Skip to main page content

• HOME
• Current Issue
• Archive
• Alerts
• About ASM
• Contact us
• Tech Support
• Journals.ASM.org

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Wei, T. Articles by Omura, T. Search for Related Content PubMed PubMed Citation Articles by Wei, T. Articles by Omura, T.

 Previous Article  |  Next Article 

Journal of Virology, September 2006, p. 8593-8602, Vol. 80, No. 17
0022-538X/06/$08.00+0     doi:10.1128/JVI.00537-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

The Spread of Rice Dwarf Virus among Cells of Its Insect Vector Exploits Virus-Induced Tubular Structures

Taiyun Wei,¹ Akira Kikuchi,^1, Yusuke Moriyasu,^1,2 Nobuhiro Suzuki,³ Takumi Shimizu,¹ Kyoji Hagiwara,¹ Hongyan Chen,^1,5 Mami Takahashi,^4, Tamaki Ichiki-Uehara,¹ and Toshihiro Omura^1*

Laboratory of Virology, National Agricultural Research Center, 3-1-1 Kannondai, Tsukuba, Ibaraki 305-8666, Japan,¹ Faculty of Agriculture, Ibaraki University, Ami, Ibaraki 300-0332, Japan,² Research Institute for Bioresources, Okayama University, 2-20-1 Chuo, Kurashiki, Okayama 710-0046, Japan,³ National Institute for Agro-Environmental Sciences, 3-1-3 Kannondai, Tsukuba, Ibaraki 305-8604, Japan,⁴ Biotechnology Center, Fujian Academy of Agricultural Sciences, Fuzhou, Fujian 350003, People's Republic of China⁵

Received 15 March 2006/ Accepted 4 June 2006

Top Abstract Introduction Materials and Methods Results Discussion References

 
Various cytopathological structures, known as inclusion bodies, are formed upon infection of cultured leafhopper cells by Rice dwarf virus, a member of the family Reoviridae. These structures include tubules of approximately 85 nm in diameter which are composed of the nonstructural viral protein Pns10 and contain viral particles. Such tubular structures were produced in heterologous non-host insect cells that expressed Pns10 of the virus. These tubules, when associated with actin-based filopodia, were able to protrude from the surface of cells and to penetrate neighboring cells. A binding assay in vitro revealed the specific binding of Pns10 to actin. Infection of clusters of cells was readily apparent 5 days after inoculation at a low multiplicity of infection with the virus, even in the presence of neutralizing antibodies. However, treatment of host cells with drugs that inhibited the elongation of actin filaments abolished the extension of Pns10 tubules from the surface of cells, with a significant simultaneous decrease in the extent of infection of neighboring cells. These results together revealed a previously undescribed aspect of the intercellular spread of Rice dwarf virus, wherein the virus exploits tubules composed of a nonstructural viral protein and actin-based filopodia to move into neighboring cells.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Various actin-based intercellular channels that connect animal cells to one another have been reported, such as filopodia (1, 4), cytonemes (15, 36), tunneling nanotubules (40), and membrane nanotubes (34). It has been proposed that these channels are involved in intercellular communication and are exploited for the transport of cellular macromolecules. Indeed, it was reported recently that some animal viruses exploit virus-induced actin-based cell-periphery protrusions to facilitate their intercellular movement. For example, vaccinia virus induces the formation of actin tails that allow its efficient egress from cells and intercellular spread (30, 38). Moreover, the alphaherpesvirus moves within virus-induced, actin-containing cell projections toward the tips of these projections and then enters uninfected cells (9), and the cell-to-cell transmission of retroviruses appears to exploit the virus-induced polarization of the actin cytoskeleton (12, 18, 19).

Rice dwarf virus (RDV) is a Phytoreovirus that multiplies both in plants and in an invertebrate insect vector, as can other plant reoviruses that belong to the genera Fijivirus, Phytoreovirus, and Oryzavirus in the family Reoviridae (3). RDV has a 12-segmented (S1 through S12) double-stranded RNA genome. Seven segments, namely, S1, S2, S3, S5, S7, S8, and S9, encode structural proteins, and the remainders encode nonstructural proteins (32, 53). RDV Pns6 was shown recently to support the cell-to-cell spread of a heterologous plus-sense single-stranded RNA virus in nonhost plants (24). RDV Pns10 was reported to act as a suppressor of RNA silencing in a nonhost plant (7), and Pns11 was reported to be a nucleic acid-binding protein (51).

A recent detailed cytopathological analysis, using antibodies against each of the 12 proteins encoded by the double-stranded RNA genome of RDV and insect vector cells in monolayers (VCMs) (50), revealed that the nonstructural proteins Pns6, Pns11, and Pns12 were the constituents of viroplasms. Pns4 was found to be the major component of fibril-like inclusion bodies, while Pns10, encoded by S10, was shown to be associated with tubule-like structures, as revealed by immunofluorescence microscopy. However, the nature of these tubular inclusions remains to be determined. In fact, no functional roles have been definitively assigned to any of these inclusion bodies, although viroplasms are believed to be sites of viral replication.

Animal viruses generally enter cells by receptor-mediated endocytosis and/or membrane fusion, and they leave cells by cytolysis and/or membrane budding, as in the case, for example, of vertebrate reoviruses (8, 31, 39). Plant reoviruses seem to utilize different strategies for spreading in their two hosts: plants and leafhoppers (13, 17). In plant viruses, they exploit so-called movement proteins (MPs) to facilitate cell-to-cell spread via a mechanism that involves plasmodesmata. Two different mechanisms for the cell-to-cell movement of viruses have been recognized and studied extensively. In one, MP is associated with viral RNA in a nucleoprotein complex that moves to the neighboring cell (10); in the other, MP modifies plasmodesmata by insertion of tubular structures, allowing the transport of entire viral particles (48).

Tubular structures that contain viral particles can frequently be observed by electron microscopy in plant and vector insect cells that have been infected with plant reoviruses (11, 44). These tubules have never been found in association with the cell wall or to be extended from plasmodesmata (3, 11, 44), suggesting that the tubules of plant reoviruses might not be involved in the cell-to-cell movement of viruses in diseased plants. By contrast, numerous tubules containing RDV particles were found in association with the microvilli of the midgut in viruliferous leafhoppers (29). These observations suggest that viruses might utilize these tubules for movement between cells of vector insects. To examine this possibility, we used confocal immunofluorescence and immunoelectron microscopy to investigate the constituents of tubules and the time course of their formation in RDV-infected VCMs. Our results showed that the nonstructural protein Pns10 of RDV was the major constituent of the tubules. Moreover, the virion-containing tubules were associated with actin-containing filopodia that made contact with and penetrated into neighboring cells. The association of Pns10 tubules of RDV with filopodia appeared to facilitate the intercellular movement of viruses in the presence of virus-neutralizing antibodies. Our results suggest a previously uncharacterized mode of intercellular viral spread wherein viruses exploit tubular structures, composed of a nonstructural protein of the virus, to move along filopodia and into adjacent invertebrate animal cells.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Cells and viruses. NC-24 cells, originally established from embryonic fragments dissected from eggs of Nephotettix cincticeps, were resuspended in the modified medium describe by Kimura and Omura (22). The O strain of RDV was purified from infected rice plants without CCl₄, as described previously (53).

Antibodies and reagents. We used rabbit polyclonal antibodies against the nonstructural protein Pns10 of RDV (45) and antibodies against purified RDV (46). Latrunculin A, jasplakinolide, ProLong antifade, Alexa Fluor 647 carboxylic acid, and rhodamine phalloidin were obtained from Invitrogen. Fluorescein isothiocyanate (FITC) was purchased from Pierce. LR Gold resin was obtained from Bioscience. Purified actin was obtained from Sigma-Aldrich.

Expression and purification of recombinant Pns10 of RDV. The sequence encoding Pns10 of RDV was cloned as a PCR-amplified EcoRI-XhoI fragment into the EcoRI-XhoI sites of pGADT7 (Clontech), and cloning of the appropriate fragment was verified by DNA sequencing. Then the fragment of RDV Pns10 cDNA was subcloned into the EcoRI-SalI sites of the bacterial expression vector pMAL-c2X (New England Biolabs) and expressed in Escherichia coli BL21(DE3) (Novagen). Pns10 was purified as a fusion protein with maltose-binding protein (MBP-Pns10) according to the instructions from New England Biolabs. The purified proteins were electrophoresed through a 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis gel.

Binding analysis by surface plasmon resonance. Surface plasmon resonance is a label-free technology for monitoring biomolecular interactions as they occur. It measures mass changes induced by association or dissociation between an immobilized analyte and a binding partner. Interactions between MBP-Pns10 and purified actin were identified and characterized by surface plasmon resonance technology using the BIAcore 2000 system (BIAcore AB, Uppsala, Sweden). Eight thousand response units of purified actin, diluted in 10 mM sodium acetate (pH 4.5), were immobilized on the dextran matrix of a CM5 sensor chip by using an amine coupling kit according to the manufacturer's instructions. Real-time interaction analysis was performed by injecting different concentrations of MBP-Pns10 (ranging from 3.5 to 28 nM). Binding experiments were carried out in HEPES-buffered saline (10 mM HEPES [pH 7.4],150 mM NaCl, 3 mM EDTA and 0.005% Tween 20), at 25°C and at a flow rate of 10 µg/ml. Ten millimolar HCl pulses were used to regenerate the chip surface. MBP, dissolved in HEPES-buffered saline buffer at 100 nM, was used as a negative control. Kinetic rate constants including association rates (K_on), dissociation rates (K_off), and affinity constants (K_D) were calculated on the basis of 1:1 (Langmuir) binding using BIAEVALUATION 3.0 software from BIAcore AB.

Expression by baculovirus of Pns10 of RDV. The construction of recombinant baculovirus, as well as the cell culture and inoculation of Spodoptera frugiperda (Sf9) cells, was performed as described previously (27). In brief, the coding region of the cDNA for RDV Pns10 that had been cloned into pGADT7 was subcloned into the pFastBac donor plasmid (Invitrogen). Then recombinant pFastBac was introduced into E. coli DH10 Bac cells (Invitrogen) for transposition into the bacmid. The recombinant bacmid was isolated and used to transfect Sf9 cells in the presence of CellFECTIN according to the instructions from Invitrogen. Then, at various times postinoculation (p.i.), Sf9 cells were collected, and the expression of proteins was examined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis and immunoblotting with Pns10-specific antibodies.

Immunofluorescence staining. Immunoglobulin G (IgG) was isolated from specific polyclonal antiserum using a protein A-Sepharose affinity column (Pierce). Eluted IgG was dialyzed exhaustively against phosphate-buffered saline. The IgG was conjugated directly to FITC or to Alexa Fluor 647 carboxylic acid according to the manufacturer's instructions. The antibodies against Pns10 were conjugated directly to FITC or Alexa Fluor 647 carboxylic acid; the antibodies against viral particles were conjugated directly to FITC. At various times after inoculation, VCMs or Sf9 cells on coverslips were fixed in 4% paraformaldehyde and processed for analysis of immunofluorescence as described previously (50). Actin was stained with rhodamine phalloidin. Samples were examined with a confocal microscope (LSM 510; Carl Zeiss, New York, NY) using a 63x oil immersion lens, and images were obtained with LSM 510 image browser software as described previously (50). Coverslips with mock-infected cells were included, as controls, in each experiment and were processed in the same way as coverslips with infected cells.

Electron microscopy. Sf9 cells and VCMs grown on coverslips were inoculated with recombinant baculovirus that encoded Pns10 and RDV, respectively. At various times after inoculation, cells were prepared for transmission electron microscopy as described previously (33). For immunoelectron microscopy, the cells were fixed and immunostained as described previously (50) with the Pns10-specific antibodies or viral particle-specific antibodies and 10-nm or 15-nm gold particle-conjugated goat antibodies against rabbit IgG (GAR10 or GAR15, respectively; British Bifocals International, Cardiff, United Kingdom). The specificity of labeling of ultrathin sections was monitored by incubating infected cells with preimmune rabbit serum or by incubating noninfected cells with Pns10-specific IgG.

Inhibition of formation of cell projections by various drugs. VCMs (1 day after seeding at low density) were inoculated with RDV at a multiplicity of infection (MOI) of 10, incubated for 2 h, washed twice, and incubated at 25°C. From 2 h p.i. onwards until fixation, cells were cultivated in the presence of 1 µM latrunculin A or 50 nM jasplakinolide. Sf9 cells that were infected with baculovirus that encoded Pns10 of RDV were also cultivated in the presence of 3 µM latrunculin A or 100 nM jasplakinolide. After fixation (18 h p.i. for VCMs or 2 day p.i. for Sf9 cells), drug-treated cells were stained with Pns10-specific IgG conjugated to FITC and rhodamine phalloidin and, finally, visualized by fluorescence microscopy as described above.

Examination of the spread of RDV in the absence and presence of various drugs. VCMs (1 day after seeding at low density) were inoculated with RDV at a low MOI of 0.001. After a 2-h adsorption period at 25°C, the inoculum was removed. Then cells were incubated in medium supplemented with antibodies (30 µg/ml of medium) against viral particles to neutralize extracellular viruses (21, 25). To study the ability of virus-neutralizing antibodies to inhibit infection by the virus, VCMs were fixed 24 h p.i. or 5 days p.i., stained with viral particle-specific IgG conjugated to FITC, and visualized by fluorescence microscopy. To study the effects of latrunculin A and jasplakinolide on the spread of RDV, infected VCMs were incubated with 1 µM latrunculin A or 50 nM jasplakinolide from 2 h p.i. until fixation. Cells were fixed 5 days p.i., stained for RDV antigens with viral particle-specific IgG conjugated to FITC, and visualized by fluorescence microscopy. The fluorescent cells were counted at a magnification of x10 by the focus count method (25). In this method, an infected cell and any adjoining infected cells were counted as one infectious unit. A minimum of four fields was counted for each sample from three or more independent experiments. Photomicrographs of representative florescent foci were obtained.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Nonstructural protein Pns10 of RDV is a constituent of tubular structures. In an earlier study, the nonstructural protein Pns10 was detected on tubule-like structures by immunofluorescence microscopy (50). To confirm the observation, VCMs were inoculated with RDV at an MOI of 10, fixed at 18 h p.i., and examined by electron microscopy. We observed tubules of approximately 85 nm in diameter protruding from the surface of leafhopper cells and surrounded by an extended plasma membrane (Fig. 1A). These tubules contained electron-dense particles of 70 nm in diameter, which correspond to the size of viral particles. To identify the constituents of the tubules, we examined the subcellular localization of Pns10 of RDV in infected VCM by immunoelectron microscopy. As shown in Fig. 1B, the tubules were evenly immunolabeled with antibodies against Pns10. Electron-dense particles in the tubules reacted with the antibodies against viral particles (Fig. 1C). No specific labeling was detected in noninfected cells after incubation with Pns10 or virus-specific IgG or in infected cells after incubation with preimmune rabbit serum (data not shown).

View larger version (88K): [in this window] [in a new window]

FIG. 1. Electron micrograph of tubules in virus-infected VCM 18 h p.i. (A) A tubule extended from the surface of the cell and surrounded by an extended plasma membrane. Immunogold labeling of Pns10 in tubule (B) and viral antigens in electron-dense particles within the tubule (C). Cells were immunostained with Pns10-specific IgG or viral particle-specific IgG as primary antibodies, followed by treatment with 10-nm gold particle-conjugated goat antibodies against rabbit IgG as second antibodies. Bars, 300 nm.

Tubular structures protrude from the surface of infected cells. To study the appearance of the tubular structures at the cellular level, we used immunofluorescence microscopy to visualize Pns10 tubules over the time course of infection of VCMs by RDV. VCMs were inoculated with RDV at an MOI of 10, stained at various times with Pns10-specific IgG conjugated to FITC, and examined by fluorescence microscopy. In infected cells, Pns10 was scattered throughout the cytoplasm in numerous thread-like structures as early as 10 h p.i. (Fig. 2A). As the infection proceeded, at 14 h p.i., Pns10 was abundant in the cytoplasm, where it appeared to form tubule-like structures at the periphery of cells or protruding from the cell surface (Fig. 2B). Later still, at 36 h p.i., many Pns10 tubule-like structures protruded from the surface of infected cells (Fig. 2C). These results suggested that Pns10 might be responsible for the extension of tubules from the surface of infected cells. No specific fluorescence was detected in noninfected cells after incubation with Pns10-specific IgG (data not shown).

View larger version (11K): [in this window] [in a new window]

FIG. 2. Subcellular localization of Pns10 of RDV in virus-infected VCMs, as visualized at 10 (A), 14 (B), and 36 (C) h p.i. VCMs were inoculated with RDV, immunostained with Pns10-specific IgG conjugated to FITC, and visualized by confocal fluorescence microscopy. Bars, 25 µm.

Aggregation of Pns10 to form tubules in vivo. To investigate whether Pns10 might have the inherent ability to form tubules, we inoculated Sf9 cells with recombinant baculovirus that encoded Pns10 and incubated cells for various periods of time. When cell extracts were analyzed by immunoblotting with Pns10-specific antibodies, Pns10 was detected first at 24 h p.i., and the level of the protein reached a maximum at 72 h p.i. (data not shown). Immunofluorescence staining of Pns10 in Sf9 cells that had been cultured on coverslips revealed the accumulation of Pns10 and the formation of numerous tubule-like structures within cells at 24 h p.i. (data not shown, refer to Fig. 2A). Later, at 48 h p.i., the Pns10-containing tubule-like structures were much more clearly defined, and some tubules even emerged from the surface of these non-host insect cells (data not shown, refer to Fig. 2B). Even later, at 72 h p.i., numerous tubule-like structures extended for considerable distances from the cell surface (data not shown, refer to Fig. 2C).

Immunoelectron microscopy allowed us to locate Pns10 more specifically in the tubular structures in the cytoplasm of Sf9 cells (data not shown, refer to Fig. 1B). These tubules resembled, in terms of the distribution of diameters and lengths, the tubules produced in VCMs that had been infected by wild-type RDV. Thus, expression of Pns10 alone, in the absence of viral proliferation, was sufficient for the formation of tubules in Sf9 cells. No reaction with cellular structures by incubation with Pns10-specific IgG in noninfected cells or viral structures with preimmune rabbit serum in infected cells was observed (data not shown).

Pns10 tubules associate with actin-based cell protrusions. In uninfected VCMs, filopodia that extended toward neighboring cells were readily observed at low cell density under the light microscope. As noted above, Pns10 tubules were able to protrude from cell surfaces, resembling the filopodia that extended from the surface of cells. These observations suggested that Pns10 tubules might attach themselves to filopodia. To examine this possibility, VCMs were inoculated with RDV at an MOI of 10, fixed at 18 h p.i., and stained with Pns10-specific IgG conjugated to FITC and rhodamine phalloidin. This double labeling of infected cells revealed that actin-based filopodia were filled with tubules of Pns10 that extended as far as the very tips of the projections (Fig. 3). Moreover, some tubules were associated with filopodia that penetrated the cytoplasm of adjacent cells, indicating that the tubules might move along filopodia to adjacent cells. In addition, the association of Pns10 tubules with filopodia seemed to not greatly cause the alterations of host actin-cytoskeleton compared with uninfected cells.

View larger version (31K): [in this window] [in a new window]

FIG. 3. Pns10 tubules of RDV associated with actin-containing cell protrusions. VCMs either mock infected or infected with RDV at an MOI of 10 and Sf9 cells mock infected or infected with a recombinant baculovirus that encoded Pns10 of RDV. Cells were fixed (at 18 h p.i. for VCMs or 2 days p.i. for Sf9 cells), stained with Pns10-specific IgG conjugated to FITC and rhodamine phalloidin, and visualized by confocal fluorescence microscopy. Bars, 10 µm.

To investigate whether Pns10 tubules have the inherent ability to associate with actin-based projections in the absence of other viral proteins, Sf9 cells were inoculated with recombinant baculovirus that encoded Pns10, fixed at 2 days p.i., and stained with Pns10-specific IgG conjugated to FITC and rhodamine phalloidin. Confocal microscopy revealed Pns10 tubules that protruded conspicuously from cell surfaces and were associated with actin-based projections from Sf9 cells (Fig. 3). Thus, Pns10 tubules were associated with actin-containing projections that extended both from host and from nonhost cells.

Binding of Pns10 to actin in vitro. Our observations by confocal microscopy indicated that Pns10 and actin might interact in vivo. To examine this possibility, we used a protein interaction assay in vitro that is based on measurements of surface plasmon resonance. For this assay, we expressed recombinant Pns10 in E. coli as a fusion protein (MBP-Pns10) and MBP as a negative control (data not shown). After affinity purification of recombinant MBP-Pns10 and MBP, we performed surface plasmon resonance measurements with a Biacore biosensor, as described in Materials and Methods. Actin was immobilized on the surface of the CM5 sensor. Then, similar amounts of purified MBP-Pns10 and MBP (data not shown), diluted in running buffer, were passed separately over the immobilized actin. Kinetic analysis of the interactions of actin with increasing concentrations of MBP-Pns10 showed a clear dose dependence (Fig. 4). The estimated values of K_on and K_off were 2.61 x 10⁴ M^–1 s^–1 and 8.84 x 10^–2 s^–1, respectively. The estimated K_D was 3.38 x 10^–6 M. The half-life for the dissociation was estimated to be less than 5 s. The rapid association and dissociation of Pns10 with actin indicate a specific but relatively low-affinity interaction. By contrast, the control assay with MBP did not show specific binding signal with immobilized actin (data not shown).

View larger version (14K): [in this window] [in a new window]

FIG. 4. Biacore surface plasmon resonance analysis of the interaction of the MBP-Pns10 with actin. Sensograms show real-time interaction of immobilized actin with increasing concentrations (3.5 nM to 28 nM, as indicated adjacent to traces) of MBP-Pns10.

Transfer of virus particles to uninfected cells in the presence of virus-neutralizing antibodies. To determine whether Pns10 tubules were related to the spread of RDV from infected to noninfected cells, we examined the spread of RDV among cells in VCMs in the presence of virus-neutralizing antibodies. VCMs were inoculated with RDV at a low MOI of 0.001, and, from 2 h p.i. onward, virus-neutralizing antibodies (30 µg/ml of medium) were added to the culture medium to inhibit infection by virus that had been released into or was present in the culture medium. At this low MOI, the infection rate was below 1% and secondary infection was easily monitored. The antibodies at this concentration used blocked the infectivity of RDV when they were added before viral inoculation in control experiments (data not shown). The spread of RDV was monitored by immunofluorescence using antibodies against intact viral particles.

For the first 1 day p.i., RDV generated similar numbers of foci of infected cells, which included one or two cells, irrespective of the presence or absence of virus-neutralizing antibodies (Fig. 5A and B). Five days after inoculation, small foci of infected cells that included 4 to 17 cells were visible in spite of the presence of virus-neutralizing antibodies, an observation that was consistent with the spread of RDV from an initially infected cell to adjacent cells in the normal inoculation condition (Fig. 5C). No single-cell infection, which indicated new infection by free virus, was observed. In parallel experiments, with medium that did not contain virus-neutralizing antibodies, large foci of infected cells that included 30 to 60 cells as well as single-cell foci in isolated regions, away from the infected cell clusters, were constantly observed (Fig. 5D). When number of foci of infected cells in the absence of neutralizing antibodies was compared 5 days p.i., viruses in the presence of neutralizing antibodies produced up to 20-fold fewer numbers of foci of infected cells, though number of infected cells in such foci increased along with the time after inoculation.

View larger version (19K): [in this window] [in a new window]

FIG. 5. Focal spread of intracellular RDV in the presence (+, panels A and C) and absence (–, panels B and D) of virus-neutralizing antibodies provides evidence of direct cell-to-cell spread. VCMs were inoculated with RDV at a low MOI of 0.001. At 2 h p.i., virus-neutralizing antibodies were added to the medium. At 1 and 5 days p.i., cells were fixed, stained with viral particle-specific IgG conjugated to FITC, and visualized by confocal fluorescence microscopy. Magnification, x10; bars, 30 µm.

Attachment of virus-containing tubules to neighboring cells in the presence of virus-neutralizing antibodies. The increase in the number of virus-infected cells in clusters in the presence of virus-neutralizing antibodies, as described above, indicated that RDV was able to spread directly from an initially infected cell to adjacent cells in the absence of infection by cell-free viruses. It is conceivable that RDV might move into adjacent cells via specific intracellular channels. To examine whether tubules might participate in the direct cell-to-cell spread of viruses, VCMs that had been seeded at low density were inoculated with RDV at a low MOI of 0.001. Then, 2 h later, virus-neutralizing antibodies were added as described above. Cells were fixed at 3 days p.i. and stained with Pns10-specific IgG conjugated to Alexa Fluor 647 carboxylic acid, viral particle-specific IgG conjugated to FITC, and rhodamine phalloidin. Confocal microscopy indicated that viral antigens were associated with several short lengths of tubules that were colocalized with filopodia and were attached to neighboring cells, which appeared either to be healthy or at an early stage of infection (Fig. 6). These results suggested that RDV particles might spread to adjacent cells from an originally infected cell, using short lengths of Pns10 tubules as transfer structures, without leaving the cytoplasm. It appears that RDV particles did not move through long contiguous tubules within filopodia.

View larger version (66K): [in this window] [in a new window]

FIG. 6. Pns10 tubules protruded from the surface of infected cells and attached to neighboring cells in the presence of virus-neutralizing antibodies. VCMs were inoculated with RDV at a low MOI of 0.001. At 2 h p.i., virus-neutralizing antibodies were added to the medium. Cells were fixed 3 days p.i., stained with Pns10-specific IgG conjugated to Alexa Fluor 647 carboxylic acid, viral particle-specific IgG conjugated to FITC, and rhodamine phalloidin, and visualized by confocal fluorescence microscopy. Bar, 10 µm.

Inhibition of the formation of filopodia and resultant effects on the intercellular spread of RDV. To examine how Pns10 tubules might associate with actin protrusions, VCMs that had been infected with RDV at an MOI of 10 were treated with chemicals that inhibit the formation of actin filament. Cells were fixed 18 h p.i. and stained with Pns10-specific IgG conjugated to FITC and rhodamine phalloidin. In preliminary experiments, a range of drug concentrations was tested (data not shown), and concentrations that exerted desired effects were determined. Treatment of cells with 1 µM latrunculin A, the inhibitor of actin filament elongation (28), resulted in the complete disassembly of actin-based filopodia (Fig. 7) in contrast to the uninfected cells (Fig. 3) and, in addition, changed the distribution of Pns10 tubules dramatically (Fig. 7), demonstrating that the inhibitor was functional. Though this inhibitory chemical had no effect on the formation of Pns10 tubules, the extension of Pns10 tubules from the surface of infected cells was almost completely inhibited (Fig. 7). Treatment with 50 nM jasplakinolide, an actin-stabilizing drug that also inhibits the elongation of actin filaments (5, 6), also suppressed the formation of actin-based filopodia, with resultant inhibition of the extension of Pns10 tubules from the surface of cells (Fig. 7). These results indicated that the association of Pns10 tubules with filopodia was mediated by the underlying actin cytoskeleton.

View larger version (36K): [in this window] [in a new window]

FIG. 7. The effects of the actin-disassembly drug latrunculin A and the actin-stabilizing drug jasplakinolide on the formation of actin-based cell projections and Pns10 tubules. VCMs were inoculated with RDV at an MOI of 10 and then cultivated in the presence of 1 µM latrunculin A or 50 nM jasplakinolide. Sf9 cells that were infected with recombinant baculovirus that encoded Pns10 of RDV were cultivated in the presence of 3 µM latrunculin A or 100 nM jasplakinolide. Cells were fixed (18 h p.i. for VCMs or 2 day p.i. for Sf9 cells), stained with Pns10-specific IgG conjugated to FITC and rhodamine phalloidin, and visualized by confocal fluorescence microscopy. Bars, 10 µm. The cell fringe corresponded to the fringe stained for actin.

In further support of the involvement of actin in efficient extension of Pns10 tubules from the cell surface, Sf9 cells that were infected with recombinant baculovirus that encoded Pns10 of RDV were also treated with latrunculin A (3 µM) or jasplakinolide (100 nM) to inhibit the formation of the actin-containing cell projection. Treated cells were fixed 2 days p.i. and stained with Pns10-specific IgG conjugated to FITC and rhodamine phalloidin. Similarly, the suppression of actin-containing cell projections formation also resulted in the inhibition of the extension of Pns10 tubules from the surface of Sf9 cells (Fig. 7).

The direct cell-to-cell spread of RDV appears to exploit Pns10 tubules that are associated with actin-based filopodia. We postulated that any inhibitors that affect the association of Pns10 tubules with actin would also affect the intercellular spread of RDV. To examine this possibility, VCMs that were seeded at low density were inoculated with RDV at a low MOI of 0.001 and, from 2 h p.i. onwards, incubated in the presence of virus-neutralizing antibodies and were treated with one of the actin disassembly or stabilizing drugs. Following 5 days of incubation, the infected cells were fixed and the effects of inhibitors were subsequently assessed using fluorescence microscopy. The virus produced foci ranging in size from 4 to 17 cells at this time (Fig. 8A). In contrast, with both the latrunculin A- and jasplakinolide-treated cells, local spread was inhibited and only single-cell or two-cell foci were visible (Fig. 8B and C). Furthermore, approximately the same numbers of infected-cell foci were produced, irrespective of the presence or absence of inhibitory chemical, demonstrating that these foci were produced from the initially infected cells. These results suggest that latrunculin A and jasplakinolide, both of which inhibited the association of Pns10 tubules with filopodia, reduced the efficiency of the intercellular spread of virus in sparsely seeded VCMs.

View larger version (5K): [in this window] [in a new window]

FIG. 8. The association of Pns10 tubules with filopodia is associated with the enhanced intercellular spread of RDV. (A) Spread of RDV in VCMs in the presence of virus-neutralizing antibodies. (B) Effects of 1 µM latrunculin A (final concentration) on the spread of RDV in VCMs in the presence of virus-neutralizing antibodies. (C) Effects of 50 nM jasplakinolide (final concentration) on the spread of RDV in VCMs in the presence of virus-neutralizing antibodies. Cells were fixed 5 days p.i., stained with viral particle-specific IgG conjugated to FITC, and visualized by confocal fluorescence microscopy. Magnification, x10; bars, 20 µm.

Taken together, our data indicate that the association of Pns10 with filopodia could facilitate the intercellular spread of RDV in the presence of virus-neutralizing antibodies.

Top Abstract Introduction Materials and Methods Results Discussion References

 
Visualization of tubules in cells infected with RDV by immunoelectron microscopy (Fig. 1B) and by immunofluorescence microscopy (Fig. 2) indicated that the tubules were composed of the nonstructural protein Pns10 that is encoded by the RDV genome. Expression of Pns10 in Sf9 cells, a nonhost of RDV, resulted in the formation of tubular structures with dimensions and appearance similar to those of tubules in RDV-infected VCMs. These observations indicated that Pns10 is responsible for the formation of the tubular structure and the extension of these tubules from the cell surface is not specific to cultured host leafhopper cells and does not require other RDV-encoded components. Viral particles were packaged in tubules from the beginning of tubule formation and they were found to be associated with extending tubules (Fig. 1, data not shown).

In the presence of neutralizing antibodies, a single cell was visibly infected 24 h p.i. with RDV at a low MOI. Then small foci of infected cells became visible 5 days after inoculation (Fig. 5). By contrast, diffuse infection of surrounding cells, mainly due to released viruses, was evident in the absence of the neutralizing antibodies 5 days after inoculation (Fig. 5). These results, together with the observation that Pns10 tubules filled with viral particles were attached to neighboring healthy cells or to cells at an early stage of infection in the presence of neutralizing antibodies (Fig. 6), demonstrate that Pns10 tubules might provide a route by which RDV particles can be transported to neighboring cells even under conditions that completely neutralize viruses released from primary infected cells.

Virion-containing tubules, when associated with actin-based filopodia, were able to protrude from the surface of infected cells and to penetrate the cytoplasm of neighboring cells (Fig. 3 and 6). This association of viral particles-containing Pns10 tubules with filopodia was believed to facilitate the intercellular spread of virus. The observations that suppression of the extension of actin filaments by inhibitory chemicals was accompanied by suppression of the extension of Pns10 tubules from the surface of infected cells (Fig. 7) and also by reduced intercellular spread of RDV in the presence of virus-neutralizing antibodies (Fig. 8) support this hypothesis. Furthermore, the inhibitory chemicals had no effect on the formation of Pns10 tubules inside infected VCMs (Fig. 7) and on the replication of RDV in VCMs (data not shown). Our results appear to reveal a previously uncharacterized aspect of the intercellular spread of RDV, wherein viruses exploit tubules composed of a nonstructural viral protein to move along actin-based dynamic filopodia to uninfected adjacent cells. RDV containing tubules observed in microvilli of the midgut in viruliferous leafhoppers (29) seem to correspond to the structure found in this study, which should be clarified in future.

The process mentioned above has been described as "F-actin flow" and has been studied extensively using artificial beads (26, 43). The way in which Pns10 tubules engage in F-actin flow remains to be determined. The on and off rates and affinities observed in the surface plasmon resonance assays with actin and MBP-Pns10 revealed a specific but relatively low-affinity interaction in vitro (Fig. 4). Moreover, the fast dissociation of Pns10 proteins from actin results in a less-stable Pns10 protein-actin binding. This instability of Pns10 protein with actin may identify the possible mechanisms of the transport of Pns10 tubules along actin filaments in infected cells. That is, it appears that strong linkage to the actin cytoskeleton of Pns10 tubules might only occur briefly as Pns10 tubules move along filopodia. Alternatively, it is possible that Pns10 tubules engage a molecular clutch and exploit a slippage type of motility rather than linking rigidly to actin filaments.

The nature of the extension of actin-based filopodia between insect cells is reflected by the current model for the actomyosin-based organization of plasmodesmata in plant cells (2, 14, 35, 52). In intact plants, MPs of several plant viruses have the ability to form tubules that extend between cells through plasmodesmata, apparently displacing the desmotubule, and provide a channel through which virions may pass (16, 20, 47, 48). In this respect, the transport of Pns10 tubules along filopodia to adjacent insect vector cells in VCMs resembles the extension of the MP tubules of several plant viruses between plant cells through plasmodesmata (23, 41). Thus, the seemingly different movement strategies exploited by plant viruses might have originated from some similar ancestral transport archetype(s) in plant hosts or insect vectors.

Intracellular endosomes/lysosomes have been studied extensively to transport along the membrane nanotubule or filopodia from cell to cell (37, 40, 42, 49). It will be interesting to investigate whether viruses can also exploit membrane nanotubules or filopodia to facilitate intercellular spread. In the case of RDV, viral particles are engulfed by a membrane enveloped structure (tubule), which accumulate in filopodia and move from cell to cell in VCMs. The tubular structures composed of Pns10 of RDV seem, functionally, reminiscent of membrane-enveloped vesicular structures. Therefore, it is intriguing to speculate that the actin cytoskeleton might serve the Pns10 tubules of RDV as "intracellular vesicles" and allow them to move into adjacent cells. However, RDV is different from several vertebrate-infecting viruses that modify and employ actin-based protrusions for cell-to-cell spread without the production of structures that resemble Pns10 tubules of RDV (9, 18, 19, 30).

Recently, Cao et al. reported that RDV Pns10 as a suppressor of RNA silencing in a nonhost plant (7). It will be interesting to examine the relationship between the ability of Pns10 to form tubules and its ability to suppress RNA silencing.

 
This project was supported by a Postdoctoral Fellowship for Foreign Researchers (15.03567) from the Japan Society for the Promotion of Science, by a Grant-in-Aid for Scientific Research (B; no. 15380038) from the Japan Society for the Promotion of Science, and by a Grant-in-Aid for Scientific Research on Priority Areas (Structures of Biological Macromolecular Assemblies) from the Ministry of Education, Culture, Sports, Science, and Technology of Japan.

 
* Corresponding author. Mailing address: National Agricultural Research Center, Tsukuba, Ibaraki 305-8666, Japan. Phone: 81 29 838 8932. Fax: 81 29 838 8929. E-mail: toomura{at}affrc.go.jp.

Present address: Gene Experiment Center, University of Tsukuba, Tsukuba, Ibaraki 305-8572, Japan.

Present address: National Agricultural Research Center, 1-2-1 Inada, Joetsu, Niigata 943-0193, Japan.

Top Abstract Introduction Materials and Methods Results Discussion References

 

1
1. Baluska, F., A. Hlavacka, D. Volkmann, and D. Menzel. 2004. Getting connected: actin-based cell-to-cell channels in plants and animals. Trends Cell Biol. 14:404-408.[CrossRef][Medline]
2
2. Baluska, F., F. Cvrckova, J. Kendrick-Jones, and D. Volkmann. 2001. Sink plasmodesmata as gateways for phloem unloading. Myosin VIII and calreticulin as molecular determinants of sink strength? Plant Physiol. 126:39-46.[Free Full Text]
3
3. Boccardo, G., and R. G. Milne. 1984. Plant reovirus group, p. 1-7. In A. F. Morant and B. D. Harrison (ed.), CM/AAB descriptions of plant viruses, no. 294. Commonwealth Mycological Institute and Association of Applied Biologists, Unwin Brothers Ltd., The Gresham Press, Old Woking, England.
4
4. Borisy, G. G., and T. M. Svitkina. 2000. Actin machinery: pushing the envelope. Curr. Opin. Cell Biol. 12:104-112.[CrossRef][Medline]
5
5. Bubb, M., A. Senderowicz, E. Sausville, K. Duncan, and E. Korn. 1994. Jasplakinolide, a cytotoxic natural product, induces actin polymerization and competitively inhibits the binding of phalloidin to F-actin J. Biol. Chem. 269:14869-14871.[Abstract/Free Full Text]
6
6. Bubb, M., I. Spector, B. Beyer, and K. Fosen. 2000. Effects of jasplakinolide on the kinetics of actin polymerization. J. Biol. Chem. 275:5163-5170.[Abstract/Free Full Text]
7
7. Cao, X., P. Zhou, X. Zhang, S. Zhu, X. Zhong, Q. Xiao, B. Ding, and Y. Li. 2005. Identification of an RNA silencing suppressor from a plant double-stranded RNA virus. J. Virol. 79:13018-13027.[Abstract/Free Full Text]
8
8. Estes, M. K. 2001. Rotaviruses and their replication, p. 1747-1786. In D. M. Knipe et al. (ed.), Fields virology, 4th ed. Lippincott Williams & Wilkins, Philadelphia, Pa.
9
9. Favoreel, H. W., G. Van Minnebruggen, D. Adriaensen, and H. J. Nauwynck. 2005. Cytoskeletal rearrangements and cell extensions induced by the US3 kinase of an alphaherpesvirus are associated with enhanced spread. Proc. Natl. Acad. Sci. USA 102:8990-8995.[Abstract/Free Full Text]
10
10. Fujiwara, T., D. Giesman-Cookmeyer, B. Ding, S. A. Lommel, and W. J. Lucas. 1993. Cell-to-cell trafficking of macromolecules through plasmodesmata potentiated by the red clover necrotic mosaic virus movement protein. Plant Cell 5:1783-1794.[Abstract]
11
11. Fukushi, I., E. Shikata, and I. Kimura. 1962. Some morphological characters of rice dwarf virus. Virology 18:192-205.[CrossRef][Medline]
12
12. Garcia, E., M. Pion, A. Pelchen-Matthews, L. Collinson, J. F. Arrighi, G. Blot, F. Leuba, J. M. Escola, N. Demaurex, M. Marsh, and V. Piguet. 2005. HIV-1 trafficking to the dendritic cell-T-cell infectious synapse uses a pathway of tetraspanin sorting to the immunological synapse. Traffic 6:488-501.[CrossRef][Medline]
13
13. Gray, S. M., and N. Banerjee. 1999. Mechanisms of arthropod transmission of plant and animal viruses. Microbiol. Mol. Biol. Rev. 63:128-148.[Abstract/Free Full Text]
14
14. Heinlein, M. 2002. Plasmodesmata: dynamic regulation and role in macromolecular cell-to-cell signaling. Curr. Opin. Plant Biol. 5:543-552.[CrossRef][Medline]
15
15. Hsiung, F., F. A. Ramirez-Weber, D. D. Iwaki, and T. B. Kornberg. 2005. Dependence of drosophila wing imaginal disc cytonemes on decapentaplegic. Nature 437:560-563.[CrossRef][Medline]
16
16. Huang, Z., Y. Han, and S. H. Howell. 2000. Formation of surface tubules and fluorescent foci in Arabidopsis thaliana protoplasts expressing a fusion between the green fluorescent protein and the cauliflower mosaic virus movement protein. Virology 271:58-64.[CrossRef][Medline]
17
17. Hull, R. 2001. Transmission 1: by in vertebrates, nematodes and fungi, p. 485-532. In R. E. F. Matthews (ed.), Matthews' plant virology, 4th ed. Academic Press, Inc., San Diego, Calif.
18
18. Igakura, T., J. Stinchcombe, P. Goon, G. Taylor, J. Weber, G. Griffiths, Y. Tanaka, M. Osame, and C. Bangham. 2003. Spread of HTLV-I between lymphocytes by virus-induced polarization of the cytoskeleton. Science 299:1713-1716.[Abstract/Free Full Text]
19
19. Jolly, C., K. Kashefi, M. Hollinshead, and Q. J. Sattentau. 2004. HIV-1 cell to cell transfer across an Env-induced, actin-dependent synapse. J. Exp. Med. 199:283-293.[Abstract/Free Full Text]
20
20. Kasteel, D. T., M. C. Perbal, J. C. Boyer, J. Wellink, R. W. Goldbach, A. J. Maule, and J. W. van Lent. 1996. The movement proteins of cowpea mosaic virus and cauliflower mosaic virus induce tubular structures in plant and insect cells. J. Gen. Virol. 77:2857-2864.[Abstract/Free Full Text]
21
21. Kimura, I. 1986. A study of rice dwarf virus in vector cell monolayers by fluorescent antibody focus counting. J. Gen. Virol. 67:2119-2124.
22
22. Kimura, I., and T. Omura. 1988. Leafhopper cell cultures as a means for phytoreovirus research. Adv. Dis. Vector Res. 5:111-135.
23
23. Lazarowitz, S. G., and R. N. Beachy. 1999. Viral movement proteins as probes for investigating intracellular and intercellular trafficking in plants. Plant Cell 11:535-548.[Free Full Text]
24
24. Li, Y., Y. M. Bao, C. H. Wei, Z. S. Kang, Y. W. Zhong, P. Mao, G. Wu, Z. L. Chen, J. Schiemann, and R. S. Nelson. 2004. Rice dwarf phytoreovirus segment S6-encoded nonstructural protein has a cell-to-cell movement function. J. Virol. 78:5382-5389.[Abstract/Free Full Text]
25
25. Liu, H. Y., and L. M. Black. 1978. Neutralization of infectivity of potato yellow dwarf virus and wound tumor virus assayed on vector-cell monolayers. Phytopathology 68:1243-1248.
26
26. Mitchison, T., and M. Kirschner. 1988. Cytoskeletal dynamics and nerve growth. Neuron 1:761-767.[CrossRef][Medline]
27
27. Miyazaki, N., K. Hagiwara, H. Naitow, T. Higashi, R. H. Cheng, T. Tsukihara, A. Nakagawa, and T. Omura. 2005. Transcapsidation and the conserved interactions of two major structural proteins of a pair of phytoreoviruses confirm the mechanism of assembly of the outer capsid layer. J. Mol. Biol. 345:229-237.[CrossRef][Medline]
28
28. Morton, W., K. Ayscough, and P. McLaughlin. 2000. Latrunculin alters the actin-monomer subunit interface to prevent polymerization. Nat. Cell Biol. 2:376-378.[CrossRef][Medline]
29
29. Nasu, S. 1965. Electron microscopy studies on transovarial passage of rice dwarf virus. Jpn. J. Appl. Entomol. Zool. 9:225-237.
30
30. Newsome, T. P., N. Scaplehorn, and M. Way. 2004. SRC mediates a switch from microtubule- to actin-based motility of vaccinia virus. Science 306:124-129.[Abstract/Free Full Text]
31
31. Nibert, M. L., and L. A. Schiff. 2001. Reoviruses and their replication, p. 1679-1728. In D. M. Knipe et al. (ed.), Fields virology, 4th ed. Lippincott Williams & Wilkins, Philadelphia, Pa.
32
32. Omura, T., and J. Yan. 1999. Role of outer capsid proteins in transmission of Phytoreovirus by insect vectors. Adv. Virus Res. 54:15-43.[Medline]
33
33. Omura, T., J. Yan, B. Zhong, M. Wada, Y. Zhu, M. Tomaru, W. Maruyama, A. Kikuchi, Y. Watanabe, I. Kimura, and H. Hibino. 1998. The P2 protein of rice dwarf phytoreovirus is required for adsorption of the virus to cells of the insect vector. J. Virol. 72:9370-9373.[Abstract/Free Full Text]
34
34. Önfelt, B., S. Nedvetzki, K. Yanagi, and D. M. Davis. 2004. Cutting edge: membrane nanotubes connect immune cells. J. Immunol. 173:1511-1513.[Abstract/Free Full Text]
35
35. Oparka, K. J. 2004. Getting the message across: how do plant cells exchange macromolecular complexes? Trends Plant Sci. 9:33-41.[CrossRef][Medline]
36
36. Ramirez-Weber, F. A., and T. B. Kornberg. 1999. Cytonemes: cellular processes that project to the principal signaling center in Drosophila imaginal discs. Cell 97:599-607.[CrossRef][Medline]
37
37. Reed, B. C., C. Cefalu, B. H. Bellaire, J. A. Cardelli, T. Louis, J. Salamon, M. A. Bloecher, and R. C. Bunn. 2005. GLUT1CBP (TIP2/GIPC1) interactions with GLUT1 and myosin VI: evidence supporting an adapter function for GLUT1CBP. Mol. Biol. Cell 16:4183-4201.[Abstract/Free Full Text]
38
38. Rietdorf, J., A. Ploubidou, I. Reckmann, A. Holmstrom, F. Frischknecht, M. Zettl, T. Zimmermann, and M. Way. 2001. Kinesin-dependent movement on microtubules precedes actin-based motility of vaccinia virus. Nat. Cell Biol. 3:992-1000.[CrossRef][Medline]
39
39. Roy, P. 2001. Orbiviruses and their replication, p. 1835-1869. In D. M. Knipe et al. (ed.), Fields virology, 4th ed. Lippincott Williams & Wilkins, Philadelphia, Pa.
40
40. Rustom, A., R. Saffrich, I. Markovic, P. Walther, and H. H. Gerdes. 2004. Nanotubular highways for intercellular organelle transport. Science 303:1007-1010.[Abstract/Free Full Text]
41
41. Scholthof, H. B. 2005. Plant virus transport: motions of functional equivalence. Trends Plant Sci. 10:376-382.[CrossRef][Medline]
42
42. Scott, G., S. Leopardi, S. Printup, and B. C. Madden. 2002. Filopodia are conduits for melanosome transfer to keratinocytes. J. Cell Sci. 115:1441-1451.[Abstract/Free Full Text]
43
43. Sheetz, M. P., S. Turney, H. Qian, and E. L. Elson. 1989. Nanometre-level analysis demonstrates that lipid flow does not drive membrane glycoprotein movements. Nature 340:284-288.[CrossRef][Medline]
44
44. Shikata, E. 1969. Electron microscopic studies on rice viruses, p. 223-240. In IRRI: the virus disease of the rice plant. Johns Hopkins University Press, Baltimore, Md.
45
45. Suzuki, N., M. Sugawara, T. Kusano, H. Mori, and Y. Matsuura. 1994. Immunodetection of rice dwarf phytoreoviral proteins in both insect and plant hosts. Virology 202:41-48.[CrossRef][Medline]
46
46. Takahashi, Y., M. Tomiyama, H. Hibino, and T. Omura. 1994. Conserved primary structures in core capsid proteins and reassembly of core particles and outer capsids between rice gall dwarf and rice dwarf phytoreoviruses. J. Gen. Virol. 75:269-275.[Abstract/Free Full Text]
47
47. Thomas, C. L., and A. J. Maule. 1999. Identification of inhibitory mutants of Cauliflower mosaic virus movement protein function after expression in insect cells. J. Virol. 73:7886-7890.[Abstract/Free Full Text]
48
48. van Lent, J., M. Storms, F. van der Meer, J. Wellink, and R. Goldbach. 1991. Tubular structures involved in movement of cowpea mosaic virus are also formed in infected cowpea protoplasts. J. Gen. Virol. 72:2615-2623.[Abstract/Free Full Text]
49
49. Vidulescu, C., S. Clejan, and K. C. O'Connor. 2004. Vesicle traffic through intercellular bridges in DU 145 human prostate cancer cells. J. Cell. Mol. Med. 8:388-396.[Medline]
50
50. Wei, T., T. Shimizu, K. Hagiwara, A. Kikuchi, Y. Moriyasu, N. Suzuki, H. Chen, and T. Omura. 2006. Pns12 protein of Rice dwarf virus is essential for formation of viroplasms and nucleation of viral-assembly complexes. J. Gen. Virol. 87:429-438.[Abstract/Free Full Text]
51
51. Xu, H., Y. Li, Z. Mao, Y. Li, Z. Wu, L. Qu, C. An, X. Ming, J. Schiemann, R. Casper, and Z. Chen. 1998. Rice dwarf phytoreovirus segment S11 encodes a nucleic acid-binding protein. Virology 240:267-272.[CrossRef][Medline]
52
52. Zambryski, P., and K. Crawford. 2000. Plasmodesmata: gatekeepers for cell-to-cell transport of developmental signals in plants. Annu. Rev. Cell Dev. Biol. 16:393-421.[CrossRef][Medline]
53
53. Zhong, B., A. Kikuchi, Y. Moriyasu, T. Higashi, K. Hagiwara, and T. Omura. 2003. A minor outer capsid protein, P9, of Rice dwarf virus. Arch. Virol. 148:2275-2280.[CrossRef][Medline]

---

Journal of Virology, September 2006, p. 8593-8602, Vol. 80, No. 17
0022-538X/06/$08.00+0     doi:10.1128/JVI.00537-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

This article has been cited by other articles:

• Wei, T., Hibino, H., Omura, T. (2008). Rice dwarf virus is engulfed into and released via vesicular compartments in cultured insect vector cells. J. Gen. Virol. 89: 2915-2920 [Abstract] [Full Text]  
• Zhou, F., Pu, Y., Wei, T., Liu, H., Deng, W., Wei, C., Ding, B., Omura, T., Li, Y. (2007). The P2 capsid protein of the nonenveloped rice dwarf phytoreovirus induces membrane fusion in insect host cells. Proc. Natl. Acad. Sci. USA 104: 19547-19552 [Abstract] [Full Text]  
• Katayama, S., Wei, T., Omura, T., Takagi, J., Iwasaki, K. (2007). Three-Dimensional Architecture of Virus-Packed Tubule. J Electron Microsc (Tokyo) 56: 77-81 [Abstract] [Full Text]  

This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Services Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Reprints and Permissions Copyright Information Books from ASM Press MicrobeWorld Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Wei, T. Articles by Omura, T. Search for Related Content PubMed PubMed Citation Articles by Wei, T. Articles by Omura, T.