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Originally published online as doi:10.1189/jlb.0507335 on August 21, 2007

Published online before print August 21, 2007
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(Journal of Leukocyte Biology. 2007;82:1301-1310.)
© 2007 by Society for Leukocyte Biology

Hepatitis C virus core protein up-regulates anergy-related genes and a new set of genes, which affects T cell homeostasis

M. Domínguez-Villar, A. Muñoz-Suano, B. Anaya-Baz, S. Aguilar, J. P. Novalbos, J. A. Giron, M. Rodríguez-Iglesias and F. Garcia-Cozar1

Puerto Real University Hospital Research Unit, School of Medicine, Department of Biochemistry (Microbiology and Immunology), University of Cadiz, Cadiz, Spain

1 Correspondence: Hospital Universitario de Puerto Real, Unidad de Investigacion, Carretera NIV, Km665, 11510 Puerto Real, Cadiz, Spain. E-mail: curro.garcia{at}uca.es


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ABSTRACT
 
Hepatitis C virus (HCV) infection is the main cause for chronic hepatitis, leading to cirrhosis and hepatic carcinoma. Virally induced immune dysfunction has been called as the cause for viral persistence. Previous results demonstrate that CD4 Jurkat cells stably expressing the HCV core protein show an increased activation of NFAT transcription factor and an impaired IL-2 promoter activity, affecting intracellular signaling pathways in a manner that mimics clonal anergy. We had shown previously that NFAT activates a transcriptional program, ensuing in immunological tolerance. In the present work, we have engineered lentiviral vectors expressing the HCV core to analyze the events, which unfold in the initial phase of HCV core-induced anergy. We show that genes initially described to be up-regulated by ionomycin-induced anergy in mice are also up-regulated in humans, not only by ionomycin but also by HCV core expression. We also show that HCV core is sufficient to cause NFAT nuclear translocation and a slow-down in cell-cycle progression, and using whole genome microarrays, we identify novel genes up-regulated in Jurkat cells expressing HCV core. The relevance of our results is highlighted by the presence of HCV in CD4 T cells from HCV chronically infected patients.

Key Words: immune evasion • HCV • gene expression • CD4 T cells • anergy • tolerance


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INTRODUCTION
 
Hepatitis C virus (HCV) is the leading cause for transfusional hepatitis in the world. Eighty percent of infected patients are not able to eliminate the virus and will develop a chronic hepatitis, which will frequently ensue in cirrhosis and hepatic carcinoma, making HCV infection the main cause for liver transplantation in Western countries [1 ]. When the infection is resolved, it has been shown that cell-mediated immune responses are pivotal for viral clearance [2 ]. It has been demonstrated that CD8 lymphocytes are able to recognize and destroy HCV-infected cells, producing cytokines, which lead to a decreased viral replication, and mounting an initial, although nonlasting, response against the virus [3 ]. The limited CD8 response has been suggested to be a result of inadequate CD4 help, which prevents an effective, cytotoxic response against HCV [4 ]. The importance of CD4 T cell responses is supported by several studies showing that spontaneous viral clearance is associated with the presence of a specific CD4 response [5 6 7 ]. HCV persistence is also best predicted by a failure to generate or sustain CD4+ T cell responses [8 ]. HCV-specific CD4 T cells have been shown to be absent [6 ] or nonfunctional during HCV infection [9 ], showing an impaired secretion of the survival cytokine IL-2 [10 ] or an increased production of anti-inflammatory cytokines such as IL-10 [11 ].

An effective response to HCV vaccination also rests on T cell function [12 ], and restoration of an adequate CD4 response is also pivotal for the success of IFN-{alpha} + ribavirine (RBV), the only effective treatment against HCV infection [13 ]. Not only does IFN-{alpha} activity rely on an enhancement of a CD4 response [14 ], but also, RBV is acting by means of an increase in CD4 Th1 cell function, able to boost cellular responses against the virus [15 ].

Efficient ligation of TCR/CD3 by high-density antigen assembled on MHC on the surface of costimulation-proficient APC can generate a productive T cell response, which is characterized by the induction of a number of genes, including cytokine IL-2. Binding of IL-2 to its receptor is essential for TCR-stimulated cell-cycle progression from G1- to S-phase, resulting in clonal expansion and eventually, in the elimination of the antigen. Suboptimal cross-linking of the TCR by antigen, in the absence of costimulation, is not sufficient to induce a productive, immune response but instead, leads to anergic and regulatory T cells (Tregs), which block responses against the antigen, preventing its efficient clearance. Tregs as well as anergic CD4 T cells, which can be coinduced in several tolerizing strategies [16 17 18 ], are of paramount importance to prevent autoimmune diseases, maintaining peripheral immune tolerance. However, their presence in infectious diseases leads to pathogen persistence and chronic infection [19 ]. HCV-specific T cell responses have been shown to be suppressed by Tregs [20 ], and HCV-infected patients show a higher percentage of CD4 Tregs able to suppress CD8 anti-HCV responses [21 ].

Immunological tolerance can be induced by the virus by altering APC-mediated costimulation or by affecting T cell signal transduction directly. Although there is contradictory evidence for an effect of HCV on APC [22 23 24 ], mounting evidence suggests that HCV proteins are able to affect CD4 signaling pathways: HCV core protein, spanning amino acids 1–199, has been shown to down-regulate T cell responses by activating the NFAT transcription factor. NFAT activation is elicited by an enhancement in its transactivation activity [25 ] and by increasing intracellular Ca2+ concentration [26 , 27 ]. An increase in Ca2+ concentration has long been implicated in T cell anergy, which is a state of lymphocyte nonresponsiveness induced by suboptimal antigen stimulation, and it is thought to be important in preventing harmful responses to self-antigens. It has also been shown by us and others [28 29 30 31 ] that NFAT activation in the absence of concurrent activation of transcription factors, such as AP1 or NF-{kappa}B, leads to anergy induction by means of the up-regulation of anergy-associated genes (AR genes). CD4 Jurkat cells, stably transfected with a HCV core, have been shown recently to be unresponsive to stimulation in a manner that mimics clonal anergy [32 ]. It has also been shown that HCV core can block NF-{kappa}B [33 ] and AP1 [34 ] activity. The fact that HCV can infect primary CD4 T cells and infect and replicate in CD4 T cell lines such as Molt4 and Jurkat [35 ] is consistent with a direct effect of viral proteins on CD4 T cell homeostasis. In this paper, we show that HCV is detected in CD4 T cells from chronically infected HCV patients and analyze the molecular events, which unfold within hours of HCV core protein expression in CD4 T cells.


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MATERIALS AND METHODS
 
Cell cultures
The Jurkat (American Type Culture Collection, Manassas, VA, USA) cell line was maintained routinely in RPMI-1640 medium supplemented with 2 mM L-glutamine, 10 mM HEPES, 10% FBS, 1% nonessential amino acids (NEAA), 1% sodium pyruvate, and 1% penicillin/streptomycin at 37°C in a 5% CO2 incubator. Human embryonic kidney (HEK) fast growing and expressing the SV40 large T antigen (FT) cells (Invitrogen, Carlsbad, CA, USA) were maintained in DMEM supplemented with 10% FBS, 1% NEAA, 1% sodium pyruvate, 10 mM HEPES, 2 mM L-glutamine, and 1% penicillin/streptomycin at 37°C, 10% CO2.

Patients and controls
A total of five HCV, chronically infected patients, who were being assisted periodically at the HUPR dialysis unit, was studied. All patients were HCV singly infected individuals (Genotype 1b) without any HCV treatment during our study. Viral loads were 1.15, 4.01, 6.9, 1.21, and 3.54 x 106 UI/ml for Patients #1–#5. As controls, PBMC, obtained from five volunteer blood donors with negative serology for HCV, were used. Plasma and white blood cells for all of the experiments were collected during routine evaluation visits. Informed consent was obtained from the patients studied according to European Union regulations.

Plasma and CD4
Blood samples were collected in tubes containing EDTA. Within 1 h of phlebotomy, the specimen tubes were centrifuged at 1500 g for 15 min. Following repeated centrifugation of plasma at 1500 g for 10 min at room temperature, the supernatant was removed and used as the plasma fraction. PBMC were recovered from the cell layer by Ficoll-Paque (Amersham Pharmacia Biotech, Piscataway, NJ, USA) density gradient. Cells were washed and resuspended in DMEM containing 10% heat-inactivated FCS. Purified CD4 cell populations were isolated from PBMC using a positive magnetic sorting system (Miltenyi Biotec, Auburn, CA, USA) with magnetic beads conjugated to CD4, according to the manufacturer’s instructions. Cell viability assessment of freshly isolated, primary CD4 cells was examined by Trypan blue dye exclusion. Total cellular RNA was extracted using Trizol (Invitrogen), according to the manufacturer’s instructions. An aliquot of isolated cells was left in culture overnight to allow for bead detachment, and the purity of CD4 cells was detected by FACS analysis using FITC-conjugated, anti-CD4 antibody (Becton Dickinson, San Jose, CA, USA).

HCV RNA viral load and genotype
HCV RNA was quantified in 1 ml EDTA-anticoagulated plasma or RNA purified from 1 million CD4 cells by RT-mediated, real-time PCR using the COBAS AmpliPrep/COBAS TaqMan HCV test (Roche Diagnostics, Mannheim, Germany), according to the manufacturer’s instructions. The lower limit of detection is 15 IU/ml [36 ]. The HCV genotype was determined by a commercial method of line blot reverse hybridization, VersantTM HCV Genotype 2.0 assay (Bayer Diagnostics, Leverkusen, Germany).

Plasmid construction
DNA, encoding the first 191 amino acids of the HCV polyprotein, was amplified by PCR from a vector containing the H77 strain (Serotype 1a) HCV genome (kindly provided by Charles M. Rice, Center for the Study of Hepatitis C, The Rockefeller University, New York, NY, USA) as a template. Amplicons were subcloned in-frame with GFP in the self-inactivating (SIN) lentiviral transfer plasmid pHR’SINcPPT CEW (pLentiGFP) by introducing an NdeI site in the forward primer and a BamHI site in the reverse primer (pLentiHCVGFP). An epitope tag from the influenza virus hemagglutinin was cloned in-frame at the 5' end.

Lentiviral production
HEK-FT packaging cells (Invitrogen) were plated in 12-well plates at a density of 2.5 x 105 cells per well the day before transfection. Cells were washed with OptiMEM (Invitrogen) prior to transfection and transfected with pLentiGFP or pLentiHCVGFP, together with gag/pol and vsv capside, using Lipofectamine 2000 (Invitrogen), according to the manufacturer’s guidelines. At 48 h and 72 h, transfection efficiency was evaluated by FACS analysis using a CyanADP-MLETM flow cytometer (DakoCytomation, Denmark). Lentiviral supernatants were collected 48 h and 72 h after transfection.

Lentiviral transduction
Jurkat cells were plated at a density of 105 cells per well (5x105 cells/ml) in 24-well plates. Lentiviral supernatant was added, and cells were cultured for 48 h in a 37°C, 5% CO2 incubator. Infection efficiency was analyzed by means of a CyanADP-MLE flow cytometer (DakoCytomation).

Immunofluorescence
Jurkat cells were mounted onto Poly-D-lysine 70,000-HBr (C6H14N2O2.HBr)n (Serva, Germany) treated coverslips by centrifugation in a cytocentrifuge (Shandon Cytospin3). Coverslips were placed in a 12-well culture dish containing RPMI medium supplemented with 2 mM L-glutamine, 10 mM HEPES, 10% FBS, 1% NEAA, 1% sodium pyruvate, and 1% penicillin/streptomycin and incubated in the presence or absence of 2 µM ionomycin for 15 min at 37°C, 5% CO2. Cells were washed subsequently three times with PBS (pH 7.0) and fixed with 4% formaldehyde in PBS for 15 min at room temperature. Fixed cells were permeabilized with 0.25% (w/v) Nonidet P-40 (NP-40), 0.01% (v/v) NaN3, 5% FBS in PBS (IF buffer) for 15 min at room temperature. Coverslips were incubated with mouse anti-NFATc2 (Acris Antibodies GMBH, Germany) or anti-NFATc1 (BD Biosciences, San Jose, CA, USA) at 1:20 dilution in IF buffer for 30 min at room temperature, followed by three washes in IF buffer. An R-PE-labeled goat anti-mouse secondary antibody (DakoCytomation) was used at a 1:20 dilution in IF buffer for 30 min at room temperature. Coverslips were washed three times in 0.25% (w/v) NP-40, 0.01% (v/v) NaN3, 5% FBS in PBS, and nuclei were stained by addition of 10 µg/ml solution of 4',6-diamidino-2-phenylindole (DAPI). After two additional washes, coverslips were mounted onto microscope slides using 10% glycerol and 0.1 M N-propylgallate to retard photobleaching [37 ]. Cells were analyzed in an Olympus fluorescent microscope using the following excitation ({lambda}ex) and emission ({lambda}em) filters: GFP ({lambda}ex: D470/20; {lambda}em: D510/20), rhodamine phycoerythrin (RPE); ({lambda}ex: D540/27; {lambda}em: D605/55), DAPI ({lambda}ex: D360/40; {lambda}em: D460/50). Pictures were obtained in an Axio Cam high-resolution camera (Zeiss, Thornwood, NY, USA) using the Axio Vision 40ACV 4.1.1.0 software. Slides were exposed for the time needed to obtain comparably exposed figures.

Cell count
Cells (5x105) transduced with HCV core-GFP or GFP were seeded onto a 24-well plate in duplicate, 48 days after transduction (Day 0). Cells were resuspended thoroughly daily, live cells were counted, and dead cells were excluded by trypan blue staining. Duplicate wells were counted at each time-point from Days 1 to 7.

Cell-cycle distribution and flow cytometry
DNA-staining dye Hoechst 33342 (Sigma Chemical Co., St. Louis, MO, USA) was added to the cultures at 17 µM final concentration, and cells were incubated for 2 h at 37°C and subsequently washed in PBS and analyzed by flow cytometry in a CyanADP-MLE (DakoCytomation) using an UV enterprise laser set at 30 mW. For G1/S arrest, thymidine (Sigma Chemical Co.) was added to the culture media to a final concentration of 2 mM. After 14 h incubation at 37°C, cells were washed twice with PBS and incubated in complete growth media for an additional 8 h at 37°C, and thymidine (2 mM) was added to the media for an additional 14 h. For M-phase block, cells were incubated with 0.1 µg/ml nocodazole (Sigma Chemical Co.) for 14 h. At the end of the blocking and at various intermediate time-points, cells were washed with PBS and analyzed by flow cytometry. The percentages of cells in various stages of the cell cycle were determined by using the Summit software package (DakoCytomation).

RNA extraction
Total RNA was extracted using TRI reagent (Sigma Chemical Co.), according to the manufacturer’s protocol. Briefly, Jurkat cells were centrifuged and lysed with TRI reagent (Sigma Chemical Co.), followed by two additional phenol/chloroform extractions and one chloroform extraction and subsequently, precipitated with isopropyl alcohol. Precipitate was washed twice with 70% ethanol and resuspended in diethylpyrocarbonate-treated water (Sigma Chemical Co.). RNA quantity and purity were measured by spectrophotometry (SmartSpec, BioRad, Hercules, CA, USA) and quality by agarose-formaldehyde gel electrophoresis.

RT-quantitative PCRs (qPCRs)
Total RNA was used to synthesize cDNA using the iScript cDNA synthesis kit (BioRad), as described by the manufacturer. Real-time qPCR was performed in an iCycler thermocycler (BioRad) using iQ SYBR Green Supermix (BioRad) and specific primers to amplify 50–55 bp. Purity of the amplified band was assessed by melting-curve analysis and agarose gel electrophoresis. For quantitation, a threshold was set in the linear zone of the amplification curve, and the number of cycles needed to reach it was calculated for each gene [comparative threshold (Ct)]. Normalization was achieved by including a sample with primers for L32 for each sample tested. RT-qPCR reactions were run in parallel with RNA from control versus ionomycin-treated cells GFP versus HCV core-transduced cells or cyclosporine A (CsA)-treated versus untreated HCV core-transduced cells. Fold induction values for each gene were calculated by subtracting the mean difference of each gene and L32 Ct number for each sample from the mean difference of the gene and L32 Ct of the corresponding controls and raising two to the power of this difference. Considering that our biological data follow a bell-shaped distribution, approximately Gaussian, a logarithmic transformation of the data was considered, and a paired t-test was used to determine statistical significance.

Microarray procedures
CodeLinkTM human whole genome microarrays (Amersham Biosciences GE Healthcare, Piscataway, NJ, USA) were used following the manufacturer’s recommendations. Briefly, biotin-labeled cRNA was prepared from 2 µg total RNA. The poly(A) RNA subpopulation was primed for RT (42°C, 2 h) by a DNA oligonucleotide containing the T7 RNA polymerase promoter 5' to a d(T)24 sequence. After second-strand cDNA synthesis (16°C, 2 h), the double-stranded cDNA was purified with QIAquick spin columns (Qiagen, Valencia, CA, USA) and used as a template for an in vitro transcription (IVT) reaction using T7 RNA polymerase (37°C, 18 h) to produce the target cRNA. The IVT was performed in the presence of biotinylated deoxy-UTP to label the target RNA. Biotin-labeled cRNA was then purified using RNeasy columns (Qiagen) and tested for quantity and purity in a spectrophotometer. Target RNA (10 µg) was fragmented (94°C, 20 min), mixed with hybridization buffer, and loaded into an array chamber (Amersham GE Healthcare). Slides were incubated for 18 h at 37°C at 300 rpm in a temperature-controlled, shaking incubator. Hybridized arrays were washed for 1 h at 46°C, with 0.75 x 75 mM Tris-HCl, pH 7.6, 112.5 mM NaCl, 0.0375% Tween 20 (TNT), and stained with Cy5-Streptavidin conjugate (22°C, 30 min), followed by 45-min washes at 22°C with 1 x 0.1 M Tris-HCl, pH 7.6, 0.15 M NaCl, 0.5% Tween 20 (TNT). Following a final, 30-s rinse with 0.1x SSC/0.05% Tween, slides were dried by centrifugation (600 g, 3 min) and scanned in a GenePix Personal 4100A analyzer (Axon Instruments, Molecular Devices, Sunnyvale, Ca, USA). A set of bacterial control mRNAs was included with the total RNA samples during target preparation to monitor each step of the procedure.


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RESULTS
 
Expression of HCV core is sufficient to cause NFAT nuclear translocation
The HCV core protein has been shown to increase expression of reported constructs driven from the NFAT promoter and NFAT binding to DNA upon treatment with 12-O-tetradecanoylphorbol 13-acetate (TPA) and TPA + ionomycin, respectively [27 , 32 ]. However, a direct effect of HCV core in NFAT nuclear translocation, the first event of NFAT activation, has not been demonstrated. We wanted to analyze whether the HCV core protein was sufficient to induce NFAT nuclear translocation in Jurkat T cells freshly transduced with lentiviral vectors expressing HCV core, fused in-frame with GFP at its C-termini. GFP-transduced Jurkat cells were used as controls (Fig. 1 ). Immunofluorescence experiments were performed using antibodies directed against NFATc2. As shown in Figure 2 , NFATc2 is localized in the nucleus of cells expressing HCV core-GFP, and it is in the cytoplasm of unstimulated, GFP-transduced control cells. Cells transduced with lentivirus expressing GFP did not cause any change in NFATc2 subcellular localization, and GFP-transduced cells are still able to localize NFATc2 in the nucleus upon ionomycin stimulation (Fig. 2) . Equivalent results were obtained when antibodies against NFATc1 were used, as well as by subcellular fractionation experiments (data not shown).


Figure 1
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  		Figure 1. Jurkat cells were transduced efficiently with HCV core-GFP. Jurkat cells were transduced with lentiviral vectors expressing GFP alone (left panel) or HCV core fused to the N terminus of GFP (right panel). Hatched histograms represent fluorescence from HCV core-GFP or GFP controls, and empty histograms represent fluorescence from untransduced cells.


Figure 2
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  		Figure 2. Nuclear translocation of NFATc2 by the HCV core protein. Jurkat cells were transduced with lentiviral vectors expressing HCV core-GFP or GFP. Cells were left untreated or treated with 1 µM ionomycin for 15 min and subsequently fixed. GFP-expressing cells were stained with anti-NFATc2 antibodies followed by RPE-labeled, goat anti-mouse secondary antibody. Nuclei were stained by addition of a 10 µg/ml solution of DAPI. NFATc2 nuclear translocation was analyzed on an Olympus fluorescence microscope with a 100x original magnification.

Decreased cell proliferation and a delay in cell-cycle progression in HCV core-transduced cells
An additional effect of HCV core expression was a steady decline in cell count of core-transduced cell cultures, 3–4 days post-transduction (Fig. 3A ). As the addition of recombinant core protein to T cell cultures inhibits cell-cycle progression through interaction with gC1q complement receptor (gC1qR) [38 ], we wanted to analyzed whether intracellularly expressed, HCV core could have an effect in cell-cycle progression in HCV-transduced cells, even if under these circumstances, HCV core could not act by interacting with gC1qR. As reported previously [39 ], only minor differences were seen in cell-cycle distribution among populations of asynchronously growing cells expressing HCV when compared with controls at any time-point of the 7-day period tested (Fig. 3B) nor were there any consistent differences in cell death measured by propidium iodine (PI), annexin, or 7-amino actinomycin (data not shown). To show differences stemming from a slowdown in cell-cycle progression, we treated cells with nocodazole or thymidine to block cell-cycle progression in G2/M and G1/S transitions, respectively. CD4+ Jurkat cells were transduced with HCV core-GFP or GFP as a control, and 48 h after transduction, cells were treated with nocodazole or thymidine. To measure DNA content, cells were stained with Hoecht at various time-points and analyzed 2 h after staining by flow cytometry; dead cells were excluded by PI staining. The time needed to reach the M-phase for HCV core-GFP versus GFP control cells was evaluated. As shown in Figure 3C , after 14 h of nocodazole treatment, more than 50 ± 0.1% of GFP control cells had already reached M-phase, and only 31.5 ± 1.5% of HCV-transduced cells had, while 24.3 ± 2.6% of cells were still in G0/G1, as opposed to 13 ± 1% for control cells. No significant differences between HCV-transduced and control cells are shown when cells are blocked in G1/S transition by thymidine treatment.


Figure 3
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  		Figure 3. Cell count and cell-cycle flow cytometry analysis of Jurkat cells expressing HCV core. (A) Cell count of Jurkat cells transduced with HCV core GFP- or GFP-expressing lentiviral vector: Jurkat cells (50x105) were seeded on Day 0 and counted daily until Day 7. Error bars were calculated from three independent experiments. (B and C) Cell-cycle distribution of HCV core-transduced Jurkat cells (hatched) versus control, GFP-transduced cells (empty). Cell-cycle distribution of asynchronous, growing cells (B) or cells blocked in G2/M transition (nocodazole treatment for 6 or 14 h) or G1/S transition (thymidine treatment for 14 h, left panels, or two successive 14-h thymidine treatments with 8 h culture without thymidine in between, right panels). Embedded graphics show statistics of three experiments.

Up-regulation of AR genes in humans
As HCV core is sufficient to activate NFATc2 [27 , 32 ], and NFATc2 is implicated in the induction of an array of AR genes [28 ], we were interested in studying the effect of HCV core on the expression of such genes. HCV infects primarily humans, and we described AR genes in mice; thus, we first needed to analyze the expression of AR genes in human T cells. As the effect of HCV core on NFAT-mediated transcription has been shown in Jurkat cells [27 , 32 ], we used such cell line to ascertain the expression of AR genes upon Ca2+ ionophore addition, the initial treatment used to identify AR genes in mice. Jurkat cells were treated with the calcium ionophore ionomycin for 16 h in equivalent conditions to those sufficient to up-regulate AR genes in mice. Total RNA was purified, and the expression of AR genes was determined by qPCR. As shown in Figure 4A , 13 of the 14 genes tested were up-regulated in human T cells under equivalent conditions, as they do in mice.


Figure 4
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  		Figure 4. AR genes are induced in human Jurkat cells by ionomycin or HCV core expression. (A) Ionomycin treatment induces expression of AR genes in human Jurkat cells, which were treated with 1 µM ionomycin for 16 h. RNA was isolated, and levels of AR genes were determined by real-time qPCR. Expression in ionomycin-treated cells is plotted relative to that in untreated controls. (B) HCV expression induces AR genes. Jurkat cells were lentivirally transduced with HCV core GFP or GFP. Expression of AR genes was determined as in A. Expression in HCV core-expressing cells is plotted relative to that in GFP-expressing controls. Results are average ± SD of three independent experiments. LDH, Lactate dehydrogenase; DAGk, diacylglycerol kinase; RPTP{kappa}, receptor-type protein tyrosine phosphatase {kappa}; TLE4, transducin-like enhancer of split four/Groucho; RGS2, regulator of G protein signaling 2; PTP-1B, protein tyrosine phosphatase 1B; SOCS2, suppressor of cytokine signaling 2; RAB10, ras-related GTP-binding protein 2; RPTP{sigma}, receptor PTP-{sigma}; TRAF5, TNFR-associated factor 5.

Up-regulation of AR genes by HCV core expression
As AR genes were also up-regulated in humans, we wanted to determine whether HCV-induced T cell unresponsiveness was enacted, at least in part, by the up-regulation of such genes. Thus, we transduced Jurkat T cells with HCV core lentiviral vectors and tested for the expression of AR genes by RT-qPCR. As shown in Figure 4B , HCV core-expressing cells had increased expression of all AR genes, including LDH, which was not up-regulated previously in human Jurkat cells by ionomycin treatment. As a control and also to evaluate the implication of Ca2+/calcineurin-mediated signals in HCV core-induced up-regulation of AR genes, we compared the expression of such genes in HCV core-transduced cells, which were left untreated or treated with CsA at 1 µM for 16 h. As shown in Figure 4C , CsA blocks the induction of all AR genes, which had been induced by HCV core.

HCV induces additional genes
The HCV core protein has been shown to modify several signaling pathways in addition to Ca2+/NFAT-mediated signals, so we were interested in identifying additional genes affected by HCV core expression. To explore these changes, we performed DNA microarray experiments, followed by a RT-qPCR confirmation. cRNA from HCV-GFP-transduced versus GFP-transduced control cells was used to hybridize CodeLink human whole genome microarrays. CodeLink microarrays allow for genome-wide gene expression analysis, targeting ~57,000 transcripts and expressed sequence tags. To prevent gene expression biases as a result of a predominance of subpopulations, we used Jurkat cells, a well characterized CD4 T cell line. To prevent gene-expression differences, stemming from a prolonged, separate tissue culture of HCV-expressing cells versus control cells, we have taken advantage of lentiviral transduction, which allowed us to analyze cells only after hours of culturing. A total of 59 genes was identified as significantly, differentially expressed (P=0.01) with 22 up-regulated genes and 37 down-regulated genes. All gene expression data and significantly differentially expressed gene lists are available as Supplemental material. Twenty-nine out of the 59 genes were of partially known function. RT-qPCR assays were used to confirm the differential expression of the genes of relevant function. Figure 5 shows differentially expressed genes, which were confirmed consistently by qPCR experiments in independent RNA preparations. Differentially expressed genes belong to several functional categories, including those that affect vesicle trafficking and endocytosis, transcription and translation, and cell-cycle progression, along with down-regulation of inflammatory cytokine receptors and up-regulation of anti-inflammatory cytokines and molecules involved in cell death by oncosis. Taken together, our results show that HCV core is able to affect human T cell signal transduction in a manner consistent with viral evasion and anergy induction.


Figure 5
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  		Figure 5. HCV core modulates the expression of additional genes. Jurkat cells were transduced with HCV core GFP- or GFP-expressing lentiviral vector. mRNA was purified and used to hybridize whole genome CodeLink microarrays (see Supplemental material). Expression of differentially expressed genes with relevant function was confirmed by qPCR experiments as in Figure 4 . Genes confirmed by three independent qPCR experiments are shown. ARF6, ADP-ribosylation factor 6; ACTG1, {gamma}-1-actin; CAP1, adenylate cyclase-associated protein 1; PLDN1, pallidin homolog; BAZ1A, bromodomain adjacent to zinc finger domain, 1A; FBXLIO3, F-box and leucine-rich repeat protein10; MINK1, mis-shapen/Nck-interacting kinase-related kinase 1; RB1, retinoblastoma 1; EIF4HG2, eukaryotic translation initiation factor 4 gamma, 2; MX1, myxovirus (influenza) resistance 1; NFATc2IP, nuclear factor of activated T cells, cytoplasmic, calcineurin-dependent 2 interacting protien; CRLF1, cytokine receptor-like factor 1; Porimin, pro-oncosis receptor inducing membrane injury gene.

HCV infects CD4 T cells in HCV chronically infected patients
HCV infects T cell lines and primary T cells in culture [35 ], but to further ascertain the clinical significance of our results, we were interested in analyzing whether circulating CD4 T cells from chronic HCV patients were infected by the virus. As shown in Figure 6 , HCV RNA was present in RNA purified from CD4+ T cells from all HCV chronic patients, and it was undetectable in CD4 cells from control individuals. To rule out a passive adsorption of the virus, we incubated CD4+ cells from control individuals with HCV-containing plasma from Patient #1. HCV RNA was undetectable in four out of eight CD4 cells from HCV-incubated controls, and the remaining four samples showed only a low-level contamination (Fig. 6B) .


Figure 6
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  		Figure 6. Detection of HCV RNA in CD4+ cells from HCV chronically infected patients. (A) HCV RNA copies (IU) detected by RT-PCR (COBAS AmpliPrep/COBAS TaqMan HCV test) in magnetically purified CD4+ cells from five HCV patients (P1–5) versus five healthy controls (C1–5). (B) HCV-containing plasma from Patient #1 was incubated with CD4 cells from eight healthy controls (C1–8) to quantify passive adsorption.


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DISCUSSION
 
Tolerance encompasses a complex interplay between positive and negative signals, which are up-regulated after a particular tolerizing stimulus [40 ], with the fate of the cell being decided in the initial stages following antigenic stimulation [19 , 40 ]. Thus, it is pivotal to analyze the molecular events, which unfold in the initial period after the tolerizing stimulus arises. In the present work, we study the molecular events associated with HCV core protein expression, which acting as a preconditioning stimulus, will direct T cells toward a nonresponding mode, allowing for viral persistence. Our work is the first to analyze the events, which unfold during the initial stages of HCV core expression in T cells. Toward such a goal, we have used lentiviral transduction, which yields expression levels similar to those seen during the course of the natural infection. In Jurkat cells, it is feasible to transduce close to 100% of the cells without the need for prolonged cell cultures required to select stably transfected clones. In addition, the use of a cell line such as Jurkat offers the homogeneity needed to extract meaningful, differential expression data. Su et al. [41 ] showed a genome-wide analysis in liver biopsies of HCV-infected chimpanzees focusing on genes, which correlated with viral clearance; here, we have a simplified approach focusing on a direct effect of HCV core by using a T lymphocyte cell line, which will eventually allow for mechanistically relevant data.

HCV core has been proposed to act primarily through up-regulation of NFAT activation [26 , 27 , 32 ], but an increase of NFAT-binding activity in the nucleus of HCV core, stably transfected Jurkat cells has only been shown after stimulation of TPA + ionomycin [32 ]; thus, a direct effect on NFAT nuclear translocation, the initial event of NFAT activation, has not been addressed yet. In this paper, we show that HCV core transduction is sufficient to cause NFAT nuclear translocation, regardless of the level of HCV core expression, indicating that low-level expression is sufficient to cause NFAT translocation. We also show a slowdown in cell-cycle progression and a decreased cell count in HCV-transduced cultures, consistent with observations from clinical infections, where a decreased, proliferative response in HCV-infected patients has been reported [9 ]. An effect of extracellularly added, recombinant HCV core in cell-cycle progression of lymphocytes has already been reported and shown to be mediated by interaction with gC1qR. Based on the fact that we obtain similar results by an intracellularly expressed HCV core, it could be hypothesized that gC1qR could be just a means by which HCV core and/or the whole virus enter T lymphocytes in vivo, as it was suggested already [38 ]. In this paper, we observe that HCV core-transduced cells show an increased expression of a set of AR genes, which was shown by us to be up-regulated in anergized murine cells, not only "in vitro" but also "in vivo." Moreover in a genome-wide analysis, we identify an additional set of genes up-regulated by HCV core in T cells. Several genes in both groups have a potential effect in hampering cell-cycle progression.

NFAT was shown to be necessary, although not sufficient, to up-regulate some AR genes [28 ]. Conversely, HCV core is sufficient to up-regulate all AR genes (Fig. 4) ; thus, HCV core is able to induce all necessary signals NFAT- and non-NFAT-dependent. HCV core-up-regulated AR genes encode diverse categories of proteins, which have been suggested to impose an anergic state in murine T cells [28 , 29 , 40 ] and can mediate a HCV-specific unresponsiveness, which ensues in chronicity.

Among AR genes, PTP-1B, a soluble receptor tyrosine phosphatase, will interfere with signaling pathways coupled to antigen receptors [42 ]. DAGK{alpha} metabolizes diacylglycerol, thus inhibiting protein kinase C activation [43 ] (an antianergizing stimulus [40 ]) and has been shown to block apoptosis [44 ]. CD98 activates Rap1 [45 ], linked to the impaired activation of the ERK-MAPK pathway, observed in anergic T cells [46 ].

AR genes involved in proteolitic pathways are strongly induced by HCV core, such as pro-caspase 3, which up-regulation has been correlated already with down-modulation of the TcR/CD3 {zeta}-chain in HCV-infected patients [47 ]. This result is consistent with a role for caspases in regulating signal transduction by limited proteolysis of signaling molecules in the absence of apoptosis [28 ], which could be even prevented by the virus as a means to maintaining potentially suppressive clones [48 ]. Among the AR genes up-regulated by HCV core, there are many negative regulators of transcription, such as Ikaros [49 , 50 ], the Groucho-related protein TLE4 [51 ], and the DNA-binding protein jumonji [52 ]. Another jumonji-related gene FBXL10 has been identified in the genome-wide analysis shown in this paper as being up-regulated by HCV core. FBXL10 protein also harnesses an Fbox motif implicated in degradation of ubiquitinated proteins [53 ] (ubiquitination is strongly related to anergy induction [31 , 40 ]). Translation is also affected by HCV core through up-regulation of the general repressor of translation eIF4GII, which is widely used as a viral evasion mechanism [54 ] and has been shown to form translationally inactive complexes favoring translation of viral RNAs [55 ].

Molecules involved in cytokine signal transduction and regulation are also modulated by HCV core: Nuclear protein NIP45 could mediate up-regulation of IL-4 transcription [56 ], consistent with the increased IL-4 secretion observed in cells expressing HCV core [32 ]. Conversely, CRLF1, an IL-6 family-soluble cytokine receptor with agonistic activity [57 ], is down-regulated by HCV core. Among the genes found to be up-regulated by HCV core in our genome-wide analysis, MINK and RB1 are implicated in blocking cell-cycle progression, consistent with the effect shown in HCV core-transduced cells. In liver cells, inconsistent results have been shown regarding Rb regulation [58 59 60 ], and Rb down-regulation is mediated by NS5B and not core protein [59 ]. In CD4+ T lymphocytes, where no HCV-mediated malignization has been reported, the predominant HCV core effect is that of an up-regulation of Rb, causing cell-cycle arrest as shown in this paper and others [38 ]. Mink, a MAP 4k, has been shown to induce a cell-cycle arrest [61 ], which could also be responsible for the cell-cycle slowdown, which we report in HCV core-transduced cells. Cytoskeleton reorganization, vesicle trafficking, and endocytosis have been shown to be pivotal for anergy induction [30 , 40 ], and some HCV core-up-regulated genes are implicated in those phenomena, such as ACTG1, a component of the cytoskeleton, the actin-binding protein, CAP1, PLDN (a molecule, which plays a role in vesicle trafficking [62 ]), and ARF6, which localization to the membrane in anergic cells has been proposed as an anergy marker [63 ].

Sustained up-regulation of Ca2+ and NFAT in the absence of a concomitant activation of NF-{kappa}B and AP1 will divert NFAT toward transcription of AR genes, whose products impose a tolerant state [28 29 30 31 , 40 ]. Early on, an alternative hypothesis was proposed [64 ], according to which anergy was the result of TCR engagement in the absence of proliferation, stating that cell-cycle arrest prevents the degradation of anergic factors [65 ] such as p27 [66 ] and linking anergy and cell-cycle progression. Consistent with both hypotheses, HCV core not only activates Ca2+ and NFAT [26 , 27 , 32 ] and blocks NF-{kappa}B [33 ] and AP1 [34 ] but also is able to block proliferation by affecting p27 degradation directly [38 ]. Here, we show that HCV core affects NFAT translocation directly, causes a decreased lymphocyte proliferation, and activates several molecules potentially involved in cell-cycle arrest. All of those HCV core-induced changes are relevant in clinical situations, as we have shown that HCV RNA is detected in CD4 cells from HCV chronically infected patients. Infection and destruction of helper lymphocytes have been proposed as possible causes for the impaired T cell response against the virus [4 ]. Nevertheless, the presence of HCV in T cells has been shown inconsistently [67 ]: Mazin et al. [68 ] reported the presence of HCV RNA in CD2+ cells from three out of five patients, and only one patient was reported to have HCV RNA in B cells, and Lerat et al. [69 ] find HCV RNA in PBMC in B cells and not T cells [70 ]. We have shown that HCV RNA is detected in CD4+ cells from all patients tested, which is in keeping with a recent report showing that HCV infects primary CD4+ T cells and infects and replicates in CD4 T cell lines [35 ]. Discrepancies are likely a result of sensitivity issues in the test used in older reports, considering that most of those tests have already been replaced. In addition, core protein, which has been shown to be circulating in blood from HCV-infected individuals [71 ], could enter CD4+ T cells through the C1qR [38 , 72 , 73 ], increasing the effect of the virus in T cell signal transduction.

Our results, showing a HCV core-mediated inhibition of CD4 T cell proliferation, are in keeping with clinical data, showing a decreased number of virus-specific T cells in HCV-infected patients compared with other viral infections [74 , 75 ]. The impaired, proliferative capacity of CD8+ cells is associated with weak, ex vivo, HCV-specific CD4+ T cell responses in HCV chronically infected patients when compared with those that have recovered [75 ]. HCV-specific CD8 T cells have also been shown to be less differentiated, displaying reduced effector functions upon antigen stimulation, and T cell effector functions are restored by the addition of IL-2, which has been proposed to be a result of a HCV core-dependent defect of CD4+ T cells [76 ], rendering CD8+ cells into a "helpless state" [77 ]. Sugimoto et al. [78 ] compared HCV-specific T cells from chronic versus recovered patients, showing that HCV persistence is associated with a global quantitative and functional suppression of HCV-specific T cells, and no differential antigenic hierarchy or cytokine phenotype was related to HCV clearance, keeping with a global effect caused by viral proteins. As we show for HCV core-expressing cells, ex vivo CD4+ defects are associated mostly with a decreased proliferation [77 , 79 ]. Indeed, an early impairment of HCV-specific T cell proliferation during acute infection is the best predictor of viral persistence, as shown in the largest cohort study performed to date [80 ]. The former clinical findings are explained best as a HCV core-mediated effect on T cells, as low concentrations of the HCV core antigen have long since been implicated in a down-regulation of cellular immune responses [81 ].

Our results could aid in explaining the differential response to HCV, where patients, who clear the virus with a subclinical infection, coexist with persistent infections, which destroy the liver of other patients. The narrow line, which separates tolerance from destruction [19 , 40 ], is bridged easily by viruses such as HCV with mild APC-stimulatory capacity carrying the ability to mingle with CD4 T cell signaling pathways. In this scenario, small differences in viral load or genetically determined host idiosyncrasies in the "strength" of any of the signaling pathways shown by us to be altered by the virus could tilt the balance toward viral clearance or persistence. Among HCV proteins, core is the best-suited to act in this initial phase, as it is the first HCV protein to be expressed [82 ] and has also been shown to be secreted onto the blood, thus amplifying its immunomodulatory effect [71 ].

Our results aid in establishing the mechanisms by which HCV alters lymphocyte homeostasis in the initial phase of infection, pivotal for understanding the tendency to chronicity observed in HCV-infected patients and to explain treatment failure. In addition, vaccination strategies for HCV have to take into account deleterious, immune-evasion mechanisms elicited by specific HCV molecules present in the vaccine [83 ].


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ACKNOWLEDGEMENTS
 
This work was supported by grants (SAF2005-00458 and Programa Ramon y Cajal) from the Ministerio de Educacion y Ciencia, (FIS-PI050715) from the Ministerio de Salud (Spain) and (0080/2005) from Consejeria de Salud de Andalucia, Spain to F. G-C. M. D-V. is a graduate fellow of the FIS program (Ministerio de Salud), and A. M-S. is a graduate fellow of the Junta de Andalucia. We thank C. M. Rice for the original HCV genome, D. Trono for the original version of the lentiviral vector, F. Martin for advice on troubleshooting lentiviral transduction, and M. M. Valdivia for advice and reagents for cell-cycle synchronization.

Received May 31, 2007; revised July 18, 2007; accepted July 24, 2007.


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K. R. Jerome
Viral Modulation of T-Cell Receptor Signaling
J. Virol., May 1, 2008; 82(9): 4194 - 4204.
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