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The interaction of mammalian reoviruses with the cytoskeleton of monkey kidney CV-1 cells
VIROLOGY
120,399-411 (1982)
The Interaction
of Mammalian Reoviruses with the Cytoskeleton Monkey Kidney CV-1 Cells
of
ARLENE H. SHARPE,* LAN BO CHEN,p$ AND BERNARD N. FIELDS*@ *Lkpartment of Mkmbiolcgg and Molecular Genetics,and ~Department of Pathology, $Sidnq Farber of Medicine Cancer Institute, Harvard Medical Scrod, Boston, Massachusetts 02115;and Wepwtmmt (Irlfectious Diseases), Brigham and Worna’s Hospital, Boston, Massachusetts 02115
Received January 13, 1982;accepted April 19, 1982 We have examined the effect of reovirus infection on the CV-1 cytoskeleton. Reovirus infection produces a major disruption of vimentin filaments without producing a discernable disorganization of microtubules or microfilament bundles in CV-1 cells. In addition to disrupting the organization of vimentin filaments, reovirus infection appears to cause a reorganization of vimentin filaments. Viral inclusions contain vimentin filamentous structures. Viral infection also alters the cytoplasmic distribution of mitochondria. consistent with the nroposed role of vimentin filaments in determining the distribution of mitochondria. - INTRODUCTION
During lytic infection, cytocidal viruses alter cell metabolism and cell structure, ultimately leading to the production of progeny viruses and cell death. The manner in which viruses interact with cells during viral replication and assembly is not well understood. It is likely that such interactions will play a central role in determining the specific ways by which viruses mediate cellular injury. To explore the mechanisms of virus-cell interactions, we have chosen to examine how reovirus, a nonenveloped, lytic animal virus with a segmented double-stranded RNA genome interacts with the CV-1 (African green monkey kidney) cell cytoskeleton. Microscopic and biochemical studies indicate that changes in host cell morphology accompany reovirus replication and assembly (Rhim et al, 1962; Spendlove et aL, 1963). Reoviruses enter cells within phagocytic vacuoles and remain within these vacuoles until the vacuoles fuse with primary or secondary lysosomes (Dales et al, 1965). Within lysosomes, reovirions are uncoated and the virion transcriptase is activated. Nascent virions are synthesized i To whom reprint requests should be addressed.
subsequently within discrete regions of the cytoplasm known as viral factories. The intracytoplasmic routes by which reoviruses move to these sites of replication are not known. In addition, the mechanism of formation of viral factories is not understood. Viral factories first appear as phase dense granular material scattered throughout the cytoplasm. As infection progresses, these discrete granules coalesce and move toward the nucleus eventually forming phase dense perinuclear inclusions. The cellular protein and structural elements which are present within viral factories remain to be clarified. Although the factory is a discrete cytoplasmic entity, it is not enclosed by a membrane. Electron microscopic studies have revealed that ribosomes are not present within factories. Viral proteins thus must be synthesized on polysomes outside the factory and transported back into the factory prior to assembly onto virus. Conversely, for viral mRNAs to be translated, the mRNAs must be synthesized within the factory and transported to adjacent polysomes within the cytoplasm. Electron microscopic studies have also
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indicated that several types of filaments are located within viral factories (Dales, 1963, 1965). Viral particles occasionally are aligned on parallel arrays of microtubules which are thought to be covered with viral protein, possibly the viral hemagglutinin (Babiss et aL, 1979). Between microtubules is a complex filament network consisting of masses of densely twisted or kinky filaments with a diameter of 50-80 A. The identity of these filaments is not known but their size is similar to that of intermediate filaments. These filaments are observed to be in intimate contact with virus particles and the viral proteins coating the microtubules. Such filaments occur regularly in wavy bundles in the cytoplasm of uninfected mouse L cells. Dales has suggested that these filaments change from the wavy to the kinky type in areas where virus is being formed (Dales et aL, 1965) and that such filaments may provide a surface for the attachment of viral mRNAs or for viral morphogenesis. The availability of specific probes to components of the cytoskeleton permitted us to examine the interactions of reoviruses with the various filament systems. In the studies reported in this paper we used immunocytochemical techniques to examine the effects of reovirus replication on the CV-1 cytoskeleton and to further define the composition of viral factories.
as described by Fields and Joklik (1969). All viruses were plaque purified and passaged in L cells twice. Second-passage virus was used for all infections. Antisera to c@oskeletal$lament proteins and reowirua Antisera to cytoskeletal filament proteins have been described previously (Sharpe et al, 1980). To detect microtubules we used rabbit antiserum prepared against tubulin, provided by Dr. Frank Solomon, Massachusetts Institute of Technology, Cambridge, Massachusetts. To detect vimentin filaments we used rabbit antisera to gerbil vimentin, prepared according to the procedure of Hynes and Destree (1978). To detect microfilaments we used mouse monoclonal antibody to actin, provided by Dr. James Lin, Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Rabbit antisera to hamster vimentin was provided by Dr. Richard 0. Hynes, Massachusetts Institute of Technology. Antiserum to reovirus type 3 Dearing was prepared by hyperimmunization of rabbits to CV-1 cells infected with reovirus type 3 Dearing. Prior to use, the antiserum was adsorbed extensively to CV1 cells. Fluorescein-conjugated goat antirabbit IgG and fluorescein-conjugated rabbit anti-mouse IgG were obtained from Meloy Laboratories, Springfield, Virginia. Indirect immunofluorescent microscopg. CV-1 cells, seeded at low density (2 X 105/ dish) in 60 X 15-mm culture dishes containing 12-mm round glass coverslips MATERIALS AND METHODS (Rochester Scientific) were infected with a m.o.i. of lo-20 PFU/cell of reovirus types Cells and growth media. CV-1 (African green monkey kidney) cells were grown as 1, 2, or 3 for the indicated times. Prior to staining cells with antibody to tubulin or monolayers in roller bottles in Richter’s improved minimal essential medium con- vimentin, the cells were fixed using the taining zinc, insulin, and 10 mM Hepes procedure of Osborn and Weber (1977). Cells were washed for 30 set at room tembuffer (N-2-hydroxyethylpiperazine-N’-2ethanesulfonic acid) (IMEMZO: Irvine Sci- perature with stabilization buffer (0.1 M piperazine-Nfl’-bis[2-ethanesulfonic acid] entific, Santa Ana, Calif.) supplemented with 10% fetal calf serum (Sterile Sys- sodium salt adjusted to pH 6.9 with KOH, 1 mM ethylene glycol bis(&aminoethyl tems, Inc., Logan, Utah) at 37”. ether)-N-N’-tetraacetic acid, 0.5 mM GTP, V&us strains. Reovirus type 1, strain Lang, and type 2, strain Jones, were ob- and 4% polyethylene glycol 6000), and then incubated for 1 min at room tempertained from the National Institute of Allergy and Infectious Diseases (type 1, ature in the same buffer containing 0.5% strain Lang, catalog No. V701-001-010). Triton X-100 (Sigma). After this treatReovirus type 3, strain Dearing, was used ment, cells were washed twice for 30 set
REOVIRUS-CYTOSKELETON
INTERACTIONS
FIG. 1. Viral antigens in CV-1 cells at 48 hr after infection with reovirus type 3. Cells were stained with rabbit antireovirus serum and fluorescein-conjugated goat anti-rabbit serum as described under Materials and Methods. Cytoplasmic inclusions are represented by the numerous globular white areas within the cytoplasm. Note the gradation of size of inclusions, from small to large, as the inclusions approach the nucleus. (Scale: bar = 20 pm.)
with stabilization buffer, fixed in chilled methanol (-20’) for 5 min, and then transferred to a humidified chamber. Ten microliters of an appropriate dilution of antiserum were applied to each coverslip. The coverslips were incubated at room temperature for 1 hr, rinsed in phosphatebuffered saline (PBS), and returned to the humidified chamber. Ten microliters of an appropriate dilution of fluorescein-conju-
gated goat anti-rabbit IgG were appliied to each coverslip. The coverslips were incubated at room temperature for 40 nlin, rinsed in PBS and water, and mounted I on glass slides in Gelvatol (Monsanto, St. Louis, MO.). Prior to staining with alntibody to actin, the cells were fixed for 10 min in 3.5% formaldehyde in PBS, rin sed in PBS, and incubated for 5 min in acct.one which had been prechilled to -20”. Ai %er
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FIG. 2. Organieation of microtubules in mock-infected and reovirus type 3-infected CV-1 cells at 48 hr after infection. (A) Fluorescence micrograph showing microtubule organization in a mockinfected CV-1 cell. (B) Phase contrast micrograph of a type 3-infected CV-1 cell showing phase dense inclusion bodies. (C) Fluorescence micrograph of the same infected cell as in (B) showing microtubule organization in a reovirus-infected CV-1 cell. Cells were subjected to the indirect immunofluorescent staining technique described under Materials and Methods using antitubulin antibody. Bar represents 25 pm.
rinsing in PBS, the fixed cells were incubated with antisera as described above. All cells were examined with a Zeiss photomicroscope III equipped with epifluorescence and a Planapochromat objective lens
(63X). Photomicrographs were made using Kodak Tri-X (ASA 400) at exposure index 1606. Loealixation of mitochondria Mitochondria were visualized in living cells using
REOVIRUS-CYTOSKELETON*INTERACTIONS
FIG. 2.--Continued
the procedure of Johnson, Walsh, and Chen (1980). CV-1 cells were grown on 12mm round glass coverslips and infected with reovirus as described above. At 12, 24, and 48 hr postinfection, the coverslips were removed from the culture dishes, transferred to IMEMZO medium containing 10 pg/ml of the laser dye rhodamine 123 (Eastman), and incubated.at 37” in a 5% COz incubator for 30 min. Following extensive rinsing, the coverslips were mounted on a microscope slide with a silicon rubber chamber suitable for live cell examination as described by Johnson et ad (1980). They were examined by epifluorescent illumination at either 546 nm (rhodamine excitation) or 485 nm (fluorescein excitation). Electron microscopy. Fixation and procedures for transmission electron microscope were according to the methods of Goldman (1971) and Goldman et cd (1975). RESULTS
Cytoskeletcm of CV-1 Cells Monkey CV-1 cells were chosen for these studies because of their capacity to support the efficient growth of reovirus. In addition, since they are well spread and
flattened cells in monolayers, they are excellent cells to use for visualizing the cell cytoskeleton. The cytoskeleton of CV-1 cells has been described (Sharpe et cd, 1980). Briefly, CV-1 cells have an extensive network of microtubules emanating from the cell nucleus (Fig. 2A) and have typical bundles of microfilaments in the cytoplasm (Fig. 3A). The vimentin filament system emanates from multiple sites surrounding the nucleus (Fig. 4A). Distribution Cells
of Reovirus Antigen in CV-1
The maturation of reovirus in the cytoplasm of CV-1 cells as studied by immunocytochemical staining with antisera to reovirus was similar to that described previously (Rhim et al, 1962; Spendlove et uL, 1963; Fields et al, 1971). Viral antigens were detected within the cytoplasm as well as within viral factories. Viral inclusions first appeared as small immunofluorescent spots typically located in the cell periphery. As the viral growth cycle progressed, viral factories increased in size and appeared to fuse to form large, irregularly shaped inclusions. The inclusions migrated from the cell periphery to a perinuclear location (Fig. 1).
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FIG. 3. Organization of microfilaments in mock-infected and reovirus type a-infected CV-1 in a at 48 hr after infection. (A) Fluorescence micrograph showing microfilament organization
REOVIRUS-CYTOSKELETON
Eflect of Reovirus Iqfection on Microtuhuh The organization of microtubules was examined in reovirus-infected cells at 12, 24, and 43 hr postinfection using antitubulin antibody. Reovirus infection did not disrupt microtubule organization even at late times postinfection (Figs. 2B and C). Microtubules were seen to span the regions of cytoplasm containing viral factories without interruption or distortion. Similar results were observed with all three serotypes. Eflect of Redrw aments
Replication on Microjl-
The distribution of microfilament bundles in reovirus-infected cells at 12,24, and 48 hr postinfection was examined using a monoclonal antibody to actin. Microfilament organization was not disturbed by reovirus infection. In addition, viral inclusions did not appear to contain actin since the monoclonal antibody to actin did not stain viral inclusions as distinct structures (Figs. 3B and C). Thus, reovirus infection does not appear to alter actin filament organization within CV-1 cells. Efect of Reovirus Replication on Intermediate Filaments The distribution of vimentin filaments was examined in cells infected with all three serotypes using antiserum to vimentin. By 12 hr postinfection, before intracytoplasmic inclusions were visible, the perinuclear organization sites of vimentin filaments disappeared. As infection progressed, there was further disruption of these filaments such that wavy filaments with no apparent organization were observed in the cytoplasm (Figs. 4B, C, and E). Similar results were observed with all three serotypes. The contrast between intermediate filament organization in cells
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containing viral inclusions and cells not containing them is illustrated clearly in Figs. 4B and C. Inclusion-bearing CV-1 cells that have disrupted intermediate filament organization are adjacent to CV-1 cells without inclusions that have the typical organization of intermediate filaments. Viral inclusions were stained to varying degrees with vimentin antibody (Figs. 4B and D). When viral inclusions were examined at a high magnification (630-fold), filamentous structures which stained with vimentin antibody were observed within the inclusions (Fig. 5). Electron microscopic examination of viral inclusions at a similar time postinfection reveals the presence of kinky filaments, which are 100 A in diameter (Fig. 6). Thus, reovirus infection disrupts vimentin filament organization within the cytoplasm of CV-1 cells and results in the presence of vimentin within viral inclusions. Eflect of Reovirms Replication on Mitt+ chondrial Llistribution CV-1 cells have a characteristic discontinuous filamentous distribution of mitochondria within the cytoplasm (Fig. 7A). Reovirus infection leads to a disorganization of mitochondria distribution in living cells as viewed with the fluorescent probe, rhodamine 123 (Figs. ‘7B and C). Mitochondria aggregated around the nucleus and occasionally appeared isolated at the cell periphery. Mitochondria were excluded from the viral factories. These findings are consistent with the observation that an intact intermediate filament system is essential for normal distribution of mitochondria in CV-1 cells (Chen et aZ., 1981). DISCUSSION
These studies show that reovirus infection produces a major disruption of vi-
mock-infected CV-1 cell. (B) Phase contrast micrograph of a type 3-infected CV-1 cell showing phase dense inclusion bodies. (C) Fluorescence micrograph of the same infected cell as in (B) showing microfilament organization in a reovirus-infected CV-1 cell. Cells were subjected to indirect immunofluorescence microscopy using a monoclonal antiactin antibody. Bar represents 25 pm.
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FIG. 4. Organization of intermediate filaments in mock-infected and reovirua type a-infected CV1 cells at 48 hr after infection. (A) Fluorescence micrograph showing intermediate filament organization in mock-infected CV-1 cells. (B and D) Phase contrast micrographs of type 3-infected CV-1 cells showing phase dense inclusion bodies. (C and E) Fluorescence micrographs of the same infected cells as in B and D, respectively, showing intermediate filament organization in reovirusinfected CV-1 cells. Note that intermediate filament organization is affected by reovirus infection. Cells were subjected to indirect immunofluorescence microscopy using antibody against vimentin. (A, B, and C, bar = 40 pm; D and E. bar = 25 pm.)
mentin filament organization without producing a discernable effect upon the organization of microtubules and microfilament bundles in CV-1 cells during the
period of observation. The distribution of mitochondria is also altered by viral infection. The disruption of vimentin filament or-
REOVIRUS-CYTOSKELETON
INTERACTIONS
FIG. 5. Fluorescence micrograph of a type 3-infected CV-1 cell at 48 hr after infection demonstrating vimentin filament organization within viral inclusions. Cell was subjected to indirect immunofluorescence microscopy using antibody against vimentin. Bar represents 20 pm.
ganization by reovirus is especially interesting in view of the finding that agents which disrupt protein synthesis such as diphtheria toxin, P. aeruginosa exotoxin A, and cycloheximide, specifically disrupt the organization of vimentin filaments without affecting microtubule or microfilament organization (Sharpe et al, 1980). The mammalian reoviruses also disrupt host cell macromolecular synthesis (Sharpe and Fields, 1981; Sharpe and Fields, 1982). It is possible that the disruption of intermediate filaments by reo-
virus is related to the capacity of reovirus to inhibit host cell protein synthesis. However, in addition to disrupting the organization of intermediate filaments, reovirus infection appears to cause a reorganization of vimentin filaments. Viral inclusions contain filamentous structures which stain with antibody to vimentin. These vimentin filamentous structures appear to be the complex filamentous network previously visualized in Dales’ electron microscopic studies of viral factories (Dales et aL, 1965). The presence of vi-
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FIG. 6. Electron micrograph of portion of a type 3-infected CV-1 cell containing a viral inclusion. Note kinky filaments present within the inclusion. These are exactly 100 A in diameter (magnification X7000).
REOVIRUS-CYTOSKELETON
mentin filaments in the viral factories suggests that these filaments may be participating in the formation of unique viral structures that are distinct from vimentin filament organization in the cytoplasm of the noninfected cells. It is possible that reovirus is using the vimentin filament system in order to create a discrete cytoplasmic entity, the viral factory, which functions as the site of viral replication and assembly. It has been proposed that vimentin filaments function in the coordination of the organization of cytoplasmic contents (Lazarides, 1980). The fact that vimentin filaments are found within viral factories suggests that they may be playing a critical role in the organization of the viral replicative process. Unlike the marked disruption of intermediate filaments, reovirus infection did not disrupt microtubule structure. Electron microscopic studies have shown that reoviruses are aligned on parallel arrays of microtubules within viral factories (Dales et CAL,1965) and have the capacity to bind to microtubules in vitro (Babiss et al, 1979). These results are consistent with our observation that microtubules course through regions of the cytoplasm containing viral factories without disrupting or distorting them. Since colchicine treatment of reovirusinfected cells does not reduce viral yield, it is uncertain whether microtubules play a role in viral growth. Disruption of microtubules by colchicine treatment, however, alters the morphology of viral inclusions. Following the treatment of virally infected cells with colchicine only small inclusions located at the cell periphery are observed and the large perinuclear inclusions usually observed in cells infected with reovirus are absent (Spendlove et al., 1964). It is thus possible that microtubules play a role in the coalescence of viral inclusions and in their migration toward the nucleus. The concomitant disruption of vimentin filament organization and mitochondrial distribution after reovirus infection is consistent with a recent report by Chen et al. (1981) that suggests a role for vimentin filaments in determining the dis-
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FIG. 7. Distribution of mitochondria in mock-infected and reovirus type 3-infected CV-1 cells at 48 hr after infection. (A) Fluorescence micrograph of a live CV-1 cell showing rhodamine W&stained mitochondria. Dotted lines indicate cell periphery. (B) Phase contrast micrograph of a live type I-infected CV-1 cell showing phase dense inclusion bodies. (C) Fluorescence micrograph of same infected cells as in (B) showing the distribution of rhodamine-stained mitochondria. Note that the distribution of mitochondria is affected by reovirus infection. Bar represents 25 pm.
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tribution of mitochondria in cultured cells. Although infection of cells with reovirus alters the distribution of mitochondria, whether reovirus infection also alters mitochondrial functions remains to be determined. The accumulation of rhodamine 123 by mitochondria reflects the transmembrane potential (Johnson et aL, 1981). Because the accumulation of rhodamine 123 in reovirus-infected cells is similar to that of uninfected cells, it is likely that the respiratory activity of infected and uninfected cells is similar. A role for the cytoskeleton in the growth and assembly of other viruses has also been proposed. For example, immunofluorescence and electron microscopic studies of pox virus interactions with the cytoskeleton of infected chick embryo fibroblasts have revealed an intimate association between mature virus particles and the cytoskeleton that involves actin-containing structures. Vaccinia virus infection appears to induce the formation of specialized microvillus-type structures, which contain a core of microfilaments and a virus particle at their tip. These microvilli may be involved in viral budding and release from infected cells (Hiller et aL, 1979). An immunofluorescence microscopic study of the fusion of cells infected with the parainfluenza virus SV-5 revealed that microtubules and intermediate filaments form large bundles during cell fusion. This observation has suggested that both types of filaments may provide a framework which directs nuclear migration during cell fusion (Wang et aL, 1979). The results presented here suggest that intermediate filaments may play an important role in the replication and assembly of reovirus. The biochemical and morphological properties of intermediate filaments suggest that these filaments are involved in the organization of components within the cytoplasmic space (Lazarides, 1980). With such a function, intermediate filaments may be expected to play a crucial role in the regulation of cell shape to be influenced by the cell’s metabolic state and response to injury. Possibly, intermediate filaments are one of the major cellular targets involved in producing the cytopathic effects resulting from
reovirus infection (rounding of cells, inclusion formation, etc.). It remains to be determined if intermediate filament disorganization (and/or reorganization) is a primary and central step in viral infection, leading ultimately to cell death. ACKNOWLEDGMENTS
We thank Lincoln V. Johnson for help with immunocytochemical staining techniques and for stimulating discussions and Karen Byers for excellent technical assistance. During the course of these studies, A.H.S. was a National Institutes of Health Predoctoral Trainee (5-T32 GM0 7196). This investigation was supported by Grants AI 13178 from the National Institutes of Health (to B.N.F.) and CA 22619 from the National Cancer Institute (to L.B.C.). L.B.C. is the recipient of an American Cancer Society Faculty Research Award. REFERENCES
BABISS, L. E., LUFTIG, R. B., WEATHERBEE, J. A., WIEHING, R. R., RAY, U. R., and FIELDS, B. N. (1979). Reovirus serotypes 1 and 3 differ in their in vitro association with microtubules. J. ViroL 30, 868-874. CHEN, L. B., SUMMERHAYES,I. C., JOHNSON,L. V., WALSH, M. L., BERNAL, S. D., and LAMPIDIS, T. J. (1981). Probing mitochondria in living cells with Rhodamine 123. Cold Sprins Harbor Synp. Quati BioL, in press. DALES, S. (1963). Association between the spindle apparatus and reovirus. Proc Nat. Ad Sci USA 50,268-275. DALES, S., GOMATOS,P. J., and Hsu, K. C. (1965). The uptake and development of reovirus in strain L cells followed with labelled viral ribonucleic acid and ferritin-antibody conjugates. Virdogy 26,193211.
FIELDS,B. N., and JOKLIK, W. K. (1969). Isolation and preliminary genetic and biochemical characterization of temperature-sensitive mutants of reovirus. Virdogy
37.335442.
FIELDS, B. N., RAINE, C. S., and BAUM, S. G. (1971). Temperature-sensitive mutants of reovirus type 3: Defects in viral maturation as studied by immunofluorescence and electron microscopy. Virology 43,569-578. GOLDMAN,R. D. (1971). The role of three cytoplasmic fibers in BHK-21 cell motility. I. Microtubules and the effects of colchicine. J. Cell Bid 51, 752-762. GOLDMAN,R. D., LAZARIDES, R., POLLACK, R., and WEBER,K. (1975). The distribution of actin in nonmuscle cells: The use of actin antibody in the localization of actin within the microfilament bundles of mouse 3T3 cells. Exp. Cell Rex 90.333-344.
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HILLER, G., WEBER, K., SCHNEIDER,L., PARAJSZ,C., and JUNGWIRTH,C. (1979). Interaction of assembled progeny pox viruses with the cellular cytoskeleton. %ob 98, 142-153. HYNES, R. G., and DESTREE,A. T. (1978). 10 nm filaments in normal and transformed cells. Cell 13, 151-163. JOHNSON,L. V., WALSH, M., and CHEN, L. B. (1980). Localization of mitochondria in living cells with Rhodamine 123. Proc. Nat. Acad Sci USA 77,990994. LAZARIDES,E. (1980). Intermediate filaments as mechanical integrators of cellular space. Nature (London) 283,249-256. OSBORN,M., and WEBER, K. (1977). The display of microtubules in transformed cells. CelE12,561-571. RHIM, J. S., JORDAN,L. E., and MAYOR, H. D. (1962). Cytochemical, fluorescent-antibody and electronmicroscopic studies on the growth of reovirus (ECHO 10) in tissue culture. Virology 17,342-355. SHARPE, A. H., and FIELDS, B. N. (1981). Reovirus
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inhibition of cellular DNA synthesis: Role of the Sl gene. J. ViroL 38,389-392. SHARPE, A. H., and FIELDS, B. N. (1982). Reovirus inhibition of cellular RNA and protein synthesis: Role of the S4 gene. Submitted for publication. SHARPE, A. H., CHEN, L. B., MURPHY, J. R., and FIELDS,B. N. (1980). Specific disruption of vimentin filament organization in monkey kidney CV-1 cells by diphtheria toxin, exotoxin A, and cycloheximide. Proc. Nat. Ad Sci USA 77.7267-7291. SPENDLOVE,R. S., LENNETTE,E. H., CHIN, J. N., and KNIGHT, C. 0. (1964). Effect of antimitotic agents on intracellular reovirus antigen. Cancer Res. 24, 1826-1833.
SPENDLOVE,R. S., LENNE’ITE, E. H., KNIGHT, C. O., and CHIN, J. H. (1963). Development of viral antigen and infectious virus in HeLa cells infected with reovirus. J. ImmunoL SO,548-553. WANG, E., CROSS,R. K., and CHOPPIN,P. W. (1979). Involvement of microtubules and 10 nm filaments in the movement and positioning of nuclei in syncytia. J. CeU Bid 83,320-337.
120,399-411 (1982)
The Interaction
of Mammalian Reoviruses with the Cytoskeleton Monkey Kidney CV-1 Cells
of
ARLENE H. SHARPE,* LAN BO CHEN,p$ AND BERNARD N. FIELDS*@ *Lkpartment of Mkmbiolcgg and Molecular Genetics,and ~Department of Pathology, $Sidnq Farber of Medicine Cancer Institute, Harvard Medical Scrod, Boston, Massachusetts 02115;and Wepwtmmt (Irlfectious Diseases), Brigham and Worna’s Hospital, Boston, Massachusetts 02115
Received January 13, 1982;accepted April 19, 1982 We have examined the effect of reovirus infection on the CV-1 cytoskeleton. Reovirus infection produces a major disruption of vimentin filaments without producing a discernable disorganization of microtubules or microfilament bundles in CV-1 cells. In addition to disrupting the organization of vimentin filaments, reovirus infection appears to cause a reorganization of vimentin filaments. Viral inclusions contain vimentin filamentous structures. Viral infection also alters the cytoplasmic distribution of mitochondria. consistent with the nroposed role of vimentin filaments in determining the distribution of mitochondria. - INTRODUCTION
During lytic infection, cytocidal viruses alter cell metabolism and cell structure, ultimately leading to the production of progeny viruses and cell death. The manner in which viruses interact with cells during viral replication and assembly is not well understood. It is likely that such interactions will play a central role in determining the specific ways by which viruses mediate cellular injury. To explore the mechanisms of virus-cell interactions, we have chosen to examine how reovirus, a nonenveloped, lytic animal virus with a segmented double-stranded RNA genome interacts with the CV-1 (African green monkey kidney) cell cytoskeleton. Microscopic and biochemical studies indicate that changes in host cell morphology accompany reovirus replication and assembly (Rhim et al, 1962; Spendlove et aL, 1963). Reoviruses enter cells within phagocytic vacuoles and remain within these vacuoles until the vacuoles fuse with primary or secondary lysosomes (Dales et al, 1965). Within lysosomes, reovirions are uncoated and the virion transcriptase is activated. Nascent virions are synthesized i To whom reprint requests should be addressed.
subsequently within discrete regions of the cytoplasm known as viral factories. The intracytoplasmic routes by which reoviruses move to these sites of replication are not known. In addition, the mechanism of formation of viral factories is not understood. Viral factories first appear as phase dense granular material scattered throughout the cytoplasm. As infection progresses, these discrete granules coalesce and move toward the nucleus eventually forming phase dense perinuclear inclusions. The cellular protein and structural elements which are present within viral factories remain to be clarified. Although the factory is a discrete cytoplasmic entity, it is not enclosed by a membrane. Electron microscopic studies have revealed that ribosomes are not present within factories. Viral proteins thus must be synthesized on polysomes outside the factory and transported back into the factory prior to assembly onto virus. Conversely, for viral mRNAs to be translated, the mRNAs must be synthesized within the factory and transported to adjacent polysomes within the cytoplasm. Electron microscopic studies have also
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indicated that several types of filaments are located within viral factories (Dales, 1963, 1965). Viral particles occasionally are aligned on parallel arrays of microtubules which are thought to be covered with viral protein, possibly the viral hemagglutinin (Babiss et aL, 1979). Between microtubules is a complex filament network consisting of masses of densely twisted or kinky filaments with a diameter of 50-80 A. The identity of these filaments is not known but their size is similar to that of intermediate filaments. These filaments are observed to be in intimate contact with virus particles and the viral proteins coating the microtubules. Such filaments occur regularly in wavy bundles in the cytoplasm of uninfected mouse L cells. Dales has suggested that these filaments change from the wavy to the kinky type in areas where virus is being formed (Dales et aL, 1965) and that such filaments may provide a surface for the attachment of viral mRNAs or for viral morphogenesis. The availability of specific probes to components of the cytoskeleton permitted us to examine the interactions of reoviruses with the various filament systems. In the studies reported in this paper we used immunocytochemical techniques to examine the effects of reovirus replication on the CV-1 cytoskeleton and to further define the composition of viral factories.
as described by Fields and Joklik (1969). All viruses were plaque purified and passaged in L cells twice. Second-passage virus was used for all infections. Antisera to c@oskeletal$lament proteins and reowirua Antisera to cytoskeletal filament proteins have been described previously (Sharpe et al, 1980). To detect microtubules we used rabbit antiserum prepared against tubulin, provided by Dr. Frank Solomon, Massachusetts Institute of Technology, Cambridge, Massachusetts. To detect vimentin filaments we used rabbit antisera to gerbil vimentin, prepared according to the procedure of Hynes and Destree (1978). To detect microfilaments we used mouse monoclonal antibody to actin, provided by Dr. James Lin, Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. Rabbit antisera to hamster vimentin was provided by Dr. Richard 0. Hynes, Massachusetts Institute of Technology. Antiserum to reovirus type 3 Dearing was prepared by hyperimmunization of rabbits to CV-1 cells infected with reovirus type 3 Dearing. Prior to use, the antiserum was adsorbed extensively to CV1 cells. Fluorescein-conjugated goat antirabbit IgG and fluorescein-conjugated rabbit anti-mouse IgG were obtained from Meloy Laboratories, Springfield, Virginia. Indirect immunofluorescent microscopg. CV-1 cells, seeded at low density (2 X 105/ dish) in 60 X 15-mm culture dishes containing 12-mm round glass coverslips MATERIALS AND METHODS (Rochester Scientific) were infected with a m.o.i. of lo-20 PFU/cell of reovirus types Cells and growth media. CV-1 (African green monkey kidney) cells were grown as 1, 2, or 3 for the indicated times. Prior to staining cells with antibody to tubulin or monolayers in roller bottles in Richter’s improved minimal essential medium con- vimentin, the cells were fixed using the taining zinc, insulin, and 10 mM Hepes procedure of Osborn and Weber (1977). Cells were washed for 30 set at room tembuffer (N-2-hydroxyethylpiperazine-N’-2ethanesulfonic acid) (IMEMZO: Irvine Sci- perature with stabilization buffer (0.1 M piperazine-Nfl’-bis[2-ethanesulfonic acid] entific, Santa Ana, Calif.) supplemented with 10% fetal calf serum (Sterile Sys- sodium salt adjusted to pH 6.9 with KOH, 1 mM ethylene glycol bis(&aminoethyl tems, Inc., Logan, Utah) at 37”. ether)-N-N’-tetraacetic acid, 0.5 mM GTP, V&us strains. Reovirus type 1, strain Lang, and type 2, strain Jones, were ob- and 4% polyethylene glycol 6000), and then incubated for 1 min at room tempertained from the National Institute of Allergy and Infectious Diseases (type 1, ature in the same buffer containing 0.5% strain Lang, catalog No. V701-001-010). Triton X-100 (Sigma). After this treatReovirus type 3, strain Dearing, was used ment, cells were washed twice for 30 set
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FIG. 1. Viral antigens in CV-1 cells at 48 hr after infection with reovirus type 3. Cells were stained with rabbit antireovirus serum and fluorescein-conjugated goat anti-rabbit serum as described under Materials and Methods. Cytoplasmic inclusions are represented by the numerous globular white areas within the cytoplasm. Note the gradation of size of inclusions, from small to large, as the inclusions approach the nucleus. (Scale: bar = 20 pm.)
with stabilization buffer, fixed in chilled methanol (-20’) for 5 min, and then transferred to a humidified chamber. Ten microliters of an appropriate dilution of antiserum were applied to each coverslip. The coverslips were incubated at room temperature for 1 hr, rinsed in phosphatebuffered saline (PBS), and returned to the humidified chamber. Ten microliters of an appropriate dilution of fluorescein-conju-
gated goat anti-rabbit IgG were appliied to each coverslip. The coverslips were incubated at room temperature for 40 nlin, rinsed in PBS and water, and mounted I on glass slides in Gelvatol (Monsanto, St. Louis, MO.). Prior to staining with alntibody to actin, the cells were fixed for 10 min in 3.5% formaldehyde in PBS, rin sed in PBS, and incubated for 5 min in acct.one which had been prechilled to -20”. Ai %er
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FIG. 2. Organieation of microtubules in mock-infected and reovirus type 3-infected CV-1 cells at 48 hr after infection. (A) Fluorescence micrograph showing microtubule organization in a mockinfected CV-1 cell. (B) Phase contrast micrograph of a type 3-infected CV-1 cell showing phase dense inclusion bodies. (C) Fluorescence micrograph of the same infected cell as in (B) showing microtubule organization in a reovirus-infected CV-1 cell. Cells were subjected to the indirect immunofluorescent staining technique described under Materials and Methods using antitubulin antibody. Bar represents 25 pm.
rinsing in PBS, the fixed cells were incubated with antisera as described above. All cells were examined with a Zeiss photomicroscope III equipped with epifluorescence and a Planapochromat objective lens
(63X). Photomicrographs were made using Kodak Tri-X (ASA 400) at exposure index 1606. Loealixation of mitochondria Mitochondria were visualized in living cells using
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FIG. 2.--Continued
the procedure of Johnson, Walsh, and Chen (1980). CV-1 cells were grown on 12mm round glass coverslips and infected with reovirus as described above. At 12, 24, and 48 hr postinfection, the coverslips were removed from the culture dishes, transferred to IMEMZO medium containing 10 pg/ml of the laser dye rhodamine 123 (Eastman), and incubated.at 37” in a 5% COz incubator for 30 min. Following extensive rinsing, the coverslips were mounted on a microscope slide with a silicon rubber chamber suitable for live cell examination as described by Johnson et ad (1980). They were examined by epifluorescent illumination at either 546 nm (rhodamine excitation) or 485 nm (fluorescein excitation). Electron microscopy. Fixation and procedures for transmission electron microscope were according to the methods of Goldman (1971) and Goldman et cd (1975). RESULTS
Cytoskeletcm of CV-1 Cells Monkey CV-1 cells were chosen for these studies because of their capacity to support the efficient growth of reovirus. In addition, since they are well spread and
flattened cells in monolayers, they are excellent cells to use for visualizing the cell cytoskeleton. The cytoskeleton of CV-1 cells has been described (Sharpe et cd, 1980). Briefly, CV-1 cells have an extensive network of microtubules emanating from the cell nucleus (Fig. 2A) and have typical bundles of microfilaments in the cytoplasm (Fig. 3A). The vimentin filament system emanates from multiple sites surrounding the nucleus (Fig. 4A). Distribution Cells
of Reovirus Antigen in CV-1
The maturation of reovirus in the cytoplasm of CV-1 cells as studied by immunocytochemical staining with antisera to reovirus was similar to that described previously (Rhim et al, 1962; Spendlove et uL, 1963; Fields et al, 1971). Viral antigens were detected within the cytoplasm as well as within viral factories. Viral inclusions first appeared as small immunofluorescent spots typically located in the cell periphery. As the viral growth cycle progressed, viral factories increased in size and appeared to fuse to form large, irregularly shaped inclusions. The inclusions migrated from the cell periphery to a perinuclear location (Fig. 1).
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FIG. 3. Organization of microfilaments in mock-infected and reovirus type a-infected CV-1 in a at 48 hr after infection. (A) Fluorescence micrograph showing microfilament organization
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Eflect of Reovirus Iqfection on Microtuhuh The organization of microtubules was examined in reovirus-infected cells at 12, 24, and 43 hr postinfection using antitubulin antibody. Reovirus infection did not disrupt microtubule organization even at late times postinfection (Figs. 2B and C). Microtubules were seen to span the regions of cytoplasm containing viral factories without interruption or distortion. Similar results were observed with all three serotypes. Eflect of Redrw aments
Replication on Microjl-
The distribution of microfilament bundles in reovirus-infected cells at 12,24, and 48 hr postinfection was examined using a monoclonal antibody to actin. Microfilament organization was not disturbed by reovirus infection. In addition, viral inclusions did not appear to contain actin since the monoclonal antibody to actin did not stain viral inclusions as distinct structures (Figs. 3B and C). Thus, reovirus infection does not appear to alter actin filament organization within CV-1 cells. Efect of Reovirus Replication on Intermediate Filaments The distribution of vimentin filaments was examined in cells infected with all three serotypes using antiserum to vimentin. By 12 hr postinfection, before intracytoplasmic inclusions were visible, the perinuclear organization sites of vimentin filaments disappeared. As infection progressed, there was further disruption of these filaments such that wavy filaments with no apparent organization were observed in the cytoplasm (Figs. 4B, C, and E). Similar results were observed with all three serotypes. The contrast between intermediate filament organization in cells
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containing viral inclusions and cells not containing them is illustrated clearly in Figs. 4B and C. Inclusion-bearing CV-1 cells that have disrupted intermediate filament organization are adjacent to CV-1 cells without inclusions that have the typical organization of intermediate filaments. Viral inclusions were stained to varying degrees with vimentin antibody (Figs. 4B and D). When viral inclusions were examined at a high magnification (630-fold), filamentous structures which stained with vimentin antibody were observed within the inclusions (Fig. 5). Electron microscopic examination of viral inclusions at a similar time postinfection reveals the presence of kinky filaments, which are 100 A in diameter (Fig. 6). Thus, reovirus infection disrupts vimentin filament organization within the cytoplasm of CV-1 cells and results in the presence of vimentin within viral inclusions. Eflect of Reovirms Replication on Mitt+ chondrial Llistribution CV-1 cells have a characteristic discontinuous filamentous distribution of mitochondria within the cytoplasm (Fig. 7A). Reovirus infection leads to a disorganization of mitochondria distribution in living cells as viewed with the fluorescent probe, rhodamine 123 (Figs. ‘7B and C). Mitochondria aggregated around the nucleus and occasionally appeared isolated at the cell periphery. Mitochondria were excluded from the viral factories. These findings are consistent with the observation that an intact intermediate filament system is essential for normal distribution of mitochondria in CV-1 cells (Chen et aZ., 1981). DISCUSSION
These studies show that reovirus infection produces a major disruption of vi-
mock-infected CV-1 cell. (B) Phase contrast micrograph of a type 3-infected CV-1 cell showing phase dense inclusion bodies. (C) Fluorescence micrograph of the same infected cell as in (B) showing microfilament organization in a reovirus-infected CV-1 cell. Cells were subjected to indirect immunofluorescence microscopy using a monoclonal antiactin antibody. Bar represents 25 pm.
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FIG. 4. Organization of intermediate filaments in mock-infected and reovirua type a-infected CV1 cells at 48 hr after infection. (A) Fluorescence micrograph showing intermediate filament organization in mock-infected CV-1 cells. (B and D) Phase contrast micrographs of type 3-infected CV-1 cells showing phase dense inclusion bodies. (C and E) Fluorescence micrographs of the same infected cells as in B and D, respectively, showing intermediate filament organization in reovirusinfected CV-1 cells. Note that intermediate filament organization is affected by reovirus infection. Cells were subjected to indirect immunofluorescence microscopy using antibody against vimentin. (A, B, and C, bar = 40 pm; D and E. bar = 25 pm.)
mentin filament organization without producing a discernable effect upon the organization of microtubules and microfilament bundles in CV-1 cells during the
period of observation. The distribution of mitochondria is also altered by viral infection. The disruption of vimentin filament or-
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FIG. 5. Fluorescence micrograph of a type 3-infected CV-1 cell at 48 hr after infection demonstrating vimentin filament organization within viral inclusions. Cell was subjected to indirect immunofluorescence microscopy using antibody against vimentin. Bar represents 20 pm.
ganization by reovirus is especially interesting in view of the finding that agents which disrupt protein synthesis such as diphtheria toxin, P. aeruginosa exotoxin A, and cycloheximide, specifically disrupt the organization of vimentin filaments without affecting microtubule or microfilament organization (Sharpe et al, 1980). The mammalian reoviruses also disrupt host cell macromolecular synthesis (Sharpe and Fields, 1981; Sharpe and Fields, 1982). It is possible that the disruption of intermediate filaments by reo-
virus is related to the capacity of reovirus to inhibit host cell protein synthesis. However, in addition to disrupting the organization of intermediate filaments, reovirus infection appears to cause a reorganization of vimentin filaments. Viral inclusions contain filamentous structures which stain with antibody to vimentin. These vimentin filamentous structures appear to be the complex filamentous network previously visualized in Dales’ electron microscopic studies of viral factories (Dales et aL, 1965). The presence of vi-
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FIG. 6. Electron micrograph of portion of a type 3-infected CV-1 cell containing a viral inclusion. Note kinky filaments present within the inclusion. These are exactly 100 A in diameter (magnification X7000).
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mentin filaments in the viral factories suggests that these filaments may be participating in the formation of unique viral structures that are distinct from vimentin filament organization in the cytoplasm of the noninfected cells. It is possible that reovirus is using the vimentin filament system in order to create a discrete cytoplasmic entity, the viral factory, which functions as the site of viral replication and assembly. It has been proposed that vimentin filaments function in the coordination of the organization of cytoplasmic contents (Lazarides, 1980). The fact that vimentin filaments are found within viral factories suggests that they may be playing a critical role in the organization of the viral replicative process. Unlike the marked disruption of intermediate filaments, reovirus infection did not disrupt microtubule structure. Electron microscopic studies have shown that reoviruses are aligned on parallel arrays of microtubules within viral factories (Dales et CAL,1965) and have the capacity to bind to microtubules in vitro (Babiss et al, 1979). These results are consistent with our observation that microtubules course through regions of the cytoplasm containing viral factories without disrupting or distorting them. Since colchicine treatment of reovirusinfected cells does not reduce viral yield, it is uncertain whether microtubules play a role in viral growth. Disruption of microtubules by colchicine treatment, however, alters the morphology of viral inclusions. Following the treatment of virally infected cells with colchicine only small inclusions located at the cell periphery are observed and the large perinuclear inclusions usually observed in cells infected with reovirus are absent (Spendlove et al., 1964). It is thus possible that microtubules play a role in the coalescence of viral inclusions and in their migration toward the nucleus. The concomitant disruption of vimentin filament organization and mitochondrial distribution after reovirus infection is consistent with a recent report by Chen et al. (1981) that suggests a role for vimentin filaments in determining the dis-
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FIG. 7. Distribution of mitochondria in mock-infected and reovirus type 3-infected CV-1 cells at 48 hr after infection. (A) Fluorescence micrograph of a live CV-1 cell showing rhodamine W&stained mitochondria. Dotted lines indicate cell periphery. (B) Phase contrast micrograph of a live type I-infected CV-1 cell showing phase dense inclusion bodies. (C) Fluorescence micrograph of same infected cells as in (B) showing the distribution of rhodamine-stained mitochondria. Note that the distribution of mitochondria is affected by reovirus infection. Bar represents 25 pm.
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tribution of mitochondria in cultured cells. Although infection of cells with reovirus alters the distribution of mitochondria, whether reovirus infection also alters mitochondrial functions remains to be determined. The accumulation of rhodamine 123 by mitochondria reflects the transmembrane potential (Johnson et aL, 1981). Because the accumulation of rhodamine 123 in reovirus-infected cells is similar to that of uninfected cells, it is likely that the respiratory activity of infected and uninfected cells is similar. A role for the cytoskeleton in the growth and assembly of other viruses has also been proposed. For example, immunofluorescence and electron microscopic studies of pox virus interactions with the cytoskeleton of infected chick embryo fibroblasts have revealed an intimate association between mature virus particles and the cytoskeleton that involves actin-containing structures. Vaccinia virus infection appears to induce the formation of specialized microvillus-type structures, which contain a core of microfilaments and a virus particle at their tip. These microvilli may be involved in viral budding and release from infected cells (Hiller et aL, 1979). An immunofluorescence microscopic study of the fusion of cells infected with the parainfluenza virus SV-5 revealed that microtubules and intermediate filaments form large bundles during cell fusion. This observation has suggested that both types of filaments may provide a framework which directs nuclear migration during cell fusion (Wang et aL, 1979). The results presented here suggest that intermediate filaments may play an important role in the replication and assembly of reovirus. The biochemical and morphological properties of intermediate filaments suggest that these filaments are involved in the organization of components within the cytoplasmic space (Lazarides, 1980). With such a function, intermediate filaments may be expected to play a crucial role in the regulation of cell shape to be influenced by the cell’s metabolic state and response to injury. Possibly, intermediate filaments are one of the major cellular targets involved in producing the cytopathic effects resulting from
reovirus infection (rounding of cells, inclusion formation, etc.). It remains to be determined if intermediate filament disorganization (and/or reorganization) is a primary and central step in viral infection, leading ultimately to cell death. ACKNOWLEDGMENTS
We thank Lincoln V. Johnson for help with immunocytochemical staining techniques and for stimulating discussions and Karen Byers for excellent technical assistance. During the course of these studies, A.H.S. was a National Institutes of Health Predoctoral Trainee (5-T32 GM0 7196). This investigation was supported by Grants AI 13178 from the National Institutes of Health (to B.N.F.) and CA 22619 from the National Cancer Institute (to L.B.C.). L.B.C. is the recipient of an American Cancer Society Faculty Research Award. REFERENCES
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