Nucleocapsid Protein Recruitment to Replication-Transcription Complexes Plays a Crucial Role in Coronaviral Life Cycle

N protein interacts with nsp3 (40, 41); therefore, we hypothesized that it is recruited to the RTCs by binding one or more nsp. We initially determined which components of the RTCs interact with the N protein, particularly whether other nsps, apart from nsp3, also bind the N protein, by using a yeast two-hybrid (Y2H) assay (42). For this purpose, the N protein of MHV and all 16 nsps were cloned into a vector carrying the activation domain (AD) or the DNA binding domain (BD), respectively, of the yeast Gal4 transcription factor (42). Each plasmid encoding a BD-nsp fusion protein was transformed together with the one expressing the AD-N chimera into the Y2H test strain, and growth of the cotransformed cells on a medium lacking histidine was used to assess interaction. As shown in Fig. 1B, this approach revealed that MHV N protein bound mainly to nsp3, in agreement with the previous findings (40, 41, 43), and to a far lesser extent to nsp4. Since this potential interaction was detected in only one replicate, we did not investigate this further.

Previous analyses revealed that the large cytosolic N-terminal part of nsp3, from amino acid 1 to amino acid 233, i.e., nsp3^N, is important for interaction with the N protein (40, 41). In those studies, however, purified nsp3^N was incubated with cell extracts from MHV-infected cells. As a result, this approach cannot exclude the possibility that N protein interacts indirectly with nsp3, i.e., via either a cellular protein or another viral protein, and thus a different factor could be involved in N protein recruitment to the RTCs. Therefore, we first examined whether the N protein and nsp3 bind directly, using in vitro binding experiments. The nsp3^N fragment was fused to glutathione transferase (GST), expressed in Escherichia coli, and purified using glutathione (GSH) conjugated to Sepharose beads. To prove that our purified nsp3^N fragment retained its overall conformation, GST or GST-nsp3^N was first incubated with cell extracts obtained from noninfected and MHV-infected LR7 cells. As shown in Fig. 1C and consistent with the literature (41), the N protein specifically bound to GST-nsp3^N but not to GST alone. We also tested whether association with RNA was influencing N protein binding to nsp3^N, because this parameter could be relevant for the follow-up in vitro studies with recombinant proteins. Therefore, cell extracts from infected cells were treated with RNase A or with this enzyme plus a specific RNase A inhibitor before being incubated with GST-nsp3^N or GST. Importantly, removal of RNA did not alter N protein interaction with nps3^N (Fig. 1C).

Next, we fused N protein with a 6×His tag and expressed this fusion protein in E. coli before preparing bacterial cell extracts and incubating them with either GST or GST-nsp3^N. As shown in Fig. 1D, recombinant N protein specifically bound to purified GST-nsp3^N but not GST. Altogether, these experiments show that nsp3, through its N terminus, is the only RTC component that binds directly to the N protein and this binding does not require N protein association with RNA.

To unravel the mechanism of N protein recruitment to the RTCs through binding via nsp3, we dissected how N protein interacts with nsp3. We generated a series of C-terminal-truncated variants of the N protein and assessed their interactions with nsp3 using the Y2H system, to identify the nsp3-binding regions of the N protein (Fig. 2A). Consistent with previous reports (40, 41, 44), we pinpointed one binding region in the N2a domain, between amino acid 233 and amino acid 250, which contains the SR-rich region. To elaborate further on this, we generated three truncated forms, i.e., N1a-N1b/NTD (amino acids 1 to 194), N2a (amino acids 195 to 257), and N2b-N3/CTD (amino acids 258 to 454) (Fig. 1A). These truncated forms were fused with the 6×His tag and expressed in E. coli before preparation of bacterial cell extracts, which were then incubated with either GST or GST-nsp3^N. As expected and as shown in Fig. 2B, none of the fusion proteins interacted with GST. In contrast, binding of GST-nsp3^N with 6×His-N1a-N1b was detected, as suggested previously (45), but not with 6×His-N2a or 6×His-N2b-N3. Because interaction between nsp3 and N2a was reported previously (40, 41) and was shown to possess weak affinity (44), we repeated the pulldown experiments with GST-nps3^N and 6×His-N2a and exposed the Western blot membranes for a longer time (Fig. 2C). This approach allowed detection of the binding between nps3^N and N2a, in agreement with our Y2H analysis of the truncated N proteins (Fig. 2A). This result shows that motifs in two different regions of the N protein, i.e., N1a-N1b and to a much smaller extent N2a, are mediating its binding to nsp3.

In order to identify these motifs and to study their relationship, we aligned the amino acid sequences of the N1-N2a segment of the N proteins from different betacoronaviruses and examined their distribution on the three-dimensional structure of the MHV N1-N2a domain (24). As highlighted in Fig. 3A, we found six stretches of amino acids that were both conserved and on the surface of the N1-N2a segment, thus making them available for interaction. Four of these amino acid stretches were localized in the N1 region and two in the N2a fragment. We changed the conserved polar and charged amino acids to alanines and the conserved nonpolar amino acids to aspartates in each one of these different sequences, creating mutated versions of the 6×His-tagged N protein truncated forms, i.e., N2a^m1 (amino acids 195 to 257, with L240E, V241E, and L242E substitutions), N2a^m2 (amino acids 195 to 257, with S197A, R202A, S203A, and S205A substitutions), N1^m3 (amino acids 1 to 194, with T73A, Q74A, and K77A substitutions), N1^m4 (amino acids 1 to 194, with K101A, Y103A, and W104A substitutions), N1^m5 (amino acids 1 to 194, with R109A, R110A, and K113A substitutions), and N1^m6 (amino acids 1 to 194, with F128A, Y129A, Y130A, and T133A substitutions) (Fig. 3A). Of note, the S205A change in the N2a^m2 variant alters the priming site that is essential for SR-rich region phosphorylation by host cell GSK-3 (32, 46). Cell extracts obtained from E. coli strains expressing these constructs were incubated with either GST or GST-nsp3^N. Although N1^m3 and N1^m5 bound GST-nsp3^N with the same affinity as wild-type N1, we found that the other mutated truncated forms displayed strong reductions in interaction with GST-nsp3^N (Fig. 3B). We concluded that amino acids at positions 101 to 104 and 129 to 133 and at positions 194 to 202 and 240 to 242 play roles in the interaction of nsp3 with N1 and N2a, respectively (Fig. 3B). This result also revealed that the SR-rich region that is phosphorylated in vivo is not important for N protein binding to nsp3.

This finding indicated two possible scenarios. In the first, the two identified nsp3 BDs act cooperatively, and disruption of one of them is sufficient to abolish the nsp3-N protein interaction. In the second, the two identified nsp3 BDs are redundant, and mutation of only one of them does not affect the association between nsp3 and N protein. Thus, to determine whether the single mutations in N1 or N2a are sufficient to block the N1-N2a interaction with nsp3^N, we expressed the N1-N2a^m1, N1-N2a^m2, N1-N2a^m4, and N1-N2a^m6 variants in E. coli before incubation of bacterial extracts with either GST or GST-nsp3^N. As shown in Fig. 3C, all constructs specifically bound GST-nsp3^N with the same affinity as wild-type N1-N2a. This finding showed that the binding of MHV N protein to nsp3 is not mediated by a single domain and thus the domains might be redundant in the binding function, as contemplated in the second scenario.

Consequently, we next assessed whether the combined mutations in the critical amino acids of N1b and N2a could block N protein-nsp3 interaction. For this purpose, we expressed full-length N, N^m4m1, N^m6m1, N^m4m2, or N^m6m2 in E. coli and incubated bacterial extracts with either GST or GST-nsp3^N. Although N^m4m2 and N^m6m2 were still able to interact with nsp3^N, N^m4m1 and N^m6m1 displayed strong reductions in their binding to nsp3^N (Fig. 4A). This finding shows that two different regions mediate MHV N protein binding to nsp3. From here, we continued with the analysis of the N^m4m1 and N^m6m1 mutants to specifically study the N protein-nsp3 interaction.

We and others recently showed that CoV N proteins form oligomers constitutively (22, 23). Therefore, a possible scenario that could not be excluded a priori was that the introduced substitutions in the N protein were affecting its organization into oligomers, which might indirectly impair binding to nsp3. To investigate whether the mutated N proteins that showed reduced nsp3 binding, i.e., N^m4m1 and N^m6m1 (Fig. 4A), had a defect in self-interaction, we analyzed their self-binding. Bacterial extracts from E. coli expressing 6×His-tagged N, N^m4m1, or N^m6m1 were incubated with GST or GST-N protein. As shown in Fig. 4B, 6×His-N, 6×His-N^m4m1, and 6×His-N^m6m1 proteins specifically interacted with GST-N protein.

To ascertain whether N^m4m1 and N^m6m1 proteins were also able to oligomerize, purified 6×His-N, 6×His-N^m4m1, and 6×His-N^m6m1 were resolved on a glycerol density gradient to determine the size of the oligomers that they formed (23). Wild-type N protein was detected mainly in fractions 4 to 6, as expected (Fig. 4D) (23). The distribution of the N^m4m1 oligomer over the gradient differed slightly, compared to that of the wild-type oligomer, i.e., the signal was present mainly in fractions 6 and 7, suggesting that the protein assembled predominately in larger oligomers. Importantly, the oligomerization of the N^m6m1 variant was similar to that of the N protein, as the oligomer was detected mainly in fractions 4 to 7 (Fig. 4C and D). These results show that the regions involved in MHV N protein binding to nsp3 are not relevant for oligomerization.

To establish whether the inability of the N protein to interact with nsp3 led to altered subcellular localization over the course of an infection, we first attempted to generate mutant MHV strains using two different approaches. First, we opted to generate recombinant MHV expressing an additional copy of green fluorescent protein (GFP)-tagged N or N^m6m1 by inserting the coding sequences of these two fusion proteins into the viral locus of the nonessential hemagglutinin esterase gene (39). The N^m6m1 mutant was chosen because it oligomerizes similarly to the wild-type N protein (Fig. 4D). We tried three times to generate an MHV-N^m6m1-GFP chimeric strain using this approach but, although we obtained the control MHV-N-GFP strain each time, we were unable to recover the mutant virus. Second, we turned to a vaccinia virus-based reverse genetics system to introduce the N^m6m1 mutation in the viral N locus (47). With this approach as well, the wild-type strain but not the mutant strain was obtained in two attempts. Because genomic expression of N^m6m1 appeared to be lethal for the virus, we decided to opt for a different strategy. We expressed mCherry, mCherry-N, or mCherry-N^m6m1 fusion proteins in mouse LR7 cells before exposing the cells to MHV. mCherry, mCherry-N, and mCherry-N^m6m1 were homogenously distributed in the cytoplasm in noninfected cells (data not shown). In contrast, MHV-infected cells displayed numerous distinct cytoplasmic puncta that were positive for both nsp2 and nsp3 (data not shown); these puncta were previously shown to represent RTCs (39). Interestingly, wild-type mCherry-N but not mCherry-N^m6m1 or mCherry alone was recruited to these nsp2/nsp3-positive RTCs (data not shown).

The low transfection efficiency of mouse LR7 cells (around 10% to 15%) did not allow examination of a sufficiently large number of cells that were both infected and expressing the analyzed chimeras for quantification. Therefore, we decided to turn to the human HeLa-CEACAM1a cell line, which expresses the MHV cell receptor CEACAM1a and is highly transfectable (48). As expected, the GFP, GFP-N, and GFP-N^m6m1 constructs were efficiently transfected into HeLa-CEACAM1a cells, with around 80% of the cells displaying the fluorescent signal. In agreement with the results obtained with LR7 cells, GFP, GFP-N, and GFP-N^m6m1 fusion proteins were homogenously distributed throughout the cytoplasm (Fig. 5A) and MHV infection caused a redistribution of GFP-N to the nsp2/nsp3-positive RTCs (Fig. 5A, fifth row). This observation confirmed that ectopically expressed GFP-tagged N protein behaved like the protein encoded by the viral gRNA over the course of an infection. Surprisingly, we found that a large number of cells expressing GFP-N^m6m1 did not display nsp2/nsp3-positive RTCs (Fig. 5B), indicating that this construct has a dominant negative effect on viral replication. In a very few cases (approximately 4% of the cells expressing GFP-N^m6m1), RTCs were present in the same number and had a similar size as in cells expressing GFP-N (data not shown) but GFP-N^m6m1 was not recruited onto the RTCs (Fig. 5A and C). Altogether, these results show that N protein binding to nsp3 is required for its recruitment to the RTCs and possibly also for MHV RNA synthesis and replication.

To determine whether blocking N protein recruitment to the RTCs led to a defect in the CoV life cycle, we performed a series of experiments to examine whether N^m6m1 indeed had a dominant negative effect on MHV replication and/or assembly/egression. We took advantage of the MHV-2aFLS strain, which carries the genes encoding firefly luciferase and allows assessment of viral replication by monitoring the activity of this enzyme (49). We transfected the plasmids expressing GFP, GFP-N, or GFP-N^m6m1 fusion protein into HeLa-CEACAM1a cells for 48 h before exposing the cells to different multiplicities of infection (MOIs) of MHV-2aFLS, i.e., MOIs of 2.5, 5, and 10, and measuring luciferase activity at 7 h postinfection (p.i.). As shown in Fig. 6A, expression of wild-type N protein promoted viral replication over time after exposure to different amounts of virus, as expected (26–29). Very interestingly, there was a significant decrease in luciferase expression in cells transfected with GFP-N^m6m1, compared to those expressing GFP-N as well as those expressing GFP alone, showing that GFP-N^m6m1 cannot promote viral replication like the wild-type protein; on the contrary, it has a dominant negative effect. Using the same approach, we explored whether the presence of N^m6m1 had a permanent negative effect on viral replication or was just retarding it. For this purpose, cells transfected with the various constructs were infected with MHV-2aFLS at a MOI of 5 and luciferase activity was measured at 5, 7, and 9 h p.i. (Fig. 6B and data not shown). In this experiment as well, the GFP-N chimera stimulated replication over time and at all the examined time points, while GFP-N^m6m1 had a pronounced inhibitory effect on MHV replication.

To confirm these observations using a wild-type MHV strain, we explored the production of viral N protein by Western blotting at different time points after inoculation with the MHV-A59 strain. HeLa-CEACAM1a cells were transfected with the plasmids carrying GFP, GFP-N, or GFP-N^m6m1 for 48 h before being infected with MHV for 5, 7, or 9 h (Fig. 6C and D and data not shown). Viral N protein synthesis was again stimulated by the presence of ectopically transfected GFP-N over time but not by GFP-N^m6m1. In the same experiment, we also collected the culture supernatants of cells and determined the virus titers, to explore whether MHV particle production and egression were impaired in cells expressing GFP-N^m6m1. This was indeed the case, as the virion titers were decreased in cells expressing GFP-N^m6m1, compared to those in cells expressing GFP or GFP-N (Fig. 6E). Altogether, these results show that N^m6m1 has a dominant negative effect on the MHV life cycle.

The principal role of CoV RTCs is in the replication and transcription of viral RNA. It remains unknown, however, whether the specific N protein recruitment to the RTCs has a regulatory role in viral RNA synthesis and whether it influences gRNA and sgmRNA transcription differently.

To examine the specific relevance of the N protein-nsp3 interaction in the transcription of gRNA and sgmRNA, HeLa-CEACAM1a cells that had been transfected with the plasmids expressing GFP, GFP-N, or GFP-N^m6m1 for 48 h were inoculated with MHV-A59 for 5, 7, or 9 h (Fig. 7A and data not shown) before the RNA was extracted. The mRNA was quantified by real-time (RT)-PCR using sets of primers that allowed specific assessment of the synthesis of gRNA and of four sgmRNA transcripts (Table 1). Expression of GFP-N stimulated the synthesis of both gRNA and sgmRNA, in comparison to cells expressing GFP only, over time and at all of the examined time points (Fig. 7A and data not shown). In contrast, GFP-N^m6m1 expression was unable upregulate gRNA and sgmRNA synthesis. This result shows that N protein association with RTCs stimulates the replication and transcription activities of these complexes.

To prove this observation with a different approach, we turned to an in vitro viral RNA synthesis (IVRS) assay using a membrane fraction enriched in RTCs, which has recently been devised to study the host factor requirements for sgmRNA transcription by SARS CoV RTCs (50). An advantage of in vitro assays is that proteins and other components can be easily added, to determine their effects on viral RNA synthesis. First, osmotically lysed MHV-infected cells were centrifuged at 13,000 × g to separate dense membranes containing RTCs (13,000 × g pellet [P13]) from the cytosol (13,000 × g supernatant [S13]). As expected, the ER integral membrane protein VAP-A was found entirely in the P13 fraction, while cytoplasmic glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was found mainly in the S13 fraction (Fig. 7B). To confirm the enrichment of RTCs in the P13 fraction, we probed the samples with anti-N antibodies and, as expected (50), RTCs were exclusively detected in the P13 fraction. Next, we tested the best experimental conditions for detection of RNA synthesis by MHV RTCs in vitro. Similar to the protocol established for SARS CoV (50), the P13 fractions from MHV-infected and noninfected cells were mixed with nucleoside triphosphates, an ATP-regenerating system, and actinomycin D, a potent inhibitor of host cellular transcription. To assess the synthesis of both sgmRNA and gRNA, we employed quantitative RT-PCR instead of labeling using radioactive nucleoside triphosphates (50). The IVRS assays showed that viral RNA was more effectively synthesized when the P13 fraction from MHV-infected cells (P13-MHV) was present in the reaction mixtures (Fig. 7C). Since it was shown that host factors that are present in the S13 fraction are required for viral RNA synthesis in vitro (50), we added S13 from infected cells and compared the viral RNA synthesis rate with that of samples not containing S13 (Fig. 7C, column 2 versus column 3). Interestingly, we did not observe an enhancement of viral RNA synthesis in the presence of S13 from infected cells, indicating that the P13 fraction was sufficient to sustain IVRS (Fig. 7C). Subsequently we explored whether adding different amounts of N protein to the IVRS assay enhanced viral transcription. As shown in Fig. 7D, we found that 1 μg of protein extract from N protein-expressing bacteria provided the maximal stimulation of MHV RTC-mediated viral RNA synthesis. Smaller amounts of the bacterial extracts had less pronounced effects, while larger amounts had an inhibitory impact.

Finally, we carried out the IVRS assay in the presence of 1 μg of bacterial extract from E. coli cells expressing equal amounts of N or N^m6m1 protein. Although wild-type N protein was able to enhance viral RNA synthesis, as shown in Fig. 7A, the addition of N^m6m1 did not increase the production of viral RNA (Fig. 7E). This result supports the notion that the N protein interaction with nsp3 stimulates the RNA synthesis activity of MHV RTCs.

LR7 (52) and HeLa-CEACAM1a (48) cells were maintained in Dulbecco’s modified Eagle medium (DMEM) (Cambrex Bioscience, Walkersville, MD) supplemented with 10% fetal calf serum (Bodinco Alkmaar, The Netherlands), 100 IU/ml penicillin, and 100 μg/ml streptomycin (both from Life Technologies, Rochester, NY). HeLa-CEACAM1a cells were cultured in the presence of 800 μg/ml Geneticin (G418 sulfate; Gibco, Waltham, MA). Wild-type MHV-A59 was propagated in LR7 cells in DMEM. The MHV strain carrying luciferase that was used in our study was MHV-2aFLS (49). Virus titers of the culture supernatants were determined on LR7 cells with a dilution factor of 3, and the 50% tissue culture infectious dose (TCID₅₀) units per milliliter of supernatant were calculated according to the Reed-Muench method (53). TCID₅₀ per milliliter values were then converted into PFU per microliter as follows: TCID₅₀ per milliliter × 0.00069 (54).

The pGBDU plasmids carrying the 16 MHV nsps and the pGAD vectors expressing the MHV N protein or its truncated variants were generated by PCR of MHV cDNA, using appropriate primers, and subsequent cloning into the pGAD-C1 and pGBDU-C1 vectors, respectively (42). The sequences coding for either full-length MHV N protein or truncated forms, i.e., N1 (amino acids 1 to 194), N2a (amino acids 195 to 257), and N2b-N3 (amino acids 258 to 454), were amplified from MHV cDNA by PCR and cloned into the pET32c vector (EMD Millipore, Amsterdam, The Netherlands) using BamHI and XhoI, creating the pET32c-N, pET32C-N1, pET32C-N2a, and pET32C-N2b-N3 plasmids, which express the 6×His-N, 6×His-N1, 6×His-N2a, and 6×His-N2b-N3 fusion proteins, respectively. The sequence coding for MHV nsp3^N was amplified from MHV cDNA by PCR and cloned into the pGEX vector (GE Healthcare, Little Chalfont, UK) using BamHI and XhoI, creating the pGEX-nsp3^N construct, which expresses the GST-nsp3^N chimera. The mutated N proteins were created from the pET32c-N construct by PCR, which generated the pET32c-N2a^m1 (amino acids 195 to 257, carrying the L240E, V241E, and L242E substitutions), pET32c-N2a^m2 (amino acids 195 to 257, carrying the S197A, R202A, S203A, and S205A substitutions), pET32c-N1^m3 (amino acids 1 to 194, carrying the T73A, Q74A, and K77A substitutions), pET32c-N1^m4 (amino acids 1 to 194, carrying the K101A, Y103A, and W104A substitutions), pET32c-N1^m5 (amino acids 1 to 194, carrying the R109A, R110A, and K113A substitutions), pET32c-N1^m6 (amino acids 1 to 194, carrying the F128A, Y129A, Y130A, and T133A substitutions), pET32c-N^m4m1 (amino acids 1 to 454, carrying the L240E, V241E, L242E, K101A, Y103A, and W104A substitutions), pET32c-N^m6m1 (amino acids 1 to 454, carrying the L240E, V241E, L242E, F128A, Y129A, Y130A, and T133A substitutions), pET32c-N^m4m2 (amino acids 1 to 454, carrying the S197A, R202A, S203A, S205A, K101A, Y103A, and W104A substitutions), and pET32c-N^m6m2 (amino acids 1 to 454, carrying the S197A, R202A, S203A, S205A, F128A, Y129A, Y130A, and T133A substitutions) plasmids (Fig. 1A and 3A). To express mCherry-tagged versions of the different N protein mutants in cells, the GFP coding sequence in the pcDNA5-GFP vector was replaced with the mCherry sequence, using StrI and BsrGI, to create pcDNA5-mCherry. PCR-generated N, N^m4m1, N^m6m1, N^m4m2, and N^m6m2 sequences were subsequently inserted into pcDNA5-mCherry as HindIII-BamHI fragments, generating pcDNA5-mCherry-N, pcDNA5-mCherry-N^m4m1, pcDNA5-mCherry-N^m6m1, pcDNA5-mCherry-N^m4m2, and pcDNA5-mCherry-N^m6m2, respectively. To express GFP-tagged versions of the mutant N proteins in cells, GFP was cloned into the pcDNA3.1 vector as a XhoI-BamHI fragment, creating pcDNA3.1-GFP. PCR-generated N, N^m4m1, N^m6m1, N^m4m2, and N^m6m2 were subsequently inserted into pcDNA3.1-GFP as BamHI-HindIII fragments, generating pcDNA3.1-GFP-N, pcDNA3.1-GFP-N^m4m1, pcDNA3.1-GFP-N^m6m1, pcDNA3.1-GFP-N^m4m2, and pcDNA3.1-GFP-N^m6m2, respectively. All point mutations were verified by DNA sequencing.

Escherichia coli BL-21 cells carrying plasmids expressing the various GST- or 6×His-tagged fusion proteins were grown in 125 ml of LB medium (0.5% yeast extract, 1% tryptone, 1% NaCl) to a late exponential phase. After induction of protein expression with the addition of 0.5 mM isopropyl-β-d-thiogalactopyranoside, cells were grown at 37°C or 20°C for 4 h or 16 h. Bacteria were harvested, resuspended in 4 ml of lysis buffer (phosphate-buffered saline [PBS] [137 mM NaCl, 10 mM phosphate [pH 7.4], 2.7 mM KCl] containing 5 mM dithiothreitol [DTT], 1 mg/ml lysozyme, 1 mM phenylmethylsulfonyl fluoride [PMSF], 10% glycerol, 1% Triton X-100, and complete protease inhibitor [Roche, Basel Switzerland]), and lysed by two 10-s rounds of sonication using a Branson sonicator (Brandon, Danbury, CT). The bacterial lysates were cleared by centrifugation at 13,000 rpm for 10 min at 4°C. The concentration of 6×His-tagged proteins was measured using a bicinchoninic acid (BCA) kit (Promega, Madison, WI). Proteins were not further purified, and bacterial lysates were stored at −80°C until used. For purification of GST fusion proteins, lysates were incubated with 125 μl of GSH-Sepharose (GE Healthcare), which had been prewashed in PBS on a rotatory wheel at 4°C for 2 h.

For the preparation of cell extracts, LR7 cells were grown in 10-cm dishes before being either mock treated or inoculated with MHV at an MOI of 1. After 8 h, cells were lysed by a 5-min sonication in 1.2 ml of lysis buffer, and supernatants were cleared by centrifugation at 13,000 rpm for 10 min at 4°C. RNase A (Invitrogen, Carlsbad, CA) treatments were carried out by incubating 200 μl of cell extract with 2.5 U of enzyme on ice for 30 min, immediately prior to pulldowns.

The Saccharomyces cerevisiae test strain PJ69 (MATa trpl-901 leu2-3,112 ura3-52 his3-200 gal4Δ gal80Δ LYS2::GALl-HIS3 GAL2-ADE2 met2::GAL7-lacZ) was used for the assay (42). The prey (AD) and bait (BD) vectors were cotransformed into the PJ69 strain using lithium acetate. Cotransformed colonies were selected on synthetic minimal medium (SMD) (0.67% yeast nitrogen base without amino acids, 2% glucose, and auxotrophic amino acids as needed) lacking uracil and leucine before they were spotted on SMD medium lacking uracil, leucine, and histidine to determine whether the tested proteins interact. The combination of empty pGAD and pGBDU vectors was used as a negative growth control (42), while the combination of pGAD-Atg19 and pGBDU-Atg11 was the control for a positive interaction (55).

For the pulldown experiments, GSH-Sepharose-bound GST fusion proteins were incubated with 200 μl of bacterial extract or 200 μl of LR7 cell extract on a rotatory wheel at 4°C for 2 h and subsequently were washed three times with PBS supplemented with 5 mM DTT, 10% glycerol, and 1% Triton X-100 and one time with PBS at 4°C. Proteins bound to the Sepharose beads were eluted with 20 μl of sample buffer (65.8 mM Tris-HCl [pH 6.8], 26.3% glycerol, 2.1% SDS, 0.01% bromophenol blue) by boiling, subjected to SDS-PAGE, blotted onto polyvinylidene difluoride (PVDF) membranes, and visualized by either membrane staining with Ponceau Red or Western blotting using an anti-6×His monoclonal antibody (product no. HIS H8; Thermo Fisher, Waltham, MA) or an anti-N protein monoclonal antibody (a kind gift of Stuart Siddell, University of Bristol) (56). Bound primary antibodies were detected using an Alexa 680-conjugated goat polyclonal anti-mouse IgG antibody (Life Technologies), and signals were visualized with an Odyssey imaging system (LI-COR, Lincoln, NE).

Bacterial extracts from E. coli cells expressing 6×His-tagged N, N^m4m1, N^m6m1, N^m4m2, or N^m6m2 fusion proteins were loaded on top of a 2.2-ml continuous glycerol gradient of 15 to 40% (wt/vol) in lysis buffer, prepared using a Gradient Master device (BioComp, Canada). After centrifugation at 135,000 × g for 75 min at 4°C in a TLS55 rotor (Beckman Coulter, Brea, CA), 11 fractions of 200 μl were collected from the top to the bottom of the gradient. After precipitation with the addition of 20 μl of trichloroacetic acid (final concentration, 10%), proteins were resolved by SDS-PAGE and analyzed by Western blotting using monoclonal antibodies against the MHV N protein.

HeLa-CEACAM1a or LR7 cells were grown on 12-mm coverslips, transfected, and infected before being fixed with 4% paraformaldehyde at the indicated p.i. times. After permeabilization using 0.2% Triton X-100 and subsequent blocking with PBS containing 1% fetal calf serum, viral nsp2 and nsp3 were detected using anti-nsp2/nsp3 antiserum, a kind gift of Susan Baker (57), followed by incubation with secondary antibody conjugated to either Alexa 488 or Alexa 568 (Life Technologies). Fluorescence signals were captured with a Leica sp8 confocal microscope (Leica, Wetzlar, Germany).

HeLa-CEACAM1a cells transfected in 96-well plates were infected with the MHV-2aFLS strain at the indicated MOI and for the indicated times, washed with PBS, and incubated with 50 μl of lysis buffer (firefly luciferase flash assay kit; Thermo Fisher) at room temperature for 15 min; the cell lysates were then stored at at −20°C. Subsequently, 25-μl aliquots of thawed cell lysates were used to assess firefly luciferase expression using the firefly luciferase flash assay kit (58). Enzymatic activities were measured with a GloMax multidetection system (Promega), using 25 μl of substrate, a 2-s delay, and 10-s measurements. Background luminescence was subtracted from each value obtained, and the results were normalized to infected cells transfected with an empty vector expressing only GFP.

HeLa-CEACAM1a cells grown in 6-well plates were infected with the MHV-A59 strain at an MOI of 5 for the indicated times, washed with cold PBS, harvested in 100 μl of 2× sample buffer on ice for 30 min, sonicated for 1 min, and boiled. Equal protein amounts were separated by SDS-PAGE and, after Western blotting, proteins were detected using specific antibodies against MHV N protein, GFP (Roche), and β-actin (Merck Millipore, Burlington, MA), with the Odyssey imaging system. Protein signal intensities were normalized and quantified using ImageJ software (NIH, Bethesda, MD).

HeLa-CEACAM1a cells grown in 96-well plates were washed with ice-cold PBS and lysed with 30 μl of lysis buffer containing DNase I for 5 min at room temperature before the addition of 3 μl of stop solution for 3 min at room temperature, according to the manufacturer’s protocol (Power SYBR green Cells-to-CT kit; Thermo Fisher). Reverse transcription of the mRNA using a poly(T) primer, cDNA synthesis, and quantitative PCR were performed in a CFX Connect thermocycler (Bio-Rad, Berkeley, CA). The primers used are described in Table 1. The levels of gRNA and the analyzed sgmRNAs were normalized to that of GAPDH according to the comparative cycle threshold method used for quantification, as recommended by the manufacturer’s protocol.

MHV- or mock-infected LR7 cells grown on five different 15-cm dishes (approximately 1.5 × 10⁷ cells/dish) were harvested by trypsinization at 8 h p.i. To inhibit cellular transcription, 2 μg/ml actinomycin D (Sigma, Saint Louis, MO) was present in all solutions used for harvesting and washing of the cells. After washing with PBS, cells were resuspended in 2 ml of ice-cold hypotonic buffer (20 mM HEPES [pH 7.4], 10 mM KCl, 1.5 mM magnesium acetate [MgOAc₂], 1 mM DTT, 133 U/ml RNase A inhibitor [Promega]) and incubated at 4°C for 10 min. Cells were disrupted using a Dounce homogenizer, with 30 strokes with a tight-fitting pestle. Isotonic conditions were restored by adding HEPES, sucrose, and DTT, which resulted in a final lysate containing 35 mM HEPES [pH 7.4], 250 mM sucrose, 8 mM KCl, 2.5 mM DTT, 1 mM MgOAc₂, 2 μg/ml actinomycin D, and 130 U/ml RNase A inhibitor. Nuclei, large debris, and any remaining intact cells were removed by two successive centrifugations at 1,000 × g for 5 min, and the resulting postnuclear supernatant (PNS) was either assayed immediately for RTC activity or stored at −80°C. P13 and S13 fractions were prepared from the PNS by centrifugation at 13,000 × g for 10 min at 4°C. The P13 was resuspended in dilution buffer (35 mM HEPES [pH 7.4], 250 mM sucrose, 8 mM KCl, 2.5 mM DTT, 1 mM MgOAc₂, 2 μg/ml actinomycin D, 130 U/ml RNase A inhibitor), in one-ninth of the original PNS volume from which the pellet had been prepared.

IVRS assay mixtures were prepared by mixing 15 μl of P13 and 30 μl of S13, supplemented with or without different amounts of bacterial extracts from E. coli cells expressing 6×His-N or 6×His-N^m6m1. Identical levels of expression of these two fusion proteins in bacterial extracts used for the IVRS assays were verified by Western blotting using an anti-6×His monoclonal antibody. Pure proteins could not be used because they were unstable and thus were partially degraded during the purification procedure. The total volume of each mixture was adjusted to 75 μl with dilution buffer, and two 25-μl aliquots were made from this mixture; 5.6 μl of 100 mM creatine phosphate (Sigma; final concentration, 20 mM) and 0.28 μl of 1,000 U/ml creatine phosphokinase (Sigma; final concentration, 10 U/ml) were added to both of these aliquots. The IVRS assay was initiated in one of these aliquots (reaction sample) with the addition of nucleoside triphosphates (final concentrations of 1 mM ATP, 0.25 mM GTP, 0.25 mM UTP, and 0.18 mM CTP). The second aliquot served as a control (background sample), and dilution buffer was added instead of the nucleoside triphosphates. The IVRS assays were carried out at 30°C for 100 min before being stopped with the addition first of 30 μl of lysis buffer containing DNase I at room temperature for 5 min and then of 3 μl of stop solution for 3 min at the same temperature (both solutions from the Power SYBR green Cells-to-CT kit; Thermo Fisher). Reverse transcription of the RNA, cDNA synthesis, and quantitative PCR were performed in a CFX Connect thermocycler using appropriate primers. The expression levels of the S gene, which is part of the viral RNA (both sgmRNA and gRNA), were normalized to that of GAPDH in each sample, according to the comparative cycle threshold method used for quantification, as recommended by the manufacturer’s protocol. Finally, the value of each background sample was subtracted from the value of the corresponding reaction sample to determine the difference due to in vitro S gene synthesis.

Statistical significance was evaluated using the two-tailed heteroscedastic t test before P values were calculated. Individual data points from each independent experiment were used for the calculation of significance. The number of independent experiments is indicated in each figure legend.