INRA, UR BIVV, 28 rue de Herrlisheim, 68021 Colmar Cedex, France
Correspondence
Véronique Brault
brault{at}colmar.inra.fr
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ABSTRACT |
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INTRODUCTION |
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The route of several luteovirids in their aphid vector during acquisition/transmission has been followed by transmission electron microscopy (TEM). These observations have led to a model for transcytosis of virus particles at the level of both the intestine and the accessory salivary glands (ASG). In epithelial cells of the gut, the virions are internalized by endocytosis via clathrin-coated vesicles at the apical side of the cell and released into the haemocoel by exocytosis at the basal side (Gildow, 1987; Reinbold et al., 2001
; Garret et al., 1993
). This polarized transcytosis operates in the reverse sense (i.e. basal to apical pole) at the ASG, where uptake from the haemocoel is followed by release of virions into the salivary canal from which they are delivered into plant phloem during subsequent feeding (Gildow, 1987
; Gildow & Gray, 1993
; Reinbold et al., 2001
). Endocytosis and exocytosis at each of these barriers are thought to involve specific interactions between capsid proteins and receptors present in the aphid's body (Gildow, 1987
, 1993
).
Different sites of virion uptake at the intestinal level have been described, depending on the combination of virus and aphid vector under examination. Hindgut cells in cereal aphids are the site of acquisition of Barley yellow dwarf virus-PAV (BYDV-PAV) and -MAV (Luteovirus), as well as Cereal yellow dwarf virus-RPV (CYDV-RPV) (Polerovirus) (Gildow, 1999). Hindgut cells have also been identified as the internalization site of Soybean dwarf virus (SbDV; unassigned member of the family Luteoviridae) in Aulacorthum solani and M. persicae (Gildow et al., 2000
). The posterior midgut of M. persicae is involved in the transport of Potato leafroll virus (PLRV; genus Polerovirus; Garret et al., 1993
) and of Beet western yellows virus (BWYV; genus Polerovirus; Reinbold et al., 2001
) from the gut lumen to the haemocoel. In the present study, ultrastructural observations were made to determine the route of CABYV virions through the aphid vectors M. persicae and A. gossypii. TEM observations were undertaken to localize CABYV particles in both gut cells and in ASG cells. These observations reveal a novel situation in which uptake of virions occurs at two distinct positions in the digestive tract.
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METHODS |
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Virus-free aphid colonies of Myzus persicae (Sulzer) were reared on caged pepper (Capsicum annuum) seedlings. Virus-free colonies of Aphis gossypii Glover were initiated from a specimen collected in a greenhouse in Colmar (Alsace, France) and reared on caged cucumber seedlings (Cucumis sativus). Both cultures were maintained in a controlled environment chamber at 20 °C with a 16 h photoperiod. For the gut observations, nymphs (third or fourth instar) or adults were given a 72 h acquisition access period (AAP) through a stretched Parafilm membrane on various concentrations of purified virions prepared in 0·1 M sodium citrate, pH 6·0, containing 20 % sucrose. Aphids fed on the artificial diet MP148 (Harrewijn, 1983) were used as non-viruliferous controls. After the AAP, some aphids were transferred to healthy M. perfoliata seedlings for a 4 day inoculation access period (IAP) to assess their capacity to transmit the virus. These test plants were assayed for CABYV infection 4 weeks later by double antibody sandwich ELISA (Clark & Adams, 1977
) using a rabbit polyclonal antiserum (H. Lecoq, INRA Avignon, France).
To visualize virions at the ASG level, aphids were allowed to acquire virus by membrane feeding, as described above, or purified virions were microinjected into the aphid's haemocoel as previously described (Bruyère et al., 1997). After injection, aphids were transferred for 24 h to M. perfoliata before being prepared for ASG ultrastructural examination (Reinbold et al., 2001
). The plants were tested for virus infection 4 weeks later by ELISA.
In order to visualize virions at the basal pole of gut cells, some aphids were microinjected with CABYV antiserum after 72 h AAP on purified CABYV. Aphids injected with BWYV anti-P19 polyclonal antiserum (Reutenauer et al., 1993) served as controls in these experiments. Microinjected aphids were fixed and embedded 3 to 5 h later (Reinbold et al., 2001
). For ultrastructural examination, aphids were bisected, fixed and embedded in Epon/Araldite plastic as previously described (Reinbold et al., 2001
). All observations were made with a Philips EM208 transmission electron microscope operating at 80 kV.
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RESULTS |
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The digestive tract of M. persicae is divided into four successive parts: the foregut, the anterior midgut (also referred to as the stomach), the posterior midgut (or tubular midgut) and the hindgut. Our TEM observations were mainly focused on the posterior midgut and the hindgut, known to be intestinal acquisition sites of viruses of the family Luteoviridae (Gildow, 1999), although the observations were also extended to the stomach. Isolated virus particles were occasionally seen in the stomach lumen in 5 out of 9 aphids examined, but no virus-like particle was ever observed within the cytoplasm or in the basal lamina of the stomach cells (Table 1
). Virions, either isolated or in rosette-like clusters of several particles, were seen free in the posterior midgut lumen, most frequently close to the microvilli of the apical plasmalemma of the epithelial cells (Fig. 1
a). Single virions were occasionally observed in shallow depressions of the apical plasmalemma (Fig. 1a
). Various membranous structures containing virus particles were observed inside posterior midgut cells in about one-third of the aphids examined (Table 1
). Virus particles were always within membrane-bound vesicles and could be differentiated from ribosomes by their sharper outline, their larger diameter and denser staining. Virions were mainly enclosed in endosome-like vesicles such as multilamellar vesicles or multivesicular bodies (Fig. 1b
). Tubular vesicles (Fig. 1b
) or coated vesicles (not shown) containing virions were also present. These different structures are known to be associated with receptor-mediated endocytosis (Roth, 1993
) and have already been described for posterior midgut cells of M. persicae fed on BWYV particles (Reinbold et al., 2001
) or on PLRV particles (Garret et al., 1993
). At the basal pole of these cells, isolated virus particles were observed embedded in the basal lamina in 3 out of 34 aphids observed (Table 1
). Feeding aphids on higher concentrations of purified CABYV increased the number of aphids (6 out of 11) in which virions were observed in the basal lamina but these virions were always isolated and dispersed along the basal lamina (Table 1
).
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Observations were also carried out on the aphid hindgut, which can be easily differentiated from the posterior midgut by its typical flat and long cells, by the low abundance of membrane invaginations formed by the apical plasmalemma and by the presence of tubules lining the apical plasmalemma (O'Loughlin & Chambers, 1972). Virus particles, both free and in rosette-like aggregates (Fig. 2
a), were frequently seen in the lumen of the hindgut, as well as close to the apical plasmalemma. Linear arrays of virions in shallow depressions or in deep invaginations on the surface of the plasmalemma were also routinely observed (Fig. 2a
). Whatever the virus concentration delivered to the aphids, a high percentage of the aphids observed contained virions within hindgut cells (Table 1
). Virions were occasionally seen clustered in larger, spherical vesicles (not shown) but they were mainly present in tubular vesicles and in small circular vesicles containing one or more virions (Fig. 2a, b
). When M. persicae was fed on CABYV particles at a concentration of 100 µg ml-1, some hindgut epithelial cells were heavily charged with virions, as shown in Fig. 2(b)
, but virus particles were never observed in the basal lamina of these cells (Table 1
), although tubular vesicles were sometimes observed in very close proximity to the basal pole of the cell (Fig. 2c
). Virions were seen between the basal plasmalemma and the basal lamina or embedded in the basal lamina of hindgut cells only when aphids were fed on higher virus concentrations or when CABYV antiserum was microinjected subsequently (Table 1
). In this latter treatment, virions were always observed in clusters (not shown). In contrast, only isolated virions were seen after microinjection of BWYV P19 antiserum (Table 1
). Virus-like particles were never observed in any cellular compartment in M. persicae fed on artificial medium lacking virus (Table 1
).
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In parallel, transmission of virus to plants by microinjected aphids was monitored by transferring 5 aphids to each test plant. When M. persicae nymphs were microinjected with a purified suspension of CABYV at 130 µg ml-1, half of the test plants became infected (5 infected plants out of 10 plants analysed). We observed that less than half of the microinjected aphids survived the treatment (22 survivors out of 50 injected aphids 24 h after microinjection). When the virus concentration delivered into the haemocoel of M. persicae was raised to 350 µg ml-1, no virus transmission to test plants was observed. However, we noticed that this treatment had a dramatic effect on aphid survival (4 survivors out of 40 injected aphids), which could explain the observed loss of virus transmissibility. Microinjection of 350 µg ml-1 CABYV suspension into A. gossypii also resulted in a reduction of aphid survival (5 survivors out of 40 microinjected), but we still observed transmission of virus to test plants for 3 plants out of 8 analysed. The reason for elevated aphid mortality following microinjection of the concentrated virus solution is not known but could be due to the presence in the purified viral suspension of a plant compound having a toxic effect when delivered directly to the haemocoel at sufficiently high concentration.
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DISCUSSION |
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Acquisition and transmission of CABYV were observed to occur similarly in its two vector species. It should be noted, however, that in order to visualize virions in the digestive tract, we were obliged to use a higher virus concentration for A. gossypii than for M. persicae, although we found that these species did not differ in transmission efficiency when fed with the same virus concentration. An opposite situation pertained at the ASG level, where virions were more numerous in cells of A. gossypii than of M. persicae when microinjected with equivalent virus concentrations. The relative efficiency of transcytosis thus appears to differ between the aphid species and between the two epithelial barriers. These observations may reflect differences in transcytosis rates or may be related to the frequency of the putative virus receptors at the corresponding epithelium. Further work using quantitative RT-PCR will help to compare the dynamics of virus transport across each type of epithelium in the two vector species (Lett et al., 2002).
Our ultrastructural observations detected virion-containing membrane-bound organelles in intestinal cells similar to those described by previous workers for other luteovirids and in accordance with the Gildow model for acquisition (Gildow, 1999). The observed virus-containing structures included clathrin-coated pits and vesicles, multilamellar and multivesicular endosome-like bodies, and tubular vesicles. Moreover, the transport of CABYV virions across the ASG cells does not seem to differ notably from previous reports for other virusvector combinations. At the cellular level, the transcytosis mechanism at the intestinal and salivary epithelia is therefore very likely to be the same for all members of the Luteoviridae and all vector species. Transcytosis is an important pathway for membrane trafficking which allows selective and rapid transcellular vesicular transport from the apical to the basolateral pole of epithelial cells (Mostov et al., 2000
), and viruses have no doubt hijacked this pathway for their own ends. Specificity of luteovirid entry into epithelial cells is thought to be mediated by receptors associated with the cell membrane, followed by endocytosis of virions, according to the so-called receptor-mediated endocytosis' (Pastan & Willingham, 1985
) mechanism, which is also widely used by animal viruses to enter host cells (for review, see Sieczkarski & Whittaker, 2002
). This step is generally followed by virus release into and replication in the cytosol, in contrast to luteovirids in their vectors (Eskandari et al., 1979
; Tamada & Harrison, 1981
). However, acquisition of luteovirids has many points in common with the manner in which human immunodeficiency viruses traffic through the gastro-intestinal wall without any uncoating and multiplication (Bomsel, 1997
).
In the midgut cells of viruliferous M. persicae and A. gossypii, typical virus-laden clathrin-coated and tubular vesicles were observed as well as endosome-like bodies. The hindgut cells of both species, on the other hand, rarely harboured multivesicular bodies; instead, tubular vesicles were predominant. This observation may reflect physiological differences in the transcytosis process between midgut and hindgut and could be related to the presence of different receptors on the two types of membranes.
Release of virions from intestinal cells into the haemolymph was confirmed after microinjection of CABYV antiserum into the haemocoel. Following this treatment, aggregated particles were seen trapped by the antibodies between the basal plasmalemma and the basal lamina or within the basal lamina. No virion-containing vesicle was ever observed fused to the basal plasmalemma, presumably indicating that, once virions have attained this site, their release from the cell occurs rapidly.
The significance of the dual tissue specificity during acquisition of CABYV virions in terms of virus receptors is not understood. One possible hypothesis is that internalization of different poleroviruses such as CABYV and BWYV relies on the same biochemical partner (i.e. a receptor or receptor complex), which would be present in both intestinal locations and in both aphid species. In this hypothesis, the inability of BWYV to enter hindgut cells could be explained by different environmental conditions in the hindgut (pH, for instance) that would inhibit the virionreceptor interaction for BWYV but not for CABYV (due to putative differences in the physico-chemical properties of the virions). Alternatively, CABYV and BWYV could utilize different receptors. The receptors would differ not only in their binding affinity for the different poleroviruses but also in their distribution in the midgut and hindgut. Whatever the case, our data support the existence of a tight association between receptors on the apical surface of aphid gut cells and determinants borne on the surface of the virus particle.
The nature of the viral determinants governing tissue specificity during acquisition is not known. Sequence comparisons of structural proteins of viruses that are acquired at the posterior midgut, at the hindgut, or at both levels, do not reveal any obvious sequence motif which could be correlated to tissue specificity (for amino acid comparisons, see Guilley et al., 1994; Mayo & Ziegler-Graff, 1996
). Further information on this question could be gained using chimeric viruses created by exchanging structural protein genes between viruses with different intestinal acquisition sites. Such chimeras between BWYV and CABYV have recently been obtained and their transmissibility by M. persicae and A. gossypii is currently being tested.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Bruyère, A., Brault, V., Ziegler-Graff, V., Simonis, M. T., van den Heuvel, J. F. J. M., Richards, K., Guilley, H., Jonard, G. & Herrbach, E. (1997). Effects of mutations in the beet western yellows virus readthrough protein on its expression and packaging and on virus accumulation, symptoms, and aphid transmission. Virology 230, 323334.[CrossRef][Medline]
Clark, M. F. & Adams, A. N. (1977). Characteristics of the microplate method of enzyme-linked immunosorbent assay for the detection of plant viruses. J Gen Virol 34, 475483.[Abstract]
Eskandari, F., Sylvester, E. S. & Richardson, J. (1979). Evidence for lack of propagation of potato leafroll virus in its aphid vector, Myzus persicae. Phytopathology 69, 4547.
Garret, A., Kerlan, C. & Thomas, D. (1993). The intestine is a site of passage for potato leafroll virus from the gut lumen to the haemocoel in the aphid vector, Myzus persicae Sulz. Arch Virol 131, 377392.[Medline]
Gildow, F. E. (1982). Coated vesicle transport of luteovirus through the salivary gland of Myzus persicae. Phytopathology 72, 12891296.
Gildow, F. E. (1985). Transcellular transport of barley yellow dwarf virus into the haemocoel of the aphid vector, Rhopalosiphum padi. Phytopathology 75, 292297.
Gildow, F. E. (1987). Virus-membrane interactions involved in circulative transmission of luteoviruses by aphids. Curr Top Vector Res 4, 93120.
Gildow, F. E. (1993). Evidence for receptor-mediated endocytosis regulating luteovirus acquisition by aphids. Phytopathology 83, 270277.
Gildow, F. E. (1999). Luteovirus transmission and mechanisms regulating vector specificity. In The Luteoviridae, pp. 88113. Edited by H. G. Smith & H. Barker. Wallingford: CAB International.
Gildow, F. E. & Gray, S. (1993). The aphid salivary gland basal lamina as a selective barrier associated with vector-specific transmission of barley yellow dwarf luteovirus. Phytopathology 83, 12931302.
Gildow, F. E. & Rochow, W. F. (1980). Role of accessory salivary glands in aphid transmission of barley yellow dwarf virus. Virology 104, 97108.
Gildow, F. E., Damsteegt, V. D., Stone, A. L., Smith, O. P. & Gray, S. M. (2000). Virusvector cell interactions regulating transmission specificity of soybean dwarf luteoviruses. J Phytopathol 148, 333342.[CrossRef]
Guilley, H., Wipf-Scheibel, C., Richards, K., Lecoq, H. & Jonard, G. (1994). Nucleotide sequence of cucurbit aphid-borne yellows luteovirus. Virology 202, 10121017.[CrossRef][Medline]
Harrewijn, P. (1983). The effect of cultural measures on behaviour and population development of potato aphids and transmission of viruses. Meded Fac Landbouwwet Rijksuniv Gent 48, 791799.
Herrbach, E. (1999). Introduction of vectorvirus interactions. In The Luteoviridae, pp. 8588. Edited by H. G. Smith & H. Barker. Wallingford: CAB International.
Lecoq, H., Bourdin, D., Wipf-Scheibel, C., Bon, M., Lot, H., Lemaire, O. & Herrbach, E. (1992). A new yellowing disease of cucurbits caused by a luteovirus, cucurbit aphid-borne yellows virus. Plant Pathol 41, 749761.
Lemaire, O., Gubler, W. D., Valencia, J., Lecoq, H. & Falk, B. W. (1993). First report of cucurbit aphid-borne yellows luteovirus in the United States. Plant Dis 77, 1169.
Lett, J.-M., Granier, M., Hippolyte, I., Grondin, M., Royer, M., Blanc, S., Reynaud, B. & Peterschmitt, M. (2002). Spatial and temporal distribution of geminiviruses in leafhoppers of the genus Cicadulina monitored by conventional and quantitative polymerase chain reaction. Phytopathology 92, 6574.
Mayo, M. A. & D'Arcy, C. J. (1999). Family Luteoviridae: a reclassification of luteoviruses. In The Luteoviridae, pp. 1522. Edited by H. G. Smith & H. Barker. Wallingford: CAB International.
Mayo, M. A. & Ziegler-Graff, V. (1996). Molecular biology of luteoviruses. Adv Virus Res 46, 413460.[Medline]
Mostov, K. E. Verges M. & Altschuler, Y. (2000). Membrane traffic in polarized epithelial cells. Curr Opin Cell Biol 12, 483490.[CrossRef][Medline]
O'Loughlin, G. T. & Chambers, T. C. (1972). Extracellular microtubules in the aphid gut. J Cell Biol 53, 575578.
Pastan, I. & Willingham, M. C. (1985). The pathway of endocytosis. In Endocytosis, pp. 144. Edited by I. Pastan & M. C. Willingham. New York: Plenum Press.
Ponsen, M. B. (1977). Anatomy of an aphid vector: Myzus persicae. In Aphids as Virus Vectors, pp. 6382. Edited by K. F. Harris & K. Maramorosch. New York: Academic Press.
Prüfer, D., Wipf-Scheibel, C., Richards, K., Guilley, H., Lecoq, H. & Jonard, G. (1995). Synthesis of a full-length infectious cDNA clone of cucurbit aphid-borne yellows virus and its use in gene exchange experiments with structural proteins from other luteoviruses. Virology 214, 150158.[CrossRef][Medline]
Reinbold, C., Gildow, F. E., Herrbach, E., Ziegler-Graff, V., Gonçalves, M. C., van den Heuvel, J. F. J. M. & Brault, V. (2001). Studies on the role of the minor capsid protein in the transport of Beet western yellows virus through Myzus persicae. J Gen Virol 82, 19952007.
Reutenauer, A., Ziegler-Graff, V., Lot, H., Scheidecker, D., Guilley, H., Richards, K. & Jonard, G. (1993). Identification of beet western yellows luteovirus genes implicated in viral replication and particle morphogenesis. Virology 195, 692699.[CrossRef][Medline]
Roth, M. G. (1993). Endocytic receptors. Adv Cell Mol Biol Membranes 1, 1950.
Sieczkarski, S. B. & Whittaker, G. R. (2002). Dissecting virus entry via endocytosis. J Gen Virol 83, 15351545.
Smith, G. R., Borg, Z., Lockhart, B. L., Braithwait, K. S. & Gibbs, M. (2000). Sugarcane yellow leaf virus: a novel member of the Luteoviridae that probably arose by inter-species recombination. J Gen Virol 81, 18651869.
Tamada, T. & Harrison, B. D. (1981). Quantitative studies on the uptake and retention of potato leafroll virus by aphids in laboratory and field conditions. Ann Appl Biol 98, 261276.
van den Heuvel, J. F. J. M., Boerma, T. M. & Peters, D. (1991). Transmission of potato leafroll virus from plants and artificial diets by Myzus persicae. Phytopathology 81, 150154.
Received 12 June 2003;
accepted 1 September 2003.