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Arenaviruses are enveloped, negative-strand RNA viruses. For several arenaviruses, virus-like particle (VLP) formation requires the viral matrix Z protein. However, the mechanism by which viral ribonucleoprotein complexes are incorporated into virions is poorly understood. Here, we show that the expression of the Z protein and nucleoprotein (NP) of Mopeia virus, a close relative of the pathogenic Lassa virus, resulted in the highly selective incorporation of the NP protein into Z protein-induced VLPs. Moreover, the Z protein promoted the association of NP with cellular membranes, suggesting that the association of NP, Z, and the cellular membranes may facilitate the efficient incorporation of NP into VLPs. By employing a series of NP deletion constructs and testing their VLP incorporation, we further demonstrated an important role for the C-terminal half of NP in its incorporation into VLPs.
The members of the family Arenaviridae are zoonotic viruses found in various rodent species, which are the natural reservoir of these viruses (http://www.cdc.gov/ncidod/dvrd/spb/mnpages/dispages/arena.htm). Of 22 recognized arenaviruses, 9 cause human illnesses, often with severe consequences (7). The diseases caused by these viruses, which include Lassa, Junin, Machupo, and Guanarito hemorrhagic fever viruses, as well as lymphocytic choriomeningitis virus (LCMV), are a major public health problem in areas of South America and West Africa (http://www.cdc.gov/ncidod/dvrd/spb/mnpages/dispages/arena.htm). In fact, a novel highly pathogenic arenavirus, Lujo virus, was recently isolated during a Lassa fever-like outbreak in South Africa (2).
Arenaviruses are enveloped viruses that possess a bisegmented, single-stranded, ambisense RNA genome (reviewed in references 3 and 21). The large segment (L segment) encodes two proteins, the viral matrix protein Z, which has both regulatory and structural roles in the viral life cycle (1, 5, 8, 9, 13, 19, 26, 31), and the viral RNA-dependent RNA polymerase (L). The small segment (S segment) encodes two major structural proteins, the external envelope glycoprotein precursor (GPC) and the internal nucleoprotein (NP) (25). The viral glycoprotein mediates viral attachment to the cellular receptors and subsequent cell entry. NP, the most abundant viral protein, encapsidates the viral genome to form viral ribonucleoprotein (RNP) (vRNP) complexes together with the L protein (28).
Viral assembly and budding are critical steps in the viral life cycle. For several enveloped negative-strand RNA viruses, viral matrix proteins play an important role in the recruitment of viral nucleocapsids to the budding sites, in the formation of virus particles, and in the incorporation of nucleocapsids into these virions (10, 14-16, 23, 37-40). For some arenaviruses, such as LCMV and Lassa virus, the expression of the viral matrix Z protein induces the formation of virus-like particles (VLPs) (26, 27, 33, 34) into which viral RNPs and/or NP are incorporated (6, 27), presumably via the interaction of NP with Z (6, 11, 31). These findings suggest a potential role for the arenavirus Z protein in NP virion incorporation. However, the specificity of the NP incorporation into arenavirus particles through the NP-Z interaction and the NP region(s) responsible for this process remain unknown. We therefore studied Mopeia virus (a close relative of Lassa virus) virion formation in cells expressing Z, or wild-type or mutant NP and Z, by investigating their colocalization, membrane association, and NP virion incorporation.
Human embryonic kidney (HEK) 293T cells and African green monkey (Vero) cells were maintained in Dulbecco's modified Eagle's medium and minimal essential medium, respectively, supplemented with 10% fetal bovine serum (FBS), l-glutamine, and penicillin-streptomycin solution. Cells were maintained at 37°C with 5% CO2.
Mouse monoclonal antibody to the hemagglutinin (HA) tag (Covance, Princeton, NJ), mouse monoclonal and rabbit polyclonal antibodies to the FLAG tag (Sigma, St. Louis, MO), rabbit anti-calnexin antibody (Santa Cruz Biotechnology, Santa Cruz, CA), and rabbit anti-enhanced green fluorescent protein (eGFP) (Sigma, St. Luis, MO) were used according to the manufacturers' instructions.
Viral RNA isolated from Vero cells infected with Mopeia virus (AN20410) or Lassa virus (Josiah) and the culture supernatant of BHK cells infected with LCMV (Armstrong) served as a template for reverse transcription (RT)-PCRs. The Mopeia virus Z gene was amplified by RT-PCR using a forward primer (5′-GCGC CCCGGG ATG GGG AAA ACG CAG TCC AAG G-3′) and a reverse primer (5′-GCGC CTCGAG TCA GGG GCT GTA GGG TGG-3′), Mopeia virus NP cDNA was amplified by using a forward primer (5′-GCGC GAATTC ATG TCC AAT TCA AAG GAG GTG AAG TCC TTC TTG-3′) and a reverse primer (5′-GCGC CCCGGG TTA CAG GAC AAC TCT GGG AGG ACC-3′), Lassa virus NP cDNA was amplified by using a forward primer (5′-GCGC GAATTC ATG AGT GCC TCA AAG GAA ATA AAA TCC TTT TTG TGG-3′) and a reverse primer (5′-GCGC CTCGAG TCA CAG AAC GAC TCT AGG TGT CGA TGT TCT GAA-3′), and LCMV cDNA was amplified by using a forward primer (5′-GCGC GAATTC ATG TCC TTG TCT AAG GAA GTT AAG AGC TTC C-3′) and a reverse primer (5′-GCGC CTCGAG TCA GAG TGT CAC AAC ATT TGG GCC TC-3′). The PCR products containing Mopeia virus Z and NP, Lassa virus NP, and LCMV NP cDNAs were cloned into the pCR2.1-TOPO vector (Invitrogen, Carlsbad, CA), and the resultant plasmids were designated pCR-Mop-Z, pCR-Mop-NP, pCR-Las-NP, and pCR-LCM-NP, respectively. After sequence confirmation, the open reading frames encoding the Z and NP proteins were subcloned into protein expression vector pCAGGS/MCS (17, 22). Plasmids encoding C-terminally FLAG-tagged NP and HA-tagged Z were constructed by using PCR and standard cloning techniques. The resulting plasmids were designated pC-MopZ-HA, pC-MopNP-FLAG, pC-LasNP-FLAG, and pC-LCMNP-FLAG.
A pCAGGS vector containing a FLAG epitope with BsmBI sites, designated N-the FLAG-pCAGGS vector, was also constructed. The NP cDNA of influenza A virus [A/WSN/33 (H1N1) strain] was amplified by PCR with plasmid pCAGGS-WSN-NP as a template and influenza A virus NP gene-specific oligonucleotide primers containing BsmBI sites. After digestion with BsmBI, the PCR product was cloned into the BsmBI sites of the N-FLAG-pCAGGS vector. The resulting plasmid, referred to as pC-WSN-FLAG-NP, encodes influenza virus NP with a FLAG tag at the N terminus. A plasmid encoding the C-terminally FLAG-tagged NP of Zaire Ebola virus (designated pCEboZ-NP-FLAG) was constructed as described elsewhere previously (37).
To generate Mopeia virus NP deletion constructs, we designed pairs of primers that flanked the regions to be deleted (the sequences of the primers are available upon request). The primers, along with the forward and reverse NP primers (see above), were used to PCR amplify two overlapping fragments from the wild-type NP coding region. These pairs of PCR products served as templates for the second round of PCR amplification to generate the final products containing the deletions. The FLAG sequences were then fused to the 3′ ends of the coding regions as described above for wild-type NP, and the products were cloned into pCAGGS/MCS. Deletions within the coding regions were verified by DNA sequencing.
All transfections were performed by using TransIT LT-1 (Mirus, Madison, WI) reagent according to the supplier's instructions. Forty-eight hours posttransfection, 293T cells were lysed in radioimmunoprecipitation assay (RIPA) buffer (50 mM Tris-Cl [pH 7.5], 150 mM NaCl, 1% Triton X-100, 0.5% deoxycholate, and 0.1% SDS). After incubation on ice for 15 min, lysates were clarified by centrifugation. Samples were then mixed with SDS-PAGE sample buffer, incubated at 100°C for 5 min, and subjected to a 4% to 20% PAGE Tris-glycine gradient gel (NuSep, Sydney, Australia). The resolved proteins were then electrotransferred onto Western polyvinylidene difluoride membranes (Whatman, Maidstone, United Kingdom) and subsequently blocked overnight at 4°C with 5% skim milk in phosphate-buffered saline (PBS)-Tween (PBS-T) (PBS containing 0.05% Tween 20 [Sigma, St. Louis, MO]). Blots were then incubated in PBS-T containing 0.5% skim milk and the corresponding primary antibodies at room temperature for 1 h and then incubated with either an anti-rabbit horseradish peroxidase (HRP)-conjugated secondary antibody (Zymed, San Francisco, CA) or an anti-mouse HRP-conjugated secondary antibody (Pierce, Rockford, IL) at room temperature for 1 h. Blots were washed three times in PBS-T after each incubation. Protein bands were developed with a chemiluminescence reagent (Roche, Branford, CT) and then exposed on Kodak Biomax films.
293T cells in 100-mm dishes were transfected with 0.5 μg of the indicated plasmids. Plasmid pCAGGS/MCS was used to adjust the total DNA amount for transfection. Culture supernatants were collected 48 h posttransfection, clarified, laid over a cushion of 20% sucrose in PBS, and ultracentrifuged at 27,000 rpm for 2 h at 4°C. The pellets were resuspended in 60 μl of STE buffer (0.01 M Tris-Cl [pH 7.5], 0.01 M NaCl, 0.001 M EDTA [pH 8.0]) overnight at 4°C. The relative amount of protein in the VLPs was estimated based on the density of the protein bands, and NP incorporation efficiency was calculated as the ratio of the amount of the NP protein relative to that of the Z protein detected in the VLPs.
Six aliquots of VLP resuspension underwent the following treatments at room temperature for 30 min: (i) none, (ii) 3 mg/ml soybean trypsin inhibitor (Biofluids, Rockville, MD), (iii) 1% Triton X-100 (Sigma, St. Louis, MO), (iv) 0.1 mg/ml trypsin (Worthington, Lakewood, NJ), (v) 1% Triton X-100 and 0.1 mg/ml trypsin, and (vi) 3 mg/ml trypsin inhibitor and 0.1 mg/ml trypsin. After 5 mg/ml trypsin inhibitor was added to each aliquot, samples were subjected to Western blot analysis as described above.
The assay was performed as described previously by Runkler et al. (30), with some modifications. 293T cells in 100-mm dishes were cotransfected with 1 μg of pC-eGFP and 2 μg of the indicated plasmids encoding viral proteins. Forty-eight hours posttransfection, cells were scraped into PBS and pelleted at 3,500 rpm for 5 min at 4°C. The pellets were resuspended in 800 μl of hypotonic buffer (20 mM Tris-HCl [pH 7.4], protease inhibitor mixture [Roche, Branford, CT]). After incubation on ice for 20 min, cells were passaged 30 times through an 18-gauge needle and clarified by centrifugation at 2,000 rpm for 3 min at 4°C to remove nuclei. The resulting supernatants were mixed with 1.6 ml of 60% iodixanol solution (OptiPrep; Sigma, St. Louis, MO), placed into Beckman SW41 ultracentrifuge tubes, overlaid with 7.2 ml of 30% iodixanol solution diluted with TNE buffer (25 mM Tris-HCl, 150 mM NaCl, 5 mM EDTA [pH 7.5], protease inhibitor mixture [Roche, Branford, CT]), and then overlaid with 2 ml of TNE buffer. The samples were ultracentrifuged at 40,000 rpm for 5 h at 4°C. Fractions were collected from the top of the gradient and analyzed with anti-HA, anti-FLAG, anti-eGFP, and anti-calnexin antibodies by Western blotting. NP percentages were calculated as the ratio of the amount of NP in a fraction to the amount of NP in the gradient. The percentages for the calnexin-positive fractions were combined to determine the value of membrane-bound NP in the gradient. Three independent experiments were performed to estimate the average percent values.
Vero cells grown on glass coverslips in 35-mm dishes were transfected with 0.25 μg of the indicated constructs. Twenty-four hours later, the cells were fixed in 4% paraformaldehyde for 15 min, permeabilized with 0.1% Triton X-100 in PBS for 5 min, and blocked with 10% goat serum in PBS. The samples were then incubated with primary antibodies for 1 h and subsequently with fluorescein isothiocyanate (FITC)-conjugated anti-mouse and Alexa Fluor 594-conjugated anti-rabbit secondary antibodies (Molecular Probes, Eugene, OR). Images were obtained by using a Zeiss LSM 510 laser scanning confocal microscope. The extent of colocalization was calculated by using LSM 510 colocalization software; 20 cell images were used for these calculations.
The Z proteins of Lassa virus and LCMV are known to facilitate the release of VLPs (27, 34). To test whether the Z protein of Mopeia virus similarly mediates the budding and release of VLPs, we first transfected 293T cells with a plasmid expressing HA-tagged Mopeia virus Z or FLAG-tagged Mopeia virus NP, as a negative control. Forty-eight hours posttransfection, culture supernatants were collected and pelleted by ultracentrifugation through a 20% sucrose cushion and were then subjected to Western blot analysis, as described previously (15). Although both the Z and NP proteins were expressed in cell lysates, only the Z protein was detected in the pellet fraction (Fig. (Fig.1A),1A), suggesting that the expression of the Mopeia virus Z protein promoted the production of Z-containing VLPs.
To test whether Mopeia virus NP was incorporated into Mopeia virus Z-induced VLPs, 293T cells were cotransfected with plasmids expressing these two proteins. As shown in Fig. Fig.1A,1A, The NP protein was detected in the pellet fraction of the culture supernatant from NP- and Z-expressing cells. To confirm that Mopeia virus NP was incorporated into the Z-induced lipid-enveloped VLPs, we performed a protease protection assay as described previously (15). When the pellet fractions from the culture supernatants of transfected cells were treated with either protease inhibitor, Triton X-100, trypsin, Triton X-100 and trypsin, or protease inhibitor and trypsin, we found that the NP protein was protected from proteolysis by trypsin treatment but that the addition of Triton X-100 disrupted the lipid envelope, resulting in NP degradation in the presence of trypsin (Fig. (Fig.1B).1B). These results suggest that NP is incorporated into the Z-containing lipid-enveloped VLPs.
To assess the specificity of the incorporation of Mopeia virus NP into Mopeia virus Z-induced VLPs, we cotransfected 293T cells with a plasmid expressing the Mopeia virus Z protein and a plasmid expressing Ebola virus NP, influenza virus NP, or Mopeia virus NP and repeated the VLP production assays. As shown in Fig. Fig.2,2, Mopeia virus NP, but not Ebola virus NP or influenza virus NP, was incorporated into the Mopeia virus Z-induced VLPs, suggesting that NP incorporation into Mopeia virus Z-induced VLPs is selective. In addition, we also tested the incorporation of Lassa virus and LCMV NPs into Mopeia virus Z-induced VLPs and found that both of these NPs were efficiently incorporated into the VLPs (data not shown).
The selective incorporation of nucleocapsids into budding virions may be mediated through the interaction of NP with Z. To assess the interaction between Mopeia virus NP and Z, we performed a coimmunoprecipitation assay. The Mopeia virus NP and Z proteins did not coprecipitate (data not shown), unlike the NP and Z proteins of Lassa virus (11). Therefore, we examined their localization in Vero cells transiently expressing these two proteins by using an immunofluorescence assay. As shown in Fig. Fig.3,3, NP was distributed throughout the cytoplasm in variable-sized aggregates in cells expressing NP alone. In contrast, the Z protein was distributed in the nucleus, to some extent, but predominantly in the perinuclear regions and in distinct punctuate structures in the cytoplasm of Z-expressing cells. Upon the coexpression of NP and Z, however, the NP localization pattern changed, and NP colocalized with Z (67.4% of NP colocalized with Z, and 79% of Z colocalized with NP, as determined by use of Zeiss LS510 colocalization software) (Fig. (Fig.3).3). These results suggest a potential association between the Mopeia virus Z and NP proteins, even though we were unable to detect their interaction by coimmunoprecipitation.
The Z proteins of LCMV and Lassa virus bind to cellular membranes (27, 34), presumably facilitating virion release. To determine whether Mopeia virus Z binds to cellular membranes, a culture supernatant of cells transfected with a plasmid expressing Mopeia virus Z was subjected to membrane flotation analysis as described previously (30), with some modifications (see Materials and Methods). In accordance with several other arenavirus studies (27, 33, 34), the distribution of the Mopeia virus Z protein paralleled that of a membrane-associated protein marker, alpha-calnexin (Fig. (Fig.4),4), confirming the association of Mopeia virus Z with cellular membranes. In contrast, most of the Mopeia virus NP appeared in the soluble fractions, although faint bands were observed in the membrane fractions (Fig. (Fig.4B4B).
Our colocalization study suggested an effect of Mopeia virus Z on the intracellular localization of NP. To examine this effect in more detail, we tested supernatants of cells cotransfected with plasmids expressing the Mopeia virus Z and NP proteins. We found that in the presence of Z, the NP level in the membrane fractions was approximately 3-fold higher than that in its absence (Fig. (Fig.4B),4B), suggesting that Mopeia virus Z facilitates an efficient association of NP with cellular membranes.
Our results suggest that the Mopeia virus Z protein may recruit NP to cellular membranes, facilitating the incorporation of NP into virion particles. To identify the NP regions responsible for the VLP incorporation of NP, we generated a series of NP deletion mutants and examined the efficiency of their incorporation into VLPs (Fig. (Fig.5A5A).
First, we examined NP protein expression in 293T cells transfected with the mutant NP constructs. The levels of mutant NP expression were almost all comparable to that of wild-type NP, except for NPΔ1-113 and NPΔ114-228; the protein expression level of NPΔ1-113 was slightly lower than that of wild-type NP (Fig. (Fig.5B),5B), and no NP expression was detected for NPΔ114-228, which was therefore not investigated further. For NPΔ1-113 and NPΔ229-341, we also observed an additional band, which migrated slower than the major band for unknown reasons.
Next, we assessed the incorporation efficiency of the mutant NPs into VLPs. As shown in Fig. Fig.5B,5B, NPΔ1-113 and NPΔ229-341 were incorporated into Z-induced VLPs more efficiently than was wild-type NP (Fig. (Fig.5B).5B). In contrast, NP mutants that lacked C-terminal regions (i.e., NPΔ342-399, NPΔ400-457, NPΔ458-514, and NPΔ515-570) were incorporated into VLPs with a very low efficiency (Fig. (Fig.5B).5B). These results indicate that C-terminal residues 342 to 570 of NP are important for NP incorporation into Z-induced VLPs.
We speculated that deletions in the C-terminal half of the Mopeia virus NP protein may affect NP membrane association and/or colocalization with Z, thus resulting in an inefficient NP incorporation into virions. To test this hypothesis, we examined the intracellular distribution of mutant NPs in the presence of the Z protein. NP mutants that were not efficiently incorporated into VLPs still colocalized with Z (i.e., >60% of NP colocalized with Z) (data not shown).
We then examined the effect of deletions in NP on the NP membrane association (Fig. 6A and B). In the presence of the Z protein, NPΔ1-113 and NPΔ229-341 were almost entirely membrane associated (98.7% and 96.4%, respectively) (Fig. (Fig.6B),6B), which may explain their efficient virion incorporation (Fig. (Fig.5B).5B). In cells expressing the Z protein, the other NP mutants associated with the membrane to levels roughly comparable to those of wild-type NP, yet these NP mutants were not efficiently incorporated into virions (Fig. (Fig.5B).5B). These findings suggest that colocalization with Z and association with cellular membranes may be necessary, but not sufficient, for efficient NP virion incorporation; rather, additional interactions (possibly with cellular factors) may be required for efficient NP virion incorporation.
For several enveloped viruses, viral matrix proteins have been shown to have an important role in virion formation and the incorporation of viral nucleocapsids into virions (10, 15, 16, 23, 37-40). In this study, we demonstrated that the viral matrix Z protein of Mopeia virus, a close relative of Lassa virus, mediates the selective incorporation of the Mopeia virus NP protein into Z protein-induced VLPs. Our data suggest that the association of NP, Z, and the cellular membranes may facilitate the efficient incorporation of NP into VLPs. Moreover, we found that the C-terminal half of NP is essential for NP incorporation into VLPs.
Interactions between Z and NP or RNP have been demonstrated for several arenaviruses, such as LCMV and Lassa virus (11, 31). In agreement with these findings, our data from immunofluorescence and membrane flotation assays also suggest an association between the Z and NP proteins of Mopeia virus (Fig. (Fig.33 and and4),4), leading to NP recruitment by Z protein, even though we were unable to detect a direct interaction between these proteins by immunoprecipitation (data not shown). Furthermore, we identified that amino acid residues 342 to 570 of NP are important for the virion incorporation of NP. Deletions of these residues, however, had no or little effect on the colocalization of NP and Z or on the Z-mediated NP membrane association, suggesting that the association between NP, Z, and the cellular membranes may not be sufficient for the VLP incorporation of NP. NP may be recruited to the budding sites, but its incorporation may be hampered by the lack of these NP residues. It may be that this region of NP is essential for the interaction of NP with other factors (i.e., host proteins), which are critical for the final steps of RNP incorporation into virions. Tortorici et al. previously described a zinc finger within the C-terminal region of the Junin virus nucleoprotein (N), although its biological relevance was not addressed (35). The Mopeia virus NP protein also contains putative zinc binding residues within the C terminus, amino acids 412 to 535, which, in our study, appear to be important for NP incorporation (Fig. (Fig.5B).5B). For HIV-1, a potential role for the zinc finger of its nucleocapsid (NC) in particle production was demonstrated previously (29). Therefore, it is possible that the zinc finger domain of Mopeia virus NP may also play a role in particle formation.
We found that the majority of the Mopeia virus Z protein associated with cellular membranes (Fig. (Fig.4),4), although the type of cellular membrane(s) with which Z associated is not yet clear: it could be the plasma membrane and/or internal membranes (e.g., the nuclear membrane, endoplasmic reticulum, Golgi apparatus, or endosomes). Our immunofluorescence study showed that the Mopeia virus Z protein was distributed predominantly in the perinuclear regions and in distinct punctuate structures in the cytoplasm rather than in the plasma membrane (Fig. (Fig.3),3), as was observed for cells expressing the Lassa virus Z protein (3, 11, 32). We also found that Z colocalized with CD63 and the mannose-6-phosphate receptor (our unpublished data), cellular markers for late endosomes/multivesicular bodies (MVBs), which are involved in the assembly and budding steps of several enveloped viruses (18, 24, 38). Moreover, host factors that function in the MVB pathway were shown to have roles in the formation of Lassa virus Z-induced VLPs (36). Taken together, it is possible that the Mopeia virus Z protein may use host functions in the endosome/MVB pathway to facilitate the efficient assembly and budding of Mopeia virus Z-induced VLPs.
The NPs of New World arenaviruses were shown to enhance Z-induced VLP formation and GP recruitment into VLPs (6, 12), whereas no such enhancement has been shown for Lassa virus VLPs (32). Although we did not observe that Mopeia virus NP promotes Mopeia virus VLP formation (Fig. (Fig.1),1), it is plausible for the Mopeia virus NP protein to participate in the recruitment of viral GP for assembly. On the other hand, although an interaction between LCMV GP and RNPs was demonstrated previously (4), the involvement of arenavirus GP in NP incorporation into VLP was not addressed. Our preliminary data suggest that the coexpression of the Mopeia virus GP and NP proteins results in the induction of enveloped vesicles containing both GP and NP, demonstrating a possible contribution of GP to the recruitment of NP/RNPs into newly forming virions (our unpublished data). Further investigation will be needed to understand how viral proteins associate to promote arenavirus particle formation.
We found that a small portion of the Mopeia virus NP protein bound to the cellular membranes, although most of the NP was soluble in cells expressing NP alone (Fig. (Fig.4),4), suggesting an inherent ability of Mopeia virus NP for membrane binding albeit to a very limited extent. Membrane proteins are categorized as integral and peripheral proteins. Since Mopeia virus NP does not contain long hydrophobic stretches sufficient for spanning the cell membrane, this protein is most likely a peripheral protein that temporarily attaches either to the lipid bilayer or to integral membrane proteins via electrostatic interactions. Interestingly, deletions of amino acids 1 to 113, 229 to 341, or 342 to 399 of NP increased the efficiency of NP membrane association in the absence of the Z protein (Fig. (Fig.6A).6A). One possible explanation for this finding is that these deletions in NP may alter the structure of the protein, exposing regions that promote the interaction between NP and the lipid bilayer or the integral membrane proteins.
In summary, we have demonstrated that the C terminus of Mopeia virus NP is required for its incorporation into Z-induced VLPs, suggesting a potential role for this region in the incorporation of RNP complexes into Mopeia virus virion particles. Furthermore, similar analyses will advance our knowledge of the replication mechanism of arenaviruses.
We thank the members of our laboratory for valuable comments, Shinji Watanabe and Susan Watson for editing of the manuscript, and Krisna Wells, Martha McGregor, Rebecca Moritz, and Kelly Moore for excellent technical assistance. We also thank Suresh Marulasiddappa and Erin Plisch of the University Wisconsin—Madison for kindly providing the LCMV stock RNA.
This work was supported by U.S. National Institutes of Health and National Institute of Allergy and Infectious Diseases Public Health Service grant AI055625.
Published ahead of print on 3 March 2010.