We have previously described that the DNA fragment of HCV ORF from genotype 1b included in the VT7-HCV_7.9 recombinant is efficiently transcribed and translated upon induction with IPTG into a viral polyprotein precursor that is correctly processed into mature structural and nonstructural (except the C-terminal part of NS5B) HCV proteins 12.

To identify the cytoplasmic compartment(s) in which viral HCV proteins accumulated during infection of HeLa cells with VT7-HCV_7.9, we performed immunofluorescence analysis using serum from an HCV-infected patient to recognize HCV proteins and antibodies specific for the Golgi apparatus (anti-gigantin), the endoplasmic reticulum (anti-calnexin) or the mitochondria (mitotracher) (Fig. 1). Inducible expression of HCV proteins caused severe disruption of the Golgi apparatus as revealed by labelling this organelle with a specific antibody (Fig. 1A). This effect was not observed in cells infected with VT7-HCV_7.9 in the absence of IPTG. There is no co-localization of HVC proteins with the disrupted Golgi, whereas in the labelling of the endoplasmic reticulum, a clear co-localization between HCV proteins expressed from VT7-HCV_7.9 and ER proteins was observed (Fig. 1B). With an in vivo mitochondrial marker (Fig. 1C), we detected partial co-localization between HCV proteins expressed from VT7-HCV_7.9 and mitochondria organelles. Moreover, the mitochondria appeared more rounded in cells infected with VT7-HCV_7.9 + IPTG, in comparison with uninfected cells or with cells infected in the absence of IPTG.

To gain more detail information on the subcellular changes induced by HCV polyprotein expression, we performed transmission electron microscopy (EM) analysis. HeLa cells were infected with VT7-HCV_7.9 in the presence or absence of IPTG, and at 16 h p.i, infected and uninfected cells were collected and ultrathin sections visualized by EM at low and high magnification. While in cells infected with VT7-HCV_7.9, in the absence of IPTG there are high number of immature (IVs) and intracellular mature (IMVs) forms of VACV virus, in cells infected with VT7-HCV_7.9 in the presence of IPTG fewer IVs and IMVs were observed, corroborating our previous finding that the expression of HCV proteins blocked VACV morphogenesis 12. In addition, several morphological alterations were detected in cells expressing the HCV polyprotein when compared with uninfected cells (Fig. 2A) or with cells only expressing VACV proteins (Fig. 2B). The first alteration seen was the loss of spatial organization of the ER, with vesicles embedded in a membrane matrix of circular or tightly undulating membranes, forming electron dense structures indicated as EDS (Fig. 2C and 2D). These cytoplasmic structures resemble those structures called "membraneous webs" that have been visualized in human hepatoma Huh7 cells expressing a subgenomic HCV replicon 51415. Other alterations observed were the presence of vacuoles (indicated as V) often surrounding the compact structures, as well as the presence of swollen mitochondria (indicated as m) (Fig. 2D and 2E). Higher magnification of the electron dense structures in cells expressing the HCV polyprotein revealed membranes and tubular structures (indicated as TS) as part of the EDS (Fig. 2E).

Since hepatocytes are the main targets of HCV virus, we next analyzed if expression of HCV polyprotein from the VT7-HCV_7.9 infected cells also affected the ultra-structure of hepatic cells. Thus, monolayers of a human hepatoblast cell line (HepG2) were infected with VT7-HCV_7.9 under the same conditions as for HeLa cells and processed at 16 h p.i for EM analysis. In this cell line, similarly as in HeLa cells, the inducible expression of HCV proteins blocks VACV morphogenesis and induces the same alterations described above. In contrast to uninfected (Fig 3A) and infected hepG2 cells in absence of IPTG (Fig. 3B), in cells expressing HCV proteins we distinguish EDS in membranous webs (Fig. 3C and 3E), formation of large vacuoles (Fig. 3C and 3D), and also identified "virus like-particles" structures surrounded by membranes and dispersed in several areas of the cell cytoplasm (Fig. 3D).

To assure that the electron dense structures appearing in the cytoplasm of infected cells are the result of HCV polyprotein expression, we performed immunogold electron microscopy analysis with antibodies against HCV structural and nonstructural proteins (Fig. 4). Thus, HeLa cells were infected with VT7-HCV_7.9 in the presence or absence of IPTG and at 16 h p.i, infected and uninfected cells were processed for immunogold labelling on ultrathin sections. Due to the fixation and embedding procedures used in immunostaining, the cell structures are less visible than by embedding in an epoxy resin. While in cells infected with VT7-HCV_7.9 in absence of IPTG there was no specific labelling detected with the serum from an HCV-infected patient (Fig. 4A), in contrast, in antibody-reacted cells expressing HCV proteins most gold particles were concentrated into electron dense and membranous structures (Fig. 4B). Discrete labelling was observed in other parts of the cell cytoplasm. The localization of some of the nonstructural HCV proteins was determined using rabbit polyclonal antibodies against NS4B or NS5A proteins. The membranous and electron dense structures were also specifically recognized by antibodies against NS4B (Fig. 4C) and NS5A (Fig. 4D), indicating that both proteins are part of electron dense membrane-associated cytoplasmic complexes.

Since by confocal microscopy we observed co-localization between ER and HCV proteins in cells infected with VT7-HCV_7.9 in the presence of IPTG (see Fig. 1B), we performed immunogold labelling using a specific ER marker (mouse anti-PDI). As seen in Fig. 4E, strong labelling of ER was found in the membranous webs. These results suggest that the membranous webs are ER-derived structures. As the staining pattern corresponds to that obtained with the NS4B or NS5A proteins, the immunogold electron microscopy indicates that the ER is a site where these proteins are localized.

The detection by confocal microscopy of the presence of HCV proteins in the mitochondria (see Fig. 1C) suggests that HCV may regulate the mitochondria homeostasis. To confirm that, we evaluated different parameters such as, release of proapototic proteins including cytochrome c, loss of mitochondrial membrane potential (ΔΨm) and production of reactive oxygen species (ROS), all hallmarks of mitochondrial dysfunction.

To determine whether HCV polyprotein expression from the VACV recombinant activates cytochrome c release, HeLa cells were infected with VT7-HCV_7.9 in the presence or absence of IPTG, or treated with staurosporine (as a positive control). The cytochrome c release was detected by confocal microscopy. As shown in Fig. 5A, the cytochrome c remained confined to the mitochondria in both uninfected cells and VT7-HCV_7.9infected cells in the absence of IPTG. However, in cells infected with VT7-HCV_7.9 in the presence of IPTG, there is a diffuse cytosolic pattern of cytochrome c staining, similarly as in cells treated with staurosporine, indicating that cytochrome c was released from the mitochondria.

Next we determine if HCV polyprotein expression affected the mitochondria membrane potential (ΔΨm). HeLa cells were infected either with VT7-HCV_7.9 in the presence or absence of IPTG, or treated with staurosporine. At 48 h p.i, cells were stained in vivo with a fluorescent mitochondrion-specific dye, TMRE 1617, and analysed by flow cytometry. The loss of the ΔΨm was assessed by the ability of the mitochondria to take up TMRE. As shown in Fig. 5B, FACS analysis demonstrated a higher proportion of cells with decreased ΔΨm (ΔΨm Low) in HCV polyprotein expressing cells and in staurosporine treated cells, in contrast with cells infected with the VT7-HCV_7.9 in the absence of IPTG or in uninfected cells. These results indicate the ability of the HCV proteins to disrupt the mitochondria membrane potential in HeLa cells.

As mitochondrial dysfunction is also characterized by the generation of reactive oxygen species (ROS) 18, we investigated whether HCV polyprotein expression triggered the generation of ROS. HeLa cells were infected with VT7-HCV_7.9 in the presence or absence of IPTG and at 48 h p.i, cells were stained with dihydroethidium (2-HE) and subjected to flow cytometry 19. As shown in Fig. 5C, there is clearly production of ROS, as revealed by an increase in ethidium staining of DNA in HeLa cells infected with VT7-HCV_7.9 in the presence of IPTG. In contrast, ROS production was significantly lower (about 10%) in both uninfected cells and VT7-HCV_7.9 infected cells in the absence of IPTG.

The above results demonstrate that HCV proteins induce mitochondrial dysfunction evidenced by the release of cytochrome c, mitochondrial membrane depolarization and generation of ROS.

It has been reported in hepatic cells that expression of structural and nonstructural proteins from HCV cDNA 20 or from full-length RNA 21, can lead to apoptotic cell death, which could be an important event in the pathogenesis of chronic HCV infection in humans. We have previously shown by an ELISA-based assay that the inducible expression of HCV proteins from VT7-HCV_7.9 triggers apoptosis 12. In view of the severe cellular damage caused by polyprotein expression in VT7-HCV_7.9 infected cells, we wished to extend our previous observation by characterizing the apoptotic pathways in this virus-cell system. We first performed a qualitative estimation of apoptosis in HeLa cells infected with VT7-HCV_7.9 in the presence or absence of IPTG. By phase contrast microscopy and DNA staining analysis we observed that cells expressing HCV polyprotein developed at 24 h p.i characteristic morphological changes of apoptosis, as defined by cell shrinkage, granulated appearance, membrane bledding and chromatin condensation (not shown). Such morphological changes were not observed in cells infected with VT7-HCV_7.9 in the absence of the inducer.

To quantify the extent of apoptosis, DNA content was analyzed by flow cytometry after permeabilization and labelling with the DNA-specific fluorochrome propidium iodide. As shown by flow cytometry, 78% of HeLa cells infected with VT7-HCV_7.9 in the presence of IPTG were apoptotic in contrast with the 26% determined when cells were infected with VT7-HCV_7.9 in the absence of the inducer (Fig. 6A).

Since DNA fragmentation represents a late apoptotic event, we analyzed the activation of effector caspases as a previous stage in the induction of apoptosis. Apoptotic caspases are activated by a proteolytic cascade resulting in the cleavage of critical cellular substrates, including lamins and poly (ADP-ribose) polymerase (PARP). By immunoblot analysis using anti-poly-(ADP-ribose) polymerase (PARP) antibody, which recognizes the full (116 kDa) and cleaved (89 kDa) form of PARP, we determined that the expression of HCV proteins induced the activation of effector caspases, as revealed by the presence of the 89 kDa cleaved form in cell extracts from VT7-HCV_7.9 infected cells in the presence of IPTG. This activation was similar to that obtained in cells expressing the apoptotic inducer protein kinase PKR used as positive control. In contrast, minimal PARP cleavage product was observed in cell extracts from both uninfected cells and VT7-HCV_7.9 infected cells in the absence of IPTG (Fig. 6B, left panel). The general caspase inhibitor zVAD-fmk blocked completely activation of caspases, as revealed by PARP cleavage and by ELISA (Fig. 6B).

Having established the activation of downstream effector caspases by HCV polyprotein expression, we set out to analyze the upstream or initiator caspases that exert regulatory roles, like caspase-8 and 9. Western blot analysis of lysates from VT7-HCV_7.9 infected cells in the presence of IPTG using an antibody that recognizes the active caspase-8, detected a cleaved product of 43 kDa, which corresponds to the active subunit of caspase-8, and a product of 57 kDa, which corresponds to pro-caspase-8 (Fig. 6C, left panel). The same size cleaved product was also observed in cell lysates from VV-PKR infected cells used as positive control. In contrast, in uninfected cells or in cells infected with VT7-HCV_7.9 in the absence of IPTG only the pro-caspase-8 product was observed. Caspase-8 activation and apoptosis induction in cells infected with VT7-HCV_7.9 in the presence of IPTG was strongly inhibited by pre-treating the cells with the specific caspase-8 inhibitor zIETD-fmk (Fig. 6C, right panel). These results showed that expression of HCV proteins induces caspase-8-mediated apoptosis.

To define if the mitochondrial route could also be involved in the apoptosis induced by HCV polyprotein expression, we analyzed by Western blot the activation of caspase-9. Lysates from uninfected and VT7-HCV_7.9infected cells in the presence or absence of IPTG were reacted with a specific antibody that detects only the cleaved product of 37 kDa corresponding to the active subunit of caspase-9 (Fig. 6D). The active caspase-9 was detected in lysates from cells expressing the HCV proteins in contrast to lysates from uninfected cells or from cells infected with VT7-HCV_7.9 in the absence of IPTG (Fig. 6D, left panel). The activation of caspase-9 was confirmed after quantification by ELISA of the extent of the apoptosis induced by HCV proteins in the presence or absence of the specific caspase-9 inhibitor zLEHD-fmk. Severe inhibition was obtained by pretreating cells infected with VT7-HCV_7.9 in the presence of IPTG with zLEHD-fmk (Fig. 6D, right panel). The above observations establish that HCV proteins activate initiator and effector caspase-dependent death processes, with involvement of the two caspase 8 and 9 pathways.

Since identification of host genes triggered in response to HCV proteins is important to understand the pathogenic effects of HCV virus, we performed a microarray analysis to profile transcripts differentially expressed in HeLa cells infected with VT7-HCV_7.9 that inducible express HCV proteins. A comparative analysis of genes regulated was done after subtracting the values obtained from cells infected in the absence of the inducer IPTG versus values obtained in the presence of IPTG. Hybridization analysis revealed that at 6 hours after induction of HCV polyprotein expression 231 genes were differentially regulated. About 71% of these genes appear upregulated whereas the remaining 29% were downregulated. At 16 h post-induction the number of transcripts that passed the filtering conditions is significantly reduced. 81 genes were differentially expressed at this time and only 43% of them appear upregulated (see Additional file 1). The reduction observed at 16 h p.i correlated with the cell damage induced by HCV proteins in the infected cells, and hence only the data from 6 h p.i will be considered. Real time RT-PCR was used to verify the transcriptional change in selected genes, as detected by microarray (Table 1) since we have previously verified that microarray data correlated well with RT-PCR quantification 2223.

Most of the biochemical and morphological changes induced by HCV proteins described in this study were reflected in the gene expression profiling. Genes involved in apoptosis, oxidative stress, mitochondrial functions or membrane transport were upregulated by HCV proteins (Table 2). Moreover, genes encoding proteins implicated in lipid metabolism, DNA binding, cell cycle, signalling and inflammatory response changed in expression throughout the infection.

Genes implicated in apoptosis such as RAD21, PPP2CA, HCA66 or PDIA3 were upregulated whereas antiapoptotic genes IGF1R or SPHK2 were downregulated by HCV proteins. Interestingly, it has been described that HCA66 is able to modulate selectively Apaf-1 dependent apoptosis increasing downstream caspase activity following cytochrome c release from the mitochondria 24, an event observed during the inducible expression of HCV proteins in our virus-cell system. Within the group of genes related with mitochondrial functions, the C1QBP and SLC25A10 transcripts have been correlated with HCV infection. C1QBP gene appears upregulated in liver biopsies from acutely HCV-infected chimpanzees whereas downregulation of SLC25A10 alters mitochondrial and cellular status resulting in altered susceptibility of hepatic cells to apoptosis 25.

HCV proteins also induced disturbance in the expression of lipid metabolism and oxidative stress. Upregulation of GPX4, PRDX1 and CYP19A1 genes have been previously detected in biopsies of HCV infected chimpanzees or in human hepatocellular carcinoma (HCC) samples 252627. In contrast, it was reported that GSTM1 null genotype may facilitate HCV infection becoming chronic 28, and also this gene was downregulated in liver cells expressing entire HCV ORF 29. Glucose-6-phosphate dehydrogenase (G6PD) activity was inhibited in hyperplastic liver as well as in HCC 30.

In agreement with the alterations and formation of electron dense structures observed in infected cells expressing the HCV polyprotein, genes such as CLTA, CENTG2 or AP3S1, which are closely related with the membrane dynamics, were upregulated. Moreover, ER-resident proteins like DNAJC10 and Reticulon 4 (RTN4), which modulate the ER morphology under stress conditions, also appear activated in HCC samples 3132. Gene encoding DNA binding proteins such as HIST1HA2M, HIST1H4H, HIST2H4A, H2AFZ and HIST1H4D, or cell cycle transcripts (CKS2 or KPNA2), were consistently upregulated. Specific increases in histones and cyclin genes were markers of proliferative changes detected in the liver of HCV infected chimpanzees 2533.

Other genes that have been associated with HCV infection and were differentially expressed in our system are CLIC1, NET1, IRAK1, DDX5, TPRKB, TCP1, OLFM1, LDOC1 and HTRA3. CLIC1 gene was upregulated in liver biopsies from infected chimpanzees 25, whereas DDX5 helicase has homology with DDX3, which plays an important role in HCV replication 34. Relative high levels of NET1 and IRAK1 were reported in HCC 3536. Genes encoding for proteins TPRKB, T-Complex 1 (TCP1), Olftactomedin 1 (OLFM1), LDOC1 and HTRA3 have been implicated positive or negatively in cancer progression. TPRKB protein acts as a potential inhibitor of the binding of p53-related protein kinase PRPK to p53 37, whereas T-Complex 1 and Olftactomedin 1 promote proliferation of cancer cells 3839. On the other hand, it has been suggested that the downregulation of LDOC1 and HTRA3 genes may play an important role in the development and/or progression of some cancers 4041.

Overall, the association of the gene expression profile obtained after induction of HCV proteins in VT7-HCV_7.9 infected HeLa cells with genomic changes in HCV pathogenesis highlights the biological significance of the morphological and biochemical events identified in this study.

Cells were maintained in a humidified air 5% CO₂ atmosphere at 37°C. Human HeLa and monkey BSC40 cells were grown in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% newborn calf serum (NCS). Human HepG2 hepatocellular carcinoma cells (ATCC HB-8065) were maintained in DMEM supplemented with 10% fetal calf serum (FCS).

The recombinant VT7-HCV_7.9, derived from the vaccinia Western Reserve strain (VACV-WR), has been previously described 12. It contains 7.9 Kb of the HCV ORF from genotype 1b inserted within the viral HA locus under the transcriptional control of the T7 promoter, and expresses the T7 RNA polymerase upon induction with IPTG. The recombinant VV-PKR expressing IPTG-inducible dsRNA-dependent protein kinase (PKR) was generated by homologous recombination of their respective pPR35-derived plasmid with the VACV-WR strain as previously described 83. Viruses were grown and titrated in BSC40 cells and purified by banding on sucrose gradients 84.

HeLa cells cultured on coverslips were infected at 5 PFU/cell with VT7-HCV_7.9 in the presence or absence of IPTG (1.5 mM final concentration). At 24 h p.i, cells were washed with PBS, fixed with 4% paraformaldehyde and permeabilized with 2% Triton X-100 in PBS (room temperature, 5 min). To detect the mitochondria, cells were stained in vivo with Mitotracker Deep Red 633 (Molecular Probes) at 500 nM in DMEM, before fixing the cells. After blockade, cells were incubated for 1 h at 37°C with the specific primary antibodies. The coverslips were then extensively washed with PBS, followed by incubation in the dark for 1 h at 37°C with specific secondary antibodies conjugated with Alexa 488 (green), Alexa 594 (red) or with the green fluorochrome Cy2 (purchased from Molecular Probes), and with the DNA staining reagent ToPro-3 (diluted 1:200). Images were obtained by the Bio-Rad Radiance 2100 confocal laser microscope at a resolution of 63X, collected by Lasersharp 2000 software and processed in LaserPix.

HeLa cells were infected as described above in the presence or absence of general and specific caspase inhibitors and harvested at 24 h p.i. The extent of apoptosis was determined using the cell death detection enzyme-linked immunosorbent assay (ELISA) kit (Roche) according to the manufacturer's instructions. Duplicate samples were measured in two independent experiments. Cells infected with VV-PKR in presence of IPTG were used as positive control. The specific inhibitors of caspase 8 (zIETD-fmk), caspase 9 (zLEHD-fmk) and the general caspases inhibitor (zVAD-fmk) were added to the cells after one hour of virus adsorption at a final concentration of 50 μM (Calbiochem).