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The Journal of Immunology, 2007, 179, 5387 -5398
Copyright © 2007 by The American Association of Immunologists, Inc.
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Bone Marrow of Persistently Hepatitis C Virus-Infected Individuals Accumulates Memory CD8+ T Cells Specific for Current and Historical Viral Antigens: A Study in Patients with Benign Hematological Disorders1

Vito Racanelli2, Maria Antonia Frassanito, Patrizia Leone, Claudia Brunetti, Simona Ruggieri and Franco Dammacco

Department of Internal Medicine and Clinical Oncology, University of Bari Medical School, Bari, Italy


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The role of virus-specific T cells in hepatitis C virus (HCV) pathogenesis is not clear. Existing knowledge on the frequency, phenotype, and behavior of these cells comes from analyses of blood and liver, but other lymphoid compartments that may be important sites for functionally mature T cells have not yet been analyzed. We studied HCV-specific T cells from bone marrow, in comparison to those from peripheral blood and liver biopsy tissue, from 20 persistently HCV-infected patients with benign hematological disorders. Bone marrow contained a sizeable pool of CD8+ T cells specific for epitopes from structural and nonstructural HCV proteins. These cells displayed the same effector memory phenotype as liver-derived equivalents and the same proliferative potential as blood-derived equivalents but had greater antiviral effector functions such as Ag-specific cytotoxicity and IFN-{gamma} production. These features were not shared by influenza virus-specific CD8+ T cells in the same bone marrow samples. Despite their highly differentiated phenotype and activated status, some bone marrow-resident HCV-specific CD8+ T cells were not directed against the infecting virus but, instead, against historical HCV Ags (i.e., viral species of a previous infection or minor viral species of the current infection). These findings provide a snapshot view of the distribution, differentiation, and functioning of virus-specific memory T cells in patients with persistent HCV infection.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The CD8+ T cells are essential for controlling most nonlytic virus infections. Upon Ag recognition, CD8+ T cells embark on a program of differentiation, acquisition of effector functions and expansion. CD8+ T cells differentiate from naive CD45RA+ T cells into memory CD45RA– T cells (1), which are further distinguished phenotypically and functionally into "central memory" and "effector memory" types (2, 3). Although memory cell pools are heterogeneous, their basic characteristics have been established in murine models. In particular, central memory T cells are associated with lymphoid organs, express the lymphoid homing marker CCR7, proliferate in response to Ag but may lack immediate immunological protective capacity. Effector memory T cells, instead, no longer express CCR7 and generally reside in peripheral tissues where they provide protection by a set of acquired effector functions. These effector functions permit them to kill virus-infected cells (e.g., by releasing granule contents such as perforin) and to inhibit viral replication (e.g., by producing IFN-{gamma}) (4). As a whole, the generation and functioning of these memory T cell populations depend on Ag-specific CD4+ T cell help (5, 6).

Defective effector function and overall low number of virus-specific CD8+ T cells are generally considered to be the immunological hallmarks of persistent hepatitis C virus (HCV)3 infection (7). Defects in the frequency, breadth of specificity, cytotoxicity, IFN-{gamma} production, and in vitro proliferative capacity of HCV-specific CD8+ T cells have all been reported to occur in persistently infected humans (8, 9, 10, 11, 12, 13, 14, 15, 16, 17) and chimpanzees (18). There is also evidence that HCV-specific CD8+ T cells exhibit phenotypic alterations characteristic of incomplete differentiation (19). These aspects of the immune response during persistent HCV infection have emerged from quantitative and qualitative analyses of circulating and intrahepatic virus-specific T cells, but not of cells residing in lymphoid organs such as bone marrow (BM). In adults, BM is the primary site for the development of both mature B cells and precursor T cells. It is part of the lymphocyte recirculation network (20) and serves as a major reservoir of memory T cells (21, 22). Moreover, BM is an autonomous priming site for T cell responses to blood-borne Ags (23, 24, 25). Despite these important roles in maintaining and priming T cells, HCV-specific T cells from BM have not yet been characterized for phenotype or function, especially because BM aspiration is not part of the routine diagnosis for chronic hepatitis. This implies that the overall T cell response during HCV infection has not been fully portrayed.

We were presented with a unique opportunity to investigate virus-specific T cells from BM of persistently HCV-infected patients who were referred for diagnosis of concomitant benign hematological disorders. In this report, we provide the first demonstration of HCV-specific T cells in BM and illustrate their phenotypic and functional characteristics in comparison to T cells from peripheral blood and liver biopsy tissue.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Study subjects

Between 2004 and 2006, a total of 47 patients with clinical and serological evidence of chronic liver disease and benign hematological disorders underwent percutaneous needle liver biopsy and posterior iliac crest BM aspiration. From this cohort, we recruited 20 HLA-A2+ consecutive individuals who were positive for HCV Abs and RNA, indicating persistent HCV infection, but negative for Abs to HIV and hepatitis B surface Ag (HBsAg). To establish background signals for flow cytometry, we also recruited five HLA-A2– patients with anti-HCV Abs and HCV RNA, as well as five HLA-A2+ patients who were HCV Ab and HCV RNA negative; control subjects were also negative for HIV and HBsAg Abs. No enrolled subjects had received antiviral or immunosuppressive treatments nor had acute influenza at the time of sampling.

The study protocol was approved by the University of Bari Medical School Ethics Committee and conformed to the good clinical practice guidelines of the Italian Ministry of Health and the ethical guidelines of the Declaration of Helsinki, as revised and amended in 2004. Informed consent was obtained from each patient.

Clinical and virological tests

HLA typing was performed by standard serological and molecular techniques. Serum alanine transferase levels were determined by routine kinetic enzymatic assay. Anti-HCV Abs were determined by ELISA (HCV 3.0; Ortho Clinical Systems) and recombinant-based immunoblot assay (Ortho Clinical Systems). HCV RNA in BM and peripheral blood (PB) plasma was measured by a RT-PCR assay (Amplicor HCV test; Roche Diagnostics) and expressed in log10 copies per ml. The lower limit of detection by this method is 2 log10 copies/ml. HCV genotypes were determined with a line-probe assay (Innogenetics) and classified as detailed by Simmonds et al. (26).

Synthetic peptides, HLA-A2 tetramers, recombinant proteins, and Abs

Synthetic peptides known to be HLA-A2-restricted HCV (genotype 1a), -flu and -HIV CD8+ T cell epitopes (Fig. 1B) were purchased from Mimotopes. They were chosen on the strength of their HLA-binding affinity and immunogenicity in infected subjects (27, 28). PE-conjugated HLA-A2 tetramers containing these peptides were purchased from Proimmune and used at the recommended dilutions. Recombinant HCV proteins (genotype 1a) and superoxide dismutase (SOD) protein were provided by B. Phelps (Chiron). The HCV proteins designated HCV core (c22, aa 2–120), NS3 (c33, aa 1192–1457), NS4 (c100, aa 1569–1931), and NS5 (NS5 aa 2054–2995) had been expressed as COOH-terminal fusion proteins with human SOD in yeast or Escherichia coli. Anti-human CD4 labeled with allophycocyanin as well as anti-human-CD13-allophycocyanin, -CD19-allophycocyanin, -CD8-allophycocyanin-Cy7, -CD45RA-PE-TxR, -CCR7-PE-Cy7, -perforin-FITC, and -IFN-{gamma}-FITC mAb and their isotype-matched fluorescent controls were purchased from BD Pharmingen or Caltag Laboratories and used at the recommended dilutions. HCVNS31073 and HCVNS31406 variant peptides for cytotoxicity assays were synthesized by Mimotopes.


Figure 1
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  		FIGURE 1. Frequency and memory phenotype of nonspecific and virus-specific CD8+ T cells in BM, PB, and liver of 20 persistently HCV-infected patients. Freshly purified BMMC, PBMC, and IHL were incubated with HCV peptide-bearing HLA-A2 tetramers and were surface stained with mAbs to CD4, CD8, CD13, CD19, CD45RA, and CCR7 before flow cytometry. A, Percentages of CD8+ T cells expressing CD45RA and CCR7 from BM (26,680 ± 8,510 CD8+ T cells analyzed), PB (17,020 ± 4,140 cells), and liver (17,250 ± 11,385 cells). B, Epitope specificities analyzed. The HCV epitopes were from structural and nonstructural (NS) proteins. C, Percentages of CD8+ T cells specifically binding HLA-A2 tetramers bearing five HCV epitopes. Circles connected by lines represent a single patient. Reported are the sums of virus-specific CD8+ T cell percentages detected by individual tetramers (BM and PB) and the total cell percentage detected by the tetramer pool (liver). CD8+ T cell percentages <0.02 in BM and 0.01 in PB and liver were arbitrarily considered 0 in the statistical analysis, Friedman test. D, Percentages of CD8+ T cells specific for individual HCV epitopes and flu epitope, Wilcoxon signed rank test. E, Percentages of CCR7-expressing cells among the overall population of HCV-specific memory (CD45RA–) CD8+ T cells, Kruskal-Wallis test. F, Percentages of CCR7-expressing cells among memory (CD45RA–) CD8+ T cells specific for individual HCV epitopes and flu epitope, Mann-Whitney U test. G and H, Flow cytometry plots for patient 1. Numbers within boxes indicate the percentages of cells within that gate or quadrant. FSC, Forward scatter.

 
Biological samples and cell preparations

BM and PB were collected into heparin-coated tubes and syringes. Plasma was obtained by centrifugation and stored at –80°C until use. BM and PB mononuclear cells were isolated by Ficoll-Hypaque (Pharmacia Biotech) density gradient centrifugation and were used immediately.

Liver biopsy specimens were washed three times with RPMI 1640 supplemented with 2 mmol/L L-glutamine (both from Sigma-Aldrich) and 10% heat-inactivated human AB serum (culture medium) to remove contaminating PBMC, and digested with 1 mg/ml collagenase type I and 25 µg/ml deoxyribonuclease I (both from Sigma-Aldrich) for 1 h at 37°C on a shaking device. Tissue was then disrupted by vigorous pipetting and forced through a cell strainer (Falcon). Intrahepatic lymphocytes (IHL) were isolated from the resulting cell suspension by Ficoll-Hypaque density gradient centrifugation. Typically, 1.5–5 x 105 IHL were recovered from each biopsy.

Expansion of IHL

From 2 x 104–6 x 104 IHL of four patients were cultured for 21 days and weekly stimulated with 0.05 µg/ml anti-human-CD3 mAb (Immunotech), 100 units/ml rIL-2 (PeproTech), and irradiated (50 Gy) autologous PBMC as previously described (9).

Preparation of APCs

Dendritic cells (DCs) were generated from PBMC as described elsewhere (29). Briefly, PBMC were resuspended in culture medium containing 100 U/ml penicillin and 10 µg/ml streptomycin, seeded in 250-ml culture flasks, and incubated at 37°C. After 1.5 h, the medium was aspirated to remove nonadherent cells, and adherent cells were fed with culture medium supplemented with GM-CSF (1000 units/ml) and rIL-4 (1000 units/ml; both from PeproTech). On day 7, cells were harvested and incubated overnight in culture medium containing 0 or 10 µM synthetic peptide HCVNS31073, HCVNS31406, or flu. These peptide-pulsed and unpulsed cells were then harvested, washed, gamma irradiated (50 Gy), and used as APCs.

Generation of peptide-specific CD8+ T cell lines

Nine patients with measurable frequencies of CD8+ T cells specific for HCVNS31073 or HCVNS31406 in BM or PB were selected. Peptide-specific cell lines were generated by a modified version of a previously reported method (30). Briefly, CD8+ T cells were isolated from BM and PB by magnetic cell sorting with anti-CD8 microbeads and MACS columns (Miltenyi Biotec). Cells were cocultured with autologous peptide-pulsed (and unpulsed) DCs in 96-well round-bottom plates. Each well contained 2 x 105 T cells and from 0.67 x 104–2 x 104 DCs (DC:T cell ratio between 1:10 and 1:30). Cells were grown in 200 µl of culture medium containing 100 U/ml penicillin and 10 µg/ml streptomycin for 14 days at 37°C, with the addition of 10 U/ml rIL-2 (PeproTech) on days 4 and 7 and 10 ng/ml rIL-7 (PeproTech) on day 7.

EBV transformation of B cells

To generate EBV-transformed B cell lines, 2 x 106 PBMC were infected for 2 h at 37°C with supernatant from the EBV producer line B95.8 (LGC Promochem). Transformed cells were expanded in 25- and 75-cm2 flasks by continuous passage in RPMI 1640 plus 10% FCS.

Surface staining

BM-derived mononuclear cells (BMMC; 5 x 105–5 x 106), PBMC (5 x 105–5 x 106), and IHL (half the yield from each biopsy specimen) were incubated for 20 min at 37°C with individual PE-conjugated tetramers or the PE-conjugated HCV tetramer mix in round-bottom 96-well plates. Cells were washed in PBS containing 1% human AB serum and incubated for 20 min at room temperature with a mixture of mAbs to six different surface molecules labeled with four different fluorophores.

For three patients for whom the yields of BMMC and PBMC were abundant (patients 6, 11, and 16), half of surface-stained BMMC and PBMC (those incubated with the HCV tetramer mix) were subjected to magnetic cell sorting with anti-PE microbeads and MACS columns (Miltenyi Biotec). These HCV-specific tetramer-binding cells were also used for IFN-{gamma} staining.

Perforin staining

Immediately after surface staining, BMMC, PBMC, and IHL were washed once in Stain Buffer (BD Pharmingen) and resuspended in BD Cytofix/Cytoperm solution (100 µl/well; BD Pharmingen). After incubation at 4°C for 20 min, they were washed twice with Perm/Wash solution (BD Pharmingen) and resuspended in 50 µl of Perm/Wash solution containing FITC-conjugated anti-perforin mAb. After incubation for 30 min at 4°C, cells were washed twice with Perm/Wash solution, resuspended in Stain Buffer, and analyzed by flow cytometry.

IFN-{gamma} staining

Immediately after surface staining, BMMC, PBMC, IHL, and purified HCV-specific tetramer-binding cells were assessed for IFN-{gamma} production as described earlier (31). Briefly, cells were washed and resuspended in 100 µl of culture medium containing 10 µg/ml synthetic peptide and 50 U/ml human rIL-2. In IHL- and tetramer-binding cell cultures, autologous gamma-irradiated (50 Gy) EBV-transformed B cells were included as APCs. After 2 h at 37°C, brefeldin A (Sigma-Aldrich) was added to a final concentration of 10 µg/ml and the cells were incubated for 4 h. Cells were then washed, fixed, permeabilized as described earlier, and stained with FITC-conjugated anti-IFN-{gamma} mAb. After 30 min at 4°C, cells were washed twice with Perm/Wash solution and resuspended in Stain Buffer for flow cytometry.

Proliferation assays

BMMC and PBMC (5 x 105–5 x 106) were resuspended at 1 x 107 cells/ml in PBS containing 0.1% human AB serum, and the division tracking dye CFSE (Molecular Probes) was added to a final concentration of 0.5 µM. After 10 min at 37°C, CFSE-labeled cells were washed with cold PBS containing 0.1% human AB serum, resuspended in culture medium containing 100 U/ml penicillin and 10 µg/ml streptomycin, and used immediately for CD8+ and CD4+ proliferation assays in round-bottom 96-well plates (2 x 105 cells/well).

CD8+ proliferation assay

Cells were stimulated with 10 ng/ml rIL-7, 10 pg/ml rIL-12 (PeproTech), and 10 µg/ml synthetic peptide. On day 4, we added 20 U/ml rIL-2. On day 7, cells were stained sequentially with PE-conjugated tetramer and allophycocyanin-Cy7-conjugated anti-CD8 mAb, as described earlier for surface staining, and resuspended in Stain Buffer for flow cytometry.

CD4+ proliferation assay

Cells were stimulated individually with 10 µg/ml recombinant SOD fusion proteins (HCV core, NS3, NS4, and NS5), 10 µg/ml control SOD, or 1 µg/ml PHA. After 7 days at 37°C, cells were stained with allophycocyanin-conjugated anti-CD4 mAb, as described earlier for surface staining, and resuspended in Stain Buffer for flow cytometry.

Flow cytometry and data analysis

Stained cells were analyzed without delay with a FACSCanto (BD Biosciences) flow cytometer and FACSDiva software (BD Biosciences). At least 100,000 events were acquired for each sample. Lymphocytes were identified by forward and side-angle light scatter characteristics. Flow cytometry data were analyzed with FlowJo software (Tree Star). The level of nonspecific tetramer-binding (background signal) was calculated from HCV or flu tetramer-stained BMMC, PBMC, and IHL from uninfected HLA-A2+ patients and from HCV-infected HLA-A2– patients. The cutoff for tetramer-positive signals was set at a level corresponding to the average background signal plus 3 SD. "Percent divided" was determined using the FlowJo Proliferation Platform.

HCV RNA sequencing

Ten patients with measurable frequencies of CD8+ T cells specific for HCVNS31073 or HCVNS31406 in BM or PB were selected. RNA was extracted from BM and PB plasma using the Viral RNA Mini Kit (Qiagen) and reverse transcribed using random hexamers and SuperScript III RT (Invitrogen Life Technologies). HCV cDNA regions containing the NS31073 and NS31406 epitopes were amplified using rTth DNA polymerase (Applied Biosystems) and nested sets of PCR primers (32). PCR products were subcloned by thymidine/adenosine (T/A) ligation into a DNA plasmid (Invitrogen Life Technologies), and 10 molecular clones each were sequenced in both directions using dye terminator chemistry and an automated Applied Biosystems Prism 310 Genetic Analyzer.

Cytotoxicity assay

HCVNS31073-, HCVNS31406- and flu-specific CD8+ T cell lines from 10 patients were surface stained (described earlier) and subjected to magnetic cell sorting with anti-PE microbeads and MACS columns (Miltenyi Biotec). These HCV-specific tetramer-binding cells were then used as effectors for cytotoxicity assessment according to a previously reported method (33). To prepare target cells, autologous EBV-transformed B cells were pulsed overnight with 10 µg/ml HCVNS31073 and HCVNS31406 prototype or variant and flu peptides, labeled for 1 h with 25 µCi of 51Cr (Amersham Pharmacia), and washed twice. Serial dilutions of effectors were incubated with 3 x 103 target cells in culture medium in round-bottom 96-well plates; the assay was performed in triplicate. After 6 h at 37° C, the radioactivity released into the supernatant was quantified. Percent specific lysis was calculated as 100 (experimental release – spontaneous release)/(maximum release – spontaneous release), where spontaneous release and maximum release reflect target cell lysis in the absence of effector cells and in the presence of 10% Triton X-100 (Sigma Aldrich), respectively.

Statistical analysis

Statistical analyses were performed with Prism (GraphPad software). Nonparametric statistics were used because many of the data were not normally distributed. Tests included the Friedman test, the Wilcoxon signed rank test, the Kruskal-Wallis test, and the Mann-Whitney U test for comparisons of groups. Two-tailed p values were calculated for all comparisons.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
We studied 20 HLA-A2+ patients with persistent HCV infection and concomitant benign hematological disorders (Table I). Nine patients had cryoglobulinemia and the purpura-weakness-arthralgia syndrome, seven had a faint serum monoclonal component, two had thrombocytopenia, and two had iron deficiency anemia. All had elevated serum alanine aminotransferase levels and liver histology consistent with chronic hepatitis. HCV genotypes varied and only four patients had genotype 1a, from which we chose epitopes to test in this study. HCV RNA was detected in serum of all patients and in BM of 10 patients. From all 20 patients, we isolated mononuclear cells from BM and PB and IHL from liver biopsy specimens.


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  		Table I. Demographic, clinical, and virological parameters of 20 patients with persistent HCV infection

 
Frequency and phenotype of virus-specific T cells

Six-color flow cytometry of BMMC, PBMC, and IHL labeled with mAbs to surface molecules revealed different patterns of CD8+ T cell differentiation (Fig. 1A). IHL contained predominantly effector memory (CD45RA–CCR7–) CD8+ T cells, while BMMC and PBMC contained mixed populations of naive (CD45RA+CCR7+), effector memory, and central memory (CD45RA–CCR7+) CD8+ T cells.

CD8+ T cells were characterized for their ability to bind PE-labeled HLA-A2 tetramers bearing five different HCV epitopes from structural and nonstructural proteins (Fig. 1B). BMMC and PBMC were incubated with the five tetramers individually and the percentages of tetramer-binding cells were summed for each patient; in contrast, IHL were labeled with a pool of five tetramers. Overall, there was a significant trend for highest percentages of HCV-specific (i.e., tetramer-binding) T cells in BM and lowest percentages in PB (Fig. 1C). For all 20 patients, HCV-specific CD8+ T cells represented a mean of 1.82% (SD = 1.87%) of all CD8+ T cells in BMMC, 0.85% (SD = 0.94%) in IHL, and 0.07% (SD = 0.13%) in PBMC (p < 0.0002). HCV-specific T cells were below the level of detection in BMMC for 5 patients, in PBMC for 10 patients, and in IHL for 7 patients (Table II). Detection of HCV-specific T cells in BM was independent of the presence of HCV RNA in BM and of the infecting HCV genotype.


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  		Table II. Frequency of tetramer-binding cells in the BM, PB, and liver of 20 patients with persistent HCV infection

 
CD8+ T cells from BM and PB of a single patient were unlikely to target the same epitope. For instance, in patient 1, the HCVNS31406 epitope was bound by 4.1% of CD8+ T cells in BM and by 0.01% of cells in PB, whereas the HCVNS5B2594 epitope was recognized by 0.01% of cells in PB but by no detectable fraction of cells in BM. For the 20 patients, the percentages of HCV-specific T cells in BM and PB were significantly different for all epitopes but HCVCORE132 (Fig. 1D). In contrast, BM and PB had similar percentages of CD8+ T cells that were labeled by the HLA-A2 tetramer bearing an influenza (flu) peptide (Fig. 1D, right panel). The flu epitope was recognized by a mean of 0.33% (SD = 0.41%) of CD8+ T cells in BM and 0.29% (SD = 0.46%) in PB. Overall, flu-specific CD8+ T cells were found in BM in 19 patients and in PB in 17 patients (Table II). Because the number of IHL obtained from biopsy was insufficient for the ex vivo analysis of the HCV-specific CD8+ T cell epitope repertoire in liver, IHL of four patients were expanded in vitro in an Ag-independent manner and growing polyclonal T cell lines were analyzed for the frequencies of tetramer-binding CD8+ T cells (Table III). Although the percentage values are not directly comparable to those of BM and PB, which were obtained ex vivo, this analysis showed that there was little correspondence between the epitope repertoire in liver and those in BM and PB. For example, when the percentage of tetramer-binding cells in liver was high (e.g., >0.4% of all CD8+ T cells), the corresponding values in BM and PB for the same patient tended to be low. Similarly, when the percentage of tetramer-binding cells in BM or PB was high, the corresponding value for the same patient tended to be low.


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  		Table III. Frequency of tetramer-binding cells in polyclonal intrahepatic T cell lines of four patients with persistent HCV infection

 
HCV-specific CD8+ T cells were then characterized for their surface expression of CD45RA and CCR7 to define their differentiation state. In the patients’ BMMC, a mean of 84.7% (SD = 9.9%) of CD8+ T cells binding any HCV epitope were CCR7– and negative or low positive for CD45RA. Thus, these cells displayed a typical effector memory phenotype. Similarly, 83.5% (SD = 10.4%) of HCV-specific CD8+ T cells in IHL had this effector memory phenotype. In sharp contrast, the major phenotype of HCV-specific CD8+ T cells in PBMC was CD45RA–CCR7+ (observed in 72.1 ± 9.3% of cells), characteristic of central memory cells. The frequencies of the CD45RA–CCR7+CD8+ phenotype among all HCV-specific T cells in BMMC, PBMC, and IHL differed significantly (Fig. 1E). When single epitopes were examined, the percentages of the CD45RA–CCR7+ phenotype among HCV-specific CD8+ T cells were always significantly lower in BM than in PB (Fig. 1F). Correction of the HCV-specific cell frequencies for the percentages of naive CD8+ T cells in BMMC, PBMC, and IHL did not noticeably change the patients’ profiles (Table II) nor did it alter the statistical significance of the phenomenon. Regarding flu-specific CD8+ T cells, the central memory phenotype (CD45RA–CCR7+) was slightly predominant in both BM (53.9 ± 17.0%) and PB (50.7 ± 20.0%), with a greater interpatient variability than that seen for HCV-specific cells (Fig. 1F, right panel). The distributions of CD8+ T cells into naive, effector memory, and central memory phenotypes for HCV and flu epitopes are illustrated for patient 1 in Fig. 1, G and H, respectively.

Effector functions of virus-specific T cells

HCV-specific CD8+ T cells in BMMC (15 patients), PBMC (10 patients), and IHL (13 patients) were then characterized for effector functions, specifically perforin stores and in vitro Ag-induced IFN-{gamma} synthesis, proliferation, and cytotoxicity. When five HCV epitopes were considered together, cytoplasmic perforin stores were immunologically detected in a mean of 80.6% (SD = 12.6%) of HCV-specific CD8+ T cells in BMMC (Fig. 2A). Perforin stores were found in only 25.3% (SD = 14.5%) of HCV-specific cells in PBMC, while IHL contained an intermediate value of 56.7% with greater interpatient variability (SD = 24.5%). Perforin stores in the three compartments were significantly different (p < 0.0001). When single epitopes were examined, the percentages of perforin-positive cells were always significantly higher in BM than in PB (Fig. 2B). In contrast, the frequencies of perforin-positive cells among flu-specific T cells from BM (21.5 ± 15.7%) and PB (30.1 ± 13.5%) were similar (Fig. 2B, right panel). Perforin positivity was regularly confined to the CCR7– subset of both HCV- and flu-specific CD8+ T cells in BM and liver, but in PB this association was not consistently observed, as illustrated for patient 1 in Fig. 2C.


Figure 2
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  		FIGURE 2. Functional characterization of virus-specific CD8+ T cells in BM, PB, and liver of persistently HCV-infected patients. Cell surface-stained BMMC, PBMC, and IHL were intracellularly stained for perforin ex vivo and for IFN-{gamma} after 6 h of in vitro stimulation with cognate peptides. A, Percentages of perforin-expressing cells among the overall population of CD8+ T cells specific for five HCV epitopes, Kruskal-Wallis test. B, Percentages of perforin-expressing cells among CD8+ T cells specific for individual HCV epitopes and flu epitope, Mann-Whitney U test. C, Flow cytometry plots from patient 1. Tetramer-binding cells gated as in Fig. 1, G and H, are displayed. D, Percentages of IFN-{gamma}-producing cells among the overall population of CD8+ T cells specific for five HCV epitopes, Kruskal-Wallis test. E, Percentage of IFN- {gamma}-producing cells among CD8+ T cells specific for individual HCV epitopes and flu epitope, Mann-Whitney U test. F, Flow cytometry plots from patient 1. Tetramer-binding cells gated as in Fig. 1, G and H, are displayed.

 
A second effector function examined was in vitro IFN-{gamma} production in response to short-term (6-h) stimulation with cognate peptide. IFN-{gamma} production was observed in a mean of 53.7% (SD = 16.7%) of HCV-specific CD8+ T cells from BMMC (Fig. 2D). IFN-{gamma} was produced by only 4.4% (SD = 2.9%) of HCV-specific cells from PBMC, while it was produced by 27.9% (SD = 17.4%) of cells from IHL. This pattern mirrored that seen for perforin content and was also significant (p < 0.0001). Significant differences between BM and PB were also found for HCV epitopes when examined singly (Fig. 2E). IFN-{gamma} was produced mainly by HCV-specific cells of the CCR7– phenotype (Fig. 2F). In contrast, IFN-{gamma} was produced by the majority of flu-specific cells from both BM (63.1 ± 16.6%) and PB (67.2 ± 18.9%) (Fig. 2E, right panel) and was independent of the CCR7 phenotype (Fig. 2F). Stimulation with the control HIV peptide did not induce IFN-{gamma} production (data not shown).

To rule out the possibility that the observed difference in IFN-{gamma} production between BM and PB was an experimental artifact due to the low frequency of HCV-specific CD8+ T cells in PB, we immunomagnetically purified these cells after surface staining and before IFN-{gamma} assay for three patients. Fig. 3A confirms that enrichment was successful and that cells selected from BM had predominantly a CD45RA–CCR7–CD8+ phenotype while those from PB were more often CD45RA–CCR7+CD8+. After stimulation with cognate (corresponding) peptide, cells purified from all three patients’ BM exhibited a strong right shift in fluorescence intensity compared with cells stimulated with control HIV Ag (Fig. 3B). Instead, cells purified from PB gave a small right-sided shoulder or modest right shift in fluorescence intensity, suggesting that a minor portion of HCV-specific CD8+ T cells from PB can produce IFN-{gamma}. Altogether, this experiment on single patients confirms the functional and phenotypic differences described earlier for the whole study population.


Figure 3
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  		FIGURE 3. Phenotype and IFN-{gamma} production of enriched HCV-specific CD8+ T cell populations from BM and PB of three patients. BMMC and PBMC were stained as described for Fig. 1. Tetramer-binding cells were selected by magnetic cell sorting and assayed as described for Figs. 2 and 3. Autologous EBV-transformed B cells were used as APCs. The unrelated HIVgag p17 76 peptide was used as negative control (Ctrl) Ag. A, Immunomagnetic purification and expression of memory markers; B, IFN-{gamma} production.

 
The division-tracking dye CFSE was used to quantify the percentage of HCV-specific CD8+ T cells induced to divide by 7 days of stimulation with cognate peptide. Depending on the HCV epitope, the mean percentage of BM-derived cells that had divided ranged from 18.7 to 37.6%; similar results were observed for PB-derived cells (Fig. 4A). Proliferation did not vary significantly with the HCV epitope recognized. Regarding flu-specific CD8+ T cells, on average 66.5% (SD = 21.3%) of those from BM entered division compared with 58.4% (SD = 24%) from PB; this difference was not significant (p = 0.112). The proliferative potential of HCV- and flu-specific cells was peptide specific, since it was not observed in response to stimulation with control HIV peptide (data not shown).


Figure 4
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  		FIGURE 4. Proliferation of virus-specific T cells from BM and PB of persistently HCV-infected patients. BMMC and PBMC were labeled with CFSE, stimulated in vitro with cognate peptides (CD8+ cells) or recombinant HCV proteins (CD4+ cells) for 7 days, surface stained, and analyzed by flow cytometry, Mann-Whitney U test. A, Proliferation of virus-specific CD8+ T cells. Top, Percentages of dividing cells among all cells specific for individual HCV epitopes and flu epitope. Bottom, CFSE profiles from patient 1. Sequential peaks of lower fluorescence intensity identify subsequent generations of proliferating daughter cells. B, Proliferation of virus-specific CD4+ T cells. Top, Percentages of dividing cells among all cells specific for four HCV proteins. Bottom, CFSE profiles from patient 1. CD4+ T cells proliferated in response to PHA but not to control SOD protein (data not shown).

 
Because CD4+ T cells are thought to be critical for the maintenance of CD8+ T cell functions during persistent infections, we also assessed the proliferative response of CD4+ T cells to 7-day stimulation with recombinant HCV proteins. As illustrated in Fig. 4B, a proliferative response was observed in both BM and PB, and for two Ags the BMMC response was substantial and significantly greater than that for PBMC. These results suggest that the CD8+ T cell response is more likely to be sustained by CD4+ T cells in BM.

T cell epitope specificities and viral peptide sequences

As stated earlier, the presence of HCV-specific CD8+ T cells in this series was independent of the infecting HCV genotype. In fact, patients with the highest frequencies of HCV-specific CD8+ T cells (>3% for the tetramer sum) were not infected with HCV genotype 1a (i.e., the genotype from which we selected epitopes) and thus were unexpected to have tetramer-binding cells. Among the possible interpretations of this phenomenon are the cross-reactivity with genotype 1a epitopes, exposure to previously cleared HCV strains, or current infection by minor viral species. We therefore cloned and sequenced HCV RNA from BM and PB of 10 patients who were positive for HCVNS31073- and HCVNS31406-specific T cells and deduced the corresponding amino acid sequences (Table IV). A total of eight different variants of epitope NS31073 was found in four patients, while six different variants of epitope NS31406 were observed in four patients. No patient with a variant sequence had HCV genotype 1a. Patients with infecting genotype 1b were found to have sequences exactly matching epitope NS31073 in three cases and matching epitope NS31406 in two cases. Patients with genotype 2a did not have sequences that could explain cross-reactivity. For patients who had HCV RNA in both compartments, there was general agreement in the sequences determined in BM and PB.


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  		Table IV. Sequences of HCV NS3 epitopes in 10 patients with persistent HCV infection

 
To further understand why patients infected with HCV genotypes different from 1a had tetramer-binding cells and to determine the functional significance of the variant sequences observed in some patients, we assayed their CD8+ T cells for cytotoxicity against autologous EBV-transformed B cells loaded with cognate prototype and variant peptides. CD8+ T cell lines specific for epitopes NS31073 and NS31406 were generated and immunomagnetically enriched (Fig. 5A). For patients in whom the infecting virus was conserved, BM-derived T cell lines lysed targets loaded with prototype peptide, as expected (Fig. 5B). For patients with one or more variant sequences, BM-derived T cell lines lysed targets loaded with prototype but not with variant peptides, with the exception of one cell line from patient 10 (Fig. 5D). The percent specific lysis tended to be higher in the group of patients with variants (Fig. 5D) than in those with conserved epitopes (Fig. 5B), but this difference was not significant (p = 0.1091 at the highest E:T ratio) The percent specific lysis was significantly lower in PB-derived T cell lines for the subgroup of patients with conserved epitopes and measurable HCV-specific T cells in PB (Fig. 5C; p = 0.0286 and p = 0.0121 for the comparisons (at the highest E:T ratio) with BM-derived T cell lines of patients with conserved and variant epitopes, respectively). Cytotoxicity of flu-specific CD8+ T cell lines was high and did not differ between BM and PB (Fig. 5, E and F).


Figure 5
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  		FIGURE 5. Cytotoxicity of in vitro-expanded virus-specific CD8+ T cells from BM and PB of persistently infected patients with measurable frequencies of HCVNS31073, HCVNS31406, and flu tetramer-binding CD8+ T cells. BMMC and PBMC were cocultured with peptide-loaded APCs for 14 days to generate a CD8+ T cell line. Cells were then stained with the corresponding tetramers, purified by magnetic cell sorting, and tested for cytotoxicity against 51Cr-labeled autologous EBV-transformed B cells. A, Representative flow cytometry plots before and after immunomagnetic selection of tetramer-binding CD8+ T cells expanded in vitro with prototype peptides. B and C, Cytotoxicity of BM-derived (B) and PB-derived (C) cells from patients in whom the HCVNS31073 or HCVNS31406 epitope was conserved in the infecting virus. Shown is the specific killing of B cells pulsed with the prototype peptide at different E:T ratios. D, Cytotoxicity of BM-derived cells from patients with one or more variant sequences at HCVNS31073 or HCVNS31406 epitope. Shown is specific killing of B cells pulsed with either prototype (filled circles) or corresponding variant ({circ}) peptides at different E:T ratios. Values are means of triplicate tests. E and F, Cytotoxicity of BM-derived (E) and PB-derived (F) cells toward B cells pulsed with the flu peptide at different E:T ratios. All cytotoxicity values are means of triplicate tests.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Our data provide a sharper picture of the distribution, differentiation and functioning of the virus-specific T cell population in chronic HCV infection. They show that a considerable pool of memory HCV-specific CD8+ T cells is "concealed" in the BM, thereby confirming our supposition that the full T cell response during HCV infection has not been completely portrayed. Somewhat surprisingly, BM is enriched with HCV-specific CD8+ T cells by ~25-fold compared with PB and 2-fold with respect to liver. Moreover, BM-residing HCV-specific CD8+ T cells display a highly differentiated phenotype (CCR7–) and retain effector functions that are instead entirely or partly lost by their liver- and blood-derived equivalents. The higher activation status of HCV-specific CD8+ T cells in BM stands out ex vivo regarding perforin content and, after short-term Ag stimulation, cytokine production. It is less evident but still present after sustained Ag stimulation in vitro, in terms of cytotoxic potential, yet vanishes in terms of cell proliferation rate. Such changes are restricted to those cells that target the infecting virus, because they are not evident in flu-specific CD8+ T cells (i.e., CD8+ T cells specific for a resolved viral infection). Of note, the epitope specificities of HCV-specific CD8+ T cells in BM do not overlap with those in PB. This inconsistency, which prevents a direct comparison between BM and PB CD8+ T cell responses against the same HCV epitope in the same subject, does not involve flu-specific T cells whose BM and PB frequencies are more homogeneous; this must be taken into account when comparisons between HCV- and flu-specific cells are made. The higher frequency and activation status of HCV-specific T cells in BM couples with the higher degree of CD4+ T cell proliferation. On the contrary, it does not reflect a higher viral titer at that site.

These data raise the questions as to where BM HCV-specific T cells come from and what their different epitope specificity and functional status mean. It can be supposed that they are immigrants from blood, because lymphatics do not drain this organ and T cells are normally produced in the thymus. They may migrate to BM after being primed elsewhere (lymph nodes, spleen, liver) or may first colonize and then be primed within the marrow, whose resident DCs capture, process, and present blood-borne Ags to naive T cells (24). Thus, HCV-specific central memory cells in the blood would become effector memory cells upon entry into BM. This pathway could be continuous, with memory cells entering and leaving the marrow, or a terminal differentiation event. The presence in BM of epitope-specific CD8+ T cells undetectable in blood suggests that virus-specific CD8+ T cells that enter the marrow are unable to traffic between the two compartments. Recruitment and retention of memory T cells in the BM would be Ag independent, since in this series not all patients with HCV-specific T cells in BM also had HCV RNA in this compartment. Although the exact mechanism to explain this finding is unknown, one hypothesis is that CD8+ T cells in BM are derived from a distinct cell population with differing migratory potential. Alternatively, the BM microenvironment, for instance the pattern of integrin expression, the cytokine milieu, and the nature of the APCs (34), may impact T cell trafficking. Nonetheless, once trapped in BM, HCV-specific T cells may then be regulated in a tissue-specific fashion.

Indeed, Ag-specific T cells in BM and their distinctive functional attributes have already been described in mice (3, 35, 36, 37, 38, 39) and chimpanzees (40) infected with other viruses. Persisting viruses, in particular, have been shown to dramatically modify the CD8+ T cell immunodominance and to induce a hierarchical loss of functional responses that follows different kinetics depending on the anatomical locations of the cells (37, 38, 39). In the same way, HCV persistence may lead to a progressive anatomical redistribution of virus-specific CD8+ T cells that alter their phenotype and effector functions, depending on the tissue of residence. Interestingly, these studies showed that down-regulation of IL-2 production and subsequent reduced proliferation represent the first stage of the loss of functions in all compartments. Our results are concordant because HCV-specific proliferation of CD8+ T cells in BM did not differ from that in PB and was lower than that of flu-specific CD8+ T cells in both compartments.

The BM scenario introduced by these studies of other viral infections is, however, further complicated in HCV infection. Our study documented the presence of prototype peptide-specific CD8+ T cells in BM of patients whose viruses encode a variant peptide. This finding, which is in agreement with a study of PB from persistently HCV-infected patients (41), may reflect cross-reactivity of the prototype and variant peptides, previous exposure to viral species encoding the prototype peptides (i.e., prior selection, historical genotype 1 infection), or concurrent exposure to a viral subpopulation encoding the prototype peptide not detected in the sequenced PCR clones (i.e., ongoing selection). Cytotoxicity experiments with HCV-specific CD8+ T cell lines from 10 patients indicated that all mechanisms are likely explanations, since one cell line from one patient was cytolytic to targets loaded with variant peptide while cell lines from other patients were not. Thus, despite the fact that HCV-specific CD8+ T cell populations in BM all express effector memory-associated molecules and functions which suggest that they are active antiviral cells (with immediate effector ability), they are not all involved in the immune response to the infecting virus.

In summary, in patients with persistent HCV infection, BM contains a heterogeneous population of virus-specific memory T cells. This includes T cells not involved in the current antiviral response, with specificity to earlier infections by different HCV strains or genotypes ("memory cells against historical Ags"). In addition, other BM-residing T cells are specific to the infecting HCV and thus may have an active role in the current antiviral response ("memory cells against current Ags"). This latter group includes CD8+ T cells specific to conserved viral sequences as well as to variant sequences when cross-reactive. We hypothesize that BM, along with PB (32) and liver (42), plays a key role in the maintenance of CD8+ memory T cell populations reactive to both historical and current Ags. What benefit is conferred by their homing to a compartment that is neither a primary HCV replication site nor a conventional secondary lymphoid organ? Because BM is rich with survival factors (43, 44), it could serve as a place for prolonged survival or as a depot for excess T cells generated elsewhere. For example, memory cells against historical Ags could use BM as a major "hub" where they could find transient lodging before engaging in immunosurveillance elsewhere (consistent with a recent study showing rapid turnover of CD8+ memory cells in BM of parabiotic mice) (45). Furthermore, memory cells against current Ags could use BM as a shelter or hideout to escape the immunosuppressive influence of the liver. In liver, virus-specific CCR7–CD8+ regulatory T cells producing considerable amounts of IL-10 prevent HCV-specific CD8+ T cells from fully acquiring the functional properties of effector memory cells (31). This suppression mechanism would be less efficient in BM, where viral Ag expression is low and Ag encounter by T cells is probably less frequent. These hypotheses must be tested in future experiments.

A final question arises as to the consequences of the presence of T cells in BM. Conversion of BM into a site of continuous recruitment of activated T cells would be detrimental for at least two reasons. First, antiviral CD8+ T cells are shifted from the main target of HCV infection. Second, the development of local immunopathology may be favored because CD4+ T cells helping CD8+ T cells may drive abnormal B cell proliferation and lead to clonal disorders such as cryoglobulinemia (46). Whether this may explain why many of the patients in this study have cryoglobulinemia remains to be determined.

There are certainly some caveats associated with the current study. In particular, only five HCV epitopes from one genotype were considered. Moreover, we only studied subjects with both persistent HCV infection and hematological disorders, introducing the possibility that factors other than HCV infection influenced T cell trafficking to BM. The sizes of some tetramer-positive populations were small and may have been underestimated because of viral variation (epitope mutations). Because cytokine production, apoptosis and TCR down-regulation are activation level dependent and therefore may change with the number of peptide restimulations (47), they may not precisely mirror the in vivo status of T cells. Nevertheless, this study demonstrates that during persistent HCV infection HCV-specific CD8+ T cells are also distributed to BM, it illustrates how their regulation depends upon the viral epitope recognized as well as the anatomical location, and it sheds new light on the role of BM as an archive of the immunological footprint of the evolutionary dynamics of HCV infection in individual patients.


   Acknowledgments
 
We are grateful to Dr. Bruce Phelps (Chiron) for generously providing reagents. Valerie Matarese provided scientific editing.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This study was supported by a grant from the Fondazione Cassa di Risparmio di Puglia (Bari, Italy) and the Associazione Italiana per la Ricerca sul Cancro (Milan, Italy). Back

2 Address correspondence and reprint requests to Dr. Vito Racanelli, Department of Internal Medicine and Clinical Oncology, University of Bari Medical School, Policlinico, 11 Piazza G. Cesare, Bari, Italy. E-mail address: v.racanelli{at}dimo.uniba.it Back

3 Abbreviations used in this paper: HCV, hepatitis C virus; BM, bone marrow; HBsAg, hepatitis B surface Ag; PB, peripheral blood; SOD, superoxide dismutase; BM-derived mononuclear cell; IHL, intrahepatic lymphocyte; DC, dendritic cell. Back

Received for publication May 21, 2007. Accepted for publication August 8, 2007.


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