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Journal of Virology, May 2008, p. 4480-4491, Vol. 82, No. 9
0022-538X/08/$08.00+0     doi:10.1128/JVI.02756-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Formation of the Arterivirus Replication/Transcription Complex: a Key Role for Nonstructural Protein 3 in the Remodeling of Intracellular Membranes

Clara C. Posthuma,^1, Ketil W. Pedersen,^2,, Zhengchun Lu,^1, Ruth G. Joosten,¹ Norbert Roos,² Jessika C. Zevenhoven-Dobbe,¹ and Eric J. Snijder^1*

Molecular Virology Laboratory, Department of Medical Microbiology, Center of Infectious Diseases, Leiden University Medical Center, PO Box 9600, 2300 RC Leiden, The Netherlands,¹ Department of Molecular Biosciences, University of Oslo, 0316 Oslo, Norway²

Received 28 December 2007/ Accepted 15 February 2008

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The replication/transcription complex of the arterivirus equine arteritis virus (EAV) is associated with paired membranes and/or double-membrane vesicles (DMVs) that are thought to originate from the endoplasmic reticulum. Previously, coexpression of two putative transmembrane nonstructural proteins (nsp2 and nsp3) was found to suffice to induce these remarkable membrane structures, which are typical of arterivirus infection. Here, site-directed mutagenesis was used to investigate the role of nsp3 in more detail. Liberation of the hydrophobic N terminus of nsp3, which is normally achieved by cleavage of the nsp2/3 junction by the nsp2 protease, was nonessential for the formation of DMVs. However, the substitution of each of a cluster of four conserved cysteine residues, residing in a predicted luminal loop of nsp3, completely blocked DMV formation. Some of these mutant nsp3 proteins were also found to be highly cytotoxic, in particular, exerting a dramatic effect on the endoplasmic reticulum. The functionality of an engineered N glycosylation site in the cysteine-containing loop confirmed both its presence in the lumen and the transmembrane nature of nsp3. This mutant displayed an interesting intermediate phenotype in terms of DMV formation, with paired and curved membranes being formed, but DMV formation apparently being impaired. The effect of nsp3 mutations on replicase polyprotein processing was investigated, and several mutations were found to influence processing of the region downstream of nsp3 by the nsp4 main protease. When tested in an EAV reverse genetics system, none of the nsp3 mutations was tolerated, again underlining the crucial role of the protein in the arterivirus life cycle.

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The replication or replication/transcription complexes (RTCs) of a wide variety of eukaryotic positive-strand RNA viruses have been found to be associated with (modified) intracellular membranes (for recent reviews, see references 1, 2, 22, 24, 25, and 31). Membrane association of the RTC is thought to be important for creating a suitable (micro)environment for viral RNA synthesis and may also aid in preventing the activation of host defense mechanisms that can be triggered by double-stranded RNA replication intermediates. For several virus groups, replicase subunits have been identified that are involved in targeting the RTC to membrane compartments and/or modifying these membranes, often resulting in vesiculation or the formation of invaginations. Frequently, parts of these nonstructural proteins, which often contain multiple hydrophobic segments, are known or thought to be embedded in the membrane. All major groups of mammalian positive-strand RNA viruses produce their replicative machinery from replicase polyproteins containing both these hydrophobic subunits and the enzymes directly involved in RNA synthesis. Consequently, both the correct proteolytic processing and the membrane association of replicase subunits are important and probably highly coordinated events during the initial stages of the viral life cycle.

Equine arteritis virus (EAV), the arterivirus prototype, is an enveloped, positive-stranded RNA virus with a 12.7-kb genome, of which about three-quarters is occupied by a large gene encoding the replicase/transcriptase (commonly referred to as "replicase" for simplicity). In terms of its analogy with other members of the order Nidovirales, which also includes corona-, toro-, and roniviruses (for reviews, see references 12 and 35), the EAV replicase gene is comprised of two large open reading frames (ORFs), ORF1a and ORF1b, with expression of the latter involving a ribosomal frameshift near the 3' end of ORF1a. Consequently, genome translation produces pp1a and pp1ab, two polyproteins of 1,727 and 3,175 amino acids, respectively, which are subject to extensive proteolytic processing by three viral proteases, resulting in 13 nonstructural proteins (nsps) (Fig. 1A) (44, 54). The mature replicase subunits then assemble into the membrane-bound RTC (28, 46) that directs viral RNA synthesis, which involves both genome amplification and the production of a nested set of subgenomic mRNAs, presumably each from their specific negative-strand RNA templates (for reviews, see references 27 and 32). The subgenomic mRNAs are used to express the viral structural proteins from a set of genes in the 3' proximal region of the genome.

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FIG. 1. (A) The domain organization of EAV replicase pp1ab. The border between ORF1a- and ORF1b-encoded residues is indicated as the RFS (ribosomal frameshift). Gray and black arrowheads represent the sites that are cleaved by accessory proteases (nsp1 papain-like cysteine protease [PCP] and nsp2 CP) and the nsp4 main protease (MP), respectively. Cleavage products are numbered, and the locations of domains that are structurally or functionally related to those in other nidovirus replicases are highlighted (12). These include four nidovirus-wide conserved domains encoded in ORF1b (RdRp, zinc-binding domain [Z], helicase [HEL], and NendoU [N]) and three putative transmembrane domains (TM1, TM2, and TM3). The and β symbols refer to the products of the recently documented internal cleavage of nsp7 by the nsp4 protease (44). (B) Predicted membrane topology of EAV nsp3. The four transmembrane helices predicted by ConPred II (http://bioinfo.si.hirosaki-u.ac.jp/ConPred2) are depicted. Open circles mark the positions of the four conserved Cys residues (see panel C) in the first (predicted) luminal loop. The position of the N glycosylation site engineered at position 873 in the Thr-873Asn mutation is also indicated. The nsp2/3 and nsp3/4 cleavage sites are indicated by arrowheads and amino acid numbers refer to those in the full-length EAV pp1ab. (C) Arterivirus-wide sequence alignment of the (predicted) luminal loop of nsp3, indicating the four conserved Cys residues (bold) and engineered N glycosylation site (see above). Fully conserved residues are highlighted in bold. LDV-P, lactate dehydrogenase-elevating virus Plagemann strain (accession no. U15146); PRRSV-EU, porcine reproductive and respiratory syndrome virus, strain Lelystad (accession no. M96262); PRRSV-NA, porcine reproductive and respiratory syndrome virus, strain VR2332 (accession no. DQ217415); SHFV, simian hemorrhagic fever virus (accession no. NC_003092); EAV, equine arteritis virus (accession no. NC_002532.).

Although many arterivirus nsps remain to be studied in detail, a number of functional domains have been identified and characterized through a combination of theoretical and experimental approaches. The ORF1b-encoded part of pp1ab contains four nidovirus-wide conserved domains (Fig. 1A), including key enzymes for viral RNA synthesis (see reference 12 and references therein) like the RNA-dependent RNA polymerase (RdRp) (3). ORF1a encodes two or three papain-like cysteine protease domains ("accessory proteases," located in nsp1 and nsp2) and a chymotrypsin-like serine protease ("main protease") in nsp4 (Fig. 1A). It also encodes substantial hydrophobic regions, residing in nsp2, nsp3, and nsp5, which were therefore proposed to be multispanning membrane proteins.

Most of the EAV nsps, including the RdRp, colocalize in the perinuclear region of the infected cell (46, 48). Electron microscopy (EM) revealed the accumulation of tightly apposed membranes and vesicles bounded by double membranes (typical diameter, 80 to 100 nm). These double-membrane vesicles (DMVs) could be labeled with antibodies specific for several replicase subunits. Br-UTP labeling of de novo-made viral RNA indicated that it was the likely site of RNA synthesis (28). In the absence of other viral components or RNA synthesis, similar membrane alterations could be induced by the expression of two of the putative transmembrane nsps of EAV in the form of a polyprotein consisting of nsp2 and nsp3 (nsp2-3), which cleaves itself due to the activity of the cysteine protease (CP) in the N-terminal domain of nsp2 (37). Partial colocalization with protein disulfide isomerase (PDI), a marker protein specific for the endoplasmic reticulum (ER), and morphological observations under EM suggested that DMVs are derived from ER membranes (28, 37, 46).

Residues Cys-270 and His-332 are thought to form the catalytic dyad of the nsp2 CP (40), which cleaves the site between nsp2 and nsp3 (nsp2/3) that is presumably downstream of Gly-832 (9). The CP belongs to the ovarian tumor domain protease superfamily (23), of which several members have been implicated in deubiquitination (for a review, see reference 43). The EAV CP was recently shown to be capable of deconjugating both ubiquitin and ISG15 from cellular proteins, potentially implicating this viral protease in immune evasion (9). In addition to its crucial role in DMV formation, nsp2 also acts as a cofactor for the nsp4 main protease, which uses alternative pathways to cleave the processing intermediate consisting of nsp3 through nsp8 (nsp3-8), depending on the association of nsp3-8 with nsp2 (44, 51).

EAV nsp3 is predicted to be a tetraspanning transmembrane protein (Fig. 1B) and contains a number of residues, including a cluster of four cysteines, that are conserved throughout the arterivirus family (Fig. 1C) and are predicted to reside in the first luminal domain of the protein (Fig. 1B). In biochemical studies (46), both nsp2 and nsp3 were recovered from the membrane fraction of infected cells and their membrane association is also supported by data from immunofluorescence assays (IFA) and EM studies. Both proteins contain internal hydrophobic domains, with one of these forming the N-terminal domain of nsp3 (amino acids 833 to 993 [Fig. 1B]), which could be made available as a signal sequence for translocation following prior cleavage of the nsp2/3 site by the nsp2 CP. Still, since both subunits are part of a larger polyprotein, the mechanism of membrane insertion remains unclear.

In this paper, we address the role of EAV nsp3 in more detail. We have used alphavirus-driven nsp2-3 expression to study DMV formation, a pp1a expression system to analyze proteolytic processing, and an EAV reverse genetics system to perform site-directed mutagenesis of nsp3. The importance of both the nsp2/3 cleavage and the cluster of conserved Cys residues in nsp3 was investigated. Conclusive evidence on the transmembrane nature of nsp3 was obtained, and a crucial role of the protein in the virus life cycle, and DMV formation in particular, was established.

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Cell lines and virus. Baby hamster kidney cells (BHK-21; ATCC CCL10) were used for pSR5 and pEAN551 transfection experiments. Rabbit kidney (RK-13) cells were used for pL1a transfection experiments following infection with vaccinia virus recombinant vTF7-3 (10), which produces the T7 RNA polymerase.

Sindbis virus-based expression vectors. A modified Sindbis virus expression vector for nsp2-3 expression, pSR5-HA23His, was engineered by in-frame fusion of the sequence encoding the influenza virus hemagglutinin (HA) tag to the 5' end of the nsp2-coding sequence in the previously described vector pSRE-nsp2+3His (37), a derivative of pSinRep5 (7). Vector pSR5-HA23His expressed a self-cleaving nsp2-3 polyprotein consisting of an N terminally tagged HA-nsp2 and a C terminally tagged nsp3-His₆. Mutations in the nsp2-3-coding sequence were introduced using appropriate shuttle vectors and standard site-directed PCR mutagenesis. The mutations engineered in the nsp2-3-coding region are listed in Table 1. BHK-21 cells were transfected by electroporation (47) with in vitro-transcribed RNA from the wild type (wt) and mutant pSR5-HA23His vectors.

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TABLE 1. Overview of nsp2 and nsp3 mutations used in this study

Analysis of pp1a proteolytic processing. To study their effect on proteolytic processing of pp1a, the nsp2 and nsp3 mutations were transferred to the previously described pp1a expression vector pL1a (40), which contains EAV ORF1a downstream of the T7 promoter and a copy of the encephalomyocarditis virus internal ribosomal entry site, which is used to enhance translation (17). RK-13 cells were infected with vaccinia virus recombinant vTF7-3 and subsequently transfected with wt or mutant pL1a, essentially as described previously (38). Transfection was carried out with Lipofectamine 2000 (Invitrogen) in Opti-MEM medium (Invitrogen) without fetal calf serum. After 2 h, the medium was replaced by Opti-MEM containing 2% fetal calf serum. Proteins were ³⁵S labeled from 5 to 8 h postinfection by using serum-free medium containing 200 mCi/ml ³⁵S-Translabel (Amersham). Protocols for cell lysis and immunoprecipitation and EAV replicase-specific antisera have previously been described (28, 38).

Proteins were analyzed by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) according to the method of Laemmli (21), and the ³⁵S-labeled protein bands were detected using a phosphorimager (Bio-Rad molecular imager FX). Deglycosylation was performed overnight using N-glycosidase F (PGNase F; New England Biolabs) at 37°C in the buffer recommended by the manufacturer, after which the samples were analyzed by SDS-PAGE as described above.

EAV reverse genetics. Using appropriate restriction sites, mutations in the nsp2-3-coding sequence were transferred into pEAN551, a derivative of the previously described EAV full-length cDNA clone (47) that contained some engineered restriction sites. Virus derived from pEAN551 displayed a wt phenotype (data not shown), and this construct was used as a wt control in the experiments. BHK-21 cells were electroporated with in vitro-derived RNA transcripts of wt and mutant EAV full-length cDNA clones as described previously (47). In order to test for the production of infectious virus progeny, plaque assays were performed as described previously (34), with supernatants harvested from transfected cells after 72 h.

Analysis of revertants was carried out by infecting fresh BHK-21 cells with supernatant harvested at 72 h posttransfection. Total RNA was isolated after 24 h by the acidic phenol method (49), and the nsp3-coding region was amplified by reverse transcription-PCR (RT-PCR) using E628 (5'-CACAGCAGCACCAGCGAG-3'; nucleotides 3081 to 3098) as antisense primer for both RT and PCR and E625 (5'-GGCGCCCATCCCAGCACCG-3'; nucleotides 1526 to 1544) as sense PCR primer. PCR products were sequenced using standard protocols to check for the presence of the mutations.

IFAs and EM. For IFAs, transfected cells were seeded on coverslips, fixed with 3% (wt/vol) paraformaldehyde at 8 h posttransfection, and processed as described previously by van der Meer et al. (46). Cells transfected with EAV full-length RNA were analyzed by double labeling IFA using antisera against EAV nsp3 (98E3) and N protein (monoclonal antibody 3E2) (46). Cells transfected with RNA derived from pSR5-HA23His and derivatives were labeled with an anti-HA monoclonal antibody (monoclonal antibody 12CA5) (8). Labeling for PDI was performed using monoclonal antibody 1D3 (50).

For EM, cells were transfected with wt or mutant pSR5-HA23His-derived RNA as described above, seeded in 10-cm² culture dishes, and fixed with 1.5% glutaraldehyde in 100 mM cacodylate buffer (pH 7.2) at 8 h posttransfection. After 1 h at room temperature, cells were postfixed in 1% osmium tetraoxide in phosphate buffer (pH 7.2) for 60 min at 4°C, followed by overnight block staining with 0.5% uranyl acetate in 70% ethanol at 4°C. The following day, cells were dehydrated using increasing concentrations of ethanol and embedded in epoxy resin X-122. Ultrathin sections were stained with lead citrate and examined using a Philips CM100 transmission electron microscope and a Philips Tecnai 12 transmission electron microscope with a MegaView III TEM soft-imaging system.

Protocols for ultrathin sectioning and for cryoimmuno-EM have previously been described by Pedersen et al. (28). Briefly, cells were fixed by using 4% paraformaldehyde and 0.1% glutaraldehyde in 200 mM HEPES, pH 7.4, scraped from the dish, pelleted, and incubated in 2.3 M sucrose. Cell pellets were flash frozen and sectioned with a Reichert Ultracut S ultramicrotome. Immunocytochemical labeling of thawed cryosections was performed essentially as described previously by Griffiths et al. (14). EM specimens were examined in a JEOL 1200EX or a Philips CM100 transmission electron microscope.

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Cleavage of the EAV nsp2/3 site is not required for DMV formation. The EAV RTC is associated with DMVs that are thought to be derived from the ER. Previously, an alphavirus RNA expression vector (SinRep5 [7]) was used to express EAV nsp2 and/or nsp3 (37). The individual expression of either nsp2 or nsp3 resulted in a diffuse cytoplasmic labeling, presumably representing localization to ER membranes, and DMV formation could not be detected (37). However, coexpression of the two proteins in the form of an nsp2-3 polyprotein, which cleaved itself due to the action of the nsp2 CP, resulted in a different and very distinct pattern. As in infected cells (46), the two subunits colocalized in the perinuclear region (Fig. 2A) and EM revealed the induction of large numbers of paired membranes and DMVs, resembling those found upon EAV infection despite being more variable in size and shape (28). Thus, both at the light microscopy and at the EM levels, the SinRep5 expression system could be used as a reporter system to study the formation of DMV-like structures (which, for simplicity, will also be termed DMVs in this paper). Below, we will use a slightly modified version of the same expression vector, in which nsp2 was N terminally tagged with an HA epitope tag in addition to the His₆ tag already present at the C terminus of nsp3 (pSR5-HA23His). The addition of the HA tag to nsp2 did not influence DMV formation (Fig. 2A and 3A; data not shown).

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FIG. 2. Immunofluorescence microscopy analysis of BHK-21 cells transfected with SinRep-5 vectors expressing wild-type and mutant versions of an HA epitope-tagged EAV nsp2-3 polyprotein. Cells were fixed at 8 h posttransfection and stained for nsp2 with a HA tag-specific antibody. Perinuclear staining was observed for the wt nsp2-3 (A) as well as for the polyprotein containing the nsp2 CP-inactivating mutation His-332Tyr (B). For the polyprotein carrying the Thr-873Asn N glycosylation mutation in the nsp3 loop, an "intermediate phenotype" was observed, with perinuclear staining and label spread through the cell (C). Expression of nsp2-3 polyproteins carrying mutations in the conserved Cys cluster in nsp3 (D to G) resulted in a dispersed labeling of the cytoplasm and the formation of many vesicles and vacuoles.

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FIG. 3. Electron micrographs of ultrathin sections of BHK-21 cells transfected with SinRep-5 vectors expressing wild-type and mutant versions of an HA-epitope-tagged EAV nsp2-3 polyprotein (8 h posttransfection). Bars, 1 µm. (A) Formation of DMVs upon expression of the wt nsp2-3 polyprotein. The arrow points to a neck-like connection to an ER cisterna, very similar to those observed previously (37). (B) Cells expressing of a nsp2-3 polyprotein containing an nsp2 CP active site mutation (His-332Tyr) show normal DMV formation. (C) Example of a cell expressing a nsp2-3 polyprotein carrying a substitution of one of the conserved Cys residues in the nsp3 loop (Cys-864Ser), which leads to the formation of single-membrane vesicles and large vacuoles that are probably derived from the ER. (D) Expression of the nsp2-3 polyprotein carrying the Thr-873Asn N glycosylation mutation in the nsp3 loop, which reduced DMV formation and produced conspicuous "balloon-shaped" structures consisting of double-membrane sheets.

Both nsp2 (572 amino acids) and nsp3 (232 amino acids) contain a hydrophobic domain (designated TM1 or TM2 in Fig. 1A) that presumably spans the membrane multiple times. Although computer predictions of the nsp2 membrane topology are not unequivocal (data not shown), arterivirus nsp3 proteins are uniformly predicted to contain four transmembrane helices, with the N and C termini of the protein residing in the cytoplasm (Fig. 1B). The most N-terminal predicted transmembrane domain of EAV nsp3 (residues 833 to 852 [numbers refer to pp1ab amino acid positions]) is found immediately downstream of the putative nsp2/3 cleavage site, and its release may, e.g., be a trigger for nsp3 translocation or membrane association. To investigate the importance of the nsp2/3 cleavage for DMV formation, the nsp2 CP was inactivated by a HisTyr replacement of active site His-332 (39). For the resulting mutant, normal perinuclear staining was observed by IFA (Fig. 2B) and EM revealed tightly apposed double membranes and DMVs that could not be discriminated from those induced by the wt nsp2-3 protein (Fig. 3B and 4A). As before, immuno-EM (Fig. 5A and B) revealed the colocalization of nsp2 and nsp3 on paired membranes and DMVs, with labeling for PDI again suggesting a link to the ER. These results demonstrated that cleavage of the nsp2/3 site is not a prerequisite for DMV formation.

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FIG. 4. Selected close-up electron micrographs of BHK-21 cells expressing mutant EAV nsp2-3 polyproteins. For details, see the legend for Fig. 3. Bars, 200 nm. (A) Typical DMV formed in cells expressing the nsp2-3 polyprotein containing an inactivated nsp2 CP. (B) Example of a single-membrane vesicle induced upon expression of a nsp2-3 polyprotein carrying a Cys-864Ser substitution in the nsp3 loop. (C) Close-up of "balloon-shaped" structures of paired membranes induced in cells expressing the Thr-873Asn N glycosylation mutant of nsp2-3.

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FIG. 5. Cryoimmuno-EM images of BHK-21 cells transfected with SinRep-5 vectors expressing mutant HA-tagged EAV nsp2-3 polyproteins. Bars, 200 nm. (A) The nsp2 CP active site mutant (H332Y), for which the nsp2-3 polyprotein remains uncleaved, produced paired membranes (arrow) which double labeled for nsp2 (anti-HA monoclonal antibody; 5 nm gold [open arrowheads]) and nsp3 (anti-nsp3 rabbit antiserum; 10 nm gold [filled arrowheads]). (B) Example of a double-membrane cisterna in cells expressing the H332Y mutant nsp2-3. The nonpaired part of the membranes is double labeled for PDI (anti-PDI monoclonal antibody; 5 nm gold [open arrowheads]) and nsp3 (anti-nsp3 rabbit antiserum; 10 nm gold [filled arrowheads]). (C) Cells expressing nsp2-3 with a Cys-864Ser replacement in the nsp3 loop. DMVs could not be detected and the labeling for nsp2 and nsp3 was localized to vesicular structures of variable size that were bounded by a single membrane (nsp2 labeled with anti-HA monoclonal antibody, 5 nm gold [open arrowheads], and nsp3 labeled with anti-nsp3 rabbit antiserum, 10 nm gold [filled arrowheads]). (D) Example of membrane structures in cells expressing the C864S mutant nsp2-3 that were double labeled for PDI (anti-PDI monoclonal antibody; 5 nm gold [open arrowheads]) and nsp3 (anti-nsp3 rabbit antiserum; 10 nm gold [filled arrowheads]). (E) Cells expressing nsp2-3 containing the Thr-873Asn glycosylation mutation in the nsp3 loop. This mutant can produce paired membranes (arrow) and DMVs similar to those seen for the wild-type control, with both structures double labeling for nsp2 (anti-HA monoclonal antibody; 5 nm gold [open arrowheads]) and nsp3 (anti-nsp3 rabbit antiserum; 10 nm gold [filled arrowheads]). However, there are also aggregates of label elsewhere in the cytoplasm (boxed in white), possibly associated with additional membranes structures.

Conserved cysteine residues in EAV nsp3 play a key role in DMV formation. The sequence immediately downstream of the first predicted nsp3 transmembrane domain, which is anticipated to be a luminal domain of about 50 amino acids (Fig. 1B), is highly conserved among arteriviruses and contains a cluster of four fully conserved Cys residues (Fig. 1C). Our attention was attracted, in particular, because of the potential for luminal interactions involving these residues, which might be involved in the formation of paired membranes. To study the importance of the conserved Cys residues (positions 855, 864, 879, and 884 of the EAV replicase), they were subjected to site-directed mutagenesis. The effect of the mutations was first analyzed in the context of the alphavirus expression system. Using IFA and EM, we tested whether the mutant nsp2-3 proteins were capable of inducing membrane modifications.

By IFA, the typical perinuclear labeling observed with wt nsp2-3, which is usually accompanied by the formation of DMVs, was absent for each of the four CysSer mutations (Fig. 2D to G). The signals for nsp2 (Fig. 2) and nsp3 (data not shown) still overlapped, but they were spread throughout the cell and the formation of many vesicles and larger vacuoles was observed. Although similar effects were observed for all four mutants, the expression of the Cys-855Ser and Cys-864Ser mutations appeared to have the most serious consequences (Fig. 2D and E). EM analysis revealed that the vesicles accumulating in transfected cells were bounded by only a single membrane. The Cys864Ser mutation is shown as an example in Fig. 3C and 4B. The initial vesicle size was similar to that of regular DMVs induced by wt nsp2-3, but whereas the latter were usually filled with cytosolic material, the interior of the single-membrane vesicles was electron lucent. Immuno-EM revealed the colocalization of nsp2 and nsp3 on these single-membrane structures (Fig. 5C and D), which also labeled positive for PDI. The lack of paired membranes and DMVs in cells transfected with the four CysSer mutations suggested that each of these conserved nsp3 residues plays a critical role in their formation.

EAV nsp3 is a transmembrane protein. In view of the importance for DMV formation of the conserved nsp3 Cys residues and the luminal domain in which they reside, we sought to obtain formal proof for their luminal localization. To this end, we replaced loop residue Thr-873 (located between the second and third conserved Cys residue) with Asn, thus engineering an Asn-X-Thr recognition signal for N-linked glycosylation. The functionality of the signal was tested by ³⁵S labeling of protein synthesis in transfected cells, first by expressing the nsp2-3/T873N mutant protein from the SinRep5 alphavirus expression vector (data not shown) and subsequently by expressing a full-length mutant pp1a/T873N using a T7 RNA polymerase-driven expression system (also see below). Following immunoprecipitation with an anti-nsp3 rabbit antiserum and SDS-PAGE, an approximately 2-kDa mobility shift was observed for nsp3 as well as for the nsp3-4 and nsp3-8 processing intermediates produced during autoprocessing of pp1a (Fig. 6).

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FIG. 6. Analysis of cleavage products of wt EAV pp1a and a mutant pp1a carrying the Thr-873Asn N glycosylation-inducing mutation, which was produced using the recombinant vaccinia virus-driven T7 expression system. Following 35S labeling and immunoprecipitation with an nsp3 antiserum, half of the precipitated proteins were treated with PGNaseF to remove N-linked sugars and proteins were separated by using a 10% SDS-PAGE gel. For nsp3 and the nsp3-4 and nsp3-8 intermediates of the T873N mutant, a mobility shift (bands labeled with asterisks) could be observed; the shift was reversed after the removal of N-linked glycans by PGNaseF. –, absence of; +, presence of.

To confirm that the mobility shift was indeed due to glycosylation of the Thr-873Asn-containing mutant nsp3, the immunoprecipitated wt and mutant proteins were treated with N-glycosidase F (PGNaseF). As anticipated, the treatment increased the mobility of the nsp3, nsp3-4, and nsp3-8 cleavage products of pp1a/T873N (Fig. 6, lane 4), which now again comigrated with the corresponding products of the wt control. The functionality of Asn-873 in N-linked glycosylation confirmed its predicted luminal localization and supported the hypothesis that pp1a residues 853 to 902 (Fig. 1B) form a luminal loop domain that also includes the four conserved Cys residues discussed above. In addition, this result constituted the first experimental evidence for the transmembrane nature of EAV nsp3.

Interestingly, when expressed using the SinRep5 vector, an intermediate phenotype was observed for the Thr-873Asn glycosylation mutant. By IFA, some concentration of the nsp2-3 labeling in the perinuclear region was observed (Fig. 2C), but overall the signal was more dispersed throughout the cell than that with the wt control labeling. Although in some EM profiles apparently normal DMVs could be observed, the presence of tightly apposed paired membranes that were arranged in extensive balloon-shaped structures was a striking deviation compared to the wt control and the samples of all other mutants in this study (Fig. 3D and 4C). Immuno-EM again revealed colocalization of nsp2 and nsp3 on these double-membrane structures, although a part of the label was found in regions containing less distinct single-membrane structures (Fig. 5E). Together, these images suggested that glycosylation of the nsp3 luminal domain somehow interfered with efficient DMV formation, possibly due to steric hindrance of the glycan in the context of interactions involving residues in the nsp3 loop domain.

Nsp3 mutations affect pp1a autoproteolysis. Immunoprecipitation after expression of a double-tagged nsp2-3 (HA-nsp23-His) protein from SinRep5 indicated that some of the Cys mutations (Cys-855Ser and Cys-864Ser) resulted in less efficient autoprocessing of the nsp2-3 site by the nsp2 CP (data not shown). To further investigate the influence of the nsp3 mutations on pp1a autoprocessing, the effects of the nsp2 CP and nsp3 loop mutations were studied by using a recombinant vaccinia virus-driven T7 RNA polymerase expression system (10, 51). This system was used in previous studies to dissect the complex processing cascade that leads to the cleavage of EAV pp1a (Fig. 7A). Cleavage products were immunoprecipitated using antisera recognizing nsp1, nsp3, nsp4, and nsp7/8, as described in Materials and Methods. An overview of the results is presented in Fig. 7.

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FIG. 7. The effect of nsp3 mutations on proteolytic processing of EAV pp1a. Replicase pp1a was expressed from vector pL1a using the recombinant vaccinia virus-driven T7 expression system and RK13 cells. Following 35S labeling, cleavage products were immunoprecipitated using a panel of antisera and separated in a 10% SDS-PAGE gel. (A) Schematic overview of the major and minor pp1a processing pathways of EAV pp1a (51). The interaction of cleaved nsp2 with nsp3 is a trigger for processing via the major pathway, which is initiated by cleavage of the nsp4/5 site. PCP, papain-like cysteine protease; MP, main protease. (B) Immunoprecipitation of nsp1, which was used to establish that the transfection efficiency was similar in all samples. (C to E) Analysis of pp1a cleavage products after immunoprecipitation with nsp3 (C), nsp4 (D), or nsp7-8 (E) antisera (-nsp). Indicated are the positions of the cleavage products, the size markers, and a novel cleavage product (arrowhead) produced by the four nsp3 Cys mutants, which was tentatively identified as nsp3-5 (see text). Bands marked with asterisks indicate products with altered mobility due to the presence of the Thr-873Asn glycosylation mutation in nsp3.

After the translation of pp1a, nsp1 rapidly cleaved itself from the replicase polyprotein (38) and the protein could be readily detected for all constructs tested (Fig. 7B). Cleavage of the nsp1/2 site was found not to be influenced by any of the mutations, and the approximately constant amounts of nsp1 that were detected confirmed that plasmid DNA transfection and pp1a expression had been equally efficient for all mutants and the wt control. After the release of nsp1, the nsp2 CP attacks the nsp2/3 site (38), resulting in the rapid release of nsp2. Further processing of pp1a is then catalyzed by the main protease (nsp4) and can follow two alternative pathways, the major or the minor pathway, depending on an interaction between cleaved nsp2 and the nsp3-8 cleavage product (Fig. 7A) (51). Nsp2 acts as a cofactor for cleavage of the nsp4/5 site, which is the first step in the "major" processing pathway. In the absence of (cleaved) nsp2, nsp3-8 processing will follow the minor pathway (Fig. 7A), in which the nsp4/5 site remains uncleaved and alternative downstream sites are targeted by the nsp4 protease.

The wt lanes in Fig. 7B to E show the main indicators for pp1a processing via the major pathway, since minor pathway products are produced only at low levels in the normal situation. These products are nsp1, nsp2, and nsp3-8, and subsequently the long-lived cleavage products nsp5-8, nsp5-7, and nsp3-4. Nsp2 has a strong interaction with nsp3 (38) and is therefore coprecipitated by antisera bringing down nsp3 or nsp3-containing intermediates (Fig. 7B to E). Figure 7E also shows some nsp7-8, the most prominent minor pathway product produced by the wt construct.

As anticipated, inactivation of the nsp2 CP by the His332Tyr active site mutation blocked the nsp2/3 cleavage (nsp2-3 intermediate shown in Fig. 7C). Due to a lack of cleaved nsp2, the major processing pathway was inactivated and also nsp3-8, nsp5-8, nsp5-7, and nsp3-4 were no longer detected. Instead nsp7-8 (minor pathway product) was overproduced (Fig. 7E), and a prominent nsp4-5 product could be observed (Fig. 7D).

The four nsp3 CysSer mutations modulated pp1a processing to variable degrees. The Cys-855 and, in particular, the Cys-864 mutants affected the efficiency of nsp2/3 cleavage (Fig. 7C). Although the Cys-855 mutant still yielded major pathway products, albeit at reduced levels, minor pathway indicators like nsp7-8 and nsp4-5 were also upregulated. The Cys-864 mutant displayed a clear defect in the further processing of the reduced amount of nsp3-8 that was produced. In contrast, the Cys-879 and Cys-884 mutants seemed more or less processing competent, producing all major pathway products and, for example, not the large amount of nsp4-5 seen for the other two Cys mutants (Fig. 7D). Still, the relatively large amounts of nsp3-8 that accumulated suggested that the kinetics of pp1a processing had been altered. Also, the coimmunoprecipitation of nsp2, and therefore likely the previously documented nsp2-3 interaction, seemed to be seriously affected. All four nsp3 Cys mutants produced a cleavage product (about 58 kDa) that had not been observed before. Its reactivity with the nsp3 and nsp4 antisera, but not the nsp7-8 antiserum, suggested that this product could be an nsp3-5 processing intermediate.

Finally, the Thr-873Asn nsp3 glycosylation mutant showed fairly normal pp1a processing, with nsp3 (Fig. 6) and nsp3-4 (Fig. 7C and D) showing the expected mobility shift due to glycosylation. Although again nsp3-8 seemed to accumulate (Fig. 7C and E), the coimmunoprecipitation of nsp2 was comparable to that observed for the wt control. Taken together, these studies confirmed the critical importance of nsp2/3 cleavage for the major processing pathway and revealed that mutations in the predicted nsp3 luminal loop can affect processing to variable degrees, with the Cys-855 and Cys-864 replacements causing the largest deviations from the wt pattern.

Reverse genetics confirms a critical role for nsp3 and the nsp2/3 cleavage in the EAV life cycle. Using site-directed mutagenesis and an EAV reverse genetics system, we investigated the role of nsp3 in the context of the complete virus life cycle. The importance of the nsp2/3 cleavage was probed by introducing the His-332Tyr nsp2 CP active site mutation into a full-length cDNA clone. In vitro-transcribed RNA was transfected into BHK-21 cells, and virus viability was monitored at different time points by using a dual-labeling IFA for replicase (nsp3) and the viral nucleocapsid (N) protein, which can be used as markers for the synthesis of genome and subgenomic mRNA7, respectively (47). Despite the fact that the nsp2/3 cleavage was nonessential for DMV formation (Fig. 2 to 4), the CP active site mutation was found to be lethal, as no signal for nsp3 and N protein could be detected in transfected cells, which were monitored up to 72 h posttransfection (data not shown). The absence of progeny virus was confirmed by a negative plaque assay when testing transfected cell culture supernatants harvested at 72 h posttransfection.

Using the same experimental design, we found that the single CysSer replacements of each of the four conserved nsp3 Cys residues (855, 864, 879, and 884) also abolished all viral RNA synthesis. Finally, the mutation specifying the Asn-873 glycosylation site in nsp3 was tested. Although cells positive by IFA were not detected for this mutant, some plaques were obtained when supernatant from this transfection was tested. To investigate the genotype of this virus, fresh BHK-21 cells were infected with the same transfection supernatant and intracellular RNA was isolated from these cells. Direct sequence analysis of an RT-PCR product covering the nsp3-coding region revealed pseudoreversion of the mutant AAC codon (Asn) to CAC (His). Despite the fact that there was no reversion to the wt ACC codon (Thr), this AC mutation inactivated the engineered nsp3 N glycosylation site, clearly suggesting a selection pressure against the original Thr873Asn mutation. Interestingly, replication of the Thr-873Asn nsp3 glycosylation mutant could also be rescued by the addition (immediately after transfection) of the N-linked glycosylation inhibitor tunicamycin (data not shown). This result further strengthened the hypothesis that the presence of a sugar moiety on nsp3 blocked efficient EAV replication, presumably because it interfered with DMV formation.

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Hijacking cellular membranes to facilitate nidovirus RNA synthesis. Viruses have adopted various strategies to coordinate the molecular interactions that are the basis for their replication in both time and intracellular space. The association of the RTC of positive-strand RNA viruses with (modified) intracellular membranes appears to be a striking example of such a strategy (1, 5, 19, 20, 22, 25, 31, 53). Specific viral replicase subunits are targeted to the membranes of particular cell organelles that are subsequently modified into characteristic structures or compartments, sometimes referred to as "viral factories," with which viral RNA synthesis is associated. The virus-induced membrane alterations can range from relatively simple spherular invaginations to elaborate networks of convoluted membranes and single-membrane vesicles or DMVs (24). For most groups of positive-strand RNA viruses, the morphogenesis, ultrastructure, and function of these replication-associated membrane structures are only beginning to be understood.

The subcellular localization of the RTC and the ultrastructure of the associated membranes have been studied in some detail for only three members of the order Nidovirales: the arterivirus EAV (28, 37, 46, 48), the coronavirus mouse hepatitis virus (6, 13, 29, 33, 45), and the severe acute respiratory syndrome coronavirus (SARS-CoV) (15, 16, 29, 36, 42). For all three viruses, the analysis of infected cells by confocal immunofluorescence microscopy revealed that the majority of replicase subunits (including the RdRp and helicase) colocalized in the perinuclear region. Also, the induction of unusual paired membranes and DMVs, which was observed in EM studies, appeared to be a common feature of cells infected with these three nidoviruses (11, 13, 28, 36, 42). For both SARS-CoV (36) and EAV (37), we have proposed that the membrane donor for these structures is the ER, a hypothesis that is firmly supported by recent electron tomography studies (K. Knoops, E. J. Snijder, et al., unpublished data). However, in particular for mouse hepatitis virus, the data have remained equivocal thus far and multiple intracellular compartments have been implicated in RTC formation, including the Golgi complex, endosomal membranes, ER, and autophagosomes (13, 29, 45).

Although the three coronavirus replicase subunits containing distinct hydrophobic domains (nsp3, nsp4, and nsp6) have been implicated in RTC formation and associated membrane modifications, these proteins remain to be characterized in detail and direct experimental evidence for this role has not been presented (18, 26, 41). Fortunately, for the corresponding arterivirus nsps, a direct assay to study their interaction with membranes became available when the expression of EAV nsp2-3 (28, 37) was found to suffice for the induction of paired membranes and DMVs that are very similar to those found upon EAV infection. Here we have used this expression system to identify replicase subunit nsp3 as a key player in the interaction of the arterivirus RTC with host cell membranes and in the virus life cycle as a whole. Site-directed mutagenesis of EAV nsp3 was found to dramatically affect the interaction of the protein with the ER, disrupt membrane pairing and DMV formation, interfere with replicase polyprotein processing, and block viral RNA synthesis. We also obtained the first direct evidence that the anchoring of the EAV RTC to membranes involves a multispanning transmembrane protein and that functionally important replicase domains reside on the luminal side of the membrane.

The mechanism of arterivirus DMV formation. Tightly paired membranes are commonly observed in nidovirus-infected cells and are generally considered to be the precursor of the abundantly formed DMVs. Most likely, nidovirus transmembrane nsps are first inserted into "regular" ER membranes, which may thus also be the site of early viral RNA synthesis. When replication leads to a rapid increase of replicase expression, interactions between the accumulating transmembrane nsps (and possibly also host proteins) may lead to membrane pairing and subsequent vesiculation, although the mechanism of the latter step remains one of the important unresolved issues (28).

Membrane pairing could be based on interactions between the luminal domains of transmembrane proteins of viral and/or host origin, which could span the lumen and thus bring opposing membranes together. In addition to the requirement for coexpression of nsp2 and nsp3 to induce DMVs (37), their direct involvement in membrane-modifying interactions is suggested by their previously documented interaction (38), their retention in the ER (46), and their abundant presence on the modified membranes (Fig. 5) (28, 37). In the EAV nsp2-3 sequence, residues 853 to 902 constitute the most prominent (predicted) luminal domain (Fig. 1), which also stands out for containing a number of arterivirus-wide conserved residues, including the four cysteines targeted here by mutagenesis. This study confirmed the luminal localization of this domain and showed that in the alphavirus-driven expression system, replacement of any of these Cys residues affected membrane pairing and DMV formation (Fig. 2 and 3). The expression of these mutants also appeared to have a dramatic effect on the overall condition of the ER, suggesting that the mutant proteins were extremely toxic to the cell, possibly due to a lack of interaction with their regular partner(s). This may explain the formation of large numbers of single-membrane vesicles and vacuoles. The interaction partner(s) of nsp3 and the nature of the interactions in which its luminal loop is engaged are the subject of ongoing investigations. Although disulfide bridge formation involving the four nsp3 Cys residues remains an obvious possibility, preliminary studies did not reveal this type of interaction within nsp3 or in the context of the previously documented nsp2-3 interaction (data not shown). Although the interaction between nsp2 and the nsp3 of the CysSer mutants was largely inhibited in the expression system (Fig. 7), it is difficult to assess whether this is a direct effect of the Cys replacements or an indirect effect resulting from, e.g., the overall changes in polyprotein processing (see below), conformational changes, or the toxicity of the mutant nsp3 proteins.

Given its multispanning transmembrane character, nsp3 is also a prime candidate for an essential role in the induction of membrane curvature that must precede DMV formation. Transmembrane nsps may induce curvature due to their specific structural features or by oligomerization. Alternatively, viral nsps might activate or recruit cellular factors involved in membrane bending. Recent studies of picornavirus-induced membrane alterations present a striking example of that strategy, documenting the involvement of various players with roles in cellular membrane trafficking (4, 52). The phenotype that was observed for the Thr-873Asn glycosylation mutant implicates the luminal loop of nsp3 directly in DMV formation. Although membrane pairing was unaffected and some regular DMVs were found, the highly curved double-membrane sheets depicted in Fig. 3D were unique for this mutant and suggested that the transition from double membranes to DMVs was somehow impaired by the presence of the sugar moiety. This observation seems most compatible with the previously proposed "protrusion-and-detachment model" for DMV formation (28). Possibly, the presence of the N-linked sugar at position 873 inhibits an interaction of nsp3 that is critical for detachment and/or sealing of the vesicles.

Coordinating translation, polyprotein processing, and membrane association. From our prior studies on EAV replicase processing, a complex interplay between three autoproteases and their substrates has emerged (54). The processing of pp1a is characterized by a combination of rapid and slow(er) cleavages and the use of two alternative pathways for nsp4-directed processing (51). The release of the replicase subunits containing (putative) transmembrane domains (nsp2, nsp3, and nsp5) is coordinated by the combined action of the nsp2 and nsp4 proteases, with nsp2 acting as cofactor for cleavage of the nsp4/5 site by nsp4. How replicase polyprotein synthesis, cleavage, and membrane insertion are coordinated in time and space remains to be resolved, but clearly the different proteolytic cleavages can be considered potential regulatory switches, as they were found to be in other virus groups. For example, for the flavivirus Kunjin virus, it was concluded that the cleavage of an NS4A-NS4B expression product is the key initiation event in the rearrangement of intracellular membranes (30). In the case of EAV particularly, the release of the N-terminal hydrophobic domains of nsp3 and nsp5 was anticipated to trigger the translocation of these subunits. However, this study showed that upon the expression of an nsp2-3 polyprotein, the functionality of nsp3 in DMV induction does not depend on prior processing of the nsp2/3 junction, suggesting that membrane insertion of nsp2-3 in fact precedes polyprotein cleavage. This finding also indicates that for the Cys-855Ser and Cys-864Ser mutations, the lack of DMV formation is unlikely to be due to their reduced processing of the nsp2/3 site (Fig. 7C).

These latter two mutations and, to a lesser extent, those at positions 879 and 884, clearly affected nsp4-driven processing (Fig. 7), in particular the major processing pathway that is initiated with cleavage of the nsp4/5 site. For the mutations at positions 855 and 864, the reduced cleavage of the nsp2/3 site, and therefore the lack of sufficient fully cleaved nsp2, may offer a straightforward explanation for this finding (51). The processing of pp1a was shifted to the minor pathway, a shift that was almost complete for the Cys-864Ser mutation. Given the fact that each of the nsp4-driven cleavages has been found to be essential for virus viability (44), it was not surprising that the Cys-855 and Cys-864 replacements were lethal to the virus when evaluated in the reverse genetics system. For the Cys-879 and Cys-884 replacements, the pp1a processing defects were somewhat less obvious, although the accumulation of nsp3-8 suggested a clear change in the kinetics of the downstream processing of this intermediate (Fig. 7). Although the Thr-873Asn nsp3 glycosylation mutant exhibited a similar defect, its general cleavage pattern approached that of the wt pp1a more closely, including the restoration of the nsp2-3 interaction, as evidenced by the coimmunoprecipitation of nsp2 depicted in Fig. 7C and E. Still, the introduction of this mutation also blocked EAV replication, although a pseudorevertant in which the glycosylation signal had been removed (Asn-873His) was picked up 3 days posttransfection. Apparently, the Thr-873Asn mutant virus was capable of a very low level of RNA synthesis that was the basis for the slow reversion, indicating that the original mutant had retained some RTC functionality, despite the fact that its DMV formation and pp1a processing were impaired.

The formation of a membrane-bound scaffold for the ORF1b-encoded core enzymes of the EAV RTC appears to be based on the interplay between ORF1a-encoded accessory protease(s), main proteases, and transmembrane domains, which all are nidovirus-wide conserved features. Although, as documented here, the integral study of this multifunctional part of the replicase requires a combination of polyprotein-based assays (to study, e.g., membrane alterations, proteolytic processing, and RTC functionality), this likely is the most productive approach to unravel the interactions and regulatory mechanisms directing this poorly understood, but well conserved, interaction between positive-strand RNA viruses and their hosts.

 
We thank Kèvin Knoops, Marjolein Kikkert, and Alexander Gorbalenya for technical assistance and/or helpful discussions.

 
* Corresponding author. Mailing address: Molecular Virology Laboratory, Department of Medical Microbiology, Leiden University Medical Center, LUMC P4-26, PO Box 9600, 2300 RC Leiden, The Netherlands. Phone: 31 71 5261657. Fax: 31 71 5266761. E-mail: e.j.snijder{at}lumc.nl

Published ahead of print on 27 February 2008.

These authors contributed equally to this work.

Present address: Invitrogen Dynal AS, Postboks 114 Smestad, 0309 Oslo, Norway.

Present address: Maxwell H. Gluck Equine Research Center, Department of Veterinary Science, University of Kentucky, Lexington, KY 40546.

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Journal of Virology, May 2008, p. 4480-4491, Vol. 82, No. 9
0022-538X/08/$08.00+0     doi:10.1128/JVI.02756-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

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