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Positive-strand RNA viruses induce modifications of cytoplasmic membranes to form replication complexes. For coronaviruses, replicase nonstructural protein 4 (nsp4) has been proposed to function in the formation and organization of replication complexes. Murine hepatitis virus (MHV) nsp4 is glycosylated at residues Asn176 (N176) and N237 during plasmid expression of nsp4 in cells. To test if MHV nsp4 residues N176 and N237 are glycosylated during virus replication and to determine the effects of N176 and N237 on nsp4 function and MHV replication, alanine substitutions of nsp4 N176, N237, or both were engineered into the MHV-A59 genome. The N176A, N237A, and N176A/N237A mutant viruses were viable, and N176 and N237 were glycosylated during infection of wild-type (wt) and mutant viruses. The nsp4 glycosylation mutants exhibited impaired virus growth and RNA synthesis, with the N237A and N176A/N237A mutant viruses demonstrating more profound defects in virus growth and RNA synthesis. Electron microscopic analysis of ultrastructure from infected cells demonstrated that the nsp4 mutants had aberrant morphology of virus-induced double-membrane vesicles (DMVs) compared to those infected with wt virus. The degree of altered DMV morphology directly correlated with the extent of impairment in viral RNA synthesis and virus growth of the nsp4 mutant viruses. The results indicate that nsp4 plays a critical role in the organization and stability of DMVs. The results also support the conclusion that the structure of DMVs is essential for efficient RNA synthesis and optimal replication of coronaviruses.
Positive-strand RNA viruses rely on host intracellular membranes to form replication complexes, defined as sites of viral RNA synthesis (11, 34, 40-42). These virus-induced membrane modifications are crucial for creating an environment that supports viral RNA synthesis, as well as protecting newly synthesized viral RNA. For many positive-strand RNA viruses, specific replicase proteins, often containing multiple hydrophobic domains, have been implicated in targeting to and modifying host membranes, ultimately leading to the formation of replication complexes.
The coronavirus murine hepatitis virus (MHV) is an enveloped, positive-strand RNA virus that contains a 31.4-kb genome, consisting of seven open reading frames (ORFs). ORF1 encodes the replicase/transcriptase polyprotein, while ORFs 2 to 7 encode structural and accessory proteins. ORF1 comprises approximately two-thirds of the genome and is translated as either polyprotein 1a (pp1a) or, due to a −1 ribosomal frameshift, pp1ab (3, 5, 6, 28, 34). pp1a and pp1ab are processed by three virus-encoded proteases to yield 16 nonstructural proteins (nsp1 to 16) (Fig. (Fig.1A)1A) (1, 3, 13, 21, 32, 48). Analysis of nsp3, nsp4, and nsp6 amino acid sequences and in vitro biochemical studies have shown that these three nsp's all have transmembrane domains that are likely important for virus-induced membrane modifications (2, 23, 28). MHV nsp4 is processed by papain-like protease 2 (PLP2) at its amino terminus, resulting in an nsp4-to-10 precursor, and after this initial processing event, nsp5 (3Clpro) mediates processing at the carboxy terminus of nsp4 (15, 17, 21, 22, 24). The predicted molecular mass of nsp4 is 56 kDa, but it is detected as a 44-kDa protein by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (22, 31).
All tested coronavirus nsp's localize to replication complexes that are located on virus-induced double-membrane vesicles (DMVs), and nsp4 has been proposed to play roles in the formation, organization, and function of these virus replication complexes (15, 38). nsp4 has been shown to associate with membrane fractions of infected cells and is resistant to membrane extraction following Triton X-114 treatment, indicating that nsp4 is an integral membrane protein (15). Bioinformatics of the MHV nsp4 amino acid sequence predicted that nsp4 has four transmembrane domains (TM1 to 4). MHV nsp4 has also been shown to be required for rescue of infectious virus (45), as have TM1 to 3, but TM4 is dispensable for recovery of infectious virus in culture. Charge-to-alanine substitutions between TM1 and TM2 of nsp4 result in viruses with phenotypes ranging from nonrecoverable to viruses that exhibit reduced virus growth, RNA synthesis, and protein processing (45).
Analysis of nsp4 from multiple coronaviruses across all coronavirus groups predicts N-linked glycosylation sites for all tested nsp4 sequences. The glycosylation sites, or sequons, Asn-X-Ser, Asn-X-Thr, and rarely Asn-X-Cys, are amino acid sequences that are recognized for glycosylation of the Asn (N) residue. Even though coronaviruses contain putative glycosylation sites within nsp4, there is little conservation of these sites between groups. Group 2a coronaviruses, such as MHV and human coronavirus HCoV-OC43, have two conserved putative N-linked glycosylation sites, N176 and N237 (Fig. (Fig.1B),1B), while the group 2b severe acute respiratory syndrome coronavirus (SARS-CoV) and group 3 avian infectious bronchitis virus (IBV), have different putative glycosylation sites, N131 and N48, respectively (29, 35). Although the glycosylation of nsp4 from group 1 coronaviruses has not been investigated, residues N176 and N237 of MHV nsp4, N131 of SARS-CoV, and N48 of IBV nsp4 have been shown to be glycosylated when nsp4 is plasmid expressed in cells or when nsp4 is expressed from nonnative locations in the coronavirus genome (10, 29, 35). Clementz et al. reported that N176 of MHV nsp4 is not required for virus replication and that an N176A mutant virus grows identically to wild-type (wt) virus (10). In that study, the N176A mutant virus-expressed nsp4 migrated faster than wt nsp4 as determined by SDS-PAGE, consistent with altered protein modification, such as loss of glycosylation. However, this was not further investigated in the study. In contrast, N237A and N176A/N237A mutant viruses could not be recovered.
Although these studies have led to an increased understanding of various aspects of nsp4, it remains unknown if N176 and/or N237 is glycosylated during infection and if the putative nsp4 glycosylation sites of MHV or other coronaviruses serve roles in membrane modifications or replication complex formation and function. In this study, we tested the glycosylation status of MHV nsp4, expressed from its native genomic location, and the role of nsp4 glycosylation sites on virus growth, viral RNA synthesis, nsp4 localization, and replication complex morphology by engineering and recovering nsp4 mutants with alanine substitutions at N176 (N176A), N237 (N237A), or both (N176A/N237A). We show that virus-expressed nsp4 is glycosylated at both N176 and N237 during infection, that glycosylation at either or both sites is dispensable for virus growth in cell culture, and that alanine substitution of N176, N237, or both results in defects in virus growth and RNA synthesis. Further, we show that loss of nsp4 glycosylation is associated with the presence of aberrant or disrupted DMVs (hereafter referred to as irregular DMVs) and increased prevalence of virus-induced convoluted membranes (CMs). The degree of irregular DMVs and increased CMs from the nsp4 mutant viruses directly correlated with an impairment in viral RNA synthesis and growth. These results demonstrate that nsp4 plays a critical role in the formation, stability, and structure of virus-induced membrane modifications. Finally, the results also support the conclusion that the physical structure and stability of DMVs are essential for efficient RNA synthesis and/or protection of viral RNAs and optimal replication of coronaviruses.
Recombinant wt MHV strain A59 (GenBank accession number AY910861) was used as the wt control for all experiments. Delayed brain tumor (DBT) cells expressing the MHV receptor carcinoembryonic antigen cell adhesion molecule-1 (9, 20, 47) and baby hamster kidney cells expressing the MHV receptor (BHK-MHVR) (8, 9, 47) were grown in Dulbecco's modified Eagle medium (Gibco) supplemented with 10% fetal calf serum for all experiments. Medium for BHK-MHVR cells was supplemented with G418 (Mediatech) at 0.8 mg/ml to maintain selection for cells expressing the MHVR. Rabbit polyclonal antibodies were used in biochemical experiments and have been described previously. Antibodies include anti-nsp4 (VU158) (45), anti-nsp8 (VU123) (4), and anti-M (J.1.3) (7).
For introduction of asparagine-to-alanine substitutions in the nsp4 coding sequence (ORF1a nucleotides 8721 to 10208), PCR was performed using the MHV-A59 infectious clone fragment B (pCR-XL-pSMART B) as a template. Fragment B of the MHV-A59 clone contains MHV ORF1a nsp4 nucleotides 8721 to 9555 (47). Asparagine-to-alanine codon changes were introduced using the ExSite/QuikChange mutagenesis kit (Stratagene) with the primers listed in Table Table1.1. Changes to the manufacturer's protocol included the use of Pfu Turbo and Pfu Ultra instead of the ExSite DNA polymerase blend. PCR was performed using the following parameters: initial denaturation at 95°C once for 2 min, denaturation at 95°C for 1 min, annealing at various temperatures depending on the primers for 1 min, extension at 72°C for 7 min, and repeating of the denaturing, annealing, and extension steps for a total of 40 cycles. Products were ligated and sequenced across the MHV genome-containing region of fragment B to ensure that PCR amplification did not introduce any unintended mutations. For introduction of both N176A and N237A, restriction endonuclease EcoN I was used to digest both single nsp4 glycosylation mutant plasmids, and ligation was used to introduce both mutations into the same plasmid.
Viruses containing the engineered mutations within nsp4 were produced using the infectious cDNA assembly strategy for MHV-A59 that has previously been described by Yount et al. (47) and modified by Denison et al. (12) and Sparks et al. (45). Briefly, plasmids containing the seven cDNA cassettes that make up the MHV genome were digested using the appropriate restriction enzymes. The correct restriction fragments were gel purified and ligated together overnight at 16°C. The ligated DNA was purified, in vitro transcribed, and electroporated with N gene transcripts into BHK-MHVR cells. The electroporated cells were then laid over a layer of 2.5 × 106 uninfected DBT cells in a 75-cm2 flask and incubated at 37°C. Virus viability was determined by cytopathic effect, in this case syncytium formation, in the electroporated cell culture. Progeny virus in the cell culture medium of electroporated cells (passage 0 [P0]) was passaged onto uninfected DBT cells (P1), and the virus released from cells in the culture medium was designated P1 stock, the titer was determined, and it was used for all experiments.
Total intracellular RNA was harvested from P1-infected cells using TRIzol (Invitrogen) according to the manufacturer's protocol. Extracted RNA was used as a template for reverse transcription (RT)-PCR. RT was performed using Superscript III reverse transcriptase (Invitrogen) and random hexamers (Roche). Primers complementary to genome nucleotides 8486 to 8502 (sense) and 10361 to 10345 (antisense) were then used to amplify the nsp4 coding region by PCR. These PCR products were sequenced to confirm the retention of the engineered mutations and the absence of additional mutations in the nsp4 coding sequence.
For radiolabeling of proteins and immunoprecipitations, cells were grown on 60-mm dishes and infected at a multiplicity of infection (MOI) of 10 PFU/cell with wt or nsp4 glycosylation mutant viruses and incubated at 37°C. At 4 h postinfection (p.i.), medium was aspirated and replaced with medium lacking methionine and cysteine and supplemented with actinomycin D (Act D; Sigma) at a final concentration of 20 μg/ml. At 5 h p.i., cells were radiolabeled with [35S]methionine-cysteine ([35S]Met-Cys) at a concentration of 0.08 mCi/ml. When cells reached ~90% involvement in syncytia, radiolabeled cells were washed once in phosphate-buffered saline (PBS), and then lysed in 1 ml of lysis buffer lacking SDS (1% NP-40, 0.5% sodium deoxycholate, 150 mM NaCl, and 50 mM Tris, pH 8.0). Lysates were then centrifuged at 6,000 × g for 3 min to remove cellular debris and nuclei, and the supernatant was collected. Immunoprecipitations were performed in a final volume of 1 ml, using protein A-Sepharose beads (Sigma), 50 μl of radiolabeled lysate, 1:200 (anti-nsp4) or 1:500 (anti-nsp8) dilutions of polyclonal antisera, and proteinase inhibitor (Roche) in lysis buffer. Immunoprecipitations were then performed as previously described (45). For endoglycosidase H (Endo H) treatment, supernatant was transferred to a new tube after heating at 70°C for 10 min. Endo H (Sigma) was added to the supernatant according to the manufacturer's protocol, and the mixture was incubated at 37°C for 3 h. Proteins were resolved by SDS-PAGE in 4 to 12% polyacrylamide gradient Bis-Tris gels (NuPage; Invitrogen) and analyzed by fluorography. 14C-labeled high-molecular-weight markers (NEB) and a full-range rainbow marker were used as molecular weight standards.
For viral growth determination (12), DBT cells were infected with wt or nsp4 glycosylation mutant viruses at the MOIs indicated. Following a 45-min absorption period at 37°C with periodic swirling, medium was aspirated, and the cells were washed three times in PBS. Prewarmed 37°C medium was then added back to the cells, and the cells were incubated at 37°C. Aliquots of medium were taken from 1 to 30 h p.i., and virus titers were determined by plaque assay as previously described (25).
DBT cells were either mock infected or infected at an MOI of 5 PFU/cell with wt or nsp4 glycosylation mutant viruses in six-well plates. Following a 45-min absorption at 37°C, medium containing virus was removed, and cells were washed twice in PBS. Cells were then incubated in growth medium at 37°C until 30 min prior to labeling, when medium was replaced with fresh medium containing 20 μg/ml Act D. After this 30-min treatment, [3H]uridine was added to a final concentration of 40 μCi/ml, and cells were incubated at 37°C for 2-h intervals from 3 to 15 h p.i. At the end of each labeling period, cells were lysed in lysis buffer (described above), and nuclei were removed by centrifugation at 14,000 × g for 3 min. RNA in 10% of each lysate was precipitated with chilled 5% trichloroacetic acid (TCA) onto glass microfiber filters (Whatman), washed twice in fresh 5% TCA and twice in 95% ethanol, and dried using vacuum filtration. Radiolabel incorporation was quantitated by liquid scintillation counting.
DBT cells grown on glass coverslips were infected with wt or nsp4 glycosylation mutant viruses at an MOI of 10 PFU/cell. At 6 h p.i., medium was aspirated from cells, and cells were fixed in 100% methanol at −20°C. Cells were rehydrated in PBS for 10 min, blocked in PBS containing 5% bovine serum albumin, and then aspirated. For indirect immunofluorescence, cells were incubated with primary antibody (anti-nsp4, 1:200; anti-M, 1:1,000) in wash solution (PBS containing 1% bovine serum albumin and 0.05% NP-40) for 1 h at room temperature. Cells were washed in wash solution three times for 5 min/wash. Cells were then incubated with secondary antibody (goat anti-rabbit Alexa 488, 1:1,000; goat anti-mouse Alexa 546, 1:1,000; Molecular Probes) for 30 min at room temperature. Cells were washed again three times for 5 min/wash, subjected to a final wash in PBS, and rinsed with distilled water. For direct immunofluorescence, anti-nsp8 was purified using HiTrap rProtein A FF columns (GE Life Sciences) for fast protein liquid chromatography. Anti-nsp8 was directly conjugated using the Alexa Fluor 546 protein labeling kit (Invitrogen) according to the manufacturer's protocol. Cells were incubated with anti-nsp8 at a concentration of 1:500, following the same procedure as above. Coverslips were mounted with Aquapolymount (Polysciences) and visualized using a Zeiss Axiovert 200 microscope with a 40× oil immersion lens. Images were processed and merged using Adobe Photoshop CS3.
DBT cells were mock infected or infected with wt or nsp4 glycosylation mutant viruses at an MOI of 5 PFU/cell in a 60-mm dish and incubated at 37°C. At 6 h p.i., medium was aspirated, and cells were washed once with PBS. The cells were then fixed in 2% glutaraldehyde for 10 min, scraped off the dishes, and centrifuged at 0.5 × g for 3 min. The initial 2% glutaraldehyde was aspirated, fresh 2% glutaraldehyde was added to the fixed cells for 1 h and aspirated, and fresh glutaraldehyde was added to the fixed cells for overnight incubation at 4°C. Cells were washed three times in PBS, transferred to 1% osmium tetroxide in distilled water (diH2O) for 1 h, and washed three times in diH2O. Cells were stained en bloc in 1% aqueous uranyl acetate for 1 h and washed three times in diH2O. Dehydration of cells was carried out gradually using a graded series of ethanol and increasing the times each remained in solution, starting with 30%, followed by 50%, 70%, 95%, and finally absolute ethanol. Propylene oxide was used as a transitional solvent to replace the dehydration solution. Cells were transferred to a 1:1 araldite-propylene oxide mixture for 1 h and then placed in pure araldite in a vacuum oven for another hour to help pull resin through the tissue. Pure resin specimens were then transferred into capsules containing fresh resin and finally placed into an oven overnight to polymerize. Ultra-thin serial sections (50 to 60 nm) from polymerized blocks were obtained using a Leica UCT Ultracut microtome (Leica Microsystems, Vienna, Austria), transferred to formvar-coated slot grids, and examined using a Phillips CM10 TEM (FEI Company, Hillsboro, OR) equipped with an Advantage Plus 2-megapixel digital charge-coupled-device system for CM10 transmission electron microscopy (TEM) (Advanced Microscopy Techniques, Danvers, MA).
For statistical analyses, DMVs were characterized into two groups, either regular (defined by inner membranes in close approximation with the outer membrane) or irregular DMVs (defined by moderate to severe disruption or separation of the inner membrane with the outer membrane). Chi-square analysis using contingency tables was performed by comparing the number of regularly formed DMVs to irregularly formed DMVs of wt and nsp4 glycosylation mutant viruses. Chi-square analysis was also performed to compare the presence of both CMs and DMVs to the presence of DMVs only. Because CMs were found only in the presence of DMVs, the presence of CMs and DMVs was compared to the presence of DMVs alone in a given TEM section. Diameters of DMVs were measured using ImageJ 1.40g (http://rsb.info.nih.gov/ij). Diameters were defined by measuring the widest diameter from the outside membrane of one side to the outside membrane of the opposite side of a single DMV. To determine whether there was a statistical difference between the diameters of DMVs, analysis of variance (ANOVA) was used to compare wt and nsp4 glycosylation mutant viruses. Because a statistical difference was indicated through ANOVA, Tukey tests were used to perform pair-wise comparisons of all viruses. P values were determined to indicate significance.
Group 2a coronaviruses contain conservation of putative glycosylation sites in nsp4 at N176 and N237 (Fig. (Fig.1B).1B). To determine if nsp4 is glycosylated at residues N176 and N237 in the context of MHV infection and what roles nsp4 glycosylation may play in the virus life cycle, viruses were engineered to contain asparagine-to-alanine substitutions at either N176, N237, or both residues N176 and N237 of nsp4 (Fig. (Fig.1C).1C). Cells were electroporated with genomic RNA for N176A, N237A, or N176A/N237A mutant viruses. All three mutant viruses induced cytopathic effect by 36 h postelectroporation, and 90 to 100% of cells were involved in syncytia by 46 to 50 h postelectroporation, similar to wt virus. Viruses were passaged and sequenced across the nsp4 coding sequence, confirming both the presence of engineered mutations and lack of any other mutations in nsp4. In contrast to previous reports, our results demonstrate that mutants with alanine substitution at N176, N237, or both are viable, demonstrating that the N176 and N237 residues are not required for replication in cell culture. To determine if compensating mutations occurred outside of the nsp4 sequence during recovery of the N237A and N176A/N237A mutant viruses, the complete genome of the N176A/N237A mutant virus was sequenced, and there were no additional mutations present in the genome. These results demonstrate that the recovery of the N237A and N176A/N237A mutant viruses was not due to second-site compensating mutations and that the Asn residues are not required for virus viability.
Previous studies have demonstrated that treatment of lysates with Endo H results in a mobility shift of nsp4 expressed from plasmid in HeLa cells (10, 35) or from nsp4-enhanced green fluorescent protein expressed in recombinant virus from an alternate location (in place of ORF2) (10, 35), consistent with glycosylation of nsp4 with mannose-rich oligosaccharides in the endoplasmic reticulum (ER) and the lack of nsp4 trafficking through Golgi. However, there has been no demonstration of N-linked glycosylation of native nsp4 in wt virus or identification of specific Asn residues subject to N-linked glycosylation. To test whether natively expressed MHV nsp4 is glycosylated during infection, immunoprecipitated nsp4 from wt MHV infection was mock treated or treated with Endo H (Fig. (Fig.2A).2A). Mock-treated nsp4 was detected as a 44-kDa protein by SDS-PAGE, while Endo H treatment resulted in a faster-migrating, 39-kDa protein. The nsp4-to-10 precursor was detected in both cases by anti-nsp4. The replicase protein nsp8 is not modified by N-linked glycosylation and was not affected by Endo H treatment (Fig. (Fig.2A).2A). The nsp4-to-10 precursor that was treated with Endo H and detected using anti-nsp8 exhibited a sharper band than that of the untreated nsp4-to-10 precursor. A possible explanation for this is that removal of N-linked glycans may alter which nsp4-to-10 precursors can be detected by anti-nsp8, e.g., nsp4-to-10 with certain posttranslational modifications.
To test whether N176 and/or N237 was targeted for glycosylation, nsp4 immunoprecipitated following infection of DBT cells with N176A, N237A, and N176A/N237A mutant viruses was treated with Endo H (Fig. (Fig.2B).2B). Untreated nsp4 from N176A and N237A mutants migrated identically and more rapidly than untreated wt nsp4 (42 kDa) but more slowly than wt nsp4 treated with Endo H (39 kDa). When nsp4 from N176A and N237A mutant viruses was treated with Endo H, both proteins were detected at 39 kDa, identical to Endo H-treated wt nsp4. nsp4 from the N176A/N237A mutant virus migrated to 39 kDa, whether untreated or treated with Endo H. The results indicate that nsp4 expressed from its native genomic location is specifically glycosylated at residues N176 and N237 and also demonstrate that no other N-linked glycosylation occurs in nsp4.
To determine whether nsp4 glycosylation mutant viruses display replication defects, DBT cells were infected with wt, N176A, N237A, and N176A/N237A viruses at an MOI of 1 PFU/cell (Fig. (Fig.3A).3A). Samples of infected cell culture medium were taken at predetermined time points from 1 to 24 h p.i., and virus titers of each sample were determined by plaque assay. The N176A mutant virus exhibited growth kinetics and peak titers indistinguishable from those of wt virus, consistent with the study by Clementz et al. (10). The N237A and N176A/N237A mutant viruses grew indistinguishably from each other and reached peak titers similar to those of wt virus; however, compared to wt and N176A, the N237A and N176A/N237A viruses exhibited a delay and decrease in growth between 4 and 12 h p.i. The N176A/N237A mutant did not appear more impaired in growth than the N237A mutant alone. Since the N237A and N176A/N237A mutant viruses exhibited growth defects, we next tested whether N176A had subtle growth defects by repeating the growth assays at an MOI of 0.01 PFU/cell (Fig. (Fig.3B).3B). Under these conditions, the N237A and N176A/N237A mutants demonstrated the same delay compared to mutants infected at a higher MOI. In contrast, for the N176A mutant virus, the lower MOI infection revealed a subtle defect in growth, displaying a delay in peak titer similar to that of N237A and N176A/N237A mutants. The experiments demonstrate that N176 and N237 both are important for exponential growth, but loss of either or both glycosylation sites still allows for wt peak titers. The contributions of N176 and N237 are independent and nonredundant, as indicated by growth defects of either N176A or N237A but are not additive or synergistic. Finally, the results suggest that glycosylation of nsp4 is important for nsp4 function during virus replication.
Since previous studies have shown that mutations in nsp4 affect viral RNA synthesis (45), we conducted experiments to determine if the growth defects of nsp4 glycosylation mutants were associated with changes in viral RNA synthesis (Fig. (Fig.4).4). DBT cells were mock infected or infected with wt, N176A, N237A, or N176A/N237A mutant viruses at an MOI of 5 PFU/cell to maximize single-round infection, and infected cells were metabolically labeled with [3H]uridine in the presence of Act D for 2-h intervals from 3 to 15 h p.i. Total RNA was extracted from harvested cells and measured for incorporation of [3H]uridine. Peak incorporation of [3H]uridine for wt MHV occurred from 9 to 11 h p.i., similar to the results from a previously published report (16). For all three nsp4 mutant viruses, peak incorporation was delayed compared to wt, occurring between 11 and 13 h p.i. Delays in the timing of peak viral RNA synthesis displayed by the nsp4 glycosylation mutant viruses were also associated with decreases in the amount of RNA synthesized over the course of the infection. The N176A mutant virus synthesized approximately 80% of the maximum amount of incorporation seen for wt over a 2-h labeling period. Both the N237A and the N176A/N237A mutant viruses exhibited a 50% reduction in peak viral RNA synthesis. These data demonstrate that there is an overall decrease in viral RNA synthesis in the nsp4 mutant viruses compared to wt virus. In addition, the delay and decrease in RNA synthesis correlated with the kinetics and peak titer of infectious viruses, suggesting that alteration of viral RNA synthesis was responsible for the growth defects from the N176A and N237A substitutions.
nsp4 colocalizes with other replicase nsp's in cytoplasmic replication complexes that are sites of viral RNA synthesis, and nsp4 has been predicted to be critical for formation of these complexes. To test if altered RNA synthesis resulting from the N176A and N237A substitutions was associated with altered nsp4 interactions with other replicase proteins, the localization of nsp4 was compared by immunofluorescence with nsp8, a well-described marker for replication complexes, and with the viral membrane protein (M), a marker for sites of virus assembly in the ER-Golgi intermediate compartment and Golgi and distinct from replication complexes. DBT cells on glass coverslips were infected with wt, N176A, N237A, or N176A/N237A viruses for 6 h, fixed, and probed for nsp4, nsp8, and M. For wt and all nsp4 mutant viruses, nsp4 colocalized extensively with nsp8 in punctate perinuclear and cytoplasmic foci (Fig. (Fig.5A).5A). However, there was a visual trend for fewer and less-intense fluorescent foci in the cells infected with the nsp4 mutants compared to wt virus, suggesting that there may be fewer-forming or altered replication complexes in the nsp4 mutant virus infections (Fig. (Fig.5A5A and data not shown). When nsp4 was compared with M (Fig. (Fig.5B),5B), wt and mutant viruses had identical patterns of noncolocalization of nsp4 with M, consistent with previous studies of MHV replicase proteins and indicating that nsp4 is not altered in its relationship to sites of assembly and not localized to the ER-Golgi intermediate compartment or Golgi. The results demonstrate that nsp4 mutant viruses are able to form cytoplasmic replication complexes and retain interactions with other replicase nsp's and that glycosylation of nsp4 is not required for this process.
Based on the replication defects and subtle visual variability observed during immunofluorescence analysis of nsp4 mutants, we next investigated whether nsp4 glycosylation mutants have altered membrane rearrangements. TEM was used to visualize the ultrastructure of membrane modifications in infected cells. DBT cells were mock infected or infected with wt or the nsp4 glycosylation mutant viruses at an MOI of 5 PFU/cell. At 6 h p.i., cells were fixed in 2% glutaraldehyde and processed for TEM analysis. For mock-infected cells, the cellular architecture and organelle morphology were intact (Fig. (Fig.6A).6A). Cells infected with wt virus exhibited clearing of cytoplasmic contents and swollen ER and Golgi (Fig. (Fig.6B).6B). Cells infected with the three nsp4 glycosylation mutant viruses also demonstrated swelling of ER and Golgi and cytoplasmic clearing, albeit less so than during wt infection (Fig. 6C to E).
In contrast, there was a striking difference between cells infected with wt and nsp4 mutants in the relationship and ultrastructure of virus-induced DMVs and CMs. wt- and nsp4 mutant-infected cells exhibited virus-induced CMs and DMVs, structures that have been identified with replication complexes and associated with viral RNA synthesis (15, 27), while no DMVs or CMs were observed with mock-infected cells. CMs were detected with wt and mutant virus-infected cells and always in close proximity to DMVs. However, DMVs were observed in the presence or absence of CMs for all viruses. The CMs were observed more frequently with electron microscopy (EM) sections of cells infected with N176A, N237A, and N176A/N237A mutant viruses compared to wt (Fig. 6C to E). The vast majority of DMVs in wt-infected cells exhibited the characteristic DMV morphology of a circular shape, regular diameter, and ultrastructure of closely approximated inner and outer membranes. A small subset of DMVs manifested a partial separation of the inner and outer membranes and exhibited a slightly larger diameter, but these were rare. In contrast, cells infected with the nsp4 glycosylation mutants demonstrated DMVs with altered shape and diameter and with increasingly aberrant (irregular) ultrastructure, consisting of severely detached and collapsed inner membranes that were not observed with any wt-infected cells. The number of irregular DMVs and the extent of DMV derangement were most profound in N237A and N176A/N237A mutant-infected cells and visibly greater than those detected with cells infected with N176A alone.
Because the EM images were originally selected based on the detection of DMVs, we used EM images to quantitatively compare (i) the prevalences of CMs, (ii) the ratios of regular (wt-like) and irregular DMVs, and (iii) the diameters of regular and irregular DMVs (Table (Table22 and Fig. Fig.7).7). Since the images were selected only for the presence of DMVs, we proposed that quantitative analysis was unbiased for these parameters. The prevalence of CMs was determined by comparing images in which CMs were observed or not observed in EM sections selected based on the presence of DMVs, since CMs were found only in the presence of DMVs. While there was no statistical difference between wt and N176A in the ratios of sections with both CMs and DMVs versus DMVs alone, the N237A and N176A/N237A mutants had significantly increased ratios of detection of both CMs and DMVs compared to DMVs alone (P < 0.01 for N237A and P < 0.001 for N176A/N237A) (Fig. (Fig.7A).7A). Analysis of the ratio of regular DMVs to total DMVs (regular plus irregular) demonstrated a significant increase in irregular DMVs in cells infected with N237A and N176A/N237A mutant viruses (P < 0.001) compared to cells infected with wt or N176A viruses (Fig. (Fig.7B).7B). We did observe more irregular DMVs with N176A than with wt, but the regular DMV/total DMV ratios were not significantly different. Finally, the measurements of the regular DMVs of both wt and all nsp4 glycosylation mutant viruses reveal no difference in their diameters (widest diameter of outer membrane) (Fig. (Fig.7C).7C). In contrast, the mean diameter of irregular DMVs in the N237A and N176A/N237A mutant viruses was significantly larger than that of either wt virus or the N176A mutant virus (Fig. (Fig.7C).7C). This analysis indicates that nsp4 is likely critical for the organization and stability of DMVs and for the relationship and evolution of membrane modifications (CMs and DMVs) over the course of infection.
Although multiple studies have investigated the roles of nsp's in inducing membrane rearrangements, understanding the role of glycosylation of nsp's from positive-strand RNA viruses remains limited. A study of the flavivirus yellow fever virus demonstrated that NS1 glycosylation was important for several functions in the virus life cycle (30, 33). NS1 interacts with membranes and is involved in replicase function (30), and removal of NS1 glycosylation by asparagine-to-alanine substitution results in impaired virus growth, RNA synthesis, and pathogenesis (33).
Coronaviruses, like other positive-strand RNA viruses, induce the formation of DMVs that serve as scaffolds for replication/transcription complexes. Exogenous expression of the poliovirus transmembrane proteins 2BC and 3A results in DMVs that are indistinguishable from those formed during wt infection (43, 46). Equine arteritis virus (EAV), which is classified with coronaviruses in the order Nidovirales, induces DMVs similar to coronaviruses (37). Exogenous plasmid expression of EAV nsp2 and nsp3 is sufficient to induce membrane modifications resulting in membrane structures similar to those seen during EAV infection, and mutations within EAV nsp3 also result in altered virus-induced membrane rearrangements (39, 44). EAV nsp3 is a tetra-spanning integral membrane protein implicated in DMV formation and organization. Of interest, an introduced Asn substitution (T873N) in an EAV nsp3 luminal domain resulted in nsp3 glycosylation in vitro but was highly detrimental when introduced into the genome and recovered only as a pseudoreversion (N873H) that abolished the glycosylation site. Thus, for another nidovirus, the glycosylation status of a membrane-modifying replicase protein is also important for DMV formation and RNA synthesis during virus replication.
Our report confirms multiple roles of MHV nsp4 in the virus life cycle, including optimal virus replication and RNA synthesis, as well as its importance in the modification and morphology of virus-induced membrane structures. In this study, we show that MHV nsp4 is glycosylated and functions as a membrane modification protein that regulates virus-induced membrane rearrangements. nsp4 glycosylation mutant viruses display highly irregular DMVs and an increased prevalence of CMs relative to DMVs alone. The extent of disrupted DMVs in the nsp4 glycosylation mutant viruses correlated directly with decreases in RNA synthesis and virus replication. These data suggest that altered membranous structures from the nsp4 glycosylation mutants result in a reduced capacity to synthesize viral RNA and/or protect viral RNA from degradation, ultimately leading to impaired virus fitness.
Previous studies have concluded that nsp4 is required for MHV replication and have identified determinants of membrane topology, subcellular localization, and function (10, 35, 45). This study is the first to recover and characterize the importance of multiple nsp4 glycosylation events to virus replication, viral RNA synthesis, and virus-induced membrane modifications during coronavirus infection. Clementz et al. recovered an nsp4 N176A mutant but were unable to recover an N237A or N176A/N237A mutant (10). Their N176A mutant grew with kinetics similar to those of wt at an MOI of 0.1 PFU/cell at 33°C and 39°C but was not further characterized in that report. In contrast to the results in the previously published report, we were able to recover and characterize the N237A and N176A/N237A mutant viruses. The reasons for the differences in recovery can only be speculated. The backgrounds of cloned MHV genome fragments should be identical since the MHV genome fragments were jointly developed by our lab and the lab of Baric and coworkers (47). In addition, we performed RT-PCR sequencing of the complete genome from the recovered N176A/N237A mutant virus, which verified the engineered mutations and also confirmed that the rest of the genome was identical, with no additional mutations of any kind, to the published recombinant MHV-A59 sequence. Thus, there were no other compensating mutations to account for or consider for the recovery of the mutant virus. We have experienced occasional mutations in the genome fragments during preparation for genome assembly that have prevented recovery of even known viable mutants and would therefore speculate that this could account for the nonrecovery of N237A and N176A/N237A mutant viruses by Clementz et al. Our results clearly demonstrate that the N176 and N137 residues and the associated glycosylation events are not required for MHV replication in cell culture. Since no other mutations in the genome RNA from the recovered N176A/N237A mutant virus were identified, we can conclude that the profound and distinct phenotypes in virus replication, RNA synthesis, and virus-induced cellular membrane modifications are due to the introduced mutations alone.
Modification of proteins by addition of N-linked glycans may result in numerous effects on protein functions (14, 19). Therefore, glycosylation of nsp4 may be important for a variety of reasons. One potential mechanism of nsp4 glycosylation is proper protein folding (18, 36). By removing N-linked glycans, the overall structure of nsp4 may be altered during protein folding. This mechanism is supported by the findings in this report, in that the nsp4 glycosylation mutant viruses displayed impairments in virus replication, viral RNA synthesis, and virus-induced membrane modifications. Other explanations are possible for the role of nsp4 glycosylation in replication complex formation and membrane modifications. For instance, glycosylation of nsp4 may be important for protein stability and prevention of nsp4 degradation (26). Lastly, it is possible that the N-linked glycans, either directly or through modification of nsp4 structure, recruit cellular factors that are involved in membrane rearrangements. Future studies are needed to distinguish between these possibilities.
Evidence from this study has led to potential models addressing the effect nsp4 has on replication complex formation, morphology, and organization. One possible model is that nsp4 may regulate the transition or formation of different membrane modifications (i.e., CMs and DMVs). The evidence from this report that there was an increased prevalence of CMs in relation to DMVs in the N237A and N176A/N237A mutant viruses suggests that MHV nsp4 may be a major player in the transition of these virus-induced membrane rearrangements from one membrane structure to another. Other findings from this report that there was an increased presence of aberrant or deranged DMVs in the N237A and N176A/N237A mutant viruses suggest another possibility that the formation of intact, functional DMVs is regulated by nsp4.
A second potential model of nsp4 function is that the curvature and size of DMVs are regulated by nsp4 (38). In N237A and N176A/N237A mutant virus-infected cells, irregular DMVs were much larger and had highly disrupted inner membranes. The N237A and N176A/N237A mutant viruses also exhibited decreases in RNA synthesis, indicating that these irregular DMVs may not be functioning properly and that curvature and size may be important for proper function. This model is supported by the fact that all virus-infected cells produced regular DMVs, although at different proportions, and that all regular DMVs were similar in size. Cells infected with wt or N176A viruses, those that had levels of RNA synthesis higher than those of the N237A and N176A/N237A mutant viruses, also had a higher percentage of regular DMVs. These data suggest that curvature and size are important for DMV function.
A third model is that nsp4 functions in tethering or “pushing” the inner membrane to the outer membrane of the DMVs. The proximity of the inner membrane to the outer membrane may be important for creating an environment optimal for RNA synthesis and/or protection of newly synthesized viral RNAs. This model is supported by the fact that the prevalence of aberrant DMVs in the nsp4 glycosylation mutants was directly related to the extent of impairment of RNA synthesis and virus growth. These results suggest that irregular DMVs have a reduced capacity to synthesize and/or protect viral RNAs and are also the first to provide direct evidence suggesting that the physical size, morphology, and stability of virus-induced DMVs are important for efficient viral RNA synthesis and optimal virus production. On the other hand, the results also show clearly that glycosylation of nsp4 is not absolutely required for formation of “regular” DMVs and that replication complex function can still ultimately allow virus replication to wt titers, albeit with delayed kinetics.
To date, all coronavirus nsp4's that were subjected to Endo H treatment have been shown to be glycosylated in the lumen of the ER between the first and second predicted transmembrane domains of nsp4 in exogenous expression experiments, including group 2a MHV nsp4, group 2b SARS-CoV nsp4, and group 3 IBV nsp4 (10, 29, 35). It will be interesting to see whether glycosylation of nsp4 is conserved among other coronaviruses, specifically group 1 coronaviruses, and what effect the loss of glycosylation sites has on virus replication, RNA synthesis, and replication complex morphology.
This study has demonstrated the importance of MHV nsp4 glycosylation sites in virus replication, replication complex morphology and organization, and viral RNA synthesis. Because nsp4 has been shown to have integral membrane characteristics and no predicted enzymatic activities, it is rational to propose that nsp4 involvement in viral RNA synthesis is due to replication complex formation, other possible membrane modifications, and/or protein interactions. The nsp4 glycosylation mutant viruses generated in this study will provide powerful tools to further dissect the definitive mechanisms of nsp4 function on replication complex formation and its roles in the virus life cycle.
We thank Elvin Woodruff for TEM assistance and image analysis. We also thank Megan Culler and Xiaotao Lu for technical assistance. We thank Michelle Becker and Lance Eckerle for advice and critical reviews of the manuscript.
Support for this work was provided by National Institutes of Health grant R01 AI50083 (M.R.D.) from the National Institute of Allergy and Infectious Diseases. M.J.G. was supported by the Training Grant in Mechanisms of Vascular Disease through the Vanderbilt University School of Medicine (T32 HL007751). J.S.S. was supported by Public Health Service award T32 CA009385. This work was also supported by the Elizabeth B. Lamb Center for Pediatric Research.
Published ahead of print on 21 October 2009.