Nematode-Borne Plant Viruses

CHAPTER 26 Nematode-Borne Plant Viruses Stuart A. MacFarlane Cell and Molecular Sciences Group, The James Hutton Institute Invergowrie, Dundee, U.K. Inga Zasada U.S. Department of Agriculture Agricultural Research Service Horticultural Crops Research Laboratory Corvallis, Oregon, U.S.A. Olivier Lemaire Gérard Demangeat French National Institute for Agricultural Research (INRA) University of Strasbourg, UMR 1131 Colmar, France Although it had been known since the early part of the twentieth century that soil could be a source of virus infection to newly established plants, the first direct demonstration that soil nematodes transmit plant viruses was not reported until 1958 (Hewitt et al., 1958). Quickly thereafter, other examples of virus transmission by nematodes were reported (Harrison and Cadman, 1959; Jha and Posnette, 1959), leading to the current acceptance of 30 nematode species that are known to transmit 15 viruses (Tables 26.1 and 26.2). The viruses are of two types that previously were referred to as either nepoviruses, with spherical particles, or tobraviruses, with rod-shaped particles. The name nepovirus originates from the contraction of nematode-transmitted virus with polyhedral particles (Cadman, 1963), while tobravirus is derived from the virus name Tobacco rattle virus. Recent changes to the classification of nematode-transmitted spherical viruses mean that not all of these viruses now are placed in the genus Nepovirus. Nevertheless, to simplify the discussion, we will continue to refer to this collection of viruses as nepoviruses. A comprehensive review detailing the biology of virus-vector nematodes was written by Taylor and Brown (1997). We build on this review to include new information about the molecular mechanism of virus transmission by nematodes, and we discuss current work that aims to control plant diseases caused by these viruses and their vector nematodes. Plant Viruses Transmitted by Nematodes Nepoviruses and tobraviruses are taxonomically unrelated to one another. However, like the majority of plant viruses, their genomes are single-stranded RNA of positive sense (i.e., the RNAs are directly translated in the host plant cell to produce virus proteins). Both types of viruses have two genomic RNAs that are contained separately within the virus particles. However, nepoviruses consist of spherical particles and tobraviruses of rod-shaped particles. Tobraviruses and viruses from the genus Nepovirus have a single coat protein (CP) that is assembled in multiple copies to form the virus particle. Some nematode-transmitted viruses that previously were classified as nepoviruses, and are highly similar to nepoviruses in both morphology and genome organization, have been found to encode two (Strawberry latent ringspot virus; unassigned genus) or even three CPs (Cherry rasp leaf virus; genus Cheravirus). In general, plant viruses have very small genomes (usually fewer than 15 kb), which, with the use of current, high-throughput RNA and DNA sequencing technologies, allows new viruses to be identified and characterized very rapidly and easily. As a consequence, the curated lists of known plant viruses are constantly being updated. Currently, the International Committee on Taxonomy of Viruses lists 36 virus species in the Nepovirus genus, four in the Cheravirus genus, and three so far unassigned to any genus (King et al., 2012). Of these 43 viruses, only 12 have been demonstrated to be transmitted by nematodes (Table 26.1). Some of the remaining viruses may be nematode transmitted, but most have not been tested to determine their transmission vector. It is also possible that some of these viruses may not have nematode vectors, for example, the nepo-like Blackcurrant reversion virus and Strawberry mottle virus, which are transmitted by mites and aphids, respectively. The nepoviruses are often also transmitted via infected seed or pollen. Therefore, focusing only on their transmission by nematodes may not be sufficient to control their spread in the environment. Individual nepoviruses tend to have restricted distributions determined by that of the individual vector nematode species, but as a group of viruses they have been reported from most parts of the world (Harrison and Murant, 1977a). There are only three members in the genus Tobravirus: Tobacco rattle virus (TRV), Pea early-browning virus (PEBV), and Pepper ringspot virus (PepRSV) (Table 26.2). TRV has a 365

366 Chapter 26 wide geographical distribution that includes Europe, North America, the former Soviet Union, Japan, and New Zealand. PEBV is found in the United Kingdom, western Europe, and North Africa, whereas PepRSV has been reported only in Brazil (Harrison and Robinson, 1986). TRV is a commercially relevant pathogen of potato and many different ornamental plants, while PEBV is limited to leguminous crop plants, in which it causes diseases of only local importance. Both viruses also infect weed plants of many species, which act as reservoirs of infection in the field, and are seed transmitted to varying degrees depending on the plant species (Harrison and Robinson, 1986). Several early reports suggested that viruses other than nepoviruses and tobraviruses, e.g., Brome mosaic virus, could be transmitted by nematodes. These reports are considered to be incorrect, resulting from the use of flawed experimental approaches, and led to the proposal of a set of criteria that should be addressed when the ability of nematodes to transmit viruses is assessed (Trudgill et al., 1983). Nematodes That Transmit Plant Viruses Only the orders Dorylaimida and Triplonchida have species that are able to transmit viruses. Within the order Dorylaimida, there are three genera in the family Longidoridae that transmit viruses: Xiphinema, Longidorus, and Paralongidorus. X. index (Fig. 26.1), X. americanum, and X. diversicaudatum have cosmopolitan distributions, while the other species in these genera are geographically isolated (Decraemer and Robbins, 2007). Species of Longidorus and Paralongidorus are most frequently encountered in Europe, with the exception of L. martini, which is found in Japan (Yagita, 1975) and L. diadecturus in North America (Robbins and Brown, 1991). Longidorid nematodes range in length from 1.3 mm (X. tarjanense) to 9.8 mm (P. maximus). Within the order Triplonchida, there are two genera in the family Trichodoridae that transmit viruses: Paratrichodorus and Trichodorus. Trichodorus is a cosmopolitan genus, but its members are predominant in temperate regions. Members of the Paratrichodorus genus are also cosmopolitan but are present mainly in tropical and subtropical regions (Decraemer and Robbins, 2007). Trichodorid nematodes (0.4 1.3 mm long) are generally much smaller than longidorid nematodes (Taylor and Brown, 1997) (Fig. 26.2). Since approximately 2000, the majority of work related to nematodes that transmit viruses has focused on improving our understanding of taxonomic relationships within these groups. The main diagnostic features of members of Longidoridae at the genus level are the structure of the feeding apparatus (odontostyle and odontophore), the structure and position of the ring that guides the feeding apparatus, and the shape and size of paired lateral sense organs near the lip region (Decraemer and Robbins, 2007). Members of this family of nematodes have life cycles that range from a few months to years (Flegg, 1968; Taylor and Brown, 1997) and can include three or four juvenile stages (Halbrendt and Brown, 1992; Robbins et al., 1995). There are seven species of Longidorus (Table 26.1) that together transmit six viruses. There is only one species of Paralongidorus, P. maximus, that transmits a virus, Raspberry ringspot virus (RpRSV); this nematode has a wide host range (Heyns, 1975). Compared with Longidorus and Paralongidorus spp., members of Xiphinema have proven to be much more difficult to define at the species level. Nine species of Xiphinema are reported to transmit viruses (Table 26.1), and of these nine species, six are part of the X. americanum group (X. americanum, X. bricolensis, X. californicum, X. intermedium, X. rivesi, and X. tarjanense) (Robbins, 1993). Since the original description of X. americanum sensu stricto (in the stricter sense) by Cobb (Cobb, 1913; Trudgill and Brown, 1978), there has been a proliferation of species, up to 38

Nematode-Borne Plant Viruses 367 (Lamberti and Carone, 1991), within this group, which is collectively referred to as X. americanum sensu lato (in the wider sense) or the X. americanum group. Because of the minimal inter- and intraspecific variation in both morphological and morphometric characters in members of the X. americanum group (Lazarova et al., 2006), taxonomic clarification of this group may not be realized until it is well characterized at the molecular level. Multiple regions of the ribosomal DNA (rdna) genes (28S, 18S, and 5.8S) and internal transcribed spacers (ITS1 and ITS2) as well as the cytochrome oxidase 1 (COX1) region of the mitochondrial DNA (mtdna) have been investigated in an attempt to better understand the taxonomic relationships within the X. americanum group. Sequence data from the 28S D2/D3 expansion region of the rdna revealed very little intraspecific variation among X. americanum group populations, including virus-transmitting X. tarjanense, from Florida (Gozel et al., 2006). The 18S region of the rdna did not differentiate the virusvectoring X. americanum, X. rivesi, and X. tarjanense (Lazarova et al., 2006) or several other non-virus-vectoring X. americanum group members (Oliveira et al., 2004). It appears that the ITS region of the rdna may be more informative. Restriction length polymorphisms in a region spanning the rdna 5.8S gene and ITS regions arranged 16 X. americanum group populations into five subgroups (Vrain et al., 1992). However, these subgroups did not group virus-transmitting populations together, with X. bricolensis and X. rivesi being found in different phylogenetically supported subgroups. Sequence data from the ITS1 region also identified subgroups placing X. americanum group populations together but separate from X. italiae and X. diversicaudatum, both of which are also virus-transmitting nematodes (Ye et al., 2004). Also of interest is the quickly evolving area of research into mtdna. Examination of the mtdna COX1 region revealed that a dichotomy exists between X. americanum group populations from North America and those from Asia, South America, and Oceania (Lazarova et al., 2006). The main diagnostic features of the members of Trichodoridae used to differentiate genera are in females (1) the reproductive system, (2) length of vagina, (3) development of vaginal sclerotized pieces, and (4) presence of advulvar lateral body pores; and in males (1) presence or absence of winglike extensions situated on or near the tail, (2) degree of development of copulatory muscles, and (3) development of spicule suspensor muscles (Decraemer and Baujard, 1998; Decraemer and Robbins, 2007). Within the members of Trichodoridae, only females having two gonads are known to transmit viruses (Decraemer and Robbins, 2007). Unlike the members of Longidoridae, which have life cycles that range from a few months to years, trichodorid nematode species are capable of multiplying rapidly at suitable temperatures and on good hosts (Taylor and Brown, 1997). The genera Trichodorus and Paratrichodorus are phylogenetically distinct on the basis of 18S rdna sequences (Boutsika et al., 2004a,b; Duarte et al., 2010). The distribution and association of the members of Trichodoridae with tobraviruses are of interest. Vector species and viruses are present in nearly all European countries, while the presence of vector species in North America is restricted and is apparently the result of introduction with infected plant material (Decraemer and Robbins, 2007). Eight species of Paratrichodorus and four species of Trichodorus (Table 26.2) transmit viruses. Of the Paratrichodorus species that transmit viruses, on the basis of rdna 18S sequence data, P. allius and P. teres formed a subcluster distinct from populations of P. anemones, P. hispanus, Fig. 26.1. Xiphinema index. (Courtesy S. A. MacFarlane APS) Fig. 26.2. Paratrichodorus allius. Inset, the head. (Courtesy K. J. Merrifield, from the files of Oregon State University Used by permission) and P. pachydermus in Portugal (Duarte et al., 2010). This difference may be the result of genetic isolation and local biotope adaptation of the populations and could reflect interspecific variability. The nematode feeding process Many different viruses are able to infect the root systems of plants, and many different nematode species feed on the roots of plants that are infected with viruses. Nevertheless, in most of these instances, even though the nematode has ingested virus particles, it is not able to pass the virus to other plants that it later feeds on. This demonstrates that there is specificity in the interaction between a particular virus and its nematode vector; some progress has been made in understanding the mechanisms involved in these interactions. Uptake (acquisition) of virus. Plant-parasitic nematodes can be divided into two groups on the basis of their feeding habits. Endoparasitic nematodes, such as the cyst nematodes (Heteroderidae), burrow into plant roots, where they become

368 Chapter 26 immobile and establish highly metabolically active feeding sites that function to direct plant nutrients to the nematodes. In contrast, the virus-vector nematodes are ectoparasitic, feeding from the outside of the root, most often targeting the region at or near to the root tip and in some instances inducing galls in this area after completion of the feeding process. Both longidorid and trichodorid nematodes feed by using a spear-shaped structure, located within the esophageal region at the anterior end of the body, to puncture the plant cell wall, allowing the cell contents to be extracted. In longidorid nematodes, the spear-shaped structure is referred to as the odontostyle. It is hollow with a narrow slit running along its length. During feeding, the odontostyle is pushed forward by protractor muscles. The nematode may repeatedly thrust the protracted odontostyle through the root surface into epidermal cells and sometimes into internally located vascular tissue cells. Nematode secretions are injected into the cell through the odontostyle, followed immediately by ingestion of the cell contents. Feeding can last from only a few minutes to up to several hours, and only at the end of feeding is the odontostyle withdrawn from the plant and retracted into the nematode esophagus. The process for trichodorid nematodes is slightly different. Here, the spear-shaped structure is referred to as the onchiostyle. It is solid rather than hollow and shorter than the longidorid odontostyle, so trichodorid nematodes feed only on epidermal cells. After penetration of the cell wall, a feeding tube consisting probably of nematode esophageal gland secretions forms around the onchiostyle. The cell contents are liquefied and then drawn into the nematode esophagus through the feeding tube. The nematode then pulls away from the root, leaving the feeding tube attached, although the tube becomes sealed, stopping further leakage of the cell contents. Virus particles are included in the liquefied cell contents, and unless they are trapped by being bound to the esophageal surface, they pass into the nematode intestine, where they are likely to be digested by proteases or excreted in the feces (McNamara, 1978). Retention and release of virus. In some cases, there are specific associations between particular nematode species and certain viruses that lead to transmission of the viruses to healthy plants. In these associations, virus particles are physically bound (i.e., retained ) at specific locations on the surfaces of the esophageal mouthparts. In Longidorus spp., virus particles bind primarily to the inner surface of the odontostyle, whereas in Xiphinema spp. and trichodorid nematodes, virus particles bind to the esophageal lining and not to the odontostyle or onchiostyle, respectively (Taylor and Robertson, 1970a,b). For transmission to occur, the retained virus must be released from the site of retention in the nematode. This is thought to occur as a result of nematode gland secretions that wash over the virus particles during feeding. However, no chemical or molecular details of the retention and release processes are known. It is suspected that in some cases the release process is more important for the success of transmission than is the retention process. For example, both Scottish (S) and English (E) strains of RpRSV were retained in Longidorus macrosoma. However, only the E strain was transmitted by this nematode (Trudgill and Brown, 1978). Another interesting finding is that the species of host plant can influence transmission. The nepovirus Grapevine fanleaf virus (GFLV) can be acquired by X. index from the roots of both grapevine and Chenopodium quinoa. However, this nematode transmits the virus only to grapevine (Trudgill and Brown, 1980) (Figs. 26.3 26.5). Both juvenile and adult nematodes are capable of transmitting viruses. However, because the internal and external cuticle (of ectodermic origin) is lost during the molt between developmental stages, any retained virus is also lost and has to be reacquired during subsequent feeding events for transmission to reoccur. Nevertheless, X. index larvae and adults are able to retain GFLV for at least 4 years in the absence of host plants (Demangeat et al., 2005), revealing an extremely long survival and viral-retention period within the nematode vector. Similar data have been obtained for X. americanum (McGuire, 1973) and X. rivesi (Bitterlin and Gonsalves, 1987) in the transmission processes of, respectively, Tobacco ringspot virus (TRSV) and Tomato ringspot virus (ToRSV). There is no Fig. 26.3. Fanleaf symptoms on Sylvaner grapes in Alsace caused by Grapevine fanleaf virus. (Courtesy O. Lemaire APS) Fig. 26.4. Grape cluster symptoms caused by Grapevine fanleaf virus. (Courtesy O. Lemaire APS)

Nematode-Borne Plant Viruses 369 nontransmissible strains were found only at locations toward the posterior of the pharynx, such that if released by secretions from any of the pharyngeal glands they would pass backward into the intestine rather than forward into any plant that was being fed upon. As in the situation described above for the nepovirus RpRSV and L. macrosoma, this suggests that release of retained particles rather than retention is the most important phase of the transmission process. Fig. 26.5. Grape leaf deformation caused by Grapevine fanleaf virus. (Courtesy O. Lemaire APS) cellular uptake of virus particles by the nematodes, so the viruses are not passed on to subsequent nematode generations through the eggs. Some studies have reported low-level, apparently nonspecific transmission of viruses (Allen and Ebsary, 1988), which might result from virus particles that are present in the esophagus but not attached to the cuticular surface (Taylor and Robertson, 1969, 1975). Specificity in the nematode virus interaction Specificity is revealed when one virus or virus isolate is found to be transmitted by only one or a few nematode species or, conversely, when one nematode species is found to transmit only one or a few viruses or virus isolates. In a study of nepoviruses, for example, L. elongatus transmitted both Tomato black ring virus (TBRV) and RpRSV, although it transmitted the S strain of RpRSV more efficiently than the E strain (Taylor and Murant, 1969). In contrast, L. macrosoma transmitted the E strain of RpRSV but did not transmit the S strain (Harrison, 1964). In another study, X. diversicaudatum transmitted Arabis mosaic virus (ArMV) and an E strain of Strawberry latent ringspot virus (SLRSV) but not two Italian SLRSV strains (Brown and Trudgill, 1983). Experiments with tobraviruses have concentrated on TRV, for which many isolates exist with different serological reactions (i.e., with a large variation in the amino acid sequences of the CP). Using viruliferous nematodes collected from the field, Ploeg et al. (1992) found that some species of nematodes appeared to transmit viruses of only one serotype, whereas other species could transmit TRV of two or more serotypes. However, in other experiments, a single nematode species, P. teres, was able to transmit two serologically different strains of TRV but was not able to transmit two other strains that were serologically the same as one of the transmitted strains (Ploeg et al., 1996). This suggests that the CP is not the only determinant of successful transmission, as was confirmed by MacFarlane et al. (1995), who replaced the CP gene from a nontransmissible PEBV strain with that from a nematode-transmissible TRV strain. An electron microscopy study of TRV particles retained in the mouthparts of feeding trichodorid nematodes (Paratrichodorus anemones, P. pachydermus, P. hispanus, and Trichodorus primitivus) showed that both transmissible and nontransmissible strains could be retained in different regions of the pharyngeal tract (Karanastasi and Brown, 2004). However, particles of the Identification and study of virus proteins involved in nematode transmission The small genomes of nepoviruses and tobraviruses have made them accessible to detailed molecular study and genetic manipulation. In early studies, by separating the two genomic RNAs from different strains of each type of nepovirus or tobravirus using gel electrophoresis or gradient centrifugation techniques and then mixing the RNAs in different combinations, researchers were able to produce pseudorecombinant isolates in which the two virus RNAs were derived from different strains possessing different transmission characteristics. With this approach, it was shown that for both nepoviruses and tobraviruses, nematode-mediated transmission is conferred by RNA2 (Harrison et al., 1974; Ploeg et al., 1993a). More recently, full-length viral cdnas of tobraviruses and some nepoviruses have been cloned that can be used to initiate infections in host plants (Belin et al., 1999; Hernández et al., 1997; MacFarlane et al., 1996). Site-directed mutagenesis of these cdnas has made it possible to identify particular virus genes and their protein products that are specifically involved in the nematode transmission process. Tobravirus proteins. RNA1 is about 6.8 kb and carries four genes encoding proteins involved in virus replication, cell-tocell movement, and suppression of RNA silencing. Interestingly, RNA1 by itself is able to infect plants, moving systemically through the vascular system, even though, in this form, the virus does not produce any CP and cannot form virus particles (Swanson et al., 2002). An RNA1-only infection is referred to as an NM infection and occurs quite frequently in some potato cultivars, such as Pentland Crown and Pentland Hawk (Harrison et al., 1983) (Fig. 26.6). NM infections are thought perhaps to arise following the delivery by nematodes of a limited number of virus particles to the plant root system, so that the long and short particles (carrying RNA1 and RNA2, respectively) become separated in the plant. It is expected that virus consisting only of unencapsidated RNA1 cannot be nematode transmitted and that the spread of such an infection through the soil is unlikely. The RNA2 molecules of different tobravirus isolates vary considerably in size (1.8 3.9 kb), which is the result of a highly variable gene content of the different isolates (MacFarlane, 1999). Many isolates have undergone recombination in which the central and 3ʹ terminal regions of RNA2 have been replaced by similarly located regions from RNA1. The extent of sequence exchange varies among isolates, but the general outcome of such recombination is that some or all of the RNA2-encoded genes involved in nematode transmission are deleted, making the virus nontransmissible. It is thought that propagation of tobraviruses in the glasshouse by mechanical inoculation encourages the selection of recombinant isolates, since the need for nematode transmission to spread the virus is removed. However, the TRV isolate PaY4, which has very little glasshouse propagation, is nevertheless a recombinant virus, but in a study by Vassilakos

370 Chapter 26 et al. (2001), deletion of nematode transmission genes did not occur and the virus could still be transmitted by nematodes. The RNA2 structure of nematode-transmissible tobravirus isolates can be divided into two groups. Each group has the virus CP gene at the 5ʹ proximal position followed by two additional genes encoding the 2b and 2c proteins. The second group has an additional gene encoding a putative small (9 kda molecular weight) protein located between the CP and 2b genes. The first group of isolates includes TRV PpK20 and TRV PaY4, and the second group includes TRV TpO1 and PEBV TpA56 (Fig. 26.7). The CP and 2b protein of different tobravirus isolates have recognizable amino acid sequence homology. However, the 2c proteins from most isolates are very different in sequence. Mutation of the 2b gene to introduce a frameshift revealed that this protein is required for the transmission of TRV isolates PpK20 (by P. pachydermus) and PaY4 (by P. pachydermus and P. anemones) and for the transmission of PEBV TpA56 (by T. primitivus) (Hernández et al., 1997; MacFarlane et al., 1996; Vassilakos et al., 2001). Mutation of the 2c gene showed that this protein is Fig. 26.6. External skin blemish (upper image) and internal spraing symptoms (lower images) in Pentland Dell potato caused by Tobacco rattle virus. (Courtesy S. A. MacFarlane APS) required for the transmission of PEBV TpA56 (by T. primitivus) but not TRV PpK20 (by P. pachydermus) or PaY4 (by P. pachydermus and P. anemones). Mutation of the PEBV TpA56 putative gene encoding the 9-kDa protein also significantly reduced transmission of this virus by T. primitivus, but, unlike the 2b and 2c proteins, the putative 9-kDa protein was not detected in infected plants (Schmitt et al., 1998). The CP and 2b protein have been shown to interact with one another in the yeast two-hybrid (Y2H) assay system (Holeva and MacFarlane, 2006; Visser and Bol, 1999), and using electron microscopy and immunogold labeling, Vellios et al. (2002) found that the 2b protein associates with virus particles. This has led to the hypothesis that the 2b protein acts as a bridge that might link the virus particles to retention sites on the surface of the nematode esophagus, although no studies have been done to try to detect the 2b protein in viruliferous nematodes. The virus CP has a disordered region at the C-terminus that is necessary for both virus transmission and interaction with the 2b protein (Brierley et al., 1993; Holeva and MacFarlane, 2006; MacFarlane et al., 1996). Release of virus particles from the nematode esophageal cuticle possibly involves nematode gland secretions that cause either a change in ionic conditions around the virus particles or perhaps carry out a proteolytic cleavage of the associated CP and 2b protein. The flexible C-terminus of the CP might be a target for such a release mechanism, although there are no data to support these hypotheses. The interaction between the CP and 2b protein appears to be specific, since in the Y2H system the CP from the TRV PpK20 strain interacted with the PpK20 2b protein but not with the PaY4 strain 2b protein. Similarly, the PaY4 strain CP interacted with the PaY4 2b protein but not with the PpK20 2b protein (Holeva and MacFarlane, 2006). Using TRV infectious clones, Vellios et al. (2002) found that the PaY4 2b protein was detectable in infected plants by western blot when the virus encoded both the PaY4 CP and PaY4 2b genes. However, when the virus encoded the PpK20 CP gene and PaY4 2b gene, the 2b protein was not detectable, suggesting that the CP 2b interaction might stabilize the 2b protein. Also, a 2b mutant of TRV PaY4 was transmitted by P. pachydermus when coinoculated to plants with a wild-type (nonmutated) TRV PaY4, showing that the 2b protein functions in trans (Vassilakos et al., 2001). However, the PaY4 2b mutant was not transmitted when coinoculated with TRV PpK20, even though both of these strains can be transmitted by P. pachydermus, underscoring the finding that strain specificity is involved in the interaction between the CP and 2b protein. The mechanism by which the 2c protein plays a role in the transmission of some tobravirus isolates is completely unknown, although the PEBV 2c protein was detected in infected plants (Schmitt et al., 1998), and using the Y2H system, Visser and Bol (1999) found that the TRV PpK20 2c protein interacted with the PpK20 CP. Nepovirus proteins. Early work comparing the transmission of pseudorecombinants constructed between S (efficiently transmitted) and E (poorly transmitted) strains of RpRSV demonstrated that RNA2 has a major influence on nepovirus transmission by nematodes (Harrison et al., 1974). In a second study, examination of pseudorecombinants of two antigenically different isolates of TBRV confirmed that the CP serotype, which was determined by RNA2, influences nematode transmissibility (Harrison and Murant, 1977b). Subsequent sequencing studies confirmed that RNA2 of GFLV (and other nepoviruses) encodes a polyprotein that is cleaved by a virus-encoded prote-

Nematode-Borne Plant Viruses 371 ase to produce three proteins: the 2A replication helper protein, the 2B virus movement protein, and the 2C virus CP (Serghini et al., 1990) (Fig. 26.8). A deeper understanding of some of the molecular aspects of the nematode transmission of nepoviruses has been obtained by studying the transmission of GFLV by X. index and by comparing GFLV with ArMV, which is genetically similar to GFLV but is not transmitted by X. index. The GFLV ArMV X. index transmission system was examined in an extensive series of studies (Andret-Link et al., 2004a; Belin et al., 1999, 2001; Fig. 26.7. Genome diagram of Tobacco rattle virus (TRV) RNA1 and RNA2 from three different TRV strains. The boxes represent the different genes encoded by each RNA. The asterisk in RNA1 denotes the leaky translation termination codon that directs the production of either the 134K or 194K replicase proteins. (Courtesy S. A. MacFarlane APS) Fig. 26.8. Genome diagram of Grapevine fanleaf virus RNA1 and RNA2. The vertical lines within the boxes denote the protease cleavage sites used for processing of the two viral polyproteins into individual smaller proteins. (Courtesy S. A. MacFarlane APS)

372 Chapter 26 Schellenberger et al., 2010, 2011). Infectious cdna clones of RNA2 of both GFLV and ArMV were constructed, and a series of recombinants was created for which the analogous ArMV RNA2 regions replaced the individual genes or parts of genes encoded by the GFLV RNA2. Difficulties were encountered because the protease cleavage sequences that separate the three virus proteins differ slightly between these two viruses and because the C-terminal part of the 2B protein and CP interact to allow virus movement. A similar situation has been observed with ArMV, so that chimeric GFLV/ArMV viruses harboring an ArMV CP are transmitted by X. diversicaudatum but not by X. index. The results of these studies demonstrate that, for both GFLV and ArMV, the CP is the sole determinant of nematode transmission. At present, similar work has not been done with other nepoviruses. Subsequent studies were conducted to identify the precise domains within the CP that are involved in transmission. Schellenberger et al. (2010) used 3D modeling based on the crystal structure of TRSV (Chandrasekar and Johnson, 1998) to identify a sequence of 11 amino acids within a loop of the B domain of the CP that differed between GFLV and ArMV. Transmission of GFLV stopped when the ArMV counterpart replaced the region. With the unraveling of the crystal structure of the GFLV CP, major progress has been made in discovering the probable domain that interacts with the nematode. This domain is composed of a positively charged pocket, whereas the main outer surface of the CP has a negative charge, surrounded by three loops (GH, BC, and CʹCʹʹ), located between the threefold and the fivefold axes of the virion. Moreover, a single mutation, Gly297 Asp, in the exposed GH loop was sufficient to dramatically lower the nematode transmission efficiency of the virus (Schellenberger et al., 2011). The nature of the side chains occupying the loops may be decisive for virus retention by the vector, given that a single mutation may modify the whole charge of the particle, since the single additional negative charge conferred by the glycine-to-aspartic acid replacement at residue 297 is multiplied by the assembly of 60 copies of the CP to form the complete virus particle. Electrostatic perturbations of the putative nematode-binding pocket and of the surrounding edges may affect the interaction of the virus particle with a specific receptor in the cuticle lining of the anterior part of the feeding apparatus of X. index. These findings show that it may be possible to decipher at the atomic level the mechanism of interaction between the viral transmission ligand (CP loop domain) and the putative cuticular receptor of the nematode, enabling the development of new control strategies aimed at breaking or inhibiting this specific linkage. Control of Nematode- Transmitted Viruses Strategies to control crop diseases caused by nematode-transmitted viruses can be directed at either the vector nematode or the virus. Difficulties arise because the virus-vectoring nematodes are protected in the soil and cannot be treated directly, have wide host ranges, and can survive in soil for long periods. Similarly, if the virus is targeted, difficulties arise because of their wide host ranges, which include many weed species. In addition, the ability of the viruses to spread via infected seed, pollen, and plant-propagation material and the natural diversity in the species and strains of nematode-transmitted viruses make management difficult. Nevertheless, a number of different approaches to nematode and virus control have been taken. Historically, practical control of nematodes has involved the application to the soil of nematicidal or nematistatic chemical treatments. However, many of the most effective fumigant and postplant nematicides have either been banned or their use has become highly regulated (European Commission, 2003, 2007; Zasada et al., 2010). While many nematicides have been shown to control nematodes that transmit viruses (Bileva et al., 2009; Hwang et al., 2010; Ingham et al., 2007), future use will depend upon the regulatory status of each nematicide in any given location. Management practices other than the use of chemicals will be relied upon heavily in the future. Alternative control measures based on exclusion, genetic resistance, biological control, and cultural practices require an extensive knowledge of nematode and virus biology to achieve satisfactory results. Specific information about the nematode and virus, including accurate identification, host range, life cycle, survival strategies, and persistence, will be needed to effectively manage this disease complex. Exclusion The easiest way to control nematodes and their associated viruses is to prevent their introduction; this applies at the international, national, and field levels. Mechanisms are in place worldwide to ensure the supply and use of nematode- and virusfree planting material. International movement of planting material is regulated through the Food and Agriculture Organization of the United Nations. These international laws empower member countries to exclude the entry of pathogens of quarantine significance, the enactment of which is the responsibility of the national plant-protection organization in each country. Examples of successful nematode- and virus-free certification programs include those operated by the California Department of Agriculture (CDFA) and by the European Union (Andret- Link et al., 2004b; Anonymous, 1994; Martelli, 2006). CDFA has specific soil treatment and handling procedures to ensure nematode cleanliness of both field- and container-grown nursery stock (Zasada et al., 2010). Detection and identification Proper nematode identification and the ability to detect viruses in nematodes are, and will continue to be, key components of disease management. These are areas in the study of nematode-transmitted viruses that have advanced significantly since 2000. Recent advancements in nematode identification have occurred because the traditional morphological techniques are laborious and time consuming and require an experienced taxonomist. Identification can also be confounded by the existence of mixed populations, in which some nematodes are virus vectors and others are not. Holeva et al. (2006) outlined the following criteria for a reliable preplant soil test for determining virus-transmitting nematodes. It should be (1) rapid and robust so that technology can be transferred among laboratories; (2) reproducible within and among samples; and (3) sensitive enough to detect the target nematode species and viruses at predetermined threshold levels. Species-specific primers located in the rdna 18S and/or ITS1 regions were developed to distinguish Paratrichodorus macrostylus, P. pachydermus, T. primitivus, and T. similis (Boutsika et al., 2004a,b)

Nematode-Borne Plant Viruses 373 and P. allius and P. teres (Riga et al., 2007). Currently, no molecular diagnostic tools are available to definitively differentiate populations within the X. americanum group. However, species-specific primers for Longidorus helveticus, L. profundorum, and L. sturhani (Hubschen et al., 2004a) and X. diversicaudatum, X. index, and X. vuittenezi (Hubschen et al., 2004b) have been developed. A multiplex PCR, in which primers located in the rdna ITS1 region are used, reliably detected two of four mixed species of X. diversicaudatum, X. index, X. italiae, and X. vuittenezi (Wang et al., 2003). It would also be highly desirable to be able to rapidly and efficiently detect the presence of viruses in collected nematodes. Such information would indicate when management is necessary and could potentially reduce the amount of field area that is currently treated. Prior to the development of molecular diagnostic tools, time-consuming transmission experiments with bait plants in the greenhouse were necessary (Taylor and Brown, 1997). Detection of TRSV by fluorescent antibodies was achieved in single specimens of X. americanum (Wang and Gergerich, 1998; Wang et al., 2002). Enzyme-linked immunosorbent assay has also been used for the detection of GFLV in X. index (Bouquet, 1983). However, this method requires the isolation and coextraction of many nematodes to produce sufficient material for successful testing. Molecular techniques such as reverse transcriptase polymerase chain reaction (RT- PCR) and real-time PCR have allowed for the rapid and sensitive detection of viruses in single nematodes. RT-PCR methods for the detection of GFLV in X. index (Demangeat et al., 2004; Esmenjaud et al., 1994), ToRSV and TRSV in X. americanum (Martin et al., 2009), and TRV in several trichodorid nematodes (Boutsika et al., 2004b; Riga et al., 2009) have been developed. A qualitative and quantitative RT-PCR assay was developed that detected P. pachydermus and T. similis as well as the associated TRV (Holeva et al., 2006). Cultural control The efficacy of cultural practices (e.g., rotation, fallow, cover crops, and tillage) to manage nematodes and viruses depends upon the target organisms. The cultural control of nematodes and viruses will likely require a combination of these approaches with other management strategies. Because several virus-transmitting nematodes are polyphagous, weed control is essential in any type of management practice (Duffus, 1971; Murant, 1970). In the U.S. state of Washington, several weeds, including kochia, prickly lettuce, henbit, nightshade, common chickweed, and annual sow thistle, were shown to be hosts of P. allius and TRV (Boydston et al., 2008; Mojtahedi et al., 2002). Many broadleaf weeds are hosts of X. americanum (Powell et al., 1982), and dispersal of ToRSV is aided by the fact that the virus is transmitted by the pollen and seeds of many of its weed hosts (Brunt et al., 1996). In Europe, special attention is given to the risk of importing ToRSV in American grapevine material, since one of its vectors, X. rivesi, is present in Slovenia (Širca et al., 2007). Rotation away from nematode- and virus-susceptible plants to other crops, including cash and cover crops, may or may not be a viable nematode management option, depending upon the nematode and virus in question. Rotation to tall fescue (Festuca arundinacea) prevented the reoccurrence of ToRSV, although population densities of X. americanum remained high (Pinkerton and Martin, 2006). In the same study, rotation with rapeseed (Brassica napus) was as effective as fumigation with methyl bromide in preventing reinfection of raspberry by ToRSV for 3 years. Two years of continuous corn, grain sorghum, or wheat followed by a soybean cultivar resistant to Soybean severe stunt virus reduced X. americanum population densities and the severity of disease (Evans et al., 2007). Conversely, wheat and corn served as inoculum reservoirs of TRV. Therefore, these crops would not be good rotation crops with potato (Mojtahedi et al., 2002). Field experiments in which mainly members of Fabaceae were used as rotation crops in association with other strategies have been evaluated in Bordeaux and Alsatian vineyards to validate this practical approach for viticulture under different agroclimatic conditions, with the aim of restricting fanleaf disease to below harmful levels (Lemaire et al., 2010). In addition, some soils are suppressive for nematodes. For example, the ectoparasitic nematode Helicotylenchus dihystera was suppressed in soil that had a high organic matter content, a low ph, and an altered bacterial community (Rimé et al., 2003). Natural resistance to vector nematodes and their viruses The deployment of resistant crop plants may be the most effective and economical means of controlling plant-parasitic nematodes. This is especially true for long-lived perennials that are affected by nematode-transmitted viruses. Trichodorids and tobraviruses. High levels of resistance to TRV are known for a number of potato cultivars and breeding lines (Borejko, 2001; Brown et al., 2000; Dale, 1989). TRV resistance has been available for many years and is not uncommon in European-type cultivars; however, few resistant cultivars are accepted by the U.S. potato industry (Khu et al., 2008). This situation is complicated by the fact that potatoes can react to TRV in three ways. Some cultivars are immune to infection, and this form of resistance is considered to exist in cultivars such as Climax, Record, Saturna, Bintje, and Arran Pilot. However, a resistance-breaking TRV isolate was recovered from Bintje (Robinson, 2004). Another group of cultivars reacts to TRV by formation of discolored, corky tissue in the affected tubers referred to as spraing or corky ringspot disease (Fig. 26.6) that makes the tubers unmarketable. However, formation of spraing, which may be a hypersensitive response by the plant, does not necessarily prevent the virus from further infecting the plant. A major quantitative trait locus for lack of spraing reaction was located on potato chromosome IX with flanking amplified fragment length polymorphism markers, which will enable marker-assisted selection (Khu et al., 2008). However, this lack of response may not indicate true resistance. Potato cultivars in the third group are fully susceptible to TRV and do not produce visible spraing symptoms. However, prolonged propagation of infected tubers results in significant reduction in tuber size and uniformity (Dale et al., 2004; Xenophontos et al., 1998). For trichodorid nematodes, studies have been done to assess resistance in various crops (Timper et al., 2007), although dominant major resistance genes have not been reported. Longidorids and nepoviruses. Natural dominant resistance genes capable of triggering a hypersensitive response or extreme resistance against GFLV or ArMV have not been found in wild or cultivated grapevines (Lahogue and Boulard, 1996). Therefore, conventional breeding for virus resistance with dominant genes cannot be developed. Alternative forms of potentially useful host plant resistance to GFLV were identified in some Vitis vinifera accessions, including a wild V. vinifera accession

374 Chapter 26 from Afghanistan, V. rotundifolia cv. Bountiful, and a V. vinifera V. rotundifolia hybrid (Harris, 1983; Walker et al., 1985). Resistance to X. index has also been identified in several Vitis spp., with high levels of resistance occurring in Muscadinia (Vitis) rotundifolia (Coiro et al., 1990; Esmenjaud et al., 2010; Staudt and Weischer, 1992). However, M. rotundifolia is not suitable as a rootstock because of its graft incompatibility with V. vinifera, its poor rooting, and its high susceptibility to limeinduced chlorosis (Bouquet, 1981). Partially nematode-resistant grapevine hybrids have been obtained, but they do not totally block transmission of virus to their rootlets (Esmenjaud and Bouquet, 2009). Other sources of resistance to X. index have been characterized in Vitis arizonica. A nucleotide binding, leucine-rich repeat sequence colocalizing with the XiR1 gene (X. index Resistance 1) and mapped on chromosome 19 has been identified as the first known dominant resistance gene against an ectoparasitic nematode (Hwang et al., 2010; Xu et al., 2008). This offers new material for breeding of grape rootstocks that are resistant to nematode-vectored GFLV. Transgenic resistance Transgenes that confer resistance to viruses or nematodes may either be engineered, for example, by utilizing pieces of the virus genome to stimulate the plant RNA-silencing mechanism, or be naturally occurring resistance genes that are moved from one species of plant into the crop that is to be protected. Tobraviruses. Studies to assess the effectiveness of genetically engineered resistance to tobraviruses have been conducted. Tobacco plants transformed with the CP genes of TRV isolate TCM or PLB were found to be resistant when inoculated with purified virus particles from the same isolate but not when inoculated with the alternative isolate or when inoculated with purified viral RNA (Angenent et al., 1990; Van Dun and Bol, 1988). The mechanism for this resistance is not known and was not effective against virus transmitted to the roots by nematodes (Ploeg et al., 1993b). Subsequently, transgenic Nicotiana benthamiana plants were produced that expressed part of the PEBV replicase gene encoding the RNA-dependent RNA polymerase domain (54K RDRP domain) (MacFarlane and Davies, 1992). These plants were resistant to mechanical inoculation with PEBV but were not challenged by nematode-mediated inoculation. Further studies suggested that the resistance in these plants operated by an RNA-silencing mechanism (van den Boogaart et al., 2001). More recently, this approach has been used with potato (cultivar Matilda) and tobacco plants transformed with the 57K RDRP domain from TRV (Melander, 2006; Vassilakos et al., 2008). With the potatoes, two selected transgenic lines were found to show reduced spraing disease symptoms when challenged with nematodes carrying TRV. However, there was no apparent resistance to TRV in these plants (Melander, 2006). With the transgenic tobacco plants, various levels of resistance were found with different plant lines. However, one line was highly resistant though not immune to infection with the PpK20 isolate of TRV but not to a Greek isolate (Vassilakos et al., 2008). Resistance was strongest in older plants, and, when the virus was delivered to the plant roots either by mechanical inoculation or by nematodes, even though virus could be detected in the roots it did not move systemically into the aerial parts of the plant. These studies suggest that obtaining transgenic resistance in plant roots is more difficult than in leaves but that this may be a useful approach to combat nematode-transmitted viruses. Nepoviruses. Several studies have expressed viral genes in grapevines, with the aim of producing virus resistance by the induction of antivirus RNA silencing. These include the CP genes of nepoviruses such as GFLV (Bardonnet et al., 1994; Bouquet et al., 2003; Gambino et al., 2005; Gribaudo et al., 2005; Maghuly et al., 2006; Vigne et al., 2004) and ArMV (Bertioli et al., 1992; Golles et al., 2000; Reustle et al., 2005; Širca et al., 2007; Spielmann et al., 2000), the movement protein gene (Valat et al., 2006), and the VPg (genome-linked viral protein) gene (Lindbo et al., 1993). In addition, grapevines have been transformed with conserved sequences of GFLV, ArMV, and RpRSV assembled in inverted repeat constructs (Gambino et al., 2010; Reustle et al., 2005, 2006) in order to induce multiple virus resistance. Evaluating the effectiveness and durability of resistance of these transformed plants is an ongoing, long-term task. Natural transmission remains the most reliable way to evaluate transgenic lines for resistance. Dual in vitro cultures of viruliferous nematodes and grapevine plantlets have been assessed. This screening system saves space and time, and an inoculation access period of 6 weeks was sufficient to infect grapevines by this means (Winterhagen et al., 2007). The highly virus-susceptible model plant N. benthamiana has been used in several studies to assess RNA-silencing-induced resistance against nepoviruses (Jardak-Jamoussi et al., 2003; Reustle et al., 2006). In another approach, expression in transgenic plants of recombinant antibodies recognizing GFLV was used, and a single-chain antibody fragment recognizing the GFLV CP conferred resistance to GFLV and partial resistance to ArMV (Nölke et al., 2009). Public acceptance of transgenic resistance Currently in Europe, the general release of genetically modified plants into the field is not permitted. At INRA (National Institute for Agricultural Research) in Colmar, France, an extensive and proactive process of communication with growers, associations, and the public through debates and a local steering committee on the risks, benefits, and general acceptance of trials that include nepovirus-resistant transgenic grapevines has been initiated. The steering committee has since evolved into a forum for debate on different alternative strategies to fight fanleaf disease (Hemmer et al., 2009; Joly and Rip, 2007; Lemaire et al., 2010). Future Research The application of molecular biological techniques has greatly advanced our knowledge of the mechanisms involved in the nematode transmission of plant viruses as well as the taxonomy of virus-transmitting nematodes. However, the intrinsic challenges of working with a disease complex that operates in the soil remain. We can expect to learn more about how the structure of virus transmission-related proteins relates to their function, but linking this information to the vector nematode is not easy. A major limitation to the advance of this area of research is the difficulty of growing pure cultures of vector nematodes, especially of Trichodorus spp. However, isogenic lines of parthenogenetically multiplying Xiphinema spp. can be obtained and reared, with the exception of X. diversicaudatum (Demangeat et al., 2010; Wang et al., 2003). The most difficult methodologi-