Requirement of cellular DDX3 for hepatitis C virus replication is unrelated to its interaction with the viral core protein

HCV core interacts with cellular DDX3 in cells replicating WT JFH1

The core–DDX3 interaction was previously demonstrated in cells ectopically expressing HCV genotype 1a core, and in the absence of virus RNA replication and productive infection (Owsianka & Patel, 1999). To test whether DDX3 interacts with JFH1 core, HuH-7 cells transfected with virus RNA were first analysed by indirect immunofluorescence. DDX3 was found to be sequestered by core in JFH1-replicating cells as detected using mAbs AO196 and AO322 [Fig. 2a and Supplementary Fig. S1a (available in JGV Online), respectively]. It should be noted that the core-bound DDX3 emitted a stronger fluorescent signal compared with free DDX3 occupying the cell cytoplasm. Thus, adjustment of the core-bound DDX3 signal to prevent overexposure resulted in the apparent loss of free DDX3 from the image. A representative image of core and DDX3 in virus-infected and surrounding non-infected cells taken at two different intensities is shown in Supplementary Fig. S1(b) (available in JGV Online). The anti-DDX3 antiserum R648 specifically co-immunoprecipitated core from JFH1-replicating cells, further confirming a direct interaction between these two proteins (Fig. 2b).

We also found that DDX3 co-localized with core protein in the NNeo/C-5B(+) cell line harbouring an autonomously replicating genome-length genotype 1b RNA (Ikeda et al., 2002), and HuH-7 cells infected with the intergenotypic 1a/2a chimeric virus [H-NS2/NS3 J(YH/QL)] (Yi et al., 2007) (Supplementary Fig. S1c, d, available in JGV Online). Thus, the recruitment of DDX3 by core is in good agreement with our previous results (Owsianka & Patel, 1999), and is not limited to one virus genotype.

DDX3 interacts with HCV core on LDs

The association of HCV core with LDs is essential for infectious virus production (Boulant et al., 2007; Miyanari et al., 2007; Shavinskaya et al., 2007). We sought to gain further insight into the core–DDX3 complex in the context of the JFH1 system by analysing the intracellular distribution of core and DDX3. HuH-7 cells electroporated with JFH1 RNA were analysed by indirect immunofluorescence for HCV core, DDX3 and adipocyte differentiation-related protein (ADRP), a protein abundant on the surface of LDs (Fujimoto et al., 2004). As shown in Fig. 2(c) left, core associated with ADRP and the localization of DDX3 was precisely coincident with both proteins, indicating its redistribution to LDs. To examine the subcellular position of DDX3 and core in relation to ADRP in greater detail, a series of Z-stacks was obtained and used after blind deconvolution to create a three-dimensional model of LDs coated by core, ADRP and DDX3. As reported before (Boulant et al., 2007), core fully coated LDs. We found that DDX3 by virtue of its interaction with core was also fully associated with LDs [Fig. 2c (i) and (ii)].

Identification of core residues required for DDX3 interaction

We previously reported that aa 1–59 of core protein are involved in its interaction with DDX3 (Owsianka & Patel, 1999). To identify critical residues within this region, a library of core 1–59 mutants containing single or multiple amino acid substitutions was generated by error-prone PCR (EP-PCR). The mutated sequences were fused in-frame with GFP and expressed in bacteria. The mutant fusion proteins were screened by incubation with GST–DDX3 fusion protein immobilized in ELISA wells, and the bound mutants detected using a rabbit polyclonal anti-GFP antiserum. Of 130 clones screened, only nine were found to be defective in binding to GST–DDX3 (data not shown). In order to confirm these results, all nine mutants were individually subcloned into a sequence encoding the HCV genotype 1a strain H77c core, E1 and E2 (pCE1E2) in a mammalian expression vector pcDNA3.1/Zeo+ (Invitrogen), and their expression was analysed by indirect immunofluorescence. The results were in accordance with the data from the initial ELISA screen in that all nine mutated core proteins failed to redistribute DDX3 (data not shown). Nucleotide sequence analysis showed that each of the mutants carried between one and four amino acid substitutions (Fig. 3). Of interest, mutant 90 had only one substitution (I30N), indicating that this residue must be required for the interaction of core with DDX3. All nine mutant proteins had at least one amino acid substitution in the region spanning residues 24–36, indicating that this region may harbour residues that are critical for the core–DDX3 interaction. To test this hypothesis, site-directed mutagenesis was carried out to revert any mutations outside of this 13 aa region back to the WT residue. These new mutant core 1–59 fragments were then subcloned into pCE1E2, and their expression analysed following transient transfection into HuH-7 cells as above. Again, none of these new core mutants redistributed DDX3 (data not shown), indicating that the 13 aa region between residues 24 and 36 of core is indeed involved in the interaction between core and DDX3. To determine which residues in this 13 aa region were essential, we carried out alanine-scanning mutagenesis across aa 24–36, individually substituting each residue in this region with alanine. As before, these alanine mutant sequences were subcloned into pCE1E2 and transiently transfected into HuH-7 cells. Immunofluorescence analysis revealed seven mutants (P25A, G26A, G28A, Q29A, V31A, G32A and L36A) that showed distinct co-localization between core and DDX3 (similar to that seen with WT HCV core), whilst the other mutants (F24A, G27A, I30A, G33A, V34A and Y35A) displayed no interaction with DDX3 at all (Supplementary Fig. S2, available in JGV Online). Thus, these results indicate that core residues F24, G27, I30, G33, V34 and Y35 are critical for its interaction with DDX3.

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Fig. 3.

Identification of core residues critical for its interaction with DDX3. Amino acid substitutions in residues 1–59 of core in nine mutants unable to interact with DDX3. Residues in the region 24–36 (shaded box) were targeted for alanine-scanning mutagenesis, which identified residues at positions 24, 27, 30, 33, 34 and 35 (underlined in the sequence at the top) that were critical for DDX3 interaction.

Disrupting the core–DDX3 interaction

Sequence comparison of core protein from strains H77c (genotype 1a) and JFH1 (genotype 2a) revealed 96.6 % identity within the first 59 residues of core, with the key residues (F24, G27, I30, G33, V34 and Y35) for interaction with DDX3 being fully conserved (data not shown). Thus, recombinant JFH1 genomes each containing one of the six mutations were constructed. All mutants were replication competent as seen by the expression of NS5A protein at 3 days post-transfection (Fig. 4). The viral core protein in these cells was detected using two different anti-core antibodies, which bound to the mutant proteins with varying affinity. mAb c7-50 bound minimally or not at all to I30A and G33A mutants, but had increased affinity for the V34A protein. The anti-core rabbit serum R308 on the other hand recognized all core forms equally except for G33A (Fig. 4). Notably, all six core mutations are located within the epitope (aa 21–40) recognized by c7-50 (Moradpour et al., 1996), which may account for the altered affinity of this mAb to some mutants. The antiserum R308 was raised against a peptide corresponding to core aa 5–25 (Hope & McLauchlan, 2000). Therefore, the smaller quantities of G33A core detected using R308 suggest either this mutation lowers the stability of the protein or simply reduces the binding efficiency of the antibody. Importantly, there was no detectable change in DDX3 protein levels in any of the transfected cell cultures tested (Fig. 4).

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Fig. 4.

Analysis of JFH1 core mutant viruses. HuH-7 cells electroporated with viral RNAs as shown were subjected to Western immunoblotting at 72 h post-incubation using anti-NS5a mAb 9E10, anti-core antibodies mAb c7-50 or R308, anti-DDX3 mAb AO196 and an anti-tubulin antibody.

We next tested by co-immunoprecipitation if these mutations had effectively disrupted the core–DDX3 interaction in virus-replicating cells. We excluded mutants I30A, G33A and V34A from this assay due to their altered affinity to mAb c7-50 (Fig. 4), which is the antibody we found to be most compatible in our co-immunoprecipitation assay. As shown in Fig. 5, the F24A and G27A core mutants were co-immunoprecipitated by the anti-DDX3 antiserum R648 in much lower amounts than the WT protein. However, the Y35A core was not co-immunoprecipated at all, indicating that its interaction with DDX3 was abrogated. Consistent with these findings, no co-localization of core and DDX3 was seen in cells replicating JFH1_Y35A by immunofluoresence analysis (Fig. 5b). Importantly, this mutant core remained associated with LDs, which is not surprising since motifs responsible for targeting core to LDs are located in the D2 domain (Hope & McLauchlan, 2000).

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Fig. 5.

Analysis of the interaction of DDX3 with core mutants. (a) HuH-7 cells were electroporated with different viral RNAs and HCV core co-immunoprecipitated at 72 h post-incubation using the anti-DDX3 serum R648 as described in the legend to Fig. 2b. (b) HuH-7 cells electroporated with JFH1_Y35A were fixed at 72 h post-incubation and analysed by confocal microscopy for the intracellular distribution of core, DDX3 and ADRP using appropriate antibodies. Bar, 10 μm.

HCV replication in the absence of the core–DDX3 interaction

To determine the importance of this interaction in the virus life cycle, we examined the phenotype of JFH1_Y35A. HuH-7 cells were electroporated with JFH1_WT, JFH1_Y35A or the replication-deficient JFH1_GND RNAs and incubated for various time points before determining the progeny virus yields released into the medium and intracellular viral RNA levels. As shown in Fig. 6(a), the number of infectious particles released by JFH1_Y35A-replicating cells was lower in comparison with JFH1_WT-transfected cells at 24 and 48 h post-transfection, although parity was achieved at 72 h. The intracellular RNA replication levels of JFH1_Y35A were similar to JFH1_WT throughout the time-course (Fig. 6b). As expected, no virus release or intracellular viral RNA replication was detected in cells harbouring JFH1_GND RNA (Fig. 6a, b​b).). We next examined the infectivity and replication of the core mutant virus released from electroporated cells. To do this, we infected naïve cells at an equal m.o.i. and quantified the released infectious virus progeny and the intracellular viral RNA at various times post-infection. As shown in Fig. 6(c, d), respectively, both the virus yields and RNA replication levels of JFH1_Y35A were very similar to JFH1_WT. We found no direct reversion or second-site mutation in JFH1_Y35A core sequence. Thus, our data indicate that the core–DDX3 interaction does not play a role in HCV morphogenesis.

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Fig. 6.

Analysis of the core mutant viruses. (a, b) Determination of virus yield and intracellular RNA levels following transfection. HuH-7 cells were electroporated with RNA derived from JFH1_WT, JFH1_Y35A or JFH1_GND cDNA. At the indicated time points (a) the released virus titres and (b) the intracellular HCV RNA levels were quantified by TCID₅₀ and RT-qPCR, respectively. (c and d) Determination of virus yield and intracellular RNA levels following infection. Naïve HuH-7 cells were infected with JFH1_WT or JFH1_Y35A (obtained from the electroporated cells above) at an m.o.i. of 0.02. The (c) titres of infectious virus released into the medium and (d) intracellular viral RNA levels were measured over 6 days as described above. Means and error ranges from duplicate electroporations and infections are shown.

Generation and characterization of anti-DDX3 antibodies.

BALB/c mice and rabbits were immunized with a bacterially expressed glutathione S-transferase-full-length DDX3 (GST–DDX3) fusion protein and antibodies generated essentially as previously described (Clayton et al., 2002). A total of two rabbit polyclonal antisera (R647 and R648) and 16 mouse hybridomas secreting mAbs (see Fig. 1) to DDX3 was generated. The mAbs were initially identified and selected on the basis of their ability to interact with GST–DDX3, but not an irrelevant GST fusion protein, in an ELISA assay (data not shown).

Plasmid constructs and mutagenesis.

The full-length DDX3 gene was cloned into the BamHI site, in-frame to sequences encoding the EGFP, in the vector pEGFP-C1 (Clontech) to generate EGFP–DDX3 fusion protein.

To identify residues within the N-terminal 59 aa of HCV core that are critical for interaction with DDX3, EP-PCR was performed to introduce random mutations into the HCV genotype 1a strain H77c (Yanagi et al., 1997) sequence encoding this region. The fidelity of Taq polymerase was decreased by altering the relative dNTP concentrations, using a high Mg²⁺ concentration and including Mn²⁺ in the reaction as described previously (Pritchard et al., 2005). The mutated sequences were cloned in-frame with (and downstream of) green fluorescence protein (GFP) in the bacterial expression vector pKK223-3 (Pharmacia) and the library of core 1–59 mutant–GFP fusion proteins generated was screened in an in vitro DDX3-binding assay as described in Results. Nucleotide sequencing subsequently identified the mutations introduced by EP-PCR in the clones of interest.

The plasmid pUC-JFH1 carries the full-length cDNA of the genotype 2a HCV strain JFH1 (Wakita et al., 2005). The plasmids pUC-GND JFH1 and pUC-JFH1 ΔE1E2 are identical except that they carry the GND mutation in the NS5B-encoding sequence, or an in-frame deletion in the E1 and E2 sequences (Wakita et al., 2005). Site-directed mutagenesis was carried out using the QuikChange-II kit (Stratagene) to introduce alanine substitutions at the target sites in core. Briefly, various alanine substitutions in the core-coding region were individually introduced into the plasmid pGEM-T (Promega) carrying nt 1–2614 (corresponding to the 5′ UTR and core to E2 coding sequence) of HCV strain JFH1 using appropriate primers (the sequences of which are available upon request). The presence of the desired mutation in the resulting clones was confirmed by nucleotide sequencing. Sequences carrying the appropriate mutation were subcloned back into pJFH1-pUC to generate mutant viruses (see below).

Generation of JFH1 virus and its mutated derivatives.

Ten micrograms of RNA synthesized in vitro using linearized plasmid templates carrying the wild-type (WT) or mutated JFH1 genomic cDNA was electroporated into HuH-7 cells as described previously (Wakita et al., 2005). The electroporated cells were seeded into appropriate tissue culture dishes and incubated at 37 °C. At the indicated time period, the medium containing the infectious virus progeny was filtered through a 0.45 μm pore-sized membrane and infectivity determined as described below.

Determination of virus infectivity and RNA replication.

Virus titres in the culture supernatants were determined as TCID₅₀ (Lindenbach et al., 2005) following immunostaining for NS5A. The intracellular and extracellular HCV RNA content was measured by reverse transcription-quantitative real-time-polymerase chain reaction (RT-qPCR) as described previously (Witteveldt et al., 2009). To determine virus replication after electroporation, cells transfected with the respective transcript were seeded into 10 cm culture dishes, incubated at 37 °C for 4 h and then trypsinized. Total RNA was prepared from 1/15 of the trypsinized cells using the RNeasy kit (Qiagen) to quantify viral RNA by qRT-PCR assay. The remaining cell suspension was split 1 : 3 into three T25 flasks. Following incubation at 37 °C for 24, 48 and 72 h, the infectious virus yields in cell culture supernatants were determined by TCID₅₀ assay and the intracellular viral RNA levels quantified by RT-qPCR. To measure virus replication after infection, 5.3×10³ naïve HuH-7 cells seeded into six-well culture dishes were infected with virus at the indicated m.o.i. Following incubation at 37 °C for 24, 48, 72, 96 and 144 h, infectious virus yields and intracellular viral RNA levels were determined as described above.

SDS-PAGE and Western blotting.

Cell lysates were subjected to SDS-PAGE followed by Western blotting using appropriate antibodies as described previously (Clayton et al., 2002). The bound antibodies were detected using enhanced chemiluminescence reagents (Amersham).

Immunoprecipitation of DDX3.

HuH-7 cells were washed with PBS, lysed in lysis buffer (20 mM Tris/HCl pH 7.4, 20 mM iodoacetamide, 150 mM NaCl, 1 mM EDTA and 0.5 % Triton X-100) and the lysate was spun briefly to remove nuclei. The clarified cell lysates were incubated with the anti-DDX3 antibodies as described in the text and the immune complexes precipitated using protein G agarose beads (Sigma). Following washes of the Sepharose beads, the immune complexes were analysed by SDS-PAGE followed by Western blotting using biotinylated R648 and anti-streptavidin-horseradish peroxidase (HRP) conjugate.

Co-immunoprecipitation of HCV core protein.

Approximately, 5×10⁶ HuH-7 cells electroporated with viral RNA were seeded into 100 mm tissue culture dishes. At 72 h post-transfection, the cells were washed and lysed in 0.5 ml lysis buffer (20 mM Tris/HCl, pH 7.4, 135 mM NaCl and 0.1 % Triton X-100) supplemented with 50 mM NAF, 5 mM Na₃VO₄ and 1 mM PMSF. The lysate was spun briefly to remove nuclei. After pre-clearing, the clarified lysate was immunoprecipitated overnight with protein G agarose beads that had been pre-incubated with the anti-DDX3 antiserum R648. The beads carrying the immune complex were spun at 2000 r.p.m. (microcentrifuge) for 2 min, washed three times with the lysis buffer and subjected to non-reducing SDS-PAGE followed by Western blotting using the anti-core mAb c7-50 and anti-mouse IgG-HRP conjugate.

Indirect immunofluorescence.

Cells on coverslips were fixed in methanol and probed with the indicated primary antibody for 1 h at room temperature. After washing with PBS, cells were incubated with anti-species antibodies conjugated with fluorescein isothiocyanate, tetramethyl rhodamine isothiocyanate or Cy5 (Invitrogen) for 1 h and then washed with PBS. The coverslips were examined with a Zeiss Laser Scanning LSM510 META inverted confocal microscope (Carl Zeiss) and the images analysed using LSM510 software. Three-dimensional (3D) reconstructions were performed from Z-stack images collected using optimum intervals. Image stacks were deconvolved by 3D-blind deconvolution using Autodeblur software (MediaCybernetics), and 3D reconstructions were generated as described previously (Boulant et al., 2007; Targett-Adams et al., 2008).

We thank Duncan McGeoch for helpful criticism of this manuscript and Charles M. Rice for the anti-NS5A mAb 9E10. This work was funded in part by grants from the Medical Research Council, UK (A. H. P.), Marie Curie Intra-European Fellowship contract 025198 (S. B.), the National Institute of Allergy and Infectious Diseases, U19-AI40035 (S. M. L.) and by the National Institute of Health, AI069285 (K. L.).