has been demonstrated in the lab and field. However, deploying such resistance that would be sufficiently robust and durable for cultivation by farmers is challenging in East Africa. Presence of multiple strains of ACMV and EACMV increases the potential for recombinations that could overcome pathogenderived, sequence-specific resistance strategies. In addition, increasing deployment of CMD-resistant cultivars carrying inherent CMD1-, CMD2- and CMD3-type resistance mechanisms has been effective for addressing CMD in this region. The opposite is true for CBSD, which significantly suppresses yields in East and Central Africa and continues to spread westwards, placing cassava production in the populous countries of West Africa at great risk. RNAi control of CBSD is effective against both causal viruses and has been shown to be capable of increasing usable yields by up to 100-fold in a susceptible cultivar. Resistance is maintained across the vegetative cropping cycle and is effective across different geographical locations, from the Lake Victoria region to southern coastal Kenya (Beyene et al., 2016a). Similar CP-derived transgenic approaches have proven successful against RNA viruses in crops such as papaya, squash and plum, with consistent field-level performance maintained across many seasons. Continued monitoring of resistance is required, along with assessment of virus evolution to ensure that the expressed CPsiRNAs remain capable of imparting required levels of resistance. Technical progress therefore indicates the significant potential for RNAi technology to bring solutions to CBSD. Delivery of CBSD resistant germplasm to farmers can occur directly via

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production of enhanced farmer-preferred cultivars. Improved tissue culture and genetic modification systems in East and West African cassava cultivars are making this a reality (Chauhan et al., 2015; Chauhan and Taylor, 2017). Alternatively, or in addition, CBSD resistant RNAi lines can be incorporated into conventional breeding programmes. Breeders can then generate a range of varieties suited for future deployment to farmers in different regions of sub-Saharan Africa which are impacted from, or threatened by, CBSD. Before farmers or breeders can benefit from such CBSD resistant planting materials, approval must be obtained from the relevant regulatory authorities. In Kenya and Uganda the developing regulatory capacity is progressing in parallel with the technical progress occurring in cassava. Removal of strict liability for field trials in Tanzania also shows promise for testing and eventual deployment of CBSD resistant planting materials in that country. Future trends and prospects Prospects for improving resistance to cassava viruses bring cause for optimism. RNAi technology offers potential near-term solutions to address CBSD. Advances in cassava genomics present new possibilities for identifying genes and the molecular mechanisms underlying virus resistance mechanisms within cultivated cassava varieties and related wild species. These could be transferred to cultivars preferred by farmers in regions impacted by CMD and CBSD (Bredeson et al., 2016). Recent reports (Beyene et al., 2016b) describing loss of resistance to CMD when CMD2-type cultivars are passed through somatic embryogenesis offer unique, previously unavailable germplasm for investigating the CMD2 function, and then by comparison, also CMD3 and CMD1 resistance mechanisms. Knowledge of the molecular mechanisms underlying inherent CMD resistance could then be exploited to modify CMD susceptible cultivars, and/or stack CMD2 and CMD1 resistance within the same germplasm. Enhancing CMD resistance in this manner could be achieved through breeding, transgenic approaches or by gene editing. Recent reports of capacity for gene editing (Odipio et al., 2017) and allele exchange in cassava (Hummel et al., 2017) open new possibilities for enhancing cassava germplasm and exploiting the increasing levels

genomic information becoming available. CRISPRCas-mediated gene editing is being applied to develop cassava with enhanced resistance to CBSD. Mutation of host eukaryotic translation initiation factor 4E (eIF4E) isoforms using CRISPR-Cas technology has been shown to generate cassava plants with reduced CBSD symptoms and suppressed virus titre (Gomez et al., 2017). This technology requires further development to generate fully resistant plants with potential for deployment to farmers. It does, however, offer an important additional strategy for control of CBSD, and could be stacked with RNAi to increase resistance and durability of deployed planting materials. Targeting the whitefly vector for CBSD and CMD is also an attractive strategy, either singly or stacked with virus resistance technologies. Recent reports of whitefly vector control in cotton hold promise for such approaches (Shukla et al., 2016). Successful application of these technologies to benefit cassava farmers will depend on effective targeting of resources and investment in capacities in regions such as East Africa. For gene editing in particular there is also urgency and importance in developing the required regulatory structures to ensure that cassava farmers benefit from what promises to be the next revolution in crop agriculture. References Ademiluyi, F.T., and Mepba, H.D. (2013). Yield and properties of ethanol biofuel produced from different whole cassava flours. ISRN Biotechnol 2013, 916481. https://doi.org/10.5402/2013/916481 Akano, O., Dixon, O., Mba, C., Barrera, E., and Fregene, M. (2002). Genetic mapping of a dominant gene conferring resistance to cassava mosaic disease. Theor. Appl. Genet. 105, 521–525. https://doi.org/10.1007/s00122-0020891-7 Alicai, T., Ndunguru, J., Sseruwagi, P., Tairo, F., OkaoOkuja, G., Nanvubya, R., Kiiza, L., Kubatko, L., Kehoe, M.A., and Boykin, L.M. (2016). Characterization by Next Generation Sequencing Reveals the Molecular Mechanisms Driving the Faster Evolutionary rate of Cassava brown streak virus Compared with Ugandan cassava brown streak virus. bioRxiv, doi: https://doi. org/10.1101/053546. Anyanwu, C.N., Ibeto, C.N., Ezeoha, S.L., and Ogbuagu, N.J. (2015). Sustainability of cassava (Manihot esculenta Crantz) as industrial feedstock, energy and food crop in Nigeria. Renew. Energy 81, 745-752. https://doi. org/710.1016/j.renene.2015.1003.1075. Aripin, A.M., Kassim, A.S.M., Daud, Z., and Hatta, M.Z.M. (2013). Cassava peels for alternative fibre in pulp and paper industry: chemical properties and morphology characterization. Int. J. Integr. Eng. 5.

Transgenic Virus-Resistant Cassava in East Africa |  173

Balagopalan, C.C.U.i.F., Feed and Industry. In Cassava: Biology, Production and Utilization, (Wallingford: CABI International), pp. 301–318. Eds R.J. Hillocks, J.M. Thresh and A.C. Bellotti (2002). Cassava Utilization in Food, Feed and Industry. In Cassava: Biology, Production and Utilization, R.J. Hillocks, J.M. Thresh, and A.C. Bellotti, eds. (Wallingford: CABI International), pp. 301–318. Beyene, G., Chauhan, R.D., Ilyas, M., Wagaba, H., Fauquet, C.M., Miano, D., Alicai, T., and Taylor, N.J. (2016a). A Virus-Derived Stacked RNAi Construct Confers Robust Resistance to Cassava Brown Streak Disease. Front. Plant Sci. 7, 2052. https://doi.org/10.3389/fpls.2016.02052 Beyene, G., Chauhan, R.D., Wagaba, H., Moll, T., Alicai, T., Miano, D., Carrington, J.C., and Taylor, N.J. (2016b). Loss of CMD2-mediated resistance to cassava mosaic disease in plants regenerated through somatic embryogenesis. Mol. Plant Pathol. 17, 1095–1110. https://doi.org/10.1111/mpp.12353 Bock, K.R. (1994). Studies on cassava brown streak virus disease in Kenya. Trop. Sci. 34, 134–145. Böttcher, B., Unseld, S., Ceulemans, H., Russell, R.B., and Jeske, H. (2004). Geminate structures of African Cassava mosaic virus. J. Virol. 78, 6758–6765. https:// doi.org/10.1128/JVI.78.13.6758-6765.2004 Bredeson, J.V., Lyons, J.B., Prochnik, S.E., Wu, G.A., Ha, C.M., Edsinger-Gonzales, E., Grimwood, J., Schmutz, J., Rabbi, I.Y., Egesi, C., et al. (2016). Sequencing wild and cultivated cassava and related species reveals extensive interspecific hybridization and genetic diversity. Nat. Biotechnol. 34, 562–570. https://doi.org/10.1038/ nbt.3535 Brown, J.K., Zerbini, F.M., Navas-Castillo, J., Moriones, E., Ramos-Sobrinho, R., Silva, J.C., Fiallo-Olivé, E., Briddon, R.W., Hernández-Zepeda, C., Idris, A., et al. (2015). Revision of Begomovirus taxonomy based on pairwise sequence comparisons. Arch. Virol. 160, 1593– 1619. https://doi.org/10.1007/s00705-015-2398-y Bull, S.E., Alder, A., Barsan, C., Kohler, M., Hennig, L., Gruissem, W., and Vanderschuren, H. (2017). FLOWERING LOCUS T Triggers Early and Fertile Flowering in Glasshouse Cassava (Manihot esculenta Crantz). Plants 6, E22. https://doi.org/10.3390/ plants6020022 Bull, S.E., Owiti, J.A., Niklaus, M., Beeching, J.R., Gruissem, W., and Vanderschuren, H. (2009). Agrobacteriummediated transformation of friable embryogenic calli and regeneration of transgenic cassava. Nat Protoc 4, 1845–1854. https://doi.org/10.1038/nprot.2009.208 Chant, S.R. (1958). Studies on the Transmission of Cassava mosaic virus By Bemisia Spp. (Aleyrodidae). Ann. Appl. Biol. 46, 210–215. DOI: 210.1111/j.1744-7348.1958. tb02198.x. Chauhan, R.D., Beyene, G., Kalyaeva, M., Fauquet, C.M., and Taylor, N. (2015). Improvements in Agrobacteriummediated transformation of cassava (Manihot esculenta Crantz) for large-scale production of transgenic plants. PCTOC 121, 591–603. https://doi.org/510.1007/ s11240-11015-10729-z. Chauhan, R.D., Beyene, G., and Taylor, N. (2017). Multiple morphogenic culture systems cause loss of resistance to cassava mosaic disease. Front. Plant Sci. Submitted.

Chauhan, R.D., and Taylor, N.J. (2017). Meta-topolin stimulates de novo shoot organogenesis and plant regeneration in cassava. PCTOC, https://doi. org/10.1007/s11240-11017-11315-11243. Chavarriaga-Aguirre, P., Brand, A., Medina, A., Prías, M., Escobar, R., Martinez, J., Díaz, P., López, C., Roca, W.M., and Tohme, J. (2016). The potential of using biotechnology to improve cassava: a review. In Vitro Cell. Dev. Biol., Plant 52, 461–478. https://doi.org/10.1007/ s11627-016-9776-3 Chellappan, P., Masona, M.V., Vanitharani, R., Taylor, N.J., and Fauquet, C.M. (2004). Broad spectrum resistance to ssDNA viruses associated with transgene-induced gene silencing in cassava. Plant Mol. Biol. 56, 601–611. https://doi.org/10.1007/s11103-004-0147-9 Curran, S.R., and Cooke, A.M. (2008). Unexpected outcomes of Thai cassava trade: a case of global complexity and local unsustainability. Globalizations 5, 111–127. https://doi.org/10.1080/14747730802057449 Dixon, A.G.O., Ogbe, F.O., and Okechukwu, R.U. (2010). Cassava mosaic disease in Sub-Saharan Africa: a feasible solution for an unsolved problem. Outlook Agric. 39, 89–94. Falck-Zepeda, J., Yorobe, J., Jr., Husin, B.A., Manalo, A., Lokollo, E., Ramon, G., Zambrano, P., and Sutrisno (2012). Estimates and implications of the costs of compliance with biosafety regulations in developing countries. GM Crops Food 3, 52-59. DOI 10.4161/ gmcr.18727. FAO (2015). FAOSTAT Database. Rome, Italy. http:// faostat3.fao.org/home/E. Fauquet, C., and Fargette, D. (1990). African Cassava mosaic virus: etiology, epidemiology and control. Plant Dis. 74, 404–411. Fregene, M., and Puonti-Kaerlas, J. (2002). Cassava biotechnology. In Cassava: Biology, production and utilization, R.J. Hillocks, J.M. Thresh, and A.C. Bellotti, eds. (CABI Publishing), pp. 179-207. Fregene, M.A., Suarez, M., Mkumbira, J., Kulembeka, H., Ndedya, E., Kulaya, A., Mitchel, S., Gullberg, U., Rosling, H., Dixon, A.G., et al. (2003). Simple sequence repeat marker diversity in cassava landraces: genetic diversity and differentiation in an asexually propagated crop. Theor. Appl. Genet. 107, 1083–1093. https://doi. org/10.1007/s00122-003-1348-3 Frischmuth, T., Engel, M., Lauster, S., and Jeske, H. (1997). Nucleotide sequence evidence for the occurrence of three distinct whitefly-transmitted, Sida-infecting bipartite geminiviruses in Central America. J. Gen. Virol. 78, 2675–2682. https://doi.org/10.1099/0022-131778-10-2675 Frischmuth, T., and Stanley, J. (1998). Recombination between viral DNA and the transgenic coat protein gene of African cassava mosaic geminivirus. J. Gen. Virol. 79, 1265–1271. https://doi.org/10.1099/0022-1317-795-1265 Gilbertson, R.L., Batuman, O., Webster, C.G., and Adkins, S. (2015). Role of the insect supervectors Bemisia tabaci and Frankliniella occidentalis in the Emergence and Global Spread of Plant Viruses. Annu Rev Virol 2, 67–93. https://doi.org/10.1146/annurevvirology-031413-085410

174  | Wagaba et al.

Gomez, M.A., Lin, Z.-J.D., Moll, T., Luebbert, C., Chauhan, R.D., Vijayaraghavan, A., Renninger, K., Beyene, G., Taylor, N.J., Carrington, J.C., et al. (2017). Simultaneous CRISPR/Cas9-mediated editing of cassava eIF4E isoforms nCBP-1 and nCBP-2 confers elevated resistance to cassava brown streak disease. bioRxiv, doi: https:// doi.org/10.1101/209874. Hahn, S.K., Terry, E.R., and Leuschner, K. (1980). Breeding cassava for resistance to cassava mosaic disease. Euphytica 29, 673–683. DOI https://doi. org/610.1007/BF00023215. Hanley-Bowdoin, L., Bejarano, E.R., Robertson, D., and Mansoor, S. (2013). Geminiviruses: masters at redirecting and reprogramming plant processes. Nat. Rev. Microbiol. 11, 777–788. https://doi.org/10.1038/ nrmicro3117 Harrison, B.D., Barker, H., Bock, K.R., Guthrie, E.J., Meredith, G., and Atkinson, M. (1977). Plant viruses with circular single-stranded DNA. Nature 270, 760– 762. https://doi.org/710.1038/270760a270760 Hillocks, R.J., and Maruthi, M.N. (2015). Post-harvest impact of cassava brown streak disease in four countries in eastern Africa. Food Chain 5, 116–122. https://doi. org/110.3362/2046-1887.2015.3008. Hong, Y., and Stanley, J. (1995). Regulation of African cassava mosaic virus complementary-sense gene expression by N-terminal sequences of the replicationassociated protein AC1. J. Gen. Virol. 76, 2415–2422. Horowitz, A.R., Antignus, Y., and D., G. (2011). Management of Bemisia tabaci whiteflies. In The Whitefly, Bemisia tabaci (Homoptera: Aleyrodidae) Interaction with Geminivirus-Infected Host Plants, W.M.O. Thompson, ed. (Dordrecht: Springer), pp. 293–322. Hummel, A.W., Chauhan, R.D., Cermak, T., Mutka, A.M., Vijayaraghavan, A., Boyher, A., Starker, C.G., Bart, R., Voytas, D.F., and Taylor, N.J. (2017). Allele exchange at the EPSPS locus confers glyphosate tolerance in cassava. Plant Biotechnol. J. [Epub ahead of print]. https://doi. org/10.1111/pbi.12868 IITA (2014). Cassava varieties tolerant to deadly viral diseases shared across five countries. IITA Bulletin 2215, 1. Jarvis, A., Ramirez-Villegas, J., Herrera Campo, B.V., and Navarro-Racines, C. (2012). Is Cassava the answer to African climate change adaptation? Trop. Plant Biol. 5, 9–29. DOI https://doi.org/10.1007/s12042-1201219096-12047. Jeremiah, S.C., Ndyetabula, I.L., Mkamilo, G.S., Haji, S., Muhanna, M.M., Chuwa, C., Kasele, S., Bouwmeester, H., Ijumba, J.N., and Legg, J.P. (2015). The dynamics and environmental influence on interactions between cassava brown streak disease and the whitefly, Bemisia tabaci. Phytopathology 105, 646–655. https://doi. org/10.1094/PHYTO-05-14-0146-R Karthikeyan, C., Patil, B.L., Borah, B.K., Resmi, T.R., Turco, S., Pooggin, M.M., Hohn, T., and Veluthambi, K. (2016). Emergence of a latent Indian Cassava mosaic virus from cassava which recovered from infection by a non-persistent Sri Lankan Cassava mosaic virus. Viruses 8, E264. Kwehangana, M., and Kasule, D. (2017). UNCST Holds the Biosafety Forum 2017. Uganda National Council

for Science and Technology. https://uncst.go.ug/uncstholds-the-biosafety-forum-2017/. Legg, J.P., Attiogbevi, E., Barker, S., Beach, L., Ceballos, H., Cuellar, W., Elkhoury, W., Gerling, D., Helsen, J., Hershey, C., et al. (2014a). A global alliance declaring war on cassava viruses in Africa. Food Sec. 6, 231. DOI https://doi.org/210.1007/s12571-12014-10340-x. Legg, J.P., Owor, B., Sseruwagi, P., and Ndunguru, J. (2006). Cassava mosaic virus disease in East and Central Africa: epidemiology and management of a regional pandemic. Adv. Virus Res. 67, 355–418. Legg, J.P., Shirima, R., Tajebe, L.S., Guastella, D., Boniface, S., Jeremiah, S., Nsami, E., Chikoti, P., and Rapisarda, C. (2014b). Biology and management of Bemisia whitefly vectors of cassava virus pandemics in Africa. Pest Manag. Sci. 70, 1446–1453. https://doi.org/10.1002/ps.3793 Lister, R.M. (1959). Mechanical transmission of cassava brown streak virus. Nature 183, 1588–1589. Manyong, V.M., Maeda, C., Kanju, E. and Legg, J.P. (2012) Economic damage of cassava brown streak disease in sub-Saharan Africa. In Tropical Root and Tuber Crops and the Challenges of Globalization and Climate Change, 11th ISTRC-AB Symposium, 4–8 2010, Kinshasa, Democratic Republic of Congo, Okechukwu, R.U., and Ntawuruhunga, P., eds. (International Institute of Tropical Agriculture, Ibadan), pp. 61–68. Marcon, M.J.A., Vieira, G.C.N., De Simas, K.N., Santos, K., Vieira, M.A., De Mello Castanho Amboni, R.D., and Amante, E.R. (2007). Effect of the improved fermentation on physicochemical properties and sensorial acceptability of sour cassava starch. Brazilian Arch. Biol. Technol. 50, 1073–1081. Maruthi, M.N., Hillocks, R.J., Mtunda, K., Raya, M.D., Muhanna, M., Kiozia, H., Rekha, A.R., Colvin, J., and Thresh, J.M. (2005). Transmission of Cassava brown streak virus by Bemisia tabaci (Gennadius). J. Phytopathol. 153, 307–312. doi:310.1111/j.14390434.2005.00974.x. Maruthi, M.N., Jeremiah, S.C., Mohammed, I.U., and Legg, J.P. (2017). The role of the whitefly, Bemisia tabaci (Gennadius), and farmer practices in the spread of cassava brown streak ipomoviruses. J Phytopathol 165, 707–717. https://doi.org/10.1111/jph.12609 Mbanzibwa, D.R., Tian, Y., Mukasa, S.B., and Valkonen, J.P. (2009b). Cassava brown streak virus (Potyviridae) encodes a putative Maf/HAM1 pyrophosphatase implicated in reduction of mutations and a P1 proteinase that suppresses RNA silencing but contains no HC-Pro. J. Virol. 83, 6934–6940. https://doi.org/10.1128/ JVI.00537-09 Mbanzibwa, D.R., Tian, Y.P., Tugume, A.K., Mukasa, S.B., Tairo, F., Kyamanywa, S., Kullaya, A., and Valkonen, J.P. (2009a). Genetically distinct strains of Cassava brown streak virus in the Lake Victoria basin and the Indian Ocean coastal area of East Africa. Arch. Virol. 154, 353–359. https://doi.org/10.1007/s00705-008-0301-9 Mbanzibwa, D.R., Tian, Y.P., Tugume, A.K., Mukasa, S.B., Tairo, F., Kyamanywa, S., Kullaya, A., and Valkonen, J.P. (2011a). Simultaneous virus-specific detection of the two cassava brown streak-associated viruses by RT-PCR reveals wide distribution in East Africa, mixed infections, and infections in Manihot glaziovii. J. Virol.

Transgenic Virus-Resistant Cassava in East Africa |  175

Methods 171, 394–400. https://doi.org/10.1016/j. jviromet.2010.09.024 Mbanzibwa, D.R., Tian, Y.P., Tugume, A.K., Patil, B.L., Yadav, J.S., Bagewadi, B., Abarshi, M.M., Alicai, T., Changadeya, W., Mkumbira, J., et al. (2011b). Evolution of cassava brown streak disease-associated viruses. J Gen Virol 92, 974-987. DOI 910.1099/vir.1090.026922026920. Mohammed, I.U., Abarshi, M.M., Muli, B., Hillocks, R.J., and Maruthi, M.N. (2012). The symptom and genetic diversity of cassava brown streak viruses infecting cassava in East Africa. Adv Virol 2012, 795697. https:// doi.org/10.1155/2012/795697 Mohammed, I.U., Ghosh, S., and Maruthi, M.N. (2016). Host and virus effects on reversion in cassava affected by cassava brown streak disease. Plant Pathol. 65, 593-600. Monger, W.A., Seal, S., Cotton, S., and Foster, G.D. (2001). Identification of different isolates of Cassava brown streak virus and development of a diagnostic test. Plant Pathol. 50, 768–775. Morgan, N.K., and Choct, M. (2016). Cassava: Nutrient composition and nutritive value in poultry diets. Anim. Nutr. 2, 253–261. Narayanan, N., Beyene, G., Chauhan, R.D., GaitánSolis, E., Grusak, M.A., Taylor, N., and Anderson, P. (2015). Overexpression of Arabidopsis VIT1 increases accumulation of iron in cassava roots and stems. Plant Sci. 240, 170–181. https://doi.org/10.1016/j. plantsci.2015.09.007 Ndunguru, J., De León, L., Doyle, C.D., Sseruwagi, P., Plata, G., Legg, J.P., Thompson, G., Tohme, J., Aveling, T., Ascencio-Ibáñez, J.T., et al. (2016). Two Novel DNAs That Enhance Symptoms and Overcome CMD2 Resistance to Cassava Mosaic Disease. J. Virol. 90, 4160–4173. https://doi.org/10.1128/JVI.02834-15 Ndunguru, J., Sseruwagi, P., Tairo, F., Stomeo, F., Maina, S., Djikeng, A., Kehoe, M., and Boykin, L.M. (2015). Analyses of Twelve New Whole Genome Sequences of Cassava Brown Streak Viruses and Ugandan Cassava Brown Streak Viruses from East Africa: Diversity, Supercomputing and Evidence for Further Speciation. PLOS ONE 10, e0139321. https://doi.org/10.1371/ journal.pone.0139321 Ndyetabula, I.L., Merumba, S.M., Jeremiah, S.C., Kasele, S., Mkamilo, G.S., Kagimbo, F.M., and Legg, J.P. (2016). Analysis of interactions between Cassava Brown Streak Disease symptom types facilitates the determination of varietal responses and yield losses. Plant Dis. 100, 1388– 1396. https://doi.org/10.1094/PDIS-11-15-1274-RE Nichols, R.F.W. (1950). The brown streak disease of cassava: distribution, climatic effects and diagnostic symptoms. East African Agric. J. 15, 154–160. Ntawuruhunga, P., Dixon, A., Kanju, E., Ssemakula, G., Okechukwu, R., Whyte, J., Obiero, H., Bigirimana, S., Gashaka, G., and Lukombo, S. (2013). Successful innovations and lessons learnt in cassava improvement and deployment by IITA in Eastern African Region. African J. Root Tuber Crops 10, 41-54. Ntui, V.O., Kong, K., Khan, R.S., Igawa, T., Janavi, G.J., Rabindran, R., Nakamura, I., and Mii, M. (2015). Resistance to Sri Lankan Cassava mosaic virus (SLCMV) in genetically engineered cassava cv. KU50 through

RNA silencing. PLOS ONE 10, e0120551. https://doi. org/10.1371/journal.pone.0120551 Nuwamanya, E., Baguma, Y., Atwijukire, E., Acheng, S., Abidrabo, P., Omongo, C.A., and Alicai, T. (2017). Cassava brown streak disease effects on leaf metabolites and pigment accumulation. African Crop Sci. J. 25, 33–45. Nuwamanya, E., Chiwona-Karltun, L., Kawuki, R.S., and Baguma, Y. (2012). Bio-ethanol production from nonfood parts of cassava (Manihot esculenta Crantz). Ambio 41, 262–270. https://doi.org/10.1007/s13280-0110183-z Nweke, F.I. (1996). Cassava: A Cash Crop in Africa. In Issue 14 of COSCA working paper (IITA). Odipio, J., Alicai, T., Ingelbrecht, I., Nusinow, D.A., Bart, R., and Taylor, N.J. (2017). Efficient CRISPR/Cas9 Genome Editing of Phytoene desaturase in Cassava. Front. Plant Sci. 8, 1780. https://doi.org/10.3389/ fpls.2017.01780 Odipio, J., Ogwok, E., Taylor, N.J., Halsey, M., Bua, A., Fauquet, C.M., and Alicai, T. (2014). RNAi-derived field resistance to Cassava brown streak disease persists across the vegetative cropping cycle. GM Crops Food 5, 16–19. https://doi.org/10.4161/gmcr.26408 Ogwok, E., Ilyas, M., Alicai, T., Rey, M.E., and Taylor, N.J. (2016). Comparative analysis of virus-derived small RNAs within cassava (Manihot esculenta Crantz) infected with cassava brown streak viruses. Virus Res. 215, 1–11. https://doi.org/10.1016/j.virusres.2016.01.015 Ogwok, E., Odipio, J., Halsey, M., Gaitán-Solís, E., Bua, A., Taylor, N.J., Fauquet, C.M., and Alicai, T. (2012). Transgenic RNA interference (RNAi)-derived field resistance to cassava brown streak disease. Mol. Plant Pathol. 13, 1019–1031. https://doi.org/10.1111/ j.1364-3703.2012.00812.x Okogbenin, E., Egesi, C.N., Olasanmi, B., Ogundapo, O., Kahya, S., Hurtado, P., Marin, J., Akinbo, O., Mba, C., Gomez, H., et al. (2012). Molecular marker analysis and validation of resistance to cassava mosaic disease in elite cassava genotypes in Nigeria. Crop Sci. 52, 2576–2586. Owor, B., Legg, J.P., Okao-Okuja, G., Obonyo, R., and Ogenga-Latigo, M.W. (2004). The effect of cassava mosaic geminiviruses on symptom severity, growth and root yield of a Cassava mosaic virus disease-susceptible cultivar in Uganda. Ann. Appl. Biol. 145, 331–337. Patil, B.L., Bagewadi, B., Yadav, J.S., and Fauquet, C.M. (2016). Mapping and identification of cassava mosaic geminivirus DNA-A and DNA-B genome sequences for efficient siRNA expression and RNAi based virus resistance by transient agro-infiltration studies. Virus Res. 213, 109–115. Patil, B.L., Legg, J.P., Kanju, E., and Fauquet, C.M. (2015). Cassava brown streak disease: a threat to food security in Africa. J Gen Virol 96, 956-968. Patil, B.L., Ogwok, E., Wagaba, H., Mohammed, I.U., Yadav, J.S., Bagewadi, B., Taylor, N.J., Kreuze, J.F., Maruthi, M.N., Alicai, T., et al. (2011). RNAi-mediated resistance to diverse isolates belonging to two virus species involved in Cassava brown streak disease. Mol. Plant Pathol. 12, 31–41. https://doi.org/10.1111/j.13643703.2010.00650.x Pita, J.S., Fondong, V.N., Sangaré, A., Otim-Nape, G.W., Ogwal, S., and Fauquet, C.M. (2001). Recombination,

176  | Wagaba et al.

pseudorecombination and synergism of geminiviruses are determinant keys to the epidemic of severe cassava mosaic disease in Uganda. J. Gen. Virol. 82, 655–665. https://doi.org/10.1099/0022-1317-82-3-655 Rabbi, I.Y., Hamblin, M.T., Kumar, P.L., Gedil, M.A., Ikpan, A.S., Jannink, J.L., and Kulakow, P.A. (2014). High-resolution mapping of resistance to cassava mosaic geminiviruses in cassava using genotypingby-sequencing and its implications for breeding. Virus Res. 186, 87–96. https://doi.org/10.1016/j. virusres.2013.12.028 Rey, C., and Vanderschuren, H. (2017). Cassava Mosaic and Brown Streak Diseases: Current Perspectives and Beyond. Annu Rev Virol 4, 429–452. https://doi. org/10.1146/annurev-virology-101416-041913 Rey, M.E., Ndunguru, J., Berrie, L.C., Paximadis, M., Berry, S., Cossa, N., Nuaila, V.N., Mabasa, K.G., Abraham, N., Rybicki, E.P., et al. (2012). Diversity of dicotyledenousinfecting geminiviruses and their associated DNA molecules in southern Africa, including the South-west Indian ocean islands. Viruses 4, 1753–1791. https://doi. org/10.3390/v4091753 Saunders, K., Salim, N., Mali, V.R., Malathi, V.G., Briddon, R., Markham, P.G., and Stanley, J. (2002). Characterisation of Sri Lankan Cassava mosaic virus and Indian Cassava mosaic virus: evidence for acquisition of a DNA B component by a monopartite begomovirus. Virology 293, 63–74. https://doi.org/10.1006/viro.2001.1251 Shukla, A.K., Upadhyay, S.K., Mishra, M., Saurabh, S., Singh, R., Singh, H., Thakur, N., Rai, P., Pandey, P., Hans, A.L., et al. (2016). Expression of an insecticidal fern protein in cotton protects against whitefly. Nat Biotechnol 34, 1046-1051. Srinivas, T. (2007). Industrial demand for cassava starch in India. Starch/Staerke 59, 477–481. Storey, H.H. (1936). Virus diseases of East African plants. VI-A progress report on studies of the disease of cassava. East African Agric. J. 2, 34–39. Taylor, N.J., Halsey, M., Gaitán-Solís, E., Anderson, P., Gichuki, S., Miano, D., Bua, A., Alicai, T., and Fauquet, C.M. (2012). The VIRCA Project: virus resistant cassava for Africa. GM Crops Food 3, 93–103. https:// doi.org/10.4161/gmcr.19144 Tiendrébéogo, F., Lefeuvre, P., Hoareau, M., Harimalala, M.A., De Bruyn, A., Villemot, J., Traoré, V.S., Konaté, G., Traoré, A.S., Barro, N., et al. (2012). Evolution of African Cassava mosaic virus by recombination between bipartite and monopartite begomoviruses. Virol. J. 9, 67. https://doi.org/10.1186/1743-422X-9-67 Tomlinson, K.R., Bailey, A.M., Alicai, T., Seal, S., and Foster, G.D. (2017). Cassava brown streak disease: historical timeline, current knowledge and future prospects. Mol. Plant Pathol. [Epub ahead of print]. https://doi. org/10.1111/mpp.12613 Tumwesigye, K.S., Oliveira, J.C., and Gallagher, M.J.S. (2016b). Integrated sustainable process design framework for cassava biobased packaging materials: Critical review of current challenges, emerging trends and prospects. Trends Food Sci. Technol. 56, 103–114. Tumwesigye, K.S., Peddapatla, R.V.G., Crean, A., Oliveira, J.C., and Sousa-Gallagher, M.J. (2016a). Integrated

process standardisation as zero-based approach to bitter cassava waste elimination and widely-applicable industrial biomaterial derivatives. Chem. Eng. Process. Process Intensif. 108, 139–150. Vanderschuren, H., Akbergenov, R., Pooggin, M.M., Hohn, T., Gruissem, W., and Zhang, P. (2007). Transgenic cassava resistance to African Cassava mosaic virus is enhanced by viral DNA-A bidirectional promoterderived siRNAs. Plant Mol. Biol. 64, 549–557. https:// doi.org/10.1007/s11103-007-9175-6 Vanderschuren, H., Alder, A., Zhang, P., and Gruissem, W. (2009). Dose-dependent RNAi-mediated geminivirus resistance in the tropical root crop cassava. Plant Mol. Biol. 70, 265–272. https://doi.org/10.1007/s11103009-9472-3 Vanderschuren, H., Moreno, I., Anjanappa, R.B., Zainuddin, I.M., and Gruissem, W. (2012). Exploiting the combination of natural and genetically engineered resistance to cassava mosaic and cassava brown streak viruses impacting cassava production in Africa. PLOS ONE 7, e45277. https://doi.org/10.1371/journal. pone.0045277 Wagaba, H., Beyene, G., Aleu, J., Odipio, J., Okao-Okuja, G., Chauhan, R.D., Munga, T., Obiero, H., Halsey, M.E., Ilyas, M., et al. (2016a). Field Level RNAi-Mediated Resistance to Cassava Brown Streak Disease across Multiple Cropping Cycles and Diverse East African Agro-Ecological Locations. Front. Plant Sci. 7, 2060. https://doi.org/10.3389/fpls.2016.02060 Wagaba, H., Beyene, G., Trembley, C., Alicai, T., Fauquet, C.M., and Taylor, N.J. (2013). Efficient transmission of cassava brown streak disease viral pathogens by chip bud grafting. BMC Res Notes 6, 516. Wagaba, H., Patil, B.L., Mukasa, S., Alicai, T., Fauquet, C.M., and Taylor, N.J. (2016b). Artificial microRNAderived resistance to Cassava brown streak disease. J. Virol. Methods 231, 38–43. https://doi.org/10.1016/j. jviromet.2016.02.004 Wang, H.L., Cui, X.Y., Wang, X.W., Liu, S.S., Zhang, Z.H., and Zhou, X.P. (2015). First Report of Sri Lankan Cassava mosaic virus Infecting Cassava in Cambodia. Plant Dis. 100, 1029. Warburg, O.H. (1894). Die kulturpflanzen usambaras (Kessinger Publishing, LLC). Were, H.K., Winter, S., and Maiss, E. (2004). Viruses infecting cassava in Kenya. Plant Dis. 88, 17–22. Winter, S., Koerbler, M., Stein, B., Pietruszka, A., Paape, M., and Butgereitt, A. (2010). Analysis of cassava brown streak viruses reveals the presence of distinct virus species causing cassava brown streak disease in East Africa. J Gen Virol 91, 1365-1372. Yadav, J.S., Ogwok, E., Wagaba, H., Patil, B.L., Bagewadi, B., Alicai, T., Gaitan-Solis, E., Taylor, N.J., and Fauquet, C.M. (2011). RNAi-mediated resistance to Cassava brown streak Uganda virus in transgenic cassava. Mol. Plant Pathol. 12, 677–687. https://doi.org/10.1111/ j.1364-3703.2010.00700.x Zhang, P., Vanderschuren, H., Fütterer, J., and Gruissem, W. (2005). Resistance to cassava mosaic disease in transgenic cassava expressing antisense RNAs targeting virus replication genes. Plant Biotechnol. J. 3, 385–397.

Viruses Infecting Rice and their Transgenic Control Gaurav Kumar, Shweta Sharma and Indranil Dasgupta*

9

Department of Plant Molecular Biology, University of Delhi, New Delhi, India. *Correspondence: [email protected] https://doi.org/10.21775/9781910190814.09

Abstract Rice is one of the world’s most important crop plants. About 16 viruses are known to infect rice worldwide. These viruses are diverse in nature and differ from each other in their molecular organization, vector transmission, symptoms produced and geographical distribution. Transgenic expression of viral coat proteins and nucleic acids has been an effective strategy for virus control in many plant species. The same methods have also been used for rice to engineer resistance to a majority of the infecting viruses. This chapter describes the genomic organization and gene functions of the viruses infecting rice and the transgenic resistance reported against them. Introduction Rice is the principal food crop, which is the main constituent for the dietary calories to a large section of the growing population in Asian countries. Undoubtedly, it is the most important food crop in the world today and will continue to be the principal food source for a large section of the human population for many years to come. Losses to rice production due to the attack of pests require the constant attention of agricultural scientists to ensure increase in rice production commensurate with the population growth. Viruses constitute a sizable section of the pests that attack the rice plant and depress its yield. Hence, protection of rice plant against viruses has been an important component of the strategy to enhance rice production

worldwide. These strategies include the generation of rice plants having transgene(s) mostly derived from the attacking viruses, using the protection strategy known as ‘pathogen-derived resistance’, which, in turn, are represented either by a transgene expressing the viral coat protein, or, more recently, triggering RNA interference (RNAi) against the attacking virus. RNAi represents a conserved defence strategy, inherent in many organisms, including plants, against the attack by viruses and is based on sequence-specific degradation of viral RNA or prevention of viral DNA expression (Ding and Voinnet, 2007). Various viruses attack the rice plant in different parts of the World. The viruses that cause disease in rice are Rice black-streaked dwarf virus, Rice bunchy stunt virus, Rice dwarf virus, Rice gall dwarf virus, Rice giallume virus, Rice grassy stunt virus, Rice hoja Blanca virus, Rice necrosis mosaic virus, Rice ragged stunt virus, Rice stripe necrosis virus, Rice stripe virus, Rice transitory yellowing virus, Rice tungro bacilliform virus, Rice tungro spherical virus and Rice yellow mottle virus, a total of fifteen (Hibino, 1996). A new virus named Southern Rice black-streaked dwarf virus, which shows distinct properties from Rice black-streaked dwarf virus is also included (Zhou et al., 2008). Except for five viruses, Rice stripe necrosis virus, Rice bunchy stunt virus, Rice giallume virus and Rice hoja Blanca virus, Southern Rice black-streaked dwarf virus, transgenic approaches to achieve resistance have been reported for all the above viruses (Table 9.1). This chapter gives a brief account of the molecular organization and gene functions

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Table 9.1  Rice viruses, their target gene/protein used as transgenes and degrees of resistance reported No.

Virus

1

Rice black-streaked dwarf virus (RBSDV)

2.

3

Target gene/ protein

Putative function

Degree of resistance

Reference

P9–-1

Major viroplasm component protein

Strong

Shimizu et al. (2011)

Southern Rice blackstreaked dwarf virus (SRBSDV)

P9-1

Major viroplasm component protein

Not yet examined

Sasaya et al. (2014)

Rice dwarf virus (RDV)

P1

RNA Polymerase

Strong

Sasaya et al. (2014)

P2

Transmission through vector

No resistance

Sasaya et al. (2014)

P3

Core capsid protein

Moderate

Sasaya et al. (2014)

Pns4

Viral movement

Strong

Sasaya et al. (2014)

P5

Capping enzyme

No resistance

Sasaya et al. (2014)

Pns6

Movement protein

Very strong

Sasaya et al. (2014)

P7

Nucleic acid binding protein

No resistance

Sasaya et al. (2014)

P8

Major outer capsid protein Very strong

Sasaya et al. (2014)

P9

Unknown function

No resistance

Sasaya et al. (2014)

Pns10

Silencing Suppressor

No resistance

Sasaya et al. (2014)

Pns11

Unknown function

Strong

Sasaya et al. (2014)

Pns12

Unknown function

Very strong

Sasaya et al. (2014)

4.

Rice gall dwarf virus (RGDV)

Pns9

Major viroplasm component protein

Very Strong

Shimizu et al. (2012)

5.

Rice grassy stunt virus (RGSV)

Pc5

Structural Protein

Strong

Shimizu et al. (2013)

Pc6

Movement protein

Strong

Shimizu et al. (2013)

6.

Rice necrosis mosaic OsHAP2E virus (RNMV)

Rice haem activator protein

Moderate

Alam et al. (2015, 2016)

7.

Rice tungro bacilliform virus (RTBV)

8.

ORF IV

Silencing suppressor

Strong

Tyagi et al. (2008)

CP

Coat protein

Moderate

Ganesan et al. (2009)

RF2a and RF2b

Host transcription factor

Strong

Dai et al. (2008)

Replication

Moderate

Huet et al. (1999)

Replicase (Antisense)

Replication

Very Strong

Huet et al. (1999)

Rice tungro spherical Replicase (Sense) virus (RTSV)

CP

Coat protein

Moderate

Sivamani et al. (1999)

9.

Rice yellow mottle virus (RYMV)

CP

Coat protein

Strong

Kouassi et al. (1997)

10.

Rice stripe virus (RSV)

CP/SP

Coat protein/structural protein

Moderate to strong

Hayakawa et al. (1992), Yan et al. (1997), Ma et al. (2011), Park et al. (2012), Shimizu et al. (2011), Zhang et al. (2012), Zhou et al. (2012)

11.

Rice ragged stunt virus (RRSV)

S5, S7, S8, S9, and S10

Structural/non-structural proteins

Moderate to very strong

Upadhyaya et al. (1996, 1998b, 2001)

12

Rice transitory yellowing virus (RTYV)

NP

Nucleocapsid protein

Moderate

Fang et al. (1996)

Note: Moderate resistance, low level of resistance with plants showing mild symptoms throughout; strong resistance, plants showing mild symptoms only at the later stages; very strong, plants attaining complete immunity with no symptoms throughout and no molecular detection of virus particle in the cell.

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of the above viruses and the transgenic resistance strategies reported in rice against them. Conventional resistance strategies, which do not include the use of transgenes have been used for some of the remaining viruses and are described briefly. Rice tungro bacilliform virus Rice tungro bacilliform virus (RTBV, genus Tungrovirus, family Caulimoviridae) is a plant pararetrovirus, which, together with Rice tungro spherical virus (RTSV) causes rice tungro disease (Hibino et al., 1978; Jones et al., 1991; Hay et al., 1991; Lockhart et al., 1990; Shen et al., 1993). Tungro disease is widespread in South and Southeast Asia, causing an average annual loss of US$1.5 billion (Rivera and Ou, 1965; Raychaudhuri et al., 1967; Herdt et al., 1991). The tungro virus complex is transmitted by green leafhopper (GLH), Nephotettix virescens (Cabauatan and Hibino, 1985). The major symptoms of tungro disease are distinct stunting of plants and yellow-orange discolouration of leaves (Rivera and Ou, 1967). RTBV particles are bacilliform in shape having dimensions of 130 × 30 nm in length and 30–35 nm in width as observed by electron microscopy. The structure of RTBV is based on T=3 icosahedral symmetry. RTBV replicates by reverse transcription, thus indicating that it is a parareterovirus. The genome of RTBV is a circular double-stranded DNA of approximately 8 kb ( Jones et al., 1991; Qu et al., 1991). Twelve complete sequences of RTBV are available in the database out of which eight are from Southeast Asia and four from India. RTBV isolates reported from India are quite distinct from their Southeast Asian counterparts due to several nucleotide insertion and deletions (indels), substitutions etc. present mostly in the intergenic region. On the basis of these differences, RTBV sequences from India belong to a separate subgroup, i.e. ‘South Asian’ type (Fan et al., 1996). Thus, the complete genomic sequences from India, i.e. pRTBV204, pRTBV203, RTBV-KK and Chinsura had smaller genome size of 7907 bp, 7934 bp, 7934 bp, 7928 bp respectively (Nath et al., 2002; Sharma et al., 2011; Banerjee et al., 2011a). RTBV genome codes for four Open Reading Frames (ORFs), the functions of most of which are not yet well defined. The ORFs encoding them

have been designated on the basis of the size of proteins they encode, namely, ORF I (P24), ORF II (P12), ORF III (P194) and ORF IV (P46) (Hay et al., 1991). As with other retroviruses and pararetroviruses the genome of RTBV is transcribed asymmetrically with all coding capacity being on one strand, the strand containing the minus-strand priming site (Hay et al., 1991; Qu et al., 1991). The first three ORFs are tightly arranged and have an overlapping start-stop signal (ATGA; Hay et al., 1991), whereas, the fourth ORF is separated by a small intergenic region and is expressed from a spliced RNA (Fütterer et al., 1994). ORF III codes for a large poly-protein, which is further cleaved to generate four proteins, i.e. a putative movement protein, a coat protein, a protease and a RT/ RNaseH or viral replicase (Hay et al., 1991). A large intergenic region separates ORF IV and ORF I and is the site of the single RTBV promoter, the polyadenylation signal, several short ORFs (sORFs, coding for 2 to 34 amino acids) and splice donor site (Qu et al., 1991). Several attempts at making tungro resistant transgenic rice have been reported. Rice varieties IR64, TN1, Taipei 309, and Kinuhikari were transformed with several antiviral constructs aimed at providing resistance to tungro disease and/or its individual causal agents. Transgenic rice lines harbouring RTBV coat protein, polymerase, protease, RNase H, ORF I, and genes designed to express antisense RTBV RNA were produced more than two decades ago. Unfortunately, none of the strategies were effective in reducing or preventing tungro infection (Azzam and Chancellor, 2002). Subsequently, several successful attempts of transgenic resistance to RTBV were reported, using same or similar strategies. Transgenic rice plants were developed capable of producing double-stranded RNA against ORF IV of an Indian isolate of RTBV. These transgenic lines showed symptom amelioration of tungro and also showed the presence of small interfering RNAs (siRNAs) specific to the target transcript, thus confirming the activation of RNAi (Tyagi et al., 2008). The RTBV resistant plants thus obtained were subsequently used for back crossing into several popular rice varieties in India, which also showed enhanced tolerance to RTD (Roy et al., 2012; Jyothsna et al., 2013; Valarmathi et al., 2016). Fig. 9.1 shows the symptoms of infection with

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Figure 9.1  Rice plants either not inoculated (UI) or 21 days after inoculation (I) with Rice tungro bacilliform virus (RTBV) and Rice tungro spherical virus, using the vector Green leafhopper. ASD16–115–3 plants (back-crossed) carry a dsRNA producing transgene against RTBV and thus show much better growth, compared to the ASD16 parental plants.

RTBV and Rice tungro spherical virus (discussed in the following section) in rice plants representing one of the back-crossed lines (ASD16–115–3), compared to the symptoms produced in the parental ASD16 plants. It is clear that the back crossed plants, bearing the transgene providing resistance to RTBV, look much healthier and grow better compared to the parental line ASD16 lacking the transgene. Transgenic rice plants expressing the RTBV CP derived from an Indian isolate were also generated which exhibited moderate levels of resistance to RTBV (Ganesan et al., 2009). Another approach that was employed for developing transgenic resistance to RTBV was based on two basic leucine zipper (b-ZIP) type rice transcription factors, RF2a and RF2b. These transcription factors were shown to interact with a cis-element and activate transcription from RTBVPhil promoter (Dai et al., 2006). Therefore, it was proposed that in order to favour transcription of the viral promoter, RTBV causes redistribution of important host transcription factors including RF2a and RF2b. This enhanced activity of the RTBV promoter may perturb the expression of genes that are important for plant growth and development and/or disease defence resulting in development of disease symptoms. Transgenic rice plants overexpressing RF2a and RF2b resulted in reduced virus accumulation and gene expression, which in turn resulted into resistance to RTBV (Dai et al., 2008).

Rice tungro spherical virus Rice tungro spherical virus (RTSV), along with RTBV, is responsible for tungro disease of rice. RTSV belongs to the family Secoviridae, which is formed as a result of amalgamation of the families Sequiviridae and Comoviridae (as well as the unassigned genera Cheravirus and Sadwavirus). The family includes the genera Comovirus, Fabavirus, Nepovirus, Sequivirus, Waikavirus, Cheravirus, Sadwavirus and Torradovirus (Sanfaçon et al., 2009). RTSV belongs to the genus Waikavirus on the basis of the presence of a poly-A tail and two short ORFs downstream of the large ORF. RTSV virions are isometric in outline with icoasahedral symmetry. They have a diameter of 30–33 nm and a buoyant density of 1551 g/ cm2 (Galvez, 1968). RTSV belongs to the order Picornavirales, which includes viruses that infect vertebrates, insects, arthropods, higher plants, fungi and algae. All picornavirales have a positive-strand RNA genome that may be monopartite or bipartite. RTSV has a monopartite genome as it contains one molecule of single-stranded positive-sense RNA of approximately 12kb (Shen et al., 1993). RTSV RNA sequence analysis revealed the presence of a large poly-protein gene encoding three coat proteins, an RNA polymerase, a proteinase and RNA-binding proteins (Shen et al., 1993, Zhang et al., 1993). The genomic RNA is 3′ polyadenylated and is speculated to have a 5’covalently linked VPg

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molecule. The basic genome organization is conserved among picornavirale members. The genome contains a typical ‘replication block’ that includes a type III helicase, the 3C-like proteinase and a type I RNA-dependent RNA polymerase [Hel-Pro-Pol] (Goldbach et al., 1987). The complete genome sequence of different isolates of RTSV from South and Southeast Asia has been reported earlier and these were found to be much more conserved unlike RTBV isolates (Shen et al., 1993; Isogai et al., 2000; Verma and Dasgupta, 2007). Among some of the first reports of transgenic resistance was the one against RTSV. Transgenic rice plants containing replicase gene in sense orientation (untranslatable) as well as replicase gene in the antisense orientation were developed. It was observed that transgenic rice lines, containing the replicase gene in sense orientation showed partial resistance to RTSV and those containing the replicase gene in antisense orientation, exhibited immunity (Huet et al., 1999). The transgenic plants could, as expected, inhibit the transmission of RTBV through GLH fed on them, because of absence of RTSV. Simultaneously, Sivamani et al. (1999) reported partial resistance to RTSV in rice plants expressing the three coat protein genes either individually or together. There was no additive effect in plants expressing more than one coat protein gene. These rice lines represented important advances in the availability of additional genes to be deployed against RTSV other than those already reported in resistant variety by conventional breeding. Transgenic rice plants containing RNA– expressing constructs based on RNAi strategy was also reported. Transmission analysis of these transgenic plants showed that these plants acted as poor sources of transmission of RTBV leading to resistance (Verma et al., 2012). Recently, Le et al. (2015) have reported creation of transgenic rice plants capable of producing small interfering RNA originating from the inverted repeat construct encoding various RTSV proteins. These transgenic plants might be resistant to RTSV, which needs to be analysed in near future. Rice yellow mottle virus Rice yellow mottle virus (RYMV), which belongs to the genus Sobemovirus and family Sobemoviridae, is a major pathogen of rice in the African continent

(Bakker, 1974; Bakker, 1975). The RYMV particles are polyhedral in shape and approx. 30 nm in diameter. RYMV genome is 4.5kb single-stranded positive-sense RNA, lacking a poly-A tail, encoding four ORFs (Yassi et al., 1994). The product of ORF1 product (P1) is involved in viral replication (Bonneau et al., 1998). The product of ORF2 is a polyprotein, which codes for viral protease, the helicase, and the RNA-dependent RNA polymerase. While the function of ORF3 product is not yet known, the ORF4 product codes for coat protein. The most common symptoms of RYMV include yellow orange discoloration of leaves, leaf mottling, stunting, reduced tillering, non-uniform flowering and sterility. RYMV is transmitted in a semi-persistent manner by chrysomelid beetles, including Sesselia pussilla, Chaetocnema pulla, and Trichispa sericea (Bakker, 1974; Bakker, 1975). RYMV is also transmitted by the longhorned grasshopper Conocephalus merumontanus. The secondary spread of infection could be by plant-plant contact. The severity of disease severity is determined by the genotype, virus strain, age of a plant at infection, and climatic factors (Bakker 1970). Additionally, dissemination by contact between healthy and infected plants, fluid from diseased plants, RYMVcontaminated hands, and chrysomelid beetles has been demonstrated (Konate et al., 1997; Abo et al., 2003a; Sarra and Peters, 2003; Traoré et al., 2006). RYMV was first reported in 1966 from Kenya (Bakker, 1970). Since then there has been several of RYMV from many rice-growing countries of East and West Africa, including Burkina Faso, Ghana, Ivory Coast, Kenya, Liberia, Mali, Niger, Nigeria, Sierra Leone and Tanzania with its severity ranging from 5% to 100% (Rossel et al., 1982; Alegbejo et al., 2006). There have been several reports of transgenic rice showing significant resistance to RYMV. Kouassi et al. (1997) developed transgenic rice ( Japonica variety Taipei 309) expressing the coat protein of RYMV under the control of a maize ubiquitin promoter (Ubi1). These transgenic lines showed accumulation of the transgene product leading to resistance to RYMV. In most of the transgenic lines that were obtained delayed symptom development was reported. Pinto et al. (1999) developed transgenic rice encoding the RNA-dependent RNA polymerase of RYMV. These transgenic lines showed resistance to RYMV strains from different

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African locations. Study of the most resistant line, using transcription analysis, indicated that the resistance is driven from an RNA-based mechanism associated with post-transcriptional gene silencing. Kouassi et al. (2006) reported the development of transgenic rice plants expressing RYMV CP gene and several of its mutants form under the control an ubiquitin promoter. Transgenic plants expressing wild-type CP (wt. CP), deleted CP (DeltaNLS. CP), untranslatable mRNA of the CP, or antisense CP sequences of the CP gene were developed. Resistance analysis of these transgenic plants revealed that lines expressing antisense CP and untranslatable CP mRNA showed a delay in virus accumulation of up to 8 days showing significantly low level of virus accumulation as compared to non-transgenic control plants. On the other hand transgenic plants expressing wtCP and deleted CP showed higher level of virus accumulation. Rice stripe virus Rice stripe virus (RSV) belongs to the genus Tenuivirus, but has not yet been assigned to a family (Toriyama et al., 1994). RSV was reported to be a serious problem for rice production in South, Southeast, and East Asian countries (Hibino, 1996). RSV mainly infects rice plants but some other hosts also reported are maize, wheat, oat, foxtail millet and several graminaceous weeds (Falk and Tsai, 1998; Lian et al., 2011). The RSV particles are circular filaments, which are approx. 510, 610, 840, or 2110 nm in length and 9 nm in width (Ramirez et al., 1995; Shimizu et al., 1996). The genome of RSV is composed of four single-stranded RNAs that encode seven proteins (Ishikawa et al., 1989). Out of the four RNAs, designated as RNA 1–4, RNA 1 is negative sense while RNAs 2–4 are ambisense in coding nature. RNA 1, which is the largest among the four RNAs, contains a single ORF in the viral-complementary sense RNA 1 (vc RNA 1). It codes for a 337-kDa protein, which is an RNAdependent RNA polymerase (RdRp) (Barbier et al., 1992; Toriyama et al., 1994). The RSV RNAs 2–4 has been shown to code for two proteins each (Takahashi et al., 1993; Zhu et al., 1991; Zhu et al., 1992). The RNA2 encodes a 22.8-kDa silencing suppressor protein P2 from the viral-sense RNA2 (vRNA2). In addition, it also encodes a 94-kDa protein PC2 from the viral complementary sense

RNA2 (vcRNA2), which is a glycoprotein (Takahashi et al., 1993). The RNA 3 of RTSV codes for a 23.9-kDa non-structural protein P3 and a 35-kDa nucleocapsid protein CP (Coat Protein). The P3 and CP are encoded from the viral sense RNA 3 (vRNA3) and viral complementary-sense RNA 3 (vcRNA3) respectively (Hayano et al., 1990; Kakutani et al., 1991). The RNA 4 of RSV has been shown to code for SP (a 20.5 kDa non-structural disease specific protein) and PC 4 (a 32 kDa movement protein, MP) from the vRNA 4 and vcRNA 4 respectively (Hayano et al., 1990; Kakutani et al., 1991). The 5′ and 3′ terminal sequences form a panhandle like structure due to sequence complementarity with each other (Takahashi et al., 1990). The symptoms induced by RSV are chlorotic stripes or mottling and necrotic streaks on leaves, and premature wilting. Chlorotic leaves are unfolded, and later droop and wilt. RSV is transmitted in rice plants by small brown planthopper (SBH) Laodelphax striatellus in a persistent manner (Falk and Tsai, 1998). RSV is transmitted transovarially, i.e. from female adults to their progeny via eggs (Hibino, 1996; Falk and Tsai, 1998). RSV is also mechanically transmissible, but with difficulty. RSV has been reported mostly from East Asian countries such as China, Japan, Taiwan and Korea leading to significant economic losses (Abo and Sy, 1997). Earlier RSV was reported mainly from Southern areas of Korea but recently it has also been reported over a large range of altitudes, suggesting that the virus is rapidly spreading (Lee et al., 2008). There had been several reports of transgenic plants showing considerable resistance to RSV. The first report was in 1992 by Hayakawa and co-workers who transformed two japonica varieties of rice (Kinuhikari and Nipponbare) with the coat protein (CP) gene of RSV. The transgenic plants were found to be expressing high levels of transgene. Out of the five transgenic plants, which were screened for viral resistance, three were found to be symptomless indicating a significant level of resistance to RSV. Yan et al. (1997) developed transgenic rice plants expressing CP gene of a Chinese isolate of RSV. These transgenic lines also showed comparable level of resistance as reported earlier (Yan et al., 1997). Subsequently, in last few years several transgenic rice plants expressing CP gene of RSV have been reported (Ma et al., 2011;

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Park et al., 2012; Shimizu et al., 2011; Zhang et al., 2012; Zhou et al., 2012). In addition, RNAi based approaches have also been demonstrated successfully against RSV. Transgenic plants developed using double-stranded (ds) RNA-mediated RNAi system carrying full length CP gene showed resistance to RSV (Park et al., 2012). In another study transgenic rice plants were obtained using RNAi vectors carrying CP, SP or chimeric CP/SP gene sequences (Ma et al., 2011). Resistance analysis of these transgenic plants revealed that chimeric CP/SP RNAi containing plants showed stronger resistance as compared to CP or SP RNAi containing plants. Zhou et al. (2012) generated transgenic plants using japonica rice varieties Suyunuo and Guanglingxiangjing with RNAi construct containing CP and SP sequences. These plants showed very strong suppression of CP and MP transcripts with no considerable developmental defects (Zhou et al., 2012). Transgenic plants have also been generated using inverted repeat (IR) RNAi construct targeting all seven genes of RSV. It was observed that transgenic plants containing IR construct specific for CP and MP showed resistance to RSV. On the other hand transgenic plants carrying IR constructs specific for glycoprotein and non-structural protein did not displayed resistance to RSV (Shimizu et al., 2011). Jiang and co-workers (2012) developed transgenic rice plants containing antisense RNA of RDR6. These plants displayed high level of RSV infection and low levels of RSV derived siRNA. Transgenic rice plants were also developed using the silencing suppressors encoded by RSV. To achieve this, NS2 and NS3, the two silencing suppressors were combined by overlapping PCR and cloned into an expression vector. The recombinant vector was used to transform japonica rice cultivar, ‘Nipponbare’. The virus resistance analysis of transgenic rice plants showed delayed onset of infection by approximately 10–20 days. In addition the transgenic plants also showed 30–50% lower level of virus accumulation, thus revealing substantial resistance to RSV (Zheng et al., 2014). Rice ragged stunt virus Rice ragged stunt virus (RRSV), a member of the group oryzavirus and family Reoviridae, causes ragged stunt disease of rice (Milne and Ling, 1982; Uyeda et al., 1995). The RRSV particles are

composed of a polyhedral core particle of about 50 nm in diameter to which are attached 20-nmwide and 10-nm-high flat spikes. Thus, the overall diameter of virus particles is approximately 70nm (Chen et al., 1997). The genome of RRSV is composed of 10 dsRNA strands and five major proteins (Chen et al., 1989; Hagiwara et al., 1986). RRSV is transmitted by the brown planthopper, Nilaparvata lugens and another Nilaparvata sp. in a persistent manner (Hibino, 1989; Milne and Ling, 1982). It is propagative in the vectors but is not transmitted via eggs. The RRSV-infected rice plants show stunting, leaves with serrated edges or twisted tips, galls on the lower surface of leaf blades or outer surface of the leaf sheaths, vein swelling etc. (Hibino, 1996). Infected cells have been shown to contain large inclusion bodies consisting of a viroplasmic matrix and numerous virus particles. Rice ragged stunt disease was first reported from Indonesia in 1976 and then subsequently was found to be widespread in several countries of Asia like Malaysia, the Philippines, Vietnam and Thailand. RRSV was reported in 1978, in China, India, Sri Lanka (Hibino, 1979), and Taiwan (Chen et al., 1979); and in 1979, in Japan (Shinkai et al., 1980). Upadhyaya and co-workers have completely sequenced the RRSV genome (10 segments) and have identified four genes encoding structural proteins (S3, S5, S8, and S9), one polymerase gene (S4), and two genes encoding nonstructural proteins (S7 and S10, Upadhyaya et al., 1995, 1998a, 2001). In order to engineer resistance to RRSV in rice, japonica (cv. Taipei 309) and indica (cv. Chinsurah Boro II) rice were transformed with plant expression vectors carrying the viral genes. For this, genes encoded by genome segments S5, S7, S8, S9, and S10 were introduced in rice plants in both sense and antisense orientations. Transgenic plants containing viral genes in sense orientation under the control of either the maize Ubi1 promoter or rice Actin1 (Act1) promoter showed accumulation of transgene protein to detectable levels. The resistance analysis of transgenic plants showed significant resistance to RRSV with the resistance levels varying from complete immunity to delayed symptom development (Upadhyaya et al., 1996, 1998b, 2001). Transgenic rice lines expressing the 39 kDa spike protein were developed which showed resistance to infection by RRSV (Chaogang, 2003).

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It was observed that the viral titres in the insect vector N. lugens, which were initially fed on transgenic plants followed by RRSV-infected plants, was inversely proportional to the levels of the 39 kDa protein expressed in the transgenic plants. Rice transitory yellowing virus Rice transitory yellowing virus (RTYV) belongs to the subgroup nucleorhabdovirus of the plant rhabdovirus group (Shikata, 1972). It is responsible for ‘Transitory yellowing disease of rice’, which was first reported in Taiwan and subsequently in Japan and Thailand (Chiu et al., 1965; Inoue et al., 1986). The particles of RTYV are bullet-shaped 180–210 nm in length and 94 nm in width. RTYV genome is composed of ssRNA, which codes for four or five proteins. The major symptoms of RTYV infection are leaf yellowing, reduced tillering, mild stunting and sterility. RTYV is transmitted in a persistent manner by Nephotettix cincticeps, N. nigropictus, and N. virescens (Hibino, 1989; Shikata, 1972). It is propagative in the vectors but is not transmitted via eggs. N. cincticeps and N. nigropictus have been found to show higher vector efficiency as compared to N. virescens. RTYV has been reported mainly from southern and central China, the Nansei Islands (Okinawa) of Japan, Taiwan, and Thailand (Hibino, 1989). Some of the severe outbreaks of RTYV were reported from Taiwan in 1960–1962, 1973–1975, and 1977–1980 (Chiu and Jean, 1969; Chen et al., 1980) and China in 1964–1966 and in 1979 in Guandong, China (Faan, 1980). RTYV is closely related to rice yellow stunt virus (RYSV), which is also a member of the group nucleorhabdovirus. RYSV causes yellow stunt disease of rice, which is mostly prevalent in central and southern China (Chen et al., 1979; Fan et al., 1965). RYSV and RTYV are transmitted by the same insect vectors. In order to deduce the relationship between RTYV and RYSV, nucleotide sequences of both the viruses were compared. Sequence analysis of RTYV and RYSV revealed an overall nucleotide and amino acid identities of 98.5% and 99.7% respectively. The complete nucleotide sequences of RTYV and RYSV indicated that both are strains of the same virus, rather than distinct viruses; however, symptoms on rice induced by RTYV infection differ slightly from those caused by RYSV infection (Hiraguri et al., 2010).

The genome organization of plant rhabdovirus is a non-segmented, single-stranded negative sense RNA genome, which contains, usually, five open reading frames (ORFs) in the order 3′ N-P-M-G-L 5′, where N encodes the nucleoprotein, P the polymerase-associated protein, M the matrix protein, G the glycoprotein, and L the polymerase (Dietzgen et al., 2011). In some rhabdoviruses, a sixth ORF that encodes a nonstructural protein of unknown function has been observed. Besides, there are leader sequences located at the 3′ and 5′ ends of the genomic RNA. Fang et al. (1992) reported that the RYSV genome contains three major structural proteins: G (84 kDa), N (60 kDa), and M (31 kDa), and two minor proteins: L (170 kDa) NS (corresponding to P protein, 42 kDa) (Fang et al., 1994). Fang et al. (1996) reported the development of transgenic rice resistant against RYSV. The transgenic rice lines were expressing a cDNA copy of RYSV nucleocapsid protein gene, or a reading frame-shift mutant of the gene has been inserted into a rice expression vector under the control of the rice Act1 promoter. Virus resistance assays of primary transgenic lines displayed resistance to the RYSV infection, indicating that this resistance might be mediated by the RNA transcripts of the nucleocapsid protein gene (Fang et al., 1996). Rice stripe necrosis virus RSNV, which belongs to the furovirus group, causes ‘crinkling disease of rice’ in West African countries (Fauquet and Thouvenel, 1983; Fauquet et al., 1993). The RSNV particles are rod shaped, having variable length of 120, 270 or 380 nm and width of 20 nm. RSNV is soil borne and is maybe transmitted by the fungal vector Polymyxa graminis, which belongs to the order Plasmodiophorales. The symptoms induced by RSNV infection are stunting, crinkling, reduced tillering, chlorotic or bright yellow stripes on leaves, necrosis of the leaves or of the whole plant. In plant cells infected with RSNV, virus particles are found as aggregates in the cytoplasm. The virus is transmitted mechanically from plant to plant (Fauquet et al., 1993). The crinkling disease of rice was first reported in 1977 from Ivory Coast (Louvel and Bidaux, 1977). Fauquet and Thouvenel (1983) revealed that this disease is associated with a rod shaped virus, which has not been reported earlier. On the basis of the symptoms produced by infection this new virus

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was designated as RSNV. Later the disease was also reported from the Ivory Coast, Liberia, Nigeria, and Sierra Leone. Estimated yield loss due to RSNV has been found to vary between 14% and 100% (Fauquet et al., 1993). Rice black-streaked dwarf virus RBSDV belongs to the genus Fijivirus of the Reoviridae family. It is transmitted over a wide host range of plants such as rice, maize, wheat and barley by small brown plant hopper (Laodelphax striatellus) in a persistent and propagative manner (Shinkai, 1962; Mertens, 2004; Wei and Li, 2016). RBSDV can be experimentally transmitted in vitro to rice, maize, wheat and sorghum (Ishii and Yoshimura, 1973). RBSDV mainly causes black streaked dwarf and maize rough dwarf diseases (Fang et al., 2001; Bai et al., 2002). In rice, the symptoms of RBSDV include drastic growth abnormalities such as severe stunting, darkening of leaves and veins with waxy gall alongside (Shikata and Kitagawa, 1977; Hibino, 1996; Fang et al., 2001). RBSDV reports first emerged from Japan in 1968 where it caused severe yield loss of rice and maize crop (Ishii and Yoshimura, 1973). In China, it was reported in 1960s and till the late 1980s or the early 1990s the virus was prevalent in most of the regions of northern Fujian and Zhejiang provinces of China. The eastern regions of China in 1997 and 1998 reported an almost 100% yield loss in rice due to RBSDV (Li et al., 1999). The virus particle is icosahedral in shape and is double layered containing a dsRNA with 10 segments (S1 to S10, Zhang et al., 2001; Wang et al., 2003). Segments S1 to S4 have one open reading frame encoding for protein P1 (RNAdependent RNA polymerase), P2 (inner core protein), P3 (capping enzyme function) and P4 (outer shell protein with spiked appearance respectively). S6, S8 and S10 again having a single ORF each, encode for P6 (viroplasm associated), P8 (minor core capsid protein), and P10 (main outer protein of the capsid, Isogai et al., 1998; Zhang et al., 2001; Liu et al., 2007 a,b). The remaining segments, S5, S7 and S9, encode two proteins each, P5-1, P7-1 and P9-1, which are viroplasm-associated (Yang et al., 2013), tubular structure forming (Isogai et al., 1998 and Liu et al., 2011) and the main viroplasm matrix protein (Zhang et al., 2008; Wang et al., 2011; Akita et al., 2012) respectively. The P5-2, P7-2 and

P9-2 have not been assigned any particular function. The quest for transgenic-based approach with RNAi as underlying mechanism is well worked in case of RBSDV. The 500 bp fragment from 5′-end of P9-1, the chief viroplasm protein of the virus was introduced as dsRNA in rice plants cv. Nipponbare (Shimizu et al., 2011). These plants when inoculated by virus remained asymptomatic even after 4 weeks post inoculation (wpi) and no virus was detected later in these transgenic plants by ELISA too, whereas the characteristic RBSDV symptoms appeared on non-transgenic plants which became more severe at 4 wpi. Southern Rice black-streaked dwarf virus Southern Rice black-streaked dwarf virus (SRBSDV) was first discovered as a novel virus in Guangdong Province of China in 2001 and is included under group 2 of the genus Fijivirus in the family Reoviridae (Zhang et al., 2008; Zhou et al., 2008). Since then the disease has spread to several areas of southern China and Vietnam and has caused severe yield loss in rice crop of those regions (Cuong et al., 2009; Guo et al., 2010; Wang et al., 2010; Hoang et al., 2011). In addition to rice, the virus has a very vast host range including maize, barnyardgrass (Echinochloa crus-galli), Chinese sorghum (Coix lacryma-jobi) and flaccid grass (Pennisetum flaccidum) (Zhou et al., 2008). In 2009–10 the SRBSDV spread from initial nine provinces of China to 13 new provinces in southern China and 28 in northern Vietnam as well as one in central Vietnam causing thousands of hectares of crop to be devastated completely resulting into huge economic loss (Guo et al., 2010). Recently, the disease has been observed in some areas of Japan too (Matsukura et al, 2013). Initially the SRBSDV was not considered as a unique virus, but a variant of RBSDV (Zhou et al., 2004), but the key findings by Zhou et al. and Zhang et al. in 2008 suggested major divergence in the sequence of S7-S9 of a Guangdong (GD) isolate and S9 and S10 of a Hainan (HN) isolate. They observed less than 80% nucleotide identity of this virus to other fijiviruses and thus suggested Fijivirus group 2 for the virus. It was named RBSDV-2 or SRBSDV by them. SRBSDV is transmitted by the white-backed planthopper (WBPH, Sogatella furcifera), order Hemiptera and class Delphacidae.

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WBPH does not transmit RBSDV, thus again making it evident that RBSDV and SRBSDV are two different but related viruses. S. furcifera also transmits to the other host plants of the Poaceae family mentioned previously except rice. In terms of symptoms also the SRBSDV do not causes white waxy outgrowths on leaves and culms, which is a characteristic feature of RBSDV; also, SRBSDV induces tiller formation in the upper parts of the infected rice plants which is usually not observed in RBSDV infection. Other SRBSDV symptoms includes stunting, dark leaf and small enations on stem and leaf back, the seriously diseased plants sometimes die of withering (Zhou et al., 2008, 2013). The intensity of infection also depends upon the growth stage of the plant; the infection at young stage of the plant causes the symptoms to be more severe (Zhou et al., 2010) as compared to the infection at later stage of the plants which do not involve stunting but produce small spikes, empty or less weight grains (Zou et al., 2013). The particle of the SRBSDV virion is icosahedral in shape with a diameter of 70–75 nm. The genome contains ten linear dsRNA segments S1-S10 (named according to the decreasing molecular weight) with sequence size ranging from 1.8–4.5 kb (Zhou et al., 2008; Wang et al., 2010). The ten genome segments contains 13 ORFs, with S5, S7 and S9 each encoding two protein while the others encoding for single protein each. S1-S4, S8 and S10 encodes for putative structural proteins which are RNA dependent RNA polymerase (RdRp), major core, capping enzyme, outer shell, minor core and major outer capsid proteins respectively. While, P6, P7-1, P7-2, P9-1 and P9-2 encode putative non-structural proteins (Zhang et al., 2008; Zhou et al., 2008; Wang et al., 2010). P6 protein is a suppressor of RNA silencing and is believed to play role in viroplasm association by self-interaction and by interacting with P5-1 and P9-1 (Lu et al., 20011; Li et al., 2013; Wang et al., 2011). P6 also interacts with rice elongation factor 1A (eEF-1A) thus inhibiting host translation (Songbai et al., 2013). P7-1 is believed to be virus movement protein and is a major component of tubules (Liu et al., 2011; Jia et al., 2014). P9-1, a major component of viroplasm helps in its formation as well as virus proliferation in insect vector ( Jia et al., 2014; Lia et al., 2015). The functions of other viral proteins of SRBSDV remain unclear. Disease management against SRBSDV includes

the control of the insect vector (Matsukura et al., 2013; Wang et al., 2015) by chemical, ecological or physical control, such as by using insect-proof nylon mesh or plastic film or variation in the crop sowing date to avoid the peak of the vector migratory population. Since the viroplasm-associated proteins in reoviruses has proved to be a vulnerable target for effective RNAi, similarly the P9-1 gene of SRBSDV, which is a major viroplasm component protein can be used to raise RNAi based transgenic plants for resistance (Sasaya et al., 2014). However, Rice cultivars with high resistance to SRBSDV are unfortunately not yet available and transgenic approaches have not yet been reported to obtain resistance to SRBSDV. Rice bunchy stunt virus Chen et al. (1978) first reported bunchy stunt disease in rice, and since then it has been reported in Fujian, Gaundong, Guanxi, Hubei, Hunan, Jiangxi, Queichou, and Yunnan provinces in China (Lishi et al., 1994). It is caused by RBSV, a member of Phytoreovirus group of family Reoviriade. The viral genome consists of 12 segments of dsRNA encapsidated in a polyhedral structure of 60 nm in diameter. The symptoms include height reduction but increase in number of tillers, excessive nodal branching and bunches of small short narrow leaflet giving a bird nest type of appearance. The disease incubation period is 8–44 days. The second crop rice shows more RBSV incidence and the occurrence is more towards the edges of the fields (Xie et al., 1979). The crop loss due to RBSV is largest when infection occurs during stages of green recovering and tillering (Xie et al., 1980, 1982, 1984; Liyan et al., 1991). RBSV is transmitted persistently but not transovarially by the leafhoppers Nephotettix cincticeps and Nephotettix virescens (Xie et al., 1996). N. cincticeps is more efficient in transmitting RBSV as compared to N. virescens (Xie et al., 1984). No transgenic approach has been used till date against RBSV; however, some rice varieties e.g. Chikuaiai 3, Baotaiai, Zhenlong 13 and Zhenlong 410 are screened for their natural genetic resistance to RBSV (Xie et al., 1984). Agronomic practices such as non-synchronizing the development stages of host rice plant and the viral vector and regulation of sowing and transplanting times in a way to avoid the maximum population stages of the vectors has

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proven to be efficient to some extent in controlling RBSV (Xie et al., 1984). Rice dwarf virus Rice dwarf virus (RDV) is a member of Phytoreovirus genus of family Reoviridae (Lida et al., 1972). It has a wide range of hosts from fungi to plants, insects, several vertebrates and even human (Boccardo and Milne, 1984). RDV causes Rice dwarf disease (RDD) with symptoms such as pronounced stunting, short dark green leaves with chlorotic specks, increased tillering, delayed and incomplete panicles (Hibino, 1996. RDV is transmitted in a persistent and propagative manner by leafhopper vector Nephotettix cincticeps, Recilia dorsalis, and few other Nephotettix sp., N. cincticeps being the principal vector (Lida et al., 1972; Hibino, 1990; Attoui et al., 2012). RDV can replicate both in the plant host, i.e. rice as well as in the vector (Suzuki et al., 1994). The particle structure of RDV icosahedral, double layered (shelled), spherical with 70 nm diameter. Its genome consists of 12 dsRNA segments (S1-S12, Mizuno and Kano, 1991; Lu et al., 1998; Naitow et al., 1999) which encode for seven structural (P1, P2, P3, P5, P7, P8 and P9) and five non-structural proteins (Pns4, Pns6, Pns10, Pns11 and Pns12) respectively (Suzuki et al., 1994, 1996; Omura and Yan, 1999; Nakagawa et al., 2003; Zhong et al., 2003; Wei et al., 2006). The P2, P8 and P9 proteins form the outer shell while the P1, P3, P5 and P7 forms the inner core shell (Mao et al., 1998; Naitow et al., 1999; Omura and Yan, 1999). The initial step towards RDV genome replication is the aggregation and packaging of the 12 viral mRNA corresponding to the 12dsRNA segments which is then followed by negative strand synthesis resulting into synthesis of new dsRNA genome segments. The genome replication is followed by assembly of the protein particle, the core proteins assembling first for the formation of viral mRNAs bulk followed by the outer shell proteins forming a protective covering to generate mature RDV virion (Zhong et al., 2003; Wei et al., 2006a,b; Miyazaki et al., 2010). P1 protein is thought to have RNA polymerase function (Suzuki et al., 1992). The P2 has its role in successful transmission of RDV from infected to un-infected plant via the vector host (Zhu et al., 2005), it also plays a key role in viral attachment and entry into the host cell involving

the mechanism of clathrin-mediated endocytosis (Tomaru et al., 1997; Omura et al., 1998; Wei et al., 2007). The electron tomographic studies indicated their presence away from the outer coat in form of spike like structures connecting the virus particle to the host cell (Miyazaki et al., 2016). The nonstructural protein Pns10 is a viral suppressor of RNA silencing (Zhou et al., 2010). The cell-to-cell movement of RDV is facilitated by another nonstructural viroplasm associated protein Pns6 (Li et al., 2004). In this regard, RNAi has been used as a strategy against RDV. To obtain RNAi-mediated resistance to RDV, transgenic plants harbouring a dsRNA construct of about 500-bp fragment of the 5′-end region of the viral genes were raised. A high degree of variability in resistance was observed (Shimizu et al., 2009; Sasaya et al., 2014). The transgenic plants with dsRNA construct corresponding to Pns4 (movement among cells), Pns6, Pns8 (chief outer capsid), Pns11 (function not known) and Pns12 (viroplasm associated) proteins of RDV were found to be highly efficient in providing resistance to RDV. The infected transgenic plants were completely asymptomatic with no detectable virus through ELISA. Whereas the transgenic plants with dsRNA construct for P2, P5, P7, P9 or Pns10 did not show significant resistance to RDV with similar symptom severity as compared to non-transgenic susceptible plants. This indicates that the proteins that are involved in attachment, movement and maintenance of viroplasm, if targeted, provide high levels of resistance to RDV as compared to protein with accessory functions. The proteins providing accessory functions include Pns10 (silencing suppressor), P2 and P9 (outer capsid proteins), the capping enzyme and nucleic acid binding protein functions being contributed by P5 and P7 respectively (Suzuki et al. 1990a,b, 1991, 1992a,b; Suzuki, 1993; Uyeda et al., 1994, Zhong et al., 2003, Li et al., 2004, Cao et al., 2005). Rice gall dwarf virus The Rice gall dwarf virus (RGDV) is a member of the group Phytoreovirus of Reoviridae family (Omura and Inoue, 1985; Uyeda et al., 1995). RGDV is transmitted by a hemipteran zigzag leafhopper of Cicadellidae family, Recilia dorsalis (Motsch) and Nephotettix nigropictus (Stal) of the same family in a persistent-propagative manner and via maternal

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inheritance, i.e. from the infective females to their progeny through the eggs (Inoue and Omura, 1982; Morinaka et al., 1982; Fan et al., 1983; Bocardo and Milne, 1984; Xie and Lin, 1984). RGDV was first reported from Thailand (Omura et al., 1980) and then from the region of China and Southeast Asia causing severe yield loss due to RGD disease (Putta et al., 1980; Ong and Omura, 1982; Fan et al., 1983). Reports have now emerged in recent years about the spread of RGDV to other provinces of China such as Guangdong, Hainan and Guangxi (Zhang et al., 2008; Fan et al., 2010). The characteristic symptoms of RGDV includes dark green discoloration of the leaf blades, with a whitish gall of 2 mm long and 0.4–0.5 mm wide dimensions along the under sides of the leaf blades and outer sides of the leaf sheaths (Putta et al., 1980; Omura et al., 1980), delayed flowering, severe stunting and incomplete panicle with half-filled or empty grains (Fan et al., 1983). The particle structure of RGDV is icosahedral and is with protective double shelled covering with an approximate diameter of 65–70nm (Boccardo and Milne, 1984). The viral genome like those of other phytoreoviruses consists of 12 segments (S1 through S12) of double-stranded RNAs (dsRNAs) (Omura et al., 1982; Hibi et al., 1984; Boccardo et al., 1985). The segments S1, S2, S3, S5, S6 and S8 encode for the structural proteins P1, P2, P3, P5, P6 and P8, respectively (Hibi et al., 1984; Boccardo et al., 1985). The remaining segments of the RGDV genome code for non-structural proteins namely Pns4, Pns7, Pns9, Pns10, Pns11 and Pns12 (Hibi et al., 1984; Boccardo et al., 1985; Koganezawa et al., 1990; Noda et al., 1991; Moriyasu et al., 2000, 2007). These non-structural proteins form viral inclusions important for propagation through the vectors (Hogenhout et al., 2008) for e.g. Pns7, Pns9 and Pns12 form important component of viroplasm, where the virus replication and progeny virions are assembled in the gut cells of insect vector (Wei et al., 2009; Akita et al., 2011). Pns11 helps in virus tubule formation and thus causing viral spread in host vector cells. Pns9 is found to be the chief viral factor for viroplasm formation in infected host cells because of its ability to form inclusions resembling viroplasm in the non-vector insect Spodoptera frugiperda (Sf9) cells (Akita et al., 2011). Pns11 encoded by RGDV segment 11 has also been shown to possess viral silencing suppressor activity (Liu et al., 2005). The function of Pns4

and Pns10 has not yet been discovered. Zheng et al. (2015) experimentally demonstrated the role of non-structural protein Pns9 in assembly of the viroplasm and its essentiality for persistent RGDV presence in its insect vector. Since the virus stays in insect gut, establishes productive infection resulting into replication and assembly of new virion particles, the control of virus in the insect vector could be the safest strategy to overcome RGDV infection spread. Successful RNAi strategies have been used to inhibit the infection of persistentpropagative viruses in insect vectors (Chen et al., 2012, 2013, 2014; Jia et al., 2012a,b, 2014; Wu et al., 2014). Further detailed studies of the mechanism of RGDV infection, spread and propagation in its vector will better equip us with innovative strategies to disrupt the efficient RGDV transmission through the vector. Although the virus vector relationship is not mutual, i.e. it is only beneficial to virus for its propagation while to the vector R. dorsalis the virus decreases the survival rate, emergence rate, reduced level of vitellogenin (Vg) in females causing reduced fecundity, and longevity of adults (Chen et al., 2016). In other words, the virus shows suicidal features in terms of its vector where it is itself responsible for reduction in vector host population, which in turn will reduce the virus proliferation. The RNAi based approach for generating transgenic plants against RGDV has proved to be very efficient in this case. For other reoviruses, such as RDV, the transgenic plants harbouring the RNAi construct against the viroplasm associated proteins showed good amount of virus resistance (Shimizu et al., 2009; Sasaya et al., 2014). The transgenic plants raised with construct against RGDV protein Pns9 which is a functional orthologue of RDV viroplasm associated protein was then thought to produce similar amount of virus resistance (Akita et al., 2009). For this, the japonica rice cultivar Nipponbare was transformed with hairpin RNA of a 500bp fragment from the 5′ end of the Pns9 protein corresponding to the S9 genome segment of RGDV and the plants were then analysed for their virus resistance (Shimizu et al., 2012). The transgenic plants showing the presence of transgene specific siRNA, showed no symptoms and until the harvest stage and the virus was undetectable by ELISA whereas the non-transgenic control plants showed all characteristic RGDV symptoms (Shimizu et al., 2012).

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Rice giallume virus RGV causes a serious disease of rice called rice giallume, prevalent in Spain and northern parts of Italy (Belly et al., 1974, Osler, 1983). It is probably developed from a strain of Barley yellow dwarf luteovirus (Osler, 1983; Belly et al., 1986). The genome consists of a single-stranded RNA (ssRNA) encapsidated in polyhedral single protein particle of less than 30 nm in size (Amici et al., 1974; Belly et al., 1986). RGV in plants is transmitted in a persistent manner by Rhopalosiphum padi, an aphid of superfamily Aphidoidea in the order Hemiptera (Osler et al., 1974). RGV is a phloem-limited virus found single or aggregated throughout the cytoplasm of phloem cells. In the rice plant it causes yelloworange leaf discoloration, reduced plant height and tillers. Phloem cells infected with RGV often show degenerated mitochondria, shows abundant fibril rich membrane vesicles and thick deposition across the cell wall (Gill and Chong, 1979; Osler, 1983; Belly et al., 1986). RGV infection also affects the plant metabolism by altering the levels of macro (N, P, K, Ca, Mg) and micro (Zn, Fe, Mn) nutrients (Mazzini et al., 1990). Although no transgenic approach has to date been used against RGV, Baldi et al. (1990) reported the involvement of an incomplete dominant gene to be responsible for RGV resistance in some rice cultivars. Rice grassy stunt virus The grassy stunt disease of rice was described by Rivera et al. (1966) and Bergonia et al. (1966). The causal organism is RGSV, a member of the genus Tenuivirus (Toriyama, 1985; Hibino, 1986; Falk and Tassai, 1998) previously known by the name Rice rosette virus (Bergonia et al., 1966). The virus is transmitted by the brown planthopper, Nilaparvata lugens, along with two other species, N. bakeri and N. muiri (Hibino, 1986, 1990). The virus is prevalent in China, Japan, and Taiwan and in parts of South and Southeast Asia (Hibino, 1996). Severe outbreaks of RGSV were reported in 1970s and 1980s from parts of Indonesia, India, Japan and Philippines (Kulshreshta et al., 1974; Mariappan et al., 1984; Iwasaki et al., 1985; Hibino, 1990). The symptoms include severe stunting, profuse tillering, short narrow pale yellow leaves with brown spot and no or few panicles with dark brown or unfilled grains. Infection at the later

stages causes severely yellow leaf, brown panicles and abortive kernels (Rivera et al., 1966; Chen and Chung, 1982; Mariappan et al., 1984). RGSV has a thread-like 6- to 8-nm-wide particle appearance; mostly the shape is circular with a varying contour length of 200–2400 nm. RGSV genome consists of six ssRNA named RNAs 1 to 6 according to their size exhibiting an ambisense coding behaviour in terms of viron sense (v) and virion complementary (vc) sequences. It has 12 ORFs, two on each RNA segment (Kormelink et al., 2011; Shirako et al., 2011). The transcription of viral mRNAs involves a 5′-endonuclease activity existing within the viral RdRp, a phenomenon often called cap snatching (Shimizu et al., 1996). The ORF of the viral negative sense RNA5 encodes for nucleoprotein (NP) (Toriyama et al., 1997). The genomic RNA of RGSV associates with the nucleoprotein (NP) to form ribonucleoprotein (RNP) complex (Falk et al. 1998). The functions of all the RGSV proteins are not yet understood. For example, RNA 1 vc strand codes for pc 1 which is believed to be an RNAdependent RNA polymerase (RdRp) and vcRNA5 and vcRNA6 encode for structural (pc 5) and movement proteins (pc 6) respectively (Hiraguri et al., 2011; Kormelink et al., 2011; Shirako et al., 2011). The p5 protein of RGSV is known to interact with the host-specific CBL–CIPK Ca2+ signalling network of rice and since CBL-CIPKs plays key role in ion metabolism in plant particularly those of potassium and sodium (Xu et al., 2006; Weinl et al., 2009; Luan et al., 2009; Wang et al., 2013), RGSV tends to hijack the CIPK/25 to manipulate the host functions as indicated by lower content of potassium in the RGSV affected host plant (Xiong et al., 2017). Two of the proteins p2 and p5 of RGSV have been shown to possess the silencing suppression activity against the host RNAi (Netsu et al., 2015; Nguyen et al., 2015). RNAi-based transgenic approach against RGSV has proved to be successful in generating virus free plants. The idea originated from an another Tenuivirus RSV, as mentioned in the previous section that the RNAi constructs against the nucleocapsid protein (pc 3) and the movement protein (pc 4) of RSV were found to be very efficient in providing resistance to RSV (Shimizu et al., 2011); their functional orthologues in RGSV pc 5 and pc 6 (Hiraguri et al., 2011) were also taken similarly as RNAi targets expecting that a similar degree of resistance would be obtained in

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case of RGSV too. The transgenic plants showed accumulation of transgene-specific siRNA. Strong resistance was observed in transgenic plants harbouring the pc 5- or pc 6-specific constructs by remaining asymptomatic even after four weeks post-inoculation as compared to the wild type, which showed typical RGSV symptoms (Shimizu et al., 2013). The findings suggested that the RGSV genes coding for the nucleocapsid protein and the movement protein are the most effective targets for producing RGSV-resistant plants. Rice hoja Blanca virus Rice hoja Blanca virus (RHBV) is a member of the genus Tenuivirus and was first reported in rice from Colombia (Garcés, 1940). The virus was isolated for the first time by Morales and Niessen (1983). The virus particles are filamentous, 3–8 nm in diameter, exhibit supercoiling with variable length (Morales and Niessen, 1983). Rice hoja blanca (RHB) disease from its first emergence in Colombia in 1935 has now spread throughout tropical and subtropical America causing severe yield loss in rice (Atkins and Adair, 1957; Garces et al., 1958; Everett and Lamey, 1969; Morales and Niessen, 1985). RHB disease symptoms includes death in young seedlings, reduction in photosynthetic activity, stunting and panicle sterility resulting into a yield loss of about 25–75% in RHBV susceptible cultivars ( Jennings et al., 1958; Everett and Lamey, 1969; Lamey, 1969). However, if the infection is delayed and occurs at the time of emergence of panicle, only slight yield reduction is observed (McGuire et al., 1960). An interesting thing about the rice–RHBV interaction is that the virus infection initiates the secondary infection of brown spot causing fungus, Helminthosporium oryzae (Lamey and Everett, 1967). The RHBV genome consists of four single-stranded linear RNA species, and the total genome is estimated to be 17.6 kb (Ramirez et al., 1992). It is transmitted in a circulative, propagative and transovarial manner by the insect vector Tagosodes orizicolus (Muir), commonly referred to as ‘sogata’ (Acuna et al., 1966; Malaguti et al., 1957; Galvez and Jennings, 1959; McGuire et al., 1960). Non-viruliferous T. orizicolus in itself is harmful for the rice crop because of its feeding behaviour and causes significant yield loss. Several cultivars possess genetic resistance to the vector; the underlying

mechanism for this resistance in the host rice plant however is quite different from that of genetic resistance in host to RHBV ( Jennings and Pineda, 1970). The genetic resistance obtained to some extent against RHBV is reported through various genetic crossing programmes in 1960s using semi dwarf indicia varieties (Morales and Jennings, 2011). Romero et al., (2014) identified three QTL in two resistant cultivars Fedearroz 50 and Fedearroz 2000; one QTL linked to RHBV resistance and the other two QTL accounts for resistance to T. orizicolus. There is no report of transgenic resistance to RHBV; however, natural RNAi mechanism in plants against RHBV is being studied in details and has revealed the involvement of NS3 protein of RHBV in binding to siRNA to suppress the host RNAi response (Goldbach et al., 2003; Hemmes et al., 2007, 2009; Yang et al., 2011) indicating that RHBV is no different from other viruses in using an RNAi suppression strategy for its multiplication in host plant (Lakatos et al., 2006; Nakahara et al., 2012). Rice necrosis mosaic virus Rice necrosis mosaic virus (RNMV) belongs to genus Bymovirus and the family Potyviridae. It is transmitted to rice roots through infection by the soil-borne plasmodiophoromycete (a fungus) Polymyxa graminis. New plants get infection from zoospores or resting spores released from the infected plants (Inouye and Fujii, 1977; Usugi et al., 1989). The genome is constituted by a positive sense ssRNA encapsidated by a coat protein (CP). It has a flexuous rod shaped particle structure with two modal lengths of 205 and 550 nm and a width of about 13–14 nm (Inouye and Fujii, 1977). The virus is serologically related to barley yellow mosaic and wheat yellow mosaic viruses (Usugi and Saito, 1976). RNMV infection in rice was first observed in parts of Japan and later at Cuttack in India (Ghosh, 1980). Rice plants infected with RNMV show reduction in height and number of tillers, a bushy shrub like growth habit, a mosaic appearance in leaf represented by yellow flecks, streaks on the lower leaves and necrotic flecks at the lower stem and sheaths (Badge et al., 1997). The RNMV genomic RNA is 7178 nt long with a single ORF encoding for a single 258kDa polyprotein, a characteristic of bymoviruses. The nucleotide sequence

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of RNA shows similarity with other members of bymoviruses, for example 46% with Oat mosaic virus and 56% identity with Barley mild mosaic virus (BaMMV, Wagh et al., 2016). RNMV infection dynamics at the cellular level has not yet been studied in details however, Wagh et al. (2016) reported that OsRDR6 of rice plays an important role in RNMV infection, as rice OsRDR6 mutants developed more virus levels as compared to the control. Recently, a probenazole (PBZ) inducible haem activator protein (HAP) gene, OsHAP2E (Os03g29760, Alam et al., 2016) was analysed for its potential role in resistance to RNMV. It was found that RNMV infection induced the expression level of OsHAP2E in both rice leaves and shoots and the T3 transgenic overexpression lines for the OsHAP2E gene showed considerably low level of RNMV, thus conferring resistance to the rice plants against RNMV (Alam et al., 2015). Conclusion In the absence of natural resistance, the only effective strategy to be used to manage viruses is to express a portion of the viral gene in the plant, such that it triggers RNAi response against the virus. Substantial resistances have been reported against most rice-infecting viruses, using this strategy. One factor to be kept in mind when using RNAi as a resistance strategy is the presence of RNAi suppressor activities in the viral proteins. The resistance strategy should, thus, preferentially target such proteins to prevent their expression, to ensure high levels of resistance. This has, in fact, been the strategy for many rice viruses. The other important factor should be the extent of variability in the viral genome or the tendency to recombine with related viruses during mixed infection. The above information should be used to choose the region of the genome as the transgene, to achieve widespread resistance in the field. Although the degree of resistance reported shows a large variation for the various viruses, it should be sufficient to manage most viral diseases of rice quite effectively. However, the deployment of this resistance under field conditions is not yet reported. The future years should see more evaluation of the resistant lines under field conditions so that the benefits of the resistance can be translated

in increasing rice production to meet the future requirements. Acknowledgements GK acknowledges the research fellowship from University Grants Commission, New Delhi. Research in the ID lab is funded by Department of Biotechnology, Government of India, under a BIPP grant and also from research grants from University of Delhi (R&D and DST-PURSE). References Abo, M.E., Alegbejo, M.D., Sy, A.A., Adeoti, A.A., and Marley, P.S. (2003). The host range of Rice yellow mottle virus genus Sobemovirus in Cote d’Ivoire. Samaru J. Agric. Res. 9, 67–78. Abo, M.E., and Sy, A.A. (1997). Rice virus diseases: epidemiology and management strategies. J. Sustain. Agric. 11, 113–134. Acuña, J., Ledón, L.R., and Cardet, Y.L. (1966). Sogata orizicola muir vector de la enfermedad virosa hoja blanca del arroz en Cuba (Centro Nacional Exp. Ext. Agric.). Akita, F., Miyazaki, N., Hibino, H., Shimizu, T., Higashiura, A., Uehara-Ichiki, T., Sasaya, T., Tsukihara, T., Nakagawa, A., and Iwasaki, K. (2011). Viroplasm matrix protein Pns9 from Rice gall dwarf virus forms an octameric cylindrical structure. J. Gen. Virol. 92, 2214–2221. Alam, M.M., Tanaka, T., Nakamura, H., Ichikawa, H., Kobayashi, K., Yaeno, T., Yamaoka, N., Shimomoto, K., Takayama, K., and Nishina, H. (2016). Overexpression of a rice heme activator protein gene (OsHAP2E) confers resistance to pathogens, salinity and drought, and increases photosynthesis and tiller number (vol 13, pg 85, 2015). PLANT Biotechnol. J. 14, 1171–1172. Alam, M.M., Nakamura, H., Ichikawa, H., Kobayashi, K., Yaeno, T., Yamaoka, N., and Nishiguchi, M. (2015). Overexpression of OsHAP2E for a CCAAT-binding factor confers resistance to Cucumber mosaic virus and Rice necrosis mosaic virus. J. Gen. plant Pathol. 81, 32–41. Alegbejo, M.D., Raji, B.A., Abubakar, I.U., and Banwo, O.O. (2006). Rice yellow mottle virus disease, a new disease of rice in Zamfara, Nigeria. NOTES FORM F. 39, 39. Amici, A., Osler, R., and Belli, G. (1974). An Isometric Virus associated with the ‘giallume’ Disease of Oryza sativa. J. Phytopathol. 79, 285–288. Atkins, J.G., and Adair, C.R. (1957). Recent discovery of hoja blanca, a new rice disease in Florida, and varietal resistance tests in Cuba and Venezuela. Plant Dis. Report. 41, 911–915. Attoui, H., Becnel, J., Belaganahalli, S., Bergoin, M., Brussaard, C.P., Chappell, J.D., Ciarlet, M., del Vas, M., Dermody, T.S., and Dormitzer, P.R. (2012). Part II: The Viruses–the double-stranded RNA viruses-family Reoviridae. Azzam, O. (2002). The biology, epidemiology, and management of rice tungro disease in Asia. Plant Dis. 86, 88–100. Bakker, W. (1970). Rice yellow mottle, a mechanically transmissible virus disease of rice in Kenya. Netherlands J. Plant Pathol. 76, 53–63.

192  | Kumar et al.

Bakker, W. (1975). Rice yellow mottle virus. C. Descr. Plant viruses 149, 1–4. Bakker, W. (1974). Characterization and ecological aspects of Rice yellow mottle virus in Kenya (Pudoc). Baldi, G., Moletti, M., and Osler, R. (1990). Inheritance of resistance to giallume in rice. In World Perspectives on Barley Yellow Dwarf International Workshop, Udine (Italy), 6–11 July 1987 (CIMMYT). Banerjee, A., Roy, S., and Tarafdar, J. (2011). Phylogenetic analysis of Rice tungro bacilliform virus ORFs revealed strong correlation between evolution and geographical distribution. Virus Genes 43, 398–408. https://doi. org/10.1007/s11262-011-0647-z Barbier, P., Takahashi, M., Nakamura, I., Toriyama, S., and Ishihama, A. (1992). Solubilization and promoter analysis of RNA polymerase from Rice stripe virus. J. Virol. 66, 6171–6174. Belli, G., Amici, A., Corbetta, G., and Osler, R. (1974). Giallume disease of rice (Oryza sativa L.). Acta Biol Iugosl, Ser B Mikrobiol. Belli, G., Cinquanta, S., Prati, S., and Soncini, S. (1986). Purification and serological identification of an isolate of Barley yellow dwarf virus causing rice giallume. Riv. di Patol. Veg. 22, 81–86. Bergonia, H.T., Capule, N.M., Novero, E.P., and Galica, C.A. (1966). Rice rosette, a new disease in the Philippines. Philipp. J. Plant Ind. 31, 47–52. Boccardo, G., and Milne, R.G. (1984). Plant reovirus group. CMI/AAB descriptions of plant viruses no. 294. Boccardo, G., and Milne, R.G. (1984). Plant reovirus group. C. Descr. plant viruses 294. Boccardo, G., Milne, R.G., Disthaporn, S., Chettanachit, D., and Putta, M. (1985). Morphology and nucleic acid of Rice gall dwarf virus. Intervirology 23, 167–171. Bonneau, C., Brugidou, C., Chen, L., Beachy, R.N., and Fauquet, C. (1998). Expression of the Rice yellow mottle virus P1 protein in vitro and in vivo and its involvement in virus spread. Virology 244, 79–86. Cabauatan, P.Q., and Hibino. (1985). Transmission of rice tungro bacilliform and spherical viruses by Nephotettix virescens Distant. Philipp. Phytopathol. v. 21. Cao, X., Zhou, P., Zhang, X., Zhu, S., Zhong, X., Xiao, Q., Ding, B., and Li, Y. (2005). Identification of an RNA silencing suppressor from a plant double-stranded RNA virus. J. Virol. 79, 13018–13027. Chaogang, S., Jianhua, W., Guoying, Z., Gang, S., Baozhen, P., Juanli, L., Dendi, J., Shenxiang, C., Upadhyaya, N.M., and Waterhouse, P. (2003). Ectopic expression of the spike protein of Rice Ragged Stunt Oryzavirus in transgenic rice plants inhibits transmission of the virus to insects. Mol. Breed. 11, 295–301. Chen, C.C., Chen, M.J., Chiu, R.J., and Hsu, H.T. (1997). Rice ragged stunt virus (Oryzavirus) possesses an outer shell and A-spikes. PLANT Prot. Bull. 39, 383–388. Chen, C.C., Chen, M.J., Chiu, R.J., and Hsu, H.T. (1989). Morphological comparisons of Echinochloa ragged stunt and Rice ragged stunt viruses by electron microscopy. Phytopathology 79, 235–241. Chen, C.C., Chiu, R.J., and Wang, E.S. (1979). Rice ragged stunt: a virus disease new to Taiwan. Plant Prot. Bull (Taiwan) 21, 447. Chen, C.C., Ko, W.H., Wang, E.S., Yu, S.M., and Hu, D.Q. (1980). Epidemiological studies on the rice transitory

yellowing with special reference to its transmission by the rice green leafhoppers. TDAIS Bull (Taiwan, ROC) 4, 1–61. Chen, C.H., Hsieh, L.H., Lin, C.Y., and Hu, F.P. (1978). Preliminary report of common dwarf disease of rice. Chung-kuo nung yeh k’o hsueh. Chen, H., Zheng, L., Jia, D., Zhang, P., Chen, Q., Liu, Q., and Wei, T. (2013). Rice gall dwarf virus exploits tubules to facilitate viral spread among cultured insect vector cells derived from leafhopper Recilia dorsalis. Front Microbiol 4, 206. https://doi.org/10.3389/fmicb.2013.00206 Chen, H., Zheng, L., Mao, Q., Liu, Q., Jia, D., and Wei, T. (2014). Development of continuous cell culture of brown planthopper to trace the early infection process of oryzaviruses in insect vector cells. J. Virol. 88, 4265– 4274. https://doi.org/10.1128/JVI.03466-13 Chen, Q., Chen, H., Mao, Q., Liu, Q., Shimizu, T., UeharaIchiki, T., Wu, Z., Xie, L., Omura, T., and Wei, T. (2012). Tubular structure induced by a plant virus facilitates viral spread in its vector insect. PLOS Pathog. 8, e1003032. https://doi.org/10.1371/journal.ppat.1003032 Chen, S.X., Yuan, Y.L., King, D.D., Chen, K.Y., Lin, R.T., and Kao, T.M. (1979). Studies on the development and epiphytotics of the rice yellow stunt disease. Acta Phytopathol. Sin. 9, 41–71. Chen, Y., Lu, C., Li, M., Wu, W., Zhou, G., and Wei, T. (2016). Adverse effects of Rice gall dwarf virus upon its insect vector Recilia dorsalis (Hemiptera: Cicadellidae). Plant Dis. 100, 784–790. Ching-Chung, C. (1982). Three symptomatologic types of rice virus diseases related to grassy stunt in Taiwan. Plant Dis. 66, 15. Chiu, R.J., and Jean, J.H. (1969). Leafhopper transmission of transitory yellowing of rice. In The Virus Diseases of the Rice Plant ( Johns Hopkins Press, Baltimore, MD), pp. 131–137. Chiu, R.J., Lo, T.-C., Pi, C.L., and Chen, M.H. (1965). Transitory yellowing of Rice and its transmission by the leaf hopper Nephotettix apicalis apicalis (Motsch). Bot. Bull. Acad. Sin. Taipei 6, 1–18. Cuong, H.V., Hai, N.V., Man, V.T., and Matsumoto, M. (2009). Rice dwarf disease in North Vietnam in 2009 is caused by Southern Rice black-streaked dwarf virus (SRBSDV). Bull. Inst. Trop. Agric. Kyushu University 32, 85–92. Dai, S., Wei, X., Alfonso, A.A., Pei, L., Duque, U.G., Zhang, Z., Babb, G.M., and Beachy, R.N. (2008). Transgenic rice plants that overexpress transcription factors RF2a and RF2b are tolerant to rice tungro virus replication and disease. Proc. Natl. Acad. Sci. U.S.A. 105, 21012–21016. https://doi.org/10.1073/pnas.0810303105 Dai, S., Zhang, Z., Bick, J., and Beachy, R.N. (2006). Essential role of the Box II cis element and cognate host factors in regulating the promoter of Rice tungro bacilliform virus. J. Gen. Virol. 87, 715–722. Dietzgen, R.G., Calisher, C.H., Kurath, G., Kuzmin, I.V., Rodriguez, L.L., Stone, D.M., Tesh, R.B., Tordo, N., Walker, P.J., and Wetzel, T. (2011). Rhabdoviridae. Everett, T.R., and Lamey, H.A. (1969). Hoja blanca. In Viruses, Vectors and Vegetation, Maramorosch, K. ( John Wiley & Sons, New York, NY), 361– 377. Faan, H.C., and Piu, W.Y. (1980). A preliminary investigation on the affecting factors and disease management of rice

Viruses Infecting Rice and their Transgenic Control |  193

yellow stunt disease in Kwangtung. J. South China Agric. Coll. 1, 1–15. Falk, B.W and Tsai, J.H. (1998). Biology and molecular biology of viruses in the genus Tenuivirus. Annu. Rev. Phytopathol. 36, 139–163. https://doi.org/10.1146/ annurev.phyto.36.1.139 Fan, H., Zeng, D.S., Ye, B.J., and Lu, B.C. (1965). The preliminary investigation on rice yellow stunt disease in Guangdong. Plant Prot 3, 143–145. Fan, H., Zhang, S.G., He, X.Z., Xie, S., Liu, C., Zhou, L., Zhu, D., and Liu, X.R. (1983). Rice gall dwarf: a new virus disease epidemic in the west of Guangdong province of south China. Acta Phytopathol. Sin 13, 1–6. Fan, Z., Dahal, G., Dasgupta, I., Hay, J., and Hull, R. (1996). Variation in the genome of Rice tungro bacilliform virus: molecular characterization of six isolates. J. Gen. Virol. 77, 847–854. https://doi.org/10.1099/0022-1317-775-847 Fang, R.X., Pang, Z., Xu, B.Y., Mang, K.Q., Li, A.M., Qin, W.S., and Chen, S.X. (1992). Partial purification of rice yellow stunt virus and characterization of virus structural proteins and genome. Chin. J. Virol. 8, 62–68. Fang, R.X., Wang, Q., Xu, B.Y., Pang, Z., Zhu, H.T., Mang, K.Q., Gao, D.M., Qin, W.S., and Chua, N.H. (1994). Structure of the nucleocapsid protein gene of rice yellow stunt rhabdovirus. Virology 204, 367–375. Fang, R.-X., Zhu, H.-T., Wang, Q., Mang, K.-Q., Gao, D.-M., Qin, W.-S., Zhang, L., Cao, S.-Y., Tian, W.-Z., and Li, L.-C. (1996). Construction of transgenic rice plants resistant to rice yellow stunt virus, a plant rhabdovirus. In Third International Rice Genetics Symposium, Manila (Philippines), 16–20 October 1995 (IRRI). Fauquet, C., and Thouvenel, J.C. (1983). Presence of a new rod-shaped virus in rice plants with stripe necrotic symptoms, in Ivory Coast. Comptes Rendus Hebd. des Seances l’Academie des Sci. Ser. 3 Sci. la Vie. Fauquet, C., Thouvenel, J.-C., Fargette, D., and Fishpool, L.D.C. (1988). Rice stripe necrosis virus: a soil-borne rod-shaped virus. Dev. Appl. Biol. II. Viruses with fungal vectors, 71–82. Fütterer, J., Potrykus, I., Valles Brau, M.P., Dasgupta, I., Hull, R., and Hohn, T. (1994). Splicing in a plant pararetrovirus. Virology 198, 663–670. Galvez, G.E., and Jennings, P.H. (1959). Transmisión de la hoja blanca del arroz en Colombia. Agric. Trop. 15, 507–515. Gálvez, G.E. (1968). Purification and characterization of rice tungro virus by analytical density-gradient centrifugation. Virology 35, 418–426. Ganesan, U., Suri, S.S., Rajasubramaniam, S., Rajam, M.V., and Dasgupta, I. (2009). Transgenic expression of coat protein gene of Rice tungro bacilliform virus in rice reduces the accumulation of viral DNA in inoculated plants. Virus Genes 39, 113–119. https://doi.org/10.1007/ s11262-009-0359-9 Garces, O.C. (1940). Report of the Commission to study and present a plan of organization of projects on Plant Pathology at the Palmira Experiment Station. Rep. Comm. to study Present a plan Organ. Proj. Plant Pathol. Palmira Exp. Station. Garces-Orejuela, C., Jennings, P.R., and Skiles, R.L. (1958). Hoja blanca of Rice and the history of the disease in Colombia. Plant Dis. Report. 42, 750–751.

Ghosh, S.K. (1980). Rice necrosis mosaic. Proc. Plant Sci. 89, 291–299. Gill, C.C., and Chong, J. (1979). Cytopathological evidence for the division of Barley yellow dwarf virus isolates into two subgroups. Virology 95, 59–69. Goldbach, R. (1987). Genome similarities between plant and animal RNA viruses. Microbiol. Sci. 4, 197–202. Goldbach, R., Bucher, E., and Prins, M. (2003). Resistance mechanisms to plant viruses: an overview. Virus Res. 92, 207–212. Guo, R., Zhou, G.H., and Zhang, S.G. (2010). Character of rice Southern black-streaked disease and its control strategy. China Plant Prot 30, 17–20. Hagiwara, K., Minobe, Y., Nozu, Y., Hibino, H., Kimura, I., and Omura, T. (1986). Component proteins and structure of Rice ragged stunt virus. J. Gen. Virol. 67, 1711–1715. Hayakawa, T., Zhu, Y., Itoh, K., Kimura, Y., Izawa, T., Shimamoto, K., and Toriyama, S. (1992). Genetically engineered rice resistant to Rice stripe virus, an insecttransmitted virus. Proc. Natl. Acad. Sci. U.S.A. 89, 9865–9869. Hayano, Y., Kakutani, T., Hayashi, T., and Minobe, Y. (1990). Coding strategy of Rice stripe virus: major nonstructural protein is encoded in viral RNA segment 4 and coat protein in RNA complementary to segment 3. Virology 177, 372–374. Hemmes, H., Kaaij, L., Lohuis, D., Prins, M., Goldbach, R., and Schnettler, E. (2009). Binding of small interfering RNA molecules is crucial for RNA interference suppressor activity of Rice hoja Blanca virus NS3 in plants. J. Gen. Virol. 90, 1762–1766. Hemmes, H., Lakatos, L., Goldbach, R., Burgyán, J., and Prins, M. (2007). The NS3 protein of Rice hoja blanca Tenuivirus suppresses RNA silencing in plant and insect hosts by efficiently binding both siRNAs and miRNAs. RNA 13, 1079–1089. Herdt, R.W. (1991). Research priorities for rice biotechnology. Rice Biotechnol. 6, 19–54. Hibi, T., Omura, T., and Saito, Y. (1984). Double-stranded RNA of Rice gall dwarf virus. J. Gen. Virol. 65, 1585– 1590. Hibino, H. (1979). Rice ragged stunt, a new virus disease occurring in tropical Asia. Rev. Plant Prot. Res. 12, 98–110. Hibino, H. (1996). Biology and epidemiology of rice viruses. Annu. Rev. Phytopathol. 34, 249–274. https:// doi.org/10.1146/annurev.phyto.34.1.249 Hibino, H. (1990). Insect-borne viruses of rice. In Advances in disease vector research (Springer), pp. 209–241. Hibino, H. (1986). Rice grassy stunt virus. Trop. Agric. Res. Ser. 19, 165–171. Hibino, H., Usugi, T., Omura, T., Tsuchizaki, T., Shohara, K., and Iwasaki, M. (1985). Rice grassy stunt virus: a planthopper-borne circular filament. Phytopathology 75, 894–899. Hiraguri, A., Hibino, H., Hayashi, T., Shimizu, T., UeharaIchiki, T., Omura, T., and Sasaya, T. (2010). Complete sequence analysis of Rice transitory yellowing virus and its comparison to rice yellow stunt virus. Arch. Virol. 155, 243–245. https://doi.org/10.1007/s00705-009-05578.

194  | Kumar et al.

Hiraguri, A., Netsu, O., Shimizu, T., Uehara-Ichiki, T., Omura, T., Sasaki, N., Nyunoya, H., and Sasaya, T. (2011). The nonstructural protein pC6 of Rice grassy stunt virus trans-complements the cell-to-cell spread of a movement-defective Tomato mosaic virus. Arch. Virol. 156, 911–916. https://doi.org/10.1007/s00705-0110939-6 Hoang, A.T., Zhang, H., Yang, J., Chen, J., Hébrard, E., Zhou, G., Vinh, V.N., and Cheng, J. (2011). Identification, characterization, and distribution of Southern Rice black-streaked dwarf virus in Vietnam. Plant Dis. 95, 1063–1069. Hsu, H.T., Hibino, H., and Cabauatan, P.Q. (1990). Development of serological procedures for rapid, sensitive, and reliable detection of Rice grassy stunt virus. Plant Dis. 74, 695–698. Huet, H., Mahendra, S., Wang, J., Sivamani, E., Ong, C.A., Chen, L., de Kochko, A., Beachy, R.N., and Fauquet, C. (1999). Near immunity to Rice tungro spherical virus achieved in rice by a replicase-mediated resistance strategy. Phytopathology 89, 1022–1027. https://doi. org/10.1094/PHYTO.1999.89.11.1022 Iida, T.T., Shinkai, A., and Kimura, I. (1972). Rice dwarf virus. CMI/AAB Descr. Pl. Viruses 102. Inoue, H., and Omura, T. (1982). Transmission of Rice gall dwarf virus by the green rice leafhopper. Plant Dis. 66, 57–59. Inoue, H., Omura, T., Hayashi, T., Morinaka, T., Putta, M., Chettanachit, D., Parejarearn, A., Disthaaporn, S., and Kadokao, S. (1986). Rice transitory yellowing virus. Virus Dis. Rice Legum. Trop. (Technical Bull. Trop. Agric. Res. Cent. Japan vol. 21), 65–77. Inouye, T., and Fujii, S. (1977). Rice necrosis mosaic virus. CMI/AAB Descr. Pl. Viruses 172. Ishikawa, K., Omura, T., and Hibino, H. (1989). Morphological characteristics of Rice stripe virus. J. Gen. Virol. 70, 3465–3468. Isogai, M., Cabauatan, P.Q., Masuta, C., Uyeda, I., and Azzam, O. (2000). Complete nucleotide sequence of the Rice tungro spherical virus genome of the highly virulent strain Vt6. Virus Genes 20, 79–85. Iwasaki, M., Nakano, M., and Shinkai, A. (1985). Detection of Rice grassy stunt virus in planthopper vectors and rice plants by ELISA. Japanese J. Phytopathol. 51, 450–458. Jennings, P.R., and Pineda, T. (1970). Screening rice for resistance to the planthopper, Sogatodes oryzicola (Muir). Crop Sci. 10, 687–689. Jennings, P.R., Rosero, M.J., and Uribe, L.J.B. (1958). Hoja blanca del arroz (Departamento de Investigación Agropecuaria). Jia, D., Chen, H., Zheng, A., Chen, Q., Liu, Q., Xie, L., Wu, Z., and Wei, T. (2012). Development of an insect vector cell culture and RNA interference system to investigate the functional role of Fijivirus replication protein. J. Virol. 86, 5800–5807. https://doi.org/10.1128/JVI.07121-11 Jia, D., Guo, N., Chen, H., Akita, F., Xie, L., Omura, T., and Wei, T. (2012). Assembly of the viroplasm by viral non-structural protein Pns10 is essential for persistent infection of Rice ragged stunt virus in its insect vector. J. Gen. Virol. 93, 2299–2309. Jia, D., Mao, Q., Chen, H., Wang, A., Liu, Y., Wang, H., Xie, L., and Wei, T. (2014). Virus-induced tubule: a vehicle for rapid spread of virions through basal lamina from

midgut epithelium in the insect vector. J. Virol. 88, 10488–10500. https://doi.org/10.1128/JVI.01261-14 Jiang, L., Qian, D., Zheng, H., Meng, L.Y., Chen, J., Le, W.J., Zhou, T., Zhou, Y.J., Wei, C.H., and Li, Y. (2012). RNAdependent RNA polymerase 6 of rice (Oryza sativa) plays role in host defense against negative-strand RNA virus, Rice stripe virus. Virus Res. 163, 512–519. https:// doi.org/10.1016/j.virusres.2011.11.016 Jyothsna, M., Manonmani, S., Rabindran, R., Dasgupta, I., and Robin, S. (2013). Introgression of transgenic resistance for rice tungro disease into mega variety, ASD 16 of Tamil Nadu through marker assisted backcross breeding. Madras Agric. J. 100, 70–74. Kakutani, T., Hayano, Y., Hayashi, T., and Minobe, Y. (1991). Ambisense segment 3 of Rice stripe virus: the first instance of a virus containing two ambisense segments. J. Gen. Virol. 72, 465–468. https://doi. org/10.1099/0022-1317-72-2-465 Koganezawa, H., Hibino, H., Motoyoshi, F., Kato, H., Noda, H., Ishikawa, K., and Omura, T. (1990). Nucleotide sequence of segment S9 of the genome of Rice gall dwarf virus. J. Gen. Virol. 71, 1861–1863. https://doi. org/10.1099/0022-1317-71-8-1861 Konate, G., Traore, O., and Coulibaly, M.M. (1997). Characterization of Rice yellow mottle virus isolates in Sudano-Sahelian areas. Arch. Virol. 142, 1117–1124. Kormelink, R., Garcia, M.L., Goodin, M., Sasaya, T., and Haenni, A.L. (2011). Negative-strand RNA viruses: the plant-infecting counterparts. Virus Res. 162, 184–202. https://doi.org/10.1016/j.virusres.2011.09.028 Kouassi, N.K., Chen, L., Siré, C., Bangratz-Reyser, M., Beachy, R.N., Fauquet, C.M., and Brugidou, C. (2006). Expression of Rice yellow mottle virus coat protein enhances virus infection in transgenic plants. Arch. Virol. 151, 2111–2122. https://doi.org/10.1007/ s00705-006-0802-3 Kouassi, N., Brugidou, C., Chen, L., Ngon, M., Yassi, A., Beachy, R.N., and Fauquet, C.M. (1997). Transgenic rice plants expressing Rice yellow mottle virus coat protein gene. Int. Rice Res. Notes 22, 14–15. Kulshreshtha, J.P., Anjaneyulu, A., and Padmanabhan, S.Y. (1974). Disastrous brown plant-hopper attack in Kerala. Indian Farming. ISSN No. 0019-4786 Lakatos, L., Csorba, T., Pantaleo, V., Chapman, E.J., Carrington, J.C., Liu, Y.P., Dolja, V.V., Calvino, L.F., López-Moya, J.J., and Burgyán, J. (2006). Small RNA binding is a common strategy to suppress RNA silencing by several viral suppressors. EMBO J. 25, 2768–2780. Lamey, H.A. (1969). Field testing. In The Virus Diseases of the Rice Plant: Proceedings of a Symposium at the International Rice Research Institute, April 1967, p. 293. Lamey, H.A., and Everett, T.R. (1967). Increased susceptibility of hoja blanca virus-infected rice leaves to Cochliobolus miyabeanus. Phytopathology 57, 227. Le, D.T., Chu, H.D., and Sasaya, T. (2015). Creation of transgenic rice plants producing small interfering RNA of Rice tungro spherical virus. GM Crops Food 6, 47–53. https://doi.org/10.1080/21645698.2015.1025188 Lee, B.C., Yoon, Y.N., Hong, S.J., Hong, Y.K., Kwak, D.Y., Lee, J.H., Yae, U.S., Kang, H.W., and Hwang, H.G. (2008). Analysis on the Occurrence of Rice stripe virus. Res. Plant Dis. 14, 210–213.

Viruses Infecting Rice and their Transgenic Control |  195

Li, J., Xue, J., Zhang, H.M., Yang, J., Lv, M.F., Xie, L., Meng, Y., Li, P.P., and Chen, J.P. (2013). Interactions between the P6 and P5-1 proteins of southern rice black-streaked dwarf Fijivirus in yeast and plant cells. Arch. Virol. 158, 1649–1659. https://doi.org/10.1007/s00705-0131660-4 Li, X., Zhang, W., Ding, Y., Wang, Z., Wu, Z., Yu, L., Hu, D., Li, P., and Song, B. (2015). Characterization of the importance of terminal residues for Southern Rice blackstreaked dwarf virus P9-1 viroplasm formations. Protein Expr. Purif. 111, 98–104. https://doi.org/10.1016/j. pep.2015.04.003 Li, Y., Bao, Y.M., Wei, C.H., Kang, Z.S., Zhong, Y.W., Mao, P., Wu, G., Chen, Z.L., Schiemann, J., and Nelson, R.S. (2004). Rice dwarf Phytoreovirus segment S6-encoded nonstructural protein has a cell-to-cell movement function. J. Virol. 78, 5382–5389. Lian, S., Jonson, M.G., Cho, W.K., Choi, H.S., Je, Y.H., and Kim, K.H. (2011). Generation of antibodies against Rice stripe virus proteins based on recombinant proteins and synthetic polypeptides. Plant Pathol. J. 27, 37–43. Lishi, L.Q.X.L.X., and Youyi, L.X.W. (1994). Studies on Rice Bunchy Stunt VII. Spectral properties of RBSV. Acta Phytopathol. Sin. 1, 1. Liu, S., Ding, Z., Zhang, C., Yang, B., and Liu, Z. (2010). Gene knockdown by intro-thoracic injection of doublestranded RNA in the brown planthopper, Nilaparvata lugens. Insect Biochem. Mol. Biol. 40, 666–671. https:// doi.org/10.1016/j.ibmb.2010.06.007 Liu, Y., Jia, D., Chen, H., Chen, Q., Xie, L., Wu, Z., and Wei, T. (2011). The P7-1 protein of Southern Rice black-streaked dwarf virus, a Fijivirus, induces the formation of tubular structures in insect cells. Arch. Virol. 156, 1729–1736. https://doi.org/10.1007/s00705-011-1041-9 Liyan, L.Q.X.L.X. (1991). Studies on the purification and properties of Rice bunchy stunt virus [ J]. Sci. Agric. Sin. 4, 8. Lockhart, B.E.L. (1990). Evidence for a double-stranded circular DNA genome in a second group of plant viruses. Phytopathology 80, 127–131. Louvel, D., and Bidaux, J.M. (1977). Observations de nouveaux symptômes pathologiques sur des variétés précoces de riz en Côte d’Ivoire. L’Agronomie Trop. 32, 257–261. Lu, G., Zhou, Z.H., Baker, M.L., Jakana, J., Cai, D., Wei, X., Chen, S., Gu, X., and Chiu, W. (1998). Structure of double-shelled Rice dwarf virus. J. Virol. 72, 8541–8549. Luan, S., Lan, W., and Chul Lee, S. (2009). Potassium nutrition, sodium toxicity, and calcium signaling: connections through the CBL-CIPK network. Curr. Opin. Plant Biol. 12, 339–346. https://doi. org/10.1016/j.pbi.2009.05.003 Ma, J., Song, Y., Wu, B., Jiang, M., Li, K., Zhu, C., and Wen, F. (2011). Production of transgenic rice new germplasm with strong resistance to two isolations of Rice stripe virus by RNA interference. Transgenic Res. 20, 1367–1377. https://doi.org/10.1007/s11248-011-9502-1 Malaguti, G., Diaz, H., and Angeles, N. (1957). La virosis ‘hoja blanca’ del arroz. Agron. Trop 6, 157–163. Mao, Z.J., Li, Y., Xu, H., Zheng, H.H., Schiemann, J., Casper, R., and Chen, Z.L. (1998). The 42K protein of Rice dwarf virus is a post-translational cleavage product of the 46K outer capsid protein. Arch. Virol. 143, 1831–1838.

Mariappan, V., Hibino, H., and Shanmugam, N. (1984). A new rice virus disease in India. Int. Rice Res. Newsl. 9, 9–10. Matsukura, K., Towata, T., Sakai, J., Onuki, M., Okuda, M., and Matsumura, M. (2013). Dynamics of Southern Rice black-streaked dwarf virus in rice and implication for virus acquisition. Phytopathology 103, 509–512. https://doi. org/10.1094/PHYTO-10-12-0261-R Mazzini, F., Moletti, M., and Feccia, S. (1990). Level of some nutrients in Rice giallume virus infected plants. In World Perspectives on Barley Yellow Dwarf International Workshop, Udine (Italy), 6–11 Jul 1987 (CIMMYT). McGuire, J.U., MCMILLAN, W.W., and Lamey, H.A. (1960). Hoja blanca disease of Rice and its insect vector. Rice J. 63, 15–28. Milne, R.G., Boccardo, G., and Ling, K.C. (1982). Rice ragged stunt virus. C. Descr. Plant Viruses 16, 248. Miyazaki, N., Higashiura, A., Higashiura, T., Akita, F., Hibino, H., Omura, T., Nakagawa, A., and Iwasaki, K. (2016). Electron microscopic imaging revealed the flexible filamentous structure of the cell attachment protein P2 of Rice dwarf virus located around the icosahedral 5-fold axes. J. Biochem. 159, 181–190. https://doi.org/10.1093/jb/mvv092 Miyazaki, N., Wu, B., Hagiwara, K., Wang, C.Y., Xing, L., Hammar, L., Higashiura, A., Tsukihara, T., Nakagawa, A., Omura, T., et al. (2010). The functional organization of the internal components of Rice dwarf virus. J. Biochem. 147, 843–850. https://doi.org/10.1093/jb/mvq017 Mizuno, H., Kano, H., Omura, T., Koizumi, M., Kondoh, M., and Tsukihara, T. (1991). Crystallization and preliminary X-ray study of a double-shelled spherical virus, Rice dwarf virus. J. Mol. Biol. 219, 665–669. Morales, F.J., and Jennings, P.R. (2011). Rice hoja blanca: a complex plant–virus–vector pathosystem. Plant Sci. Rev. 2010, 163. Morales, F.J., and Niessen, A.I. (1985). Rice hoja Blanca virus. AAB Descr. Pl. Viruses 299. Morales, F.J., and Niessen, A.I. (1983). Association of spiral filamentous viruslike particles with rice hoja blanca. Phytopathology 73, 971–974. Morinaka, T., Putta, M., Chettanachit, D., Parejarearn, A., Disthaporn, S., Omura, T., and Inoue, H. (1982). Transmission of Rice gall dwarf virus by cicadellid leafhoppers Recilia dorsalis and Nephotettix nigropictus in Thailand. Plant Dis. 66, 703–704. Moriyasu, Y., Maruyama-Funatsuki, W., Kikuchi, A., Ichimi, K., Zhong, B., Yan, J., Zhu, Y., Suga, H., Watanabe, Y., Ichiki-Uehara, T., et al. (2007). Molecular analysis of the genome segments S1, S4, S6, S7 and S12 of a Rice gall dwarf virus isolate from Thailand; completion of the genomic sequence. Arch. Virol. 152, 1315–1322. https://doi.org/10.1007/s00705-007-0948-7 Moriyasu, Y., Ishikawa, K., Kikuchi, A., Imanishi, S., Tomita, S., Akutsu, K., and Omura, T. (2000). Sequence analysis of Pns11, a nonstructural protein of Rice gall dwarf virus, and its expression and detection in infected rice plants and vector insects. Virus Genes 20, 237–241. Naitow, H., Morimoto, Y., Mizuno, H., Kano, H., Omura, T., Koizumi, M., and Tsukihara, T. (1999). A low-resolution structure of Rice dwarf virus determined by ab initio phasing. Acta Crystallogr. Sect. D Biol. Crystallogr. 55, 77–84.

196  | Kumar et al.

Nakagawa, A., Miyazaki, N., Taka, J., Naitow, H., Ogawa, A., Fujimoto, Z., Mizuno, H., Higashi, T., Watanabe, Y., Omura, T., et al. (2003). The atomic structure of Rice dwarf virus reveals the self-assembly mechanism of component proteins. Structure 11, 1227–1238. Nakahara, K.S., Masuta, C., Yamada, S., Shimura, H., Kashihara, Y., Wada, T.S., Meguro, A., Goto, K., Tadamura, K., Sueda, K., et al. (2012). Tobacco calmodulin-like protein provides secondary defense by binding to and directing degradation of virus RNA silencing suppressors. Proc. Natl. Acad. Sci. U.S.A. 109, 10113–10118. https://doi.org/10.1073/ pnas.1201628109 Nath, N., Mathur, S., and Dasgupta, I. (2002). Molecular analysis of two complete Rice tungro bacilliform virus genomic sequences from India. Arch. Virol. 147, 1173– 1187. https://doi.org/10.1007/s00705-002-0801-y Netsu, O., Hiraguri, A., Uehara-Ichiki, T., Komatsu, K., and Sasaya, T. (2015). Functional comparison of RNA silencing suppressor between the p5 protein of Rice grassy stunt virus and the p3 protein of Rice stripe virus. Virus Res. 203, 10–19. https://doi.org/10.1016/j. virusres.2015.03.010 Nguyen, T.D., Lacombe, S., Bangratz, M., Ta, H.A., Vinh, d.o.N., Gantet, P., and Brugidou, C. (2015). P2 of Rice grassy stunt virus (RGSV) and p6 and p9 of Rice ragged stunt virus (RRSV) isolates from Vietnam exert suppressor activity on the RNA silencing pathway. Virus Genes 51, 267–275. https://doi.org/10.1007/s11262015-1229-2 Noda, H., Ishikawa, K., Hibino, H., Kato, H., and Omura, T. (1991). Nucleotide sequences of genome segments S8, encoding a capsid protein, and S10, encoding a 36K protein, of Rice gall dwarf virus. J. Gen. Virol. 72, 2837– 2842. https://doi.org/10.1099/0022-1317-72-11-2837 Omura, T. (1980). Observations on rice gall dwarf, a new virus disease. Int. Rice Res. Newsl. 5, 11–12. Omura, T., and Inoue, H. (1985). Rice gall dwarf virus. CMI/ AAB Descr. Pl. Viruses 299. Omura, T., Morinaka, T., Inoue, H., and Saito, Y. (1982). Purification and some properties of Rice gall dwarf virus, a new Phytoreovirus. Phytopathology 72, 241–246. Omura, T., and Yan, J. (1999). Role of outer capsid proteins in transmission of Phytoreovirus by insect vectors. Adv. Virus Res. 54, 15–43. Omura, T., Yan, J., Zhong, B., Wada, M., Zhu, Y., Tomaru, M., Maruyama, W., Kikuchi, A., Watanabe, Y., Kimura, I., et al. (1998). The P2 protein of rice dwarf Phytoreovirus is required for adsorption of the virus to cells of the insect vector. J. Virol. 72, 9370–9373. Osler, R. (1984). Rice giallume, a disease related to barley yellow dwarf in Italy. In Barley yellow dwarf, a proceedings of the workshop December 6-8, 1983 CIMMYT Mexico, sponsored by the United Nations Development Programme and CIMMYT (Centro Internacional de Mejoramiento de Maíz y Trigo), pp. 125–131. Osler, R., Amici, A., and Belli, G. (1974). Transmission of Rice ‘Giallume’ by an aphid, Rhopalosiphum padi. Riv. di Patol. Veg. 10, 5–17. Park, H.M., Choi, M.S., Kwak, D.Y., Lee, B.C., Lee, J.H., Kim, M.K., Kim, Y.G., Shin, D.B., Park, S.K., and Kim, Y.H. (2012). Suppression of NS3 and MP is important

for the stable inheritance of RNAi-mediated Rice stripe virus (RSV) resistance obtained by targeting the fully complementary RSV-CP gene. Mol. Cells 33, 43–51. https://doi.org/10.1007/s10059-012-2185-5 Pinto, Y.M., Kok, R.A., and Baulcombe, D.C. (1999). Resistance to Rice yellow mottle virus (RYMV) in cultivated African rice varieties containing RYMV transgenes. Nat. Biotechnol. 17, 702–707. https://doi. org/10.1038/10917 Qiliang, X.L.L.Q.Z. (1982). On the Bunchy stunt disease of rice II: The distribution, loss, hosts and overwintering of Rice bunchy stunt [ J]. Acta Phytopathol. Sin. 4, 1. Qu, R.D., Bhattacharyya, M., Laco, G.S., De Kochko, A., Rao, B.L., Kaniewska, M.B., Elmer, J.S., Rochester, D.E., Smith, C.E., and Beachy, R.N. (1991). Characterization of the genome of Rice tungro bacilliform virus: comparison with Commelina yellow mottle virus and caulimoviruses. Virology 185, 354–364. Ramirez, B.C., Macaya, G., Calvert, L.A., and Haenni, A.L. (1992). Rice hoja Blanca virus genome characterization and expression in vitro. J. Gen. Virol. 73, 1457–1464. https://doi.org/10.1099/0022-1317-73-6-1457 Raychaudhuri, S.P., Mishra, M.D., and Ghosh, A. (1967). Preliminary note on transmission of virus disease resembling tungro of rice in India and other virus-like symptoms. Plant Dis. Rep. 51, 300–301. Rivera, C.T., and Ou, S.H. (1967). Transmission studies of the two strains of rice tungro virus. Plant Dis. Rep. 51, 877–881. Rivera, C.T., and Ou, S.H. (1967). Strains of rice tungro virus. Philipp. Phytopathol. 3, 19–20. Rivera, C.T., and Ou, S.H. (1965). Leafhopper transmission of ‘tungro’ disease of rice. Plant Dis. Report. 49. Rivera, C.T., Ou, S.H., and Iida, T.T. (1966). Grassy stunt disease of Rice and its transmission by the planthopper Nilaparvata lugens Stal. Plant Dis. Rep. 50, 453–456. Romero, L.E., Lozano, I., Garavito, A., Carabali, S.J., Triana, M., Villareal, N., Reyes, L., Duque, M.C., Martinez, C.P., Calvert, L., et al. (2014). Major QTLs control resistance to Rice hoja Blanca virus and its vector Tagosodes orizicolus. G3: Genes, Genomes, Genetics 4, 133–142. https://doi.org/10.1534/g3.113.009373 Rossel, H.W., Thottapilly, G., and Buddenhagen, I.W. (1982). Occurrence of Rice yellow mottle virus in two important rice-growing areas of Nigeria. FAO Plant Prot. Bull. 30, 137–139. Roy, S., Banerjee, A., Tarafdar, J., Senapati, B.K., and Dasgupta, I. (2012). Transfer of transgenes for resistance to rice tungro disease into high-yielding rice cultivars through gene-based marker-assisted selection. J. Agric. Sci. 150, 610–618. Sanfaçon, H., Wellink, J., Le Gall, O., Karasev, A., van der Vlugt, R., and Wetzel, T. (2009). Secoviridae: a proposed family of plant viruses within the order Picornavirales that combines the families Sequiviridae and Comoviridae, the unassigned genera Cheravirus and Sadwavirus, and the proposed genus Torradovirus. Arch. Virol. 154, 899–907. https://doi.org/10.1007/s00705009-0367-z Sarra, S., and Peters, D. (2003). Rice yellow mottle virus is transmitted by cows, donkeys, and grass rats in irrigated rice crops. Plant Dis. 87, 804–808.

Viruses Infecting Rice and their Transgenic Control |  197

Sasaya, T., Nakazono-Nagaoka, E., Saika, H., Aoki, H., Hiraguri, A., Netsu, O., Uehara-Ichiki, T., Onuki, M., Toki, S., Saito, K., et al. (2014). Transgenic strategies to confer resistance to viruses in rice plants. Front. Microbiol. 4, 409. https://doi.org/10.3389/ fmicb.2013.00409 Sharma, S., Rabindran, R., Robin, S., and Dasgupta, I. (2011). Analysis of the complete DNA sequence of Rice tungro bacilliform virus from southern India indicates it to be a product of recombination. Arch. Virol. 156, 2257–2262. https://doi.org/10.1007/s00705-0111092-y Shikata, E. (1972). Rice transitory yellowing virus. C. Descr. Plant Viruses. Shimizu, T., Nakazono-Nagaoka, E., Akita, F., Wei, T., Sasaya, T., Omura, T., and Uehara-Ichiki, T. (2012). Hairpin RNA derived from the gene for Pns9, a viroplasm matrix protein of Rice gall dwarf virus, confers strong resistance to virus infection in transgenic rice plants. J. Biotechnol. 157, 421–427. https://doi.org/10.1016/j. jbiotec.2011.12.015 Shimizu, T., Nakazono-Nagaoka, E., Uehara-Ichiki, T., Sasaya, T., and Omura, T. (2011). Targeting specific genes for RNA interference is crucial to the development of strong resistance to Rice stripe virus. Plant Biotechnol. J. 9, 503–512. https://doi.org/10.1111/j.14677652.2010.00571.x Shimizu, T., Ogamino, T., Hiraguri, A., Nakazono-Nagaoka, E., Uehara-Ichiki, T., Nakajima, M., Akutsu, K., Omura, T., and Sasaya, T. (2013). Strong resistance to Rice grassy stunt virus is induced in transgenic rice plants expressing double-stranded RNA of the viral genes for nucleocapsid or movement proteins as targets for RNA interference. Phytopathology 103, 513–519. https:// doi.org/10.1094/PHYTO-07-12-0165-R Shimizu, T., Toriyama, S., Takahashi, M., Akutsu, K., and Yoneyama, K. (1996). Non-viral sequences at the 5’ termini of mRNAs derived from virus-sense and viruscomplementary sequences of the ambisense RNA segments of rice stripe Tenuivirus. J. Gen. Virol. 77, 541– 546. https://doi.org/10.1099/0022-1317-77-3-541 Shimizu, T., Yoshii, M., Wei, T., Hirochika, H., and Omura, T. (2009). Silencing by RNAi of the gene for Pns12, a viroplasm matrix protein of Rice dwarf virus, results in strong resistance of transgenic rice plants to the virus. Plant Biotechnol. J. 7, 24–32. https://doi.org/10.1111/ j.1467-7652.2008.00366.x Shinkai, A., Nakano, M., and Iwasai, M. (1980). Occurrence of rice ragged stunt disease in Kyushu, Japan. Ann. Phytopathol. Soc. Jpn 46, 411. Shirako, Y., Falk, B.W., and Haenni, A.L. (2011). Genus Tenuivirus. Virus Taxon. Classif. Nomencl. Ninth Rep. Int. Commitee Taxon. Viruses, 771–776. Sivamani, E., Huet, H., Shen, P., Ong, C.A., de Kochko, A., Fauquet, C., and Beachy, R.N. (1999). Rice plant (Oryza sativa L.) containing Rice tungro spherical virus (RTSV) coat protein transgenes are resistant to virus infection. Mol. Breed. 5, 177–185. Songbai, Z., Zhenguo, D., Liang, Y., Zhengjie, Y., Kangcheng, W., Guangpu, L., Zujian, W., and Lianhui, X. (2013). Identification and characterization of the interaction between viroplasm-associated proteins from two different plant-infecting reoviruses and eEF-1A of rice.

Arch. Virol. 158, 2031–2039. https://doi.org/10.1007/ s00705-013-1703-x Suzuki, N. (1993). In vitro translation of rice dwarf Phytoreovirus genome segments S4 to S10. Arch. Virol. 130, 201–208. Suzuki, N., Harada, M., and Kusano, T. (1991). Molecular analysis of rice dwarf Phytoreovirus segment S11 corresponding to wound tumour Phytoreovirus segment S12. J. Gen. Virol. 72, 2233–2237. https://doi. org/10.1099/0022-1317-72-9-2233 Suzuki, N., Kusano, T., Matsuura, Y., and Omura, T. (1996). Novel NTP binding property of rice dwarf Phytoreovirus minor core protein P5. Virology 219, 471–474. Suzuki, N., Sugawara, M., and Kusano, T. (1992). Rice dwarf Phytoreovirus segment S12 transcript is tricistronic in vitro. Virology 191, 992–995. Suzuki, N., Sugawara, M., Kusano, T., Mori, H., and Matsuura, Y. (1994). Immunodetection of rice dwarf phytoreoviral proteins in both insect and plant hosts. Virology 202, 41–48. Suzuki, N., Tanimura, M., Watanabe, Y., Kusano, T., Kitagawa, Y., Suda, N., Kudo, H., Uyeda, I., and Shikata, E. (1992). Molecular analysis of rice dwarf Phytoreovirus segment S1: interviral homology of the putative RNA-dependent RNA polymerase between plant and animal-infecting reoviruses. Virology 190, 240–247. Suzuki, N., Watanabe, Y., Kusano, T., and Kitagawa, Y. (1990). Sequence analysis of rice dwarf Phytoreovirus genome segments S4, S5, and S6: comparison with the equivalent wound tumor virus segments. Virology 179, 446–454. Suzuki, N., Watanabe, Y., Kusano, T., and Kitagawa, Y. (1990). Sequence analysis of the rice dwarf Phytoreovirus segment S3 transcript encoding for the major structural core protein of 114 kDa. Virology 179, 455–459. Takahashi, M., Toriyama, S., Hamamatsu, C., and Ishihama, A. (1993). Nucleotide sequence and possible ambisense coding strategy of Rice stripe virus RNA segment 2. J. Gen. Virol. 74, 769–773. https://doi.org/10.1099/00221317-74-4-769 Takahashi, M., Toriyama, S., Kikuchi, Y., Hayakawa, T., and Ishihama, A. (1990). Complementarity between the 5′- and 3′-terminal sequences of Rice stripe virus RNAs. J. Gen. Virol. 71, 2817–2821. https://doi. org/10.1099/0022-1317-71-12-2817 Tomaru, M., Maruyama, W., Kikuchi, A., Yan, J., Zhu, Y., Suzuki, N., Isogai, M., Oguma, Y., Kimura, I., and Omura, T. (1997). The loss of outer capsid protein P2 results in nontransmissibility by the insect vector of rice dwarf Phytoreovirus. J. Virol. 71, 8019–8023. Toriyama, S. (1985). Purification and biochemical properties of Rice grassy stunt virus. Ann. Phytopathol. Soc. Japan 51, 59. Toriyama, S., Kimishima, T., and Takahashi, M. (1997). The proteins encoded by Rice grassy stunt virus RNA5 and RNA6 are only distantly related to the corresponding proteins of other members of the genus Tenuivirus. J. Gen. Virol. 78, 2355–2363. https://doi.org/10.1099/00221317-78-9-2355 Toriyama, S., Takahashi, M., Sano, Y., Shimizu, T., and Ishihama, A. (1994). Nucleotide sequence of RNA 1, the largest genomic segment of Rice stripe virus, the prototype of the Tenuiviruses. J. Gen. Virol. 75,

198  | Kumar et al.

3569–3579. https://doi.org/10.1099/0022-1317-7512-3569 Traoré, O., Pinel, A., Hébrard, E., Dieudonné Gumedzoé, M.Y., Fargette, D., Traoré, A.S., and Konaté, G. (2006). Occurrence of resistance-breaking isolates of Rice yellow mottle virus in West and Central Africa. Plant Dis. 90, 259–263. Tyagi, H., Rajasubramaniam, S., Rajam, M.V., and Dasgupta, I. (2008). RNA-interference in rice against Rice tungro bacilliform virus results in its decreased accumulation in inoculated rice plants. Transgenic Res. 17, 897–904. https://doi.org/10.1007/s11248-008-9174-7 Upadhyaya, N.M., Li, Z., Ramm, K., Gaudron, J., Waterhouse, P.M., Kositratana, W., Wang, M.-B., and Gong, Z.-X. (1997). Production of transgenic rice with Rice ragged stunt virus synthetic resistance genes. In International Symposium on Biotechnology of Tropical and Subtropical Species Part 2 461, pp. 393–400. Upadhyaya, N.M., Li, Z., Wang, M.B., Chen, S., Gong, Z.X., and Waterhouse, P.M. (2001). Engineering for virus resistance in rice. Rice Genet. IV, 405. Upadhyaya, N.M., Ramm, K., Gellatly, J.A., Li, Z., Kositratana, W., and Waterhouse, P.M. (1998). Rice ragged stunt oryzavirus genome segment S4 could encode an RNA dependent RNA polymerase and a second protein of unknown function. Arch. Virol. 143, 1815–1822. Upadhyaya, N.M., Ramm, K., Yang, M., Kositratana, W., and Waterhouse, P.M. (1996). Rice ragged stunt virus synthetic resistance genes and japonica rice transformation. In Third International Rice Genetics Symposium, Manila (Philippines), 16-20 Oct 1995 (IRRI). Upadhyaya, N.M., Yang, M., Kositratana, W., Ghosh, A., and Waterhouse, P.M. (1995). Molecular analysis of rice ragged stunt oryzavirus segment 9 and sequence conservation among isolates from Thailand and India. Arch. Virol. 140, 1945–1956. Usugi, T., Kashiwazaki, S., Omura, T., and Tsuchizaki, T. (1989). Some properties of nucleic acids and coat proteins of soil-borne filamentous viruses. Japanese J. Phytopathol. 55, 26–31. Usugi, T., and Saito, Y. (1976). Purification and serological properties of barley yellow mosaic virus and wheat yellow mosaic virus. Japanese J. Phytopathol. 42, 12–20. Uyeda, I., Kimura, I., and Shikata, E. (1995). Characterization of genome structure and establishment of vector cell lines for plant reoviruses. Adv. Virus Res. 45, 249–279. Uyeda, I., and Milne, R.G. (1995). Introduction: Genomic organization, diversity and evolution of plant reoviruses. In Seminars in virology (Academic Press), pp. 85–88. Uyeda, I., Suda, N., Yamada, N., Kudo, H., Murao, K., Suga, H., Kimura, I., Shikata, E., Kitagawa, Y., and Kusano, T. (1994). Nucleotide sequence of rice dwarf Phytoreovirus genome segment 2: completion of sequence analyses of Rice dwarf virus. Intervirology 37, 6–11. Valarmathi, P., Kumar, G., Robin, S., Manonmani, S., Dasgupta, I., and Rabindran, R. (2016). Evaluation of virus resistance and agronomic performance of rice cultivar ASD 16 after transfer of transgene against Rice tungro bacilliform virus by backcross breeding. Virus Genes 52, 521–529. https://doi.org/10.1007/s11262016-1318-x

Verma, V., and Dasgupta, I. (2007). Sequence analysis of the complete genomes of two Rice tungro spherical virus isolates from India. Arch. Virol. 152, 645–648. https:// doi.org/10.1007/s00705-006-0861-5 Verma, V., Sharma, S., Devi, S.V., Rajasubramaniam, S., and Dasgupta, I. (2012). Delay in virus accumulation and low virus transmission from transgenic rice plants expressing Rice tungro spherical virus RNA. Virus Genes 45, 350–359. https://doi.org/10.1007/s11262-0120787-9 Wagh, S.G., Kobayashi, K., Yaeno, T., Yamaoka, N., Masuta, C., and Nishiguchi, M. (2016). Rice necrosis mosaic virus, a fungal transmitted Bymovirus: complete nucleotide sequence of the genomic RNAs and subgrouping of bymoviruses. J. Gen. Plant Pathol. 82, 38–42. Wang, Q., Tao, T., Zhang, Y., Wu, W., Li, D., Yu, J., and Han, C. (2011). Rice black-streaked dwarf virus P6 selfinteracts to form punctate, viroplasm-like structures in the cytoplasm and recruits viroplasm-associated protein P9-1. Virol. J. 8.1, 24. https://doi.org/10.1186/1743422X-8-24 Wang, Q., Yang, J., Zhou, G., Zhang, H., Chen, J., and Adams, M.J. (2010). The complete genome sequence of two isolates of southern rice black‐streaked dwarf virus, a new member of the genus Fijivirus. J. Phytopathol. 158, 733–737. Wang, Y., and Wu, W.H. (2013). Potassium transport and signaling in higher plants. Annu. Rev. Plant Biol. 64, 451–476. https://doi.org/10.1146/annurevarplant-050312-120153 Wei, T., Chen, H., Ichiki-Uehara, T., Hibino, H., and Omura, T. (2007). Entry of Rice dwarf virus into cultured cells of its insect vector involves clathrin-mediated endocytosis. J. Virol. 81, 7811–7815. Wei, T., Shimizu, T., Hagiwara, K., Kikuchi, A., Moriyasu, Y., Suzuki, N., Chen, H., and Omura, T. (2006). Pns12 protein of Rice dwarf virus is essential for formation of viroplasms and nucleation of viral-assembly complexes. J. Gen. Virol. 87, 429–438. Wei, T., Uehara-Ichiki, T., Miyazaki, N., Hibino, H., Iwasaki, K., and Omura, T. (2009). Association of Rice gall dwarf virus with microtubules is necessary for viral release from cultured insect vector cells. J. Virol. 83, 10830–10835. https://doi.org/10.1128/JVI.01067-09 Weinl, S., and Kudla, J. (2009). The CBL-CIPK Ca(2+)decoding signaling network: function and perspectives. New Phytol. 184, 517–528. Wu, W., Zheng, L., Chen, H., Jia, D., Li, F., and Wei, T. (2014). Nonstructural protein NS4 of Rice stripe virus plays a critical role in viral spread in the body of vector insects. PLOS ONE 9, e88636. https://doi. org/10.1371/journal.pone.0088636 Xie, L.H., Chen, Z.X., and Lin, Q.Y. (1979). Studies on the dwarf-like disease of rice plants. Acta Phytopathol. Sin 9, 93–100. Xie, L.H., Lin, J.Y., Xie, L.M., and Nai, G.B. (1984). On the bunchy stunt disease of rice IV: The experiments of occurrence, development and control of rice bunchy stunt. Acta Phytopathol. Sin. 14, 33–38. Xie, L.H., and Lin, Q.Y. (1984). Progress in the research of virus diseases of rice in China. Sci. Agric. Sin. 6, 58–64.

Viruses Infecting Rice and their Transgenic Control |  199

Xie, L.H., and Lin, Q.Y. (1980). Studies on bunchy stunt disease of rice, a new virus disease of rice plant. Kexue Tongbao 25, 785–789. Xie, L.H., Lin, Q.Y., and Xie, L.Y. (1996). Rice bunchy stunt virus: A new member of Phytoreovirus. J.Fujian Agri. Univ. 25, 312–319. Xiong, G., Liu, X., Qiu, P., Wu, X., Du, Z., Zhang, J., Yang, L., and Wu, Z. (2017). Rice grassy stunt virus p5 interacts with two protein components of the plant-specific CBLCIPK Ca+2 signaling network of rice. Virus Genes. 53, 446–453. https://doi.org/10.1007/s11262-017-1437-z Xu, J., Li, H.D., Chen, L.Q., Wang, Y., Liu, L.L., He, L., and Wu, W.H. (2006). A protein kinase, interacting with two calcineurin B-like proteins, regulates K+ transporter AKT1 in Arabidopsis. Cell 125, 1347–1360. Yan, Y., Wang, J., Qiu, B., and Tian, B. (1996). Resistance to Rice stripe virus couferred by expression of coat protein in transgenic indica rice plants regenerated from bombarded suspension culture. Virol. Sin. 12, 260–269. Yang, X., Tan, S.H., Teh, Y.J., and Yuan, Y.A. (2011). Structural implications into dsRNA binding and RNA silencing suppression by NS3 protein of Rice Hoja Blanca Tenuivirus. RNA 17, 903–911. https://doi. org/10.1261/rna.2552811 Yassi, M.N., Ritzenthaler, C., Brugidou, C., Fauquet, C., and Beachy, R.N. (1994). Nucleotide sequence and genome characterization of Rice yellow mottle virus RNA. J. Gen. Virol. 75, 249–257. https://doi.org/10.1099/00221317-75-2-249 Zhang, C., Song, Y., Jiang, F., Li, G., Jiang, Y., Zhu, C., and Wen, F. (2012). Virus resistance obtained in transgenic tobacco and rice by RNA interference using promoters with distinct activity. Biol. Plant. 56, 742–748. Zhang, H.M., Yang, J., Chen, J.P., and Adams, M.J. (2008). A black-streaked dwarf disease on rice in China is caused by a novel Fijivirus. Arch. Virol. 153, 1893–1898. https://doi.org/10.1007/s00705-008-0209-4 Zhang, S., Jones, M.C., Barker, P., Davies, J.W., and Hull, R. (1993). Molecular cloning and sequencing of coat protein-encoding cDNA of Rice tungro spherical virus a plant picornavirus. Virus Genes 7, 121–132.

Zheng, L.P., Lin, C., Xie, L.Y., Wu, Z.J., and Xie, L.H. (2014). Construction of Rice stripe virus NS2 and NS3 Co-RNAi transgenic rice and disease-resistance analysis. Bing du xue bao= Chinese journal of virology. 30.6, 661–667. Zhong, B., Kikuchi, A., Moriyasu, Y., Higashi, T., Hagiwara, K., and Omura, T. (2003). A minor outer capsid protein, P9, of Rice dwarf virus. Arch. Virol. 148, 2275–2280. https://doi.org/10.1007/s00705-003-0160-3 Zhou, G.H., Zhang, S.G., Zou, S.F., Xu, Z.W., and Zhou, Z.Q. (2010). Occurrence and damage analysis of a new rice dwarf disease caused by Southern Rice black-streaked dwarf virus. Plant Prot 36, 144–146. Zhou, G., Wen, J., Cai, D., Li, P., Xu, D., and Zhang, S. (2008). Southern Rice black-streaked dwarf virus: A new proposed Fijivirus species in the family Reoviridae. Chinese Sci. Bull. 53, 3677–3685. Zhou, G., Xu, D., Xu, D., and Zhang, M. (2013). Southern Rice black-streaked dwarf virus: a white-backed planthoppertransmitted Fijivirus threatening rice production in Asia. Front. Microbiol. 4, 270. https://doi.org/10.3389/ fmicb.2013.00270 Zhou, Y., Yuan, Y., Yuan, F., Wang, M., Zhong, H., Gu, M., and Liang, G. (2012). RNAi-directed down-regulation of RSV results in increased resistance in rice (Oryza sativa L.). Biotechnol. Lett. 34, 965–972. https://doi. org/10.1007/s10529-012-0848-0 Zhu, S., Gao, F., Cao, X., Chen, M., Ye, G., Wei, C., and Li, Y. (2005). The Rice dwarf virus P2 protein interacts with ent-kaurene oxidases in vivo, leading to reduced biosynthesis of gibberellins and rice dwarf symptoms. Plant Physiol. 139, 1935–1945. Zhu, Y., Hayakawa, T., and Toriyama, S. (1992). Complete nucleotide sequence of RNA 4 of Rice stripe virus isolate T, and comparison with another isolate and with maize stripe virus. J. Gen. Virol. 73, 1309–1312. https://doi. org/10.1099/0022-1317-73-5-1309 Zhu, Y., Hayakawa, T., Toriyama, S., and Takahashi, M. (1991). Complete nucleotide sequence of RNA 3 of Rice stripe virus: an ambisense coding strategy. J. Gen. Virol. 72, 763–767. https://doi.org/10.1099/0022-1317-724-763

Whitefly-transmitted Begomoviruses and Advances in the Control of their Vectors

10

Surapathrudu Kanakala* and Murad Ghanim

Department of Entomology, Volcani Center, Bet Dagan, Israel. *Correspondence: [email protected] https://doi.org/10.21775/9781910190814.10

Abstract Whitefly-transmitted begomoviruses infect a large number of dicotyledonous hosts, causing many diseases of economic importance. Members of the genus Begomovirus are transmitted by only one whitefly species, Bemisia tabaci; and the symptoms include severe leaf curling, yellow mosaic, and yellow vein mosaic in several important agricultural crops. The insect vector B. tabaci plays a major role in the spread of many members of this virus group, and as a consequence, it also contributes to the emergence of new strains and species. In the last decade, 360 Begomovirus species and 41 morphologically indistinguishable B. tabaci species that were characterized across the globe have dramatically increased. Together, both the complex members of B. tabaci species and begomovirus species are spreading around the world; however, little information exists about the geographical distribution of this group of viruses, the relationships between its species, and the dependence for transmission on the only whitefly vector. The ability of multiple virus transmissions, the analysis of the B. tabaci species, and the identification of insect proteins implicated in virus replication and transmission will open up novel avenues for the management of this group of important diseases. This chapter provides an overview of the global diversity and geographical distribution of the B. tabaci species complex and their association with begomoviruses and recent advances in their control.

Introduction The whitefly, Bemisia tabaci (Gennadius) (Hemiptera: Aleyrodidae), has emerged as an important insect pest in the past two decades owing to its increasing infestation habits, and its role in causing tremendous damage to agricultural crops and ornamentals. B. tabaci was described as Aleyrodes tabaci based on its presence on tobacco collected from Greece in 1889 (Gennadius, 1889). It was later described as A. inconspicua by Quaintance in 1900 (Quaintance, 1900), and in 1957, it was named as B. tabaci (Russell, 1957). According to Misra and Lamba (1929), in the late 1920s, B. tabaci was first reported to be a serious agricultural insect pest in the northern parts of India. At present, B. tabaci has a worldwide distribution and has been documented in all continents, except Antarctica (Martin et al., 2000). B. tabaci is an extremely polyphagous insect that develops through the egg and several nymphal stages that start in the instar (crawler) stage, followed by three nymphal stages (2nd, 3rd and 4th instar) until the pupal stage. The whitefly’s major impact on agriculture is due to the facts that (1) the whitefly feeds on over 700 plant species from 86 families; and (2) it damages plants by direct feeding which weakens plants and through the secretion of honeydew. The indirect damage caused by B. tabaci is due to the fact that following aphids; B. tabaci is the most important vector for plant viruses and transmits virus species belonging to the

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to be acquired and transmitted by the whitefly for the successful expression of the disease symptoms. Examples of transmitted viruses include the Tomato leaf curl New Delhi virus (ToLCNDV) and the Mungbean yellow mosaic India virus (MYMIV). The monopartite genome only consists of DNA-A. DNA-A alone is infectious and produces typical symptoms in experimental assay hosts and on primary hosts (for example: Tomato yellow leaf curl virus (TYLCV) (Fig. 10.1). In recent years, there has been a tremendous increase in the number of B. tabaci putative cryptic species and B. tabacitransmitted begomoviruses across the world. Begomoviruses are vectored by B. tabaci in a persistent, circulative manner. Over the past three decades, the emergence and the spread of these viruses have been a consequence of (1) the global spread of the B. tabaci cryptic complex species, (2) the acquisition of the complex begomoviruses species/pseudo-recombinants by the indigenous and invasive species of B. tabaci, (3) the introduction of begomoviruses to new crops by the B. tabaci species that feed on unusually broad range of plant families, (4) the natural history of B. tabaci, which has a high reproduction rate and a short life cycle, (5) human activity through the movement of virusinfected and whitefly-harbouring plant materials, and (6) the adaptation of the B. tabaci species to

Begomovirus, Crinivirus, Closterovirus, Carlavirus, and the Torradovirus groups. Among the whiteflytransmitted viruses, 90% belong to the genus Begomovirus, which belongs to the family Geminiviridae ( Jones, 2003); this family includes more than 360 species and has emerged globally as the most destructive group of plant viruses during the past two decades, particularly in the tropical and subtropical regions. The family Geminiviridae is a group of plant viruses with single-stranded DNA genomes, encapsidated in geminate particles that infect monocot or dicot plants. Members of the family are grouped into nine genera (this includes: Mastrevirus, Begomovirus, Curtovirus, Topocuvirus, Becurtovirus, Eragrovirus, Capulavirus, Grablovirus, and Turncurtovirus) based on the genome organization, vector transmission, and host range (Zerbini et al., 2017). Members of the genus Begomovirus are transmitted by the whitefly B. tabaci, and cause severe leaf curling, yellow mosaic, and yellow vein mosaic in several important crops and weeds. Begomoviruses can have either a bipartite or monopartite genome. The bipartite genome consists of two circular single-stranded DNA (2.5 to 2.7 kb), which are referred to as DNA-A or DNA-B. Both of these components are independently encapsidated, and the geminate particles encapsidating A and B need (A)

(B)

(C)

(D)

(E)

(F)

(G)

(H)

Figure 10.1 Diseases caused by whitefly-transmitted begomoviruses. (A) Whiteflies feeding on field grown soybean plants. (B) B. tabaci (MEAM1). (C) Tomato yellow leaf curl virus (TYLCV) symptoms on tomato. (D) Tomato leaf curl disease in field-grown tomato in Tanzania. (E) Mixed infections of Tomato leaf curl New Delhi virus (ToLCNDV) and Tomato leaf curl Palampur virus (ToLCPalV) in the fields of Hisar, India. (F) Yellow mosaic symptoms of field-grown mungbean plants in India. (G) Mixed infection of Squash leaf curl virus (SqLCV) and Watermelon chlorotic stunt virus (WmCSV) in the field-grown squash plants in Israel. (H) Cassava mosaic disease symptoms.

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climatic changes, and its ability to quickly develop resistance to pesticides. These factors have favoured the rapid spread of begomoviruses across the globe. This spread with the rapid evolution of this group of viruses pose a serious threat to a variety of cultivated crops. This chapter highlights the emergence and the worldwide spread of the B. tabaci complex species, begomoviruses, which are transmitted by the indigenous and invasive species of B. tabaci in selected crops, and the recent advances in vector control and management. Global diversity of the B. tabaci species complex B. tabaci is complex of biotypes that differ in their genetic make-up, behaviour, host range, their ability to transmit plant viruses, and the bacterial endosymbionts that they harbour and the ability to develop resistance to pesticides (Ghanim, 2014). In 2007, for the first time, the global phylogenetic reconstruction of the B. tabaci complex species was conducted with 366 sequences; and as a result, around 12 major genetic groups were identified in this study (Boykin et al., 2007). Later, De Barro et al. (2011) and Dinsdale et al. (2010) suggested that term ‘biotype’ which has been used to distinguish the whitefly population based on 3.5% mitochondrial cytochrome oxidase gene sequence divergence is no longer appropriate to highlight the actual differences in this species complex. Based on this criterion, 24 B. tabaci cryptic species were determined and nominated. Recently, Lee et al. (2013) observed that a 4.0% genetic boundary was more realistic than 3.5% in distinguishing the B. tabaci species. Though a few controversies did occur (Mouton et al., 2015, Qin et al., 2016), reclassification based on the 3.5% and 4% genetic divergence revealed that 41 groups are clearly distinct (Fig. 10.2). The newly described morphologically indistinguishable species (which include Africa, Asia I, Asia I-India, Asia II 1–12, Asia III, Asia IV, Asia V, Australia, Australia/Indonesia, China 1–5, Indian Ocean, Ru, Middle East Asia Minor I-II (MEAM), Mediterranean (MED), New World 1–2, Japan 1–2, Uganda, Italy 1, and sub-Saharan Africa 1–5) have been currently delimited at the global level (Boykin et al., 2007; Dinsdale et al., 2010; De Barro et al., 2011; Firdaus et al., 2013; Hu et al., 2017; Lee et al., 2013).

Geographic distribution of the B. tabaci species complex A worldwide spread, emergence, displacement, and movement of the native and the invasive B. tabaci species has apparently resulted in the spread of begomoviruses into new areas. This includes the severe outbreak of the B. tabaci species in tropical and subtropical regions over the past two decades with the emergence of the native and the invasive B. tabaci-transmitted begomoviruses in several cultivated crops (Briddon and Markham, 2000; Rybicki and Pietersen, 1999; Thresh et al., 1998). The worldwide genetic diversity of the B. tabaci species complex and its association with the begomoviruses are not well understood. On the basis of our comprehensive analyses of the B. tabaci species from 82 countries (Kanakala and Ghanim, unpublished), it was revealed that Asia contains the highest number of B. tabaci species, with a total of 29 native and invasive species (including Asia I, Asia I-India, Asia II 1–12, Asia III, Asia IV, Asia V, MED, MEAM1, China 1–5, Australia/Indonesia, India, and Japan 1–2) distributed across 13 countries (i.e. India, Pakistan, Bangladesh, Nepal, Cambodia, Indonesia, Malaysia, Myanmar, Singapore, China, Japan, South Korea, and Taiwan). So far, MEAM1 species was reported in 42 countries, while MED was reported in 44 countries, suggesting that these species are the most invasive and have been distributed on a global scale. The species that are endemic to China were described in four countries, the Asian species were described in 14 countries, the Indian Ocean species were described in six countries, the Sub-Saharan African species were described in 19 countries, and the New World species were described in 12 countries. Each newly described species (including Asia I-India, Asia II 2, Asia II 3, Asia II 4, Asia II 8, Asia II 9, Asia II 10, Asia II 11, Asia II 12, Asia IV, Asia V, Australia, Australia/Indonesia, China 4, China 5, Italy 1, Japan 1, Japan 2, sub Saharan Africa 5 and Uganda) were described in only one country (Table 10.1). The association of B. tabaci complexes with begomoviruses Viral diseases by begomoviruses, criniviruses, carlaviruses, ipomoviruses, and torradoviruses have been mainly associated with the exotic B.

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Figure 10.2  Bayesian tree inferred from global B. tabaci mtCOI data showing the B. tabaci species complex members. Numbers above branches represent the percentage of 1000 bootstraps and number below branches represents genetic distance.

tabaci native and invasive species, which is the most important whitefly species in terms of virus transmission ( Jones, 2003). The diverse agroclimatic conditions under which different crop genotypes are grown ensure a continuous build-up of the whitefly population and the perpetuation of the virus inoculum. Human activities also indirectly enhance the introduction of viral diseases by introducing viruliferous whiteflies with new introduced viruses ( Jones, 2009). Some of the whitefly-transmitted begomoviruses affect agricultural crops such as cotton, cassava, tomatoes, beans,

and ornamentals. Due to this fact, during the last two decades, begomoviruses have been expanding their host range and even crossing the boundaries of their place of origin. Several studies have shown that the B. tabaci species are also geographically distributed in the New and the Old World. Among the species reported, both the MEAM1 and MED have the highest expansion rate on a worldwide scale. The intensification of cropping systems and the plant-material spread resulted in increase in the population of whiteflies, and thus the emergence and the spread of the native begomovirus species

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Table 10.1 List of B. tabaci species and begomoviruses from each country No. Country

B. tabaci species

Begomoviruses

AbMV (HQ588899), AEV (EU867513); BYVBhV (FJ589571); BYVMV (AF241479); BYVV (GU181356); BYVMMaV (HM590503); CToLCV (DQ629102); ChiLCINV (FM877858); ChiLCVV (HM007121); ChiLCV (DQ629103); ClYMV (EF408037); CoGMV (EU636712); CoYV (KC196077); CLCuBaV (AY705380); CLCuKoV (AY456683); CLCuMuV (AF363011); ChiLCGV (KJ957157); ChiLCAV (KM880103); CoMoV (KM244719); JMLV (KC513823); JLYMKV (JN698954); JLCrIV (KM066975); MLCV (LK054801); DoYMV (AJ968370); FbLCV (JQ866297); HemYMV (KC898543); HoLCV (KC476655); HgYMV (AJ627904); ICMV (AY730035); JLCuV (EU798996); JMINV (HM230683); JYMINV (FJ177030); MaYVV (EU366903); MeYVMBaV (EU360303); MYMIV (AF416742); MYMV (AJ421642); OELCuV (GU111996); PaLCrV (HM140367); PaLCuV (Y15934); PeLCV J(N807764); PepLCBV (HM007096); PepLCLaV (JN135234); RaLCuV (EF175733); RhYMIV (HM777508); RoLCuV (KF584008); SiLCuV (AM259382); SpiYVSKV (KF660223); SLCCNV (AM286794); SLCMV (AJ579307); SHLDV (EU194914); SPLCV (FN432356); TbCSV (GQ994095); TbLCPuV (HQ180391); ToLCBaV (KF551588); ToLCBV (JQ765395); ToLCJV (HM991146); ToLCKeV (EU910141); ToLCNDV (AM286433); ToLCPalV (AM884015, AM992534); ToLCPatV (EU862323); ToLCPuV (AY754814); ToLCRaV (DQ339117); ToLCV (AF413671).

OLD WORLD South Asia 1

India

Asia I, Asia I-India, Asia II 1, Asia II 5, Asia II 7, Asia II 8, Asia II 11, China 3, MEAM1

2

Pakistan

Asia I, Asia II 1, Asia II 5, Asia II ACMV (FJ751234); AEV (AM261836); AlYVV (FN432361); 7, MEAM1 BYVMV (AJ002451); CaYMV (HE580234); ChiLCPKV (DQ116877); ChiLCV (AF336806); ClYMV (HE863667); CLCuAlV (AJ002452); CLCuGeV (FR751142); CLCuKoV (AJ002448); CLCuMuV (EU365615); MeYVMV (FR772081); MYMIV (AJ512495); MYMV (AY269991); OELCuV (HF567945); PaLCrV (HE580236); PaLCuV (AJ436992); PeLCV (AM948961); PepLCBV (AM404179); RaLCuV (GU732204); RhYMV (AM999981); RoLCuV (GQ478342); SLCCNV (AM286794); ToLCNDV (AF448058, AY150305); ToLCPalV (AM494976); ToLCV (FR819708)

3

Bangladesh

Asia I, Asia II 1, Asia II 5, China ChiLCV (AJ875159); DoYMV (AY271891); MYMIV 3 (AF314145); PepLCBV (AF314531); ToLCBV (AF188481); ToLCNDV (AJ875157, AJ875158)

4

Nepal

Asia II 1

AEV (AJ437618); AYVV (KC28264); MYMIV (AY271895); ToLCV (AY234383)

Southeast Asia 5

Cambodia

MED1, Asia I

TYLCKaV (KR073086); SLCMV (KT861468); MYMV (AY271892); MaYVCaV (KP188831)

6

Indonesia

Australia/Indonesia, Asia II 7, Asia II 12

AYVV (AB100305); MYMIV (JN368432); PepYLCIV (AB267834); TYLCIDV (AF189018), TYLCKaV (KF446661), ToLCSuV (FJ237615), ToLCNDV (AB613825), ToLCJaV-A (AB100304), ToLCJaV-B (AB162141)

7

Malaysia

Asia I, MED, China 2, Asia II 7

PepLCV (AF414287); PuYMV (EF197941); SLCCNV (EF197940); ToLCMYV (AF327436)

8

Myanmar

Asia II 5

TYLCTHV-D (AF206674)

9

Singapore

Asia I

AYVV (X74516)

206  | Kanakala and Ghanim

Table 10.1 Continued No. Country

B. tabaci species

Begomoviruses

ALCuV (AJ851005); AllLCV (EF602306); ChiLCPKV (HM587709); ClGMCNV (FJ011668); ClGMJsV (FN396966); ClGMV (HQ317134); CLCuMuV (EF465535); CraYVV (EF165536); EYVV (EU377539); EuLCGxV (AM411424); EuLCuV (AJ558121); KuMV (FJ539014); LaYVV (AY795900); LuYVV (AJ965539); MaLCuV (AJ971263); MaYMV (AM236755); MaYVYnV (AJ786711); PaLCuCNV (AJ558116); PaLCuGdV (FJ495184); PepLCYnV (EU585781); PepYLCCNV (KC149938); PepYVMLV (AM691547); PouGMV (JX183732); SeYMV (AJ876550); SiLCuV (AM050730); SiYMCNV (AJ810096); SgYVGxV (AM238692); SgYVV (AM183224); SLCCNV (AB027465); SLCuYV (AJ420319); StaLCuV (AJ495814); SPLCCNV (DQ512731); SPLCGV (JX448368); SPLCHnV (KC907406); SPLCSiV (KC488316); SPLCV (EU253456); TbCSV (AF240675); TbLCYnV (AF240674); ToLCCNV (AJ558118); ToLCGdV (AY602165); ToLCGxV (AM236784); ToLCHaiV (KF150142); ToLCHsV (EF125190); ToLCTV (AM698111); TYLCCNV (AF311734); TYLCTHV-C (AJ495812); TYLCV (AM282874); VeYVFV (JF265670); LyYMV (KT582302)

East Asia 10

China

MED, MEAM1, Asia I, Asia II 1, Asia II 2, Asia II 3, Asia II 4, Asia II 6, Asia II 7, Asia II 9, Asia II 10, Asia IV, Asia V, China 1, China 2, China 3, China 4, China 5

11

Japan

MED, MEAM1, MEAM2, Asia I, AYVV (AB050781); EpYVV (AB007990); HYVV (AB020781); Asia III, Japan 1, Japan 2, Asia SPLCV (AB433786); TYLCV (AB014346) II 1, Asia II 6

12

South Korea

MED, MEAM1

HYVV (FJ434943); SPLCCNV (JX961671); SPLCV (JX961670); TYLCV (HM856919)

13

Taiwan

MED1, MEAM1, Asia I, Asia III, Asia II 1, Asia II 6, Asia II 7

AYVV (AF307861); EuLCuV (KC161185); PaLCuGdV (JN703795); PouMGDV (KF414123); SLCuPV (DQ866135); SLCuPV (JF746195); ToLCHsV (DQ866131); ToLCHsV (DQ866131); ToLCNDV (GU180095, GU180096); ToLCTV (AM698111); TYLCTHV-A(AF141922, AF141897); TYLCTHV-B (EF577264, EF577265)

Middle East 14

Egypt

MED, MEAM1

CLCuGeV (AF014881); SLCuV (DQ285019); TYLCV (AY594174)

15

Iran

MEAM1

MaYSV (JN419015); ToLCPalV (EU547682, FJ660442); ToLCV (AY297924); TYLCV (AJ132711); ToLCNDV (KP793719; WmCSV (KT272765); OELCuV (KJ397527)

16

Iraq

MEAM1

TYLCV (JQ354991)

17

Israel

MED, MEAM1

SLCuV (HQ184436); TYLCV (DQ845787), TYLCSV (DQ845787); WmCSV (EF201809)

18

Jordan

MEAM1

CLCuGeV (GU945265); SLCuV (EF532620); TYLCV (AY594175); WmCSV (EU561237); TYLCAxV (KM215610)

19

Kuwait

MEAM1

TYLCV (JF451352)

20

Saudi Arabia

MEAM1

ToLCSDV (KC845301); WmCSV (KM066100)

21

Syria

Asia II 1, MED, MEAM1

–

22

Turkey

MED, MEAM1, Asia I

TYLCV (AJ812277)

23

United Arab MEAM1 Emirates (UAE)

CLCuGeV (KJ939446)

24

Yemen

MEAM1

WmCSV (AJ012081); ToCSDV (EF110891)

MED

–

Africa 25

Algeria

Whitefly-Transmitted Begomoviruses and Control of their Vectors |  207

Table 10.1 Continued No. Country

B. tabaci species

Begomoviruses

26

Benin

MED, sub-Saharan Africa 1

ACMV (KR476371); CYMV (KU683747); ChaYMV (KT454819)

27

Burkina Faso

MED

ACMV (FM877473); CLCuGeV (EU432374); EACMV (FM877474); PepYVMLV (FM876847); ToLCMLV (LM651404); TYLCMLV (LM651401); ToLCBFV (KX853168) –

28

Burundi

sub-Saharan Africa 1

29

Cameroon

ACMV (AF112352); CLCuGeV (FM210276); EACMV Africa, MED, sub-Saharan Africa 2, sub-Saharan Africa 3, (AF112354); OYCrV (FM164724); TelMV (KU683744); ToLCGV (FM210277); TYLCMLV (FM212660); SbCBV sub-Saharan Africa 4 (KT444613), WAAV1 (KT444603)

30

Democratic Republic of the Congo

sub-Saharan Africa 1, subSaharan Africa 3

ACMV (FN435277); EACMV (JX910240)

31

Ghana

MED, sub-Saharan Africa 3, sub-Saharan Africa 1

ACMV (JN165088); EACMV (JN165089); ToLCKuV (KT382325); ToLCGV (EU350585); TYLCMLV (EU847740)

32

Ivory Coast

MED

–

33

Kenya

sub-Saharan Africa 1, subSaharan Africa 2

ACMV (J02057); EACMKV (AJ717569); EACMV (AJ006458); EACMZV (AJ717560); DMV (KT878831)

34

Madagascar

Indian Ocean

AMMGV (KP663483); CMMGV (HE617299); CLCuGeV (AM701757); EACMKV (HE984147); SACMV (AJ422132); ToLCDiV (AM701765); ToLCMGV (AJ865338); ToLCYTV (AM701761); ToLCToV (AM701768)

35

Malawi

Sub-Saharan Africa 1

ACMV (JX658682); EACMMV (AJ006459)

36

Mali

sub-Saharan Africa 2

CLCuGeV (EU024120); OYCrV (DQ875879); PepYVMLV (AY502935); ToLCMLV (AY502936); TYLCMLV (DQ358913)

37

Mauritius

Indian Ocean

TYLCV (HM448447)

38

Morocco

MED, MEAM1

TYLCV (EF060196), TYLCSV (AY702650)

39

Mozambique

sub-Saharan Africa 1

–

40

Nigeria

MED, sub-Saharan Africa 2

ACMV (AJ427910); ChaYMV (AJ223191); CPGMV (AF029217); EACMV (AJ867444); JMNV (JX025358); SbCBV (GQ472985); SbMMoV (GQ472984); WAAV1 (KT444601); ToLCGV (FJ685621)

41

Senegal

MED, MEAM1

–

42

Seychelles

Indian Ocean

EACMV (JF909151); ToLCSCV (AM491778)

43

Sudan

MED, New World

CLCuGeV (AF260241); WmCSV (AJ245650); ToLCSDV (AY044137); TYLCV (AY044138);

44

Swaziland

Sub-Saharan Africa 1

–

45

South Africa

MED, MEAM1, sub-Saharan Africa 1

SACMV (AF155806); SPLCSPV (JQ621844); SPMV (JQ621843); ToCSV (AF261885)

46

Tanzania

MED, sub-Saharan Africa 1

ACMV (AY795982); EACMV (AY795983); EACMZV (AF422174); ToLCArV (DQ519575); DMV (KT878829); ToLCArV (DQ519575)

47

Togo

MED, sub-Saharan Africa 3

ACMV (KR476372); ChaYMV (KT454820); SbCBV (KT454811); ToLCGV (FJ685620)

48

Tunisia

MED, MEAM1, sub-Saharan Africa 2

TYLCV (AY736854), TYLCSV (AY736854)

49

Uganda

ACMV (AF126800); EACMV (AF126804); DesMOV Uganda, Indian Ocean, MED, (KY294724); SPLCUV (FR751068); ToLCUV (DQ127170); sub-Saharan Africa 2, subSaharan Africa 5, sub-Saharan VeCrV (KX831132) Africa 1

50

Zambia

sub-Saharan Africa 1

EACMV (KT869123); EACMMV (KT869119); ACMV (KT869127)

51

Zimbabwe

MED

SACMV (AJ575560); TbLCZV (AF350330)

208  | Kanakala and Ghanim

Table 10.1 Continued No. Country

B. tabaci species

Begomoviruses

52

Mayotte

MEAM1

EACMKV (JF909172); EACMV (JF909163); ToLCYTV (AJ865340)

53

Reunion

MED, MEAM1, MEAM2, Indian TYLCV (AM409201) Ocean

Europe 54

Bosnia and Herzegovina

MED

–

55

Croatia

MED, MEAM1

–

56

Cyprus

MED, MEAM1

–

57

Czech Republic

MED

–

58

France

MED, MEAM1, Indian Ocean

–

59

Greece

MED, MEAM1

SPLCV (KF697069)

60

Italy

MED, MEAM1, Italy 1, Ru

SPLCV (AJ586885); TYLCAxV (EU734831); TYLCSV (GU951759); ToLCNDV (KU145141); TYLCV (DQ144621), TYLCSV (GU951759), TYLCAxV (EU734832)

61

Netherlands

MED

TYLCV (FJ439569)

62

Netherlands Antilles

MEAM1

–

63

Portugal

MED, sub-Saharan Africa 2

TYLCV (AF105975), TYLCSV (JN859134), TYLCAxV (JN859138); TYLCMaV (JN859135)

64

Spain

MEAM1, MED,sub-Saharan Africa2, sub-Saharan Africa 3

SPLCCV (EF456742); SPLCV (AJ132548); TYLCMaV (AF271234); TYLCSV (AJ519675); TYLCSV (AJ519675), TYLCV (AF071228); ToLCNDV (KF749223); TYLCAxV (AY227892)

Australia, MEAM1

HYVV (JX416174); ToLCV (AF084006); TYLCV (GU178813)

Australia 65

Australia

NEW WORLD South America 66

Argentina

MEAM1, New World 2, MED

SbBMV (EF016486); SiMBoV2 (HM585443); SPLCV (JQ349087); ToMoWV (JQ714137); TRYLCV (KC132844); ToYSV (FJ538207); ToYVSV (GQ387369); ToDfLV (JN564749)

67

Bolivia

New World 2

AbMBoV (HM585445); SiMMV (HM585431); SiMBoV1 (HM585441); SiMBoV2 (HM585443); SoMBoV (HM585435)

68

Brazil

MEAM1,MED, New World 1, New World 2

AbMBV (FN434438); BGMV (FJ665283); BlYSV (EU710756); CeYSV (JN419002); CdTAV (HM357461); CleGMV (HQ396465); CoChSpV (KF358470); EuYMV (FJ619507); MaYSV (JN419005); MaYVV (JN419021); OMoV (EU914817); PSLDV (FJ972767); SiCMV (EU710751); SiGMBRV (FN436001); SiMMV (AJ557451); SiMoV (AJ557450); SiYBV (JX871380); SiYLCV (EU710750); SiYMAV (JX871383); SiYMV (AY090558); SidGLSV (HM357458); SPLCSPV (HQ393477); SPLCV (FJ969829); SPMV (FJ969831); ToBYMV (KC791690); ToBYMoV (KC791691); ToCMoV (AF490004, AF491306); ToCmMV (EU710754, EU710755); ToGLDV (HM357456); TGMV (JF694488); ToICV (JF803252); ToLDV (EU710749); ToMMV (EU710752, EU710753); ToMoLCV (JF803246); ToRMV (AF291705, AF291706); ToSRV (AY029750); ToYSV (DQ336350, DQ336351); ToYVSV (EF417915, EF417916)

69

Colombia

New World 1

BDMV (M88179); PYMV (EU518935); PLDV (KT899302); SPLCV (KC253242)

Whitefly-Transmitted Begomoviruses and Control of their Vectors |  209

Table 10.1 Continued No. Country

B. tabaci species

Begomoviruses

70

Uruguay

MED

TRYLCV (JN381813); ToYVSV (KR024026); ToGLSV (KC626021);

71

Venezuela

New World 1, MEAM1

BChV (JN848770); BChMV (JN848772); DaChMV (JN848775); DaLDV (JN848773); MClMV (HM163576); MerMV (AY508991); PYMV (D00940); SicGMV (JX857691); SiGMLaV (JX857693); ToClLDV (HQ171986); ToMYLCV (AY927277, EF547938); ToMLCuV (AY508991, AY508992); ToYMLCV (AY508993)

72

Trinidad and Tobago

MEAM1

PYMV (AF039031)

North America 73

USA

MED, MEAM1, New World 1

ChLCV (HM626515); ClGMCNV (JQ305797); EuMV (JQ963887); ToMoV (L14460, L14461); TYLCV (AY530931)

74

Canada

MED, MEAM1

–

Central America 75

Honduras

New World 1

RnGMV (AF239671); SiYVV (Y11097)

76

Belize

New World 1

–

77

Mexico

MED, MEAM1, New World 1

AbGMYV (KC430935); BCaMV (AF110189); BGYMV (AF173555); BYMMxV (FJ944023); BleICV (JX827487); BoYSV (EF121755); CabLCV (DQ406672); CarYSYV (KC426927); CdTV (AF101476); CoYSV (DQ875868); CLCrV (AF480940); DesLDV (DQ875870); EuMV (DQ318937); JacMYuV (JQ821386); OYMV (DQ022611); PepGMV (GU128148); PHYVV (AY044162); RhGMV (AF408199); SiMSiV (DQ520944); SiYMYuV (DQ875872); ToChLPV-A (AY339618); ToChLPV-B (DQ347948) ToGMoV (DQ520943, DQ406674); ToSLCV (JN676151); TYLCV (DQ631892); WmCSV (KY124280)

78

Panama

New World 1

PYMV (Y15034)

79

Dominican Republic

MEAM1

BGYMV (D00200); TYLCV (AF024715); JMV (KJ174330)

80

Guatemala

MED, MEAM1, New World 1

BGYMV (M91604); MCLCuV (AF325497); TYLCV (GU355941); ToGMoV (AF132852), ToSLCV (AF130415)

Asia II 5

–

Micronesia, Oceania 81

Nauru

Greater Antilles, Caribbean 82

Puerto Rico

MEAM1, New World 1

BGYMV (M10070); EuMV (AF068642); MacMPRV (AF449192); MerLCV (DQ644561); MerMPRV (FJ944021); PYMV (AY965897); RhMMV (FJ944019); SPLCV (DQ644562); ToMLCuV (AF068636, AY965899); TYLCV (AY134494)

83

Cuba

MEAM1

BGYMV (AJ544531); DiYMV (HE806446); EuMV (FJ807782); MacYMV (AJ344452); RhGMHaV (HM236368); RhRGMV (HM236370 HM236371); SiGMFlV (HE806442); SiGYVV (AJ577395); SiYMoV (HE806448); TbLCuCV (AM050143); TbLRV (AJ488768); TbMoLCV (FM160943); TbYCV (FJ213931); ToMHaV (Y14874); ToMoTaV (AF012300); ToMHaV (Y14874, Y14875); ToMoTaV (AF012300, AF012301); TYLCV (AJ223505); ToYLDV (FJ174698)

1GenBank numbers are not available for MED biotype from Cambodia (Chiel et al., 2009; Gueguen et al., 2010) and Taiwan (Hsieh et al., 2011). Begomovirus species abbreviations are available at the ICTV website (https://talk. ictvonline.org/ictv_wikis/Geminiviridae/m/files_gemini/5120/download). Note: Only B. tabaci species reported countries till date were listed in this table.

210  | Kanakala and Ghanim

into new geographic regions. MEAM1 and MED are distributed worldwide, and have invaded many countries mainly through international trade and changes in agricultural practices, which sometimes leads to the displacement of one indigenous species by an invasive one. TYLCV and its major vector B. tabaci MEAM I have been the most studied system in the last six decades. In the last two decades, however, begomoviruses have been noticed for their role in infecting vast host ranges worldwide. For example, a recent survey conducted by Papayiannis et al. (2011) in Cyprus enabled the detection of TYLCV in 49 different species belonging to 15 plant families. On the other hand, multiple viruses causing diseases have been detected in tomatoes. TYLCV has also spread to many provinces in China during the last decade through the invasive B. tabaci species. The rapid displacement of the MEAM1 biotype by the MED, however, has only been recently observed, suggesting an enhanced TYLCV spread in the southern provinces of China due to this invasive species (Chu et al., 2010; Hu et al., 2011; Luo et al., 2010; Pan et al., 2011). MED has rapidly displaced the MEAM1 in most locations across China (Pan et al., 2012; Zhang et al., 2014) and the United states (Dennehy et al., 2006), which caused many virus outbreaks. One of the major causes for this MED high invasiveness could be its exceptional ability to develop resistance to insecticides (Horowitz and Ishaaya, 2014; Xie et al., 2014), and in many countries, its ability to transmit TYLCV even more than MEAM1 (Ning et al., 2015; Pan et al., 2012). Similarly, in the Indian sub-continent, begomoviruses infecting beans, cassava, cotton, and tomato were associated and transmitted by the indigenous ‘Asia’ cryptic species. Recent reports revealed the presence of the ‘Asia’ species in Southeast Asia (Götz and Winter, 2016; Firdaus et al., 2013; Hu et al., 2017) and Turkey (de la Rua et al., 2006), and the cryptic species ‘China’, which originated in China, in India (Ellango et al., 2015). Alternatively, the presence and spread of begomoviruses between neighbouring countries or continents, suggests their global spread by the insect vector or other factors discussed above. This includes, for example, (1) MYMIV, MYMV (Mungbean yellow mosaic virus), ToLCNDV, ToLCPalV (Tomato leaf curl Palampur virus), and ToLCV (Tomato leaf curl virus), which are native to Asia, in the Middle

East, Europe, Southeast Asia, and East Asia; (2) Ageratum begomoviruses in Nepal, China, India, Indonesia, Singapore, Taiwan, and Thailand; (3) Chilli begomoviruses in India, Sri Lanka, Bangladesh, Pakistan, and Oman; (4) cotton begomoviruses in India, Pakistan, and China; and (5) Bhendi begomoviruses in India, Thailand, and Pakistan through the infection of a vast host range, suggesting their spread across boundaries (Table 10.1). In addition to TYLCV, other cassava-begomovirus species causing the mosaic disease in cassava in Africa were associated with the sub-Saharan Africa (1–5) species; these are rapidly spreading to neighbouring countries (Legg et al., 2014). Similarly, the African Cassava mosaic virus (ACMV) is infecting Gossypium species in Pakistan (Nawaz-ul-Rehman et al., 2012) and the East African Cassava mosaic Zanzibar virus (EACMZV) is infecting cassava in Oman (Khan et al., 2013); all the above reports suggest that begomoviruses, which infect agricultural crops in India and Africa, are expanding their host range and crossing boundaries into new geographic areas. This movement is driven by the following contributing factors: (a) the distribution of the B. tabaci cryptic species, (b) the movement of the infected plant material, and (c) the introduction of crops or genotypes susceptible to the native begomoviruses. Begomovirus mixed infections are also a prerequisite for the evolution of pseudo-recombinants. The transportation of infected plants and the translocation of viruliferous whiteflies by wind can be a fertile ground for the evolution of the pseudorecombination of begomoviruses, which lead to coadaptation between viruses and vectors from the same origin or geographical region, which results in recombination. Recent reports suggest that the rapid displacement of the whitefly biotypes are also supported by the new virus recombinants that became prevalent and led to the displacement of native begomoviruses (Sánchez-Campos et al., 1999; Monci et al., 2002; Davino et al., 2006). Recombination among begomovirus species may either occur in the host plant or the insect vector. When mixed infections occur in plants, begomoviruses interact synergistically or antagonistically with one another. Many recent reports showed particularly inter- and intra-begomovirus species mixed infections in agricultural crops across the world (Blawid

Whitefly-Transmitted Begomoviruses and Control of their Vectors |  211

et al., 2008; Chakraborty et al., 2008; Fondog et al., 2000; Kanakala et al., 2013; Pita et al., 2001; Silva et al., 2014). The synergism between cassava (Fondog et al., 2000; Pita et al., 2001; Patil and Fauquet, 2009) and tomato begomoviruses (Alves-Júnior et al., 2009; Blawid et al., 2008; Kanakala et al., 2013; Silva et al., 2014) in field conditions have revealed the possibility of the emergence of recombinants, and have demonstrated their role in the outbreak of epidemics. The rapid evolutionary potential of these begomoviruses combined with the polyphagous feeding behaviour of B. tabaci can lead to the emergence of new strains (Rosario et al., 2015). Rosario et al. (2015) performed high-throughput sequencing of whiteflies from the Old and the New World, and identified seven new different species from Guatemala, Puerto Rico, and Spain. There is, however, no direct evidence of these new viruses that are known to be transmitted to or infected in plants in nature. The coat protein (CP) has a redundant function in the transmission of begomoviruses by B. tabaci. Interestingly, replacement of few amino acids in the CP of begomoviruses restricts virus transmission by the whitefly. Briefly, only two amino acids, Q124K and H147Q of Sida golden mosaic Costa Rica virus (SiGMCRV), were found to be sufficient to render Abutilon mosaic virus (AbMV) transmission by whiteflies (Höhnle et al., 200; Höfer et al., 1997). The region of the CP between the amino acids 129 and 152 (Q129, Q134, and D152) of the Monopartite Tomato yellow leaf curl Sardinia virus (TYLCSV) was found to be relevant for virion assembly, systemic infection, and transmission by the vector (Noris et al., 1998). Similarly, the double-mutant Q129P and Q134H, and a further D152E of TYLCSV were acquired by B. tabaci, and circulated within the insect for up to 10 days as a wild-type virus; it was detected in the salivary glands, but it was not transmissible (Caciagli et al., 2009). Another mutant, N130D, was also found to be non-transmissible by B. tabaci (Caciagli et al., 2009). Similar results were confirmed in another bipartite begomovirus Watermelon chlorotic stunt virus (WmCSV) (Kheyr-Pour et al., 2000). Kollenberg et al. (2014) performed a comparison of the transmission of the WmCSV and the TYLCV in vector and non-vector populations of the MEAM1 species and found that the two populations acquired similar virus amounts; in the latter,

virus-localization studies revealed only negligible virus amounts in the primary salivary glands, which suggested an absence or a modified receptor implication in virus translocation. In addition, a recent report phylogenetically and structurally investigated the amino acids of CP, and suggested a biological relevance of K124 and Q149 in whitefly transmission (Fischer et al., 2015). Furthermore, hetero-encapsidation and the whitefly-transmission of the pseudo-recombinants, ToLCNDV and ToLCPalV, was demonstrated, indicating the possibility of hetero-encapsidation and the whitefly transmissions of the pseudo-recombinants of DNA-A of ToLCNDV and the DNA-B of ToLCPalV, and vice versa (Kanakala et al., 2013). Additionally, circular ssDNA satellites, beta-satellite (Ueda et al., 2012; Tabein et al., 2013), and delta-satellites (Hassan et al., 2016) have been shown to be transmitted by B. tabaci in the presence of helper viruses. Expanding host ranges, recombination among inter- and intrabegomoviruses, and multiple virus transmissions by its insect vector have been shown to contribute significantly to the evolution of new begomovirus species or strains. Worldwide scenario The first whitefly-transmitted geminivirus (yellow vein virus symptoms in Eupartorium chinense) was referred to in a poem written by Empress Koken in Manyoshu, a Japanese anthology that was prepared in 752 ad; later, it was identified as the Tobacco leaf curl virus belonging to the genus Begomovirus (Saunders et al., 2003). After the 1980s, the complete nucleotide sequencing of bipartite begomoviruses Bean golden mosaic virus (BGMV) (Goodman, 1978) and the monopartite TYLCV-Israel (Navot et al., 1991) revealed the existence of bipartite and monopartite begomoviruses. Since these findings, 360 begomovirus species have been characterized to date (https://talk.ictvonline.org/taxonomy). Together, both the B. tabaci species complex members and the begomovirus species have spread around the world; however, little information exists on the geographical distribution of this group of viruses, and its relationships and dependence for its transmission by whitefly vector populations. Agricultural intensification has been proposed as the main cause behind the spread of begomoviruses/mixed infections/the evolution of new virus

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species through recombination with increasing numbers; which led to their worldwide spread. This chapter is combining a comprehensive database that encompasses the most up-to-date sequence information published for all begomoviruses and B. tabaci species around the world. The distribution of different begomoviruses and B. tabaci species from various agriculturally important crops across the different countries of South Asia is summarized in Table 10.1. Surprisingly, South Asia, East Asia, and Southeast Asia comprise higher numbers of distinct begomovirus species and cryptic B. tabaci species. Mechanisms and proteins involved in whitefly– begomovirus interactions Begomoviruses are transmitted by B. tabaci in a circulative manner. When ingesting the sap from the infected tissue during feeding, virions reach the mid gut and localize in the filter chamber from which they are absorbed to the haemolymph. TYLCV and other begomoviruses cross the midgut and are transported through the haemolymph to salivary gland. Once in the salivary gland, the secretary cells mediate the transmission of the virions to plants (Ghanim, 2014; Wei et al., 2014). Furthermore, TYLCV may replicate in the insect vector (Pakkianathan et al., 2015; Sinisterra et al., 2005)

and trigger complex interactions with the insect vector (Wang et al., 2016). Several factors are involved in the transmission of the begomovirus. For example, two members of the heat-shock protein family, BtHSP16 and HSP70, interact with the tomato begomovirus (Götz et al., 2012; Ohnesorge and Bejarano, 2009). A small number of identified insect proteins appear to be implicated in the virus circulation and transmission within the whitefly’s body (Fig. 10.3). For example, Cyclophilin B (Kanakala and Ghanim, 2016), the Knottin-1 Gene (Hariton Shalev et al., 2016), the Peptidoglycan Recognition Proteins (PGRPs) (Wang et al., 2016), and the midgut protein (Rana et al., 2016) were also shown to interact with the begomovirus and play a role in virus transmission. More recently, clathrin-mediated endocytosis and endosomes have been detected to play a role in TYLCV transport across the whitefly midgut (Pan et al., 2017). However, more studies need to be carried out in order to clarify the virus circulation within the whitefly’s body. B. tabaci secondary symbionts have also been shown to facilitate the vector acquisition, retention, and transmission of begomoviruses (Gottlieb et al., 2010; Rana et al., 2012; Kliot et al., 2014). The symbiont GroEL (63kDa) protein was the first protein that has been shown to interact with TYLCV. Gottlieb et al. (2010) showed that only the GroEL protein from

Figure 10.3  Proteins implicated in Tomato yellow leaf curl virus transmission by B. tabaci. After being acquired from the plant phloem (p), virions pass through the stylet (s) and the oesophagus (e) to reach the filter chamber (fc). In the filter chamber, TYLCV interacts with HSP 16 or 70 (green), Cyclophilin B (yellow), Midgut protein (black), Peptidoglycan recognition proteins (PGRPs) and Clathrin (Purple particles). After being exocytosed from the midgut (mg) they circulate in the hemolymph where they interact with the GroEL protein (blue) and Knottin-1 (orange). The virions reach the primary salivary glands (psg) and enter their cells via endocytosis, and then they are injected into the host plant with salivary secretions. hg, hindgut; bc, bacteriocytes.

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Hamiltonella interacted with the TYLCV protein, and not the GroEL proteins from other B. tabaci symbionts. It was further shown that the release of virions protected by GroEL occur adjacent to the primary salivary gland. Similarly, Rana et al. (2012) showed that the Arsenophonus GroEL interacts with CLCuV, and it is localized in the midgut and the salivary gland of the Asia II species. A recent study (Kliot et al., 2014) showed that the Rickettsiainfected B. tabaci acquired more TYLCV virions from the infected plants and increases TYLCV transmission efficacy compared to Rickettsia-uninfected B. tabaci genetic sister strains. Few recent studies have shown that the virus infection can improve the performance of the vector whiteflies by repressing the plant defences (Li et al., 2014; Luan et al., 2013; Zhang et al., 2012). It was also recently shown that the pathogenicity factor βC1, encoded in the beta-satellite of TYLCCNV, can repress the tobacco JA-mediated defence against whiteflies and promote their performance on virus-infected plants (Zhang et al., 2012). Similarly, TYLCCNVsuppressing terpene biosynthesis in tobacco plants enhanced the performance of the whitefly (Li et al., 2014; Luan et al., 2013). Whitefly pre-infested plants reduced the plant susceptibility to virus transmission via triggering the Salicylic Acid (SA) pathway, thereby leading to the deposition of callose, which inhibited begomovirus replication or movement (Li et al., 2017). Advances in control of whiteflies B. tabaci is considered to be one of the most important invasive pests and virus vectors among several important families of plant viruses ( Jones, 2003; Hogenhout et al., 2008). The development of effective methods to control whiteflies is a major challenge owing to its wide host range, high reproductive rate, and its rapid evolution of resistance to insecticides. Chemical control At present, the use of chemical control is the primary approach employed to control whitefly populations. A number of insecticides have been used to control B. tabaci including organochlorines, Organophosphates (OPs), carbamates, pyrethroids, Insect Growth Regulators (IGRs; these include: buprofezin, pyriproxyfen, and novaluron), neonicotinoids,

diafenthiuron, pymetrozine, spiromesifen, spirotetramat, ryanodine receptors, and cyantraniliprole (Castle et al., 2010; Horowitz et al., 2011). However, in these cropping systems, chemical control must be considered as a temporary control owing to the ability of whiteflies developing a resistance to insecticides (Kontsedalov et al., 2012; Wang et al., 2017; Xie et al., 2014). The majority of the insecticides that are used to control whiteflies threaten the environment, beneficial organisms, and human health. Other methods, combining different tactics such as resistant varieties to the virus or the vector and biological control, plant-mediated resistance, RNA interference, and crop management offer potential control at minimal cost; however, they do not form major methods for whitefly control at present, and some of these methods are found in the development stage. Biological control Several parasitoids and other biological control agents can be used as natural enemies against whiteflies. These include fungi (Mycotal® (Verticillium lecanii-m), Botanigard® (Beauveria bassiana), and PreFeRal® (Paecilomyces fumosoroseus) (Stansly and Natwick (2010); parasitoids (Encarsia and Eretmocerus spp.) (Liu et al., 2015); and few predators (Amblyseius swirskii, Macrolophus caliginosus, Nesidiocoris tenuis, Delphastus spp. (Stansly and Natwick, 2010). These biological controls have been successfully implemented through the application of classic biological control approaches; however they are limited in their effectiveness. Host- plant resistance Host plant resistance has proven to be efficient and possesses a great potential to conduct successful Integrated Pest Management (IPM) against whiteflies in order to prevent viral diseases in plants (Nombela and Muñiz, 2009). Plant resistance, mediated by specific genes, also showed repercussions on whitefly fitness and development. In general, whiteflies often prefer to feed on plants bearing higher number of glandular trichomes (Nombela et al., 2000). Nombela et al. (2003) evaluated the response of transgenic tomato plants that carry the Mi-1.2 gene resistance to B. tabaci. The results showed a level of resistance to both, the MEAM1 and the MED species (Nombela et

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al., 2000, 2001, and 2003; Rodríguez-Álvarez et al., 2017). A few more studies showed the presence of glandular trichomes and the production of allelochemicals (including acylsugars, sesquiterpenes, and methyl ketones), which have higher levels of resistance to whiteflies (Bleeker et al., 2009; Firdaus et al., 2012; Freitas et al., 2002; Muigai et al., 2002). Another study reported that the wild tomato species’ (Solanum pimpinellifolium) accession, TO-937, was resistant to B. tabaci (Rodríguez-López et al., 2011). Well-known plant defence hormones such as salicylic acid (SA), jasmonic acid ( JA), and ethylene (ET) are involved in many biological processes including pathogen/pest attack. Whiteflies feeding on plants can suppress JA signalling by eliciting SA signalling (Zhang et al., 2013). A recent study showed that the phloem-feeding silver leaf whitefly induces SA defences and suppresses the effectual JA defences on Arabidopsis; the results revealed a dramatic delay in nymph development (Zarate et al., 2007). Rodríguez-Álvarez et al. (2015) also showed that SA has no role in basal defence; while SA is an important component of the Mi-1-mediated resistance to B. tabaci in tomatoes. More recently, two more studies showed that treatment with JA and its methyl ester, methyl jasmonate (MeJA), in tomatoes, enhanced the resistance to whiteflies and led to a decrease in the virus disease incidence (Escobar-Bravo et al., 2016; Sun et al., 2017). RNA interference The diseases caused by whitefly-transmitted begomoviruses influence agricultural productivity and continue to be challenges in the tropical and sub-tropical regions of the world. A high percentage of yield loss may occur even after spraying insecticides/biological agents. To date, however, no 100% whitefly-resistant commercial plant varieties are available. Recently, several transgenic approaches (RNA interference or RNAi, etc.) and genome-engineering platforms [Artificial Zinc-finger (AZF), Transcription Activator-like Effectors (TALEs), and clustered regularly interspaced short palindromic repeats (CRISPRs)/CRISPRassociated 9 (Cas9)] have also been employed for controlling insect pests and vectored viruses. RNA interference (RNAi) was recently suggested as a promising strategy as pesticide-free method to control insect vectors by silencing their genes.

Primarily, the RNAi mechanism involves the synthesis of double-stranded RNA (dsRNA), which is cleaved by the ribonuclease III-type Dicer into small RNAs (short interfering RNA, siRNA/ microRNA, miRNA/piwi-interacting RNA, and piRNA), which range between 19 and 25 bp in size, with 2 nucleotide 3′ overhangs. These fragments are assembled by Argonaute proteins to form the RNA-Induced Silencing Complex (RISC), which is a multiprotein complex where the destruction of the complementary sequence or the messenger RNA occurs (Simmer et al., 2002). The list of insect genes that have not been reported include those that cause mortality/fecundity/interference with pathogen transmission in insects (Kanakala and Ghanim, 2016). Ghanim et al. (2007) reported successful silencing in whiteflies by injecting dsRNA of against midgut and salivary gland genes, and their expression was significantly reduced down to 70% compared to control (Ghanim et al., 2007). Similarly, in another study, whiteflies feeding on the ds/ siRNA of the ribosomal protein L9 (RPL9) and the Vacuolar-type ATPase A subunit (v-ATPase A) have been reported to cause mortality (Upadhyay et al., 2011). A significant reduction in the transcript levels of the target genes and mortality in whiteflies have been achieved by expressing dsRNA in transgenic tobacco plants that target the v-ATPase A (Thakur et al., 2014), osmoregulators (Raza et al., 2016), P450 CYP6M1 (Li et al., 2015), and acetylcholinesterase and the ecdysone receptors (Malik et al., 2016). A recent study identified a fern (Tectaria macrodonta) protein Tma12 that exhibits insecticidal activity against B. tabaci (Shukla et al., 2016). Chen et al. (2015) used the RNAi strategy to cause mortality of B. tabaci by expressing the Toll-Like Receptor 7 (TLR7) in the entomapthogenic fungus Isaria fumosorosea. To maximize protection, Javaid et al. (2016) targeted two genes from the onion leaf lectin and a neurotoxin (Hvt) from the venom of a spider (Hadronyche versuta), targeting multiple sap-sucking pests, including B. tabaci. Whitefly fitness has been also affected by suppressing terpenoid synthesis in transgenic tobacco plants via plant-mediated gene silencing (Luan et al., 2013). Another report has demonstrated the suppression of silencing protein kinase (GhMPK3) through virusinduced gene silencing (VIGS), which resulted in the suppression of the MPK-WRKY-jasmonic acid ( JA) and the Ethylene (ET) pathways, and resulted in enhanced whitefly susceptibility (Li et al., 2016).

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These recent studies suggest that the employment of these genes targeting whitefly development/ pathways with the above approaches will reduce the use of chemical pesticides and achieve broadspectrum resistance to different groups of insect pests and vectors. Conclusion and future perspectives In this chapter, we summarized the worldwide reported begomoviruses and the B. tabaci cryptic species; the mechanisms and proteins involved in whitefly–begomovirus interactions; and the recent advances in whitefly control. Very few hosts and symbiont proteins involved in whitefly–pathogen interactions were investigated. These studies add more evidence for the potential uses of these genes in controlling insect vectors and their transmitted viruses. So far, there has been no universal set of target genes available for broad spectrum protection against hemipteran insects; however, recent studies will open up new avenues for insect vectors control using genetic methods. Like using Bacillus thuringiensis toxins against lepidopterans, the RNAi approach can be used as an insecticide-free method to control insect vectors, however, this will greatly depend upon the identification and selection of suitable genes for silencing. References

Alves-Júnior, M., Alfenas-Zerbini, P., Andrade, E.C., Esposito, D.A., Silva, F.N., F da Cruz, A.C., Ventrella, M.C., Otoni, W.C., and Zerbini, F.M. (2009). Synergism and negative interference during co-infection of tomato and Nicotiana benthamiana with two bipartite begomoviruses. Virology 387, 257–266. https://doi. org/10.1016/j.virol.2009.01.046 Blawid, R., Van, D.T., and Maiss, E. (2008). Transreplication of a Tomato yellow leaf curl Thailand virus DNA-B and replication of a DNAbeta component by Tomato leaf curl Vietnam virus and Tomato yellow leaf curl Vietnam virus. Virus Res. 136, 107–117. https://doi. org/10.1016/j.virusres.2008.04.025 Bleeker, P.M., Diergaarde, P.J., Ament, K., Guerra, J., Weidner, M., Schütz, S., de Both, M.T., Haring, M.A., and Schuurink, R.C. (2009). The role of specific tomato volatiles in tomato-whitefly interaction. Plant Physiol. 151, 925–935. https://doi.org/10.1104/pp.109.142661 Boykin, L.M., Shatters, R.G., Rosell, R.C., McKenzie, C.L., Bagnall, R.A., De Barro, P.J., and Frohlich, D.R. (2007). Global relationships of Bemisia tabaci Hemiptera, Aleyrodidae revealed using Bayesian analysis of mitochondrial CO1 DNA sequence. Mol Phylogenet Evol 44, 1306–1319.

Briddon, R.W., and Markham, P.G. (2000). Cotton leaf curl virus disease. Virus Res. 71, 151–159. Caciagli, P., Medina Piles, V., Marian, D., Vecchiati, M., Masenga, V., Mason, G., Falcioni, T., and Noris, E. (2009). Virion stability is important for the circulative transmission of Tomato yellow leaf curl Sardinia virus by Bemisia tabaci, but virion access to salivary glands does not guarantee transmissibility. J. Virol. 83, 5784–5795. https://doi.org/10.1128/JVI.02267-08 Castle, S.J., Palumbo, J.C., Prabhaker, N., Horowitz, A.R., and Denholm, I. (2010). Ecological determinants of Bemisia tabaci resistance to insecticides. In Bemisia, bionomics and management of a global pest, P.A. Stansly, S.E. Naranjo, eds. (Dordrecht: Springer), pp. 423-465. Chakraborty, S., Vanitharani, R., Chattopadhyay, B., and Fauquet, C.M. (2008). Supervirulent pseudorecombination and asymmetric synergism between genomic components of two distinct species of begomovirus associated with severe tomato leaf curl disease in India. J. Gen. Virol. 89, 818–828. Chen, X., Li, L., Hu, Q., Zhang, B., Wu, W., Jin, F., and Jiang, J. (2015). Expression of dsRNA in recombinant Isaria fumosorosea strain targets the TLR7 gene in Bemisia tabaci. BMC Biotechnol. 15, 64. https://doi. org/10.1186/s12896-015-0170-8 Chiel, E., Inbar, M., Mozes-Daube, N., White, J.A., Hunter, M.S., and Zchori-Fein, E. (2009). Assessments of Fitness Effects by the Facultative Symbiont Rickettsia in theSweetpotato Whitefly (Hemiptera: Aleyrodidae). Ann. Entomol. Soc. Am. 102, 413–418. Chu, D., Zhang, Y.J., and Wan, F.H. (2010). Cryptic invasion of the exotic Bemisia tabaci biotype Q occurred widespread in Shandong Province of China. Flo Entomol 93, 203-207. Davino, S., Napoli, C., Davino, M., and Accotto, G.P. (2006). Spread of Tomato yellow leaf curl virus in Sicily, partial displacement of another geminivirus originally present. Eur J Plant Pathol 114, 293-299. De Barro, P.J., Liu, S.S., Boykin, L.M., and Dinsdale, A.B. (2011). Bemisia tabaci: a statement of species status. Annu. Rev. Entomol. 56, 1–19. https://doi. org/10.1146/annurev-ento-112408-085504 De la Rúa, P., Simon, B., Cifuentes, D., Martinez‐Mora, C.L.C.J., and Cenis, J.L. (2006). New insights into the mitochondrial phylogeny of the whitefly Bemisia tabaci (Hemiptera, Aleyrodidae) in the Mediterranean Basin. J Zool Syst Evol Res 44, 25-33. Dennehy, T.J., DeGain, B., Harpold, G., Brown, J.K., Byrne, F., Morin, S., and Nichols, R. (2006). First New World report of Q biotype of Bemisia tabaci (Gennadius) reveals high levels of resistance to insecticides. Resistant Pest Management Newsletter 15, 18-20. Dinsdale, A., Cook, L., Riginos, C., Buckley, Y.M., and De Barro, P. (2010). Refined global analysis of Bemisia tabaci(Hemiptera, Sternorrhyncha, Aleyrodoidea, Aleyrodidae) mitochondrial cytochrome oxidase 1 to identify species level genetic boundaries. Ann. Entomol. Soc. Am. 103, 196-208. Ellango, R., Singh, S.T., Rana, V.S., Gayatri Priya, N., Raina, H., Chaubey, R., Naveen, N.C., Mahmood, R., Ramamurthy, V.V., Asokan, R., et al. (2015). Distribution of Bemisia tabaci Genetic Groups in India. Environ.

216  | Kanakala and Ghanim

Entomol. 44, 1258–1264. https://doi.org/10.1093/ee/ nvv062 Escobar-Bravo, R., Alba, J.M., Pons, C., Granell, A., Kant, M.R., Moriones, E., and Fernández-Muñoz, R. (2016). A jasmonate-inducible defense trait transferred from wild into cultivated tomato establishes increased whitefly resistance and reduced viral disease incidence. Front. Plant Sci. 7, 1732. https://doi.org/10.3389/ fpls.2016.01732 Firdaus, S., van Heusden, A.W., Hidayati, N., Supena, E.D.J., Visser, R.G., and Vosman, B. (2012). Resistance to Bemisia tabaci in tomato wild relatives. Euphytica 187, 31-45. Firdaus, S., Vosman, B., Hidayati, N., Jaya Supena, E.D., Visser, R.G., and van Heusden, A.W. (2013). The Bemisia tabaci species complex: additions from different parts of the world. Insect Sci. 20, 723–733. https://doi. org/10.1111/1744-7917.12001 Fischer, A., Strohmeier, S., Krenz, B., and Jeske, H. (2015). Evolutionary liberties of the Abutilon mosaic virus cluster. Virus Genes 50, 63–70. https://doi.org/10.1007/ s11262-014-1125-1 Fondong, V.N., Pita, J.S., Rey, M.E.C., De Kochko, A., Beachy, R.N., and Fauquet, C.M. (2000). Evidence of synergism between African Cassava mosaic virus and a new double-recombinant geminivirus infecting cassava in Cameroon. J. Gen. Virol. 81, 287-297. Freitas, J.A., Maluf, W.R., das Graças Cardoso, M., Gomes, L.A., and Bearzotti, E. (2002). Inheritance of foliar zingiberene contents and their relationship to trichome densities and whitefly resistance in tomatoes Euphytica 127, 275–287. Gennadius, P. (1889). Disease of tobacco plantations in the Trikonia. The aleyrodid of tobacco. Ellenike Georgia 5, 1–3. Ghanim, M., Kontsedalov, S., and Czosnek, H. (2007). Tissue-specific gene silencing by RNA interference in the whitefly Bemisia tabaci (Gennadius). Insect Biochem. Mol. Biol. 37, 732–738. Ghanim, M. (2014). A review of the mechanisms and components that determine the transmission efficiency of Tomato yellow leaf curl virus (Geminiviridae; Begomovirus) by its whitefly vector. Virus Res. 186, 47–54. https://doi.org/10.1016/j.virusres.2014.01.022 Goodman, R.M., and Bird, J. (1978). Bean golden mosaic virus. CMI/AAB Descriptions of Plant Viruses. 192. Gottlieb, Y., Zchori-Fein, E., Mozes-Daube, N., Kontsedalov, S., Skaljac, M., Brumin, M., Sobol, I., Czosnek, H., Vavre, F., Fleury, F., et al. (2010). The transmission efficiency of Tomato yellow leaf curl virus by the whitefly Bemisia tabaci is correlated with the presence of a specific symbiotic bacterium species. J. Virol. 84, 9310–9317. https://doi. org/10.1128/JVI.00423-10 Götz, M., and Winter, S. (2016). Diversity of Bemisia tabaci in Thailand and Vietnam and indications of species replacement. J. Asia Pac. Entomol. 19, 537-543. Götz, M., Popovski, S., Kollenberg, M., Gorovits, R., Brown, J.K., Cicero, J.M., Czosnek, H., Winter, S., and Ghanim, M. (2012). Implication of Bemisia tabaci heat shock protein 70 in Begomovirus-whitefly interactions. J. Virol. 86, 13241–13252. https://doi.org/10.1128/JVI.0088012 Gueguen, G., Vavre, F., Gnankine, O., Peterschmitt, M., Charif, D., Chiel, E., Gottlieb, Y., Ghanim, M., .

Zchori-Fein, E., and Fleury, F. (2010). Endosymbiont metacommunities, mtDNA diversity and the evolution of the Bemisia tabaci (Hemiptera: Aleyrodidae) species complex. Mol. Ecol. 19, 4365–4376. https://doi. org/10.1111/j.1365-294X.2010.04775.x Hariton Shalev, A., Sobol, I., Ghanim, M., Liu, S.S., and Czosnek, H. (2016). The Whitefly Bemisia tabaci Knottin-1 Gene Is Implicated in Regulating the Quantity of Tomato yellow leaf curl virus Ingested and Transmitted by the Insect. Viruses 8, E205. https://doi. org/10.3390/v8070205 Hassan, I., Orílio, A.F., Fiallo-Olivé, E., Briddon, R.W., and Navas-Castillo, J. (2016). Infectivity, effects on helper viruses and whitefly transmission of the deltasatellites associated with sweepoviruses (genus Begomovirus, family Geminiviridae). Sci Rep 6, 30204. https://doi. org/10.1038/srep30204 Höfer, P., Bedford, I.D., Markham, P.G., Jeske, H., and Frischmuth, T. (1997). Coat protein gene replacement results in whitefly transmission of an insect nontransmissible geminivirus isolate. Virology 236, 288–295. Hogenhout, S.A., Ammar, e.l.-.D., Whitfield, A.E., and Redinbaugh, M.G. (2008). Insect vector interactions with persistently transmitted viruses. Annu Rev Phytopathol 46, 327–359. https://doi.org/10.1146/ annurev.phyto.022508.092135 Höhnle, M., Höfer, P., Bedford, I.D., Briddon, R.W., Markham, P.G., and Frischmuth, T. (2001). Exchange of three amino acids in the coat protein results in efficient whitefly transmission of a nontransmissible Abutilon mosaic virus isolate. Virology 290, 164–171. Horowitz, A.R., and Ishaaya, I. (2014). Dynamics of biotypes B and Q of the whitefly Bemisia tabaci and its impact on insecticide resistance. Pest Manag. Sci. 70, 1568–1572. https://doi.org/10.1002/ps.3752 Horowitz, A.R., Antignus, Y., and Gerling, D. (2011). Management of Bemisia tabaci whiteflies. In The Whitefly, Bemisia tabaci (Homoptera, Aleyrodidae) Interaction with Geminivirus-Infected Host Plants, W.M. Thompson, ed. (Netherlands: Springer). pp. 293322. Hsieh, C.H., Wang, C.H., and Ko, C.C. (2011). Population genetic structure of the newly invasive Q biotype of Bemisiatabaci in Taiwan. Entomol Exp Appl 138, 263–171. Hu, J., De Barro, P., Zhao, H., Wang, J., Nardi, F., and Liu, S.S. (2011). An extensive field survey combined with a phylogenetic analysis reveals rapid and widespread invasion of two alien whiteflies in China. PLOS ONE 6, e16061. https://doi.org/10.1371/journal. pone.0016061 Hu, J., Zhang, X., Jiang, Z., Zhang, F., Liu, Y., Li, Z., and Zhang, Z. (2017). New putative cryptic species detection and genetic network analysis of Bemisia tabaci (Hempitera: Aleyrodidae) in China based on mitochondrial COI sequences. Mitochondrial DNA A DNA Mapp Seq Anal [Epub ahead of print]. https:// doi.org/10.1080/24701394.2017.1307974 Javaid, S., Amin, I., Jander, G., Mukhtar, Z., Saeed, N.A., and Mansoor, S. (2016). A transgenic approach to control hemipteran insects by expressing insecticidal genes under phloem-specific promoters. Sci Rep 6, 34706. https://doi.org/10.1038/srep34706

Whitefly-Transmitted Begomoviruses and Control of their Vectors |  217

Jones, D.R. (2003). Plant viruses transmitted by whiteflies. Eur J Plant Pathol 109, 195-219. Jones, R.A. (2009). Plant virus emergence and evolution: origins, new encounter scenarios, factors driving emergence, effects of changing world conditions, and prospects for control. Virus Res. 141, 113–130. https:// doi.org/10.1016/j.virusres.2008.07.028 Kanakala, S., and Ghanim, M. (2016). Implication of the whitefly Bemisia tabaci cyclophilin B protein in the transmission of Tomato yellow leaf curl virus. Front. Plant Sci. 7, 1702. https://doi.org/10.3389/fpls.2016.01702 Kanakala, S., and Ghanim, M. (2016). RNA interference in insect vectors for plant viruses. Viruses 8, E329. Kanakala, S., Jyothsna, P., Shukla, R., Tiwari, N., Veer, B.S., Swarnalatha, P., Krishnareddy, M., and Malathi, V.G. (2013). Asymmetric synergism and heteroencapsidation between two bipartite begomoviruses, Tomato leaf curl New Delhi virus and tomato leaf curl Palampur virus. Virus Res. 174, 126–136. https://doi.org/10.1016/j. virusres.2013.03.011 Khan, A.J., Akhtar, S., Al-Matrushi, A.M., Fauquet, C.M., and Briddon, R.W. (2013). Introduction of East African cassava mosaic Zanzibar virus to Oman harks back to ‘Zanzibar, the capital of Oman’. Virus Genes 46, 195–198. https://doi.org/10.1007/s11262-012-0838-2 Kheyr-Pour, A., Bananej, K., Dafalla, G.A., Caciagli, P., Noris, E., Ahoonmanesh, A., Lecoq, H., and Gronenborn, B. (2000). Watermelon chlorotic stunt virus from the Sudan and Iran: Sequence Comparisons and Identification of a Whitefly-Transmission Determinant. Phytopathology 90, 629–635. https://doi.org/10.1094/ PHYTO.2000.90.6.629 Kliot, A., Cilia, M., Czosnek, H., and Ghanim, M. (2014). Implication of the bacterial endosymbiont Rickettsia spp. in interactions of the whitefly Bemisia tabaci with Tomato yellow leaf curl virus. J. Virol. 88, 5652–5660. https://doi. org/10.1128/JVI.00071-14 Kollenberg, M., Winter, S., and Götz, M. (2014). Quantification and localization of Watermelon chlorotic stunt virus and Tomato yellow leaf curl virus (Geminiviridae) in populations of Bemisia tabaci (Hemiptera, Aleyrodidae) with differential virus transmission characteristics. PLOS ONE 9, e111968. https://doi.org/10.1371/journal.pone.0111968 Kontsedalov, S., Abu-Moch, F., Lebedev, G., Czosnek, H., Horowitz, A.R., and Ghanim, M. (2012). Bemisia tabaci biotype dynamics and resistance to insecticides in Israel during the years 2008–2010. Journal of Integrative Agriculture 11, 312-320. Lee, W., Park, J., Lee, G.S., Lee, S., and Akimoto, S. (2013). Taxonomic status of the Bemisia tabaci complex (Hemiptera: Aleyrodidae) and reassessment of the number of its constituent species. PLOS ONE 8, e63817. https://doi.org/10.1371/journal.pone.0063817 Legg, J.P., Shirima, R., Tajebe, L.S., Guastella, D., Boniface, S., Jeremiah, S., Nsami, E., Chikoti, P., and Rapisarda, C. (2014). Biology and management of Bemisia whitefly vectors of cassava virus pandemics in Africa. Pest Manag. Sci. 70, 1446–1453. https://doi.org/10.1002/ps.3793 Li, J., Zhu, L., Hull, J.J., Liang, S., Daniell, H., Jin, S., and Zhang, X. (2016). Transcriptome analysis reveals a comprehensive insect resistance response mechanism in cotton to infestation by the phloem feeding insect

Bemisia tabaci (whitefly). Plant Biotechnol. J. 14, 1956– 1975. https://doi.org/10.1111/pbi.12554 Li, J., Zhu, L., Hull, J.J., Liang, S., Daniell, H., Jin, S., and Zhang, X. (2016). Transcriptome analysis reveals a comprehensive insect resistance response mechanism in cotton to infestation by the phloem feeding insect Bemisia tabaci (whitefly). Plant Biotechnol. J. 14, 1956– 1975. https://doi.org/10.1111/pbi.12554 Li, P., Shu, Y.N., Fu, S., Liu, Y.Q., Zhou, X.P., Liu, S.S., and Wang, X.W. (2017). Vector and nonvector insect feeding reduces subsequent plant susceptibility to virus transmission. New Phytol. 215, 699–710. https://doi. org/10.1111/nph.14550 Liu, T.X., Stansly, P.A., and Gerling, D. (2015). Whitefly parasitoids: distribution, life history, bionomics, and utilization. Annu. Rev. Entomol. 60, 273–292. https:// doi.org/10.1146/annurev-ento-010814-021101 Luan, J.B., Yao, D.M., Zhang, T., Walling, L.L., Yang, M., Wang, Y.J., and Liu, S.S. (2013). Suppression of terpenoid synthesis in plants by a virus promotes its mutualism with vectors. Ecol. Lett. 16, 390–398. https://doi.org/10.1111/ele.12055 Luan, J.B., Yao, D.M., Zhang, T., Walling, L.L., Yang, M., Wang, Y.J., and Liu, S.S. (2013). Suppression of terpenoid synthesis in plants by a virus promotes its mutualism with vectors. Ecol. Lett. 16, 390–398. https://doi.org/10.1111/ele.12055 Luo, C., Jones, C.M., Devine, G., Zhang, F., Denholm, I., and Gorman, K. (2010). Insecticide resistance in Bemisia tabaci biotype Q (Hemiptera, Aleyrodidae) from China. Crop Prot 29, 429-434. Malik, H.J., Raza, A., Amin, I., Scheffler, J.A., Scheffler, B.E., Brown, J.K., and Mansoor, S. (2016). RNAimediated mortality of the whitefly through transgenic expression of double-stranded RNA homologous to acetylcholinesterase and ecdysone receptor in tobacco plants. Sci Rep 6, 38469. https://doi.org/10.1038/ srep38469 Martin, J.H., Mifsud, D., and Rapisarda, C. (2000). The whiteflies (Hemiptera: Aleyrodidae) of Europe and the Mediterranean Basin. Bull. Entomol. Res. 90, 407–448. Misra, C.S., and Lamba, S.K. (1929). The cotton whitefly (Bemisia gossypiperda n. sp.). Agric Res Inst Pusa 196, 7. Monci, F., Sánchez-Campos, S., Navas-Castillo, J., and Moriones, E. (2002). A natural recombinant between the geminiviruses Tomato yellow leaf curl Sardinia virus and Tomato yellow leaf curl virus exhibits a novel pathogenic phenotype and is becoming prevalent in Spanish populations. Virology 303, 317–326. Mouton, L., Gnankiné, O., Henri, H., Terraz, G., Ketoh, G., Martin, T., Fleury, F., and Vavre, F. (2015). Detection of genetically isolated entities within the Mediterranean species of Bemisia tabaci: new insights into the systematics of this worldwide pest. Pest Manag. Sci. 71, 452–458. https://doi.org/10.1002/ps.3834 Muigai, S.G., Schuster, D.J., Snyder, J.C., Scott, J.W., Bassett, M.J., and McAuslane, H.J. (2002). Mechanisms of resistance in Lycopersicon germplasm to the whitefly Bemisia argentifolii. Phytoparasitica 30, 347-360. Navot, N., Pichersky, E., Zeidan, M., Zamir, D., and Czosnek, H. (1991). Tomato yellow leaf curl virus: a whitefly-transmitted geminivirus with a single genomic component. Virology 185, 151–161.

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Nawaz-ul-Rehman, M.S., Briddon, R.W., and Fauquet, C.M. (2012). A melting pot of Old World begomoviruses and their satellites infecting a collection of Gossypium species in Pakistan. PLOS ONE 7, e40050. https://doi. org/10.1371/journal.pone.0040050 Ning, W., Shi, X., Liu, B., Pan, H., Wei, W., Zeng, Y., Sun, X., Xie, W., Wang, S., Wu, Q., et al. (2015). Transmission of Tomato yellow leaf curl virus by Bemisia tabaci as Affected by Whitefly Sex and Biotype. Sci Rep 5, 10744. https:// doi.org/10.1038/srep10744 Nombela, G., and Muñiz, M. (2009). Host plant resistance for the management of Bemisia tabaci, a multi-crop survey with emphasis on tomato. In Bemisia, bionomics and management of a global pest, P.A. Stansly, S.E. Naranjo, eds. (Dordrecht: Springer), pp. 357-383. Nombela, G., Beitia, F., and Muñiz, M. (2000). Variation in tomato host response to Bemisia tabaci (Hemiptera: Aleyrodidae) in relation to acyl sugar content and presence of the nematode and potato aphid resistance gene Mi. Bull. Entomol. Res. 90, 161–167. Nombela, G., Beitia, F., and Muñiz, M. (2001). A differential interaction study of Bemisia tabaci Q‐biotype on commercial tomato varieties with or without the Mi resistance gene, and comparative host responses with the B‐biotype. Entomol Exp Appl 98, 339-344. Nombela, G., Williamson, V.M., and Muñiz, M. (2003). The root-knot nematode resistance gene Mi-1.2 of tomato is responsible for resistance to the whitefly Bemisia tabaci. Mol. Plant Microbe Interact. 16, 645–649. https://doi. org/10.1094/MPMI.2003.16.7.645 Noris, E., Vaira, A.M., Caciagli, P., Masenga, V., Gronenborn, B., and Accotto, G.P. (1998). Amino acids in the capsid protein of Tomato yellow leaf curl virus that are crucial for systemic infection, particle formation, and insect transmission. J. Virol. 72, 10050–10057. Ohnesorge, S., and Bejarano, E.R. (2009). Begomovirus coat protein interacts with a small heat-shock protein of its transmission vector (Bemisia tabaci). Insect Mol. Biol. 18, 693–703. https://doi.org/10.1111/j.13652583.2009.00906.x Pakkianathan, B.C., Kontsedalov, S., Lebedev, G., Mahadav, A., Zeidan, M., Czosnek, H., and Ghanim, M. (2015). Replication of Tomato yellow leaf curl virus in Its Whitefly Vector, Bemisia tabaci. J. Virol. 89, 9791–9803. https:// doi.org/10.1128/JVI.00779-15 Pan, H., Chu, D., Ge, D., Wang, S., Wu, Q., Xie, W., Jiao, X., Liu, B., Yang, X., Yang, N., et al. (2011). Further spread of and domination by Bemisia tabaci (Hemiptera: Aleyrodidae) biotype Q on field crops in China. J. Econ. Entomol. 104, 978–985. Pan, H., Chu, D., Yan, W., Su, Q., Liu, B., Wang, S., Wu, Q., Xie, W., Jiao, X., Li, R., et al. (2012). Rapid spread of Tomato yellow leaf curl virus in China is aided differentially by two invasive whiteflies. PLOS ONE 7, e34817. https:// doi.org/10.1371/journal.pone.0034817 Pan, L.L., Chen, Q.F., Zhao, J.J., Guo, T., Wang, X.W., Hariton-Shalev, A., Czosnek, H., and Liu, S.S. (2017). Clathrin-mediated endocytosis is involved in Tomato yellow leaf curl virus transport across the midgut barrier of its whitefly vector. Virology 502, 152–159. Papayiannis, L.C., Katis, N.I., Idris, A.M., and Brown, J.K., (2011). Identification of weed hosts of Tomato yellow leaf curl virus in Cyprus. Plant Dis. 95, 120–125.

Patil, B.L., and Fauquet, C.M. (2009). Cassava mosaic geminiviruses: actual knowledge and perspectives. Mol. Plant Pathol. 10, 685–701. https://doi.org/10.1111/ j.1364-3703.2009.00559.x Pita, J.S., Fondong, V.N., Sangare, A., Otim-Nape, G.W., Ogwal, S., and Fauquet, C.M. (2001). Recombination, pseudorecombination and synergism of geminiviruses are determinant keys to the epidemic of severe cassava mosaic disease in Uganda. J Gen Virol 82, 655-665. Qin, L., Pan, L.L., and Liu, S.S. (2016). Further insight into reproductive incompatibility between putative cryptic species of the Bemisia tabaci whitefly complex. Insect Sci. 23, 215–224. https://doi.org/10.1111/17447917.12296 Quaintance, A.L. (1900). Contribution towards a monograph of the American Aleurodidae. US Department of Agriculture. Technica Series. Bur Entomol 8, 9–64. Rana, V.S., Popli, S., Saurav, G.K., Raina, H.S., Chaubey, R., Ramamurthy, V.V., and Rajagopal, R. (2016). A Bemisia tabaci midgut protein interacts with begomoviruses and plays a role in virus transmission. Cell. Microbiol. 18, 663–678. https://doi.org/10.1111/cmi.12538 Rana, V.S., Singh, S.T., Priya, N.G., Kumar, J., and Rajagopal, R. (2012). Arsenophonus GroEL interacts with CLCuV and is localized in midgut and salivary gland of whitefly B. tabaci. PLOS ONE 7, e42168. https://doi. org/10.1371/journal.pone.0042168 Raza, A., Malik, H.J., Shafiq, M., Amin, I., Scheffler, J.A., Scheffler, B.E., and Mansoor, S. (2016). RNA interference based approach to down-regulate osmoregulators of whitefly (Bemisia tabaci): potential technology for the control of whitefly. PLOS ONE 11, e0153883. https:// doi.org/10.1371/journal.pone.0153883 Rodríguez-Álvarez, C.I., Muñiz, M., and Nombela, G. (2017). Effect of plant development (age and size) on the Mi-1-mediated resistance of tomato to whitefly Bemisia tabaci. Bull. Entomol. Res. 107, 768–776. https://doi.org/10.1017/S0007485317000281 Rodríguez-López, M.J., Garzo, E., Bonani, J.P., Fereres, A., Fernández-Muñoz, R., and Moriones, E. (2011). Whitefly resistance traits derived from the wild tomato Solanum pimpinellifolium affect the preference and feeding behavior of Bemisia tabaci and reduce the spread of Tomato yellow leaf curl virus. Phytopathology 101, 1191–1201. https://doi.org/10.1094/PHYTO-01-110028 Rosario, K., Seah, Y.M., Marr, C., Varsani, A., Kraberger, S., Stainton, D., Moriones, E., Polston, J.E., Duffy, S., and Breitbart, M. (2015). Vector-enabled metagenomic (VEM) surveys using whiteflies (Aleyrodidae) reveal novel begomovirus species in the New and Old Worlds. Viruses 7, 5553–5570. https://doi.org/10.3390/ v7102895 Russell, L.M. (1957). Synonyms of Bemisia tabaci (Gennadius) (Homoptera, Aleyrodidae). Bull. Brooklyn Entomol Soc. 52, 122–123. Rybicki, E.P., and Pietersen, G. (1999). Plant virus disease problems in the developing world. Adv. Virus Res. 53, 127–175. Sánchez-Campos, S., Navas-Castillo, J., Camero, R., Soria, C., Díaz, J.A., and Moriones, E. (1999). Displacement of Tomato yellow leaf curl virus (TYLCV)-Sr by TYLCV-Is

Whitefly-Transmitted Begomoviruses and Control of their Vectors |  219

in Tomato Epidemics in Spain. Phytopathology 89, 1038–1043. https://doi.org/10.1094/ PHYTO.1999.89.11.1038 Saunders, K., Bedford, I.D., Yahara, T., and Stanley, J. (2003). Aetiology: The earliest recorded plant virus disease. Nature 422, 831. https://doi.org/10.1038/422831a Shukla, A.K., Upadhyay, S.K., Mishra, M., Saurabh, S., Singh, R., Singh, H., Thakur, N., Rai, P., Pandey, P., Hans, A.L., and Srivastava, S. (2016). Expression of an insecticidal fern protein in cotton protects against whitefly. Nature Biotechnol. 34, 1046–1051. Silva, F.N., Lima, A.T., Rocha, C.S., Castillo-Urquiza, G.P., Alves-Júnior, M., and Zerbini, F.M. (2014). Recombination and pseudorecombination driving the evolution of the begomoviruses Tomato severe rugose virus (ToSRV) and Tomato rugose mosaic virus (ToRMV): two recombinant DNA-A components sharing the same DNA-B. Virol. J. 11, 66. https://doi. org/10.1186/1743-422X-11-66 Simmer, F., Tijsterman, M., Parrish, S., Koushika, S.P., Nonet, M.L., Fire, A., Ahringer, J., and Plasterk, R.H. (2002). Loss of the putative RNA-directed RNA polymerase RRF-3 makes C. elegans hypersensitive to RNAi. Curr. Biol. 12, 1317–1319. Sinisterra, X.H., McKenzie, C.L., Hunter, W.B., Powell, C.A., and Shatters Jr, R.G. (2005). Differential transcriptional activity of plant-pathogenic begomoviruses in their whitefly vector (Bemisia tabaci, Gennadius, Hemiptera Aleyrodidae). J Gen Virol, 86, 1525-1532. Stansly, P.A., and Natwick, A. (2010). Integrated systems for managing Bemisia tabaci in protected and open field agriculture. In Bemisia, bionomics and management of a global pest, P.A. Stansly, S.E. Naranjo, eds. (Dordrecht: Springer), pp. 467-497. Sun, Y.C., Pan, L.L., Ying, F.Z., Li, P., Wang, X.W., and Liu, S.S. (2017). Jasmonic acid-related resistance in tomato mediates interactions between whitefly and whitefly-transmitted virus. Sci Rep 7, 566. https://doi. org/10.1038/s41598-017-00692-w Tabein, S., Behjatnia, S.A., Anabestani, A., and Izadpanah, K. (2013). Whitefly-mediated transmission of cotton leaf curl Multan betasatellite: evidence for betasatellite encapsidation in coat protein of helper begomoviruses. Arch. Virol. 158, 19–26. https://doi.org/10.1007/ s00705-012-1448-y Thakur, N., Upadhyay, S.K., Verma, P.C., Chandrashekar, K., Tuli, R., and Singh, P.K. (2014). Enhanced whitefly resistance in transgenic tobacco plants expressing double-stranded RNA of v-ATPase A gene. PLOS ONE 9, e87235. https://doi.org/10.1371/journal. pone.0087235 Thresh, J.M., Otim-Nape, G.W., Thankappan, M., and Muniyappa, V. (1998). The mosaic diseases of cassava in Africa and India caused by whitefly-borne geminiviruses. Rev Plant Pathol 77, 935–945. Ueda, S., Onuki, M., Yamashita, M., and Yamato, Y. (2012). Pathogenicity and insect transmission of a begomovirus complex between Tomato yellow leaf curl virus and Ageratum yellow vein betasatellite. Virus Genes 44, 338–344. https://doi.org/10.1007/s11262-011-0691-8

Upadhyay, S.K., Chandrashekar, K., Thakur, N., Verma, P.C., Borgio, J.F., Singh, P.K., and Tuli, R. (2011). RNA interference for the control of whiteflies (Bemisia tabaci) by oral route. J. Biosci. 36, 153–161. Wang, L.L., Wang, X.R., Wei, X.M., Huang, H., Wu, J.X., Chen, X.X., Liu, S.S., and Wang, X.W. (2016). The autophagy pathway participates in resistance to Tomato yellow leaf curl virus infection in whiteflies. Autophagy 12, 1560–1574. https://doi.org/10.1080/15548627.2 016.1192749 Wang, S., Zhang, Y., Yang, X., Xie, W., and Wu, Q. (2017). Resistance Monitoring for Eight Insecticides on the Sweetpotato Whitefly (Hemiptera: Aleyrodidae) in China. J. Econ. Entomol. 110, 660–666. https://doi. org/10.1093/jee/tox040 Wang, Z.Z., Shi, M., Huang, Y.C., Wang, X.W., Stanley, D., and Chen, X.X. (2016). A peptidoglycan recognition protein acts in whitefly (Bemisia tabaci) immunity and involves in Begomovirus acquisition. Sci Rep 6, 37806. https://doi.org/10.1038/srep37806 Wei, J., Zhao, J.J., Zhang, T., Li, F.F., Ghanim, M., Zhou, X.P., Ye, G.Y., Liu, S.S., and Wang, X.W. (2014). Specific cells in the primary salivary glands of the whitefly Bemisia tabaci control retention and transmission of begomoviruses. J. Virol. 88, 13460–13468. https://doi. org/10.1128/JVI.02179-14 Xie, W., Liu, Y., Wang, S., Wu, Q., Pan, H., Yang, X., Guo, L., and Zhang, Y. (2014). Sensitivity of Bemisia tabaci (Hemiptera: Aleyrodidae) to several new insecticides in China: effects of insecticide type and whitefly species, strain, and stage. J. Insect Sci. 14, 261. https://doi. org/10.1093/jisesa/ieu123 Zarate, S.I., Kempema, L.A., and Walling, L.L. (2007). Silverleaf whitefly induces salicylic acid defenses and suppresses effectual jasmonic acid defenses. Plant Physiol. 143, 866–875. Zerbini, F.M., Briddon, R.W., Idris, A., Martin, D.P., Moriones, E., Navas-Castillo, J., Rivera-Bustamante, R., Roumagnac, P., and Varsani, A. (2017). ICTV virus taxonomy profiles, Geminiviridae. J Gen Virol 98, 131–133. Zhang, P.J., Broekgaarden, C., Zheng, S.J., Snoeren, T.A., van Loon, J.J., Gols, R., and Dicke, M. (2013). Jasmonate and ethylene signaling mediate whitefly-induced interference with indirect plant defense in Arabidopsis thaliana. New Phytol. 197, 1291–1299. https://doi. org/10.1111/nph.12106 Zhang, T., Luan, J.B., Qi, J.F., Huang, C.J., Li, M., Zhou, X.P., and Liu, S.S. (2012). Begomovirus-whitefly mutualism is achieved through repression of plant defences by a virus pathogenicity factor. Mol. Ecol. 21, 1294–1304. https://doi.org/10.1111/j.1365-294X.2012.05457.x Zhang, W.M., Fu, H.B., Wang, W.H., Piao, C.S., Tao, Y.L., Guo, D., and Chu, D. (2014). Rapid spread of a recently introduced virus (Tomato yellow leaf curl virus) and its vector Bemisia tabaci (Hemiptera: Aleyrodidae) in Liaoning Province, China. J. Econ. Entomol. 107, 98–104.

Virus-resistant Transgenic Tomato: Current Status and Future Prospects S.V. Ramesh1 and Shelly Praveen2*

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1ICAR-Central Plantation Crops Research Institute (ICAR-CPCRI), Kasaragod, Kerala, India.

2Division of Biochemistry, ICAR-Indian Agricultural Research Institute (ICAR-IARI), New Delhi, India.

*Correspondence: [email protected] https://doi.org/10.21775/9781910190814.11

Abstract Virus-resistant transgenic plants form an essential constituent of crop protection measures. Tomato is an important vegetable crop grown throughout the world for its nutritional benefits and tomatobased processed food consumption. However, the production levels of tomato are threatened by many viral infections. In the absence of resistant tomato genotypes (where available genetic sources or resistance are scarce), development of transgenic resistance to pathogenic viruses is indispensable. The last couple of decades have witnessed substantial progress in incorporating virus resistance trait in tomato. This chapter provides overview of the strategies and successful instances of transgenic virus resistance with special emphasis on prominent viruses infecting tomato. Various approaches to incorporate virus resistance in tomato from antisense RNA expression, through various RNA interference (RNAi) based strategies and foray in to genome editing techniques are discussed. The significant achievements made in developing transgenic resistance to combat Tomato leaf curl viruses, ground nut bud necrosis virus and Cucumber mosaic virus are presented. Also, the utility of employing recently emerging genome editing tool in incorporating resistance to tomato viruses is also discussed. Introduction Tomato is one of the important vegetable crops having tremendous global popularity. Due to its

delicious taste, rich in micronutrients and antioxidants, tomato is the most valuable crop grown around the world. The antioxidant potential of tomato owing to its lycopene content protects against oxidative damage to the cells and slows the process of ageing and provides protection against many stress related diseases. With a worldwide production of 17 million metric tonnes and occupying 72% of total value of fresh vegetable market, tomato crop has potential economic value. However, the cultivation and the production status of tomato are threatened due to many diverse species of infectious, pathogenic viruses. The number of described viral species that infect tomato crops amounts to 136, whereas this number is notably lower for other vegetable crops (Hanssen et al., 2010). One of the reasons for more number of viral species affecting tomato is the sensitivity of tomato crop to members of the genus Begomovirus, which comprises a large variety of species. With a limited genetic resource for disease resistance, intensive breeding for improved production and emergence of viral variants, tomato cultivation faces serious threat across the globe. Changing climate conditions also contribute to a successful spread of the virus or its vector in the areas that were previously unfavourable, thus enhancing viral spread in different geographical regions. Considering the most important viral diseases of tomato inflicting damage in the last two decades, and efforts of genetic engineering in developing transgenic crops are discussed in this chapter. Thus,

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this chapter encompasses (i) prominent viruses infecting tomato, (ii) approaches to develop resistance/tolerance to important viruses of tomato and (iii) possibilities of using gene editing tools for management of tomato viruses. Viruses infecting tomato Diverse group of viruses are known to naturally infect tomato and hence it is considered an ideal host for countless phytopathogenic viruses. Among the various groups of viruses that infect tomato, at least 10 viruses have been considered economically important and these viruses show great genetic variation due to distinct strain characteristic features (Brunt et al., 1996). Tomato leaf curl disease (ToLCD) is an economically important disease of tomato grown worldwide. It is caused by group of viruses belonging to family Geminiviridae, genus Begomovirus. The viruses are classified into Tomato leaf curl viruses (ToLCVs) and Tomato yellow leaf curl viruses (TYLCVs) based on the disease symptoms (Abhary et al., 2007). ToLCV and its variants are predominantly found in south Asia (Varma and Malathi, 2003). TYLCVs were discovered in Israel and Jordan in 1939 but the virus has wide spread presence now in Middle East, Mediterranean region, East Asia, Central America, Australia (Abhary et al., 2007; Boulton, 2003), Iran (Fazeli et al., 2009), South America (Melgarejo et al., 2013; and Europe ( Juarez et al., 2014). Besides differences in disease symptoms, TYLCV and ToLCV differ in their genomic components. TYLCVs are having monopartite single-stranded DNA as their genome, whereas ToLCVs are of two different types; some of them are monopartite while others have bipartite genome. Among ToLCVs, Tomato leaf curl New Delhi virus (ToLCNDV) is a bipartite virus prevalent in northern India. Recent reports (Fortes et al., 2016) suggest that ToLCNDV is not only extending its host range but also the geographical reach. Bud necrosis disease in tomato is another most severe disease in India caused by groundnut bud necrosis virus (GBNV), first reported by Heinze et al. (1995). It is the type member of the Orthotospovirus genus of Bunyaviridae family of virus. It is fast spreading and emerging as a serious pathogen of tomato in India (Umamaheswaran et al., 2003; Akram and Naimuddin, 2010). Diseases caused

by tospoviruses cause great losses in agriculture throughout the world (Goldbach and Kuo, 1996). Among them, Tomato spotted wilt virus (TSWV) is causing huge losses in different crops including tomato. It is prevalent in temperate, tropical and subtropical region. These tospoviruses carry tripartite single-stranded RNA as genome with ambisense polarity. Next to leaf curl and bud necrosis disease in tomato, Cucumber mosaic virus (CMV) is another important threat to tomato cultivation. Infection by CMV causes reduction in fruit production and quality. Cucumber mosaic virus (CMV) is the type member of the genus Cucumovirus, family Bromoviridae (Palukaitis et al., 1992). Cucumber mosaic disease, first described in 1916, was one of the earliest plant diseases caused by virus (Doolittle, 1916). CMV occurs worldwide especially in temperate region and causes severe damages in many vegetable crops including tomato. CMV carries multipartite (varying between 3–5 segments) single-stranded RNA as genome with positive sense polarity (Fig. 11.1). Management of viruses infecting tomato Natural source of resistance in tomato genotypes against viruses are very scanty. Mapping studies of tomato genotypes led to the identification of some resistance/tolerance genes, which are being exploited by breeders for developing resistance to viruses. The resistant loci (Ty-based) that are known for tomato yellow leaf curl viruses are also providing resistance to tomato leaf curl viruses although, same is not applicable in case of tospoviruses. The resistant loci Sw5 provide resistance to Tomato spotted wilt virus but not to groundnut bud necrosis virus. There are no resistance sources available to provide resistance to Cucumber mosaic virus. The resistance loci for TYLCVs and ToLCVs (Dominant gene: Ty-1, Ty-2, Ty-3, Ty-4 and Ty-6; recessive gene: ty-5) have different origins; Ty-1, Ty-3, Ty-4 and Ty-6 are introgressed from different accessions of Solanum chilense; Ty-2 is from S. habrochaites; ty-5 is from S. peruvianum. Instead of classical R gene mediated hyper-sensitive response (HR), these Ty loci were found to adopt a different mechanism of imparting resistance. The resistance loci Ty-1 and Ty-3 are allelic and were found to

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Figure 11.1 Phenotypic symptoms during viral infection in tomato and genomic organization of viruses infecting tomato. (A) (i) Leaf curl symptoms in tomato; (ii) single-stranded circular DNA genome (bipartite) of ToLCV; DNA-A and DNA-B. (B) (i) Bud necrosis symptoms in tomato, chlorotic rings on tomato fruit; (ii) singlestranded RNA genome (tripartite) of GBNV; L, large; M, medium; S, small; ambisense + and – polarity. (C) (i) Mosaic symptoms on tomato leaves, shoe-string symptoms in leaves; (ii) single-stranded RNA genome (multipartite) of CMV; RNA1-4 with satellite RNA, having + polarity.

belong to DFDGD class of RDR genes, which are involved in the biogenesis and amplification of siRNA required for transcriptional gene silencing by AGO4-mediated siRNA-directed DNA methylation (RdDM).

Due to lack of resistant sources available for different viruses in tomato, from the last three decades, genetic engineering has become a powerful tool in combating viral diseases of plants, through the transfer of alien genes, particularly those of

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viral origin, to the desired genotypes of plants. Pathogen-derived resistance (PDR) is an important disease-resistance strategy of engineering a virus gene into a plant. The molecular basis of PDR has been proposed to involve protein mediated defence response and also at nucleic acids based resistance to invading viruses. Further, protein– protein interactions has been the mainstay for coat protein-mediated resistance wherein expression of viral derived proteins in plants has provided resistance to invading homologous viruses and in some cases against heterologous viruses too. The defence response mediated at the nucleic acids level generally involves production of RNA molecules against specific target viruses. In other words, protection at nucleic acid level is sequence specific hence, homologous viruses are targeted. The concept of PDR (Sanford et al., 1985) has stimulated research on obtaining virus resistance through genetic engineering both at protein as well as at RNA level. Since, PDR is mediated either by the viral protein (protein-mediated) or by the viral transcript (RNAmediated), selection of viral protein or transcript for developing resistance is crucial and varies from virus to virus. Expression or suppression of viral protein affects the viral cycle by differential mechanisms (Table 11.1). The most favoured mode of transgenic resistance in crops was coat protein-mediated resistance (CPMR) because this strategy showed considerable increase in yields over non-transgenic or wildtype genotypes of Papaya and Squash (Gonsalves, 1996). Hence, transgenic tomato lines have been developed with a view to incorporate resistance to Tobacco mosaic virus (TMV), Tomato mosaic virus (ToMV) (Nelson et al., 1988; Sanders et al., 1992) and Cucumber mosaic virus (CMV) (Mayo, 1992; Tumer et al., 1987). Recent developments in the field of molecular biology have led to the identification of phenomenon of RNA silencing in plants. RNA silencing is a sequence dependent gene silencing process that plays significant role in repression of invading nucleic acids such as viruses, regulation of transposons and other cellular processes. RNA silencing that occurs in the nucleus is called as transcriptional gene silencing (TGS) and apparently affects the transcription of a gene by modulating cis-acting elements. However, gene silencing also occurs in the cytoplasm, wherein sequence specific

degradation of RNA transcripts cause shut down of its expression. This phenomenon of gene silencing affecting RNA turnover in cytoplasm is referred as post-transcriptional gene silencing (PTGS). The ultimate effector molecules of TGS and PTGS are small non-coding RNAs such as small interfering RNAs (siRNAs), microRNAs (miRNAs), transacting siRNAs (tasiRNAs), etc. Later, various other strategies, such as ectopic expression of viral derived siRNAs, artificial miRNAs (amiRNAs), artificial tasiRNAs (atasiRNAs), etc., have been employed to engineer virus resistance in tomato. In Indian context, transgenic resistance to leaf curl disease caused by Tomato leaf curl New Delhi virus is a significant achievement. The coat protein-mediated resistance was developed at National Botanical Research Institute (NBRI, Lucknow) whereas antisense RNA mediated resistance for ToLCD was achieved in ICAR-IARI, New Delhi (Praveen et al., 2005b, Raj et al., 2005). Coat protein-mediated resistance As discussed elaborately in one of the chapters by Patil BL in this book coat protein-mediated resistance (CPMR) is one of the most successful interventions of genetic engineering for virus resistance. Coat protein (CP) is an important structural protein that is involved not only in encapsidation of viral nucleic acid but also it plays significant role in acquisition of viral particles by insect vectors, intercellular and long distance movement of viral nucleic acids, etc. In fact, CPMR was the first strategy employed to develop antiviral resistance in plants when Powell et al., 1986 demonstrated its utility in tobacco plants to counter Tobacco mosaic virus (TMV) through the expression coat protein gene of TMV. Since then CPMR strategy has been widely employed for engineering resistance to wide range of phytopathogenic viruses including DNA viruses. In tomato, CPMR was successfully shown to impede the progress or infectivity of Alfalfa mosaic virus (Tumer et al., 1987), and Tomato yellow leaf curl virus (TYLCV) (Kunik et al., 1994), Cucumber mosaic virus (Xue et al., 1994; Provvidenti et al., 1995; Gielen et al., 1996). It has also been proven that the level of protection against virus infection was directly correlated to the level of CP expression. Further, resistance in CPMR strategy

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Table 11.1 Approaches for transgenic resistance to phytopathogenic viruses infecting tomato Transgene or No. strategy

Transgenic interference or mode of action

1

Vector transmission, Alfalfa mosaic virus uncoating, assembly of virus particle

2

3

Coat protein (CP)

5

References Tumer et al. (1987)

Tobacco mosaic virus (TMV),

Nelson et al. (1988)

Tomato mosaic virus (ToMV),

Sanders et al. (1992)

Tomato yellow leaf curl virus (TYLCV)

Kunik et al. (1994)

Cucumber mosaic virus (CMV)

Mayo (1992), Tumer et al. (1987), Xue et al. (1994), Provvidenti et al. (1995), Gielen et al. (1996)

Tomato leaf curl New Delhi virus (ToLCNDV)

Raj et al. (2005)

Cucumber mosaic virus (CMV)

Gal-On et al. (1998)

Defective replicase

Tomato yellow leaf curl virus

Lapidot and Friedmann (2002)

Full length replicase

Tomato yellow leaf curl Sardinia virus (TYLCSV)

Brunetti et al. (2001); Lucioli et al. (2003); Prins et al. (2008)

Truncated replicase

Tomato yellow leaf curl virus

Antignus et al. (2004)

Tomato leaf curl virus

Chatterji et al. (2001)

Tomato yellow leaf curl virus (TYLCV)

Bendahmane and Gronenborn (1997)

Tomato yellow leaf curl virus (TYLCV)

Yang et al. (2004)

Replication associated protein (rep)

Antisense RNA

Viral replication, inhibition of viral gene transcription, protein–protein interaction

DNA replication, translation, assembly, RNA silencing

dsRNA threshold for siRNA based gene silencing 4

target virus

Small interfering RNA silencing RNAs (siRNAs)

Artificial microRNAs (amiRNAs)

Tomato leaf curl virus (ToLCV) Praveen et al. (2005a) Tomato yellow leaf curl Sardinia virus

Noris et al. (2004)

Tomato yellow leaf curl virus (TYLCV),

Fuentes et al. (2006)

Multiple siRNA production strategies

Tomato leaf curl virus (ToLCV) Ramesh et al. (2007)

Varied arm length of hairpin RNA

Tomato leaf curl virus (ToLCV) Praveen et al. (2010)

RNA silencing

Tomato leaf curl virus (ToLCV) Praveen et al. (2007)

Cucumber mosaic virus (CMV), Tobacco mosaic virus and Tomato yellow leaf curl virus

Zhang et al. (2011)

Tomato leaf curl virus (ToLCV) Van Tu et al. (2013) 6

Artificial transacting siRNAs (atasiRNAs)

RNA silencing

Tomato leaf curl New Delhi virus (ToLCNDV) and tomato leaf curl Gujarat virus (ToLCGV)

Singh et al. (2015)

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Table 11.1 Continued Transgene or No. strategy

Transgenic interference or mode of action

7

RNA silencing

8

Chimeric gene construct

target virus

References

Tomato leaf curl virus (ToLCV) Praveen et al. (2006) and Cucumber mosaic virus (CMV) Tomato leaf curl Taiwan virus (ToLCTWV) and Tomato spotted wilt virus (TSWV)

Lin et al. (2011)

Tomato spotted wilt virus (TSWV), Groundnut ringspot virus (GRSV), Tomato chlorotic spot virus (TCSV) and Watermelon silver mottle virus (WSMoV),

Bucher et al. (2006)

Artificial zinc finger protein (AZP)

Tomato yellow leaf curl virus (TYLCV)

Takenaka et al. (2007)

Zinc finger nuclease (ZFN)

Tomato yellow leaf curl China Chen et al. (2014) virus (TYLCCNV),

Transcription activator like effector nucleases (TALE)

Tomato yellow leaf curl China Cheng et al. (2015) virus (TYLCCNV)

Genome editing Sequence specific platforms nucleases are used to alter or induce mutations in viral genomic region

Clustered, regularly Tomato yellow leaf curl virus interspaced (TYLCV) short palindromic repeats-CRISPR-associated proteins (CRISPR/Cas-9)

has been ascribed to the inhibition of uncoating of viral particles inside the cell in the initial stages of infection (Nelson et al., 1988). In Indian context, CP mediated resistance was developed at National Botanical Research Institute (NBRI), Lucknow. Rep gene mediated resistance Replication being an indispensable process in the viral infection cycle, numerous attempts was aimed at crippling the virus replication process by targeting replicase or rep protein. Similar to CPMR, rep-mediated resistance (RMR) has also been demonstrated in Tobacco mosaic virus by Golemboski et al. (1990). Furthermore, virus resistance trait has been found to be expressed even by expression of full length, partial, mutated or wild type replication protein (Beachy, 1997; Palukaitis and Zaitlin, 1997). Transgenic expression of defective replicase gene in tomato conferred resistance to CMV by inhibiting its long

Ali et al. (2016)

distance movement (Gal-On et al., 1998). Resistance to tomato yellow leaf curl disease broke down when the plants were infected at early stage or when virus inoculum load was high (Lapidot and Friedmann, 2002). Hence, transgenic expression of replication initiator protein (rep) has been a utilized approach. Expression of oligomerization domain of rep protein alone was found to inhibit heterologous viral DNA accumulation as shown by Chatterji et al. in 2001. Truncated c1 gene of Tomato yellow leaf curl Sardinia virus (TYLCSV), when potentially co-expressed with the C4 protein, conferred resistance to the homologous virus (Brunetti et al., 2001). Rep-mediated resistance was incorporated in tomato using truncated TYLCSV rep protein (210 amino acids); however, the study also documented the breakdown of resistance owing to transgene silencing (Lucioli et al., 2003). Similar approaches of using truncated rep gene encoding 129 amino acids showed

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transgenic resistance to TYLCV (Antignus et al., 2004) and TYLCSV (Prins et al., 2008). The study by Lucioli et al. (2003) also proposed models for rep-mediated resistance. Two model for resistance were proposed: (a) when homologous virus is challenge inoculated, inhibition of viral gene (C1) transcription was proposed; (b) when challenge inoculation was carried out by non-homologous virus, protein–protein interaction was proposed as the basis for rep-mediated resistance. Antisense RNA Besides, protein-mediated resistance, tomato transgenic lines have been developed that functions on the principle of RNA mediated repression of viral genes. In this approach either antisense RNA of the viral gene was expressed or defective RNAs, i.e. alterations in the gene leading to non-translatable RNAs were also employed to achieve antisense RNA mediated resistance. Furthermore, defective interfering RNAs and satellite molecules associated with viruses were expressed to confer resistance in tomato (Saito et al., 1992; Mc Garvey et al., 1994). Transgenic tomato plants have been obtained by expressing satellite RNA (satRNA) associated with CMV and this system provided resistance to some strains of CMV (Saito et al., 1992; Mc Garvey et al., 1994). In an instance, antisense RNAs of Rep protein was expressed in Nicotiana benthamiana to interfere with the disease caused by Tomato yellow leaf curl virus (TYLCV) (Bendahmane and Gronenborn, 1997). Transgenic tomato resistant to TYLCV was developed using antisense rep gene construct of the virus (Yang et al., 2004). Successful use of antisense approach was demonstrated at IARI, New Delhi (Praveen et al., 2005b) (Fig. 11.2A–D). The investigators demonstrated variable degrees of resistance to the single-stranded DNA Tomato leaf curl virus (ToLCV) in transgenic tomato plants (Praveen et al. 2005a). In order to attain stable resistance to the leaf curl disease of tomato, RNA mediated repression of replication associated protein was attempted by expressing rep gene in antisense orientation (Praveen et al., 2005b). Tomato transgenics resistant to ToLCD using replicase (rep) gene sequences of the Tomato leaf curl virus in antisense orientation, via Agrobacterium-mediated transformation were developed at IARI, New Delhi. A binary vector carrying the antisense rep gene (untranslatable full length sequence 1086 bp) along with the npt II gene

was used for transformation. This is demonstration of RNA-mediated silencing, since plants carry the untranslatable antisense rep gene, and have no detectable protein expression. Progeny analysis of these plants showed a classical Mendelian pattern of inheritance. It is important to note that two of the transgenic lines with a single transgene insertion have shown more than 80% resistance compared to the non-transformed control plants. These were selfed to produce progeny for resistance evaluation at the T2 stage, which followed the same pattern of resistance as the T1 stage (Praveen et al., 2004; www.isb.vt.edu/news/2006/news06.jun. htm#jun0603). Further, transgenic introduction of antisense rep gene in ToLCV infected plants recovered the plants from viral infection in vitro and this kind of recovery requires a threshold level of dsRNA production culminating in the production of siRNAs to counter viral replicase gene (Praveen et al., 2005b). Small interfering RNAs (siRNAs) The molecular basis of antisense RNA mediated viral gene silencing started emerging with the advent of phenomenon of RNA interference (Waterhouse et al., 1998). Later, the effector molecules of RNA silencing strategy were found to be small non-coding RNAs such as small interfering RNAs (siRNAs), and microRNAs (miRNAs). Further, the finding that simultaneous expression of sense and antisense arm of RNA separated by an intron provides efficient silencing platform provided impetus to hairpin RNA (hpRNA) expression based antiviral strategies (Smith et al., 2000; Wesley et al., 2001). The trigger for the biochemical process of virus resistance in plants is initiated by the expression of inverted repeats transgenes that is capable of generating self-complementary hairpin RNA cognate to viral RNAs (Smith et al., 2000). Intron spliced hairpin RNA mediated silencing of Tomato yellow leaf curl virus (TYLCV), in tomato plants were achieved by transforming the tomato with a gene construct targeting TYLCV rep gene (C1). The extreme resistance to the virus by these transgenic plants was ascertained with dot blot hybridization and it was found that plant exhibited no TYLCV DNA presence even after 60 days post challenge inoculation with virus (Fuentes et al., 2006). Tomato leaf curl viruses derived replication initiator protein (rep) nucleotide sequences were

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Figure 11.2  Development of transgenic tomato resistant to Tomato leaf curl disease (ToLCD) using replication associated protein (rep) gene sequence in antisense orientation. (A) (i) DNA-A of ToLCV carrying AV1 ORF coding for replicase protein; (ii) important domains and motifs of replicase protein. (B) Binary construct of replicase gene (truncated; Trep) in antisense orientation. (C) (i) Tomato transformants under tissue culture (T0); (ii) tomato transformant showing flowering and fruiting; (iii) tomato transformants under Phytotron (T1). (C) Field testing of transgenic tomatoline PED-AR-26 (T2) under controlled condition; (i) field layout; (ii) non-transgenic tomato under field conditions; (iii) transgenic line PED-AR-26 under field condition. (D) Histological analysis of gut tissues: haematoxylin and eosin staining of gastric mucosa: (a) Phosphate Buffer saline (control); (b) wild type tomato (non-transgenic); (c) GE (transgenic line PED-AR-26) tomato fed mice: sections showing jejunum villi; (d) PBS; (e) wild type tomato; (f) GE tomato fed mice. Sections showing liver sections: (g) PBS; (h) wild type tomato; (i) GE tomato fed mice.

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analysed and a conserved core of 318 nucleotides at the 3′ end of the gene encoding for Motif III, playing a vital role in viral replication was identified. Then multiple siRNAs have been used to target the conserved rep gene, including a small overlapping AC4 gene essential for pathogenicity and RNAi suppression. These strategies imply that ToLCV rep-driven RNAi, targeting AC4 and conserved viral sequences, provides a promising approach to suppress a wide spectrum ToLCV infection in the tomato (Fig. 11.2) (Ramesh et al., 2007). In addition, the length of hpRNA arm was analysed for its efficiency in providing virus resistance. Four different gene construct strategies, with varying lengths of hpRNA arm, were developed and their silencing efficiency against ToLCNDV infection was ascertained in tomato transgenics. The study proved that greater hpRNA arm length is required for efficient viral gene silencing as number of in vivo diced siRNAs produced in such cases were higher (Praveen et al., 2010). Furthermore, constitutive expression of short hairpin RNA expression 21 nt siRNA as hpRNA caused phenotypic abnormalities in tomato (Praveen et al., 2010). Although these results suggest the applicability of the RNAi based viral gene silencing, the breakdown of silencing could not be ignored. In Tomato yellow leaf curl Sardinia virus RNA mediated silencing of the rep –mRNA of the TYLCSV, either by simultaneous expression of sense and antisense RNA or expression of multiple copies of sense RNAs, could not be sustained. With the high pressure of the viral inoculum on whitefly mediated transmission of the virus the resistance through RNA silencing could be overcome (Noris et al., 2004). Artificial miRNA (amiRNAs) The host endogenous gene regulatory mechanism involving microRNAs (miRNAs) has been manipulated to develop antiviral resistance in tomato. The advantages of employing artificial (amiRNA) triggered gene silencing strategy over siRNA mediated mechanism are a) high degree of target specificity is achieved as amiRNAs are of 21 nt sequences in length b) amiRNA mediated gene silencing mechanism does not involve genetic recombination and the chances of generating off-targets are very less hence suitable in terms of biosafety aspects too and c) miRNAs are not immediate targets of viral suppressors of RNA silencing. All these features

and the possibility of incorporating multiple amiRNAs each targeting specific virus makes it a better alternative for siRNA based antiviral resistance. A comparative analysis of siRNA and amiRNA mediated gene silencing strategies was performed by developing tomato transgenics resistant to ToLCNDV (Praveen et al., 2007). Based on these studies miRNA vector has been designed to effect gene silencing in plants (Koundal and Praveen, 2010). The study revealed that amiRNA mediated strategy yielded superior gene silencing efficiency thereby virus resistance (Praveen et al., 2007). Tomato transgenic developed using a two amiRNAs targeting the shared sequences of two coding regions (2a and 2b) and conserved 3′UTR regions of Cucumber mosaic virus showed resistance to CMV and nontarget viruses such as Tobacco mosaic virus and Tomato yellow leaf curl virus (Zhang et al., 2011). Similarly, two amiRNA constructs have been designed to target coat protein (AV1) and pre-coat protein (AV2) genes of Tomato leaf curl virus and transgenic tomato lines were generated. Analysis of putative transgenic tomato lines indicate that the level of resistance varied from extreme to moderate depending upon the target viral gene. However, the study demonstrated that development of amiRNA is a viable strategy to control ToLCV (VanTu et al., 2013). Artificial trans-acting siRNAs (ata-siRNAs) In the plant genomes, endogenous trans-acting siRNA producing loci (TAS loci) have been identified which are characterized with the production of novel class of non-coding RNAs called transacting (ta-siRNAs). ta-siRNAs characteristically cleave non-identical transcripts and effect gene silencing. Production of ta-siRNAs and its effect on RNA silencing pathway also involves concerted action of miRNAs (Yoshikawa et al., 2005). Non-coding gene could be modified to produce artificial tasiRNAs by altering the miRNA binding site. Vectors were developed to generate artificial tasiRNAs directed against AC2 and AC4 genes of ToLCNDV. Successful alleviation of leaf curl disease symptoms and reduction of virus quantity were demonstrated in tomato following ToLCNDV agro-inoculation. This study thus demonstrated the utility of tasiRNAs in conferring resistance to ToLCNDV and tomato leaf curl Gujarat virus (ToLCGV)

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in transgenic tobacco plants (Singh et al., 2015). However, whether the transgenic expression of atasiRNAs in tomato confers resistance to multiple viruses is yet to be investigated. Dual/multiple virus resistance Multiple viral infections are not uncommon in the field conditions hence, genetic engineering approaches are required to develop plant genotype that can withstand more than one viral infections at a given point of time. Tomato is infected by Tomato leaf curl virus (ToLCV) and Cucumber mosaic virus (CMV) hence it was decided to incorporate dual resistance in tomato. Later the stacking of two viral sequences for developing dual resistance in tomato was demonstrated by our group at IARI (Praveen et al. 2006). A chimeric gene construct was developed comprising the conserved core region of rep gene encoding replicase protein (AC1) of ToLCV along with, 830 bp ORF of NP gene of GBNV. Around 65–85% of transgenic tomato plants developed using this chimeric construct showed complete resistance to both the viruses (Praveen et al., 2006). Also, dual resistance has been incorporated in tomato against the infection DNA [(Tomato leaf curl Taiwan virus (ToLCTWV)] and RNA [Tomato spotted wilt virus (TSWV)] containing viruses using a single chimeric transgene construct suggesting the versatility of RNA based viral gene silencing (Lin et al., 2011). In order to engineer multiple tospovirus-resistant tomato line, genetic fragments of N gene sequences of the four major tomato-infecting viruses, Tomato spotted wilt virus (TSWV), Groundnut ringspot virus (GRSV), Tomato chlorotic spot virus (TCSV) and Watermelon silver mottle virus (WSMoV), were incorporated in a single chimaeric hairpin (hp) RNA construct. Furthermore, more than 80% of the transformed plants showed multiple virus resistance in this approach (Bucher et al., 2006). Thus, RNA silencing based virus resistance strategy has been effectively employed to confer dual and multiple virus resistance in tomato. Interestingly, marker assisted breeding strategy has been successfully used to develop tomato lines (UMH1200 and UMH 1203) that were resistant to multiple viruses such as Tomato mosaic virus (ToMV), Tomato spotted wilt virus (TSWV) and Tomato yellow leaf curl virus (TYLCV) (García-Martínez et al., 2011; García-Martínez et al., 2012).

Safety assessment of virusresistant transgenic tomato The safety assessment of virus-resistant transgenic tomato have been carried out based on the principle of substantial equivalence enunciated by organizations such as OECD (1993) and FAO/WHO (2001). The toxicity, allergenicity and nutritional imbalance tests of tomato transgenics resistant to ToLCV (Praveen et al., 2005a) was assessed in Balb/c mice following standard procedures (Fig. 11.2E). Prior to analysing the effect of toxicity and allergenicity in mice, it was proven that the viral derived sequences used in the transgenic development do not have any proclivity to silence mouse or human gene (Praveen et al., 2007). Ovalbumin was used to develop the allergic mice model and hypersensitive patients’ sera were used to analyse tomato extract for IgE binding. Even though antisense RNA based transgenics do not produce functional proteins, the transgene viral sequences were analysed in silico for its potential allergen inducing property and it was found that the translated protein did not show any match with allergens listed in the SDAP or Farrp databases. The study unequivocally concluded that genetically engineered tomato with ToLCV derived rep gene was found to be safe with respect to toxicity and allergenicity (Singh et al., 2009). Way forward The rapid progress made in the field of virusresistant transgenics has helped in development of tomato genotypes resistant to virus infections. However, in order to obtain stable, broad spectrum resistance, identification of conserved viral genomic sequences employing next generation sequencing (NGS) technologies could help design better transgene for countering emergence and reemergence of resistance breaking virus strains in field conditions. In general, the development of transgenic resistance to tomato infecting viruses involved use of viral genome derived sequences and hence pathogen derived resistance has been a predominant approach. However, transgenic virus resistance approaches, especially nucleotide sequence specific RNA based silencing, have some potential pitfalls. Mixed viral infections are common in the field conditions hence, potential resistance breakdown is

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envisaged because the RNA based resistance mechanism is negated in the presence of viral suppressor proteins derived from even the unrelated viruses. In this regard wild relatives of tomato are reservoir for R-genes that have potential use in developing virus-resistant genotypes. Genetic investigations have identified many loci and mapped various TYLCV tolerance or resistance conferring genes such as Ty-1, Ty-3 and Ty-4 from S. chilense (Zamir et al., 1994; Ji et al., 2007, 2009). Similarly, Ty-2 and Ty-5 were identified in S. habrochaites and S. peruvianum respectively (Hanson et al., 2006; Hutton et al., 2012). However, Verlaan et al. (2013) have fine mapped Ty-1 and Ty-3 genes and found them to be allelic and the genes encode for RNA dependent RNA polymerase (RDR gamma). The tolerance to TYLCV conferred by these genes was explained based on the role of RDRs in RNA silencing pathway. Further, integration of transcriptomic and metabolomic approaches have unearthed the biochemical and molecular response of susceptible and tomato cultivars towards TYLCV infection. Further the study revealed that the resistant genotype is intermediate between susceptible cultivar and the wild relative S. habrochaites (Sade et al., 2015). The next generation gene alteration tools such as genome editing has been considered to play significant role in developing virus-resistant genotypes. Various genome targeting platforms have been utilized to show the effectiveness of this technology in providing antiviral resistance. The four major genome editing platforms based on sequence specific nucleases are (i) zinc finger nucleases (ZFNs), (ii) transcription activator like effector nucleases (TALENs) (iii) clustered, regularly interspaced short palindromic repeatsCRISPR-associated proteins (CRISPR/Cas-9) (iv) artificial zinc finger protein (AZP). Interestingly, all these four platforms have been successfully used for resistance to tomato infecting viruses (Takenaka et al., 2007; Chen et al., 2014; Cheng et al., 2015; Ali et al., 2016). However, considering the multiplicity of viruses infecting tomato in the field conditions, it is to be seen if the genome altering technologies could provide broad spectrum resistance to phytopathogenic viruses of tomato. Besides the regulatory system required for monitoring the crop genotypes generated through genome targeting/ editing technologies, potential drawbacks including

the development and introduction of even more challenging virus isolates in the environment could not be ruled out. Hence, NGS techniques could complement genome editing efforts to identify any potential off-target effects in the genomes of virus and host plants and gain molecular insights. References Abhary, M., Patil, B.L., and Fauquet, C.M. (2007). Molecular biodiversity, taxonomy, and nomenclature of Tomato yellow leaf curl-like viruses.In Tomato Yellow Leaf Curl Disease. Management, Molecular Biology, Breeding for Resistance, H. Czosnek, ed., (Springer, Dordrecht), pp.85–118. Akram, M., and Naimuddin. (2010). First report of Groundnut bud necrosis virus infecting pea (Pisum sativum) in India. New Dis. Rep. 21, 10. https://doi.org /10.5197/j.2044-0588.2010.021.010 Ali, Z., Abulfaraj, A., Idris, A., Ali, S., Tashkandi, M., and Mahfouz, M.M. (2015). CRISPR/Cas9-mediated viral interference in plants. Genome Biol. 16, 238. https:// doi.org/10.1186/s13059-015-0799-6 Antignus, Y., Vunsh, R., Lachman, O., Pearlsman, M., Maslenin, L., Hananya, U., and Rosner, A. (2004). Truncated Rep gene originated from Tomato yellow leaf curl virus-Israel [Mild] confers strain-specific resistance in transgenic tomato. Ann. Appl. Biol. 144, 39–44. https://doi.org/10.1111/j.1744-7348.2004.tb00314.x Beachy, R.N. (1997). Mechanisms and applications of pathogen-derived resistance in transgenic plants. Curr. Opin. Biotechnol. 8, 215–220. Bendahmane, M., and Gronenborn, B. (1997). Engineering resistance to Tomato yellow leaf curl virus (TYLCV) using antisense RNA. Plant Mol. Biol. 33, 351–357. Boulton, M.I. (2003). Geminiviruses: major threats to world agriculture. Ann. Appl. Biol. 142, 143–143. https://doi. org/10.1111/j.1744-7348.2003.tb00239.x Brunetti, A., Tavazza, R., Noris, E., Lucioli, A., Accotto, G.P., and Tavazza, M. (2001). Transgenically expressed T-Rep of Tomato yellow leaf curl Sardinia virus acts as a transdominant-negative mutant, inhibiting viral transcription and replication. J. Virol. 75,10573–10581. https://doi. org/10.1128/JVI.75.22.10573-10581.2001 Brunt, A., Crabtree, K., Dallwitz, M., Gibbs, A., and Watson, L. (1996). Viruses of Plants. Description and Lists from the VIDE Database (CAB International, Wallingford, UK). Bucher, E., Lohuis, D., van Poppel, P.M., GeertsDimitriadou, C., Goldbach, R., and Prins, M. (2006). Multiple virus resistance at a high frequency using a single transgene construct. J. Gen. Virol. 87, 3697-3701. https://doi.org/10.1099/vir.0.82276-0 Chatterji, A., Beachy, R.N., and Fauquet, C.M. (2001). Expression of the oligomerisation domain of the replication-associated protein (Rep) of Tomato leaf curl New Delhi virus interferes with DNA accumulation of heterologous geminiviruses. J. Biol. Chem. 276, 2563125638. https://doi.org/10.1074/jbc.M100030200 Chen, W., Qian, Y., Wu, X., Sun, Y., Wu, X., and Cheng, X. (2014). Inhibiting replication of begomoviruses using

232  | Ramesh and Praveen

artificial zinc finger nucleases that target viral-conserved nucleotide motif. Virus Genes 48, 494–501. https://doi. org/10.1007/s11262-014-1041-4 Cheng, X., Li, F., Cai, J., Chen, W., Zhao, N., Sun, Y., Guo, Y., Yang, X., and Wu, X. (2015). Artificial TALE as a convenient protein platform for engineering broadspectrum resistance to Begomoviruses. Viruses 7, 4772–4782. https://doi.org/10.3390/v7082843 Doolittle, S.P. (1916). A new infectious mosaic disease of cucumber. Phytopathology 6, 145–147. FAO/WHO (2001). Evaluation of allergenicity of genetically modified foods. Report of a joint FAO/ WHO expert consultation on allergenicity of foods derived from biotechnology, 22–25 January 2001: Food and Agriculture Organization of the United Nations, Rome. Available from ftp://ftp.fao.org/es/esn/food/ gmreport.pdf Fazeli, R., Heydarnejad, J., Massumi, H., Shaabanian, M., and Varsani, A. (2009). Genetic diversity and distribution of tomato-infecting begomoviruses in Iran. Virus Genes 38,311–319. https://doi.org/10.1007/ s11262-008-0310-5 Fortes, I.M., Sánchez-Campos, S., Fiallo-Olivé, E., DíazPendón, J.A., Navas-Castillo, J., and Moriones, E. (2016). A novel strain of Tomato leaf curl New Delhi virus has spread to the Mediterranean basin. Viruses 8, E307. Fuentes, A., Ramos, P.L., Fiallo, E., Callard, D., Sánchez, Y., Peral, R., Rodríguez, R., and Pujol, M. (2006). Intronhairpin RNA derived from replication associated protein C1 gene confers immunity to Tomato yellow leaf curl virus infection in transgenic tomato plants. Transgenic Res. 15, 291–304. https://doi.org/10.1007/s11248-0055238-0 Gal-On, A., Wolf, D., Wang, Y., Faure, J.E., Pilowsky, M., and Zelcer, A. (1998). Transgenic resistance to Cucumber mosaic virus in tomato: blocking of long-distance movement of the virus in lines harboring a defective viral replicase gene. Phytopathology 88, 1101–1107. https:// doi.org/10.1094/PHYTO.1998.88.10.1101 García-Martínez, S., Grau, A., Alonso, A., Rubio, F., Valero, M., and Ruiz, J.J. (2012). UMH 1203, a multiple virus-resistant fresh-market tomato breeding line for open-field conditions. HortScience 47, 124-125. García-Martínez, S., Grau, A., Alonso, A., Rubio, F., Valero, M., and Ruiz, J.J. (2011). UMH 1200, a breeding line within the Muchamiel tomato type resistant to three viruses. HortScience 46,1054–1055. Gielen, J., Ultzen, T., Bontems, S., Loots, W., van Schepen, A.,Westerbroek, A., de Haan, P., and van Grinsven, M. (1996). Coat protein mediated protection to Cucumber mosaic virus infections in cultivated tomato. Euphytica 88, 139–149. https://doi.org/10.1007/BF00032445 Gielen, J.J.L., de Haan, P., Kool, A.J., Peters, D., van Grinsven, M.Q.J.M., and Goldbach, R.W. (1991). Engineered resistance to Tomato spotted wilt virus, a negative-strand RNA virus. Nature Biotechnol 9,1363–1367. https:// doi.org/10.1038/nbt1291-1363 Goldbach, R., and Kuo, G. (1996). Introduction. Proceedings International symposium on tospoviruses and thrips of floral and vegetable crops. Taiwan Agricultural Research Institute, Taichung, Taiwan, November 7–10, 1995. Acta Hortic. 431, 21–26.

Golemboski, D.B., Lomonossoff, G.P., and Zaitlin, M. (1990). Plants transformed with a Tobacco mosaic virus nonstructural gene sequence are resistant to the virus. Proc. Natl. Acad. Sci. U.S.A. 87, 6311–6315. Gonsalves, D. (1996). Cornell Community Conference on Biological Control. Hanson, P., Green, S., and Kuo, G. (2006). Ty-2, a gene on chromosome 11 conditioning geminivirus resistance in tomato. Tomato Genet Coop Rep56, 17–18 Hanssen, I.M., Lapidot, M., and Thomma, B.P. (2010). Emerging viral diseases of tomato crops. Mol. Plant Microbe Interact. 23, 539–548. https://doi. org/10.1094/MPMI-23-5-0539 Heinze, C., Maiss, E., Adam, G., and Casper, R. (1995). The complete nucleotide sequence of the S RNA of a new Tospovirus species, representing serogroup IV. Phytopathology 85, 683–690. Hutton, S.F., Scott, J.W., and Schuster, D.J. (2012). Recessive resistance to Tomato yellow leaf curl virus from the tomato cultivar Tyking is located in the same region as Ty-5 on chromosome 4. HortScience 47, 324–327. Ji, Y., Schuster, D.J., and Scott, J.W. (2007). Ty-3, a begomovirus resistance locus near the Tomato yellow leaf curl virus resistance locus Ty-1 on chromosome 6 of tomato. Mol. Breeding 20, 271–284. Ji, Y., Scott, J.W., Schuster, D.J., and Maxwell, D.P. (2009). Molecular mapping of Ty-4, a new Tomato yellow leaf curl virus resistance locus on chromosome 3 of tomato. J. Am. Soc. Hortic. Sci. 134, 281–288. Juárez, M., Tovar, R., Fiallo-Olivé, E., Aranda, M.A., Gosálvez, B., Castillo, P., Moriones, E., and NavasCastillo, J.,(2014). First detection of Tomato leaf curl New Delhi virus infecting zucchini in Spain. Plant Dis. 98, 857. Koundal, V., and Praveen, S. (2010). MicroRNA-based RNA interference vector for gene silencing in plants. J. Plant Biochem. Biotechnol. 19, 79–82. Kunik, T., Salomon, R., Zamir, D., Navot, N., Zeidan, M., Michelson, I., Gafni, Y., and Czosnek, H. (1994). Transgenic tomato plants expressing the Tomato yellow leaf curl virus capsid protein are resistant to the virus. Biotechnology 12, 500–504. Lapidot, M., and Friedmann, M. (2002). Breeding for resistance to whitefly transmitted geminiviruses. Ann. Appl. Biol. 140, 109–127. doi: 10.1111/j.17447348.2002.tb00163.x Lin, C.Y., Ku, H.M., Tsai, W.S., Green, S.K., and Jan, F.J. (2011). Resistance to a DNA and a RNA virus in transgenic plants by using a single chimeric transgene construct. Transgenic Res. 20, 261–270. https://doi. org/10.1007/s11248-010-9412-7 Lucioli, A., Noris, E., Brunetti, A., Tavazza, R., Ruzza, V., Castillo, A.G., Bejarano, E.R., Accotto, G.P., and Tavazza, M. (2003). Tomato yellow leaf curl Sardinia virus rep-derived resistance to homologous and heterologous geminiviruses occurs by different mechanisms and is overcome if virus-mediated transgene silencing is activated. J. Virol. 77, 6785–6798. Mayo, M.A. (1992). Organization of Viral Genomes: the potential of virus -genes in the production of transgenic virus-resistant plants, In Biotechnology for Crop Improvement in Asia, ed., Moss, J.P. (ICRISAT, Hyderabad), pp. 251-264.

Virus-resistant Transgenic Tomato |  233

McGarvey, P.B., Montasser, M.S., and Kaper, J.M. (1994). Transgenic tomato plants expressing satRNA are tolerant to some strains of Cucumber mosaic virus. J. Am. Soc. Hortic. Sci. 119, 642–647. Melgarejo, T.A., Kon, T., Rojas, M.R., Paz-Carrasco, L., Zerbini, F.M., and Gilbertson, R.L. (2013). Characterization of a new world monopartite begomovirus causing leaf curl disease of tomato in Ecuador and Peru reveals a new direction in geminivirus evolution. J. Virol. 87, 5397–5413. https://doi. org/10.1128/JVI.00234-13 Nelson, R.S., McCormick, S.M., Delannay, X., Dubé, P., Layton, J., Anderson, E.J., Kaniewska, M., Proksch, R.K., Horsch, R.B., Rogers, S.G., Fraley, R.T., and Beachy, R.N. (1988). Virus tolerance, plant growth, and field performance of transgenic tomato plants expressing coat protein from Tobacco mosaic virus. Nature Biotechnol. 6, 403–409. doi:10.1038/nbt0488-403 Noris, E., Lucioli, A., Tavazza, R., Caciagli, P., Accotto, G.P., and Tavazza, M. (2004). Tomato yellow leaf curl Sardinia virus can overcome transgene mediated RNA silencing of two essential viral genes. J. Gen. Virol. 85, 1745–1749. OECD (1993a) Safety evaluation of foods derived by modern biotechnology, concepts and principles. Organization for Economic Cooperation and Development, Paris. www.oecd.org/pdf/Mooo33000/ M00033002.pdf33. Palukaitis, P., Roossinck, M.J., Dietzgen, R.G., and Francki, R.I. (1992). Cucumber mosaic virus. Adv. Virus Res. 41, 281–348. Palukaitis, P., and Zaitlin, M. (1997). Replicase-mediated resistance to plant virus disease. Adv. Virus Res. 48, 349–377. Abel, P.P., Nelson, R.S., De, B., Hoffmann, N., Rogers, S.G., Fraley, R.T., and Beachy, R.N. (1986). Delay of disease development in transgenic plants that express the Tobacco mosaic virus coat protein gene. Science 232, 738–743. Praveen, S., Kushwaha, C.M., Mishra, A.K., Singh, V., Jain, R.K., and Varma, A. (2005a) Engineering tomato for resistance to Tomato leaf curl disease using viral rep gene sequences. Plant Cell Tiss. Organ Cult. 83, 311-318. https://doi.org/10.1007/s11240-005-7858-8 Praveen, S., Mishra, A.K., and Antony, G. (2006). Viral gene suppression in transgenic plants expressing chimeric transgene from Tomato leaf curl virus and Cucumber mosaic virus. Plant Cell Tiss. Organ Cult. 84, 49-55. Praveen, S., Mishra, A.K., and Dasgupta, A. (2005b) Antisense suppression of replicase gene expression recovers tomato plants from leaf curl infection. Plant Science 168, 1011-1014. https://doi.org/10.1016/j. plantsci.2004.12.008 Praveen, S., Dasgupta, A., and Varma, A. (2004). Phylogenetic analysis and homologies of the replicase of tomato leaf curl geminiviruses: implications for obtaining pathogen derived resistance. Virus Genes 28, 195–199. https://doi.org/10.1023/B:VIRU.0000016858.66565. f8 Praveen, S., Ramesh, S.V., Koundal, V., Mishra, A.K., and Jain, R.K. (2007). Small RNA mediated silencing to target Tomato leaf curl virus. J Plant Interact. 2, 213-218. https://doi.org/10.1080/17429140701684731

Praveen, S., Ramesh, S.V., Mishra, A.K., Koundal, V., and Palukaitis, P. (2010). Silencing potential of viral derived RNAi constructs in Tomato leaf curl virus-AC4 gene suppression in tomato. Transgenic Res. 19, 45–55. https://doi.org/10.1007/s11248-009-9291-y Prins, M., Laimer, M., Noris, E., Schubert, J., Wassenegger, M., and Tepfer, M. (2008). Strategies for antiviral resistance in transgenic plants. Mol. Plant Pathol. 9, 73–83. https://doi.org/10.1111/j.13643703.2007.00447.x Provvidenti, R., and Gonsalves, D. (1995). Inheritance of resistance to Cucumber mosaic virus in transgenic tomato line expressing the coat protein gene of the white leaf strain. J. Hered. 86, 85–88. Raj, S.K., Singh, R., Pandey, S.K., and Singh, B.P. (2005). Agrobacterium-mediated tomato transformation and regeneration of transgenic lines expressing Tomato leaf curl virus coat protein gene for resistance to TLCV infection. Curr. Sci. 88, 1674-1679. Ramesh, S.V., Mishra, A.K., and Praveen, S. (2007). Hairpin RNA-mediated strategies for silencing of Tomato leaf curl virus AC1 and AC4 genes for effective resistance in plants. Oligonucleotides 17, 251–257. https://doi. org/10.1089/oli.2006.0063 Sade, D., Shriki, O., Cuadros-Inostroza, A., Tohge, T., Semel, Y., Haviv, Y., Willmitzer, L., Fernie, A.R., Czosnek, H., and Brotman, Y. (2015). Comparative metabolomics and transcriptomics of plant response to Tomato yellow leaf curl virus infection in resistant and susceptible tomato cultivars. Metabolomics 11, 81–97. https://doi. org/10.1007/s11306-014-0670-x Saito, Y., Komari, T., Masuta1, C., Hayashi, Y., Kumashiro, T., and Takanami, Y. (1992). Cucumber mosaic virustolerant transgenic tomato plants expressing a satellite RNA. Theor. Appl. Genet. 83, 679–683. https://doi. org/10.1007/BF00226684 Sanders, P.R., Sammons, B., Kaniewski, W., Haley, L., Layton, J., LaVallee, B.J., Delannay, X., and Tumer, N.E. (1992). Field resistance of transgenic tomatoes expressing the Tobacco mosaic virus or Tomato mosaic virus coat protein genes. Phytopathology 82, 683–690. Sanford, J.C., and Johnston, S.A. (1985). The concept of parasite-derived resistance-deriving resistance genes from the parasite’s own genome. J. Theor. Biol. 113(2), 395–405. Sangaré, A., Deng, D., Fauquet, C.M., and Beachy, R.N. (1999). Resistance to African Cassava mosaic virus conferred by a mutant of the putative NTP-binding domain of the Rep gene (AC1) in Nicotiana benthamiana. Mol. Breeding 5, 95-102. Singh, A., Taneja, J., Dasgupta, I., and Mukherjee, S.K. (2015). Development of plants resistant to tomato geminiviruses using artificial trans-acting small interfering RNA. Mol. Plant Pathol. 16, 724–734. https://doi.org/10.1111/mpp.12229 Singh, A.K., Praveen, S., Singh, B.P., Varma, A., and Arora, N. (2009). Safety assessment of leaf curl virus resistant tomato developed using viral derived sequences. Transgenic Res. 18, 877–887. https://doi.org/10.1007/ s11248-009-9274-z Smith, N.A., Singh, S.P., Wang, M.B., Stoutjesdijk, P.A., Green, A.G., and Waterhouse, P.M. (2000). Total

234  | Ramesh and Praveen

silencing by intron-spliced hairpin RNAs. Nature 407, 319–320. https://doi.org/10.1038/35030305 Takenaka, K., Koshino-Kimura, Y., Aoyama, Y., and Sera, T. (2007). Inhibition of Tomato yellow leaf curl virus replication by artificial zinc-finger proteins. Nucleic Acids Symp Ser51, 429–430. Tumer, N.E., O’connell, K.M., Nelson, R.S., Sanders, P.R., Beachy, R.N., Fraley, R.T., and Shah, D.M. (1987). Expression of Alfalfa mosaic virus coat protein gene confers cross-protection in transgenic tobacco and tomato plants. EMBO J. 6, 1181–1188. Umamaheswaran, K., Jain, R.K., Bhat, A.I., and Ahlawat, Y.,S. (2003). Biological and molecular characterization of tospovirus isolate from tomato and its relationship with other tospoviruses. Indian Phytopathol. 56, 163–173. Vu, T.V., Choudhury, N.R., and Mukherjee, S.K. (2013). Transgenic tomato plants expressing artificial microRNAs for silencing the pre-coat and coat proteins of a begomovirus, Tomato leaf curl New Delhi virus, show tolerance to virus infection. Virus Res. 172, 35–45. https://doi.org/10.1016/j.virusres.2012.12.008 Varma, A., and Malathi, V.G. (2003). Emerging geminivirus problems: a serious threat to crop production. Ann. App. Biol. 142, 145–164. doi 10.1111/j.1744-7348.2003. tb00240.x Verlaan, M.G., Hutton, S.F., Ibrahem, R.M., Kormelink, R., Visser, R.G., Scott, J.W., Edwards, J.D., and Bai, Y. (2013). The Tomato yellow leaf curl virus resistance genes Ty-1 and Ty-3 are allelic and code for DFDGDclass RNA-dependent RNA polymerases. PLOS Genet. 9, e1003399. https://doi.org/10.1371/journal. pgen.1003399

Waterhouse, P.M., Graham, M.W., and Wang, M.B. (1998). Virus resistance and gene silencing in plants can be induced by simultaneous expression of sense and antisense RNA. Proc. Natl. Acad. Sci. U.S.A. 95, 13959–13964. Wesley, S.V., Helliwell, C.A., Smith, N.A., Wang, M.B., Rouse, D.T., Liu,Q., Gooding, P.S., Singh, S.P., Abbott, D., Stoutjesdijk, P.A., et al. (2001). Construct design for efficient, effective and high-throughput gene silencing in plants. Plant J. 27, 581–590. Xue, B., Gonsalves, C., Provvidenti, R., Slightom, J.L., Fuchs, M., and Gonsalves, D. (1994). Development of transgenic tomato expressing a high level of resistance to Cucumber mosaic virus strains of subgroups I and II. Plant Dis. 78, 1038-1041. Yang, Y., Sherwood, T.A., Patte, C.P., Hiebert, E., and Polston, J.E. (2004). Use of Tomato yellow leaf curl virus (TYLCV) Rep gene sequences to engineer TYLCV resistance in tomato. Phytopathology 94, 490–496. https://doi.org/10.1094/PHYTO.2004.94.5.490 Yoshikawa, M., Peragine, A., Park, M.Y., and Poethig, R.S. (2005). A pathway for the biogenesis of trans-acting siRNAs in Arabidopsis. Genes Dev. 19, 2164–2175. Zamir, D., Ekstein-Michelson, I., Zakay, Y., Navot, N., Zeidan, M., Sarfatti, M., Eshed, Y., Harel, E., Pleban, T., van-Oss, H., et al. (1994). Mapping and introgression of a Tomato yellow leaf curl virus tolerance gene, TY-1. Theor. Appl. Genet. 88, 141–146. https://doi.org/10.1007/ BF00225889 Zhang, X., Li, H., Zhang, J., Zhang, C., Gong, P., Ziaf, K., Xiao, F., and Ye, Z. (2011). Expression of artificial microRNAs in tomato confers efficient and stable virus resistance in a cell-autonomous manner. Transgenic Res. 20, 569–581. https://doi.org/10.1007/s11248-0109440-3

12

Management of Geminiviruses Focusing on Small RNAs in Tomato Archana Singh1 and Sunil Kumar Mukherjee2*

1School of Biological Sciences, University of East Anglia, Norwich, UK. 2Division of Plant Pathology, IARI, New Delhi, India.

*Correspondence: [email protected] https://doi.org/10.21775/9781910190814.12

Abstract Geminiviruses are plant-infecting, ssDNAcontaining viruses that have twinned geminate particles. The majority of geminiviruses belong to the genus Begomovirus, having eithera monopartite or bipartite genome (DNA-A and DNA-B). Tomato leaf curl New Delhi virus (ToLCNDV), a pathogenic member of genus Begomovirus, causes Tomato leaf curl disease (ToLCD). ToLCD is responsible for economic losses of up to 100% in many regions of world. Control of ToLCV is very difficult as the classical vector control method turns out to be an unworthy option and Ty-based breeding yields partial success. Our laboratory has employed RNAi-based pathogen derived resistance (PDR) approach to successfully tackle ToLCD. We have overproduced artificial microRNA (a-miR) and artificial tasiRNA (a-tasiRNA) in tomato to silence the RNAi-suppressors encoded by ToLCVs. A few of the tomato transgenics show high level of virus resistance when challenged with broad range of ToLCVs. Introduction Tomato is an herbaceous plant belonging to the family Solanaceae. After potatoes, tomato is one of the most consumed vegetables in the world (FAOSTAT, http://faostat3.fao.org/home/index. html), and probably the most preferred garden crop. Tomato contains lycopene pigment, which is a carotenoid, one of the most potent antioxidants

among dietary carotenoids. Dietary intake of tomatoes has been shown to decrease risk of chronic diseases, such as cardiovascular disease and cancer (Friedman et al., 2013). Because of its prolific growth, relative cheap availability and other agronomic characters, tomato is also called poor person’s apple. However, such a nice crop is also very sensitive to a number of phytopathogens, begomoviruses being the most potent amongst them. Begomoviruses belong to the largest genus (Begomovirus) of the virus family Geminiviridae. The geminiviruses (family: Geminiviridae) are plant-infecting ssDNA-containing viruses that have twinned geminate particles. Depending on genome organization, host range, insect vector, and phylogenetic relationships the family is divided into seven genera, four of which are named as Curtovirus, Topocuvirus, Mastrevirus and Begomovirus, etc. (Brown et al., 2012). Geminiviruses together with potyviruses (family Potyviridae), constitute the two largest and most important plant virus families (Gibbs and Ohshima, 2010; Scholthof et al., 2011). However, here our major attention will be only on begomoviruses. The circular, singlestranded genome of begomoviruses can be either monopartite or bipartite (components known as DNA-A and DNA-B) with a twinned icosahedral particle morphology for each of the DNA components. Begomoviral diseases cause major crop loss in tropical and subtropical regions (Van et al., 2013). Tomatoes are particularly seriously affected

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by begomoviruses worldwide (Abhary et al., 2007; Hanssen et al., 2010; Lefeuvre et al., 2010). Disease caused by begomoviruses have significantly impacted tomato production in America since 1980s (Polston and Anderson., 1997; Ribeiro et al., 2003). Tomato leaf curl virus (ToLCV) is a potent phyto-pathogen causing Tomato leaf curl disease (ToLCD). This is the most important viral disease of tomato in India and other Asian countries (Van et al., 2013; Nakhla and Maxwell, 1998; Czosnek et al., 1990; Polston et al., 1999). Traditional approaches to control geminivirus infection rely on vector control, mainly by pesticides or physical barriers (Palumbo et al., 2001; Hilje et al., 2001). But most of the insecticides are carcinogens, environmental pollutants and help evolve vectors for insecticide tolerance. Thus these methods cause more harm than good to the agro-ecology and are not very effective. Physical barriers such as screens and UVabsorbing plastic sheets has been found effective in reducing virus spread by inhibiting penetration of whiteflies (Antignus et al., 2001). However, it adds to overheating, poor ventilation and cost for production. Therefore, breeding technique was adopted and crop tolerant/resistant to geminiviruses are used to introduce trait/genes from one variety, or line into new genetic background through sexual crosses (Pico et al., 1996). However, breeding for durable resistance to begomoviruses has not been successful so far. In Latin America around 20,000 different accessions of common bean were screened for immunity against BGMV but none of them were found to be immune (Lucioli et al., 2008). Similar studies were done in Mungbean for Mungbean yellow mosaic virus (MYMV), but resistant accessions could not be found. However, a few resistance genes have been characterized against tomato infecting begomoviruses; breeding tomato for virus resistance is somewhat successful. Genetic engineering could be another option where genes can be transferred to a plant by means other than the usual sexual crosses, including gene transfer by Agobacterium tumefaciens, a plant pathogenic bacterium that is competent to naturally transfer DNA to plants (Bundock et al., 2002). Moreover, approaches based on genes derived from viral pathogens that are known as pathogen derived resistance (PDR) have also been reported abundantly in literature. Using the latter gene transfer methods, Thomson, 2014 reported the production of maize plants

resistant to Maize streak virus, which harboured a viral gene transferred to it. Overall increase in 20% of bean production was achieved in Brazil with the commercial release of genetically engineered bean against BGMV (Tollefson et al., 2011). These days many modern versions of pdr involving principles of RNAi have been reported and these offer a new paradigm of resistance to begomoviruses. These approaches rely on generating host plants that overproduce small RNAs capable of silencing the important transcripts of the infecting viruses. Plants defend themselves against viruses using the host-coded RNAi factors which help generate siRNAs from all over the viral genomes. These siRNAs in turn down regulate the viral transcripts and proteins. The down-regulation can occur both at post-transcriptional and/or transcriptional stages. On the other hand, the viruses have also evolved to battle host RNAi activities and encode RNAi-suppressors proteins as counter defence principles. These suppressors are important for virus replication and transmission and are also crucial pathogenic factors. So the hosts, harbouring the small RNAs to silence the viral RNAi-suppressors, are presumed to be virus resistant. We have engineered tomato that can over express various forms of small RNAs to silence the transcripts of RNAisuppressors of Tomato leaf curl virus (ToLCV) and a few of the transgenic show virus resistance to the immunity level (Tien et al., 2013; Singh et al., 2015). Geminiviruses: classification and metabolism Genome organization Geminiviruses are circular single-stranded DNA viruses which infect dicot as well as monocot plants including economically important crops such as bean, beet, cotton, cassava, pepper, maize and tomato (Hanley-Bowdoin et al., 2013). Genome contains one or two 2.5–3.0 kb circular DNA molecules, each of which is encapsidated in a twinned (quasi)-icosahedral virion (so-called Gemini). The genome of geminiviruses encodes for a few proteins and thus these viruses are dependent mostly on host factors for DNA replication, transcription and movement across various host cells. Fig. 12.1 shows the gene organization of mono- and bipartite

Management of Geminiviruses Focusing on Small RNAs in Tomato |  237

(A)

(B)

(C)

Figure 12.1  Genome map of type species of geminiviruses. (A) DNA-A component and the encoded transcripts shown by arrows. CR represents the common region containing TAATATTAC sequence, which serves as the origin of DNA replication. (B) DNA-B component and the associated transcripts. (C) β-satellite

viruses. Monopartite viruses generally contain a satellite DNA (β-DNA) in their virion. These viruses encode the crucial protein (Rep) for their rolling circle DNA replication (RCR) and also the coat-protein (CP) for establishing their virion coat. Bipartite viruses encode movement proteins (MP) to facilitate intracellular and inter cellular transfer of viral genome. The origin of RCR is conserved across all geminiviruses and the replication is initiated at a specific point within a conserved 9-mer DNA sequence by the action of the sequence specific binding of and nicking by the Rep protein. Classification Geminiviridae is the largest plant virus family, which currently includes 325 species (www.ictvonline. org/virusTaxonomy. asp). The family Geminiviridae has been divided into seven genera by International Committee on Taxonomy of Viruses (ICTV) (www.ictvonline.org/virusTaxonomy. asp) on the basis of their genomic organization, sequence relatedness and insect vector (Varsani et al., 2014). Recently an eighth genus Capulavirus is being proposed to accommodate two highly divergent geminiviruses with no known vector. The third member identified as Capulavirus is Alfalfa leaf curl virus is transmitted by Aphis craccivora (Aphid). This is the first report of a geminivirus transmission by aphids (Roumagnac et al., 2015). Mastrevirus They have monopartite genomes with a conserved arrangement and contain four open reading frames. ORF encoded on virion strand [V1 and V2, code

coat protein (CP) and movement protein (MP) respectively] and two ORFs are transcribed from the complementary sense strand (C1 and C2, also known as RepA and RepB respectively). The virion and complementary strands are separated by small intergenic region (SIR) and large intergenic region (LIR). They are transmitted by leafhopper. The majority infects monocotyledonous plants of the family Poaceae. The type members include MSV and Wheat dwarf virus (WDV). Some of them infect only dicotyledonous plants, for example Tobacco yellow dwarf virus (TYDV) and Chickpea chlorotic dwarf virus (CpCDV) (Nahid et al., 2008). Curtovirus Curtoviruses also have circular single-stranded monopartite genomes. They have commonly seven open reading frames (ORFs). V1 to V3 are encoded by sense strand and C1 to C4 are encoded by antisense or complementary strand. The capsid or coat protein (CP), which is highly conserved, is encoded by V1 ORF and is necessary for systemic infection. V2 and V3 being unique to Curtoviruses helps in regulating genomic single -tranded (ss) and dsDNA production and cell to cell movement of the virus respectively. C1 encodes for replication-associated protein (Rep), C2 protein helps in movement of the virus, C3 stands for Replication enhancer and C4 affects cell division and symptom expression. Virus such as Beet curly top virus (BCTV) are transmitted by beet leafhopper (Circulifer tenellus) and can infect a wide variety of dicotyledonous plant species like beet, tomato, spinach (Lam et al., 2009).

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Topocuvirus Tomato pseudo-curly top virus (TPCTV) has a monopartite genome and has six ORFs. Sense strand encodes V1 and V2 and the complementary strand encodes for C1, C2, C3 and C4 proteins. Function of these proteins is known to be same as Curtovirus. They are transmitted by a treehopper (Micrutalis malleifera) (Fondong, 2013). Begomovirus The majority of geminiviruses belongs to the genus Begomovirus, of which the type member is Bean golden mosaic virus (BGMV), although Tomato leaf curl virus (ToLCV), Tomato golden mosaic virus (TGMV) and Mung bean yellow mosaic virus (MYMV) have all been studied extensively. In India ToLCV causing ToLCD has been first reported in 1948 (Vasudeva and Samraj, 1948) and is known to continuously emerge in new areas. The genome consists of six ORFs, V1 and V2 are in the virion strand. V2 encodes for coat protein. Complimentary strand encodes for C1, C2 and C3 proteins and these ORFs are partially overlapping and a small ORF C4 is located within the C1 ORFs but in different reading frame. All begomoviruses are transmitted by whiteflies (Bemisia tabaci) and collectively infect a wide range of dicotyledonous plants. They can be either monopartite or bipartite genomes (DNA-A and DNA-B). In the New World (NW), the vast majority of begomoviruses are bipartite. BGMV and TGMV are the only known viruses having monopartite genome. In addition, some are associated with ssDNA known as alpha and betasatellites. Becurtovirus This genus contains two species. One of them is Beet curly top Iran virus (BCTIV), which is leafhopper (Circulifer haematoceps) transmitted and is the better characterized of the two. Genome encodes three viral sense gene V1, V2 and V3 and two complementary sense genes C1 and C2. Hosts identified so far include sugarbeet, tomato, common bean (Phaseolus vulgaris) and cowpea (Vigna unguiculata). Eragrovirus Contains a single species, Eragrostis curvula streak virus (ECSV), which has so far only been identified in a monocotyledonous weed for which vector has

yet to be identified (Varsani et al., 2014). It encodes for four ORFs: C1 and C2 from viral sense strand and V1 and V2 from the complementary strand. Turncurtovirus Contains a single species, Turnip curly top virus, which is leafhopper (Circulifer haematoceps) transmitted and has been identified in a wide range of dicotyledonous hosts. The genome encodes for four proteins from viral sense strand (C1, C2, C3 and C4) and two from complementary strand (V1 and V2). DNA replication Geminiviruses replicate via rolling circle replication (RCR) in nuclei of the host plant cell using host DNA polymerases. Replication starts with double-stranded DNAs as an intermediate in the host nucleus. Viruses encode AL1 and Rep protein which is essential for the replication. After the virus penetrates into the host cell, it releases the ssDNA which finds its way into the nucleus. The ssDNA interacts with several host proteins including DNA replication protein PCNA (proliferation cellular nuclear antigen), cell cycle regulator retinoblastoma-related protein (RBR), and replicative DNA polymerases and eventually gets converted to double-stranded (ds) DNA or RF (replicative forms) forms etc. Rep protein nicks viral DNA at specific position to initiate RCR. The Rep protein acts as both the initiator and terminator of the RCR-fork. Both the DNA component (DNA-A and DNA-B) replicate using same principle. Although DNA-A and DNA-B are very different in the sequence they share a common region of around 200 nt, known as intergenic region/common region (CR/IR) (Revington et al., 1989; Lazarowitz et al., 1992; Fontes et al., 1994) This CR has repeat sequence that makes hairpin structure known as ‘iterons’. Rep recognizes and binds specifically to the iterons to make nick in the specific nonamer region (TAATATT↓AC; ↓; site of nick) present in iterons to initiate RCR (Pant et al., 2001). Rep also has helicase activity that helps in the extension of the RCR fork (Choudhury et al., 2006). One round of RCR ends with one viral dsDNA along with one single-stranded viral DNA. The new viral ssDNA is circularized using Rep protein. This can be further used as a template for replication making several viral ssDNA. The ssDNA can either be transported to the neighbouring cell

Management of Geminiviruses Focusing on Small RNAs in Tomato |  239

AC1 (Rep gene), AC2, AC3 and AC4 are transcribed in antisense direction. The DNA-B encodes two genes, one (BC1) on the complementary strand and the other (BV1) on the virion strand.

with the help of movement protein (MP) through plasmodesmata or packaged in virions. Transport to the neighbouring cells helps in local transport of virus and finally leading to long-distance spread (Fig. 12.2).

Coat protein The coat protein (CP) (MW = 30.3 kDa; 260 amino acids) is encoded by the AV1 gene. It is the lateexpression gene and is the only structural protein of geminivirus particles and is essential for the systemic infection in monopartite begomoviruses (Noris et al., 1996). In addition to virus genome packaging, CP has been known to be involved in other functions like insect transmission (Boulton et al., 1993), systemic spread of virus (Liu et al., 1997); and in case of monopartite virus, it helps in

Gene expression and function Most studies on geminiviral gene functions have been conducted using begomoviruses. Transcription of the geminivirus genome is bidirectional to produce 3′ co-terminal virion-sense and complementary-sense transcripts (Fig. 12.1). DNA-A codes for six proteins mainly required for virus DNA replication and encapsidation. They are AV1 (coat-protein gene) and AV2 (Pre-coat protein gene), being transcribed in sense strand, whereas

Recombination dependent replication

Rolling circle replication

Whitefly carrying geminivirus particles

Uncoating ssDNA

Infection of plant cell

ssDNA

CP

Replication

dsDNA Entry in plasmodesmata for virus spread

Encapsidation

n criptio Trans

MP

AAA AAAA AAAA A Entry of viral proteins back into nucleus

Translation

Viral mRNAs Viral proteins

Whitefly infection with geminivirus

Figure 12.2  The geminivirus lifecycle: Whitefly infected with geminivirus releases viral ssDNA to the plant cells cytoplasm which is carried to the nuclus for amplification of the DNA. The ssDNA is first converted into dsDNA using host machinery, followed by rolling circle replication (RCR). Viral mRNA are exported to the cytoplasm for translation of different viral protein. These viral DNA with the help of movement protein can also enter into neighbouring cell for viral spread. MP represents Movement protein.

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shuttling of viral DNA into and out of nucleus (Liu et al., 1999). Pre-coat protein The protein of MW = 13.5 kDa with 116 amino acids is encoded by the AV2 gene (V2 in monopartite). In ToLCV, AV2 protein helps in accumulation of ssDNA (Rigden et al., 1993). It is known to be essential for the viral movement to the monopartite virus. AV2 protein of ToLCV was reported to be involved in suppression of gene silencing (Zrachya et al., 2007) and to interact with the host cell SGS3 protein (Glick et al., 2008, 2009). Rep protein The replication-associated protein (Rep) encoded by the AC1 (also called AL1) in bipartite and C1 (also called L1) in monopartite (MW = 41 kDa; 357 amino acids) viruses. It is moderately conserved in sequence, position and function and expressed under the control of bidirectional promoter in the inverted repeat region in the genome (IR) (Hanley-Bowdoin et al., 2013). This protein initiates viral DNA replication by rolling circle amplification mechanism and possesses sequence specific DNA binding activity (Singh et al., 2015). This is the only protein essential for replication and is likely involved in recruitment and assembly of the viral replisome (complex including viral protein and host factors). Replication enhancer protein (REn), also known as AL3/C3, interact with Rep and host factors (Settlage et al., 2005) to enhance viral DNA replication. Both Rep and REn bind to proliferation cell nuclear antigen (PCNA) (Castillo et al., 2003; Bagewadi et al., 2004) which is a processivity factor for host DNA polymerase-δ. TGMV Rep has also been reported to interact with small ubiquitin-related modifier (SUMO) conjugation enzyme (SCE-1) and modulate SUMO level in host, and therefore create suitable environment for virus replication (Sanchez-Duran et al., 2011). In addition to the above function, Reps regulate viral gene expression by blocking complementary-sense gene expression of begomoviruses (Eagle et al., 1994; Shivaprasad et al., 2005) and mastreviruses (Hefferon et al., 2006). Rep was also shown to act as a replicative helicase in case of Mungbean yellow mosaic India virus (MYMIV) (Choudhury et al., 2006).

TrAP protein The transcriptional activator protein (TrAP) (MW = 15.6 kDa; 135 amino acids) is encoded by the AC2 gene in bipartite and C2 in monopartite viruses. TrAP is a positional analogue of C2 (or L2) protein in curtoviruses and topocuviruses and is a multifunctional protein. It is reported as a suppressor of post-transcriptional gene silencing (PTGS) in plant (Kumar et al., 2015). AC2 encodes for transcriptional activator protein (TrAP). AC2 along with AC3 is expressed from a dicistronic transcript (Shivaprasad et al., 2005). TrAP activates the transcription of coat protein and movement protein (MP) encoding gene. Transactivation is dependent on its zinc-finger and C-terminal acidic domains (Hanley-Bowdoin et al., 1999). Recently the AC2 function has been shown as replication brake in CSR, RCR and/or RDR with its DNA binding capacity of the zinc finger motif (Krenz et al., 2015). TrAP is also found to be involved virus pathogenicity (Hong et al., 1996) and suppression of gene silencing (Kumar et al., 2015). AC3 protein The replication enhancer protein (REn) is encoded by AC3 ORF (MW = 15.9 kDa; 134 amino acids) in bipartite and by C3 (or L3) in Curtovirus and monopartite virus. AC3, although not essential for infectivity, interacts with Rep and act as replication enhancer. Although REn, encoded by AC3, is not essential for virus replication, it helps in viral DNA accumulation and symptom development. AC4 protein The AC4 protein (or AL4) (MW = 10.9 kDa; 98 amino acids) is encoded by the AC4 gene. AC4 are contained completely within the AC1 ORF, but is expressed in different reading frame. This is the least conserved geminiviral protein and has different functions in monopartite and bipartite geminivirus. AC4 is considered an important symptom determinant (Krake et al., 1998) and is also known as silencing suppressor. TGMV AC4 is involved in virus movement (Pooma and Petty, 1996) but not so critical for virus infection. The ability of AC4 and C4 to suppress RNAi is conserved in many monopartite and bipartite virus example in East African Cassava mosaic virus (EACMV) (Fondong et al., 2007) either by binding miRNA and siRNA (Chellappan et al., 2004).

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Nuclear shuttle protein (BV1) BV1 is one of the two protein encoded by B component of bipartite begomovirus. It encodes for nuclear shuttle protein (NSP), which is required for movement of viral ssDNA between nucleus and cytoplasm (Noueiry et al., 1994). In addition to this, it is an avirulence determinant in some host. Recent studies have shown histone H3 to interact with MP and NSP and form a complex along with viral DNA and help the movement of nascent viral DNA to the cytoplasm from nucleus for cell to cell and long distance movement. Movement protein (BC1, V2) The MP is encoded by the BC1 (or BL1) ORF in case of bipartite virus from DNA-B component. With cooperative interaction with NSP, it is required for long distance and cell to cell movement (Briddon, 2015). Begomovirus-associated satellites Alpha and betasatellites are small ssDNA components. They are nearly half the size of the genome of their helper viruses and are associated with majority of monopartite begomoviruses; and requires helper virus for replication (Briddon and Stanley, 2006). Betasatellites have a conserved structure encoding a single gene product known as βC1. However they are highly diverse and have little sequence similarity to their helper DNA. βC1 is known to be involved in virus movement and pathogenicity determinant (Yang et al., 2011). In contrast, unlike betasatellite, alphasatellites are capable of autonomous replication but like betasatellite needs helper virus for movement and encapsidation (Mansoor et al., 1999). However the biological function of alphasatellites is unclear and are established to be non-essential for infection. These satellite DNAs depend on helper virus for systemic infection (Briddon and Stanley, 2006). Betasatellites have been demonstrated to be essential for disease symptom production (Briddon et al., 2015; Cui et al., 2004). The βC1 proteins and βC1 promoters can also affect symptom induction (Ding et al., 2009). Transgenic Nicotiana benthamiana and Nicotiana tabacum plants expressing TYLCCNB-βC1 was found to develop severe developmental abnormalities which imply that

βC1 interferes with plant cell division (Cui et al., 2004) and other host functions involved in RNAi activities. βC1 proteins are also known to function as RNA silencing suppressors. For example, TYLCCNB-βC1 associated with Tomato leaf curl China virus (ToLCCNV) and βC1 proteins associated with Bhendi yellow vein mosaic virus (BYVMV) (Gopal et al., 2007; Sharma et al., 2010) are well-studied RNAi suppressors. Further βC1 was also found to contribute in methylation inhibition and suppression of TGS (Yang et al., 2011) of viral nucleosomes. Until recently, alphasatellite have been only found in monopartite virus associated with betasatellite (Xie et al., 2010). However, only few betasatellite-containing viruses were found to contain alphasatellite. Recently alphasatellites have been found associated with bipartite begomovirus in Brazil (Paprotka et al., 2010) and Venezuela (Romay et al., 2010). They are mostly found in the Old World but the first report for alphasatellite of New World was identified in Singapore (referred as DNA-2) (Saunders et al., 2002). More recently DNA-2 was found in India (Zaffalon et al., 2012). Developing resistance to geminiviruses Breeding for resistance and developing agronomic practice to reduce the infecting virus load are generally applied to protect crops. Here we focus mostly on tomato and the leaf curl viruses of tomato. Virus resistance using plant resistance genes (breeding) Disease resistance (R) genes in plants code for proteins that recognize incompatible pathogens (bacteria, fungi or virus) and as a result trigger events that lead to disease resistance in the host plant. There are many known R-genes which have been used in past for crop improvement programmes. Based on their membrane-spanning domain and motifs, they are divided into eight groups: (a) leucine-rich repeats (LRR); (b) nucleotide-binding site (NBS); (c) protein degradation domain (PEST); (d) endocytosis cell signalling domain (ECS); (e) Helminthosporium carbonum toxin reductase enzyme (HM1); (f) nuclear

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localization signal (NLS) (g) transmembrane domain (TrD); (h) amino acid domain (WRKY); (i) TIRToll/interleukin-1- receptors; and (j) C-C – coiled coil. The advantage of using R genes in resistance breeding programmes is that the input of pesticides is reduced, with efficient reduction of pathogen growth. However, breeding for resistance from one species to gene pool of another by repeated backcrossing takes several generations and is time-consuming (Gururani et al., 2012). The use of these dominant R genes appears to be an attractive option for breeders when available. Most of them fall to nucleotide binding site-leucine-rich repeat (NBS-LRR) class of resistance genes. These genes recognize pathogen avr factors related to diverse viral gene product. The NBS-LRR virus resistance genes mostly confer complete resistance (qualitative) and are sometimes regulated by cascade of small RNAs via trans-acting siRNAs, which requires miRNA as a trigger. (Zhai et al., 2011; Shivaprasad et al., 2012; Li et al., 2012). There are currently about 12 genes available providing viral resistance (Table 12.1). N gene of tobacco against TMV and Rx gene of potato against Potato virus X (PVX; Potexvirus) are the best-characterized

virus-resistant genes (Liu, 2010). As apparent in Table 12.1, ssDNA virus does not have dominant resistant gene. Most of them provide resistance to (+) ssRNA. The breeding programmes continuously work upon transfer of the wild type resistance genes to the cultivated tomato varieties. But due to complex genetics of these resistant gene and interspecific barriers between wild type and cultivated varieties (Lapidot, 2007) breeding for TYLCV resistance is very slow. Moreover most of the genes identified for resistance in tomato conferred delayed symptom development only (Galvez et al., 2014). Recently, two TYLCV resistance genes, Ty-1 and Ty-3 is being characterized as resistance gene against TYLCV. These resistance genes are allelic to each other and encodes for DFDGD-class of RNA-dependent RNA polymerase (RdRps) unlike other resistance genes against viruses that encode NBS-LRR proteins (Verlaan et al., 2013). Both Ty-1 and TY-3 loci are derived from the chromosome #6 of wild tomato species S. chilense. Besides these, Ty-2, Ty-4 and ty-5 loci also confer resistance to TYLCV. The Ty-2 locus is derived from chromosome #11 of the wild species S.

Table 12.1  Resistance genes of hosts known to act against plant viruses R gene

Virus

Strand

Genome

AVR

Host

N

Tomato mosaic virus

+

ssRNA

Replicase

Tobacco

HRT

Turnip crinkle virus

+

ssRNA

Coat protein

A.thaliana

Rx1

Potato virus X (PVX)

+

ssRNA

Coat protein

Potato

Rx2

PVX

+

ssRNA

Coat protein

Potato

Rsv1

Soybean mosaic virus

+

ssRNA

Unknown

Soybean

RT4-4

Cucumber mosaic virus

+

ssRNA

2a gene

Phaseolus

RTM1

Tobacco etch virus (TEV)

+

ssRNA

Unknown

A.thaliana

RTM2

TEV

+

ssRNA

Unknown

A.thaliana

RCY1

CMV

+

ssRNA

Coat protein

A.thaliana

Rep

TYLCSV

+

ssDNA

AC4

N.bentamian

Sw5

Tomato spotted wilt virus

_

ssRNA

Movement protein

Tomato

Tm22

Tomato mosaic virus

+

ssRNA

Movement protein

Tomato

Y-1

Potato virus Y

+

ssRNA

Unknown

Potato

Management of Geminiviruses Focusing on Small RNAs in Tomato |  243

habrochaites and two candidate genes within this locus might play a role in resistance to TYLCV (Yang et al., 2014). Both Ty-1 and Ty-2 are dominant markers and are widely used against various strains of monopartite begomoviruses by breeders. But neither of them is effective against bipartite begomoviruses and some strains of TYLCV can bypass the individual resistances offered by either Ty-1 or Ty-2 locus. However, when both of them are pyramided together, a high level of resistance to bipartite begomoviruses could be achieved (Mejia et al., 2010). The Ty-4 locus is also derived from the wild species S. chilense and is located in the chromosome #3. The recessive ty-5 loss of function mutation is most probably derived from a complex of wild species of S. peruvianum accessions (Anbinder, 2009). Transgenic approaches for virus resistance in plants Breeding for resistance has many challenges, including availability of geminivirus resistance genes, development of dominant molecular markers linked to it and introgression of resistance gene into susceptible cultivars. Genetic engineering approach involves transferring gene of interest from one organism to another by means other than sexual crosses involving Agrobacterium tumefaciens bacterium that naturally transfer DNA during disease process to plant (Bundock et al., 2002). This approach can be used to express virus derived genes directly or in the form of hairpin and miRNA to make plant resistant against virus. Transgenic plant developed expressing truncated Rep gene was shown to be resistant against Tomato yellow leaf curl Sardinia virus (TYLCSV) up to 15 weeks post virus inoculation (Noris et al., 1996; Brunetti et al., 1997). RNAi has evolved to protect plants against viruses and application of this technology can lead to development of virus resistance in the engineered plants. Overexpression of artificial microRNAs (a-miRs) to silence the virus encoded pathogenic RNAi suppressors, namely AC2 or AC4, in tomato plant has been shown to confer resistance to ToLCNDV (Yadava et al., 2010; Yadava and Mukherjee, 2012). Recently transgenic plants overproducing amiRs against AV2/AV1 transcripts were also reported to be tolerant/resistant against various accessions of ToLCV (Tien et al., 2013).

RNA interference The loss of anthocyanin in transgenic petunia heralded the era of RNAi science. RNA interference (RNAi) in its present form was first described in nematode, Caenorhabditis elegans (C. elegans) and it was found to be at least 100-fold more efficient in silencing the gene compared to antisense RNAs (Fire et al., 1998; Agrawal et al., 2003). Subsequently RNAi was also observed in insects (Kennerdell and Carthew, 1998), frog (Oelgeschlager et al., 2000), and other animals including mice (Wianny and Zernicka-Goetz, 2000; Svoboda et al., 2000). In plants, pathogen A. tumefaciens has been exploited long back to transfer known DNA fragment into the plants. However, sometimes when more than one transgene copy is incorporated, low expression of transgene was observed. The same effect was observed when extra copy of endogenous gene was added leading to so called co-suppression of endogenous gene. All of such phenomena can be grouped together as reports of plant RNAi (Napoli et al., 1990). RNAi is a conserved eukaryotic mechanism involving small non-coding RNAs mediating repression of transcription causing either Transcriptional Gene Silencing (TGS) or degradation of mRNA called post-transcriptional gene silencing (PTGS). In plants, PTGS is a sequence specific defence mechanism that can silence both cellular and viral mRNAs (Hamilton and Baulcombe, 1999) whereas TGS involves pairing based on sequence similarity resulting in DNA secondary structure formation, that attracts heterochromatin components, leading to transcription inhibition by methylation (Vaucheret and Fagard, 2001). Methylation in case of TGS is not limited to CpG or CpXpG sequences. If methylation occurs in the coding sequence, there could be nil or minimal effect on the transcription whereas if it occurs in the promoter sequence, it can result in TGS and unlike PTGS, the former is heritable and stable (Rodríguez‐Negrete et al., 2013). PTGS can take place by mainly two processes: Target transcript degradation and translational repression. Highly complementary binding of small RNAs to the target normally triggers target transcript degradation. Translational repression occurs in case of partial complementary binding of small RNAs to coding region or untranslated region (UTR) of target transcripts. The binding does not lead to splicing but it inhibits the assembly of translational apparatus,

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hence silences the protein expression (Huntzinger and Izaurralde, 2011). The small RNAs, regulating gene expression, are mainly classified into two major categories as microRNAs (miRNAs) arise from imperfectly base paired foldback structures and small interfering RNAs (siRNAs) arising from dsRNA hairpin structures. MiRNA precursor called pri-miRNA generally arises due to Pol II activity whereas siRNA generally arise by the activity of RNA dependent RNA polymerase (RdRp) on suitable RNA transcript. Drosha and Dicer (DCR) process pri-miRNAs in animals and DCR-Like proteins (DCL) along with partner proteins carry out the same job in plants, leading to generation of 21–24 nt duplex from miRNA and siRNA precursors. The duplex are then methylated at 2′-OH position on ribose ring of the 3′ terminal nucleotide by HEN1 protein in plants to protect duplex from uridylation, which may otherwise lead to degradation by exonuclease (Ramachandran and Chen, 2008). Mature miRNA/siRNA duplex are then exported to the cytoplasm by HASTY (HST). The mature duplex are then recruited by AGO proteins and other protein partners to form RISC complex where one of the strand is selected for targeting mRNA based on complementarity. The mechanism of selecting the strand for degradation is mostly thermodynamically controlled although a few protein components are also reportedly involved. The sorting into RISC is probably directed by the 5′ terminal nucleotide since most of miRNAs containing 5′ uridine are loaded into AGO1, 5′ adenine into AGO2 and 5′ cytosine into AGO5 (Mi et al., 2008). Although gene families encoding common components involved in the biogenesis are conserved across eukaryotic kingdoms, small RNA pathways are specialized and diversified among and between kingdoms. This diversification can be seen in Arabidopsis thaliana harbouring 4 DCLs, 10 AGOs and at least 6 functional RDR genes (Agrawal et al., 2003). Among these, AGO1 and DCL1 are required to guide cleavage in case of miRNA. The siRNAs are of several classes and they are made by distinct pathways. Hetrochromatin siRNAs which are 24 nt are transcribed from RNA polymerase IV and involve RDR2, DCL3, and AGO4 for their activity. These are responsible for cytosine methylation of DNA and Lys-9 methylation on histone H3 (chromatin modification). The siRNAs from exogenous source

such as viral and transgenic may involve RDR1 and RDR6 and DCL2 in case of some viruses. DCL4 and DCL2 are involved in production of 20 to 22-nt siRNA with perfect complementarity (Hamilton and Baulcombe, 1999; Hamilton et al., 2002; Bouche et al., 2006). Loading of small RNAs onto specific AGOs, is strongly determined by 5′-terminal nucleotide of plant small RNAs. For example, in A. thaliana, AGO1 and AGO10 binds to small RNAs with 5′-uracil whereas AGO2, AGO4, AGO6, AGO7 and AGO9 has bias for adenine and AGO5 prefers cytosine (Havecker et al., 2010; Takeda et al., 2008; Mi et al., 2008). Additional layer of complexity is determined by miRNA-loading which is partially regulated by miRNA-duplex structure based on complementarity and bulges. For example, miR166 interacts with AGO10 with one internal base mismatch flanked by two paired bases within the mature duplex (Zhu et al., 2011; Liu et al., 2009; Ji et al., 2011) and prevent miR166 from loading onto AGO1 and thus silencing of transcription factors (homeodomain Leu zipper) in the shoot apical meristem (Zhu et al., 2011; Liu et al., 2009). Trans acting siRNAs(tasiRNA) which is a class of endogenous siRNA arise from polIII genes requires RNA dependent RNA polymerase (RDR6) which is needed to make double strand RNA from the transcript and suppressor of gene silencing 3 (SGS3). MicroRNAs Plant microRNAs (miRNAs), transcribed mostly by polymerase II, are a class of endogenous small regulatory RNAs (20–22 nt) having partially double-stranded stem loop structure ( Jones-Rhoades et al., 2006). Initially primary miRNAs(pri-miRNA) are transcribed as long primary transcripts. These pri-miRNAs are cleaved by DCL1 in two-step process: first into pre-miRNA and then into mature miRNA duplex (miRNA/miRNA*). The mature miRNAs are then loaded in the RISC complex and target mRNA for silencing either through cleavage or translational repression (Chen, 2004; Brodersen et al., 2008). The repression also occurs through transcriptional inhibition (Khraiwesh et al., 2010). In plants, where miRNAs usually are 20–24 nt long and targets most often in the coding sequence, cleaving the target. However in animals, miRNAs are usually 20–22 nt, mostly acting as translational inhibitor, often targeting motifs in 3′UTRs

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(Untranslated regions) with several mismatches (Kamthan et al., 2015). Plant miRNA are less conserved compared to animal miRNAs (Bartel, 2004). Plant miRNA-targets mostly include transcription factor genes, some hormone receptor genes (Navarro et al., 2006) few biosynthetic genes (Fujii et al., 2005). Many plant miRNA are known to be involved in PTGS and play role in abiotic stress, pathogen response and many developmental processes etc (Khraiwesh et al., 2010). For example up-regulation of miR393, miR160 and miR167 has been reported during drought and/or salt stress in several plant species (Sunkar et al., 2012). The artificial microRNAs (a-miRs) has found wide usage to silence desired target mRNAs both in animals and plants (Fowler et al., 2016). Short-interfering RNAs Short-interfering RNAs (SiRNAs) arise from shorthairpin RNA or long dsRNA precursors. These are homologous to the target genes for silencing to take place (Fire et al., 1998). Transposons, introduction of transgene or the entry of virus into the plant can trigger this RNAi pathway. Double-stranded RNA is then cleaved by dicer enzyme into siRNA (21–25 nt), which are 5′-phosphorylated and with two nucleotides 3′ end overhang (Hamilton and Baulcombe, 1999). Degradation of sense strand, having same sequence as target strand, occurs in siRNA-induced silencing complex (siRISC). Antisense strand of siRNA is incorporated into RISC which cause cleavage of target mRNA. The cleavage occurs due to the slicing activity of AGO and other effector proteins (Kamthan et al., 2015). The siRNAs are mainly divided into heterochromatic siRNAs, secondary siRNAs and natural antisense transcript siRNAs (NAT-siRNAs). Secondary siRNA are further divided into trans-acting siRNA if they are phased, and NAT-siRNA that can be cis or trans (Axtell, 2013). Using hairpin RNA, genes can be silenced leading to the identification of their functions by examining the silenced phenotype. Following entry of plant viruses in host cells, the viral transcripts are converted to dsRNA forms by various means. These dsRNAs are substrates of DCL proteins and thus siRNAs (VsiRNAs) emerge from all over the viral genome. These siRNAs do not allow the viral proteins to form, thus acting as defence factors of the host (Fig. 12.3). Transgenic

beans have been developed making use of this natural defence activity to resist Bean golden mosaic virus (BGMV). These transgenic beans are grown and consumed in Southern America (Aragao, 2014). Trans-acting siRNA Trans-acting small interfering RNAs (tasiRNAs) belong to the class of siRNAs (21 nt). They are transcribed from TAS loci by RNA polymerase II. After cleavage with specific miRNA, they are made double-stranded by RDR6/SGS3 proteins. Double-stranded transcripts are processed in subsequent 21 nt phased manner by DCL4. They suppress the expression of target sequence by hybridizing with the target mRNA through 21-nt sequence generated. A. thaliana has four families, i.e. TAS1, TAS2, TAS3 and TAS4 (Allen et al., 2005). Among all known families, TAS3 is the most conserved and it is known to be present in rice, maize, moss and gymnosperms (Fei et al., 2013). TAS1 (loci: TAS1a, TAS1b, TAS1c) and TAS2 families are targeted by miR173, whereas the TAS3 family (loci: TAS3a, TAS3b and TAS3c) is targeted by miR390 and TAS4 by miR828 (Rajagopalan et al., 2006; Yoshikawa et al., 2005; Allen et al., 2005). TAS transcript may require one miRNA binding site (BS), for example, in case of TAS1, TAS2, TAS4 and TAS5 or two miRNA BSs (TAS3) for the maturation of tasiRNA and are transcribed from the non-coding regions of the genome (Yoshikawa et al., 2005). Based on the 5′ base on the miRNA miR173, in case of TAS1 and TAS2,is loaded onto AGO1/ AGO4 whereas miR390, which is involved in production of trans-acting siRNAs from TAS3 transcript, is preferentially loaded onto AGO7 (Axtell et al., 2006). The association is preferred by 5′-adenine and mismatch at position 11 of the miRNA duplex. Tomato has been reported to have three families of TAS genes: TAS3, TAS4 and TAS5. TAS1/TAS2 derived ta-siRNAs are known to target pentatricopeptide repeat gene transcripts (Allen et al., 2005). TAS3 are known to target Auxin response transcription factors ARF2, ARF3, ARF4 (Garcia et al., 2006). TAS4siRNAs are known to regulate MYB transcription factor including MYB113, MYB75 and MYB90 involved in biogenesis of anthocyanin (Rajagopalan et al., 2006; Singh et al., 2016).

246  | Archana and Mukherjee Cytoplasm

VsiRNA

Nuleus

Me-RISC Me

ati Re pli c

Viral ssDNA

on

VdsDNA

Viral ssDNA

VdsDNA

Transcription V-mRNA

(TGS)

RdRP

DCL VsiRNA Or Endo-VsiRNA

V-RISC VsiRNA

VsiRNA

Endo VsiRNA

PTGS Endo AC2/AC4 siRNA

Figure 12.3  Biogenesis and Function of VsiRNAs of begomoviruses. Viral transcripts and V-siRNAs are coloured in blue. Endogenously produced siRNAs of transgenic plants are coloured in red. RdRP is RNA dependent RNA polymerase, mostly of type6 (RDR6). DCL represent Dicerlike proteins, mostly of type 2 and 4 (i.e. DCL2/DCL4). PTGS and TGS represent post-transcriptional gene silencing and transcriptional Gene silencing respectively. Me is the methyl moiety shown in red inverted arrow on double-stranded (ds) begomoviral DNA. The ssDNA is the single-stranded viral genome shown in large black empty circles. C and N respectively represent the cytosol and nucleus of the plant cell, the wall of which is shown with plasmodesmatal channels.

RNA interference approach to generate resistance to geminiviruses At present, RNAi is the most widely investigated strategy to combat geminivirus infections. However homology between the transgene and target sequence decides the effectiveness of this approach and its limits for broad-spectrum resistance. Until this date only a single engineered resistance to a geminivirus has been commercialized which is the transgenic bean plant with hairpin RNAi construct containing BGMV replication associated protein (Rep) (Aragão et al., 2013). Since geminiviruses strictly require Rep gene for replication, it is considered as very important RNAi target (HanleyBowdoin et al., 2013). In most cases dsRNA or hpRNA constructs have been used for silencing. The miRNA precursors have also been used as structure for producing artificial miRNAs, which target geminiviral transcripts. Recently it has been shown that miRNA-encoding transcripts can be modified

specifically and successfully to produce an artificial miRNA (amiRNA) in Arabidopsis, C. elegans, S. lycopersicum and other organisms (Schwab et al., 2006; Niu et al., 2006). This is achieved by replacing the endogenous miRNA and its complementary sequence in the RNA fold-back structure, referred to as the miRNA precursor, so that the amiRNA is processed as if it were the endogenous miRNA and the processed amiR guides cleavage and suppression of the selected target genes. Transgenic tomato plants had been developed that overproduce either AC2 or AC4 based amiRs and challenged the transgenics with ToLCNDV. Some transgenic lines were found to be significantly resistant against ToLCNDV (Yadava et al., 2010; Yadava and Mukherjee, 2012). Transgenic tomato using replicase (Rep) gene (untranslatable full length sequence, 1086 bp) of ToLCV in antisense orientation was made. Inheritability of transgene and high level of resistance was observed up to the T2 generation against

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the virus (Praveen et al., 2005). Another example of TGS was observed in cassava resistance to ACMV, transformed with full-length Rep gene in antisense orientation. At lower infection load (100 ng viral DNA/plant), several lines were found to be symptomless while at higher dose, symptoms were found to be reduced (Zhang et al., 2005; Fuentes et al., 2006). Tomato plants transformed with a ‘stem–loop’ construct containing castor bean catalase intron and 726 nt fragments from TYLCV Rep gene and transgenic resistant lines were observed to express ≈ 25 nt siRNA. Transgenic plants showed immunity even at high infection pressure. Artificial miRNA construct was designed also againstAV1 (Van et al., 2013) of ToLCNDV; and the transgenic tomato plants engineered using these constructs, were found to be highly tolerant against the virus until T2 generation. tasiRNA approach to control geminivirus In Arabidopsis artificial tasiRNAs (ata-siRNAs) have been shown to target RNAs that are produced from TAS1a (Felippes and Weigel, 2009), TAS1c (de la Luz Gutiérrez-Nava et al., 2008; Montgomery et al., 2008b), and TAS3a (Montgomery et al., 2008a; Felippes and Weigel, 2009) transcripts or from gene fragments linked to an upstream miR173 binding site (de-Felippes et al., 2012). The production of ta-siRNA seems to be complex but this system can be modified in a more flexible way to construct artificial ta-siRNA compared to miRNA system to suppress target sequences. Most recently ata-siRNA vector was designed and constructed. This vector can be used to clone any sequence of interest to produce ata-siRNA. The advantage of using this vector is that it uses a simple single step cloning process which is an alternative to multistep cloning for constructing hairpin RNA and the second advantage being use of this vector to target multiple gene or multiple target on a single insert. This vector was used to target two ToLCNDV genes separately namely AC2 and AC4. Tomatoes harbouring the construct were found to be effective against ToLCNDV to large extent (Singh et al., 2015). Other control strategies Several non-virus derived approaches have been reported which are found to be effective in

controlling geminiviruses. In A. thaliana or tomato plant, zinc-finger DNA binding protein was artificially designed to bind replication origin of either TYLCV or BCTV sequences and found to protect plants from virus infection (Briddon, 2015). Expression of GroEL, a protein binding to virus particles that are produced by insect endosymbiotic bacteria in tomato was found to ameliorate symptoms of TYLCV infection (Edelbaum et al., 2009). Transgenic tomato lines expressing aptamers were found to exhibit very mild or no symptoms on TYLCV or ToMoV infection (Reyes et al., 2013). Ribosome inactivating proteins (RIPs) are plant toxins that are naturally expressed in plants and exhibit antiviral activity against many animal and plant viruses. Dianthus caryophyllus has a potent RIP dianthin, which was exploited to engineer resistance in transgenic N. benthamiana against African Cassava mosaic virus (ACMV) by expressing dianthin from ACMV virion-sense promoter, transactivated by AC2 gene. The viral DNA accumulation was found to be significantly reduced in the tissue expressing dianthin (Hong et al., 1996). A model of DNA virus-resistance was created using artificial zinc finger protein (AZP) targeting the replication origin of Beet severe curly top virus (BSCTV). AZP efficiently blocked binding of Rep at the replication origin in vitro and all transgenic plant made in Arabidopsis expressing this AZP showed strong resistance phenotypically (Sera, 2005). Another approach based on ability of whitefly endosymbiotic GroEL to bind viruses. GroEL gene was expressed in N. benthamiana plants to interfere pathogenesis upon viral infection postulating that GroEL will bind to the virus. The transgenic plant was checked against TYLCV and CMV and was found to be tolerant. As a control Trichovirus grapevine virus A (GVA) and the Tobacco mosaic virus (TMV) were used which are not known to interact with GroEL and was found to be susceptible (Edelbaum et al., 2009). Conclusion Begomovirus-induced diseases lead to major crop losses worldwide. Many strategies have been used to combat the diseases. Earlier, to block the progression of the virus infectious process, viral protein over expression based approaches relied mostly on the expression of transgenic coat protein

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(CP). Other strategies exploit the plant’s innate defence mechanisms to combat viral pathogens. Prior inoculation of mild viral strain in plant confers resistance to severe strain in cross-protection. Of all the existing strategies, recently discovered RNAi based approaches are more successful to generate virus resistance, which include artificial miRNA and hairpin approach. ToLCV turns out to be a deadly virus damaging total yield and control of this virus poses severe challenges because of its fast changing biodiversity and its potential to encode several RNAi suppressors. Hence hairpin based approaches do not confer broad spectrum ToLCV resistance. However, a-miR and a-tasiRNA approaches are relatively more successful in achieving the goal of ToLCV resistance in tomato. Design of artificial miRNA involves multiple cloning steps and can be used to target one sequence at a time. Various studies that have been reported are artificial miRNA were used against RNAi suppressors AC2, AC4, AV1 and AV2 to combat geminivirus infections (Van et al., 2013). In principle, constructs harbouring multiple a-miRs can be designed to simultaneously silence multiple viral ORFs and generate robust virus resistance but such approaches have not been reported yet in tomato. The miRNA target site in TAS1 transcript was shown to be sufficient for the formation of tasiRNA and the remaining sequence in backbone of TAS1 play only a minor role in biogenesis of tasiRNA (Felippes and Weigel, 2009). With this view, most recently gene-silencing vector employing single step cloning process was constructed to target ToLCNDV. Since use of a tasiRNA may lead to off target effects, the DNA sequence for target selection is crucial to minimize off target related troubles. As the viral ORFs frequently overlap with each other, one can use this to silence two viral genes by targeting the overlapping region. For example, in the case of ToLCNDV, two regions from DNA-A, which include AC1 (Rep) and two RNA silencing suppressors AC2 and AC4 were selected. AC2 partially overlaps with AC1 (Rep) gene whereas AC4 completely overlaps with the AC1. Both AC2 and AC4 are transcribed from the same promoter as Rep. Targeting the overlapping regions for silencing led to successful inhibition of virus infection in tomato (Yadav et al., 2012; Tien, et al., 2013; Singh et al., 2015). Normally in field, infection involves mixture of viruses (Saunders and

Stanley., 1995; Rentería-Canett et al., 2011). It also includes some interactive viruses that can increase severity of the disease (Chakraborty et al., 2008). In such cases, both or multiple viral ORFs require to be targeted. The advantages of using atasiRNAs are: (1) tedious and difficult cloning steps can be avoided (2) the vector can be used to target multiple sequences in a facile manner (Ossowski et al., 2008). Though ToLCV is a difficult virus to control, pyramiding together several principles will confer robust resistance in tomato in near future. Acknowledgement SM is a recipient of INSA Senior Scientist Fellowship. References Abhary, M., Patil, B.L., and Fauquet, C.M. (2007). Molecular biodiversity, taxonomy, and nomenclature of Tomato yellow leaf curl-like viruses. In Tomato yellow leaf curl virus Disease: Management, Molecular Biology, Breeding for Resistance, Czosnek, H., ed. (Springer, Dordrecht), pp. 85–118. Agrawal, N., Dasaradhi, P.V., Mohmmed, A., Malhotra, P., Bhatnagar, R.K., and Mukherjee, S.K. (2003). RNA interference: biology, mechanism, and applications. Microbiol. Mol. Biol. Rev. 67, 657–685. Allen, E., Xie, Z., Gustafson, A.M., and Carrington, J.C. (2005). microRNA-directed phasing during trans-acting siRNA biogenesis in plants. Cell 121, 207–221. Anbinder, I., Reuveni, M., Azari, R., Paran, I., Nahon, S., Shlomo, H., Chen, L., Lapidot, M., and Levin, I. (2009). Molecular dissection of Tomato leaf curl virus resistance in tomato line TY172 derived from Solanum peruvianum. Theor. Appl. Genet. 119, 519–530. https:// doi.org/10.1007/s00122-009-1060-z Antignus, Y., Nestel, D., Cohen, S., and Lapidot, M. (2001). Ultraviolet-deficient greenhouse environment affects whitefly attraction and flight-behaviour. Environ. Entomol, 30, 394­399. Aragão, F.J.L. (2014). GM plants with RNAi - golden mosaic resistant bean. BMC Proc. 8, O24. Aragão, F.J., Nogueira, E.O., Tinoco, M.L., and Faria, J.C. (2013). Molecular characterization of the first commercial transgenic common bean immune to the Bean golden mosaic virus. J. Biotechnol. 166, 42–50. https://doi.org/10.1016/j.jbiotec.2013.04.009 Axtell, M.J., Jan, C., Rajagopalan, R., and Bartel, D.P. (2006). A two-hit trigger for siRNA biogenesis in plants. Cell 127, 565–577. Bagewadi, B., Chen, S., Lal, S.K., Choudhury, N.R., and Mukherjee, S.K. (2004). PCNA interacts with Indian mung bean yellow mosaic virus rep and down-regulates Rep activity. J. Virol. 78, 11890–11903. Bartel, D.P. (2004). MicroRNAs: genomics, biogenesis, mechanism, and function. Cell 116, 281–297. Boulton, M.I., Pallaghy, C.K., Chatani, M., MacFarlane, S., and Davies, J.W. (1993). Replication of maize streak

Management of Geminiviruses Focusing on Small RNAs in Tomato |  249

virus mutants in maize protoplasts: evidence for a movement protein. Virology 192, 85–93. Bouché, N., Lauressergues, D., Gasciolli, V., and Vaucheret, H. (2006). An antagonistic function for Arabidopsis DCL2 in development and a new function for DCL4 in generating viral siRNAs. EMBO J. 25, 3347–3356. Brown, J.K., Zerbini, F.M., Navas-Castillo, J., Moriones, E., Ramos-Sobrinho, R., Silva, J.C., Fiallo-Olivé, E., Briddon, R.W., Hernández-Zepeda, C., Idris, A., et al. (2015). Revision of Begomovirus taxonomy based on pairwise sequence comparisons. Arch. Virol. 160, 1593– 1619. https://doi.org/10.1007/s00705-015-2398-y Briddon, R.W. (2015). Geminiviridae. eLS. Online Publication. doi: 10.1002/ 9780470015902.a0000750 Briddon, R.W., and Stanley, J. (2006). Subviral agents associated with plant single-stranded DNA viruses. Virology 344, 198–210. Brodersen, P., Sakvarelidze-Achard, L., Bruun-Rasmussen, M., Dunoyer, P., Yamamoto, Y.Y., Sieburth, L., and Voinnet, O. (2008). Widespread translational inhibition by plant miRNAs and siRNAs. Science 320, 1185–1190. https://doi.org/10.1126/science.1159151 Brunetti, A., Tavazza, M., Noris, E., Tavazza, R., Caciagli, P., Ancora, G., and Accotto, G.P. (1997). High expression of truncated viral Rep protein confers resistance to Tomato yellow leaf curl virus in transgenic tomato plants. Mol. Plant Microbe Interact. 10, 571–579. Bundock, P., van Attikum, H., den Dulk-Ras, A., and Hooykaas, P.J. (2002). Insertional mutagenesis in yeasts using T-DNA from Agrobacterium tumefaciens. Yeast 19, 529–536. Castillo, A.G., Collinet, D., Deret, S., Kashoggi, A., and Bejarano, E.R. (2003). Dual interaction of plant PCNA with geminivirus replication accessory protein (Ren) and viral replication protein (Rep). Virology 312, 381–394. Chellappan, P., Vanitharani, R., Pita, J., and Fauquet, C.M. (2004). Short interfering RNA accumulation correlates with host recovery in DNA virus-infected hosts, and gene silencing targets specific viral sequences. J. Virol. 78, 7465–7477. https://doi.org/10.1128/JVI.78.14.74657477.2004 Chen, X. (2004). A microRNA as a translational repressor of APETALA2 in Arabidopsis flower development. Science 303, 2022–2025. https://doi.org/10.1126/ science.1088060 Choudhury, N.R., Malik, P.S., Singh, D.K., Islam, M.N., Kaliappan, K., and Mukherjee, S.K. (2006). The oligomeric Rep protein of Mungbean yellow mosaic India virus (MYMIV) is a likely replicative helicase. Nucleic Acids Res. 34, 6362–6377. Cui, X., Tao, X., Xie, Y., Fauquet, C.M., and Zhou, X. (2004). A DNAbeta associated with Tomato yellow leaf curl China virus is required for symptom induction. J. Virol. 78, 13966–13974. Czosnek, B., Czosnek, H., Navot, N., and Laterrot, H. (1990). Geographical distribution of Tomato yellow leaf curl virus. A first survey using a specific DNA probe. Phytopathol. Mediterr. 6, 1–6. Felippes, F.F., Wang, J.W., and Weigel, D. (2012). MIGS: miRNA-induced gene silencing. Plant J. 70, 541–547. https://doi.org/10.1111/j.1365-313X.2011.04896.x

de la Luz Gutiérrez-Nava, M., Aukerman, M.J., Sakai, H., Tingey, S.V., and Williams, R.W. (2008). Artificial transacting siRNAs confer consistent and effective gene silencing. Plant Physiol. 147, 543–551. https://doi. org/10.1104/pp.108.118307 Ding, C., Qing, L., Li, Z., Liu, Y., Qian, Y., and Zhou, X. (2009). Genetic determinants of symptoms on viral DNA satellites. Appl. Environ. Microbiol. 75, 5380– 5389. https://doi.org/10.1128/AEM.01193-09 Eagle, P.A., Orozco, B.M., and Hanley-Bowdoin, L. (1994). A DNA sequence required for geminivirus replication also mediates transcriptional regulation. Plant Cell 6, 1157–1170. https://doi.org/10.1105/tpc.6.8.1157 Edelbaum, D., Gorovits, R., Sasaki, S., Ikegami, M., and Czosnek, H. (2009). Expressing a whitefly GroEL protein in Nicotianabenthamiana plants confers tolerance to Tomato yellow leaf curl virus and Cucumber mosaic virus, but not to grapevine virus A or Tobacco mosaic virus. Arch. Virol. 154, 399–407. Fei, Q., Xia, R., and Meyers, B.C. (2013). Phased, secondary, small interfering RNAs in posttranscriptional regulatory networks. Plant Cell 25, 2400–2415. https://doi. org/10.1105/tpc.113.114652 Felippes, F.F., and Weigel, D. (2009). Triggering the formation of tasiRNAs in Arabidopsis thaliana: the role of microRNA miR173. EMBO Rep. 10, 264–270. https://doi.org/10.1038/embor.2008.247 Fire, A., Xu, S., Montgomery, M.K., Kostas, S.A., Driver, S.E., and Mello, C.C. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391, 806–811. https:// doi.org/10.1038/35888 Fontes, E.P., Gladfelter, H.J., Schaffer, R.L., Petty, I.T., and Hanley-Bowdoin, L. (1994). Geminivirus replication origins have a modular organization. Plant Cell 6, 405–416. Fondong, V.N. (2013). Geminivirus protein structure and function. Mol. Plant Pathol. 14, 635–649. https://doi. org/10.1111/mpp.12032 Fondong, V.N., Reddy, R.V., Lu, C., Hankoua, B., Felton, C., Czymmek, K., and Achenjang, F. (2007). The consensus N-myristoylation motif of a geminivirus AC4 protein is required for membrane binding and pathogenicity. Mol. Plant Microbe Interact. 20, 380–391. https://doi. org/10.1094/MPMI-20-4-0380 Fowler, D.K., Williams, C., Gerritsen, A.T., and Washbourne, P. (2016). Improved knockdown from artificial microRNAs in an enhanced miR-155 backbone: a designer’s guide to potent multi-target RNAi. Nucleic Acids Res. 44, e48. https://doi.org/10.1093/nar/ gkv1246 Friedman, M. (2013). Anticarcinogenic, cardioprotective, and other health benefits of tomato compounds lycopene, α-tomatine, and tomatidine in pure form and in fresh and processed tomatoes. J. Agric. Food Chem. 61, 9534–9550. https://doi.org/10.1021/jf402654e Fuentes, A., Ramos, P.L., Fiallo, E., Callard, D., Sánchez, Y., Peral, R., Rodríguez, R., and Pujol, M. (2006). Intronhairpin RNA derived from replication associated protein C1 gene confers immunity to Tomato yellow leaf curl virus infection in transgenic tomato plants. Transgenic Res. 15, 291–304. https://doi.org/10.1007/s11248-0055238-0

250  | Archana and Mukherjee

Fujii, H., Chiou, T.J., Lin, S.I., Aung, K., and Zhu, J.K. (2005). A miRNA involved in phosphate-starvation response in Arabidopsis. Curr. Biol. 15, 2038–2043. Galvez, L.C., Banerjee, J., Pinar, H., and Mitra, A. (2014). Engineered plant virus resistance. Plant Sci. 228, 11–25. https://doi.org/10.1016/j.plantsci.2014.07.006 Garcia, D., Collier, S.A., Byrne, M.E., and Martienssen, R.A. (2006). Specification of leaf polarity in Arabidopsis via the trans-acting siRNA pathway. Curr. Biol. 16, 933–938. Gibbs, A., and Ohshima, K. (2010). Potyviruses and the digital revolution. Annu. Rev. Phytopathol. 48, 205–223. https://doi.org/10.1146/annurevphyto-073009-114404 Glick, E., Levy, Y., and Gafni, Y. (2009). The viral etiology of tomato yellow leaf curl disease-a review. Plant Prot. Sci. 45, 81–97. Glick, E., Zrachya, A., Levy, Y., Mett, A., Gidoni, D., Belausov, E., Citovsky, V., and Gafni, Y. (2008). Interaction with host SGS3 is required for suppression of RNA silencing by Tomato yellow leaf curl virus V2 protein. Proc. Natl. Acad. Sci. U.S.A. 105, 157–161. https://doi.org/10.1073/pnas.0709036105 Gopal, P., Pravin Kumar, P., Sinilal, B., Jose, J., Kasin Yadunandam, A., and Usha, R. (2007). Differential roles of C4 and betaC1 in mediating suppression of post-transcriptional gene silencing: evidence for transactivation by the C2 of Bhendi yellow vein mosaic virus, a monopartite begomovirus. Virus Res. 123, 9–18. Gururani, M.A., Venkatesh, J., Upadhyaya, C.P., Nookaraju, A., Pandey, S.K., and Park, S.W. (2012). Plant disease resistance genes: current status and future directions. Plant J. Plant Mol. Biol, 78, 51–65. Hamilton, A.J., and Baulcombe, D.C. (1999). A species of small antisense RNA in posttranscriptional gene silencing in plants. Science 286, 950–952. Hanley-Bowdoin, L., Bejarano, E.R., Robertson, D., and Mansoor, S. (2013). Geminiviruses: masters at redirecting and reprogramming plant processes. Nat. Rev. Microbiol. 11, 777–788. https://doi.org/10.1038/ nrmicro3117 Hanley-Bowdoin, L., Settlage, S.B., Orozco, B.M., Nagar, S., and Robertson, D. (1999). Geminiviruses: models for plant DNA replication, transcription, and cell cycle regulation. CRC Crit. Rev. Plant Sci. 18, 71–106. Hanssen, I.M., Lapidot, M., and Thomma, B.P. (2010). Emerging viral diseases of tomato crops. Mol. Plant Microbe Interact. 23, 539–548. https://doi. org/10.1094/MPMI-23-5-0539 Hefferon, K.L., Moon, Y.S., and Fan, Y. (2006). Multitasking of nonstructural gene products is required for bean yellow dwarf geminivirus transcriptional regulation. FEBS J. 273, 4482–4494. Hilje, L., Costa, H.S., and Stansly, P.A. (2001). Cultural practices for managing Bemisia tabaci and associated viral diseases. Crop Prot. 20, 801–812. Hong, Y., Saunders, K., Hartley, M.R., and Stanley, J. (1996). Resistance to geminivirus infection by virus-induced expression of dianthin in transgenic plants. Virology 220, 119–127. Huntzinger, E., and Izaurralde, E. (2011). Gene silencing by microRNAs: contributions of translational repression and mRNA decay. Nat. Rev. Genet. 12, 99–110. https:// doi.org/10.1038/nrg2936

Hamilton, A., Voinnet, O., Chappell, L., and Baulcombe, D. (2002). Two classes of short interfering RNA in RNA silencing. EMBO J. 21, 4671–4679. Havecker, E.R., Wallbridge, L.M., Hardcastle, T.J., Bush, M.S., Kelly, K.A., Dunn, R.M., Schwach, F., Doonan, J.H., and Baulcombe, D.C. (2010). The Arabidopsis RNAdirected DNA methylation argonautes functionally diverge based on their expression and interaction with target loci. Plant Cell 22, 321–334. https://doi. org/10.1105/tpc.109.072199 Ji, L., Liu, X., Yan, J., Wang, W., Yumul, R.E., Kim, Y.J., Dinh, T.T., Liu, J., Cui, X., Zheng, B., et al. (2011). ARGONAUTE10 and ARGONAUTE1 regulate the termination of floral stem cells through two microRNAs in Arabidopsis. PLOS Genet. 7, e1001358. https://doi. org/10.1371/journal.pgen.1001358 Jones-Rhoades, M.W., Bartel, D.P., and Bartel, B. (2006). MicroRNAS and their regulatory roles in plants. Annu Rev Plant Biol 57, 19–53. https://doi.org/10.1146/ annurev.arplant.57.032905.105218 Kamthan, A., Chaudhuri, A., Kamthan, M., and Datta, A. (2015). Small RNAs in plants: recent development and application for crop improvement. Front. Plant Sci. 6, 208. https://doi.org/10.3389/fpls.2015.00208 Kennerdell, J.R., and Carthew, R.W. (1998). Use of dsRNAmediated genetic interference to demonstrate that frizzled and frizzled 2 act in the wingless pathway. Cell 95, 1017–1026. Khraiwesh, B., Arif, M.A., Seumel, G.I., Ossowski, S., Weigel, D., Reski, R., and Frank, W. (2010). Transcriptional control of gene expression by microRNAs. Cell 140, 111–122. https://doi.org/10.1016/j.cell.2009.12.023 Krake, L.R., Rezaian, M.A., and Dry, I.B. (1998). Expression of the tomato leaf curl geminivirus C4 gene produces viruslike symptoms in transgenic plants. Mol. Plant Microbe Interact. 11, 413–417. Krenz, B., Deuschle, K., Deigner, T., Unseld, S., Kepp, G., Wege, C., Kleinow, T., and Jeske, H. (2015). Early function of the Abutilon mosaic virus AC2 gene as a replication brake. J. Virol. 89, 3683–3699. https://doi. org/10.1128/JVI.03491-14 Kumar, V., Mishra, S.K., Rahman, J., Taneja, J., Sundaresan, G., Mishra, N.S., and Mukherjee, S.K. (2015). Mungbean yellow mosaic Indian virus encoded AC2 protein suppresses RNA silencing by inhibiting Arabidopsis RDR6 and AGO1 activities. Virology 486, 158–172. https://doi.org/10.1016/j.virol.2015.08.015 Lam, N., Creamer, R., Rascon, J., and Belfon, R. (2009). Characterization of a new Curtovirus, pepper yellow dwarf virus, from chile pepper and distribution in weed hosts in New Mexico. Arch. Virol. 154, 429–436. https://doi.org/10.1007/s00705-009-0320-1 Lapidot, M. (2007). Screening for TYLCV-resistance plants using whitefly-mediated inoculation. In Tomato yellow leaf curl virus Disease, Czosnek, H. (Springer, Netherlands), pp. 329–342. Lazarowitz, S.G., Wu, L.C., Rogers, S.G., and Elmer, J.S. (1992). Sequence-specific interaction with the viral AL1 protein identifies a geminivirus DNA replication origin. Plant Cell 4, 799–809. https://doi.org/10.1105/ tpc.4.7.799 Lefeuvre, P., Martin, D.P., Harkins, G., Lemey, P., Gray, A.J., Meredith, S., Lakay, F., Monjane, A., Lett, J.M.,

Management of Geminiviruses Focusing on Small RNAs in Tomato |  251

Varsani, A., et al. (2010). The spread of Tomato yellow leaf curl virus from the Middle East to the world. PLOS Pathog. 6, e1001164. https://doi.org/10.1371/journal. ppat.1001164 Li, F., Pignatta, D., Bendix, C., Brunkard, J.O., Cohn, M.M., Tung, J., Sun, H., Kumar, P., and Baker, B. (2012). MicroRNA regulation of plant innate immune receptors. Proc. Natl. Acad. Sci. U.S.A. 109, 1790–1795. https:// doi.org/10.1073/pnas.1118282109 Liu, Q., Yao, X., Pi, L., Wang, H., Cui, X., and Huang, H. (2009). The ARGONAUTE10 gene modulates shoot apical meristem maintenance and establishment of leaf polarity by repressing miR165/166 in Arabidopsis. Plant J. 58, 27–40. https://doi.org/10.1111/j.1365313X.2008.03757.x Liu, H., Boulton, M.I., and Davies, J.W. (1997). Maize streak virus coat protein binds single- and double-stranded DNA in vitro. J. Gen. Virol. 78, 1265–1270. https://doi. org/10.1099/0022-1317-78-6-1265 Liu, L., Saunders, K., Thomas, C.L., Davies, J.W., and Stanley, J. (1999). Bean yellow dwarf virus RepA, but not Rep, binds to maize retinoblastoma protein, and the virus tolerates mutations in the consensus binding motif. Virology, 256, 270–279. Liu, P.P., Bhattacharjee, S., Klessig, D.F., and Moffett, P. (2010). Systemic acquired resistance is induced by R gene-mediated responses independent of cell death. Mol. Plant Pathol. 11, 155–160. https://doi. org/10.1111/j.1364-3703.2009.00564.x Lucioli, A., Sallustio, D.E., Barboni, D., Berardi, A., Papacchioli, V., Tavazza, R., and Tavazza, M. (2008). A cautionary note on pathogen-derived sequences. Nat. Biotechnol. 26, 617–619. https://doi.org/10.1038/ nbt0608-617 Mansoor, S., Khan, S.H., Bashir, A., Saeed, M., Zafar, Y., Malik, K.A., Briddon, R., Stanley, J., and Markham, P.G. (1999). Identification of a novel circular single-stranded DNA associated with cotton leaf curl disease in Pakistan. Virology 259, 190–199. https://doi.org/10.1006/ viro.1999.9766 Mejia, L., Teni, R.E., García, B.E., Fulladolsa, A.C., Méndez, L., Melgar, S., and Maxwell, D.P. (2010). Preliminary observations on the effectiveness of five introgressions for resistance to begomoviruses in tomato. TGC Report 60, 41–53. Mi, S., Cai, T., Hu, Y., Chen, Y., Hodges, E., Ni, F., Wu, L., Li, S., Zhou, H., Long, C., et al. (2008). Sorting of small RNAs into Arabidopsis argonaute complexes is directed by the 5’ terminal nucleotide. Cell 133, 116–127. https://doi.org/10.1016/j.cell.2008.02.034 Montgomery, T.A., Howell, M.D., Cuperus, J.T., Li, D., Hansen, J.E., Alexander, A.L., Chapman, E.J., Fahlgren, N., Allen, E., and Carrington, J.C. (2008). Specificity of ARGONAUTE7-miR390 interaction and dual functionality in TAS3 trans-acting siRNA formation. Cell 133, 128–141. https://doi.org/10.1016/j. cell.2008.02.033 Montgomery, T.A., Yoo, S.J., Fahlgren, N., Gilbert, S.D., Howell, M.D., Sullivan, C.M., Alexander, A., Nguyen, G., Allen, E., Ahn, J.H., et al. (2008). AGO1-miR173 complex initiates phased siRNA formation in plants. Proc. Natl. Acad. Sci. U.S.A. 105, 20055–20062. https:// doi.org/10.1073/pnas.0810241105

Nahid, N., Amin, I., Mansoor, S., Rybicki, E.P., van der Walt, E., and Briddon, R.W. (2008). Two dicot-infecting Mastreviruses (family Geminiviridae) occur in Pakistan. Arch. Virol. 153, 1441–1451. https://doi.org/10.1007/ s00705-008-0133-7 Nakhla, M.K., and Maxwell, D.P. (1998). Epidemiology and management of tomato yellow leaf curl disease. In Hadidi, A., Khetarpal, R.K., and Koganezawa, H., eds., Plant Virus Disease Control. (APS Press, Minnesota), pp. 565–583. Napoli, C., Lemieux, C., and Jorgensen, R. (1990). Introduction of a Chimeric Chalcone Synthase Gene into Petunia Results in Reversible Co-Suppression of Homologous Genes in trans. Plant Cell 2, 279–289. https://doi.org/10.1105/tpc.2.4.279 Navarro, L., Dunoyer, P., Jay, F., Arnold, B., Dharmasiri, N., Estelle, M., Voinnet, O., and Jones, J.D. (2006). A plant miRNA contributes to antibacterial resistance by repressing auxin signaling. Science 312, 436–439. Niu, Q.W., Lin, S.S., Reyes, J.L., Chen, K.C., Wu, H.W., Yeh, S.D., and Chua, N.H. (2006). Expression of artificial microRNAs in transgenic Arabidopsis thaliana confers virus resistance. Nat. Biotechnol. 24, 1420–1428. Noris, E., Accotto, G.P., Tavazza, R., Brunetti, A., Crespi, S., and Tavazza, M. (1996). Resistance to tomato yellow leaf curl geminivirus in Nicotiana benthamiana plants transformed with a truncated viral C1 gene. Virology 224, 130–138. Noueiry, A.O., Lucas, W.J., and Gilbertson, R.L. (1994). Two proteins of a plant DNA virus coordinate nuclear and plasmodesmal transport. Cell 76, 925–932. Oelgeschläger, M., Larraín, J., Geissert, D., and De Robertis, E.M. (2000). The evolutionarily conserved BMP-binding protein Twisted gastrulation promotes BMP signalling. Nature 405, 757–763. https://doi. org/10.1038/35015500 Palumbo, J.C., Horowitz, A.R., and Prabhaker, N. (2001). Insecticidal control and resistance management for Bemisia tabaci. Crop Prot. 20, 739–765. Pant, V., Gupta, D., Choudhury, N.R., Malathi, V.G., Varma, A., and Mukherjee, S.K. (2001). Molecular characterization of the Rep protein of the blackgram isolate of Indian mungbean yellow mosaic virus. J. Gen. Virol. 82, 2559–2567. Paprotka, T., Metzler, V., and Jeske, H. (2010). The first DNA 1-like alpha satellites in association with New World begomoviruses in natural infections. Virology 404, 148–157. https://doi.org/10.1016/j.virol.2010.05.003 Picó, B., Díez, M.J., and Nuez, F. (1996). Viral diseases causing the greatest economic losses to the tomato crop. II. The Tomato yellow leaf curl virus—a review. Sci. Hort. 67, 151–196. Polston, J.E., and Anderson, P.K. (1997). The emergence of whitefly-transmitted geminiviruses in tomato in the western hemisphere. Plant Dis. 81, 1358–1369. Pooma, W., and Petty, I.T. (1996). Tomato golden mosaic virus open reading frame AL4 is genetically distinct from its C4 analogue in monopartite geminiviruses. J. Gen. Virol. 77, 1947–1951. https://doi.org/10.1099/00221317-77-8-1947 Rajagopalan, R., Vaucheret, H., Trejo, J., and Bartel, D.P. (2006). A diverse and evolutionarily fluid set of microRNAs in Arabidopsis thaliana. Genes Dev. 20, 3407–3425.

252  | Archana and Mukherjee

Ramachandran, V., and Chen, X. (2008). Degradation of microRNAs by a family of exoribonucleases in Arabidopsis. Science 321, 1490–1492. https://doi. org/10.1126/science.1163728 Revington, G.N., Sunter, G., and Bisaro, D.M. (1989). DNA sequences essential for replication of the B genome component of Tomato golden mosaic virus. Plant Cell 1, 985–992. https://doi.org/10.1105/tpc.1.10.985 Reyes, M.I., Nash, T.E., Dallas, M.M., Ascencio-Ibáñez, J.T., and Hanley-Bowdoin, L. (2013). Peptide aptamers that bind to geminivirus replication proteins confer a resistance phenotype to Tomato yellow leaf curl virus and tomato mottle virus infection in tomato. J. Virol. 87, 9691–9706. https://doi.org/10.1128/JVI.01095-13 Ribeiro, S.G., Ambrozevícius, L.P., Avila, A.C., Bezerra, I.C., Calegario, R.F., Fernandes, J.J., Lima, M.F., de Mello, R.N., Rocha, H., and Zerbini, F.M. (2003). Distribution and genetic diversity of tomato-infecting begomoviruses in Brazil. Arch. Virol. 148, 281–295. https://doi. org/10.1007/s00705-002-0917-0 Rigden, J.E., Dry, I.B., Mullineaux, P.M., and Rezaian, M.A. (1993). Mutagenesis of the virion-sense open reading frames of tomato leaf curl geminivirus. Virology 193, 1001–1005. Rodríguez-Negrete, E., Lozano-Durán, R., Piedra-Aguilera, A., Cruzado, L., Bejarano, E.R., and Castillo, A.G. (2013). Geminivirus Rep protein interferes with the plant DNA methylation machinery and suppresses transcriptional gene silencing. New Phytol. 199, 464–475. https://doi. org/10.1111/nph.12286 Romay, G., Chirinos, D., Geraud-Pouey, F., and Desbiez, C. (2010). Association of an atypical alphasatellite with a bipartite New World begomovirus. Arch. Virol. 155, 1843–1847. https://doi.org/10.1007/s00705-0100760-7 Roumagnac, P., Granier, M., Bernardo, P., Deshoux, M., Ferdinand, R., Galzi, S., Fernandez, E., Julian, C., Abt, I., Filloux, D., et al. (2015). Alfalfa leaf curl virus: an AphidTransmitted Geminivirus. J. Virol. 89, 9683–9688. https://doi.org/10.1128/JVI.00453-15 Sánchez-Durán, M.A., Dallas, M.B., Ascencio-Ibañez, J.T., Reyes, M.I., Arroyo-Mateos, M., Ruiz-Albert, J., Hanley-Bowdoin, L., and Bejarano, E.R. (2011). Interaction between geminivirus replication protein and the SUMO-conjugating enzyme is required for viral infection. J. Virol. 85, 9789–9800. https://doi. org/10.1128/JVI.02566-10 Saunders, K., Bedford, I.D., and Stanley, J. (2002). Adaptation from whitefly to leafhopper transmission of an autonomously replicating nanovirus-like DNA component associated with ageratum yellow vein disease. J. Gen. Virol. 83, 907–913. Scholthof, K.B., Adkins, S., Czosnek, H., Palukaitis, P., Jacquot, E., Hohn, T., Hohn, B., Saunders, K., Candresse, T., Ahlquist, P., et al. (2011). Top 10 plant viruses in molecular plant pathology. Mol. Plant Pathol. 12, 938–954. https://doi.org/10.1111/j.13643703.2011.00752.x Schwab, R., Ossowski, S., Riester, M., Warthmann, N., and Weigel, D. (2006). Highly specific gene silencing by artificial microRNAs in Arabidopsis. Plant Cell 18, 1121–1133.

Sera, T. (2005). Inhibition of virus DNA replication by artificial zinc finger proteins. J. Virol. 79, 2614–2619. Settlage, S.B., See, R.G., and Hanley-Bowdoin, L. (2005). Geminivirus C3 protein: replication enhancement and protein interactions. J. Virol. 79, 9885–9895. Sharma, P., Ikegami, M., and Kon, T. (2010). Identification of the virulence factors and suppressors of posttranscriptional gene silencing encoded by Ageratum yellow vein virus, a monopartite begomovirus. Virus Res. 149, 19–27. https://doi.org/10.1016/j. virusres.2009.12.008 Shivaprasad, P.V., Akbergenov, R., Trinks, D., Rajeswaran, R., Veluthambi, K., Hohn, T., and Pooggin, M.M. (2005). Promoters, transcripts, and regulatory proteins of Mungbean yellow mosaic geminivirus. J. Virol. 79, 8149–8163. Shivaprasad, P.V., Chen, H.M., Patel, K., Bond, D.M., Santos, B.A., and Baulcombe, D.C. (2012). A microRNA superfamily regulates nucleotide binding site-leucinerich repeats and other mRNAs. Plant Cell 24, 859–874. https://doi.org/10.1105/tpc.111.095380 Singh, A., Taneja, J., Dasgupta, I., and Mukherjee, S.K. (2015). Development of plants resistant to tomato geminiviruses using artificial trans-acting small interfering RNA. Mol. Plant Pathol. 16, 724–734. https://doi.org/10.1111/mpp.12229 Singh, A., Saraf, S., Dasgupta, I., and Mukherjee, S.K. (2016). Identification and validation of a virus-inducible ta-siRNA-generating TAS4 locus in tomato. J. Biosci. 41, 109–118. Sunkar, R., Li, Y.F., and Jagadeeswaran, G. (2012). Functions of microRNAs in plant stress responses. Trends Plant Sci. 17, 196–203. https://doi.org/10.1016/j. tplants.2012.01.010 Svoboda, P., Stein, P., Hayashi, H., and Schultz, R.M. (2000). Selective reduction of dormant maternal mRNAs in mouse oocytes by RNA interference. Development 127, 4147–4156. Thomson, J.A., Mundree, S.G., Shepherd, D.M., and Rybicki, E.P. (2014). The use of African indigenous genes in the development of transgenic maize tolerant to drought and resistant to maize streak virus. In Biotechnology in Africa, Wambugu, F., and Kamanga, D., eds. (Springer International Publishing, Switzerland), pp. 135–155. Tollefson, J. (2011). Brazil cooks up transgenic bean. Nature 478, 168. https://doi.org/10.1038/478168a Vu, T.V., Choudhury, N.R., and Mukherjee, S.K. (2013). Transgenic tomato plants expressing artificial microRNAs for silencing the pre-coat and coat proteins of a begomovirus, Tomato leaf curl New Delhi virus, show tolerance to virus infection. Virus Res. 172, 35–45. https://doi.org/10.1016/j.virusres.2012.12.008 Vasudeva, R.S., and Sam Raj, J. (1948). A leaf-curl disease of tomato. Phytopathology 38, 364–369. Varsani, A., Navas-Castillo, J., Moriones, E., HernándezZepeda, C., Idris, A., Brown, J.K., Murilo Zerbini, F., and Martin, D.P. (2014). Establishment of three new genera in the family Geminiviridae: Becurtovirus, Eragrovirus and Turncurtovirus. Arch. Virol. 159, 2193–2203. https:// doi.org/10.1007/s00705-014-2050-2 Vaucheret, H., and Fagard, M. (2001). Transcriptional gene silencing in plants: targets, inducers and regulators. Trends Genet. 17, 29–35.

Management of Geminiviruses Focusing on Small RNAs in Tomato |  253

Verlaan, M.G., Hutton, S.F., Ibrahem, R.M., Kormelink, R., Visser, R.G., Scott, J.W., Edwards, J.D., and Bai, Y. (2013). The Tomato yellow leaf curl virus resistance genes Ty-1 and Ty-3 are allelic and code for DFDGDclass RNA-dependent RNA polymerases. PLOS Genet. 9, e1003399. https://doi.org/10.1371/journal. pgen.1003399 Wianny, F., and Zernicka-Goetz, M. (2000). Specific interference with gene function by double-stranded RNA in early mouse development. Nat. Cell Biol. 2, 70–75. https://doi.org/10.1038/35000016 Xie, Y., Wu, P., Liu, P., Gong, H., and Zhou, X. (2010). Characterization of alphasatellites associated with monopartite begomovirus/betasatellite complexes in Yunnan, China. Virol. J. 7, 178. https://doi. org/10.1186/1743-422X-7-178 Yadava, P., Suyal, G., and Mukherjee, S.K. (2010). Begomovirus DNA replication and pathogenicity. Curr. Sci. 98. Yadava, P., and Mukherjee, S.K. (2012). Artificial microRNA and its applications. In Regulatory RNAs, Bibekanand, M., and Zhumer, G., eds (Springer, Berlin) pp. 505–521. Yang, X., Caro, M., Hutton, S.F., Scott, J.W., Guo, Y., Wang, X., Rashid, M.H., Szinay, D., de Jong, H., Visser, R.G., et al. (2014). Fine mapping of the Tomato yellow leaf curl virus resistance gene Ty-2 on chromosome 11 of tomato. Mol. Breed. 34, 749–760. https://doi.org/10.1007/ s11032-014-0072-9 Yang, X., Xie, Y., Raja, P., Li, S., Wolf, J.N., Shen, Q., Bisaro, D.M., and Zhou, X. (2011). Suppression of methylationmediated transcriptional gene silencing by βC1-SAHH protein interaction during geminivirus-betasatellite

infection. PLOS Pathog. 7, e1002329. https://doi. org/10.1371/journal.ppat.1002329 Yoshikawa, M., Peragine, A., Park, M.Y., and Poethig, R.S. (2005). A pathway for the biogenesis of trans-acting siRNAs in Arabidopsis. Genes Dev. 19, 2164–2175. Zaffalon, V., Mukherjee, S.K., Reddy, V.S., Thompson, J.R., and Tepfer, M. (2012). A survey of geminiviruses and associated satellite DNAs in the cotton-growing areas of northwestern India. Arch. Virol. 157, 483–495. https:// doi.org/10.1007/s00705-011-1201-y Zhai, J., Jeong, D.H., De Paoli, E., Park, S., Rosen, B.D., Li, Y., González, A.J., Yan, Z., Kitto, S.L., Grusak, M.A., et al. (2011). MicroRNAs as master regulators of the plant NB-LRR defense gene family via the production of phased, trans-acting siRNAs. Genes Dev. 25, 2540– 2553. https://doi.org/10.1101/gad.177527.111 Zhang, P., Vanderschuren, H., Fütterer, J., and Gruissem, W. (2005). Resistance to cassava mosaic disease in transgenic cassava expressing antisense RNAs targeting virus replication genes. Plant Biotechnol. J. 3, 385–397. Zhu, H., Hu, F., Wang, R., Zhou, X., Sze, S.H., Liou, L.W., Barefoot, A., Dickman, M., and Zhang, X. (2011). Arabidopsis Argonaute10 specifically sequesters miR166/165 to regulate shoot apical meristem development. Cell 145, 242–256. https://doi. org/10.1016/j.cell.2011.03.024 Zrachya, A., Kumar, P.P., Ramakrishnan, U., Levy, Y., Loyter, A., Arazi, T., Lapidot, M., and Gafni, Y. (2007). Production of siRNA targeted against TYLCV coat protein transcripts leads to silencing of its expression and resistance to the virus. Transgenic Res. 16, 385–398. https://doi.org/10.1007/s11248-006-9042-2

Viruses Infecting Banana and their Transgenic Management Ramasamy Selvarajan*, Chelliah Anuradha, Velusamy Balasubramanian, Sivalingam Elayabalan and Kanicheluam Prasanya Selvam

13

Molecular Virology Lab, ICAR-National Research Centre for Banana, Tiruchirapalli, India. *Correspondence: [email protected] https://doi.org/10.21775/9781910190814.13

Abstract Viral pathogens cause serious diseases in banana and plantains leading to a greater loss on the production and productivity. Banana bunchy top disease, banana streak, banana bract mosaic and infectious chlorosis or banana mosaic pose serious concern in banana cultivation across the world. BBTV infection can result into 100% yield loss whereas other three diseases cause ca. 9 −70% yield loss. Breeding for virus disease resistance is not attempted probably owing to lack of resistance source, inherent problem in Musa due to sterility, parthenocarpic nature and compatibility. At present the management is mainly through timely identification of the diseased plants and eradication, and supply of certified virus free quality planting material to the farmers. Transgenic approaches have been attempted in a few labs across the world and came out with promising results for BBTV resistance but for other viruses attempts were not yet made. Transgenic resistance lines using RNAi has been developed for BBTV successfully but field level release and deployment in the farmers field is yet to take place. Multiple virus resistance targeting the suppressor of gene silencing with RNAi strategy is being attempted at ICAR-National Research Centre on Banana (ICAR-NRCB). CRISPR-Cas9 approach of genome editing for resistance and removal of eBSVs are the future goals of banana virologists to achieve the better management through molecular approaches. In this paper, we

have reviewed and discussed about the important viruses of banana and transgenic management of them. Introduction Banana (Musa spp.) and plantains are the most important fruit crops of tropical and subtropical parts of the world. These fruit crops are cultivated in an area covering about 10.3 million ha, and annually 139 million tonnes are produced with an average productivity of 13.4 million tonnes/ha in the world (FAOSTAT, 2014). Banana yield and productivity are subjected to several biotic and abiotic stress factors. Among biotic stresses, viruses cause the most significant loss on production and productivity. They can also have important indirect effects by restricting germplasm movement and by predisposing plants to damage by other biotic and abiotic stress factors. Currently, there are four major viral diseases: banana bunchy top disease, banana streak, banana bract mosaic and infectious chlorosis or banana mosaic posing causing very fundamental problem in banana cultivation across the world. These viruses are of different size and shapes having DNA and RNA as their genome and the brief characters and their vector and detection methods are furnished in Table 13.1. Although conventional method of eradication and supply of certified virusfree stock have successfully been used to reduce crop losses caused by these viruses, they were not

Table 13.1  Important banana infecting viruses with their taxonomy and shape, size, genome, vector and detection techniques Virus species

Size of virus particle (nm)

Family

Genus

Shape

BBTV

Nanoviridae

Babuvirus

Icosahedral 18–20

BBrMV

Potyviridae

Potyvirus

Flexuous 750  filamentous

BSOLV

Caulimoviridae

Badnavirus

Bacilliform

Size (bp) of genome or its Genome segments ssDNA

11 ssRNA

130–150 × 30 dsDNA

Banana black aphid-Pentalonia nigronervosa (Coquerel)

ELISA, PCR,MPCR,IC-PCR,NASH, Q-PCR,LAMP,RCA and RPA

9711

Aphids- Pentanlonia nigronervosa, Rhopalosiphummaidis, Aphis gossypii, A. craccivora

ELISA, RT-PCR, IC-RTPCR, M-RTPCR,NASH, Q-PCR,RTLAMP and IC-RTLAMP

6950;7389

Mealybugs-Planococcuscitri, Pseudococcus spp, Dysmicoccus spp, Planococcusmusa, Ferrisiavirgata, D. brevipes, P. ficus, Paracoccus burnerae

ELISA, IC-PCR,MRTPCR,NASH, LAMP and RCA

A. gossypii, A. craccivora, R. maidis, R. prunifolium, Myzuspersicae

ELISA,RT-PCR, IC-RTPCR, M-RTPCR, NASH, Q-PCR,RTLAMP

7650

BSGFV

7263

BSVNV

7801

BSAcYNV

7722

BSCAV

7408

BSIMV

7769

BSUAV

7519

BSUIV

7458

BSULV

7401

CMV

Detection techniques reported

DNA-R- 1111 DNA-U-1064 DNA-S-1074 DNA-M-1046 DNA-N-1087 DNA-C-1018

BSMYV

BSUMV

Vector

7532 Bromoviridae

Cucumovirus

Isometric

28–30

Three, positive sense ssRNAs

RNA1–3358 RNA2–2982 RNA3–2219

ELISA, enzyme linked immunosorbent assay; IC-PCR, immunocapture PCR; LAMP, loop-mediated isothermal amplification; M-PCR, multiplex-PCR; NASH, nucleic acid spot hybridization; PCR, polymerase chain reaction; Q-PCR, quantitative PCR; RCA, rolling circle amplification; RPA, recombinase polymerase amplification; RT-LAMP, reverse transcription-LAMP; RTPCR, reverse transcription PCR.

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sufficient enough for controlling banana viruses. As no genes have been identified within the Musa gene pool that confers resistance to banana viruses, the conventional breeding approach has never been attempted and it may be a difficult option for banana crop due to their polyploidy, sterility and parthenocarpic nature. Genetic transformation, in conjunction with pathogen-derived resistance is one of the potential strategies for developing virus resistance in bananas, which has proven to be difficult to obtain by conventional breeding. In transgenics, there are two approaches based on the source of the genes used. The genes can be either from the pathogenic virus itself where a complete or partial viral gene is introduced into the plant, which, subsequently, interferes with one or more essential steps in the life cycle of the virus based on the concept of pathogen-derived resistance (PDR). The second approach is the non-pathogen-derived resistance, which is based on utilizing host resistance genes and other genes responsible for adaptive host processes, elicited in response to pathogen attack, to obtain transgenics resistant to the virus. Recently CRISPR (Clustered Regularly Inter-Spaced Palindromic Repeats)-Cas9 approach is being followed to obtain resistance for viral diseases in plants. Banana bunchy top virus Banana bunchy top disease (BBTD) caused by Banana bunchy top virus (BBTV) has become a major threat to banana and plantain cultivation in several tropical and sub tropical regions of the world including India and causes significant yield losses (Dale, 1987; Selvarajan and Balasubramanian, 2013a; Kumar et al., 2015). The Invasive Species Specialist Group (ISSG) of the International Union for the Conservation of Nature (IUCN) listed BBTV in the World’s 100 Worst Invasive Alien Species. Though BBTD is present in number of countries in Africa, Asia and the south Pacific but its geographical distribution is erratic within regions and as on date it is not reported in Central and South America. BBTV infection can result into 100% yield loss (Dale, 1987). It was extrapolated that BBTV caused an annual loss of production worth US$50 million in 2009–2010 probably due to inadvertent spread of virus through tissue culture plants which were not tested and certified in India (Selvarajan and Balasubramanian, 2014).

BBTV is a multi-component, circular singlestranded DNA virus belonging to the genus Babuvirus and family Nanoviridae. The genome of BBTV consists of at least six integral components each approximately 1 kb in size is individually encapsidated within icosahedral virions 18–20 nm in diameter (Thomas and Dietzgen, 1991; Wu and Su, 1990a; Burns et al., 1995). Each genome segment encode for a single open reading frame (ORF) for different functional proteins from the virion sense strand such as master replication initiation protein (DNA R) (Beetham et al., 1997), a protein with unknown function (DNA U3) (Beetham et al., 1999), a capsid protein (DNA S) (Wanitchakorn et al., 1997, 2000a), a movement protein (DNA M) (Wanitchakorn et al., 2000b), a cell cycle link protein (DNA C) (Wanitchakorn et al., 2000b), and a nuclear shuttle protein (DNA N) (Wanitchakorn et al., 2000b). All the components contain a highly conserved, major common region (CR-M) and a stem–loop common region (CR-SL) downstream of CR-M (Burns et al., 1995). Characteristic symptoms BBTV infected plants shows characteristic discontinuous dark green flecks and streaks of variable length on the leaf sheath, midrib, leaf veins, and petioles (Fig. 13.1A and B). New leaves that are produced after infection are progressively shorter, narrow, brittle in texture, display marginal yellowing or chlorosis and bunched at the top, hence the name ‘bunchy top’ disease (Thomas et al., 1994). BBTV infected plants usually fail to produce bunch. However, in late infections the plant may produce bunch but the fingers are malformed (Nelson, 2004). Bracts of male flower buds may sometimes turn to leafy structure with dark green dots and streaks (Thomas et al., 1994). Side suckers emerging from infected plants show severe symptoms. Transmission and host range BBTV primarily spreads through the infected planting material such as corms, suckers and tissue culture plants (Drew et al., 1989) and secondarily transmitted by the banana black aphid, Pentalonia nigronervosa (Coquerel) in a persistent circulative but non-replicative manner (Anhalt and Almeida, 2008). P. caladii van der Goot has also been shown to transmit BBTV under experimental inoculation conditions with low efficiency (Watanabe et al., 2013). The only confirmed hosts of BBTV

258  | Selvarajan et al.

Figure 13.1 (A) A clump of banana plant infected with Banana bunchy top virus exhibiting typical stunted growth and narrow leaves with chlorosis on leaf margins; (B) short dark green morse code symptoms on the midribs and petiole, a characteristic symptoms of BBTV; (C) reddish discolouration and mosaic symptom of banana bract mosaic virus disease on the pseudostem; (D) typical spindle shaped reddish coloured mosaic on bracts of banana emerging flower; (E), Golden yellowish streaks parallel to veins of banana leaf due to BSMYV infection in Mysore bananas; (F) cigar leaf showing chlorotic streaks due to BSMYV; (G) line pattern, spindle or diamond shaped mosaic pattern along the veins of leaf blade due to CMV infection in banana; (H) a young banana plant showing mosaic symptoms in parts of leaf blade with the infection of CMV.

Viruses Infecting Banana and their Transgenic Management |  259

are species within the genus Musa (M. acuminata, M. balbisiana, M. sinensis, M. paradisica and their hybrids) (Manser, 1982; Magee, 1948), abaca (M. textilis) (Manila hemp) (Sharman et al., 2008) and Ensete ventricosum (Selvarajan and Balasubramanian, 2013a). Molecular characterization of Banana bunchy top virus Characterization of BBTV isolates from different parts of the world has revealed a homology of >85% (Banerjee et al., 2014). Genetic diversity studies from India, Pakistan, Africa, Democratic Republic of Congo (DRC), Indonesiaand Oceania shows very low diversity within a country (Selvarajan et al., 2010a; Vishnoi et al., 2009; Amin et al., 2008; Kumar et al., 2011; Stainton et al., 2012; 2015; Chiaki et al., 2015; Mukwa et al., 2016). However, greater diversity of BBTV was observed in the northeastern region of India (Banerjee et al., 2014) and China (Yu et al., 2012). Recently, two groups of BBTV were identified: the Pacific-Indian Oceans (PIO) group (comprising the isolates from Australia, Egypt, Hawaii, India, Myanmar, Pakistan, Sri Lanka, DRC and Tonga) and the South-East Asian (SEA) group (comprising the isolates from China, Indonesia, Japan, Philippines, Taiwan and Vietnam) (Fig. 13.2) (Yu et al., 2012; Banerjee et al., 2014; Wickramaarachchi et al., 2016). Detection of Banana bunchy top virus To detect BBTV in field-grown plants, TC plants and viruliferous aphids, various forms of enzyme linked immunosorbent assay (ELISA) using monoclonal and polyclonal antibodies have been adopted (Thomas and Dietzgen, 1991; Wu and Su, 1990b; Geering and Thomas, 1996; Selvarajan et al., 2010b). Polymerase chain reaction-based (PCR) detection protocol has been developed for the sensitive detection of BBTV (Selvarajan et al., 2008; Selvarajan et al., 2010b; Xie and Hu, 1995; Mansoor et al., 2005; Selvarajan and Balasubramanian, 2008) or by combining immuno-PCR techniques (Selvarajan and Balasubramanian, 2008; Sharman et al., 2000a; Bressan and Watanabe, 2011). Multiplex PCR approaches developed using two or more sets of gene-specific or component specific primers (Chandrasekar et al., 2011; Sasireka and Selvarajan, 2014). Nucleic acid spot hybridization (NASH)

using radioactive or non-radioactive labelled DNA probes have been applied for BBTV (Xie and Hu, 1995; Selvarajan and Balasubramanian, 2008). Highly sensitive quantitative PCR assay has been reported for detection of BBTV in both plants and aphid tissues with SYBR green (Watanabe and Bressan, 2013; Selvarajan et al., 2015) and TaqManTM chemistries (Bressan and Watanabe, 2011; Chen and Hu, 2013). Peng et al. (2012) reported an isothermal DNA amplification method, i.e. loopmediated isothermal amplification (LAMP) and Stainton et al. (2012) used a rolling circle amplification (RCA), another isothermal amplification technique which targets only the circular genome of DNA. LAMP products can be detected using either conventional agarose gel electrophoresis or by visual observation of turbidity/colour changes using dyes (Peng et al., 2012). LAMP assay is reported to be 100-fold more sensitive than PCR, rapid and simpler, convenient for quick in field or on-site diagnosis in areas where molecular laboratory facilities are not available. Recently a novel Recombinase Polymerase Amplification has been developed for the detection of BBTV (Kapoor et al., 2017). Non-transgenic management and exploiting the virus for management of fungal diseases Management of viral diseases of banana is mainly through timely identification of the diseased plants through visual or applying molecular diagnostic tools and eradication of infected plants would be the primary method of management. Supply of virus free certified planting material is one of the best methods to manage the disease spread. Effective way of controlling this disease is by using disease resistant cultivars. Till date resistant source has not been reported against this virus. Mutation breeding using gamma-irradiation has also been exploited to develop banana variants for BBTV resistance (Damasco et al., 2006; Dizon et al., 2012; Abustan, 2012). However, mutants may lead to more aberrations, owing to this limitation; generation of stable transgenic resistant lines is the best viable way to combat the disease. BBTV infected plants were found to be antagonistic to Fusarium oxysporum f. sp. cubense (Foc) there by offering more resistant towards Foc and thus improving the survivability of plants against blight. B4 protein

260  | Selvarajan et al. KM607603:RC JQ820453:Malawi KM607649:DRC KM607660:US-Hawaii AB252641:Myanmar DQ656118:India-Kanpur JQ820459:Rwanda AF416467:Tonga KP876489:India-Arunachal Pradesh HQ259074:Egypt JN243751:India-Bangalore KP876497:India-Assam EU140342:India-Tamilnadu NC003479:Australia FJ605506:India-Bihar AF416466:Fiji DQ656119:India-Etawah JN250593:Srilanka KP876493:India-Manipur

Pacific and Indian Ocean Group

KM607602:Burundi

AY996562:Pakisthan KM607673:Samoa KR350591:India-Tripura KC119098:India-Umiam

98

9 9 JQ911667:India-Meghalaya KP876496:India-Mizoram HQ616074:Haikou AF246123:China KM607593:Indonesia 90 66

AB108458:Japan AB189067:Philippines

South East Asia Group

AB113659:Vietnam 69

AF416468:Taiwan

JX867550:CBDV EF546813:ABTV

Out Group

0 .0 2

Figure 13.2  Phylogenetic analysis based on the 861 base pairs of nucleotide sequence of BBTV DNA-R master replication protein of 33 BBTV isolates originated from different countries. Places of origin are preceded by NCBI GenBank accession numbers. Bootstrap analysis was applied using 1000 replicates. The phylogenetic tree was generated using maximum likelihood method in MEGA6 (Tamura et al., 2013). Abaca bunchy top virus (ABTV) and Cardamom bushy dwarf virus (CBDV) are included as out-group species.

Viruses Infecting Banana and their Transgenic Management |  261

which is the suppressor of RNA silencing showed fungicidal properties in vitro (Zhuang et al., 2016). This viral–fungal interaction studies throw light on these organisms dynamics within banana and this can be further exploited for transgenic resistance to Fusarium-related blight/wilt in other crops. Banana streak viruses Banana streak disease (BSD) caused by a cryptic virus species complex known as Banana streak viruses (BSVs) are becoming a major threat to banana cultivation worldwide (Hull et al., 2000; James et al., 2011). BSD has become a major threat to production, international exchange and crop improvement programmes. Estimated yield losses range from 7 to 90% in various banana cultivars across the world (Harper et al., 2004; Lockhart et al., 1998; Davis et al., 2000; Daniells et al., 2001; Selvarajan et al., 2011a). BSVs are plant pararetroviruses belonging to the genus Badnavirus and family Caulimoviridae having non-enveloped bacilliform virus particles of 30 × 130–150 nm with a doublestranded non-covalently closed circular DNA (dsDNA) genome approximately 6.9–7.8 kb long which uses a virus encoded reverse transcriptase (RT) to replicate (King et al., 2012). BSV genome has three consecutive ORFs on one strand (King et al., 2012). ORF I potentially encode a protein of unknown function of 20.8 kDa and ORF II encodes a viral-associated protein (VAP) of 14.5 kDa. ORF III encodes a multifunction large polyprotein of 220 kDa consists of a putative cell-to-cell movement protein (MP), the coat protein (CP), an aspartic protease, and the viral replicase consisting of RT and ribonuclease H (RNase H) function (Harper and Hull, 1998; Geering et al., 2005; King et al., 2012). This polyprotein is likely to be cleaved into functional units by the aspartic protease following full-length translation. BSV naturally exists in two states: an episomal form that infects plant cells and integrated copies of BSV named endogenous BSV (eBSV) are integrated within the banana B genome (Musa balbisiana) millions of years ago and the integrated BSV genomes that can release infectious pararetrovirus (Chabannes et al., 2013) and cause spontaneous infection. BSVspecies are proved to have integration of their genome into the banana host especially B genome of Musa and cause disease by expressing

the episomal virus spontaneously through two homologous recombination. Till date three species of endogenous banana infecting badna viruses (BIBs) can get activated, resulting in the infective episomal form (Chabannes et al., 2013) which has been attributed to recent outbreaks. BSV particles from both origins can be transmitted by mealybugs in semipersistent manner. Currently, BSD due to eBSV is one of the important constraints to plant breeding, international movement ofbanana germplasm, genetic improvement and also for mass propagation. Characteristic symptoms of banana streak disease The most common symptoms of banana streak disease include narrow, discontinuous or continuous golden yellow chlorotic and/or necrotic streaks (Fig. 13.1E and F) which are parallel to the veins of leaf lamina (Lockhart and Jones, 2000). The leaf streaks become darker as the leaves age, eventually turns to dark brown or black, hence necrotic streaks. Temperature is considered to influence disease expression of symptoms and the typical symptoms can regress with periods of symptomless growth and vice versa (Harper and Hull, 1998; Dahal et al., 1998; Dahal et al., 2000). Other symptoms include cigar leaf necrosis, internal necrosis of the pseudostem, leaf/fruit distortion, pseudostem splitting, seediness in fruits, choking of bunches and abnormal bunch development (Lockhart and Jones, 2000; Dahal et al., 2000), Though the symptomatic and symptomless stages exists during the growth of infected plants, the virus could be detected at all stages (Lockhart and Jones, 2000; Harper et al., 2002). Transmission and host range BSV is primarily transmitted through the use of infected planting materials and it is spread in the field by mealybugs in a semi-persistent-mediated transmission. BSVs are not mechanically transmitted. Two main mealybug species reported to transmit BSV are Planococcus citri and Pseudococcus spp. (Dahal et al., 2000; Kubiriba et al., 2001). Other species reported to transmit BSVs are Dysmicoccus spp. in West Africa and South America, Planococcus musa in Nigeria, Ferrisia virgata (striped mealybug) in India (Selvarajan et al., 2006), D. brevipes and P. ficus (Meyer et al., 2008). BSV is also transmitted

262  | Selvarajan et al.

to new hybrids through the activation of integrated viral sequences present in the genomes of the parents having B genome (Harper et al., 1999a). Such activation may occur during tissue culture process adopted for propagation. BSMYV severely infects bananas of Mysore subgroup (AAB), Cavendish (AAA) subgroup and Musa zebrina. Many wild bananas also found to have BSV genome as integrants in their genomes but they are not known to gives rise to episomal virus needed for infection. A high degree of heterogeneity exists among isolates of BSV and they differ serologically, genomically and biologically (Lockhart and Olszewski, 1993; Geering et al., 2000). This great variability found in BSVs possibly hinders the development of stable transgenic difficult for obtaining resistance. Since the principal method of disease spread is by vegetative propagation, disease control must be based on the use of virus-free stock plants for propagation by suckers or in vitro plantlets. It is also important to avoid the introduction of BSV into banana breeding lines. De novo infection of BSV from its genome integrated in the host during hybridization and tissue culture processes pose a great problem, hence the detection of episomal BSV genome or activatable eBSVs is therefore the key to the control of these viruses. Molecular characterization The genomes of 11 distinct BSV species have been fully sequenced, and partial genomic sequences are also available for numerous strains of these and other species. At present, a cryptic BSV complex comprising 11 species of BSV are known to be associated with the streak disease of banana worldwide (King et al., 2012; James et al., 2011). Phylogenetic analyses show that BSVs display a high level of molecular diversity (Geering et al., 2000; Harper et al., 2004, 2005; Jaufeerally-Fakim et al., 2006). BSVs are grouped into three main clades based on phylogenetic analysis (Gayral and Iskra-Caruana., 2009; Iskra-Caruana et al., 2014). Detection Detection methods such as ELISA, immunosorbent electron microscopy (ISEM) (Nodowora, 1998) and double antibody sandwich (DAS) ELISA (Geering et al., 2000; Meyer, 2005; Thottappilly et al., 1998) have been employed for detection of BSV. The viral genome integrated in host chromosome is

often amplified in standard PCR, which leads to false positives. To allow detection of episomal virus DNA, IC-PCR has been successfully employed (Harper et al., 1999b; Le Provost et al., 2006; Sharma et al., 2014; Selvarajan et al., 2016). Le Provost et al. (2006) used Musa sequence tagged microsatellite site primers to detect genomic DNA contamination in PCR to confirm thefalse positives. Multiplex reverse transcription (RT)–PCR has been used for detecting BSV to avoid detecting eBSV sequences (Selvarajan et al., 2011a; Liu et al., 2012). However, exclusive detection of episomal BSV by this method is not robust and appears to be difficult because eBSV sequences are sometimes known to be transcribed resulting in RNA transcripts that can be detected by RT–PCR and leading to wrong interpretations (Kumar et al., 2015). A novel and sensitive techniques, namely RCA ( James et al., 2011) and LAMP were developed to detect the BSV (Peng et al., 2012). Recently, Selvarajan et al. (2016) successfully detected the virus in an IC-PCR technique using antiserum to viral associated protein of BSMYV. Banana bract mosaic disease Banana bract mosaic disease (BBrMD), caused by Banana bract mosaic virus (BBrMV), was first reported in 1979 in the Philippines at Davao on the island of Mindanao and subsequently, reported from India, Sri Lanka, Western Samoa, Thailand, Vietnam, and Ecuador (Bateson and Dale, 1995; Thomas et al., 1997; Rodoni et al., 1997, 1999; Balasubramanian and Selvarajan, 2012, 2014a; Selvarajan and Balasubramanian, 2017). BBrMV was also reported to infect ornamental ginger plants (Alpinia purpurata) in Hawaii (Wang et al., 2010). Recently this disease has been reported to occur in the Assam, north eastern region, a place where wild bananas having B genome are believed to be one of the origins (Selvarajan and Balasubramanian, 2017). BBrMD cause a yield loss of 30–70% in French plantain cultivar Nendran in India and the Philippines (Cherian et al., 2002; Magnaye and Espino, 1990; Selvarajan and Jeyabaskaran., 2006). BBrMV, a member of the genus Potyvirus, family Potyviridae, and have flexuous filamentous particles of approximately 725 nm long which encapsidate a monopartite ssRNA genome of 9,711 bp in length (Thomas et al., 1997). A major open reading

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frame (ORF) encodes for a large polyprotein of 3,125 amino acids which is further cleaved into ten mature functional proteins by viral proteinase (Ha et al., 2008; Balasubramanian and Selvarajan, 2012). In addition, an overlapping PIPO is present in the +2 reading frame within the encoding region of P3 protein. Molecular diversity analysis of Banana bract mosaic virus The complete genome of BBrMV-TRY from India and BBrMV-PHI from the Philippines showed 94% similarity in the nucleotide sequence and 88–98% amino acid sequence identities for the individual proteins (Balasubramanian and Selvarajan, 2012). The coat protein-encoding gene along with 3′-untranslated region of BBrMV isolates from India, the Philippines, Samoa, and Vietnam, were reported to be 95.4–99% identical (Rodoni et al., 1999). The CP gene of 49 isolates of BBrMV were studies for the genetic diversity which revealed a greater variation among them and the recombination analysis revealed eleven putative recombinants and two of the 49 isolates from Tamil Nadu were found distinct with 18–21% divergence at nucleotide level and 12–20% divergence at amino acid level (Balasubramanian and Selvarajan, 2014a) whereas the analysis of HC-Pro gene of BBrMV isolates showed that 92–100% similarity both at the nucleotide (nt) and amino acid level (Balasubramanian et al., 2014). Partial gene sequences of BBrMV from Hawaii and Ecuador were 99% identical, compared with the corresponding fragment of BBrMV isolate from the Philippines (Quito-Avila et al., 2013; Wang et al., 2010). Further study with a greater population will reveal the real population structure of the virus. Characteristic symptoms Banana bract mosaic disease (BBrMD), caused by BBrMV, is characterized by the presence of dark reddish-brown spindle-shaped, discontinuous, mosaic on pseudostem, midrib, peduncle, bracts of inflorescence and fingers (Fig. 13.1C and D). The characteristic mosaic symptoms on the flower bracts give the disease its common name; however, the numerous reddish spindle-shaped streaks on the emerging suckers and the pseudostem are the earliest signs of the infection. Due to changes in leaf orientation in French plantains like cultivar Nendran, the infected bananas often resemble

traveller’s palm, Ravenala madagascariensis (Balakrishnan et al., 1996; Jones, 2000). Infected plants have bunches with unusually long or very short peduncle, and bunches sometimes get choked and fingers get malformed or distorted (Selvarajan and Jeyabaskaran, 2006). Transmission and host range BBrMV is primarily transmitted through infected planting material including suckers, corms, and tissue-cultured plantlets. The virus is non–persistently transmitted through several aphid species, namely Pentanlonia nigronervosa, Rhopalosiphum maidis, Aphis gossypii (Magnaye and Espino, 1990; Munez, 1992), A. craccivora (Selvarajan et al., 2006). The main host of BBrMV is Musa but it also infects Abacá (M. textilis) (Sharman et al., 2000) and has been reported to infect small cardamom (Elettaria cardamomum) in India (Siljo et al., 2012) and flowering ginger, Alpinia purpurata (Vieill.) K. Schum, in Hawaii (Wang et al., 2010). Detection Serological techniques such as ELISA, Western blots and dot-blot immune-binding assays have been reported to detect BBrMV using polyclonal and monoclonal antisera (Espino et al., 1990; Thomas et al., 1997; Rodoni et al., 1999). Reverse transcription PCR (RT-PCR) and multiplex RT–PCR/IC-RT–PCR were found to be more sensitive in detecting BBrMV (Bateson and Dale,1995; Rodoni et al., 1997; 1999; Thomas et al., 1997; Sharman et al., 2000a; Iskra-Caruana et al., 2008; Balasubramanian and Selvarajan, 2012; Selvarajan and Balasubramanian, 2013b). Real-time quantitative PCR assay was developed for the detection and quantification of BBrMV using the SYBR Green and TaqMan chemistries (WeiGang et al., 2009; Siljo et al., 2014; Balasubramanian and Selvarajan, 2014b). Reverse transcription loop-mediated isothermal amplification (RT-LAMP) and IC-RTLAMP have been developed for rapid detection of BBrMV. The detection sensitivity of the RT-LAMP assay was 100 times higher than that of the conventional RT-PCR (Siljo and Bhat, 2014; Zhang et al., 2016). Non-transgenic management Variability among the BBrMV isolates is relatively low, so development of stable transgenic resistance will help in managing this disease. Sources of

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resistance in germplasm of banana have not been reported for BBrMV. It is recommended to follow standard control measures including the removal and destruction of infected plants along with the rhizome and replace with virus-free tissue culture planting material. Application of fertilizer was reported to mitigate the symptoms of BBrMV in certain commercial banana cultivars (Selvarajan et al., 2009, 2012, 2017). Dizon and co-workers (2012) developed mutant abaca cvs. Tinawagan Pula and Tangongon with putative resistance to BBrMV through in vitro mutagenesis by gammairradiation. Infectious chlorosis (banana mosaic) Banana mosaic is one of the common viral diseases to affect bananas and plantains in all tropical and subtropical regions of the world. This disease, also called infectious chlorosis, is caused by Cucumber mosaic virus (CMV) (Lockhart and Jones, 2000). CMV infects over 1200 species of plant hosts in tropical, subtropical and temperate climates, including members of 85 plant families and approximately 365 genera, making it the most common of all plant viruses reported (Edwardson and Christie, 1991). The host range includes both dicots and monocots and it has been recorded virtually in every country with extensive agriculture. CMV is the type species of the genus Cucumovirus and family Bromoviridae (Palukaitis and García-Arenal, 2003). The virus particles are isometric in shape, measure 28–30 nm in diameter and each particle is composed of 180 subunits (Palukaitis et al., 1992). CMV has tripartite, positive-sense single-stranded RNA genome with 5′ cap and 3′ tRNA-like structures. CMV has three genomic RNAs, namely RNA 1, RNA 2 and RNA 3, and two subgenomic RNAs, namely RNA 4 and RNA 4A, which are transcribed from the 3′ portions of RNA 3 and RNA 2, respectively (Ding et al., 1994). RNAs 1 and 2 encode the nonstructural proteins involved in viral replication. RNA 3 contains two genes, 3a encodes a movement protein and the 3b encodes a coat protein. It is reported that the subgenomic RNA 4 get translated into CP, while the RNA 2-derived RNA 4A encodes the 2b protein which is involved in the long-distance movement, suppression of gene silencing, and

expression of systemic symptoms (Ding et al., 1995; Brigneti et al., 1998). Characteristic symptoms of banana mosaic A range of symptoms due to CMV infection on banana are reported. Diffused leaf mosaic to severe chlorosis, chlorotic streaking or flecking, stripes, line patterns, ring spots, leaf curling, distortion, cigar leaf or heart leaf rotting (Fig. 13.1G and H), rosette appearance of leaf arrangement and stunting of plant (Niblett et al., 1994). Uneven ripening has been associated with this virus infection (Lockhart and Jones, 2000). Mosaic symptoms are most pronounced during cool weather but do not persist all along the growth stages of plant. In contrast, the severe heart rot strains of CMV can cause damaging symptoms which include chlorosis, cigar leaf necrosis, internal pseudostem necrosis and ultimately leads to plant death (Srivastava et al., 1995). Transmission and host range The virus is transmitted through planting material and non-persistently by several aphids, such as A. gossypii, A. craccivora, R. maidis, R. prunifolium and Myzus persicae (Rao, 1980). Numerous strains of CMV have been described as it has an extremely broad host range numbering 1200 plant species. Genetic diversity Several strains of CMV have been classified into subgroups I and II based on serological properties and nucleotide sequence homology (Palukaitis et al., 1992). The subgroup I has been further divided into two groups (IA and IB) by phylogenetic analyses (Roossinck et al., 1999). One of the subgroups (IB) is restricted to Asia, whereas the other two subgroups (IA and II) are distributed worldwide. Most of the isolates of CMV from banana (CMVB) belong to subgroup I, which includes most of the CMV isolates from the tropics. CP gene of several CMV isolates infecting banana in India has been sequenced and compared (Selvarajan et al., 2007; Khan et al., 2011). The sequence analysis showed 93–98% nt and 94–99% aa sequence identity, and phylogenetic analysis revealed that all banana-infecting isolates belong to subgroup IB (Vishnoi et al., 2013). Qin et al. (2012) reported that also the subgroup II of CMV infects banana in China.

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Detection of Cucumber mosaic virus Serological methods such as direct antigen coating (DAC) ELISA, DAS-ELISA and dot immunobinding assay (Kawano et al., 1995; Kiranmai et al., 1996), and nucleic acid-based assays such as RT–PCR (Selvarajan et al., 2007; Singh et al., 1995; Hu et al., 1995) and dot blot hybridization technique using radio-labelled and DIG probes (Srivastava et al., 1995) to detect banana infecting CMV isolates have been reported. IC- multiplex PCR has been successfully used to detect BBrMV, CMV and BBTV (Selvarajan and Balasubramanian, 2008; Sharman et al., 2000a). Kouassi et al. (2010) reported that to screen banana planting material against CMV in Ivory Coast, real-time PCR and FTA technologies have been developed. Peng et al. (2012) developed real-time LAMP for the effective and accurate detection of CMV in banana.

mother plants and tissue culture derived plants. In Australia, Queensland Banana Accredited Nursery (QBAN) generates pathogen-free stocks, which are used as foundation stock in the TC industryfor mass propagation (QPPR, 2002). Of late, BBTV is managed through the use of virus-free tissue culture, certified plants in Taiwan, the Philippines and India (Molina et al., 2009; Su et al., 2007; Selvarajan et al., 2011b). At present in India approximately100 million tissue culture raised banana plants are certified annually and 90 TCPUs are involved in raising quality certified banana plantlets. This will go long way to control and eradicate the banana viruses in a greater extent.

Management of banana viruses through certification of tissue culture plants Supply of high-quality and virus-free plants in banana is the foremost step in the management of viruses in any vegetatively propagated plants. Preceding this, identification and eradication of infected clumps or plants are needed to eliminate the inoculum. Availability of virus-free certified banana plants for the replacement of eradicated clumps is very important. In India, tissue culture banana plants are covered by the Seed Act 1966, with effect from 2006. Accordingly, the Department of Biotechnology, Government of India, has set a standard operating procedures and guidelines to ensure quality banana plants free from viruses and genetically uniform under the National Certification System for Tissue Culture Plants (NCS-TCP). There are five accredited laboratories including Molecular Virology lab of Indian Council Agricultural Research-NRC for Banana (ICARNRCB), Tiruchirappalli, India, for virus testing of tissue culture raised plants of banana. All the DBT recognized banana tissue culture production units (TCPUs) laboratories in India have to get certification of quality and labels for their TC raised banana plants ensuring freedom from viruses and genetic uniformity that qualify the TCPU’s to sell or dispatch their plants to the farmers. At ICARNRCB several validated detection techniques developed are being used to detect the viruses from

Transgenic resistance to Banana bunchy top virus The best option for managing plant viruses is using genetic engineering to obtain resistant transgenics, and this is more applicable where there are no opportunities of developing resistant plants through conventional breeding to obtain resistance. Such genetic engineering technological intervention includes generating plants with a foreign gene either from the pathogen itself which is called as pathogen derived resistance (Sanford and Johnston, 1985). The whole of part of viral gene, either sense or antisense, can be moved to the plant of interest by transforming them either through Agrobacterium-mediated approach or through direct gene delivery system using gene gun. There are many plant species in which resistance to RNA viruses has been achieved through transformation of viral derived genes especially (Goldbach et al., 2003). This concept is known as pathogen-derived resistance (PDR), and has been employed successfully in various plant species to confer resistance to many RNA viruses. In case of DNA viruses of Geminiviridae, similar PDR strategy has been partially successful in obtaining the resistance or tolerance (Vanderschuren et al., 2007; Hanley-Bowdoin and Settlage, 2004; Hanley-Bowdoin et al., 2004; Goldbach et al., 2003). These reports have given some hope that it will be possible to develop resistance to other DNA viruses, such as BBTV, a babuvirus belonging to the family Nanoviridae.

Current status in viral resistant transgenics in banana

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To overcome the current limitations of lack of resistant source to BBTV in the Musa germplasm and breeding challenges for edible Musa spp., transgenic approaches based on the PDR strategies are explored in many laboratories. Using PDR approach attempts were made to develop transgenic banana, with resistance to BBTV, in Australia, Hawaii and India. Coat protein and Rep gene of BBTV has been used for developing transgenics. At the University of Hawaii several putative transgenic lines have been developed expressing either mutated or anti-sense Rep genes with partial resistance to BBTV. A few lines were symptomless for up to a year; however they were not immune to BBTV as they all developed symptoms later in their crop cycle. Banana cultivars of Cavendish sub-group and Lady finger were transformed with the mutated Rep under the control of high level of constitutive promoter and they have also developed the virus activated cell death strategy where in which the BBTV intergenic region has been embedded into an intron and inserted into a split barnase gene construct to develop transgenics (Dale and Harding, 2003). Putative transgenic Hill banana lines resistant for BBTV using embryogenic cell suspensions were developed with cp gene (Selvarajan et al., 2014) and Replicase gene (Selvarajan, unpublished) (Fig. 13.3A and B). Borth et al. (2011) have succeeded in generating banana plants resistant to BBTV using the viral Rep gene. Strategies using BBTV DNA-R gene or satellite DNA (DNA-S4) resulted partial resistance to BBTV (Tsao, 2008). The latest trend is to adopt gene silencing through RNA interference (RNAi) strategy in bananas for a stable BBTV resistance. More recent efforts using RNAi have resulted in the development of clones with near immunity to infection (Borth et al., 2011; Shekhawat et al., 2012; Elayabalan et al., 2013). Shekhawat et al. (2012) explored the concept of using ihpRNA (intron containing hairpin) transcripts corresponding to viral master replication initiation protein (Rep) to generate BBTV-resistant transgenic banana plants. This study indicated that the use of an intron between the two complementary domains of Rep-derived sequences makes for efficient siRNA synthesis, and they have obtained 100% resistance to BBTV infection in transgenic plants. Elayabalan et al. (2013) generated hill banana resistant to BBTV targeting master replicase initiation protein coding gene

using a RNAi-approach with a construct containing intron and partial master replicase initiation protein coding gene of BBTV (Fig. 13.4A). The transformed plants were symptomless, and the replication of challenged BBTV was almost completely suppressed. This approach was shown to be effective in the management of BBTV in hill banana. Self-forming hairpin using full length Master rep gene of BBTV and an inverted partial master rep gene constructs (Fig. 13.4B) were used to generate putative transgenic lines of Hill banana (Selvarajan, unpublished). Recently, IITA and Queensland University of Technology (QUT) have initiated an RNAi-based transgenic programme to develop banana and plantain resistant to BBTV and aphids for Africa (Kumar et al., 2015). They have generated a large number of Cavendish lines with a range of RNAi constructs targeting different BBTV genes that have shown levels of BBTV resistance in the glasshouse in Australia (Mwari, 2016). Many of these lines have been progressed through to a field trial recently planted in Malawi. Transgenic resistance to Banana bract mosaic virus BBrMV genome has relatively less variability and development of horizontal resistance is necessary. PTGS mechanism can be used for developing transgenics against this virus as the suppressor gene silencing in viral genome has been reported in other species of potyviruses. Dale and Harding (2003) have successfully developed transgenic ‘Cavendish’ banana by transforming with the coat protein gene of a Philippines isolate under the control of the maize poly-ubiquitin promoter using microprojectile bombardment and obtained resistance to BBrMV at QUT under the World Bank-sponsored Banana Improvement Program (BIP). Tavanlar (2007) has also reported the use of coat protein gene of BBrMV for developing transgenic SABA resistant to BBrMV. Transgenic resistance to banana streak viruses Owing to hypervariability of episomal BSVs reported in banana, it appears to be a difficult to develop a sustainable strategy to get broad resistance in banana for various species of BSV. To date there are no successful reports of generation of a high level of resistance to badna or caulimovirus

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(A)

(B)

Figure 13.3  (A) Development of ECS for Hill banana cv. Virupakshi, having Geographical Indications (GI 124) for the use in the transgenic development against BBTV, a serious viral pathogen devastated the variety in Lower Puney Hills of Tamil Nadu, India. (B) A view of transgenic Hill banana plants developed using cp gene of BBTV.

except for a few attempts against Rice tungro bacilliform virus (RTBV) in case of rice, in which the cp gene has been exploited for developing resistance (Ganesan et al., 2009). Dr Dasgupta’s group from

the University of Delhi (Tyagi et al., 2008) used RNAi for the control of RTBV infection where in transgenic rice plants expressing hpRNA targeting ORF IV of RTBV, resulted in the formation of

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(A)

(B)

Figure 13.4  (A) Intron containing RNAi vector using partial mater replicase initiation protein coding gene of BBTV, reprinted by permission from Springer Nature (Development of Agrobacterium-mediated transformation of highly valued hill banana cultivar Virupakshi (AAB) for resistance to BBTV disease, World Journal of Biotechnology and Microbiology 29, by R Selvarajan, 2018); (B) self-forming hairpin using full length master rep gene of BBTV and an inverted parial master rep gene (source: Selvarajan, unpublished).

double-stranded (ds) RNA. However, in case of banana it will be difficult to generate resistance to BSV due to the presence of viral genome integrations and hypervariability of episomal BSVs. The genome editing tools like TALENS, Zinc finger nuclease, and CRISPR-Cas9 with short guide RNA-based methods have been used to mutate or delete unwanted genes or gene fragments in model plant species. Now, it becomes possible to delete the unwanted endogenous integrated viral sequences, which are capable of infecting the plants spontaneously and produce episomal viral forms, which can get transmitted to other healthy plants by mealybug

vectors. Efforts to unravel the significance of integrated badnaviral sequences would lead to a better understanding of acquired immunity and natural RNAi in plant crops (Bhat et al., 2016). Transgenic resistance to Cucumber mosaic virus Generation of broad spectrum transgenic resistance to CMV will be challenging as there is a wide genetic variability among different isolates. Coat proteinmediated resistance and post-transcriptional gene silencing (PTGS) has been exploited for generating transgenic resistance to CMV in other crops. First

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engineered resistance to CMV utilizing the coat protein (CP) gene was demonstrated by Cuozzo et al. (1988). Since then, many examples of CPmediated resistance (CPMR) in varying degrees have been described, with different constructs in different hosts (Beachy et al., 1990; Srivastava and Raj, 2008; Kumar et al., 2012). Till date no attempts have been made for generating transgenics against CMV isolates affecting banana. Future prospects Now novel techniques are available for developing viral resistance transgenics in banana. In case of BBTV in addition to DNA-R, gene products encoded by other DNA components also play pivotal roles in the life cycle of BBTV. In particular, the proteins encoded by DNA-M and DNA-S have been reported as silencing suppressors (Amin et al., 2011; Niu et al., 2009), and may be involved in modulating the hosts RNAi response. To date, these genes as well as DNA-N and –C have not been assessed for pathogen derived resistance through RNAi in banana. Targeting these viral suppressors of PTGS for silencing themselves may improve the effectiveness of RNA-mediated virus resistance. Internal ribosome entry site (IRES) can be used for stable expression of transgenes in plants (RenaudGabardos et al., 2015). CRISPR–Cas9 construct targeting conserved regions of BBTV can be developed for generating non-chimeric line with high expression levels in banana as reported in other crops for virus resistance (Baltes et al., 2015; Ji et al., 2015; Ali et al., 2015; Dale et al., 2017). Unique CRISPR–Cas constructs that target conserved BBTV sequences may be designed and engineered into banana plants to produce transgenic banana plants that are resistant to BBTV (Green et al., 2016). Eukaryotic translation initiation factor (eIF) gene family, including eIF4E and its paralogue eIF(iso)4E, have previously been identified as recessive resistance alleles against various potyviruses in a range hosts (Pyott et al., 2016). eIF4E of banana was found to interact with the VPg of BBrMV in yeast two-hybrid system (Anuradha and Selvarajan, unpublished). Hence, CRISPR-Cas9 technology could be exploited for introducing sequence specific deleterious point mutations at the eIF(iso)4E locus of banana to engineer and obtain stable resistance to BBrMV.

This same technology can also be used to generate BSV free plants by genome editing of the integrated sequences which on stress pop out to form episomal virus leading to spontaneous infection. These are some of the novel approaches which can also be explored for developing stable transgenics against the banana virus. References Abustan, S.A.-F.A. (2012). IPB-UPLB rallies against banana disease, gets PCAARRD support. Available online: www.pcaarrd.dost.gov.ph/home/portal/index.php/ quick-information-dispatch/2066-ipb-uplb-ralliesagainst-banana-disease-gets-pcaarrd-support. Accessed 20 March 2014. Ali, Z., Abulfaraj, A., Idris, A., Ali, S., Tashkandi, M., and Mahfouz, M.M. (2015). CRISPR/Cas9-mediated viral interference in plants. Genome Biol. 16, 238. https:// doi.org/10.1186/s13059-015-0799-6 Amin, I., Qazi, J., Mansoor, S., Ilyas, M., and Briddon, R.W. (2008). Molecular characterisation of Banana bunchy top virus (BBTV) from Pakistan. Virus Genes 36, 191–198. https://doi.org/10.1007/s11262-007-0168-y Anhalt, M.D., and Almeida, R.P. P. (2008). Effect of temperature, vector life stage, and plant access period on transmission of Banana bunchy top virus to banana. Phytopathology 98, 743–748. https://doi.org/10.1094/ PHYTO-98-6-0743 Balakrishnan, S., Gokulapalan, C., and Paul, S. (1996). A widespread banana malady in Kerala, India. Infomusa 5, 28–29. Balasubramanian, V., and Selvarajan, R. (2012). Complete genome sequence of a banana bract mosaic virus isolate infecting the French plantain cv. Nendran in India. Arch. Virol. 157, 397–400. https://doi.org/10.1007/s00705011-1179-5 Balasubramanian, V., and Selvarajan, R. (2014a). Genetic diversity and recombination analysis in the coat protein gene of Banana bract mosaic virus. Virus Genes 48, 509–517. https://doi.org/10.1007/s11262-014-1056-x Balasubramanian, V., and Selvarajan, R. (2014b). Taq Man real-time PCR for detection and quantitation of banana bract mosaic virus in banana and plantain. In: Abstracts of the XXIII National Conference of Indian Virological Society (IVS), ‘VIROCON 2014’ on ‘Recent Trends in Virology Research in the Omics Era’, Tamil Nadu Agricultural University, Coimbatore, Tamil Nadu, India, 18–20 December, 2014. pp. 85. Balasubramanian, V., Sukanya, R.S., Anuradha, C., and Selvarajan, R. (2014). Population structure of Banana bract mosaic virus reveals recombination and negative selection in the helper component protease (HCPro) gene. VirusDisease 25, 460–466. https://doi. org/10.1007/s13337-014-0241-9 Baltes, N.J., Hummel, A.W., Konecna, E., Cegan, R., Bruns, A.N., Bisaro, D.M., and Voytas, D.F. (2015). Conferring resistance to geminiviruses with the CRISPR–Cas prokaryotic immune system. Nat. Plants 1, 15145. Banerjee, A., Roy, S., Behere, G.T., Roy, S.S., Dutta, S.K., and Ngachan, S.V. (2014). Identification and characterization of a distinct Banana bunchy top virus

270  | Selvarajan et al.

isolate of Pacific-Indian Oceans group from North-East India. Virus Res.183, 41–49. https://doi.org/10.1016/j. virusres.2014.01.017 Bateson, M.F., and Dale, J.L. (1995). Banana bract mosaic virus:characterisation using potyvirus specific degenerate PCR primers. Arch. Virol. 140, 515–527. Beachy, R.N., Loesch-Fries, S., and Tumer, N.E. (1990). Coat protein-mediated resistance to virus infection. Annu. Rev. Phytopathol. 28, 451–74. Beetham, P.R., Harding, R.M., and Dale, J.L. (1999). Banana bunchy top virus DNA-2 to 6 are monocistronic. Arch.Virol. 144, 89–105. Beetham, P.R., Hafner, G.J., Harding,R.M., and Dale, J.L. (1997). Two mRNAs are transcribed from Banana bunchy top virus DNA-1. J. Gen. Virol. 78, 229–236. https://doi.org/10.1099/0022-1317-78-1-229 Bhat, A.I., Hohn, T., and Selvarajan, R. (2016). Badnaviruses: The Current Global Scenario. Viruses 8, E177. https:// doi.org/10.3390/v8060177 Borth, W., Perez, E., Cheah, K., Chen, Y., Xie, W.S., Gaskill, D., et al. (2011). Transgenic banana plants resistant to Banana bunchy top virus infection. Acta Horticulturae, 897, 449–457. Bressan, A., and Watanabe, S. (2011). Immunofluorescence localization of Banana bunchy top virus (family Nanoviridae) within the aphid vector, Pentalonia nigronervosa, suggests a virus tropism distinct from aphid-transmitted luteoviruses. Virus Res. 155, 520– 525. Brigneti, G., Voinnet, O., Li, W.X., Ji, L.H., Ding, S.W., and Baulcombe, D.C. (1998). Viral pathogenicity determinants are suppressors of transgene silencing in Nicotiana benthamiana. EMBO J. 17, 6739–6746. https://doi.org/10.1093/emboj/17.22.6739 Burns, T.M., Harding, R.M., and Dale, J.L. (1995). The genome organization of Banana bunchy top virus: analysis of six ssDNA components. J. Gen. Virol. 76, 1471–1482. https://doi.org/10.1099/0022-1317-76-6-1471 Chabannes, M., Baurens, F.C., Duroy, P.O., Bocs, S.,Vernerey, M.S., Rodier-Goud, M., Barbe, V., Gayral, P., and Iskra-Caruana, M.L. (2013). Three infectious viral species lying in wait in the banana genome. J.Virol. 87, 8624–8637. https://doi.org/10.1128/JVI.00899-13 Chandrasekar, A., Kalaiponmani, K., Elayabalan, S., Kumar, K.K., Angappan, K., and Balasubramanian, P. (2011). Screening of Banana bunchy top virus through multiplex PCR approach. Archives of Phytopathology and Plant Protection 44, 1920–1925. Chen, Y., and Hu, X. (2013). High-throughput detection of Banana bunchy top virus in banana plants and aphids using real-time TaqMan(®) PCR. J. Virol. Methods 193, 177– 183. https://doi.org/10.1016/j.viromrt.2013.06.013 Cherian, A.K., Menon, R., Suma, A., Nair, S., And Sudheesh, M.V. (2002). Impact of banana bract mosaic diseases on the yield of commercial banana varieties of Kerala. Global Conference on Banana and Plantain, Bangalore, 28–31 October 2002, Abstract p. 155. Chiaki, Y., Nasir, N., Herwina, H., Jumjunidang, Sonoda, A., Fukumoto, T., Nakamura, M., and Iwai H. Genetic structure and diversity of the Banana bunchy top virus population on Sumatra Island, Indonesia. European J. of Plant Pathology 143, 113-122.

Cuozzo M., O’Connell, K.M., Kaniewski, W., Fang, R., Chua, N., and Tumer, N.E. (1988). Viralprotection in transgenic tobacco plants expressing the Cucumber mosaic virus coat protein or its antisense RNA. Biotechnology 6, 549–557. Dahal, G., Hughes, Jd’A ., Thottappilly, G., and Lockhart, B.E.L.(1998). Effect of temperature on symptom expression and reliability of banana streak badnavirus detection in naturally infected plantain and banana (Musa spp.). Plant Disease 82, 16–21. Dahal, G., Ortiz, R., Tenkouano, A., Hughes, J. d’A ., Thottappilly, G., Vuylsteke, D., etal. (2000). Relationship between natural occurrence of banana streak badnavirus and symptom expression, relative concentration of viral antigen, and yield characteristics of some micropropagated Musa spp. Plant Pathol. 49, 68–79. Damasco, O.P., Estrella, J.B., Caymo, L.C., Dizon, T.O., Rabara, R.C., de la Cruz, F.S., Jr., et al. (2006). Banana bunchy top virus (BBTV) resistance in cultivar ‘Lakatan’ developedby gamma irradiation of shoot tips. Philippine J. Crop Sci. 31, 21–34. Daniells, J.W., Geering, A.D. W., Bryde, N.J., and Thomas, J.E. (2001). The effect of Banana streak virus on the growth and yield of dessert bananas in tropical Australia. Ann. Appl. Biol. 139, 51–60. Dale, J.L. (1987). Banana bunchy top: an economically important tropical plant virus disease. Adv. Virus Res. 33, 301–325. Dale, J., Paul, J.Y., Dugdale, B., and Harding, R. (2017). Modifying bananas: from transgenics to organics? Sustainability 9, 333. Dale, J., and Harding, R. (2003). Strategies for the generation of virus resistance banana. In Genetic Transformation Strategies to Address the Major Constraints to Banana and Plantain Production in Africa: Atkinson, H.J, Dale, J., Harding, R., Kiggundu, A., Kunert, K., Muchwezi J.M., et al., eds. (INIBAP, Montpellier), pp. 108–118. Davis, R.I., Geering, A.D.W., Thomas, J.E., Gunua, T.G., and Rahamma, S. (2000). First records of Banana streak virus on the island of New Guinea. Aus. Plant Pathol. 29, 281. Ding, S.W., Anderson, B.J., Haase, H.R., and Symons, R.H. (1994). New overlapping gene encoded by the Cucumber mosaic virus genome. Virology 198, 593–601. Ding, S.W., Li, W.X., and Symons, R.H. (1995). A novel naturally occurring hybrid gene encoded by a plant RNA virus facilitates long distance virus movement. EMBO J. 14, 5762–5772. Dizon, O.T., Damasco, O.P., Lobina, I.T., Pinili, M.S., Lalusin, A.G., and Natsuaki, K.T. (2012). Induction of putative resistant lines of abaca (Musa textiles Nee) to bananabunchytop virus and banana bract mosaic virus through in vitro mutagenesis. J. ISSAAS 18, 87–99. Drew, R.A., Moisander, J.A., and Smith, M.K. (1989). The transmission of banana bunchytop virus in micropropagated bananas. Plant Cell Tissue and Organ Culture 16, 187–193. Edwardson, J.R., and Christie, R.G. (1991).Cucumoviruses: CRC Handbook of Viruses Infecting Legumes (CRC Press, Boca Raton), pp. 293–319. Elayabalan, S., Kalaiponmani, K., Subramaniam, S., Selvarajan, R., Panchanathan, R., Muthuvelayoutham, R., Kumar, K.K., and Balasubramanian, P. (2013).

Viruses Infecting Banana and their Transgenic Management |  271

Development of Agrobacterium-mediated transformation of highly valued hill banana cultivar Virupakshi (AAB) for resistance to BBTV disease. World J. Microbiol. Biotechnol. 29, 589–596. https:// doi.org/10.1007/s11274-012-1214-z Espino, T.M., Exconde, S.B., Zipagan, F.B., and Espino, R.R.C. (1990). Banana bract mosaic, a new disease of banana II. Isolation andpurification for monoclonal antibody production. The Philippine Agriculture 73, 61–68. FAOStat (2014). FAO production statistics for banana and plantain 2012. Available online: http://faostat.fao.org/ site/567/DesktopDefault.aspx?PageID¼567#ancor. Accessed 20 March 2014. Gayral, P., and Iskra-Caruana, M.L. (2009). Phylogeny of Banana Streak Virus reveals recent and repetitive endogenization in the genome of its banana host (Musa sp.). J. Mol. Evol. 69, 65–80. https://doi.org/10.1007/ s00239-009-9253-2 Geering, A.D.W., and Thomas, J.E. (1996). A comparison of four serological tests for the detection of Banana bunchy top virus in banana. Austral. J. Agric. Res. 47, 403–412. Geering, A.D., McMichael, L.A., Dietzgen, R.G., and Thomas, J.E. (2000).Genetic diversity among Banana streak virus isolates from Australia. Phytopathology 90, 921–927. https://doi.org/10.1094/PHYTO.2000.90.8.921 Geering, A.D., Pooggin, M.M., Olszewski, N.E., Lockhart, B.E., and Thomas,J.E. (2005). Characterisation of Banana streak Mysore virus and evidence that its DNA is integrated in the B genome of cultivated Musa. Arch. Virol. 150, 787–796. https://doi.org/10.1007/s00705004-0471-z Ganesan, U., Suri, S.S., Rajasubramaniam, S., Rajam, M.V., and Dasgupta, I. (2009). Transgenic expression of coat protein gene of Rice tungro bacilliform virus in rice reduces the accumulation of viral DNA in inoculated plants. Virus Genes 39, 113–119. https://doi.org/10.1007/ s11262-009-0359-9 Goldbach, R., Bucher, E., and Prins, M. (2003). Resistance mechanisms to plant viruses: an overview. Virus Res. 92, 207–212. Green, J.C, Borth, W., and Hu, J.S. (2017). Engineering resistance to viruses. In Banana: Genomics and Transgenic Approaches for Genetic Improvement. Mohandas, S., Ravishankar, K.V., eds. (Springer, Singapore), pp. 237–246. Ha, C., Coombs, S., Revill, P.A., Harding, R.M., Vu, M., and Dale, J.L. (2008). Design and application of two novel degenerate primer pairs for the detection and complete genomic characterization of potyviruses. Arch. Virol. 153, 25–36. https://doi.org/10.1007/s00705-0071053-7. Hanley-Bowdoin, L., and Settlage, S.B. (2004). Transgenic plants expressing mutant geminivirus AL3/C3 coding sequence. North Carolina State University. U.S. Patent.6747188 B2. Hanley-Bowdoin, L., Orozco, B.M., Kong, L.J., and Gruissem, W. (2004). Geminivirus transgenic plants. North Carolina State University and Reagents of the University of Carolina. U.S. Patent 6800793 B2. Harper, G., and Hull, R. (1998). Cloning and sequence analysis of banana streak virus DNA. Virus Genes 17, 271–278.

Harper, G., Osuji, J.O., Heslop-Harrison, J.S., and Hull, R. (1999a). Integration of banana streak badnavirus into the Musa genome: molecular and cytogenetic evidence. Virology 255, 207–213. Harper, G., Ganesh, D., Thottappilly, G., and Hull, R. (1999b). Detection of episomal banana streak virus by IC-PCR. J. Virol. Meth., 79, 1–8. Harper, G., Hart, D., Moult, S., and Hull, R. (2002). Detection of Banana streak virus in field samples of bananas from Uganda. Ann. Appl. Biol., 141, 247–257. Harper, G., Hart, D., Moult, S., and Hull, R. (2004). Banana streak virus is very diverse in Uganda. Virus Res. 100, 51–56. https://doi.org/10.1016/j.virusres.2003.12.024 Harper, G., Hart, D., Moult, S., Hull, R., Geering, A., and Thomas, J. (2005). The diversity of Banana streak virus isolates in Uganda. Arch. Virol. 150, 2407–2420. https://doi.org/10.1007/s00705-005-0610-1 Hu, J.S., Li, H.P., Barry, K., Wang, M., and Jordan, R. (1995). Comparison of dot blot, ELISA, and RT-PCR assays for detection of two Cucumber mosaic virus isolates infecting banana in Hawaii. Plant Dis.79, 902–906. Hull, R., Harper, G., and Lockhart, B. (2000). Viral sequences integrated into plant genomes. Trends Plant Sci. 5, 362–365. Iskra-Caruana, M.L., Chabannes, M., Duroy, P.O., and Muller, E. (2014). A possible scenario for the evolution of Banana streak virus in banana. Virus Res. 186, 155– 162. https://doi.org/10.1016/j.virusres.2014.01.005 Iskra-Caruana, M.L., Galzi, S., and Laboureau, N. (2008). A reliable IC One-step RT-PCR method for the detection of BBrMV to ensure safe exchange of Musa germplasm. J.Virol. Methods 153, 223–231. https://doi. org/10.1016/j.jviromet.2008.06.028 James, J.A., Geijskes, R.J., Dale, J.L., and Harding, R.M. (2010). Development of a novel rolling-circle amplification technique to detect banana streak virus which also discriminates between integrated and episomal virus sequences. Plant Dis. 95, 57–62. James, A.P., Geijskes, R.J., Dale, J.L., and Harding, R.M. (2011). Molecular characterisation of six badnavirusspeciesassociated with leaf streak disease of banana in EastAfrica. Ann. Appl. Biol. 158, 346–353. Jaufeerally-Fakim, Y., Khorugdharry, A., and Harper, G. (2006). Genetic variants of Banana streak virus in Mauritius. Virus Res.115, 91–98. Ji, X., Zhang, H., Zhang, Y., Wang, Y., and Gao, C. (2015). Establishing a CRISPR-Cas-like immune system conferring DNA virus resistance in plants. Nat. Plants 1, 15144. https://doi.org/10.1038/nplants.2015.144 Jones, D.R. (2000). Diseases of Banana, Abaca and Enset. (CAB Int., Wallingford), pp. 263–274. Kapoor, R., Srivastava, N., Kumar, S., Saritha, R.K., Sharma, S.K., Jain, R.K., and Baranwal, V.K. (2017). Development of a recombinase polymerase amplification assay for the diagnosis of Banana bunchy top virus in different banana cultivars. Arch. Virol. 162, 2791–2796. https://doi. org/10.1007/s00705-017-3399-9 Kawano, S., Ikeshiro, I., Yonaha, T., Hanada, K., and Tokashiki, I. (1995). Occurrence of banana mosaic disease caused by Cucumber mosaic virus in Okinawa. Jap. J. Phytopathol. 61, 597–598. Khan, S., Jan. A.T., Aquil, B., and Haq, Q.M.R. (2011). Coat protein gene based characterization of Cucumber

272  | Selvarajan et al.

mosaic virus isolates infecting banana in India. J. Phytol. 3, 94–101. King, A.M.Q., Adams, M.J., Lefkowitz, E.J., Carstens, E.B. (2012). Virus Taxonomy: Ninth Report of the International Committee on Taxonomy of Viruses (Academic Press, San Diego), p. 1463. Kiranmai, G., Sreenivasulu, P., and Nayudu, M.V. (1996). Comparison of three different tests for detection of cucumber mosaic cucumovirus in banana (Musa paradisica). Curr. Sci. 71, 764–767. Kouassi, N.K., Wendy, M., Boonham, N., and Smith, J. (2010). Development of a diagnostic protocol for Cucumber mosaic virus for screening banana (Musa spp.) planting material in Ivory Coast. In Proc. IC on banana and plantain in Africa, T. Dubois, et al. Acta Hort. 879, ISHS 2010. Kubiriba, J., Legg, J.P., Tushemereirwe, W., and Adipala, E. (2001). Vector transmission of banana streak virus in the screenhouse in Uganda. Ann. Appl.Biol. 139, 37–43. Kumar, P.L., Hanna, R., Alabi, O.J., Soko, M.M., Oben, T.T., Vangu, G.H., and Naidu, R.A. (2011). Banana bunchy top virus in sub-Saharan Africa: investigations on virus distribution and diversity. Virus Res. 159, 171–182. https://doi.org/10.1016/j.virusres.2011.04.021 Kumar, S., Raj, S.K., Sharma, A.K., and Varma. H.N. (2012). Genetic transformation and development of Cucumber mosaic virus resistant transgenic plants of Chrysanthemum morifolium cv. Kundan. Scientia Horticulturae 134, 40–45. Kumar, P.L., Selvarajan, R., Iskra-Caruana, M.L., Chabannes, M.,and Hanna, R. (2015). Biology, etiology, and control of virus diseases of banana and plantain. Adv. Virus Res. 91, 229–269. https://doi.org/10.1016/ bs.aivir.2014.10.006 Le Provost, G., Iskra-Caruana, M.L., Acina, I., and Teycheney, P.Y. (2006). Improved detection of episomal Banana streak viruses by multiplex immunocapture PCR. J. Virol. Methods 137, 7–13. Liu, F., Feng, L., Chen, X., Han, Y., Li, W., Xu, W., et al. (2012). Simultaneous detection of four banana viruses by multiplex PCR. J. Phytopathol. 160, 622–627. Lockhart, B.E.L., and Olszewski, N.E. (1993). Serological and genomic heterogeneity of banana streak badnavirus: implications for virus detection in Musa germplasm. In Breeding Banana and Plantain for Resistance to Diseases and Pests J. Ganry Ed. (CIRAD/INIBAP, Montpellier), pp. 105–113. Lockhart, B.E.L., Ndowora, T.C., Olszewski, N.E.,and Dahal, G. (1998). Studies on integration of banana streak Badnavirus sequences in Musa: identification of episomally expressible badnaviralintegrants in Musa genotypes. In Banana Streak Virus: A Unique VirusMusa Interaction? Frison, E.A., and Sharrock, S.E., eds. Proceedings of a workshop of the PROMUSA Virol. working group held in Montpellier, France. Lockhart, B.E.L., and Jones, D.R. (2000). Banana streak mosaic. In Diseases of Banana, Abaca and Enset. Jones, D.R., ed. (CAB International, Wallingford), pp. 263–274. Lockhart, B.E.L., and Jones, D.R. (2000). Banana mosaic. In Diseases of Banana, Abaca and Enset. Jones, D.R., ed. (CAB International, Wallingford), pp.256–63.

Magee, C.J. P. (1948). Transmission of banana bunchy top to banana varieties. J. Austral. Instit. Agric.Sci. 14, 18–24. Magnaye, L.V., and Espino, R.R.C. (1990). Note: Banana bract mosaic, a new disease of banana I. Symptomatology. Philippine Agriculturist, 73, 55–59. Manser, P.D. (1982). Bunchy top disease of plantain. F.A.O. Plant Protect. Bull. 30, 78–79. Mansoor, S., Qazi, J., Amin, I., Khatri, A., Khan, I.A., Raza, S., Zafar, Y., and Briddon, R.W. (2005). A PCR-based method, with internal control, for the detection of Banana bunchy top virus in banana. Mol. Biotechnol.30, 167–170. Meyer, J.B. (2005). Banana streak badnavirus (BSV) in South Africa: incidence, transmission, and the development of an antibody-based detection system. (University of Pretoria, Pretoria). Meyer, J.B., Kasdorf, G.G.F., Nel, L.H., and Pietersen, G. (2008). Transmission of activated episomal banana streak OL (badna)virus (BSOLV) to cv. Williams banana (Musa sp.) by three mealybug species. Plant Dis. 92, 1158–1163. Molina, A.B., Sinohin, V.G.O., de la Cueva, F.M., Esguerra, A.V., Crucido, S.S., Vida, E., et al. (2009). Managing Banana bunchy top virus in smallholdings in the Philippines. Acta Horticulturae 828, 383–387. Mukwa, L.F.T., Gillis, A., Vanhese, V., Romay, G., Galzi, S., Laboureau, N., Kalonji-Mbuyi, A., Iskra-Caruana, M L., and Bragard, C. (2016). Low genetic diversity of Banana bunchy top virus, with a sub-regional pattern of variation, in Democratic Republic of Congo Virus Genes 52, 900–905. Munez, A.R. (1992). Symptomatology, transmission and purification of banana bract mosaic virus (BBMV) in ‘giant cavendish’ banana (University of Philippines, Los Baños). Mwari, B. (2016). Development of Banana bunchy top virus resistance in bananas: RNAi approach. Ph.D. thesis, Queensland University of Technology, Brisbane, Australia. Ndowora, T.C. (1998). Banana streak virus: development of an immunoenzymatic assay fordetection and characterisation of sequences that are integrated into the genome of the host Musa sp. PhD thesis, University of Minnesota. Nelson, S.C. (2004). Banana bunchy top: detailed signs and symptom. Cooperative extension service college of tropical agriculture and human resources (Mānaoa University of Hawai’i, Hawai’i). Niblett, C.L., Pappu, S.S., Bird, J., and Lastra, R. (1994). Infectious chlorosis,mosaic and heart rot. In Compendium of Tropical Fruit Disease, Ploetz, R.C., Zentmyer, G.A., Nishijima, W.T., Rohrback, K.G., Ohr, H.D., eds. (APS Press, Minnesota), pp. 18–19. Niu, S., Wang, B., Guo, X., Yu, J., Wang, X., Xu, K., Zhai, Y., Wang, J., and Liu, Z. (2009). Identification of two RNA silencing suppressors from Banana bunchy top virus. Arch.Virol. 154, 1775–1783. https://doi.org/10.1007/ s00705-009-0515-5 Palukaitis, P., and García-Arenal, F. (2003). Cucumoviruses. Adv.Virus Res. 62, 241–323. Palukaitis, P., Roossinck, M.J., Dietzgen, R.G., and Francki, R.I. (1992). Cucumber mosaic virus. Adv. Virus Res.41, 281–348.

Viruses Infecting Banana and their Transgenic Management |  273

Peng, J., Fan, Z., and Huang, J. (2012). Rapid detection of banana streak virus by loop-mediated isothermal amplification assay in South China. J. Phytopathol., 160, 248–250. Peng, J., Shi, M., Xia, Z., Huang, J., and Fan, Z. (2012). Detection of Cucumber mosaic virus isolates from banana by one-step reverse transcription loop-mediated isothermal amplification. Arch. Virol. 157, 2213–2217. https://doi.org/10.1007/s00705-012-1376-x Peng, J., Zhang, J., Xia, Z., Li, Y., Huang, J., and Fan, Z. (2012). Rapid and sensitive detection of Banana bunchy top virus by loop-mediated isothermal amplification. J. Virol. Methods 185, 254–258. https://doi.org/10.1016/j. jviromet.2012.06.026 Pyott, D.E., Sheehan, E., and Molnar, A. (2016). Engineering of CRISPR/Cas9-mediated potyvirus resistance in transgene-free Arabidopsis plants. Mol. Plant Pathol. 17, 1276–1288. https://doi.org/10.1111/mpp.12417 Qin, Z., Jin, L., Xu, F., Zhang, J., Ouyang, P., Chen, J. (2012). RNA3 of a Cucumber mosaic virus strain infecting Musa basjoo based on the molecular characterization of its 50 UTR. Advances in Intelligent and Soft Computing 134, 919–924. QPPR (2002). Queensland Plant Protection (Banana Pest Quarantine) Regulation 1999. Quito-Avila, D.F., Ibarra, M.A., Alvarez, R.A., Ratti, M.F., Espinoza, L., Cevallos-Cevallos, J.M., et al. (2013). First report of banana bract mosaic virus in ‘Cavendish’ banana in Ecuador. Plant Dis. 97, 1003. Rao, D.G. (1980). Studies on a new strain of banana mosaic virus in South India. In Proc. National Seminar on Banana Production Technology. Muthukrishnan, C.R., and Abdul Khader, J.B.M., eds. (Tamilnadu Agriculturual University, Coimbatore), pp. 155–159. Renaud-Gabardos, E., Hantelys, F., Morfoisse, F., Chaufour, X., Garmy-Susini, B., and Prats, A.C. (2015). Internal ribosome entry site-based vectors for combined gene therapy. World J. Exp. Med. 5, 11–20. https://doi. org/10.5493/wjem.v5.i1.11 Rodoni, B.C., Ahlawat, Y.S., Varma, A., Dale, J.L., and Harding, R.M. (1997). Identification and characterization of banana bract mosaicvirus in India. Plant Dis. 81, 669–672. Rodoni, B.C., Dale, J.L., and Harding, R.M. (1999). Characterization and expression of the coat proteincoding region of banana bract mosaic potyvirus, development of diagnostic assays and detection of the virus in banana plants from five countries in southeast Asia. Arch. Virol. 144, 1725–1737. Roossinck, M.J., Bujarski, J., Ding, S., Hajimorad, R., Hanad, K., Scott, S., et al. (1999).Family Bromoviridae. In Virus Taxonomy-Seventh Report of the International Committee on Taxonomy of Viruses. van Regenmorte, l.M.H.V., Fauquet, C.M., Bishop, D.H.L, eds. (Academic Press, San Diego), pp. 923–935. Sanford, J.C., and Johnston, S.A. (1985). The concept of parasite-derived resistance – deriving resistance genes from the parasite’s own genome. J. Theoret. Biol. 113, 395-405. Sasireka. T., and Selvarajan, R.(2014). Simultaneous detection of multi components of Banana bunchy top virus (BBTV). In XXIII National Conference of Indian Virological Society (IVS), ‘VIROCON 2014’ on ‘Recent

Trends in Virology Research in the Omics Era’, 18–20th December 2014, Tamil Nadu Agricultural University, Coimbatore, 2014. p. 80. Selvarajan, R., and Jeyabaskaran, K.J. (2006). Effect of banana bract mosaic virus (BBrMV) on growth and yield of cultivar nendran (plantain, AAB). Ind. Phytopathol. 59, 496–500. Selvarajan, R., Balasubramanian, V., and Sathiamoorthy, S. (2006). Vector transmission of banana bract mosaic and banana streak viruses in India. In Abstracts of XVI annual convention and international symposium on management of vector-borne viruses, ICRISAT, 7–10th February, 2006. p. 110. Selvarajan, R., Mary Sheeba, M., Balasubramanian, V., and Mustaffa, M.M. (2007). Molecular characterization and expression of coat protein gene of Cucumber mosaic virus infecting banana -TRY isolate. In National Conference on Banana: ‘Production and Utilization of Banana for Economic Livelihood and Nutritional Security’, 25–28th October 2007, Trichy, Tamil Nadu, 2007. p. 203. Selvarajan, R., and Balasubramanian, V. (2008). Banana viruses. In Characterization, Diagnosis and Management of Plant Viruses: 2. Rao, G.P., Myrta, A., and Ling, K, eds., (Studium press LLC, Texas), pp. 109–124. Selvarajan, R., Balasubramanian, V., Kavitha, K., Kavitha, K.S., Sathiamoorthy, S., and Ahlawat. Y.S. (2008). Detection of Banana bunchy top virus (BBTV) and Banana Streak Mysore Virus (BSMysV) by PCR: impact of storing virus infected banana samples. Ind. J. Virol. 19, 28–33. Selvarajan,R., Balasubramanian, V., Jeyabaskaran, K.J., Pandey, S.D., and Mustaffa, M.M. (2009). Management of banana bract mosaic disease through higher dose of fertilizer application in banana cv. Ney Poovan (AB). Ind. J. Horticulture 66, 301–305. Selvarajan, R., Mary Sheeba, M., Balasubramanian, V., Rajmohan, R., Lakshmi Dhevi, N., and Sasireka, T. (2010a). Molecular characterization of geographically different Banana bunchy top virus isolates in India. Ind. J. Virol. 21, 110–116. https://doi.org/10.1007/s13337010-0020-1 Selvarajan, R., Balasubramanian, V., Dayakar, S., Sathiamoorthy, S., and Ahlawat, Y.S. (2010b). Evaluation of immunological and molecular techniques for the detection of different isolates of Banana bunchy top virus in India. Ind.n Phytopath.y 63, 333. Selvarajan, R., Mary Sheeba, M., and Balasubramanian, V. (2011a). Simultaneous detection of episomal Banana streak Mysore virus and Banana bunchytop virus using multiplex RT–PCR. Curr. Sci. 10, 31–34. Selvarajan, R., Balasubramanian, V., Sheeba, M.M., Raj Mohan, R., and Mustaffa, M.M. (2011b). Virusindexing technology for production of quality banana planting material: a boon to the tissue-culture industry and banana growers in India. In Acta Horticulturae: 897. Van den Bergh, I., et al. eds., Proceedings from the International ISHS-ProMusasymp. on global perspectives on Asian challenges, pp. 463–469. Selvarajan, R., Balasubramanian,V., Jeyabaskaran, K.J., Pandey, S.D., and Mustaffa, M.M. (2012). Mitigating the effect of banana bract mosaic disease through application of increased doses of fertilizer in banana cultivar Robusta (AAA). J. Plant Dis. Scie. 7, 158–161.

274  | Selvarajan et al.

Selvarajan, R., and Balasubramanian, V. (2013a). Natural occurrence of Banana bunchy top virus in Ensete superbum in India. Ind. J. Virol. 24, 97–98. https://doi. org/10.1007/s13337-012-0123-y Selvarajan, R., and Balasubramanian, V. (2013b) Immunodiagnosis of Banana bract mosaic virus (BBrMV) using the polyclonal antiserum developed against recombinant coat protein. In Abstracts of the National Symposiumon Pathogenomics for diagnosis and Management of Plant Disease. Indian Phytopathology Society, Central Tuber Crops Research Institute, Thiruvananthapuram, India, 24–25 October, 2013, p. 77. Selvarajan, R., and Balasubramanian, V. (2014). Host interaction host–virus interactions inbanana-infecting viruses. In Plantvirus–Host Interaction Molecular Approaches And Viral Evolution, Gaur, R.K., ed. (Elsevier, Singapore), pp. 57–78. Selvarajan, R., Anuradha, C., Rajmohan, R., Mary Sheeba, M., and Balasubramanian. V. (2014). Agrobacteriummediated in planta transformation of hill banana for developing resistance to Banana bunchy top virus. Madras Agric. J. 101, 374–378. Selvarajan. R., Balasubramanian, V., and Sasireka, T. (2015). A simple, rapid and solvent free nucleic acid extraction protocol for detection of Banana bunchy top virus by polymerase chain reaction and loop-mediated isothermal amplification. Eur. J. Plant Pathol. 142, 389–396. Selvarajan. R., Balasubramanian, V., and Gayathrie, T. (2016).Highly efficient immunodiagnosis of episomal Banana Streak Mysore Virus (BSMYV) using polyclonal antibodies raised against recombinant viral associated protein. J. Phytopathol. 64, 497–508. Selvarajan. R., and Balasubramanian, V. (2017). First report of banana bract mosaic virus in banana in Assam, India. J. Plant Pathol. 99, 534. Selvarajan,R., Balasubramanian, V., Jeyabaskaran, K.J., and Mustaffa, M.M. (2017). Yield loss management through fertilizers in Banana bract mosaic virus affected French plantain cv. Nendran (AAB). Ind. J. Agric. Sci. 87,1055–1061. Sharma, S.K., Kumar, P.V., and Baranwal, V.K. (2014). Immunodiagnosis of episomal Banana streak MY virus using polyclonal antibodies to an expressed putative coat protein. J. Virol. Methods 207, 86–94. https://doi. org/10.1016/j.jviromet.2014.06.018 Sharman, M., Thomas, J.E., and Dietzgen, R.G. (2000a). Development of a multiplex immunocapture PCR with colorimetric detection for viruses of banana. J. Virol. Methods 89, 75–88. Sharman, M., Gambley, C.F., Oloteo, E.O., Abgona, R.V. J., and Thomas, J.E. (2000b). First record of natural infection of abaca (Musa textilis) with banana bract mosaic potyvirus in the Philippines. Australasian Plant Pathol.29, 69. Sharman, M., Thomas, J.E., Skabo, S., and Holton, T.A. (2008). Abacá bunchy top virus, a new member of the genus Babuvirus (family Nanoviridae). Arch. Virol. 153, 135–147. https://doi.org/10.1007/s00705-007-1077-z Shekhawat, U.K.S., Ganapathi, T.R., and Hadapad, A.B. (2012). Transgenic banana plants expressing siRNAs targeted against viral replication initiation gene display

high-level resistance to Banana bunchy top virus infection. J. Gen. Virol. 93, 1804–1813. Singh, Z., Jones, R.A.C., and Jones, M.G.K.(1995). Identification of Cucumber mosaic virus subgroup 1 isolates from banana plants affected by infectious chlorosis disease using RT–PCR. Plant Dis. 79, 713–716. Siljo, A., Bhat, A.I., Biju, C.N., and Venugopal, M.N. (2012). Occurrence of Banana bract mosaic virus on cardamom. Phytoparasitica 40, 77–85. Siljo, A., Bhat, A.I., and Biju, C.N. (2014). Detection of Cardamom mosaic virus and Banana bract mosaic virus in cardamom using SYBR Green based reverse transcription-quantitative PCR. Virus Disease 25, 137–141. https://doi.org/10.1007/s13337-013-0170-z Siljo, A., and Bhat, A.I. (2014). Reverse transcription loop-mediated isothermal amplification assay for rapid and sensitive detection of banana bract mosaic virus in cardamom (Elettariacardamomum). Eur. J. Plant Pathol. 138, 209–214. Srivastava, A., and Raj, S.K. (2008). Coat protein-mediated resistance to an Indian isolate of the Cucumber mosaic virus subgroup IB in Nicotiana benthamiana. J. Biosci. 33, 249–257. Srivastava, A., Raj, S.K., Haq, Q.M., Srivastava, K.M., Singh, B.P., and Sane, P.V. (1995). Association of a Cucumber mosaic virus strain with mosaic disease of banana, Musa paradisiaca – an evidence using immuno/nucleic acid probe. Ind. J. Exp. Biol. 33, 986–988. Stainton, D., Kraberger, S., Walters, M., Wiltshire, E.J., Rosario, K., Halafihi, M., Lolohea, S., Katoa, I., Faitua, T. H., Aholelei, W., et al. (2012). Evidence of inter-component recombination, intra-component recombination and reassortment in Banana bunchy top virus. J. Gen. Virol., 93, 1103–1119. Stainton, D., Martin, D.P., Muhire, B.M., Lolohea, S., Halafihi, M., Lepoint, P., Blomme, G., Crew, K.S., Sharman, M., Kraberger, S., et al. (2015). The global distribution of Banana bunchy top virus reveals little evidence for frequent recent, human-mediated long distance dispersal events. Virus Evol. 1, vev009. https:// doi.org/10.1093/ve/vev009 Su, H.J., Hwang, A.S., Lee, S.Y., and Chao, C.P. (2007). Conservation, disease indexing and utilization of pathogen free citrus and banana genetic resources in Taiwan. In International training workshop on the conservation and utilization of Tropical/Subtropical plant genetic resources, pp. 1–24. Tamura, K., Stecher, G., Peterson, D., Filipski, A., and Kumar, S. (2013). MEGA6: Molecular Evolutionary Genetics Analysis version 6.0. Mol. Biol. Evol. 30, 2725–2729. https://doi.org/10.1093/molbev/mst197 Thomas, J.E., and Dietzgen, R.G. (1991). Purification, characterization and serological detection of virus-like particles associated with banana bunchy top disease in Australia. J. Gen. Virol. 72, 217–224. https://doi. org/10.1099/0022-1317-72-2-217 Thomas, J.E., Iskra-Caruana, M.L., and Jones, D.R. (1994). Banana bunchy top disease. Musa disease fact sheet No. 4. Montpellier, France: INIBAP, pp. 2. Thomas, J.E., Geering, A.D., Gambley, C.F., Kessling, A.F., and White, M. (1997). Purification, properties, and diagnosis of banana bract mosaic potyvirus

Viruses Infecting Banana and their Transgenic Management |  275

and its distinction from abaca mosaic potyvirus. Phytopathology 87, 698–705. https://doi.org/10.1094/ PHYTO.1997.87.7.698 Thottappilly, G., Dahal, G., and Lockhart, B.E.L. (1998). Studies on a Nigerian isolate of banana streak badnavirus. Purification and enzyme-linked immunoassay. Ann. Appl. Biol. 132, 253–261. Tavanlar, M.A.T. (2007). Development of a banana (Musa sp.) transformation system for the production of transgenes banana expressing viral genes conferring resistance to banana bract mosaic virus. University Library, University of the Philippines at Los Baños. Tsao, T.T.H. (2008). Towards the development of transgenic Banana bunchy top virus (BBTV)- resistant banana plants: Interference with replication. PhD thesis, Queensland University of Technology, Brisbane. Tyagi, H., Rajasubramaniam, S., Rajam, M.V., and Dasgupta, I. (2008).RNA-interference in rice against Rice tungro bacilliform virus results in its decreased accumulation in inoculated rice plants. Transgenic Res. 17, 897–904. https://doi.org/10.1007/s11248-008-9174-7 Vanderschuren, H., Akbergenov, R., Pooggin, M.M., Hohn, T., Gruissem, W., and Zhang, P. (2007). Transgenic cassava resistance to African Cassava mosaic virus is enhanced by viral DNA-A bidirectional promoterderived siRNAs. Plant Mol. Biol. 64, 549–557. https:// doi.org/10.1007/s11103-007-9175-6 Vishnoi, R., Raj, S.K., and Prasad, V. (2009). Molecular characterization of an Indian isolate of Banana bunchy top virus based on six genomic DNA components. Virus Genes 38, 334–344. https://doi.org/10.1007/s11262009-0331-8 Vishnoi, R. Kumar, S.,and Raj, S.K. (2013). Molecular characterization of a Cucumber mosaic virus isolate associated with mosaic disease of banana in India. Phytoparasitica 41, 545–555. Wanitchakorn, R., Harding, R.M., and Dale, J.L. (1997). Banana bunchy top virus DNA-3 encodes the viral coat protein. Arch. Virol. 142, 1673–1680. Wanitchakorn, R., Harding, R.M., and Dale, J.L. (2000a). Sequence variability in the coat protein gene of two groups of banana bunchy top isolates. Arch. Virol. 145, 593–602. Wanitchakorn, R., Hafner, G.J., Harding, R.M., and Dale, J.L. (2000b). Functional analysis of proteins encoded by bananabunchy top virus DNA-4 to-6. J. Gen. Virol. 81, 299–306.

Wang, I.C., Sether, D.M., Melzer, M.J., Borth, W.B., and Hu, J.S. (2010). First report of banana bract mosaic virus in flowering ginger in Hawaii. Plant Dis. 94, 921. Watanabe, S., Greenwell, A.M., and Bressan, A. (2013). Localization, concentration, and transmission efficiency of Banana bunchy top virus in four asexual lineages of Pentalonia aphids. Viruses 5, 758–776. https://doi. org/10.3390/v5020758 WeiGang, W., Zhang, T., and Ying, Y. (2009).Detection of banana bract mosaic virus by real time fluorescent RT-PCR. Acta Phytophylacica Sinica 36, 417–420. Wickramaarachchi, W.A.R.T., Shankarappa, K.S., Rangaswamy, K.T., Maruthi, M.N., Rajapakse, R.G.A.S., and Ghosh, S. (2016). Molecular characterization of Banana bunchy top virus isolate from Sri Lanka and its genetic relationship with other isolates. Virus Disease 27, 154–160. https://doi.org/10.1007/s13337-0160311-2 Wu, R.Y., and Su, H.J. (1990a). Purification and characterization of Banana bunchy top virus. J. Phytopathol. 128, 153–160. Wu, R.Y., and Su, H.J. (1990b). Production of monoclonal antibodies against Banana bunchy top virus and their use in enzyme-linked immunosorbent assay. J. Phytopathol. 128, 203–208. Xie, W.S., and Hu, J.S. (1995). Molecular cloning, sequence analysis and detection of Banana bunchy top virus in Hawaii. Phytopathology 85, 339–347. Yu, N.T., Zhang, Y.L., Feng, T.C., Wang, J.H., Kulye, M., Yang, W.J., Lin, Z.S., Xiong, Z., and Liu, Z.X. (2012). Cloning and sequence analysis of two Banana bunchy top virus genomes in Hainan. Virus Genes 44, 488–494. https://doi.org/10.1007/s11262-012-0718-9 Zhang, J., Borth, W.B., Lin, B., Dey, K.K., Melzer, M.J., Shen, H., Pu, X., Sun, D., and Hu, J.S. (2016). Deep sequencing of banana bract mosaic virus from flowering ginger (Alpinia purpurata) and development of an immunocapture RT-LAMP detection assay. Arch. Virol.161, 1783–1795. https://doi.org/10.1007/ s00705-016-2830-y Zhuang, J., Coates, C.J., Mao, Q., Wu, Z., and Xie, L. (2016). The antagonistic effect of Banana bunchy top virus multifunctional protein B4 against Fusarium oxysporum. Mol. Plant Pathol. 17, 669–679. https:// doi.org/10.1111/mpp.12319

Virus-induced Gene Silencing (VIGS) and its Applications

14

Deep Ratan Kumar, Tejbhan Saini and Radhamani Anandalakshmi*

Plant–Virus Interactions Laboratory, Mahyco Research Centre, Jalna, India. *Correspondence: [email protected] https://doi.org/10.21775/9781910190814.14

Abstract During the last two decades, virus-induced gene silencing (VIGS) has emerged as the most powerful and cost effective tool to determine gene functions in a relatively shorter period of time. VIGS has been used for both forward and reverse genetics studies for functional analysis of a large number of genes in both monocot and dicot plants. Various VIGS vectors have been developed, tested and modified to down-regulate the endogenous gene expression effectively in a sequence-specific manner in model plants, cereals, vegetables, fibre crops, fruits and timber trees. This review focuses on VIGS as a technique for inducing gene silencing in plants and its applications in understanding the role of genes controlling cellular and metabolic pathways, development, abiotic and biotic stress in economically important crop species. Introduction Advances in genome sequencing technologies such as next generation sequencing (NGS), expressed sequence tag (EST) and microarrays analysis have generated large-scale genome information of different plant species. However, the characterization of genes from whole genome sequences is essential to understand their biological function and to exploit potential applications. Even in a widely studied model plant like Arabidopsis, having a relatively small genome (≈ 135 Mbp), less than 10% of the predicted genes have been identified and functionally characterized (Matthew, 2004). To investigate

the biological roles of genes, they could be either over expressed or knocked down and their effects studied at phenotypic and molecular levels. Conventional methods employed to knock off gene expression by forward genetics include chemical mutagenesis (EMS, MMS), physical mutagenesis (radiation mutagenesis like X-rays, gamma rays, fast neutrons), and T-DNA insertional mutagenesis. However, these methods are time-consuming and difficult to deploy in many crop plants due to polyploidy, large genome size, low transformation efficiency, low levels of inheritance and unwanted off-target effects. Often, mutations fail to either completely inactivate the gene or result in embryo lethality. Mutations induced in genes belonging to a multigene family, frequently fail to show associated phenotype due to complementation by proteins encoded by other members of the gene family having redundant function. Moreover, mutations occur randomly in a genome, it is difficult to assign plant phenotypes to a mutation in a particular gene. RNA interference (RNAi) or RNA silencing is one effective way to study gene function. RNAi is a conserved phenomenon in plants, animals and lower organisms which targets and degrades RNAs in a nucleotide sequence specific manner (Waterhouse et al., 2001; Vance and Vaucheret, 2001; Voinnet, 2001, 2005). This involves silencing the expression of the targeted genetic elements, improperly matured RNAs and foreign nucleic acids including viruses and transposons (Lacomme, 2015). RNA silencing is exploited to knock out gene expression in plants, both in transient as well as stable manner

278  | Kumar et al.

(Baulcombe, 1999). Virus-induced gene silencing (VIGS) exploits RNA silencing mediated natural antiviral defence mechanism of plants to silence genes of interest in hosts (Baulcombe, 1999; Liu et al., 2002a; Dinesh-Kumar et al., 2003; Burch-Smith et al., 2004; Robertson, 2004; Becker and Lange, 2010; Senthil-Kumar and Mysore, 2011a; Dawson and Folimonova, 2013; Dolja and Koonin, 2014). Virus-infected plants trigger RNA silencing in hosts that is specifically targeted against the invader. By engineering a virus to carry a fragment of a host gene transcript, RNA based defence against the virus can be directed to target any endogenous gene. The infectious clones derived from viruses are called VIGS vectors. By VIGS, expression of a host gene could be switched off in a very short period and it has the potential to silence multi copy genes at once. Partial homologues of genes that vary up to 20% like that of the tobacco ribulose bisphosphate carboxylase (Rubisco) family (Dean et al., 1989) or MAP kinase genes (Liu et al., 2002a,b) can also be targeted by this method. VIGS finds application in both reverse and forward genetic screens (Baulcombe, 1999; Robertson, 2004). It has accelerated the identification of genes involved in cellular, developmental, metabolic processes, biotic and abiotic stress pathways. Among the various conventional and RNAi based methods, in plants VIGS is most effective for knocking off gene function as it neither requires stable transformation nor selection. It has made possible gene function analysis in a multitude of annual and perennial plants with limited transformation potential and high ploidy levels. Mechanism of VIGS VIGS is a homology-dependent RNA-mediated gene silencing technique (Baulcombe, 1999). The term ‘VIGS’ was first coined by A. van Kammen to describe the phenomenon of recovery from virus infection in plants (van Kammen, 1997). Later on studies have shown that recovery from virus infections occur when plants are able to mount strong RNA silencing based defence and degrade the infecting viral genome, thereby, diminishing the viral titre and the disease symptoms. This process involves the recognition of double-stranded RNA (dsRNA) made during the replication of the infecting DNA or RNA viruses. Most plant pathogenic

viruses belong to single-stranded positive sense RNA group. Double-stranded RNA is formed either from RNA intermediates pairing between positive (+) and negative (-) ssRNA strands, or overlap of sense and antisense RNAs, or folded secondary structures of viral RNAs. DNA viruses also induce gene silencing via aberrant RNAs formed during replication. VIGS is initiated when dsRNA intermediates of viral replication is recognized by Dicer-like endonucleases, RNAse III like enzymes and degraded into small interfering (si) RNAs of 21 to 25 nucleotides. Plant RNA-dependent RNA polymerase 1 (RDRP1), RDRP2 and RDRP6 use single-stranded RNA as a template and siRNA as primers to produce more dsRNA to amplify the process. Guide strand of the viral siRNAs then associate with argonaute AGO1 family proteins and form a complex known as RNA-induced silencing complex (RISC). The guide strand in RISC complex is complementary to the target RNA and helps in identification, priming and cleavage of complementary RNA sequences within the cell (Fagard et al., 2000; Morel et al., 2002). Although initially it was believed that post-transcriptional gene silencing (PTGS) is involved in RNA silencing based defence, transcriptional gene silencing via methylation is also associated with VIGS ( Jones et al., 2001). Once triggered, VIGS stays on for a short period of 3–4 weeks after which the plants begin to recover partially or completely from silencing (Marathe et al., 2000; Ratcliff et al., 2001; Hiriart et al., 2003; Ryu et al., 2004). However, by suitably modifying plant growth conditions that favour viral multiplication, VIGS can be maintained for several months (Fu et al., 2006; Tuttle et al., 2008; SenthilKumar and Mysore, 2011b, 2014) and silencing may even get transmitted to the next generation (Senthil-Kumar and Mysore, 2011b). VIGS vectors Several RNA and DNA viruses have been modified into VIGS vectors for gene expression in plant species. These vectors are usually standard binary Ti-plasmids into which the viral genome and a homologous fragment of the host gene to be targeted are cloned. One of the challenges in developing VIGS system is to identify those viruses that can move systemically but produce very mild or no symptoms in host plants. Occasionally, viral

Virus-Induced Gene Silencing and its Applications |  279

genomes are modified by deleting or altering genes which are responsible for symptoms. DNA (DNA viruses) or cDNA (RNA viruses) of viral genomes are cloned into binary vectors under a constitutive promoter like CaMV35S promoter with addition of convenient multiple cloning sites (MCS) to facilitate insertion of host gene fragments to be targeted for silencing (Voinnet, 2001; Liu et al., 2002a,b). The first successful application of VIGS was reported in 1995 using a Tobacco mosaic virus (TMV) derived vector where silencing of the endogenous phytoene desaturase (PDS) gene in Nicotiana benthamiana, resulted in photo bleached phenotype in leaves (Kumagai et al., 1995). Potato virus X (PVX) based vectors were developed by Chapman et al., 1992 and Ruiz et al., 1998 to silence genes in Nicotiana sp. VIGS vectors have also been derived from different DNA viruses like Tomato golden mosaic virus (TGMV) and RNA viruses such as Barley stripe mosaic virus (BSMV), etc. Some viruses produce severe symptoms in certain hosts while others have a very narrow host range. Hence it is essential to derive VIGS vectors from viruses that have a wide host range but are symptomless in most plants. It has been observed that viruses that do not encode suppressors of gene silencing or have weak suppressors, make better VIGS vectors that are capable of maintaining silencing in host plants for longer periods of time (Baulcombe, 1999; Li and Ding, 2001; Cao et al., 2005). Potato virus X (PVX), Barley stripe mosaic virus (BSMV), Tobacco rattle virus (TRV), etc. are few such RNA viruses. On the other hand, Tobacco etch virus (TEV), Potato virus Y (PVY), Cucumber mosaic virus (CMV) and Tomato bushy stunt virus (TBSV) would make poor VIGS vectors as they encode strong suppressors of silencing. For example: TEV and PVY encode HC-Pro, a potent silencing suppressor that directly interferes with the PTGS allowing virus to escape silencing (Anandalakshmi et al., 1998; Kassachu and Carrington, 1998). Similarly, CMV encodes 2b (Li et al., 1999) and TBSV encodes P19 (Lakatos et al., 2004; Scholthof, 2006) which are strong silencing suppressors. In addition, viruses that are capable of silencing the expression of genes in the meristematic tissues are the most desirable. Tomato black ring virus (TBRV) and TRV are seed and pollen transmitted viruses and are able to penetrate the meristems (Lister and Murani, 1967; Baulcombe, 1999).

Both RNA and DNA viruses are capable of inducing homology-dependent systemic silencing of endogenous genes. However, they vary in infection patterns, vector transmissibility and interactions with the host’s defence mechanism. Similarly, the extent of spread of silencing and the severity of the symptoms induced by a recombinant virus also have significant variations in different host plants depending on the plant–virus interactions. More number of VIGS vectors are derived from positive, single-stranded RNA viruses than negative stranded RNA or DNA viruses. DNA viruses were not preferred as expression vectors of full length cDNAs due to their size constraints for movement. However, in 1998, Kjemtrup et al. demonstrated silencing of the sulphur (su) and luciferase (luc) gene by modifying the DNA-A component of a single-stranded DNA virus – Tomato golden mosaic virus (TGMV). In subsequent years, VIGS vectors of geminiviruses such as, Cabbage leaf curl virus (CaLCuV), Cotton leaf crumple virus (CLCrV), African Cassava mosaic virus (ACMV), and Tomato yellow leaf curl virus (TYLCV) and their associated satellites have been developed for silencing genes in plants like N. benthamiana, Arabidopsis, cassava, cotton and tomato (Turnage et al., 2002; Fofana et al., 2004; Tao and Zhou, 2004; Tuttle et al., 2008). A detailed list of various VIGS vectors developed so far, and host plants and endogenous genes targeted is given in Table 14.1. TRV and BSMV based VIGS vectors are extensively used for functional analysis and are described in more detail here. VIGS vectors derived from CMV, Bean pod mottle virus (BPMV), Apple latent spherical virus (ALSV), Pea early browning virus (PEBV), Tobacco ringspot virus (TRSV), White clover mosaic virus (WCIMV), Subterranean clover mottle virus (SCMoV), Soybean yellow common mosaic virus (SYCMV) and Sunhemp mosaic virus (SHMV) are described in detail in Chapter 15. Tobacco rattle virus (TRV) based VIGS system that induces efficient silencing of endogenous genes was developed and modified by two research groups (Ratcliff et al., 2001; Liu et al., 2002b). TRV belongs to the genus Tobravirus and is mostly transmitted by soil nematodes (Matthews, 1991; Visser and Bol, 1999). TRV is a single-stranded, positivesense RNA virus with bipartite genome (RNA1 and RNA2). TRV-based VIGS system uses both RNA1 and RNA2 components. The RNA1 encodes

Table 14.1  VIGS vector and plant species used for gene silencing No.

VIGS vector

Genus

Host species silenced

Target gene(s) silenced

Cheravirus

N. tabacum, N. benthamiana GFP

References

RNA virus 1

2

Apple latent spherical virus (ALSV)

Barley stripe mosaic virus (BSMV)

Hordeivirus

Yaegashi et al. (2007)

A. thaliana, Cucurbits, Legumes, Tobacco, Tomato

CH42, PDS, RC33Y1, SU, PCNA, GFP

Igarashi et al. (2009)

Rosa rugosa

PDS

Ito et al. (2012)

Prunus (apricot and Japanese apricot)

PDS

Kawai et al. (2014)

Glycine max, apple

PDS; ERA1A, ERA1B

Yamagishi and Yoshikawa (2009, 2013), Ogata et al. (2017)

Apple, pear and Japanese pear

RbcS, CPN60a, EF-1a, Actin, TFL1; SQS1

Sasaki et al. (2011), Navarro Gallón et al. (2017)

H. vulgare

PDS, P23k, NecS1; RAR1, SGT1, HSP90; MLO, EXPB7, Bln1

Holzberg et al. (2002), Lacomme et al. (2003), Hein et al. (2005), Bruun-Rasmussen et al. (2007), Oikawa et al. (2007), Zhang et al. (2009a), Delventhal et al. (2011), He et al. (2015b), Xu et al. (2015), Barciszewska-Pacak et al. (2016)

T. aestivum, H. vulgare

PDS, ChlH, 20S-β7; Lr21, RAR1, SGT1, HSP90; GFP, DMC1; Mla13

Scofield et al. (2005), Tai et al. (2005), Cakir et al. (2010), Bennypaul et al. (2012), Lee et al. (2015)

Costus spicatus, Zingiber officinale; Z. zerumbet

PDS

Renner et al. (2009), Mahadevan et al. (2015b)

Brachypodium distachyon, H. vulgare, T. aestivum; Zea mays, A. sativa, A. strigosa, N. benthamiana

PDS, IPS1, PHR1, PHO2; GTF1, GTF2; PDS, ChlH, TK, PMR5

Nowara et al. (2010), Pacak et al. (2010), Yuan et al. (2011)

T. aestivum

WRKY53, PAL, GFP, PDS, 1Bx14, Myb3, TBF3, GLY1, EIL1, GLI1, Era1, Cyp707a, Sal1, SCL14, CL14010, CL12788, CL176, Unigene 16777, CL8746, Unigene 10196, NAC1, PR1, Pm21, RSR1

van Eck et al. (2010), Tufan et al. (2011), Ma et al. (2012), Scofield and Brandt (2012), Yang et al. (2013), Duan et al. (2013), Kang et al. (2013), Feng et al. (2013), Manmathan et al. (2013), Chen et al. (2015), Feng et al. (2015), Wang et al. (2015a), Buhrow et al. (2016), He et al. (2016), Liu et al. (2016a)

Haynaldia villosa

GFP, PDS, LRR

Wang et al. (2010b)

Secale cereale

Bx1

Groszyk et al. (2017)

Table 14.1  Continued No.

VIGS vector

Genus

Host species silenced

Target gene(s) silenced

3

Bean pod mottle virus (BPMV)

Comovirus

Glycine max

PDS, MPK4, SGT1A, SGT1B, RPS6, Zhang et al. (2006a, 2009a, 2010), Juvale et al. RPS13, Actin, GFP, Bar, RBOH (2012), Kandoth et al. (2013), Ranjan et al. (2017)

Phaseolus vulgaris

GFP, Nodulin 22, Stearoyl-acyl carrier protein desaturase, PDS

Zhang et al. (2010), Diaz-Camino et al. (2011), Pflieger et al. (2014)

Pisum sativum

PDS, KOR1

Meziadi et al. (2016)

O. sativa, H. vulgare, Z. mays, Festuca arundinacea, N. benthamiana

PDS, Actin 1, Rubisco activase

Ding et al. (2006), Scofield and Nelson (2009)

Z. mays

PDS, YFP, terpene synthase (tps6 ⁄ 11), ecb, bti, bi-1, Elc, GFP

van der Linde et al. (2011, 2013), Benavente et al. (2012), Zhu et al. (2014)

Sorghum

Cs1A, Cs2A, LTP1, ZnTF1, CD1, DEFL1, CK2

Biruma et al. (2012)

4

5

Brome mosaic virus (BMV)

Cucumber mosaic virus (CMV)

Bromovirus

Cucumovirus

References

N. benthamiana

GFP

Otagaki et al. (2006)

Glycine max

CHS, F3’H

Nagamatsu et al. (2007)

Petunia, Tomato

LeSPL-CNR

Kanazawa et al. (2011)

Antirrhinum majus

ANT

Kim et al. (2011)

Tomato, Chilli pepper, N. benthamiana

PDS

Hong et al. (2012)

Z. mays

PDS, IspH, ATG3, ATG8a,

Wang et al. (2016b)

Lilium leichtlinii

PDS

Tasaki et al. (2016)

Phalaenopsis orchids; P. aphrodite

UFGT3, MADS5, MADS6, TF15

Lu et al. (2007, 2012), Hsieh et al. (2013)

H. vulgare, T. aestivum, Setaria italica

PDS, ChlH, CLA1, IspH

Robertson et al. (2000), Liu et al. (2016b)

6

Cymbidium mosaic virus (CymMV)

7

Foxtail mosaic virus (FoMV) Potexvirus

Sweetcorn

PDS, les22, ij, bm3

Mei et al. (2016)

8

Grapevine virus A (GVA)

Vitivirus

Vitis vinifera

PDS

Muruganantham et al. (2009)

9

Grapevine Algerian latent virus (GALV)

Tombusvirus

Grapevine, N. benthamiana

GFP, ChlH

Park et al. (2016)

10

Grapevine leafrollassociated virus-2 (GLRaV-2)

Closterovirus

V. vinifera

PDS, GFP, Chl1

Kurth et al. (2012)

Potexvirus

Table 14.1  Continued No.

VIGS vector

Genus

Host species silenced

Target gene(s) silenced

References

11

Pea early browning virus (PEBV)

Tobravirus

P. sativum, M. truncatula, L. odorata

PDS, UNI, Nin, KOR1, PsPIP2;1, PEAM4

Constantin et al. (2004, 2008), Grønlund et al. (2008, 2010), Chen et al. (2014), Grønlund (2015), Song et al. (2016)

12

Poplar mosaic virus (PopMV)

Carlavirus

N. benthamiana

GFP

Naylor et al. (2005)

13

Potato virus A (PVA)

Potyvirus

N. benthamiana

GFP

Gammelgard et al. (2007)

14

Potato virus X (PVX)

Potexvirus

N. benthamiana

PDS, SSU, GFP, HSP90, CYP51, FtsH, 14-3-3, GUS, Chl1, RLIh

Ruiz et al. (1998), Angell & Baulcombe (1999), Braunstein et al. (2002), Saitoh and Terauchi (2002), Burger et al. (2003), Kanzaki et al. (2003), Lu et al. (2003a), Petersen and Albrechtsen (2005), Hirano et al. (2007)

N. benthamiana, S. tuberosum, S. bulbocastanum

PDS

Faivre-Rampant et al. (2004), Lacomme and Chapman (2008)

Tomato

CRY2

Giliberto et al. (2005)

15

Plum pox virus (PPV)

Potyvirus

N. benthamiana

RDR6, Replicase transgene

Voinnet (2001), Vaistij et al. (2009)

16

Prunus necrotic ringspot virus (PNRSV)

Ilarvirus

N. benthamiana, peach

eIF(iso)4E

Cui and Wang (2017)

17

Sun-hemp mosaic virus (SHMV)

Tobamovirus

N. benthamiana, M. truncatula

GFP, PDS, Chlorata 42

Várallyay et al. (2010)

18

Sweet potato feathery mottle potyvirus (SPFMV)

Potyvirus

Sweet potato

GFP transgene

Sonoda and Nishiguchi (2000a, b)

19

Tobacco etch virus (TEV)

Potyvirus

N. benthamiana

PDS

Voinnet (2001)

20

Tobacco mosaic virus (TMV)

Tobamovirus

Allium chinense

PDS

Chen et al. (1996)

21

Tobacco necrosis virus A (TNV-A)

Necrovirus

N. benthamiana

PDS, Su, ChlH

Gao et al. (2011c)

22

Tobacco rattle virus (TRV)

Tobravirus

N. benthamiana

PDS, RAR1, EDS1, NPR1/NIM1, DEF, MADS1, PMT, SERK1, Chl, LCY1, LCY2, Bip4, Bip5, Ubiquitin E2, FRO1; RRP41, RRP43

Ratcliff et al. (2001), Liu et al. (2002b, 2004b), Dong et al. (2007), Senthil-Kumar et al. (2007), Liu and Page (2008), Velasquez et al. (2009), Hayward et al. (2011), Mantelin et al. (2011), Yan et al. (2012), Shi et al. (2014), Moon et al. (2016), Conti et al. (2017), Gama et al. (2017), Zhou and Zeng (2017)

N. benthamiana, N. tabacum PDS, ChlH

Metzlaff (2002), Hiriart et al. (2003), Robertson (2004)

Table 14.1  Continued No.

VIGS vector

Genus

Host species silenced

Target gene(s) silenced

References

22

Tobacco rattle virus (TRV)

Tobravirus

Tomato

PDS, CTR1, CTR2, RbcS, EIN2, LEA4, SGT1, RAR1, HSP90, TAPGs, EXP11, EXP12, CEL1, CEL2, TomloxC, SlODO1, HT1, SERK1, CHS, RIN, ACS, ACO1, Chl, Del, Ros1, NCED, CYP707A

Liu et al. (2002a), Dinesh-kumar et al. (2003), Ryu et al. (2004), Fu et al. (2005, 2006), Rotenberg et al. (2006), Senthil-Kumar and Uday-Kumar (2006), Bhattarai et al. (2007), Jiang et al. (2008, 2011), Orzaez et al, (2009), Velasquez et al. (2009), Eybishtz et al. (2010), Li et al. (2011b), Mantelin et al. (2011), Quadrana et al. (2011), Bachan and Dinesh-kumar (2012), Yan et al. (2012), FernandezMoreno et al. (2013), Ji et al. (2014)

Chilli pepper

CYP1, PDS, RbcS, PDS, Chl, Ccs, Psy, Lcyb, Crtz, An2, Capsaicin synthase

Chung et al. (2004), Ryu et al. (2004), Kim et al. (2006b, 2017), Yan et al. (2012), Tian et al. (2014b)

N. attenuate, N. tabacum, N. PMT, TI, Germin like protein, Indole- Saedler and Baldwin (2004), Mase et al. (2012), 3-acetic acid-amido synthetase Zhang and Thomma (2014), Groten et al. (2015) umbratica GH3.9, CCaMK, Ve1, MEK2, SIPK S. bulbocastanum, S. nigrum, S. okadae, S. pimpinellifolium, S. rostratum; S. tuberosum, S. venturii, Egg plant

ChIH, PDS, MPK1, RB, R1, Rx, StWIPK, StMKK6,

Brigneti et al. (2004), Ryu et al. (2004), Hartl et al. (2008), Yan et al. (2012), Li et al. (2014), Dobnik et al. (2016), Meng et al. (2016)

Petunia hybrida

PDS, ChlH, CHS, MKS1, PhOBF1

Chen et al. (2004), Spitzer et al. (2007), Jiang et al. (2011), Broderick and Jones (2014), Gargul et al. (2015), Sun et al. (2017a)

Opium and California poppy PDS, CYP719A21

Hileman et al. (2005), Wege et al. (2007), Dang and Facchini (2014)

Arabidopsis

PDS, CH42, GFP, Cullin 1, RPS2, LOX, TGG, AGO1, AGO2, AGO4, NRPD1a

Burch-Smith et al. (2006), Wang et al. (2006), Cai et al. (2007), Quadrana et al. (2011), Zheng et al. (2011), Ma et al. (2015), Bilichak and Kovalchuk (2017)

Hyoscyamus sp. (Egyptian henbane)

CYP80F1

Li et al. (2006b)

Aquilegia vulgaris

PDS, ANS, PISTILLATA, APETALA3, PISTILLATA

Gould and Kramer (2007)

Rosa hybrida

PDS, PIP

Ma et al. (2008, Tian et al. (2014a)

Jatropha curcas

PDS, PCNA, CH42

Ye et al. (2009)

Vaccinium myrtillus

TDR4

Jaakola et al. (2010)

Table 14.1  Continued No.

VIGS vector

Genus

Host species silenced

Target gene(s) silenced

References

22

Tobacco rattle virus (TRV)

Tobravirus

Brassica nigra

PDS

Zheng et al. (2010)

Thalictrum dioicum

PDS

Di Stilio et al. (2010)

Gossypium sp, G. raimondii, NDR1, MKK2, KATANIN, WRINKLED1, CLA1, ECR, Cla1, G. barbadense MPK9, MPK13, MPK25, PDS, CLA1, ANS, ANR

23

Tobacco ringspot virus (TRSV)

Nepovirus

Gao et al. (2011b), Qu et al. (2012), Gao and Shan (2013), Pang et al. (2013), Zhang et al. (2014b), Mustafa et al. (2016, 2017)

Fragaria ananassa

PYR1, GFP

Catharanthus roseus

Vindoline biosynthetic genes, Iridoid Liscombe and, O’Connor (2011), Carqueijeiro et al. oxidase, PDS (2015)

Gerbera hybrida

PDS, Chl, CHS, GLO1

Deng et al. (2012)

Cysticapnos vesicaria

PDS, FLO

Hidalgo et al. (2012)

Aquilegia coerulea

ANS, PDS

Sharma and Kramer (2013)

Cherry

PacMYBA, PacNCED1, PacCYP707A1-4

Shen et al. (2014), Li et al. (2015b)

Dhatura stramonium

PDS

Eftekhariyan et al. (2014)

Dendranthema grandiflorum

GFP

Tian et al. (2014a)

Gladiolus hybridus

PDS

Zhong et al. (2014)

Mimulus guttatus

PDS, CYC1, CYC2

Preston et al. (2014)

Phelipanche aegyptiaca

CCD7, CCD8

Aly et al. (2014)

Physalis floridana

PDS, MPF2, MPF3

Zhang et al. (2014a)

Striga hermonthica

PDS

Kirigia et al. (2014)

Populus euphratica, P.x canescens

PDS

Shen et al. (2015)

Piper colubrinum

Osmotin

Anu et al. (2015)

Rauwolfia tetraphylla, Rauwolfia serpentina

PDS

Corbin et al. (2017)

Spinacia oleracea

PDS

Lee et al. (2017)

Wheat, Maize

PDS, MLO

Zhang et al. (2017)

N. benthamiana, Arabidopsis, Cucurbits, Legumes

PDS

Zhao et al. (2016)

Chai et al. (2011), Tian et al. (2014a)

Table 14.1  Continued No.

VIGS vector

Genus

Host species silenced

Target gene(s) silenced

References

24

Tomato bushy stunt virus (TBSV)

Tombusvirus

L. esculentum; N. benthamiana

GFP

Hou and Qiu (2003), Pignatta et al. (2007)

25

Tomato mosaic virus (ToMV)

Tobamovirus

N. benthamiana

PDS

Kumagai et al. (1995)

26

Turnip yellow mosaic virus (TYMV)

Tymovirus

Arabidopsis

PDS, LEAFY

Pflieger et al. (2008), Jupin (2013)

27

White clover mosaic virus (WClMV)

Potexvirus

P. sativum, Vicia faba, M. littoralis, M. truncatula

PDS

Ido et al. (2012)

RNA satellite virus 28

Bamboo mosaic virus (BaMV) and its satellite RNA (satBaMV)

Potexvirus

N. benthamiana; Brachypodium distachyon

SU, GFP, HSP90, HSP70

Liou et al. (2014)

29

STMV (Satellite of Tobacco mosaic virus) vector with TMV helper virus (SVISS)

Tobamovirus

N. tabacum

PDS, ChsA, RbcS, CesA, TK, ALS, PPX, Gln, RpII, Act, NPK1, PARP

Gossele et al. (2002)

DNA virus 30

Abutilon mosaic virus (AbMV)

Begomovirus

N. benthamiana

PDS

Krenz et al. (2010)

31

African Cassava mosaic virus (ACMV)

Begomovirus

Manihot esculenta; N. benthamiana

PDS, SU, CYP79D2, SPINDLY

Fofana et al. (2004), Beyene et al. (2017)

32

Beet curly top virus (BCTV)

Begomovirus

Spinacia oleracea, S. esculentum

RbcS, Transketolase, ChlI

Golenberg et al. (2009)

33

Cabbage leaf curl virus (CaLCuV/CbLCV)

Begomovirus

Arabidopsis

ChlI,CH42, PDS

Turnage et al. (2002), Muangsan et al. (2004), Flores et al. (2015)

N. benthamiana

PDS, SU, CLA1, SGT1

Tang et al. (2010, 2013), Huang et al. (2011)

34

Cotton leaf crumple virus (CLCrV)

Begomovirus

G. hirsutum

ChlI, PDS, GFP, EF-1α, cry1A

Tuttle et al. (2008, 2012, 2015), Gu et al. (2014)

35

Grapevine virus A (GVA)

Vitivirus

N. benthamiana; V. vinifera

PDS, GFP, ChlH

Muruganantham et al. (2009), Park et al. (2016)

36

Pepper huasteco yellow vein virus (PHYVV)

Begomovirus

Capsicum sp, Tobacco, Tomato

Comt, pAmt, Kas, SU

Carrillo-Tripp et al. (2006), del Rosario et al. (2008)

37

Rice tungro bacilliform virus Tungrovirus (RTBV)

O. sativa

PDS, ChlH, Xa21

Dasgupta et al. (1991), Purkayastha et al., (2010, 2013), Kant et al. (2015), Kant and Dasgupta (2017)

Table 14.1  Continued No.

VIGS vector

Genus

Host species silenced

Target gene(s) silenced

References

38

Tobacco yellow dwarf virus (TYDV)

Mastrevirus

Petunia hybrida

Chalcone synthase transgene

Atkinson et al. (1998), Voinnet et al. (2001)

39

Tomato golden mosaic virus (TGMV)

Begomovirus

N. benthamiana

SU, LUC; PCNA, ChlI, RBR1

Kjemtrup et al. (1998), Peele et al. (2001), Turnage et al. (2002), Carrillo-Tripp et al. (2006), Jordan et al. (2007)

40

Tomato leaf curl virus (ToLCV)

Begomovirus

N. benthamiana, S. lycopersicum

PCNA

Pandey et al. (2009), Pasumarthy et al. (2011)

Satellite DNA 41

Tomato leaf curl virus (TLCV) satellite DNA (satDNA)

Begomovirus

Tobacco, Petunia

GUS, ChsA

Li et al. (2008)

42

Tomato yellow leaf curl china necrotic virus (TYLCCNV) beta satellite

Begomovirus

N. benthamiana, L. esculentum, N. glutinosa, N. tabacum

PCNA, PDS, SU, GFP, FRO1

Tao et al. (2004a,b), Qian et al. (2006), Cai et al., (2007), He et al.,(2008), Zhou and Huang (2012)

43

Tobacco curly shoot virus (TbCSV) Betasatellite

Begomovirus

N. benthamiana

GFP, SU

Qian et al. (2006)

44

Tobacco curly shoot virus (TbCSV) Alphasatellite

Begomovirus

S. lycopersicum and Nicotina spp.; Petunia hybrida

GUS, SU, EDS1

Huang et al. (2009, 2011)

45

Bhendi yellow vein mosaic virus (BYMV)β-satellite

Begomovirus

N. benthamiana

GFP, SU, PDS, PCNA and AGO1

Jeyabharathy et al. (2015)

Virus-Induced Gene Silencing and its Applications |  287

RNA-dependent RNA polymerase (RdRp), movement protein (MP) and 16K cysteine rich protein (Macfarlane, 1999). The RNA2 encodes coat protein (CP) and restriction sites for cloning the target genes (Ratcliff et al., 2001, Liu et al., 2002b). TRV infects a wide range of host plants systemically including meristems, without producing severe virus associated symptoms. In addition, TRV does not encode any strong suppressor of silencing making it the most efficient VIGS vector developed so far (Ratcliff et al., 2001; Valentine et al., 2004; Martin-Hernandez and Baulcombe, 2008). TRV with 2b protein was developed for VIGS of root specific genes (Valentine et al., 2004). TRV-based VIGS vector has been successfully used in N. benthamiana (Ratcliff et al., 2001), Arabidopsis (Burch-Smith et al., 2006; Wang et al., 2006), tomato (Liu et al., 2002a; Ekengren et al., 2003), chilli pepper (Chung et al., 2004; Ryu et al., 2004), potato (Brigneti et al.,2004; Ryu et al.,2004; Du et al., 2013), wheat and maize (Zhang et al., 2017), Spinacea (Lee et al., 2017), opium poppy (Hileman et al., 2005), Aquilegia vulgaris (Gould and Kramer, 2007), cotton (Gao et al., 2011a; Qu et al., 2012; Zhang et al., 2014b), gladiolus (Zhong et al., 2014), petunia ( Jiang et al., 2011), woody plants (Vernicia, Populus, Camellia) ( Jiang et al., 2014b) and white poplar (Shen et al., 2015). TRV-VIGS vector allows cloning of DNA inserts up to ≈ 1 kb in size and produces very mild or no symptoms upon infection (Lu et al., 2003b, Burch-Smith et al., 2004). Viruses like TRV with wide host range, have enabled the exploitation of a single VIGS vector for gene silencing in numerous plant species. Barley stripe mosaic virus (BSMV) is a singlestranded, tripartite, positive-sense RNA virus that infects monocots like barley, oats, wheat, maize and rye. BSMV derived VIGS vectors have provided important information about host gene functioning, development and pathogenesis in monocots. Using this VIGS vector, the expression of several genes like phytoene desaturase (PDS), plastid transketolase (TK), magnesium chelatase subunit H (ChlH), and benzoxacin (Bx1) were down-regulated in wheat, barley, Brachypodium distachyon, N. benthamiana and rye (Holzberg et al., 2002; Scofield et al., 2005; Tai et al., 2005; Pacak et al., 2010; Groszyk et al., 2017). In order to extend the utility of VIGS to all economically important plants, few of the proven

virus-based silencing vectors were modified by different research groups. For example, a second generation TRV-VIGS vector was developed for high-throughput functional genomics by adopting GATEWAY technology (Thermo Fisher) (Liu et al., 2002a). However, this proved to be expensive and had proprietary issues, hence a modified ligation-independent cloning (LIC) approach that does not require either restriction digestion or ligation was used to develop a third generation TRV-VIGS vector for larger scale gene cloning (Dong et al., 2007). Recently, TRV vector with a green fluorescent protein (GFP) tag was generated and tested in several plants. Visualization of VIGS vector movement in non-solanaceous plants like rose, strawberry and chrysanthemum which do not support efficient infection of TRV at all times will help the researchers to track the movement of the vector in host plants (Tian et al., 2014a). VIGS methodology The basic principle of VIGS involves delivering recombinant viruses containing partial endogenous gene sequences and inducing silencing of the host gene via RNA silencing machinery of the plant (Fig. 14.1). Delivery of the recombinant viral genome in plants can be done by biolistic methods or via Agrobacterium tumefaciens by engineering them into T-DNA plasmids. Agrobacterium-mediated delivery is cheaper, easier and can be achieved by different methods described in section – inoculation methods. Agrobacterium mediated VIGS has advantages over other methods like in vitro transcription (Vaghchhipawala and Mysore, 2008) for obtaining infectious viral RNA, because steps such as cDNA synthesis that are time-consuming and costly are avoided. The Agrobacterium strain and method of inoculation must be standardized for each plant species. At first, few plant cells at the site of inoculation that are transformed with the virus vector would serve as a reservoir of infection, which then spreads systemically throughout the plant. Recognition of virus by the plant triggers PTGS (Lu et al., 2003b; Burch-Smith et al., 2004) first locally and then systemically. The co-expression of viral RNAs of the multi component viruses is achieved by mixing the Agrobacterium cultures harbouring different viral genomes. For example, Agrobacterium

288  | Kumar et al.

Figure 14.1  Schematic diagram of VIGS methodology.

harbouring RNA1 and RNA2 of TRV is mixed prior to inoculation. Vector insert The size of the host gene insert cloned into the VIGS vector affects the efficiency of silencing. Studies have shown that, when host fragments that are equal to or more than 1500 bp or bigger in size are cloned into VIGS vectors, they failed to induce silencing irrespective of the virus from which the vector was derived. Some studies have shown that while a 23 bp insertion was able to induce VIGS, higher silencing efficiency was obtained by choosing fragments of 200–350 bp in length (Lacomme et al., 2003). Usually, most VIGS vectors can carry a fragment of length between 150 and 800 bp. TRV RNA2 can tolerate up to 1kb size insert. For example, when TRV-based VIGS constructs carrying a 266-bp or 558-bp fragment of the phytoene desaturase (PDS) gene were Agrobacterium-infiltrated

into the leaves of the two Populus species (Populus euphratica and P. canescens), it was found that the shorter insert of 266 bp, but not the longer insert of 558 bp, resulted in the expected photobleaching in both tree species (Shen et al., 2015). Target gene The utmost care has to be taken while choosing the host gene sequences for targeting by VIGS. DNA sequences selected for cloning into the vector should be specific to the gene of interest to avoid off-target silencing and to obtain accurate phenotype caused by the lack of gene expression. Gene sequences that are part of conserved domains of multigene families can result in silencing of several genes leading to faulty results. On the other hand, if the aim is to study the effect of silencing of genes from a multigene family, VIGS is the ideal method to realize the same in the shortest possible time.

Virus-Induced Gene Silencing and its Applications |  289

Inoculation methods The efficiency of gene silencing is affected to a great extent by the inoculation methods. In most cases, Agrobacterium-carrying VIGS vectors are directly inoculated into the plants to induce silencing. However, in some cases, virus susceptible plants are agro inoculated at first to multiply the virus, and then the crude sap or the viral RNA extract is inoculated onto plants (Lee et al., 2015b). The commonly used Agro inoculation methods are – infiltration with a needleless syringe (Liu et al., 2002a; Dinesh-Kumar et al., 2003; Lu et al., 2003b), pin-prick method, agro drenching (Ryu et al., 2004), paste application in the root–shoot junction (Deshmukh et al., 2010) and high pressure sprays using air brush (Padmanabhan and Dinesh-Kumar, 2009; Sasaki et al., 2011). Particle bombardment of infectious clones (Li et al., 2015b), rub-inoculation with RNA transcripts (Diaz-camino et al., 2011) and DNA (Pflieger et al., 2008), and vasculature puncture inoculation (Louie, 1995; Benavente et al., 2012) are also reported by some researchers. Leaf infiltration of agroinfectious clones using a needleless syringe is a very efficient method of delivering Agrobacterium for inducing silencing in plants, particularly in dicots. This method has been proven successful for VIGS vector delivery in Arabidopsis, Nicotiana sp., tomato, chilli pepper, potato, cucurbits, cotton, petunia, etc. Direct inoculation with an agro-culture in fruits produced a more obvious silencing phenotype than inoculation of cotyledons or seedlings (Fu et al., 2005, Orzaez et al., 2006). Silencing gene expression in fruits at pre-harvest and post-harvest stages was better achieved by direct injection of agrocultures in fruits such as tomato, strawberry and bilberry when compared with inoculation of seedlings or young plantlets (Jaakola et al., 2010, Chai et al., 2011, Romero et al., 2011). Leaf and fruit infiltrations are laborious for large-scale screening as leaves and fruits of some plants are difficult to infiltrate and individually inoculating plants or fruits is labour intensive. In several solanaceous species like N. benthamiana, tomato, pepper, tobacco, potato and petunia, high efficiency silencing is reported by drenching roots with a suspension of agroinfectious clones (Ryu et al., 2004). Agro drench is advantageous over leaf

infiltration as it can be utilized for VIGS in very young seedlings and does not require multiple fully expanded leaves. Agrobacterium-mediated inoculation of VIGS vectors in a small herbaceous plant like Arabidopsis could be achieved by simply rubbing the leaves with an abrasive – Celite that gives uniform silencing (Manhaes et al., 2015). Previously, Ding et al. (2006) reported that agro-infiltration under vacuum was effective in plants like barley, rice and maize that are difficult to inoculate by conventional methods. Later, vacuum infiltration of sprouts was shown to be efficient for inducing VIGS in Solanaceous species like tomato, aubergine, pepper and N. benthamiana (Yan et al., 2012). Recently, Zhang et al. (2017) achieved whole-plant level gene silencing with TRV-VIGS system wherein wheat and maize (germinated) seeds were inoculated under vacuum with agro cocultivation. Liu et al. 2002a used an artist’s airbrush to deliver TRV vectors into tomato by spraying agro-cultures and an abrasive under pressure. Among the different types of infection methods, true leaf infection was found to be more efficient than cotyledon and sprout infiltration for long-term VIGS in multiple plant organs (Meng et al., 2016). The co-inoculation of viral suppressors along with VIGS vectors was also found to increase virus titre in inoculated cells leading to better systemic silencing. By inventing new and combining different established VIGS-vector delivery methods, VIGS could be expanded to more plant species. Growth conditions Efficiency of gene silencing is affected by environmental factors that affect plant growth and development (Patil and Fauquet, 2015). They are also modulated by factors that affect multiplication and systemic spread of the virus. TRV replication and infection is favoured at temperatures between 17°C and 21°C (Fu et al., 2006; Tuttle et al., 2008). VIGS by TRV is effective at temperatures less than 26°C. At the same time, a higher temperature is required for efficient replication of geminiviral DNA A and DNA B-derived VIGS vectors and they induce very good silencing between 22°C to 32°C (Tao and Zhou, 2004; He et al., 2008). VIGS efficiency was highest at a photoperiod of 16/8 h (light/dark) and at a growth temperature of approximately 27°C following syringe infiltration

290  | Kumar et al.

of Soybean yellow common mosaic virus (SYCMV) based vectors in soybean (Kim et al., 2016). Applications of VIGS Abiotic stress (drought, salt, oxidative, nutrient deficiency) Abiotic and biotic stresses are major factors that limit the productivity of agricultural crops worldwide. Abiotic stress responses, being complex in nature, are controlled by networks of genetic and environmental factors. Several abiotic stressresponsive genes have been previously identified and their functions exploited. Recent development of vectors with higher silencing efficiency has facilitated the application of VIGS for the characterization of genes that are responsive to abiotic-stress like drought, salinity, heat, cold, nutrient deficiency, etc. Drought stress Eighty per cent (1.2 billion ha) of the total cultivated area world wide is rainfed. Intensive cultivation of crops and development of industries have unscrupulously exploited water resources. Developing drought-tolerant varieties and hybrids of crop plants is crucial to meet the food demands of an ever growing world population. Data on drought responsive genes shortlisted by functional validation of candidate genes in model plants such as A. thaliana are an excellent source of information for breeding varieties of non-model staple crops. VIGS has been used successfully to screen and validate potential candidate genes from model plants and using selected genes for imparting drought tolerance in crops that are recalcitrant to transformation. How these drought-responsive genes influence plant metabolism to mitigate moisture stress is investigated using VIGS in both dicots and monocots. The roles of many genes that are differentially regulated by moisture stress in different stages of plant growth have been confirmed by VIGS. Late embryogenesis abundant (LEA) protein gene family functions in cellular protection during abiotic stress tolerance. TRV-VIGS mediated silencing of LEA4 gene in tomato increased the susceptibility of plants to moisture stress. LEA4 silenced plants wilted faster and recovered more slowly upon

re-watering than controls. These plants exhibited a reduced osmotic adjustment, higher superoxide radical levels and low cell viability indicating a role for this gene in drought mitigation (Senthil-Kumar and Udayakumar, 2006, 2010). VIGS of two other LEA genes – Hordeum vulgaris abundant protein (HvHVA1) and dehydrin 6 (HvDhn6) in Tibetan hull-less barley (H. vulgare) – under drought stress showed lower survival rates due to higher rates of water loss (Liang et al., 2012). In another study, VIGS of CaLEA1 in pepper plants led to enhanced sensitivity to both drought and salt stresses accompanied by high levels of lipid peroxidation in dehydrated and salt [sodium chloride (NaCl)]treated leaves (Lim et al., 2015). Expression of dehydrins (DHNs), a subfamily of Group-2 LEA genes is up-regulated by cold, salt, osmotic stress and salicylic acid treatments. Loss of function analysis of chilli pepper dehydrin (CaDHN1) by VIGS led to decreased tolerance to cold, salt and osmoticinduced stresses. Although many LEA genes have been identified and analysed in angiosperms, very few have been studied in gymnosperms (Gao and Lan, 2016). VIGS is an ideal tool for investigating the biological roles of heterologous genes in species where gene function analysis is lacking. Responses of plants to reduced water availability and developmental processes are coordinated by the hormone abscisic acid (ABA). Many key regulators of ABA signalling have been identified using both forward and reverse genetic approaches. The repression of farnesyl transferase genes (Enhanced Response to ABA1 – ERA1) has been shown to enhance drought resistance in Arabidopsis, canola, wheat, rice and soybean. For example, BSMV-VIGS mediated silencing of Era1 and inositol polyphosphate 1-phosphatase (Sal1) genes in wheat also enhanced drought tolerance by decreasing sensitivity to ABA (Manmathan et al., 2013). PacCYP707A1, a gene regulating ABA content and response of sweet cherry to drought during fruit development was identified by TRV-VIGS (Li et al., 2015b). ABA-Insensitive RING protein 1 gene (CaAIR1) is needed for drought stress induced hypersensitive response (HR) in hot pepper (Capsicum annuum) and its expression levels are up-regulated in leaves by ABA, drought and NaCl treatments. Silencing of this gene in pepper reduced water loss by transpiration and enhanced drought tolerance (Park et al., 2015). ASLV-mediated VIGS

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of Arabidopsis ERA1 homologues – GmERA1A and GmERA1B – in soybean confirmed that they encode negative regulators of ABA signalling and controls stomatal movements under low moisture conditions. Down-regulation of ERA1 genes has the potential to impart drought tolerance in soybean (Ogata et al., 2017). The Pepper WPP Domain Protein (CaWDP1), a novel negative regulator of ABA-mediated stress signalling, was recently identified using VIGS (Park et al., 2017). Increased production and accumulation of proline is one of the ways by which, plants acclimatize to water stress. VIGS of genes involved in proline synthesis pathways- Δ (1)-pyrroline 5-carboxylate synthase (P5CS) and ornithine-δ-aminotransferase (OAT) in N. benthamiana showed that proline accumulation is inhibited in drought and PEG treatments when P5CS was silenced. With ABA treatment also, proline failed to accumulate in P5CS silenced plants. However, OAT silenced plants were unaffected. Both genes were also observed to have roles in chlorophyll degradation (Ku et al., 2011). BSMV-VIGS-mediated silencing of the Basic Transcription Factors 3 (TaBTF3) and autophagyrelated 8 (TdAtg8) genes established a role for these genes in drought and osmotic stress response in wheat (Kuzuoglu-Ozturk et al., 2012; Kang et al., 2013). BSMV-VIGS of a putative H. vulgare ATG6 gene (HvATG6) led to accelerated yellowing of leaves with dark and hydrogen peroxide treatments, showing its importance in stress induced autophagy (Zeng et al., 2017). When HvEXPB7, a novel β-expansin gene was silenced using BSMV vector in a drought tolerant cultivar X25, root hair growth and K+ uptake were severely suppressed under control and drought conditions. These findings confirmed the significance of HvEXPB7 induced root hair growth and provided a novel insight into the genetic basis for drought tolerance in Tibetan wild barley (He et al., 2015b). The mitogen-activated protein kinase (MAPK) is an evolutionarily conserved signal transduction module consisting of MAP kinase kinase kinases (MAPKKKs/MEKKs/MKKKs), MAP kinase kinases (MAPKKs/MEKs/MKKs) and MAP kinases (MAPKs/MPKs), which sequentially phosphorylate the corresponding downstream substrates (Mishra et al., 2006; Rodriguez et al., 2010; Xu and Zhang, 2015). MAPK cascades control

signalling pathways and are activated by different abiotic stresses such as heat, cold, touch, wounding, UV, osmotic shock, salt and heavy metals (Tena et al., 2001; Zhang et al., 2006b; Colcombet and Hirt, 2008; Liu, 2012). VIGS approach was adopted to study the role of MAP kinases -SpMPK1, SpMPK2, and SpMPK3 in drought tolerant wild relative of tomato, S. pimpinellifolium using TRV vector (Li et al., 2013). Silencing of SlMPK4 led to early wilting and reduced the tolerance to drought (Virk et al., 2013). VIGS of a Raf-like MAPKKK gene – GhRaf19 in cotton (Gossypium hirsutum) showed that it negatively regulates tolerance to drought and salt and positively regulates resistance to cold stress by modulating cellular reactive oxygen species (ROS) ( Jia et al., 2016). Silencing of GhMKK3, a MAPKK in cotton, increased the rate of water loss and resulted in decreased tolerance to drought (Wang et al., 2016a). Cotton plants silenced for sucrose non-fermenting 1-related protein kinase2 (GHSnRK2) gene also showed alleviated drought tolerance (Bello et al., 2014). Transcription factors (TF) play essential roles in the adaptation of plants to abiotic stresses. Many TFs are encoded by multigene families like NAC, MYB and WRKY. Expression of several of these genes is influenced by drought stress. NAC factor JUNGBRUNNEN1 ( JUB1) is a regulator of drought tolerance in cultivated tomato. Silencing of SlJUB1 by VIGS drastically lowered the degree of drought tolerance in tomato. Silenced tissue showed an increase in ion leakage, accumulation of hydrogen peroxide (H2O2) and a decrease in the expression of several drought-responsive genes (Thirumalkumar et al., 2017). MYB and WRKY transcription factors are vital to plants for adapting to abiotic stress. Down-regulation of a MYB transcription factor, PbrMYB21 in Pyrus betulaefolia by TRV-VIGS reduced arginine decarboxylase (ADC) and polyamine levels along with increased sensitivity to drought (Li et al., 2017d). VIGS of a WRKY gene (GhWRKY27a) imparted enhanced tolerance to drought stress in cotton (Yan et al., 2015). GhWRKY59 is yet another gene that regulates the drought stress response in cotton (Li et al., 2017b). VIGS of SR/CAMTA transcription factor, SlSR1L, in tomato resulted in increased water loss from leaves and reduction in the root biomass indicating that SISR1L is a positive regulator of drought stress tolerance (Li et al., 2014b).

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Gene knockouts using VIGS is an effective way of deciphering gene function in vegetatively propagated plants and in tree species. Inhibition of petal expansion due to moisture stress often leads to abnormal flower opening and reduction in the marketability of cut flowers. NAC transcription factors 2 and 3 (RhNAC2 and RhNAC3) and a putative expansin gene type expansin 4 (RhEXPA4) were found to be up-regulated by dehydration in rose (Rosa hybrida). VIGS of RhNAC2 or RhEXPA4 in rose petals significantly decreased the recovery of plants after rehydration. Interestingly, RhEXPA4 expression was repressed in RhNAC2-silenced rose petals (Dai et al., 2012). Silencing of RhNAC3 also significantly decreased the cell expansion in rose petals when rehydrated ( Jiang et al., 2014a). A role for ethylene in dehydration-induced inhibition of cell expansion was also demonstrated by VIGS of an ethylene receptor, RhETR3 (Liu et al., 2013). Senthil-Kumar et al. (2007) functionally characterized water deficit-induced genes from peanut using VIGS. Silencing of peanut flavonol 3-O-glucosyltransferase (F3OGT), alcohol dehydrogenase, salt inducible protein and heat shock protein 70 (HSP70) in N. benthamiana showed visible wilting symptoms as compared to controls during water deficit, demonstrating applications of heterologous VIGS. Salt stress Genes differentially regulated by salt stress also show changes with other types of abiotic stresses, especially drought. The role of genes involved in salt stress tolerance such as glutaredoxin (SlGRX1) in tomato (Guo et al., 2010), related-toABI3/VP1 (Abscisic acid insensitive 3/viviparous 1) (CaRAV1) and oxidoreductase (CaOXR1) in chilli pepper (Lee et al., 2010) was established by VIGS. Silencing of these genes resulted in yellowing or severe bleaching of leaves due to reduction in the chlorophyll content. Silencing of peroxidise 2 (CaPO2) in chilli pepper using TRV-vector caused a reduction in chlorophyll pigments, an increase in lipid peroxidation and made the plants more susceptible to salt stress (Choi and Hwang, 2012). Silencing of a WRKY TF and a mitogen-activated protein kinase (MAPK) by TRV-VIGS significantly increased the susceptibility of cotton to salt stress (Cai et al., 2017).

Similarly VIGS of S. pimpenellifolium RING finger E3 ligase (SpRing) led to an increased sensitivity to salt (Qi et al., 2016). Jia et al. (2016) showed that, in cotton, GhRaf19 negatively regulates salt tolerance and positively regulates cold stress tolerance by modulating cellular reactive oxygen species. Bahieldin et al. (2016) detected the role of ethylene response transcription factor, ERF109, which is co-expressed during programmed cell death (PCD) and salt stress. The same group also identified and functionally characterized a PCD related anti apoptotic Bax inhibitor-1 (BI-1) and apoptotic mildew resistance locus O (Mlo) gene using VIGS. Their results showed that higher level of salt tolerance in plants can be achieved by retarding the expression of PCD-inducing BI-1 gene. γ-aminobutyric acid (GABA) accumulates in many plant species in response to environmental stress. Bao et al. (2015) studied the functional role of glutamate decarboxylases (GADs), GABA transaminases (GABA-Ts) and succinic semialdehyde dehydrogenase (SSADH) using VIGS in tomato. VIGS of SlGADs and SlGABA-Ts led to increased accumulation of ROS and salt sensitivity. Oxidative stress Reactive oxygen species (ROS) are redox signals essential to many physiological processes including growth and development and stress tolerance in plants. However, at very high concentrations, ROS cause damage to macromolecules and disrupt normal signalling. Glutaredoxin (GRXs) are ubiquitous small heat-stable disulfide oxidoreductases that reduce cell damages during oxidative stress. SlGRX1 is expressed in tomato leaf, root, stem and flower and is induced by oxidative, drought and salt stresses. The biological function of SlGRX gene was validated using VIGS in tomato. Silencing this gene led to increased sensitivity to oxidative and salt stresses with decreased relative chlorophyll content, and reduced tolerance to drought associated with decreased relative water content (Guo et al., 2010). Chloroplast thioredoxins (TRXs) and glutathione function as redox messengers in the regulation of photosynthesis. PEBV mediated cosilencing of thioredoxin genes, TRX-F/TRX-M, in pea showed a significant down-regulation of

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magnesium-chelatase activity and 5-aminolaevulinic acid synthesizing capacity. Photosynthesis was affected due to decline in chlorophyll and carotenoid pigments. ROS concentration in tissues was high due to lower levels of expression of genes governing tetrapyrrole biosynthesis (Luo et al., 2012). VIGS of the TRXs genes-TRX-f, TRX-m2, TRX-m1/4 and TRX-y was carried out individually in tomato to study their roles in brassinosteroids induced changes in cellular redox homeostasis (Cheng et al., 2014); it led to an increase in membrane lipid peroxidation and accumulation of 2-Cys peroxiredoxin dimers in leaves. Chen et al. (2015) isolated a novel wheat GRAS gene, TaSCL14 and functionally characterized using VIGS. Silencing of TaSCL14 in wheat resulted in plant growth inhibition, decreased photosynthetic capacity and photooxidative stress tolerance suggesting its role as multifunctional regulator. Plant peroxidases encoded by a large number of super family genes exhibit diverse expression patterns in response to stresses such as wounding, ethylene, pathogen infection, drought, low-temperature, iron deficiency, light and plant growth regulators (Hiraga et al., 2000; Ito et al., 2000; Kim et al., 2000; Park et al., 2003; Yoshida et al., 2003; Valério et al., 2004; Passardi et al., 2005; Choi et al., 2007; Matin et al., 2010; Sang et al., 2010). VIGS of extracellular peroxidase 2 (CaPO2) in chilli pepper resulted in an increased susceptibility to mannitolinduced osmotic stress showing severe bleaching (Choi and Hwang, 2012). Plants in which CaRAV1 was silenced alone or together with CaOXR1 by TRV-VIGS exhibited reduced tolerance to mannitol-induced osmotic stress (Lee et al., 2010) and enhanced lipid peroxidation (Lee et al., 2010; Choi and Hwang, 2012). The expression levels of antimicrobial protein (CaAMP1) and osmotin (CaOSM1) were also reduced during oxidative stress (Hong et al., 2004; Lee and Hwang, 2009) in chilli pepper. Silencing of CaMLO2 using TRV-based VIGS resulted in lower malondialdehyde (MDA) levels (Lim and Lee, 2014), whereas silencing of TdAtg8 using BSMV-based VIGS resulted in higher MDA levels suggesting a role of these genes in drought stress tolerance. Atg8 appears to be a positive regulator in osmotic and drought stress response (Kuzuoglu-Ozturk et al., 2012).

Nutrient deficiency The function of few genes during mineral nutrient deficiency and toxicity in plants has also been determined by using VIGS. Iron (Fe) is an essential micronutrient required by plants and its uptake, translocation, distribution and utilization are maintained in a complex manner. Silencing of a ferric reductase oxidase (FRO1) gene that encodes ferric-chelate reductase enzyme (FCR) for chlorophyll synthesis and Fe partitioning, via a modified geminiviral satellite (DNA β) associated with Tomato yellow leaf curl China virus (TYLCCNV), down regulated FCR activity in tomato roots (He et al., 2008). PEBV based VIGS vector was used to study arbuscular-mycorrhizal-fungi (AMF)-associated phosphate acquisition by silencing of symbiotic gene, PsSym19, and Pi transporter gene, PsPT4, that resulted in reduction of phosphate uptake in pea plants (Grønlund et al., 2010). Soybean plants failed to respond to an increased Fe nutrition when replication protein A (GmRPA3) gene was silenced using BPMV-based VIGS vector, suggesting a role of the GmRPA3 gene in iron acquisition (Atwood et al., 2014). Du et al. (2015) down regulated the expression of basic helix loop helix (bHLH), SlbHLH068 in tomato using VIGS and showed that silencing of SlbHLH068 resulted in reduction of LeFRO1 and LeIRT1 expression and iron accumulation in leaves and roots. Gama et al. (2017) used TRV-VIGS to silence FRO1 gene in N. benthamiana and showed its importance in chlorophyll synthesis and Fe partition. VIGS of FRO1 considerably reduced FCR activity, consequently preventing Fe uptake and resulted in chlorotic symptoms in plants. Biotic stress: plant defence response against pathogens and pests VIGS is effectively utilized for unravelling the functions of novel genes involved in plant disease resistance to bacteria, fungi, viruses, nematodes and insect pests. VIGS has led to the functional analysis of numerous host genes involved in plant resistance signalling. Viruses Several cDNAs from both model and crop plants were silenced using VIGS to determine their functional significance in susceptibility and resistance to

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viruses. VIGS technology has led to identification of novel genes involved in viral resistance pathways and HR in different plant species. By VIGS, identification and confirmation of new roles for previously known genes and identification of new factors in defence signalling was achieved. Tobacco mosaic virus (TMV) is an economically important virus as well as model virus for classroom teaching. Resistance to TMV is governed by N gene in Nicotiana sp. VIGS has been successfully applied to determine the functional role of several genes like enhanced disease susceptibility 1 (EDS1), Non-Expresser of Pathogenesis Related Genes 1 (NPR1/NIM1) (Liu et al., 2002b; Peart et al., 2002), MAPKK, NTF6 MAPK, WRKY and MYB transcription factors, coronatine-insensitive 1 (COI1) and constitutive triple response protein (CTR1) (Liu et al., 2004a), transcription factorNRG1, MAP ERK kinase (MEK1) (Peart et al., 2005), heat shock protein (HSP) 90 genes (Lu et al., 2003a) and RAN GTPase activating protein 2 (RanGAP2) (Tameling et al., 2007) involved in the N-gene mediated resistance to TMV. Silencing of several kinases like NPK1 ( Jin et al., 2002), wound-induced protein kinase (WIPK), salicylic acid-induced protein kinase (SIPK), NtMEK1 and NtMEK2 ( Jin et al., 2003; Liu et al., 2003, 2004a; Sharma et al., 2003) impacted N-gene mediated resistance to TMV in N. benthamiana. VIGS of AUTOPHAGY 6 (ATG6/BECLIN-1) (Liu et al., 2005) and Nuclear receptor interacting protein 1 (NRIP1) (Caplan et al., 2008) in N-gene containing resistant plants led to an unrestricted PCD upon TMV infection. VIGS of microRNA (MiRVIGS) of N. benthamiana suppressor-of-G2-allele-of-skp1 (NbSGT1) led to the loss of N-mediated resistance to TMV (Tang et al., 2013). VIGS was deployed to study the biological function of endogenous miRNAs (Tang et al., 2013). VIGS was also used to determine the function of mitochondrial alternative oxidase (AOX) in N-mediated resistance. When inoculated onto NbAOX1a-silenced N-transgenic N. benthamiana plants, TMV was able to move systemically resulting in HR-PCD and death of whole plant (Zhu et al., 2015). Suppression of a fatty acid desaturase 1 (FAD1) gene expression in tobacco led to the blocking of cell death induced by Bcl2associated X (Bax) protein. When its homologue in C. annuum (CaFAD1) gene was silenced in pepper, resistance to TMV-P0 infection was significantly

weakened when compared to vector alone control plants (Kim et al., 2007). Potato virus Y (PVY) is an aphid transmitted single-stranded, positive sense RNA virus that infects mainly Solanaceous crops like potato, chilli pepper, tomato, etc. and causes huge devastation worldwide. PVY and several other members of the Potyviridae family of viruses cause synergistic interactions with heterologous viruses like PVX, CMV, TMV etc. in mixed infections leading to severe crop loss. Several genes governing PVX-PVY synergism and Rx gene mediated resistance to PVX were identified and functionally characterized by means of VIGS. Silencing of N. benthamiana MAPK genes, SIPK, WIPK, and the MAPK kinase (MAPKK) genes MEK1 and MKK1, partially compromised HR-like cell death and necrosis induced by the synergistic interaction of PVX with PVY. Ameliorated cell death induced by PVX-PVY in the MAPKK-silenced plants did not facilitate virus accumulation in systemically infected leaves. Dual silencing of SIPK and oxylipin biosynthetic gene 9-Lipoxygenase showed that the latter was epistatic to SIPK in response to PVX-PVY infection. These findings demonstrate that SIPK, WIPK, MEK1 and MKK1 function as positive regulators of PVXPVY-induced cell death (Aguilar et al., 2017). Pepper mottle virus (PepMoV) is a major pathogen on chilli pepper. Pepper potyvirus resistance gene, Pvr9, confers HR in N. benthamiana during PepMoV infection. Requirement of components of this signal transduction pathway like HSP90, SGT1, NDR1 and SA for Pvr9-mediated HR was revealed by VIGS (Tran et al., 2016). Recently, Cui and Wang (2017) demonstrated that silencing of host factor – eukaryotic translation initiation factor 4E isoform (elF(iso)4E) using Prunus necrotic ringspot virus (PNRSV)-based vector, induced resistance to Sharka disease caused by Plum pox virus (PPV) in N. benthamiana and Prunus. Silencing of translation initiation factors, eIF4E and eIFiso4E by TRV-VIGS in pepper resistant to Chilli veinal mottle virus (ChiVMV) showed reduction in virus titre, suggesting that, by making simultaneous mutations or knockouts in eIF4E and eIFiso4E, ChiVMV infection in pepper can be controlled (Hwang et al., 2009). As seen with N-mediated resistance to TMV, silencing of MAPKK kinase, NPK1 also interfered

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with the function of the disease-resistance governed by CC-NBS-LRR (coiled coil, nucleotide binding site, leucine rich repeat) resistance gene Rx against PVX ( Jin et al., 2002). VIGS of Ran GTPase activating (RanGAP2) in Rx-gene carrying plants induced PCD against PVX infection (Tameling et al., 2007). When a homologue of the pepper nucleotide exchange unit eEF1Bβ of plant translation elongation factor 1(eEF1) was suppressed in N. benthamiana using VIGS, the accumulation of PVX was greatly affected suggesting that eEF1Bβ could be a potential target for engineering virus-resistant plants (Hwang et al., 2015). Geminiviruses are single-stranded DNA viruses transmitted by insects like whiteflies and leafhoppers that limit the production of quite large number of vegetable crops in tropics and subtropics. Germin-like proteins (GLPs) were identified and functionally characterized in geminivirus-resistant Capsicum chinense Jacq accession BG-3821. VIGS of GLP in resistant pepper increased susceptibility to single and mixed infections by geminiviruses: Pepper huasteco yellow vein virus (PHYVV) and Pepper golden mosaic virus (PepGMV) (MejiaTeniente et al., 2015). Using TRV-VIGS, the role of hexose transporter LeHT1 in resistance of tomato to Tomato yellow leaf curl virus (TYLCV) was studied. Inoculation of LeHT1-silenced plants with the virus, compromised resistance and resulted in an increase in virus replication, systemic spread, tissue necrosis along with growth inhibition of plants (Eybishtz et al., 2010). Czosnek et al. (2013) employed TRV-VIGS as a reverse genetics tool for the identification of host genes that impart resistance to the virus in tomato as well as those genes that are essential for TYLCV infection. TYLCV resistance genes- Ty-1 and Ty-3 derived from different Solanum chilense accessions are allelic and code for RNA-dependent RNA polymerase (RDR) with a conserved DFDGD catalytic domain. Ty-1/Ty-3 gene was silenced in various Solanum lycopersicon (cultivated tomato) lines into which resistance from S. chilense accessions was introgressed to decipher the resistance mechanism operating in these lines against the virus (Caro et al., 2015). Silencing of MAP kinase genes, SlMAPK1, SlMAPK2 and SlMAPK3 in tomato, reduced tolerance to TYLCV, increased leaf H2O2 concentrations, and attenuated expression of defence-related genes after TYLCV infection, especially in SlMAPK3-silenced

plants (Li et al., 2017f). The response of members of WRKY group III TFs was identified during TYLCV infection in tomato. VIGS of WRKY41 and WRKY54 showed low level of viral DNA in TYLCV resistant plants when compared to controls (Huang et al., 2016). TRV-mediated silencing of autophagy (ATG) genes- ATg-6/Beclin1, phosphatidyl inositol 3-kinase (PI3K) and VPS15 which constitute phosphatidylinositol 3-kinase (PI3K) complex, in N. benthamiana confirmed the involvement of these genes in calmodulin-like protein-Rgs-CaM-mediated degradation of Suppressor of gene silencing 3 (SGS3) and an increase in the symptom induced by geminivirus associated with beta satellite (Li et al., 2017a). Similarly, a rapid VIGS-based screening system was developed to determine the role of genes involved in resistance and susceptibility to cassava mosaic disease (CMD) within 2–4 weeks. VIGS of Manihot esculenta SPINDLY (MeSPY) using East African Cassava mosaic virus (EACMVK201) vector resulted in shoot-tip necrosis die back of whole plant in CMD-susceptible cassava lines (Beyene et al., 2017). HRT (HR to TCV-Turnip crinkle virus) is CCNB-LRR disease resistance protein that activates HR-mediated cell death on identification of avirulence factor – coat protein (CP) of TCV in N. benthamiana. Capsicum annum gene BLP5 encodes an ER-associated immunoglobulin-binding protein (BiP). VIGS of CaBLP5 decreased HRT-mediated HR significantly and silenced BiP4/5. HRTmediated HR induced ER stress-responsive genes. Further, co-expressing TCV-CP in BiP4/5-silenced plants stopped induction of HRT completely (Moon et al., 2016). Tamura et al. (2013) showed that plants preinfected with ALSV vector harbouring fragments of the target viral genome sequences inhibited the replication and spread of challenged viruses like Bean yellow mosaic virus (BYMV), Zucchini yellow mosaic virus (ZYMV) and CMV. Zhang et al. (2012) used VIGS for identification of multiple defence related genes required for Rsv-1 mediated extreme resistance to Soybean mosaic virus (SMV). Arabidopsis tolerant to Tobacco ringspot virus 1 (TTR1) encodes a TIR-NBS-LRR protein and develops lethal systemic necrosis phenotype upon TRSV infection. VIGS studies showed that TTR1 mediated resistance is dependent on SGT1, while EDS1 was not required (Nam et al., 2011).

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Study of the interaction of Rice stripe virus (RSV) p2 and plant nucleolar protein fibrillarin showed that nuclear localization and cytoplasm distribution of RSVp2 is affected in fibrillarin silenced N. benthamiana (Zheng et al., 2015). RSVp2 interacted with fibrillarin to manipulate nuclear function to promote systemic movement of virus. Fungi VIGS was utilized for functional validation of genes involved in defence response and R-gene mediated resistance to several fungal pathogens in cereals, vegetables and fibre crops. Cladosporium fulvum is a biotrophic fungus that causes leaf mould disease in tomato. C. fulvum and tomato is a model for R (Resistance)–Avr (Avirulent) protein interaction studies. VIGS has contributed significantly to the understanding of resistance signalling pathways governed by Cf proteins that upon recognition of any one of the several Avr products from the fungus trigger resistance response. VIGS was employed by several groups to identify and characterize various host genes such as ACIK1 (Avr9/Cf-9 involved kinase 1) (Rowland et al., 2005), HSP90, nuclear GTPase, L19 ribosomal protein, NRC-1 (an NB-LRR-type protein required for HR associated cell death) (Gabriëls et al., 2006, 2007) and tomato PLC4 (Phospholipase C) (Vossen et al., 2010) involved in Cf-gene-familymediated resistance to leaf mould pathogen C. fulvum in tomato. CITRX (Cf-9-interacting thioredoxin) was found to negatively regulate Cf-9/Avr 9 induced HR (Rivas et al., 2004), while LeMPK1, LeMPK2, and LeMPK3 positively regulates Cf-4/ Avr4 induced HR and resistance in tomato (Stulemeijer et al., 2007). Phytophthora blight, which is caused by the oomycete pathogen Phytophthora capsici Leonian, is a severe threat to chilli pepper production worldwide. P. capsici resistance gene, CaRGA2 (Resistance gene analogs 2), from a highly resistant pepper (C. annuum CM334), was isolated and characterized. When CaRGA2 gene was silenced, via VIGS, the resistance level was suppressed, showing that this gene is critical for P. capsici resistance. Several other genes which are involved in resistance response of plants to P. capsici were also identified using VIGS. Ethylene-responsive factors (ERF), CaPTI1 silenced pepper plants were more susceptible to P. capsici. The expression of

defence genes such as pathogenesis related (PR1), defensin1 (DEF1) and SAR8.2 and activity of signalling molecules SA, methyl jasmonate, ethephon, and hydrogen peroxide in roots were reduced in CaPTI1 silenced plants, suggesting a critical role for this gene in defence responses to the fungus ( Jin et al., 2016). Further, the results of VIGS of ChiIV3a chitinase encoding gene in pepper showed that it is required for inhibition of P. capsici growth by triggering defence signalling and up-regulation of PR proteins (Liu et al., 2017). Silencing of the MYB transcription factor CaMYB in detached leaves of pepper rendered more sporulation of P. capsici Leonian, indicating that CaMYB is important in the defence response to pathogens (Zhang et al., 2013). VIGS revealed WIPK and SIPK genes function in mediating HR response to INF1 by P. infestans and defence response to P. cichorii in N. benthamiana. However, WIPK and SIPK when silenced showed no effect on HR induced by INF1 and reduced resistance to P. cichorii (Sharma et al., 2003). The U-box E3 ubiquitin ligase PUB17 was silenced in potato to demonstrate that it acts in the nucleus and is a positive regulator of PCD and specific immune pathways triggered by P. infestans (He et al., 2015a). Identification of new resistance genes against P. capsici will aid in breeding for broad spectrum durable resistance in pepper cultivars (Zhang et al., 2013). Botrytis is a necrotrophic fungus which causes grey mould disease of tomato, grapes, strawberry, cotton, etc. VIGS of two MAPK genes in tomato (SlMKK2 and SlMKK4) compromised the resistance to B. cinerea (Li et al., 2014c). Similarly, VIGS of individual matrix metalloproteinases (SlMMPs) and disease assays indicated that silencing of Sl3-MMP also resulted in reduced resistance (Li et al., 2015a). In another study by Zhang et al. (2015), TRV-VIGS-based knockdown of SlHUB1 and SlHUB2-H2B monoubiquitination E3 ligases increased the susceptibility of tomato to grey mould. Silencing of SlERF.A1, SlERF.A3, SlERF. B4, or SlERF.C3 belonging to the B3 group of tomato ERF family of transcription factors, also led to increased susceptibility to B. cinerea and promoted fungus induced H2O2 accumulation (Ouyang et al., 2016). On the other hand, knockdown of SlSR1 and SlSR3L genes encoding SR/ CAMTA transcription activators, individually, in tomato resulted in enhanced resistance to B. cinerea which was accompanied by accumulation of H2O2,

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elevated expression of defence genes and marker genes for pathogen-associated molecular pattern (PAMP) triggered immunity, and regulatory genes involved in the SA and ethylene-mediated signalling pathways (Li et al., 2014b). Rusts caused by Puccinia species are major fungal diseases of wheat and barley. Wheat stripe rust caused by Puccinia striiformis f. sp. tritici (Pst), stem rust caused by Puccinia graminis f. sp. tritici and leaf rust caused by Puccinia triticina are some of the most destructive fungal diseases worldwide. Scofield et al. (2005) used BSMV- VIGS assay to study genes that contribute to leaf rust resistance mediated by Lr21, an NBS-LRR class of resistance protein in wheat. Silencing of RAR1, SGT1 and HSP90 genes involved in resistance pathways governed by NBSLRR proteins resulted in fungus susceptibility of the R lines, showing that these genes are also critical for rust resistance. Suppression of TaHsp90.2 or TaHsp90.3 genes encoding heat shock protein 90 (Hsp90) compromised the HR of the wheat variety Suwon 11 to stripe rust fungi (Wang et al., 2011). Knocking down the expression of an ethylene-insensitive transcription factor (TaEIL1) through BSMV-VIGS attenuated the growth of Pst, due to reduction in the size of hyphae and hyphal branches, haustorial mother cells and colony size. Enhanced necrosis by Pst avirulent race CYR23 meant that HR was strengthened inTaEIL1-silenced wheat plants suggesting that by the suppression of TaEIL1 the resistance of wheat to stripe rust fungus could be enhanced (Duan et al., 2013). Similarly, silencing the expression of TaLSD1- a wheat homologue of Arabidopsis lesion simulating disease 1 (lsd1) also increased wheat resistance to Pst accompanied by faster HR, increase in PR1 gene expression and reduction in the growth of fungal hyphae (Guo et al., 2013). Feng et al. (2014) employed VIGS to down-regulate TaMDHAR4, a mono dehydroascorbate in wheat and showed that it enhanced wheat resistance to Puccinia striiformis f. sp .tritici by inhibiting fungal sporulation, increased necrotic area at the infection site and suppressed hyphal elongation. Using VIGS, a contig 4211 was identified as a candidate for NecS1 gene encoding a cation/proton exchange protein in barley. Rpg5 was confirmed as a resistance gene after VIGS of this gene in resistant barley resulted in susceptibility to stem rust fungi (Zhang et al., 2009b). Jiang et al. (2017) functionally characterized a putative small

GTP binding protein, TaRab18 from wheat using VIGS and showed that TaRab18 positively regulates stripe rust resistance in wheat by regulating HR. Trihelix is a plant transcription factor family involved in regulation of growth and development, morphogenesis and response to stress. A Trihelix subfamily gene, TaGTγ-3 from Triticum urartu was functionally characterized by Ding et al. (2016) using BSMV-VIGS and showed that TaGTγ-3 positively regulates resistance to stripe rust. Gordon et al. (2016) identified an abscisic acid receptor, pyrabactin resistant-like (Ta_PYL4AS_A) from wheat and found that down-regulation of Ta_PYL4AS_A resulted in an increase in early stage resistance to fusarium head blight and decreased mycotoxin accumulation in wheat. Li et al. (2011a) identified an H2O2 responsive wheat fatty acid desaturase gene (TaFAD) and confirmed its role in powdery mildew resistance by VIGS. Down-regulation of Mlo-transcripts in barley led to higher resistance to Blumeria graminis f. sp. hordei and enhanced susceptibility against M. oryzae (Delventhal et al., 2014). Verticillium wilt is one of the most devastating diseases of cotton worldwide caused by the soilborne fungus Verticillium dahlia. This pathogen is difficult to control due to its broad host range and long-term survival as microsclerotia in soil. Plant respond to Verticillium wilt by activating various signalling pathways like phenylpropanoid, ethylene, brassinosteroids, salicylic acid, jasmonic acid and lignin biosynthesis. Several genes that play a critical role in Verticillium wilt resistance in cotton have been identified and functionally characterized using VIGS. GhNDR1, GhMKK2 (Gao et al., 2011b), GhBAK1 (Gao et al., 2013), MPK9, MPK13, MPK25 (Zhang et al., 2014b), GbRVd (Yang et al., 2016), GhWRKY22, GhWRKY33, GhChitinase, GhCML, GhDirigent (Zhang et al., 2016), GbNAC1 (Wang et al., 2016c), CG02 (Li et al., 2017e) genes functionally characterized using VIGS in cotton are required for Verticillium wilt resistance while GbCAD1, a key enzyme in gossypol biosynthesis (Gao et al., 2013) and HDTF1 (Gao et al., 2016) are found to negatively regulate resistance. Bacteria Pseudomonas syringae causes bacterial speck of tomato. The tomato Pto kinase confers resistance to

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P. syringae pv. tomato by recognizing the pathogen effector proteins AvrPto or AvrPtoB. Involvement of genes- MEK1, MEK2, NTF6 NPR1, TGA1a and TGA2.2 in Pto-mediated disease resistance to P. syringae in tomato was elucidated via TRV-VIGS (Ekengren et al., 2003; Zhu et al., 2010). The role of tomato protein phosphatase type 2A (PP2A) catalytic subunits in the up-regulation of plant defence responses and formation of localized cell death lesions in leaves and stems of N. benthamiana to P. syringae was also determined via VIGS (He et al., 2004). The suppression of salicylic acid accumulation in tomato by coronatine (COR), an effector molecule produced by P. syringae (Uppalapati et al., 2007) was established using VIGS. Involvement of SGT1 and SlALC1 in coronatine/jasmonate ( JA) pathway, and inducible NO synthase (iNOS) in resistance to P. syringae in tomato, was revealed by VIGS (Chandok et al., 2004). Suppression of tomato APR134 gene encoding a CaM-related induced protein in P. syringae pv. tomato resistant tomato, compromised immune response (Chiasson et al., 2005). VIGS was effectively employed for functional analysis of RPS2-dependent resistance to P. syringae in Arabidopsis (Burch-Smith et al., 2006; Cai et al., 2006). N. benthamiana silenced for acotinase, an iron-sulphur protein catalysing the inter-conversion of citrate and isocitrate, delayed HR induced by co-injection of Pto and Avrpto and supported increased levels of P. syringae (Moeder et al., 2007). VIGS of the catalytic subunit of ferredoxin:thioredoxin reductase (FTR) designated -cSlFTR-c led to the formation of necrotic lesions and ROS accumulation in tomato leaves. Moreover, these SlFTR-c-silenced plants displayed enhanced disease resistance to P. syringae pv. tomato DC3000, by the induction of defence-related genes (Lim et al., 2010). VIGS of SGT1 in N. benthamiana positively regulated the process of cell death during both host and non-host interactions with P. syringae (Wang et al., 2010a). Silencing of aconitase gene in N. benthamiana resulted in stunting, necrotic lesions and provided resistance to paraquat, a superoxide generating chemical. Ishiga et al. (2013) showed that silencing of Jasmonate Zim Domain ( JAZ) gene S1JAZ2, S1JAZ6 and S1JAZ7 have no effect on coronatine induced chlorosis in tomato and N. benthamiana and enhanced disease associated cell death to Pst DC3000. During a VIGS based functional analysis of tomato, SR/CAMTA transcription

factors S1SR1L and S1SR3L were found to negatively regulate plant defence response to P. syringae pv. tomato DC3000 (Li et al., 2014b). Silencing of genes encoding LRR receptor kinase called somatic embryogenesis receptor kinase 3 (SlSERK3A or SlSERK3B) resulted in enhanced susceptibility to P. syringae pv. tomato (Pst) DC3000 hrcC indicating that SlSERK3s are positive regulators of defence (Peng and Kaloshian, 2014). Li et al. (2015a) identified and functionally characterized matrix metalloproteinase (MMPs) gene using VIGS and showed that VIGS based down-regulation of S13MMP gene in tomato resulted in reduced resistance to P. syringae pv. tomato DC3000. Xanthomonas campestris pv. vesicatoria causes bacterial spot in different cultivars of chilli pepper (Capsicum annum). Several genes required for defence responses against X. campestris in plants were characterized including a predicted TIR-NBSLRR protein- Bs4 (Schorneck et al., 2004), SGT1 (Leister et al., 2005), cytochrome P450 enzymeCaCYP450A (Hwang and Hwang, 2010), E3 ubiquitin ligase gene CaRING1 (Lee et al., 2011), SlMKK2 and SlMPK2 (Melech-Bonfil et al., 2011) and CCRF-4 associated factor 1 (CAF1) protein belongs to the CCR4–NOT complex, a protein complex controlling transcription and mRNA decay in yeast and mammals. VIGS of CaCAF1 homologue in pepper caused significant growth retardation of plants and enhanced susceptibility to the pepper bacterial spot pathogen X. axonopodis pv. vesicatoria (Sarowar et al., 2007). Silencing of membrane located receptor like protein- CaMRP1 in pepper conferred enhanced basal resistance to Xcv infection, accompanied by induction of genes encoding basic PR1 (CaBPR1), defensin (CaDEF1) and SAR8.2 (CaSAR8.2A) (An et al., 2008). Down-regulation of an endogenous lipid transfer proteins (CALTPI) by VIGS in pepper also enhanced the susceptibility to X. campestris pv. vescatoria (Sarowar et al., 2009). The results of VIGS of gene encoding leucine-rich repeat protein CaLRR1 and a hypersensitive induced reaction protein CaHIR1 suggest that these proteins are probably cell -death regulators associated with response to X. campestris in pepper (Choi et al., 2011). C. annuum mildew resistance locus O (CaMLO2) and calmodulin (CaCaM1) genes are required, respectively for disease associated cell death and hypersensitive cell death. CaMLO2 silencing in pepper (C.

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annuum L. cv. Nockwang) significantly enhanced ROS accumulation, cell death, and resistance responses to X. campestris pv. vesicatoria Ds1 and Ds1 (avrBsT). This was accompanied by induction of CaCaM1, CaPR1 and CaPO2 (Kim et al., 2014). ROS are responsible for mediating cellular defence responses in plants. Local or systemic expression of an extracellular peroxidase gene CaPO2 is induced in pepper by avirulent X. campestris pv. vesicatoria (Xcv) infection. Function of CaPO2 gene in plant defence using the VIGS technique and gain-offunction transgenic plants has been examined. CaPO2-silenced pepper plants were highly susceptible to Xcv infection. VIGS of the CaPO2 gene also compromised hydrogen peroxide (H2O2) accumulation and hypersensitive cell death in leaves, both locally and systemically, during avirulent Xcv infection (Choi et al., 2007). Bacterial wilt (BW) caused by Ralstonia solanacearum is a serious threat to crop production due to its large host range, worldwide occurrence and persistence in fields. It is a soil-borne pathogen which infects tomato, potato, chilli pepper, aubergine and banana. Down-regulation of CaWRKY58 in pepper plants by TRV-VIGS enhanced resistance to a highly virulent strain of R. solanacearum, supporting that it is a transcriptional activator of negative regulators in the resistance of pepper to this bacterium (Wang et al., 2013). VIGS of CaWRKY27, a subgroup of WRKY proteins in bacterial wilt resistant pepper cultivar compromised resistance of plants to R. solanacearum infection. CaWRKY27 is reported to modulate SA-, JA- and ET-mediated signalling pathways and is a positive regulator of plant resistance to R. solanacearum (Dang et al., 2014). Similarly VIGS of CaCDPK15 (Calcium dependent protein kinase), a protein involved in the activation of another WRKY protein – CaWRKY40, significantly increased the susceptibility of pepper to bacterial wilt and repressed the expression of the immunity-associated markers CaNPR1, CaPR1, and CaDEF1 (Shen et al., 2016b). CaCDPK15 is thought to form a positive feedback loop with CaWRKY40. CabZIP63, a pepper bZIP transcription factor, another regulator of CaWRKY40, when silenced also significantly attenuated the resistance of chilli pepper plants to BW, accompanied by down-regulation of CaPR1, CaNPR1 and CaDEF1 (Shen et al., 2016a). VIGS of CaHDZ27, a homeo domain leucine zipper

class I (HD-Zip I) transcription factor in chilli pepper significantly attenuated the resistance to R. solanacearum and down regulated several defencerelated marker genes including CaHIR1, CaACO1, CaPR1, CaPR4, CaPO2 and CaBPR1 suggesting its role as a positive regulator in pepper resistance to bacterial wilt (Mou et al., 2017). Silencing of Hsp90 and SGT-1 in N. benthamiana resulted in significant suppression of wilting symptoms, due to decrease in growth of R. solanacearum and increase in PR proteins, whereas RAR1-silenced plants showed an increase in bacterial growth and multiplication, resulting in the suppression of disease tolerance (Ito et al., 2015). Silencing of ACO1/3, EIN2, ERF3, NPR1, TGA2.2, TGA1a, MKK2, MPK1/2 and MPK3 caused proliferation of Ralstonia in tomato. Plants silenced for TGA2.2, TGA2.1a, MKK2 and MPK1/2 exhibited partial wilting symptoms indicating that ethylene and SA mediated defence pathways have a role in BW resistance of tomato (Chen et al., 2009). Transgenic tomato (Solanum lycopersicon) overexpressing A. thaliana CBF1 (AtCBF1), which confers tolerance to infection by R. solanacearum, constitutively expresses transcription factors RAV, ethylene-responsive factor (ERF) family genes and several pathogenesis-related (PR) genes. VIGS of SlRAV2 and SlERF5 in AtCBF1 transgenic and BW-resistant cultivar Hawaii 7996 plants gave rise to plants with enhanced susceptibility to BW. Knockdown of expression of SlRAV2 inhibited SlERF5 and PR5 gene expression under pathogen infection and significantly decreased BW tolerance. Other genes with functions confirmed using VIGS include NbCD1, a class II ethyleneresponsive element binding factor-like protein that regulates plant cell death and non-host resistance (Nasir et al., 2005). Rice Xa21 gene confers resistance to bacterial leaf blight (BLB) disease. VIGS of Xa21 using DNA virus RTBV vector showed reduction in the transcript level upon challenge inoculation with Xanthomonas oryzae (Kant and Dasgupta, 2017). VIGS was used to identify genes needed for N. benthamiana mediated resistance to X. oryzae pv. Oryzae (Xoo). Identification of ACE (Avr/Cf-elicited) genes and their VIGS analysis results revealed that oxidative burst and calcium-dependent signalling pathways are required for non-host resistance to Xoo (Li et al., 2012). VIGS has also been utilized to identify the role of Dsp A/E, a type III effector

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of Erwinia amylarora in N. benthamiana SGT1dependent cell death (Oh et al., 2007). Nematode and herbivores Mi-1 encodes a CC-NBS-LRR resistance protein and confers resistance to root-knot nematodes (Meloidogyne spp.) and phloem-feeding insects (potato aphids, Macrosiphum euphorbiae, and sweet potato whitefly, Bemisia tabaci). Valentine et al. (2004) demonstrated efficient systemic VIGS in roots of N. benthamiana, Arabidopsis and tomato using a modified TRV retaining helper protein 2b, required for transmission by specific vector nematode. The plant silenced for root development genes-IRT1, transparent testa glabra (TTG1), root hairless1 (RHL1) and beta-tubulin; root meristemless 1 (RML1) and nematode resistance (Mi) had low level of target mRNA. By TRV-based VIGS, factors critical for Mi-1-mediated resistance to nematodes and aphids like HSP90, Sgt1 and WRKY72 (Bhattarai et al., 2007, 2010), MPK1, MPK2, MPK3 (Li et al., 2006a) and somatic embryogenic receptor kinase (SlSERK1) (Mantelin, 2011) were delineated. The functional analysis of Mi-9 using VIGS technology, confirmed the role of this gene in imparting heat-stable resistance to root-knot nematode ( Jablonska et al., 2007). Mantelin et al. (2013) also employed VIGS to demonstrate the role of ethylene (ET) in Mi-1-mediated resistance to M. incognita in tomato. The hormone perception weakened in ET-insensitive Never ripe (Nr) mutant plants, enhanced susceptibility to M. incognita indicating the role of ETR3 in limiting nematode infection. Using BPMV based VIGS vector, Kandoth et al. (2013) identified genes involved in resistance to soybean cyst nematode (SCN) in Glycine max. VIGS also confirmed the role of germin-like protein, threonine deaminase and JAR4 in N. attenuata in conferring resistance to herbivore Manduca sexta (Lou and Baldwin, 2006; Kang et al., 2006). Silencing of tomato MPK1 and MPK2 together by VIGS confirmed their role in systemin-mediated defence response against M. sexta (Kandoth et al., 2007). Involvement of SGT1 (Meldau et al., 2011), MKK1 and MEK2 (Heinrich et al., 2011) and BAK1 (Yang et al., 2011) were also shown to be critical in herbivore M. sexta-induced responses by regulating jasmonic acid ( JA) biosysnthesis. VIGS was used to silence S-nitrosoglutathione

reductase (GSNOR) activity thereby reducing accumulation of JA and ET induced by feeding of M. sexta in N. attenuata. GSNOR mediates methyl jasmonate (MeJA)-induced accumulation of defence related secondary metabolites (Wunsche et al., 2011). Silencing of cytochrome P450 of CYP94 family doubled herbivore induced phytohormone jasmonoyl-l-isoleucine ( JA-Ile) levels. COI1 regulates hydroxylation of JA-Ile, decreasing resistance of N. attenuata plants to Spodoptera litura (Luo et al., 2016). The functional role of gene encoding arabinogalactan protein (AGP) in tomato during attack by plant parasite Cuscuta reflexa was also shown by VIGS (Albert et al., 2006). Zheng et al. (2011) silenced lipoxygenase (AtLOX2), an enzyme of the JA pathway and thioglucoside glucohydrolase: myrosinase (AtTGG1/TGG2), a hydrolytic enzyme responsible for the release of defensive volatile products originating from glucosinolates by heterologous VIGS using sequences from Brassica oleracea. Knocking down of BoLOX and BoMYR, caused better development of the specialist pest, small cabbage white butterfly (Pieris rapae) on plants silenced for both genes while generalist cabbage moth (Mamestra brassicae) larvae grew better on TGG1/TGG2 silenced plants. Galis et al. (2013) described the use of VIGS to screen genes involved in defence response of plants to herbivores, effects of gene silencing on the accumulation of phytohormones, signal transduction, defence metabolites and bioassays with specialist and generalist herbivores. VIGS was used to silence HR-associated candidate genes for Dn1-mediated resistance to Russian wheat aphid (Diuraphis noxia). Phi-class glutathione-S-transferase (TaGSTF6) silenced plants had low H2O2 and increased aphid population, hence involved in Dn1-mediated D. noxia resistance in wheat (Schultz et al., 2015). Plant development VIGS has facilitated the functional characterization of a large number of genes responsible for plant growth and development. VIGS, being a transient assay, is one of the most powerful tools for creating knockouts of genes whose loss-of-function mutations cause embryo- and seedling-lethal phenotypes. Many genes controlling cell division, differentiation, and tissue and organ development have been identified and characterized using VIGS.

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A modified TRV- vector was used to successfully knock out the expression of genes involved in root development (IRT1, TTG1-transparent testa glabra, RHL1-root hairless1 and beta-tubulin) and lateral root-meristem function (RML1-root meristemless1) (Valentine et al., 2004). Calpain (calcium-dependent cysteine protease) homologue of N. benthamiana (NbDEK) controlling the proliferation and differentiation of cells when silenced, arrested organ development and caused hyperplasia (Ahn et al., 2004). Knocking off the expression of PEX12, a peroxisome biogenesis gene, reduced the number of peroxisomes in cells (Fan et al., 2005). Retinoblastoma protein (Rb) is a key factor controlling cell division, cell differentiation, and apoptosis. Retinoblastoma-related gene (RBR) function was identified in plants by heterologous VIGS. Silencing of Rb gene by VIGS retarded plant growth in N. benthamiana (Park et al., 2005). VIGS of a Proliferating Cell Nuclear Antigen (PCNA) meristem gene via ASLV vector led to chlorosis, malformation, and severe dwarfing in newly developed leaves of tobacco. P23K is a monocot-unique protein that is highly expressed in the scutellum of germinating barley seed. VIGS of this gene in barley led to abnormal leaf development, asymmetric orientation of main veins, and cracked leaf edges caused by mechanical weakness establishing its involvement in cell wall polysaccharides and secondary wall formation in leaves (Oikawa et al., 2007). VIGS of genes expressed in roots, leaves and meiotic tissues of wheat was carried out using BSMV vector (Bennypaul et al., 2012). Cellulose synthase catalytic subunits (CESAs) are essential for plant growth, development and disease resistance. Cell wall biosynthesis was investigated by silencing gene encoding Cellulose synthase (Ces A) (Burton et al., 2000; Held et al., 2008) and UDP-D-apiose/UDP-D-xylose synthase (AXS1) in N. benthamiana resulted in yellowing of leaves, growth arrest, cell lysis, disintegration of cellular organelles and excessive thickening of cell walls (Ahn et al., 2006b). Characterization of several cellulose synthase genes in flax (Linum usitatissimum), an economically important fibre crop, became possible due to VIGS technology (Chantreau et al., 2015). The VIGS of two genes NbDUF579 and NbKNAT7 involved in cellulose, hemicellulose and lignin synthesis during secondary cell wall formation in tobacco led to an increase in

proliferation of xylem but with thinner walls. Lines silenced for both the genes exhibited an increase in saccharification. By silencing selected genes in the lignocellulosic pathway, plants can be made to produce higher quantities of bioethanol (Pandey et al., 2016). Silencing of PhCESA3 gene induced in Petunia hybrida via VIGS resulted in swollen stems, pedicels, filaments, styles, epidermal hairs, thickened leaves and corollas; also reduced the length and increased the width of cells suggesting its role in inhibition of elongation and stimulating radial expansion in petunia (Yang et al. 2017a). RPN9, a subunit of 26S proteasome when silenced in N. benthamiana led to the formation of extra leaf veins with increased xylem and decreased phloem partly through the regulation of brassinosteroid and auxin signalling pathways governing vascular formation ( Jin et al., 2006). Silencing the expression of SAMT1 in N. benthamiana resulted in severe growth retardation suggesting that this methylation-related protein has a critical role in plant development (Bouvier et al., 2006). Suppression of Bypass (BPS1) gene also resulted in growth retardation, abnormal leaf development and cell death in N. benthamiana indicating its role in cell division and differentiation in the cambium (Kang et al., 2008). VIGS of an AAA ATPase NgCDC48 gene in tobacco led to severe abnormalities in leaf and shoot development (Bae et al., 2009). VIGS of Ran binding protein NbRanBP1 in N. benthamiana caused stunted growth, yellowing and abnormal morphology of leaves. Confirmation of the involvement of Dt1, an orthologue of pea (Pisum sativum) Terminal Flower 1 (GmTFL1b) in soybean stem growth habit determination was carried out by VIGS (Liu et al., 2010). Silencing of putative RPC5-like subunit of the RNA polymerase III in N. benthamiana adversely affected growth, development and response of plants to the environment. VIGS of RPC5L transcripts brought about pleiotropic defects including severe stunting, vein clearing, chlorosis, shortening of internodes, abnormal leaf shape and defective flowers (Nemchinov et al., 2016). VIGS of late meristem identity 1 gene in G. hirsutum showed that modifications to this gene are responsible for the leaf shapes in cotton (Andres et al., 2017). Wang et al. (2015c) studied the regulatory role of Argonaute 1 (AGO1) in tomato compound leaf development. Morphological defects in

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adaxial-abaxial sides of leaves and trichome development were observed in AGO1 silenced tomato. When RNA exosome components (RRP41 and RRP43) were knocked down using TRV-VIGS, N. benthamiana plants exhibited developmental defects like mosaic and leaf shape distortions, similar to TMV symptoms and were associated with altered levels of different microRNAs (miRNAs) (Conti et al., 2017). The transition from vegetative to reproductive phases at the shoot apical meristem (SAM) is controlled by the interaction of positive and negative regulators, such as LEAFY (LFY), APETALA1 (AP1), FRUITFULL (FUL) and TERMINAL FLOWER1 (TFL1). Ratcliff et al., 2001 successfully silenced Nicotiana FLO (homologue of Antirrhinum majus FLORICAULA) and Arabidopsis LFY using TRV. The role of many flowering genes in tobacco and Petunia hybrida such as flowering time control gene (FY) (Henderson et al., 2005), floral organ identity genes like APETALA3 (AP3) and DEFICIENS (DEF) (Liu et al., 2004b; Kramer et al., 2007), prohibitin (PhPHB1 and PhPHB2) (Chen et al., 2005; Ahn et al., 2006b), cell cycle genes like cell cycle division 5 (CDC5) (Lin et al., 2007) and flower development genes (NbMADS4-1, NbMADS4-2) (Dong et al., 2007) have been deciphered by VIGS. A role for Sqamosa Promoter Binding (SBP)-box protein transcription factor – SPB-1 in the regulation of FUL and LFY homologues was established by VIGS in A. majus. SPB1 silenced plants either flowered very late or failed to flower (Preston and Hileman, 2010). The role of PapsAG-1 in stamen and carpel identity of the opium poppy was also confirmed by VIGS (Hands et al., 2011). Inhibition of an anther-specific lipid transfer protein (LTP) gene called CaMF2 by VIGS in C. annuum resulted in shriveled pollen grains with low fertility and germination indicating that CaMF2 had a vital role in pollen development and may be exploited for developing genetic male sterility (Chen et al., 2011). Chen et al. (2014) efficiently silenced Pea MADS4 (PEAM4) gene using PEBV-VIGS in Pea and showed that it has crucial role in maintaining floral organ identity and flower development. Ripening of fruits is a complex developmental process that depends on coordinated regulation of various genes. Several families of transcription factors such as MADS-box, MYB, AP2/ETHYLENE

RESPONSE FACTOR (ERF), HD-zip, basic helix– loop–helix (bHLH) and auxin response factors (ARFs) and SBP/SPL families DNA methylation and chromatin remodelling factors, participate in the transcriptional regulatory network that modulates ripening (reviewed in Li et al., 2016). Role of ethylene in fruit ripening process and interaction between ethylene and ripening associated developmental factors are reviewed in detail in Liu et al. (2015). VIGS was used to functionally characterize NF-Y (Nuclear Transcription Factor Y) in tomato fruit ripening (Li et al., 2016). Chen et al. (2011) functionally characterized ABA receptor gene FaPYR1 from strawberry and showed that the down-regulation of FaPYR1 gene significantly delayed fruit ripening. Agro infiltration of TRVVIGS vectors into flesh of ripening peach fruits for silencing carotenoid cleavage dioxygenase (CCD4) resulted in yellow coloration (Zhou et al., 2015; Bai et al., 2016). Sepallata genes have essential role in floral organ development and fruit ripening belongs to a subfamily of MADS-box transcription factors. VIGS was used to down regulate the expression of SEP1 gene in melting fresh (MF) peach. SEP1 silenced MF peach fruits remained firm and fruit softening was delayed (Li et al., 2017c). Tsaballa et al., 2011 down-regulated the expression of OVATE like gene CaOvate through VIGS in a pepper cultivar with round fruits and obtained oblong fruits indicating that CaOvate plays a critical role in fruit shape in pepper and is required for the regulation of CaGA20ox1. Utilization of heterosis is a promising approach to improve the yield and quality of wheat. Yang et al. (2017b) identified two MADS-box genes, TaAG-A and TaAG-B, from wheat K-type cytoplasmic male sterile (CMS) line and functionally characterized them using VIGS. Silencing of TaAG-A and TaAG-B in fertile wheat line resulted in green and yellow striped leaves, emanciated spikes and decreased selfing seed set rates. VIGS of a ATP binding cassette (ABC) family C transporter gene TaABCC3 from wheat resulted in reduced grain number by 28% (Walter et al., 2015). Sun et al. (2017b) identified a homologue of auxin response factor 19 ( JcARF19) from Jatropha curcas and functionally characterized its role using VIGS in controlling seed size and seed yield. Qu et al. (2012) investigated the functional role of KATANIN and WRINKLED1 in cotton fibre development using TRV-VIGS and

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showed that KATANIN positively regulates, while WRINKLED1 negatively regulates the cotton fibre development. Cellular functions and metabolic pathways Virus-induced gene silencing has facilitated functional verification of various plant genes involved in cellular functions and metabolic pathways. VIGSbased approach was also used to characterize the functional role of genes- CAF1 and VIP2 involved in transcription (Anand et al., 2007; Sarowar et al., 2007), flavin (Asai et al., 2010), CHMP1 in chromatin structure and modification (Yang et al., 2004), histone H3 (Anand et al., 2007), histone H4 (Lu et al., 2003a), importins-Impα1 and Impα2 in nuclear trafficking (Kanneganti et al., 2007) and AGO1 and AGO4 in systemic RNA silencing ( Jones et al., 2006). Park et al. (2011) investigated physiological role of plastid RNA binding protein (PRBP) using VIGS in N. benthamiana and suggested its role in chloroplast RNA processing and chloroplast biogenesis. Ahn et al. (2015) used VIGS to study the in vivo function of the COPI coatomer complex in plants. In early secretory pathway, COPI vesicles are very essential to the retrograde transport of proteins. Their results showed that COPI vesicles are very essential in growth and survival of plants by modulation of cell plate formation and by maintaining the Golgi apparatus. VIGS of basic transcription factor 3 (TaBTF3) from wheat suggested its role in development of wheat chloroplast, mitochondria and mesophyll cells (Ma et al., 2012). Carotenoids are accessory pigments required in the photosynthesis. Phytoene desaturase (PDS) and phytoene synthase (Psy) are key enzymes of the plant carotenoid biosynthesis pathway. Initial testing and standardization of most VIGS vectors is done by knocking out the expression of PDS gene which causes photobleaching in plants in light (Fig. 14.2). Silencing of an R2R3-MYB transcription factorCaMYB in leaves of pepper cultivar Z1 resulted not only in the loss of anthocyanin but also the repression of MYC and several genes including chalcone synthase (CHS), chalcone isomerase (CHI), flavanone 3-hydroxylase (F3H), flavonoid 3′,5′-hydroxylase (F3′5′’H), dihydroflavonolo 4-reductase (DFR), anthocyanin synthase (ANS), UDP-glucose flavonoid 3-glycosyl transferase (UFGT), anthocyanin

permease (ANP) and glutathione S-transferase (GST), indicating that CaMYB is a key player in the regulation of anthocyanin biosynthesis (Zhang et al., 2015). The production of Capsanthin, the red carotenoid that gives mature pepper fruits their red colour, is regulated mainly by the genes capsanthin/ capsorubin synthase (Ccs), phytoene synthase (Psy), lycopene-β-cyclase (Lcyb) and β-carotene hydroxylase(Crtz). Ccs, Psy, Lcyb and Crtz genes were individually silenced through VIGS technology, to explore the molecular mechanism of pepper fruit colour formation (Tian et al., 2014b). In another study, the function of the CarbcL gene involved in chlorophyll metabolism and the maintenance of balance between chlorophyll and capsanthin levels was determined by VIGS (Wang et al., 2015b). Kim et al. (2017) used TRV-LIC VIGS system to silence an endogenous An2 MYB transcription factor which is responsible for purple colour or anthocyanin-rich pepper, resulting in plants that failed to accumulate purple pigments in leaves, flowers and fruits. Co-silencing of PDS and capsaicin synthase along with An2 resulted in typical photo bleaching in leaves. Co-silencing of endogenous An2 and capsaicin synthase in fruits resulted in decreased levels of capsaicin and hydrocapsaicin and absence of purple pigment in fruits. Recently, VIGS of FaMYB5, a transcription factor responsible for flavonoid metabolism in strawberry fruits, demonstrated that FaMYB5 is a negative regulator of pro anthocyanidins metabolic pathway (Wang et al., 2017b). Silencing of Pun1, involved in regulating pungency in pepper resulted in decreased accumulation of capsaicinoids (Steward et al., 2005). The function of a wheat starch regulator 1 (TaRSR1) in regulating the synthesis of grain storage starch was determined using by BSMV-VIGS. TaRSR1-silenced wheat plants showed a significant increase in the grain starch contents, one thousand kernel weight, grain length and width suggesting that TaRSR1 is a negative regulator and plays a vital role in starch synthesis in wheat grains (Liu et al., 2016a). George et al. (2012) used VIGS to silent genes in starch degradation pathway. Silencing of disproportionating enzyme 1 (DPE1) and disproportionating enzyme 2 (DPE2) in N. benthamiana resulted in a near complete suppression in starch and malto-oligosachharides accumulation. ADPglucose pyrophosphorylase (AGPase) is another

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Figure 14.2  VIGS of PDS in (A) chilli, (B) tomato, (C) cotton and (D) N. benthamiana.

gene that regulates starch biosynthesis in storage organs and is an important determinant of sink strength. VIGS of an AGPase gene (GhAGPS1) from gladiolus resulted in the lowered corm quality and cormel yield. VIGS was also used to elucidate the cellular functions of genes involved in chloroplasts and mitochondria biogenesis (Cho et al., 2004; Ahn et al., 2005, 2006a; Kim et al., 2005; Arsova et al., 2010; Jeon et al., 2010; Kang et al., 2010; Park et al., 2011), plastid biogenesis (Kim et al., 2005; Bouvier et al., 2006), peroxisome biogenesis (Fan et al., 2005) and membrane biogenesis (Park et al., 2005). Benzoxazinoids (BXs) are a group of natural chemical compounds with insecticidal, antimicrobial, and allelopathic activities synthesized in grasses including crops such as maize, wheat and rye, as well as a few dicot species (Adhikari et al., 2015). Bx1 is one of the three genes that encodes an enzyme with indole-3-glycerol phosphate lyase activity that

catalyses the first step in the BXs pathway and its expression is developmentally regulated. Transcriptional and post-transcriptional silencing of the rye ScBx1 gene using BSMV-VIGS lowered the transcript levels and BXs accumulation. Infection with BSMV induced the expression of ScBx1 and stimulated BXs accumulation indicating that Bx1 expression is induced by biotic stress (Groszyk et al., 2017). In recent study, VIGS was used to silence two squalene synthase isoforms of apple tree to determine the biological function of phytosterols in plastid pigmentation and leaf development (Navarro-Gallón et al., 2017). VIGS has been utilized for functional analysis of several metabolic pathways including alkaloid biosynthesis (Li et al., 2006b; Todd et al., 2010; Liscombe and O’Connor, 2011), isoprenoid biosynthesis (Page et al., 2004; Ahn et al., 2008), ascorbic acid biosynthesis (Qian et al., 2007) and sterol biosynthesis (Burger et al., 2003; Darnet et al., 2004).

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Singh et al. (2017) silenced WRKY1 transcription factor in Withania somnifera and demonstrated that WRKY1 has positive regulatory role in withanolides biosynthesis, defence against biotic stress and phytosterol biosynthesis. Dang and Facchini (2014) isolated and characterized a cytochrome P450 (CYP719A21) from opium poppy. VIGS of CYP719A21 resulted in significant increase in (S)-tetrahydrocolumbamine accumulation and decrease in level of downstream intermediates and noscapine in opium poppy. Wijekoon and Facchini (2012) employ VIGS to investigate the regulation of morphine biosynthesis in opium poppy by confirming the physiological function of the enzymes responsible for the final steps in the pathway. Liscombe and O’Connor (2011) used VIGS approach to investigate the function of vindoline biosynthetic genes in Catharanthus roseus. Limonoides are group of highly bioactive secondary metabolites produced by citrus which provides health benefits for humans. Wang et al. (2017a) identified putative genes involved in limonoid biosynthesis using VIGS in citrus. Silencing of CiOSC gene in citrus significantly reduce limonoid content showing its role in biosynthesis of limonoids. Jeon et al. (2012) used VIGS to examine the physiological and biochemical function of S1 domain containing transcription stimulating factor (STF) from N. benthamiana. Their results showed that in regulation of plastid transcription and chloroplast biosynthesis, STF plays an important role as an auxiliary factor of plastid encoded multimeric RNA polymerase (PEP) transcription complex. VIGS was used to identify the genes involved in programmed cell death and apoptosis, including mitochondrial-associated hexokinase Hxk1 gene (Kim et al., 2006a), two prohibitin subunit genes PHB1 and PHB2 (Ahn et al., 2006a); the larger sub complexes, the 20S proteasome and the 19S regulatory complex of the 26S proteasome (Kim et al., 2005); and a Myb-related gene CDC5 (Lin et al., 2007). Advantages of VIGS • VIGS needs much less time and effort to achieve gene knockouts than to generate mutants or RNAi transgenic lines. VIGS vectors that use

Gateway based/LIC are available making this technique inexpensive and high throughput. Depending on the type and number of plants that are to be targeted one can choose different VIGS methods such as infiltration in leaves or agro spray (Liu et al., 2002a) or agro drench (Ryu et al., 2004; Senthil-Kumar, 2007) or toothpick inoculations (Lu et al., 2003a; BurchSmith et al., 2004) or a combination of methods (Zhang et al., 2017). • By carefully selecting the target sequences cloned into the VIGS vector, it is possible to silence multiple genes from the same family to overcome functional redundancy. Similarly, among closely related sequences, silencing of a specific gene is possible by selecting the sequences that do not produce 21bp nucleotide having 100% similarity with other related genes. • VIGS only modulates transcript levels of the target genes and therefore it is highly suitable for studying biological functions of genes without altering the genome sequence of the plant. A small number of plants are required to elucidate gene function using VIGS. Therefore, repeating an experiment is cost effective and easy. • VIGS is very useful in plant species that are recalcitrant and difficult to transform. Gene knockdowns using VIGS are dominant whereas the insertion or loss of function mutations are recessive. Gene knockdown in polyploid genomes that contains four or more orthologs of a gene can be achieved easily by VIGS when compared to traditional mutagenesis approaches. Limitations of VIGS and its remedies • VIGS can only be used if the sequence information of the specific target gene is available. • VIGS only down regulate 75–90% transcript level of the target gene, complete loss of function by VIGS is not achieved. • VIGS is usually performed on seedlings, hence genes expressed during germination or in early seedling stage cannot be analysed. VIGS is transient and phenotype is not normally inherited. • VIGS phenotype is not very stable and its efficiency may vary with plant species.

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Conclusion and future prospects VIGS is a rapid, simple and versatile tool for plant functional genomics employed to characterize the genes involved in various processes. VIGS has made possible the identification and unravelling the function of hitherto unknown genes involved in plant-environment and plant–pathogen interactions, plant development, metabolic processes and other cellular processes. Efforts to develop new VIGS vectors for silencing genes in more plant species of interest are under way by plant virologists and molecular biologists. Efforts to widen the host range and silencing efficiency of VIGS vectors by modify the existing vectors and by combining more than one inoculation methods have also proved to be effective. The applicability of VIGS in plant biology will continue to increase as new vectors and protocols are developed for gene silencing in more plant species. Acknowledgements The authors regret that it was not possible to cite all the excellent work done by various scientific groups including our colleagues in the area of virusinduced gene silencing in plants owing to space limitations. References

Adhikari, K.B., Tanwir, F., Gregersen, P.L., Steffensen, S.K., Jensen, B.M., Poulsen, L.K., Nielsen, C.H., Høyer, S., Borre, M., and Fomsgaard, I.S. (2015). Benzoxazinoids: Cereal phytochemicals with putative therapeutic and health-protecting properties. Mol. N. Food Res. 59, 1324–1338. https://doi.org/10.1002/mnfr.201400717 Aguilar, E., Del Toro, F.J., Canto, T., and Tenllado, F. (2017). Identification of MAPKs as signal transduction components required for the cell death response during compatible infection by the synergistic pair Potato virus X - Potato virus Y. Virology 509, 178–184. Ahn, C.S., and Pai, H.S. (2008). Physiological function of IspE, a plastid MEP pathway gene for isoprenoid biosynthesis, in organelle biogenesis and cell morphogenesis in Nicotiana benthamiana. Plant Mol. Biol. 66, 503–517. https://doi.org/10.1007/s11103007-9286-0 Ahn, C.S., Lee, J.H., and Pai, H.S. (2005). Silencing of NbNAP1 encoding a plastidic Suf B-like protein affects chloroplast development in Nicotiana benthamiana. Mol. Cells 20, 112–118. Ahn, C.S., Lee, J.H., Reum Hwang, A., Kim, W.T., and Pai, H.S. (2006a). Prohibitin is involved in mitochondrial biogenesis in plants. Plant J. 46, 658–667. Ahn, H.K., Kang, Y.W., Lim, H.M., Hwang, I., and Pai, H.S. (2015). Physiological functions of the COPI complex in higher plants. Mol. Cells 38, 866–875. https://doi. org/10.14348/molcells.2015.0115

Ahn, J.W., Kim, M., Lim, J.H., Kim, G.T., and Pai, H.S. (2004). Phytocalpain controls the proliferation and differentiation fates of cells in plant organ development. Plant J. 38, 969–981. https://doi.org/10.1111/j.1365313X.2004.02102.x Ahn, J.W., Verma, R., Kim, M., Lee, J.Y., Kim, Y.K., Bang, J.W., Reiter, W.D., and Pai, H.S. (2006b). Depletion of UDP-D-apiose/UDP-D-xylose synthases results in rhamnogalacturonan-II deficiency, cell wall thickening, and cell death in higher plants. J. Biol. Chem. 281, 13708–13716. Albert, M., Belastegui-Macadam, X., and Kaldenhoff, R. (2006). An attack of the plant parasite Cuscuta reflexa induces the expression of attAGP, an attachment protein of the host tomato. Plant J. 48, 548–556. Aly, R., Dubey, N.K., Yahyaa, M., Abu-Nassar, J., and Ibdah, M. (2014). Gene silencing of CCD7 and CCD8 in Phelipanche aegyptiaca by Tobacco rattle virus system retarded the parasite development on the host. Plant Signal Behav 9, e29376. https://doi.org/10.4161/ psb.29376 An, S.H., Choi, H.W., Hwang, I.S., Hong, J.K., and Hwang, B.K. (2008). A novel pepper membrane-located receptor-like protein gene CaMRP1 is required for disease susceptibility, methyl jasmonate insensitivity and salt tolerance. Plant Mol. Biol. 67, 519–533. https:// doi.org/10.1007/s11103-008-9337-1 Anand, A., Vaghchhipawala, Z., Ryu, C.M., Kang, L., Wang, K., del-Pozo, O., Martin, G.B., and Mysore, K.S. (2007). Identification and characterization of plant genes involved in Agrobacterium-mediated plant transformation by virus-induced gene silencing. Mol. Plant Microbe Interact. 20, 41–52. https://doi. org/10.1094/MPMI-20-0041 Anandalakshmi, R., Pruss, G.J., Ge, X., Marathe, R., Mallory, A.C., Smith, T.H., and Vance, V.B. (1998). A viral suppressor of gene silencing in plants. Proc. Natl. Acad. Sci. U.S.A. 95,13079–13084. Andres, R.J., Coneva, V., Frank, M.H., Tuttle, J.R., Samayoa, L.F., Han, S.W., Kaur, B., Zhu, L., Fang, H., Bowman, D.T., et al. (2017). Modifications to a LATE MERISTEM IDENTITY1 gene are responsible for the major leaf shapes of Upland cotton (Gossypium hirsutum L.). Proc. Natl. Acad. Sci. U.S.A. 114, E57–E66. https:// doi.org/10.1073/pnas.1613593114 Angell, S.M., and Baulcombe, D.C. (1999). Technical advance: potato virus X amplicon-mediated silencing of nuclear genes. Plant J. 20, 357–362. Anu, K., Jessymol, K.K., Chidambareswaren, M., Gayathri, G.S., and Manjula, S. (2015). Down-regulation of osmotin (PR5) gene by virus-induced gene silencing (VIGS) leads to susceptibility of resistant Piper colubrinum Link. to the oomycete pathogen Phytophthora capsici Leonian. Indian J. Exp. Biol. 53, 329–334. Arsova, B., Hoja, U., Wimmelbacher, M., Greiner, E., Ustün, S., Melzer, M., Petersen, K., Lein, W., and Börnke, F. (2010). Plastidial thioredoxin z interacts with two fructokinase-like proteins in a thiol-dependent manner: evidence for an essential role in chloroplast development in Arabidopsis and Nicotiana benthamiana. Plant Cell 22, 1498–1515. https://doi.org/10.1105/tpc.109.071001 Asai, S., Mase, K., and Yoshioka, H. (2010). A key enzyme for flavin synthesis is required for nitric oxide and

Virus-Induced Gene Silencing and its Applications |  307

reactive oxygen species production in disease resistance. Plant J. 62, 911–924. https://doi.org/10.1111/j.09607412.2010.04206.x Atkinson, R.G., Gleave, L.R.F.B.A.P., Janssen, B.J., and Morris, B.A.M. (1998). Post-transcriptional silencing of chalcone synthase in petunia using a geminivirusbased episomal vector. Plant J. 15, 593–604. https://doi. org/10.1046/j.1365-313x.1998.00211.x Atwood, S.E., O’Rourke, J.A., Peiffer, G.A., Yin, T., Majumder, M., Zhang, C., Cianzio, S.R., Hill, J.H., Cook, D., Whitham, S.A., et al. (2014). Replication protein A subunit 3 and the iron efficiency response in soybean. Plant Cell Environ. 37, 213–234. https://doi. org/10.1111/pce.12147 Bachan, S., and Dinesh-Kumar, S.P. (2012). Tobacco rattle virus (TRV)-based virus-induced gene silencing. Methods Mol. Biol. 894, 83–92. https://doi. org/10.1007/978-1-61779-882-5_6 Bae, H., Choi, S.M., Yang, S.W., Pai, H.S., and Kim, W.T. (2009). Suppression of the ER-localized AAA ATPase NgCDC48 inhibits tobacco growth and development. Mol. Cells 28, 57–65. https://doi.org/10.1007/s10059009-0101-4 Bahieldin, A., Alqarni, D.A., Atef, A., Gadalla, N.O., Al-matary, M., Edris, S., Al-Kordy, M.A., Makki, R.M., Al-Doss, A.A., Sabir, J.S., et al. (2016). Suppression of PCD-related genes affects salt tolerance in Arabidopsis. C. R. Biol. 339, 105–114. https://doi.org/10.1016/j. crvi.2016.02.004 Bai, S., Tuan, P.A., Tatsuki, M., Yaegaki, H., Ohmiya, A., Yamamizo, C., and Moriguchi, T. (2016). Knockdown of Carotenoid Cleavage Dioxygenase 4 (CCD4) via Virusinduced gene silencing confers yellow coloration in peach fruit: evaluation of gene function related to fruit traits. Plant Mol Biol Rep. 34, 257–264. https://doi. org/10.1007/s11105-015-0920-8 Bao, H., Chen, X., Lv, S., Jiang, P., Feng, J., Fan, P., Nie, L., and Li, Y. (2015). Virus-induced gene silencing reveals control of reactive oxygen species accumulation and salt tolerance in tomato by γ-aminobutyric acid metabolic pathway. Plant Cell Environ. 38, 600–613. https://doi. org/10.1111/pce.12419 Barciszewska-Pacak, M., Jarmołowski, A., and Pacak, A. (2016). Virus-Induced gene silencing for gene function studies in barley. methods. Mol Biol. 1398, 293–308. https://doi.org/10.1007/978-1-4939-3356-3_22 Baulcombe, D.C. (1999). Fast forward genetics based on virus-induced gene silencing. Curr. Opin. Plant Biol. 2, 109–113. Becker, A., and Lange, M. (2010). VIGS – genomics goes functional. Trends Plant Sci. 15, 1–4. https://doi. org/10.1016/j.tplants.2009.09.002 Bello, B., Zhang, X., Liu, C., Yang, Z., Yang, Z., Wang, Q., Zhao, G., and Li, F. (2014). Cloning of Gossypium hirsutum sucrose non-fermenting 1-related protein kinase 2 gene (GhSnRK2) and its overexpression in transgenic Arabidopsis escalates drought and low temperature tolerance. PLOS ONE9, e112269. https:// doi.org/10.1371/journal.pone.0112269 Benavente, L.M., Ding, X.S., Redinbaugh, M.G., Nelson, R.S., and Balint-Kurti, P. (2012). Virus-induced gene silencing in diverse maize lines using the Brome mosaic virus-based silencing vector. Maydica 57, 206–214.

Bennypaul, H.S., Mutti, J.S., Rustgi, S., Kumar, N.,Okubara, P.A.,and Gill, K.S. (2012). Virus-induced gene silencing (VIGS) of genes expressed in root, leaf, and meiotic tissues of wheat. Funct. Integr. Genomics 12, 143–156. https://doi.org/10.1007/s10142-011-0245-0 Beyene, G., Chauhan, R.D., and Taylor, N.J. (2017). A rapid virus-induced gene silencing (VIGS) method for assessing resistance and susceptibility to cassava mosaic disease. Virol. J. 14, 47. https://doi.org/10.1186/ s12985-017-0716-6 Bhattarai, K.K., Atamian, H.S., Kaloshian, I., and Eulgem, T. (2010). WRKY72-type transcription factors contribute to basal immunity in tomato and Arabidopsis as well as gene-for-gene resistance mediated by the tomato R gene Mi-1. Plant J. 63, 229–240. https://doi.org/10.1111/ j.1365-313X.2010.04232.x Bhattarai, K.K., Li, Q., Liu, Y., Dinesh-Kumar, S.P., and Kaloshian, I. (2007). The MI-1-mediated pest resistance requires Hsp 90 and Sgt1. Plant Physiol. 144, 312–323. Bilichak, A., and Kovalchuk, I. (2017). Increasing a Stable Transformation Efficiency of Arabidopsis by Manipulating the Endogenous Gene Expression Using Virus-Induced Gene Silencing. Methods Mol. Biol. 1456, 225–236. https://doi.org/10.1007/978-1-48997708-3_17 Biruma, M., Martin, T., Fridborg, I., Okori, P., and Dixelius, C. (2012). Two loci in sorghum with NB-LRR encoding genes confer resistance to Colletotrichum sublineolum. Theor. Appl. Genet. 124, 1005–1015. https://doi. org/10.1007/s00122-011-1764-8 Bouvier, F., Linka, N., Isner, J.C., Mutterer, J., Weber, A.P., and Camara, B. (2006). Arabidopsis SAMT1 defines a plastid transporter regulating plastid biogenesis and plant development. Plant Cell 18, 3088–3105. Braunstein, T.H., Moury, B., Johannessen, M., and Albrechtsen, M. (2002). Specific degradation of 3’ regions of GUS mRNA in posttranscriptionally silenced tobacco lines may be related to 5’-3’ spreading of silencing. RNA 8, 1034–1044. Brigneti, G., Martín-Hernández, A.M., Jin, H., Chen, J., Baulcombe, D.C., Baker, B., and Jones, J.D. (2004). Virus-induced gene silencing in Solanum species. Plant J. 39, 264–272. https://doi.org/10.1111/j.1365313X.2004.02122.x Broderick, S.R., and Jones, M.L. (2014). An Optimized protocol to increase virus-induced gene silencing efficiency and minimize viral symptoms in petunia. Plant Mol. Biol. Rep. 32, 219–233. https://doi.org/10.1007/ s11105-013-0647-3 Bruun-Rasmussen, M., Madsen, C.T., Jessing, S., and Albrechtsen, M. (2007). Stability of Barley stripe mosaic virus-induced gene silencing in barley. Mol. Plant Microbe Interact. 20,1323–1331. https://doi. org/10.1094/MPMI-20-11-1323 Buhrow, L.M., Clark, S.M., and Loewen, M.C. (2016). Identification of an attenuated barley stripe mosaic virus for the virus-induced gene silencing of pathogenesisrelated wheat genes. Plant Methods 12, 12. https://doi. org/10.1186/s13007-016-0112-z Burch-Smith, T.M., Anderson, J.C., Martin, G.B., and Dinesh-Kumar, S.P. (2004). Applications and advantages of virus-induced gene silencing for gene

308  | Kumar et al.

function studies in plants. Plant J. 39, 734–746. https:// doi.org/10.1111/j.1365-313X.2004.02158.x Burch-Smith, T.M., Schiff, M., Liu, Y., and Dinesh-Kumar, S.P. (2006). Efficient virus-induced gene silencing in Arabidopsis. Plant Physiol. 142, 21–27. Burger, C., Rondet, S., Benveniste, P., and Schaller, H. (2003). Virus-induced silencing of sterol biosynthetic genes: identification of a Nicotiana tabacum L. obtusifoliol-14alpha-demethylase (CYP51) by genetic manipulation of the sterol biosynthetic pathway in Nicotiana benthamiana L. J. Exp. Bot. 54, 1675–1683. https://doi.org/10.1093/jxb/erg184 Burton, R.A., Gibeaut, D.M., Bacic, A., Findlay, K., Roberts, K., Hamilton, A., Baulcombe, D.C., and Fincher, G.B. (2000). Virus-induced silencing of a plant cellulose synthase gene. Plant Cell 12, 691–706. Cai, C., Wu, S., Niu, E., Cheng, C., and Guo, W. (2017). Identification of genes related to salt stress tolerance using intron-length polymorphic markers, association mapping and virus-induced gene silencing in cotton. Sci. Rep. 7, 528. https://doi.org/10.1038/s41598-01700617-7 Cai, X., Wang, C., Xu, Y., Xu, Q., Zheng, Z., and Zhou, X. (2007). Efficient gene silencing induction in tomato by a viral satellite DNA vector. Virus Res. 125, 169–175. Cai, X.Z., Xu, Q.F., Wang, C.C., and Zheng, Z. (2006). Development of a virus-induced gene-silencing system for functional analysis of the RPS2-dependent resistance signalling pathways in Arabidopsis. Plant Mol. Biol. 62, 223–232. https://doi.org/10.1007/s11103-006-9016-z Cakir, C., Gillespie, M.E., and Scofield, S.R. (2010). Rapid determination of gene function by virus-induced gene silencing in wheat and barley. Crop Sci. 50, S-77–S-84. https://doi.org/10.2135/cropsci2009.10.0567 Cao, X., Zhou, P., Zhang, X., Zhu, S., Zhong, X., Xiao, Q., Ding, B., and Li, Y. (2005). Identification of an RNA silencing suppressor from a plant double-stranded RNA virus. J. Virol. 79, 13018–13027. Caplan, J.L., Mamillapalli, P., Burch-Smith, T.M., Czymmek, K., and Dinesh-Kumar, S.P. (2008). Chloroplastic protein NRIP1 mediates innate immune receptor recognition of a viral effector. Cell 132, 449–462. https://doi.org/10.1016/j.cell.2007.12.031 Caro, M., Verlaan, M.G., Julián, O., Finkers, R., Wolters, A.A., Hutton, S.F., Scott, J.W., Kormelink, R., Visser, R.G.F., Díez, M.J., et al. (2015). Assessing the genetic variation of Ty-1 and Ty-3 alleles conferring resistance to Tomato yellow leaf curl virus in a broad tomato germplasm. Mol. Breed. 35, 132. https://doi.org/10.1007/s11032-0150329-y Carqueijeiro, I., Masini, E., Foureau, E., Sepúlveda, L.J., Marais, E., Lanoue, A., Besseau, S., Papon, N., Clastre, M., Dugé de Bernonville, T., et al. (2015). Virus-induced gene silencing in Catharanthus roseus by biolistic inoculation of Tobacco rattle virus vectors. Plant Biol. 17,1242–1246. https://doi.org/10.1111/plb.12380 Carrillo-Tripp, J., Shimada-Beltrán, H., and RiveraBustamante, R. (2006). Use of geminiviral vectors for functional genomics. Curr. Opin. Plant Biol. 9, 209–215. Chai, Y.M., Jia, H.F., Li, C.L., Dong, Q.H., and Shen, Y.Y. (2011). FaPYR1 is involved in strawberry fruit ripening.

J. Exp. Bot. 62, 5079–5089. https://doi.org/10.1093/ jxb/err207 Chandok, M.R., Ekengren, S.K., Martin, G.B., and Klessig, D.F. (2004). Suppression of pathogen-inducible NO synthase (iNOS) activity in tomato increases susceptibility to Pseudomonas syringae. Proc. Natl. Acad. Sci. U.S.A. 101, 8239–8244. https://doi.org/10.1073/ pnas.0402344101 Chantreau, M., Chabbert, B., Billiard, S., Hawkins, S., and Neutelings, G. (2015). Functional analyses of cellulose synthase genes in flax (Linum vsitatissimum) by virusinduced gene silencing. Plant Biotechnol. J. 13, 1312–24. https://doi.org/10.1111/pbi.12350 Chapman, S., Kavanagh, T., and Baulcombe, D. (1992). Potato virus X as a vector for gene expression in plants. Plant J. 2, 549–557. Chen, C., Chen, G., Hao, X., Cao, B., Chen, Q., Liu, S., and Lei, J. (2011). CaMF2, an anther-specific lipid transfer protein (LTP) gene, affects pollen development in Capsicum annuum L. Plant Sci. 181, 439–448. https:// doi.org/10.1016/j.plantsci.2011.07.003 Chen, J., Watanabe, Y., Sako, N., Ohshima, K., and Okada, Y. (1996). Complete nucleotide sequence and synthesis of infectious in vitro transcripts from a full-length cDNA clone of a rakkyo strain of Tobacco mosaic virus. Arch. Virol. 141, 885–900. Chen, J.C., Jiang, C.Z., and Reid, M.S. (2005). Silencing a prohibitin alters plant development and senescence. Plant J. 44, 16–24. Chen, J.C., Jiang, C.Z., Gookin, T.E., Hunter, D.A., Clark, D.G., and Reid, M.S. (2004). Chalcone synthase as a reporter in virus-induced gene silencing studies of flower senescence. Plant Mol. Biol. 55, 521–530. Chen, K., Li, H., Chen, Y., Zheng, Q., Li, B., and Li, Z. (2015). TaSCL14, a novel wheat (Triticum aestivum L.) GRAS gene, regulates plant growth, photosynthesis, tolerance to photooxidative stress, and senescence. J. Genet. Genomics 42, 21–32. https://doi.org/10.1016/j. jgg.2014.11.002 Chen, Y.Y., Lin, Y.M., Chao, T.C., Wang, J.F., Liu, A.C., Ho, F.I., and Cheng, C.P. (2009). Virus-induced gene silencing reveals the involvement of ethylene-, salicylic acid- and mitogen-activated protein kinaserelated defense pathways in the resistance of tomato to bacterial wilt. Physiol. Plant. 136, 324–335. https://doi. org/10.1111/j.1399-3054.2009.01226.x Chen, Z.H., Jia, F.F., Hu, J.Q., Pang, J.L., Xu, L., and Wang, L.L. (2014). Virus-induced gene silencing of PEAM4 affects floral morphology by altering the expression pattern of PsSOC1a and PsPVP in pea. J. Plant Physiol. 171, 148–153. https://doi.org/10.1016/j. jplph.2013.08.010 Cheng, F., Zhou, Y.H., Xia, X.J., Shi, K., Zhou, J., and Yu, J.Q. (2014). Chloroplastic thioredoxin-f and thioredoxin-m1/4 play important roles in brassinosteroids-induced changes in CO2 assimilation and cellular redox homeostasis in tomato. J. Exp. Bot. 65, 4335–4347. https://doi.org/10.1093/jxb/eru207 Chiasson, D., Ekengren, S.K., Martin, G.B., Dobney, S.L., and Snedden, W.A. (2005). Calmodulin-like proteins from Arabidopsis and tomato are involved in host defense

Virus-Induced Gene Silencing and its Applications |  309

against Pseudomonas syringae pv. tomato. Plant Mol. Biol. 58, 887–897. https://doi.org/10.1007/s11103-0058395-x Cho, H.S., Lee, S.S., Kim, K.D., Hwang, I., Lim, J.S., Park, Y.I., and Pai, H.S. (2004). DNA gyrase is involved in chloroplast nucleoid partitioning. Plant Cell 16, 2665– 2682. https://doi.org/10.1105/tpc.104.024281 Choi, H.W., and Hwang, B.K. (2012). The pepper extracellular peroxidase CaPO2 is required for salt, drought and oxidative stress tolerance as well as resistance to fungal pathogens. Planta 235, 1369–1382. https://doi.org/10.1007/s00425-011-1580-z Choi, H.W., Kim, Y.J., and Hwang, B.K. (2011). The hypersensitive induced reaction and leucine-rich repeat proteins regulate plant cell death associated with disease and plant immunity. Mol. Plant Microbe Interact. 24, 68–78. https://doi.org/10.1094/MPMI-02-10-0030 Choi, H.W., Kim, Y.J., Lee, S.C., Hong, J.K., and Hwang, B.K. (2007). Hydrogen peroxide generation by the pepper extracellular peroxidase CaPO2 activates local and systemic cell death and defense response to bacterial pathogens. Plant Physiol. 145, 890–904. Chung, E., Seong, E., Kim, Y.C., Chung, E.J., Oh, S.K., Lee, S., Park, J.M., Joung, Y.H., and Choi, D. (2004). A method of high frequency virus-induced gene silencing in chili pepper (Capsicum annuum L. cv. Bukang). Mol. Cells 17, 377–380. Colcombet, J., and Hirt, H. (2008). Arabidopsis MAPKs: a complex signalling network involved in multiple biological processes. Biochem. J. 413, 217–226. https:// doi.org/10.1042/BJ20080625 Constantin, G.D., Grønlund, M., Johansen, I.E., Stougaard, J., and Lund, O.S. (2008). Virus-induced gene silencing (VIGS) as a reverse genetic tool to study development of symbiotic root nodules. Mol. Plant Microbe Interact. 21, 720–727. https://doi.org/10.1094/MPMI-21-6-0720 Constantin, G.D., Krath, B.N., MacFarlane, S.A., Nicolaisen, M., Johansen, I.E., and Lund, O.S. (2004). Virusinduced gene silencing as a tool for functional genomics in a legume species. Plant J. 40, 622–631. Conti, G., Zavallo, D., Venturuzzi, A.L., Rodriguez, M.C., Crespi, M., and Asurmendi, S. (2017). TMV induces RNA decay pathways to modulate gene silencing and disease symptoms. Plant J. 89, 73–84. https://doi. org/10.1111/tpj.13323 Corbin, C., Lafontaine, F., Sepúlveda, L.J., Carqueijeiro, I., Courtois, M., Lanoue, A., Dugé de Bernonville, T., Besseau, S., Glévarec, G., Papon, N., et al. (2017). Virusinduced gene silencing in Rauwolfia species. Protoplasma 254, 1813–1818. https://doi.org/10.1007/s00709017-1079-y Cui, H., and Wang, A. (2017). An efficient viral vector for functional genomic studies of Prunus fruit trees and its induced resistance to Plum pox virus via silencing of a host factor gene. Plant Biotechnol. J. 15, 344–356. https://doi.org/10.1111/pbi.12629 Czosnek, H., Eybishtz, A., Sade, D., Gorovits, R., Sobol, I., Bejarano, E., Rosas-Díaz, T., and Lozano-Durán, R. (2013). Discovering host genes involved in the infection by the Tomato yellow leaf curl virus complex and in the establishment of resistance to the virus using Tobacco

rattle virus-based post transcriptional gene silencing. Viruses 5, 998–1022. https://doi.org/10.3390/ v5030998 Dai, F., Zhang, C., Jiang, X., Kang, M., Yin, X., Lü, P., Zhang, X., Zheng, Y., and Gao, J. (2012). RhNAC2 and RhEXPA4 are involved in the regulation of dehydration tolerance during the expansion of rose petals. Plant Physiol. 160, 2064–2082. https://doi.org/10.1104/ pp.112.207720 Dang, F., Wang, Y., She, J., Lei, Y., Liu, Z., Eulgem, T., Lai, Y., Lin, J., Yu, L., Lei, D., et al. (2014). Overexpression of CaWRKY27, a subgroup IIe WRKY transcription factor of Capsicum annuum, positively regulates tobacco resistance to Ralstonia solanacearum infection. Physiol Plant 150, 397–411. https://doi.org/10.1111/ ppl.12093 Dang, T.T., and Facchini, P.J. (2014). Cloning and characterization of canadine synthase involved in noscapine biosynthesis in opium poppy. FEBS Lett. 588, 198–204. https://doi.org/10.1016/j. febslet.2013.11.037 Darnet, S., and Rahier, A. (2004). Plant sterol biosynthesis: identification of two distinct families of sterol 4alphamethyl oxidases. Biochem J. 378, 889–898. https://doi. org/10.1042/BJ20031572 Dasgupta, I., Hull, R., Eastop, S., Poggi-Pollini, C., Blakebrough, M., Boulton, M.I., and Davies, J.W. (1991). Rice tungro bacilliform virus DNA independently infects rice after Agrobacterium-mediated transfer. J. Gen. Virol. 72, 1215–1221. https://doi.org/10.1099/0022-131772-6-1215 Dawson, W.O., and Folimonova, S.Y. (2013). Virusbased transient expression vectors for woody crops: a new frontier for vector design and use. Annu Rev Phytopathol 51, 321–337. https://doi.org/10.1146/ annurev-phyto-082712-102329 Dean, C., Pichersky, E., and Dunsmuir, P. (1989). Structure, evolution, and regulation of Rbcs genes in Higher-Plants. Annu. Rev. Pl. Physiol. Plant Mol. Biol. 40, 415–439. del Rosario Abraham-Juárez, M., del Carmen RochaGranados, M., López, M.G., Rivera-Bustamante, R.F., and Ochoa-Alejo, N. (2008). Virus-induced silencing of Comt, pAmt and Kas genes results in a reduction of capsaicinoid accumulation in chili pepper fruits. Planta 227, 681–695. https://doi.org/10.1007/s00425-0070651-7 Delventhal, R., Falter, C., Strugala, R., Zellerhoff, N., and Schaffrath, U. (2014). Ectoparasitic growth of Magnaporthe on barley triggers expression of the putative barley wax biosynthesis gene CYP96B22 which is involved in penetration resistance. BMC Plant Biol. 14, 26. https://doi.org/10.1186/1471-2229-14-26 Delventhal, R., Zellerhoff, N., and Schaffrath, U. (2011). Barley stripe mosaic virus-induced gene silencing (BSMV-IGS) as a tool for functional analysis of barley genes potentially involved in nonhost resistance. Plant Signal Behav. 6, 867–869. Deng, X., Elomaa, P., Nguyen, C.X., Hytönen, T., Valkonen, J.P., and Teeri, T.H. (2012). Virus-induced gene silencing for Asteraceae – a reverse genetics approach for functional genomics in Gerbera hybrida. Plant

310  | Kumar et al.

Biotechnol. J. 10, 970–978. https://doi.org/10.1111/ j.1467-7652.2012.00726.x Deshmukh, Y., Kunkalikar, S., Reddy, R., Gulhane, S., Rajagopalan, P., and Anandalakshmi, R. (2010). Use of agro infectious clones of tomato leaf curl geminiviruses for tomato screening. conference on ‘whitefly and thrips transmitted viruses’ at University of Delhi-South Campus, New Delhi. Di Stilio, V.S., Kumar, R.A., Oddone, A.M., Tolkin, T.R., Salles, P., and McCarty, K. (2010). Virus-induced gene silencing as a tool for comparative functional studies in Thalictrum. PLOS ONE 5, e12064. https://doi. org/10.1371/journal.pone.0012064 Díaz-Camino, C., Annamalai, P., Sanchez, F., Kachroo, A., and Ghabrial, S.A. (2011). An effective virus-based gene silencing method for functional genomics studies in common bean. Plant Methods 7, 16. https://doi. org/10.1186/1746-4811-7-16 Dinesh-Kumar, S.P., Anandalakshmi, R., Marathe, R., Schiff, M., and Liu, Y. (2003). Virus-induced gene silencing. Methods Mol. Biol. 236, 287–294. Ding, L.J., Pu, M.Y., Wei, B., Wang, X.P., Fan, R.C., and Zhang, X.Q. (2016). Transcription factor gene TuGTγ-3 is involved in the stripe rust resistance in Triticum urartu. Yi Chuan 38, 1090–1101. https://doi.org/10.16288/j. yczz.16-133 Ding, X.S., Schneider, W.L., Chaluvadi, S.R., Mian, M.A., and Nelson, R.S. (2006). Characterization of a Brome mosaic virus strain and its use as a vector for gene silencing in monocotyledonous hosts. Mol. Plant Microbe Interact. 19, 1229–1239. https://doi.org/10.1094/MPMI-191229 Dobnik, D., Lazar, A., Stare, T., Gruden, K., Vleeshouwers, V.G., and Žel, J. (2016). Solanum venturii, a suitable model system for virus-induced gene silencing studies in potato reveals StMKK6 as an important player in plant immunity. Plant Methods 12, 29. https://doi. org/10.1186/s13007-016-0129-3 Dolja, V.V., and Koonin, E.V. (2013). The closterovirusderived gene expression and RNA interference vectors as tools for research and plant biotechnology. Front. Microbiol. 4, 83. https://doi.org/10.3389/ fmicb.2013.00083 Dong, Y., Burch-Smith, T.M., Liu, Y.L., Mamillapalli, P., and Dinesh-Kumar, S.P. (2007). A ligation-independent cloning TRV vector for high-throughput virus-induced gene silencing identifies roles for NbMADS4-1 and -2 in floral development. Plant Physiol. 145, 1161–70. https://doi.org/10.1104/pp.107.107391 Du, J., Huang, Z., Wang, B., Sun, H., Chen, C., Ling, H.Q., and Wu, H. (2015). SlbHLH068 interacts with FER to regulate the iron-deficiency response in tomato. Am. Bot. 116, 23–24. https://doi.org/10.1093/aob/mcu058 Du, J., Tian, Z., Liu, J., Vleeshouwers, V.G., Shi, X., and Xie, C. (2013). Functional analysis of potato genes involved in quantitative resistance to Phytophthora infestans. Mol. Biol. Rep. 40, 957–967. https://doi.org/10.1007/ s11033-012-2137-3 Duan, X., Wang, X., Fu, Y., Tang, C., Li, X., Cheng, Y., Feng, H., Huang, L., and Kang, Z. (2013). TaEIL1, a wheat homologue of AtEIN3, acts as a negative regulator in the wheat-stripe rust fungus interaction. Mol. Plant Pathol. 14, 728–739. https://doi.org/10.1111/mpp.12044

Eftekhariyan Ghamsari, M.R., Karimi, F., Mousavi Gargari, S.L., Hosseini Tafreshi, S.A., and Salami, S.A. (2014). Assessing the tobacco-rattle-virus-based vectors system as an efficient gene silencing technique in Datura stramonium (Solanaceae). Virus Genes 49, 512–516. https://doi.org/10.1007/s11262-014-1104-6 Ekengren, S.K., Liu, Y., Schiff, M., Dinesh-Kumar, S.P., and Martin, G.B. (2003). Two MAPK cascades, NPR1, and TGA transcription factors play a role in Pto-mediated disease resistance in tomato. Plant J. 36, 905–917. Eybishtz, A., Peretz, Y., Sade, D., Gorovits, R., and Czosnek, H. (2010). Tomato yellow leaf curl virus infection of a resistant tomato line with a silenced sucrose transporter gene LeHT1 results in inhibition of growth, enhanced virus spread, and necrosis. Planta 231, 537–548. https:// doi.org/10.1007/s00425-009-1072-6 Fagard, M., Boutet, S., Morel, J.B., Bellini, C., and Vaucheret, H. (2000). AGO1, QDE-2, and RDE-1 are related proteins required for post-transcriptional gene silencing in plants, quelling in fungi, and RNA interference in animals. Proc. Natl. Acad. Sci. U.S.A. 97, 11650–11654. https://doi.org/10.1073/pnas.200217597 Faivre-Rampant, O., Gilroy, E.M., Hrubikova, K., Hein, I., Millam, S., Loake, G.J., Birch, P., Taylor, M., and Lacomme, C. (2004). Potato virus X-induced gene silencing in leaves and tubers of potato. Plant Physiol. 134, 1308–1316. https://doi.org/10.1104/ pp.103.037507 Fan, J., Quan, S., Orth, T., Awai, C., Chory, J., and Hu, J. (2005). The Arabidopsis PEX12 gene is required for peroxisome biogenesis and is essential for development. Plant Physiol. 139, 231–239. Feng, H., Liu, W., Zhang, Q., Wang, X., Wang, X., Duan, X., Li, F., Huang, L., and Kang, Z. (2014). TaMDHAR4, a mono dehydro ascorbate reductase gene participates in the interactions between wheat and Puccinia striiformisf. sp. tritici. Plant Physiol. Biochem. 76, 7–16. https://doi. org/10.1016/j.plaphy.2013.12.015 Feng, H., Zhang, Q., Li, H., Wang, X., Wang, X., Duan, X., Wang, B., and Kang, Z. (2013). vsiRNAs derived from the miRNA-generating sites of pri-tae-miR159a based on the BSMV system play positive roles in the wheat response to Puccinia striiformis f. sp. tritici through the regulation of taMyb3 expression. Plant Physiol. Biochem. 68, 90–95. https://doi.org/10.1016/j. plaphy.2013.04.008 Feng, Y.L.,Wang, K.T., Ma, C., Zhao, Y.Y., and Yin, J. (2015). Virus-induced gene silencing-based functional verification of six genes associated with vernalization in wheat. Biochem. Biophys. Res. Commun. 458, 928–933. https://doi.org/10.1016/j.bbrc.2015.02.064 Fernandez-Moreno, J.P., Orzaez, D., and Granell, A. (2013). VIGS: a tool to study fruit development in Solanum lycopersicum. Methods Mol. Biol. 975, 183–196. https:// doi.org/10.1007/978-1-62703-278-0_14 Flores, M.A., Reyes, M.I., Robertson, D.N., and Kjemtrup, S. (2015). Persistent virus-induced gene silencing in asymptomatic accessions of Arabidopsis. Methods Mol. Biol. 1284, 305–322. https://doi.org/10.1007/978-14939-2444-8_15 Fofana, I.B., Sangaré, A., Collier, R., Taylor, C., and Fauquet, C.M. (2004). A geminivirus-induced gene silencing system for gene function validation in cassava. Plant

Virus-Induced Gene Silencing and its Applications |  311

Mol. Biol. 56, 613–624. https://doi.org/10.1007/ s11103-004-0161-y Fu, D.Q., Zhu, B.Z., Zhu, H.L., Jiang, W.B., and Luo, Y.B. (2005). Virus-induced gene silencing in tomato fruit. Plant J. 43, 299–308. Fu, D.Q., Zhu, B.Z., Zhu, H.L., Zhang, H.X., Xie, Y.H., Jiang, W.B., Zhao, X.D., and Luo, K.B. (2006). Enhancement of virus-induced gene silencing in tomato by low temperature and low humidity. Mol. Cells 21, 153–160. Gabriëls, S.H., Takken, F.L., Vossen, J.H., de Jong, C.F., Liu, Q., Turk, S.C., Wachowski, L.K., Peters, J., Witsenboer, H.M., de Wit, P.J., et al. (2006). CDNA-AFLP combined with functional analysis reveals novel genes involved in the hypersensitive response. Mol. Plant Microbe Interact. 19, 567–576. https://doi.org/10.1094/ MPMI-19-0567 Gabriëls, S.H., Vossen, J.H., Ekengren, S.K., van Ooijen, G., Abd-El-Haliem, A.M., van den Berg, G.C., Rainey, D.Y., Martin, G.B., Takken, F.L., de Wit, P.J., et al. (2007). An NB-LRR protein required for HR signalling mediated by both extra- and intracellular resistance proteins. Plant J. 50, 14–28. Galis, I., Schuman, M.C., Gase, K., Hettenhausen, C., Hartl, M., Dinh, S.T., Wu, J., Bonaventure, G., and Baldwin, I.T. (2013). The use of VIGS technology to study plant-herbivore interactions. Methods Mol. Biol. 975, 109–137. https://doi.org/10.1007/978-1-62703-2780_9 Gama, F., Saavedra, T., Dandlen, S., de Varennes, A., Correia, P.J., Pestana, M., and Nolasco, G. (2017). Silencing of the FRO1 gene and its effects on iron partition in Nicotiana benthamiana. Plant Physiol. Biochem. 114, 111–118. Gammelgard, E., Mohan, M., and Valkonen, J.P.T. (2007). Potyvirus-induced gene silencing: the dynamic process of systemic silencing and silencing suppression. J. of General Virology 88, 2337–2346. https://doi. org/10.1099/vir.0.82928-0. Gao, J., and Lan, T. (2016). Functional characterization of the late embryogenesis abundant (LEA) protein gene family from Pinus tabuliformis (Pinaceae) in Escherichia coli. Sci Rep 6, 19467. https://doi.org/10.1038/ srep19467 Gao, W., Long, L., Xu, L., Lindsey, K., Zhang, X., and Zhu, L. (2016). Suppression of the homeobox gene HDTF1 enhances resistance to Verticillium dahliae and Botrytis cinerea in cotton. J Integr Plant Biol 58, 503–513. https://doi.org/10.1111/jipb.12432 Gao, X., and Shan, L. (2013). Functional genomic analysis of cotton genes with Agrobacterium-mediated virusinduced gene silencing. Methods Mol. Biol. 975, 157–165. https://doi.org/10.1007/978-1-62703-2780_12 Gao, X., Britt, R.C., Shan, L., and He, P. (2011a). Agrobacterium-mediated virus-induced gene silencing assay in cotton. J Vis Exp 54, 2938. https://doi. org/10.3791/2938 Gao, X., Wheeler, T., Li, Z., Kenerley, C.M., He, P., and Shan, L. (2011b). Silencing GhNDR1 and GhMKK2 compromises cotton resistance to Verticillium wilt. Plant J. 66, 293–305. https://doi.org/10.1111/j.1365313X.2011.04491.x Gao, Y., Zhang, Y., Zhang, X., Han, C., Yu, J., and Li, D. (2011c). Development and optimization of Tobacco

necrosis virus A induced gene silencing in Nicotiana benthamiana. Prog. Biochem. Biophys. 38, 919–928. https://doi.org/10.3724/SP.J.1206.2011.00129 Gargul, J.M., Mibus, H., and Serek, M. (2015). Manipulation of MKS1 gene expression affects Kalanchoë blossfeldiana and Petunia hybrida phenotypes. Plant Biotechnol. J. 13, 51–61. https://doi.org/10.1111/pbi.12234 George, G.M., Bauer, R., Blennow, A., Kossmann, J., and Lloyd, J.R. (2012). Virus-induced multiple gene silencing to study redundant metabolic pathways in plants: silencing the starch degradation pathway in Nicotiana benthamiana. Biotechnol. J. 7, 884–890. https://doi.org/10.1002/biot.201100469 Giliberto, L., Perrotta, G., Pallara, P., Weller, J.L., Fraser, P.D., Bramley, P.M., Fiore, A., Tavazza, M., and Giuliano, G. (2005). Manipulation of the blue light photoreceptor cryptochrome 2 in tomato affects vegetative development, flowering time, and fruit antioxidant content. Plant Physiol. 137, 199–208. Golenberg, E.M., Sather, D.N., Hancock, L.C., Buckley, K.J., Villafranco, N.M., and Bisaro, D.M. (2009). Development of a gene silencing DNA vector derived from a broad host range geminivirus. Plant Methods 5, 9. https://doi.org/10.1186/1746-4811-5-9 Gordon, C.S., Rajagopalan, N., Risseeuw, E.P., Surpin, M., Ball, F.J., Barber, C.J., Buhrow, L.M., Clark, S.M., Page, J.E., Todd, C.D., et al. (2016). Characterization of Triticum aestivum Abscisic Acid Receptors and a Possible Role for These in Mediating Fusairum Head Blight Susceptibility in Wheat. PLOS ONE 11, e0164996. https://doi.org/10.1371/journal.pone.0164996 Gosselé, V., Faché, I., Meulewaeter, F., Cornelissen, M., and Metzlaff, M. (2002). SVISS - a novel transient gene silencing system for gene function discovery and validation in tobacco plants. Plant J. 32, 859–866. Gould, B., and Kramer, E.M. (2007). Virus-induced gene silencing as a tool for functional analyses in the emerging model plant Aquilegia (columbine, Ranunculaceae). Plant Methods 3, 6. Grønlund, M. (2015). VIGS for dissecting mechanisms involved in the symbiotic interaction of microbes with plants. Methods Mol. Biol. 1287, 255–265. https://doi. org/10.1007/978-1-4939-2453-0_19 Grønlund, M., Constantin, G., Piednoir, E., Kovacev, J., Johansen, I.E., and Lund, O.S. (2008). Virus-induced gene silencing in Medicago truncatula and Lathyrus odorata. Virus Res. 135, 345–349. https://doi. org/10.1016/j.virusres.2008.04.005 Grønlund, M., Olsen, A., Johansen, E.I., and Jakobsen, I. (2010). Protocol: using virus-induced gene silencing to study the arbuscular mycorrhizal symbiosis in Pisum sativum. Plant Methods 6, 28. https://doi. org/10.1186/1746-4811-6-28 Groszyk, J., Kowalczyk, M., Yanushevska, Y., Stochmal, A., Rakoczy-Trojanowska, M., and Orczyk, W. (2017). Identification and VIGS-based characterization of Bx1 ortholog in rye (Secale cereale L.). PLOS ONE 12, e0171506. https://doi.org/10.1371/journal. pone.0171506 Groten, K., Pahari, N.T., Xu, S., Miloradovic van Doorn, M., and Baldwin, I.T. (2015). Virus-induced gene silencing using Tobacco rattle virus as a tool to study the interaction between Nicotiana attenuata and Rhizophagus irregularis.

312  | Kumar et al.

PLOS ONE 10, e0136234. https://doi.org/10.1371/ journal.pone.0136234 Gu, Z., Huang, C., Li, F., and Zhou, X. (2014). A versatile system for functional analysis of genes and microRNAs in cotton. Plant Biotechnol. J. 12, 638–649. https://doi. org/10.1111/pbi.12169 Guo, J., Bai, P., Yang, Q., Liu, F., Wang, X., Huang, L., and Kang, Z. (2013). Wheat zinc finger protein TaLSD1, a negative regulator of programmed cell death, is involved in wheat resistance to stripe rust fungus. Plant Physiol. Biochem. 71, 164–172. https://doi.org/10.1016/j. plaphy.2013.07.009 Guo, Y., Huang, C., Xie, Y., Song, F., and Zhou, X. (2010). A tomato glutaredoxin gene SlGRX1 regulates plant responses to oxidative, drought and salt stresses. Planta 232, 1499–1509. https://doi.org/10.1007/s00425010-1271-1 Hands, P., Vosnakis, N., Betts, D., Irish, V.F., and Drea, S. (2011). Alternate transcripts of a floral developmental regulator have both distinct and redundant functions in opium poppy. Ann. Bot. 107, 1557–1566. https://doi. org/10.1093/aob/mcr045 Hartl, M., Merker, H., Schmidt, D.D., and Baldwin, I.T. (2008). Optimized virus-induced gene silencing in Solanum nigrum reveals the defensive function of leucine aminopeptidase against herbivores and the shortcomings of empty vector controls. New Phytol. 179, 356–365. Hayward, A., Padmanabhan, M., and Dinesh-Kumar, S.P. (2011). Virus-induced gene silencing in Nicotiana benthamiana and other plant species. Methods Mol. Biol. 678, 55–63. https://doi.org/10.1007/978-1-60761682-5_5 He, H., Zhu, S., Jiang, Z., Ji, Y., Wang, F., Zhao, R., and Bie, T. (2016). Comparative mapping of powdery mildew resistance gene Pm21 and functional characterization of resistance-related genes in wheat. Theor. Appl. Genet. 129, 819–829. https://doi.org/10.1007/s00122-0162668-4 He, Q., McLellan, H., Boevink, P.C., Sadanandom, A., Xie, C., Birch, P.R., and Tian, Z. (2015a). U-box E3 ubiquitin ligase PUB17 acts in the nucleus to promote specific immune pathways triggered by Phytophthora infestans. J. Exp. Bot. 66, 3189–3199. https://doi.org/10.1093/jxb/ erv128 He, X., Anderson, J.C., del Pozo, O., Gu, Y.Q., Tang, X., and Martin, G.B. (2004). Silencing of subfamily I of protein phosphatase 2A catalytic subunits results in activation of plant defense responses and localized cell death. Plant J. 38, 563–577. https://doi.org/10.1111/j.1365313X.2004.02073.x He, X., Jin, C., Li, G., You, G., Zhou, X., and Zheng, S.J. (2008). Use of the modified viral satellite DNA vector to silence mineral nutrition-related genes in plants: silencing of the tomato ferric chelate reductase gene, FRO1, as an example. Sci. China, C, Life Sci. 51, 402–409. He, X., Zeng, J., Cao, F., Ahmed, I.M., Zhang, G., Vincze, E., and Wu, F. (2015b). HvEXPB7, a novel β-expansin gene revealed by the root hair transcriptome of Tibetan wild barley, improves root hair growth under drought stress.

J. Exp. Bot. 66, 7405–7419. https://doi.org/10.1093/ jxb/erv436 Hein, I., Barciszewska-Pacak, M., Hrubikova, K., Williamson, S., Dinesen, M., Soenderby, I.E., Sundar, S., Jarmolowski, A., Shirasu, K., and Lacomme, C. (2005). Virus-induced gene silencing-based functional characterization of genes associated with powdery mildew resistance in barley. Plant Physiol. 138, 2155–2164. Heinrich, M., Baldwin, I.T., and Wu, J. (2011). Two mitogenactivated protein kinase kinases, MKK1 and MEK2, are involved in wounding- and specialist lepidopteran herbivore Manduca sexta-induced responses in Nicotiana attenuata. J. Exp. Bot. 62, 4355–4365. https://doi. org/10.1093/jxb/err162 Held, M.A., Penning, B., Brandt, A.S., Kessans, S.A., Yong, W., Scofield, S.R., and Carpita, N.C. (2008). Smallinterfering RNAs from natural antisense transcripts derived from a cellulose synthase gene modulate cell wall biosynthesis in barley. Proc. Natl. Acad. Sci. U.S.A. 105, 20534–20539. https://doi.org/10.1073/ pnas.0809408105 Henderson, I.R., Liu, F., Drea, S., Simpson, G.G., and Dean, C. (2005). An allelic series reveals essential roles for FY in plant development in addition to flowering-time control. Development 132, 3597–3607. Hidalgo, O., Bartholmes, C., and Gleissberg, S. (2012). Virus-induced gene silencing (VIGS) in Cysticapnos vesicaria, a zygomorphic-flowered Papaveraceae (Ranunculales, basal eudicots). Ann. Bot. 109, 911–920. https://doi.org/10.1093/aob/mcs008 Hileman, L.C., Drea, S., Martino, G., Litt, A., and Irish, V.F. (2005). Virus-induced gene silencing is an effective tool for assaying gene function in the basal eudicot species Papaver somniferum (opium poppy). Plant J. 44, 334–341. Hiraga, S., Yamamoto, K., Ito, H., Sasaki, K., Matsui, H., Honma, M., Nagamura, Y., Sasaki, T., and Ohashi, Y. (2000). Diverse expression profiles of 21 rice peroxidase genes. FEBS Lett. 471, 245–250. Hirano, T., Ito, A., Berberich, T., Terauchi, R., and Saitoh, H. (2007). Virus-induced gene silencing of 14-3-3 genes abrogates dark repression of nitrate reductase activity in Nicotiana benthamiana. Mol. Genet. Genomics 278, 125–133. https://doi.org/10.1007/s00438-007-0234-0 Hiriart, J.B., Aro, E.M., and Lehto, K. (2003). Dynamics of the VIGS-mediated chimeric silencing of the Nicotiana benthamiana ChlH gene and of the Tobacco mosaic virus vector. Mol. Plant Microbe Interact. 16, 99–106. https://doi.org/10.1094/MPMI.2003.16.2.99 Holzberg, S., Brosio, P., Gross, C., and Pogue, G.P. (2002). Barley stripe mosaic virus-induced gene silencing in a monocot plant. Plant J. 30, 315–327. Hong, J.K., Jung, H.W., Lee, B.K., Lee, S.C., Lee, Y.K., and Hwang, B.K. (2004). An osmotin-like protein gene, CAOSM1, from pepper: differential expression and insitu localization of its mRNA during pathogen infection and abiotic stress. Physiol. Mol. Plant 64, 301– 310. https://doi.org/10.1016/j.pmpp.2004.10.004 Hong, J.S., Rhee, S.J., Kim, E.J., Kim, T.S., Ryu, K.H., Masuta, C., and Lee, G.P. (2012). Application of a reassortant Cucumber mosaic virus vector for gene silencing in

Virus-Induced Gene Silencing and its Applications |  313

tomato and chili pepper plants. Plant Pathol. J. 28, 81–86. https://doi.org/10.5423/PPJ.NT.11.2011.0220 Hou, H., and Qiu, W. (2003). A novel co-delivery system consisting of a Tomato bushy stunt virus and a defective interfering RNA for studying gene silencing. J. Virol. Methods 111, 37–42. Hsieh, M.H., Lu, H.C., Pan, Z.J., Yeh, H.H., Wang, S.S., Chen, W.H., and Chen, H.H. (2013). Optimizing virus-induced gene silencing efficiency with Cymbidium mosaic virus in Phalaenopsis flower. Plant Sci. 201-202, 25–41. https://doi.org/10.1016/j.plantsci.2012.11.003 Huang, C., Xie, Y., and Zhou, X. (2009). Efficient virusinduced gene silencing in plants using a modified geminivirus DNA1 component. Plant Biotechnol. J. 7, 254–265. https://doi.org/10.1111/j.14677652.2008.00395.x Huang, C.J., Zhang, T., Li, F.F., Zhang, X.Y., and Zhou, X.P. (2011). Development and application of an efficient virus-induced gene silencing system in Nicotiana tabacum using geminivirus alphasatellite. J. Zhejiang Univ. Sci. B 12, 83–92. https://doi.org/10.1631/jzus. B1000157 Huang, Y., Li, M.Y., Wu, P., Xu, Z.S., Que, F., Wang, F., and Xiong, A.S. (2016). Members of WRKY Group III transcription factors are important in TYLCV defense signaling pathway in tomato (Solanum lycopersicum). BMC Genomics 17, 788. https://doi.org/10.1186/ s12864-016-3123-2 Hwang, I.S., and Hwang, B.K. (2010). Role of the pepper cytochrome P450 gene CaCYP450A in defense responses against microbial pathogens. Planta 232, 1409–1421. https://doi.org/10.1007/s00425-0101266-y Hwang, J., Lee, S., Lee, J.H., Kang, W.H., Kang, J.H., Kang, M.Y., Oh, C.S., and Kang, B.C. (2015). Plant Translation Elongation Factor 1Bβ Facilitates Potato Virus X (PVX) Infection and Interacts with PVX Triple Gene Block Protein 1. PLOS ONE 10, e0128014. https://doi. org/10.1371/journal.pone.0128014 Hwang, J., Li, J., Liu, W.Y., An, S.J., Cho, H., Her, N.H., Yeam, I., Kim, D., and Kang, B.C. (2009). Double mutations in eIF4E and eIFiso4E confer recessive resistance to Chilli veinal mottle virus in pepper. Mol. Cells 27, 329–336. https://doi.org/10.1007/s10059-009-0042-y Ido, Y., Nakahara, K.S., and Uyeda, I. (2012). White clover mosaic virus-induced gene silencing in pea. J. Gen. Plant Pathol. 78, 127. https://doi.org/10.1007/s10327-0120360-3 Igarashi, A., Yamagata, K., Sugai, T., Takahashi, Y., Sugawara, E., Tamura, A., Yaegashi, H., Yamagishi, N., Takahashi, T., Isogai, M., et al. (2009). Apple latent spherical virus vectors for reliable and effective virus-induced gene silencing among a broad range of plants including tobacco, tomato, Arabidopsis thaliana, cucurbits, and legumes. Virology 386, 407–416. https://doi. org/10.1016/j.virol.2009.01.039 Ishiga, Y., Ishiga, T., Uppalapati, S.R., and Mysore, K.S. (2013). Jasmonate ZIM-domain ( JAZ) protein regulates host and nonhost pathogen-induced cell death in tomato and Nicotiana benthamiana. PLOS ONE 8, e75728. https://doi.org/10.1371/journal.pone.0075728

Ito, H., Hiraga, S., Tsugawa, H., Matsui, H., Honma, M., Otsuki, Y., Murakami, T., and Ohashi, Y. (2000). Xylemspecific expression of wound-inducible rice peroxidase genes in transgenic plants. Plant Sci. 155, 85–100. Ito, H., Ochiai, M., Kato, H., Shiratake, K., Takemoto,D., Otagaki, S., and Matsumoto, S. (2012). Rose Phytoene Desaturase gene silencing by Apple latent spherical virus vector. Hortscience 47, 1278–1282. Ito, M., Ohnishi, K., Hikichi, Y., and Kiba, A. (2015). Molecular chaperons and co-chaperons, Hsp90, RAR1, and SGT1 negatively regulate bacterial wilt disease caused by Ralstonia solanacearum in Nicotiana benthamiana. Plant Signal Behav 10, e970410. https:// doi.org/10.4161/15592316.2014.970410 Jaakola, L., Poole, M., Jones, M.O., Kämäräinen-Karppinen, T., Koskimäki, J.J., Hohtola, A., Häggman, H., Fraser, P.D., Manning, K., King, G.J., et al. (2010). A SQUAMOSA MADS box gene involved in the regulation of anthocyanin accumulation in bilberry fruits. Plant Physiol. 153, 1619–1629. https://doi. org/10.1104/pp.110.158279 Jablonska, B., Ammiraju, J.S., Bhattarai, K.K., Mantelin, S., Martinez de Ilarduya, O., Roberts, P.A., and Kaloshian, I. (2007). The Mi-9 gene from Solanum arcanum conferring heat-stable resistance to root-knot nematodes is a homolog of Mi-1. Plant Physiol. 143, 1044–1054. Jeon, Y., Hwang, A.R., Hwang, I., and Pai, H.S. (2010). Silencing of NbCEP1 encoding a chloroplast envelope protein containing 15 leucine-rich-repeats disrupts chloroplast biogenesis in Nicotiana benthamiana. Mol. Cells 29, 175–183. https://doi.org/10.1007/s10059010-0011-5 Jeon, Y., Jung, H.J., Kang, H., Park, Y.I., Lee, S.H., and Pai, H.S. (2012). S1 domain-containing STF modulates plastid transcription and chloroplast biogenesis in Nicotiana benthamiana. New Phytol. 193, 349–363. https://doi.org/10.1111/j.1469-8137.2011.03941.x Jeyabharathy, C., Shakila, H., and Usha, R. (2015). Development of a VIGS vector based on the β-satellite DNA associated with bhendi yellow vein mosaic virus. Virus Res. 195, 73–78. https://doi.org/10.1016/j. virusres.2014.08.009 Ji, K., Kai, W., Zhao, B., Sun, Y., Yuan, B., Dai, S., Li, Q., Chen, P., Wang, Y., Pei, Y., et al. (2014). SlNCED1 and SlCYP707A2: key genes involved in ABA metabolism during tomato fruit ripening. J. Exp. Bot. 65, 5243–5255. https://doi.org/10.1093/jxb/eru288 Jia, H., Hao, L., Guo, X., Liu, S., Yan, Y., and Guo, X. (2016). A Raf-like MAPKKK gene, GhRaf19, negatively regulates tolerance to drought and salt and positively regulates resistance to cold stress by modulating reactive oxygen species in cotton. Plant Sci. 252, 267–281. Jiang, C.Z., Chen, J.C., and Reid, M. (2011). Virus-induced gene silencing in ornamental plants. Methods Mol. Biol. 744, 81–96. https://doi.org/10.1007/978-1-61779123-9_6 Jiang, C.Z., Lu, F., Imsabai, W., Meir, S., and Reid, M.S. (2008). Silencing polygalacturonase expression inhibits tomato petiole abscission. J. Exp. Bot. 59, 973–979. https://doi.org/10.1093/jxb/ern023 Jiang, X., Zhang, C., Lü, P., Jiang, G., Liu, X., Dai, F., and Gao, J. (2014a). RhNAC3, a stress-associated NAC

314  | Kumar et al.

transcription factor, has a role in dehydration tolerance through regulating osmotic stress-related genes in rose petals. Plant Biotechnol. J. 12, 38–48. https://doi. org/10.1111/pbi.12114 Jiang, Y., Ye, S., Wang, L., Duan, Y., Lu, W., Liu, H., Fan, D., Zhang, F., and Luo, K. (2014b). Heterologous gene silencing induced by Tobacco rattle virus (TRV) is efficient for pursuing functional genomics studies in woody plants. Plant Cell Tissue Organ Cult 116, 163–174. https://doi.org/10.1007/s11240-013-0393-0 Jiang, Z., Wang, H., Zhang, G., Zhao, R., Bie, T., Zhang, R., Gao, D., Xing, L., and Cao, A. (2017). Characterization of a small GTP-binding protein gene TaRab18 from wheat involved in the stripe rust resistance. Plant Physiol. Biochem. 113, 40–50. Jin, H., Axtell, M.J., Dahlbeck, D., Ekwenna, O., Zhang, S., Staskawicz, B., and Baker, B. (2002). NPK1, an MEKK1like mitogen-activated protein kinase kinase kinase, regulates innate immunity and development in plants. Dev. Cell 3, 291–297. Jin, H., Li, S., and Villegas, A. (2006). Down-regulation of the 26S proteasome subunit RPN9 inhibits viral systemic transport and alters plant vascular development. Plant Physiol. 142, 651–661. Jin, H., Liu, Y., Yang, K.Y., Kim, C.Y., Baker, B., and Zhang, S. (2003). Function of a mitogen-activated protein kinase pathway in N gene-mediated resistance in tobacco. Plant J. 33, 719–731. Jin, J.H., Zhang, H.X., Tan, J.Y., Yan, M.J., Li, D.W., Khan, A., and Gong, Z.H. (2015). A New Ethylene-Responsive Factor CaPTI1 Gene of Pepper (Capsicum annuum L.) Involved in the Regulation of Defense Response to Phytophthora capsici. Front. Plant Sci. 6, 1217. https:// doi.org/10.3389/fpls.2015.01217 Jones, L., Keining, T., Eamens, A., and Vaistij, F.E. (2006). Virus-induced gene silencing of argonaute genes in Nicotiana benthamiana demonstrates that extensive systemic silencing requires Argonaute1-like and Argonaute4-like genes. Plant Physiol. 141, 598–606. Jones, L., Ratcliff, F., and Baulcombe, D.C. (2001). RNAdirected transcriptional gene silencing in plants can be inherited independently of the RNA trigger and requires Met1 for maintenance. Curr. Biol. 11, 747–757. Jordan, C.V., Shen, W., Hanley-Bowdoin, L.K., and Robertson, D.N. (2007). Geminivirus-induced gene silencing of the tobacco retinoblastoma-related gene results in cell death and altered development. Plant Mol. Biol. 65, 163–175. https://doi.org/10.1007/s11103007-9206-3 Jupin, I. (2013). A protocol for VIGS in Arabidopsis thaliana using a one-step TYMV-derived vector. Methods Mol. Biol. 975, 197–210. https://doi.org/10.1007/978-162703-278-0_15 Juvale, P.S., Hewezi, T., Zhang, C., Kandoth, P.K., Mitchum, M.G., Hill, J.H., Whitham, S.A., and Baum, T.J. (2012). Temporal and spatial Bean pod mottle virusinduced gene silencing in soybean. Mol. Plant Pathol. 13, 1140–1148. https://doi.org/10.1111/j.13643703.2012.00808.x Kanazawa, A.,Inaba, J.I.,Shimura, H.,Otagaki, S.,Tsukahara, S.,Matsuzawa, A.,Kim, B.M.,Goto, K.,and Masuta, C. (2011). Virus-mediated efficient induction of epigenetic modifications of endogenous genes with phenotypic

changes in plants. Plant J. 65, 156–168. https://doi. org/10.1111/j.1365-313X.2010.04401.x Kandoth, P.K., Heinz, R., Yeckel, G., Gross, N.W., Juvale, P.S., Hill, J., Whitham, S.A., Baum, T.J., and Mitchum, M.G. (2013). A virus-induced gene silencing method to study soybean cyst nematode parasitism in Glycine max. BMC Res Notes 6, 255. https://doi.org/10.1186/17560500-6-255 Kandoth, P.K., Ranf, S., Pancholi, S.S., Jayanty, S., Walla, M.D., Miller, W., Howe, G.A., Lincoln, D.E., and Stratmann, J.W. (2007). Tomato MAPKs LeMPK1, LeMPK2, and LeMPK3 function in the systeminmediated defense response against herbivorous insects. Proc. Natl. Acad. Sci. U.S.A. 104, 12205–12210. Kang, G., Li, G., Ma, H., Wang, C., and Guo, T. (2013). Proteomic analysis on the leaves of TaBTF3 gene virusinduced silenced wheat plants may reveal its regulatory mechanism. J Proteomics 83, 130–143. https://doi. org/10.1016/j.jprot.2013.03.020 Kang, J.H., Wang, L., Giri, A., and Baldwin, I.T. (2006). Silencing threonine deaminase and JAR4 in Nicotiana attenuata impairs jasmonic acid-isoleucine-mediated defenses against Manduca sexta. Plant Cell 18, 3303– 3320. Kang, Y.W., Kim, R.N., Cho, H.S., Kim, W.T., Choi, D., and Pai, H.S. (2008). Silencing of a BYPASS1 homolog results in root-independent pleiotrophic developmental defects in Nicotiana benthamiana. Plant Mol. Biol. 68, 423–437. https://doi.org/10.1007/s11103-008-9384-7 Kang, Y.W., Lee, J.Y., Jeon, Y., Cheong, G.W., Kim, M., and Pai, H.S. (2010). In vivo effects of NbSiR silencing on chloroplast development in Nicotiana benthamiana. Plant Mol. Biol. 72, 569–583. https://doi.org/10.1007/ s11103-009-9593-8 Kanneganti, T.D., Bai, X., Tsai, C.W., Win, J., Meulia, T., Goodin, M., Kamoun, S., and Hogenhout, S.A. (2007). A functional genetic assay for nuclear trafficking in plants. Plant J. 50, 149–158. Kant, R., and Dasgupta, I. (2017). Phenotyping of VIGSmediated gene silencing in rice using a vector derived from a DNA virus. Plant Cell Rep. 36, 1159–1170. https://doi.org/10.1007/s00299-017-2156-6 Kant, R., Sharma, S., and Dasgupta, I. (2015). Virusinduced gene silencing (VIGS) for functional genomics in rice using Rice tungro bacilliform virus (RTBV) as a vector. Methods Mol. Biol. 1287, 201–217. https://doi. org/10.1007/978-1-4939-2453-0_15 Kanzaki, H., Saitoh, H., Ito, A., Fujisawa, S., Kamoun, S., Katou, S., Yoshioka, H., and Terauchi, R. (2003). Cytosolic HSP90 and HSP70 are essential components of INF1-mediated hypersensitive response and nonhost resistance to Pseudomonas cichorii in Nicotiana benthamiana. Mol. Plant Pathol. 4, 383–391. https:// doi.org/10.1046/j.1364-3703.2003.00186.x Kasschau, K.D., and Carrington, J.C. (1998). A counter defensive strategy of plant viruses: suppression of post transcriptional gene silencing. Cell 95, 461–470. Kawai, T., Gonoi, A., Nitta, M., Kaido, M., Yamagishi, N., Yoshikawa, N., and Tao, R. (2014). Virus-induced gene silencing in apricot (Prunus armeniaca L.) and Japanese apricot (P. mume Siebold and Zucc.) with the Apple latent spherical virus vector system. J. Jpn. Soc. Hortic. Sci. 83, 23–31. https://doi.org/10.2503/jjshs1.CH-091

Virus-Induced Gene Silencing and its Applications |  315

Kim, B.M., Inaba, J., and Masuta, C. (2011). Virus induced gene silencing in Antirrhinum majus using the Cucumber mosaic virus vector: Functional analysis of the AINTEGUMENTA (Am-ANT) gene of A. majus. Horti. Enviro. Biotech. 52, 176–182. Kim, D.S., Choi, H.W., and Hwang, B.K. (2014). Pepper mildew resistance locus O interacts with pepper calmodulin and suppresses Xanthomonas AvrBsTtriggered cell death and defense responses. Planta 240, 827–839. https://doi.org/10.1007/s00425-014-2134-y Kim, J., Park, M., Jeong, E.S., Lee, J.M., and Choi, D. (2017). Harnessing anthocyanin-rich fruit: a visible reporter for tracing virus-induced gene silencing in pepper fruit. Plant Methods 13, 3. https://doi.org/10.1186/s13007016-0151-5 Kim, K.H., Lim, S., Kang, Y.J., Yoon, M.Y., Nam, M., Jun, T.H., Seo, M.J., Baek, S.B., Lee, J.H., Moon, J.K., et al. (2016). Optimization of a virus-induced gene silencing system with Soybean yellow common mosaic virus for gene function studies in soybeans. Plant Pathol. J. 32, 112– 122. https://doi.org/10.5423/PPJ.OA.04.2015.0063 Kim, K.J., Lim, J.H., Lee, S., Kim, Y.J., Choi, S.B., Lee, M.K., Choi, D., and Paek, K.H. (2007). Functional study of Capsicum annuum fatty acid desaturase 1 cDNA clone induced by Tobacco mosaic virus via microarray and virus-induced gene silencing. Biochem. Biophys. Res. Commun. 362, 554–561. Kim, K.Y., Kwon, H.K., Kwon, S.Y., Lee, H.S., Hur, Y., Bang, J.W., Choi, K.S., and Kwak, S.S. (2000). Differential expression of four sweet potato peroxidase genes in response to abscisic acid and ethephon. Phytochemistry 54, 19–22. Kim, M., Lim, J.H., Ahn, C.S., Park, K., Kim, G.T., Kim, W.T., and Pai, H.S. (2006a). Mitochondria-associated hexokinases play a role in the control of programmed cell death in Nicotiana benthamiana. Plant Cell 18, 2341–2355. Kim, Y.C., Kim, S.Y., Paek, K.H., Choi, D., and Park, J.M. (2006b). Suppression of CaCYP1, a novel cytochrome P450 gene, compromises the basal pathogen defense response of pepper plants. Biochem. Biophys. Res. Commun. 345, 638–645. Kim, Y.K., Lee, J.Y., Cho, H.S., Lee, S.S., Ha, H.J., Kim, S., Choi, D., and Pai, H.S. (2005). Inactivation of organellar glutamyl- and seryl-tRNA synthetases leads to developmental arrest of chloroplasts and mitochondria in higher plants. J. Biol. Chem. 280, 37098–37106. Kirigia, D., Runo, S., and Alakonya, A. (2014). A virusinduced gene silencing (VIGS) system for functional genomics in the parasitic plant Striga hermonthica. Plant Methods 10, 16. https://doi.org/10.1186/1746-481110-16 Kjemtrup, S., Sampson, K.S., Peele, C.G., Nguyen, L.V., Conkling, M.A., Thompson, W.F., and Robertson, D. (1998). Gene silencing from plant DNA carried by a Geminivirus. Plant J. 14, 91–100. Kramer, E.M., Holappa, L., Gould, B., Jaramillo, M.A., Setnikov, D., and Santiago, P.M. (2007). Elaboration of B gene function to include the identity of novel floral organs in the lower eudicot Aquilegia. Plant Cell 19, 750–766. Krenz, B., Wege, C., and Jeske, H. (2010). Cell-free construction of disarmed Abutilon mosaic virus-based

gene silencing vectors. J. Virol. Methods 169, 129–137. https://doi.org/10.1016/j.jviromet.2010.07.010 Ku, H.M., Hu, C.C., Chang, H.J., Lin, Y.T., Jan, F.J., and Chen, C.T. (2011). Analysis by virus induced gene silencing of the expression of two proline biosynthetic pathway genes in Nicotiana benthamiana under stress conditions. Plant Physiol. Biochem. 49, 1147–1154. https://doi.org/10.1016/j.plaphy.2011.07.003 Kumagai, M.H., Donson, J., della-Cioppa, G., Harvey, D., Hanley, K., and Grill, L.K. (1995). Cytoplasmic inhibition of carotenoid biosynthesis with virus-derived RNA. Proc. Natl. Acad. Sci. U.S.A. 92, 1679–1683. Kurth, E.G., Peremyslov, V.V., Prokhnevsky, A.I., Kasschau, K.D., Miller, M., Carrington, J.C., and Dolja, V.V. (2012). Virus-derived gene expression and RNA interference vector for grapevine. J. Virol. 86, 6002–6009. https:// doi.org/10.1128/JVI.00436-12 Kuzuoglu-Ozturk, D., Cebeci Yalcinkaya, O., Akpinar, B.A., Mitou, G., Korkmaz, G., Gozuacik, D., and Budak, H. (2012). Autophagy-related gene, TdAtg8, in wild emmer wheat plays a role in drought and osmotic stress response. Planta 236, 1081–1092. https://doi. org/10.1007/s00425-012-1657-3 Lacomme, C. (2015). Strategies for altering plant traits using virus-induced gene silencing technologies. Methods Mol. Biol. 1287, 25–41. https://doi.org/10.1007/9781-4939-2453-0_2 Lacomme, C., and Chapman, S. (2008). Use of potato virus X (PVX)-based vectors for gene expression and virus-induced gene silencing (VIGS). Curr Protoc Microbiol Chapter, Unit 16I.1. https://doi. org/10.1002/9780471729259.mc16i01s8 Lacomme, C., Hrubikova, K., and Hein, I. (2003). Enhancement of virus-induced gene silencing through viral-based production of inverted-repeats. Plant J. 34, 543–553. Lakatos, L., Szittya, G., Silhavy, D., and Burgyan, J. (2004). Molecular mechanism of RNA silencing suppression mediated by p19 protein of tombus viruses. EMBO J. 23, 876–884. Lee, D.H., Choi, H.W., and Hwang, B.K. (2011). The pepper E3 ubiquitin ligase RING1 gene, CaRING1, is required for cell death and the salicylic acid-dependent defense response. Plant Physiol. 156, 2011–2025. https://doi. org/10.1104/pp.111.177568 Lee, J., Cao, D.V., Kim, J., Pamplona, R.S., Ahn, J., Cho, S.K., Yang, S.W., and Boo, K.H. (2017). Development of a virus-induced gene silencing (VIGS) system for Spinacia oleracea L. In Vitro Cell. Dev. Biol. Plant. 53, 97–103. https://doi.org/10.1007/s11627-017-9806-9. Lee, S.C., and Hwang, B.K. (2009). Functional roles of the pepper antimicrobial protein gene, CaAMP1, in abscisic acid signaling, and salt and drought tolerance in Arabidopsis. Planta 229, 383–391. https://doi. org/10.1007/s00425-008-0837-7 Lee, S.C., Choi, D.S., Hwang, I.S., and Hwang, B.K. (2010). The pepper oxidoreductase CaOXR1 interacts with the transcription factor CaRAV1 and is required for salt and osmotic stress tolerance. Plant Mol. Biol. 73, 409–424. https://doi.org/10.1007/s11103-010-9629-0 Lee, W.S., Rudd, J.J., and Kanyuka, K. (2015). Virus induced gene silencing (VIGS) for functional analysis of wheat genes involved in Zymoseptoria tritici susceptibility and

316  | Kumar et al.

resistance. Fungal Genet. Biol. 79, 84–88. https://doi. org/10.1016/j.fgb.2015.04.006 Leister, R.T., Dahlbeck, D., Day, B., Li, Y., Chesnokova, O., and Staskawicz, B.J. (2005). Molecular genetic evidence for the role of SGT1 in the intramolecular complementation of Bs2 protein activity in Nicotiana benthamiana. Plant Cell 17, 1268–1278. Li, A., Zhang, R., Pan, L., Tang, L., Zhao, G., Zhu, M., Chu, J., Sun, X., Wei, B., Zhang, X., et al. (2011a). Transcriptome analysis of H2O2-treated wheat seedlings reveals a H2O2-responsive fatty acid desaturase gene participating in powdery mildew resistance. PLOS ONE 6, e28810. https://doi.org/10.1371/journal. pone.0028810 Li, C., Yan, J.M., Li, Y.Z., Zhang, Z.C., Wang, Q.L., and Liang, Y. (2013). Silencing the SpMPK1, SpMPK2, and SpMPK3 genes in tomato reduces abscisic acid-mediated drought tolerance. Int. J. Mol. Sci.14, 21983–21996. https://doi.org/10.3390/ijms141121983 Li, C., Zhang, Z.C., Ghebremariam, K.M., Wang, L.H., Wu, L., and Liang, Y. (2014a). A novel method for the evaluation of virus-induced gene silencing efficiency. Genet. Mol. Res. 13, 9443–9452. https://doi. org/10.4238/2014.November.11.9 Li, D., Behjatnia, S.A., Dry, I.B., Walker, A.R., Randles, J.W., and Rezaian, M.A. (2008). Tomato leaf curl virus satellite DNA as a gene silencing vector activated by helper virus infection. Virus Res. 136, 30–34. https:// doi.org/10.1016/j.virusres.2008.04.011 Li, D., Zhang, H., Song, Q., Wang, L., Liu, S., Hong, Y., Huang, L., and Song, F. (2015a). Tomato Sl3-MMP, a member of the Matrix metalloproteinase family, is required for disease resistance to Botrytis cinerea and Pseudomonas syringae pv. tomato DC3000. BMC Plant Biol. 15, 143. https://doi.org/10.1186/s12870-0150536-z Li, F., Li, M., Wang, P., Cox, K.L., Duan, L., Dever, J.K., Shan, L., Li, Z., and He, P. (2017b). Regulation of cotton (Gossypium hirsutum) drought responses by mitogenactivated protein (MAP) kinase cascade-mediated phosphorylation of GhWRKY59. New Phytol. 215, 1462–1475. https://doi.org/10.1111/nph.14680 Li, F., Zhao, N., Li, Z., Xu, X., Wang, Y., Yang, X., Liu, S.S., Wang, A., and Zhou, X. (2017a). A calmodulinlike protein suppresses RNA silencing and promotes geminivirus infection by degrading SGS3 via the autophagy pathway in Nicotiana benthamiana. PLOS Pathog. 13, e1006213. https://doi.org/10.1371/ journal.ppat.1006213 Li, H.W., Lucy, A.P., Guo, H.S., Li, W.X., Ji, L.H., Wong, S.M., and Ding, S.W. (1999). Strong host resistance targeted against a viral suppressor of the plant gene silencing defense mechanism. EMBO J., 18, 2683–2691. Li, J., Li, F., Qian, M., Han, M., Liu, H., Zhang, D., Ma, J., and Zhao, C. (2017c). Characteristics and regulatory pathway of the PrupeSEP1 SEPALLATA gene during ripening and softening in peach fruits. Plant Sci. 257, 63–73. Li, K., Xing, C., Yao, Z., and Huang, X. (2017d). PbrMYB21, a novel MYB protein of Pyrus betulaefolia, functions in drought tolerance and modulates polyamine levels by regulating arginine decarboxylase gene. Plant

Biotechnol. J. 15, 1186–1203. https://doi.org/10.1111/ pbi.12708 Li, L., Zhu, B., Fu, D., and Luo, Y. (2011). RIN transcription factor plays an important role in ethylene biosynthesis of tomato fruit ripening. J. Sci. Food Agric. 91, 2308–2314. https://doi.org/10.1002/jsfa.4475 Li, Q., Chen, P., Dai, S., Sun, Y., Yuan, B., Kai, W., Pei, Y., He, S., Liang, B., Zhang, Y., et al. (2015b). PacCYP707A2 negatively regulates cherry fruit ripening while PacCYP707A1 mediates drought tolerance. J. Exp. Bot. 66, 3765–3774. https://doi.org/10.1093/jxb/erv169 Li, Q., Xie, Q.G., Smith-Becker, J., Navarre, D.A., and Kaloshian, I. (2006a). Mi-1-Mediated aphid resistance involves salicylic acid and mitogen-activated protein kinase signaling cascades. Mol. Plant Microbe Interact. 19, 655–664. https://doi.org/10.1094/MPMI-19-0655 Li, R., Reed, D.W., Liu, E., Nowak, J., Pelcher, L.E., Page, J.E., and Covello, P.S. (2006b). Functional genomic analysis of alkaloid biosynthesis in Hyoscyamus niger reveals a cytochrome P450 involved in littorine rearrangement. Chem. Biol. 13, 513–520. Li, S., Li, K., Ju, Z., Cao, D., Fu, D., Zhu, H., Zhu, B., and Luo, Y. (2016). Genome-wide analysis of tomato NF-Y factors and their role in fruit ripening. BMC Genomics 17, 36. https://doi.org/10.1186/s12864-015-2334-2 Li, T., Ma, X., Li, N., Zhou, L., Liu, Z., Han, H., Gui, Y., Bao, Y., Chen, J., and Dai, X. (2017e). Genome-wide association study discovered candidate genes of Verticillium wilt resistance in upland cotton (Gossypium hirsutum L.). Plant Biotechnol. J. 15, 1520–1532. https://doi.org/10.1111/pbi.12734 Li, W., Xu, Y.P., Zhang, Z.X., Cao, W.Y., Li, F., Zhou, X., Chen, G.Y., and Cai, X.Z. (2012). Identification of genes required for nonhost resistance to Xanthomonas oryzae pv. oryzae reveals novel signaling components. PLOS ONE 7, e42796. https://doi.org/10.1371/journal. pone.0042796 Li, W.X., and Ding, S.W. (2001). Viral suppressors of RNA silencing. Curr. Opin. Biotechnol. 12, 150–154. https:// doi.org/10.1016/S0958-1669(00)00190-7 Li, X., Huang, L., Zhang, Y., Ouyang, Z., Hong, Y., Zhang, H., Li, D., and Song, F. (2014b). Tomato SR/CAMTA transcription factors SlSR1 and SlSR3L negatively regulate disease resistance response and SlSR1L positively modulates drought stress tolerance. BMC Plant Biol. 14, 286. https://doi.org/10.1186/s12870014-0286-3 Li, X., Zhang, Y., Huang, L., Ouyang, Z., Hong, Y., Zhang, H., Li, D., and Song, F. (2014c). Tomato SlMKK2 and SlMKK4 contribute to disease resistance to Botrytis cinerea. BMC Plant Biol. 14, 166. https://doi. org/10.1186/1471-2229-14-166 Li, Y., Qin, L., Zhao, J., Muhammad, T., Cao, H., Li, H., Zhang, Y., and Liang, Y. (2017f). SlMAPK3 enhances tolerance to Tomato yellow leaf curl virus (TYLCV) by regulating salicylic acid and jasmonic acid signaling in tomato (Solanum lycopersicum). PLOS ONE 12, e0172466. https://doi.org/10.1371/journal.pone.0172466 Liang, J., Deng, G., Long, H., Pan, Z., Wang, C., Cai, P., Xu, D., Nima, Z., and Yu, M. (2012). Virus-induced silencing of genes encoding LEA protein in Tibetan hulless barley (Hordeum vulgare ssp. vulgare) and their relationship to

Virus-Induced Gene Silencing and its Applications |  317

drought tolerance. Mol. Breed. 30, 441–451. https:// doi.org/10.1007/s11032-011-9633-3. Lim, C.J., Kim, W.B., Lee, B.S., Lee, H.Y., Kwon, T.H., Park, J.M., and Kwon, S.Y. (2010). Silencing of SlFTR-c, the catalytic subunit of ferredoxin:thioredoxin reductase, induces pathogenesis-related genes and pathogen resistance in tomato plants. Biochem. Biophys. Res. Commun. 399, 750–754. https://doi.org/10.1016/j. bbrc.2010.08.016 Lim, C.W., and Lee, S.C. (2014). Functional roles of the pepper MLO protein gene, CaMLO2, in abscisic acid signaling and drought sensitivity. Plant Mol. Biol. 85, 1–10. https://doi.org/10.1007/s11103-013-0155-8 Lim, C.W., Lim, S., Baek, W., and Lee, S.C. (2015). The pepper late embryogenesis abundant protein CaLEA1 acts in regulating abscisic acid signaling, drought and salt stress response. Physiol Plant 154, 526–542. https:// doi.org/10.1111/ppl.12298 Lin, Z., Yin, K., Wang, X., Liu, M., Chen, Z., Gu, H., and Qu, L.J. (2007). Virus induced gene silencing of AtCDC5 results in accelerated cell death in Arabidopsis leaves. Plant Physiol. Biochem. 45, 87–94. Liou, M.R., Huang, Y.W., Hu, C.C., Lin, N.S., and Hsu, Y.H. (2014). A dual gene-silencing vector system for monocot and dicot plants. Plant Biotechnol. J. 12, 330–343. https://doi.org/10.1111/pbi.12140 Liscombe, D.K., and O’Connor, S.E. (2011). A virusinduced gene silencing approach to understanding alkaloid metabolism in Catharanthus roseus. Phytochemistry 72, 1969–1977. https://doi. org/10.1016/j.phytochem.2011.07.001 Lister, R.M., and Murant, A.F. (1967). Seed transmission of nematode-borne viruses. Ann. Appl. Biol. 59, 49. Liu, B., Watanabe, S., Uchiyama, T., Kong, F., Kanazawa, A., Xia, Z., Nagamatsu, A., Arai, M., Yamada, T., Kitamura, K., et al. (2010). The soybean stem growth habit gene Dt1 is an ortholog of Arabidopsis TERMINAL FLOWER1. Plant Physiol. 153, 198–210. https://doi. org/10.1104/pp.109.150607 Liu, D., Liu, X., Meng, Y., Sun, C., Tang, H., Jiang, Y., Khan, M.A., Xue, J., Ma, N., and Gao, J. (2013). An organspecific role for ethylene in rose petal expansion during dehydration and rehydration. J. Exp.Bot. 64, 2333–2344. https://doi.org/10.1093/jxb/ert092 Liu, E., and Page, J.E. (2008). Optimized cDNA libraries for virus-induced gene silencing (VIGS) using Tobacco rattle virus. Plant Methods 4, 5. https://doi. org/10.1186/1746-4811-4-5 Liu, G., Wu, Y., Xu, M., Gao, T., Wang, P., Wang, L., Guo, T., and Kang, G. (2016a). Virus-Induced gene silencing identifies an important role of the TaRSR1 transcription factor in starch synthesis in bread Wheat. Int. J. Mol. Sci. 17, E1557. https://doi.org/10.3390/ijms17101557 Liu, M., Pirrello, J., Chervin, C., Roustan, J.P., and Bouzayen, M. (2015). Ethylene control of fruit ripening: revisiting the complex network of transcriptional regulation. Plant Physiol. 169, 2380–2390. https://doi.org/10.1104/ pp.15.01361 Liu, N., Xie, K., Jia, Q., Zhao, J., Chen, T., Li, H., Wei, X., Diao, X., Hong, Y., and Liu, Y. (2016b). Foxtail Mosaic Virus-induced gene silencing in monocot plants. Plant Physiol. 171, 1801–1807. https://doi.org/10.1104/ pp.16.00010

Liu, Y. (2012). Roles of mitogen-activated protein kinase cascades in ABA signaling. Plant Cell Rep. 31, 1–12. https://doi.org/10.1007/s00299-011-1130-y Liu, Y., Jin, H., Yang, K.Y., Kim, C.Y., Baker, B., and Zhang, S. (2003). Interaction between two mitogen-activated protein kinases during tobacco defense signaling. Plant J. 34, 149–160. Liu, Y., Nakayama, N., Schiff, M., and Dinesh-Kumar, S.P. (2004a). Involvement of MEK1, MAPKK, NTF6, MAPK, WRKY/MYB transcription factors, CO11 and CTR1 in N-mediated resistance to Tobacco mosaic virus. Plant J. 38, 800–9. https://doi.org/10.1111/j.1365313X.2004.02085.x Liu, Y., Nakayama, N., Schiff, M., Litt, A., Irish, V.F., and Dinesh-Kumar, S.P. (2004b). Virus induced gene silencing of a DEFICIENS ortholog in Nicotiana benthamiana. Plant Mol. Biol. 54, 701–711. https://doi. org/10.1023/B:PLAN.0000040899.53378.83 Liu, Y., Schiff, M., and Dinesh-Kumar, S.P. (2002a). Virusinduced gene silencing in tomato. Plant J. 31, 777–786. Liu, Y., Schiff, M., Czymmek, K., Tallóczy, Z., Levine, B., and Dinesh-Kumar, S.P. (2005). Autophagy regulates programmed cell death during the plant innate immune response. Cell 121, 567–577. Liu, Y., Schiff, M., Marathe, R., and Dinesh-Kumar, S.P. (2002b). Tobacco Rar1, EDS1 and NPR1/NIM1 like genes are required for N-mediated resistance to Tobacco mosaic virus. Plant J. 30, 415–429. Liu, Z., Shi, L., Yang, S., Lin, Y., Weng, Y., Li, X., Hussain, A., Noman, A., and He, S. (2017). Functional and promoter analysis of ChiIV3, a chitinase of pepper plant, in response to Phytophthora capsici infection. Int. J. Mol. Sci. 18, E1661. Lou, Y., and Baldwin, I.T. (2006). Silencing of a germin-like gene in Nicotiana attenuata improves performance of native herbivores. Plant Physiol. 140, 1126–1136. Louie, R. (1995). Vascular puncture of maize kernels for the mechanical transmission of maize white line mosaic virus and other viruses of maize. Phytopathology 85, 139–143. Lu, H.C., Chen, H.H., Tsai, W.C., Chen, W.H., Su, H.J., Chang, D.C., and Yeh, H.H. (2007). Strategies for functional validation of genes involved in reproductive stages of orchids. Plant Physiol. 143, 558–569. Lu, H.C., Hsieh, M.H., Chen, C.E., Chen, H.H., Wang, H.I., and Yeh, H.H. (2012). A high-throughput virus-induced gene-silencing vector for screening transcription factors in virus-induced plant defense response in orchid. Mol. Plant Microbe Interact. 25, 738–746. https://doi. org/10.1094/MPMI-10-11-0266 Lu, R., Malcuit, I., Moffett, P., Ruiz, M.T., Peart, J., Wu, A.J., Rathjen, J.P., Bendahmane, A., Day, L., and Baulcombe, D.C. (2003a). High throughput virus-induced gene silencing implicates heat shock protein 90 in plant disease resistance. EMBO J. 22, 5690–5699. https:// doi.org/10.1093/emboj/cdg546 Lu, R., Martin-Hernandez, A.M., Peart, J.R., Malcuit, I., and Baulcombe, D.C. (2003b). Virus-induced gene silencing in plants. Methods 30, 296–303. https://doi. org/10.1016/S1046-2023(03)00037-9. Luo, J., Wei, K., Wang, S., Zhao, W., Ma, C., Hettenhausen, C., Wu, J., Cao, G., Sun, G., Baldwin, I.T., et al. (2016). COI1regulated hydroxylation of jasmonoyl-L-isoleucine

318  | Kumar et al.

impairs Nicotiana attenuata’s resistance to the generalist herbivore Spodoptera litura. J. Agric. Food Chem. 64, 2822–2831. https://doi.org/10.1021/acs.jafc.5b06056 Luo, T., Fan, T., Liu, Y., Rothbart, M., Yu, J., Zhou, S., Grimm, B., and Luo, M. (2012). Thioredoxin redox regulates ATPase activity of magnesium chelatase CHLI subunit and modulates redox-mediated signalling intetra pyrrole biosynthesis and home- ostasis of reactive oxygen species in pea plants. Plant Physiol. 159, 118–130. https://doi.org/10.1104/pp.112.195446. Ma, M.,Yan, Y.,Huang, L.,Chen, M.,and Zhao, H. (2012). Virus-induced gene-silencing in wheat spikes and grains and its application in functional analysis of HMW-GSencoding genes. BMC Plant Biol. 12, 141. https://doi. org/10.1186/1471-2229-12-141 Ma, N., Xue, J., Li, Y., Liu, X., Dai, F., Jia, W., Luo, Y., and Gao, J. (2008). Rh-PIP2;1, a rose aquaporin gene, is involved in ethylene-regulated petal expansion. Plant Physiol. 148, 894–907. https://doi.org/10.1104/ pp.108.120154 Ma, X., Nicole, M.C., Meteignier, L.V., Hong, N., Wang, G., and Moffett, P. (2015). Different roles for RNA silencing and RNA processing components in virus recovery and virus-induced gene silencing in plants. J. Exp. Bot. 66, 919–932. https://doi.org/10.1093/jxb/eru447 MacFarlane, S.A. (1999). Molecular biology of the tobraviruses. J. Gen.Virol. 80, 2799–2807. https://doi. org/10.1099/0022-1317-80-11-2799 Magnani, E., Bartling, L., and Hake, S. (2006). From Gateway to MultiSite Gateway in one recombination event. BMC Mol. Biol. 7, 46. Mahadevan, C., Jaleel, A., Deb, L., Thomas, G., and Sakuntala, M. (2015). Development of an Efficient Virus Induced Gene Silencing Strategy in the Non-Model Wild GingerZingiber zerumbet and Investigation of Associated Proteome Changes. PLOS ONE 10, e0124518. https:// doi.org/10.1371/journal.pone.0124518 Manhães, A.M., de Oliveira, M.V., and Shan, L. (2015). Establishment of an efficient virus-induced gene silencing (VIGS) assay in Arabidopsis by Agrobacteriummediated rubbing infection. Methods Mol. Biol. 1287, 235–241. https://doi.org/10.1007/978-1-4939-24530_17 Manmathan, H., Shaner, D., Snelling, J., Tisserat, N., and Lapitan, N. (2013). Virus-induced gene silencing of Arabidopsis thaliana gene homologues in wheat identifies genes conferring improved drought tolerance. J. Exp.Bot. 64, 1381–1392. https://doi.org/10.1093/ jxb/ert003 Mantelin, S., Bhattarai, K.K., Jhaveri, T.Z., and Kaloshian, I. (2013). Mi-1-mediated resistance to Meloidogyne incognita in tomato may not rely on ethylene but hormone perception through ETR3 participates in limiting nematode infection in a susceptible host. PLOS ONE8, e63281. https://doi.org/10.1371/journal. pone.0063281 Mantelin, S., Peng, H.C., Li, B., Atamian, H.S., Takken, F.L., and Kaloshian, I. (2011). The receptor-like kinase SlSERK1 is required for Mi-1-mediated resistance to potato aphids in tomato. Plant J. 67, 459–471. https:// doi.org/10.1111/j.1365-313X.2011.04609.x Marathe, R., Anandalakshmi, R., Smith, T.H., Pruss, G.J., and Vance, V.B. (2000). RNA viruses as inducers,

suppressors and targets of post-transcriptional gene silencing. Plant Mol. Biol. 43, 295–306. Martín-Hernández, A.M., and Baulcombe, D.C. (2008). Tobacco rattle virus 16- Kilo Dalton protein encodes a suppressor of RNA silencing that allows transient viral entry in meristems. J. Virol. 82, 4064–4071. https://doi. org/10.1128/JVI.02438-07 Mase, K., Mizuno, T., Ishihama, N., Fujii, T., Mori, H., Kodama, M., and Yoshioka, H. (2012). Ethylene signaling pathway and MAPK cascades are required for AAL toxin-induced programmed cell death. Mol. Plant Microbe Interact. 25, 1015–1025. https://doi. org/10.1094/MPMI-02-12-0036-R Matin, M.N., Pandeya, D., Baek, K.H., Lee, D.S., Lee, J.H., Kang, H., and Kang, S.G. (2010). Phenotypic and genotypic analysis of rice lesion mimic mutants. Plant Pathology J. 26, 159–169. Matthew, L. (2004). RNAi for plant functional genomics. Comp. Funct. Genomics 5, 240–244. https://doi. org/10.1002/cfg.396 Matthews, R.E.F. (1991). Plant Virology, 3rd edn. (Academic Press, San Diego). Mei, Y., Zhang, C., Kernodle, B.M., Hill, J.H., and Whitham, S.A. (2016). A Foxtail mosaic virus Vector for virusinduced gene silencing in maize. Plant Physiol. 171, 760–772. https://doi.org/10.1104/pp.16.00172 Mejía-Teniente, L., Joaquin-Ramos, A.d.e.J., TorresPacheco, I., Rivera-Bustamante, R.F., Guevara-Olvera, L., Rico-García, E., and Guevara-Gonzalez, R.G. (2015). Silencing of a germin-like protein gene (CchGLP) in geminivirus-resistant pepper (Capsicum chinense Jacq.) BG-3821 increases susceptibility to single and mixed infections by geminiviruses PHYVV and PepGMV. Viruses 7, 6141–6151. https://doi.org/10.3390/ v7122930 Meldau, S., Baldwin, I.T., and Wu, J. (2011). SGT1 regulates wounding- and herbivory-induced jasmonic acid accumulation and Nicotiana attenuata’s resistance to the specialist lepidopteran herbivore Manduca sexta. New Phytol. 189, 1143–1156. https://doi.org/10.1111/ j.1469-8137.2010.03558.x Melech-Bonfil, S., and Sessa, G. (2011). The SlMKK2 and SlMPK2 genes play a role in tomato disease resistance to Xanthomonas campestris pv. vesicatoria. Plant Signal Behav 6, 154–156. Meng, L.H., Wang, R.H., Zhu, B.Z., Zhu, H.L., Luo, Y.B., and Fu, D.Q. (2016). Efficient Virus-Induced Gene Silencing in Solanum rostratum. PLOS ONE 11, e0156228. https://doi.org/10.1371/journal.pone.0156228 Metzlaff, M. (2002). RNA-mediated RNA degradation in transgene- and virus-induced gene silencing. Biol. Chem. 383, 1483–1489. https://doi.org/10.1515/ BC.2002.170 Meziadi, C., Blanchet, S., Richard, M.M., Pilet-Nayel, M.L., Geffroy, V., and Pflieger, S. (2016). Bean pod mottle virus: a new powerful tool for functional genomics studies in Pisum sativum. Plant Biotechnol. J. 14, 1777–1787. https://doi.org/10.1111/pbi.12537 Mishra, N.S., Tuteja, R., and Tuteja, N. (2006). Signaling through MAP kinase networks in plants. Arch. Biochem. Biophys. 452, 55–68. Moeder, W., Del Pozo, O., Navarre, D.A., Martin, G.B., and Klessig, D.F. (2007). Aconitase plays a role in regulating

Virus-Induced Gene Silencing and its Applications |  319

resistance to oxidative stress and cell death in Arabidopsis and Nicotiana benthamiana. Plant Mol. Biol. 63, 273– 287. https://doi.org/10.1007/s11103-006-9087-x Moon, J.Y., Lee, J.H., Oh, C.S., Kang, H.G., and Park, J.M. (2016). Endoplasmic reticulum stress responses function in the HRT-mediated hypersensitive response in Nicotiana benthamiana. Mol. Plant Pathol. 17, 1382– 1397. https://doi.org/10.1111/mpp.12369 Morel, J.B., Godon, C., Mourrain, P., Béclin, C., Boutet, S., Feuerbach, F., Proux, F., and Vaucheret, H. (2002). Fertile hypomorphic ARGONAUTE (ago1) mutants impaired in post-transcriptional gene silencing and virus resistance. Plant Cell 14, 629–639. Mou, S., Liu, Z., Gao, F., Yang, S., Su, M., Shen, L., Wu, Y., and He, S. (2017). CaHDZ27, a Homeodomain-Leucine Zipper I (HD-Zip I) protein, positively regulates the resistance to Ralstonia solanacearum infection in pepper. Mol. Plant Microbe Interact. https://doi.org/10.1094/ MPMI-06-17-0130-R Muangsan, N., Beclin, C., Vaucheret, H., and Robertson, D. (2004). Geminivirus VIGS of endogenous genes requires SGS2/SDE1 and SGS3 and defines a new branch in the genetic pathway for silencing in plants. Plant J. 38, 1004–1014. https://doi.org/10.1111/ j.1365-313X.2004.02103.x Muruganantham, M., Moskovitz, Y., Haviv, S., Horesh, T., Fenigstein, A., Preez, J.d., Stephan, D., Burger, J.T., and Mawassi, M. (2009). Grapevine virus A-mediated gene silencing in Nicotiana benthamiana and Vitis vinifera. J. Virol. Methods 155, 167–174. https://doi. org/10.1016/j.jviromet.2008.10.010 Mustafa, R., Hamza, M., Kamal, H., Mansoor, S., Scheffler, J., and Amin, I. (2017). Tobacco rattle virus-Based Silencing of Enoyl-CoA Reductase Gene and Its Role in Resistance to Cotton Wilt Disease. Mol. Biotechnol. 59, 241–250. https://doi.org/10.1007/s12033-017-0014-y Mustafa, R., Shafiq, M., Mansoor, S., Briddon, R.W., Scheffler, B.E., Scheffler, J., and Amin, I. (2016). Virusinduced gene silencing in cultivated cotton (Gossypium spp.) using Tobacco rattle virus. Mol. Biotechnol. 58, 65–72. https://doi.org/10.1007/s12033-015-9904-z Nagamatsu, A., Masuta, C., Senda, M., Matsuura, H., Kasai, A., Hong, J.S., Kitamura, K., Abe, J., and Kanazawa, A. (2007). Functional analysis of soybean genes involved in flavonoid biosynthesis by virus-induced gene silencing. Plant Biotechnol. J. 5, 778–790. Nam, M., Koh, S., Kim, S.U., Domier, L.L., Jeon, J.H., Kim, H.G., Lee, S.H., Bent, A.F., and Moon, J.S. (2011). Arabidopsis TTR1 causes LRR-dependent lethal systemic necrosis, rather than systemic acquired resistance, to Tobacco ringspot virus. Mol Cells. 32, 421–9. https://doi. org/10.1007/s10059-011-0101-z Nasir, K.H., Takahashi, Y., Ito, A., Saitoh, H., Matsumura, H., Kanzaki, H., Shimizu, T., Ito, M., Fujisawa, S., Sharma, P.C., et al. (2005). High-throughput inplanta expression screening identifies a class II ethylene-responsive element binding factor-like protein that regulates plant cell death and non-host resistance. Plant J. 43, 491–505. Navarro Gallón, S.M., Elejalde-Palmett, C., Daudu, D., Liesecke, F., Jullien, F., Papon, N., Dugé de Bernonville, T., Courdavault, V., Lanoue, A., Oudin, A., et al. (2017).

Virus-induced gene silencing of the two squalene synthase isoforms of apple tree (Malus × domestica L.) negatively impacts phytosterol biosynthesis, plastid pigmentation and leaf growth. Planta 246, 45–60. https://doi.org/10.1007/s00425-017-2681-0 Naylor, M., Reeves, J., Cooper, J.I., Edwards, M.L., and Wang, H. (2005). Construction and properties of a gene-silencing vector based on Poplar mosaic virus (genus Carlavirus). J. Virol. Methods 124, 27–36. Nemchinov, L.G., Boutanaev, A.M., and Postnikova, O.A. (2016). Virus-induced gene silencing of the RPC5like subunit of RNA polymerase III caused pleiotropic effects in Nicotiana benthamiana. Sci Rep 6, 27785. https://doi.org/10.1038/srep27785 Nowara, D., Gay, A., Lacomme, C., Shaw, J., Ridout, C., Douchkov, D., Hensel, G., Kumlehn, J., and Schweizer, P. (2010). HIGS: host-induced gene silencing in the obligate biotrophic fungal pathogen Blumeria graminis. Plant Cell 22, 3130–3141. https://doi.org/10.1105/ tpc.110.077040 Ogata, T., Nagatoshi, Y., Yamagishi, N., Yoshikawa, N., and Fujita, Y. (2017). Virus-induced down-regulation of GmERA1A and GmERA1B genes enhances the stomatal response to abscisic acid and drought resistance in soybean. PLOS ONE 12, e0175650. https://doi. org/10.1371/journal.pone.0175650 Oh, C.S., Martin, G.B., and Beer, S.V. (2007). DspA/E, a type III effector of Erwinia amylovora, is required for early rapid growth in Nicotiana benthamiana and causes NbSGT1-dependent cell death. Mol. Plant Pathol. 8, 255–265. https://doi.org/10.1111/j.13643703.2007.00390.x Oikawa, A., Rahman, A., Yamashita, T., Taira, H., and Kidou, S. (2007). Virus-induced gene silencing of P23k in barley leaf reveals morphological changes involved in secondary wall formation. J. Exp. Bot. 58, 2617–2625. Orzaez, D., Medina, A., Torre, S., Fernández-Moreno, J.P., Rambla, J.L., Fernández-Del-Carmen, A., Butelli, E., Martin, C., and Granell, A. (2009). A visual reporter system for virus-induced gene silencing in tomato fruit based on anthocyanin accumulation. Plant Physiol. 150, 1122–1134. https://doi.org/10.1104/pp.109.139006 Orzaez, D., Mirabel, S., Wieland, W.H., and Granell, A. (2006). Agroinjection of tomato fruits. A tool for rapid functional analysis of transgenes directly in fruit. Plant Physiol. 140, 3–11. Otagaki, S., Arai, M., Takahashi, A., Goto, K., Hong, J.S., Masuta, C., and Kazazawa, A. (2006). Rapid induction of transcriptional and post-transcriptional gene silencing using a novel Cucumber mosaic virus vector. Plant Biotechnol. 23, 259–265. Ouyang, Z., Liu, S., Huang, L., Hong, Y., Li, X., Huang, L., Zhang, Y., Zhang, H., Li, D., and Song, F. (2016). Tomato SlERF.A1, SlERF.B4, SlERF.C3 and SlERF.A3, members of B3 group of ERF family, are required for resistance to Botrytis cinerea. Front. Plant Sci. 7, 1964. https://doi.org/10.3389/fpls.2016.01964 Pacak, A., Geisler, K., Jørgensen, B., Barciszewska-Pacak, M., Nilsson, L., Nielsen, T.H., Johansen, E., Grønlund, M., Jakobsen, I., and Albrechtsen, M. (2010). Investigations of barley stripe mosaic virus as a gene silencing vector in

320  | Kumar et al.

barley roots and in Brachypodium distachyon and oat. Plant Methods 6, 26. https://doi.org/10.1186/17464811-6-26 Padmanabhan, M., and Dinesh-Kumar, S.P. (2009). Virusinduced gene silencing as a tool for delivery of dsRNA into plants. Cold Spring Harb Protoc pdb.prot5139. https://doi.org/10.1101/pdb.prot5139 Page, J.E., Hause, G., Raschke, M., Gao, W., Schmidt, J., Zenk, M.H., and Kutchan, T.M. (2004). Functional analysis of the final steps of the 1-deoxy-D-xylulose 5-phosphate (DXP) pathway to isoprenoids in plants using virus-induced gene silencing. Plant Physiol. 134, 1401–1413. https://doi.org/10.1104/pp.103.038133 Pandey, P., Choudhury, N.R., and Mukherjee, S.K. (2009). A geminiviral amplicon (VA) derived from Tomato leaf curl virus (ToLCV) can replicate in a wide variety of plant species and also acts as a VIGS vector. Virol. J. 6, 152. https://doi.org/10.1186/1743-422X-6-152 Pandey, S.K., Nookaraju, A., Fujino, T., Pattathil, S., and Joshi, C.P. (2016). Virus-induced gene silencing (VIGS)-mediated functional characterization of two genes involved in lignocellulosic secondary cell wall formation. Plant Cell Rep. 35, 2353–2367. https://doi. org/10.1007/s00299-016-2039-2 Pang, J., Zhu, Y., Li, Q., Liu, J., Tian, Y., Liu, Y., and Wu, J. (2013). Development of Agrobacterium-mediated virusinduced gene silencing and performance evaluation of four marker genes in Gossypium barbadense. PLOS ONE 8, e73211. https://doi.org/10.1371/journal. pone.0073211 Park, C., Chae Lim, W., Baek, W., Kim, J.H., Lim, S., Kim, H.S., Kim, K.N., and Lee, C.S. (2017). The Pepper WPP Domain Protein, CaWDP1, Acts as a Novel Negative Regulator of Drought Stress via ABA Signaling. Plant Cell Physiol. 58, 1118. https://doi.org/10.1093/pcp/ pcx055 Park, C., Lim, C.W., Baek, W., and Lee, S.C. (2015). RING Type E3 ligase CaAIR1 in pepper acts in the regulation of ABA signaling and drought stress response. Plant Cell Physiol. 56, 1808–1819. https://doi.org/10.1093/pcp/ pcv103 Park, J.A., Ahn, J.W., Kim, Y.K., Kim, S.J., Kim, J.K., Kim, W.T., and Pai, H.S. (2005). Retinoblastoma protein regulates cell proliferation, differentiation, and endoreduplication in plants. Plant J. 42, 153–163. Park, S.H., Choi, H., Kim, S., Cho, W.K., and Kim, K.H. (2016). Development of virus-induced gene expression and silencing vector derived from Grapevine Algerian Latent Virus. Plant Pathol. J. 32, 371–376. https://doi. org/10.5423/PPJ.NT.11.2015.0237 Park, S.Y., Ryu, S.H., Kwon, S.Y., Lee, H.S., Kim, J.G., and Kwak, S.S. (2003). Differential expression of six novel peroxidase cDNAs from cell cultures of sweetpotato in response to stress. Mol. Genet. Genomics 269, 542–552. https://doi.org/10.1007/s00438-003-0862-y Park, Y.J., Cho, H.K., Jung, H.J., Ahn, C.S., Kang, H., and Pai, H.S. (2011). PRBP plays a role in plastid ribosomal RNA maturation and chloroplast biogenesis in Nicotiana benthamiana. Planta 233, 1073–1085. https://doi. org/10.1007/s00425-011-1362-7 Passardi, F., Cosio, C., Penel, C., and Dunand, C. (2005). Peroxidases have more functions than a Swiss army knife.

Plant Cell Rep. 24, 255–265. https://doi.org/10.1007/ s00299-005-0972-6 Pasumarthy, K.K., Mukherjee, S.K., and Choudhury, N.R. (2011). The presence of Tomato leaf curl Kerala virus AC3 protein enhances viral DNA replication and modulates virus induced gene-silencing mechanism in tomato plants. Virol. J. 8, 178. https://doi.org/10.1186/1743422X-8-178 Patil, B.L., and Fauquet, C. (2015). Light intensity and temperature affect systemic spread of silencing signal in transient agro-infiltration studies. Mol. Plant Pathol. 16, 484–494. Peart, J.R., Cook, G., Feys, B.J., Parker, J.E., and Baulcombe, D.C. (2002). An EDS1 orthologue is required for N-mediated resistance to Tobacco mosaic virus. Plant J. 29, 569–579. Peart, J.R., Mestre, P., Lu, R., Malcuit, I., and Baulcombe, D.C. (2005). NRG1, a CC-NB-LRR protein, together with N, a TIR-NB-LRR protein, mediates resistance to Tobacco mosaic virus. Curr. Biol. 15, 968–973. Peele, C., Jordan, C.V., Muangsan, N., Turnage, M., Egelkrout, E., Eagle, P., Hanley-Bowdoin, L., and Robertson, D. (2001). Silencing of a meristematic gene using geminivirus-derived vectors. Plant J. 27, 357–366. Peng, H.C., and Kaloshian, I. (2014). The tomato leucinerich repeat receptor-like kinases SlSERK3A and SlSERK3B have overlapping functions in bacterial and nematode innate immunity. PLOS ONE 9, e93302. https://doi.org/10.1371/journal.pone.0093302 Petersen, B.O., and Albrechtsen, M. (2005). Evidence implying only unprimed RdRP activity during transitive gene silencing in plants. Plant Mol. Biol. 58, 575–583. https://doi.org/10.1007/s11103-005-7307-4 Pflieger, S., Blanchet, S., Camborde, L., Drugeon, G., Rousseau, A., Noizet, M., Planchais, S., and Jupin, I. (2008). Efficient virus-induced gene silencing in Arabidopsis using a ‘one-step’ TYMV-derived vector. Plant J. 56, 678–690. https://doi.org/10.1111/j.1365313X.2008.03620.x Pflieger, S., Blanchet, S., Meziadi, C., Richard, M.M., Thareau, V., Mary, F., Mazoyer, C., and Geffroy, V. (2014). The ‘one-step’Bean pod mottle virus (BPMV)derived vector is a functional genomics tool for efficient overexpression of heterologous protein, virus-induced gene silencing and genetic mapping of BPMV R-gene in common bean (Phaseolus vulgaris L.). BMC Plant Biol. 14, 232. https://doi.org/10.1186/s12870-014-0232-4 Pignatta, D., Kumar, P., Turina, M., Dandekar, A., and Falk, B.W. (2007). Quantitative analysis of efficient endogenous gene silencing in Nicotiana benthamiana plants using tomato bushy stunt virus vectors that retain the capsid protein gene. Mol. Plant Microbe Interact. 20, 609–618. https://doi.org/10.1094/MPMI-20-6-0609 Preston, J.C., and Hileman, L.C. (2010). SQUAMOSAPROMOTER BINDING PROTEIN 1 initiates flowering in Antirrhinum majus through the activation of meristem identity genes. Plant J. 62, 704–712. https:// doi.org/10.1111/j.1365-313X.2010.04184.x Preston, J.C., Barnett, L.L., Kost, M.A., Oborny, N.J., and Hileman, L.C. (2014). Optimization of virusinduced gene silencing to facilitate evo-devo studies in the emerging model species Mimulus guttatus

Virus-Induced Gene Silencing and its Applications |  321

(Phrymaceae). Annals of the Missouri Bot. Garden 99, 301–312. https://doi.org/10.3417/2010120 Purkayastha, A., Mathur, S., Verma, V., Sharma, S., and Dasgupta, I. (2010). Virus-induced gene silencing in rice using a vector derived from a DNA virus. Planta 232, 1531–1540. https://doi.org/10.1007/s00425010-1273-z Purkayastha, A., Sharma, S., and Dasgupta, I. (2013). Virusinduced gene silencing for rice using agroinoculation. Methods Mol. Biol. 975, 33–45. https://doi. org/10.1007/978-1-62703-278-0_3 Qi, S., Lin, Q., Zhu, H., Gao, F., Zhang, W., and Hua, X. (2016). The RING finger E3 ligase SpRing is a positive regulator of salt stress signaling in salt-tolerant wild tomato species. Plant Cell Physiol. 57, 528–539. https:// doi.org/10.1093/pcp/pcw006 Qian, W., Yu, C., Qin, H., Liu, X., Zhang, A., Johansen, I.E., and Wang, D. (2007). Molecular and functional analysis of phosphomannomutase (PMM) from higher plants and genetic evidence for the involvement of PMM in ascorbic acid biosynthesis in Arabidopsis and Nicotiana benthamiana. Plant J. 49, 399–413. Qian, Y., Mugiira, R.B., and Zhou, X. (2006). A modified viral satellite DNA-based gene silencing vector is effective in association with heterologous begomoviruses. Virus Res. 118, 136–142. Qu, J., Ye, J., Geng, Y.F., Sun, Y.W., Gao, S.Q., Zhang, B.P., Chen, W., and Chua, N.H. (2012). Dissecting functions of KATANIN and WRINKLED1 in cotton fiber development by virus-induced gene silencing. Plant Physiol. 160, 738–748. https://doi.org/10.1104/ pp.112.198564 Quadrana, L., Rodriguez, M.C., López, M., Bermúdez, L., Nunes-Nesi, A., Fernie, A.R., Descalzo, A., Asis, R., Rossi, M., Asurmendi, S., et al. (2011). Coupling virusinduced gene silencing to exogenous green fluorescence protein expression provides a highly efficient system for functional genomics in Arabidopsis and across all stages of tomato fruit development. Plant Physiol. 156, 1278–1291. https://doi.org/10.1104/pp.111.177345 Ranjan, A., Jayaraman, D., Grau, C., Hill, J.H., Whitham, S.A., Ané, J.M., Smith, D.L., and Kabbage, M. (2017). The pathogenic development of Sclerotinia sclerotiorum in soybean requires specific host NADPH oxidases. Mol. Plant Pathol. [Epub ahead of print]. https://doi. org/10.1111/mpp.12555 Ratcliff, F., Martin-Hernandez, A.M., and Baulcombe, D.C. (2001). Tobacco rattle virus as a vector for analysis of gene function by silencing. Plant J 25, 237–245. Renner, T., Bragga, J., Driscoll, H.E., Cho, J., Jackson, A.O., and Specht, C.D. (2009). Virus-induced gene silencing in the culinary ginger (Zingiber officinale): an effective mechanism for down-regulating gene expression in tropical monocots. Mol Plant 2, 1084–1094. https:// doi.org/10.1093/mp/ssp033 Rivas, S., Rougon-Cardoso, A., Smoker, M., Schauser, L., Yoshioka, H., and Jones, J.D. (2004). CITRX thioredoxin interacts with the tomato Cf-9 resistance protein and negatively regulates defence. EMBO J. 23, 2156–2165. https://doi.org/10.1038/sj.emboj.7600224 Robertson, D. (2004). VIGS vectors for gene silencing: many targets, many tools. Annu Rev Plant Biol

55, 495–519. https://doi.org/10.1146/annurev. arplant.55.031903.141803 Robertson, N.L., French, R., and Morris, T.J. (2000). The open reading frame 5A of foxtail mosaic virus is expressed in vivo and is dispensable for systemic infection. Arch. Virol. 145, 1685–1698. Rodriguez, M.C., Petersen, M., and Mundy, J. (2010). Mitogen-activated protein kinase signaling in plants. Annu. Rev. Plant Biol. 61, 621–649. https://doi. org/10.1146/annurev-arplant-042809-112252 Romero, I., Tikunov, Y., and Bovy, A. (2011). Virus-induced gene silencing in detached tomatoes and biochemical effects of phytoene desaturase gene silencing. J. Plant Physiol. 168, 1129–1135. https://doi.org/10.1016/j. jplph.2010.12.020 Rotenberg, D., Thompson, T.S., German, T.L., and Willis, D.K. (2006). Methods for effective real-time RT-PCR analysis of virus-induced gene silencing. J. Virol. Methods 138, 49–59. Rowland, O., Ludwig, A.A., Merrick, C.J., Baillieul, F., Tracy, F.E., Durrant, W.E., Fritz-Laylin, L., Nekrasov, V., Sjölander, K., Yoshioka, H., et al. (2005). Functional analysis of Avr9/Cf-9 rapidly elicited genes identifies a protein kinase, ACIK1, that is essential for full Cf-9dependent disease resistance in tomato. Plant Cell 17, 295–310. Ruiz, M.T., Voinnet, O., and Baulcombe, D.C. (1998). Initiation and maintenance of virus-induced gene silencing Plant Cell 10, 937–946. Ryu, C.M., Anand, A., Kang, L., and Mysore, K.S. (2004). Agrodrench: a novel and effective agroinoculation method for virus-induced gene silencing in roots and diverse solanaceous species. Plant J. 40, 322–331. https://doi.org/10.1111/j.1365-313X.2004.02211.x Saedler, R., and Baldwin, I.T. (2004). Virus-induced gene silencing of jasmonate-induced direct defences, nicotine and trypsin proteinase-inhibitors in Nicotiana attenuata. J. Exp. Bot. 55, 151–157. https://doi.org/10.1093/jxb/ erh004 Saitoh, H., and Terauchi, R. (2002). Virus-induced silencing of FtsH gene in Nicotiana benthmiana causes a striking bleached leaf phenotype. Genes Genet. Syst. 77, 335–340. Sang, M.K., Kim, J.G., and Kim, K.D. (2010). Biocontrol activity and induction of systemic resistance in pepper by compost water extracts against Phytophthora capsici. Phytopathology 100, 774–783. https://doi. org/10.1094/PHYTO-100-8-0774 Sarowar, S., Kim, Y.J., Kim, K.D., Hwang, B.K., Ok, S.H., and Shin, J.S. (2009). Overexpression of lipid transfer protein (LTP) genes enhances resistance to plant pathogens and LTP functions in long-distance systemic signaling in tobacco. Plant Cell Rep. 28, 419–427. https://doi. org/10.1007/s00299-008-0653-3 Sarowar, S., Oh, H.W., Cho, H.S., Baek, K.H., Seong, E.S., Joung, Y.H., Choi, G.J., Lee, S., and Choi, D. (2007). Capsicum annuum CCR4-associated factor CaCAF1 is necessary for plant development and defence response. Plant J. 51, 792–802. Sasaki, S., Yamagishi, N., and Yoshikawa, N. (2011). Efficient virus-induced gene silencing in apple, pear and Japanese pear using Apple latent spherical virus vectors.

322  | Kumar et al.

Plant Methods 7, 15. https://doi.org/10.1186/17464811-7-15 Scholthof, H.B. (2006). The Tombusvirus-encoded P19: from irrelevance to elegance. Nat. Rev. Microbiol. 4, 405–411. Schornack, S., Ballvora, A., Gürlebeck, D., Peart, J., Baulcombe, D., Ganal, M., Baker, B., Bonas, U., and Lahaye, T. (2004). The tomato resistance protein Bs4 is a predicted non-nuclear TIR-NB-LRR protein that mediates defense responses to severely truncated derivatives of AvrBs4 and overexpressed AvrBs3. Plant J. 37, 46–60. Schultz, T., van Eck, L., and Botha, A.M. (2015). Phi-class glutathione-S-transferase is involved in Dn1-mediated resistance. Physiol Plant 154, 1–12. https://doi. org/10.1111/ppl.12284 Scofield, S.R., and Nelson, R.S. (2009). Resources for virusinduced gene silencing in the grasses. Plant Physiol. 149, 152–157. https://doi.org/10.1104/pp.108.128702 Scofield, S.R., Huang, L., Brandt, A.S., and Gill, B.S. (2005). Development of a virus-induced gene-silencing system for hexaploid wheat and its use in functional analysis of the Lr21-mediated leaf rust resistance pathway. Plant Physiol. 138, 2165–2173. Scofield, S.R., and Brandt, A.S. (2012). Virus-induced gene silencing in hexaploid wheat using barley stripe mosaic virus vectors. Methods Mol. Biol. 894, 93–112. https:// doi.org/10.1007/978-1-61779-882-5_7 Senthil-Kumar, M., and Mysore, K.S. (2011a). New dimensions for VIGS in plant functional genomics. Trends Plant Sci. 16, 656–665. https://doi. org/10.1016/j.tplants.2011.08.006 Senthil-Kumar, M., and Mysore, K.S. (2011b). Virusinduced gene silencing can persist for more than 2 years and also be transmitted to progeny seedlings in Nicotiana benthamiana and tomato. Plant Biotechnol. J. 9, 797–806. https://doi.org/10.1111/j.1467-7652.2011.00589.x Senthil-Kumar, M., and Mysore, K.S. (2014). Tobacco rattle virus-based virus-induced gene silencing in Nicotiana benthamiana. Nat Protoc 9, 1549–1562. https://doi. org/10.1038/nprot.2014.092 Senthil-Kumar, M., and Udayakumar, M. (2006). Highthroughput virus-induced gene-silencing approach to assess the functional relevance of a moisture stressinduced cDNA homologous to lea4. J. Exp. Bot. 57, 2291–2302. Senthil-Kumar, M., and Udayakumar, M. (2010). Post transcriptional gene silencing methods for functional characterization of abiotic stress responsive genes in plants. In Gene Silencing: Theory, Techniques and Applications, Catalano, A.J., ed. (Nova Sci Publishing Inc, New York). Senthil-Kumar, M., Hema, R., Anand, A., Kang, L., Udayakumar, M., and Mysore, K.S. (2007). A systematic study to determine the extent of gene silencing in Nicotiana benthamiana and other Solanaceae species when heterologous gene sequences are used for virusinduced gene silencing. New Phytol. 176, 782–791. Sharma, B., and Kramer, E.M. (2013). Virus-induced gene silencing in the rapid cycling columbine Aquilegia coerulea ‘Origami’. Methods Mol. Biol. 975, 71–81. https://doi.org/10.1007/978-1-62703-278-0_6

Sharma, P.C., Ito, A., Shimizu, T., Terauchi, R., Kamoun, S., and Saitoh, H. (2003). Virus-induced silencing of WIPK and SIPK genes reduces resistance to a bacterial pathogen, but has no effect on the INF1-induced hypersensitive response (HR) in Nicotiana benthamiana. Mol. Genet. Genomics 269, 583–591. https://doi. org/10.1007/s00438-003-0872-9 Shen, L., Liu, Z., Yang, S., Yang, T., Liang, J., Wen, J., Liu, Y., Li, J., Shi, L., Tang, Q., et al. (2016a). Pepper CabZIP63 acts as a positive regulator during Ralstonia solanacearum or high temperature-high humidity challenge in a positive feedback loop with CaWRKY40. J. Exp. Bot. 67, 2439–2451. https://doi.org/10.1093/jxb/erw069 Shen, L., Yang, S., Yang, T., Liang, J., Cheng, W., Wen, J., Liu, Y., Li, J., Shi, L., Tang, Q., et al. (2016b). CaCDPK15 positively regulates pepper responses to Ralstonia solanacearum inoculation and forms a positive-feedback loop with CaWRKY40 to amplify defense signaling. Sci Rep 6, 22439. https://doi.org/10.1038/srep22439 Shen, X., Zhao, K., Liu, L., Zhang, K., Yuan, H., Liao, X., Wang, Q., Guo, X., Li, F., and Li, T. (2014). A role for PacMYBA in ABA-regulated anthocyanin biosynthesis in red-colored sweet cherry cv. Hong Deng (Prunus avium L.). Plant Cell Physiol. 55, 862–880. https://doi. org/10.1093/pcp/pcu013 Shen, Z., Sun, J., Yao, J., Wang, S., Ding, M., Zhang, H., Qian, Z., Zhao, N., Sa, G., Zhao, R., et al. (2015). High rates of virus-induced gene silencing by Tobacco rattle virus in Populus. Tree Physiol. 35, 1016–1029. https://doi. org/10.1093/treephys/tpv064 Shi, Y., Wang, R., Luo, Z., Jin, L., Liu, P., Chen, Q., Li, Z., Li, F., Wei, C., Wu, M., et al. (2014). Molecular cloning and functional characterization of the lycopene ε-cyclase gene via virus-induced gene silencing and its expression pattern in Nicotiana tabacum. Int J Mol Sci 15, 14766– 14785. https://doi.org/10.3390/ijms150814766 Singh, A.K., Kumar, S.R., Dwivedi, V., Rai, A., Pal, S., Shasany, A.K., and Nagegowda, D.A. (2017). A WRKY transcription factor from Withania somnifera regulates triterpenoid withanolide accumulation and biotic stress tolerance through modulation of phytosterol and defense pathways. New Phytol. 215, 1115–1131. https://doi.org/10.1111/nph.14663 Song, J., Ye, G., Qian, Z., and Ye, Q. (2016). Virus-induced plasma membrane aquaporin PsPIP2;1 silencing inhibits plant water transport of Pisum sativum. Bot Stud 57, 15. https://doi.org/10.1186/s40529-016-0135-9 Sonoda, S., and Nishiguchi, M. (2000a). Delayed activation of posttranscriptional gene silencing and de novo transgene methylation in plants with the coat protein gene of sweet potato feathery mottle potyvirus. Plant Sci. 156, 137–144. https://doi.org/10.1016/S01689452(00)00239-9 Sonoda, S., and Nishiguchi, M. (2000b). Graft transmission of posttranscriptional gene silencing: target specificity for RNA degradation is transmissible between silenced and non-silenced plants, but not between silenced plants. Plant J. 21, 1–8. Spitzer, B., Zvi, M.M., Ovadis, M., Marhevka, E., Barkai, O., Edelbaum, O., Marton, I., Masci, T., Alon, M., Morin, S., et al. (2007). Reverse genetics of floral scent: application of Tobacco rattle virus-based gene silencing in Petunia. Plant Physiol. 145, 1241–1250.

Virus-Induced Gene Silencing and its Applications |  323

Stewart, C., Kang, B.C., Liu, K., Mazourek, M., Moore, S.L., Yoo, E.Y., Kim, B.D., Paran, I., and Jahn, M.M. (2005). The Pun1 gene for pungency in pepper encodes a putative acyltransferase. Plant J. 42, 675–688. Stulemeijer, I.J., Stratmann, J.W., and Joosten, M.H. (2007). Tomato mitogen-activated protein kinases LeMPK1, LeMPK2, and LeMPK3 are activated during the Cf-4/ Avr4-induced hypersensitive response and have distinct phosphorylation specificities. Plant Physiol. 144, 1481–1494. Sun, D., Li, S., Niu, L., Reid, M.S., Zhang, Y., and Jiang, C.Z. (2017a). PhOBF1, a petunia ocs element binding factor, plays an important role in antiviral RNA silencing. J. Exp. Bot. 68, 915–930. https://doi.org/10.1093/jxb/ erw490 Sun, Y., Wang, C., Wang, N., Jiang, X., Mao, H., Zhu, C., Wen, F., Wang, X., Lu, Z., Yue, G., et al. (2017b). Manipulation of Auxin Response Factor 19 affects seed size in the woody perennial Jatropha curcas. Sci Rep 7, 40844. https://doi.org/10.1038/srep40844 Tai, Y.S., Bragg, J., and Edwards, M.C. (2005). Virus vector for gene silencing in wheat. BioTechniques 39, 310–314. Tameling, W.I., and Baulcombe, D.C. (2007). Physical association of the NB-LRR resistance protein Rx with a Ran GTPase-activating protein is required for extreme resistance to Potato virus X. Plant Cell 19, 1682–1694. Tamura, A., Kato, T., Taki, A., Sone, M., Satoh, N., Yamagishi, N., Takahashi, T., Ryo, B.S., Natsuaki, T., and Yoshikawa, N. (2013). Preventive and curative effects of Apple latent spherical virus vectors harboring part of the target virus genome against potyvirus and cucumovirus infections. Virology 446, 314–324. https://doi.org/10.1016/j. virol.2013.08.019 Tang, Y., Lai, Y., and Liu, Y. (2013). Virus-induced gene silencing using artificial miRNAs in Nicotiana benthamiana. Methods Mol. Biol. 975, 99–107. https:// doi.org/10.1007/978-1-62703-278-0_8 Tang, Y., Wang, F., Zhao, J., Xie, K., Hong, Y., and Liu, Y. (2010). Virus-based microRNA expression for gene functional analysis in plants. Plant Physiol. 153, 632– 641. https://doi.org/10.1104/pp.110.155796 Tao, X., and Zhou, X. (2004). A modified viral satellite DNA that suppresses gene expression in plants. Plant J. 38, 850–860. https://doi.org/10.1111/j.1365313X.2004.02087.x Tao, X.R., Zhou, X.P., Cui, X.F., and Qian, Y.J. (2004). Virus-induced gene silencing and its application for analysis of genomic function in plants. Prog Biochem Biophys 31, 777–783. Tasaki, K., Terada, H., Masuta, C., and Yamagishi, M. (2016). Virus-induced gene silencing (VIGS) in Lilium leichtlinii using the Cucumber mosaic virus vector. Plant Biotech. 33, 1–9. https://doi.org/10.5511/ plantbiotechnology.16.1018a Tena, G., Asai, T., Chiu, W.L., and Sheen, J. (2001). Plant mitogen-activated protein kinase signaling cascades. Curr. Opin. Plant Biol. 4, 392–400. Thirumalaikumar, V.P., Devkar, V., Mehterov, N., Ali, S., Ozgur, R., Turkan, I., Mueller-Roeber, B., and Balazadeh, S. (2017). NAC transcription factor JUNGBRUNNEN1 enhances drought tolerance in tomato. Plant Biotechno.l J. 16, 354–366. https://doi.org/10.1111/pbi.12776

Tian, J., Pei, H., Zhang, S., Chen, J., Chen, W., Yang, R., Meng, Y., You, J., Gao, J., and Ma, N. (2014a).TRV-GFP: a modified Tobacco rattle virus vector for efficient and visualizable analysis of gene function.J. Exp. Bot. 65, 311–322. https://doi.org/10.1093/jxb/ert381 Tian, S.L., Li, L., Chai, W.G., Shah, S.N., and Gong, Z.H. (2014b). Effects of silencing key genes in the capsanthin biosynthetic pathway on fruit color of detached pepper fruits. BMC Plant Biol. 14, 314. https://doi. org/10.1186/s12870-014-0314-3 Todd, A.T., Liu, E., Polvi, S.L., Pammett, R.T., and Page, J.E. (2010). A functional genomics screen identifies diverse transcription factors that regulate alkaloid biosynthesis in Nicotiana benthamiana. Plant J. 62, 589–600. https:// doi.org/10.1111/j.1365-313X.2010.04186.x Tran, P.T., Choi, H., Choi, D., and Kim, K.H. (2016). Virus-induced gene silencing reveals signal transduction components required for the Pvr9-mediated hypersensitive response in Nicotiana benthamiana. Virology 495, 167–172. https://doi.org/10.1016/j. virol.2016.05.011 Tsaballa, A., Pasentsis, K., Darzentas, N., and Tsaftaris, A.S. (2011). Multiple evidence for the role of an Ovate-like gene in determining fruit shape in pepper. BMC Plant Biol. 11, 46. https://doi.org/10.1186/1471-2229-11-46 Tufan, H.A., Stefanato, F.L., McGrann, G.R., MacCormack, R., and Boyd, L.A. (2011). The Barley stripe mosaic virus system used for virus-induced gene silencing in cereals differentially affects susceptibility to fungal pathogens in wheat. J. Plant Physiol. 168, 990–994. https://doi. org/10.1016/j.jplph.2010.11.019 Turnage, M.A., Muangsan, N., Peele, C.G., and Robertson, D. (2002). Geminivirus-based vectors for gene silencing in Arabidopsis. Plant J.30, 107–114. Tuttle, J.R., Haigler, C.H., and Robertson, D. (2012). Method: low-cost delivery of the cotton leaf crumple virus-induced gene silencing system. Plant Methods 8, 27. https://doi.org/10.1186/1746-4811-8-27 Tuttle, J.R., Haigler, C.H., and Robertson, D.N. (2015). Virus-induced gene silencing of fiber-related genes in cotton. Methods Mol. Biol. 1287, 219–234. https://doi. org/10.1007/978-1-4939-2453-0_16 Tuttle, J.R., Idris, A.M., Brown, J.K., Haigler, C.H., and Robertson, D. (2008). Gemini virus-mediated gene silencing from Cotton leaf crumple virus is enhanced by low temperature in cotton. Plant Physiol. 148, 41–50. https://doi.org/10.1104/pp.108.123869 Uppalapati, S.R., Ishiga, Y., Wangdi, T., Kunkel, B.N., Anand, A., Mysore, K.S., and Bender, C.L. (2007). The phytotoxin coronatine contributes to pathogen fitness and is required for suppression of salicylic acid accumulation in tomato inoculated with Pseudomonas syringae pv. tomato DC3000. Mol. Plant Microbe Interact. 20, 955–965. https://doi.org/10.1094/ MPMI-20-8-0955 Vaghchhipawala, Z.E., and Mysore, K.S. (2008). Agroinoculation: a simple procedure for systemic infection of plants with viruses. Methods Mol. Biol. 451, 555–562. https://doi.org/10.1007/978-1-59745-1024_38 Vaistij, F.E., and Jones, L. (2009). Compromised virusinduced gene silencing in RDR6-deficient plants. Plant

324  | Kumar et al.

Physiol. 149, 1399–1407. https://doi.org/10.1104/ pp.108.132688 Valentine, T., Shaw, J., Blok, V.C., Phillips, M.S., Oparka, K.J., and Lacomme, C. (2004). Efficient virus-induced gene silencing in roots using a modified Tobacco rattle virus vector. Plant Physiol. 136, 3999–4009. Valério, L., De Meyer, M., Penel, C., and Dunand, C. (2004). Expression analysis of the Arabidopsis peroxidase multigenic family. Phytochemistry 65, 1331–1342. https://doi.org/10.1016/j.phytochem.2004.04.017 van der Linde, K., and Doehlemann, G. (2013). Utilizing virus-induced gene silencing for the functional characterization of maize genes during infection with the fungal pathogen Ustilago maydis. Methods Mol. Biol. 975, 47–60. https://doi.org/10.1007/978-1-62703278-0_4 van der Linde, K., Kastner, C., Kumlehn, J., Kahmann, R., and Doehlemann, G. (2011). Systemic virus-induced gene silencing allows functional characterization of maize genes during biotrophic interaction with Ustilago maydis. New Phytol. 189, 471–483. https://doi. org/10.1111/j.1469-8137.2010.03474.x Van Eck, L., Schultz, T., Leach, J.E., Scofield, S.R., Peairs, F.B., Botha, A.M., and Lapitan, N.L. (2010). Virusinduced gene silencing of WRKY53 and an inducible phenylalanine ammonia-lyase in wheat reduces aphid resistance. Plant Biotechnol. J. 8, 1023–1032. https:// doi.org/10.1111/j.1467-7652.2010.00539.x van Kammen, A. (1997). Virus-induced gene silencing in infected and transgenic plants. Trends Plant Sci. 2, 409–411. https://doi.org/10.1016/S13601385(97)01128-X Vance, V., and Vaucheret, H. (2001). RNA silencing in plants – defense and counterdefense. Science 292, 2277–2280. https://doi.org/10.1126/science.1061334 Várallyay, E., Lichner, Z., Sáfrány, J., Havelda, Z., Salamon, P., Bisztray, G., and Burgyán, J. (2010). Development of a virus induced gene silencing vector from a legumes infecting tobamovirus. Acta. Biol. Hung. 61, 457–469. https://doi.org/10.1556/ABiol.61.2010.4.9 Velásquez, A.C., Chakravarthy, S., and Martin, G.B. (2009). Virus-induced gene silencing (VIGS) in Nicotiana benthamiana and tomato. J Vis Exp 28, 1292. https:// doi.org/10.3791/1292 Virk, N., Liu, B., Zhang, H., Li, X., Zhang, Y., Li, D., and Song, F. (2013). Tomato SlMPK4 is required for resistance to Botrytis cinerea and tolerance to drought stress. Acta Physiol. Plant 35, 1211–1221. https://doi. org/10.1007/s11738-012-1160-2 Visser, P.B., and Bol, J.F. (1999). Nonstructural proteins of Tobacco rattle virus which have a role in nematodetransmission: expression pattern and interaction with viral coat protein. J. Gen. Virol. 80, 3273–3280. https:// doi.org/10.1099/0022-1317-80-12-3273 Voinnet, O. (2001). RNA silencing as a plant immune system against viruses. Trends Genet. 17, 449–459. Voinnet, O. (2005). Induction and suppression of RNA silencing: insights from viral infections. Nat. Rev. Genet. 6, 206–220. Vossen, J.H., Abd-El-Haliem, A., Fradin, E.F., van den Berg, G.C., Ekengren, S.K., Meijer, H.J., Seifi, A., Bai, Y., ten Have, A., Munnik, T., et al. (2010). Identification of tomato phosphatidylinositol-specific phospholipase-C

(PI-PLC) family members and the role of PLC4 and PLC6 in HR and disease resistance. Plant J. 62, 224–239. https://doi.org/10.1111/j.1365-313X.2010.04136.x Walter, S., Kahla, A., Arunachalam, C., Perochon, A., Khan, M.R., Scofield, S.R., and Doohan, F.M. (2015). A wheat ABC transporter contributes to both grain formation and mycotoxin tolerance. J. Exp. Bot. 66, 2583–2593. https://doi.org/10.1093/jxb/erv048 Wang, C., Cai, X., Wang, X., and Zheng, Z. (2006). Optimisation of Tobacco rattle virus-induced gene silencing in Arabidopsis. Func. Plant Biol. 33, 347–355. https://doi.org/10.1071/FP05096. Wang, C., Lu, W., He, X., Wang, F., Zhou, Y., Guo, X., and Guo, X. (2016a). The Cotton Mitogen-Activated Protein Kinase Kinase 3 Functions in Drought Tolerance by Regulating Stomatal Responses and Root Growth. Plant Cell Physiol. 57, 1629–1642. https://doi.org/10.1093/ pcp/pcw090 Wang, F., Wang, M., Liu, X., Xu, Y., Zhu, S., Shen, W., and Zhao, X. (2017a). Identification of Putative Genes Involved in Limonoids Biosynthesis in Citrus by Comparative Transcriptomic Analysis. Front. Plant Sci. 8, 782. https://doi.org/10.3389/fpls.2017.00782 Wang, F., Lin, R., Feng, J., Chen, W., Qiu, D., and Xu, S. (2015a). TaNAC1 acts as a negative regulator ofstriperust resistance in wheat, enhances susceptibility to Pseudomonas syringae, and promotes lateral root development in transgenic Arabidopsis thaliana. Front. Plant Sci. 6, 108. https://doi.org/10.3389/ fpls.2015.00108 Wang, G.F., Wei, X., Fan, R., Zhou, H., Wang, X., Yu, C., Dong, L., Dong, Z., Wang, X., Kang, Z., et al. (2011). Molecular analysis of common wheat genes encoding three types of cytosolic heat shock protein 90 (Hsp90): functional involvement of cytosolic Hsp90s in the control of wheat seedling growth and disease resistance. New Phytol. 191, 418–431. https://doi.org/10.1111/ j.1469-8137.2011.03715.x Wang, K., Uppalapati, S.R., Zhu, X., Dinesh-Kumar, S.P., and Mysore, K.S. (2010a). SGT1 positively regulates the process of plant cell death during both compatible and incompatible plant-pathogen interactions. Mol. Plant Pathol. 11, 597–611. https://doi.org/10.1111/j.13643703.2010.00631.x Wang, L., Tang, H., Wang, X., Chen, Q., Jiang, L., and Lin, Y. (2017b). Virus-induced gene silencing as a tool for FaMYB5 gene functional studies in strawberry. Acta Horticulturae Sinica 44, 33–42. https://doi. org/10.16420/j.issn.0513-353x.2016-0346 Wang, R., Yang, X., Wang, N., Liu, X., Nelson, R.S., Li, W., Fan, Z., and Zhou, T. (2016b). An efficient virusinduced gene silencing vector for maize functional genomics research. Plant J. 86, 102–115. https://doi. org/10.1111/tpj.13142 Wang, S.B., Tian, S.L., Shah, S.N., Pan, B.G., Diao, W.P., and Gong, Z.H. (2015b). Cloning and characterization of the CarbcL gene related to chlorophyll in pepper (Capsicum annuum L.) under fruit shade stress. Front Plant Sci6, 850. https://doi.org/10.3389/fpls.2015.00850 Wang, T., Li, R., Wen, L., Fu, D., Zhu, B., Luo, Y., and Zhu, H. (2015c). Functional Analysis and RNA Sequencing Indicate the Regulatory Role of Argonaute1 in Tomato Compound Leaf Development. PLOS ONE

Virus-Induced Gene Silencing and its Applications |  325

10, e0140756. https://doi.org/10.1371/journal. pone.0140756 Wang, W., Yuan, Y., Yang, C., Geng, S., Sun, Q., Long, L., Cai, C., Chu, Z., Liu, X., Wang, G., et al. (2016c). Characterization, Expression, and Functional Analysis of a Novel NAC Gene Associated with Resistance to Verticillium Wilt and Abiotic Stress in Cotton. G3 6, 3951–3961. https://doi.org/10.1534/g3.116.034512 Wang, X., Cao, A., Yu, C., Wang, D., Wang, X., and Chen, P. (2010b). Establishment of an effective virus induced gene silencing system with BSMV in Haynaldia villosa. Mol. Biol. Rep. 37, 967–972. https://doi.org/10.1007/ s11033-009-9766-1 Wang, Y., Dang, F., Liu, Z., Wang, X., Eulgem, T., Lai, Y., Yu, L., She, J., Shi, Y., Lin, J., et al. (2013). CaWRKY58, encoding a group I WRKY transcription factor of Capsicum annuum, negatively regulates resistance to Ralstonia solanacearum infection. Mol. Plant Pathol. 14, 131–144. https://doi.org/10.1111/j.13643703.2012.00836.x Waterhouse, P.M., Wang, M.B., and Lough, T. (2001). Gene silencing as an adaptive defence against viruses. Nature 411, 834–842. https://doi.org/10.1038/35081168 Wege, S., Scholz, A., Gleissberg, S., and Becker, A. (2007). Highly efficient virus-induced gene silencing (VIGS) in California poppy (Eschscholzia californica): an evaluation of VIGS as a strategy to obtain functional data from nonmodel plants. Ann. Bot. 100, 641–649. Wijekoon, C.P., and Facchini, P.J. (2012). Systematic knockdown of morphine pathway enzymes in opium poppy using virus-induced gene silencing. Plant J. 69, 1052–1063. https://doi.org/10.1111/j.1365313X.2011.04855.x Wünsche, H., Baldwin, I.T., and Wu, J. (2011). S-Nitrosoglutathione reductase (GSNOR) mediates the biosynthesis of jasmonic acid and ethylene induced by feeding of the insect herbivore Manduca sexta and is important for jasmonate-elicited responses in Nicotiana attenuata. J. Exp. Bot. 62, 4605–4616. https://doi. org/10.1093/jxb/err171 Xu, J., and Zhang, S. (2015). Mitogen-activated protein kinase cascades in signaling plant growth and development. Trends Plant Sci. 20, 56–64. https://doi. org/10.1016/j.tplants.2014.10.001 Xu, W., Meng, Y., Surana, P., Fuerst, G., Nettleton, D., and Wise, R.P. (2015). The knottin-like Blufensin family regulates genes involved in nuclear import and the secretory pathway in barley-powdery mildew interactions. Front. Plant Sci. 6, 409. https://doi. org/10.3389/fpls.2015.00409 Yaegashi, H., Yamatsuta, T., Takahashi, T., Li, C., Isogai, M., Kobori, T., Ohki, S., and Yoshikawa, N. (2007). Characterization of virus-induced gene silencing in tobacco plants infected with apple latent spherical virus. Arch. Virol. 152, 1839–1849. https://doi.org/10.1007/ s00705-007-1011-4 Yamagishi, N., and Yoshikawa, N. (2009). Virus-induced gene silencing in soybean seeds and the emergence stage of soybean plants with Apple latent spherical virus vectors. Plant Mol. Biol. 71, 15–24. https://doi.org/10.1007/ s11103-009-9505-y Yamagishi, N., and Yoshikawa, N. (2013). Highly efficient virus-induced gene silencing in apple and soybean

by apple latent spherical virus vector and biolistic inoculation. Methods Mol. Biol. 975, 167–181. https:// doi.org/10.1007/978-1-62703-278-0_13 Yan, H.X., Fu, D.Q., Zhu, B.Z., Liu, H.P., Shen, X.Y., and Luo, Y.B. (2012). Sprout vacuum-infiltration: a simple and efficient agroinoculation method for virus-induced gene silencing in diverse solanaceous species. Plant Cell Rep. 31, 1713–1722. https://doi.org/10.1007/s00299012-1285-1 Yan, Y., Jia, H., Wang, F., Wang, C., Liu, S., and Guo, X. (2015). Overexpression of GhWRKY27a reduces tolerance to drought stress and resistance to Rhizoctonia solani infection in transgenic Nicotiana benthamiana. Front Physiol 6, 265. https://doi.org/10.3389/ fphys.2015.00265 Yang, D.H., Hettenhausen, C., Baldwin, I.T., and Wu, J. (2011). BAK1 regulates the accumulation of jasmonic acid and the levels of trypsin proteinase inhibitors in Nicotiana attenuata’s responses to herbivory. J. Exp. Bot. 62, 641–652. https://doi.org/10.1093/jxb/erq298 Yang, J., Ma, Q., Zhang, Y., Wang, X., Zhang, G., and Ma, Z. (2016). Molecular cloning and functional analysis of GbRVd, a gene in Gossypium barbadense that plays an important role in conferring resistance to Verticillium wilt. Gene. 575, 687–94. https://doi.org/10.1016/j. gene.2015.09.046 Yang, K.S., Jin, U.H., Kim, J., Song, K., Kim, S.J., Hwang, I., Lim, Y.P., and Pai, H.S. (2004). Molecular characterization of NbCHMP1 encoding a homolog of human CHMP1 in Nicotiana benthamiana. Mol. Cells 17, 255–261. Yang, W., Cai, Y., Hu, L., Wei, Q., Chen, G., Bai, M., Wu, H., Liu, J., and Yu, Y. (2017a). PhCESA3 silencing inhibits elongation and stimulates radial expansion in petunia. Sci. Rep. 7, 41471. https://doi.org/10.1038/srep41471 Yang, W., Lou, X., Li, J., Pu, M., Mirbahar, A.A., Liu, D., Sun, J., Zhan, K., He, L., and Zhang, A. (2017b). Cloning and Functional Analysis of MADS-box Genes, TaAG-A and TaAG-B, from a Wheat K-type Cytoplasmic Male Sterile Line. Front. Plant Sci. 8, 1081. https://doi.org/10.3389/ fpls.2017.01081 Yang, Y.,Zhao, J.,Liu, P.,Xing, H.,Li, C.,Wei, G.,and Kang, Z. (2013). Glycerol-3-phosphate metabolism in wheat contributes to systemic acquired resistance to Puccinia striiformis f. sp. tritici. PLOS ONE 8, e81756. https:// doi.org/10.1371/journal.pone.0081756 Ye, J., Qu, J., Bui, H.T., and Chua, N.H. (2009). Rapid analysis of Jatropha curcas gene functions by virusinduced gene silencing. Plant Biotechnol. J. 7, 964–976. https://doi.org/10.1111/j.1467-7652.2009.00457.x Yoshida, K., Kaothien, P., Matsui, T., Kawaoka, A., and Shinmyo, A. (2003). Molecular biology and application of plant peroxidase genes. Appl. Microbiol. Biotechnol. 60, 665–670. https://doi.org/10.1007/s00253-0021157-7 Yuan, C., Li, C., Yan, L., Jackson, A.O., Liu, Z., Han, C., Yu, J., and Li, D. (2011). A high throughput barley stripe mosaic virus vector for virus induced gene silencing in monocots and dicots. PLOS ONE 6, e26468. https:// doi.org/10.1371/journal.pone.0026468 Zeng, X., Zeng, Z., Liu, C., Yuan, W., Hou, N., Bian, H., Zhu, M., and Han, N. (2017). A barley homolog of yeast ATG6 is involved in multiple abiotic stress responses

326  | Kumar et al.

and stress resistance regulation. Plant Physiol. Biochem. 115, 97–106. Zhang, C., and Ghabrial, S.A. (2006a). Development of Bean pod mottle virus-based vectors for stable protein expression and sequence-specific virus-induced gene silencing in soybean. Virology 344, 401–411. Zhang, C., Grosic, S., Whitham, S.A., and Hill, J.H. (2012). The requirement of multiple defense genes in soybean Rsv1-mediated extreme resistance to Soybean mosaic virus. Mol. Plant Microbe Interact. 25, 1307–1313. https://doi.org/10.1094/MPMI-02-12-0046-R Zhang, C., Yang, C., Whitham, S.A., and Hill, J.H. (2009a). Development and use of an efficient DNA-based viral gene silencing vector for soybean. Mol. Plant Microbe Interact. 22, 123–131. https://doi.org/10.1094/ MPMI-22-2-0123 Zhang, C., Bradshaw, J.D., Whitham, S.A., and Hill, J.H. (2010). The development of an efficient multipurpose bean pod mottle virus viral vector set for foreign gene expression and RNA silencing. Plant Physiol. 153, 52–65. https://doi.org/10.1104/pp.109.151639 Zhang, J., Yu, D., Zhang, Y., Liu, K., Xu, K., Zhang, F., Wang, J., Tan, G., Nie, X., Ji, Q., et al. (2017). Vacuum and co-cultivation agroinfiltration of (germinated) seeds results in Tobacco rattle virus (TRV) Mediated whole-plant virus-induced gene silencing (VIGS) in wheat and maize. Front. Plant Sci. 8, 393. https://doi. org/10.3389/fpls.2017.00393 Zhang, J.S., Zhao, J., Zhang, S., and He, C. (2014a). Efficient gene silencing mediated by Tobacco rattle virus in an emerging model plant physalis. PLOS ONE 9, e85534. https://doi.org/10.1371/journal.pone.0085534 Zhang, L., Lavery, L., Gill, U., Gill, K., Steffenson, B., Yan, G., Chen, X., and Kleinhofs, A. (2009b). A cation/ proton-exchanging protein is a candidate for the barley NecS1 gene controlling necrosis and enhanced defense response to stem rust. Theor. Appl. Genet. 118, 385–397. https://doi.org/10.1007/s00122-008-0910-4 Zhang, T., Liu, Y., Yang, T., Zhang, L., Xu, S., Xue, L., and An, L. (2006b). Diverse signals converge at MAPK cascades in plant. Plant Physiol. Biochem. 44, 274–283. Zhang, W., Zhang, H., Qi, F., and Jian, G. (2016). Generation of transcriptome profiling and gene functional analysis in Gossypium hirsutum upon Verticillium dahliae infection. Biochem. Biophys. Res. Commun. 473, 879–885. Zhang, X., Wang, L., Xu, X., Cai, C., and Guo, W. (2014b). Genome-wide identification of mitogen-activated protein kinase gene family in Gossypium raimondii and the function of their corresponding orthologs in tetraploid cultivated cotton. BMC Plant Biol. 14, 345. https://doi.org/10.1186/s12870-014-0345-9 Zhang, Y., Li, D., Zhang, H., Hong, Y., Huang, L., Liu, S., Li, X., Ouyang, Z., and Song, F. (2015). Tomato histone H2B monoubiquitination enzymes SlHUB1 and SlHUB2 contribute to disease resistance to Botrytis cinerea through modulating the balance between SA- and JA/ET-mediated signaling pathways. BMC Plant Biol. 15, 252. https://doi.org/10.1186/s12870-015-0614-2 Zhang, Y.L., Jia, Q.L., Li, D.W., Wang, J.E., Yin, Y.X., and Gong, Z.H. (2013). Characteristic of the pepper CaRGA2 gene in defense responses against Phytophthora

capsici Leonian. Int. J. Mol. Sci. 14, 8985–9004. https:// doi.org/10.3390/ijms14058985 Zhang, Z., and Thomma, B.P. (2014). Virus-induced gene silencing and Agrobacterium tumefaciens-mediated transient expression in Nicotiana tabacum. Methods Mol. Biol. 1127, 173–181. https://doi.org/10.1007/978-162703-986-4_14 Zhang, Z., Li, D.W., Jin, J.H., Yin, Y.X., Zhang, H.X., Chai, W.G., and Gong, Z.H. (2015). VIGS approach reveals the modulation of anthocyanin biosynthetic genes by CaMYB in chili pepper leaves. Front. Plant Sci. 6, 500. https://doi.org/10.3389/fpls.2015.00500 Zhao, F., Lim, S., Igori, D., Yoo, R.H., Kwon, S.Y., and Moon, J.S. (2016). Development of Tobacco ringspot virus-based vectors for foreign gene expression and virus-induced gene silencing in a variety of plants. Virology 492, 166–178. https://doi.org/10.1016/j.virol.2016.02.025 Zheng, L., Du, Z., Lin, C., Mao, Q., Wu, K., Wu, J., Wei, T., Wu, Z., and Xie, L. (2015). Rice stripe Tenuivirus p2 may recruit or manipulate nucleolar functions through an interaction with fibrillarin to promote virus systemic movement. Mol. Plant Pathol. 16, 921–930. https://doi. org/10.1111/mpp.12220 Zheng, S.J., Snoeren, T.A., Hogewoning, S.W., van Loon, J.J., and Dicke, M. (2010). Disruption of plant carotenoid biosynthesis through virus-induced gene silencing affects oviposition behaviour of the butter fly Pieris rapae. New Phytol. 186, 733–45. https://doi. org/10.1111/j.1469-8137.2010.03213.x Zheng, S.J., Zhang, P.J., van Loon, J.J., and Dicke, M. (2011). Silencing defense pathways in Arabidopsis by heterologous gene sequences from Brassica oleracea enhances the performance of a specialist and a generalist herbivorous insect. J. Chem. Ecol. 37, 818–829. https:// doi.org/10.1007/s10886-011-9984-6 Zhong, X., Yuan, X., Wu, Z., Khan, M.A., Chen, J., Li, X., Gong, B., Zhao, Y., Wu, J., Wu, C., et al. (2014). Virusinduced gene silencing for comparative functional studies in Gladiolus hybridus. Plant Cell Rep. 33, 301–312. https://doi.org/10.1007/s00299-013-1530-2 Zhou, B., and Zeng, L. (2017). Elucidating the role of highly homologous Nicotiana benthamiana ubiquitin E2 gene family members in plant immunity through an improved virus-induced gene silencing approach. Plant Methods 13, 59. https://doi.org/10.1186/s13007-017-0210-6 Zhou, H., Lin-Wang, K., Wang, H., Gu, C., Dare, A.P., Espley, R.V., He, H., Allan, A.C., and Han, Y. (2015). Molecular genetics of blood-fleshed peach reveals activation of anthocyanin biosynthesis by NAC transcription factors. Plant J. 82, 105–121. https://doi.org/10.1111/ tpj.12792 Zhou, X., and Huang, C. (2012). Virus-induced gene silencing using begomovirus satellite molecules. Methods Mol. Biol. 894, 57–67. https://doi.org/10.1007/978-161779-882-5_4 Zhu, F., Deng, X.G., Xu, F., Jian, W., Peng, X.J., Zhu, T., Xi, D.H., and Lin, H.H. (2015). Mitochondrial alternative oxidase is involved in both compatible and incompatible host-virus combinations in Nicotiana benthamiana. Plant Sci. 239, 26–35. https://doi.org/10.1016/j. plantsci.2015.07.009

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Zhu, M., Chen, Y., Ding, X.S., Webb, S.L., Zhou, T., Nelson, R.S., and Fan, Z. (2014). Maize Elongin C interacts with the viral genome-linked protein,VPg, of Sugarcane mosaic virus and facilitates virus infection. New Phytol. 203, 1291–1304. https://doi.org/10.1111/nph.12890

Zhu, X., Caplan, J., Mamillapalli, P., Czymmek, K., and Dinesh-Kumar, S.P. (2010). Function of endoplasmic reticulum calcium ATPase in innate immunity-mediated programmed cell death. EMBO J. 29, 1007–1018. https://doi.org/10.1038/emboj.2009.402

Possible Strategies for Establishment of VIGS Protocol in Chickpea Ranjita Sinha and Muthappa Senthil-Kumar*

15

National Institute of Plant Genome Research, New Delhi, India. *Correspondence: [email protected], [email protected] https://doi.org/10.21775/9781910190814.15

Abstract Chickpea is the second most common legume in the world. The worldwide production of chickpea is far below its potential because of the factors like nitrogen deficiency, low nutrient absorption, flower or seed abortion and its vulnerability to the abiotic and biotic stresses. Consequently, it is important to understand the key molecular factors involved in stress tolerance, growth, flowering and seed development for the genetic improvement of the existing varieties. Currently, the whole-genome sequencing data and transcriptome information are widely facilitating functional genomic studies in chickpea. Further, marker-based trait association mapping information is available for assistance in breeding programme. However, information about exact function of genes is still lacking because of the absence of genetic mutants and difficult genetic transformation in this crop. Hence, the current scenario demands the establishment of virus-induced gene silencing (VIGS) technique in chickpea. VIGS would serve as an important tool for the functional characterization of large number of genes. Despite attempts by several research teams, the VIGS protocol is not yet available till date, though VIGS has been successfully applied for the gene characterization in other legumes. In this chapter we propose some strategies that can be attempted for development of successful VIGS protocol. We also describe our experience from present and past research projects aimed to study VIGS in chickpea.

Introduction Chickpea is an important legume crop that is a rich source of proteins and carbohydrates. With a worldwide production of 12.2 metric tonnes during 2011–13 (FAO, 2016), chickpea production is still far below its potential (Bhatia et al., 2006). Soil nutrient deficiency, terminal drought, temperature stress, Helicoverpa infestation, dry root rot, black root rot, Fusarium wilt and Ascochyta blight severely restricts its potential yield (Gaur et al. 2010; Nene et al., 2012; Pang et al., 2017; Farooq et al., 2017). In this regard, chickpea productivity can be increased by improving nutrient absorption, grain filling, reproductive growth, abiotic and biotic tolerance where understanding the associated molecular mechanisms is very important. The draft sequences of the chickpea genome (Varshney et al., 2013; Jain et al., 2013; Parween et al., 2015; Gupta et al., 2017) and transcriptome (Garg et al., 2011; Hiremath et al., 2011) are available. Further, the genomic information have been used to develop genome wide simple sequence repeat (SSR) markers (Parida et al., 2015), genome wide single nucleotide polymorphism (SNP) identification and QTL association mapping (Diapari et al., 2014; Deokar et al., 2014; Gaur et al., 2015; Upadhyaya et al., 2016), to accelerate the genotyping in the chickpea breeding programmes. Transcriptomic information facilitated the comparative transcriptomic analysis for various physiological aspects (Agarwal et al., 2012; Jhanwar

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et al., 2012; Pradhan et al., 2014; Garg et al., 2016; Sinha et al., 2017). However, to assess the true relevance and biological function of genes, suitable functional genomics techniques are essential. Functional validation can be done by either overexpression of genes or knockdown of genes using RNAi or mutation, both of which however, require stable transformation. Genetic transformation is difficult in chickpea where transformation frequency for chickpea is very low (around 1%) and it takes around 3 months to generate a full-grown T0 transgenic chickpea transformants. Moreover, the chickpea transformants further require micrografting for rooting (Sanyal et al., 2003, 2005). Also, chickpea do not have genetic mutant resources for functional genomics analysis. With these limitations, the generation of large scale chickpea mutant population and identification of mutated genes is a lengthy and laborious process. Functional validation of chickpea genes on the other hand can be done in heterologous plants like medicago (Medicago truncatula), Arabidopsis (Wardhan et al., 2016) and tobacco yet not all the genes can be studied in heterologous system because of the limitation with sequence homology. Overall, lack of reverse and forward genetics tools is a serious constraint for gene manipulation and thus functional validation in chickpea. In this preview, to overcome these hurdles, establishing virus-induced gene silencing (VIGS), an effective post-transcriptional gene silencing (PTGS)-mediated gene knockdown technique for the assessment of gene function (Burch-Smith et al., 2004; Robertson, 2004), in chickpea will be important. VIGS has been successfully implemented for gene silencing in legumes (Pflieger et al., 2013). VIGS technique is based on a natural defence mechanism employed by plants to fight against invading viruses (Voinnet, 2001), where virus entry into plant cell induces PTGS-mediated degradation of viral genome (Waterhouse et al., 2001). First step in VIGS-mediated silencing of candidate gene is the cloning of an efficient (in siRNA generation), off-target free target gene fragment (300–500 bp) (http://plantgrn.noble.org/pssRNAit/) into viral vector (Liu and Page, 2008; Senthil-Kumar and Mysore, 2014). Various VIGS vectors have been developed by cloning of viral genome into binary vectors (for Agrobacterium mediated inoculation)

or other non-binary expression vectors thus making them as infectious clones (for in vitro transcription mediated inoculation or direct plasmid inoculation for in vivo transcription) (Kumagai et al., 1995; Ruiz et al., 1998; Ratcliff et al., 2001; Liu et al., 2002a). In many of the VIGS vectors, genes responsible for disease symptoms and suppression of gene silencing were deleted from the native viral genome to facilitate better gene silencing, and restriction sites were inserted for easy gene cloning (Voinnet, 2001; Liu et al., 2002a,b). Second step, is the introduction of viral vector by either rub inoculation of in vitro transcribed RNA/plasmid DNA or by Agrobacterium tumefaciens-mediated inoculation of viral plasmids. Viral DNA or RNA initiates VIGS via synthesis of aberrant dsRNA. The dsRNAs are produced in plant cell after viral infection in the form of replicative intermediates by viral or plant RNA dependent RNA polymerase (RdRp) or by annealing of positive and negative strand of ssRNA to make replicative form. It can also be generated by folding of ssRNA or terminal dsRNA structures synthesized during transcription of viral RNA (Voinnet, 2005; Dalmay et al., 2000; Mourrain et al., 2000). Thus produced dsRNA structures are processed by DICER-like enzyme (DCL4) into 21to 25-nucleotides small interfering RNA (siRNA) (Deleris et al., 2006). The siRNAs are then incorporated into RNA-induced silencing complexes (RISC). RISC is a multiprotein complex with Argonaute (AGO1) and uses siRNA as template to find complementary target gene transcript for endonucleolytic cleavage (Fagard et al., 2000; Morel et al., 2002). VIGS is advantageous over other genome analysis tools as this is a rapid method to obtain loss-of-function phenotype. It does not require generation of transgenics and it can be implemented to target all the homologues of a gene at a time. VIGS can be used to characterize the gene’s function in multiple genetic backgrounds. It does not require complete gene sequence information and it can be implemented to study the function of genes which are otherwise lethal in stable RNAi transgenics or mutants. VIGS allows the high-throughput analysis of genes (Burch-Smith et al., 2004). However, VIGS protocol is not yet available for chickpea. In this chapter, we propose some of the possible strategies to develop VIGS protocol in chickpea to stimulate research in this direction.

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We suggest two ways to establish VIGS protocol in chickpea: (1) VIGS vectors widely used in other legumes can be exploited; and (2) we suggest developing disarmed VIGS vector from chickpea host viruses. VIGS vectors available for some leguminous plants VIGS have been successfully employed in legume plants, namely medicago (M. truncatula), shore medick (M. littoralis), common bean (Phaseolus vulgaris), pea (Pisum sativum), soybean (Glycine max), sweet pea (Lathyrus odorata), adzuki bean (Vigna angularis), cowpea (V. unguiculata), broad bean (Vicia faba) and Lotus japonicus. The VIGS vector developed from Cucumber mosaic virus (CMV; Nagamatsu et al., 2007), Bean pod mottle virus (BPMV; Zhang and Ghabrial, 2006), Apple latent spherical virus (ALSV; Igarashi et al., 2009), Pea early browning virus (PEBV; Constantin et al., 2004), Tobacco ringspot virus (TRSV; Zhao et al., 2016), White clover mosaic virus (WClMV; Ido et al., 2012), Sunnhemp mosaic virus (SHMV; Várallyay et al., 2010) and Soybean yellow common mosaic virus (SYCMV; Lim et al., 2016) have been extensively used for the gene silencing in legumes. These vectors could be tried for developing VIGS protocol in chickpea. CMV derived VIGS vector CMV is a tripartite, positive-sense, single-stranded RNA virus with widest host range, including chickpea (Palukaitis et al., 1992; Jones and Coutts, 1996). It belongs to genus Cucumovirus. RNA 1 and RNA 2 of CMV encodes for replication protein 1a and 2a proteins, respectively. RNA 2 also translates to silencing-suppressor protein (2b) which overlaps ORF 2a. RNA3 encodes movement protein (MP) and coat protein (CP) (Palukaitis and Garcia-Arenal, 2003). Suzuki and group (1991) cloned the full-length cDNAs of RNAs 1, 2, and 3 of CMV-Y strain into pUC118 vector and named them as pCY1, pCY2, and pCY3, respectively. Later, Otagaki and group (2006) modified the pCY2 for VIGS by deleting 3′ portion of ORF 2b and simultaneously introduced MluI and SnaBI restriction sites. They named modified CMV RNA2 (pCY2) vector as CMV2-A1 and tested the

silencing effect of CMV VIGS vectors in Nicotiana benthamiana. Later, Nagamatsu and group (2007) examined CMV mediated silencing in soybean by down-regulation of the chalcone synthase (CHS) genes involved in flavonoid biosynthesis. They inoculated in vitro transcripts of CMV vectors in N. benthamiana for viral multiplication and used the leaf sap for the inoculation into soybean. This vector is appropriate for silencing the genes in vegetative meristem tissues, pods and seed coats of legumes (Nagamatsu et al., 2007, 2009; Liu et al., 2010). Chickpea is a natural host plant for CMV ( Jones and Coutts, 1996) and thus this vector can directly be checked for its gene silencing effect in chickpea. Subterranean clover mottle virus (SCMoV)-derived VIGS vectors SCMoV is a single-stranded, positive-sense RNA virus from Sobemovirus genus. It infects clover and other legumes, including chickpea. It was found to infect 36 legume hosts in total, of which 22 were systemic hosts (Fosu-Nyarko et al., 2002). SCMoV contains 68 nt 5′ non-coding region (NCR) and 174 nt 3′ NCR and four overlapping ORFs. ORF1 encodes a putative protein which has similarity to suppressor of silencing. ORF2a encodes a putative protein with motif like serine protease. ORF2b contains GDD motif representative of RNA virus polymerases and ORF3 encodes CP (Dwyer et al., 2003). Fosu-Nyarko (2005) developed five constructs by fusing pFL transcription factors and Green fluorescent protein (GFP) gene from jelly fish to different parts of SCMoV genome. Bombardment of SCMoV vectors with gold particles was used to inoculate legumes (Fosu-Nyarko, 2005). In order to use this VIGS vector for silencing endogenous chickpea genes, GFP in the vector can be replaced with the candidate gene. Bean pod mottle virus (BPMV)derived VIGS vectors BPMV is a bipartite, positive-sense RNA virus from genus Comovirus. The RNA1 (≈ 6.0 kb) and RNA2 (≈ 3.6 kb) of BPMV are encapsidated in separate isometric particles. Both the RNAs are expressed as polyproteins. RNA1 polyprotein upon proteinase processing produces five mature replication related proteins. RNA2 polyprotein carries two CPs (L-CP

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and S-CP) and a cell-to-cell MP. The pathogenicity component for the severe foliar symptoms caused by BPMV infection is carried by RNA1 (Gu and Ghabrial, 2005). This virus was first isolated from common bean plant (Zaumeyer and Thomas, 1948) and is known to infect several other legume species (Brunt et al., 1996; Giesler et al., 2002). Zhang and Ghabrial (2006) developed the first BPMV VIGS vectors and later modified that into second and third generation VIGS vectors. The first generation VIGS vector was constructed by inserting a restriction site between MP and L-CP coding region (Zhang and Ghabrial, 2006). BPMV-VIGS was demonstrated in soybean (Zhang and Ghabrial, 2006) by rub-inoculation of in vitro transcripts of RNA1 and RNA2 containing phytoene desaturase (PDS) reporter gene on primary leaves. Photobleaching in soybean leaves was observed 2 weeks after inoculation (Zhang and Ghabrial, 2006). In the third generation vector, one more restriction site was introduced for cloning of candidate gene fragments after the RNA2 ORF translation stop codon (Zhang et al., 2010; 2013). The latest vectors have increased VIGS efficiency, less viral symptoms and suitable for high-throughput VIGS based functional studies (Zhang et al., 2009; 2010). These BPMV vectors are called ‘one-step’ viral vector because even plasmid DNA construct can be successfully infected into plant curtailing the need of inoculation techniques like Agrobacteriummediated transformation, in vitro transcription or biolistics (Pflieger et al., 2014). They were considered as appropriate for high-throughput silencing in soybean (Zhang et al., 2010; Pflieger et al., 2014). The BPMV VIGS vectors have been used to evaluate the function of genes involved in plant–microbe interactions (Kachroo et al., 2008; Fu et al., 2009; Zhang et al., 2009; Liu et al., 2011; Singh et al., 2011) and disease resistance (Meyer et al., 2009; Liu et al., 2012; Zhang et al., 2012). Apple latent spherical virus (ALSV)derived VIGS vector ALSV is a bipartite positive-sense single-stranded RNA virus of the genus Cheravirus. The two RNA molecules, RNA1 (≈ 6.8 kb) and RNA2 (≈ 4.4 kb), are enclosed in three distinct isometric particles. RNA1 contains a single ORF coding for a polyprotein for viral RNA polymerase, helicase and protease proteins. RNA2 encodes a polyprotein

for MP and three CPs named Vp25, Vp20, and Vp24. ALSV was initially isolated from apple trees (Koganezawa et al., 1985) that were symptomless. ALSV has a wide host range including Fabaceae. In Fabaceae, it infects pea, soybean, adzuki bean and cowpea and does not cause symptoms (Igarashi et al., 2009). Li and group (2004) constructed the infectious cDNA clones of ALSV by cloning the RNA1 and RNA2 between 35S promoter and NOS terminator in pE18PGT plasmid vector and named them as pEALSR1 and pEALSR2, respectively. They added Xho I, Sma I and BamHI restriction sites in RNA2 while creating a duplicate Q/G protease cleavage site for cloning. Plant gene fragment targeted for VIGS can be cloned in between the MP and Vp25 CP coding regions in RNA2 (pEALSR2) (Li et al., 2004). Before inoculation, the ALSV needs primary multiplication in Chenopodium quinoa leaves. The virus DNA (infectious clone) can be rubbed over C. quinoa leaves. The infected C. quinoa leaves can be harvested after 2–3 weeks and can be used for direct inoculation into host plant by rubbing (Igarashi et al., 2009). Silencing of PDS gene using this vector resulted in photo-bleaching (Igarashi et al., 2009) and silencing existed for more than a month. This VIGS in soybean was also shown to be inheritable to the next generation and the plant infected with ALSV-PDS showed highly uniform photo-bleaching symptoms in almost 33% of the seedlings in the next generation (Yamagishi and Yoshikawa, 2009). Pea early-browning virus (PEBV)derived VIGS vector PEBV is a bipartite, positive-sense, single-stranded RNA virus from Tobravirus genus. The two RNAs of PEBV are encapsidated in separate virus particles. RNA1 (≈ 6.8 kb) encodes the replicase and MP of the virus. RNA2 (≈ 4.5 kb, variable size in different isolates) encodes the CP and nematode transmission protein. It was first isolated from pea and later it has been found to infect various other legumes (Grønlund et al., 2008). RNA1 and RNA2 were cloned into binary vector pCAMBIA1300 under the regulation of 35S promoter and NOS terminator and named as pCAPE1 and pCAPE2 respectively (Constantin et al., 2004). GFP was then cloned downstream of the CP ORF translation stop codon in RNA2 by replacing the gene encoding for nematode transmission protein. PEBV VIGS vectors are

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amenable for Agrobacterium-mediated inoculation into plants. PEBV-mediated VIGS has been confirmed in pea by demonstrating PDS silencing and then further efficient silencing was demonstrated for UNIFOLIATA (UNI) and pea homologue of Arabidopsis KORRIGAN1 (KOR1). PEBV VIGS showed silencing of endogenous target genes in 9–10 days after inoculation (Constantin et al., 2004). PEBV has been employed to understand the functional genomics in various biological processes in different plant organs (Pflieger et al., 2013). Tobacco ringspot virus (TRSV)derived VIGS vector TRSV is a bipartite, single-stranded, positive-sense polyadenylated RNA virus from genus Nepovirus. Each TRSV RNA encodes a large polyprotein where RNA1 polyprotein is cleaved into proteinase cofactor, helicase, genome-linked protein, protease and RdRp. RNA2 polyprotein contains RNA replication protein, MP and CP (Zhao et al., 2016). It has a wide host range and causes bud blight disease in soybean (Demski et al., 1989). Zhao and group (2015) constructed two infectious clones, pTRSR1 and pTRSR2, by cloning RNA1 and RNA2 of TRSV, respectively, in binary vector pPZP211 in between the 35S promoter and NOS terminator. Later Zhao and group (2016) developed pT2C2A from pTRSR2 by inserting a gene encoding FMDV 2A catalytic peptide and BamHI-MIuI-SpeI restriction sites into the N-terminus of the CP. They checked TRSV-mediated silencing in soybean using PDS (GmPDS) gene. Photo-bleaching was visible at approximately 14 days after inoculation. The photo-bleaching was visible on complete leaves and axillary buds throughout plant development (Zhao et al., 2016). White clover mosaic virus (WClMV)derived VIGS vector WClMV is a monopartite, single-stranded, positive-sense RNA virus from the genus Potexvirus. It has been isolated from white clover, red clover, milk vetch (Astragalus sinicus) and pea plant (Iizuka and Iida,1965; Matsumoto et al., 1989; Tsuchizaki et al.,1981). The single RNA has five ORFs, where ORF1 encodes RdRp, ORFs 2–4 code for triple gene block (TGB protein, 26, 13, 7 kDa) and ORF 5 encodes CP proteins. Ido and group (2012) developed the infectious clone of WClMV-RC

strain by placing the cDNA clone of it in-between 35S promoter and 20 nucleotides of the poly(A) tail in plasmid vector pBluescript II Sk(–) background. The multiple cloning site was introduced downstream of the CP initiation site in the subgenomic RNA. The biolistic inoculation of this construct in broad bean cv. Kawachi-issun showed symptoms in upper leaf one week after inoculation. The sap from upper leaves of broad bean was then used as an inoculum for introducing viral particles in pea plants. Chlorosis, mosaic, and leaf deformation symptoms were observed in pea, 20 days after inoculation (Ido et al., 2012). PDS gene was successfully silenced in pea, faba bean, shore medick and medicago, but silencing was limited to areas of stems and upper leaves (Ido et al., 2012). The WClMV VIGS vector requires less time for the establishment of VIGS (8–10 days) than other vectors (Ido et al., 2012). Sunn-hemp mosaic virus (SHMV)derived VIGS vector SHMV is a single-stranded, positive-sense RNA virus of the genus Tobamovirus. It contains a 6-kblong RNA genome with 4 ORFs which encodes for replicase, MP and CP. Várallyay and group (2010) showed systemic infection of SHMV in medicago and cloned the virus genome into pUC18 under T7 RNA polymerase promoter for in vitro transcription. Further, they created two SHMV vectors for VIGS namely, SHMVProSHMV with a duplicate CP promoter downstream of the first CP promoter and SHMVProTOMV with ToMV CP subgenomic promoter in place of second SHMV CP promoter. They tested silencing effect of both the vectors in medicago by using CH42 (CHLI subunit of magnesium chelatase) gene fragment. Ten per cent of the medicago plants inoculated with SHMVProSHMVCH4240 showed yellowing on the emerging new leaves at 30 days after inoculation (Várallyay et al., 2010). In legumes, rub inoculation of this vector can be used to achieve silencing of gene of interest. Soybean yellow common mosaic virus (SYCMV)-derived VIGS vector SYCMV belongs to the genus Sobemovirus. It contains a single, positive-sense RNA genome. It has four putative ORFs. ORF 1 translates to a polyprotein which separates into MP and viral suppressor of RNA silencing (VSR). ORF 4

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encodes CP and ORF 2a and ORF 2b codes for polyproteins containing serine protease, viral protein genome-linked (VPg) and RdRp (Nam et al., 2012). SYCMV has a narrow host range. The complete cDNA of SYCMV genome was cloned into the pPZP211 binary vector between CaMV 35S promoter and an Rz sequence (self-cleaving ribozyme) and NOS terminator. A BsrGI site was introduced in the 3′ UTR region after stop codon of the CP for the cloning of gene fragment (Lim et al., 2016). PDS and RbcS genes were silenced using SYCMV vector in soybean. Three inoculation methods, such as Agrobacterium-mediated inoculation, particle bombardment and direct DNA-rubbing inoculation, were tried in soybean. All three methods showed persistent VIGS phenotypes in upper leaves starting 14 days after inoculation (Lim et al., 2016). The last seven viruses mentioned above have not been reported to infect chickpea, however, from our preliminary experiment results; these VIGS vectors may potentially infect chickpea upon artificial inoculation and can be candidate VIGS vectors. Selection of reporter genes Development of VIGS protocol in chickpea can be tried using above mentioned viral vectors. Selection of appropriate reporter or marker gene is important to study the silencing effect. The marker genes should facilitate the easy identification of silencing effects and additionally, the phenotype of marker gene should not be interfered by viral symptoms. Phytoene desaturase (PDS) and Mg-protoporphyrin chelatase (ChlH) are widely used marker genes. PDS enzyme converts phytoene to coloured carotene in carotenoid synthesis pathway (Bartley and Scolnik, 1995). This is an important rate-limiting enzyme and thus, its silencing is expected to produce photo-bleaching phenotype in leaves (Kumagai et al., 1995, Wang et al., 2009). ChlH is an important enzyme in chlorophyll biosynthesis. It catalyses the addition of magnesium ion to protoporphyrin IX to produce Mg-protoporphyrin IX and thus, its silencing is known to produce yellow leaf phenotype (Hiriart et al., 2002). Apart from these genes, actin and ubiquitin can be also be used for the protocol establishment. Silencing of the actin gene results in severely stunted shoot and roots (Ryu et al., 2004). Chalcone synthase is yet another marker gene to

study silencing in flowers and pods (Nagamatsu et al., 2007). Selection of appropriate gene fragment for cloning in VIGS vector A gene fragment of 200–400 nucleotides from the coding region or UTR with fewer off-target effects is recommended for effective gene silencing (Liu and Page, 2008; Senthil-Kumar and Mysore, 2014). The reporter gene selected for the VIGS establishment should be analysed for the efficient siRNA production and minimum probable offtarget effect using pssRNAit (http://plantgrn. noble.org/pssRNAit/). A fragment of target gene having maximum number of target gene specific siRNA with higher efficiency and minimum number of siRNA with off-target are suitable for establishing VIGS. SGN-VIGS tools (http://vigs. solgenomics.net/) can also be employed using M. truncatula database reference to scan chickpea candidate genes. SGN VIGS tool simulates VIGS process in-silico to predict the best gene fragment for silencing. It processes target genes in all possible 21 nt fragment or in the size of siRNA selected by user. Further, this tool maps virtual siRNA onto all genes of the selected transcriptome by Bowtie (ultrafast, memory-efficient short read aligner). Finally, SGN VIGS tool provides a query sequence (300 nt long or length selected by user) with minimum match to off targets and with maximum coverage for the target gene. It displays all the alignments with targets and off-targets with score values and also suggests best possible construct (Fernandez-Pozo et al., 2015). The selected fragment can then be cloned into VIGS vectors. Some of the previous studies suggest that cloning of gene fragment in inverted repeat (Lacomme et al., 2003) or antisense direction in VIGS vectors results in more efficient silencing (Lacomme et al., 2003; Zhang et al., 2010). Suggestions for virus inoculation methods Agrobacterium infiltration, leaf sap inoculation or rub inoculation of DNA or transcripts have been so far used as methods for VIGS vector delivery into plants. Below we are describing suitable inoculation

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methods for VIGS vectors mentioned in this chapter. Agrobacterium-mediated VIGS vector inoculation PEBV and SYCMV based VIGS vectors are suitable for the Agrobacterium-mediated inoculation. For the Agrobacterium-mediated inoculation, Agrobacterium-containing VIGS vector plasmids can be vacuum infiltrated (Wang et al., 2006) into the chickpea leaves (Fig. 15.1A). Leaf sap inoculation ALSV, TRSV and WClMV vectors need primary multiplication in intermediate host C. quinoa, N. benthamiana and faba beans respectively. Before rub inoculation of sap from intermediate host to chickpea, evaluation of the viral load in the intermediate host is necessary. The viral load can be tested by immunocapture RT-PCR (Nolasco et al., 1993).

Alternatively, viral load can also be evaluated by leaf lesion assay in C. amaranticolor (Senthil-Kumar and Mysore, 2014). Leaves exhibiting abundant viral load from intermediate host can then be utilized for sap inoculation. The Agrobacterium containing TRSV vector plasmids with chickpea marker gene (pTRSR1 and pT2C2A; Zhao et al., 2016) need to be agro-infiltrated into N. benthamiana leaves by syringe infiltration of Agrobacterium culture mix for primary multiplication. The N. benthamiana leaves after 6–7 days of infiltration can be crushed and sap (Zhao et al., 2016) can be applied to chickpea leaves. ALSV VIGS vector plasmids (pEALSR1 and pEALSR2L5R5, Li et al., 2004; Igarashi et al., 2009) need two times multiplication in intermediate host C. quinoa leaves (Li et al., 2004; Igarashi et al., 2009). The sap from secondary infection can be rubbed over chickpea leaves. WCIMV vector plasmid can be first inoculated into intermediate host faba bean leaves by particle bombardment and

Figure 15.1 Proposed methods for inoculation of VIGS vectors in chickpea leaf. (A) Vacuum infiltration of Agrobacterium tumefaciens harbouring VIGS binary plasmid into chickpea leaves. First step in vacuum infiltration is to invert the chickpea plant in A. tumefaciens solution. A vacuum pressure of 60 Pa should to be then applied for 10 mins for infiltration of Agrobacterium in chickpea. (B–D) Rub-inoculation of plasmid DNA/in vitro transcripts/leaf sap over chickpea leaves. Carborundum should be dusted over chickpea leaflets before application of VIGS plasmid/in vitro transcript or leaf sap onto chickpea leaflet. DNA/RNA or leaf sap should be gently rubbed over chickpea leaflet.

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one week after inoculation, sap (Ido et al., 2012) can be rub inoculated onto chickpea leaves (Fig. 15.1B–D). Alternatively, the virion particles can be enriched from the intermediate host by precipitation in polyethylene glycol and can be rub-inoculated to chickpea leaves (Lu et al., 2003). This will enable direct entry of virus particle with high titre in chickpea leaves. Direct rub inoculation of plasmid DNA/in vitro transcripts BPMV vector has been developed with the ease of one step inoculation (Pflieger et al., 2014). The BPMV vector plasmids harbouring chickpea marker gene can be rubbed directly over the chickpea leaves to induce gene silencing. Similarly, SYCMV vector plasmid can also be directly rubbed (Lim et al., 2016) over chickpea leaves. SHMV and CMV based vectors are driven under T7 promoter so that viral RNA with cloned gene fragment can be generated by in vitro transcription. The resultant transcripts can be rub inoculated (Várallyay et al., 2010; Nagamatsu et al., 2007) over chickpea leaves to test successful infection (Fig. 15.1B–D). Assessment of gene silencing Before assessing if the gene silencing has happened, confirmation of viral infection in chickpea is necessary. Many VIGS vectors mentioned in this chapter shows silencing in legumes 2–3 weeks after virus inoculation, except WClMV which shows early silencing (8–10 days). Therefore, presence of virus should be assessed around 2 weeks after inoculation. The viral titre in chickpea can be assessed by quantitative RT-PCR using primers specific to viral genes. Next, it is important to know whether the virus infection is systemic. This can be studied by assessing presence of virus in the systemic upper newly developed noninoculated leaves by RT-PCR. To assess the titre of the viral particle, immunocapture RT-PCR and leaf lesion assays on C. amaranticolor should be performed (Nolasco et al., 1993; Senthil-Kumar and Mysore, 2014). Leaf lesion assay involves the detection of visible lesions on C. amaranticolor leaves upon viral infection as a result of plant response (Senthil-Kumar and Mysore, 2011;

2014). Often, deletion of gene insert from viral vector is reported during virus multiplication and systemic movement in plants (Guo et al., 1998; Bruun-Rasmussen et al., 2007) therefore it is necessary to confirm that insert is intact with viral genome in systemically infected leaves. This can be assessed by semi-quantitative RT-PCR with primers flanking candidate gene. The detection of siRNA generated against marker gene would confirm the VIGS (Kauppinen and Havelda, 2008). Next, the silencing of viral transcripts should be confirmed by estimating the decrease in level of viral genome by the RT-PCR analysis using primers specific to viral genome. Similarly, endogenous target marker gene silencing can also be confirmed by RT-PCR using gene specific primers designed for region outside of gene fragment cloned in the VIGS vector. The systemic silencing effect can be assessed at phenotypic levels. The systemic silencing of PDS and ChlH can be known by presence of photo-bleaching and yellowing respectively in the non-infected leaves (Kumagai et al., 1995; Hiriart et al., 2002). PDS silencing can be confirmed by biochemical evaluation of carotenoids. The silencing of actin and ubiquitin genes can be assessed by the shoot and root length (Ryu et al., 2004) compared to plants inoculated with empty VIGS vector (VIGS vector without candidate gene). Development of VIGS vectors from chickpea host viruses Construction of VIGS vector from the chickpea host viruses can be the best strategy to establish efficient and high-throughput gene silencing in chickpea. Chickpea is host for Alfalfa mosaic virus (AMV, Bromoviridae), Bean yellow mosaic virus (BYMV, Potyviridae), Pea seed-borne mosaic virus (PSbMV, Potyviridae), Beet western yellows virus (BWYV, Luteoviridae), Chickpea chlorotic dwarf virus (CpCDV) (genus Mastrevirus, family Geminiviridae), Lettuce necrotic yellows virus (LNYV, Rhabdoviridae), Bean (pea) leaf roll virus (BLRV, Luteoviridae) and Subterranean clover red leaf (SCRLV, Luteoviridae) (see complete list in Table 15.1). These wide range of viruses representing various genera provide scope for developing suitable VIGS vectors. Earlier, Nakahara and group (2015) developed the infectious cDNA clone of BYMV for gene expression and assessed its

Establishment of VIGS in Chickpea |  337

Table 15.1 List of viruses reported to infect chickpea No. Virus name

Virus family

Symptom description

Virus genome

Reference

RNA viruses 1

Alfalfa mosaic Bromoviridae virus

Necrosis of shoot tips

Jones and Coutts ssRNA, positive (1996) strand Multipartite (four particles: three bacilliform and 1 spheroidal)

2

Cucumber mosaic virus

Bromoviridae

Reddening and chlorosis of leaf and plant stunting

ssRNA, tripartite

3

Bean yellow mosaic virus

Potyviridae

Apical necrosis, reddening and plant stunting

ssRNA, monopartite Chalam (1982)

4

Potyviridae Pea seedborne mosaic virus

Brown ring patterns and spots on the seeds and plant stunting

ssRNA

Makkouk et al. (1993)

5

Lettuce mosaic virus

Potyviridae

Yellowing and leaf tip wilting

ssRNA positivestrand

Bosque-Perez et al. (1989)

6

Beet western yellows virus

Luteoviridae

Yellowing, rolling and stiffening of leaves, and plant stunting

ssRNA, monopartite Bosque-Perez and Buddenhagen (1990)

7

Bean leaf roll virus

Luteoviridae

Interveinal chlorosis, yellowing and rolling of leaf and plant stunting

ssRNA, positive Ashby (1984) strand, monopartite

8

Subterranean Luteoviridae clover red leaf

Stunted plant

ssRNA, positive Bosque-Perez and strand, monopartite Buddenhagen (1990)

9

Pea enation mosaic virus

Luteoviridae

Twisted and malformed leaves, severe plant stunting, poor pod filling and premature plant death

ssRNA positivestrand

Makkouk et al. (2003a)

10

Phasey bean mild yellows virus

Luteoviridae

Mild leaf yellowing

ssRNA positivestrand

Sharman et al. (2016)

11

Chickpea chlorotic stunt virus

Luteoviridae

Slight plant stunting and yellowing, andveinalchlorosis

ssRNA positivestrand

Abraham et al. (2006)

12

Red clover vein mosaic virus

Betaflexiviridae Severe plant stunting, mosaic, proliferation of axillary buds, malformation of leaves and branches

ssRNA positivestrand

Larsen and Miklas (2001)

13

Lettuce necrotic yellows virus

Rhabdoviridae

14

Subterranean Sobemovirus clover mottle virus

Bleached, necrotic tip burn. reddish-brown necrosis of stem,chlorosis and death of leaflet and premature plant death

Jones and Coutts (1996)

ssRNA, monopartite Behncken (1983)

Leaf mottling and distortion and ssRNA, monopartite Fosu-Nyarko et al. plant stunting (2002)

DNA viruses 15

Chickpea chlorosis virus

Geminiviridae

Plant stunting, chlorosis or reddening of leaves

ssDNA

Thomas et al. (2010)

16

Chickpea redleaf virus

Geminiviridae

Plant stunting, chlorosis or reddening of leaves

ssDNA

Thomas et al. (2010)

17

Chickpea yellow dwarf virus

Geminiviridae

Plant stunting, chlorosis or reddening of leaves

ssDNA

Kraberger et al. (2015)

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Table 15.1 Continued No. Virus name

Virus family

Symptom description

Virus genome

Reference

18

Chickpea yellows virus

Geminiviridae

Plant stunting, chlorosis or reddening of leaves

ssDNA

Hadfield et al. (2012)

19

Chickpea chlorotic dwarf virus

Geminiviridae

Yellowing, chlorosis or reddening of leaves and plant stunting

ssDNA

Horn et al. (1993)

20

Tobacco yellow dwarf virus

Geminiviridae

Plant stunting, chlorosis or reddening of leaves

ssDNA

Thomas et al. (2010)

21

Faba bean necrotic yellows virus

Nanoviridae

Leaf rolling, yellowing and plant stunting

ssDNA

Makkouk et al. (2003b)

Note: list and details compiled from cited references and other resources, namely virus–host database (www. genome.jp/virushostdb/12472), description of plant viruses (www.dpvweb.net/dpv/showdpv.php?dpvno=387) and international committee on taxonomy of viruses (ICTV) (https://talk.ictvonline.org/).

multiplication in bean and pea. This virus has potential for modification as VIGS vector for chickpea. Selection of suitable chickpea genotype A suitable host, apart from efficient VIGS vector, is needed for effective VIGS. Efficient host should have strong PTGS-mediated defence that lead to prolific silencing, yet allow systemic movement of virus to new tissues. This will lead to long duration and systemic silencing. Host must not show severe viral infection symptoms which can not only mask the silencing symptoms but also negatively impact physiology of the plant. VIGS vector should induce silencing throughout the developmental stages (Lacomme, 2015). Therefore, various genotypes of the chickpea need to be evaluated for first viral susceptibility or tolerance and host with the moderate susceptibility would be best to use for VIGS establishment. For instance, Kanakala and group (2013) studied the susceptible and resistance genotype of chickpea for Chickpea chlorotic dwarf virus (CpCDV) by Agrobacterium-mediated delivery of the viral genome. They found genotype GOURI1499, ICC-8923 and RSG-931 were resistance to CpCDV infection whereas genotypes BG-1053, BGD-112 and BG-3003 were susceptible. Genotypes DCP 92–3, ICCV-06302, JG-130 and JG-6 were moderately susceptible (Kanakala et al., 2013) and therefore, these moderately susceptible genotypes would be helpful to study the silencing

by CpCDV vectors. Similar approaches can be employed for screening chickpea genotypes and identify susceptible genotypes to other chickpea host viruses (Martin et al., 1999; Kanakala et al., 2013). Harries and group (2008) demonstrated that mild expression of viral silencing suppressor 126kDa protein increases the susceptibility of tobacco (N. tabacum) to many otherwise resistant viruses and at the same time also increases the VIGS-mediated silencing. Kheyr-Pour and group (1994) could achieve the Tomato yellow leaf curl virus (TYLCV) infection in wild tomato species by agroinoculation of tandem repeat of TYLCV genome. Similar strategies can also be tried to increase the viral infection in resistant chickpea genotypes. Future prospects Gene function characterization and understanding its biological mechanism is not currently feasible in chickpea owing to non-availability of functional genomics and gene manipulation tools and related resources. Currently, VIGS protocol is not available in chickpea. Moreover not much information is available on molecular mechanism of virus–chickpea interaction. In this chapter we explained some of the possible steps that can be tried to establish VIGS protocol in chickpea. We have also provided information about some of the native viruses infecting chickpea. The future strategies to develop VIGS in chickpea should include the following: (1) Selection of suitable virus infecting chickpea

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from natural host range. 2) Understanding the symptomatology of naturally infecting viruses and development of infectious clone. Preparation of disarmed virus that cause mild or no symptoms but with systemic infection. 3) Selection of appropriate chickpea genotype that produce systemic infection with very mild viral symptoms. Apart from these strategies, suitability of currently available legume infecting VIGS vectors can be tested. Acknowledgements Projects at MS-K lab are supported by National Institute of Plant Genome Research (NIPGR) core funding. We thank Science and Engineering Research Board (SERB) India, for providing grant and fellowship (SB/YS/LS-237/2013) to RS. We thank Drs. Bendangchuchang Longchar and Aarti Gupta for critically reading the manuscript. We thank DBT e-Library Consortium (DeLCON) for providing access to e-resources. References

Abraham, A.D., Menzel, W., Lesemann, D.E., Varrelmann, M., and Vetten, H.J. (2006). Chickpea chlorotic stunt virus: a new polerovirus infecting cool-season food legumes in Ethiopia. Phytopathology 96, 437–446. https://doi.org/10.1094/PHYTO-96-0437 Agarwal, G., Jhanwar, S., Priya, P., Singh, V.K., Saxena, M.S., Parida, S.K., Garg, R., Tyagi, A.K., and Jain, M. (2012). Comparative analysis of kabuli chickpea transcriptome with desi and wild chickpea provides a rich resource for development of functional markers. PLOS ONE 7, e52443. https://doi.org/10.1371/journal. pone.0052443 Ashby, J.W. (1984). Bean leaf roll virus. In: Descriptions of plant viruses. CMI/AAB Descr. PI. Viruses. 286, 4. Bartley, G.E., and Scolnik, P.A. (1995). Plant carotenoids: pigments for photoprotection, visual attraction, and human health. Plant Cell 7, 1027–1038. https://doi. org/10.1105/tpc.7.7.1027 Behncken, G.M. (1983). A disease of chickpea caused by lettuce necrotic yellows virus. Australasian Plant Pathol. 12, 64–65. Bhatia, V.S., Singh, P., Wani, S.P., Kesava Rao, A.V.R., and Srinivas, K. (2006). Yield gap analysis of soybean, groundnut, pigeonpea and chickpea in India using simulation modeling. Global Theme on Agroecosystems Report no. 31. Andhra Pradesh, India: International Crops Research Institute for the Semi-Arid Tropics (ICRISAT). 156. Bosque-Perez, N.A., and Buddenhagen, I.W. (1989). First report of lettuce mosaic virus on chickpea. Plant Dis. 73. Bosque-Perez, N.A., and Buddenhagen, I.W. (1990). Studies on epidemiology of virus disease of chickpea in California. Plant Dis. 74, 372–378. Brunt, A.A., Crabtree, K., Dallwitz, M.J., Gibbs, A.J., Watson, L., and Zurcher, E.J. (1996). Plant viruses

online: descriptions and lists from the VIDE database. ‘Available at http://biology.anu.edu.au/Groups/MES/ vide/. Bruun-Rasmussen, M., Madsen, C.T., Jessing, S., and Albrechtsen, M. (2007). Stability of Barley stripe mosaic virus-induced gene silencing in barley. Mol. Plant Microbe Interact. 20, 1323–1331. https://doi. org/10.1094/MPMI-20-11-1323 Burch-Smith, T.M., Anderson, J.C., Martin, G.B., and Dinesh-Kumar, S.P. (2004). Applications and advantages of virus-induced gene silencing for gene function studies in plants. Plant J. 39, 734–746. https:// doi.org/10.1111/j.1365-313X.2004.02158.x Chalam, V. (1982). Identification and characterization of Cucumber Mosaic and Bean yellow mosaic viruses affecting chickpea (Cicer arietinum L.) in India. Ph. D Thesis, Andhra Pradesh Agricultural University, Rajendranagar, Hyderabad, 1–92. Constantin, G.D., Krath, B.N., MacFarlane, S.A., Nicolaisen, M., Johansen, I.E., and Lund, O.S. (2004). Virusinduced gene silencing as a tool for functional genomics in a legume species. Plant J. 40, 622–631. Dalmay, T., Hamilton, A., Rudd, S., Angell, S., and Baulcombe, D.C. (2000). An RNA-dependent RNA polymerase gene in Arabidopsis is required for posttranscriptional gene silencing mediated by a transgene but not by a virus. Cell 101, 543–553. Deleris, A., Gallego-Bartolome, J., Bao, J., Kasschau, K.D., Carrington, J.C., and Voinnet, O. (2006). Hierarchical action and inhibition of plant Dicer-like proteins in antiviral defense. Science 313, 68–71. Demski, J.W., and Kuhn, C.W. (1989). Tobacco ringspot virus. Hartman G.L. (Ed.), Compendium of Soybean Diseases (3rd edn.), Am. Phytopath. Soc., St. Paul, MN, pp. 57–59. Deokar, A.A., Ramsay, L., Sharpe, A.G., Diapari, M., Sindhu, A., Bett, K., Warkentin, T.D., and Tar’an, B. (2014). Genome wide SNP identification in chickpea for use in development of a high density genetic map and improvement of chickpea reference genome assembly. BMC Genomics 15, 708. https://doi. org/10.1186/1471-2164-15-708 Diapari, M., Sindhu, A., Bett, K., Deokar, A., Warkentin, T.D., and Tar’an, B. (2014). Genetic diversity and association mapping of iron and zinc concentrations in chickpea (Cicer arietinum L.). Genome 57, 459–468. https://doi.org/10.1139/gen-2014-0108 Dwyer, G.I., Njeru, R., Williamson, S., Fosu-Nyarko, J., Hopkins, R., Jones, R.A., Waterhouse, P.M., and Jones, M.G. (2003). The complete nucleotide sequence of Subterranean clover mottle virus. Arch. Virol. 148, 2237– 2247. https://doi.org/10.1007/s00705-003-0144-3 Fagard, M., Boutet, S., Morel, J.B., Bellini, C., and Vaucheret, H. (2000). AGO1, QDE-2, and RDE-1 are related proteins required for post-transcriptional gene silencing in plants, quelling in fungi, and RNA interference in animals. Proc. Natl. Acad. Sci. U.S.A. 97, 11650–11654. https://doi.org/10.1073/pnas.200217597 Farooq, M., Hussain, M., Nawaz, A., Lee, D.J., Alghamdi, S.S., and Siddique, K.H.M. (2017). Seed priming improves chilling tolerance in chickpea by modulating germination metabolism, trehalose accumulation and

340  | Sinha and Senthil-Kumar

carbon assimilation. Plant Physiol. Biochem. 111, 274–283. Fernandez-Pozo, N., Rosli, H.G., Martin, G.B., and Mueller, L.A. (2015). The SGN VIGS tool: user-friendly software to design virus-induced gene silencing (VIGS) constructs for functional genomics. Mol Plant 8, 486– 488. https://doi.org/10.1016/j.molp.2014.11.024 Fosu-Nyarko, J. (2005). Studies on Subterranean clover mottle virus towards development of a gene silencing vector. PhD thesis, Murdoch University). Fosu-Nyarko, J., Jones, R.A.C., Smith, L.J., Jones, M.G.K., and Dwyer, G.I. (2002). Host range and symptomatology of Subterranean clover mottle virus in alternative pasture, forage and crop legumes. Australasian Plant Pathol. 31, 345–350. Fu, D.Q., Ghabrial, S., and Kachroo, A. (2009). GmRAR1 and GmSGT1 are required for basal, R gene-mediated and systemic acquired resistance in soybean. Mol. Plant Microbe Interact. 22, 86–95. https://doi.org/10.1094/ MPMI-22-1-0086 Garg, R., Patel, R.K., Tyagi, A.K., and Jain, M. (2011). De novo assembly of chickpea transcriptome using short reads for gene discovery and marker identification. DNA Res. 18, 53–63. https://doi.org/10.1093/dnares/ dsq028 Garg, R., Shankar, R., Thakkar, B., Kudapa, H., Krishnamurthy, L., Mantri, N., Varshney, R.K., Bhatia, S., and Jain, M. (2016). Transcriptome analyses reveal genotype- and developmental stage-specific molecular responses to drought and salinity stresses in chickpea. Sci Rep 6, 19228. https://doi.org/10.1038/srep19228 Gaur, P.M., Tripathi, S., Gowda, C.L.L., Ranga Rao, G.V., Sharma, H.C., Pande, S., and Sharma, M. (2010). Chickpea seed production manual. ICRISAT. 502, 1–28. Gaur, R., Jeena, G., Shah, N., Gupta, S., Pradhan, S., Tyagi, A.K., Jain, M., Chattopadhyay, D., and Bhatia, S. (2015). High density linkage mapping of genomic and transcriptomic SNPs for synteny analysis and anchoring the genome sequence of chickpea. Sci Rep 5, 13387. https://doi.org/10.1038/srep13387 Giesler, L.J., Ghabrial, S.A., Hunt, T.E., and Hill, J.H. (2002). Bean pod mottle virus: a threat to US soybean production. Plant Dis. 86, 1280–1289. Grønlund, M., Constantin, G., Piednoir, E., Kovacev, J., Johansen, I.E., and Lund, O.S. (2008). Virus-induced gene silencing in Medicago truncatula and Lathyrus odorata. Virus Res. 135, 345–349. https://doi. org/10.1016/j.virusres.2008.04.005 Gu, H., and Ghabrial, S.A. (2005). The Bean pod mottle virus proteinase cofactor and putative helicase are symptom severity determinants. Virology 333, 271–283. Guo, H.S., López-Moya, J.J., and García, J.A. (1998). Susceptibility to recombination rearrangements of a chimeric plum pox potyvirus genome after insertion of a foreign gene. Virus Res. 57, 183–195. Gupta, S., Nawaz, K., Parween, S., Roy, R., Sahu, K., Kumar Pole, A., Khandal, H., Srivastava, R., Kumar Parida, S., and Chattopadhyay, D. (2017). Draft genome sequence of Cicer reticulatum L., the wild progenitor of chickpea provides a resource for agronomic trait improvement. DNA Res. 24, 1–10. https://doi.org/10.1093/dnares/ dsw042

Hadfield, J., Thomas, J.E., Schwinghamer, M.W., Kraberger, S., Stainton, D., Dayaram, A., Parry, J.N., Pande, D., Martin, D.P., and Varsani, A. (2012). Molecular characterisation of dicot-infecting Mastreviruses from Australia. Virus Res. 166, 13–22. https://doi. org/10.1016/j.virusres.2012.02.024 Harries, P.A., Palanichelvam, K., Bhat, S., and Nelson, R.S. (2008). Tobacco mosaic virus 126-kDa protein increases the susceptibility of Nicotiana tabacum to other viruses and its dosage affects virus-induced gene silencing. Mol. Plant Microbe Interact. 21, 1539–1548. https://doi. org/10.1094/MPMI-21-12-1539 Hiremath, P.J., Farmer, A., Cannon, S.B., Woodward, J., Kudapa, H., Tuteja, R., Kumar, A., Bhanuprakash, A., Mulaosmanovic, B., Gujaria, N., et al. (2011). Largescale transcriptome analysis in chickpea (Cicer arietinum L.), an orphan legume crop of the semi-arid tropics of Asia and Africa. Plant Biotechnol. J. 9, 922–931. https:// doi.org/10.1111/j.1467-7652.2011.00625.x Hiriart, J.B., Lehto, K., Tyystjärvi, E., Junttila, T., and Aro, E.M. (2002). Suppression of a key gene involved in chlorophyll biosynthesis by means of virus-inducing gene silencing. Plant Mol. Biol. 50, 213–224. Horn, N.M., Reddy, S.V., Roberts, I.M., and Reddy and D.V.R. (1993). Chickpea chlorotic dwarf virus, a new leafhopper‐transmitted geminivirus of chickpea in India. Ann. Appl. Biol. 122, 467–479. Ido, Y., Nakahara, K.S., and Uyeda, I. (2012). White clover mosaic virus-induced gene silencing in pea. J. Gen. Plant Pathol. 78, 127–132. Igarashi, A., Yamagata, K., Sugai, T., Takahashi, Y., Sugawara, E., Tamura, A., Yaegashi, H., Yamagishi, N., Takahashi, T., Isogai, M., et al. (2009). Apple latent spherical virus vectors for reliable and effective virus-induced gene silencing among a broad range of plants including tobacco, tomato, Arabidopsis thaliana, cucurbits, and legumes. Virology 386, 407–416. https://doi. org/10.1016/j.virol.2009.01.039 Iizuka, N., and Iida, W. (1965). Studies on the white clover mosaic virus. Ann. Phytopathol. Soc. Japan. 30, 46–53. Jain, M., Misra, G., Patel, R.K., Priya, P., Jhanwar, S., Khan, A.W., Shah, N., Singh, V.K., Garg, R., Jeena, G., et al. (2013). A draft genome sequence of the pulse crop chickpea (Cicer arietinum L.). Plant J. 74, 715–729. https://doi.org/10.1111/tpj.12173 Jhanwar, S., Priya, P., Garg, R., Parida, S.K., Tyagi, A.K., and Jain, M. (2012). Transcriptome sequencing of wild chickpea as a rich resource for marker development. Plant Biotechnol. J. 10, 690–702. https://doi. org/10.1111/j.1467-7652.2012.00712.x Jones, R.A.C., and Coutts, B.A. (1996). Alfalfa mosaic and Cucumber mosaic virus infection in chickpea and lentil: incidence and seed transmission. Ann. Appl. Biol. 129, 491–506. Kachroo, A., Fu, D.Q., Havens, W., Navarre, D., Kachroo, P., and Ghabrial, S.A. (2008). An oleic acid-mediated pathway induces constitutive defense signaling and enhanced resistance to multiple pathogens in soybean. Mol. Plant Microbe Interact. 21, 564–575. https://doi. org/10.1094/MPMI-21-5-0564 Kanakala, S., Verma, H.N., Vijay, P., Saxena, D.R., and Malathi, V.G. (2013). Response of chickpea genotypes

Establishment of VIGS in Chickpea |  341

to Agrobacterium-mediated delivery of Chickpea chlorotic dwarf virus (CpCDV) genome and identification of resistance source. Appl. Microbiol. Biotechnol. 97, 9491–9501. https://doi.org/10.1007/s00253-0135162-9 Kauppinen, S., and Havelda, Z. (2008). Detection of siRNAs and miRNAs. Plant Virology Protocols: From Viral Sequence to Protein Function. pp. 217–227. Kheyr-Pour, A., Gronenborn, B., and Czosner, H. (1994). Agroinoculation of Tomato yellow leaf curl virus (TYLCV) overcomes the virus resistance of wild Lycopersicon species. Plant Breed. 112, 228–233. Koganezawa, H., Yanase, H., Ochiai, M., and Sakuma, T. (1985). An isometric virus-like particle isolated from russet ring-diseased apple. Ann. Phytopathol. Soc. Japan. 51, 363. Kraberger, S., Mumtaz, H., Claverie, S., Martin, D.P., Briddon, R.W., and Varsani, A. (2015). Identification of an Australian-like dicot-infecting Mastrevirus in Pakistan. Arch. Virol. 160, 825–830. https://doi.org/10.1007/ s00705-014-2299-5 Kumagai, M.H., Donson, J., della-Cioppa, G., Harvey, D., Hanley, K., and Grill, L.K. (1995). Cytoplasmic inhibition of carotenoid biosynthesis with virus-derived RNA. Proc. Natl. Acad. Sci. U.S.A. 92, 1679–1683. Lacomme, C., Hrubikova, K., and Hein, I. (2003). Enhancement of virus-induced gene silencing through viral-based production of inverted-repeats. Plant J. 34, 543–553. Lacomme, C. (2015). Strategies for altering plant traits using virus-induced gene silencing technologies. Plant Gene Silencing: Methods and Protocols. pp. 25–41. Larsen, R.C., and Miklas, P.N. (2001). Effect of red clover vein mosaic carlavirus infection on seed production and biomass yield in chickpea. Phytopathology 91, 186. Li, C., Sasaki, N., Isogai, M., and Yoshikawa, N. (2004). Stable expression of foreign proteins in herbaceous and apple plants using Apple latent spherical virus RNA2 vectors. Arch. Virol. 149, 1541–1558. https://doi. org/10.1007/s00705-004-0310-2 Lim, S., Nam, M., Kim, K.H., Lee, S.H., Moon, J.K., Lim, H.S., Choung, M.G., Kim, S.M., and Moon, J.S. (2016). Development of a new vector using Soybean yellow common mosaic virus for gene function study or heterologous protein expression in soybeans. J. Virol. Methods 228, 1–9. https://doi.org/10.1016/j. jviromet.2015.11.005 Liu, B., Watanabe, S., Uchiyama, T., Kong, F., Kanazawa, A., Xia, Z., Nagamatsu, A., Arai, M., Yamada, T., Kitamura, K., et al. (2010). The soybean stem growth habit gene Dt1 is an ortholog of Arabidopsis TERMINAL FLOWER1. Plant Physiol. 153, 198–210. https://doi. org/10.1104/pp.109.150607 Liu, E., and Page, J.E. (2008). Optimized cDNA libraries for virus-induced gene silencing (VIGS) using Tobacco rattle virus. Plant Methods 4, 5. https://doi. org/10.1186/1746-4811-4-5 Liu, H., Reavy, B., Swanson, M., and MacFarlane, S.A. (2002a). Functional replacement of the Tobacco rattle virus cysteine-rich protein by pathogenicity proteins from unrelated plant viruses. Virology 298, 232–239. Liu, Y., Schiff, M., and Dinesh-Kumar, S.P. (2002b). Virusinduced gene silencing in tomato. Plant J. 31, 777–786.

Liu, J.Z., Horstman, H.D., Braun, E., Graham, M.A., Zhang, C., Navarre, D., Qiu, W.L., Lee, Y., Nettleton, D., Hill, J.H., et al. (2011). Soybean homologs of MPK4 negatively regulate defense responses and positively regulate growth and development. Plant Physiol. 157, 1363–1378. https://doi.org/10.1104/pp.111.185686 Liu, S., Kandoth, P.K., Warren, S.D., Yeckel, G., Heinz, R., Alden, J., Yang, C., Jamai, A., El-Mellouki, T., Juvale, P.S., et al. (2012). A soybean cyst nematode resistance gene points to a new mechanism of plant resistance to pathogens. Nature 492, 256–260. https://doi. org/10.1038/nature11651 Lu, R., Martin-Hernandez, A.M., Peart, J.R., Malcuit, I., and Baulcombe, D.C. (2003). Virus-induced gene silencing in plants. Methods 30, 296–303. Makkouk, K.M., Kumari, S.G., and Bos, L. (1993). Pea seedborne mosaic virus: occurrence in Faba bean (Vicia faba) and lentil (Lensculinaris) in West Asia and North Africa and further information on host range, transmission characteristics and purification. Netherlands J. Plant Pathol. 99, 115–124. Makkouk, K.M., Kumari, S.G., Shahraeen, N., Fazlali, Y., Farzadfar, S., Ghotbi, T., and Mansouri, A.R. (2003a). Identification and seasonal variation of viral diseases of chickpea and lentil in Iran. J. Plant Dis. Protection. 110, 157–169. Makkouk, K.M., Hamed, A.A., Hussein, M., and Kumari, S.G. (2003b). First report of Faba bean necrotic yellows virus (FBNYV) infecting chickpea (Cicerarietinum) and Faba bean (Viciafaba) crops in Sudan. Plant Pathol. 52, 412–412. Martin, D.P., Willment, J.A., and Rybicki, E.P. (1999). Evaluation of maize streak virus pathogenicity in differentially resistant Zeamays genotypes. Phytopathology 89, 695–700. Matsumoto, T., Ookawa, T., Hashegawa, S., and Inoue, M. (1989). Occurrence of viral disease caused by white clover mosaic virus in pea (abstract in Japanese). Ann. Phytopathol. Soc. Japan. 55, 90. Meyer, J.D., Silva, D.C., Yang, C., Pedley, K.F., Zhang, C., van de Mortel, M., Hill, J.H., Shoemaker, R.C., Abdelnoor, R.V., Whitham, S.A., et al. (2009). Identification and analyses of candidate genes for rpp4-mediated resistance to Asian soybean rust in soybean. Plant Physiol. 150, 295–307. https://doi.org/10.1104/pp.108.134551 Morel, J.B., Godon, C., Mourrain, P., Béclin, C., Boutet, S., Feuerbach, F., Proux, F., and Vaucheret, H. (2002). Fertile hypomorphic ARGONAUTE (ago1) mutants impaired in post-transcriptional gene silencing and virus resistance. Plant Cell 14, 629–639. Mourrain, P., Béclin, C., Elmayan, T., Feuerbach, F., Godon, C., Morel, J.B., Jouette, D., Lacombe, A.M., Nikic, S., Picault, N., et al. (2000). Arabidopsis SGS2 and SGS3 genes are required for posttranscriptional gene silencing and natural virus resistance. Cell 101, 533–542. Nagamatsu, A., Masuta, C., Matsuura, H., Kitamura, K., Abe, J., and Kanazawa, A. (2009). Down-regulation of flavonoid 3’-hydroxylase gene expression by virusinduced gene silencing in soybean reveals the presence of a threshold mRNA level associated with pigmentation in pubescence. J. Plant Physiol. 166, 32–39. https://doi. org/10.1016/j.jplph.2008.02.002

342  | Sinha and Senthil-Kumar

Nagamatsu, A., Masuta, C., Senda, M., Matsuura, H., Kasai, A., Hong, J.S., Kitamura, K., Abe, J., and Kanazawa, A. (2007). Functional analysis of soybean genes involved in flavonoid biosynthesis by virus-induced gene silencing. Plant Biotechnol. J. 5, 778–790. Nakahara, K.S., Nishino, K., and Uyeda, I. (2015). Construction of infectious cDNA clones derived from the potyviruses Clover yellow vein virus and Bean yellow mosaic virus. Plant Virology Protocols: New Approaches to Detect Viruses and Host Responses. pp. 219–227. Nam, M., Kim, J.S., Park, S.J., Park, C.Y., Lee, J.S., Choi, H.S., Kim, J.S., Kim, H.G., Lim, S., Moon, J.S., et al. (2012). Biological and molecular characterization of Soybean yellow common mosaic virus, a new species in the genus Sobemovirus. Virus Res. 163, 363–367. https://doi. org/10.1016/j.virusres.2011.08.005 Nene, Y.L., Reddy, M.V., Haware, M.P., Ghanekar, A.M., Amin, K.S., and Pande, S. (2012). Field diagnosis of chickpea diseases and their control. Information Bulletin No. 28 (revised). Technical Report, International Crops Research Institute for the Semi-Arid Tropics, Patancheru. Nolasco, G., de Blas, C., Torres, V., and Ponz, F. (1993). A method combining immunocapture and PCR amplification in a microtiter plate for the detection of plant viruses and subviral pathogens. J. Virol. Methods 45, 201–218. Otagaki, S., Arai, M., Takahashi, A., Goto, K., Hong, J.S., Masuta, C., and Kanazawa, A. (2006). Rapid induction of transcriptional and post-transcriptional gene silencing using a novel Cucumber mosaic virus vector. Plant Biotechnol. 23, 259–265. Palukaitis, P., and García-Arenal, F. (2003). Cucumoviruses. Adv. Virus Res. 62, 241–323. Palukaitis, P., Roossinck, M.J., Dietzgen, R.G., and Francki, R.I. (1992). Cucumber mosaic virus. Adv. Virus Res. 41, 281–348. Pang, J., Turner, N.C., Khan, T., Du, Y.L., Xiong, J.L., Colmer, T.D., Devilla, R., Stefanova, K., and Siddique, K.H.M. (2017). Response of chickpea (Cicer arietinum L.) to terminal drought: leaf stomatal conductance, pod abscisic acid concentration, and seed set. J. Exp. Bot. 68, 1973–1985. https://doi.org/10.1093/jxb/erw153 Parida, S.K., Verma, M., Yadav, S.K., Ambawat, S., Das, S., Garg, R., and Jain, M. (2015). Development of genomewide informative simple sequence repeat markers for large-scale genotyping applications in chickpea and development of web resource. Front. Plant Sci. 6, 645. https://doi.org/10.3389/fpls.2015.00645 Parween, S., Nawaz, K., Roy, R., Pole, A.K., Venkata Suresh, B., Misra, G., Jain, M., Yadav, G., Parida, S.K., Tyagi, A.K., et al. (2015). An advanced draft genome assembly of a desi type chickpea (Cicer arietinum L.). Sci Rep 5, 12806. https://doi.org/10.1038/srep12806 Pflieger, S., Blanchet, S., Meziadi, C., Richard, M.M., Thareau, V., Mary, F., Mazoyer, C., and Geffroy, V. (2014). The ‘one-step’ Bean pod mottle virus (BPMV)derived vector is a functional genomics tool for efficient overexpression of heterologous protein, virus-induced gene silencing and genetic mapping of BPMV R-gene in common bean (Phaseolus vulgaris L.). BMC Plant Biol. 14, 232. https://doi.org/10.1186/s12870-014-0232-4

Pflieger, S., Richard, M.M.S., Blanchet, S., Meziadi, C., and Geffroy, V. (2013). VIGS technology: an attractive tool for functional genomics studies in legumes. Funct. Plant Biol. 40, 1234−1248. Pradhan, S., Bandhiwal, N., Shah, N., Kant, C., Gaur, R., and Bhatia, S. (2014). Global transcriptome analysis of developing chickpea (Cicer arietinum L.) seeds. Front. Plant Sci. 5, 698. https://doi.org/10.3389/ fpls.2014.00698 Ratcliff, F., Martin-Hernandez, A.M., and Baulcombe, D.C. (2001). Technical Advance. Tobacco rattle virus as a vector for analysis of gene function by silencing. Plant J. 25, 237–245. Robertson, D. (2004). VIGS vectors for gene silencing: many targets, many tools. Annu. Rev. Plant Biol. 55, 495–519. https://doi.org/10.1146/annurev. arplant.55.031903.141803 Ruiz, M.T., Voinnet, O., and Baulcombe, D.C. (1998). Initiation and maintenance of virus-induced gene silencing Plant Cell 10, 937–946. Ryu, C.M., Anand, A., Kang, L., and Mysore, K.S. (2004). Agrodrench: a novel and effective agroinoculation method for virus-induced gene silencing in roots and diverse Solanaceous species. Plant J. 40, 322–331. https://doi.org/10.1111/j.1365-313X.2004.02211.x Sanyal, I., Singh, A.K., and Amla, D.V. (2003). Agrobacteriumtumefaciens-mediated transformation of chickpea (Cicerarietinum L.) using mature embryo axes and cotyledonary nodes. Ind. J. Biotechnol. 2, 524–532. Sanyal, I., Singh, A.K., Kaushik, M., and Amla, D.V. (2005). Agrobacterium-mediated transformation of chickpea (Cicerarietinum L.) with Bacillusthuringiensis cry1Ac gene for resistance to pod borer insect Helicoverpa armigera. Plant Sci. 168, 135–1146. Senthil-Kumar, M., and Mysore, K.S. (2011). Virus-induced gene silencing can persist for more than 2 years and also be transmitted to progeny seedlings in Nicotiana benthamiana and tomato. Plant Biotechnol. J. 9, 797–806. https://doi.org/10.1111/j.1467-7652.2011.00589.x Senthil-Kumar, M., and Mysore, K.S. (2014). Tobacco rattle virus-based virus-induced gene silencing in Nicotiana benthamiana. Nat Protoc. 9, 1549–1562. https://doi. org/10.1038/nprot.2014.092 Sharman, M., Kehoe, M., Coutts, B., van Leur, J., Filardo, F., and Thomas, J. (2016). Two complete genome sequences of phasey bean mild yellows virus, a novel member of the luteoviridae from Australia. Genome Announc 4, e01569–15. https://doi.org/10.1128/ genomeA.01569-15 Singh, A.K., Fu, D.Q., El-Habbak, M., Navarre, D., Ghabrial, S., and Kachroo, A. (2011). Silencing genes encoding omega-3 fatty acid desaturase alters seed size and accumulation of Bean pod mottle virus in soybean. Mol. Plant Microbe Interact. 24, 506–515. https://doi. org/10.1094/MPMI-09-10-0201 Sinha, R., Gupta, A., and Senthil-Kumar, M. (2017). Concurrent Drought stress and vascular pathogen infection induce common and distinct transcriptomic responses in chickpea. Front. Plant Sci. 8, 333. https:// doi.org/10.3389/fpls.2017.00333 Suzuki, M., Kuwata, S., Kataoka, J., Masuta, C., Nitta, N., and Takanami, Y. (1991). Functional analysis of deletion

Establishment of VIGS in Chickpea |  343

mutants of Cucumber mosaic virus RNA3 using an in vitro transcription system. Virology 183, 106–113. Thomas, J.E., Parry, J.N., Schwinghamer, M.W., and Dann, E.K. (2010). Two novel Mastreviruses from chickpea (Cicer arietinum) in Australia. Arch. Virol. 155, 1777– 1788. https://doi.org/10.1007/s00705-010-0763-4 Tsuchizaki, T., Goto, T., Fujisawa, I., and Yoshida, K. (1981). Virus disease occurring on legumes and vegetables in Hokkaido. Res. Bull. Hokkaido Natl. Agric. Exp. Stn. 131, 71–93. Upadhyaya, H.D., Bajaj, D., Das, S., Kumar, V., Gowda, C.L., Sharma, S., Tyagi, A.K., and Parida, S.K. (2016). Genetic dissection of seed-iron and zinc concentrations in chickpea. Sci. Rep. 6, 24050. https://doi.org/10.1038/ srep24050 Várallyay, E., Lichner, Z., Sáfrány, J., Havelda, Z., Salamon, P., Bisztray, G., and Burgyán, J. (2010). Development of a virus induced gene silencing vector from a legumes infecting tobamovirus. Acta. Biol. Hung. 61, 457–469. https://doi.org/10.1556/ABiol.61.2010.4.9 Varshney, R.K., Song, C., Saxena, R.K., Azam, S., Yu, S., Sharpe, A.G., Cannon, S., Baek, J., Rosen, B.D., Tar’an, B., et al. (2013). Draft genome sequence of chickpea (Cicer arietinum) provides a resource for trait improvement. Nat. Biotechnol. 31, 240–246. https:// doi.org/10.1038/nbt.2491 Voinnet, O. (2001). RNA silencing as a plant immune system against viruses. Trends Genet. 17, 449–459. Voinnet, O. (2005). Induction and suppression of RNA silencing: insights from viral infections. Nat. Rev. Genet. 6, 206. Wang, C., Cai, X., Wang, X., and Zheng, Z. (2006). Optimisation of Tobacco rattle virus-induced gene silencing in Arabidopsis. Funct. Plant Biol. 33, 347–355. Wang, R.Y., Stork, J., and Nagy, P.D. (2009). A key role for heat shock protein 70 in the localization and insertion of tombusvirus replication proteins to intracellular membranes. J. Virol. 83, 3276–3287. https://doi. org/10.1128/JVI.02313-08 Wardhan, V., Pandey, A., Chakraborty, S., and Chakraborty, N. (2016). Chickpea transcription factor CaTLP1 interacts with protein kinases, modulates ROS accumulation and promotes ABA-mediated stomatal closure. Sci. Rep. 6, 38121. https://doi.org/10.1038/ srep38121

Waterhouse, P.M., Wang, M.B., and Lough, T. (2001). Gene silencing as an adaptive defence against viruses. Nature 411, 834–842. https://doi.org/10.1038/35081168 Yamagishi, N., and Yoshikawa, N. (2009). Virus-induced gene silencing in soybean seeds and the emergence stage of soybean plants with Apple latent spherical virus vectors. Plant Mol. Biol. 71, 15–24. https://doi.org/10.1007/ s11103-009-9505-y Zaumeyer, W.J., and Thomas, H.R. (1948). Pod mottle, a virus disease of beans. J. Agricult. Res. 77, 81–96. Zhang, C., Bradshaw, J.D., Whitham, S.A., and Hill, J.H. (2010). The development of an efficient multipurpose bean pod mottle virus viral vector set for foreign gene expression and RNA silencing. Plant Physiol. 153, 52–65. https://doi.org/10.1104/pp.109.151639 Zhang, C., Grosic, S., Whitham, S.A., and Hill, J.H. (2012). The requirement of multiple defense genes in soybean Rsv1-mediated extreme resistance to Soybean mosaic virus. Mol. Plant Microbe Interact. 25, 1307–1313. https://doi.org/10.1094/MPMI-02-12-0046-R Zhang, C., and Ghabrial, S.A. (2006). Development of Bean pod mottle virus-based vectors for stable protein expression and sequence-specific virus-induced gene silencing in soybean. Virology 344, 401–411. Zhang, C., Whitham, S.A., and Hill, J.H. (2013). Virusinduced gene silencing in soybean and common bean. Methods Mol. Biol. 975, 149–156. https://doi. org/10.1007/978-1-62703-278-0_11 Zhang, C., Yang, C., Whitham, S.A., and Hill, J.H. (2009). Development and use of an efficient DNA-based viral gene silencing vector for soybean. Mol. Plant Microbe Interact. 22, 123–131. https://doi.org/10.1094/ MPMI-22-2-0123 Zhao, F., Hwang, U.S., Lim, S., Yoo, R.H., Igori, D., Lee, S.H., Lim, H.S., and Moon, J.S. (2015). Complete genome sequence and construction of infectious fulllength cDNA clones of tobacco ringspot Nepovirus, a viral pathogen causing bud blight in soybean. Virus Genes 51, 163–166. https://doi.org/10.1007/s11262-015-1221-x Zhao, F., Lim, S., Igori, D., Yoo, R.H., Kwon, S.Y., and Moon, J.S. (2016). Development of Tobacco ringspot virus-based vectors for foreign gene expression and virus-induced gene silencing in a variety of plants. Virology 492, 166–178. https://doi.org/10.1016/j.virol.2016.02.025

Index

A Abaca bunchy top virus (ABTV)  260 ABA-insensitive RING protein 1 gene (CaAIR1)  290, 320 Abscisic acid (ABA)  7, 8, 290, 292, 297 Abutilon mosaic virus (AbMV)  10, 70, 211, 285 AC1/C1 (replication initiation protein or Rep)  68, 71, 106, 145, 165, 225, 227, 228, 237, 240, 246, 257, 266 AC2/C2 (transcription activator protein or TrAP)  26, 58, 68– 1, 74, 75, 145, 165, 240 AC3/C3 (replication enhancer or REn)  68–71, 145, 240 Adenosine kinase (ADK)  59, 75 African cassava mosaic virus (ACMV)  12, 106, 160, 163, 210, 247, 279 Alfalfa mosaic virus (AMV)  43, 107, 124, 224, 225, 336, 337 amiRNAs (artificial microRNAs)  13, 108, 122, 151, 152, 166, 224, 225, 229, 246 Amplified fragment length polymorphism (AFLP)  87, 88, 90–95 Antisense RNA (asRNA)  108, 183, 221, 224, 225, 227, 230, 243, 278 Apetela2/ethylene-responsive element binding (AP2/ ERF)  8, 25–27, 36–38, 43, 118 Apple latent spherical virus (ALSV)  279, 280, 295, 331–332, 335 Arabidopsis nitric oxide-associated protein 1 (AtNOA1) 10 Arabidopsis thaliana  2, 34, 57, 97, 121, 244 Arabidopsis transcription activator factor (ATAF1/2)  26 Argonaute (AGO)  2, 61, 122, 147, 214, 278, 301, 330 Argonaute 4 (AGO4)  3, 59, 60, 62–67, 76 Artichoke mottled crinkle virus (AMCV)  111 Auxin (Aux)  6, 8, 245, 301, 302

B Babuvirus  256, 257, 266 Bacillus thuringiensis 215 Bacterial artificial chromosomes (BACs)  91 Badnavirus  256, 261 Banana bract mosaic virus (BBrMV)  256, 262–266, 269 Banana bunchy top virus (BBTV)  255–260, 265–275 Banana streak virus (BSV)  73, 261, 262, 266–269 Basic helix–loop–helix (bHLH) domain  8, 39, 43, 293, 302 Basic leucine zipper (bZIP)  8, 25–28, 38–51, 299

Basic leucine zipper (bZIP) type rice transcription factors, RF2a and RF2b  39, 178, 180 Bean golden mosaic virus (BGMV)  68, 106, 151, 208, 211, 236, 238, 245, 246 Bean golden yellow mosaic virus (BGYMV)  94, 95, 209 Bean pod mottle virus (BPMV)  279, 281, 293, 300, 331, 332, 336 Beet curly top virus (BCTV)  3, 4, 70, 75, 237, 247, 285 Begomovirus  14, 35, 36, 66, 68, 70, 96, 98, 107, 108, 121, 143–146, 151–157, 160, 163, 201–219, 221–222, 235–253, 285, 286 Bemisia tabaci  163, 201 Brassinosteroid (BR)  8, 28, 35, 293, 297, 301 Bulk segregant analysis (BSA)  91

C Cabbage leaf curl virus (CaLCuV)  2, 3, 61, 74, 109, 279, 285 CaMV35S promoter  165, 168, 279, 334 Cardamom bushy dwarf virus (CBDV) 260 Cassava (Manihot esculenta)  1, 5, 12, 95, 159–176, 204, 210, 211, 236, 247, 279, 295 Cassava brown streak disease (CBSD)  159–172 Cassava brown streak virus (CBSV)  160, 163–170 Cassava mosaic disease (CMD)  159–172 Cassava mosaic geminiviruses (CMGs)  1, 160, 163, 166 Cauliflower mosaic virus (CaMV)  2, 3, 8, 61, 149 Caulimoviridae  107, 179, 256, 261 Chickpea chlorotic dwarf virus (CpCDV)  237, 336, 338 Chromatin immunoprecipitation (ChIP)  44, 75, 77 Chromomethylase 3 (CMT3)  59, 60, 64–67, 73–76 Coat protein (AV1/V1/CP)  1, 4, 42, 43, 66, 68, 69, 71, 104, 105, 144–150, 164–166, 177–182, 190, 211, 224, 225, 229, 237–242, 247, 261–266, 269, 287, 295, 331 Coat protein mediated resistance (CPMR)  105, 106, 108, 224, 226, 269 Coiled-coil (CC) – or Toll/interleukin-1 receptor (TIR) – nucleotide-binding leucine-rich repeat (NLR) protein families 5 Confined field trials (CFTs)  167, 169, 171 Cotton leaf crumple virus (CLCrV)  209, 279, 285 Cowpea mosaic virus (CPMV)  97, 107 CRISPR (clustered regulatory interspaced short palindromic repeats)  5, 13, 14, 77, 109, 121, 123, 135, 136, 151, 152, 172, 214, 226, 231, 255, 257, 268, 269

346  | Index

CRISPR-associated nuclease 9 (Cas9) (CRISPR-Cas9) system  13, 14, 109, 255, 268, 269 Cucumber mosaic virus (CMV)  4, 39, 62, 97, 105, 108, 124, 125, 133, 147, 149, 221, 222, 224–226, 229, 230, 242, 264, 268, 279, 281, 331, 337 Cup-shaped cotyledon (CUC)  26 Cyclophilin B 212 Cylindrical inclusion protein (C1)  145 Cymbidium ring spot virus 2

D Defective interfering (DI) RNAs or DNAs  104, 107 Dehydrins (DHNs)  290 Dicer-like proteins (DCL)  2, 61, 72, 108, 121, 147, 278, 330 DNA methyltransferase 1 (MET1)  59 DNA-dependent RNA polymerase II (Pol II)  70 Double antibody sandwich (DAS) ELISA  262 Double-stranded RNA (dsRNA)  2–6, 59–67, 72, 108, 119–137, 146, 165, 168, 179, 188, 214, 245, 278 DRB3 3 DRM1  3, 59, 60 DRM2  3, 59, 60, 61–67, 76

E East African cassava mosaic Cameroon virus (EACMCV) 163 East African cassava mosaic Malawi virus (EACMMV)  163, 207 East African Cassava mosaic virus (EACMV)  160, 163–168, 171, 207, 208, 240, 295 East African cassava mosaic Zanzibar virus (EACMZV)  163, 207, 210 Effector-triggered immunity (ETI)  1, 5, 6, 9, 26, 33 Endocytosis cell signaling domain (ECS)  241 Endogenous BSV (eBSV)  255 Endogenous Petunia vein clearing virus (ePVCV)  58 Enhanced response to ABA1 – ERA1  280, 290, 291 Enzyme-linked immunosorbent assays (ELISAs)  144, 185, 187, 188, 256, 259, 262, 263, 265 Ethylene response factors (ERFs)  37, 110 Expressed sequence tag (EST)  93, 277 Extreme resistance (ER)  89, 97, 98, 227, 295

F Flowering Locus C (FLC)  57 Food and Agriculture Organization (FAO)  142 Friable embryogenic callus (FEC)  165

G Geminiviridae  66, 107, 143, 144, 160, 163, 202, 222, 235, 237, 265, 336–338 Genetically modified crops (GMOs)  122 Gibberellins (GA)  8 Green fluorescent protein (GFP)  280–287, 331, 332 Groundnut bud necrosis virus (GBNV)  222, 223, 230

H Hairpin RNA (hpRNA)  108, 227, 229, 246, 266, 268 Helix–turn–helix (HTH)  34 Helper component proteinase (HC-Pro)  124, 125, 132–134, 145, 149, 263, 279 Histone acetyltransferases (HATs)  57

Histone deacetylase (HDAC)  57 Histone lysine methyltransferases (HKMTs)  57 Homology-directed repair (HDR)  109 Host induced gene silencing (HIGS)  119, 122 HUA Enhancer 1 (HEN1)  4, 60–67, 76, 244 HY5 Homologue (HYH)  57 Hydrogen peroxide (H2O2)  112 Hypersensitive reaction/response (HR)  89, 109–111

I IC-RT-LAMP  256, 263 Immunosorbent electron microscopy (ISEM)  262 Indian Cassava mosaic virus (ICMV)  74, 205 Integrated pest management (IPM)  120, 151, 213 Intellectual property rights (IPR)  112 Intergenic region (IR)  58, 63, 66, 74, 163, 179, 237, 238, 266 Internal ribosome entry site (IRES)  269 International Committee on Taxonomy of Viruses (ICTV)  66, 209, 211, 237, 338 Inter-simple sequence repeats (ISSR)  90, 92 Ipomovirus  108, 160, 163, 164, 203

J JAX1 (jacalin-type lectin)  97, 110

L Leucine-rich repeat receptor-like kinase (LRR-RLK)  5, 12, 36, 87, 96–98, 110, 242, 280, 294–300 Ligation-independent cloning (LIC)  287 Lipid transfer protein (LTP)  10, 298, 302 Long Hypocotyl 5 (HY5)  57 Loop-mediated isothermal amplification (LAMP)  256, 259, 262, 263, 265 Lycopersicon chilense 90 Lysine of histone H3 (H3K9)  57–64

M Magnesium chelatase subunit H (ChlH)  280–287, 292, 333, 334, 336 MAP kinase kinase kinases (MAPKKKs/MEKKs/ MKKKs) 291 MAP kinase kinases (MAPKKs/MEKs/MKKs)  291 MAP kinases (MAPKs/MPKs)  32, 291 MAPK (Mitogen-activated protein kinase)  5, 6, 291, 292 Marker-assisted selection (MAS)  87, 88 Mesta yellow vein mosaic virus (MYVMV)  9 Methyl jasmonate (MeJA)  214, 296, 300 Methylated DNA immunoprecipitation (MeDIP)  77 microRNA (miRNAs)  13, 63, 107, 108, 151, 166, 214, 224, 225, 227, 229, 235, 243–245, 294, 302 miRNA-induced gene silencing (MIGS)  108 Movement protein (BC1/MP)  7, 68–71, 106, 107, 120, 145, 178, 186–190, 237–242, 257, 261, 264, 287, 331 Myeloblastosis-related proteins (MYB)  8, 25–28, 34–36, 40, 43, 45–53, 61, 245, 280, 284, 291, 294, 296, 302–305 Myelocytomatosis-related family (MYC)  26–27, 39–41, 43, 303

N N receptor interacting protein (NPRI)  10 Nanoviridae  256, 257, 266, 338 Natural antisense transcript siRNAs (NAT-siRNAs)  245

Index |  347

New World (NW)  68, 203, 207–211, 238, 241 Next generation sequencing (NGS)  59, 119, 122, 123, 230, 277 No apical meristem (NAM)  26 Non-expressor of pathogenesis related proteins 1 (NPR1)  7–9, 29–32, 38, 39, 42, 43 Non-homologous end joining (NHEJ)  109 Non-host resistance  89, 299 Nonsense-mediated decay (NMD)  121 Nuclear inclusion protein a (NIa)  145 Nuclear inclusion protein b (NIb)  145, 149, 150 Nuclear shuttle protein (BV1/NSP)  14, 68, 69, 70, 71, 121, 145, 241, 257 Nucleoprotein (NP)  7, 184, 189

O Old World (OW)  68, 204, 205, 241

P PAMP-triggered immunity (PTI)  5–7 Papaya leaf crumple virus (PaLCrV)  143, 144, 205 Papaya leaf curl virus  143, 144 Papaya leaf distortion mosaic virus (PLDMV)  143–146, 149–151 Papaya lethal yellowing virus (PLYV)  143, 144 Papaya meleira virus (PMeV)  143, 144, 146 Papaya mosaic virus (PapMV)  143–145 Papaya ringspot virus (PRSV)  104, 141–157 Pathogen-associated molecular patterns (PAMPs)  1, 2, 5–7, 26, 296 Pathogen-derived resistance (PDR)  104, 121, 147, 149, 152, 224, 235, 236, 257, 265, 266 Pathogen-triggered immunity (PTI)  1, 5–7, 9, 14 Pathogenesis-related (PR) proteins  8, 11, 30, 37, 111, 144, 296, 299 Pathogenesis-related protein (AC4)  69 Pattern recognition receptors (PRRs)  1, 5, 7 Pea early browning virus (PEBV)  279, 282, 292, 293, 331, 332, 333, 335 Peanut chlorotic streak virus 107 Pentalonia nigronervosa  256, 257, 263 Pepper golden mosaic virus (PepGMV)  58, 73, 209, 295 Pepper mild mottle virus (PMMoV)  4, 110, 124 Peptidoglycan recognition proteins (PGRPs)  212 Phased secondary siRNAs (phasiRNAs)  5, 6 Phytoene desaturase (PDS)  279, 287, 288, 303, 332, 334 Plum pox virus (PPV)  8, 97, 105, 108, 124, 282, 294 Pokeweed (Phytolacca americana) antiviral protein (PAP) 111 Pol IV  3, 62–67, 76, 123 Pol V  3, 60–67, 123 Polycomb repressive complex 2 (PRC2)  57 Ponciru strifoliata 90 Post-transcriptional gene silencing (PTGS)  2, 7, 59, 70, 106, 107, 121, 147, 163, 165, 182, 224, 240, 243, 246, 269, 278, 330 Potato leaf roll virus (PLRV)  89, 105, 107, 111 Potato spindle tuber viroid (PSTVd)  73, 124 Pre-coat protein (AV2)  68, 69, 145, 163, 168, 229, 239, 240, 243, 248 Programmed cell death (PCD)  5, 7, 9, 292, 294, 296 Protein arginine methyltransferases (PRMTs)  57 Protein degradation domain (PEST)  241

R Random amplified polymorphic DNA (RAPD)  87, 88, 90–102 Reactive nitrogen species (RNS)  7 Reactive oxygen species (ROS)  5, 7, 9, 291, 292 Recombination dependent replication (RDR)  239 Regulatory field trials (RFTs)  167 Resistance gene homologue (RGH)  91 Restriction fragment length polymorphism (RFLP)  87, 88, 90–94 Reverse-transcriptase polymerase chain reaction (RTPCR)  132–135, 160, 170, 256, 262, 263, 265, 335, 336 Ribonucleoprotein (RNP) complex  1, 7, 61, 106, 189 Ribosomal protein L9 (RPL9)  214 Ribosome-inactivating proteins (RIP)  104, 110 Rice black-streaked dwarf virus (RBSDV)  42, 178, 185, 186 Rice bunchy stunt virus  177, 186 Rice dwarf virus (RDV)  3, 31, 42, 177, 178, 187 Rice gall dwarf virus  177, 178, 187 Rice giallume virus  177, 189 Rice grassy stunt virus (RGSV)  177, 178, 189 Rice hoja Blanca virus  177, 190 Rice necrosis mosaic virus  177, 178, 190 Rice ragged stunt virus (RRSV)  42, 178, 183, 184 Rice stripe necrosis virus  177, 184 Rice stripe virus (RSV)  3, 90, 177, 178, 182 Rice transitory yellowing virus (RTYV)  31, 42, 177, 178, 184 Rice tungro bacilliform virus  177, 178–180 Rice tungro spherical virus  90, 177, 178–180 Rice yellow mottle virus  3, 177, 178–181 Rice yellow stunt virus (RYSV)  3, 184 RNA-antisense purification with mass spectrometry (RAPMS) 77 RNA-dependent RNA polymerase (RDR)  2, 61, 63, 98, 106, 145, 181, 242, 278, 295 RNA-dependent RNA polymerase 2 (RDR2)  62 RNA-directed DNA methylation (RdDM)  3, 4, 56, 59–66, 72–74, 123, 136, 223 RNA-directed DNA Methylation 1 (RDM1)  60, 62, 63 RNA-induced silencing complex (RISC)  3, 4, 61, 108, 121, 122, 147, 151, 214, 244–246, 278, 280, 330 RNA quality control (RQC)  120, 121 RNA silencing (RNA-interference or RNAi)  1, 44, 61, 69, 98, 104, 107, 108, 119–127, 131–140, 147, 151–156, 159, 166–177, 179, 181, 183, 185, 186–191, 214, 215, 221, 229, 235, 236, 240–249, 255, 266, 268, 269, 277–278, 305, 330, 334 Rolling-circle replication (RCR)  66, 69, 70, 237–240 RuBisCO small subunit (RbCS)  11, 280, 283, 285, 334

S S-adenosyl homocysteine hydrolase (SAHH)  74–76, 110 S-adenosyl-methionine (SAM) synthesis  4, 75 S-adenosyl-methionine decarboxilase 1 (SAMDC1) 74–76 Salicylic acid (SA)  5, 7, 8, 14, 25, 32, 33, 37–39, 42, 43, 111, 112, 213, 214, 290, 294, 297, 298 Sequence characterized amplified regions (SCARs)  88 Sequence enhancing geminivirus symptoms 1 and 2 (SEGS-1and SEGS-2)  161 Shoot apical meristem (SAM)  244, 302 shRNAs (short hairpin RNAs)  107

348  | Index

Sida golden mosaic Costa Rica virus (SiGMCRV)  211 Single Nucleotide Polymorphisms (SNPs)  13, 94 Single-cell restriction analysis of methylation (SCRAM) 77 Single-chain variable fragment (scFv) antibodies  111 siRNAs (small interfering RNAs)  2, 7, 107, 147, 179, 224, 225, 227, 244, 245 Solanum chilense  98, 222, 295 Solanum habrochaites  89, 222, 231, 242 Solanum peruvianum  11, 89, 98, 222, 231, 243 Solanum stoloniferum  90, 98 South African Cassava mosaic virus (SACMV)  12, 163, 207 Soybean mosaic virus (SMV)  89, 90, 96, 97, 242, 295 Soybean yellow common mosaic virus (SYCMV)  279, 289, 331, 333–336 Spray-induced gene silencing (designated as SIGS)  130 Sri Lankan Cassava mosaic virus (SLCMV)  12, 163, 168, 205 Subterranean clover mottle virus (SCMoV)  279, 331, 337 Sunhemp mosaic virus (SHMV)  279 Sweetpotato mild mottle virus (SPMMV)  160 Systemic acquired resistance (SAR)  5, 7–9, 32, 39, 110, 111

T Target region amplified polymorphism (TRAP)  68–71, 74, 87, 91–94 Targeting Induced Local Lesions IN Genomes (TILLING) 13 tasiRNAs (trans-acting siRNAs)  13, 60, 63–65, 107–108, 224, 225, 229, 230, 235, 244, 245, 247–249 Tobacco mosaic virus (TMV)  4, 30–34, 36, 38, 42, 91, 96, 98, 104, 120, 121, 124, 134, 147, 224–226, 229, 247, 279, 282, 285, 294 Tobacco rattle virus (TRV)  3, 107, 279, 282–284 Tobacco ringspot virus (TRSV)  12, 107, 279, 284, 295, 331, 333 Tomato golden mosaic virus (TGMV)  70, 73–75, 238, 279, 286 Tomato leaf curl New Delhi virus (ToLCNDV)  69, 73, 144, 202, 205–208, 210, 211, 222, 225, 229, 230, 235, 243, 246–248 Tomato leaf curl viruses (ToLCVs)  70, 205, 206, 208, 210, 222, 223, 225–230, 235–240, 243, 246, 248, 286 Tomato spotted wilt virus (TSWV)  110, 222, 226, 230 Tomato yellow leaf curl China virus (TYLCCNV)  75, 206, 213, 226, 286, 293

Tomato yellow leaf curl Sardinia virus (TYLCSV)  73–75, 106, 206–208, 211, 225–229, 243 Tomato yellow leaf curl virus (TYLCV)  3, 8–11, 70, 89, 92, 109, 121, 136, 144, 202, 206–213, 222, 224–227, 230–231, 242, 243, 247, 279, 295, 338 Trait selection trials (TST)  167 Transcription activator-like effector nucleases (TALENs)  109, 231, 268 Transcription factors (TFs)  3, 6–9, 12, 14, 25–53, 58, 60, 62, 69, 75, 109, 178, 180, 244–245, 291–304, 331 Transcriptional gene silencing (TGS)  2, 3, 7, 56, 59, 62, 69, 70, 98, 106, 107, 121, 123, 223, 224, 243, 246, 278 Trans-kingdom RNAi (tkRNAi)  131, 136 Translation initiation factors (eIFs)  109, 110, 121, 294 Transmembrane immune receptor NIK1 (nuclear shuttle protein (NSP)-Interacting Kinase1)  14 Transposable elements (TEs)  56

U Ubiquitin proteasome system (UPS)  10 Ugandan cassava brown streak virus (UCBSV)  160, 163–170

V Vacuolar-type ATPase A subunit (v-ATPase A) 214 Virus-associated molecular patterns (VAMPs)  5 Virus-induced gene silencing (VIGS)  41, 98, 132, 136, 166, 277–342 VPg (viral protein-genome linked)  109, 144, 145, 152, 180, 334

W Watermelon chlorotic stunt virus (WmCSV)  202, 206–209, 211 Watermelon mosaic virus (WMV)  97, 105, 108, 147 White clover mosaic virus (WCIMV)  107, 279, 285, 331, 333, 335

Y Yield selection trials (YST)  167

Z Zinc finger nucleases (ZFNs) technology  109, 231 Zucchini yellow mosaic virus (ZYMV)  4, 97, 104–105, 108, 109, 121, 125, 132, 295

Genes, Genetics and Transgenics for Virus Resistance in Plants Viral diseases of crop plants cause significant yield and economic losses and this poses a major threat to global food security. To make matters worse, there are no effective antiviral chemicals available and, although naturally resistant host genotypes exist, they are so rare that conventional breeding techniques cannot be used reliably to create resistant plants. The most effective option to combat phytopathogenic viruses is through biotechnological intervention, such as the use of genetic engineering to develop transgenic plants or the topical use of RNA-silencing technologies to prevent or modulate the severity of the viral infection. Since the first report on the virus resistance of transgenic tobacco plants in 1986, enormous progress has been made in this field. In addition, great strides have been made in our ability to genetically manipulate plants and viruses, leading to a plethora of novel applications. This has prompted the need for this book, which distills the most important research and provides a timely overview. This authoritative volume contains fifteen chapters, the breadth of which reflects the diversity of this research area. Topics covered range from understanding the mechanisms of virus resistance in plants, and the management of whitefly-transmitted viruses, to the principles and methods involved in genetic engineering of virus-resistant plants. Other chapters cover individual crops such as papaya, cassava, rice, tomato and banana, for which virus resistance has been accomplished by employing different transgenic technologies. This book is essential reading for everyone working in this field, both students and specialists, from academia, research institutes/organizations and industries. 

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