Department of Microbiology and Immunology, Emory University School of Medicine, Atlanta, GA 30322, USA1
Author for correspondence: Richard Compans. Fax +1 404 727 8250. e-mail compans{at}microbio.emory.edu
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Abstract |
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Introduction |
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The cellular cytoskeleton has been reported to play an important role in transcription, maturation, morphogenesis and budding of a number of enveloped viruses, including Newcastle disease virus (NDV), RSV, SeV and measles virus in the family Paramyxoviridae (Cudmore et al., 1997 ). Paramyxoviruses were among the first viruses to be reported to contain actin as a component within the virion (Örvell, 1978
; Sundqvist & Ehrnst, 1976
; Tyrrell & Norrby, 1978
; Wang et al., 1976
); however, its function in virus structure and replication is not understood. Actin occurs in two forms, a globular, monomeric form that represents the soluble pool of actin and a filamentous form that constitutes the actin microfilaments of the cytoskeletal framework. Actin microfilaments have been reported to be involved in the movement of viral envelope proteins to the cell surface, in communication between envelope proteins and the nucleocapsids in the plasma membrane and in virus budding (Bohn et al., 1986
; Stallcup et al., 1983
; Tyrrell & Ehrnst, 1979
). It was also suggested that the M protein of paramyxoviruses is essential for the incorporation of actin within the virion (Giuffre et al., 1982
). In measles virus infection, actin-like microfilaments were reported to project into developing particles at the cell membrane and actin was also found to be present in released measles virions (Bohn et al., 1986
). Therefore, it was of interest to investigate further whether the cellular cytoskeleton has any effect on PIV morphology and virus release.
In this study, surface immunofluorescence was used to analyse PIV filament formation. We compared the formation of filamentous virus particles by different PIVs in polarized and non-polarized epithelial cells. Furthermore, we studied the effects of cytochalasin D (CD), an actin-disrupting agent, and jasplakinolide, a potent inducer of actin polymerization, on virus morphology and virus release.
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Methods |
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Haemagglutination (HA) assay.
HA titres of the released virus particles were determined by incubating equal volumes of serial twofold dilutions of culture medium in PBS-def (PBS deficient in Mg2+ and Ca2+) with guinea pig red blood cells (final concentration 0·25%) for 1 h at 4 °C.
Plaque assay.
Serial tenfold dilutions of the culture media collected at designated time-points were added to monolayers of Vero 76 cells. After 1 h incubation at 37 °C, the inoculum was removed and overlaid with 2% white agar mixed at a 1:1 ratio with 2xDMEM containing 4% FBS. Four days after overlaying, 0·05% neutral red stain was added onto the agar. Plaques were counted after 6 h.
Virus infection.
Cells were grown to 80% confluence on 12 mm glass coverslips for immunofluorescence studies or 60 mm plastic dishes for virus yield studies and were inoculated with 1 p.f.u. of HPIV2 or HPIV3 stocks per cell. After adsorption for 1 h at 37 °C, the inoculum was removed and the cells were incubated in DMEM supplemented with 2% FBS. A stock solution of CD (5 mg/ml in DMSO, Sigma) was diluted 100-fold and jasplakinolide (500 mM in DMSO, Molecular Probes) was diluted 10-fold in medium just prior to use, added to the virus culture medium after the 1 h inoculation period and maintained throughout the infection period. Control cells were incubated with the respective amount of DMSO.
35S-radiolabelling of HPIV2 and SDSPAGE.
Vero C1008 cells were grown to 80% confluence on 100 mm tissue culture dishes and were inoculated with 1 p.f.u. of HPIV2 stock per cell. After adsorption for 1 h at 37 °C, the inoculum was removed and the cells were incubated in DMEM supplemented with 2% FBS. CD (5 µg/ml) was added to the culture medium after the 1 h inoculation period and maintained throughout the infection period. At 12 h post-infection (p.i.), cells were labelled continuously with 50 µCi [35S]methionine/[35S]cysteine per ml of a mixture of 75% methionine-deficient medium and 25% complete medium in the presence of CD. After 24 h labelling, the supernatant was collected and centrifuged at 2500 r.p.m. in a desktop centrifuge to spin down cell debris. Virus was then pelleted at 120000 g for 1 h and resuspended at 4 °C overnight in a small volume of PBS. Viruses were further purified through a 3060% sucrose cushion at 240000 g for 1 h. Purified virions were collected from the interface of the sucrose cushion, diluted in PBS and pelleted at 120000 g for 1 h. Pelleted virions were lysed in RIPA buffer (150 mM NaCl, 50 mM TrisHCl, pH 7·5, 1% Triton X-100, 1% SDS, 1 mM EDTA) and the viral protein profile was analysed by SDSPAGE under reducing conditions followed by autoradiography.
Surface immunofluorescence.
Vero 76, Vero C1008 or MDCK cells grown on 12 mm glass coverslips were infected with HPIV2, HPIV3 or SeV at an m.o.i. of 1 p.f.u. per cell. At 24 or 48 h p.i., cells were washed three times with ice-cold PBS. Guinea pig anti-HPIV2 antiserum at a dilution of 1:500, guinea pig anti-HPIV3 antiserum at a dilution of 1:50 or rabbit anti-SeV antiserum at a dilution of 1:50 was added onto cell monolayers and cells were then incubated at 4 °C for 30 min. Cells were washed three times with ice-cold PBS and then FITC-conjugated goat anti-guinea pig IgG antibody or rhodamine-conjugated goat anti-rabbit IgG antibody (Southern Biotechnology Associates), at a dilution of 1:100, was added and the monolayers were incubated at 4 °C for 30 min. Cells were then washed and fixed with 2% paraformaldehyde in PBS. Cell surface fluorescence was examined by fluorescence microscopy with a Nikon Optiphot microscope.
Electron microscopy.
HPIV2- or HPIV3-infected Vero C1008 cells were examined by thin-section electron microscopy at 48 h p.i. Cells were fixed with buffered 1% glutaraldehyde for 30 min, post-fixed for 1 h with 1% osmium tetroxide, dehydrated with a graded ethanol series and embedded for electron microscopy. Thin sections were prepared on a Reichert ultramicrotome, mounted on 300-mesh copper grids, stained with uranyl acetate and lead citrate and examined with a Philips CM10 electron microscope (EM). For negative staining, virions in culture medium were allowed to adhere to carbon/formvar grids and stained with 1% ammonium phosphotungstate, pH 7·4. Specimens were viewed with a Philips CM10 EM.
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Results |
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The formation of long filamentous HPIV2 virions was found to be host cell dependent. As can be seen in Fig. 1(a), long filamentous HPIV2 particles were observed predominantly in polarized epithelial cells, and the lengths of particles were up to 15 µm. In contrast, only short, elongated virus particles were seen in Vero 76 cells (Fig. 1c
). This result indicates that the polarized cells preferentially support the production of long, filamentous HPIV2 virions.
Absence of filament formation in HPIV3 and SeV virions
In order to determine whether filament formation is a general property of other paramyxoviruses, Vero 76, Vero C1008 and MDCK cells were infected with HPIV3 or SeV and were examined by indirect surface immunofluorescence. Fig. 2(a) shows that HPIV3-infected Vero C1008 cells exhibited only punctate surface fluorescence, and a similar fluorescence pattern was also seen in HPIV3-infected MDCK and Vero 76 cells. When EM sections of HPIV3-infected cells were examined, spherical viral particles 0·2 to 0·5 µm in diameter were observed (Fig. 2b
). Fig. 2(c)
shows that only punctate surface fluorescence could be seen in SeV-infected Vero C1008 cells as well. No budding of filamentous particles was observed in SeV-infected Vero 76 or MDCK cells (not shown). Therefore, the formation of filamentous virions is a property of at least two PIVs in the genus Rubulavirus, but was not observed in two members of the genus Respirovirus.
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CD and jasplakinolide affect virus morphology but not virus release
To investigate whether the inhibition of HPIV2 filament formation by CD or jasplakinolide reflected an inhibition of virion assembly or budding at the cell surface, released virus HA and infectivity titres were determined. HPIV2-infected culture media were collected at 24 and 48 h p.i. in the presence or absence of CD treatment. The HA titres of the released viruses were not affected by the CD treatment at 24 h p.i., but were reduced at 48 h p.i. (Table 1). Virus infectivity titres in the culture in the presence of CD were only about twofold lower at 24 h p.i. compared with the control culture infected in the absence of CD. HPIV2 virus production in the untreated culture increased further by about fivefold from 24 to 48 h p.i., whereas there was no corresponding increase in virus yield in the CD-treated culture. The absence of a further increase in titre after 24 h may result from an effect of CD on the viability of the cells.
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To determine whether there were any differences in viral protein profiles in the presence of CD treatment, HPIV2 virus produced in the presence or absence of CD was radiolabelled with [35S]methionine/[35S]cysteine, purified and analysed by SDSPAGE. By comparing control HPIV2 protein profiles with those of HPIV2 produced in the presence of CD, the major viral proteins present in both preparations were found to be similar in profile and relative amounts (data not shown). This result indicates that the change in virus morphology observed as a result of altering the actin microfilament network does not involve changes in parainfluenza virion protein incorporation or virus infectivity.
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Discussion |
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Although influenza virus morphology is known to be determined largely genetically (Choppin, 1963 ; Kilbourne, 1963
; Mitnaul et al., 1996
; Roberts et al., 1998
; Smirnov et al., 1991
), recent studies (Roberts & Compans, 1998
) have provided evidence that host cell factors are also important determinants of the formation of filamentous particles. It has been reported that filamentous influenza particles exhibit a higher specific infectivity and a higher RNA content than spherical virions (Ada et al., 1958
; Smirnov et al., 1991
). One or more structural gene products of influenza virus has been linked to the formation of virus filaments (Choppin, 1963
; Kilbourne, 1963
; Mitnaul et al., 1996
; Smirnov et al., 1991
; Nishimura et al., 1990
; Roberts et al., 1998
). Our observation that HPIV2 forms filamentous particles preferentially in polarized epithelial cells is similar to recent results with influenza viruses (Roberts & Compans, 1998
), indicating that the host cell is a determinant of filamentous virus production by both of these viruses. It is interesting that the primary site of PIV infection is the mucosal surface of the upper respiratory tract. Thus, it is likely that the polarized phenotype of cells in the respiratory tract is compatible with virus filament formation. Furthermore, we found that CD, an actin microfilament-depolymerizing agent, or the macrocyclic peptide jasplakinolide, a potent inducer of actin polymerization, inhibited HPIV2 filamentous particle formation but had little effect on the HA or p.f.u. titres of released virus particles. These results indicate that the host cell type and the integrity of the cytoskeleton play important roles in PIV morphology.
Filamentous virus morphology may give some advantage to a virus in infecting cells or evading immune responses of the host. Many respiratory tract virus pathogens are filamentous, such as influenza virus and RSV. By their elongated particle size, filamentous particles may be able to infect neighbouring cells prior to release and therefore resist upward mucociliary removal of fluids from the respiratory tract (Roberts & Compans, 1998 ). A recent report demonstrated that intracellular vaccinia virions can utilize host actin microfilaments to project themselves out of the cytoplasm and into neighbouring cells (Cudmore et al., 1995
). Filamentous paramyxovirus particles similarly may be able to mediate infection of neighbouring cells. In contrast, spherical particles may be more readily incorporated into aerosols and may play a more important role in person-to-person transmission, while filamentous particles may be important in cell-to-cell transmission, as suggested for influenza virus (Roberts & Compans, 1998
).
The finding of actin in many enveloped viruses (Damsky et al., 1977 ; Naito & Matsumoto, 1978
; Wang et al., 1976
) triggered studies of the role of the cellular cytoskeleton in virus assembly. Although the cytoskeleton is thought to be involved at some stage in the morphogenesis of enveloped viruses, its exact role has not been defined. It has been reported that treatment of infected cells with CD has little direct effect upon the assembly and release of vesicular stomatitis virus (Genty & Bussereau, 1980
) or influenza virus (Griffin & Compans, 1979
; Griffin et al., 1983
; Roberts & Compans, 1998
). However, treatment of influenza virus-infected cells with CD was found to abolish filamentous particle formation (Roberts & Compans, 1998
). Also, release of measles virus particles was inhibited in CD-treated infected cells (Stallcup et al., 1983
). CD also caused a blockage in the final release of enveloped vaccinia virus from the cell surface (Payne & Kristensson, 1982
) and inhibited the release of murine leukaemia virus (Mousa et al., 1978
) and New World hantaviruses (Ravkov et al., 1998
). Other studies have reported that CD can stimulate release of some viruses, such as NDV and rotavirus (Bass et al., 1995
; Bedows et al., 1983
; Morrison & McGinnes, 1985
). The role of actin as a determinant of virus assembly may also depend on the virus and cell type. There was no obvious structural rearrangement of F-actin in influenza virus-infected cells (Roberts & Compans, 1998
), which contrasts with results in human immunodeficiency virus (HIV)- and vaccinia virus-infected cells. In HIV-infected epithelial cells, F-actin was redistributed into pseudopods and HIV was preferentially released from the pseudopod (Pearce-Pratt et al., 1994
). Actin filaments in vaccinia virus-infected cells formed actin tails and projections and the virus particles were propelled to the neighbouring cells on their tips (Cudmore et al., 1995
). We did not observe disrupted actin microfilament arrays in HPIV2-infected cells, nor did we observe co-localization of actin with the filamentous virus particles on cell surfaces. This may indicate why there was no marked inhibition of virus release by CD treatment.
Jasplakinolide is a membrane-permeable, F-actin-stabilizing drug. It competes with phalloidin for binding and enhances actin polymerization by inhibiting the depolymerization of actin filaments, leading to a change in actin filament dynamics (Bubb et al., 1994 ; Senderowicz et al., 1995
; Lee et al., 1998
). Cells that are treated with jasplakinolide form F-actin aggregates because of their inability to depolymerize the stabilized actin filaments at a normal rate. It was also reported that rearrangement of the actin cytoskeleton of plant cells resulted from treatment with jasplakinolide (Sawitzky et al., 1999
). Therefore, treatment with jasplakinolide will alter the normal cellular actin cytoskeleton functions. However, no effects of jasplakinolide on virus morphology or virus assembly have been reported. Here, we found that jasplakinolide can also abolish the filamentous morphology of HPIV2. The patchy fluorescence pattern of viral antigens in the CD- or jasplakinolide-treated, HPIV2-infected cells may also result from disrupted or aggregated cytoskeletal structures. The normal cytoskeleton plays an important role in establishment and maintenance of epithelial cell polarity, and polarized cells preferentially support filamentous virus particle formation. We therefore suggest that CD and jasplakinolide abolish filamentous particle formation by altering the polarized phenotype of the epithelial host cells and the normal function of the cellular cytoskeleton.
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Acknowledgments |
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References |
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Received 3 November 1999;
accepted 7 January 2000.