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Virus-specific CD8+ T cells probably mediate control over HIV replication in rare individuals, termed long-term nonprogressors (LTNPs) or elite controllers. Despite extensive investigation, the mechanisms responsible for this control remain incompletely understood. We observed that HIV-specific CD8+ T cells of LTNPs persisted at higher frequencies than those of treated progressors with equally low amounts of HIV. Measured on a per-cell basis, HIV-specific CD8+ T cells of LTNPs efficiently eliminated primary autologous HIV-infected CD4+ T cells. This function required lytic granule loading of effectors and delivery of granzyme B to target cells. Defective cytotoxicity of progressor effectors could be restored after treatment with phorbol ester and calcium ionophore. These results establish an effector function and mechanism that clearly segregate with immunologic control of HIV. They also demonstrate that lytic granule contents of memory cells are a critical determinant of cytotoxicity that must be induced for maximal per-cell killing capacity.
Encouragement for the ultimate development of effective vaccines and immunotherapies that would limit HIV replication can be drawn from patients who are naturally occurring examples of immune system-mediated control. These rare individuals contain HIV for many years without antiretroviral therapy (ART) (Migueles et al., 2000). Although definitions and terms have varied, individuals within our cohort (referred to as long-term non-progressors [LTNPs] or elite controllers) have been infected for a median of 17 years yet have remained clinically well, with stable CD4+ T cell counts and viral loads of <50 copies/ml. Direct and indirect lines of evidence in humans and animal models suggest that virus-specific CD8+ T cells mediate this control, although the mechanisms by which this occurs remain unknown (reviewed in Migueles et al., 2004) (Friedrich et al., 2007; Maness et al., 2008).
Over the past 10 years, new tools have permitted an extensive characterization of the HIV-specific T cell response of LTNPs (reviewed in Migueles et al., 2004) (Bailey et al., 2006; Emu et al., 2005; Migueles et al., 2000; Navis et al., 2007; Tilton et al., 2007). The HIV-specific CD8+ T cells of LTNPs maintain greater frequencies of “polyfunctional” cells, named for their ability to degranulate and to produce several cytokines, including interleukin-2 (IL-2) (Betts et al., 2006; Zimmerli et al., 2005). However, these cells make up an extremely small subset of the total HIV-specific CD8+ T cell response, and many LTNPs demonstrate few or no such cells. HIV-specific CD8+ T cells of LTNPs maintain greater proliferative capacity than those of progressors and upregulate perforin upon stimulation (Arrode et al., 2005; Horton et al., 2006; Migueles et al., 2002). However, the relevance of differences in in vitro proliferative capacity to the in vivo situation is unclear given the similar frequencies of HIV-specific CD8+ T cells between LTNPs and untreated progressors (Migueles et al., 2002).
Cytotoxicity has been explored in some prior work but has not thus far distinguished patients with immunologic control of HIV (Cao et al., 1995; Klein et al., 1995; Pantaleo et al., 1995). Interpretation of these early studies is complicated by the lack of sensitive viral-load assays that would permit recruitment of homogeneous cohorts with clear immunologic control of HIV, by comparisons to patients with a global decline in immunity, and by the use of chromium-release-based assays that are neither highly quantitative nor highly reproducible in humans. More recently, the ability of CD8+ T cells of LTNPs to diminish HIV replication in humanized mice or in vitro was used as a measure of suppression of HIV replication (de Quiros et al., 2000; Saez-Cirion et al., 2007). However, such assays are not sufficiently powerful to establish whether the mechanism responsible for differences in function is determined by precursor frequency, precursor proliferation, preferential target or effector cell death, Fas-mediated killing, or secretion of chemokines, TNF, or suppressor factors. For these reasons, HIV-specific cytotoxicity is not being measured in most laboratories in this field or in current vaccine trials.
Furthermore, the relationship between changes in lytic granule contents after stimulation and killing by virus-specific memory CD8+ T cells has remained poorly understood. It has been an accepted paradigm for some time that memory cells maintain full cytotoxic capacity (reviewed in Seder and Ahmed, 2003; Trambas and Griffiths, 2003). Barber et al. observed that lymphocytic choriomeningitis virus (LCMV)-specific memory cells in immune mice retained the ability to kill peptide-pulsed targets in 1–4 hr, suggesting that upregulation of effector molecules is not necessary for killing by memory cells (Barber et al., 2003). On the basis of this model, re-expansion of memory cells does not involve qualitative changes in cytotoxic capacity but rather is primarily quantitative. More recently, other results have suggested that cytolytic capacity may be related to the effector molecule content of memory CD8+ T cells (Wolint et al., 2004; Meng et al., 2006; Migueles et al., 2002; Sandberg et al., 2001). However, this has been largely unexplored, in part because of the lack of reagents for staining of some lytic granule proteins in mice and the need for assays that measure killing on a per-cell basis. Thus, whether changes in lytic granule contents translate into qualitative changes in killing capacity has not been established.
In the present study, we examine the frequency and cytotoxic function of HIV-specific CD8+ T cells and their mechanism of killing autologous HIV-infected CD4+ T cells. We observed that HIV-specific CD8+ T cells of LTNPs exhibited extraordinary cytotoxic capacity on a per-cell basis against HIV-infected cells. CD8+ T cells of progressors lysed HIV-infected cells poorly, even at high true effector:target (E:T) ratios. These findings suggest that CD8+ T cell loading and delivery of cytotoxic proteins to HIV-infected CD4+ T cells is an HIV-specific effector mechanism that clearly segregates with LTNPs. Perhaps most importantly, they demonstrate that lytic granule contents of memory cells are a critical determinant of cytotoxicity that must be restimulated for maximal per-cell killing capacity.
We first examined the relationship between frequency of HIV-specific CD8+ T cells and amount of viral antigen. The frequency of HIV-specific CD8+ T cells in the peripheral blood of LTNPs is no different from that of untreated progressors (Betts et al., 2001; Gea-Banacloche et al., 2000; Migueles et al., 2002). In contrast, progressors with HIV RNA amounts suppressed to <50 copies/ml plasma by ART (Rx < 50) have considerably lower HIV-specific CD8+ T cell frequencies in the peripheral blood than do LTNPs or untreated progressors (Gray et al., 1999; Migueles et al., 2002). It has been presumed that these differences between LTNPs and Rx < 50 are due to greater virus replication in LTNPs below the detection threshold in standard assays. We examined the relationship between viral replication and CD8+ T cell frequency in these patient populations (Tables S1 and S2 available online) by using a newer assay with a lower detection limit of 1 copy/ml (Palmer et al., 2003). We observed that the median HIV-1 RNA amount of Rx < 50 was not significantly different from those of LTNPs (1 versus 2 copies/ml, respectively, p = 0.3; Figure 1A), consistent with some recent observations (Dinoso et al., 2008).
The median frequency of HIV-specific CD8+ T cells producing interferon-γ (IFN-γ) in response to autologous HIVSF162-infected CD4+ T cells was 20-fold lower in Rx < 50 than in LTNPs (0.14% versus 2.8%, respectively, p < 0.001; Figure 1B). HIV RNA was not detected in this assay in some LTNPs or Rx < 50. However, even when the analysis was limited to 19 LTNPs and 28 ART recipients with plasma viremia ≥1 copy/ml, the median frequency of HIV-specific CD8+ T cells was still significantly lower in Rx < 50 than in LTNPs (0.13% versus 3.24%, respectively, p < 0.001), despite similar amounts of plasma viral RNA (medians: 5 versus 4.7 copies/ml plasma, respectively, p > 0.5; Figures 1C and 1D). These data suggest that the HIV-specific CD8+ T cells of LTNPs persist at markedly higher frequencies than those in progressors with equally low antigen levels in vivo.
Expression of effector proteins by HIV-specific CD8+ T cells in LTNPs and untreated progressors was compared (Figure 2A). In unstimulated peripheral blood mononuclear cells (PBMCs), we and others have not detected differences in perforin or granzyme B content of HIV-specific cells between patient groups (Appay et al., 2002; Migueles et al., 2002; Sandberg et al., 2001; Zhang et al., 2003). However, upon stimulation, the HIV-specific CD8+ T cells of LTNPs proliferated and upregulated perforin, with substantial increases observed by day 3 and peak values measured by day 6 (data not shown). In the present study, we extended analyses of effector protein expression to include measurements of Ki67, granzymes A and B, and IFN-γ. In addition, the ability to transport CD107a (lysosome-associated membrane protein-1) to the cell surface was used as a marker of degranulation capacity (Betts et al., 2006). Significantly higher frequencies of HIV Gag-tetramer+ CD8+ T cells were detected in eight LTNPs compared with eight progressors after a 6 day stimulation (medians: 33.7% [17.4%–53.3%] versus 3.77% [0.4%–5.76%], respectively, p < 0.001; Figure 2A and data not shown). Only the fraction of tetramer+ cells expressing granzyme B (GrB) or perforin (medians: 87.9% [68.9%–98.4%] versus 61.5% [40.4%–85.3%], p < 0.01, and 85.6% [69.7%–96.6%] versus 46.4% [27.7%–75.4%], p = 0.004, respectively) and the mean fluorescence intensity (MFI) of perforin (medians: 3565.5 [1639–6348] versus 1268 [303–2728], p = 0.01) were significantly higher in LTNPs compared with those of progressors (Figures 2A–2C). Furthermore, GrB and perforin expression were very strongly correlated (R = 0.87, p < 0.001; data not shown). Differences in the ability of HIV-specific CD8+ T cells to degranulate were not observed between LTNPs and progressors (p > 0.5; Figures 2B and 2C). These findings support the existence of functional differences between the CD8+ T cells of LTNPs and progressors, not in the ability to degranulate but rather in the cytotoxic granule content (Meng et al., 2006; Migueles et al., 2002; Wolint et al., 2004).
We next explored whether differences in cytotoxic granule content of HIV-specific CD8+ T cells translated into differences in granule-exocytosis-mediated cytotoxicity. Traditional assays of cytotoxicity are unable to differentiate whether differences in killing are due to differences in CD8+ T cell proliferation. They are also unable to discern the mechanism of target cell killing. To determine whether the increased GrB and perforin expression in HIV-specific CD8+ T cells of LTNPs were associated with increased granule-exocytosis-mediated HIV-specific cytotoxicity on a per-cell basis, we adapted a flow-cytometry-based cytotoxicity assay that measures GrB-mediated intracellular cleavage of a cell-permeable fluorogenic substrate in live targets (Packard et al., 2007). Using this technique, we measured only functional GrB that had been delivered to the target cell, not inactive GrB stored within cytotoxic granules. Cytotoxicity mediated by PBMCs that were either rested overnight (“day 0” cells) or incubated with Gag peptides for 6 days (“day 6” cells) was assessed (Figure 3A). Target PBMCs were unpulsed or pulsed with immunodominant histocompatibility leukocyte antigen (HLA) B27- or B57-restricted Gag optimal epitope peptides. GrB-mediated substrate cleavage was associated with characteristics of early death by light scatter, as shown by the increased numbers of low forward-scatter events in the samples with greater GrB activity (Figure 3B) (Packard et al., 2007). Similar changes in light scatter and GrB substrate cleavage were previously associated with apoptosis and death of target cells on the basis of annexin-V staining, morphologic changes such as membrane blebbing, and cell death on the basis of staining with propidium iodide or 7-amino-actinomycin D (Packard et al., 2007). To precisely quantitate the actual numbers of antigen-specific CD8+ T cells present in the cultures, we stained aliquots of day 0 and day 6 cells with the appropriate HLA class I tetramers (Figure 3C). Because the responses of progressors are broader than those of LTNPs (Migueles et al., 2000), progressors were further screened with tetramers containing non-B27-restricted or non-B57-restricted subdominant epitopes (Table S3). Responses were expressed as a sum of the individual cytotoxic responses when more than one epitope was recognized.
The total cytotoxic response of day 0 cells (at a ratio of 25 total PBMC effector cells to 1 target cell) was low and not significantly different between LTNPs and progressors (medians: 3.1% versus 1.9%, respectively, p > 0.5; Figure 3D). However, at the same E:T ratio, day 6 cells of LTNPs mediated dramatically more potent cytotoxicity compared to that of progressors (medians: 67% versus 8.1%, respectively, p < 0.001; Figure 3E). Cytotoxicity was remarkably rapid and complete, with day 6 HIV-specific CD8+ T cells of LTNPs frequently killing >70% of targets during the 1 hr incubation. Examination of the outliers suggested an association among proliferation, upregulation of effector molecules, and cytotoxicity; i.e., cells from the one LTNP with a low cytotoxic response had undergone less peptide-induced expansion, and the cells of the three HLA B*27+ or 57+ progressors (patients 21, 132, and 144) with relatively high cytotoxic responses had exhibited greater proliferation after a 6 day peptide stimulation. Cytotoxic capacity on a per-cell basis was similar with day 0 cells from LTNPs or progressors over the shared range of measured E:T ratios (p = 0.27; Figure 3F). In contrast, with day 6 cells, the cytotoxic-response curves were significantly different between LTNPs and progressors (p = 0.03; Figure 3G). The potent ability of LTNP CD8+ T cells to lyse target cells was highly efficient and observed down to E:T ratios as low as 2–3:1.
A clear determination of whether the diminished cytotoxic responses of progressors relative to LTNPs were due to lower CD8+ T cell numbers or reduced per-cell cytotoxic capacity was difficult to establish at the low E:T ratios observed with peptide-pulsed targets. Therefore, we developed a system to measure CD8+ T cell-mediated cytotoxicity of autologous, acutely HIV-infected CD4+ T cells (Migueles et al., 2002; Sacha et al., 2007), permitting a sampling of a broader array of cells specific for other HIV-encoded peptides and, thereby, the total cytotoxic responses at higher E:T ratios. In addition, we used a LIVE DEAD reagent that enabled separation of three cell populations: effectors, targets, and targets that were dead prior to the 1 hr coincubation (see Experimental Procedures; Figure 4A). In this assay, CD8+ effector frequency (based upon IFN-γ secretion) and target cell GrB activity were measured in parallel (Figures 4B and 4C). After analysis for GrB activity, the fraction of targets expressing HIV p24 was quantified in the same samples by flow cytometry, and the infected CD4+ T cell elimination (ICE) was determined as another measure of cytotoxic T cell efficacy (Figure 4D). The association between GrB target cell activity and cell death in this system was verified by the observation of increased membrane permeability to propidium iodide only in the infected target cells exhibiting increased GrB activity (Figure S1). Furthermore, the responses measured by GrB activity or ICE were abrogated when CD8+ T cells were incubated with autologous uninfected or heterologous, HLA-mismatched infected targets, confirming that cytotoxicity was mediated by HIV-specific CD8+ T cells in an HLA-restricted fashion (Figure S1). The median amount of GrB or ICE activity in six uninfected controls was 1.2% and 0.3%, respectively. Thus, this combination of techniques permitted accurate measurements of HIV-specific CD8+ T cell frequency, delivery of functional granzyme B into infected lymphoblast targets, and infected target cell frequency and elimination.
With day 0 CD8+ T cells (at a ratio of 25 total CD8+ T cells to 1 target cell), the cytotoxic responses measured by GrB activity or ICE were comparable between patient groups, except for significant differences in ICE between LTNPs and Rx < 50 (p < 0.001; Figures 5A and 5B). Background GrB activity was low in response to day 0 or day 6 CD8+ T cells (4.7% and 7.4%, respectively). Although cytotoxicity measured by either method was significantly greater with day 6 cells compared with day 0 cells for each patient group, day 6 CD8+ T cells derived from LTNPs had markedly greater cytotoxic capacity than either progressors or Rx < 50 (p < 0.001 for all comparisons; Figures 5A and 5B). A very strong correlation was noted between GrB target cell activity and ICE when day 6 cells were used (R = 0.79, p < 0.001; Figure 5C). In a subset of 17 patients, perforin content was tightly correlated with GrB target cell activity and ICE (R = 0.92, p < 0.001 and R = 0.83, p < 0.001, respectively; Figures 5D and 5E). These results suggest that cytotoxicity measured by either method is highly dependent upon memory cell lytic granule loading.
We also analyzed the above data on a per-cell basis by using measured E:T ratios. With day 0 cells, target cell GrB activity was not significantly different between LTNPs and progressors (p > 0.5; Figure 6A). ICE was modestly but significantly greater in LTNPs than in progressors over the common range of E:T ratios (p = 0.03; Figure 6B). In contrast to the results with day 0 cells, GrB activity and ICE mediated by day 6 cells were significantly greater for LTNPs than progressors by a constant 18% and 40%, respectively, over the common range of E:T ratios (p < 0.001; Figures 6C and 6D). In Figure 6C, the slopes of the trend lines were not significantly different from zero. The GrB activity observed probably reached a plateau for different reasons for the two patient groups. Considering that 40%–60% of the total target cells are infected, this plateau probably occurred for LTNP-derived day 6 cells because GrB activity approached the theoretical maximum at very low measured E:T ratios (Figure 6C). However, cytotoxicity mediated by day 6 cells of progressors, measured either by GrB activity or ICE, never reached the levels observed in LTNPs, even at high E:T ratios. The minimal overlap of E:T ratios between Rx < 50 and the other groups precluded meaningful comparisons. However, the median measured E:T for Rx < 50 was 1.1:1, and at this ratio, neither GrB activity nor ICE approached the levels observed in LTNPs, but rather was similar to those of progressors. Diminished per-cell cytotoxicity in progressors compared to LTNPs was not due to death of HIV-specific CD8+ T cells, given the persistence of high frequencies of IFN-γ+ cells 6 hr later. The differences in cytotoxic responses between LTNPs and other patient groups were consistent with, and more dramatic than, those measured against peptide-pulsed targets. These data suggest that the cytotoxic capacity of HIV-specific CD8+ T cells of LTNPs is attributable not merely to increases in cell numbers, but also to qualitative changes in effector cells. Furthermore, the observation that the cytotoxic capacities of these cells are not substantially restored in patients with suppressed viremia due to ART supports the idea that diminished cytotoxicity of untreated progressors’ cells is not simply a consequence of high amounts of antigen in the chronic phase of infection. These results suggest that elimination of autologous HIV-infected CD4+ T cells mediated by the granule-exocytosis pathway segregates with immunologic control of HIV. They also demonstrate that lytic granule contents of memory cells are an important determinant of cytotoxicity that must be induced for maximal per-cell killing capacity.
Recent data have suggested that disrupted HIV-specific CD8+ T cell proliferation in progressors is associated with increased surface programmed death-1 (PD-1) (Day et al., 2006; Petrovas et al., 2006; Trautmann et al., 2006) and decreased CD127 (IL-7Rα chain) (Paiardini et al., 2005) expression. For this reason, these surface markers were examined in our cohorts. We found significant increases in the percentage of PD-1+ HIV HLA-tetramer+ cells (p = 0.02) and significant reductions in the percentage expressing CD127 (p = 0.004) in 18 viremic progressors compared with 16 LTNPs (Figure S2A). Although these markers were weakly but significantly correlated with viral-load measurements in these 34 patients (Figure S2B), PD-1 and CD127 expression in eight Rx < 50 approached the levels observed in LTNPs (p > 0.5; Figure S2A). Because immune-mediated restriction of HIV replication is not restored in Rx < 50 (Davey et al., 1999; Ortiz et al., 2001), these data suggest that the changes in PD-1 or CD127 expression on HIV-specific CD8+ T cells with therapy are an effect of viremia, consistent with some recent observations (Rehr et al., 2008; Streeck et al., 2008).
Several features of HIV-specific CD8+ T cells of progressors are consistent with some states of anergy, including diminished proliferative capacity and IL-2 production (Betts et al., 2006; Migueles et al., 2002; Zimmerli et al., 2005). A number of stimuli have been reported to overcome the anergic state (reviewed in Schwartz, 2003). Stimulation with phorbol-12-myristate-13-acetate and ionomycin (PMA+Io), followed by a period of rest prior to restimulation with HIV peptides and IL-2 (day 24 PMA+Io), produced frequencies of HIV-specific CD8+ T cells that were greater than those produced by treatment with anti-CD3 and anti-CD28, a period of rest, and restimulation with HIV peptides and IL-2 (day 24 CD3+CD28, p = 0.03; Figure S3, Figures 7A and 7B, and data not shown). To analyze whether these frequencies translated into changes in cytotoxic capacity, we performed a regression analysis of GrB activity on the log of the E:T ratio for the three sets of conditions (day 24 PMA+Io, day 24 CD3+CD28, and cells stimulated for 6 days with Gag peptides [day 6 Gag]; Figures 7C–7E). For a fixed E:T ratio of 1, there was a statistically significant difference for day 24 PMA+Io versus day 24 CD3+CD28 (51.4% versus 28.6%, p < 0.05) and for day 24 PMA+Io versus day 6 Gag (51.4% versus 19.8%, p < 0.001), but not for day 24 CD3+CD28 versus day 6 Gag (p > 0.5). Therefore, unlike cells treated with anti-CD3, anti-CD28, and IL-2, PMA+Io-treated cells mediated significantly greater cytotoxicity of peptide-pulsed targets compared with D6 Gag cells, in a manner that overlapped with activity of cells from LTNPs (Figures 7C–7E).
The recovery of proliferation of HIV-specific CD8+ T cells of progressors by PMA+Io and prior descriptions of diminished IL-2 production by these cells (Betts et al., 2006; Zimmerli et al., 2005) suggested that there may be some disruption of the calcineurin-nuclear factor of activated T cells (NFAT) pathway. Nuclear translocation of NFAT is an important early signal leading to cell division and IL-2 transcription (reviewed in Sundrud and Rao, 2007). We examined this pathway by using a technique that allows for quantitative image analysis and flow cytometry in a single platform (Figure 7F). We observed greater NFAT nuclear translocation in the HIV-specific cells of LTNPs (median: 77.2%) than those of untreated (52.3%, p = 0.05) or treated progressors (26.1%, p = 0.002; Figure 7G). Although only a limited number of specificities can be examined by this technique and there is some overlap between patient groups, these data suggest that a greater fraction of HIV-specific cells of LTNPs maintain the ability to translocate NFAT to the nucleus upon antigen encounter.
Although HIV-specific CD8+ T cells have long been suspected of mediating immunologic control in LTNPs, qualitative features of the CD8+ T cell response that clearly segregate with LTNPs and might mediate effective control of HIV have remained poorly understood. The efficient elimination of an HIV-infected CD4+ T cell via the granule-exocytosis pathway involves a series of events that include T cell activation, expansion, upregulation of effector molecules, degranulation, and delivery of lytic granule contents to the target. In the present study, we examined these functions in the HIV-specific CD8+ T cells of patients with and without immunologic control of HIV. To control for differences in proliferation or death of effector or target cells or effects not mediated by CD8+ T cells or lytic granules, we applied assays that measured effector cell frequency, infected target frequency, delivery of functional GrB into targets, and HIV-infected CD4+ T cell elimination. The HIV-specific CD8+ T cells of LTNPs were found to have extraordinary cytotoxic capacity on a per-cell basis. The potent ability of HIV-specific CD8+ T cells of LTNPs to eliminate HIV-infected CD4+ T cells was mediated by delivery of GrB to target cells. These data provide an effector function and mechanism that clearly segregate with LTNPs.
Perhaps as importantly, these results have implications that may extend to other chronic viral infections of experimental animals or humans. It has been the accepted paradigm for some time that memory cells maintain full cytotoxic capacity (reviewed in Seder and Ahmed, 2003; Trambas and Griffiths, 2003). On the basis of this model, re-expansion does not involve qualitative changes in cytotoxic capacity but rather is primarily quantitative, merely reflecting increases in the actual numbers of effector memory cells. However, a determination of whether increases in lytic granule content translate into qualitative changes has been limited by the lack of antibodies for some lytic granule proteins in mice and a lack of assays that measure cytotoxicity on a per-cell basis. In the present study, there were dramatic increases in per-cell killing capacity over 6 days associated with increases in perforin and GrB. This observation strongly suggests that restimulation of memory cells involves not only quantitative expansion, but also increases in lytic granule content that provide large qualitative increases in cytotoxic capacity.
At first inspection, it was surprising that the greatest differences in cytotoxic function between patient groups were observed after several days of exposure to autologous HIV-infected CD4+ T cells. However, the function of CD8+ T cells should probably be interpreted in the context of levels of antigen and likelihood of recent stimulation in vivo. The measurement of the CD8+ T cell response in LTNPs, treated progressors, or vaccinees is performed under conditions of low or absent antigen. Thus, an examination of such a recall response involves measurements of activation and cytokine secretion over several hours, cell-cycle progression and loading of effector molecules over several days, and target cell recognition and degranulation over 1–4 hr. Our results suggest that, under conditions of low amounts of antigen, measurements of virus-specific cell frequency and direct ex vivo cytotoxicity may not provide the best measure of the ability to restrict virus replication. Rather, proliferative capacity, upregulation of effector molecules, and target cell elimination are all important functions of the recall response of memory cells that should be considered.
This study also documents qualitative defects of the HIV-specific CD8+ T cells of progressors. We observed that the HIV-specific CD8+ T cells of Rx < 50, unlike those of LTNPs, require ongoing exposure to antigen for persistence, reminiscent of some recent findings in mice acutely or chronically infected with LCMV (Shin et al., 2007). In the present study and prior work, we did not detect defects in early events of HIV-specific CD8+ T cell activation in progressors, such as recognition of autologous virus-infected cells and cytokine production. Diminished HIV-specific cytotoxicity of viremic progressor or Rx < 50 CD8+ T cells relative to those of LTNPs was also not caused by defects in degranulation. HIV-infected CD4+ T cell elimination by progressors’ cells did not exceed 60%, even at very high measured E:T ratios. Diminished killing by progressors’ CD8+ T cells was not due to death of cytotoxic cells because they produced IFN-γ 6 hr later. Taken together, these data suggest that diminished killing capacity of HIV-specific CD8+ T cells of progressors is due to deficient loading of lytic granules and is likely to be an important mechanism accounting for the loss of immunologic control of HIV in these patients.
We also observed that considerable expansion and cytotoxic potential can be restimulated from the PBMCs of progressors, suggesting that these properties are not permanently lost by deletion, anergy, or replicative senescence. The recovery of proliferation by PMA+Io described here and the previously reported decreases in IL-2 production by HIV-specific CD8+ T cells of progressors (Betts et al., 2006; Zimmerli et al., 2005) suggested that there might be some disruption of calcium flux or the calcineurin-NFAT pathway. Nuclear translocation of NFAT is an important early event in cell proliferation and IL-2 transcription (reviewed in Sundrud and Rao, 2007). Although we observed that a significantly greater proportion of HIV-specific CD8+ T cells of LTNPs translocated NFAT upon peptide stimulation, there was considerable overlap between patient groups. It is possible that larger differences in NFAT translocation would be observed between patient groups, similar to differences observed with the ICE assay, if a larger number of specificities were sampled in assays using infected cells instead of peptide stimuli.
These results begin to address some of the cause-and-effect relationships between the cellular immune response and HIV viremia. Lowering of viral antigen levels by antiretroviral therapy is sufficient to restore HIV-specific CD4+ T cell proliferation, IL-2 production, and surface phenotype to levels observed in LTNPs (Tilton et al., 2007). It is also sufficient to restore PD-1, CD127, or IL-2 expression by HIV-specific CD8+ T cells in results presented here and some recent studies (Rehr et al., 2008; Streeck et al., 2008). Because immunologic control of HIV replication is not restored in treated-progressor cohorts, these results suggest changes in these parameters are a consequence of viremia and not a cause of the loss of immunologic control. However, we have not observed a similar restoration of HIV-specific CD8+ T cell proliferative capacity, lytic granule loading, or cytotoxic potential here or in prior work. It remains possible that one or several host or viral factors may increase peak viremia during acute infection, resulting in disruption of proliferative or cytotoxic capacity. However, this is a separate issue because the subject of the present study is to determine those functions that are disrupted during the chronic phase of infection. The results of the present study suggest that proliferative and cytotoxic capacity are disrupted functions of the HIV-specific CD8+ T cell response. A formal demonstration that cells with these functions intact cause immunologic control of lentiviral replication would require passive-transfer studies in humans or experimental animals.
Perhaps most importantly, data from the present study suggest that induction of cells able to undergo rapid expansion and mediate cytotoxicity upon antigen encounter are possible goals for HIV vaccines. Most vaccine trials and studies of chronic infection have relied heavily upon detection of HIV-specific cells by cytokine secretion in response to high concentrations of non-optimal peptides with excess costimulatory antibodies and varying degrees of intercell contact. The HIV-infected cell assay used in the present study permits measurement of the response in the context of maintained major histocompatibility complex (MHC)-peptide stoichiometry, Nef-mediated MHC downregulation, and naturally processed peptides. The findings of the present study demonstrate the qualitative features of an effective virus-specific CD8+ T cell response that should be considered in the testing of the next generation of prophylactic and therapeutic vaccines for HIV.
Subjects signed Investigational Review Board-approved informed consent documents. HIV infection was documented by HIV-1 and 2 immunoassay. LTNP criteria include the following: clinically healthy status, negative history for opportunistic diseases, stable T cell counts, set point HIV-1 RNA amounts < 50 copies/ml (bDNA assay, Bayer Diagnostics, Tarrytown, NY, USA), and no ongoing ART (Table S1). Progressors, defined as patients with a progressive decline in CD4+ T cell counts and/or current or previously documented poor restriction of virus replication (HIV-1 RNA amounts > 1000 copies/ml) when not receiving antiretroviral therapy, were divided into subgroups based on duration of infection, HIV-1 RNA set points, and treatment status (Table S2). Untreated patients were either ART naive or had been off ART for at least six months prior to leukapheresis. Treated patients received continuous ART, and patients with <50 HIV RNA copies/ml had been suppressed for a median of 5 years (range 2–11 years). HLA typing was performed by sequence-specific hybridization as described previously (Migueles et al., 2000).
In a subgroup of LTNPs and treated patients with <50 copies/ml, single-copy assays were performed as described (Palmer et al., 2003).
PBMCs were obtained as described previously (Migueles et al., 2002). CD4+ T cells were positively selected from cryopreserved PBMCs by magnetic automated cell sorting (AutoMACS, Miltenyi Biotec, Auburn, CA, USA) and polyclonally stimulated prior to infection as previously described (Migueles et al., 2002; Sacha et al., 2007).
In peptide-based assays, cryopreserved PBMC targets were resuspended at 2 × 106 cells/ml in 0.5% human serum (HAB; Gem Cell Gemini Bio-Products, Sacramento, CA, USA) RPMI medium and incubated in medium or medium containing optimal peptides (2.5–5 μM for each peptide, Multiple Peptide Systems, San Diego, CA, USA; Table S3) for 1 hr at 37°C. During the final 15 min, targets were stained with the TFL4 label (GranToxiLux, OncoImmunin, Gaithersburg, MD, USA) (Packard et al., 2007). Targets were then labeled with a LIVE DEAD Fixable Violet Stain Kit (Molecular Probes, Invitrogen Detection Technologies, Eugene, OR, USA) per the manufacturer’s instructions. Effector PBMCs, which had either been rested overnight (day 0, D#0) or stimulated with pooled HIV-1 Gag, Pol, Nef, or Env clade B 15-mer peptides (2 μg/ml of each peptide, National Institutes of Health AIDS Research and Reference Program, Bethesda, MD, USA) corresponding to the relevant optimal epitopes (day 6, D#6), were combined with targets at an E:T ratio of 25:1 at 37°C for 1 hr. The GranToxiLux killing assay was conducted per the manufacturer’s protocols (OncoImmunin) except where otherwise noted. The GrB substrate was used at a 4× dilution. Aliquots of D#0 and D#6 cells were stained with the appropriate HLA class I tetramers for determining E:T ratios on the basis of actual numbers of peptide-specific cells.
In assays using CD4+ targets infected with HIVSF162, D#0 cells and D#6 cells (coincubated with infected targets for 6 days) underwent negative selection of CD8+ T cells (CD8+ T cell Isolation Kit II, Miltenyi Biotec). Targets were labeled with the LIVE DEAD violet stain as described above. After analysis of GrB activity by flow cytometry, cells were treated with Cytofix/Cytoperm (BD Biosciences, San Jose, CA, USA) prior to staining for confirmation of infection and measurement of elimination of p24-expressing cells. The measured effector numbers were the frequencies of IFN-γ+ CD8+ T cells detected in parallel. The true target numbers were measured on the basis of the total percentages of p24+ cells. Infected CD4+ T cell elimination (ICE) was calculated as follows: ([% p24 expression of infected targets − % p24 expression of infected targets mixed with D#0 or D#6 cells] ÷ % p24 expression of infected targets) × 100.
PBMCs were resuspended in 10% HAB medium to a concentration of 4 × 106 cells/ml and stimulated with phorbol-12-myristate-13-acetate (PMA, 6.5 nM; Calbiochem, Darmstadt, Germany, USA) and ionomycin (Io, 0.2 μM; Sigma Aldrich, St. Louis, MO, USA) or CD3 antibodies (Orthoclone OKT3, 1 μg/ml; Ortho Biotech, Bridgewater, NJ, USA) and CD28 antibodies (1 μg/ml; BD Biosciences) at 37°C. At 6 hr, cells were incubated with DNase I (10 U/ml; Invitrogen, Carlsbad, CA, USA) at 37°C, washed, resuspended in 10% HAB medium without (in the case of PMA+Io stimulated) or with CD3+CD28 antibodies and plated in culture plates with 96 1-ml-deep wells (PGC Scientifics, Frederick, MD, USA) for 6, 12, or 18 days at 37°C. Unstimulated PBMCs were also plated as controls. Medium was replaced every 6 days. Pooled HIV-1 Gag peptides with or without IL-2 (2 or 20 IU/ml; Roche Diagnostics, Manheim, Germany) were added for 6 additional days prior to tetramer staining. Some wells from each condition were labeled with CFSE and analyzed for proliferation as described previously (Migueles et al., 2002).
Measurements of intracellular accumulation of lytic granule contents, degranulation, and proliferation were performed as previously described (Betts et al., 2006; Migueles et al., 2002). Surface and/or intracellular staining was done with the following antibodies: fluorescein isothiocyanate (FITC)-conjugated anti-CD3, phycoerythrin (PE)-conjugated anti-CD8, peridinine chlorophyll protein (PerCP)-conjugated anti-CD3, allophycocyanin (APC)-conjugated anti-IFN-γ or anti-CD4, Pacific Blue-conjugated anti-PD-1 and anti-GrA, PE Cy7-conjugated anti-perforin, Alexa 700-conjugated anti-GrB, and APC Cy7-conjugated anti-Ki67 (BD Biosciences). APC or PE-labeled HLA class I tetramers (Table S3), p24 antibodies (Kc57 RDI), and PE Alexa 700-conjugated anti-CD127 were purchased from Beckman Coulter (Fullerton, CA, USA). Samples were analyzed on a FACSAria multilaser cytometer (BD Biosciences) with FACSDiva software, and 50,000–2 × 106 CD3+ CD8+ lymphocyte events were collected. In cytotoxicity experiments, 5,000–8,000 target events were collected. Data were analyzed with FlowJo software (TreeStar, San Carlos, CA, USA).
PBMCs were stimulated at 2 ×106 cells per well in a total volume of 500 μl. Cells were incubated in medium alone, in medium containing optimal peptides (final concentration 2 μg/ml each), or in PMA+Io (final concentration 400 nM each). Plates were incubated at 37°C for 30 min, and then cells were transferred to V-bottom tubes and chilled on ice for 10 min. Cells were then centrifuged and stained with anti-CD8 PE-Alexa 610 (Invitrogen) and tetramer for 30 min at 4°C. Cells were then washed, fixed with 4% paraformaldehyde, and permeabilized with 500 μl of a 1:1 mix of Phosflow buffers II and III (BD Biosciences) according to the manufacturer’s protocol. The cells were then stained with Alexa 488-labeled anti-NFAT antibody (BD Biosciences), washed, and resuspended in 120 μl of PBS+BSA containing 5 μM of DRAQ-5 nuclear stain (Alexis, Lausen, Switzerland).
Cell images were collected with an Image Stream 100 (Amnis, Seattle, WA, USA), and analysis was performed as recently described (George et al., 2006). Briefly, the ratio of the NFAT integral in the nuclear mask to the NFAT integral in the cytoplasmic mask was then used to create the similarity score. The percent of tetramer+ cells that translocated on the basis of similarity score was measured on 140,000 images per condition and expressed as %M, %P, and %PMA+Io (cells incubated in medium alone, with peptide, or with PMA+Io, respectively). The percent of maximum translocation was calculated as follows: (%P − %M)/(%PMA+Io − %M) ×100 = percentage of maximum translocation.
The Wilcoxon signed-rank test was used for comparing paired data. Independent groups were compared by the Wilcoxon two-sample test. Correlation was determined by the Spearman rank method. The Bonferroni method was used for adjusting p values for multiple testing. Analysis of covariance with appropriately transformed variables was used for quantifying the difference in GrB activity and ICE of peptide-pulsed and HIV-infected CD4+ T cell targets in LTNPs and progressors over the range of E:T ratios. Linear mixed models and generalized estimating equations were used for analysis of the PMA+Io or anti-CD3+anti-CD28 reversal experiments.
Supplemental Data include three tables and three figures and can be found with this article online at http://www.immunity.com/supplemental/S1074-7613(08)00501-3.
We thank Marta Catalfamo for her thoughtful review of this manuscript. B.Z.P. and A.K. are owners of Oncoimmunin, makers of Grantoxilux. This research was supported by the Intramural Research Program of the National Institute of Allergy and Infectious Diseases, National Institutes of Health.
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