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Recently, claudin-1 (CLDN1) was identified as a host protein essential for hepatitis C virus (HCV) infection. To evaluate CLDN1 function during virus entry, we searched for hepatocyte cell lines permissive for HCV RNA replication but with limiting endogenous CLDN1 expression, thus permitting receptor complementation assays. These criteria were met by the human hepatoblastoma cell line HuH6, which (i) displays low endogenous CLDN1 levels, (ii) efficiently replicates HCV RNA, and (iii) produces HCV particles with properties similar to those of particles generated in Huh-7.5 cells. Importantly, naïve cells are resistant to HCV genotype 2a infection unless CLDN1 is expressed. Interestingly, complementation of HCV entry by human, rat, or hamster CLDN1 was highly efficient, while mouse CLDN1 (mCLDN1) supported HCV genotype 2a infection with only moderate efficiency. These differences were observed irrespective of whether cells were infected with HCV pseudoparticles (HCVpp) or cell culture-derived HCV (HCVcc). Comparatively low entry function of mCLDN1 was observed in HuH6 but not 293T cells, suggesting that species-specific usage of CLDN1 is cell type dependent. Moreover, it was linked to three mouse-specific residues in the second extracellular loop (L152, I155) and the fourth transmembrane helix (V180) of the protein. These determinants could modulate the exposure or affinity of a putative viral binding site on CLDN1 or prevent optimal interaction of CLDN1 with other human cofactors, thus precluding highly efficient infection. HuH6 cells represent a valuable model for analysis of the complete HCV replication cycle in vitro and in particular for analysis of CLDN1 function in HCV cell entry.
Hepatitis C virus (HCV) is a liver-tropic plus-strand RNA virus of the family Flaviviridae that has chronically infected about 130 million individuals worldwide. During long-term persistent virus replication, many patients develop significant liver disease which can lead to cirrhosis and hepatocellular carcinoma (54). Current treatment of chronic HCV infection consists of a combination of pegylated alpha interferon and ribavirin. However, this regimen is not curative for all treated patients and is associated with severe side effects (37). Therefore, an improved therapy is needed and numerous HCV-specific drugs targeting viral enzymes are currently being developed (47). These efforts have been slowed down by a lack of small-animal models permissive for HCV replication since HCV infects only humans and chimpanzees. Among small animals, only immunodeficient mice suffering from a transgene-induced disease of endogenous liver cells and repopulated with human primary hepatocytes are susceptible to HCV infection (39).
The restricted tropism of HCV likely reflects very specific host factor requirements for entry, RNA replication, assembly, and release of virions. Although HCV RNA replication has been observed in nonhepatic human cells and even nonhuman cells, its efficiency is rather low (2, 11, 59, 67). In addition, so far, efficient production of infectious particles has only been reported with Huh-7 human hepatoma cells, Huh-7-derived cell clones, and LH86 cells (33, 61, 65, 66). Although murine cells sustain HCV RNA replication, they do not produce detectable infectious virions (59). Together, these results suggest that multiple steps of the HCV replication cycle may be blocked or impaired in nonhuman or nonhepatic cells.
HCV entry into host cells is complex and involves interactions between viral surface-resident glycoproteins E1 and E2 and multiple host factors. Initial adsorption to the cell surface is likely facilitated by interaction with attachment factors like glycosaminoglycans (4, 31) and lectins (13, 35, 36, 51). Beyond these, additional host proteins have been implicated in HCV entry. Since HCV circulates in the blood associated with lipoproteins (3, 43, 57), it has been postulated that HCV enters hepatocytes via the low-density lipoprotein receptor (LDL-R), and evidence in favor of an involvement of LDL-R has been provided (1, 40, 42, 44). Direct interactions between soluble E2 and scavenger receptor class B type I (SR-BI) (53) and CD81 (49) have been reported, and firm experimental proof has accumulated that these host proteins are essential for HCV infection (5, 6, 16, 26, 28, 33, 41, 61). Finally, more recently, claudin-1 (CLDN1) and occludin, two proteins associated with cellular tight junctions, have been identified as essential host factors for infection (20, 34, 50) and an interaction between E2 and these proteins, as revealed by coimmunoprecipitation assays, was reported (7, 34, 63). Although the precise functions of the individual cellular proteins during HCV infection remain poorly defined, based on kinetic studies with antibodies blocking interactions with SR-BI, CD81, or CLDN1, these factors are likely required subsequent to viral attachment (14, 20, 31, 64). Interestingly, viral resistance to antibodies directed against CLDN1 seems to be slightly delayed compared to resistance to antibodies directed against CD81 and SR-BI (20, 64), suggesting that there may be a sequence of events with the virus encountering first SR-BI and CD81 and subsequently CLDN1. Moreover, in Huh-7 cells, engagement of CD81 by soluble E1/E2 induces Rho GTPase-dependent relocalization of these complexes to areas of cell-to-cell contact, where these colocalized with CLDN1 and occludin (9). Together, these findings are consistent with a model where HCV reaches the basolateral, sinusoid-exposed surface of hepatocytes via the circulation. Upon binding to attachment factors SR-BI and CD81, which are highly expressed in this domain (52), the HCV-receptor complex may be ferried to tight-junction-resident CLDN1 and occludin and finally be endocytosed in a clathrin-dependent fashion (8, 38). Once internalized, the viral genome is ultimately delivered into the cytoplasm through a pH-dependent fusion event (24, 26, 31, 58). Recently, Ploss et al. reported that expression of human SR-BI, CD81, CLDN1, and occludin was sufficient to render human and nonhuman cells permissive for HCV infection (50). These results indicate that these four factors are the minimal cell type-specific set of host proteins essential for HCV entry. Interestingly, HCV seems to usurp at least CD81 and occludin in a very species-specific manner since their murine orthologs permit HCV infection with limited efficiency only (22, 50). Recently, it was shown that expression of mouse SR-BI did not fully restore entry function in Huh-7.5 cells with knockdown of endogenous human SR-BI, suggesting that also SR-BI function in HCV entry is, to some extent, species specific (10).
In this study, we have developed a receptor complementation system for CLDN1 that permits the assessment of functional properties of this crucial HCV host factor with cell culture-derived HCV (HCVcc) and a human hepatocyte cell line. This novel model is based on HuH6 cells, which were originally isolated from a male Japanese patient suffering from a hepatoblastoma (15). These cells express little endogenous CLDN1, readily replicate HCV RNA, and produce high numbers of infectious HCVcc particles with properties comparable to those of Huh-7 cell-derived HCV. In addition, we identified three mouse-typic residues of CLDN1 that limit receptor function in HuH6 cells. These results suggest that besides CD81 and occludin, and to a minor degree SR-BI, CLDN1 also contributes to the restricted species tropism of HCV.
Huh-7.5 and HuH6 cells were grown in Dulbecco's modified Eagle medium (DMEM; Invitrogen, Karlsruhe, Germany) supplemented with 2 mM l-glutamine, nonessential amino acids, 100 U of penicillin per ml, 100 μg of streptomycin per ml, and 10% fetal calf serum (DMEM complete). Culture medium of HuH6-derived cell pools transduced with lentiviral vectors was supplemented with 5 μg blasticidin (Invivo Gen, San Diego, CA) per ml.
The HuH6 cell lines were created by lentiviral gene transfer using the lentiviral self-inactivating vector pWPI-BSD (56) transducing individual CLDN1 genes and subsequent selection of transduced cells with DMEM complete supplemented with blasticidin (5 μg/ml). Briefly, lentiviral particles were generated by cotransfection of the human immunodeficiency virus Gag-Pol expression construct pCMV-ΔR8.74 (19), the envelope protein expression construct pcz-VSV-G (27), and the respective lentiviral vector at a ratio of 3:1:3 into 293T cells (18) seeded 24 h prior to transfection at a density of 1.4 × 106/6-cm-diameter culture dish. Forty-eight hours later, the cell-free culture fluids were harvested and directly used to inoculate HuH6 cells. Subsequently, transduced cells were passaged in the presence of 5 μg/ml blasticidin.
Plasmids encoding the genotype 2a/2a chimera Jc1 (pFK-Jc1) and the Luc-NS3-NS5B/JFH1 reporter replicon (pFKi389Luc-EI/NS3-3_JFH1_dg) have been described recently (48, 56). To generate lentiviral vectors encoding the CLDN1 orthologs, RNAs from Huh-7.5 (human origin), Hep-56.1D (a murine liver-derived cell line ), CHO (Chinese hamster ovary), and Sprague-Dawley rat liver cells were prepared by using the Nucleo Spin RNAII kit (Macherey-Nagel, Düren, Germany). The total RNA prepared was used for reverse transcription (RT)-PCR-based cloning of the CLDN1 cDNAs employing the Expand RT-PCR kit (Roche, Mannheim, Germany) as indicated by the manufacturer. The RT reaction was conducted using an oligo(dT) primer. Subsequently, 10% of the RT reaction mixture was used for a PCR using oligonucleotides S-BamH-NcoI-hCLDN1 (5′-TCT GGA TCC ATG GCC AAC GCG GGG CTG CAG-3′) and A-Spe-hCLDN1 (5′-AGA ACT AGT TCA CAC GTA GTC TTT CCC GCT G-3′). For cloning of mouse, hamster, and rat CLDN1, S-BamH-NcoI-hCLDN1 was used in conjunction with A-Spe-mCLDN1 (5′-AGA ACT AGT TCA CAC ATA CTC TTT CCC ACT AG-3′). The PCR fragments were restricted with BamHI and SpeI and inserted into the BamHI- and SpeI-restricted pWPI vector. All PCR-based inserts were verified by sequence analysis (MWG, Martinsried, Germany). Detailed sequences are available on request. Notably, two different CLDN1 variants from the rat are deposited in the NCBI protein database which deviate from each other at four amino acid positions and that encode either V or I, respectively, at residue 155 (AAF04850 and NP_113887, respectively). The CLDN1 protein obtained from the liver of a Sprague-Dawley rat (for the primary sequence, see the alignment in Fig. Fig.8)8) has a valine at position 155.
To generate in vitro transcripts of Jc1 and the Luc-NS3-5B/JFH1 replicon, 10 μg of pFK-Jc1 or pFKi389Luc-EI/NS3-3_JFH1_dg plasmid DNA was linearized using MluI. Subsequently, DNA was extracted with phenol and chloroform and, after precipitation with ethanol, dissolved in RNase-free water. In vitro transcription reaction mixtures contained 80 mM HEPES (pH 7.5), 12 mM MgCl2, 2 mM spermidine, 40 mM dithiothreitol (DTT), a 3.125 mM concentration of each ribonucleoside triphosphate, 1 U of RNasin (Promega, Mannheim, Germany) per μl, 0.1 μg of plasmid DNA/μl, and 0.6 U of T7 RNA polymerase (Promega, Mannheim, Germany) per μl. After incubation for 2 h at 37°C, an additional 0.3 U of T7 RNA polymerase/μl of reaction mixture was added, followed by incubation for another 2 h at 37°C. Transcription was terminated by the addition of 1.2 U of RNase-free DNase (Promega) per μg of plasmid DNA and a 30-min incubation at 37°C. The RNA was extracted with acidic phenol and chloroform, precipitated with isopropanol, and dissolved in RNase-free water. The concentration was determined by measurement of the optical density at 260 nm. For electroporation of HCV RNA into cells, single-cell suspensions were prepared by trypsinization of monolayers and subsequent resuspension with DMEM complete. Cells were washed with phosphate-buffered saline (PBS), counted, and resuspended at 1.5 × 107/ml in Cytomix (60) containing 2 mM ATP and 5 mM glutathione. Five micrograms of in vitro-transcribed RNA was mixed with 400 μl of cell suspension by pipetting and then electroporated with a Gene Pulser system (Bio-Rad, Munich, Germany) in a cuvette with a gap width of 0.4 cm (Bio-Rad) at 975 μF and 270 V. Cells were immediately transferred to 10 ml of DMEM complete, and 2 ml of the cell suspension was seeded per well of a six-well plate.
Cells were washed once with PBS, lysed in NPB buffer (50 mM Tris-HCl [pH 7.5], 150 mM NaCl, 1% Nonidet P-40, 1% sodium deoxycholate, 0.1% sodium dodecyl sulfate [SDS], 1 mM phenylmethylsulfonyl fluoride, 0.01 U/ml aprotinin, and 0.02 U/ml leupeptin). The total protein content of each sample was determined by Bradford assay to ensure equal sample loading. An equal volume of 2× sample buffer (400 mM Tris [pH 8.8], 10 mM EDTA, 0.2% bromophenol blue, 20% sucrose, 3% SDS, 2% mercaptoethanol) was added, and samples were heated for 5 min at 95°C. The samples were loaded onto an 11% polyacrylamide-SDS gel. After electrophoresis, proteins were transferred to a polyvinylidene difluoride membrane using a semidry blotter (Carl Roth, Karlsruhe, Germany) according to the manufacturer's instructions. The membrane was blocked in PBS supplemented with 0.5% Tween 20 (PBS-T) and 3% milk for at least 1 h. Upon blocking, the membrane was incubated for 1 h with primary antibodies diluted in PBS-T supplemented with 3% milk. CLDN-1-specific antibodies were diluted 1:500, occludin-specific antibodies were diluted 1:200 (both from Zymed, San Francisco, CA), and SR-BI-specific antibodies were diluted 1:1,000 (Novus Biologicals, Littleton, CO), while anti-β-actin antibodies were diluted 1:2,000 (Sigma-Aldrich, Steinheim, Germany). After extensive washing with PBS-T, the membrane was incubated with horseradish peroxidase-conjugated secondary antibodies (anti-mouse immunoglobulin G [IgG] and anti-rabbit IgG; Sigma Aldrich) diluted in PBS-T supplemented with 3% milk. Bound antibodies were detected after a washing step with the ECL Plus Western blotting detection system (GE Healthcare Europe, Freiburg, Germany).
Cells were seeded at a density of 2 × 105 per well of a six-well plate. Twenty-four hours after seeding, cells were washed with PBS and labeled with 100 μCi/ml [35S]methionine/cysteine-containing culture fluid for 16 h (Easytag Express Protein labeling mix; Perkin-Elmer, Mechelen, Belgium). After extensive washing with PBS, cells were incubated with freshly prepared PBS supplemented with a cell membrane-nonpermeating biotinylation probe (Biotin-NHS; Calbiochem, Darmstadt, Germany) at a concentration of 0.6 mg/ml for 30 min at 4°C. The biotinylation reaction was quenched by addition of 1/10 volume of PBS containing 1 M glycine and incubation for 10 min at 4°C. Subsequently, cells were lysed using NPB buffer supplemented with 0.1 M glycine. Lysates were cleared by centrifugation at 13,000 × g for 15 min at 4°C. The cleared lysates were used for immunoprecipitation using a 1:1 mixture of protein A-agarose (Bio-Rad, Hercules, CA) and protein G-agarose (Roche, Mannheim, Germany) coupled to CLDN1-specific polyclonal antibodies (Neomarker, Fremont, CA) or glyceraldehyde 3-phosphate dehydrogenase (GAPDH)-specific antibodies (Sigma-Aldrich, Steinheim, Germany). Beads were washed three times with NPB buffer, and bound proteins were eluted by boiling the samples in SDS-PAGE sample buffer. The proteins were resolved by 11% SDS-PAGE. After electrophoresis, proteins were transferred to a polyvinylidene difluoride membrane (GE Healthcare, Freiburg, Germany) using a semidry blotter (Carl Roth, Karlsruhe, Germany) according to the manufacturer's instructions. The membrane was blocked in 10% skim milk in PBS for 1 h at room temperature. After several washing steps, biotinylated proteins were detected by incubation of the membrane with streptavidin-coupled horseradish peroxidase (Pierce, Rockford, IL) for 45 min at 4°C and subsequent staining with the ECL Plus Western blotting detection system (GE Healthcare Europe, Freiburg, Germany). The chemiluminescent biotin signal was allowed to fade overnight. Thereafter, the blot was exposed to MS film (Kodak, Rochester, NY) for detection of total CLDN1 expression.
Briefly, cells were washed once with PBS, lysed directly on the plate with 350 μl of ice-cold lysis buffer (0.1% Triton X-100, 25 mM glycylglycine, 15 mM MgSO4, 4 mM EGTA, 1 mM DTT [pH 7.8]), and frozen. After being thawed, lysates were resuspended by being pipetted up and down. For each well, 100 μl of lysate was mixed with 360 μl of assay buffer (25 mM glycylglycine, 15 mM MgSO4, 4 mM EGTA, 1 mM DTT, 2 mM ATP, 15 mM K2PO4, [pH 7.8]) and, after addition of 200 μl of a luciferin solution (200 μM luciferin, 25 mM glycylglycine [pH 8.0]), measured for 20 s in a luminometer (Lumat LB9507; Berthold, Freiburg, Germany).
Virus titers were determined as previously described, with minor modifications (33). In brief, cells were seeded into 96-well plates at a density of 1 × 104 per well 24 h prior to inoculation with dilutions of filtered cell culture supernatant (six parallel wells were used per dilution). After 72 h, cells were fixed for 20 min with ice-cold methanol at −20°C, washed once with PBS, and then permeabilized for 5 min with 0.5% Triton X-100 in PBS. After three washes with PBS, NS5A was detected with a 1:2,000 dilution of hybridoma supernatant 9E10 (33) in PBS for 1 h at room temperature. Subsequently, cells were washed as described above and bound 9E10 was detected by incubation with peroxidase-conjugated antibodies specific to murine IgG (Sigma-Aldrich, Steinheim, Germany) diluted 1:200 in PBS. After a 1-h incubation at room temperature, cells were washed as specified above. Finally, peroxidase activity was detected by using carbazole substrate. To this end, 0.32% (wt/vol) 3-amino-9-ethylcarbazole (Sigma) in N,N-dimethylformamide was diluted at a ratio of 1:3.3 with 15 mM acetic acid-35 mM sodium acetate (pH 8.0)-0.4% H2O2 and incubated with the cells for 15 min at room temperature until brown staining of infected cells was clearly visible. Subsequently, the carbazole substrate was aspirated and cells were kept in distilled H2O. Virus titers (50% tissue culture infective doses per milliliter) were calculated based on the methods of Kärber and Spearman (29, 55).
Cell culture-derived HCV particles were harvested 24 h after electroporation of appropriate HCV RNA into cells and passed through 0.45-μm-pore-size filters. One milliliter of the preparation was layered under a 0 to 30% continuous iodixanol gradient (Optiprep; Axis-Shield, Oslo, Norway) prepared in a cell suspension medium containing 0.85% (wt/vol) NaCl and 10 mM Tricine-NaOH (pH 7.4). Gradients were centrifuged for 15 to 18 h at 154,000 × g in a TH-641 swing-out rotor at 4°C using a Sorvall Ultra WX80 centrifuge. Twenty fractions of 0.5 ml each were collected from the bottom, and virus infectivity and the quantity of core protein were determined using a limiting-dilution assay and a core-specific enzyme-linked immunosorbent assay, respectively. The density of each fraction was quantified by refractometry.
HCV core protein was measured using an HCV core antigen kit (Wako Chemicals, Neuss, Germany) according to the instructions of the manufacturer. Cell culture medium was filtered through 0.45-μm-pore-size filters and either directly used for enzyme-linked immunosorbent assay or diluted with PBS prior to measurement.
Viral RNA was isolated from infected cells or from cell culture supernatant, respectively, using a Nucleo Spin RNAII kit (Macherey-Nagel, Düren, Germany) as recommended by the manufacturer. Two microliters of the RNA sample was used for quantitative RT-PCR analysis using a LightCycler 480 (Roche, Mannheim, Germany). HCV-specific RT-PCRs were conducted in duplicate utilizing a one-step RT-PCR LightCycler 480 RNA Master Hydrolysis Probes kit (Roche, Mannheim, Germany) and the following JFH1-specific probe (TIB Molbiol, Berlin, Germany) and primers (MWG-Biotech, Martinsried, Germany): A-195 [5′-6FAM (6-carboxyfluorescein)-AAA GGA CCC AGT CTT CCC GGC AAT T-TAMRA (tetrachloro-6-carboxyfluorescein)-3′], S-146 (5′-TCT GCG GAA CCG GTG AGT A-3′), and A-219 (5′-GGG CAT AGA GTG GGT TTA TCC A-3′). To normalize for equal quantities of total RNA in the samples, the GAPDH-specific mRNA was detected in parallel employing GAPDH-specific oligonucleotides (S-GAPDH, 5′-GAA GGT GAA GGT CGG AGT C-3′; A-GAPDH, 5′-GAA GAT GGT GAT GGG ATT TC-3′) and a GAPDH-specific probe, 5′-TET (6-carboxy-4,7,2′,7′-tetrachlorofluorescein)-CAA GCT TCC CGT TCT CAG CCT-TAMRA-3′. Reactions were performed in three stages by using the following conditions: stage 1 (RT), 3 min at 63°C; stage 2 (initial denaturation), 30 s at 95°C; stage 3 (amplification), 35 cycles of 15 s at 95°C and 30 s at 60°C. The amount of HCV RNA was calculated by comparison to serially diluted in vitro transcripts and normalized to the amount of GAPDH, which served as a housekeeping gene.
Cells were detached by using PBS supplemented with 0.2% (wt/vol) EDTA and washed twice with PBS. Approximately 1 × 106 cells per ml were stained for 1 h at 4°C with CD81-specific antibody (188.8.131.52; Santa Cruz, Santa Cruz, CA) diluted 1:200 in PBS containing 0.2% bovine serum albumin and 0.02% sodium azide (fluorescence-activated cell sorter [FACS] sample buffer). Subsequently, cells were washed with PBS and bound antibodies were detected by incubation for 1 h at 4°C with mouse-specific secondary antibodies conjugated with phycoerythrin (eBioscience, San Diego, CA) at a dilution of 1:100 in FACS sample buffer. Stained cells were washed with PBS, resuspended in 500 μl FACS sample buffer, and analyzed immediately using a FACScalibur apparatus (Becton Dickinson Biosciences) and the FlowJo software (Tree Star, Ashland, OR).
Murine leukemia virus (MLV)-based pseudotypes bearing vesicular stomatitis virus glycoproteins (VSV-G) or the HCV J6-derived E1 and E2 proteins were generated by cotransfection of 293T cells. Briefly, 1.5 ×106 293T cells were seeded into 6-cm-diameter plates 1 day before transfection with 2.6 μg envelope protein expression construct pczVSV-G, pcDNA3ΔcE1E2-J6, or an empty-vector control, 2.6 μg MLV Gag-Pol expression construct pHIT60, and 2.6 μg firefly luciferase transducing vector by using Lipofectamine 2000 (Invitrogen). The medium was replaced 6 h after transfection, and the supernatants containing the pseudoparticles were harvested 48 h later. The supernatants were cleared of cells by passage through a 0.45-μm-pore-size filter and used to infect Huh-7.5, HuH6, or 293T cells. The efficiency of pseudotype virus infection was evaluated by luciferase assays at 72 h postinfection.
Cells were seeded onto glass coverslips in 24-well plates at a density of 2 × 104 per well. Infection was performed 24 h after seeding by inoculation with HCV Jc1 particles for 4 h. At 72 h later, cells were fixed in PBS supplemented with 3% paraformaldehyde for 20 min at room temperature. Subsequently, cells were washed with PBS, permeabilized with PBS supplemented with 0.5% Triton X-100, and incubated with NS5A-specific primary antibody 9E10 diluted 1:2,000 in PBS supplemented with 5% normal goat serum. After 1 h, cells were washed extensively with PBS and bound primary antibodies were detected using a goat antibody specific for murine IgG conjugated with Alexa-Fluor 488 (Invitrogen) at a dilution of 1:1,000 for 1 h in the dark. DNA was stained with DAPI (4′,6-diamidino-2-phenylindole; Invitrogen) for 1 min at room temperature. After washing with PBS and once in water, cells were mounted on slides using Fluoromount G (Southern Biotechnology Associates, Birmingham, AL).
To establish a model for assessment of HCV receptor function in an authentic cellular environment, we searched for host cells (i) of human hepatic origin (ii) that support HCV RNA replication and (iii) that have limiting amounts of a crucial HCV entry factor in order to permit systematic receptor complementation studies. Since HuH6 cells, a human hepatoblastoma cell line, support replication of subgenomic Con1 (genotype 1b) replicons (62), we first assessed if these cells sustain replication and virus production of the highly efficient infectious genotype 2a chimera Jc1 (48). To this end, we transfected HuH6 cells either with a subgenomic genotype 2a JFH1 luciferase replicon (Luc-NS3-5B/JFH1) for evaluation of HCV RNA replication (Fig. (Fig.1A)1A) or with Jc1 RNA for analysis of production of virus particles and their properties (Fig. 1B to E). For comparison, we transfected Huh-7.5 cells, a subclone of the Huh-7 hepatocarcinoma cell line which is highly permissive for HCV and is widely used for HCV in vitro studies. These experiments revealed that Luc-NS3-NS5B/JFH1 replicates in transfected HuH6 cells with high efficiency, reaching reporter gene expression levels comparable to those in transfected Huh-7.5 cells 72 h posttransfection (Fig. (Fig.1A).1A). However, reporter gene activity 4 h posttransfection was slightly lower and the kinetics of luciferase accumulation was clearly delayed in HuH6 cells, indicating that transfection efficiency, translation efficiency, and/or RNA replication is somewhat lower in these cells. Likely as a consequence, yields of infectious HCV particles obtained after transfection of the Jc1 genome were approximately 10-fold lower than those obtained with Huh-7.5 cells (Fig. (Fig.1B).1B). The properties of Huh-7.5- and HuH6-derived HCV particles, however, were indistinguishable regarding distribution in density gradients and specific infectivity (Fig. 1C to E). Together, these results establish that HuH6 cells sustain efficient HCV RNA replication and virus production.
When we challenged HuH6 cells with a large dose of infectious Jc1 particles (multiplicity of infection [MOI], ca. 10) and monitored infection 72 h later using an NS5A-specific indirect immunofluorescence assay, we observed only a very low percentage of NS5A-expressing cells (less than 0.01%), while the same dose of Jc1 particles had almost completely infected the Huh-7.5 cell culture (Fig. (Fig.2A).2A). These data indicated that HuH6 cells may be resistant to infection by genotype 2a HCV. In line with this interpretation, high copy numbers of HCV RNA were associated with both HuH6 and Huh-7.5 cells directly subsequent to inoculation with Jc1 particles (Fig. (Fig.2B,2B, ,44 h), while HCV RNA copy numbers subsequently declined in HuH6 cells and rapidly rose in Huh-7.5 cells. This finding argues that HCV particles adsorbed to HuH6 cells but were unable to efficiently infect them. Finally, when we determined the titer of a Jc1 virus preparation by using a limiting-dilution assay with either Huh-7.5 or HuH6 host cells, no infectivity was detectable in the latter case (Fig. (Fig.2C).2C). Taken together, these data indicate that HuH6 cells are either resistant to HCV genotype 2a infection or at least very poorly infectible.
To find out if this defect is linked to low expression or absence of any of the four key host factors required for HCV infection, we compared SR-BI, CD81, CLDN1, and occludin expression between Huh-7.5 and HuH6 cells using cell surface FACS analysis where applicable or Western blotting. As depicted in Fig. Fig.3A,3A, Huh-7.5 and HuH6 cells display similar amounts of CD81 at the cell surface and express comparable quantities of SR-BI (Fig. (Fig.3B).3B). Mouse NIH 3T3 cells, which were used as a control for the expression analyses, display some endogenous SR-BI, which was detected due to our use of a polyclonal serum that, according to the manufacturer, reacts with mouse, rat, mink, hamster, and human SR-BI. In addition, HuH6 cells express high levels of occludin clearly exceeding those observed in Huh-7.5 cells (Fig. (Fig.3C).3C). However, with the sensitivity of this assay, we were unable to detect CLDN1 expression in HuH6 cells (Fig. (Fig.3D),3D), suggesting that the poor permissiveness for HCV infection may be due to limiting amounts of this HCV entry factor.
Next we assessed (i) if naïve HuH6 cells are indeed nonpermissive for HCV infection due to lack of endogenous CLDN1 and (ii) whether CLDN1 orthologs from different rodent species can substitute for human CLDN1 (hCLDN1) during HCV entry. To this end, we cloned human, mouse, hamster, and rat CLDN1 and transduced naïve HuH6 cells to ectopically express these proteins by lentiviral gene transfer. Subsequently, we determined the surface expression of CD81 by FACS (Fig. (Fig.4A)4A) and SR-BI and occludin levels by Western blotting (Fig. 4B and C). As expected, transduction of CLDN1 did not modulate the expression of these proteins (Fig. 4A to C). In parallel, we quantified the total CLDN1 expression and cell surface levels between the HuH6 cell lines by using a cell surface biotinylation and radioimmunoprecipitation assay (for details, see Materials and Methods). As expected, naïve HuH6 cells expressed very little endogenous CLDN1, with no protein detectable at the cell surface (Fig. (Fig.4D).4D). In contrast, all of the HuH6-derived cell lines transduced to express CLDN1 produced large amounts of the respective CLDN1 homologs and displayed a large quantity of these proteins at the cell surface. As a control for specific surface staining, we also performed the biotinylation and radioimmunoprecipitation assay using a GAPDH-specific antibody. As expected, we were unable to detect GAPDH at the surface of the cells, whereas a strong signal for the total expression level could be observed. We also determined HCV RNA replication after transfection of Luc-NS3-5B/JFH1 RNA into these cell lines to rule out the possibility that transduction with the individual CLDN1 orthologs had altered the permissiveness of these cells for HCV RNA replication (Fig. (Fig.5).5). Finally, we evaluated the susceptibility of these different cell lines to HCV infection by three complementary approaches. First, we challenged these cells with a large dose of infectious Jc1 particles and determined HCV RNA copy numbers associated with the cells 72 h postinoculation by quantitative RT-PCR (Fig. (Fig.6A).6A). Second, we determined the titer of a Jc1 preparation on each of these cell lines using the limiting-dilution assay (Fig. (Fig.6B).6B). Third, we used HCVpp carrying J6CF-derived glycoproteins and transducing a luciferase reporter gene and assessed infection efficiency using luciferase assays (Fig. (Fig.6C).6C). A combination of all three approaches was chosen to distinguish if possible differences in the susceptibilities of the cell lines to HCV are related to differences during virus entry only (determined in the HCVpp assay) and/or due to differences in other steps of the viral replication cycle (HCVcc infection).
Ectopic expression of hCLDN1 in HuH6 cells dramatically increased infection by genotype 2a HCVcc and HCVpp, indicating that the resistance of naïve HuH6 cells to HCV is primarily due to limiting CLDN1 expression. The infection efficiency of HCVpp in HuH6-hCLDN1 cells was ca. 2.5-fold lower than in Huh-7.5 cells (Fig. (Fig.6C),6C), whereas HCVcc infected HuH6-hCLDN1 cells ca. 100-fold less efficiently (Fig. 6A and B). Together, these data argue that HuH6-hCLDN1 cells sustain moderately lower virus entry than do Huh-7.5 cells and that additional life cycle steps such as, for instance, HCV RNA replication and/or translation (compare Fig. Fig.1A)1A) are likely also impaired in Huh6-hCLDN1 cells.
Beyond this, we observed a considerable difference in the ability of CLDN1 orthologs from rodent species to confer susceptibility to genotype 2a HCVcc and HCVpp infection on HuH6 cells. More specifically hamster- and rat-derived CLDN1 sustained HCVpp infection like the human homolog (Fig. (Fig.6C),6C), while mouse CLDN1 (mCLDN1) was about fivefold less efficient. Similarly HCVcc infection was impaired in cells expressing mCLDN1, as is evident from the accumulation of ca. 10-fold less HCV RNA than in cells expressing human, rat, or hamster CLDN1 (Fig. (Fig.6A)6A) and ca. 20-fold lower titers in the limiting-dilution assay (Fig. (Fig.6B).6B). Although the expression levels of these proteins was somewhat variable, these results nevertheless argue that CLDN1 properties specific to the mouse homolog may limit its utility as an HCV entry factor in human hepatocytes (see also below). Notably, Evans et al. and Ploss et al. did not observe less-efficient usage of mCLDN1 in human embryonic kidney (293T) cells or nonhuman cell lines, respectively (20, 50). Therefore, we introduced the CLDN1 orthologs described above into 293T cells, which lack endogenous CLDN1 (20), and reassessed the species-specific usage of CLDN1 in these cells. In line with the report by Evans et al., HCVpp-infected 293T cells expressing mCLDN1 or hCLDN1 with comparable efficiency (Fig. (Fig.7).7). Thus, we concluded that usage of CLDN1 in HCV entry is cell type dependent, revealing species-specific differences in HuH6, but not 293T, cells.
To identify such mouse-specific determinants of CLDN1 which may be responsible for the lower efficiency of HCV infection, we performed a sequence alignment of the human, rat, hamster, and mouse orthologs of CLDN1 (Fig. (Fig.8)8) and searched for polymorphic sites that are conserved among the former but differ in the mouse homolog. Using this criterion, we identified residues M152, V155, and A180 of CLDN1, which are shared among the human, rat, and hamster proteins but are different in the mouse protein (L152, I155, and V180). To analyze whether these amino acid residues, located in the second extracellular loop and the fourth transmembrane region of CLDN1, are responsible for the comparatively inefficient usage of the mouse protein, we introduced into HuH6 cells variants of mCLDN1 in which any of these three residues had been replaced with the one found in the human protein individually or all three together (mCLDN1-LIV/MVA). Next, we made sure that HCV RNA replication (Fig. (Fig.9A)9A) and receptor expression (Fig. (Fig.10)10) were comparable between these cells and then determined susceptibility to HCV infection using HCVcc and HCVpp. As shown in Fig. Fig.9D,9D, mCLDN1 mutations I155V and V180A, as well as the triple mutation (mCLDN1-LIV/MVA), conferred susceptibility to genotype 2a HCVpp infection comparable to that of hCLDN1. In contrast, the L152M mutation only slightly enhanced mCLDN1 function as an HCV entry factor (Fig. (Fig.9D).9D). Similarly also for Jc1 infection, the former proteins were used with high efficiency, while the L152M mutation only slightly increased entry function over mCLDN1. Although slight differences in HCV RNA replication efficiency in these cells and some variability in CLDN1 expression levels may also influence HCV entry, together these results indicate that three mouse-specific residues of CLDN1 residing in the second extracellular loop and the fourth transmembrane domain of the protein limit HCV genotype 2a entry.
HCV requires at least four proteins, SR-BI, CD81, CLDN1, and occludin, to infect human or nonhuman cells (50). Likely, a number of additional host factors contribute to initial virus binding, subsequent uptake into endosomes, and final delivery of the viral genome into host cells. However, these seem to be (i) fairly conserved among different species and (ii) expressed in many cell types, since transfer of the four entry factors alone into various human and nonhuman cells was sufficient to render them susceptible to HCV infection (50). Notably, receptor complementation assays indicate that at least CD81 and occludin usage by HCV is highly species specific, as the mouse versions of these proteins only inefficiently support HCV entry (22, 50). In addition, mouse SR-BI also seems to support HCV entry slightly less efficiently than the human ortholog does (10).
Here we took advantage of the human hepatoblastoma cell line HuH6, which expresses little endogenous hCLDN1, and found evidence that HCV genotype 2a infection via mCLDN1 is limited due to three mouse-typic residues within the second extracellular loop and the most C-terminal transmembrane domain of the protein. This moderate yet reproducible difference suggests that, besides the major contribution by CD81, and occludin, and a minor influence of SR-BI, mouse-specific properties of CLDN1 also limit HCV entry and may thus preclude efficient propagation of HCV in mice.
Notably, these findings are in disagreement with recent observations by Evans et al. and Ploss et al. (20, 50). These authors complemented SR-BI in CHO cells or CLDN1 in CHO and human embryonic kidney (293T) cells. By inoculating these cells with HCVpp, they noted comparable infections, irrespective of whether human or mouse SR-BI or CLDN1 was expressed, suggesting that the mouse versions of these proteins are at least as effective for HCV entry as the human orthologs (20, 50). Congruent with these studies, expression of CLDN1 orthologs in 293T cells did not reveal species-specific differences, whereas these were evident for the mouse version in the context of HuH6 cells. Although the reason for this discrepancy is not known, it is important to keep in mind that HCV infection efficiency in vitro is influenced by receptor abundance. More specifically, it has been shown that downregulation of entry factor surface levels via RNA interference limits HCV entry (20, 30, 50), while overexpression of, for instance, CD81 or SR-BI increases infection (23). However, a certain threshold concentration of CD81 at the Huh7-Lunet cell surface is sufficient to render these cells fully permissive and additional CD81 does not further increase susceptibility to infection, even with a low dose of HCV (30). Therefore, it is conceivable that high, possibly saturating, surface expression of CLDN1 in 293T or CHO cells may mask species-specific difference in CLDN1 usage. In addition, we cannot exclude the possibility that subtle cell type-specific differences in the composition of the HCV receptor complex or the localization of individual factors between HuH6 cells on the one hand and CHO and 293T cells on the other may influence CLDN1 usage. For instance, differential recruitment of CLDN1 into tight junctions between these cells may influence the availability of this host factor, revealing species-specific differences in CLDN1 usage in one cell type but not the other. Clearly, more work, ideally using highly differentiated cells, is needed to better understand the contributions of the individual HCV entry factors in the course of infection and to dissect how species-specific properties of individual entry factors may limit infection.
With regard of the latter, two principal mechanisms can be envisioned. First, variation of the entry factor sequence in the area of the viral binding site may decrease the affinity of the interaction at the expense of viral entry. Alternatively, species-specific determinants of the entry factor may limit its interplay with other host factors needed for infection, thus impairing virus infection. In the case of CD81, for instance, there is firm evidence that HCV in the course of infection directly binds to CD81 via an interaction between a complex conformational epitope within E2 and the large extracellular loop (LEL) of CD81 (12, 17, 21, 25, 45, 46, 49). Notably, the LEL of CD81 comprises a subloop that is stabilized by two absolutely conserved disulfide bridges, a structural signature of the “tetraspanin fold.” Interestingly, key residues important for efficient binding of soluble E2 localize to this subdomain, which is hypervariable between tetraspanin family members and between CD81 orthologs from different species. This pronounced variability at the viral binding site may explain the very inefficient usage of murine CD81 by HCV.
Recently, coimmunoprecipitation of HCV E1/E2 and CLDN1 coexpressed in 293T cells provided evidence that HCV glycoprotein complexes may also directly interact with CLDN1 (63). However, a putative direct CLDN1 interaction domain has not been mapped. Using chimeras of CLDN1 and CLDN7, discrete residues (I32, E48) within the first extracellular loop of CLDN1 were identified that are critical for infection (20), and it is conceivable that this could be due to an involvement in a direct interaction with the virus. In this study, we have identified three mouse-typic CLDN1 residues that limit the utility of the murine protein for infection by HCV genotype 2a in human HuH6 hepatoblastoma cells. While the L152M mutation only slightly increased the entry function of the protein, the I155V or the V180A substitution conferred full HCV entry function on the murine protein. These data suggest that the second extracellular domain of CLDN1 and the fourth transmembrane helix of the protein comprise species-specific determinants of HCV infection. In the absence of precise structural information on CLDN1 and without a clearly mapped binding site for HCV, it is impossible to predict if these mouse-specific residues may influence the affinity or exposure of a putative HCV binding site. Since at least one of the key residues that compensate for the poor entry factor function of mCLDN1 is resident within a transmembrane-spanning helix, at least in this case, an effect on virus binding is likely indirect. Alternatively, the residues identified by us may modulate interactions with additional host factors important for HCV infection. Since these are of human origin in HuH6 cells, such contacts may be inefficient for the murine protein and could be restored by the respective amino acid substitutions. Further work is needed to distinguish by which mechanism the mouse-typic CLDN1 residues identified in this work limit HCV receptor function.
In summary, we have characterized HCV replication and virus production in the human hepatoblastoma cell line HuH6 and noted that, due to limited endogenous CLDN1 expression, naïve HuH6 cells are refractory to HCV infection. Since HCV infection is restored by ectopic expression of hCLDN1 and as these cells replicate and produce large amounts of virus particles, HuH6-hCLDN1 cells, next to Huh-7 derivatives, are an attractive cellular model to dissect the complete HCV replication cycle. In addition, we have identified mouse-specific CLDN1 residues that limit its function during HCV infection. Therefore, moderately reduced entry through mCLDN1 may further limit HCV infection of mouse cells due to highly inefficient usage of mouse CD81 and occludin and somewhat lower SR-BI function. Receptor complementation studies with HuH6 cells may, in the future, reveal further details of CLDN1 function during HCV infection.
We are grateful to Takaji Wakita for the gift of the JFH1 isolate and to Jens Bukh for the J6CF strain, to Dirk Lindemann for provision of pcz-VSV-G, to Didier Trono for pWPI and pCMVΔR8.74, to Charles Rice for Huh-7.5 cells and the 9E10 monoclonal antibody, and to Brent E. Korba for providing HuH6 cells. We also thank Sandra Ciesek, Thomas von Hahn, and Eike Steinmann for critical readings of the manuscript and all of the members of our laboratories for comments on and discussions of this work.
This work was supported by an Emmy Noether fellowship from the Deutsche Forschungsgemeinschaft to T.P. (PI 734/1-1), by grants from the Helmholtz Association SO-024, and by grants from the EU to R.B. (Marie Curie: EI-HCV, contract 035599, and ERC-2008-AdG 233130 HEPCENT). TWINCORE is a joint venture between the Medical School Hannover (MHH) and the Helmholtz Center for Infection Research (HZI).
Published ahead of print on 4 November 2009.