Molecular Biology of Potyviruses

ARTICLE IN PRESS

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

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

102 103 105 106 107 114 115 116 117 118 119 119 121 125 128 128 132 138 138 142 144 145 152 155 165 166 167 167

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

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2015 Elsevier Inc. All rights reserved.

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

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

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

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

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HCpro

P3

6K2

P1

6K1

Potyviruses, Rymoviruses Cl

Nla

Nlb

CP

poly A

VPg-pro P3N + PIPO

HCpro

P3

6K2

P1

6K1

Ipomoviruses Cl

Nla

Nlb

poly A

CP

SPMMV

VPg-pro P3

P1b

6K2

P1a

6K1

P3N + PIPO

Cl

Nla

Nlb

CP

poly A CV YV,SqVYV

CP

poly A

VPg-pro 6K2

P3

P1

6K1

P3N + PIPO

Cl

Nla

Nlb

HAM

CBSV

VPg-pro

P3N + PIPO

P3

6K2

P1 HCpro

6K1

Tritimoviruses, Poaceviruses Cl

Nla

Nlb

poly A

CP

VPg-pro

P3N + PIPO

HCpro

P3

6K2

P1

6K1

Brambyviruses Cl

Nla

Nlb

poly A

CP

VPg-pro

AlkB P3N + PIPO

P3

Cl

6K2

6K1

Macluraviruses HC pro

Nla

Nlb

CP

poly A

VPg-pro P3N + PIPO

P2-2

poly A

P3

Cl

6K2

P2-1

6K1

Bymoviruses Nla

Nlb

CP

poly A

VPg-pro P3N + PIPO

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

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

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

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HCpro

6K1

P1

P3

Cl

P3N + PIPO

Virus movement

Serine proteinase Accessory factor for virus amplification Host adaptation

Membrane vesicles proliferation Membrane targeting

6K2

Modulation of P3 activity?

Virus amplification Cysteine proteinase Host adaptation Helper factor for aphid transmission RNA silencing suppression Enhancement of yield of virus particles

RNA helicase RNA replication Pinwheel formation Virus movement

Nla

Nlb

RNA replicase

CP

poly A

VPg-pro Cysteine proteinase DNAse

Protection of genomic RNA Virus movement Aphid transmission

Genome-linked protein Primer of RNA replication RNA translation Virus movement

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

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

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

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

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

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

P1

Chloroplastic Rieske Fe/S protein

Y2H, Co-IP

SMV/Pinellia ternata Unknown

60S ribosomal subunit

Affinity purification

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

eIF4E/eIF(iso)4E

Y2H, BiFC

PVA-PVY-TEV/ tobacco-potato

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

Interaction associated with viral replication vesicles

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

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

Regulation of RNA silencing suppressor activity

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

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

TEV/tobacco; TuMV/A. thaliana

Mediating HCPro silencing suppressor activity

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

RNA methyltransferase HEN1

ELISA-binding assay

ZYMV/A. thaliana

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

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

Y2H, in vitro binding assays, BiFC

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

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

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HCPro

References

Y2H, BiFC

PVA/potato– tobacco

Reduced PVA accumulation in inoculated leaves of HIP2-silenced plants

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

RING finger protein HIP1

Y2H

PVA/potato

Unknown

Guo, Spetz, Saarma, and Valkonen (2003)

Chloroplast precursor of ferredoxin-5

Y2H, BiFC

SCMV/maize

Symptom development? Cheng et al. (2008)

Calreticulin

Y2H, BiFC

PRSV/papaya

Plant calcium signaling pathways?

Shen et al. (2010)

Chloroplast division-related factor NtMinD

Y2H, BiFC

PVY/tobacco

Chloroplast division?

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

P3

RubisCO subunits RbcL and RbcS

Y2H, Co-IP

SYSV-OYDVUnknown SMV-TuMV/onion

P3N– PIPO

Cation-binding protein PCaP1

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

Cell-to-cell movement

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

RubisCO subunits RbcL and RbcS

Y2H

SYSV-TuMV/ onion

Unknown

Lin et al. (2011)

eIF4E

ELISA-based assays, BiFC

LMV/lettuce

Cell-to-cell movement?

Tavert-Roudet et al. (2012)

CI

Lin et al. (2011)

Continued

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Microtubuleassociated protein HIP2

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

Jime´nez et al. (2006)

Chloroplastic photosystem I, PSI-K

Y2H, pull-down

PPV-TVMV/ N. benthamiana– A. thaliana

P58IPK

Y2H, pull-down

TEV/N. benthamiana Virulence factor

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

eIF4E/eIF(iso)4E

Y2H, in vitro binding assays, BiFC

Several potyvirus/ plant pathosystems

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

Fibrillarin

Y2H, BiFC

PVA/N. benthamiana Unknown

Rajama¨ki and Valkonen (2009)

PABP

ELISA-based binding assay, Co-IP

TuMV/Brassica perviridis

Viral RNA translation/ replication

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

eEF1A

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

Viral RNA translation/ replication

Thivierge et al. (2008)

Viral RNA translation/ replication

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VPg

Antiviral defense?

References

Y2H, BiFC

PPV/PeachA. thaliana

Viral RNA translation/ replication?

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

PVIP

Y2H

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

Movement?

Dunoyer, Thomas, et al. (2004)

NIaPro

Methionine sulfoxide reductase B1

Y2H

PRSV/papaya

Unknown

Gao et al. (2012)

NIb

eEF1A

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

Viral RNA translation/ replication

Thivierge et al. (2008)

PABP

Y2H, TAP

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

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

Hsc70-3

TAP

TuMV/A. thaliana

Dufresne, Thivierge, et al. (2008)

SUMOconjugating enzyme SCE1

Y2H, BiFC, FRET

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

Xiong and Wang (2013)

RubisCO-LSU

ELISA-based binding assay, bacterial twohybrid system

PVY/Tobacco

Feki et al. (2005)

CP

Viral RNA translation/ replication

Continued

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AtRH8

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

CP degradation

References

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

Co-IP, Y2H

PVA, PVY/ N. benthamianatobacco

HSP70

Co-IP

PVA/N. benthamiana CP degradation

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

Chloroplastic 37 kDa protein

ELISA-based binding assay

TuMV/lettuce

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

Unknown

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

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CPIP

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

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

References

50 -NCR

Chlorotic mosaic symptoms

PPV/ N. clevelandii

Simo´n-Buela, Guo, and Garcı´a (1997)

P1

Mild/severe symptoms

PPV/Nicotiana– Prunus persica

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

HCPro

Severe symptoms

Several pathosystems

See in the text

P3

Systemic necrosis

SMV/soybean

Hajimorad et al. (2005)

Systemic HR

TuMV/ A. thaliana

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

P3 N- and Severe C-terminal parts symptoms

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

Yellow mosaic

TuMV/Brassica juncea

Jenner et al. (2003)

P3 C-terminal part and 6K1

Chlorotic mottle symptoms

PPV/ N. benthamiana

Sa´enz et al. (2000)

P3/6K1 junction

Severe symptoms

PPV/ N. clevelandii

Riechmann et al. (1995)

P3 and CI– 6K2–VPg encompassing regions

Wilting response

TEV/tabasco pepper

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

VPg associated with P3

Severe symptoms

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

CI N-terminal part

Severe symptoms

SMV/soybean

Zhang et al. (2009)

NIa and NIb

Vein-clearing symptoms

PSbMV/pea

Johansen et al. (1996)

30 -NCR

Vein-mottling and blotch symptoms

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

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

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

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

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

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

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

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

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

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

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

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

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

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