VP40 Octamers Are Essential for Ebola Virus Replic
http://www.100md.com
病菌学杂志 2005年第3期
Institut für Virologie, Marburg, Germany
Claude Bernard University Lyon 1, INSERM U 412, IFR 128, Lyon
European Molecular Biology Laboratory, Grenoble, France
ABSTRACT
Matrix protein VP40 of Ebola virus is essential for virus assembly and budding. Monomeric VP40 can oligomerize in vitro into RNA binding octamers, and the crystal structure of octameric VP40 has revealed that residues Phe125 and Arg134 are the most important residues for the coordination of a short single-stranded RNA. Here we show that full-length wild-type VP40 octamers bind RNA upon HEK 293 cell expression. While the Phe125-to-Ala mutation resulted in reduced RNA binding, the Arg134-to-Ala mutation completely abolished RNA binding and thus octamer formation. The absence of octamer formation, however, does not affect virus-like particle (VLP) formation, as the VLPs generated from the expression of wild-type VP40 and mutated VP40 in HEK 293 cells showed similar morphology and abundance and no significant difference in size. These results strongly indicate that octameric VP40 is dispensable for VLP formation. The cellular localization of mutant VP40 was different from that of wild-type VP40. While wild-type VP40 was present in small patches predominantly at the plasma membrane, the octamer-negative mutants were found in larger aggregates at the periphery of the cell and in the perinuclear region. We next introduced the Arg134-to-Ala and/or the Phe125-to-Ala mutation into the Ebola virus genome. Recombinant wild-type virus and virus expressing the VP40 Phe125-to-Ala mutation were both rescued. In contrast, no recombinant virus expressing the VP40 Arg134-to-Ala mutation could be recovered. These results suggest that RNA binding of VP40 and therefore octamer formation are essential for the Ebola virus life cycle.
INTRODUCTION
The filoviruses Ebola virus and Marburg virus belong to the order of nonsegmented negative-strand RNA viruses (Mononegavirales) and are the causative agents of a severe and often fatal hemorrhagic fever in humans (6). Genomes of most enveloped RNA viruses encode a matrix protein, which is indispensable for efficient assembly and budding of virus particles (7). Viral matrix proteins interact with membranes and most likely with viral glycoproteins and the ribonucleoprotein complex in a process which drives assembly and positions the matrix proteins underneath the viral membranes.
Accordingly, Ebola virus matrix protein VP40 is the most abundant protein in virus particles, which are filamentous and show distinct variations in length (5, 8). The filamentous morphology is determined by VP40, consistent with the appearance of virus-like particle (VLP) formation upon VP40 expression (11, 31). Notably, VLP formation is improved by the coexpression of VP40 and glycoprotein GP (1, 17, 25), which is recruited into lipid raft microdomains that serve as an assembly platform for a number of enveloped viruses (1, 19, 20, 25, 26, 28, 37).
The role of VP40 in budding is consistent with the presence of late domain sequences at its N terminus, which have been reported to interact with cellular factors (10, 16, 21, 32, 36), such as Tsg101. Such factors participate in multiprotein complex formations that drive enveloped virus budding processes (22, 30, 35) as well as the sorting of plasma membrane receptors into multivesicular bodies at late endosomal membranes (12). The involvement of late endosomal membranes in filovirus assembly was also demonstrated by the localization of Marburg virus VP40 to the late endosome, the presence of endosomal markers in virus particles (13), and vesicular transport of VP40 from late endosomes to the plasma membrane (14), where assembly and budding take place (8).
VP40 forms monomers in solution that are composed of two structurally related beta sandwich domains, which associate in a metastable conformation (4). In vitro, the monomeric conformation of VP40 can be switched to an oligomeric ring-like structure by several means, including urea treatment (29), in agreement with the metastability of VP40. We have further shown that the N-terminal domain of VP40 is sufficient for oligomerization, while the C-terminal domain is necessary for membrane interaction (27). Single-particle electron microscopy analysis suggested that the C-terminal membrane binding domain is flexibly linked to the ring-like structure formed by the N-terminal domain (29). A recent report also suggested that oligomeric VP40 is enriched in lipid rafts (26), consistent with the role of lipid microdomains in virus assembly (19, 20, 28, 37). In addition, these data may indirectly implicate membrane targeting in the conversion of monomers to oligomers, as suggested by in vitro studies (29).
We previously reported that oligomeric ring-like VP40 structures consist of either hexamers or octamers in vitro and that octamer formation is critically dependent on RNA binding (9, 33). Accordingly, the crystal structure of the octameric ring-like structure formed by the N-terminal domain revealed that the Ebola virus matrix protein is a sequence-specific single-stranded RNA (ssRNA) binding protein (9). The central pore of octameric VP40 binds eight copies of a short RNA with the sequence 5'-UGA-3' at the dimer-dimer interface (9). As suggested by the crystal structure and confirmed biochemically, RNA binding plays an important structural role, as no octamers could be observed in the absence of RNA (33). The structure also suggested that two conserved residues, Phe125 and Arg134, are the most important residues for RNA binding (9). In addition, RNA binding confers sodium dodecyl sulfate (SDS) resistance to the oligomer composed of either the N-terminal domain alone or full-length wild-type VP40 when separated under nonboiling conditions by SDS-polyacrylamide gel electrophoresis (PAGE) (9, 33).
Here we confirm that octamer formation is also driven by RNA binding in HEK 293 cells expressing VP40. Interestingly, octamers are not necessary for the formation of VP40-containing VLPs, although they appear to be critical for Ebola virus replication, as a single mutation that blocks RNA binding inhibits the formation of infectious particles. This report shows for the first time that the sequence-specific RNA binding of a matrix protein may be essential for the virus life cycle.
MATERIALS AND METHODS
Cell lines. HEK 293, HUH-T7, and Vero E6 cells were cultured at 37°C in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal calf serum (FCS). BSR T7/5 cells (BHK 21 cells stably expressing T7 RNA polymerase) were maintained in Glasgow medium supplemented with 10% newborn calf serum.
Molecular cloning. VP40 was subcloned from plasmid pTM-VP40 (14) into vector pCAGGS by using restriction endonucleases EcoRI and XhoI. The VP40-FA, VP40-RA, and VP40-RFA mutants were constructed by using a QuikChange site-directed mutagenesis kit (Stratagene) according to the supplier's instructions with pTM-VP40 serving as a template. For constructing the VP40-FA mutant, primers 1271 (5'-CAC TAT CAC CCA TGC CGG CAA GGC AAC-3') and 1272 (5'-GTT GCC TTG CCG GCA TGG GTG ATA GTG-3') were used; the mutated nucleotides are underlined. To construct the VP40-RA mutant, primers 1273 (5'-CAA TCC ACT TGT CGC AGT CAA TCG GCT GGG-3') and 1274 (5'-CCC AGC CGA TTG ACT GCG ACA AGT GGA TTG-3') were used. The VP40-RFA mutant was constructed by using primers 1273 and 1274 together with pTM-VP40-FA as a template. All constructs then were subcloned into vector pCAGGS by using EcoRI and XhoI and were verified by sequencing.
Transfection of cells. HEK 293 or HUH-T7 cells were grown in six-well plates (7 cm2) to a confluence of 70%. Transfection was performed with Lipofectamine Plus (Invitrogen) according to the manufacturer's instructions. Briefly, 1 μg of DNA was diluted in 100 μl of serum-free DMEM, and 6 μl of Lipofectamine Plus solution was added. In parallel, 4 μl of Lipofectamine was diluted in 100 μl of serum-free DMEM. After 15 min of incubation, the two solutions were combined. After an additional 15 min of incubation, the transfection mixture was added to the cells, which had been washed twice with serum-free DMEM and then brought to a medium volume of 800 μl. After 3 h of incubation at 37°C, 2.5 ml of DMEM supplemented with 10% FCS was added to the cells. For harvesting, the cells were placed on ice, washed twice with ice-cold phosphate-buffered saline (PBS; 0.8% [wt/vol] NaCl, 0.02% [wt/vol] KCl, 0.115% [wt/vol] Na2HPO4, 0.02% [wt/vol] KH2PO4), and then scraped into 100 μl of PBS.
Release of VLPs. At 48 h after cell transfection, the supernatant was harvested and centrifuged at 5,000 x g and 4°C for 10 min to remove cellular debris. The supernatant then was loaded onto a 20% sucrose cushion and centrifuged at 250,000 x g and 4°C for 3 h. The pellet was resuspended in PBS and either subjected to immunoelectron microscopy analysis as described previously (14) or examined for the presence of SDS-resistant VP40 octamers as described below.
Detection of SDS-resistant VP40 octamers. Harvested cells or VLPs were lysed with 1% Triton X-100 for 5 min on ice. Cell lysates were cleared by centrifugation at 830 x g and 4°C for 5 min. The postnuclear supernatant or the lysed VLPs were mixed at room temperature with reducing SDS loading buffer (40% glycerol, 20% ?-mercaptoethanol, 0.5% [wt/vol] SDS, 20% 1 M Tris-HCl [pH 6.8], 0.2% [wt/vol] bromophenol blue) and separated by noncontinous gradient (8 to 12%) SDS-PAGE. Subsequently, Western blotting was performed as described previously (2). Ebola virus VP40 was detected with mouse monoclonal antibody 2C4 (18) at a dilution of 1:100. As a secondary antibody, a horseradish peroxidase-conjugated goat anti-mouse antibody (Dako) at a dilution of 1:20,000 was used. Bound secondary antibody was detected by using a SuperSignal Ultra kit (Pierce).
Detection of RNA bound to VP40. Cells were transfected as described above, with the exception that no additional medium was added at 3 h posttransfection. At 6 h posttransfection, the cells were labeled with 25 μCi of [3H]uridine (Amersham) per dish. At 24 h posttransfection, the cells were placed on ice, washed twice with ice-cold PBS, and lysed with 1% Triton X-100 for 20 min at 37°C in the presence or in the absence of 10 μg of RNase A (Qiagen). Subsequently, nuclei were pelleted at 830 x g, and the supernatant was subjected to SDS-PAGE and Western blotting as described above. The blots were dried overnight, and radioactivity was detected by exposure to BioImager plates for 4 weeks.
Recombinant VP40. The expression, purification, chemical cross-linking, and analysis of SDS resistance of recombinant VP40 were performed as described previously (4, 9, 33).
Indirect immunofluorescence. Cells were transfected as described above and fixed with 4% paraformaldehyde at 24 h posttransfection. Immunofluorescence analysis was performed as described previously (3) with mouse monoclonal antibody 2C4 at a dilution of 1:10 and a rhodamine-coupled goat anti-mouse secondary antibody (Dianova) at a dilution of 1:100. Nuclei were stained with 4',6'-diamidino-2-phenylindole.
Construction of plasmids and recovery of recombinant Ebola viruses. Full-length antigenomic cDNAs of Ebola virus and the protocol used for virus rescue were described elsewhere (34). VP40 mutations R125A and F134A were introduced by site-directed mutagenesis of intermediate plasmid pKSN4 (34). Full-length antigenomic cDNAs of Ebola virus mutants then were generated and analyzed for the presence of mutations by sequencing of the genome. The recovery of recombinant Ebola viruses was performed in three independent experiments.
RNA extraction and RT-PCR of recovered Ebola viruses. The recovered viruses were passaged three times in Vero E6 cells, and culture supernatants were harvested at 6 days postinfection and cleared by low-speed centrifugation. Viral RNA was extracted from the recovered viruses by using an RNeasy kit (Qiagen) according to the manufacturer's instructions. Reverse transcription (RT) and PCR amplification were carried out with forward (5'-AAA AAC CTA CCT CGG CTG AGA GAG T-3') and reverse (5'-AAG CAT GCA GGC AAT TTG AGG ATA AG-3') primers by using a Titan one-tube RT-PCR kit (Roche). To confirm the presence of the introduced mutations, the VP40 gene was sequenced.
Kinetics of multicycle growth of recombinant viruses. Monolayers of Vero E6 cells were infected at a multiplicity of infection of 0.001 PFU per cell, and aliquots of culture media were harvested at 24-h intervals for 6 days. Samples were analyzed by plaque assays in duplicate. Briefly, Vero E6 cells were infected with serial dilutions of cell-free supernatants and 1.6% carboxymethyl cellulose (BDH Laboratory Supplies) in DMEM supplemented with 2.5% FCS. Infectious foci were detected after 5 days of culturing by incubation with mouse anti-Ebola virus NP monoclonal antibodies followed by horseradish peroxidase-conjugated goat polyclonal anti-mouse immunoglobulin G (Sigma-Aldrich, Saint Quentin Fallavier, France) and TrueBlue peroxidase substrate (Kirkegaard & Perry Laboratories). Virus titers were expressed as focus-forming units per milliliter of culture medium. Culture medium and cell lysates collected 6 days postinfection were also analyzed by Western blotting with rabbit polyclonal antibodies directed against VP40 or VP24.
RESULTS
Specific RNA binding of VP40 expressed in HEK 293 cells. The crystal structure of octameric VP40 showed that the N-terminal domain of VP40 binds a short ssRNA with the sequence 5'-UGA-3'. Analysis of the interactions indicated that Arg134 and Phe125 contributed most to the coordination of the ssRNA fragment (9). In addition, both residues were exposed in the monomeric conformation of VP40 (Fig. 1). Based on these findings, we engineered three VP40 mutants, an Arg134-to-Ala mutant (VP40-RA), a Phe125-to-Ala mutant (VP40-FA), and a mutant with a double mutation of Phe125 and Arg134 to Ala (VP40-RFA), and analyzed them in a mammalian expression system.
We previously demonstrated that VP40 octamers show SDS resistance on SDS-PAGE when loaded under nonboiling conditions and that both resistance and octamer formation are dependent on the presence of nucleic acids (33). In order to test whether the mammalian expression of wild-type VP40 leads to the incorporation of RNA, we labeled cellular RNA with [3H]uridine. Analysis of VP40 expression by Western blotting showed a band for the monomer (37 kDa) and a band which migrated above the 220-kDa marker protein (Fig. 2A, lane 3), indicating SDS resistance and therefore the octameric form of VP40. Although we could detect a very faint band corresponding to the oligomer for VP40-FA (Fig. 2A), no octamers were detectable for VP40-RA and VP40-RFA. Monomeric mutant VP40, however, was expressed (Fig. 2A). In accordance with octamer formation, the amount of monomeric wild-type VP40 detected was smaller than that of the mutants (Fig. 2A).
Next, the same blot was exposed to a BioImager plate in order to detect radioactively labeled bands derived from VP40 expression in [3H]uridine-labeled mammalian cells. This experiment revealed a major radioactive band migrating above the 220-kDa marker protein and corresponding to the SDS-resistant high-molecular-mass form of VP40 shown in Fig. 2A (Fig. 2B, lane 3). Therefore, octameric VP40 also binds RNA upon expression in HEK 293 cells. Two additional radioactively labeled bands could be observed approximately between the 97- and 66-kDa marker proteins in all experiments. These bands appeared at the same intensities in all lanes, including the negative control. These bands therefore serve as an internal control and are consistent with the fact that the same amounts of labeled material were loaded in the lanes. As these additional bands were unrelated to VP40 octamerization, we suggest that they might represent abundant cellular RNA(s).
We next tested whether RNase A treatment of total cell lysates would affect VP40 octamerization. Following treatment, there was a decrease in the amount of detectable SDS-resistant wild-type VP40 octamers (Fig. 2C, lane 2). Similarly, an increase in the total amount of monomers could be detected (Fig. 2C, lanes 1 and 2). As indicated in Fig. 2A, VP40-FA still formed octamers at low levels, which were further reduced upon RNase A treatment (Fig. 2C, lanes 3 and 4). Only monomers were detected for VP40-RA and VP40-RFA (Fig. 2C, lanes 5 to 8).
In order to better understand the decrease in VP40 SDS resistance in the presence of RNase A, we performed in vitro SDS resistance experiments with long and short RNA molecules. We previously showed that monomeric full-length VP40 can be converted to SDS-resistant octamers in the presence of RNA and urea (9). Here we incubated VP40 with only urea, with urea in the presence of Escherichia coli RNA, or with urea in the presence of a short synthetic ssRNA with the sequence 5'-UGA-3', which is present in the crystal structure of octameric VP40 (9). Then we tested the oligomerization state of VP40 by SDS resistance and chemical cross-linking analyses. These analyses showed that VP40 alone did not produce SDS-resistant oligomers, as only bands corresponding to monomers were visible (Fig. 2D, lane 2). Urea treatment of VP40 in the absence of RNA led to the formation of high-molecular-mass aggregates that barely entered the gel, as detected by chemical cross-linking, indicating nonspecific aggregation (Fig. 2D, lane 3). VP40 treated with urea in the presence of E. coli nucleic acids showed a new SDS-resistant band that migrated above the 250-kDa marker protein, as shown previously (9) (Fig. 2D, lane 4). This oligomer could also be cross-linked, resulting in a ladder of bands, with the major species migrating more slowly than the SDS-resistant oligomer, as shown previously for VP40 octamers (33) (Fig. 2D, lane 5). Reconstitution of the octamer in the presence of short synthetic ssRNA with the sequence 5'-UGA-3', the minimal binding sequence detected in the crystal structure, did not result in SDS-resistant octamers. However, octamer formation could be shown by chemical cross-linking and was represented by a band migrating at the same position as VP40 cross-linked in the presence of E. coli RNA (Fig. 2D, lanes 6 and 7). These data are consistent with the reduction of SDS-resistant VP40 by RNase treatment and indicate that the proposed SDS resistance of octameric VP40 depends on the presence of longer ssRNA molecules. To further confirm that the Arg mutants no longer bound RNA, we expressed and purified recombinant VP40-RFA, which did not bind RNA; therefore, no octamers could be detected by SDS resistance and chemical cross-linking analyses in vitro (data not shown).
VLP formation induced by wild-type or mutant VP40 expression. We next tested whether the mutations affected VP40 release into cell culture supernatants and the formation of VLPs. These analyses showed that all mutant VP40 constructs induced the release of equal amounts of VP40 into supernatants upon expression in HEK 293 cells (Fig. 3A). Negative staining electron microscopy revealed filamentous particles produced by wild-type VP40 and mutant VP40; these particles showed similar morphologies and no significant difference in size. Particle size analysis revealed diameters (mean and standard deviation) of 41 ± 10 nm for wild-type VP40, 41 ± 9 nm for VP40-FA, 44 ± 10 for VP40-RA, and 40 ± 7 nm for VP40-RFA VLPs (Fig. 3B). Wild-type VLPs contained SDS-resistant VP40 octamers similar to those detected in cells (Fig. 3C, lanes 1 and 2) and reported previously (26). However, VLPs generated from VP40-RA did not contain VP40 octamers (Fig. 3C, lane 3), consistent with the absence of octamers in cells expressing VP40-RA (Fig. 3C, lane 4). In addition, VP40-RA VLPs contained dimers of VP40 (Fig. 3C, lane 3) similar to those detected in Ebola virus particles (9).
Cellular localization of wild-type or mutant VP40. Indirect immunofluorescence analysis indicated that the intracellular localization of the three mutant constructs differed from that of wild-type VP40, although the transfection efficiencies (20% for all constructs) and the amounts of VP40 expressed inside cells and released into cell culture supernatants were similar in all experiments, as judged by Western blot analysis (data not shown). While wild-type VP40 was distributed mainly in small patches along the plasma membrane (Fig. 4), mutant VP40-FA was detected in larger aggregates along the plasma membrane as well as in the perinuclear region (Fig. 4). In contrast, mutant VP40-RA and double mutant VP40-RFA were found mostly in large aggregates in the perinuclear region and at the plasma membrane. In addition to the larger intracellular aggregates formed by VP40-RA and VP40-RFA, the extent of cytoplasmic VP40 distribution seemed to be reduced compared to that of wild-type VP40 and VP40-FA (Fig. 4).
Effects of VP40 mutations on Ebola virus replication. Finally, we used an established reverse-genetics system for Ebola virus (34) to examine the significance of the VP40 mutations for virus replication. VP40 mutations F125A and R134A were introduced into Ebola virus cDNA in an otherwise identical genomic background, and supernatants from transfected BSR T7/5 cells were passaged in Vero E6 cells. Clear cytopathic effects were observed with constructs containing wild-type VP40 as well as mutant VP40-FA. To demonstrate the recovery of recombinant viruses, viral RNA from culture supernatants derived from the third consecutive passage of virus in Vero E6 cells was assayed by RT-PCR, followed by sequence analysis. Nucleotide sequencing confirmed the presence of the F125A mutation in recovered recombinant viruses. Subsequent analysis of virus replication in multicycle virus growth revealed no significant differences in endpoint titers between wild-type and mutant viruses carrying the VP40-FA mutation. However, the growth curves indicated that the VP40-FA-containing recombinant virus had a slightly increased ability to replicate in Vero E6 cells, as demonstrated by higher virus titers at an early stage of infection (Fig. 5A). For the VP40-FA-containing mutant virus, Western blot analysis of total cell lysates and cell culture supernatants with anti-VP40 and anti-VP24 antibodies revealed that the culture supernatants contained slightly higher levels of VP40 and VP24 (Fig. 5B). In contrast, for the wild-type Ebola virus, slightly higher levels of intracellular VP40 and VP24 were detected (Fig. 5B).
Several attempts to recover recombinant virus containing the VP40 R134A mutation (VP40-RA or VP40-RFA) failed. We therefore suggest that Arg at position 134 has an important role in virus replication. As the same mutant showed a defect in RNA binding upon expression in HEK 293 cells, the rescue experiments strongly indicated that RNA binding, which confers VP40 octamer formation, is important for the life cycle of Ebola virus.
DISCUSSION
The crystal structure of the N-terminal domain of Ebola virus VP40 in complex with ssRNA indicated that specific RNA binding is responsible for the formation of SDS-resistant octameric VP40 (9, 33). Here we established that octameric VP40 binds cellular RNA labeled with [3H]uridine when expressed in HEK 293 cells. Detection of SDS-resistant VP40 is sensitive to RNase treatment, in agreement with our in vitro data showing that VP40 octamer formation with short ssRNA molecules with the sequence 5'-UGA-3' is detectable only by chemical cross-linking and no longer by SDS resistance. As the 5' and 3' ends of the bound ssRNA molecules are freely accessible in the crystal structure, it is conceivable that only longer RNA molecules contribute to SDS stability (9). Furthermore, in the octamer, the central pore containing the RNA binding site carries a mostly basic charge (9), which might allow for additional nonspecific RNA binding that could further stabilize the octamer and thus contribute to its stability in the presence of SDS. Therefore, SDS resistance is only an indication of octamer formation and cannot be used as a quantitative measurement of octamer formation.
The atomic details of the octameric crystal structure indicated an important role for Arg134 and Phe125 in RNA binding (Fig. 1); this finding was confirmed by our mutational analysis, which showed reduced RNA binding for VP40-FA and no RNA binding for VP40-RA and VP40-RFA. The conclusion that the abolishment of RNA binding leads to the blockage of octamer formation is also in agreement with the described structural role of ssRNA in VP40 octamer formation (9, 33). The complete absence of RNA binding and thus of octamer formation in the presence of the Arg134 mutation was also confirmed by analysis of recombinant VP40-RFA in vitro.
The cellular localization of the VP40-FA mutant was still similar to that of wild-type VP40 (1, 10, 21, 26, 31). However, the intracellular aggregates formed were larger, and they were even more pronounced for VP40-RA and VP40-RFA. Although the differences in localization might be due to the presence or absence of VP40 octamers, it is possible that the mutations themselves interfere with the active transport of VP40 (14). Furthermore, the mutations, which are both located in or close to a basic patch at the interface of the N- and C-terminal domains of monomeric VP40 (Fig. 1A) (4), might interfere with a proposed switch from the monomeric form to an oligomeric form (29) or a polymeric form of VP40. Therefore, mutation of Phe125 to Ala and, to an even greater extent, of Arg134 to Ala might lead to increased monomer destabilization, resulting in larger intracellular aggregates containing VP40 oligomers or polymers and a shorter half-life of cytoplasmic monomers, compared to the properties of wild-type VP40.
It was previously shown that VP40 expression as well as VP40 and GP coexpression in mammalian cells induces the release of filamentous VLPs (1, 11, 17, 25, 31). Here we show that the VLPs produced by wild-type VP40 expression as well as the expression of mutant VP40 have similar filamentous morphologies, with particle diameters ranging from 40 to 50 nm. However, the VLP diameters reported here are smaller than those described for released VLPs containing VP40 alone (65 nm) (25) but are similar to those of filamentous particles inside cells expressing VP40 (45 nm) (25). In addition, VLPs produced by VP40 and GP coexpression had diameters ranging from 50 to 70 nm (1) or an outer diameter of 80 nm (25); these data indicate that the diameters of VLPs can vary greatly. The final conserved 80-nm diameter of infectious viruses (5) is therefore most likely driven by the presence of the nucleocapsid or other unknown viral or cellular factors. Our data indicate that RNA binding and octamer formation, which are completely abolished in VP40-RA and VP40-RFA, are not necessary for VLP formation, although SDS-resistant octamers are present in VLPs produced by wild-type VP40, consistent with previously published data (26).
When the VP40 mutations were introduced into the Ebola virus genome, only recombinant wild-type virus and virus carrying the VP40-FA mutation could be rescued (34). These data strongly indicate that VP40 RNA binding and octamer formation might be important for the life cycle of Ebola virus, as VP40-RA showed a complete block in RNA binding and thus octamer formation. In contrast, when expressed alone, the second mutation, VP40-FA, still resulted in reduced RNA binding and octamer formation, which seemed to be sufficient for virus replication. Interestingly, when growth curves for wild-type and recombinant viruses carrying the VP40-FA mutation were compared, the VP40-FA-carrying virus showed a slight increase in replication kinetics. These data suggest that while RNA binding seems to be important for virus replication, the VP40-FA-carrying mutant virus might have a slight advantage in assembly over the wild-type virus. We propose that this effect is not dependent on VP40 RNA binding but rather is due to a potential destabilizing effect of the F125A mutation, as discussed above, which might facilitate the proposed transition from a monomeric form to an assembly-activated form of VP40.
In summary, our data indicate that the octamerization deficient mutants is still active in VLP formation, a finding which implies that they are competent to facilitate virion assembly and promote the egress of newly produced virus particles from the plasma membrane. Therefore, octameric VP40 must exert another, as-yet-unknown function during Ebola virus replication, a function which we propose to be important for the virus life cycle.
ACKNOWLEDGMENTS
We thank M. Rossi for expert technical help and Allison Groseth for critical reading of the manuscript. All experiments involving Ebola virus were carried out in biosafety level 4 laboratories at the Institute of Virology in Marburg, Germany, and at the Jean Merieux P4 Center in Lyon, France.
This work was supported by EMBL (W.W.), by INSERM (V.V.), and by funds from Deutsche Forschungsgemeinschaft SFB 593 Marburg (to W.W., S.B., and V.V.). T.H. was supported by a scholarship from Verband der Chemischen Industrie.
REFERENCES
Bavari, S., C. M. Bosio, E. Wiegand, G. Ruthel, A. B. Will, T. W. Geisbert, M. Hevey, C. Schmaljohn, A. Schmaljohn, and M. J. Aman. 2002. Lipid raft microdomains: a gateway for compartmentalized trafficking of Ebola and Marburg viruses. J. Exp. Med. 195:593-602.
Becker, S., H. D. Klenk, and E. Muhlberger. 1996. Intracellular transport and processing of the Marburg virus surface protein in vertebrate and insect cells. Virology 225:145-155.
Becker, S., C. Rinne, U. Hofsass, H. D. Klenk, and E. Muhlberger. 1998. Interactions of Marburg virus nucleocapsid proteins. Virology 249:406-417.
Dessen, A., V. Volchkov, O. Dolnik, H. D. Klenk, and W. Weissenhorn. 2000. Crystal structure of the matrix protein VP40 from Ebola virus. EMBO J. 19:4228-4236.
Ellis, D. S. 1987. The Filoviridae, p. 313-321. In M. V. Nermut and A. C. Steven (ed.), Animal virus structure, 3rd ed. Elsevier Science Publishing, Inc., New York, N.Y.
Feldmann, H., V. E. Volchkov, V. A. Volchkova, U. Stroher, and H. D. Klenk. 2001. Biosynthesis and role of filoviral glycoproteins. J. Gen. Virol. 82:2839-2848.
Garoff, H., R. Hewson, and D. J. Opstelten. 1998. Virus maturation by budding. Microbiol. Mol. Biol. Rev. 62:1171-1190.
Geisbert, T. W., and P. B. Jahrling. 1995. Differentiation of filoviruses by electron microscopy. Virus Res. 39:129-150.
Gomis-Rüth, F. X., A. Dessen, J. Timmins, A. Bracher, L. Kolesnikova, S. Becker, H. D. Klenk, and W. Weissenhorn. 2003. The matrix protein VP40 from Ebola virus octamerizes into pore-like structures with specific RNA binding properties. Structure 11:423-433.
Harty, R. N., M. E. Brown, G. Wang, J. Huibregtse, and F. P. Hayes. 2000. A PPxY motif within the VP40 protein of Ebola virus interacts physically and functionally with a ubiquitin ligase: implications for filovirus budding. Proc. Natl. Acad. Sci. USA 97:13871-13876.
Jasenosky, L. D., G. Neumann, I. Lukashevich, and Y. Kawaoka. 2001. Ebola virus VP40-induced particle formation and association with the lipid bilayer. J. Virol. 75:5205-5214.
Katzmann, D. J., G. Odorizzi, and S. D. Emr. 2002. Receptor downregulation and multivesicular-body sorting. Nat. Rev. Mol. Cell. Biol. 3:893-905.
Kolesnikova, L., H. Bugany, H. D. Klenk, and S. Becker. 2002. VP40, the matrix protein of Marburg virus, is associated with membranes of the late endosomal compartment. J. Virol. 76:1825-1838.
Kolesnikova, L., S. Bamberg B. Berghofer, and S. Becker. 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.
Kraulis, P. 1991. MOLSCRIPT: a program to produce both detailed and schematic plots of protein structures. J. Appl. Crystallogr. 24:924-950.
Licata, J. M., M. Simpson-Holley, N. T. Wright, Z. Han, J. Paragas, and R. N. Harty. 2003. Overlapping motifs (PTAP and PPEY) within the Ebola virus VP40 protein function independently as late budding domains: involvement of host proteins TSG101 and VPS-4. J. Virol. 77:1812-1819.
Licata, J. M., R. F. Johnson, Z. Han, and R. N. Harty. 2004. Contribution of Ebola virus glycoprotein, nucleoprotein, and VP24 to budding of VP40 virus-like particles. J. Virol. 78:7344-7351.
Lucht, A., R. Grunow, P. Moller, H. Feldmann, and S. Becker. 2003. Development, characterization and use of monoclonal VP40-antibodies for the detection of Ebola virus. J. Virol. Methods 111:21-28.
Manes, S., G. del Real, R. A. Lacalle, P. Lucas, C. Gomez-Mouton, S. Sanchez-Palomino, R. Delgado, J. Alcami, E Mira, and A. C. Martinez. 2000. Membrane raft microdomains mediate lateral assemblies required for HIV-1 infection. EMBO Rep. 1:190-196.
Manie, S. N., S. Debreyne, S. Vincent, and D. Gerlier. 2000. Measles virus structural components are enriched into lipid raft microdomains: a potential cellular location for virus assembly. J. Virol. 74:305-311.
Martin-Serrano, J., T. Zang, and P. D. Bieniasz. 2001. HIV-1 and Ebola virus encode small peptide motifs that recruit Tsg101 to sites of particle assembly to facilitate egress. Nat. Med. 7:1313-1319.
Martin-Serrano, J., A. Yarovoy, D. Perez-Caballero, and P. D. Bieniasz. 2003. Divergent retroviral late-budding domains recruit vacuolar protein sorting factors by using alternative adaptor proteins. Proc. Natl. Acad. Sci. USA 100:12414-12419.
Merritt, E. A., and D. J. Bacon. 1997. Raster3D photorealistic graphics. Methods Enzymol. 277:505-524.
Nicholls, A., K. A. Sharp, and B. Honig. 1991. Protein folding and association: insights from the interfacial and thermodynamic properties of hydrocarbons. Proteins 11:281-296.
Noda, T., H. Sagara, E. Suzuki, A. Takada, H. Kida, and Y. Kawaoka. 2002. Ebola virus VP40 drives the formation of virus-like filamentous particles along with GP. J. Virol. 76:4855-4865.
Panchal, R. G., G. Ruthel, T. A. Kenny, G. H. Kallstrom, D. Lane, S. S. Badie, L. Li, S. Bavari, and M. J. Aman. 2003. In vivo oligomerization and raft localization of Ebola virus protein VP40 during vesicular budding. Proc. Natl. Acad. Sci. USA 100:15936-15941.
Ruigrok, R. W., G. Schoehn, A. Dessen, E. Forest, V. Volchkov, O. Dolnik, H. D. Klenk, and W. Weissenhorn. 2000. Structural characterization and membrane binding properties of the matrix protein VP40 of Ebola virus. J. Mol. Biol. 300:103-112.
Scheiffele, P., A. Rietveld, T. Wilk, and K. Simons. 1999. Influenza viruses select ordered lipid domains during budding from the plasma membrane. J. Biol. Chem. 274:2038-2044.
Scianimanico, S., G. Schoehn, J. Timmins, R. H. Ruigrok, H. D. Klenk, and W. Weissenhorn. 2000. Membrane association induces a conformational change in the Ebola virus matrix protein. EMBO J. 19:6732-6741.
Strack, B., A. Calistri, S. Craig, E. Popova, and H. G. G?ttlinger. 2003. AIP1/ALIX is a binding partner for HIV-1 p6 and EIAV p9 functioning in virus budding. Cell 114:689-699.
Timmins, J., S. Scianimanico, G. Schoehn, and W. Weissenhorn. 2001. Vesicular release of Ebola virus matrix protein VP40. Virology 283:1-6.
Timmins, J., G. Schoehn, S. Ricard-Blum, S. Scianimanico, T. Vernet, R. W. Ruigrok, and W. Weissenhorn. 2003. Ebola virus matrix protein VP40 interaction with human cellular factors Tsg101 and Nedd4. J. Mol. Biol. 326:493-502.
Timmins, J., G. Schoehn, C. Kohlhaas, H. D. Klenk, R. W. Ruigrok, and W. Weissenhorn. 2003. Oligomerization and polymerization of the filovirus matrix protein VP40. Virology 312:359-368.
Volchkov, V. E., V. A. Volchkova, E. Muhlberger, L. V. Kolesnikova, M. Weik, O. Dolnik, and H. D. Klenk. 2001. Recovery of infectious Ebola virus from complementary DNA: RNA editing of the GP gene and viral cytotoxicity. Science 291:1965-1969.
von Schwedler, U. K., M. Stuchell, B. Muller, D. M. Ward, H. Y. Chung, E. Morita, H. E. Wang, T. Davis, G. P. He, D. M. Cimbora, A. Scott, H. G. Krausslich, J. Kaplan, S. G. Morham, and W. I. Sundquist. 2003. The protein network of HIV budding. Cell 114:701-713.
Yasuda, J., M. Nakao, Y. Kawaoka, and H. Shida. 2003. Nedd4 regulates egress of Ebola virus-like particles from host cells. J. Virol. 77:9987-9992.
Zhang, J., A. Pekosz, and R. A. Lamb. 2000. Influenza virus assembly and lipid raft microdomains: a role for the cytoplasmic tails of the spike glycoproteins. J. Virol. 74:4634-4644.(Thomas Hoenen, Viktor Vol)
Claude Bernard University Lyon 1, INSERM U 412, IFR 128, Lyon
European Molecular Biology Laboratory, Grenoble, France
ABSTRACT
Matrix protein VP40 of Ebola virus is essential for virus assembly and budding. Monomeric VP40 can oligomerize in vitro into RNA binding octamers, and the crystal structure of octameric VP40 has revealed that residues Phe125 and Arg134 are the most important residues for the coordination of a short single-stranded RNA. Here we show that full-length wild-type VP40 octamers bind RNA upon HEK 293 cell expression. While the Phe125-to-Ala mutation resulted in reduced RNA binding, the Arg134-to-Ala mutation completely abolished RNA binding and thus octamer formation. The absence of octamer formation, however, does not affect virus-like particle (VLP) formation, as the VLPs generated from the expression of wild-type VP40 and mutated VP40 in HEK 293 cells showed similar morphology and abundance and no significant difference in size. These results strongly indicate that octameric VP40 is dispensable for VLP formation. The cellular localization of mutant VP40 was different from that of wild-type VP40. While wild-type VP40 was present in small patches predominantly at the plasma membrane, the octamer-negative mutants were found in larger aggregates at the periphery of the cell and in the perinuclear region. We next introduced the Arg134-to-Ala and/or the Phe125-to-Ala mutation into the Ebola virus genome. Recombinant wild-type virus and virus expressing the VP40 Phe125-to-Ala mutation were both rescued. In contrast, no recombinant virus expressing the VP40 Arg134-to-Ala mutation could be recovered. These results suggest that RNA binding of VP40 and therefore octamer formation are essential for the Ebola virus life cycle.
INTRODUCTION
The filoviruses Ebola virus and Marburg virus belong to the order of nonsegmented negative-strand RNA viruses (Mononegavirales) and are the causative agents of a severe and often fatal hemorrhagic fever in humans (6). Genomes of most enveloped RNA viruses encode a matrix protein, which is indispensable for efficient assembly and budding of virus particles (7). Viral matrix proteins interact with membranes and most likely with viral glycoproteins and the ribonucleoprotein complex in a process which drives assembly and positions the matrix proteins underneath the viral membranes.
Accordingly, Ebola virus matrix protein VP40 is the most abundant protein in virus particles, which are filamentous and show distinct variations in length (5, 8). The filamentous morphology is determined by VP40, consistent with the appearance of virus-like particle (VLP) formation upon VP40 expression (11, 31). Notably, VLP formation is improved by the coexpression of VP40 and glycoprotein GP (1, 17, 25), which is recruited into lipid raft microdomains that serve as an assembly platform for a number of enveloped viruses (1, 19, 20, 25, 26, 28, 37).
The role of VP40 in budding is consistent with the presence of late domain sequences at its N terminus, which have been reported to interact with cellular factors (10, 16, 21, 32, 36), such as Tsg101. Such factors participate in multiprotein complex formations that drive enveloped virus budding processes (22, 30, 35) as well as the sorting of plasma membrane receptors into multivesicular bodies at late endosomal membranes (12). The involvement of late endosomal membranes in filovirus assembly was also demonstrated by the localization of Marburg virus VP40 to the late endosome, the presence of endosomal markers in virus particles (13), and vesicular transport of VP40 from late endosomes to the plasma membrane (14), where assembly and budding take place (8).
VP40 forms monomers in solution that are composed of two structurally related beta sandwich domains, which associate in a metastable conformation (4). In vitro, the monomeric conformation of VP40 can be switched to an oligomeric ring-like structure by several means, including urea treatment (29), in agreement with the metastability of VP40. We have further shown that the N-terminal domain of VP40 is sufficient for oligomerization, while the C-terminal domain is necessary for membrane interaction (27). Single-particle electron microscopy analysis suggested that the C-terminal membrane binding domain is flexibly linked to the ring-like structure formed by the N-terminal domain (29). A recent report also suggested that oligomeric VP40 is enriched in lipid rafts (26), consistent with the role of lipid microdomains in virus assembly (19, 20, 28, 37). In addition, these data may indirectly implicate membrane targeting in the conversion of monomers to oligomers, as suggested by in vitro studies (29).
We previously reported that oligomeric ring-like VP40 structures consist of either hexamers or octamers in vitro and that octamer formation is critically dependent on RNA binding (9, 33). Accordingly, the crystal structure of the octameric ring-like structure formed by the N-terminal domain revealed that the Ebola virus matrix protein is a sequence-specific single-stranded RNA (ssRNA) binding protein (9). The central pore of octameric VP40 binds eight copies of a short RNA with the sequence 5'-UGA-3' at the dimer-dimer interface (9). As suggested by the crystal structure and confirmed biochemically, RNA binding plays an important structural role, as no octamers could be observed in the absence of RNA (33). The structure also suggested that two conserved residues, Phe125 and Arg134, are the most important residues for RNA binding (9). In addition, RNA binding confers sodium dodecyl sulfate (SDS) resistance to the oligomer composed of either the N-terminal domain alone or full-length wild-type VP40 when separated under nonboiling conditions by SDS-polyacrylamide gel electrophoresis (PAGE) (9, 33).
Here we confirm that octamer formation is also driven by RNA binding in HEK 293 cells expressing VP40. Interestingly, octamers are not necessary for the formation of VP40-containing VLPs, although they appear to be critical for Ebola virus replication, as a single mutation that blocks RNA binding inhibits the formation of infectious particles. This report shows for the first time that the sequence-specific RNA binding of a matrix protein may be essential for the virus life cycle.
MATERIALS AND METHODS
Cell lines. HEK 293, HUH-T7, and Vero E6 cells were cultured at 37°C in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal calf serum (FCS). BSR T7/5 cells (BHK 21 cells stably expressing T7 RNA polymerase) were maintained in Glasgow medium supplemented with 10% newborn calf serum.
Molecular cloning. VP40 was subcloned from plasmid pTM-VP40 (14) into vector pCAGGS by using restriction endonucleases EcoRI and XhoI. The VP40-FA, VP40-RA, and VP40-RFA mutants were constructed by using a QuikChange site-directed mutagenesis kit (Stratagene) according to the supplier's instructions with pTM-VP40 serving as a template. For constructing the VP40-FA mutant, primers 1271 (5'-CAC TAT CAC CCA TGC CGG CAA GGC AAC-3') and 1272 (5'-GTT GCC TTG CCG GCA TGG GTG ATA GTG-3') were used; the mutated nucleotides are underlined. To construct the VP40-RA mutant, primers 1273 (5'-CAA TCC ACT TGT CGC AGT CAA TCG GCT GGG-3') and 1274 (5'-CCC AGC CGA TTG ACT GCG ACA AGT GGA TTG-3') were used. The VP40-RFA mutant was constructed by using primers 1273 and 1274 together with pTM-VP40-FA as a template. All constructs then were subcloned into vector pCAGGS by using EcoRI and XhoI and were verified by sequencing.
Transfection of cells. HEK 293 or HUH-T7 cells were grown in six-well plates (7 cm2) to a confluence of 70%. Transfection was performed with Lipofectamine Plus (Invitrogen) according to the manufacturer's instructions. Briefly, 1 μg of DNA was diluted in 100 μl of serum-free DMEM, and 6 μl of Lipofectamine Plus solution was added. In parallel, 4 μl of Lipofectamine was diluted in 100 μl of serum-free DMEM. After 15 min of incubation, the two solutions were combined. After an additional 15 min of incubation, the transfection mixture was added to the cells, which had been washed twice with serum-free DMEM and then brought to a medium volume of 800 μl. After 3 h of incubation at 37°C, 2.5 ml of DMEM supplemented with 10% FCS was added to the cells. For harvesting, the cells were placed on ice, washed twice with ice-cold phosphate-buffered saline (PBS; 0.8% [wt/vol] NaCl, 0.02% [wt/vol] KCl, 0.115% [wt/vol] Na2HPO4, 0.02% [wt/vol] KH2PO4), and then scraped into 100 μl of PBS.
Release of VLPs. At 48 h after cell transfection, the supernatant was harvested and centrifuged at 5,000 x g and 4°C for 10 min to remove cellular debris. The supernatant then was loaded onto a 20% sucrose cushion and centrifuged at 250,000 x g and 4°C for 3 h. The pellet was resuspended in PBS and either subjected to immunoelectron microscopy analysis as described previously (14) or examined for the presence of SDS-resistant VP40 octamers as described below.
Detection of SDS-resistant VP40 octamers. Harvested cells or VLPs were lysed with 1% Triton X-100 for 5 min on ice. Cell lysates were cleared by centrifugation at 830 x g and 4°C for 5 min. The postnuclear supernatant or the lysed VLPs were mixed at room temperature with reducing SDS loading buffer (40% glycerol, 20% ?-mercaptoethanol, 0.5% [wt/vol] SDS, 20% 1 M Tris-HCl [pH 6.8], 0.2% [wt/vol] bromophenol blue) and separated by noncontinous gradient (8 to 12%) SDS-PAGE. Subsequently, Western blotting was performed as described previously (2). Ebola virus VP40 was detected with mouse monoclonal antibody 2C4 (18) at a dilution of 1:100. As a secondary antibody, a horseradish peroxidase-conjugated goat anti-mouse antibody (Dako) at a dilution of 1:20,000 was used. Bound secondary antibody was detected by using a SuperSignal Ultra kit (Pierce).
Detection of RNA bound to VP40. Cells were transfected as described above, with the exception that no additional medium was added at 3 h posttransfection. At 6 h posttransfection, the cells were labeled with 25 μCi of [3H]uridine (Amersham) per dish. At 24 h posttransfection, the cells were placed on ice, washed twice with ice-cold PBS, and lysed with 1% Triton X-100 for 20 min at 37°C in the presence or in the absence of 10 μg of RNase A (Qiagen). Subsequently, nuclei were pelleted at 830 x g, and the supernatant was subjected to SDS-PAGE and Western blotting as described above. The blots were dried overnight, and radioactivity was detected by exposure to BioImager plates for 4 weeks.
Recombinant VP40. The expression, purification, chemical cross-linking, and analysis of SDS resistance of recombinant VP40 were performed as described previously (4, 9, 33).
Indirect immunofluorescence. Cells were transfected as described above and fixed with 4% paraformaldehyde at 24 h posttransfection. Immunofluorescence analysis was performed as described previously (3) with mouse monoclonal antibody 2C4 at a dilution of 1:10 and a rhodamine-coupled goat anti-mouse secondary antibody (Dianova) at a dilution of 1:100. Nuclei were stained with 4',6'-diamidino-2-phenylindole.
Construction of plasmids and recovery of recombinant Ebola viruses. Full-length antigenomic cDNAs of Ebola virus and the protocol used for virus rescue were described elsewhere (34). VP40 mutations R125A and F134A were introduced by site-directed mutagenesis of intermediate plasmid pKSN4 (34). Full-length antigenomic cDNAs of Ebola virus mutants then were generated and analyzed for the presence of mutations by sequencing of the genome. The recovery of recombinant Ebola viruses was performed in three independent experiments.
RNA extraction and RT-PCR of recovered Ebola viruses. The recovered viruses were passaged three times in Vero E6 cells, and culture supernatants were harvested at 6 days postinfection and cleared by low-speed centrifugation. Viral RNA was extracted from the recovered viruses by using an RNeasy kit (Qiagen) according to the manufacturer's instructions. Reverse transcription (RT) and PCR amplification were carried out with forward (5'-AAA AAC CTA CCT CGG CTG AGA GAG T-3') and reverse (5'-AAG CAT GCA GGC AAT TTG AGG ATA AG-3') primers by using a Titan one-tube RT-PCR kit (Roche). To confirm the presence of the introduced mutations, the VP40 gene was sequenced.
Kinetics of multicycle growth of recombinant viruses. Monolayers of Vero E6 cells were infected at a multiplicity of infection of 0.001 PFU per cell, and aliquots of culture media were harvested at 24-h intervals for 6 days. Samples were analyzed by plaque assays in duplicate. Briefly, Vero E6 cells were infected with serial dilutions of cell-free supernatants and 1.6% carboxymethyl cellulose (BDH Laboratory Supplies) in DMEM supplemented with 2.5% FCS. Infectious foci were detected after 5 days of culturing by incubation with mouse anti-Ebola virus NP monoclonal antibodies followed by horseradish peroxidase-conjugated goat polyclonal anti-mouse immunoglobulin G (Sigma-Aldrich, Saint Quentin Fallavier, France) and TrueBlue peroxidase substrate (Kirkegaard & Perry Laboratories). Virus titers were expressed as focus-forming units per milliliter of culture medium. Culture medium and cell lysates collected 6 days postinfection were also analyzed by Western blotting with rabbit polyclonal antibodies directed against VP40 or VP24.
RESULTS
Specific RNA binding of VP40 expressed in HEK 293 cells. The crystal structure of octameric VP40 showed that the N-terminal domain of VP40 binds a short ssRNA with the sequence 5'-UGA-3'. Analysis of the interactions indicated that Arg134 and Phe125 contributed most to the coordination of the ssRNA fragment (9). In addition, both residues were exposed in the monomeric conformation of VP40 (Fig. 1). Based on these findings, we engineered three VP40 mutants, an Arg134-to-Ala mutant (VP40-RA), a Phe125-to-Ala mutant (VP40-FA), and a mutant with a double mutation of Phe125 and Arg134 to Ala (VP40-RFA), and analyzed them in a mammalian expression system.
We previously demonstrated that VP40 octamers show SDS resistance on SDS-PAGE when loaded under nonboiling conditions and that both resistance and octamer formation are dependent on the presence of nucleic acids (33). In order to test whether the mammalian expression of wild-type VP40 leads to the incorporation of RNA, we labeled cellular RNA with [3H]uridine. Analysis of VP40 expression by Western blotting showed a band for the monomer (37 kDa) and a band which migrated above the 220-kDa marker protein (Fig. 2A, lane 3), indicating SDS resistance and therefore the octameric form of VP40. Although we could detect a very faint band corresponding to the oligomer for VP40-FA (Fig. 2A), no octamers were detectable for VP40-RA and VP40-RFA. Monomeric mutant VP40, however, was expressed (Fig. 2A). In accordance with octamer formation, the amount of monomeric wild-type VP40 detected was smaller than that of the mutants (Fig. 2A).
Next, the same blot was exposed to a BioImager plate in order to detect radioactively labeled bands derived from VP40 expression in [3H]uridine-labeled mammalian cells. This experiment revealed a major radioactive band migrating above the 220-kDa marker protein and corresponding to the SDS-resistant high-molecular-mass form of VP40 shown in Fig. 2A (Fig. 2B, lane 3). Therefore, octameric VP40 also binds RNA upon expression in HEK 293 cells. Two additional radioactively labeled bands could be observed approximately between the 97- and 66-kDa marker proteins in all experiments. These bands appeared at the same intensities in all lanes, including the negative control. These bands therefore serve as an internal control and are consistent with the fact that the same amounts of labeled material were loaded in the lanes. As these additional bands were unrelated to VP40 octamerization, we suggest that they might represent abundant cellular RNA(s).
We next tested whether RNase A treatment of total cell lysates would affect VP40 octamerization. Following treatment, there was a decrease in the amount of detectable SDS-resistant wild-type VP40 octamers (Fig. 2C, lane 2). Similarly, an increase in the total amount of monomers could be detected (Fig. 2C, lanes 1 and 2). As indicated in Fig. 2A, VP40-FA still formed octamers at low levels, which were further reduced upon RNase A treatment (Fig. 2C, lanes 3 and 4). Only monomers were detected for VP40-RA and VP40-RFA (Fig. 2C, lanes 5 to 8).
In order to better understand the decrease in VP40 SDS resistance in the presence of RNase A, we performed in vitro SDS resistance experiments with long and short RNA molecules. We previously showed that monomeric full-length VP40 can be converted to SDS-resistant octamers in the presence of RNA and urea (9). Here we incubated VP40 with only urea, with urea in the presence of Escherichia coli RNA, or with urea in the presence of a short synthetic ssRNA with the sequence 5'-UGA-3', which is present in the crystal structure of octameric VP40 (9). Then we tested the oligomerization state of VP40 by SDS resistance and chemical cross-linking analyses. These analyses showed that VP40 alone did not produce SDS-resistant oligomers, as only bands corresponding to monomers were visible (Fig. 2D, lane 2). Urea treatment of VP40 in the absence of RNA led to the formation of high-molecular-mass aggregates that barely entered the gel, as detected by chemical cross-linking, indicating nonspecific aggregation (Fig. 2D, lane 3). VP40 treated with urea in the presence of E. coli nucleic acids showed a new SDS-resistant band that migrated above the 250-kDa marker protein, as shown previously (9) (Fig. 2D, lane 4). This oligomer could also be cross-linked, resulting in a ladder of bands, with the major species migrating more slowly than the SDS-resistant oligomer, as shown previously for VP40 octamers (33) (Fig. 2D, lane 5). Reconstitution of the octamer in the presence of short synthetic ssRNA with the sequence 5'-UGA-3', the minimal binding sequence detected in the crystal structure, did not result in SDS-resistant octamers. However, octamer formation could be shown by chemical cross-linking and was represented by a band migrating at the same position as VP40 cross-linked in the presence of E. coli RNA (Fig. 2D, lanes 6 and 7). These data are consistent with the reduction of SDS-resistant VP40 by RNase treatment and indicate that the proposed SDS resistance of octameric VP40 depends on the presence of longer ssRNA molecules. To further confirm that the Arg mutants no longer bound RNA, we expressed and purified recombinant VP40-RFA, which did not bind RNA; therefore, no octamers could be detected by SDS resistance and chemical cross-linking analyses in vitro (data not shown).
VLP formation induced by wild-type or mutant VP40 expression. We next tested whether the mutations affected VP40 release into cell culture supernatants and the formation of VLPs. These analyses showed that all mutant VP40 constructs induced the release of equal amounts of VP40 into supernatants upon expression in HEK 293 cells (Fig. 3A). Negative staining electron microscopy revealed filamentous particles produced by wild-type VP40 and mutant VP40; these particles showed similar morphologies and no significant difference in size. Particle size analysis revealed diameters (mean and standard deviation) of 41 ± 10 nm for wild-type VP40, 41 ± 9 nm for VP40-FA, 44 ± 10 for VP40-RA, and 40 ± 7 nm for VP40-RFA VLPs (Fig. 3B). Wild-type VLPs contained SDS-resistant VP40 octamers similar to those detected in cells (Fig. 3C, lanes 1 and 2) and reported previously (26). However, VLPs generated from VP40-RA did not contain VP40 octamers (Fig. 3C, lane 3), consistent with the absence of octamers in cells expressing VP40-RA (Fig. 3C, lane 4). In addition, VP40-RA VLPs contained dimers of VP40 (Fig. 3C, lane 3) similar to those detected in Ebola virus particles (9).
Cellular localization of wild-type or mutant VP40. Indirect immunofluorescence analysis indicated that the intracellular localization of the three mutant constructs differed from that of wild-type VP40, although the transfection efficiencies (20% for all constructs) and the amounts of VP40 expressed inside cells and released into cell culture supernatants were similar in all experiments, as judged by Western blot analysis (data not shown). While wild-type VP40 was distributed mainly in small patches along the plasma membrane (Fig. 4), mutant VP40-FA was detected in larger aggregates along the plasma membrane as well as in the perinuclear region (Fig. 4). In contrast, mutant VP40-RA and double mutant VP40-RFA were found mostly in large aggregates in the perinuclear region and at the plasma membrane. In addition to the larger intracellular aggregates formed by VP40-RA and VP40-RFA, the extent of cytoplasmic VP40 distribution seemed to be reduced compared to that of wild-type VP40 and VP40-FA (Fig. 4).
Effects of VP40 mutations on Ebola virus replication. Finally, we used an established reverse-genetics system for Ebola virus (34) to examine the significance of the VP40 mutations for virus replication. VP40 mutations F125A and R134A were introduced into Ebola virus cDNA in an otherwise identical genomic background, and supernatants from transfected BSR T7/5 cells were passaged in Vero E6 cells. Clear cytopathic effects were observed with constructs containing wild-type VP40 as well as mutant VP40-FA. To demonstrate the recovery of recombinant viruses, viral RNA from culture supernatants derived from the third consecutive passage of virus in Vero E6 cells was assayed by RT-PCR, followed by sequence analysis. Nucleotide sequencing confirmed the presence of the F125A mutation in recovered recombinant viruses. Subsequent analysis of virus replication in multicycle virus growth revealed no significant differences in endpoint titers between wild-type and mutant viruses carrying the VP40-FA mutation. However, the growth curves indicated that the VP40-FA-containing recombinant virus had a slightly increased ability to replicate in Vero E6 cells, as demonstrated by higher virus titers at an early stage of infection (Fig. 5A). For the VP40-FA-containing mutant virus, Western blot analysis of total cell lysates and cell culture supernatants with anti-VP40 and anti-VP24 antibodies revealed that the culture supernatants contained slightly higher levels of VP40 and VP24 (Fig. 5B). In contrast, for the wild-type Ebola virus, slightly higher levels of intracellular VP40 and VP24 were detected (Fig. 5B).
Several attempts to recover recombinant virus containing the VP40 R134A mutation (VP40-RA or VP40-RFA) failed. We therefore suggest that Arg at position 134 has an important role in virus replication. As the same mutant showed a defect in RNA binding upon expression in HEK 293 cells, the rescue experiments strongly indicated that RNA binding, which confers VP40 octamer formation, is important for the life cycle of Ebola virus.
DISCUSSION
The crystal structure of the N-terminal domain of Ebola virus VP40 in complex with ssRNA indicated that specific RNA binding is responsible for the formation of SDS-resistant octameric VP40 (9, 33). Here we established that octameric VP40 binds cellular RNA labeled with [3H]uridine when expressed in HEK 293 cells. Detection of SDS-resistant VP40 is sensitive to RNase treatment, in agreement with our in vitro data showing that VP40 octamer formation with short ssRNA molecules with the sequence 5'-UGA-3' is detectable only by chemical cross-linking and no longer by SDS resistance. As the 5' and 3' ends of the bound ssRNA molecules are freely accessible in the crystal structure, it is conceivable that only longer RNA molecules contribute to SDS stability (9). Furthermore, in the octamer, the central pore containing the RNA binding site carries a mostly basic charge (9), which might allow for additional nonspecific RNA binding that could further stabilize the octamer and thus contribute to its stability in the presence of SDS. Therefore, SDS resistance is only an indication of octamer formation and cannot be used as a quantitative measurement of octamer formation.
The atomic details of the octameric crystal structure indicated an important role for Arg134 and Phe125 in RNA binding (Fig. 1); this finding was confirmed by our mutational analysis, which showed reduced RNA binding for VP40-FA and no RNA binding for VP40-RA and VP40-RFA. The conclusion that the abolishment of RNA binding leads to the blockage of octamer formation is also in agreement with the described structural role of ssRNA in VP40 octamer formation (9, 33). The complete absence of RNA binding and thus of octamer formation in the presence of the Arg134 mutation was also confirmed by analysis of recombinant VP40-RFA in vitro.
The cellular localization of the VP40-FA mutant was still similar to that of wild-type VP40 (1, 10, 21, 26, 31). However, the intracellular aggregates formed were larger, and they were even more pronounced for VP40-RA and VP40-RFA. Although the differences in localization might be due to the presence or absence of VP40 octamers, it is possible that the mutations themselves interfere with the active transport of VP40 (14). Furthermore, the mutations, which are both located in or close to a basic patch at the interface of the N- and C-terminal domains of monomeric VP40 (Fig. 1A) (4), might interfere with a proposed switch from the monomeric form to an oligomeric form (29) or a polymeric form of VP40. Therefore, mutation of Phe125 to Ala and, to an even greater extent, of Arg134 to Ala might lead to increased monomer destabilization, resulting in larger intracellular aggregates containing VP40 oligomers or polymers and a shorter half-life of cytoplasmic monomers, compared to the properties of wild-type VP40.
It was previously shown that VP40 expression as well as VP40 and GP coexpression in mammalian cells induces the release of filamentous VLPs (1, 11, 17, 25, 31). Here we show that the VLPs produced by wild-type VP40 expression as well as the expression of mutant VP40 have similar filamentous morphologies, with particle diameters ranging from 40 to 50 nm. However, the VLP diameters reported here are smaller than those described for released VLPs containing VP40 alone (65 nm) (25) but are similar to those of filamentous particles inside cells expressing VP40 (45 nm) (25). In addition, VLPs produced by VP40 and GP coexpression had diameters ranging from 50 to 70 nm (1) or an outer diameter of 80 nm (25); these data indicate that the diameters of VLPs can vary greatly. The final conserved 80-nm diameter of infectious viruses (5) is therefore most likely driven by the presence of the nucleocapsid or other unknown viral or cellular factors. Our data indicate that RNA binding and octamer formation, which are completely abolished in VP40-RA and VP40-RFA, are not necessary for VLP formation, although SDS-resistant octamers are present in VLPs produced by wild-type VP40, consistent with previously published data (26).
When the VP40 mutations were introduced into the Ebola virus genome, only recombinant wild-type virus and virus carrying the VP40-FA mutation could be rescued (34). These data strongly indicate that VP40 RNA binding and octamer formation might be important for the life cycle of Ebola virus, as VP40-RA showed a complete block in RNA binding and thus octamer formation. In contrast, when expressed alone, the second mutation, VP40-FA, still resulted in reduced RNA binding and octamer formation, which seemed to be sufficient for virus replication. Interestingly, when growth curves for wild-type and recombinant viruses carrying the VP40-FA mutation were compared, the VP40-FA-carrying virus showed a slight increase in replication kinetics. These data suggest that while RNA binding seems to be important for virus replication, the VP40-FA-carrying mutant virus might have a slight advantage in assembly over the wild-type virus. We propose that this effect is not dependent on VP40 RNA binding but rather is due to a potential destabilizing effect of the F125A mutation, as discussed above, which might facilitate the proposed transition from a monomeric form to an assembly-activated form of VP40.
In summary, our data indicate that the octamerization deficient mutants is still active in VLP formation, a finding which implies that they are competent to facilitate virion assembly and promote the egress of newly produced virus particles from the plasma membrane. Therefore, octameric VP40 must exert another, as-yet-unknown function during Ebola virus replication, a function which we propose to be important for the virus life cycle.
ACKNOWLEDGMENTS
We thank M. Rossi for expert technical help and Allison Groseth for critical reading of the manuscript. All experiments involving Ebola virus were carried out in biosafety level 4 laboratories at the Institute of Virology in Marburg, Germany, and at the Jean Merieux P4 Center in Lyon, France.
This work was supported by EMBL (W.W.), by INSERM (V.V.), and by funds from Deutsche Forschungsgemeinschaft SFB 593 Marburg (to W.W., S.B., and V.V.). T.H. was supported by a scholarship from Verband der Chemischen Industrie.
REFERENCES
Bavari, S., C. M. Bosio, E. Wiegand, G. Ruthel, A. B. Will, T. W. Geisbert, M. Hevey, C. Schmaljohn, A. Schmaljohn, and M. J. Aman. 2002. Lipid raft microdomains: a gateway for compartmentalized trafficking of Ebola and Marburg viruses. J. Exp. Med. 195:593-602.
Becker, S., H. D. Klenk, and E. Muhlberger. 1996. Intracellular transport and processing of the Marburg virus surface protein in vertebrate and insect cells. Virology 225:145-155.
Becker, S., C. Rinne, U. Hofsass, H. D. Klenk, and E. Muhlberger. 1998. Interactions of Marburg virus nucleocapsid proteins. Virology 249:406-417.
Dessen, A., V. Volchkov, O. Dolnik, H. D. Klenk, and W. Weissenhorn. 2000. Crystal structure of the matrix protein VP40 from Ebola virus. EMBO J. 19:4228-4236.
Ellis, D. S. 1987. The Filoviridae, p. 313-321. In M. V. Nermut and A. C. Steven (ed.), Animal virus structure, 3rd ed. Elsevier Science Publishing, Inc., New York, N.Y.
Feldmann, H., V. E. Volchkov, V. A. Volchkova, U. Stroher, and H. D. Klenk. 2001. Biosynthesis and role of filoviral glycoproteins. J. Gen. Virol. 82:2839-2848.
Garoff, H., R. Hewson, and D. J. Opstelten. 1998. Virus maturation by budding. Microbiol. Mol. Biol. Rev. 62:1171-1190.
Geisbert, T. W., and P. B. Jahrling. 1995. Differentiation of filoviruses by electron microscopy. Virus Res. 39:129-150.
Gomis-Rüth, F. X., A. Dessen, J. Timmins, A. Bracher, L. Kolesnikova, S. Becker, H. D. Klenk, and W. Weissenhorn. 2003. The matrix protein VP40 from Ebola virus octamerizes into pore-like structures with specific RNA binding properties. Structure 11:423-433.
Harty, R. N., M. E. Brown, G. Wang, J. Huibregtse, and F. P. Hayes. 2000. A PPxY motif within the VP40 protein of Ebola virus interacts physically and functionally with a ubiquitin ligase: implications for filovirus budding. Proc. Natl. Acad. Sci. USA 97:13871-13876.
Jasenosky, L. D., G. Neumann, I. Lukashevich, and Y. Kawaoka. 2001. Ebola virus VP40-induced particle formation and association with the lipid bilayer. J. Virol. 75:5205-5214.
Katzmann, D. J., G. Odorizzi, and S. D. Emr. 2002. Receptor downregulation and multivesicular-body sorting. Nat. Rev. Mol. Cell. Biol. 3:893-905.
Kolesnikova, L., H. Bugany, H. D. Klenk, and S. Becker. 2002. VP40, the matrix protein of Marburg virus, is associated with membranes of the late endosomal compartment. J. Virol. 76:1825-1838.
Kolesnikova, L., S. Bamberg B. Berghofer, and S. Becker. 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.
Kraulis, P. 1991. MOLSCRIPT: a program to produce both detailed and schematic plots of protein structures. J. Appl. Crystallogr. 24:924-950.
Licata, J. M., M. Simpson-Holley, N. T. Wright, Z. Han, J. Paragas, and R. N. Harty. 2003. Overlapping motifs (PTAP and PPEY) within the Ebola virus VP40 protein function independently as late budding domains: involvement of host proteins TSG101 and VPS-4. J. Virol. 77:1812-1819.
Licata, J. M., R. F. Johnson, Z. Han, and R. N. Harty. 2004. Contribution of Ebola virus glycoprotein, nucleoprotein, and VP24 to budding of VP40 virus-like particles. J. Virol. 78:7344-7351.
Lucht, A., R. Grunow, P. Moller, H. Feldmann, and S. Becker. 2003. Development, characterization and use of monoclonal VP40-antibodies for the detection of Ebola virus. J. Virol. Methods 111:21-28.
Manes, S., G. del Real, R. A. Lacalle, P. Lucas, C. Gomez-Mouton, S. Sanchez-Palomino, R. Delgado, J. Alcami, E Mira, and A. C. Martinez. 2000. Membrane raft microdomains mediate lateral assemblies required for HIV-1 infection. EMBO Rep. 1:190-196.
Manie, S. N., S. Debreyne, S. Vincent, and D. Gerlier. 2000. Measles virus structural components are enriched into lipid raft microdomains: a potential cellular location for virus assembly. J. Virol. 74:305-311.
Martin-Serrano, J., T. Zang, and P. D. Bieniasz. 2001. HIV-1 and Ebola virus encode small peptide motifs that recruit Tsg101 to sites of particle assembly to facilitate egress. Nat. Med. 7:1313-1319.
Martin-Serrano, J., A. Yarovoy, D. Perez-Caballero, and P. D. Bieniasz. 2003. Divergent retroviral late-budding domains recruit vacuolar protein sorting factors by using alternative adaptor proteins. Proc. Natl. Acad. Sci. USA 100:12414-12419.
Merritt, E. A., and D. J. Bacon. 1997. Raster3D photorealistic graphics. Methods Enzymol. 277:505-524.
Nicholls, A., K. A. Sharp, and B. Honig. 1991. Protein folding and association: insights from the interfacial and thermodynamic properties of hydrocarbons. Proteins 11:281-296.
Noda, T., H. Sagara, E. Suzuki, A. Takada, H. Kida, and Y. Kawaoka. 2002. Ebola virus VP40 drives the formation of virus-like filamentous particles along with GP. J. Virol. 76:4855-4865.
Panchal, R. G., G. Ruthel, T. A. Kenny, G. H. Kallstrom, D. Lane, S. S. Badie, L. Li, S. Bavari, and M. J. Aman. 2003. In vivo oligomerization and raft localization of Ebola virus protein VP40 during vesicular budding. Proc. Natl. Acad. Sci. USA 100:15936-15941.
Ruigrok, R. W., G. Schoehn, A. Dessen, E. Forest, V. Volchkov, O. Dolnik, H. D. Klenk, and W. Weissenhorn. 2000. Structural characterization and membrane binding properties of the matrix protein VP40 of Ebola virus. J. Mol. Biol. 300:103-112.
Scheiffele, P., A. Rietveld, T. Wilk, and K. Simons. 1999. Influenza viruses select ordered lipid domains during budding from the plasma membrane. J. Biol. Chem. 274:2038-2044.
Scianimanico, S., G. Schoehn, J. Timmins, R. H. Ruigrok, H. D. Klenk, and W. Weissenhorn. 2000. Membrane association induces a conformational change in the Ebola virus matrix protein. EMBO J. 19:6732-6741.
Strack, B., A. Calistri, S. Craig, E. Popova, and H. G. G?ttlinger. 2003. AIP1/ALIX is a binding partner for HIV-1 p6 and EIAV p9 functioning in virus budding. Cell 114:689-699.
Timmins, J., S. Scianimanico, G. Schoehn, and W. Weissenhorn. 2001. Vesicular release of Ebola virus matrix protein VP40. Virology 283:1-6.
Timmins, J., G. Schoehn, S. Ricard-Blum, S. Scianimanico, T. Vernet, R. W. Ruigrok, and W. Weissenhorn. 2003. Ebola virus matrix protein VP40 interaction with human cellular factors Tsg101 and Nedd4. J. Mol. Biol. 326:493-502.
Timmins, J., G. Schoehn, C. Kohlhaas, H. D. Klenk, R. W. Ruigrok, and W. Weissenhorn. 2003. Oligomerization and polymerization of the filovirus matrix protein VP40. Virology 312:359-368.
Volchkov, V. E., V. A. Volchkova, E. Muhlberger, L. V. Kolesnikova, M. Weik, O. Dolnik, and H. D. Klenk. 2001. Recovery of infectious Ebola virus from complementary DNA: RNA editing of the GP gene and viral cytotoxicity. Science 291:1965-1969.
von Schwedler, U. K., M. Stuchell, B. Muller, D. M. Ward, H. Y. Chung, E. Morita, H. E. Wang, T. Davis, G. P. He, D. M. Cimbora, A. Scott, H. G. Krausslich, J. Kaplan, S. G. Morham, and W. I. Sundquist. 2003. The protein network of HIV budding. Cell 114:701-713.
Yasuda, J., M. Nakao, Y. Kawaoka, and H. Shida. 2003. Nedd4 regulates egress of Ebola virus-like particles from host cells. J. Virol. 77:9987-9992.
Zhang, J., A. Pekosz, and R. A. Lamb. 2000. Influenza virus assembly and lipid raft microdomains: a role for the cytoplasmic tails of the spike glycoproteins. J. Virol. 74:4634-4644.(Thomas Hoenen, Viktor Vol)