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Journal of Virology, August 2004, p. 8732-8745, Vol. 78, No. 16
0022-538X/04/$08.00+0     DOI: 10.1128/JVI.78.16.8732-8745.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.

Putative Autocleavage of Outer Capsid Protein µ1, Allowing Release of Myristoylated Peptide µ1N during Particle Uncoating, Is Critical for Cell Entry by Reovirus

Amy L. Odegard,1,2 Kartik Chandran,1,{dagger} Xing Zhang,3 John S. L. Parker,1,{ddagger} Timothy S. Baker,3 and Max L. Nibert1*

Department of Microbiology and Molecular Genetics, Harvard Medical School, Boston, Massachusetts 02115,1 Department of Biochemistry, University of Wisconsin—Madison, Madison, Wisconsin 53706,2 Department of Biological Sciences, Purdue University, West Lafayette, Indiana 479073

Received 24 January 2004/ Accepted 8 April 2004


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ABSTRACT
 
Several nonenveloped animal viruses possess an autolytic capsid protein that is cleaved as a maturation step during assembly to yield infectious virions. The 76-kDa major outer capsid protein µ1 of mammalian orthoreoviruses (reoviruses) is also thought to be autocatalytically cleaved, yielding the virion-associated fragments µ1N (4 kDa; myristoylated) and µ1C (72 kDa). In this study, we found that µ1 cleavage to yield µ1N and µ1C was not required for outer capsid assembly but contributed greatly to the infectivity of the assembled particles. Recoated particles containing mutant, cleavage-defective µ1 (asparagine -> alanine substitution at amino acid 42) were competent for attachment; processing by exogenous proteases; structural changes in the outer capsid, including µ1 conformational change and {sigma}1 release; and transcriptase activation but failed to mediate membrane permeabilization either in vitro (no hemolysis) or in vivo (no coentry of the ribonucleotoxin {alpha}-sarcin). In addition, after these particles were allowed to enter cells, the {delta} region of µ1 continued to colocalize with viral core proteins in punctate structures, indicating that both elements remained bound together in particles and/or trapped within the same subcellular compartments, consistent with a defect in membrane penetration. If membrane penetration activity was supplied in trans by a coinfecting genome-deficient particle, the recoated particles with cleavage-defective µ1 displayed much higher levels of infectivity. These findings led us to propose a new uncoating intermediate, at which particles are trapped in the absence of µ1N/µ1C cleavage. We additionally showed that this cleavage allowed the myristoylated, N-terminal µ1N fragment to be released from reovirus particles during entry-related uncoating, analogous to the myristoylated, N-terminal VP4 fragment of picornavirus capsid proteins. The results thus suggest that hydrophobic peptide release following capsid protein autocleavage is part of a general mechanism of membrane penetration shared by several diverse nonenveloped animal viruses.


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INTRODUCTION
 
The mechanisms by which nonenveloped animal viruses mediate membrane penetration to invade the host cytoplasm remain less well understood than the mechanisms of membrane fusion by enveloped viruses. Since nonenveloped viruses lack a membrane, they must traverse the cellular membrane barrier by a mechanism other than fusion. Nonenveloped animal viruses contain "penetration proteins, " analogous to the fusion proteins of enveloped viruses, that accomplish membrane penetration, perhaps either by forming a membrane-spanning pore or by locally disrupting the membrane bilayer. For example, capsid proteins VP1 and VP4 of poliovirus, each present in 60 copies per virion, are thought to play the critical roles in membrane penetration by that nonenveloped virus (reviewed in reference 38).

In many viruses, the fusion or penetration proteins are primed to adopt their membrane-seeking forms by posttranslational cleavage, which is mediated in some cases by the proteins themselves (autocatalytic cleavage, or "autocleavage"). Cleavage may be necessary to remove conformational restraints, to expose terminal hydrophobic sequences for membrane insertion, or to allow peptide release. Examples of enveloped virus fusion proteins primed by cleavage include influenza virus HA (Orthomyxoviridae), Sendai virus F (Paramyxoviridae), and human immunodeficiency virus gp160 (Retroviridae) (reviewed in reference 37). Examples of nonenveloped virus penetration proteins primed by autocleavage include polio- and rhinovirus VP0 (Picornaviridae), flock house virus {alpha} (Nodaviridae), and Nudaurelia capensis omega virus {alpha} (Tetraviridae) (reviewed in reference 41). Polio- and rhinovirus VP0 proteins are cleaved into fragments VP4 and VP2, allowing VP4 release during cell entry (6, 24, 32, 46, 47). Similarly, flock house virus and N. capensis omega virus {alpha} proteins are cleaved into fragments ß and {gamma}, allowing {gamma} release during cell entry (8, 35, 53, 59, 64, 68).

Mammalian orthoreoviruses (reoviruses), members of the Reoviridae family, are nonenveloped viruses comprising a 10-segment, double-stranded RNA genome surrounded by two concentric, icosahedral protein capsids. The 10 genome segments encode eight structural proteins, which constitute the T=1 inner and T=13 (laevo) outer capsid layers, and three nonstructural proteins not present in the mature virion. The inner capsid proteins possess the enzymatic activities necessary for viral transcription (reviewed in reference 63). In addition, the outer capsid protein {lambda}2 is responsible for capping the 5' end of each viral plus-strand RNA as it exits the particle during transcription (34; reviewed in reference 58). The other outer capsid proteins—µ1, {sigma}3, and {sigma}1—are involved in cell entry. The majority of the outer capsid lattice is formed by 600 copies of µ1, the putative membrane penetration protein (reviewed in reference 16), which are organized as trimers on the virus particle (29, 48). The {sigma}3 protein, also present in 600 copies, closely associates with µ1 to coat the particle surface (29, 48). The remainder of the outer capsid is made up of 36 copies of the attachment protein {sigma}1, arranged as trimers at the fivefold axes of symmetry (20, 22, 33; reviewed in reference 44). The double-layered nature of the reovirus capsid (diameter, ~85 nm) makes it distinct from—and much larger than—the T=3 single capsids of poliovirus and flock house virus (diameter, 30 to 35 nm) (31, 39) and the T=4 single capsid of N. capensis omega virus (diameter, 40 to 45 nm) (53).

To initiate infection, the {sigma}1 protein in reovirus virions binds to host cell surface receptors (reviewed in references 5 and 44), and the particles are then internalized, most likely by receptor-mediated endocytosis (10, 62). Within the endocytic vesicles, lysosomal proteases cleave the outer capsid proteins (30, 62) to generate particles very similar to the infectious subvirion particles (ISVPs) that can be generated in vitro by protease treatment (42, 54). In these particles, {sigma}3 is degraded, leaving µ1 as the major surface protein (29). Through a process that we continue to dissect, ISVPs can undergo a structural transformation to yield a related particle form, the ISVP* (15, 42). Generation of the ISVP*, which contains an altered conformer of µ1 and has shed {sigma}1, appears necessary for membrane penetration, promoting particle release into the host cytoplasm (10, 11, 15, 18, 43). Either during the process of membrane penetration or after release into the cytoplasm, a large piece of µ1, the central "{delta}" fragment (see below), is also lost from the particle (18). Furthermore, once in the cytoplasm, the now transcriptionally activated particle can produce the viral plus-strand RNAs that are used as templates for new protein synthesis and duplex RNA genome synthesis. This cytoplasmically delivered "payload" of reovirus—a partially uncoated particle with transcriptase activity—is distinct from those of poliovirus, flock house virus, and N. capensis omega virus, which are thought to enter the cytoplasm by delivering only their translation-competent, plus-sense RNA genomes across the membrane (reviewed in references 38 and 41).

The putative membrane penetration protein of reoviruses, µ1 (76 kDa; 708 amino acids), is known to undergo at least three different proteolytic cleavages in vitro and in vivo (Fig. 1A). Upon exposure to host proteases in the intestinal lumen (7) or in endocytic vesicles (30, 62), particle-bound µ1 is cleaved near amino acid 580 to generate two fragments, µ1{delta} and {phi}, that remain particle bound (54) (Fig. 1A). The cleavage of µ1 into µ1{delta} and {phi} has been shown to be dispensable for membrane permeabilization and transcriptase activation, as well as for infection, by ISVPs (17, 19). As recently identified (51; M. L. Nibert, unpublished data), removal of ~10 amino acids from the C terminus of µ1 (i.e., from the {phi} region) can also accompany protease treatment, but the significance of this cleavage remains unclear. Lastly, µ1 is cleaved near its N terminus, between amino acids 42 and 43, to generate two fragments, a small N-terminal fragment, µ1N (4 kDa), and a large C-terminal fragment, µ1C (72 kDa), which are also both present in particles (55, 61) (Fig. 1A). No known protease has been linked to the µ1N/µ1C cleavage, and this cleavage is instead considered to be autocatalytic (48, 55). Interestingly, µ1N contains the N-myristoyl group of µ1 at its N terminus, making it further reminiscent of polio- and rhinovirus VP4, which has been implicated in membrane penetration by those nonenveloped viruses (23, 25, 32, 46, 47, 52).



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  		FIG. 1. Expression of µ1N42A, generation of recoated particles, and protease treatment of recoated particles to generate ISVPs. (A) The µ1 protein and its cleavage products are diagrammed. The µ1 N-terminal and C-terminal amino acid positions are labeled 2 and 708, respectively. The autocleavage products µ1N and µ1C are indicated. The N-terminal myristoyl group (myr) is present in both full-length µ1 and µ1N. Cleavage of full-length µ1 by exogenous proteases (e.g., chymotrypsin or trypsin) yields the fragments µ1{delta} and {phi}. Cleavage of µ1C by exogenous proteases yields the fragments {delta} and {phi}. (B) SDS-PAGE and immunoblotting of insect cell lysates containing either coexpressed µ1WT and wild-type {sigma}3 (lane 3) or µ1N42A and wild-type {sigma}3 (lane 4) was performed. The samples were probed with either the µ1-specific mouse MAb 10H2 (top) or a {sigma}3-specific rabbit polyclonal antiserum (bottom). Virions (lane 1) and cores (lane 2) were included for comparison. (C) Samples of purified µ1WT r-cores (lane 3) and µ1N42A r-cores (lane 4) were examined by SDS-PAGE followed by either Coomassie staining (top) or immunoblotting with µ1-specific MAb 10H2 (bottom). Virions (lane 1) and cores (lane 2) were included for comparison. (D) Samples of purified µ1WT r-cores plus {sigma}1 (lane 3) and µ1N42A r-cores plus {sigma}1 (lane 4) were examined by SDS-PAGE followed by either Coomassie staining (top) or immunoblotting with a {sigma}1-specific rabbit polyclonal antiserum (bottom). Virions (lane 1) and cores (lane 2) were included for comparison. (E) Samples of chymotrypsin-generated µ1WT pr-cores (lane 3) and µ1N42A pr-cores (lane 4) were examined by SDS-PAGE and Coomassie staining. Untreated µ1WT r-cores (lane 1) and µ1N42A r-cores (lane 2) were included for comparison.

The significance of the µ1N/µ1C cleavage in reovirus infection has not been directly addressed, largely because of an inability to block this cleavage in viral particles. We overcame this obstacle by generating a point mutation in µ1 to prevent µ1N/µ1C cleavage and then studying its effects in "recoated cores" (r-cores), a versatile system developed for studies of reovirus outer capsid assembly and functions in cell entry (19, 20). The residue immediately N-terminal to the µ1N/µ1C cleavage site, Asn42, was specifically chosen for mutagenesis based on a previous report that it is required for the cleavage to occur (65). In this report, we describe our results with this approach and the new insights we obtained into the role of µ1 in membrane penetration by reoviruses. The findings suggest that hydrophobic peptide release following capsid protein autocleavage is part of a general mechanism of membrane penetration shared by several diverse nonenveloped viruses.


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MATERIALS AND METHODS
 
Mouse and insect cells and native reovirus particles. Spinner-adapted mouse L929 cells were grown in suspension in Joklik's modified minimal essential medium (Irvine) supplemented to contain 2% fetal and 2% neonatal bovine sera (HyClone), as well as 2 mM glutamine, 100 U of penicillin/ml, and 100 µg of streptomycin (Irvine)/ml. Spodoptera frugiperda clone 21 (Sf21) and Trichoplusia ni High Five insect cells (Invitrogen) were grown in TC-100 medium (Invitrogen) supplemented to contain 10% heat-inactivated fetal bovine serum. Mv1Lu and CV1 cells were grown in Dulbecco's modified Eagle's medium (Invitrogen) supplemented to contain 10% fetal bovine serum and 10 µg of gentamicin (Invitrogen)/ml. Purified reovirus type 1 Lang (T1L) virions, ISVPs, and top component particles (33), as well as cores (19), were obtained as previously described. The purified particles were stored at 4°C in virion buffer (150 mM NaCl, 10 mM MgCl2, 10 mM Tris, pH 7.5).

Generation of recombinant baculovirus to express mutant µ1. Site-directed mutagenesis of the plasmid pBKS-M2L (19) was performed using the QuikChange protocol (Stratagene). The missense mutation that resulted in changing Asn42 to alanine (µ1N42A) was introduced using the complementary mutagenic primers 5'-CTATCACCTGGAATGTTAGCGCCTGGAGGAGTACCATGG-3' and 5'-CCATGGTACTCCTCCAGGCGCTAACATTCCAGGTGATAG-3' (the changes are underlined). The nucleotide changes also introduced a HaeII restriction site that was used to screen for plasmids containing the mutant clone. Mutant clones were sequenced from nucleotides 1 to 1120 to confirm that they contained the desired changes and to rule out any additional mutations. The NotI-Bsu36I restriction fragment (nucleotides 1 to 727) containing the desired changes was subcloned into pFb-M2L (19). Clones containing the subcloned insert were then further subcloned into pFbD-M2L/S4L (19) using the Bsu36I-SalI restriction sites (nucleotides 1 to 727). All restriction enzymes were obtained from New England BioLabs.

Expression of reovirus proteins from baculovirus vectors. The recombinant baculovirus containing the T1L M2 and S4 genes used to coexpress wild-type µ1 (µ1WT) and wild-type {sigma}3 was described previously (20). The recombinant baculovirus containing the T1L S1 gene used to express wild-type {sigma}1 has been described (20, 21). To express these wild-type proteins or µ1N42A, T. ni High Five cells were infected at a multiplicity of infection of 5 to 10 PFU/cell with fourth-passage baculovirus stocks. Baculovirus-infected cells expressing {sigma}1 or coexpressing µ1 and {sigma}3 were harvested 65 h postinfection (p.i.), and cytoplasmic extracts were prepared as previously described (19, 20).

Protein gel electrophoresis and immunoblotting. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) of reduced or nonreduced samples was carried out as previously described (56). Samples were reduced unless otherwise indicated in the text. For immunoblot analyses, proteins were transferred to a nitrocellulose membrane in 25 mM Tris-192 mM glycine (pH 8.3). The µ1-specific mouse monoclonal antibody (MAb) 10H2 (67), T1L {sigma}1-specific rabbit polyclonal antiserum (15), and {sigma}3-specific rabbit polyclonal antiserum (40) were each used at a 1:2,000 dilution. The µNS-specific rabbit polyclonal antiserum (14) was used at a 1:5,000 dilution. The binding of these primary antibodies was detected with alkaline phosphatase-coupled goat anti-mouse or goat anti-rabbit antibodies (Bio-Rad) and the colorimetric reagents p-nitroblue tetrazolium chloride and 5-bromo-4-chloro-3-indolylphosphate p-toluidine salt (Bio-Rad).

R-cores. R-cores and r-cores plus {sigma}1 were generated as described previously (19, 20). Briefly, cytoplasmic insect cell extracts containing baculovirus-expressed µ1 and {sigma}3 or µ1, {sigma}3, and {sigma}1 were incubated with purified T1L cores at 37°C for 2 to 4 h. Following incubation, the reaction mixtures were chilled and loaded atop a 14-ml CsCl step gradient (1.25 to 1.45 g/cm3) with a 2-ml sucrose cushion (20% [wt/vol]). The gradients were subjected to centrifugation at 25,000 rpm in a Beckman SW28 rotor for 2 to 20 h at 4°C. The resulting particles were harvested and concentrated on a second CsCl step gradient and centrifuged at 40,000 rpm in a Beckman SW50.1 rotor for 2 h at 4°C. The r-cores were harvested from the second gradient and dialyzed extensively in virion buffer. Particle concentrations were determined by densitometry of gels stained with Coomassie brilliant blue R250 (Sigma) as described previously (19).

Electron cryomicroscopy and image reconstruction. Vitrified samples of µ1N42A r-cores were prepared on holey carbon films and maintained at –176°C for electron cryomicroscopy as described previously (4). Electron micrographs were recorded under low-dose conditions (~20 e–/Å2) in an FEI-Philips CM300 FEG microscope at a calibrated magnification of x47,440. The micrographs were digitized at 7-µm intervals on a Zeiss PHODIS scanner, and the data were then bin-averaged to yield pixels corresponding to 2.95 Å in the specimen. A total of 22 micrographs, recorded with defocus settings ranging between 1.9- and 3.6-µm underfocus, were selected for processing. From these micrographs, 973 particle images were masked and extracted using the program RobEM (http://bilbo.bio.purdue.edu/~baker/programs/programs.html), and each particle image was stored as a 363- by 363-pixel array. Origin and orientation parameters for each particle were determined and refined using model-based procedures (3). Eleven cycles of refinement and data screening led to a three-dimensional reconstruction at 15.6-Å resolution, computed from 807 particle images. The effects of the microscope contrast transfer function were compensated for in part for each image (12). A difference density map was computed by subtracting the map of µ1N42A r-cores from an existing 17.6-Å map of µ1WT r-cores plus {sigma}1 (20), after first reducing the resolution and scaling the magnification and contrast of the former against the latter using RobEM. High-spatial-frequency features in all maps were sharpened through the application of a 1/600-Å2 inverse temperature factor (36).

Infectivity and hemagglutination. Determinations of numbers of PFU per milliliter of reovirus particle preparations were performed as previously described (33). Particle/PFU ratios were used to compare relative infectivities. Hemagglutination titers were determined as previously described (26).

Immunostaining and immunofluorescence (IF) microscopy. For immunostaining, the following primary antibody dilutions were used: µ1-specific mouse MAb 10H2 (67) (1:2,000 dilution) and µNS-specific rabbit polyclonal antiserum (14) and core-specific rabbit polyclonal antiserum (19) (1:5,000 dilutions). Goat anti-mouse immunoglobulin G and goat anti-rabbit immunoglobulin G, each conjugated to either Alexa 488 or Alexa 594, were obtained from Molecular Probes. CV1 cells were plated directly on glass coverslips at a density of 1.2 x 104 cells/cm2 and allowed to adhere overnight. When indicated, cycloheximide (50 µg/ml; Sigma) was added at 37°C 1 h prior to infection. Cells were infected at 2 x 104 to 5 x 104 particles/cell for 1 h at 4°C. Following virus attachment, the cells were either fixed immediately in 2% paraformaldehyde for 10 min at room temperature or incubated at 37°C to allow infection to proceed and fixed in 2% paraformaldehyde at the appropriate time p.i. Unless otherwise indicated, the cells were permeabilized, blocked, incubated with antibodies and 4',6-diamidino-2-phenylindole (Molecular Probes), and mounted as described previously (57). Microscopy images were collected digitally using a Nikon TE-300 inverted microscope equipped with phase and fluorescence optics and analyzed using Metamorph version 5.1 (Universal Imaging) as described previously (57). Images were processed and presented using Photoshop and Illustrator software (Adobe).

Detection of µNS by immunoblotting. CV1 cells (2.5 x 106) were infected at 1,000 particles/cell. Lysates were collected at 0, 2, or 24 h p.i. by scraping the cells into 2 ml of phosphate-buffered saline (PBS) (137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.5). The cells were pelleted by centrifugation and resuspended in 30 µl of PBS. The cells were then lysed in sample buffer, boiled for 10 min, and subjected to SDS-PAGE. Following transfer to nitrocellulose, µNS protein was detected using a µNS-specific rabbit polyclonal antiserum (14) and an alkaline phosphatase-coupled goat anti-rabbit secondary antibody (Bio-Rad).

Flow cytometry. Confluent monolayers of Mv1Lu cells were incubated with virus at 2 x 104 particles/cell for 1 h at 4°C to allow virus attachment to occur. Following attachment, 0-h p.i. time point specimens were collected by scraping the cells into 2 ml of PBS, pelleting the cells at 500 x g in a Beckman Allegra 6R centrifuge, and fixing the cells in 1 ml of 2% paraformaldehyde for 10 min at room temperature. Cells analyzed at later times p.i. were first incubated at 37°C to allow infection to proceed for the desired time and then processed as described above. Following fixation, the cells were washed with PBS containing 1% bovine serum albumin (PBSA), permeabilized with PBSA containing 0.1% Triton X-100, and incubated for 1 h at 4°C with either µNS-specific rabbit polyclonal antiserum (14) (1:5,000 dilution) or µ1-specific mouse MAb 10H2 (67) (1:2,000 dilution). The cells were then washed with PBSA to remove unbound antibody and incubated with goat anti-rabbit immunoglobulin G or goat anti-mouse immunoglobulin G, each conjugated to Alexa 488, to detect the rabbit or mouse primary antibodies, respectively. The cells were again washed with PBSA to remove unbound antibodies and resuspended in PBS. Flow cytometry was performed using a Becton Dickinson FACS Calibur.

Hemolysis and other assays of the ISVP -> ISVP* transition. Hemolysis experiments were performed as described elsewhere (17). µ1 protease sensitivity, bis-ANS (Molecular Probes) fluorescence, and {sigma}1 release assays were performed as described previously (15). For all of the described experiments, the final particle concentration was in the range of 4 x 1012 to 7.5 x 1012 particles/ml. Cs+ was added to promote the ISVP -> ISVP* transition and associated activities relative to Na+ in these types of experiments (15, 17-20). K+ also demonstrates this effect and may be physiologically relevant in this regard as the predominant IA cation inside cells (9-11, 15). Available data indicate that the effect of larger IA cations is to accelerate the rate of the transition, not to affect its outcome.

Transcriptase activation. Transcription reactions containing 9 x 1010 particles in a total volume of 15 µl were performed as described elsewhere (27), with slight modifications. Briefly, 32P-labeled transcripts were generated by incubating NaCl-treated ISVPs (or chymotrypsin-treated r-cores with or without {sigma}1) or CsCl-treated ISVPs (or chymotrypsin-treated r-cores with or without {sigma}1) with a transcription reaction mixture (100 mM Tris, pH 8; 10 mM MgCl2; 1 mM [each] ATP, UTP, and CTP; 10 mM phosphoenolpyruvate; 0.4 U of pyruvate kinase {sigma}; 0.33 µU of RNasin [Promega]; 2 mM dithiothreitol; and 2.5 µCi of [{alpha}-32P]GTP [Perkin-Elmer]) at 37°C for 105 min. Transcription levels were measured by trichloroacetic acid precipitation followed by scintillation counting.

{alpha}-Sarcin coentry. The {alpha}-sarcin coentry assay was performed as previously described (18).

µ1N release. ISVPs labeled with [3H]myristate (Perkin-Elmer) were generated as previously described (55). These particles were then incubated for 30 min at 32°C in buffer containing 50 mM Tris (pH 7.5), 0.5% Triton X-100, and either 300 mM NaCl or 300 mM CsCl. Following incubation, the samples were cooled on ice for 10 min and then layered atop a prechilled 1.25- to 1.45-g/cm3 CsCl density gradient. The gradients were subjected to centrifugation in a Beckman SW60 rotor at 50,000 rpm and 4°C for 2 h and then fractionated using a peristaltic pump (5 drops/fraction). Ready Safe scintillation fluid (20 ml; Beckman) was added to each fraction, and the radioactivity (in counts per minute) was measured using a Beckman LS 6500 scintillation counter.


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RESULTS
 
Alanine substitution for Asn42 abrogates µ1N/µ1C cleavage. Mutant µ1 containing an asparagine -> alanine substitution at position 42 (µ1N42A) was expressed in insect cells using a recombinant baculovirus. This vector was designed to coexpress µ1 with its "protector" protein, {sigma}3, which binds to µ1 in solution and is required for its assembly into particles (19, 60). Insect cell lysates containing either µ1N42A or µ1WT coexpressed with wild-type {sigma}3 were analyzed by SDS-PAGE and immunoblotting to estimate the levels of protein expression and µ1N/µ1C cleavage. The levels of µ1 and {sigma}3 expression with µ1N42A were comparable to those with µ1WT (Fig. 1B). However, µ1N42A exhibited no detectable µ1N/µ1C cleavage, as evidenced by the absence of µ1C; most of the mutant protein comigrated with uncleaved µ1 (Fig. 1B, lane 4).

µ1N/µ1C cleavage is not required for outer capsid assembly. To test for outer capsid assembly with µ1N42A, insect cell lysates containing wild-type {sigma}3 coexpressed with either µ1N42A or µ1WT were incubated with cores, and the resulting particles (µ1N42A and µ1WT r-cores, respectively) were isolated on CsCl gradients. In both cases, a single particle band was observed, at a density consistent with virion-like particles containing the µ1-{sigma}3 outer capsid. SDS-PAGE and Coomassie staining (Fig. 1C, top) or immunoblotting (Fig. 1C, bottom) confirmed that µ1 and {sigma}3 had been incorporated into the particles in both samples. As expected for the µ1WT particles (19), most µ1 underwent µ1N/µ1C cleavage (µ1C was visible in these gels), while only a small portion (~5%) remained uncleaved. In the µ1N42A particles, however, µ1 remained wholly uncleaved (Fig. 1C, lane 4), suggesting that µ1N/µ1C cleavage is not required for outer capsid assembly. Densitometry of Coomassie-stained gels confirmed that recoating was largely complete, since the levels of µ1 and {sigma}3 relative to other structural proteins in µ1N42A r-cores approximated those of µ1 plus µ1C and {sigma}3 in native virions or µ1WT r-cores (data not shown). Use of insect cell lysates containing [3H]myristate confirmed that the µ1 protein in both µ1N42A and µ1WT r-cores was myristoylated (data not shown), as is that in native virions (55).

R-cores including the wild-type attachment protein {sigma}1 (r-cores plus {sigma}1) were also generated (20). When purified on CsCl gradients, both µ1N42A and µ1WT r-cores plus {sigma}1 migrated at a density similar to that of virions. SDS-PAGE and Coomassie staining (Fig. 1D, top), followed by densitometry (data not shown), confirmed that both µ1N42A and µ1WT r-cores plus {sigma}1 contained a nearly full complement of µ1 (or µ1 plus µ1C) and {sigma}3. The amount of {sigma}1 could not be easily quantified due to the usual low-copy number of this protein. However, the presence of {sigma}1 was confirmed by immunoblotting (Fig. 1D, bottom), and those results suggested that the amounts of {sigma}1 in both types of r-cores plus {sigma}1 were similar to that in native virions.

Evidence for proper outer capsid assembly by µ1N42A. To determine whether the outer capsid had been properly assembled, µ1N42A and µ1WT r-cores were treated with protease (trypsin [Fig. 1E] or chymotrypsin [data not shown]) to generate partially uncoated particles resembling ISVPs (pr-cores), which were then isolated on CsCl gradients. SDS-PAGE of the pr-cores (Fig. 1E, lanes 3 and 4) alongside untreated r-cores (Fig. 1E, lanes 1 and 2) indicated that {sigma}3 was degraded and most µ1 was cleaved at the {delta}-{phi} junction, as expected for ISVPs (54). The cleavage of µ1N42A at the {delta}-{phi} junction generated two fragments: a large N-terminal fragment, µ1{delta} (Fig. 1E, lane 4), and a small C-terminal fragment, {phi} (not visible on this gel, but observed on others). This differs from native ISVPs (54) and µ1WT pr-cores (Fig. 1E, lane 3) (19), in which the larger µ1 cleavage product is primarily {delta} instead of µ1{delta}. Since µ1N/µ1C cleavage is necessary to convert µ1{delta} to {delta} (Fig. 1A), the presence of µ1{delta} confirmed that this cleavage had not occurred in the µ1N42A particles. Essentially the same results were observed with µ1N42A and µ1WT pr-cores plus {sigma}1 as with pr-cores (data not shown). The finding that digestion of the outer capsid with protease generates ISVP-like particles from µ1N42A particles provides evidence that µ1N42A and its associated {sigma}3 protein are conformationally similar to wild-type µ1 and {sigma}3 in native virions and µ1WT r-cores. SDS-PAGE of µ1N42A r-cores disrupted under nonreducing conditions additionally revealed a disulfide bond between the {phi} regions of adjacent µ1 monomers (data not shown), as was also found in virions and µ1WT r-cores (56).

As a further test for proper outer capsid assembly, we subjected µ1N42A r-cores to electron cryomicroscopy and three-dimensional image reconstruction. In micrographs the individual particles were well dispersed and had an appearance similar to those of virions and µ1WT r-cores (data not shown). A final reconstruction, computed using data from 807 particles, was determined to be reliable to at least 15.6-Å resolution (4). The resolution was subsequently reduced to 17.6 Å (Fig. 2, top row) for quantitative comparisons with other reconstructed density maps. Compared to reconstructions of µ1WT r-cores (data not shown) (19) and µ1WT r-cores plus {sigma}1 (Fig. 2, middle row) (20), few if any differences were apparent. In a difference map obtained by subtracting the density map of µ1N42A r-cores from that of µ1WT r-cores plus {sigma}1, the only clear features were densities attributable to {sigma}1 at the icosahedral vertices (Fig. 2, bottom row). We conclude that the µ1N42A mutation has minimal effects on outer capsid structure and assembly.



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  		FIG. 2. Three-dimensional-image reconstruction of µ1N42A recoated particles. Shown are surface-shaded representations (left) and central, density-coded sections (right) of µ1N42A r-cores (top row) and µ1WT r-cores plus {sigma}1 (middle row) and a difference map obtained by subtracting the densities of the former from those of the latter (bottom row). Both reconstructions, as well as the difference map revealing {sigma}1- but not µ1-related densities, are shown at 17.6-Å resolution. Scale bar, 200 Å.

µ1N42A particles are poorly infectious. Consistent with previous findings (20), µ1WT r-cores plus {sigma}1 were nearly as infectious as native virions in L929 cells (Table 1). In contrast, µ1N42A r-cores plus {sigma}1 were much less infectious than either native virions or µ1WT r-cores plus {sigma}1 (Table 1) (relative [per-particle] infectivity, {approx}1/3,200 that of virions). This suggests that the µ1N/µ1C cleavage is critical for productive infection, although infection can still occur at very low efficiency (~0.03% that of virions) in the absence of this cleavage. Similarly, µ1N42A pr-cores plus {sigma}1 were much less infectious than native ISVPs or µ1WT pr-cores plus {sigma}1 (Table 1) (relative infectivity, {approx}1/1,600 that of ISVPs), suggesting that proteolytic removal of {sigma}3 and cleavage of µ1 at the {delta}-{phi} junction are not sufficient to enhance the relative infectivity of µ1N42A r-cores plus {sigma}1 to any substantial degree. Also consistent with previous findings (19), r-cores containing the outer capsid proteins µ1WT and {sigma}3, but lacking the attachment protein {sigma}1, were ~200 times more infectious than core particles in L929 cells (Table 1). Although the addition of µ1WT and {sigma}3 enhanced the relative infectivity of r-cores to this large extent, the µ1WT r-cores were still only 1/2,500 as infectious as virions (Table 1). This is consistent with previous evidence that the attachment protein {sigma}1 is needed, in addition to µ1 and {sigma}3, for r-cores to approach virion-like levels of infectivity (20). µ1N42A r-cores, in comparison, showed little if any enhancement in relative infectivity over cores and were only ~1/100 as infectious as µ1WT r-cores and ~1/27,500 as infectious as native virions. These findings suggest that the µ1N/µ1C cleavage is required to confer enhanced infectivity on r-cores. Experiments were next performed to identify the step(s) in infection at which µ1N42A particles are defective.


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  		TABLE 1. Relative infectivities of native and recoated particles

Barrier to infectivity of µ1N42A particles occurs after attachment but before or during protein synthesis. The capacity of µ1N42A pr-cores plus {sigma}1 to attach to cells was addressed by IF microscopy. CV1 cells were incubated with µ1N42A pr-cores plus {sigma}1, µ1WT pr-cores plus {sigma}1, or native ISVPs for 1 h at 4°C. The cells were then washed extensively to remove free viral particles and fixed. IF microscopy using the µ1-specific MAb 10H2 (67) revealed that binding of the µ1N42A pr-cores plus {sigma}1 to CV1 cells (Fig. 3A, bottom left) was comparable to that of µ1WT pr-cores plus {sigma}1 (Fig. 3A, top left) or ISVPs (data not shown). Essentially the same results were observed in L929 and Mv1Lu cells (data not shown). To compare the levels of binding more quantitatively, flow cytometry using Mv1Lu cells was performed. Mv1Lu cells were incubated with µ1N42A pr-cores plus {sigma}1, µ1WT pr-cores plus {sigma}1, or native ISVPs for 1 h at 4°C to allow attachment. The cells were then washed extensively to remove free particles, fixed, and stained using µ1 MAb 10H2. Cells incubated with any of the three particle types showed essentially identical levels of increased 10H2 staining in comparison to uninfected cells (data not shown), providing further evidence that the µ1N42A pr-cores plus {sigma}1 are fully competent for attachment. We also used a hemagglutination assay (26) to test cell-binding actvity. µ1N42A r-cores plus {sigma}1 displayed a relative (per-particle) capacity for hemagglutination very similar to those of µ1WT r-cores plus {sigma}1 and native virions (data not shown), indicating that the µ1N42A r-cores plus {sigma}1 are largely functional for binding to erythrocytes. In sum, these findings indicate that the infectivity barrier for µ1N42A particles follows attachment.



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  		FIG. 3. Attachment and protein synthesis during infection of cells by recoated particles. (A) CV1 cells were infected with µ1WT pr-cores (top row) or µ1N42A pr-cores (bottom row) (50,000 particles per cell) and then fixed at either 0 (left column) or 2 (right column) h p.i. The fixed cells were coimmunostained with a µNS-specific rabbit polyclonal antiserum followed by Alexa 594-conjugated goat anti-rabbit immunoglobulin G (red) and the µ1-specific mouse MAb 10H2 followed by Alexa 488-conjugated goat anti-mouse immunoglobulin G (green). Cell nuclei were counterstained with 4',6-diamidino-2-phenylindole (blue). Samples were examined by fluorescence microscopy. (B) Mv1Lu cells were infected with µ1WT pr-cores (top) or µ1N42A pr-cores (bottom) (20,000 particles per cell) and then fixed at either 0 (green), 4 (cyan), or 8 (magenta) h p.i. The fixed cells were stained with µNS-specific rabbit polyclonal antiserum followed by Alexa 488-conjugated goat anti-rabbit immunoglobulin G. Samples were examined by flow cytometry.

The synthesis of new viral proteins in infected cells was first evaluated by IF microscopy using a µNS-specific polyclonal antiserum (14). µNS was used as a marker of new protein synthesis, since it is a nonstructural protein that is produced during infection but is not present in input virions or r-cores plus {sigma}1. In CV1 cells infected with µ1N42A pr-cores plus {sigma}1 (Fig. 3A, bottom left), µ1WT pr-cores plus {sigma}1 (Fig. 3A, top left), or native ISVPs (data not shown), no µNS was detected at 0 h p.i. By 2 h p.i., however, µNS was clearly evident in cells infected with µ1WT r-cores plus {sigma}1 (Fig. 3A, top right) or ISVPs (data not shown), but not in cells infected with µ1N42A r-cores plus {sigma}1 (Fig. 3A, bottom right). Essentially the same differences among these particle types were observed in L929 cells at 0 and 2 h p.i. (data not shown). Similarly, µNS was detected by immunoblotting in lysates harvested as early as 2 h p.i. from CV1 cells infected with µ1WT pr-cores plus {sigma}1 but was not detected in lysates harvested as late as 24 h p.i. from CV1 cells infected with µ1N42A pr-cores plus {sigma}1 (data not shown). Flow cytometry after staining with µNS-specific polyclonal antiserum revealed that Mv1Lu cells infected with native ISVPs (data not shown) or µ1WT pr-cores plus {sigma}1 (Fig. 3B, top) showed increased levels of µNS at 4 h p.i and that nearly every cell was strongly µNS positive by 8 h p.i. In Mv1Lu cells infected with µ1N42A r-cores plus {sigma}1, in contrast, µNS remained virtually undetectable at both 4 and 8 h p.i. (Fig. 3B, bottom). These findings suggest that the infectivity barrier for µ1N42A particles occurs before or during protein synthesis. We next attempted to pinpoint the step between attachment and protein synthesis at which µ1N42A r-cores are blocked.

µ1N42A particles fail to permeabilize membranes in vitro. A membrane permeabilization defect of µ1N42A particles was demonstrated in vitro using a hemolysis assay (17, 19). When incubated with erythrocytes, ISVPs can permeabilize the cells' plasma membranes, leading to hemoglobin release and providing a useful model for membrane penetration during cell entry. Consistent with previous reports (15, 17-20), we found that both native ISVPs and µ1WT pr-cores (with or without {sigma}1) displayed robust hemolytic activity (Fig. 4A). µ1N42A pr-cores (with or without {sigma}1), in contrast, did not mediate hemolysis (Fig. 4A). Time course experiments confirmed the rapidity with which both ISVPs and µ1WT pr-cores (with or without {sigma}1) mediated hemolysis, compared to the failure of µ1N42A pr-cores (with or without {sigma}1) to mediate hemolysis over extended periods (Fig. 4B and data not shown). These findings suggest that µ1N/µ1C cleavage is an important step in the membrane permeabilization pathway and that the barrier to infectivity of µ1N42A particles may involve a specific defect in membrane penetration.



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  		FIG. 4. Hemolysis by µ1N42A recoated particles. (A) Samples of native ISVPs, µ1WT pr-cores (with or without {sigma}1), or µ1N42A pr-cores (with or without {sigma}1) were incubated with bovine calf erythrocytes (final concentration, 3% [vol/vol]) at 32°C for 30 min in the presence of 200 mM CsCl. The reactions were terminated by incubation on ice for 10 min, and the cells were pelleted by centrifugation at 300 x g for 5 min. The extent of hemolysis was determined by measuring the A405 of the supernatant and was expressed as a percentage (hemolysis by 1% Triton X-100 = 100%). Each bar represents the mean ± standard deviation from three trials. (B) Samples of µ1WT or µ1N42A pr-cores were evaluated for hemolysis as described for panel A, except that samples were harvested for analysis after different periods at 32°C. The results of a representative experiment are shown. Use of CsCl as a promoting agent is discussed in Materials and Methods.

µ1N42A particles exhibit other hallmarks of the ISVP -> ISVP* transition in vitro. ISVPs can transform into a distinct particle form, the ISVP*, that contains a protease-sensitive conformer of µ1, is more hydrophobic, has released {sigma}1, and has been activated for synthesis of viral plus-strand RNAs (15). The transition from ISVP to ISVP* precedes or accompanies hemolysis (15) and appears to precede or accompany membrane penetration during productive cell entry as well (18). In an attempt to determine why µ1N42A particles are defective for hemolysis, we examined whether they exhibit other hallmarks of the ISVP -> ISVP* transition in vitro.

(i) Increase in µ1 protease sensitivity. The µ1 conformer in ISVP*s (µ1*) is highly sensitized to protease degradation at 4°C (15). Upon conversion to ISVP*-like particles, µ1WT pr-cores also contained a protease-sensitive conformer of µ1 (Fig. 5A), as should occur if pr-cores are mimicking the behaviors of native ISVPs (15). Interestingly, samples of µ1N42A pr-cores incubated under the same conditions to promote the ISVP -> ISVP* transition also contained a protease-sensitive conformer of µ1 (Fig. 5A). None of the particle samples incubated under conditions that do not promote the ISVP -> ISVP* transition showed increased protease sensitivity of µ1 (Fig. 5A), indicating that they had not been converted to ISVP*s. We performed identical experiments in the presence of erythrocytes and looked for µ1 protease sensitivity following a hemolysis reaction. After immunoblotting for µ1, we obtained essentially the same findings with regard to changes in µ1 protease sensitivity in the three particle types that we obtained in the absence of erythrocytes (data not shown). We conclude that µ1N42A particles can exhibit at least one structural hallmark of the ISVP -> ISVP* transition, increased protease sensitivity of µ1, in the absence of µ1N/µ1C cleavage. Since the ISVP*-like µ1N42A particles are defective at hemolysis, however, the findings suggest that some missing element(s) of the transition in these particles is necessary for membrane permeabilization.



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  		FIG. 5. Capacity of µ1N42A recoated particles to undergo the ISVP -> ISVP* transition. (A) Protease sensitivity of µ1. Samples of µ1WT pr-cores plus {sigma}1 (lanes 1 and 2) or µ1N42A pr-cores plus {sigma}1 (lanes 3 and 4) were incubated with bovine erythrocytes, and hemolysis was performed as for Fig. 4 in the presence of CsCl or NaCl (200 mM). Following incubation at 32°C and removal to ice, the samples were incubated with trypsin (100 µg/ml) for 45 min on ice. The trypsin was then inactivated by addition of soybean trypsin inhibitor (300 µg/ml). Samples were subjected to SDS-PAGE, followed by immunoblotting with the µ1-specific MAb 10H2. (B) Hydrophobicity. Samples of µ1WT or µ1N42A pr-cores plus {sigma}1 were incubated in reaction buffer containing bis-ANS (25 µM) and either NaCl or CsCl (300 mM) for 30 min at 32°C. The levels of bis-ANS fluorescence were then measured on a fluorescence microplate reader (Spectramax; Molecular Dynamics) (excitation, 405 nm; emission, 485 nm). The results of a representative experiment are shown. (C) Release of {sigma}1. Samples of µ1WT pr-cores plus {sigma}1 (lanes 1 and 2) or µ1N42A pr-cores plus {sigma}1 (lanes 3 and 4) were incubated in reaction buffer containing either CsCl or NaCl (300 mM) for 30 min at 32°C. The samples were removed to ice for 10 min and then loaded atop a sucrose cushion (20% [wt/vol]; 500 µl). Following centrifugation in a Beckman TLA 100.2 rotor (90,000 rpm for 1 h at 5°C), the top fraction (200 µl) of the sucrose cushion was removed, concentrated by precipitation with trichloroacetic acid, and subjected to SDS-PAGE, followed by immunoblotting with a {sigma}1-specific rabbit polyclonal antiserum. (D) Transcriptase activation. Samples of µ1WT or µ1N42A pr-cores plus {sigma}1 were incubated in reaction buffer containing either CsCl or NaCl (300 mM) for 30 min at 32°C. The samples were removed to ice for 10 min and then incubated in transcription reaction buffer (see Materials and Methods) containing [{alpha}-32P]GTP at 37°C for 105 min. RNA products were concentrated by precipitation with trichloroacetic acid, and the radioactivity in each sample was measured by scintillation counting. Each bar represents the mean ± standard deviation from three separate trials. Use of CsCl as a promoting agent is discussed in Materials and Methods.

(ii) Increase in particle hydrophobicity. To address changes in hydrophobicity following the ISVP -> ISVP* transition, we used a fluorescent probe, bis-ANS, that binds to exposed hydrophobic regions of proteins (15). Consistent with previous findings, ISVPs showed increased binding of bis-ANS upon conversion to ISVP*s (data not shown). Both µ1WT pr-cores and µ1N42A pr-cores showed similarly increased binding of bis-ANS as well (Fig. 5B). In contrast, the control samples held under conditions that do not promote the ISVP -> ISVP* transition showed only limited increases in bis-ANS binding (Fig. 5B and data not shown), indicating they had not been converted to ISVP*s. We conclude that µ1N42A particles can exhibit a second structural feature of the ISVP -> ISVP* transition, increased hydrophobicity, and that µ1N/µ1C cleavage is not required for this increase to occur.

(iii) Release of {sigma}1. To assay for loss of attachment protein {sigma}1 that accompanies the ISVP -> ISVP* transition, we subjected particles to centrifugation through a sucrose cushion, during which particle-associated proteins should pellet while released proteins remain near the top of the cushion (15). Following centrifugation, the top portion of the sucrose cushion was collected and analyzed by SDS-PAGE and immunoblotting to detect the presence of released {sigma}1. Consistent with previous findings, {sigma}1 was released from ISVPs upon conversion to ISVP*s (data not shown). Both µ1WT pr-cores and µ1N42A pr-cores also showed release of {sigma}1 (Fig. 5C). In contrast, none of the control samples held under conditions that do not promote the ISVP -> ISVP* transition showed {sigma}1 release (Fig. 5C and data not shown), indicating that they had not been converted to ISVP*s. The {delta} or µ1{delta} fragment of µ1 remained particle associated in each of the ISVP* samples (data not shown), consistent with previous results (15). We conclude that µ1N42A particles can exhibit a third structural hallmark of the ISVP -> ISVP* transition, release of {sigma}1, and that µ1N/µ1C cleavage is not required for this release to occur.

(iv) Transcriptase activation. µ1 must adopt the hydrophobic, protease-sensitive conformation now known to be associated with ISVP*s in order for the particle-associated transcriptases to be activated from their latent state and become capable of synthesizing the full-length viral plus-strand RNAs (9, 15, 27, 42). To test if µ1N/µ1C cleavage is required for transcriptase activation in vitro, samples of µ1N42A or µ1WT pr-cores plus {sigma}1 were incubated under standard conditions either to promote or not to promote the ISVP -> ISVP* transition. The particles were then added to an in vitro transcription reaction mixture that included [{alpha}-32P]GTP. The RNA products were isolated by trichloroacetic acid precipitation, and 32P incorporation was measured by scintillation counting. In this assay, after incubation under conditions to promote the ISVP -> ISVP* transition, µ1N42A pr-cores plus {sigma}1 produced RNA transcripts at high levels, similar to those of µ1WT pr-cores plus {sigma}1 (Fig. 5D) or native ISVPs (data not shown). Only low levels of transcripts were produced, however, after each of the particle types was incubated under nonpromoting conditions (Fig. 5D and data not shown). Essentially the same results were obtained in experiments with µ1N42A and µ1WT pr-cores lacking {sigma}1 (data not shown). We conclude that µ1N42A particles can exhibit a major functional hallmark of the ISVP -> ISVP* transition, transcriptase activation, and thus that µ1N/µ1C cleavage is not required for this activation to occur.

In summary, the results indicate that µ1N42A pr-cores with or without {sigma}1 (ISVP-like particles) are competent to undergo important elements of the ISVP -> ISVP* transition in vitro but are nevertheless not functional for in vitro membrane permeabilization.

µ1N42A particles are also defective at membrane permeabilization during cell entry. Since µ1N42A pr-cores (with or without {sigma}1) failed to mediate membrane permeabilization in vitro, we hypothesized that the barrier to infectivity of µ1N42A particles may involve the membrane penetration step during cell entry. To test this hypothesis, we assessed the capacity of µ1N42A pr-cores plus {sigma}1 to mediate cytoplasmic coentry of the ribonucleotoxin {alpha}-sarcin, which in turn inhibits both cellular and viral protein synthesis (18, 50). L929 cells were infected with µ1WT pr-cores plus {sigma}1 or µ1N42A pr-cores plus {sigma}1 in the presence of {alpha}-sarcin, and levels of protein synthesis were measured at 20-min intervals p.i. In cells infected with µ1WT pr-cores plus {sigma}1 (Fig. 6A), inhibition of protein synthesis occurred within 20 min p.i. at 37°C, consistent with entry of the toxin along with penetrating viral particles and also consistent with previous results (18). Similar results were seen in cells infected with ISVPs (data not shown). In contrast, cells infected with µ1N42A pr-cores plus {sigma}1 showed no decrease in protein synthesis relative to uninfected cells (Fig. 6A), supporting the conclusion that the µ1N42A particles are defective at membrane permeabilization, and {alpha}-sarcin-based intoxication, during cell entry.



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  		FIG. 6. Behaviors of µ1N42A recoated particles in cells. (A) Coentry of {alpha}-sarcin during viral infection. L929 cells were incubated with µ1WT or µ1N42A pr-cores plus {sigma}1 at 4°C to allow attachment, washed with methionine-free medium containing [35S]methionine-cysteine with or without {alpha}-sarcin (50 µg/ml), and incubated at 37°C for different times, as indicated. The cells were then lysed, and macromolecules were concentrated by trichloroacetic acid precipitation. The radioactivity (in counts per minute) in the washed precipitates was measured by scintillation counting. The values shown represent the average of two experiments, each done in duplicate. (B) Antibody-detected structural changes in infected cells. CV1 cells were infected with µ1WT pr-cores plus {sigma}1 (top row) or µ1N42A pr-cores plus {sigma}1 (bottom row) (50,000 particles per cell) and then fixed at either 0 (left column) or 2 (right column) h p.i. The fixed cells were coimmunostained with a core-specific rabbit polyclonal antiserum followed by Alexa 594-conjugated goat anti-rabbit immunoglobulin G (red) and the µ1-specific mouse MAb 4A3 followed by Alexa 488-conjugated goat anti-mouse immunoglobulin G (green). Cell nuclei were counterstained with 4',6-diamidino-2-phenylindole (blue). Samples were examined by fluorescence microscopy. (C) Rescue of infectivity by top component. L929 cell monolayers were coinfected with µ1WT or µ1N42A r-cores plus {sigma}1 (1,000 per cell) plus top component particles (0, 1,000, 10,000, or 100,000 per cell). After 24 h at 37°C, the cells were lysed, and infectious titers were determined by plaque assay. Samples of cells infected with each amount of top component particles alone were identically generated and analyzed in parallel. Within each experiment, the latter titers were subtracted from the respective former titers to correct for any residual infectivity of the top component. Each bar represents the mean ± standard deviation of the results from three separate experiments.

µ1N42A particles exhibit hallmarks of the ISVP -> ISVP* transition in cells. The µ1-specific MAb 4A3 (67) recognizes a conformer of the {delta} region of µ1 that is present in ISVP*s both in vitro and in infected cells at early times p.i (18). This or associated changes in particle structure also allow the recognition of particles by a polyclonal antiserum specific for core surface proteins (18). The capacity of the {delta} region of µ1 in µ1N42A pr-cores plus {sigma}1 to undergo these conformation-specific changes in infected cells was addressed using IF microscopy. CV1 cells were incubated with µ1N42A or µ1WT pr-cores plus {sigma}1 in the presence of cycloheximide for 1 h at 4°C. At 0 and 2 h p.i., the infected cells were washed to remove unattached virus and fixed prior to being immunostained. At 0 h p.i., little staining was observed with either particle type and either antibody reagent (Fig. 6B, left), consistent with prior evidence that neither of these antibody reagents is effective at binding to ISVP-like particles (18). Also consistent with prior evidence (18), at 2 h p.i., cells infected with µ1WT pr-cores plus {sigma}1 displayed diffuse staining with the MAb 4A3 and punctate staining with the core-specific antiserum (Fig. 6B, upper right), indicating that the {delta} region of µ1 had undergone a conformational change, was now largely separated from the particles, and was probably free in the cytoplasm and nucleus. Results similar to these were obtained with native ISVPs in this (data not shown) and the previous study (18). In contrast, at 2 h p.i., cells infected with µ1N42A pr-cores plus {sigma}1 displayed punctate staining with both MAb 4A3 and the core-specific antiserum in colocalizing spots (Fig. 6B, lower right). These findings provide evidence that µ1N42A pr-cores plus {sigma}1 undergo structural changes associated with ISVP* formation during cell entry but that the conformationally altered {delta} region of µ1N42A fails to separate from particles in the same manner as µ1WT. Two possible explanations are that (i) the {delta} region of µ1N42A remains bound to particles and (ii) the {delta} region of µ1N42A dissociates from particles but both remain trapped within the same subcellular compartments (e.g., endocytic vesicles) so that their separation cannot be resolved by IF microscopy.

Infectivity of µ1N42A particles is rescued by coinfection with genome-deficient particles. Since µ1N42A particles can be activated for transcription, we reasoned that other viral particles may be capable of supplying membrane penetration activity in trans, enhancing the infectivity of µ1N42A particles. For this purpose, we chose reovirus top component particles, which have protein contents and structures essentially identical to those in virions but which cannot replicate in cells because they lack the viral genome (28, 61). Moreover, top component particles are known to mediate membrane penetration in vivo (13, 49), as well as in the hemolysis assay in vitro (M. L. Nibert, unpublished data). L929 cells were coinfected with either µ1N42A or µ1WT r-cores plus {sigma}1, along with increasing amounts of top component particles. The infections were allowed to proceed for 24 h, and viral growth was then measured by plaque assay. Cells infected with matching amounts of top component alone were analyzed in parallel to permit correction for any growth attributable to those particles. With increasing amounts of top component, growth attributable to µ1N42A r-cores plus {sigma}1 was greatly enhanced while growth attributable to µ1WT r-cores plus {sigma}1 was not affected (Fig. 6C). With the largest amount of top component tested, growth attributable to µ1N42A r-cores plus {sigma}1 was enhanced by >100-fold and approached 10% of the levels attained by µ1WT r-cores plus {sigma}1 (Fig. 6C). The finding that coinfections with top component particles greatly enhanced the infectivity of µ1N42A particles supports the hypothesis that µ1N42A particles are defective at the membrane penetration step but are otherwise functional for subsequent steps in the onset of viral replication.

Release of µ1N from ISVP*s. A likely possibility would be that µ1N/µ1C cleavage is critical to allow release of the N-myristoylated µ1N fragment (600 copies per virion) (29, 48, 55) before or during membrane penetration. We therefore tested for µ1N release from particles at the ISVP -> ISVP* transition in vitro, which had not been addressed in previous studies. ISVPs labeled with [3H]myristate, which is specifically incorporated as the N-terminal myristoyl group of µ1 and µ1N (55, 65), were used to monitor the particle association of µ1N. Separate samples of ISVPs were incubated under conditions that either converted them to ISVP*s or retained them as ISVPs. Upon centrifugation in CsCl gradients at 4°C, the particles should migrate to their known densities (~1.38 g/cm3) (15, 42) while released proteins remain near the top of the gradient. Following centrifugation, the gradients were fractionated, and scintillation counting was performed to identify fractions containing the radiolabel. In initial experiments, we found that the label could be efficiently recovered from ISVP* samples only when a nonionic detergent (e.g., Triton X-100) was included in the reaction mixture (data not shown). This was presumably due to the hydrophobic nature of µ1N, which caused it to stick to surfaces, such as that of the centrifuge tube, once released. In later experiments, we therefore included this detergent in both samples and consistently found that µ1N was retained in ISVPs but was indeed released from ISVP*s (Fig. 7A). This finding is consistent with the hypothesis that µ1N/µ1C cleavage is required during cell entry to allow the release of the myristoylated µ1N fragment from particles.



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  		FIG. 7. µ1N release from ISVP*s, model, and sequences. (A) µ1N release from ISVP*s. [3H]myristate-labeled ISVPs were incubated with either 300 mM NaCl (solid circles) or 300 mM CsCl (open circles) in the presence of 0.5% Triton X-100 for 30 min at 32°C. The particles were then purified on a 1.25- to 1.45-g/cm3 CsCl density gradient. The gradients were fractionated, and the radioactivity (in counts per minute) in each sample was measured by scintillation counting. The results of a representative experiment are shown. (B) Updated model of reovirus uncoating in vitro and during cell entry. Interactions of viral particles and proteins with cell surface receptors and possible localizations to specific subcellular compartments are not included in the diagram. The proposed uncoating intermediates and fates of the outer capsid proteins—µ1N (blue), µ1C (cyan), {sigma}1 (magenta), and {sigma}3 (purple)—are shown. (Step a) The virion (V) undergoes proteolytic processing to generate the ISVP (I). In ISVPs, the {sigma}3 protein has been degraded, and its differently sized fragments have been released. (Step b) The ISVP then undergoes a major structural transition to the ISVP' (I'). ISVP's lack {sigma}1 and contain an altered, hydrophobic conformer of µ1. Particles containing µ1HS are blocked at step b (18). (Step c) The ISVP' then undergoes a further transition to the ISVP* (I*), during which the remaining µ1N/µ1C cleavage occurs and the µ1N peptide is released. As suggested in the diagram (by the increase in particle diameter), further conformational changes in µ1C seem likely to accompany this transition. Particles containing µ1N42A are blocked at step c. (Steps d and e) The released µ1N peptides, putatively in concert with other portions of µ1C remaining in the ISVP*, effects membrane penetration (step d), and transcriptionally active core particles (C) are ultimately released into the cytoplasm (step e). The capacity of each particle type to mediate viral transcription is indicated below the diagram (+, able to mediate; –, unable to mediate). Note that cleavage of µ1C at the {delta}-{phi} junction during generation of ISVPs is not shown in this diagram, because the fate of {phi} during and after membrane penetration remains unknown. The cyan lines accompanying the core after step e specifically represent the released {delta} fragment (18). (C) Sequence comparison of the N-terminal regions of orthoreovirus (ORV) µ1 and aquareovirus (AqRV) VP4 proteins. Amino acids 2 to 43 of µ1 are shown on top, with corresponding residues of VP4 aligned below. Identical residues are shown in uppercase. The myristoylation consensus sequence in both proteins is indicated by the line. The N-terminal N-myristoyl group of µ1 is labeled (myr), and the site of the putative autocleavage is indicated by the arrowhead. The myristoyl group plus amino acids 2 to 42 constitute the µ1N peptide.


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DISCUSSION
 
Consistent with results of previous studies, in which Asn42 in µ1 was mutated to threonine, glutamine, glutamate, or histidine (65), we found that an alanine substitution at Asn42 completely blocked µ1N/µ1C cleavage. Autolytic capsid proteins from several other nonenveloped animal viruses also require an asparagine at the cleavage site (1, 46, 59, 64, 68; reviewed in reference 41), suggesting that this particular amino acid plays a key role in the cleavage mechanism. The current model for the cleavage mechanism of both flock house virus and N. capensis omega virus {alpha} proteins is that the nitrogen of the asparagine side chain nucleophilically attacks the carbonyl group of the peptide backbone immediately C-terminal to the asparagine residue (64). We are currently working to determine a crystal structure of the reovirus µ1N42A-{sigma}3 heterohexamer (L. Zhang, A. L. Odegard, M. L. Nibert, and S. C. Harrison, unpublished data) in preparation for further studies to dissect the µ1N/µ1C cleavage mechanism.

This report further illustrates the utility of r-cores (19, 20) for studies of cell entry by reovirus. Using this system, we were able to show that µ1N42A can assemble into a largely complete outer capsid surrounding the core, despite not having undergone the µ1N/µ1C cleavage. Most of the previous literature would have led us not to predict this, since µ1N/µ1C cleavage appears to occur upon association of µ1 and {sigma}3 and thus has been thought to be important for assembly (e.g., see references 45 and 65). Based partly on evidence in this study, we now believe that µ1N/µ1C cleavage must occur before or during membrane penetration, but not necessarily as a step in assembly. Other data indicate that in a fraction of the µ1 subunits, cleavage in fact does not occur during assembly but rather during disassembly of the µ1-{sigma}3 heterohexamer (48) or the particle-associated outer capsid. After disruption of virions under certain conditions preceding SDS-PAGE, a much larger fraction of the µ1 subunits remain uncleaved at the µ1N/µ1C junction, suggesting that at least a portion of this cleavage accompanies disassembly (M. L. Nibert, A. L. Odegard, M. A. Agosto, K. Chandran, and L. A. Schiff, unpublished data). The coupling of µ1N/µ1C cleavage to disassembly provides a plausible explanation for why the cleavage-defective mutant µ1N42A can assemble normally into r-cores.

Since proteolytic cleavage is required to prime many enveloped virus fusion proteins (reviewed in reference 37) and nonenveloped virus penetration proteins (reviewed in references 38 and 41) for membrane interactions, we reasoned that the µ1N/µ1C cleavage may allow µ1 to adopt its membrane-seeking conformation during cell entry. In this study, we obtained multiple points of evidence that µ1N/µ1C cleavage is indeed critical for membrane permeabilization and penetration by reovirus particles, but not for some earlier steps in entry, such as attachment. If membrane penetration activity was supplied in trans by genome-deficient top component particles containing µ1WT, the µ1N42A particles displayed much higher levels of infectivity. The large number of top component particles needed to maximize this effect may reflect the inefficiency with which appropriate forms of the two particles were colocalized to the same subcellular compartment, from which membrane penetration could then occur. Thus, although we hypothesize that the defect in µ1N42A particles involves the membrane penetration step per se, we recognize that this mutation could alternatively or additionally affect particle delivery to the appropriate subcellular location.

Recent studies have proposed a model in which ISVPs undergo a series of structural and functional changes to yield a new particle form, the ISVP*, before or during membrane penetration (15, 18). Structurally, ISVP*s contain an altered conformer of µ1 and have lost the attachment protein {sigma}1 (15, 18, 42). Functionally, these particles not only are associated with membrane permeabilization in vitro and in vivo but also have been activated to transcribe the viral plus-strand RNAs (15, 18). The evidence in this study that µ1N42A particles can progress only partially through the transition to ISVP*s leads us to propose a new uncoating intermediate, the ISVP', at which particles are trapped in the absence of µ1N/µ1C cleavage (Fig. 7B). The existence of this intermediate is supported by evidence that in a fraction of the µ1 subunits, µ1N/µ1C cleavage directly accompanies the ISVP -> ISVP* transition (Nibert et al., unpublished). The ISVP' is thus proposed to be the particle form that immediately precedes this cleavage. Another mutant, µ1HS, has also recently been characterized using the r-core system (18) and shown to be blocked at an earlier step, between the ISVP and the newly defined ISVP' (Fig. 7B). The results indicate that µ1HS particles do not mediate membrane permeabilization, but neither do they exhibit any hallmarks of the ISVP -> ISVP* transition. Thus, through studies with r-cores containing different mutations in µ1, we appear to be dissecting the uncoating and membrane penetration processes of reoviruses into a series of discrete steps. Trapped uncoating intermediates obtained with these mutants may be useful for other biochemical and structural studies to characterize the changes in µ1 that allow it to interact with and alter the membrane for particle translocation.

Whether the released µ1N peptide plays an active role in membrane penetration or must simply be discarded before other regions of µ1 can act remains to be determined. The very low residual infectivity of µ1N42A particles (Table 1) is consistent with either possibility. For example, the µ1N peptide might be capable of playing an active role, but with greatly reduced efficiency, even if it remains tethered to the {delta} region of µ1C. If µ1N indeed plays an active role, there are several possible reasons why it must be cleaved and released for maximal activity. The freed C-terminal end of µ1N may be necessary for the released peptide to fold into a particular conformation for its activity. The cleavage and release may also be necessary to allow the peptide to diffuse farther from the particle than would be possible if it remained tethered to the {delta} region of µ1C (Fig. 1A). Yet another possibility is that the cleavage and release may be necessary to allow large numbers of µ1N peptides to associate, either in solution or in the membrane. One simple idea is that of a "µ1N bullet, " which is ejected from the particle and interacts with the nearby membrane, directly participating in the penetration process that allows the particle to enter the cytoplasm. The N-terminal N-myristoyl group is an obvious candidate to play a key role in this proposed function, although the µ1N peptide sequence is itself quite hydrophobic (Fig. 7C). Preliminary evidence obtained with r-cores containing a myristoylation-defective µ1 protein suggest that these particles are indeed poorly infectious (I. S. Kim, M. A. Agosto, K. Chandran, and M. L. Nibert, unpublished data). Notably, the µ1N region (including the myristoylation signal and autocleavage site) is highly conserved with those parts of the 69-kDa outer capsid protein VP4 of aquareoviruses (Fig. 7C) and much more conserved than is the µ1C region with the rest of VP4 (2, 48), suggesting that µ1N and the N-terminal portion of the aquareovirus protein share conserved functions.

In several other nonenveloped animal viruses, small and sometimes N-myristoylated peptides are known or thought to play critical roles in cell entry. One well-known example is the autocleavage product VP4 of poliovirus. After binding of a poliovirus virion to its cell surface receptors, and either before or after uptake and acidification along the endocytic pathway, the 65- to 70-residue, N-terminally N-myristoylated VP4 peptide is released from the particle and the previously buried N terminus of VP1 is externalized (reviewed in reference 38). A favored hypothesis for polio- and rhinovirus membrane penetration is that the newly exposed N terminus of VP1, possibly in concert with VP4, interacts with the plasma or endosomal membrane to form a pore through which the genomic RNA is extruded into the cytoplasm (6, 24, 46, 47, 66). As with µ1N, the exact role of VP4 is not known. However, Danthi et al. and Moscufo et al. have described mutations in VP4 that severely impact infection even though the virus particle binds to the cell surface and undergoes all of its normal entry-related structural changes, including VP0 cleavage and VP4 release, suggesting that the released VP4 plays a direct role in entry (25, 52). Similarly, in flock house virus, the 44-residue autocleavage product, {gamma}, is thought be directly involved in the membrane penetration process (8, 59; reviewed in reference 41). Mutations that block the autocleavage needed to generate {gamma} render the virus very poorly infectious (59). In addition, an {alpha}-helical region of the {gamma} peptide has been shown to interact with and permeabilize membranes, supporting the hypothesis that {gamma} is directly involved in penetrating the endosomal membrane during cell entry (8). A similar role has been proposed for the 74-residue {gamma} peptide of N. capensis omega virus (41, 53).

Further studies of these and other systems should provide additional comparisons for obtaining a more complete mechanistic description of how different nonenveloped viruses traverse the membrane barrier during cell entry. In such comparisons, it is important to note that reovirus virions are much larger than those of poliovirus, flock house virus, or N. capensis omega virus (85 versus 30 to 45 nm); that reovirus µ1 occupies a different structural position than does poliovirus VP0, flock house virus {alpha}, or N. capensis omega virus {alpha} (T=13 outer capsid versus T=3 or 4 single capsid); and that the entry payload of reovirus is substantially different from those of the other viruses (transcriptase particle versus plus-strand RNA). Despite these differences, these diverse nonenveloped viruses have now been suggested to share hydrophobic-peptide release following capsid-protein autocleavage as part of a general mechanism of membrane penetration during entry.


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ACKNOWLEDGMENTS
 
We thank H. Ploegh and his laboratory members for access to and assistance with flow cytometry instrumentation and experiments; E. C. Freimont and J. B. Dinoso for superb technical assistance; other members of our laboratory, D. Bubeck, M. Ehrlich, S. C. Harrison, J. M. Hogle, T. Kirchhausen, and L. Zhang, for helpful discussions; and M. A. Agosto, I. S. Kim, and K. S. Myers for comments on the manuscript.

This work was supported in part by NIH grants K08 AI52209 to J.S.L.P., R01 GM33050 to T.S.B., and R01 AI46440 to M.L.N.; a Keck Foundation award to the Purdue Structural Biology group for purchase of the CM300 FEG microscope; and a Purdue University reinvestment grant to the Structural Biology group. K.C. was additionally supported by a Fields postdoctoral fellowship made available to the Department of Microbiology and Molecular Genetics through the generosity of Ruth Peedin Fields.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Microbiology and Molecular Genetics, Harvard Medical School, 200 Longwood Ave., Boston, MA 02115. Phone: (617) 645-3680. Fax: (617) 738-7664. E-mail: mnibert{at}hms.harvard.edu. Back

{dagger} Present address: Hematology Division, Department of Medicine, Brigham and Women's Hospital, Boston, MA 02115. Back

{ddagger} Present address: James A. Baker Institute for Animal Health, College of Veterinary Medicine, Cornell University, Ithaca, NY 14853. Back


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Journal of Virology, August 2004, p. 8732-8745, Vol. 78, No. 16
0022-538X/04/$08.00+0     DOI: 10.1128/JVI.78.16.8732-8745.2004
Copyright © 2004, American Society for Microbiology. All Rights Reserved.



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