Localizing viruses in their insect vectors.

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Localizing Viruses in their Insect Vectors Annu. Rev. Phytopathol. 2014.52. Downloaded from www.annualreviews.org by University of Waikato on 06/17/14. For personal use only.

Stéphane Blanc,∗ Martin Drucker, and Marilyne Uzest INRA, UMR BGPI, CIRAD-INRA-SupAgro, CIRAD TA-A54K, Campus International de Baillarguet, 34398 Montpellier Cedex 05, France; email: [email protected], [email protected], [email protected]

Annu. Rev. Phytopathol. 2014. 52:18.1–18.23

Keywords

The Annual Review of Phytopathology is online at phyto.annualreviews.org

plant virus, vector transmission, arbovirus, virus-host interaction, virus-vector interaction

This article’s doi: 10.1146/annurev-phyto-102313-045920 c 2014 by Annual Reviews. Copyright All rights reserved ∗

Corresponding author

Abstract The mechanisms and impacts of the transmission of plant viruses by insect vectors have been studied for more than a century. The virus route within the insect vector is amply documented in many cases, but the identity, the biochemical properties, and the structure of the actual molecules (or molecule domains) ensuring compatibility between them remain obscure. Increased efforts are required both to identify receptors of plant viruses at various sites in the vector body and to design competing compounds capable of hindering transmission. Recent trends in the field are opening questions on the diversity and sophistication of viral adaptations that optimize transmission, from the manipulation of plants and vectors ultimately increasing the chances of acquisition and inoculation, to specific “sensing” of the vector by the virus while still in the host plant and the subsequent transition to a transmissionenhanced state.

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INTRODUCTION Transmission from one host to another is a vital step in the life cycle of a virus, determining the survival of viruses in a given environment, the number of host species colonized, and viral incidence, and thus the extent to which a virus species directly or indirectly affects ecosystems. It is therefore not surprising that more than a century of effort has been directed at understanding the diversity of transmission strategies and elucidating the molecular mechanisms involved. In plants, most viruses are transmitted from host to host by an additional player that is able to create a breach in the plant cell wall and to move efficiently within the host community. This player, or vector, can be a fungus, a nematode, a mite, or a biting/chewing insect, but vectors are most often sap-feeding hemipteran insects.

Historical Description of Virus Routes Within the Vector Annu. Rev. Phytopathol. 2014.52. Downloaded from www.annualreviews.org by University of Waikato on 06/17/14. For personal use only.

Early work on virus transmission by hemipteran insects focused on the description of compatible virus-vector pairs and on quantitative parameters, such as the time required for virus acquisition by a vector on an infected plant, the time during which the infectious virus is retained, and the time required for efficient inoculation into a new healthy plant. Accordingly, three vector-transmission categories have been described (Figure 1): (a) nonpersistent (132), (b) persistent (132), and (c) intermediate semipersistent (116) transmission. Nonpersistent transmission takes place when the virus is acquired on an infected plant within seconds and needs to be immediately inoculated into a new host because it is not retained in an infectious form within the vector for more than a few minutes. In contrast, in persistent transmission, acquisition and inoculation times are much longer (hours to days), retention time is indefinite, and a so-called latent period of several hours to days post acquisition is required before efficient inoculation becomes possible. Further analysis, helped by the development of electron microscopy, provided qualitative clues distinguishing these transmission categories. Nonpersistent viruses were seen to restrict the reversible association with their insect vectors to the cuticle of the mouthparts or the foregut, whereas persistent viruses were found circulating across the gut, the hemolymph, and the salivary glands. The latent period could then be interpreted as the time required for persistent viruses to complete their cycle within the vector body. The more logical terms noncirculative and circulative were proposed to substitute for nonpersistent and persistent (63), as they better describe the actual route of the virus within its vector (Figure 1). Intermediate semipersistent viruses, with acquisition and inoculation periods of several minutes to hours and no requirement for a latent period prior to inoculation, associate with their vector externally, on the cuticle lining the mouthparts or the foregut, and are thus generally included in the noncirculative category (56). Titration of the virus within the vector body, incorporation of radioactive isotopes into viral nucleic acid, and quantification of viral genomes and/or coat protein (CP) demonstrated that some plant viruses can replicate within their insect vectors (56a). In contrast, some species of plant viruses can complete a circulative cycle through the gut cells into the hemolymph and through the salivary gland cells of their vectors without replicating. This distinguishes propagative and nonpropagative viruses in circulative/persistent viruses (Figure 1). All of the above data paved the way in this field of research by defining the main categories of virus-vector interactions and a terminology that is still in use today. Although defined for vectors in the group of hemipteran insects, this classification also applies to other virus-vector pairs involving biting-chewing insects, such as beetles, mites, and even nematodes (22).

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Common food/ salivary canal Figure 1 Different routes of plant viruses in their aphid-vectors. In the aphid graphic, the gut is represented in blue and the salivary glands and salivary duct in brown. The white arrows represent the cycle of circulative viruses ( yellow hexagons) within the aphid body, across the gut epithelium to the hemolymph and/or other organs, and ultimately to the salivary glands. Noncirculative (nonpersistent) viruses appear at their attachment sites at the tip of the stylets as red hexagons. Abbreviations: FG, foregut; HG, hindgut; MG, midgut. The inset at the bottom right represents the common food/salivary canal located at the tip of the aphid maxillary stylets. Noncirculative viruses interact with putative receptors embedded in the stylet cuticle. In the capsid strategy, viruses directly bind putative receptors via a domain of their capsid protein (for example, the genus Cucumovirus). , In the helper strategy, the virus-receptor binding is mediated by additional viral proteins that are designated as “helper components” (HCs; blue). Best-known cases are the genera Potyvirus (3; the HC is designated as HC-Pro) and Caulimovirus (2; the HC is designated as P2). Figure adapted with permission from References 16 and 55.

Molecular Interactions Between Virus and Vector Our view of virus-vector interactions has widened impressively during the past three decades. This period has witnessed the identification and biochemical characterization of viral proteins and complexes responsible for virus-vector compatibility. A novel distinction of transmission modes was defined by the discovery that some viruses can interact directly with vectors via their CP or envelope proteins, whereas others exploit an intermediate viral protein—a helper component (HC)—linking virus particles to attachment sites within vectors (Figure 1). Two molecular viral strategies—capsid and helper—have thus been proposed (99), the latter exclusively for noncirculative viruses.

www.annualreviews.org • Plant-Virus-Vector Interactions


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Ecological Impacts of Virus-Insect-Plant Interactions


More recently, research trends have focused on more integrative approaches and ecological considerations. It has become clear that viruses can manipulate their vectors, either directly or through modifications induced in the host plant (reviewed in 55). Virus infection can modify plant attractiveness or repellence (81) and/or augment the fitness of specific vector species (60). More directly, once inside the vector, viruses can also affect vector behavior in a way that increases opportunities for virus inoculation. For example, viruliferous thrips carrying Tomato spotted wilt virus (TSWV) spend more time using nondestructive cell feeding concomitant with augmented salivation—a feeding behavior favoring virus inoculation with the saliva (113). Another remarkable behavioral switch is that of aphids carrying Barley yellow dwarf virus (BYDV). Although initially attracted by infected plants, when aphids become viruliferous they turn their preference to healthy ones, clearly boosting virus dispersion in the host community (18, 59). These virus-related effects on insect vector populations are ecologically relevant (76). Beyond their evident impact on virus, insect, and plant communities, they support large-scale biological invasion by insect species boosted by the virus, at the expense of other species whose fitness does not increase on infected plants (60, 94).

Scope of the Review The mechanisms of virus-vector interactions and their implications on a broader ecological scale have both been reviewed extensively in the past five years (11, 14, 16, 22, 26, 55, 57, 76, 114, 143). Here, we restrict our focus to the precise localization of the virus within both the vector and the host plant, as these localizations govern efficient acquisition, retention, and inoculation. Targeting the transmission step has long been seen as a possible alternative to classical measures for controlling viruses (22), but development of such novel control strategies requires conquering of technical and conceptual bottlenecks. On the one hand, the concept of blocking interaction between the virus and its receptor within the vector is simple (66), but technical hindrances conspire to keep these receptor molecules hidden, and none have been definitively identified to date. On the other hand, the idea that virus-vector interactions can be blocked only in the vector is restrictive, and the possibility of blocking the virus at sites nonoptimal for uptake by the vector can now be envisaged (78). Here, we review the localization of viruses within insect vectors, covering the most recent progress as well as ongoing limitations in the search for viral receptors. We also discuss the novel concept of vector-sensing by viruses, which induces a transient positioning of viral transmissible units within specific host sites, facilitating acquisition and transmission.

VIRAL DETERMINANTS OF VECTOR TRANSMISSION Viral proteins (ligands) regulating the specificity of vector transmission are of prime technical/ practical interest as useful probes in the search for attachment sites and viral receptors within vectors, and ultimately as putative competing agents capable of impairing transmission.

Enveloped Viruses Enveloped viruses infecting plants are found in the families Bunyaviridae and Rhabdoviridae. Among bunyaviruses, the only genus with member species that infect plants is Tospovirus. The type member species, TSWV, encodes two transmembrane glycoproteins (GN and GC ), which are probably 18.4

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the first viral compounds to contact the thrips vector Frankliniella occidentalis (reviewed in 57). Repeated passage in plants by mechanical inoculation induces the accumulation of mutations in these glycoproteins, with concomitant loss of insect transmission (91). Consistently, a point mutation in the GN /GC open reading frame suppressed thrips transmission without affecting plant infection (111), and a GN protein produced in vitro inhibited TSWV transmission when fed to thrips prior to transmission assays (137, 139). Rhabdoviruses have two genera described in plants, Cytorhabdovirus and Nucleorhabdovirus, differing in the site of maturation within infected cells [endoplasmic reticulum or nucleus, respectively (reviewed in (57)]. Although most plant virus genera are associated with a single group of insect vectors, a remarkable feature of plant rhabdoviruses is that different species can be transmitted by vectors as diverse as aphids, leafhoppers, planthoppers, and mites. In all cases, the first viral protein to contact the gut of the insect vector is likely the transmembrane glycoprotein G, which both recognizes the vector specifically and controls the viral endocytosis and membrane fusion, resulting in cell infection. Plant rhabdovirus transmission remains poorly studied, and current understanding of virus-vector interactions in this group is based largely on data from animal systems (see Reference 6 for further details).

Nonenveloped Virus: Capsid Strategy Numerous viruses lacking a membranous envelope can interact directly with their insect vectors via specific domains on their CP. A simple approach exploiting artificial feeding of insect vectors through stretched parafilm membranes (83, 98) allowed successful transmission of purified virus particles, whereas an alternative and complementary approach using synthetic variants encapsidating the genome of one strain (and/or species) with the CP of another showed that it is the CP that determines transmission by the vector species (29, 32, 62). Thus, there is convincing evidence of the sole involvement of CPs in transmission for a number of noncirculative [e.g., Cucumovirus (32, 47, 62), Alfamovirus (83), and Carlavirus (133)] and circulative [e.g., Luteoviridae (25, 28, 53, 143) and Geminiviridae (29)] viruses. A similar capsid-based strategy is also implicated in poorly characterized cases of virus transmission by beetles (48, 84). Detailed information on the virus CP subdomains that bind directly to the vector is available for only a handful of viral species. Solving the atomic structure of the Cucumber mosaic virus (CMV) particle has revealed the metal ion–binding βH-βI loop exposed at the surface of the virion (72, 112). Directed mutagenesis at various positions within this loop generated nontransmissible mutants without obvious structural alteration of the particle (72). Charge alterations in the βHβI loop of these mutants are thought to be responsible for the disrupted interaction between CMV and its aphid vector. In various species of the family Luteoviridae, both the CP and an extension of this protein resulting from a conditional read-through of the stop codon appear to define compatibility between virus and aphid vectors (23, 27). Disentangling the respective roles of the CP and the read-through protein proved difficult and would benefit greatly from structural data on the conformation of mature virions (26). The best-described plant reovirus species is Rice dwarf virus (RDV) (reviewed in 57), for which the atomic structure of the mature virion has been solved (61, 93). The outer capsid protein P2, which protrudes from the surface of the outer shell of virions, is responsible for initial contact and infection of the leafhopper vector (135). Indeed, both mutations in P2 and chemical treatments removing it from the virion outer shell preclude attachment to, and infection of, gut cells (see citations in Reference 57). Nevertheless, despite the mounting number of cases in which the CP (or a CP component) has been shown to recognize the vector directly, the precise protein motifs binding to putative counterpart receptors in vectors remain extremely poorly characterized. www.annualreviews.org • Plant-Virus-Vector Interactions


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Nonenveloped Virus: Helper Strategy


The absence of transmission of purified virus particles from various viral species initially suggested that the virion itself was not sufficient for successful uptake, retention, and inoculation. The fact that transmission of purified particles could be rescued by prefeeding insect vectors onto plants infected with the corresponding virus indicated that a virus-induced HC was present in infected plants and that this HC could be acquired efficiently prior to virions. Originally developed in the seminal work of Govier & Kassanis (52) for the study of potyvirus transmission by aphids, this approach later demonstrated the involvement of HCs in the transmission of caulimoviruses by aphids (75) and of waika- and sequiviruses by leafhoppers and aphids (99), respectively (remarkably, all are noncirculative viruses). HC molecules have been characterized in depth only for potyviruses and caulimoviruses. The HC of potyviruses (HC-Pro) is a 50-kDa protein composed of two helix-rich domains, respectively located near the N- and C-terminal ends of the protein and linked by a hinge (54, 102, 109). In these domains, two conserved motifs, KITC (12) and PTK (96), are thought to bind to aphids and to the consensus DAG motif of the CP, respectively, reversibly bridging the two (15). In Cauliflower mosaic virus (CaMV), the type member of the genus Caulimovirus, the HC is an 18-kDa protein designated P2 (13, 142). The N-terminal region of P2 most probably binds to aphids (86), whereas the α-helices at the C terminus engage in coiled-coil interactions with α-helices of the N-terminal end of the virion-associated protein P3 (68, 103). Notably, an intriguing observation (46) suggests the involvement of an as yet unidentified and uncharacterized HC in the circulative transmission of the nanovirus Faba bean necrotic yellows virus (FBNYV). If confirmed, this would be the first report of an HC-mediated transmission for a circulative virus.

LOCALIZATION OF VIRUSES INSIDE THEIR INSECT VECTORS Noncirculative Viruses in the Stylets of Insect Vectors Sixty years ago, Bradley and colleagues first proposed that potyvirus Potato virus Y (PVY) was retained at the tip of the stylets of its aphid vector Myzus persicae. Formalin treatment or UV irradiation of the distal 15 μm of the stylets bundle abolished PVY transmission (20, 21). Because a negative impact of these treatments on aphid feeding behavior could not be excluded, these data remained controversial for more than 20 years (100). Several studies, combining electron microscopy, membrane acquisition of radio-labeled virions and HC-Pro, light microscopy, γ-counting, and autoradiography, later confirmed the stylet localization of potyviruses (Figure 2). Nevertheless, virus particles were sometimes found distributed erratically all along the length of the maxillary stylets (3, 10) and sometimes preferentially retained at their distal extremity (120) in the food canal (127) (Figure 2). A totally different approach (77, 106), based on correlative studies of aphid feeding behavior and virus inoculation, provided compelling evidence that the transmitted PVY and CMV particles are most likely retained at the very tip of the maxillary stylets, in the tiny distal region where the food and salivary canal fuse to form the common duct (Figure 2). Inoculation of PVY and CMV occurs during the first phase of the feeding process (77), when watery saliva is injected into the cytoplasm of the visited plant cell (106). Thus, although potyviruses could be observed all along the maxillary stylets, only those retained in the common duct can be flushed out by watery saliva and efficiently inoculated. Definitive proof that the receptors of noncirculative viruses can be located at the stylet tips of hemipteran insects came from the development of a novel tool allowing in vitro interaction 18.6

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between viruses and/or HC molecules and dissected stylets (124). The technique involved fusing HCs of CaMV (P2) to green fluorescent protein (GFP) and incubating the P2-GFP fusion with individual stylets adsorbed onto a microscopy slide. Binding of P2-GFP was readily observed by epifluorescence microscopy, revealing its exclusive attachment at the far end of the maxillary stylets, in the bed of the common duct (Figure 2). Using nonvector aphid species and nonfunctional P2 mutants allowed the authors to correlate binding of CaMV P2 at this precise location with successful transmission. Further analyses indicated that the corresponding receptor is a nonglycosylated protein that is deeply embedded in the chitin (124) and confined to an area paving the bottom of the common duct, where the surface of the cuticle appears swollen in high-resolution scanning electron microscopy (Figure 2). This newly discovered area is termed the acrostyle and has been demonstrated to contain cuticular proteins with a conserved RR2 motif (123) representing potential candidate receptors for CaMV. The idea that noncirculative viruses other than CaMV are also specifically hooked onto the acrostyle is tempting, and even sometimes assumed as confirmed fact (17, 95). To the best of our knowledge, however, no direct experimental proof for the involvement of the acrostyle in the transmission of potyviruses, cucumoviruses, or any noncirculative virus other than CaMV has been published, and further work is needed to confirm or refute this possibility.

Noncirculative Viruses in the Foregut of Insect Vectors Other than the stylets, possible localizations of noncirculative viruses include the precibarial pump or the foregut, although very few reports are available. In the family Sequiviridae, viral particles of the Anthriscus yellows virus (AYV; genus Waikavirus) were observed in the pharynx of the aphid vector Cavariella aegopodii, where the foregut joins the sucking pump (90), and Maize chlorotic dwarf virus (MCDV; genus Sequivirus) was seen in the precibarium and in the sucking pump of its leafhopper vector Graminella nigrifons (35). Somewhat confusingly, others reported virions in the cibarium and even in the food canal of the maxillary stylets of three leafhopper vectors: G. nigrifons, Graminella sonora, and Amblysellus grex, and also inside a nonvector insect, the cicadellid Dalbulus maidis (5). These conflicting data call for a clear correlation between viral location and successful transmission, which remains to be established for viruses in this family. Chen and coworkers (31) developed an elegant protocol to determine the retention sites of Lettuce infectious yellows virus (LIYV), a crinivirus (family Closteroviridae), in its whitefly vector Bemisia tabaci biotype A. LIYV uses the capsid strategy and does not require any HCs. Insects were sequentially fed a first diet containing purified virus particles, a second containing LIYV antibodies, and finally a third containing a secondary antibody conjugated to a fluorophore. Specific retention of the crinivirus in the anterior foregut and/or cibarium of the whitefly vectors was convincingly observed (Figure 2) and further shown to be mediated by the minor capsid protein CPm. Most importantly, parallel transmission assays demonstrated that virus retention at these sites correlates with successful transmission. In summary, two long-suspected possible locations for noncirculative viruses within their insect vectors have now been confirmed: the stylet tip for CaMV (124) and the foregut for LIYV (31).

Circulative Viruses in the Gut of Insect Vectors The first barrier encountered by circulative viruses is the gut epithelium. Its relevance for virus transmission was first recognized in the early 1930s, when it was shown that a nonvector leafhopper species could transmit the geminivirus Maize streak virus (MSV) solely when its gut was punctured with a needle or when the virus was injected directly into the hemocoel—the virus artificially www.annualreviews.org • Plant-Virus-Vector Interactions


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bypassing the gut epithelium in both cases (115). This method was later used extensively to confirm that the gut epithelium is the first specific checkpoint in nearly all interactions between a circulative virus and its insect vector, suggesting the existence of specific receptors at this site. For circulative nonpropagative virus species in the family Luteoviridae, internalization inside aphid vectors has been studied extensively by electron microscopy (reviewed in 24, 53). In compatible virus-vector pairs, the virus crosses the gut epithelium at the hindgut or posterior midgut level (108), via a transcytosis process, mediated partly by clathrin-coated vesicles (24). Virus particles are initially incorporated at the apical plasmalemma in spherical clathrin-coated vesicles and delivered to the endosomal compartment. Tubular uncoated vesicles containing virions then bud from the endosome and migrate to the basal plasmalemma for virus release (Figure 3). Strikingly, virus particles never come into contact with the cell cytoplasm during this process, and this observation strongly reinforces the belief that luteovirids do not replicate or express any of their genes inside aphid vectors.

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Species of the two families Geminiviridae and Nanoviridae are most often classified in the group of circulative nonpropagative viruses [with the exception of Tomato yellow leaf curl virus (TYLCV), see below], and the cycle within their insect vector is inferred mostly from what is described for luteoviruses. However, experimental data on how both nano- and geminiviruses actually cross the gut epithelium are nonexistent. Contrary to luteovirids, even which viral form passes though the insect cells remains uncertain, and distinctive viral particles could not be tracked clearly inside the vector body. Ultrastructural observation of a nano- or geminivirus inside its vector has been reported only for MSV (geminivirus) in the cytoplasm of midgut cells of Cicadulina mbila (1). The authors noted large cytoplasmic aggregates of MSV CPs arranged into paracrystalline arrays and enclosed in membranous vesicles. These structures were interpreted as virion arrays, but individual virions could barely be distinguished and the origin and actual role of such large CP or virion arrays remains mysterious. The nanovirus Banana bunchy top virus (BBTV) was recently localized by immunofluorescence (against coat protein) in cells of the anterior midgut of the aphid Pentalonia nigronervosa (130) (Figure 3). In whiteflies and leafhoppers, geminiviruses appear to cross the gut at the level of the filter chamber, which is the point of contact between midgut, hindgut, and Malpighian tubes, allowing water to bypass most of the digestive tract and be excreted rapidly (1). In particular, TYLCV (genus Begomovirus) could be located by immunofluorescence and FISH within the cytoplasm of gut cells of the filter chamber of the whitefly vector Bemisia tabaci (49, 82) (Figure 3). Intriguingly, several studies have suggested that TYLCV expresses some of its genes and even replicates within its vector Bemisia tabaci (41). Although this latter point remains under debate, the understanding of the vector transmission of geminiviruses (and likely that of nanoviruses) would certainly benefit from further investigation on the viral forms crossing the gut cellular barriers and their actual pathway within the insect cells. Entry into midgut cells via receptor-mediated endocytosis is also likely for circulative propagative viruses, such as tospo-, reo-, and rhabdoviruses (Figure 3) (6, 57). The best-characterized case is that of the reovirus RDV accumulating in midgut cells of the leafhopper Nephotettix cincticeps (33). The establishment of cell cultures from insect vectors allowed parallel microscopy and ←−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−− Figure 2 Localization of noncirculative viruses in their insect vectors. (a) Schematic drawing of the anterior alimentary tract of hemipteran insects: the food canal (FC) in the stylets is green, as are the precibarium (preCb), and cibarium (Cb). The salivary canal (SC) and principal (PSG) and accessory (ASG) salivary glands are in blue. (b) Two mandibular stylets (ma) are protecting two interlocked maxillary stylets (mx). Only the mandibular stylets contain dendrites (DE). The cross sections further reveal the salivary (SC) and common (CC) canals. Panels adapted with permission from Reference 120. (c) The helper components of Cauliflower mosaic virus (CaMV) [P2; green fluorescence in the common canal (CC)] specifically binds to the extreme tip of the maxillary stylets of the aphid Acyrthosiphon pisum, (d ) but not of the nonvector species Acyrthosiphon lactucae. (e) A nonfunctional mutant of P2 is not retained by any aphid species. ( f ) Specific antibodies reveal cuticular proteins with RR2 motif at this site, which correspond to the acrostyle further described in panels p–s. Panels adapted with permission from Reference 123 and 124. In panels g–i, Lettuce infectious yellows virus particles are immunolabeled in the Cb of the ( g) whitefly Bemisia tabaci biotype A, (h) but not in that of the nonvector biotype B. (i ) The minor capsid protein CPm binds specifically in the Cb and is thus directly involved in virus-vector interaction. Panels adapted with permission from reference 31. In panels j–o, potyvirus particles are retained in various parts of the stylet bundle: ( j,k) Radiolabeled virions were revealed by autoradiography, and filamentous virus-like particles (VLP) could be visualized and gold-labeled in the (l ) presence but not in the (m) absence of the helper HC-Pro. Another study reported potyvirus VLPs (arrowheads) in the (n) stylets and, although rarely, also in the (o) Cb. Panels j–m adapted with permission from Reference 127. Panels n–o adapted with permission from Reference 3. ( p–s) The acrostyle stands at the tip of aphids’ maxillary stylets and is delineated by white arrowheads. ( p; enlarged in q and r) It appears as an electron-dense region of the cuticle lining the bottom bed of the CC. (q) Two spherical CaMV particles are visible in the distal part of the acrostyle. (s) In scanning electron microscopy, the acrostyle appears as a swollen surface of the cuticle delineated by white arrowheads. Panels adapted with permission from References 123 and 124. Abbreviations: EC, epicuticule; L, leg; M, matrix; S, stylet bundle; V, filamentous virus-like particles. www.annualreviews.org • Plant-Virus-Vector Interactions


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pharmacological experiments, together showing that RDV enters cells through receptor-mediated clathrin-dependent endocytosis (134). Subsequently, the virus moves inside the cells along microtubules (136) and from cell-to-cell through spectacular tubular formations (reviewed recently in 85). Virus particles are tightly engulfed in nanotubes formed by the viral protein Pns10 that were observed in association with actin-based filopodia of epithelial cells and with muscle fibers of visceral muscle tissues in the alimentary canal (Figure 3). These structures are instrumental in the actin-dependent progression of RDV within the insect vector and ulterior transmission to plants.

Circulative Viruses in the Hemolymph of Insect Vectors


Different viral routes from the gut to the salivary glands have been described; these obviously differ for propagative and nonpropagative viruses, with all organs other than the salivary glands

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representing dead-end traps for the latter. Once released from the gut, nonpropagative luteoviruses are believed to diffuse passively into the hemolymph until they encounter putative receptors located specifically at the basal lamina of the salivary gland cells (24). In the hemolymph, several indirect lines of evidence suggest the involvement of a protein, symbionin, produced by bacterial endosymbionts that could act as a chaperone to protect virions from the insect immune system (126). Symbionin was shown to interact with Potato leafroll virus (PLRV) and BYDV (45, 125), and similar chaperone proteins were reported to enhance the transmission of geminiviruses by whitefly (88, 107) but not that of nanoviruses by aphids (129). The role of symbionin-like proteins in the transmission of luteo- and geminiviruses has been queried for two reasons: (a) the association between symbionins and viruses is largely nonspecific and occurs similarly with transmissible and nontransmissible viruses (87) and (b) symbionin might not be secreted by symbiotic bacteria in the hemolymph of the insect, casting doubts on its functional association with virions in vivo (19). For propagative viruses, routes within the vector can be extremely diversified, depending on which organs they actually infect. Although poorly characterized in many instances, some viruses may be released in the hemolymph and secondarily infect different organs, including the salivary glands, whereas others may have a very low viremia and rather pass from organ to organ by direct contact as described below.

Circulative Viruses Replicating in Various Organs of Insect Vectors Despite the fact that they replicate in their vector, propagative viruses do not invade all organs indifferently. Rhabdoviruses can infect nearly all tissues within the vector but are preferentially neurotropic, and their route from gut to salivary glands is via the nervous system (2). The reovirus RDV also seems to have a preferential sequence of organ infection within its leafhopper vector (33). From the initially infected midgut cells in the filter chamber, the virus progresses rapidly in the anterior midgut and associated muscles, presumably through tubule-assisted intercellular spread (34). The nervous system is infected rapidly, prior to other organs, including salivary ←−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−−− Figure 3 Localization of circulative viruses in the gut of their insect vectors. (a) Schematic representation of the alimentary tracts of aphids and cicadellids showing differences in their respective anatomies. Panel adapted with permission from Reference 42. (b–e) Luteovirid particles in the gut cells of the aphid Myzus persicae observed under transmission electron microscopy. (b) Virions attach to the apical plasmalemma (APL), (c) are internalized into diverse types of spherical vesicles [multivesicular body (MVB), multilamellar vesicle (MLV), smaller uncoated vesicles (VS), (d,e) transported into tubular vesicles (TV), and ( f ) finally delivered to the basal lamina (BL). Arrows in panel d show virions singly enclosed in vesicles. Panels adapted with permission from Reference 24. ( g) The geminivirus Tomato yellow leaf curl virus is visualized in the filter chamber (FC) and the ceca (CA) of its vector Bemisia tabaci biotype A by bluefluorescent in situ hybridization (FISH). Panel adapted with permission from Reference 49. (h,i ) The nanovirus Banana bunchy top virus is visualized in cells of the anterior midgut of the aphid Pentalonia nigronervosa by red immunofluorescence of the coat protein, (h) 4 and (i ) 10 days post ingestion. The cell nuclei are DAPI-stained in blue. Panels adapted with permission from Reference 131. ( j,k) Rhabdoviruses can be observed in many parts of their vectors’ gut. (j) Maize mosaic virus is immunolabelled in green in the esophagus (ES), the anterior diverticulum (AD), and the midgut (MG) of its planthopper vector. (k) Maize fine streak virus is immunolabelled in green in the ES, filter chamber (FC), anterior (AMG) and posterior (PMG) midgut, and hindgut (HG) of its leafhopper vector. In these images, actin is stained purple with Phallo¨ıdin. Panels adapted with permission from Reference 6. (l,m) The reovirus Rice dwarf virus moves from cell to cell in the vector through tubular structures of viral origin, (l ) associated with microvilli (Mv) of the anterior midgut (arrow). The high-resolution image in panel m was obtained with electron tomography and 3D model reconstruction. The model shows virus particles ( yellow) enclosed into tubules (red ), and plasma membranes (blue). Panels reproduced from Reference 85. Abbreviations: AM, ascending midgut; BPL, basal plasmalemma; DM, descending midgut; EC, epithelial cell; FG, foregut; GL, gut lumen; LU, lumen; MG; midgut; MT, Malpighian tubules; R, ribosomes; RE, rectum. www.annualreviews.org • Plant-Virus-Vector Interactions


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glands and ovarioles. The fact that rhabdo- and reoviruses can ultimately colonize many different organs of their vector is certainly a condition that enables these viruses to undergo transovarial transmission [as is also the case for the poorly characterized tenuiviruses (57)]. In contrast, there is no report of transovarial transmission for tospoviruses and marafiviruses, and this might be explained by the strict tissue specificity that targets the gut and salivary glands of their respective vectors. Accordingly, the tissue tropism of the tospovirus TSWV has been shown to be extremely limited. TSWV cannot be detected in the hemolymph (122), suggesting direct infection of thrips from midgut and surrounding muscle cells, the only two locations where the virus accumulates to detectable levels, to the salivary glands (92).

Circulative Viruses in the Salivary Glands of Insect Vectors Annu. Rev. Phytopathol. 2014.52. Downloaded from www.annualreviews.org by University of Waikato on 06/17/14. For personal use only.

Insects have two types of salivary glands, the principal (PSG) and accessory (ASG) glands (104), and viruses are generally restricted to one type. Circulative nonpropagative luteoviruses have been found solely in ASG—a more selective barrier than the gut (97). Viral particles are transported through the salivary glands via a transcytosis mechanism similar to that in gut cells but in the opposite direction from basal to apical plasmalemma (50). Other viruses, such as geminiviruses (30, 39, 51) and nanoviruses (130, 131), have been found exclusively in the PSG. Most circulative propagative viruses invade the PSG, but dual localization in both PSG and ASG has sometimes been reported. For example, the rhabdovirus MMV has been detected in various acini of both ASG and PSG (6), where virions bud from the plasma membrane to accumulate in the intercellular spaces that connect the salivary ducts (4). Likewise, different pairs of salivary glands, the tubular and ovoid glands, have been described in thrips (an insect taxa unrelated to hemipterans) and both can be infected by TSWV (92, 121, 140).

Receptors of Viruses Within Insect Vectors As with transmission of noncirculative viruses, specific receptors involved in initial contact between circulative viruses and insect vectors have eluded researchers for more than two decades. Identifying such molecules is tricky because insect compounds interacting with circulative viruses are not only receptors but may also be involved in other steps of the virus cycle within the vector. During the past 15 years, biochemical (9, 65, 70, 110), proteomic (37), transcriptomic (138), and genetic/genomic (69, 118, 119) approaches have been applied to various virus groups, such as luteoviruses (37, 70, 110, 118, 119), tospoviruses (9, 65), rhabdoviruses (138), and geminiviruses (69). Many insect proteins (not listed here) have been identified as being possibly involved in the regulation of the virus-vector interaction. Unfortunately, thus far, the main bottleneck is at the level of functional validation of these candidates, and no receptor of any circulative virus has yet been definitely confirmed. A noteworthy finding used phage display technology to isolate a 12 amino acid peptide that was able to impede the uptake of Pea enation mosaic virus (PEMV; Luteoviridae) in the midgut of its aphid vectors (73). This peptide, designated GBP3.1, was screened and isolated on dissected aphid guts, where it proved able to outcompete virus attachment. This result led to GBP3.1 being used to specifically target a fused Bt toxin to the aphid gut, significantly enhancing its toxicity (36). In this latter study, the authors also mentioned a specific interaction between GBP3.1 and alanyl aminopeptidase-N (APN) of the aphid vector, which represents a likely candidate receptor for PEMV.

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IN PLANTA VIRUS LOCALIZATION AND IMPACT ON UPTAKE BY THE INSECT VECTOR A trivial but nonetheless noteworthy point is that for successful transmission, viruses must be accessible to the vector, i.e., virus localization in plants or other hosts must match the feeding sites of the vector. The question is whether this is simply incidental and viruses are taken up whenever present in hosts fluids ingested by vectors or whether true viral adaptation exists, with sophisticated mechanisms placing the virus at the right place at the right time (16). Although the situation appears simple for viruses transmitted by biting vectors (reviewed in 48), in the most frequent cases of viral transmission by phloem feeders (aphids, whiteflies, hoppers) the situation is multifarious. There, viruses can be acquired and released either during test punctures in epidermal and parenchymal cells, during phloem sap ingestion, or during both. Many circulative, but also some noncirculative, viruses have a tissue tropism restricted strictly to phloem tissues and have been shown to be acquired by their phloem-feeding vectors during sustained sap ingestion in sieve tubes (44, 114). In these cases, the virus titer within the sap is likely the key determinant of efficient acquisition. However, many noncirculative viruses (i.e., poty-, cucumo-, alfamo-, carlavirus, etc.) with no restricted tissue tropism are acquired not during phloem-sap ingestion but during testprobing punctures in epidermis and parenchyma (44, 77, 114). As reported for aphids and, to a lesser extent, for whiteflies, these test probes sample minute amounts of cell content, leaving the cell alive after stylet withdrawal. Such delicate feeding behavior further poses the problem of virus location at the intracellular scale, where viruses sequestered in inclusion bodies are not readily accessible to vectors just anywhere in the cytosol. It thus appears reasonable to question the possible existence of sophisticated viral adaptations that maximize uptake by the vector through targeted positioning within the host cell. In general, parasites do not simply colonize the host and passively await transmission. Instead, they control the differentiation of specific transmissible morphs with specific localization, or with various extraordinary manipulations of host behavior, to ensure higher success in transmission (80). For example, the protozoan parasite Toxoplasma gondii blocks the innate aversion of predators in their rodent secondary hosts, making them more prone to be eaten by cats—the primary hosts in which this parasite can reproduce sexually (58). Certain hairworms cause infested crickets to dive into water, a deadly habitat otherwise carefully avoided by the insects but necessary for the worms to mate (105). Additional fascinating examples are discussed in Reference 67. Comparable phenomena have also been reported specifically for viruses. Rabies virus, for example, changes host behavior (reviewed in 67) to increase biting contact with a new host. Baculoviruses ensure host-to-host transmission by forming specific morphs, the so-called polyhedra—stable inclusions that can persist in the environment until a new host feeds on them (reviewed in 8). In plants, viruses may change their vector behavior directly or indirectly through changes of the volatile aura, the color and/or the nutrient content of the host plant (reviewed in 55, 76). However extraordinary, all these examples represent constitutive optimization of a virus-infected host for efficient ulterior transmission. A very recent study on the transmission of CaMV advanced this concept significantly by demonstrating that a virus-host interaction optimizing vector transmission can be transient and conditional, and sparked off specifically by the arrival of an insect vector on the host plant. CaMV forms a viral inclusion [termed the transmission body (TB)] specialized for vector transmission in infected plant cells (43, 64, 79). The TB is not the form acquired by the aphid vector; rather, the TB reacts to the puncture of the aphid vector into the infected leaf and immediately spreads transmission-specific viral morphs throughout the cell. More precisely, Martinière and collaborators (78) showed that the TB can dissociate within seconds of vector contact, instantly relocalizing its contents (basically the viral HC P2; see Figure 4) onto microtubules throughout



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the cell. At the same time, CaMV viral factories (the sites of virus replication and virion accumulation) dispatch virus particles into the cytosol to associate immediately with P2 on the microtubules (7). This phenomenon was demonstrated to enhance the acquisition of CaMV markedly and to totally revert after departure of the aphid vector, resetting the infected cell for another round of transmission. Because viruses do not themselves possess a proper sensory system, these findings suggest that CaMV interferes with a pre-existing host plant defense system against aphid infestation, highjacking it for its own purposes. To date, no other similar viral reaction has been clearly identified. However, an interesting parallel can be found in the literature for a virus that infects mammals. Bluetongue virus (BTV) can survive several months (during winters) in the absence of Culicoides midge vectors and with no detectable cases of viremia into the host population. This phenomenon, designated overwintering, has long intrigued scientists in the field (74), and one of the proposed explanations is that the virus re-emergence in hosts with latent infection could be triggered by the bites of insect vectors, when reappearing after the cold season (141). Cultures of ovine γδT cells can be latently infected, and contact with skin fibroblasts results in conversion to a lytic infection and increased BTV release (117). Because feeding of midges induces skin inflammation, recruiting activated γδT cells, an appealing hypothesis has been proposed: Upon feeding of midge vectors, latently infected γδT cells would come into contact with skin fibroblasts, triggering conversion of BTV into a lytic cycle, which would result in de novo virus accumulation precisely at the biting sites. Although controversial (141), this explanation for BTV overwintering might represent another example of viruses reacting to the presence of their vector through the highjack of host defense pathways to enable timely and efficient acquisition.


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CONCLUDING REMARKS AND FUTURE PROSPECTS From the above, it is striking that detailed molecular information on the initial recognition factors in both virus and host is limited to a very restricted number of cases. Despite known HC molecules from poty- and caulimoviruses, CP proteins from cucumo- and reoviruses, and glycoprotein from rhabdo- and tospoviruses, the actual peptide sequences binding to vector receptors, and their structures, have not been elucidated. Continued efforts in this direction are needed because such protein motifs will likely form the basis for the design of antagonistic molecules capable of outcompeting viruses for attachment to vectors. The search for the identity of plant virus receptors in their insect vectors is undoubtedly one of the major challenges in this field in the coming years. Despite nearly two decades of research, serious technical bottlenecks have as yet proved intractable, likely because of the paucity of genetic tools/resources on the vector side. Although a number of protein partners of viruses have been identified and/or isolated successfully from various insect vector species, validation and determination of their actual function remains problematic. However, breakthroughs can be expected soon, given the increasing genomic data for aphids (38) and whiteflies, and the promising development of directed RNAi in several hemipteran insect species (71). That a virus can perceive the vector and respond by switching to a transmission-enhanced configuration is a totally new concept worthy of further exploration. Convincingly demonstrated by the example of CaMV (Figure 4), there are good reasons to believe that other viruses may similarly control timely and efficient encounter with vectors. A possible illustration of this can be seen in a study published nearly 20 years ago that monitored the accumulation of the circulative PLRV in plant leaves infested with aggregated aphid colonies (89). The presence of aphids on an infected plant enhanced virus accumulation specifically at the feeding site, potentially enhancing acquisition and transmission. The author speculated that the virus was more likely to move toward 18.14

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Plant cell

P3 VF

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P2 TB Virion

P6 Virion

Microtubule


a

b

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Activated TB

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c

Mixed networks

Figure 4 Model of Cauliflower mosaic virus acquisition during test punctures by aphid vectors. (a) In an infected cell in the standby state, there are numerous virus factories (VF) containing most of the replicated virions ( yellow circles), which are enclosed within a matrix of viral protein P6 ( gray). This is accompanied in the cell by a mostly single transmission body (TB), composed of a matrix containing all of the cell’s P2 (red ), coaggregated with P3 (blue) and some virus particles. Microtubules are represented in green. (b) An aphid landing on an infected plant inserts its stylets into a cell to test the plant. This causes a mechanical stress (stylet movement) and/or a chemical stress (e.g., elicited by saliva components). This stress, symbolized by the yellow lightning bolts, is immediately perceived by the plant and can induce subsequent defense responses. The initial aphid recognition signal is transduced simultaneously in a TB response, characterized by an influx of tubulin ( green) into the TB. (c) In the second step, the TB disintegrates rapidly (within seconds), and all the P2 as well as numerous virus particles originating from the VFs relocalize on the cortical microtubules as mixed networks (shown in the foreground ). Transmissible P2-virus complexes are now homogeneously distributed throughout the cell periphery, which significantly increases the chances of successful binding of P2 and virus to the stylets and thus transmission. (d) After departure of the aphid vector (loaded with P2 and virus), a new TB is reformed from the mixed networks and is ready for another round of transmission. Figure adapted with permission from Reference 78.

nutrient sinks created at the sites of aggregated aphid feeding than to locally increase their replication rate. This observation, together with that discussed above for CaMV and BTV, may unravel an interesting diversity of viral reactions when contacting their vectors: from an instantaneous reshuffling of viral compounds inside a cell (CaMV) to a translocation of virions at the leaf or whole host level (PLRV) or even a locally resumed replication (BTV). www.annualreviews.org • Plant-Virus-Vector Interactions


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Finally, a perusal of the literature on virus location within vectors might lead one to wonder whether the question is being adequately or comprehensively addressed. Indeed, the most common approach is to localize virus particles and/or sites where the virus possibly replicates within its vector, but there are other viral compounds that may play a key role in the transmission process. These include nonstructural viral proteins and viral siRNAs. It has been shown that antibodies fed to whiteflies can pass the gut barrier and make their way inside the body to their target organ (40). Likewise, siRNAs can penetrate the aphid body and silence their target genes (101). It is tempting to imagine that nonstructural viral proteins, as well as viral siRNA with microhomology to some genes of insect vectors, could accumulate in phloem sap, be amply ingested by insect vectors, and remodel their physiology to facilitate virus transmission. SUMMARY POINTS Annu. Rev. Phytopathol. 2014.52. Downloaded from www.annualreviews.org by University of Waikato on 06/17/14. For personal use only.

1. Although the route of viruses within the bodies of their vectors is well established, the actual molecular interactions between both remain poorly understood. 2. The viral proteins or protein domains that are responsible for specific recognition of the vector are documented in only a handful of cases. 3. Whatever the virus-vector system, the virus receptor within the vector could not as yet be definitely confirmed. 4. Viruses can evolve exquisitely sophisticated interactions with their host plants to switch to a transmissible mode specifically when the vector is present.

FUTURE ISSUES 1. Further biochemical and structural characterization of the viral determinants of vector recognition is needed both in the search for counterpart receptors in vectors and for designing putatively interfering molecules. 2. The identification and characterization of receptors of viruses within vectors is a major challenge. 3. A new research horizon lies in the investigation of viral mechanisms transiently switching the plant-virus system to a transmission-enhanced mode. 4. Research on the localization of viruses within their vectors should widen to the localization of nonstructural proteins and noncoding RNAs susceptible of impacting on both physiology and behavior of vectors to facilitate transmission.

DISCLOSURE STATEMENT The authors are not aware of any affiliations, memberships, funding, or financial holdings that might be perceived as affecting the objectivity of this review.

ACKNOWLEDGMENTS We are grateful to H. Rothnie for English editing of the manuscript. S.B., M.D., and M.U. acknowledge support from INRA Department SPE. This work is also funded by ANR grant 18.16

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number 2010BLAN170401 (S.B.), ANR grant number 12-BSV7-005-01 (M.D.), and a grant from the region LR Chercheurse d’Avenir 2011 (M.U.).


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22. Bragard C, Caciagli P, Lemaire O, Lopez-Moya JJ, MacFarlane S, et al. 2013. Status and prospects of plant virus control through interference with vector transmission. Annu. Rev. Phytopathol. 51:177–201 23. Brault V, Bergdoll M, Mutterer J, Prasad V, Pfeffer S, et al. 2003. Effects of point mutations in the major capsid protein of beet western yellows virus on capsid formation, virus accumulation, and aphid transmission. J. Virol. 77:3247–56 24. Brault V, Herrbach E, Reinbold C. 2007. Electron microscopy studies on luteovirid transmission by aphids. Micron 38:302–12 25. Brault V, Perigon S, Reinbold C, Erdinger M, Scheidecker D, et al. 2005. The polerovirus minor capsid protein determines vector specificity and intestinal tropism in the aphid. J. Virol. 79:9685–93 26. Brault V, Uzest M, Monsion B, Jacquot E, Blanc S. 2010. Aphids as transport devices for plant viruses. C. R. Biol. 333:524–38 27. Brault V, van den Heuvel JF, Verbeek M, Ziegler-Graff V, Reutenauer A, et al. 1995. Aphid transmission of beet western yellows luteovirus requires the minor capsid read-through protein P74. EMBO J. 14:650– 59 28. Brault V, Ziegler-Graff V, Richards KE. 2001. Viral determinants involved in luteovirus-aphid interactions. See Ref. 56a, pp. 207–32 29. Briddon RW, Pinner MS, Stanley J, Markham PG. 1990. Geminivirus coat protein replacement alters insect specificity. Virology 177:85–94 30. Brown JK, Czosnek H. 2002. Whitefly transmission of plant viruses. Adv. Bot. Res. 36:65–100 31. Chen AY, Walker GP, Carter D, Ng JC. 2011. A virus capsid component mediates virion retention and transmission by its insect vector. Proc. Natl. Acad. Sci. USA 108:16777–82 32. Chen B, Francki RIB. 1990. Cucumovirus transmission by the aphid Myzus persicae is determined solely by the viral coat protein. J. Gen. Virol. 71:939–44 33. Chen H, Chen Q, Omura T, Uehara-Ichiki T, Wei T. 2011. Sequential infection of Rice dwarf virus in the internal organs of its insect vector after ingestion of virus. Virus Res. 160:389–94 34. Chen Q, Chen H, Mao Q, Liu Q, Shimizu T, et al. 2012. Tubular structure induced by a plant virus facilitates viral spread in its vector insect. PLoS Pathog. 8:e1003032 35. Childress SA, Harris KF. 1989. Localization of virus-like particles in the foreguts of viruliferous Graminella nigrifrons leafhoppers carrying the semi-persistent maize chlorotic dwarf virus. J. Gen. Virol. 70:247–51 36. Chougule NP, Li H, Liu S, Linz LB, Narva KE, et al. 2013. Retargeting of the Bacillus thuringiensis toxin Cyt2Aa against hemipteran insect pests. Proc. Natl. Acad. Sci. USA 110:8465–70 37. Cilia M, Tamborindeguy C, Fish T, Howe K, Thannhauser TW, Gray S. 2011. Genetics coupled to quantitative intact proteomics links heritable aphid and endosymbiont protein expression to circulative polerovirus transmission. J. Virol. 85:2148–66 38. Consortium TIAG. 2010. Genome sequence of the pea aphid Acyrthosiphon pisum. PLoS Biol. 8:e1000313 39. Czosnek H, Ghanim M, Ghanim M. 2002. The circulative pathway of begomoviruses in the whitefly vector Bemisia tabaci: insights from studies with Tomato yellow leaf curl virus. Ann. Appl. Biol. 140:215–31 40. Czosnek H, Morin S, Rubinstein M, Fridman V, Zeidan M, Ghanim M. 2001. Tomato yellow leaf curl virus: a disease sexually transmitted by whiteflies. See Ref. 56a, pp. 1–27 41. Diaz-Pendon JA, Canizares MC, Moriones E, Bejarano ER, Czosnek H, Navas-Castillo J. 2010. Tomato yellow leaf curl viruses: menage a trois between the virus complex, the plant and the whitefly vector. Mol. Plant Pathol. 11:441–50 42. Engel P, Moran NA. 2013. The gut microbiota of insects: diversity in structure and function. FEMS Microbiol. Rev. 37:699–735 43. Espinoza AM, Medina V, Hull R, Markham PG. 1991. Cauliflower mosaic virus gene II product forms distinct inclusion bodies in infected plant cells. Virology 185:337–44 44. Fereres A, Moreno A. 2009. Behavioural aspects influencing plant virus transmission by homopteran insects. Virus Res. 141:158–68 45. Filichkin SA, Brumfield S, Filichkin TP, Young MJ. 1997. In vitro interactions of the aphid endosymbiotic SymL chaperonin with barley yellow dwarf virus. J. Virol. 71:569–77 46. Franz AW, van der Wilk F, Verbeek M, Dullemans AM, van den Heuvel JF. 1999. Faba bean necrotic yellows virus (genus Nanovirus) requires a helper factor for its aphid transmission. Virology 262:210–19


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47. Gera A, Loebenstein G, Raccah B. 1979. Protein coats of two strains of cucumber mosaic virus affect transmission of Aphis gossypii. Phytopathology 69:369–99 48. Gergerich RC. 2001. Elucidation of transmission mechanisms: mechanism of virus transmission by leaffeeding beetles. See Ref. 56a, pp. 133–40 49. Ghanim M, Brumin M, Popovski S. 2009. A simple, rapid and inexpensive method for localization of Tomato yellow leaf curl virus and Potato leafroll virus in plant and insect vectors. J. Virol. Methods 159:311–14 50. Gildow F. 1999. Luteovirus transmission mechanisms regulating vector specificity. In The Luteoviridae, ed. HG Smith, H Barker, pp. 88–111. Wallingford, UK: CABI 51. Goldman V, Czosnek H. 2002. Whiteflies (Bemisia tabaci ) issued from eggs bombarded with infectious DNA clones of Tomato yellow leaf curl virus from Israel (TYLCV) are able to infect tomato plants. Arch. Virol. 147:787–801 52. Govier DA, Kassanis B. 1974. A virus induced component of plant sap needed when aphids acquire potato virus Y from purified preparations. Virology 61:420–26 53. Gray S, Gildow FE. 2003. Luteovirus-aphid interactions. Annu. Rev. Phytopathol. 41:539–66 54. Guo B, Lin J, Ye K. 2011. Structure of the autocatalytic cysteine protease domain of potyvirus helpercomponent proteinase. J. Biol. Chem. 286:21937–43 55. Gutiérrez S, Michalakis Y, Van Munster M, Blanc S. 2013. Plant feeding by insect vectors can affect life cycle, population genetics, and evolution of plant viruses. Funct. Ecol. 27:610–22 56. Harris KF. 1977. An ingestion-egestion hypothesis of non circulative virus transmission. In Aphids as Virus Vectors, ed. KF Harris, K Maramorosch, pp. 166–208. New-York: Academic 56a. Harris KF, Smith OP, Duffus JE, eds. 2001. Virus-Insect-Plant Interactions. San Diego, CA: Acad. Press 57. Hogenhout SA, Ammar ED, Whitfield AE, Redinbaugh MG. 2008. Insect vector interactions with persistently transmitted viruses. Annu. Rev. Phytopathol. 46:327–59 58. Ingram WM, Goodrich LM, Robey EA, Eisen MB. 2013. Mice infected with low-virulence strains of Toxoplasma gondii lose their innate aversion to cat urine, even after extensive parasite clearance. PLoS ONE 8:e75246 59. Ingwell LL, Eigenbrode SD, Bosque-Perez NA. 2012. Plant viruses alter insect behavior to enhance their spread. Sci. Rep. 2:578 60. Jiu M, Zhou X-P, Tong L, Xu J, Yang X, et al. 2007. Vector-virus mutualism accelerates population increase of an invasive whitefly. PLoS ONE 2:e182 61. Johnson JE. 2003. An atomic model of a plant reovirus: rice dwarf virus. Structure 11:1193–94 62. Kaper JM. 1969. Reversible dissociation of cucumber mosaic virus (strain S). Virology 37:134–39 63. Kennedy JS, Day MF, Eastop VF. 1962. A Conspectus of Aphids as Vectors of Plant Viruses. London: Commonwealth Inst. Entomol. 114 pp. 64. Khelifa M, Journou S, Krishnan K, Gargani D, Esperandieu P, et al. 2007. Electron-lucent inclusion bodies are structures specialized for aphid transmission of cauliflower mosaic virus. J. Gen. Virol. 88:2872– 80 65. Kikkert M, Meurs C, van de Wetering F, Dorfmuller S, Peters D, et al. 1998. Binding of Tomato spotted wilt virus to a 94-kDa thrips protein. Phytopathology 88:63–69 66. Killiny N, Rashed A, Almeida RP. 2012. Disrupting the transmission of a vector-borne plant pathogen. Appl. Environ. Microbiol. 78:638–43 67. Lefevre T, Adamo SA, Biron DG, Misse D, Hughes D, Thomas F. 2009. Invasion of the body snatchers: the diversity and evolution of manipulative strategies in host-parasite interactions. Adv. Parasitol. 68:45– 83 68. Leh V, Jacquot E, Geldreich A, Hermann T, Leclerc D, et al. 1999. Aphid transmission of cauliflower mosaic virus requires the viral PIII protein. EMBO J. 18:7077–85 69. Leshkowitz D, Gazit S, Reuveni E, Ghanim M, Czosnek H, et al. 2006. Whitefly (Bemisia tabaci ) genome project: analysis of sequenced clones from egg, instar, and adult (viruliferous and non-viruliferous) cDNA libraries. BMC Genomics 7:79 70. Li C, Cox-Foster D, Gray SM, Gildow F. 2001. Vector specificity of barley yellow dwarf virus (BYDV) transmission: identification of potential cellular receptors binding BYDV-MAV in the aphid, Sitobion avenae. Virology 286:125–33 www.annualreviews.org • Plant-Virus-Vector Interactions


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18.23

Ecology, evolution and control of plant viruses and their vectors.

Insect-specific viruses and their potential impact on arbovirus transmission.

Water deficit enhances the transmission of plant viruses by insect vectors.

Circadian rhythms in insect disease vectors.

Acoustic communication in insect disease vectors.

Enzyme synergistic for insect viruses.

The speed of tubule formation of two fijiviruses corresponds with their dissemination efficiency in their insect vectors.

Defective interfering viruses and their impact on vaccines and viral vectors.

Insect vector-mediated transmission of plant viruses.

Gene expression using insect cells and viruses.

Targeting renal epithelial channels for the control of insect vectors.

Chemosterilants for Control of Insects and Insect Vectors of Disease.

Insect Vectors of Disease: Untapped Reservoirs for New Antimicrobials?

Heritable strategies for controlling insect vectors of disease.

Olfactory disruption: toward controlling important insect vectors of disease.

Insect vectors as drivers of plant virus emergence.

Retroviral vectors: from cancer viruses to therapeutic tools.

Insect-specific viruses detected in laboratory mosquito colonies and their potential implications for experiments evaluating arbovirus vector competence.

The diversity of insect antiviral immunity: insights from viruses.

Comparative ultrastructural studies of insect granulosis and nuclear polyhedrosis viruses.

Full-genome characterisation of Orungo, Lebombo and Changuinola viruses provides evidence for co-evolution of orbiviruses with their arthropod vectors.

Comparative measurements of size and polydispersity of several insect viruses.

Localizing factors in atherosclerosis.

Respiratory viruses and their impact in healthcare.