Rice dwarf virus is engulfed into and released via
vesicular compartments in cultured insect vector
Taiyun Wei, Hiroyuki Hibino and Toshihiro Omura
National Agricultural Research Center, 3-1-1 Kannondai, Tsukuba, Ibaraki 305-8666, Japan
Received 12 March 2008
Accepted 24 June 2008
Vector insect cells infected with Rice dwarf virus had vesicular compartments containing viral
particles located adjacent to the viroplasm when examined by transmission electron and confocal
microscopy. Such compartments were often at the periphery of infected cells. Inhibitors of
vesicular transport, brefeldin A and monensin, and an inhibitor of myosin motor activity,
butanedione monoxime, abolished the formation of such vesicles and prevented the release of
viral particles from infected cells without significant effects on virus multiplication. Furthermore,
the actin-depolymerizing drug, cytochalasin D, inhibited the formation of actin filaments without
significantly interfering with formation of vesicular compartments and the release of viruses from
treated cells. These results together revealed intracellular vesicular compartments as a mode for
viral transport in and release from insect vector cells infected with a plant-infecting reovirus.
In vector cells grown in monolayers (VCMs), Rice dwarf
virus (RDV), a phytoreovirus, multiplies and spreads from
primarily infected cells to neighbouring cells (Wei et al.,
2006a) in addition to spreading via mature, free viral
particles. Infection via free viral particles was protected by
the addition of neutralizing antibodies to the cell culture
medium. As part of integrated studies on RDV prolifera-
tion in vector insects, we have focused our study on the
accumulation of the virus in cells of the insect vector and
on the subsequent release of the virus.
In electron micrographs of thin sections of insect tissues
infected with plant-pathogenic reoviruses, viral particles
were sequestered in spherical vesicular compartments
(Fukushi et al., 1962; Shikata & Maramorosch, 1965;
Shikata, 1969; Vidano, 1970; Omura et al., 1985). However,
the biological significance of these inclusions in RDV
infection has not been clarified because, for the most part,
the tissues examined were in the late stage of infection
which made it difficult to gather details on the formation
of these compartments. Similar vesicular compartments
appear to play a role in the release of viral particles from
cultured cells infected with animal viruses, such as severe
acute respiratory syndrome coronavirus (Ng et al., 2003)
and human immunodeficiency virus (HIV) (Nydegger
et al., 2003).
A method for the continuous culture of cells from the
leafhopper vector Nephotettix cincticeps, consisting of
VCMs, has allowed the study of infection with RDV
because such infection results in asymptomatic but
persistent infection (Peterson & Nuss, 1985; Kimura,
1986). VCMs have been used to reveal fundamental aspects
of viral activity at the cellular level, which suggests details
of the transmission, multiplication and cytopathology of
RDV in vector insects (Wei et al., 2006a, b, c, 2007, 2008).
In the present study with this system, we investigated the
role of vesicular compartments in the transport of RDV
and its release from infected VCMs by confocal and
electron microscopy, and the use of drugs that inhibit
We examined the intracellular distribution of RDV during
the late stages of viral infection when viral particles would
be released from the cells. VCMs grown on a coverslip
(15 mm in diameter) were inoculated with RDV at an
m.o.i. of 10 and fixed 36 h post-inoculation (p.i.) for
transmission electron microscopy as described previously
(Omura et al., 1998). In these figures, RDV particles,
confirmed by immunoelectron microscopy in our earlier
study (Wei et al., 2006a), were easily distinguished by their
spherical appearance and diameter (70 nm), and they were
not found in uninoculated controls (Fig. 1). Further, the
viroplasm, the site of viral replication and assembly,
reacted specifically with Pns12-specific antibodies as
reported earlier (Wei et al., 2006b, c), was easily recognized
by its characteristic appearance (Fig. 1a). As reported in
our earlier studies (Wei et al., 2006c), viral particles
distributed at the periphery of the viroplasm (Fig. 1a). We
sometimes observed viral particles in vesicular compart-
ments other than the viroplasm in the cytoplasm of RDV-
infected VCMs (Fig. 1a). These compartments varied in
size and often reached a diameter of 2 mm. Occasionally,
we observed viral particles in vesicular compartments at the
periphery of infected cells (Fig. 1b, c). Furthermore, viral
Journal of General Virology (2008), 89, 2915–2920
2008/002063G2008 SGMPrinted in Great Britain2915
particles and cellular remnants were often observed outside
of infected cells (Fig. 1c). Similar features were never seen
in uninfected cells.
To study the appearance of virus-containing vesicular
compartments in relation to the viroplasm at the cellular
level, we used confocal fluorescence microscopy to visualize
the relative localization of the viral antigen and the
viroplasm in RDV-infected cells. VCMs were inoculated
at an m.o.i. of 10 with RDV and fixed 36 h p.i. for 30 min
in 2% paraformaldehyde. Cells were immunostained with
RDV-specific antibodies conjugated with fluorescein iso-
thiocyanate (FITC) and with antibodies raised against the
non-structural protein Pns12, a constituent of the vir-
oplasm, conjugated with rhodamine and then examined by
confocal fluorescence microscopy as described previously
(Wei et al., 2006c). Viral particles were assembled in the
peripheral region of the viroplasm and formed ring-like
structures (Fig. 2, untreated panel), as described previously
(Wei et al., 2006c). RDV-specific antibodies also reacted
with numerous spherical structures in the cytoplasm or
outside of infected cells (Fig. 2, untreated panel). The
viroplasm and the spherical structure often located beside
it corresponded to the appearance in Fig. 1(a). These
observations revealed that the viroplasms differed distinctly
from the spherical structures. In our thorough examination
of immunoelectron micrographs of thin sections of RDV-
infected cells, we did not find any inclusions other than the
spherical structures we observed with immunofluorescence
staining that corresponded to the virus-containing vesi-
cular compartments. The similarity in the sizes of the
vesicular compartments observed with electron microscopy
and the spherical structures observed after immunofluor-
escence staining support these results. Thus, all our results
together suggest a possible pathway whereby viral particles,
assembled at the periphery of the viroplasm, are engulfed
by vesicular compartments. The occurrence of such
compartments at the periphery of infected cells (Fig. 1b,
c) also suggests that the virus may be released from the cells
upon fusion of the compartments with the cell membranes,
rather than be degraded within these compartments.
To confirm the existence of such a pathway, we studied the
effects of inhibitors of vesicular transport applied during
the viral propagation period. Brefeldin A (BFA) is a fungal
metabolite that disrupts the Golgi complex in many cell
types, thus inhibiting normal cellular sorting and transport
functions (Klausner et al., 1992; Pelham, 1991). Monensin
is a carboxylic ionophore that arrests vesicular transport at
a site distal to the proximal portion of the Golgi complex
(Tartakoff, 1983). These two inhibitors have been exten-
sively used to inhibit vesicular transport of several viruses
(Blank et al., 2000; Boulanger et al., 2000; Suikkanen et al.,
2003; Kolesnikova et al., 2004; Bugarcic & Taylor, 2006).
Two hours p.i. of VCMs with RDV at an m.o.i. of 10, either
0.1 or 0.5 mg BFA (Sigma) ml21, or 5 or 20 mM monensin
(Sigma) was added and incubation was continued for a
further 34 h. In preliminary experiments, we tested a range
of concentrations of the drugs to determine effective
concentrations and to avoid toxic effects of BFA and
monensin (data not shown; Wei et al., 2008). Treatment of
VCMs with 0.5 mg BFA ml21and 20 mM monensin did not
interfere with the formation of the viroplasm, composed
mainly of Pns12 (Fig. 2; Wei et al., 2006c), while both
chemicals reduced the number and size of virus-positive
spherical structures in the cytoplasm (Fig. 2).
Next, we examined whether these two drugs had any effects
on the replication of RDV in VCMs. Two hours p.i. of
VCMs with RDV at an m.o.i. of 10, BFA or monensin was
added at a range of concentrations and incubation was
continued for a further 34 h. The extracellular medium and
the cells were collected. The medium was centrifuged for
30 min at 15000 g, and the supernatant was collected and
frozen at –70 uC before analysis. The cells were subjected to
three cycles of freeze–thaw to release viral particles and
stored at –70 uC before analysis. The viral titre of each
sample was determined in duplicate using VCMs at a
Fig. 1. Transmission electron micrographs of virus-containing
vesicular compartments in vector cell monolayer (VCM) infected
with RDV at 36 h p.i. (a) Typical appearance of viral particles at the
periphery of viroplasm (VP) or within vesicular compartments
(arrow) inside cells. (b) Virus-containing vesicular compartment
(arrow) at the periphery of an infected cell. (c) Virus-containing
vesicular compartments (arrows) inside and at the periphery of a
cell. Arrowhead points to a cellular remnant containing viral
particles outside of the infected cell. Bars, 400 nm.
T. Wei, H. Hibino and T. Omura
2916 Journal of General Virology 89
magnification of 610 using a fluorescent focus assay as
described previously (Kimura, 1986). End-point titres were
calculated as means with standard deviations. As shown in
Fig. 3, BFA (0.1 and 0.5 mg ml21) and monensin (5 and
20 mM) caused more than 50-fold decrease in viral titre of
the medium. By contrast, at doses that significantly affected
Fig. 2. Confocal immunofluorescence micrographs of VCMs inoculated with RDV and treated with or without inhibitors. VCMs
were inoculated with RDV at an m.o.i. of 10, then cultured in the absence or presence of drug in the culture medium with
concentrations as follows: 0.5 mg BFA ml”1, 20 mM monensin, 2 mg CytD ml”1or 40 nM BDM. Cells were fixed at 36 h p.i.,
treated with RDV-specific antibodies conjugated to FITC to stain spherical structures and Pns12-specific antibodies
conjugated to rhodamine to stain viroplasm. The green colour in BDM-treated cells was adjusted to the level of that in BFA- and
monensin-treated cells to view better the distribution of viral antigen. The ring-like staining of viral antigens and spherical
structures in Pns12 showing viroplasms (Wei et al., 2006c) is indicated by arrows. Arrowheads show spherical structures
beside the viroplasm. Lines show the smaller virus-containing spherical structures outside the infected cells. Bars, 10 mm.
Release of Rice dwarf virus
the release of viral particles, the inhibitors did not
significantly reduce the titres of cell-associated viruses.
These results demonstrated that RDV had proliferated in
the infected cells but particle release from the cells had been
impeded by the inhibitors. Taken together, the evidence
that release of RDV from cells was suppressed to a
significant extent (Fig. 3) in the presence of the drugs,
whereby the formation of virus-positive spherical struc-
tures was significantly inhibited (Fig. 2), supported the
hypothesis that RDV particles, generated in the viroplasm,
were packaged in the spherical structures that correspond
to the vesicular compartments and then released from cells.
The actin cytoskeleton is crucial for the intracellular
trafficking of vesicular compartments (Gottlieb et al.,
1993; Radtke et al., 2006) and for the transport of RDV
between cells of the vector insect (Wei et al., 2006a, 2008).
To evaluate the involvement of the actin cytoskeleton in the
replication process of RDV, we examined the effects of
cytochalasin D (CytD), an actin-depolymerizing drug
(Sigma) (Goddette & Frieden, 1986; Sampath & Pollard,
1991), and butanedione monoxime (BDM; Sigma), a
myosin motor inhibitor (Cramer & Mitchison, 1995), on
the assembly of viral particles and their release from infected
cells. Fluorescence analysis of the materials revealed well-
organized actin filaments in untreated RDV-infected cells,
while the majority of cellular filamentous actin was
disrupted inCytD(2 mg ml21)-treated
(40 nM) did not affect formation of actin filaments, as
described previously (Wei et al., 2008). Immunostaining of
viral antigens and Pns12 in infected cells revealed viroplasms
and a number of prominent virus-associated spherical
structures in the presence of CytD (Fig. 2). Furthermore,
smaller virus-associated spherical structures occurred out-
side of infected cells in the presence of CytD (Fig. 2). By
(Fig. 2). Taken together, these results indicated that BDM
inhibited the association of viral particles with the spherical
structures, whereas CytD seemed only to change the pattern
of distribution of virus-associated vesicles although it was
previously found to interfere with the intercellular transport
of RDV (Wei et al., 2008).
Next, we collected the cells and medium separately at 36 h
p.i., and the amount of virus in each fraction was
quantified by virus infection assay as mentioned. An
analysis of extracellular RDV revealed that CytD (0.5 and
2 mg ml21) slightly reduced the release of RDV (10% of
control yield), whereas BDM (20 and 40 nM) reduced the
level of detectable virus in the medium to 98% of that in
the control (Fig. 3). Both inhibitors had negligible effects
on the production of cell-associated viruses (Fig. 3),
suggesting that they had little effect on viral replication.
These results suggested that myosin has a major role in the
release of RDV, in contrast to a less prominent role for
actin filaments. Similarly, myosin also plays more import-
ant roles than actin in vesicle trafficking (Dura ´n et al.,
2003) and HIV release from infected host cells (Sasaki et al.,
1995). All our results clearly showed a relationship between
the formation of virus-associated spherical structures and
the release of progeny viruses from cells, demonstrating
that the spherical structures that correspond to vesicular
compartments play an important role in the accumulation
of viral particles and their release from cells.
As shown in Fig. 3, none of the inhibitors at the various
concentrations substantially reduced the titre of cell-associated
viruses. These results suggest that the inhibitors had no
Fig. 3. Effects of inhibitors on the accumula-
tion of cell-associated (open bars) and extra-
cellular (black bars) viruses in VCMs infected
with RDV (m.o.i. of 10) at 36 h p.i. as
percentage of titre in untreated cells (titre in
treated cells/titre in untreated cells ?100).
Viral titre was determined in duplicate by a
fluorescent focus assay (Kimura, 1986). Error
bars indicate the standard deviation.
T. Wei, H. Hibino and T. Omura
2918 Journal of General Virology 89
significant effect on the RDV replication. Therefore, the
reduction in the release of RDV in the medium in the presence
of BFA, monensin and BDM was due specifically to
interference with post-replication phenomena. CytD, an
transport of viral particles to neighbouring cells through Pns10
tubules, which need actin for extension (Wei et al., 2008). This
phenomenon was confirmed in this study (data not shown)
and this drug did not affect spherical structure formation (Fig.
2), viral replication (Fig. 3) or release of viral particles from
infected cells (Fig. 3), suggesting that this chemical also worked
in specific manners under our conditions.
The apparent restriction of viral particles in spherical
structures in untreated and CytD-treated cells was not
observed in BFA-, monensin- and BDM-treated cells (Fig.
2), although the virus in BFA-, monensin- and BDV-treated
cells proliferated to a level similar to that in CytD-treated
and untreated cells (Fig. 3). These results suggest that viruses
are concentrated in the spherical structures in untreated and
CytD-treated cells, but such accumulation was prevented in
the presence of BFA, monensin and BDM. Electron
microscopic observation showed that nascent viruses were
engulfed in vesicles and released from the surface of cells
(Fig. 1). In confocal microscopy, the spherical structure
formation (Fig. 2) was accompanied by viral release (Fig. 3),
and inhibition of the spherical structure formation (Fig. 2)
resulted in the failure of viral release from infected cells (Fig.
3). All these results suggest that the spherical structure, i.e.
the vesicular compartment, plays an important role in the
release of viral particles from infected leafhopper cells.
It is unclear whether the mechanism involved in the release
of viral particles from infected cells is also involved in
sequestering a large fraction of infectious viruses within
infected VCMs because a high titre of infectious RDV
remains associated with infected cells. However, this type
of sequestration might be a reason for the longevity of
virus-infected cultured cells and the absence of any
apparent deleterious effect of viral infection on viruliferous
vector insects (Peterson & Nuss, 1985; Kimura, 1986). This
hypothesis might also hold for viruses in the genera
Fijivirus, Phytoreovirus and Oryzavirus, which also multiply
in both plants and vector insects.
This project was supported by a Postdoctoral Fellowship for Foreign
Researchers (15.03567) 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, and by the Program for Promotion of Basic Research Activities
for Innovative Biosciences (PROBRAIN).
Blank, C. A., Anderson, D. A., Beard, M. & Lemon, S. M. (2000).
Infection of polarized cultures of human intestinal epithelial cells with
hepatitis A virus: vectorial release of progeny virions through apical
cellular membranes. J Virol 74, 6476–6484.
Boulanger, D., Smith, T. & Skinner, M. A. (2000). Morphogenesis and
release of fowlpox virus. J Gen Virol 81, 675–687.
Bugarcic, A. & Taylor, J. A. (2006). Rotavirus nonstructural
glycoprotein NSP4 is secreted from the apical surfaces of polarized
epithelial cells. J Virol 80, 12343–12349.
Cramer, L. P. & Mitchison, T. J. (1995). Myosin is involved in
postmitotic cell spreading. J Cell Biol 131, 179–189.
Dura ´n, J. M., Valderrama, F., Castel, S., Magdalena, J., Toma ´s, M.,
Hosoya, H., Renau-Piqueras, J., Malhotra, V. & Egea, G. (2003).
Myosin motors and not actin comets are mediators of the actin-based
Golgi-to-endoplasmic reticulum protein transport. Mol Biol Cell 14,
Fukushi, T., Shikata, E. & Kimura, I. (1962). Some morphological
characters of rice dwarf virus. Virology 18, 192–205.
Goddette, D. W. & Frieden, C. (1986). Actin polymerization. The
mechanism of action of cytochalasin D. J Biol Chem 261, 15974–15980.
Gottlieb, T. A., Ivanov, I. E., Adesnik, M. & Sabatini, D. D. (1993). Actin
microfilaments play a critical role inendocytosis at theapical but notthe
basolateral surface of polarized epithelial cells. J Cell Biol 120, 695–710.
Kimura, I. (1986). A study of rice dwarf virus in vector cell
monolayers by fluorescent antibody focus counting. J Gen Virol 67,
Klausner, R. D., Donaldson, J. G. & Lippincott-Schwartz, J. (1992).
Brefeldin A: insights into control of membrane traffic and organelle
structure. J Cell Biol 116, 1071–1080.
Kolesnikova, L., Bamberg, S., Bergho ¨fer, B. & Becker, S. (2004). The
matrix protein of Marburg virus is transported to the plasma
membrane along cellular membranes: exploiting the retrograde late
endosomal pathway. J Virol 78, 2382–2393.
Ng, M. L., Tan, S. H., See, E. E., Ooi, E. E. & Ling, A. E. (2003).
Proliferative growth of SARS coronavirus in vero E6 cells. J Gen Virol
Nydegger, S., Foti, M., Derdowski, A., Spearman, P. & Thali, M.
(2003). HIV-1 egress is gated through late endosomal membranes.
Traffic 4, 902–910.
Omura, T., Hibino, H., Inoue, H. & Tsuchizaki, T. (1985). Particles of
rice gall dwarf virus in thin sections of diseased rice plants and insect
vector cells. J Gen Virol 66, 2581–2587.
Omura, T., Yan, J., Zhong, B., Wada, M., Zhu, Y., Tomaru, M.,
Maruyama, W., Kikuchi, A., Watanabe, Y. & other authors (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.
Pelham, H. R. B. (1991). Multiple targets for brefeldin A. Cell 67,
Peterson, A. J. & Nuss, D. L. (1985). Wound tumor virus polypeptide
synthesis in productive noncytopathic infection of cultured insect
vector cells. J Virol 56, 620–624.
Radtke, K., Dohner, K. & Sodeik, B. (2006). Viral interactions with the
cytoskeleton: a hitchhiker’s guide to the cell. Cell Microbiol 8, 387–400.
Sampath, P. & Pollard, T. D. (1991). Effects of cytochalasin, phall-
oidin, and pH on the elongation of actin filaments. Biochemistry 30,
Sasaki, H., Nakamura, M., Ohno, T., Matsuda, Y., Yuda, Y. &
Nonomura, Y. (1995). Myosin-actin interaction plays an important
role in human immunodeficiency virus type 1 release from host cells.
Proc Natl Acad Sci U S A 92, 2026–2030.
Shikata, E. (1969). Electron microscopic studies on rice viruses. In
The Virus Disease of the Rice Plant, pp. 223–240. Baltimore: Johns
Release of Rice dwarf virus