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Previous Article | Next Article 
Journal of Virology, April 2002, p. 4008-4021, Vol. 76, No. 8
0022-538X/02/$04.00+0 DOI: 10.1128/JVI.76.8.4008-4021.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.
Gabriel Rutter,3 Dennis Strand,4 and Ralf Bartenschlager1*
Institute for Virology,1 Institute for Internal Medicine, Johannes-Gutenberg University Mainz, 55131 Mainz,4 Department of Molecular Pathology, Heinrich-Pette Institut, 20251 Hamburg, Germany,3 Virco Ltd., Cambridge CB4 0G1, United Kingdom2
Received 31 October 2001/ Accepted 16 January 2002
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ABSTRACT |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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HCV particles are enveloped, have a diameter of 55 to 65 nm, and harbor an 
9,600-nucleotide-long plus-strand RNA genome. It carries a single long open reading frame (ORF), which is flanked by highly conserved and structured nontranslated regions (NTRs), both of which have been shown to be required for RNA replication (25, 42, 70). The 5' NTR also harbors an internal ribosome entry site (IRES) which directs the expression of a large polyprotein that is co- and posttranslationally cleaved by cellular and viral proteases into at least 10 mature viral proteins (3, 59). Analogous to other flaviviruses, the nonstructural proteins presumably form an ordered replicase complex, which associates with intracellular membranes. Even though the exact conformation of this complex is currently unknown, extensive in vitro studies have identified several enzymatic activities within the nonstructural proteins shedding light on the organization and function of the replication machinery. The NS3 protein is the key protease of HCV which in conjunction with NS4A mediates all cleavages in the NS3-to-NS5B region (4, 28, 69). Furthermore, NS3 harbors nucleoside triphosphatase and helicase activities (40, 66). While the function of NS4B in the viral replication cycle remains to be defined, NS5A was found to be a highly phosphorylated polypeptide that may be involved in the resistance to the antiviral effects elicited by alpha interferon (23, 24, 26, 27). Interestingly, a large number of adaptive mutations that increase the replication efficiency of subgenomic HCV replicons in Huh-7 cells map within the NS5A gene, in part affecting potential phosphorylation sites (9, 30, 43, 48). Although this suggests a direct role of this protein in RNA replication, the exact mechanism is currently obscure. NS5B, which is the most C-terminal cleavage product of the polyprotein, constitutes the RNA-dependent RNA polymerase (RdRp) (8, 47).
The viral structural proteins core, E1, and E2 are located in the N-terminal third of the ORF separated from the nonstructural proteins by a short hydrophobic polypeptide (p7) of unknown function. The HCV core protein has been shown to bind RNA and is believed to be responsible for genome packaging (44). Two major core species with apparent molecular masses of 23 and 21 kDa (p21 and p23) have been described, corresponding to the unprocessed precursor and the processed form that lacks at its C terminus the signal sequence of E1 (36, 46, 53, 61, 71). As p21 predominates both in transfected tissue culture cells and in virus particles from infected sera, it is believed to be the mature form (71). The glycoproteins E1 and E2 are liberated by signalase cleavages and associate to form noncovalently linked heterodimers which are retained in the endoplasmic reticulum (ER) due to the presence of specific retention signals within their respective transmembrane domains (15-18, 20, 22, 29, 58, 64; for a recent review, see reference 19). In addition to these native heterodimeric glycoprotein complexes which are stabilized by noncovalent interactions, the production of disulfide-linked E1-E2 aggregates is well documented (18, 20). Although it is generally thought that the native complexes are the prebudding form, the specific contribution of either glycoprotein form to virus assembly is currently unknown.
Due to low virus titers, isolation and characterization of HCV particles from the sera of infected individuals are difficult. Furthermore, the virus population in these sera is very heterogeneous, as viral particles associate with immunoglobulins (12, 31, 68) and ß-lipoproteins (52, 57, 67), complicating direct morphological studies. Different buoyant densities of HCV derived from patient sera have been reported. HCV-immunoglobulin complexes from the sera of chronically infected patients exhibit an intermediate density ranging between 1.18 and 1.21 g/ml (12). A second population of HCV with a very low density of 1.08 to 1.11 g/ml in sucrose was observed and most likely represents the infectious form of the virus (10, 31, 52). Recent studies revealed the presence of significant amounts of viral particles with a buoyant density of 1.32 to 1.34 g/ml in CsCl density gradients and the physicochemical, morphological, and antigenic properties of HCV nucleocapsids in plasma samples from infected individuals (50). These structures closely resemble nonenveloped virus-like particles produced in insect cells upon infection with recombinant baculoviruses harboring cDNA for the structural region of an HCV genotype 1b isolate (7). Using this system, large amounts of both nonenveloped and enveloped virus-like particles can be prepared, which are important surrogate models that can be used to investigate virus receptor usage and entry into the host cell.
A variety of different approaches, mainly based on infection of primary human liver cells or diverse cell lines of hepatic or lymphoid origin with human patient sera have been explored in order to establish a robust in vitro infection system for HCV (5, 39). However, so far the success of these attempts has been limited due to the extremely low replication levels that do not allow detailed studies. The recent development of subgenomic HCV replicons, which allow high-level replication of HCV RNAs in cell culture, has overcome this hurdle (49). However, since the complete structural region was deleted, virus assembly and particle release could not be studied with this system. Moreover, important information about a potential influence of the structural proteins on the host cell could not be obtained.
To establish a system that allows efficient replication of the complete HCV genome in cell culture, in this study we constructed cell culture-adapted selectable full-length HCV genomes (sfl genomes). We show that these genomes stably replicate and express all viral proteins for prolonged cultivation periods. Furthermore, an authentic HCV genome that carries cell culture-adaptive mutations was generated and shown to replicate to high levels in transiently transfected cells. However, despite the release of nuclease-resistant HCV RNA into the cell culture supernatant, no evidence for virus particle assembly was found by either approach, suggesting that host cell factors important for virus production are not present in Huh-7 cells.
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MATERIALS AND METHODS |
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Plasmid construction. Full-length HCV constructs were generated based on the consensus genotype 1b isolate Con1 obtained from a German patient (41) (EMBL database accession number AJ238799). To create a construct with the complete HCV ORF, we first introduced an NcoI site at the beginning of the HCV ORF into the pTMC1 vector harboring the Con1 isolate downstream of the T7 promoter (41). Due to this PCR-based mutagenesis, the second amino acid in core was changed from serine to glycine. The entire HCV ORF and the 3' NTR were then transferred to the subgenomic replicon vector pFK-I389neo/NS3-3'/wt (48, 49) (EMBL accession number AJ242654) using NcoI and SpeI restriction sites. The resulting plasmid pFK-I389/Core-3'/wt harbors a bicistronic full-length genome with the authentic HCV 5' NTR directing the expression of the neomycin phosphotransferase gene (neo) and an IRES element derived from the encephalomyocarditis virus (EMCV) permitting expression of the complete HCV polyprotein. To increase the replication efficiency, the SfiI fragment (positions 3615 to 8492; numbering according to the published Con1 sequence) derived from the highly adapted subgenomic replicon rep 5.1 (43) was introduced into the full-length construct. The transferred DNA carries three adaptive mutations (E1202G, T1280I, and S2197P; numbering according to the polyprotein of the published Con1 sequence) and four further mutations that do not affect replication efficiency in cell culture (43). The Con1-ET construct used for transient-replication assays is a derivative of HCV strain Con1 that was modified by PCR-based techniques to possess three cell culture-adaptive mutations (E1202G, T1280I, and K1846T; V. Lohmann et al., unpublished results). Similarly, a subgenomic replicon (replicon-ET) that carries these mutations and the sh ble gene (Zeozine resistance gene) in place of neo was created.
Electroporation, generation of cell lines bearing sfl genomes, and transient-replication assay. In vitro transcription was performed as described previously (49). To generate cell lines harboring sfl genomes, 4 x 106 Huh-7 cells were electroporated with 1 µg of purified in vitro transcripts. Cells were seeded into three 10-cm-diameter-dishes and after 24 h were selected with G418 at a concentration of 100, 250, or 500 µg/ml, respectively. Approximately 3 to 5 weeks postelectroporation, small colonies were visible; these colonies were isolated and expanded for further analysis. For transient-replication assays, 2.4 x 107 Huh-7 cells were electroporated with 60 µg of HCV RNA. At the time points given, cells were harvested and analyzed by Northern and Western blotting. At 96 h posttransfection, supernatants were harvested, cleared of cellular debris, and subjected to density gradient centrifugation as described below.
Northern blot analysis and quantification of HCV RNA. The methods used for preparation of total RNA and Northern blotting have been described recently (56). HCV- and ß-actin-specific bands were quantified by phosphorimaging using a BAS 2500 scanner (Fuji), and the number of replicon molecules was determined by comparison with a serial dilution of in vitro transcripts loaded in parallel onto the gel. ß-Actin was used to correct for different amounts of total RNA loaded in each lane of the gel.
Amplification of sfl genome RNA, cloning of amplified DNA fragments, and sequence analysis. The complete HCV ORF from cell lines 20-1 and 21-5 (passages 52 and 57, respectively) was amplified in two overlapping fragments by reverse transcription-PCR (RT-PCR) as described recently (48) with minor modifications. One microgram of total RNA was mixed with 50 pmol of primer A6103 (5'-GCT ATC AGC CGG TTC ATC CAC TG-3') or A9412 (5'-CAG GAT GGC CTA TTG GCC TGG AG-3') in a total volume of 10.5 µl. Samples were denatured for 10 min at 65°C and then subjected to reverse transcription with the Expand-RT system (Roche Biochemicals, Mannheim, Germany) as recommended by the manufacturer in a total volume of 20 µl. After 1 h at 42°C, 1/10 of the reaction mixture was used for PCR with the Expand Long Template PCR kit (Roche Biochemicals) according to the instructions of the manufacturer. Cycle conditions were as follows: an initial denaturation step for 2 min at 94°C, followed by 40 cycles, with 1 cycle consisting of 10 s at 94°C, 90 s at 54°C, and 5 min at 68°C. After 10 cycles, the extension time was increased by 10 s for each additional cycle. Finally, the reaction mixture was incubated for 10 min at 68°C. The 5' PCR was performed with primers S59 (5'-TGT CTT CAC GCA GAA AGC GTC TAG-3') and A4919 (5'-AGC ACA GCC CGC GTC ATA GCA CTC G-3'), and the PCR product was inserted into pFK-I389neo/NS3-3'/wt after restriction with AseI and SalI. The 3' PCR was performed with primers S4542-HindIII (5'-CCA AGC TTG ATG AGC TCG CCG CGA AGC TGT CC-3') and A9386-SpeI (5'-CGT ACT AGT TAG CTC CCC GTT CAT CGG TTG G-3'), and after restriction with HindIII and SpeI, the fragment was inserted into a modified pUC18 vector. Sequencing was performed with a set of 44 primers, the ABI PRISM Big Dye Terminator Cycle Sequencing Ready Reaction Kit, and ABI PRISM DNA Analyzer 3700 or 377 (PE Applied Biosystems, Warrington, United Kingdom). The Sequencher software (Gene Codes Corporation, Ann Arbor, Mich.) was used for the assembly and analysis of the sequences.
Western blotting and immunofluorescence. Cells were washed once with phosphate-buffered saline (PBS) and detached from the plate by treatment with 0.05% trypsin-0.02% EDTA. Cells contained in a small aliquot of the suspension were counted, and the remaining cells were lysed by sonification in denaturing protein sample buffer (200 mM Tris-HCl [pH 8.8], 5 mM EDTA, 0.1% bromophenol blue, 10% sucrose, 3.3% sodium dodecyl sulfate [SDS], 2% 2-mercaptoethanol [2-ME]). Aliquots of cell lysates corresponding to 4 x 105 cells were analyzed by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) (11% polyacrylamide) and transferred to a polyvinylidene difluoride membrane (PolyScreen; NEN Life Science Products, Zaventem, Belgium) using a semidry blotter (Bio-Rad, Munich, Germany) according to the manufacturer's instructions. NS5B and core protein were detected as described previously using monoclonal antibodies that were kindly provided by D. Moradpour (56). For immunofluorescence, cells were grown on glass coverslips. Three days postseeding, cells were incubated with medium containing 20 nM brefeldin A (Molecular Probes, Eugene, Oreg.) for 3 h or left untreated. Subsequently, cells were fixed with methanol-acetone, and HCV proteins were detected essentially as described previously (56). Briefly, either a core-specific mouse monoclonal antibody or a monoclonal rat antibody directed against E2, kindly provided by J. McKeating, was used at a dilution of 1:100 or 1:500, respectively. Anti-human Golgin 97 mouse monoclonal antibodies (Molecular Probes) were used at a dilution of 1:300 to stain the Golgi complex. Bound antibodies were detected with Cy3-conjugated goat anti-mouse immunoglobulin G (IgG) (Dianova, Hamburg, Germany) or fluorescein isothiocyanate (FITC)-conjugated rabbit anti-rat IgG (Sigma, Deisenhofen, Germany) at a concentration of 1:1,000 or 1:500, respectively. The cell nucleus was counterstained employing bisbenzimide (Hoechst 33342; Sigma) at a concentration of 10 µg/ml. Cells were examined with a Zeiss 510 laser scan microscope.
Metabolic radiolabeling of proteins, immunoprecipitation, and deglycosylation. A total of 7 x 104 cells were seeded in a 3-cm-diameter culture dish in complete DMEM supplemented with Geneticin. Four days later, cells were washed, starved in methionine-free medium for 60 min, and incubated for 3 h in methionine-free DMEM supplemented with Express Labeling Mix (200 µCi/ml; NEN Life Science Products). Cells were lysed directly. For pulse-chase analysis, the cells were washed several times and incubated in complete DMEM for the time periods given in Results. Cell lysis and immunoprecipitation under nondenaturing conditions have been described previously (6). For deglycosylation studies, washed immunocomplexes were denatured for 10 min at 95°C in denaturing buffer (0.5% SDS, 1% 2-ME) and divided into two aliquots. Samples were incubated with or without 2,000 U of endoglycosidase H (New England Biolabs, Frankfurt, Germany) for 1 h at 37°C. Deglycosylation was stopped by the addition of an equal volume of protein sample buffer. Samples were heated at 95°C for 5 min, and proteins were analyzed by SDS-PAGE (10% polyacrylamide). For immunoprecipitations performed under nonreducing conditions, lysis buffer (50 mM Tris-Cl [pH 7.5], 150 mM NaCl, 1% Nonidet P-40 [NP-40], 1% sodium deoxycholate, 0.1% SDS) was supplemented with 20 mM iodoacetamide (Sigma) to prevent oxidation of thiol groups during the immunoprecipitation procedure. Prior to SDS-PAGE, samples were denatured in protein sample buffer lacking 2-ME.
EM. The procedures used for electron microscopy (EM) and immuno-EM have been described recently (56) and were applied with the following modifications. The grids were kept overnight at 4°C in a mixture of rabbit polyclonal antiserum against E2 glycoprotein (41) and a mouse monoclonal antibody specific for the core polypeptide of HCV, kindly provided by D. Moradpour. After the grids were washed five times in TCG (150 mM NaCl, 20 mM Tris-HCl [pH 7.5], 0.5% casein, 0.2% gelatin), they were treated with a cocktail consisting of goat anti-rabbit antibodies and anti-mouse antibodies coupled to colloid gold particles with a diameter of 10 and 6 nm, respectively (Dianova). After 1 h of incubation at room temperature, the reaction was terminated by repeated rinsing in TCG. Finally, the grids were washed with distilled water, stained with 4% aqueous uranyl acetate and Reynold's lead citrate, air dried, and examined at 60 kV with a Philips CM120 EM. For controls, we used ultrathin sections of Huh-7 cells treated as described above and thin sections of cells with full-length replicons that were incubated with a cocktail consisting of irrelevant antibodies (polyclonal rabbit antiserum directed against the capsid protein of human immunodeficiency virus type 1 [HIV-1] and mouse monoclonal antibodies specific for measles virus phosphoprotein). Gold-labeled secondary antibodies were the same as described above. Additional controls were done by omitting the primary antibodies from the labeling procedure.
Ultracentrifugations and nuclease treatment of cell culture supernatants.
Supernatant (10 to 30 ml) derived from 1 x 107 to 3 x 107 cells was harvested 4 to 7 days postseeding, and cellular debris was removed by low-speed centrifugation at 13,000 x g for 30 min at 4°C, followed by filtration with 0.45-µm-pore-size filters (Millipore, Molsheim, France). Supernatant bearing infectious bovine viral diarrhea virus (BVDV) was harvested 24 h postinfection of R.D.420 bovine testicle cells (kindly provided by R. Donis) with virus stocks of cytopathic BVDV strain NADL (kindly provided by R. Donis). For nuclease resistance assays, samples were spun in an L8-80 M Beckman ultracentrifuge (Beckman, Palo Alto, Calif.) at 110,000 x g at 4°C for 6 h. Sediments were resuspended in PBS, and different aliquots were supplemented with various amounts of RNase A and NP-40 as indicated in Results. After 4 h of incubation at room temperature, RNA was prepared by phenol-chloroform extraction as described below. For density gradient analyses, 30 ml of cleared culture fluid was spun over a 40% (wt/vol) iodixanol (Optiprep; Invitrogen, Karlsruhe, Germany) density cushion (
= 1.215 g/ml), prepared in CSM (0.85% [wt/vol] NaCl, 10 mM Tricine-NaOH [pH 7.4]; = 1,006 g/ml) for 6 h at 52,000 x g at 4°C. The supernatant was discarded, and the interface and density cushion (2.5 ml) were transferred to the bottom of a fresh tube. Samples were overlaid with a linear iodixanol gradient (28 to 0%) and spun for 16 h at 110,000 x g at 4°C. Twelve fractions (1 ml each) were taken from the top of the tube, and RNA was prepared as described below.
RNA preparation and quantitative Taqman RT-PCR. Gradient fractions and resuspended sediments were supplemented with 0.1% SDS and extracted with an equal volume of a phenol-chloroform mixture (1:1). After 15 min of centrifugation at 18,000 x g, the nucleic acids in the aqueous phase were transferred to a new tube, precipitated with isopropanol, washed once with 70% ethanol, and resuspended in 20 µl of double-distilled water. Five microliters of the sample was used for quantitative RT-PCR analysis using an ABI PRISM 7700 Sequence Detector (Taqman; Perkin-Elmer, Wellesley, Mass.). Amplifications were conducted in duplicate with the One Step RT-PCR Kit (Qiagen, Hilden, Germany) using the following primers and 3'-phosphate-blocked, 6-carboxyfluorescine (6-FAM)- and tetrachloro-6-carboxyfluorescine (TAMRA)-labeled probes (TIB Molbiol, Berlin, Germany): BVDV Taqman probe, 5'-6FAM-ATA GCA GCT CTG AGC TTT GGT TGA GGC AT-TAMRA-3'; BVDV-S6291, 5'-CCG GGA TAA CCT ATG CAT CAT ACG-3'; BVDV-A6401, 5'-TGG CAC AAT GGT ATT CAT CTA AGA-3'; HCV Taqman probe, 5'-6FAM-TCC TGG AGG CTG CAC GAC ACT CAT-TAMRA-3'; HCV-S66, 5'-ACG CAG AAA GCG TCT AGC CAT-3'; and HCV-A165, 5'-TAC TCA CCG GTT CCG CAG A-3'. Reactions were performed in three stages under the following conditions: stage 1, 60 min at 50°C (reverse transcription reaction); stage 2, 15 min at 95°C (heat inactivation of reverse transcriptase and activation of Taq polymerase); stage 3, 40 cycles, with 1 cycle consisting of 15 s at 95°C and 1 min at 60°C (PCR amplification). The total volume of the reaction mix was 15 µl and contained the following components: 2.66 µM 6-carboxy-X-rhodamine (Rox, passive reference), 4 mM MgCl2, 0.66 mM deoxynucleoside triphosphates, 0.266 µM probe, 1 µM (each) sense and antisense primer, and 0.6 µl of enzyme mix. The amounts of HCV and BVDV RNA were calculated by comparison to serially diluted in vitro transcripts included in the RT-PCR analysis.
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RESULTS |
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1.5 x 107 to 2.5 x 107 molecules per µg of total RNA (data not shown), which is about fivefold lower than the copy number of a subgenomic replicon present in cell line 9-13 (108 molecules per µg of total RNA) (49). Stability of sfl genomes and genetic drift. To ensure stable high-level replication of sfl genomes in established cell lines, the cells were continuously maintained in the presence of G418. Under these conditions, only those cells with sufficient amplification of the RNA that carries the selectable marker (neo) could propagate. Hence, selective pressure assured only the maintenance of a functional replicase complex, whereas the viral structural proteins that did not contribute to viral replication might even pose a disadvantage because of potential cytopathogenicity or the increased length of the RNA. In view of these considerations, we wanted to assess whether the sfl genome was stable over extended periods of time. Therefore, six different cell lines carrying an sfl genome were cultured under continuous selection for almost 6 months, corresponding to about 50 passages. Total RNA was isolated from cells at various time points and probed by HCV-specific Northern blotting. As shown in Fig. 2A, sfl genomes in all cell lines included in this follow-up were stable. Interestingly, in cell lines 20-1 and 22-1, shorter RNA molecules coreplicated with the full-length genomes, but at later time points these shorter RNAs disappeared, evidence that the deletions had no selective advantage (data not shown). In summary, these data show that the sfl genome is stably maintained for at least 6 months.
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6 months. HCV RNA was amplified by RT-PCR in two overlapping fragments, and several clones of each fragment were sequenced. For both the structural and nonstructural regions of the genome, only 40% of all nucleotide substitutions detected were silent. An example of the amino acid substitutions found in sfl genomes that were recovered from cell line 21-5 that illustrates the distribution of amino acid changes throughout the complete HCV polypeptide is shown in Fig. 2B. The coding region for the structural proteins appeared to be slightly more likely to acquire mutations than the nonstructural portion of the genome with the exception of the NS5A protein. Interestingly, six substitutions were present in both sequenced clones (K6R, V1282A, G1308S, E1709V, S2327L, and W2405R), which implies that they are dominant in the population of RNA molecules replicating in the cell line at the time the cells were analyzed (passage 57) (Fig. 2B). In summary, our sequencing analysis revealed a high genetic variability throughout the entire polyprotein with an average mutation rate of 8.5 x 10-3 substitutions per amino acid position per 6 months. The highest drift rates (12.2 x 10-3 to 17.4 x 10-3 amino acid changes) were found with the core protein, E1, and p7, implying that the structural region is more variable than the remainder of the sfl genome. This result is in keeping with the notion that only the viral replicase complex is under selective pressure, whereas the structural proteins are subject to free genetic drift. Characterization of structural protein expression in cell lines with an sfl genome. Having developed cell lines that harbor stably replicating HCV sfl genomes, we next investigated the expression and maturation of the structural proteins because this is a critical prerequisite for virus formation. As shown in Fig. 3A, the core protein was efficiently expressed in four different cell lines with sfl genomes as a single species with an apparent molecular mass of 21 kDa. This protein most likely corresponded to the processed form of core (positions 1 to 174) from which the signal sequence of E1 had been removed. As evidenced by the pulse-labeling experiment performed on cell line 21-5 and subsequent nondenaturing immunoprecipitations, E2 was expressed as a rather stable E2-p7-NS2 precursor, which could be detected with a chase period of 6 h (Fig. 3B). Furthermore, E2 released from the precursor and small amounts of coprecipitating E1 and NS2 were observed for at least 12 h. Deglycosylation of immunocomplexes with endoglycosidase H revealed the absence of terminally glycosylated forms of both E1 and E2 in the cell line (Fig. 3B). This observation suggests that the glycoproteins were retained in the ER and/or early cis-Golgi compartment and is consistent with earlier reports which report ER retention of both envelope proteins due to the presence of retention signals within their respective transmembrane domains (15, 16, 18, 22, 51).
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Morphology of cells with an sfl genome and subcellular localization of structural proteins. In order to investigate potential cytopathic effects exerted by the replication of sfl genomes and the expression of all HCV proteins, we first analyzed cultures of parental Huh-7 control cells by EM. Naive Huh-7 cells displayed a loose inconspicuous cytoplasmic architecture with variable numbers of vesicles of various sizes (Fig. 4B). The degree of vesiculation was apparently dependent on the cell density and correlated with the culture time after seeding. The vesicles were heterogeneous in size and contained homogenous material that exhibited gray staining. Similarly, cell lines 20-1 and 21-5 with an sfl genome and cells with a subgenomic replicon revealed strong cytoplasmic vesiculation. Since the frequency and distribution of vesicles in the cytoplasm did not depend on the presence of replicons, we concluded that these structures represented a general feature of Huh-7 cells (56) (data not shown). However, vesicles in cells bearing an sfl genome were delimited to different degrees by an electron-dense region resembling a thickened membrane (Fig. 4A) that distinguished them from the structures observed in parental Huh-7 cells and cells bearing subgenomic replicons, which are devoid of this electron-dense rim (compare Fig. 4A and B). Additionally, some cells exhibited discrete hyperplastic modifications of ER cisternae or altered mitochondria of abnormal, irregular shape which were sometimes vacuolized or with thinned and fragmented cristae similar to those observed by others (not shown) (2). Particular morphological alterations of nuclei potentially caused by apoptosis as described by others for HepG2 cell transfected with full-length HCV RNA (37) could not be detected.
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To clarify whether core and/or E2 localize to the Golgi complex, cells were treated for 3 h with brefeldin A, which is known to cause the disintegration of the Golgi complex. Subsequently, cells were fixed and analyzed by indirect immunofluorescence. As can be seen in Fig. 4H, brefeldin A treatment was effective and abrogated the detection of Golgin 97. In contrast, the subcellular distributions of E2 and core were not significantly altered under these conditions (Fig. 4F and G, respectively), suggesting that E2 and core do not localize mainly to the Golgi apparatus. However, it should be mentioned that the overall intensity of the E2 staining was considerably reduced after brefeldin A treatment (data not shown), which may be caused by the toxicity of the drug or the loss of the Golgi compartment. As regards the granular distribution of core, a very similar staining pattern has been observed for core-expressing cell lines, in which an association of core with cytoplasmic lipid storage vesicles has been described (1, 34, 53). In order to determine whether the same is true in cell lines with an sfl genome and to analyze the subcellular distribution of core and E2 more precisely, we conducted immuno-EM using a rabbit antiserum directed against E2 and a mouse monoclonal antibody specific for core. Bound antibodies were detected by matched secondary antibodies complexed to colloidal gold particles with diameters of 10 nm for E2 and 6 nm for core. Since we omitted osmium tetroxide to improve the preservation of antigenicity, under these conditions improperly fixed lipids were extracted. Interestingly, by using this method, i.e., EM contrast, the vesicular structures in cells embedded for immunostaining appeared devoid of any content (Fig. 5A and C), suggesting that they did indeed contain lipids. The 6-nm-diameter gold particles specific for core localized primarily to the border of these empty vesicles (Fig. 5A), consistent with the studies performed by Barba and coworkers showing the association of core with lipid droplets in transiently transfected cells (1). In contrast, E2 was not found to associate with these vesicles but rather at stacked cisternae, suggesting that E2 localizes primarily to the ER and early Golgi compartments (Fig. 5B). It was shown previously, that HCV replication and/or translation are tightly coupled to the cell cycle (56). Therefore, in an asynchronously growing replicon cell population, HCV antigen levels can differ dramatically from cell to cell. Consequently, in our immunogold labeling experiments, not all labeled sfl genome-containing cells carried detectable levels of gold particles, which indicates the specificity of our staining (data not shown). This specificity was confirmed by the result shown in Fig. 5C in which cells of the sfl genome-bearing cell line 21-5 were incubated with isotype-matched primary antibodies directed against HIV and measles virus proteins, followed by the same gold-labeled secondary antibodies. In conclusion, these data show that cells with an sfl genome exhibit several morphological peculiarities, most notably the presence of large amounts of lipid vesicles that are delimited by an electron-dense rim. The majority of core did not colocalize with E2, since only minor staining was detected at the ER, while most of the antigen was tightly associated with cytoplasmic lipid vesicles.
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5 x 104 molecules per ml of medium). Interestingly, in the absence of detergent, this RNA was resistant to RNase A up to a concentration of 100 µg/ml but was completely degraded upon the addition of 0.5% NP-40 (compare Fig. 6C and A). While the detergent-labile nature of the otherwise protected HCV RNA was principally consistent with the notion that it is encased in a virus particle, we were surprised to detect HCV RNA with similar characteristics in the supernatant of cells carrying subgenomic replicons that lack the HCV structural proteins (105 molecules per ml of medium [Fig. 6B]). This result indicated that these cell lines were capable of releasing large amounts of nuclease-resistant HCV RNAs by as yet unknown nonspecific mechanisms.
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109 RNA molecules per ml of gradient fraction) were found in the range between 1.04 to 1.11 g/ml with a peak at a density corresponding to 1.06 g/ml. Interestingly, HCV RNA structures derived from sfl genome-bearing cell lines 20-1 and 21-5 had similar low densities with peaks of 1.04 and 1.05 g/ml, respectively. This shows that HCV RNA released into the supernatants of sfl genome-bearing cell lines is most likely incorporated into lipid-containing structures that have biophysical properties comparable to those of infectious virus particles of BVDV. However, the amount of HCV RNA was about 3 orders of magnitude lower than that observed 24 h after a productive infection with cytopathic BVDV (106 copies/ml for HCV versus 109 copies/ml for BVDV [compare Fig. 7B and C]). Furthermore, the HCV RNAs present in supernatants of cell lines with subgenomic replicons had similar low densities (1.06 and 1.07 g/ml for cell lines 5-15 and 9-13, respectively), and the RNA amounts were at least fourfold higher, reflecting their higher replication efficiency. These results suggest the nonspecific release of lipid- and HCV RNA-containing structures (vesicles) into the medium of cell lines harboring subgenomic replicons and sfl genomes.
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DISCUSSION |
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Recently, Chung and coworkers described the replication of HCV in CV-1 and HepG2 cells after transfection of a plasmid that carries the full-length H77 genome flanked at the 3' end by the hepatitis delta virus ribozyme (14). This system is based on transcription of the HCV genomic RNA by T7 RNA polymerase that is provided by infection with a recombinant vaccinia virus (vTF7-3). Although this is an interesting approach, it has some limitations. For instance, infections with vTF7-3 are cytopathic and may have pleiotropic effects. Furthermore, the specific detection of the minus-strand RNA is complicated due to possible copy-back RNA synthesis by T7 RNA polymerase and the presence of large amounts of double-stranded plasmid DNA used for the transfection. In addition, since T7 RNA polymerase-based transcription is error prone, it is difficult to differentiate between genetic drift caused by error-prone RNA synthesis of NS5B RdRp and errors introduced by the low fidelity of T7 polymerase. Finally, vTF7-3 itself can be blocked by alpha interferon, making the distinction between inhibition of HCV RNA replication and the recombinant vaccinia virus problematic (14). Therefore, we think that the self-replication of full-length HCV genomes after transfection of RNA as described in the present study is superior. The availability of the cell lines described in this study for the first time permitted the analysis of HCV structural proteins expressed from autonomously replicating full-length HCV RNAs which closely mimics the in vivo situation within an infected hepatocyte. We found that the dominant core protein within cells bearing an sfl genome is p21, which presumably results from a cleavage event around amino acid residue 174. This observation is in direct accordance with previous data which clearly established that the major core species both in tissue culture cells expressing HCV core and in virus particles derived from human sera is p21 (71). Immunolocalization experiments revealed an exclusively cytoplasmic distribution of the polypeptide, with a strikingly granular pattern that was independent of brefeldin A treatment. This is in contrast to a report by Yasui et al. (71) that reports some core staining also within the nuclei of cell lines stably or transiently expressing HCV core. However, since we used the anti-core monoclonal antibody C7-50, which according to Yasui and coworkers recognizes only core protein expressed in the cytoplasm (71), we cannot rule out the possibility that a certain proportion of core also localizes to the nucleus in cells with the sfl genome.
The increased resolution provided by immuno-EM established that the majority of core protein localized to the boundary of cytoplasmic lipid storage vesicles, whereas only minor staining was found at the ER. This finding directly correlates with previous studies which show a tight association of the core protein with lipid droplets. Since this observation was made using core protein from different HCV isotypes expressed by alternative approaches in various cell lines (1, 53, 71) and as the association of HCV core with lipid droplets was also observed in specimens obtained from chronically infected chimpanzees, this appears to be a general phenomenon that also takes place in infected hepatocytes in vivo.
It was shown that folding of HCV E1 and E2 glycoproteins can proceed by two pathways: (i) a slow and inefficient productive pathway that leads to the assembly of noncovalently linked heterodimers which are believed to be the prebudding forms of E1 and E2 (18-21) and (ii) a nonproductive pathway that leads to the formation of disulfide-linked heterogeneous E1-E2 aggregates (18, 20). In either case, the complexes are retained in the ER, the presumed site of virus budding, due to retention signals present in the transmembrane domains of both E1 and E2 (15-17, 22). As shown in this report, the majority of E1-E2 complexes present in cell lines with an sfl genome folded by the productive pathway. The absence of glycoprotein aggregates in these cells might be due to a lowered propensity of the Con1-derived glycoproteins to fold nonproductively or to preferential binding of our antiserum to native E1-E2 complexes. However, it should be kept in mind that the expression level of the HCV glycoproteins from the sfl genome is significantly lower than in the vaccinia virus-based expression systems frequently employed to analyze HCV glycoproteins (data not shown). Since productive folding of E1-E2 heterodimers is a slow process that involves the ER chaperone calnexin (20, 21), it is conceivable that the high expression levels reached in these systems may overload cellular factors required for proper folding, while the lower expression in our system may not exceed the capacities of these factors in the folding pathway. Although the majority of the glycoproteins detected appeared to fold along the productive pathway, only a weak coprecipitation of E1 with E2 was found. Again, this could be an inherent feature of our Con1 HCV isolate, or it could be due to the antibodies used. However, this is unlikely, because we observed much more efficient coprecipitation of E1 with E2 after expression of the glycoproteins in Huh-7 cells with recombinant vaccinia virus, employing the same antiserum for immunoprecipitation (41). Therefore, the weak association between E1 and E2 may be ascribed to the presence of mutations that have accumulated during prolonged passage of these cell lines. This notion is further supported by the high genetic drift observed with the sfl genomes.
Encouraged by the high replication levels, efficient expression of the structural proteins, and formation of native glycoprotein complexes, we examined whether cells carrying the sfl genome produced and released HCV particles. To this end, we directly transferred cleared cell culture supernatant to parental Huh-7 cells but failed to detect specific infectivity. The same was found using cell lysates generated after repetitive freeze-thaw cycles. However, since it is not known whether Huh-7 cells are susceptible to HCV infection, we cannot rule out the possibility that we may have missed infectious virus due to a flawed assay system. On the other hand, cell lines that have been reported to be permissive for HCV infection do not support replication of sfl genomes and are thus not suitable as target cells for infectivity assays. Therefore, as an alternative, we examined whether cells with an sfl genome released nuclease-resistant HCV RNA. Interestingly, such RNAs were indeed found in the supernatants of these cells (>104 copies per ml). Moreover, these RNAs had a very low density similar to the density reported for infectious HCV (31) and, as shown here, also to that of the pestivirus BVDV. However, in supernatants of cell lines with subgenomic HCV replicons, HCV RNA-containing structures with equivalent characteristics were found, indicating that their release is a general feature of replicon-bearing Huh-7 cells that is not mediated by the viral structural proteins. The exact nature of these RNA-containing structures is currently unknown, but their biophysical properties suggest that they correspond to membranous vesicles. In addition, the overall yield of these structures is rather low. While intracellular HCV RNA levels range from 200 to 500 copies per cell, the number of extracellular HCV RNA-containing vesicles amounts to 0.1 to 1 vesicle per cell in supernatants harvested 5 to 7 days postseeding. Although we cannot strictly rule out the possibility that sfl genome cells also produce HCV particles, this is unlikely for two reasons. First, the overall low level of released HCV RNA suggests a nonspecific process, and second, the amount of HCV RNA in the supernatant did not increase due to the presence of the structural proteins, which would be expected for productive assembly and release. In fact, both in transient-replication assays and in cell lines carrying sfl genomes or subgenomic replicons, we consistently found slightly higher levels of HCV RNA in the supernatant of the latter. Thus, the amount of these RNA species correlated with the intracellular replication level, rather than the presence or absence of the structural proteins, suggesting that RNAs were released by nonspecific mechanisms.
There are several factors or reasons that may cause the inability of the cell lines with an sfl genome to produce HCV particles. First, one could imagine that the serine-to-glycine mutation that was introduced at the second position of the core protein due to the cloning procedure may inactivate the protein and thus abrogate virus assembly. However, this is unlikely, as second-generation cell lines with an sfl genome that harbor constructs with an authentic core protein displayed similar phenotypes in our assays and thus apparently also did not yield virus particles (unpublished results). Alternatively, the presence of the heterologous sequences (EMCV IRES and neo) may disrupt a packaging signal or increase the size of the RNA beyond the packaging limit. Finally, the high genetic variability observed with sfl genomes may quickly lead to the accumulation of mutations that inactivate the assembly competence of the structural proteins. However, our transient-replication assays with full-length unmodified Con1 RNA and a variant of this RNA bearing only three adaptive mutations in the nonstructural region also did not reveal evidence for virus assembly in Huh-7 cells. Thus, since these RNA molecules did not carry heterologous sequences and because genetic variability can be ruled out due to the transient nature of these assays, we conclude that the lack of virus assembly is likely attributable to other factors. Recently, it was shown that the Con1 isolate is infectious in vivo (Bukh, unpublished), indicating that the structural proteins are assembly competent. Therefore, we think that the lack of virus production might be due to the inability of Huh-7 cells to support virus assembly and/or release perhaps due to the lack of certain host cell factor(s). Such a dependence of particle formation on cellular factors has been reported recently for other viruses. For instance, in the morphogenesis of adenovirus, cellular proteins are thought to be part of the viral packaging machinery (62). Furthermore, it was recently reported that cellular ubiquitin plays a vital role in a late step of the retroviral budding process (55, 63, 65). It is to be hoped that future research will soon unravel such cellular factors needed to support HCV assembly and release in cell culture, as this would finally allow construction of a model of the entire HCV replication cycle in cell culture. Furthermore, these findings may point to new virus-host interactions which may prove to be attractive targets for future antiviral treatment.
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ACKNOWLEDGMENTS |
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This work was supported in part by a grant from the European Community (QLK2-1999-00356).
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FOOTNOTES |
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Present address: Visible Genetics Ltd., 184 Cambridge Science Park, Cambridge CB4 0GA, United Kingdom.
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