ABSTRACT
Accumulated evidence indicates that immune cells can support the replication of hepatitis C virus (HCV) in infected patients and in culture. However, there is a scarcity of data on the degree to which individual immune cell types support HCV propagation and on characteristics of virus assembly. We investigated the ability of authentic, patient-derived HCV to infect in vitro two closely related but functionally distinct immune cell types, CD4+ and CD8+ T lymphocytes, and assessed the properties of the virus produced by these cells. The HCV replication system in intermittently mitogen-stimulated T cells was adapted to infect primary human CD4+ or CD8+ T lymphocytes. HCV replicated in both cell types although at significantly higher levels in CD4+ than in CD8+ T cells. Thus, the HCV RNA replicative (negative) strand was detected in CD4+ and CD8+ cells at estimated mean levels ± standard errors of the means of 6.7 × 102 ± 3.8 × 102 and 1.2 × 102 ± 0.8 × 102 copies/μg RNA, respectively (P < 0.0001). Intracellular HCV NS5a and/or core proteins were identified in 0.9% of CD4+ and in 1.2% of CD8+ T cells. Double staining for NS5a and T cell type-specific markers confirmed that transcriptionally competent virus replicated in both cell types. Furthermore, an HCV-specific protease inhibitor, telaprevir, inhibited infection in both CD4+ and CD8+ cells. The emergence of unique HCV variants and the release of HCV RNA-reactive particles with biophysical properties different from those of virions in plasma inocula suggested that distinct viral particles were assembled, and therefore, they may contribute to the pool of circulating virus in infected patients.
IMPORTANCE Although the liver is the main site of HCV replication, infection of the immune system is an intrinsic characteristic of this virus independent of whether infection is symptomatic or clinically silent. Many fundamental aspects of HCV lymphotropism remain uncertain, including the degree to which different immune cells support infection and contribute to virus diversity. We show that authentic, patient-derived HCV productively replicates in vitro in two closely related but functionally distinct types of T lymphocytes, CD4+ and CD8+ cells. The display of viral proteins and unique variants, the production of virions with biophysical properties distinct from those in plasma serving as inocula, and inhibition of replication by an antiviral agent led us to ascertain that both T cell subtypes supported virus propagation. Infection of CD4+ and CD8+ T cells, which are central to adaptive antiviral immune responses, can directly affect HCV clearance, favor virus persistence, and decisively influence the development and progression of hepatitis C.
INTRODUCTION
Hepatitis C virus (HCV) is a positive-strand RNA virus of approximately 9.4 kb that causes chronic hepatitis C (CHC) in up to 80% of those with clinically identifiable infection. The disease affects over 170 million people globally, with up to one-fifth of those advancing to liver cirrhosis, who are at a greater risk of developing primary hepatocellular carcinoma (1, 2). In addition to symptomatic chronic infection, which is normally accompanied by circulating HCV RNA and anti-HCV antibodies, HCV also persists as an essentially clinically silent (occult) infection accompanied by very low levels of HCV RNA in serum (usually below 100 to 200 virus genome copies or virus genome equivalents [vge]/ml), liver, and peripheral blood mononuclear cells (PBMC) (3). This occult HCV infection (OCI) may continue for decades after spontaneous (self-limited) or antiviral therapy-induced resolution of hepatitis C (4–11). In this context, the accumulated experimental data indicate that HCV propagates not only in the liver but also in the immune system, where it can modify the proliferation and function of affected cells (12–18). It also became apparent that immune cells are a site of long-term persistence of HCV, where the virus may hide from immune surveillance and elimination, as other lymphotropic viruses do (19, 20). The ability of HCV to infect the immune system is consistent with a significantly greater prevalence of certain lymphoproliferative disorders in patients chronically infected with HCV, including non-Hodgkin's lymphoma, mixed cryoglobulinemia, and marginal zone lymphoma (21–24). Notably, the regression of these diseases in considerable numbers of patients treated with anti-HCV therapies provides persuasive evidence for a direct role of HCV in the pathogenesis of these disorders (25, 26).
Previous studies on HCV compartmentalization in the immune system showed the presence of the virus in all main types of circulating lymphomononuclear cells, i.e., B and T lymphocytes and monocytes, in both symptomatically and silently infected individuals (4, 12, 15, 27, 28). The data acquired have shown that HCV lymphotropism is reflected not just in the mere identification of the virus RNA positive (nonreplicative) strand but also in the detection of the virus RNA negative (replicative) strand and intracellularly located structural and nonstructural proteins, including E2 envelope, core, NS3, or NS5a, in both immune cells of HCV-positive patients and cells infected in vitro with patient-derived virus (3, 7, 15, 29–32). Other evidence came from the identification of unique HCV variants in immune cells distinct from those in plasma or livers of the same patients and from the emergence of similar variants in immune cells infected in vitro (15, 27, 29, 33, 34). The eradication of HCV replication from PBMC of patients with OCI and CHC after ex vivo treatment with exogenous interferon alpha (IFN-α) or due to stimulation of the production of endogenous IFN-γ, respectively, and from in vitro-infected T cells treated with IFN-α or telaprevir (TLPV), an HCV-specific protease inhibitor (35), further verified that these cells are the site of active HCV propagation (7, 10, 29, 30). This was congruent with the demonstration that HCV released from naturally infected or in vitro-infected PBMC transmitted infection to primary T cells in culture, resulting in the production of HCV virions that could be precipitated with an antibody against the virus envelope protein and visualized by immunoelectron microscopy (7, 29).
The recent identification of CD5, a lymphocyte-specific glycoprotein belonging to the scavenger receptor cysteine-rich family, as a molecule essential for infection of human T lymphocytes by authentic HCV (30), and the finding that costimulatory receptor B7.2 (CD86) is involved in infection of human memory B cells by a HCV “SB” variant (36), advanced the understanding of the nature of HCV lymphotropism. These findings imply that cell type-specific surface molecules, rather than a combination of molecules naturally displayed on many cell types, mediate HCV lymphotropism. Regarding viral determinants facilitating HCV tropism toward immune cells, the most recent findings elegantly showed that a specific E1-E2 protein epitope presented by a distinct subpopulation of viral particles is likely responsible for virus entry into B cells (37).
Although the authenticity of HCV lymphotropism is now evident, and sizable progress has been made in deciphering this event, there remains a substantial paucity in the data on several fundamental aspects of this infection, including (i) susceptibility and the degree to which individual immune cell subsets support HCV propagation, (ii) genetic and biochemical features predisposing the virus to infection of immune cells, and (iii) molecular, biophysical, and pathogenic properties of the virus produced. We have previously shown that HCV can replicate in CD4+ and CD8+ T lymphocytes in patients with CHC and OCI (15). This was based on the demonstration of the HCV RNA negative strand and the virus NS5a protein within the cells and was supported by clonal sequencing analysis showing that different HCV variants were harbored in cells compared to those occurring in patients' plasma. At the same time, an in vitro HCV replication system in normal human primary T cells was established (7, 29). This cell culture model supports the complete cycle of HCV propagation although at much lower level than with the HCV JFH-1–Huh7.5 cell system. However, it needs to be emphasized that authentic, patient-derived HCV but not laboratory-created HCV clones, such as JFH-1, infects PBMC and primary T cells in culture (31, 38). Nonetheless, it remained unknown whether the two closely related but functionally distinct T lymphocyte subpopulations, CD4+ and CD8+ cells, are receptive to HCV infection and capable of supporting HCV replication to the same degree. It was expected that a different intracellular microenvironment, including the augmented expression of IFN-γ in activated CD8+ T cells, might predispose them differently (39, 40). The aims of the present study were to (i) establish an in vitro infection model in which authentic, patient-derived HCV would infect primary CD4+ and CD8+ T lymphocytes; (ii) determine whether productive replication of HCV takes place in both cell types; (iii) recognize the biophysical properties of HCV RNA-reactive particles released by de novo-infected CD4+ and CD8+ T cells and compare them to those of virions in the plasma inocula used to infect the cells; and (iv) recognize whether unique HCV variants not accountable for in the plasma inocula may arise in de novo-infected CD4+ and CD8+ T lymphocytes.
RESULTS
Selection of HCV inocula for infection experiments.When we investigated the susceptibility of PBMC-derived immune cells to infection with authentic, patient-derived HCV, we noticed that the following features help to identify patient plasma samples that are most suitable for infection experiments: (i) donors should be treatment naive, (ii) the plasma HCV RNA load should preferentially be ≥1 × 106 copies or vge/ml, and (iii) PBMC of the plasma donors should be HCV RNA reactive (7, 29, 30, 41). For the present study, these criterions were supplemented with the demonstration that the potential plasma HCV inocula were infectious to the total pool of T lymphocytes. These T cells were derived from PBMC of healthy donors by intermittent stimulation with a T cell proliferation-promoting mitogen, phytohemagglutinin (PHA), in the presence of human interleukin-2 (IL-2), a T cell survival-supporting cytokine, as reported previously (7, 29). Accordingly, the identification of suitable HCV inocula was a three-step process, except when patients' PBMC were not available for examination. Hence, in the first step, the HCV load was quantified in plasma by real-time reverse transcription (RT)-quantitative PCR (qRT-PCR) (sensitivity of 1 × 102 vge/reaction) in 16 randomly selected treatment-naive patients with CHC (Table 1). The HCV RNA positive strand was detected in all patients and was quantifiable for 15 patients. The loads ranged from below 100 vge/ml (unquantifiable), as in patient 5/M, to 3.1 × 107 vge/ml, in patient 10/M (Table 1).
Clinical characteristics and detection of HCV RNA in plasma and PBMC of patients with chronic hepatitis C examined in this studye
In the second step, PBMC available from 13 out of the 16 patients were tested for the HCV RNA positive strand by qRT-PCR. HCV was detected in 5 out of 13 (38.5%) PBMC samples and was quantifiable in 2 of them (Table 1). In three cases with unquantifiable signals, HCV RNA detection was confirmed by nucleic acid hybridization (NAH) of qRT-PCR amplicons by Southern blotting (not shown). PBMC negative for HCV RNA by qRT-PCR (n = 8) were tested by endpoint nested RT-PCR (nRT-PCR)/NAH with a sensitivity that was ∼10-fold higher than that of the former assay (4, 10). This assay identified HCV in 5 of the 8 patients (62.5%) (Table 1). The combined results from qRT-PCR and nRT-PCR/NAH analyses showed that virus was detected in 10 out of 13 CHC patients (76.9%) whose PBMC were available for analysis (Table 1). It is of note that PBMC were examined without prior ex vivo mitogen stimulation that normally augments HCV RNA expression and virus replication in PBMC of patients with low HCV loads in these cells (4, 9, 10, 15, 29, 30, 42).
In the final step, plasma samples with the highest HCV loads (i.e., ≥1 × 106 vge/ml), from patients 10/M and 11/M, whose PBMC were HCV RNA reactive, as well as plasma from patient 16/F, whose PBMC were not available for examination, were tested for infectivity toward total T cells derived from PBMC of healthy donor A/M or B/F. The HCV RNA positive strand was detectable in the cells following the completion of the different phases of infection with all three inocula. However, the HCV RNA negative strand, indicative of active virus replication, was detected only after the cells were exposed to plasma samples from donors 11/M and 16/F (Fig. 1; also see Fig. 9). As a consequence, plasma specimens from these two patients, designated inocula HCV-11/M and HCV-16/F, were used in infection experiments with primary CD4+ and CD8+ T cells. Both inocula carried HCV genotype 1a (Table 1).
Detection of HCV RNA positive and negative strands in total T cells after in vitro infection by HCV-positive plasma from patients 11/M and 16/F. Monocyte-depleted PBMC from healthy donor A/M were pretreated with PHA, exposed to test inocula, and cultured as indicated in Materials and Methods. RNA from the cells collected at different days (d) postinfection was analyzed by RT-PCR/NAH for the expression of the HCV RNA positive strand (A) and the HCV RNA negative strand (B). Synthetic HCV RNA (HCV sRNA) positive (pos) and negative (neg) strands at 1 × 105 copies/reaction were used to confirm the specificity of detection. The rHCV UTR-E2 fragment at 1 × 105 copies/reaction served as an additional positive control for the detection of the HCV RNA positive strand. Water instead of cDNA amplified in direct (D/W) and nested (N/W) reactions as well as a mock-treated sample without RNA (Mock) were included as negative and contamination controls. Positive signals showed the expected 244-bp bands.
HCV genome expression and replication status in de novo-infected CD4+ and CD8+ T cells.Six separate experiments were completed, in which purified CD4+ or CD8+ T cells were exposed to the HCV-11/M or HCV-16/F inoculum, cultured under the same conditions, and evaluated (Tables 2 and 3 and Fig. 2). The HCV RNA positive strand was detected in CD4+ T cells at all time points postinfection, apart from 14 days postinfection (dpi) in experiment 1 (Fig. 2). With regard to CD8+ T cells, the HCV RNA positive strand was identified at all time points in all experiments, except at 14 dpi in experiment 3 (Table 2 and Fig. 2). Taken together, the HCV RNA positive strand was detected equally as often in CD4+ (15/16; 93.7%) and CD8+ (13/14; 92.8%) cells (Table 3). Estimated levels of expression of this strand in CD4+ cells ranged from 30 to 1 × 104 vge/μg RNA, with a mean level of 2.7 × 103 ± 1.1 × 103 vge/μg. The levels in CD8+ T cells were within the same range but with a mean value of 4.4 × 103 ± 1.3 × 103 vge/μg (Table 3). This difference was not statistically significant (P = 0.30).
Detection of HCV RNA positive and negative strands in CD4+ and CD8+ T cells exposed to authentic, patient-derived HCVc
Parameters for detection of HCV RNA positive and negative strands in CD4+ and CD8+ T cells exposed to authentic, patient-derived HCV
Detection of HCV RNA positive and negative strands in CD4+ and CD8+ T lymphocytes infected with authentic HCV. Affinity-purified CD4+ and CD8+ T cells from healthy donors B/F and A/M pretreated with PHA were exposed to HCV-11/M or HCV-16/F plasma or to healthy donor plasma (HDP). (A) Cells from experiment 1 infected with HCV-11/M or exposed to HDP. (B) Cells from experiment 3 infected with HCV-11/M. (C) Cells from experiment 5 infected with HCV-11/M or exposed to HDP. (D) Cells from experiment 6 infected with HCV-16/F. CD4+ and CD8+ T cells were cultured under intermittent stimulation with PHA as described in Materials and Methods. When the numbers of cells recovered allowed, CD4+ and CD8+ T cells were also exposed to HDP and cultured under the same conditions as those for infected cells (mock infection). RNA was analyzed for HCV RNA positive and negative (replicative) strands by RT-PCR/NAH assays. Contamination and specificity controls included water added instead of cDNA and amplified by direct PCR (D/W) and, if applicable, nested PCR (N/W) and an RNA/cDNA-free mock (Mock) sample. The synthetic HCV RNA positive (HCV sRNA pos) strand at 1 × 106 copies was used as the positive and specificity control, and the rHCV UTR-E2 fragment at 1 × 106 copies was used as an additional positive control for the detection of the HCV RNA positive strand. Serial dilutions of the synthetic HCV RNA negative (HCV sRNA neg) strand and the HCV sRNA positive strand were used to confirm the specificity of detection and as quantitative standards for the detection of the HCV RNA negative strand. Positive signals showed the expected 442-bp (direct RT-PCR/NAH) or 244-bp (nested RT-PCR/NAH) oligonucleotide fragments.
Considering the HCV RNA negative strand, this replicative intermediate was identified in both CD4+ and CD8+ T cells in all six experiments although not at all time points postinfection where the HCV RNA positive strand was reactive (Tables 2 and 3). Specifically, the strand was identified in 8 of 15 CD4+ T cell samples (53.3%) at estimated levels ranging from ≤25 to 5 × 103 copies/μg RNA and with a mean level of 6.7 × 102 ± 3.8 × 102 copies/μg RNA. In CD8+ T cells, the strand was detected in 6 of 13 cell samples (46.1%), and the load ranged from ≤25 to 1 × 103 copies/μg, with a mean of 1.2 × 102 ± 0.8 × 102 copies/μg RNA (Tables 2 and 3). Overall, the virus RNA negative strand was detected more often in CD4+ than in CD8+ T cells (53.3% versus 46.1% of evaluations completed) and at a significantly higher mean level in CD4+ than in CD8+ cells (P < 0.0001). Not surprisingly, HCV RNA negative-strand positivity usually coincided with higher levels of positive-strand expression in either CD4+ or CD8+ T lymphocytes (Tables 2 and 3).
The irregular detection of HCV RNA positive and negative strands during intermittent stimulation of CD4+ and CD8+ T cells with PHA/IL-2 was consistent with previous observations (7, 29). There might be several reasons behind this uneven detection, among which multiple rounds of infection of primary T cells, which are not fully synchronized considering the proliferation cycle; the status of intracellular antiviral innate responses and expression of virus receptor molecules; and the amount and variant composition of the virus released during repeated cell stimulation could be the most influential. In addition, the ≥10-fold-higher sensitivity of the assay detecting the HCV RNA positive strand than that of the assay identifying the negative strand has to be taken under consideration when the expression levels of these two infection markers are compared (4, 15, 30).
Identification of HCV NS5a and core proteins in de novo-infected CD4+ and CD8+ T cells.To determine whether the detection of the HCV genome and its replication in CD4+ and CD8+ T cells was accompanied by the synthesis of viral proteins, lymphocytes exposed to the HCV-11/M inoculum (experiment 5, 10 and 14 dpi) were examined for the presence of HCV core or NS5a (Fig. 3). In addition, the cell subsets exposed to HCV-16/F (experiment 6, 10 and 14 dpi) were assessed for the presence of both virus NS5a and CD4 or CD8 by double staining with antibodies conjugated with contrasting fluorochromes (Fig. 4). As shown by confocal microscopy, both HCV proteins were detected in the cytoplasm of CD4+ and CD8+ T lymphocytes. Their staining was usually homogeneous throughout the cytoplasm, while occasionally seen granular staining tended to accumulate at the cytoplasmic side of the plasma membrane in both cell types (Fig. 3 and 4). Percentages of NS5a- or core-reactive cells enumerated by confocal microscopy were 0.9% and 1.2% for CD4+ and CD8+ cells, respectively, and were similar at different time points after infection with HCV-11/M. CD4+ and CD8+ T cells exposed to healthy donor plasma (HDP) and stained for the HCV NS5a or core protein (Fig. 3 and 4) as well as CD4+ and CD8+ T cells exposed to HCV-11/M and incubated with the mouse IgG1k isotype control (data not shown) revealed no staining. In summary, HCV core and NS5a were clearly displayed in both CD4+ and CD8+ T cells infected with HCV in culture. In addition, the NS5a protein was detected in CD4+ or CD8+ T lymphocytes after double staining with anti-NS5a and anti-CD4 or anti-CD8, conclusively documenting that both T cell subtypes supported the replication of a translationally competent virus after in vitro infection.
Identification of HCV core and NS5a proteins by confocal microscopy in CD4+ and CD8+ T cells infected with authentic, patient-derived HCV. Affinity-purified CD4+ T cells (A) or CD8+ T cells (B) from healthy donor B/F were exposed to HDP or the HCV-11/M inoculum and cultured as described in Materials and Methods. The cells collected at 10 or 14 dpi (experiment 5) were stained with either anti-HCV core (anti-core) or anti-HCV NS5a (anti-NS5A) antibody and counterstained with DAPI. The images were captured at an original magnification of ×60. Positive cells show cytoplasmic staining of either the HCV core or NS5a protein, while T cells exposed to HDP are negative.
Double staining of CD4+ and CD8+ T lymphocytes infected with the HCV-16/F inoculum with anti-NS5a and anti-CD4 or anti-CD8 antibodies. Affinity-purified CD4+ cells (A) and affinity-purified CD8+ T cells (B) from healthy donor A/M were exposed to HCV-16/F or HDP, as a negative mock control, if numbers of recovered cells allowed, and cultured as indicated in Materials and Methods. The cells collected at 10 or 14 dpi (experiment 6) were stained with anti-CD4 or anti-CD8 antibody, counterstained with DAPI, and then incubated with anti-NS5a antibody. The images were captured at an original magnification of ×60.
Inhibition of HCV replication in CD4+ and CD8+ T lymphocytes by telaprevir.Treatment of CD4+ and CD8+ T cells with 4 μM TLPV, under conditions established in our previous studies (10, 30), inhibited the replication of HCV-11/M and HCV-16/F in both cell subtypes, as revealed by the absence of the HCV RNA negative strand (Fig. 5). In contrast, the HCV RNA replicative strand remained detectable in infected T cell subsets treated with 0.5% dimethyl sulfoxide (DMSO) alone, which was used as a vehicle to solubilize TLPV. These results further ensured the authenticity of active HCV replication in both CD4+ and CD8+ T cells (Fig. 5).
Inhibition of HCV replication in CD4+ and CD8+ T lymphocytes by treatment with the HCV-specific protease inhibitor TLPV. Affinity-purified CD4+ and CD8+ T cells from donor A/M were exposed to authentic HCV-11/M or HCV-16/F in the presence of 4 μM TLPV (T, treated) or 0.5% DMSO alone (UT, TLPV untreated) for 72 h. Experiments with CD4+ and CD8+ T cells exposed to HCV-16/F and TLPV or 0.5% DMSO were done in duplicate, whereas those with HCV-11/M and TLPV or 0.5% DMSO were done once because to sparsity of the inoculum. The HCV RNA negative strand in CD4+ T cells (A) and CD8+ T cells (B) was identified by strand-specific RT-PCR/NAH in which sHCV RNA positive (pos) and negative (neg) strands at 1 × 104 copies/reaction were used as specificity controls. Water amplified in direct (DW) and nested (NW) reactions and a mock (M) extraction served as contamination controls. Positive signals demonstrated the expected 244-bp oligonucleotide fragments.
HCV particles released by T cell subsets biophysically differ from those harbored in plasma used as inocula and remain infectious.To characterize the biophysical properties of HCV RNA-reactive particles released by de novo-infected CD4+ or CD8+ T cells and to compare them to those of virions carried in plasma serving as HCV inocula, culture supernatants of the infected cell subsets and the respective infectious plasma samples were ultracentrifuged over iodixanol gradients. The resulting fractions were analyzed for HCV RNA reactivity and iodixanol density. The examination of fractions of HCV-11/M plasma showed HCV RNA-reactive particles in fractions 8 and 10 to 16, corresponding to iodixanol densities of 1.138 and 1.104 to 1.017 g/cm3, respectively (Fig. 6A). Peak HCV RNA positivity was detected in fractions 8, 13, and 14 at buoyant densities of 1,138, 1.060, and 1.039 g/cm3, respectively, with corresponding relative pixel density units (PDU) of the signals of 2.8, 4.2, and 3.9, respectively (Fig. 6A). In the culture supernatant from HCV-11/M-infected CD4+ T cells collected at 10 dpi (experiment 3), HCV RNA-reactive particles were found in fractions 10 and 16 at densities of 1.104 and 1.017 g/cm3, with peak HCV RNA positivity at fraction 16 with a PDU of 1.0 (Fig. 6A). In the supernatant collected from HCV-11/M-infected CD8+ T cells obtained at 7 dpi in the same experiment, HCV RNA-positive particles were detected only in fraction 4 at a density of 1.225 g/cm3 and a PDU of 0.7 (Fig. 6A).
Buoyant density in an iodixanol gradient of HCV particles produced by in vitro-infected CD4+ and CD8+ T cells. (A) Culture supernatants collected at 10 days postinfection from CD4+ T cells and at 7 dpi from CD8+ T cells infected with HCV-11/M (experiment 3). (B) Culture supernatants collected at 14 dpi from CD4+ T cells and at 10 dpi from CD+ T cells infected with HCV-16/F (experiment 6). The cell culture supernatants and samples of HCV-11/M and HCV-16/F were concentrated by ultracentrifugation. The resulting pellets were suspended, layered over 10 to 50% iodixanol gradients, and ultracentrifuged, as described in Materials and Methods. Sixteen 300-μl fractions were collected, starting from the top of each gradient, and evaluated for HCV RNA by RT-PCR/NAH. In parallel, the same volume of AIM-V medium alone was fractionated under identical conditions, and the collected fractions were used for measuring iodixanol density (grams per cubic centimeter). The RT-PCR contamination and specificity controls were the same as those outlined in the legends to Fig. 1 and 2. The intensity of hybridization signals was measured by using a Pharos FX plus molecular imager system and is expressed a pixel density units (PDU).
Analysis of the HCV-16/F plasma inoculum showed that HCV RNA-reactive particles were present in fractions 1, 3, and 9 to 16, corresponding to iodixanol densities of 1.348, 1.252, and 1.114 to 1.017 g/cm3, respectively (Fig. 6B). Peak HCV RNA reactivities were detected in fractions 9, 13, and 14 at densities of 1.114, 1.060, and 1.039 g/cm3, respectively, with PDU of 17.0, 17.9, and 19.5, respectively. In the culture supernatant of CD4+ T cells exposed to HCV-16/F (experiment 6) and collected at 14 dpi, HCV RNA was identified in fractions 3 and 9 at densities of 1.252 and 1.114 g/cm3, respectively, with PDU of 1.0 and 1.5, respectively. With regard to the culture supernatant from CD8+ T cells obtained at 10 dpi, after exposure to the same inoculum (experiment 6), HCV RNA was detected in fractions 11 and 16 at densities of 1.088 and 1.017 g/cm3, respectively, and PDU of 0.5 and 1.8, respectively (Fig. 6B). Overall, the HCV RNA-reactive particles harbored by the HCV-11/M and HCV-16/F inocula exhibited similar buoyant density profiles, with the main peaks of HCV RNA reactivity at 1,114 to 1,138, 1,060, and 1,039 g/cm3. However, the levels of HCV RNA-positive particles released by CD4+ and CD8+ T cells infected with these inocula tended to peak at different iodixanol densities. These differences in biophysical properties between HCV particles released from infected T cell subsets and those occurring in HCV inocula were consistent with the de novo production of virus within both T cell subtypes.
To assess whether HCV released from CD4+ and CD8+ T cells remained infectious, HCV RNA-reactive iodixanol gradient fractions of culture supernatants from CD4+ and CD8+ lymphocytes infected with HCV-11/M were separately concentrated by ultracentrifugation, and the recovered virus was quantified and used to infect CD4+ or CD8+ T cells freshly isolated from a healthy donor. The results showed that the virus produced by the CD4+ or CD8+ subset was able to establish replication in the respective T cell type, as demonstrated by the detection of the virus RNA negative strand (Fig. 7).
Transmission of infection by HCV released into the culture supernatants from CD4+ or CD8+ T cells infected with authentic plasma-derived HCV. Freshly affinity-purified CD4+ or CD8+ T cells were incubated with HCV recovered by ultracentrifugation from HCV RNA-reactive fractions obtained after iodixanol gradients of culture supernatants from CD4+ or CD8+ T lymphocytes infected with HCV-11/M or HCV-16/F, as described in Materials and Methods. The expression level of the HCV RNA negative strand, indicative of progressing virus replication, was determined by strand-specific RT-PCR/NAH. sHCV RNA positive (pos) and negative (neg) strands at 1 × 104 copies/reaction served as specificity controls. Other controls are described in the legend to Fig. 5.
HCV genome variants in de novo-infected CD4+ and CD8+ T cells.To assess whether HCV replication in in vitro-infected CD4+ and CD8+ T cells led to the emergence of virus variants distinct from those occurring in the plasma serving as inocula, 5′-untranslated region (UTR) amplicons derived from plasma and PBMC of patient 11/M and from affinity-purified CD4+ and CD8+ T cells infected with HCV-11/M were cloned. Twenty clones from each amplicon were bidirectionally sequenced and compared. Clonal sequencing showed that apart from an insertion of cysteine (C) at position 126 (126insC), which was found in both the HCV-11/M inoculum (5% of clones) and HCV-11/M-infected CD4+ T cells (experiment 1) obtained at 10 dpi (5% of clones), the remaining variants identified in infected CD4+ and CD8+ cells represented unique nucleotide changes not found in the HCV-11/M inoculum (Table 4 and Fig. 8). Among 30 unique nucleotide substitutions identified in infected CD4+ and CD8+ T cells, 17 were detected in CD4+ T cells. Most variants (24 of 30) were found only in 1 out of 20 clones (5%) examined, while the percentage of clones carrying a unique variant (i.e., single nucleotide polymorphism [SNP]) was relatively high; e.g., a C-to-T change at position 115 occurred in 40% of clones (8 of 20) derived from CD8+ T cells collected at 7 dpi (experiment 3). The majority of the variants (27 of 30) identified were detected in one immune cell subtype; nonetheless, three SNPs occurred in both CD4+ and CD8+ T cells. Thus, in experiment 5, a T-to-C change at position 144 was found in both CD4+ and CD8+ T cells at 7 dpi, and an A-to-G substitution at position 252 was identified in both cell subtypes collected at 7 and 14 dpi, respectively. Furthermore, a T-to-C substitution at position 282 was detected in CD4+ T cells obtained at 10 dpi (experiment 3) and in CD8+ T cells acquired at 7 dpi (experiment 5). Finally, no variants were identified among the clones derived from the in vivo-infected PBMC of patient 11/M, as the sequences of all 20 clones were identical to the predominant sequence detected in HCV-11/M plasma. In summary, unique HCV variants were identified in de novo-infected CD4+ and CD8+ cells at all time points postinfection examined and at various frequencies. Some of the variants occurred in both CD4+ and CD8+ T cells in the same or different experiments. All of the variants, with the exception of an insertion at position 126, were unique to the de novo-infected T cells and were not found in the plasma inoculum used to infect them.
Identification of single nucleotide polymorphisms in the HCV 5′-UTR sequences detected in CD4+ or CD8+ T lymphocytes infected in vitro with authentic genotype 1a inoculum HCV-11/M
Nucleotide sequence alignment of the clones from HCV 5′-UTR fragments amplified from plasma and PBMC of the HCV-11/M inoculum donor and from CD4+ and CD8+ T lymphocytes infected in culture with this inoculum. The 5′-UTR amplicons from each sample were cloned, and 20 randomly selected clones were sequences bidirectionally. The resulting sequence reads were compared with the sequence identified in patient 11/M plasma (inoculum), shown in the top line. Representative sequences are shown, and the number of clones in which a given variant was found is depicted at the end of each sequence (B). Nucleotide positions are numbered according to the HCV genotype 1a prototype (GenBank accession number M67463). Dots indicate sequence identity with the sequence under GenBank accession number M67463, underlining indicates a gap introduced to preserve sequence alignment when an insertion was identified in one or more clones at a given position, dashes indicate deletions, and nucleotide differences are shown as letters. Single nucleotide polymorphisms found commonly between CD4+ and CD8+ T cells are marked with squares. Abbreviations: Exp., experiment; d.p.i., days postinfection; nt, nucleotide.
DISCUSSION
The ability of HCV to infect primary CD4+ and CD8+ T lymphocytes under culture conditions has been expected but not conclusively demonstrated, and the properties of the virus produced by these cells were not recognized prior to this study. For the present work, we adopted an in vitro HCV replication system in which intermittently mitogen-stimulated T cells from healthy donors served as targets for infection with authentic, patient-derived HCV (29). However, instead of the total population of primary T cells enriched from PBMC in culture, purified CD4+ and CD8+ T lymphocytes were used as targets. In this system, stimulation with PHA prior to HCV exposure and subsequent intermittent stimulation of the virus-exposed cells with the same mitogen in the presence of IL-2 have been found to be important (29, 42). As shown by data reported previously, the activation of primary T cells enhanced their susceptibility to HCV infection, while their subsequent periodic stimulation augmented the expression of HCV negative-strand and viral proteins, indicating increases in both cell receptiveness to virus and virus replication (4, 7, 29, 42, 43). The increased T cell susceptibility to HCV was consistent with the upregulation of the expression of CD5, a molecule identified as being pivotal for infection of human primary T cells and selected T cell lines with naturally occurring HCV (30, 41). Another factor influencing the infectivity of CD4+ and CD8+ cells were characteristics of HCV-positive plasma serving as the inoculum. It became apparent that high virus loads (≥1 × 106 vge/ml) in plasma originating from a treatment-naive patient combined with the ability of the virus to infect total T cells were indicators of the most desirable source of virus for infection experiments. This finding together with the high sensitivity and specificity of the molecular assays used to identify HCV RNA positive and negative strands allowed the detection and, in many instances, quantification of HCV that typically occurred at very low levels in infected T cells. It might be of importance to note that the encountered need to use plasma with a high virus load suggested that the ability to infect T cells in vitro was not a universal characteristic of all viral particles in plasma serving as inocula.
It is necessary to emphasize that the existence of HCV replication in primary CD4+ or CD8+ T cells in the present study was considered only when the HCV RNA replicative (negative) strand was detected. The finding of the virus RNA positive (vegetative) strand was used merely as an indicator of whether a given cell sample should be further investigated. This approach proved to be successful in identifying cultures in which T cells expressed viral proteins and produced virus. In addition, inhibition of HCV infection in both cell subtypes by a HCV-specific protease inhibitor, telaprevir, which requires active virus replication to exert its antiviral effect; the finding of distinctive biophysical properties of HCV RNA-reactive particles released by de novo-infected CD4+ and CD8+ T cells; and the identification of unique HCV variants arising within these cells considerably strengthened the conclusion that both T cell subtypes supported the productive replication of HCV. Overall, the data conclusively showed that both CD4+ and CD8+ T lymphocytes are susceptible to HCV infection and capable of propagating the virus in vitro.
In this context, it has been reported that HCV carried in the serum or plasma of HCV-positive patients was able to infect selected T cell lines. Accordingly, HCV in plasma was found to be infectious to Molt-4 T cells (30, 41, 44), HPB-Ma T cells (45), as well as Jurkat and prestimulated PM1 T cells (30, 41). It was also shown that HCV released from SB, a B cell line established from the splenocytes of an HCV-positive patient with type II mixed cryoglobulinemia and monocytoid lymphoma, was able to infect human primary CD4+ T cells (46) and Molt-4 and Jurkat T cells (47). As mentioned above, studies from this laboratory demonstrated that HCV occurring in plasma could infect total T cells enriched in culture by mitogen stimulation of PBMC from healthy donors, resulting in the production of infectious HCV particles displaying ultrastructural characteristics of the complete virus (7, 29). Importantly, we have also uncovered that both CD4+ and CD8+ T cells support HCV replication in patients with CH or OCI, although there were significant differences with regard to virus load and the detection of the virus RNA negative strand in both forms of infection (15). Others also reported infection of T lymphocytes in patients with CHC and OCI but without dissecting of the T cell subtype (5, 32).
When we compared the amounts of HCV identified in CD4+ or CD8+ T cells infected in vitro in the present study with those estimated for the same cell types isolated from patients with CHC or OCI by using the same detection approaches (15), it became apparent that the virus RNA positive strand occurred at the same mean levels in CD4+ T cells infected in culture (2.7 × 103 ± 1.1 × 103 vge/μg RNA; range, 30 to 1 × 104 vge/μg RNA) and those from patients with CHC (2.6 × 103 ± 1.6 × 103 vge/μg RNA; range, 10 to 1 × 104 vge/μg RNA) (P = 0.42). However, the mean level of the HCV RNA negative strand was significantly higher in CD4+ T cells infected in vitro (6.7 × 102 ± 3.8 × 102 vge/μg RNA; range, ≤25 to 5 × 103 vge/μg RNA) than in the cells circulating in CHC patients (62 ± 24 vge/μg RNA; range, 50 to 1 × 102 vge/μg RNA) (P < 0.0001). Regarding CD8+ T cells, the HCV RNA positive strand was identified at a mean level of 4.4 × 103 ± 1.3 × 103 vge/μg RNA (range, 30 to 1 × 104 vge/μg RNA) in cells infected in culture versus 1.7 × 102 ± 1.2 × 102 vge/μg RNA (range, 20 to 1 × 104 vge/μg RNA) in cells of individuals with CHC. This difference was not statistically significant (P = 0.17). In contrast, the virus replicative strand was detected at a significantly higher mean level in CD8+ T cells infected in vitro (1.2 × 102 ± 0.8 × 102 copies/μg RNA; range ≤25 to 1 × 103 copies/μg RNA) than in vivo (42 ± 30 vge/μg RNA; range, 25 to 100 vge/μg RNA) (P < 0.03). Note that the mean levels of HCV RNA positive and negative strands identified in CD4+ and CD8+ cells from patients with CHC were not meaningfully different (P = 0.25 and P = 0.41, respectively) (15). For completeness of presentation, the mean levels of HCV RNA positive and negative strands in both lymphocyte subtypes from individuals with OCI were significantly lower than those in cells from CHC patients (15) and in cells infected in culture. This comparison revealed that while infection of primary CD4+ and CD8+ cells in culture was essentially indistinguishable from that of the same cell types from patients with CHC, considering the mean levels of the virus RNA positive strand, molecular evidence of HCV replication was significantly more pronounced for the cell subtypes infected in vitro than for those infected in vivo. Certainly, there are important differences between in vitro and in vivo infection conditions, among which the usage of highly purified CD4+ or CD8+ T cells as infection targets and perhaps a relative enrichment of virus in the culture milieu could be major ones. Nonetheless, this comparison provided a valuable reciprocal validation of the authenticity of the observations made in culture and in HCV-infected patients.
The existence of active replication of HCV in the two cell types investigated in this study was further supported by the detection of virus NS5a and/or core proteins in infected cells, as identified by confocal microscopy. Infection resulted in the display of either one or both proteins in CD4+ and CD8+ T cells (Fig. 3 and 4), verifying that the virus was transcriptionally active. The estimated percentages of positive cells after infection with HCV-11/M were 0.9% for CD4+ cells and 1.2% for CD8+ cells. These values were much higher than those estimated for the respective cell types isolated from individuals with OCI after their ex vivo stimulation with PHA/IL-2, which were 0.1% and 0.02%, respectively (15). As previously reported (15), no apparent correlation was seen between the display of HCV proteins and virus RNA negative-strand expression, possibly due to a relatively low level of HCV replication. It is notable that the detection of the NS5a protein provided further evidence for ongoing HCV replication within CD4+ and CD8+ T cells, since the nonstructural proteins are not constitutive elements of HCV virions, and therefore, their detection cannot be due simply to the uptake or cell surface adhesion of circulating virions.
The ability of de novo-infected CD4+ and CD8+ T cells to support infection by HCV was further confirmed when the biophysical properties of HCV RNA-reactive particles released by the infected cells were compared to those harbored in the plasma inocula. More specifically, ultracentrifugation of the HCV-11/M and HCV-16/F plasmas resulted in the recovery of HCV RNA-positive particles throughout a wide variety of densities. Such variability was expected, as it has been extensively reported that the buoyant densities of HCV virions can vary greatly among patients (48–50). This heterogeneity can be due to several factors, including a variable degree of association with host lipoproteins and immunoglobulins (48, 50). However, we observed a different and much less heterogeneous buoyant density profile for HCV RNA-reactive particles in the supernatants of in vitro-infected CD4+ and CD8+ cells than in the infectious plasma used for their infection. Thus, most of the particles occurring in the plasma displayed buoyant densities not observed in the CD4+ and CD8+ T cell culture supernatants, while particles of other densities were more predominant in gradient fractions after ultracentrifugation of plasma. Such a difference implies that the viral particles released from infected cells exhibited different biophysical properties than those in plasma. This might be due to the possibility that circulating HCV predominantly originated from infected liver and contained a greater variety of particles interacting with different host lipids and proteins. Previous studies came to the same conclusion, although those researchers applied different fractionation procedures (7, 29). Overall, the finding that the HCV RNA-reactive particles released from de novo-infected primary CD4+ and CD8+ T lymphocytes differed in their biophysical properties from those carried in plasma used to infect these cells supports the conclusion that new HCV virions with distinctive biophysical characteristics were produced in these cells. Importantly, the virus produced by both CD4+ and CD8+ T cells remained infectious, as implied by the detection of HCV replication in de novo-infected CD4+ and CD8+ T cell subsets obtained from a healthy donor (Fig. 7).
The emergence of unique HCV variants in the sequence of the highly conserved 5′ UTR of the virus genomes detected in de novo-infected primary CD4+ and CD8+ T cells lends further support to the fact that the cells were susceptible to infection with HCV. Evolution in the virus sequence can take place only when the virus is replicating. It is notable that some of the SNPs were found in both CD4+ and CD8+ cell subtypes, while other SNPs occurred in one of the subsets. Our findings are in agreement with previously reported data where distinct variants appeared in cultured total T cells enriched from PBMC and infected with authentic HCV under the same experimental conditions as the ones in the present study (7). Furthermore, others have also reported some of the variants identified in this study. Thus, the insertion of C at nucleotide position 126 (i.e., 126insC), which we observed among the clones originating from the HCV-11/M inoculum as well as the clones derived from in vitro-infected CD4+ T cells (experiment 1) (Table 4), has also been identified in the lymph nodes and cerebellum of a patient with CHC (51). The 126insC variant has also been found in pretransplant PBMC but not in serum samples of two patients with CHC (52). In one of these patients, this mutation was identified in serum after transplant, where it was maintained for up to 2 weeks after liver transplantation, suggesting its extrahepatic origin. Furthermore, the C-to-T substitution at nucleotide position 183 (C183T), which we found in CD8+ T cell-derived clones (experiment 1) (Table 4), has also been seen in monocytes, but not in serum, of an HCV-positive patient, with the concomitant detection of the HCV RNA negative strand in these cells (50). In another study, the C183T mutation was found in PBMC but not in serum of an HCV-infected patient and in matching serum samples and PBMC from two other HCV-positive patients (53). Taken together, the variants identified in vivo in the studies mentioned above and in vitro in the present study suggest that they may reflect the evolution of the HCV sequence over the course of replication in nonhepatic cells.
It would be reasonable to expect that the replication of HCV in CD4+ and CD8+ T cells or even the exposure of the cells to HCV could affect their functions and proliferation kinetics. However, the data in this regard remain sparse. Nevertheless, it has been shown that in vitro infection of primary CD4+ T cells with a lymphotropic HCV SB strain affected the cells' IFN-γ/STAT-1/T-bet signaling pathway, leading to the inhibition of IFN-γ production (46, 54). It was also reported that infection with this virus strain suppressed the proliferation of primary CD4+ T cells and their development toward the Th1 lineage and inhibited the proliferation of Molt-4 T cells and their enhanced apoptosis (47). A study from our laboratory showed that authentic, patient-derived HCV inhibited CD4+ but not CD8+ T cell proliferation in a total T cell-HCV infection model without augmenting cell death (55). Interestingly, the results suggested that just exposure to HCV in the absence of molecularly evident replication might be sufficient to inhibit CD4+ T cell proliferation. It has also been shown that the HCV core protein upregulated the expression of anergy-related genes in Jurkat T cells stably expressing this protein (56). This was accompanied by the activation of nuclear factor of activated T cells (NFAT) and the suppression of IL-2 promoter activity.
Apart from the studies mentioned above, HCV infection of CD4+ and CD8+ T cells may directly impair HCV-specific T cell effector reactivity, which is a hallmark of CHC and a principal factor underlying the pathogenesis of this disease (57, 58). Hence, by infecting T cells, HCV may affect the efficacy of the virus-specific immune cell response, impair virus clearance, and favor its persistence. This, combined with infection of other immune cell types, including B cells and monocytes, may create an array of functional alterations further supporting HCV persistence even in the absence of liver disease, as is the case in OCI continuing after the resolution of hepatitis C (3–10). The high likelihood of a direct induction of CD4+ and CD8+ T cell immune dysfunctions by HCV, the mechanisms of this interference, and the role of impairment in the development and progression of liver and extrahepatic diseases coinciding with HCV infection strongly validate the importance of further investigations of HCV lymphotropism and its pathogenic significance.
MATERIALS AND METHODS
HCV inocula.Plasma from 16 treatment-naive patients with CHC who were free of coinfection with either HIV or hepatitis B virus (HBV), as established by clinical laboratory testing, were available for this study (Table 1). PBMC from 13 of these patients were also available for investigation. In addition, plasma samples from two HCV-naive healthy donors were used in control experiments (see below). Samples were collected after study approval by the Institutional Health Research Ethic Authority and after obtaining written informed consent.
Cells.PBMC from two healthy donors (donors A/M and B/F) provided targets for the identification of the most suitable HCV inocula and the isolation of CD4+ and CD8+ T lymphocytes for the main infection experiments. Both donors have no history or molecular evidence of HCV exposure, as their sera were negative for HCV RNA by a RT-PCR/NAH assay (sensitivity of <10 vge/ml or <3 IU/ml) (4, 10) and nonreactive for anti-HCV antibody by an enzyme immunoassay (Abbott Molecular, Mississauga, Ontario, Canada). PBMC from healthy individuals and patients with CHC were isolated from acid-citrate-dextrose-treated blood over a Ficoll-Paque Plus density gradient (Amersham Pharmacia Biotech AB, Uppsala, Sweden) immediately after blood collection, as reported previously (4, 59). The recovered cells were treated with ammonium-chlorate-potassium (ACK) (Invitrogen Life Technologies, Burlington, Ontario, Canada) to remove residual red blood cells and washed, and their numbers and viability were determined with a Countess automated cell counter (Invitrogen).
CD4+ and CD8+ T lymphocytes were affinity purified from PBMC isolated from 80 to 130 ml of donor A/M or B/F blood by applying a sequential cell sorting approach and an automatic AutoMACSPro separator (Miltenyi Biotech Inc., Auburn, CA). In the preceding studies, a protocol maximizing the sequential recovery of different immune cell subsets from the same PBMC was established (60; C. P. Corkum, S. A. MacParland, D. P. Ings, C. Simonds, G. Skardasi, C. Burgess, S. Karwowska, W. Kroll, and T. I. Michalak, unpublished data). Briefly, B cells were first removed from total PBMC using CD19 microbeads by immunoaffinity on two sequential magnetic columns, essentially according to the manufacturer's instructions (Miltenyi). The CD19-negative fraction was further used for CD8+ T cell isolation using CD8 microbeads. After positive selection of CD8+ T cells, the isolation of CD14+ monocytes from the CD19/CD8-negative fraction continued prior to the separation of CD4+ T cells on CD4 microbeads from the CD19/CD8/CD14-negative cell pool. The recovered CD8+ and CD4+ T lymphocytes were counted, their viability was measured by using an automated cell counter (Countess; Invitrogen), and the cells were immediately used as targets for infection experiments. The remaining cell subsets isolated during this protocol were cryopreserved for other ongoing studies. In preliminary experiments, CD4+ and CD8+ cells isolated by negative selection from PBMC of healthy donors according to the manufacturer's protocol (Miltenyi) were also used to compare the efficacies of HCV infection in cells purified by either positive or negative selection. There was no measurable difference in the efficiencies of HCV infection between the cell subtypes isolated by these two methods (data not shown). The purity of the resulting CD4+ and CD8+ T cells was consistently >96%, as determined by flow cytometry (see below).
HCV infection.For the initial infection experiments to determine the suitability of HCV-positive plasma samples as inocula, PBMC depleted of monocytes via adhesion to plastic were prepared as reported previously (7, 29). Monocyte-depleted PBMC as well as affinity-purified CD4+ and CD8+ T cells for the main infection experiments were plated at 3.2 × 106 to 5.0 × 106 cells/well on a 6-well plate in 5 ml of complete AIM-V medium (Invitrogen) and exposed to 5 μg/ml of PHA (ICN Biomedicals Inc., Aurora, OH) for 48 h at 37°C. PHA-pretreated cells were then supplemented with infectious plasma that was heat inactivated at 56°C for 30 min and precleared at 400 × g for 30 min or concentrated at virus/cell ratios of between 1:1 and 1:16. After 24 h of inoculation, the HCV inoculum was removed, and the cells were washed three times, counted, and cultured under alternating stimulation with 5 μg/ml PHA and/or 20 U/ml of human recombinant IL-2 (Roche Molecular Diagnostics, Laval, Quebec, Canada) for 14 dpi, as reported previously (29). More specifically, after the removal of the inoculum, cells were cultured with IL-2 for 72 h (phase A), followed by a 72-h stimulation with PHA in the presence of IL-2 (phase B). Cells were then cultured with IL-2 alone for another 72 h (phase C) and, finally, with PHA and IL-2 for 96 h (phase D). At the end of each phase, 1 × 106 cells were washed three times and returned to culture. The remaining cells collected at the end of each phase were used for the identification of HCV RNA positive and negative strands and the detection of HCV NS5A and core proteins. Cell culture supernatants were also collected at the end of each phase and stored at −80°C until analysis.
For infection of CD4+ or CD8+ T cells with HCV recovered from culture supernatants of CD4+ or CD8+ T cells infected with the HCV-11/M inoculum, HCV RNA-reactive fractions obtained after ultracentrifugation in iodixanol gradients of the supernatants from either CD4+ or CD8+ T cells were separately pooled, and the virus was concentrated over 20% sucrose cushions at 273,000 × g for 22 h at 4°C by using a TH-641 rotor and a Sorvall Discovery 100SE ultracentrifuge (Mandel Scientific Company Inc., Guelph, Ontario, Canada). The HCV content in the resulting pellets suspended in AIM-V medium was quantified by qRT-PCR (see below), and the virus was used for infection of CD4+ or CD8+ T cells freshly isolated from healthy donor A/F. The infection conditions were adjusted to accommodate low levels of the recovered virus. Thus, cells were cultured at a virus/cell ratio of ∼1:100 in the presence of PHA (5 μg/ml) and IL-2 (20 U/ml) for 72 h. They were then extensively washed, RNA was extracted, and the expression of the HCV RNA negative strand was determined (see below).
Treatment of HCV-infected CD4+ and CD8+ T cells with telaprevir.TLPV or VX-950 (Vertex Pharmaceuticals, Cambridge, MA), an HCV-specific protease inhibitor (35), is highly effective in the inhibition of HCV replication in both de novo-infected cells in culture and primary T cells and PBMC from HCV-positive patients (10, 30). A previously established protocol was applied to test whether HCV replication in CD4+ and CD8+ T cells can be suppressed by TLPV. It was previously found that TLPV concentrations of ≤4 μM in 0.5% DMSO were nontoxic to both primary and cultured T cells (30). For the present study, 2 × 105 affinity-purified CD4+ or CD8+ T cells were incubated with the HCV-11/M or HCV-16/F inoculum in the presence of 4 μM TLPV for 72 h in complete AIM-V medium supplemented with 5 μg/ml PHA and 20 U/ml IL-2. The same cells treated with 0.5% DMSO alone under the same conditions served as controls. Subsequently, the cells were harvested, and the expression of the HCV RNA negative strand was evaluated in TLPV-treated and control CD4+ and CD8+ T cells as described below.
Iodixanol gradient analysis of HCV released from lymphocytes.To examine the biophysical properties of the viral particles released from in vitro-infected CD4+ or CD8+ T cells and to compare their properties to those of the virions carried in the plasma samples serving as inocula, ultracentrifugation on an iodixanol gradient was performed (61). Thus, cell culture supernatants (10 ml) collected at different time points (phases A to D) after infection of CD4+ or CD8+ T cells, which were identified to be HCV RNA negative-strand reactive, were analyzed. More specifically, culture supernatants collected at 10 and 14 dpi for CD4+ T cells and at 7 dpi for CD8+ T cells infected with HCV-11/M (experiment 3) (Tables 2 and 3) were examined. In parallel, 200 μl of HCV-11/M plasma in 9.8 ml of phosphate-buffered saline (PBS) (pH 7.4) was fractionated. Furthermore, 10 ml of culture supernatants collected at 14 dpi for CD4+ T cells and at 10 dpi for CD8+ T cells infected with HCV-16/F was analyzed (experiment 6) (Tables 2 and 3). Again, 200 μl of HCV-16/F plasma in 9.8 ml PBS was fractionated in parallel. Prior to ultracentrifugation, all supernatants and the diluted plasma samples were preclarified at 400 × g for 30 min at 4°C after supplementation with a protease inhibitor cocktail (Sigma-Aldrich, Oakville, Ontario, Canada). They were then spun down at 150,000 × g for 22 h at 4°C in a TH-641 rotor using a Sorvall Discovery 100SE ultracentrifuge (Mandel Scientific Company Inc., Guelph, Ontario, Canada). After the removal of the supernatant, pellets normally not visible by the naked eye were resuspended in 500 μl of AIM-V culture medium without supplements. Suspensions were layered over 4.5-ml continuous 10 to 50% iodixanol gradients prepared by using OptiPrep density gradient medium (Sigma-Aldrich) and Hanks' balanced salt solution (HBSS) (Invitrogen). A 500-μl sample consisting solely of culture AIM-V medium layered over a 10 to 50% iodixanol gradient was also prepared. After ultracentrifugation at 100,000 × g for 16 h at 4°C in a Beckman SW55 Ti rotor (Beckman Coulter Inc., Pasadena, CA), 16 300-μl fractions were collected, starting from the top of each tube. The fractions recovered after ultracentrifugation of the culture medium alone were used to measure the iodixanol density with a UriSystem refractometer (Fisher Scientific International Inc., Hampton, NH).
RNA extraction and cDNA synthesis.For extraction of total RNA from cells, 1 ml of TRIzol reagent (Invitrogen) was added to ∼1 × 107 cells. For RNA isolation from 250-μl plasma or 300-μl iodixanol gradient fractions, each aliquot was supplemented with 750 μl or 700 μl of TRIzol LS (Invitrogen), respectively. The extractions were essentially done according to procedures recommended by the manufacturer. The final RNA pellets were suspended in RNase-free Tris-EDTA (TE) buffer (pH 8.0) (Ambion by Life Technologies, Carlsbad, CA), and the concentration and quality of the RNA were evaluated by spectrophotometric analyses. On average, 1 × 107 PBMC or T cells yielded between 20 and 30 μg of RNA that was aliquoted in 5-μg amounts and stored at −80°C until use. Mock samples containing UltraPure DNase/RNase-free distilled water (Invitrogen) treated with TRIzol in place of test cells, plasma, or iodixanol gradient fractions were included as contamination controls. In addition, when cell numbers allowed, affinity-purified CD4+ (experiments 1, 3, and 6) (Tables 2 and 3) and CD8+ (experiment 3) (Tables 2 and 3) T cells exposed to HDP and treated exactly the same as the cells exposed to HCV inocula were used as contamination and infection controls. RNA was transcribed to cDNA with Moloney murine leukemia virus reverse transcriptase (Invitrogen), as reported previously (4, 30).
Detection of the HCV genome and replication.Enumeration of HCV RNA copy numbers in the plasma and PBMC of CHC patients was done by qRT-PCR, as reported previously (4, 10, 30). For this purpose, 50 ng of cDNA transcribed from total cell RNA or from all RNA recovered from 250-μl plasma samples or 300-μl gradient fractions was amplified in a LightCycler 480 instrument (Roche Diagnostics). Reactions were performed in triplicates in a volume of 10 μl containing 2 μl of SoFast EvaGreen Supermix (Bio-Rad Laboratories Inc., Hercules, CA), 2 μl of cDNA at 25 ng/μl, and 5 pmol of sense (UTR4 [5′-GCAGAAAGCGTCTAGCCAT]) and antisense (RTU3 [5′-CTCGCAAGCACCCTATCAG]) primers, as reported previously (4). Enumeration of the viral load was based on 10-fold serial dilutions of recombinant HCV (rHCV) UTR-E2 (4). With each run, a mock extraction and water samples instead of test cDNA were included as specificity controls, while the synthetic HCV (sHCV) RNA positive strand at 1 × 104 to 1 × 106 copies/μl served as a positive control. The detection limit of the assay was 1 × 102 vge/reaction. In some cases, the qRT-PCR products were further tested by NAH with 32P-labeled rHCV UTR-E2 to augment the sensitivity of detection and confirm the validity of the results (4, 10, 30).
The HCV RNA positive strand was also detected by endpoint nRT-PCR/NAH using cDNA transcribed from 1 μg or, in the case of a negative result, from 3 μg of total RNA. HCV cDNA was amplified by employing direct and, if required, subsequent nested rounds of PCR. Primers used to amplify the HCV 5′ UTR, cycling conditions, and controls were previously reported (4, 15). For nested PCR, 10 μl of the direct PCR product served as the template. The rHCV UTR-E2 fragment at 1 × 104 to 1 × 106 copies/μl served as a positive control. A mock extraction and a water sample that had undergone reverse transcription were included as contamination controls, while the synthetic HCV RNA positive strand at 1 × 106 copies/μl was used as a positive control. The specificity of amplifications and the validity of controls were routinely confirmed by NAH with 32P-labeled rHCV UTR-E2 as a probe. The sensitivity of this assay was ≤10 vge/ml (≤3 IU/ml) or ≤5 vge/μg of total RNA (4, 30).
The HCV replication status was determined by the detection of the HCV RNA negative (replicative) strand using a strand-specific RT-PCR assay (62) and recombinant Thermus thermophilus DNA polymerase (Promega Corp., Madison, WI), as reported in detail previously (4, 30). More specifically, as the RNA template, 1 to 4 μg of test RNA diluted in UltraPure DNase/RNase-free distilled water (Invitrogen) was used. The amount of the template RNA tested in this assay depended on the estimated copy number of the HCV RNA positive strand. When the copy number of the HCV RNA positive strand was 10 to 100 vge/μg in the test sample, 4 μg of test RNA was used as a template. When the copy number of the HCV RNA positive strand was >100 vge/μg, 1 to 2 μg of test RNA was used for the RNA negative-strand assay. PCR signals were routinely validated by NAH (4, 30). As a specificity control, serial dilutions of the synthetic HCV RNA positive strand were used, while serial dilutions of the synthetic HCV RNA negative strand served as a sensitivity control. These standards showed that the assay detected as few as 1 × 102 copies of the negative strand per reaction and maintained the specificity of detection for up to 1 × 106 copies/reaction, as shown in Fig. 9. This was in agreement with data previously reported (4, 30). As negative controls, water instead of cDNA amplified in direct and nested reactions and mock samples extracted and treated as test cDNA were included.
Specificity of the assay for HCV RNA negative (replicative)-strand detection. As a specificity control for negative-strand detection, serial 10-fold dilutions of the synthetic HCV RNA positive strand were applied, while serial 10-fold dilutions of the synthetic HCV RNA negative strand were used as the assay sensitivity control. Using these standards, the negative-strand RT-PCR/NAH assay detected as few as 1 × 102 copies/reaction of the correct (negative) strand while nonspecifically identifying the positive strand at concentrations of ≥1 × 106 copies. Water instead of cDNA amplified in direct (D/W) and nested (N/W) reactions was included as a contamination control. Positive signals showed the expected 442-bp (direct RT-PCR) or 244-bp (nested RT-PCR) nucleotide fragments. The specificity of the RT-PCR products was verified by NAH using a 32P-labeled rHCV 5′-UTR-E2 fragment as a probe.
Flow cytometry.The purity of CD4+ and CD8+ T cell subsets was evaluated immediately following isolation from PBMC by flow cytometric analysis. For each labeling reaction, 5 × 105 cells were incubated with a mixture of peridinin chlorophyll protein (PerCP) complex-conjugated anti-human CD4, fluorescein isothiocyanate (FITC)-conjugated anti-human CD8, allophycocyanin (APC)-conjugated anti-human CD14, and phycoerythrin (PE)-conjugated anti-human CD20. Parallel incubations with appropriate isotype controls were performed by using PerCP-conjugated mouse IgG2a, FITC-conjugated mouse IgG2a, APC-conjugated mouse IgG2a, and PE-conjugated mouse IgG1. All antibodies and isotype controls were purchased from Miltenyi Biotec. Incubation took place in the dark at 4°C for 30 min. The cells were then washed with 0.25% Tween 20 (Sigma-Aldrich) in PBS, fixed in 2% paraformaldehyde, washed again, and examined with a FACSCalibur cytometer (Becton Dickinson Biosciences, Mountain View, CA). Analysis of the data was done with FlowJo v.X.0.7 software (Tree Star, Ashland, OR). Using forward versus side scatter, lymphocytes were selected. Gates for determining positivity were established by using isotype controls so that 99.0% of events were negative.
Confocal microscopy.To detect intracellular HCV NS5A and core proteins, CD4+ and CD8+ B/F cells infected with the HCV-11/M inoculum and collected at 10 and 14 dpi (experiment 5) (Tables 2 and 3) were fixed with 2% paraformaldehyde and permeabilized with 0.1% saponin at ambient temperature. Blocking of potential nonspecific binding was done with 5% normal donkey serum (Jackson ImmunoResearch Laboratories Inc., West Grove, PA) in PBS (blocking buffer) for 1 h at room temperature. The cells were then incubated with mouse anti-HCV NS5A monoclonal antibody (MAb) (Chemicon International, Temecula, CA) or with mouse anti-HCV core MAb (Fisher Scientific) in blocking buffer. As controls, cells exposed to the mouse IgG1k isotype control (eBioscience, Affymetrix, Santa Clara, CA) were used. The following day, cells were washed with PBS containing 0.25% Tween 20 and exposed to Cy3-conjugated donkey anti-mouse IgG (Jackson ImmunoResearch) in PBS containing 0.25% Tween 20 for 1 h at room temperature.
To simultaneously detect CD4 or CD8 and HCV NS5A in T cells, cells obtained at 10 and 14 dpi with HCV-16/F (experiment 6) (Tables 2 and 3) were stained first with anti-HCV NS5a MAb and then with Cy3-conjugated donkey anti-mouse antibody, as described above. In the next step, the cells were treated with 5% normal donkey serum for 1 h at room temperature and exposed to either rabbit anti-human CD4 MAb (Abcam, Cambridge, United Kingdom) in blocking buffer for 30 min on ice or the rabbit IgG polyclonal isotype control (Abcam). CD8+ T cells were stained with rabbit anti-human CD8 MAb (Abcam) in blocking buffer or exposed to the rabbit IgG polyclonal isotype control (Abcam) for 30 min on ice. After incubation, cells were washed and exposed to Alexa 488-conjugated donkey anti-rabbit IgG(H+L) (Jackson ImmunoResearch) in PBS containing 0.25% Tween 20 for 1 h in the dark at room temperature. Finally, the cells were resuspended in 20% glycerol, mounted onto slides, counterstained with 4′,6-diamidino-2-phenylindole (DAPI) (Vector Laboratories Inc., Burlingame, CA), and examined under an Olympus BX50W1 microscope with a FluoView FV300 confocal system (Olympus America Inc., Melville, NY). Approximately 1,000 cells from each preparation were examined, and cells displaying HCV protein were enumerated. The cells collected 10 and 14 days after exposure to HDP were subjected to the same treatment procedure and served as negative controls.
Cloning, sequencing, and HCV sequence analysis.HCV genome amplicons from in vitro-infected CD4+ and CD8+ T cells (experiments 1, 3, and 5) (Tables 2 and 3) at the time of detection of the HCV RNA negative strand were analyzed by clonal sequencing. Cloning of 5′-UTR HCV amplicons was done by using the TOPO TA cloning kit (Invitrogen), as reported previously (10, 15). DNA from 20 clones derived from each amplicon was sequenced bidirectionally by using universal forward and reverse M13 primers and an ABI 3730xl DNA analyzer (Applied Biosystems by Life Technologies, Foster City, CA) at The Centre for Applied Genomics (TCAG) at The Hospital for Sick Kids, Toronto, Ontario, Canada. Sequence analysis was done with the help of Sequencer software version 5.0 (Gene Codes Corp., Ann Arbor, MI). The sequences were compared with the sequence of HCV carried in the plasma inoculum used to infect the cells and, when feasible, with that of the virus harbored by PBMC of the donor of the inoculum.
Statistical analysis.Statistical analysis was done with GraphPad Prism software (GraphPad Software Inc., La Jolla, CA). The two-tailed, unpaired Student t test, with 95% confidence intervals, was used to compare the means for the groups. P values of ≤0.05 were considered to be statistically significant.
ACKNOWLEDGMENTS
We thank Patricia M. Mulrooney-Cousins, Christopher P. Corkum, and Norma D. Churchill for their excellent laboratory support and technical advice.
This study was supported by operating grant MOP-126056 from the Canadian Institutes of Health Research (CIHR) awarded to T.I.M. G.S. received a graduate student fellowship from the National CIHR Research Training Program in Hepatitis C. T.I.M. was a recipient of the Senior (Tier 1) Canada Research Chair in Viral Hepatitis/Immunology sponsored by the Canada Research Chair Program and funds from the CIHR, the Canada Foundation for Innovation, and Memorial University, St. John's, NL, Canada.
We declare that we have no competing interests.
FOOTNOTES
- Received 13 October 2017.
- Accepted 9 November 2017.
- Accepted manuscript posted online 22 November 2017.
- Copyright © 2018 American Society for Microbiology.
