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Prepublished online as a Blood First Edition Paper on August 22, 2002; DOI 10.1182/blood-2002-03-0791.
IMMUNOBIOLOGY
From the Center for Blood Research, Department of
Pediatrics, and Department of Biostatistical Science/Dana Farber Cancer
Institute, Harvard Medical School, Boston, MA; and the Center for AIDS
Research, University Hospitals of Cleveland, Case Western Reserve
University, OH.
Despite the frequency of HIV-specific CD8 T cells, most
HIV-infected patients do not control viral replication without
antiviral drugs. Although CD8 T cells are important in containing acute HIV and simian immunodeficiency virus (SIV) infection, CD8
T-cell functions are compromised in chronic infection. To investigate whether functional deficits are specific to HIV, the phenotypic and
functional properties of HIV, Epstein-Barr virus (EBV), and cytomegalovirus (CMV)-specific CD8 T cells, labeled with HLA A2.1 or
B8 tetramers, were compared in 35 HIV-infected and 9 healthy donors.
Cytotoxic T lymphocytes express the cytolytic molecules perforin and
granzymes, and are thought to be CD45RA+CD27 Antigen-specific CD8 T cells suppress viral
replication in vitro by direct cytotoxicity and by secretion of soluble
factors.1-3 There is compelling evidence that antiviral
CD8 T cells are important in containing HIV and simian immunodeficiency
virus (SIV) replication during acute infection. Acute HIV-1 viremia
resolves with the appearance of HIV-specific cytotoxic T
lymphocytes (CTLs).4,5 Moreover, in rhesus
macaques, the elimination of CD8 T cells results in a dramatic increase
in SIV viral load.6,7 The role CD8 T cells play during
chronic infection is less clear. Cytotoxicity and interferon- CTL lysis of HIV-infected cells occurs primarily through granule
exocytosis, which requires perforin to facilitate the entry of
apoptosis-inducing serine protease granzymes into the cytosol of
infected target cells.9 However, perforin expression at the site of infection in lymphoid tissues during chronic HIV infection is rare, despite a high frequency of antigen-experienced CD8 T cells
expressing granzyme A (GzmA).10 After recent infection, perforin expression in the lymph nodes is greater, but still
substantially less, than GzmA expression.11 In addition,
though a large proportion of circulating CD8 T cells are perforin
positive (34% ± 20% in HIV-infected donors compared with
6% ± 2% in healthy donors in one study12),
circulating tetramer-stained HIV-specific CD8 T cells generally lack
perforin expression.13 Some studies have suggested that
effector CTLs in healthy donors are primarily
CD27 Because HIV infection specifically targets the critical CD4 T-helper
compartment and also infects CD4dim professional
antigen-presenting cells, including macrophages and dendritic cells,
impaired cytotoxic function by HIV-specific CD8 T cells could be a
special property of HIV infection. In fact, in a few subjects in one
study, impaired cytotoxicity and low perforin expression were more
pronounced in response to HIV than to CMV, suggesting that CD8 T-cell
dysfunction is a special feature of HIV-specific cells.13
Immune responses are tightly regulated, however, in patients with
chronic antigenic exposure to prevent autoimmune reactions to
self-antigens. This has been studied mostly for CD4 T cells, but
similar mechanisms likely regulate CD8 responses. Therefore, it would
not be surprising if CD8 T cells repeatedly exposed to antigen during
other chronic infections also have reduced cytotoxicity. To address
this question, we compared the functional and phenotypic properties of
HIV-specific CD8 T cells, identified by tetramer staining, with those
of EBV- and CMV-specific cells in 35 HIV-infected donors and 9 healthy
donors expressing HLA A2.1 or B8.
Study population
Tetramers
Flow cytometry Peripheral blood mononuclear cells (PBMCs) were resuspended in 500 µL fluorescence-activated cell sorter (FACS) blocking buffer (Hanks balanced salt solution [HBSS] with 10% human AB serum, 0.5% human immunoglobulin G [IgG], 5 mM EDTA [ethylenediaminetetraacetic acid]) for 15 minutes at 4°C and then incubated with streptavidin-PE-conjugated tetramer for an additional 40 minutes at 4°C. For external staining, the cells were washed and resuspended in FACS buffer, and aliquots of the suspension were stained with 2 µL fluorescein isothiocyanate (FITC)-conjugated mAbs to CD27 or CD45RA and Cy5-conjugated CD8 mAb or with IgG-FITC and PE and Cy5 isotype-matched controls (Immunotech). After incubation for 30 minutes at 4°C, cells were washed and resuspended in FACS buffer with 1% formaldehyde for analysis. For internal staining with GzmA mAb CB9 or perforin mAb G9 (PharMingen, San Diego, CA), tetramer-stained cells
were resuspended in 50 µL FACS buffer and were permeabilized using
the Caltag Laboratories (Burlingame, CA) Fix and Perm kit according to
the manufacturer's protocol. Fixed cells were incubated for 15 minutes
at room temperature with 2 µL respective antibodies conjugated to
FITC, washed, and resuspended in 50 µL FACS buffer. Cells were then
stained with CD8-Cy5 for 15 minutes and fixed in FACS buffer with 1%
formaldehyde. All samples were analyzed on a FACScalibur with Cell
Quest software (Becton Dickinson, San Jose, CA) on a lymphocyte-gated
population. In a subset of samples, instead of tetramer staining, PBMCs
were costained for CD8-Cy5, CD27-PE, or CD45RA-PE and were stained for
perforin-FITC or GzmA-FITC, as above.
Modulation of perforin and GzmA expression by agents that block granule exocytosis, lysosomal acidification, protein secretion, or proteolysis Peptide-specific CD8 T-cell lines were generated by incubating PBMCs from HLA A2.1- or B8-expressing healthy donors with 1 µg/mL epitopic peptide and adding recombinant human (rh) IL-2 (1-100 IU/mL) 1 to 2 days later. Cultures were fed biweekly to maintain a cell density of 5 × 105 cells/mL and were used in experiments 5 to 10 days after stimulation. Cell lines were incubated for 2 hours at 37°C with the following agents, alone or in combination, at the stated final concentration: cathepsin B inhibitors (CI), 50 µM CA074me and 50 µM z-FA-fmk (CalBiochem, San Diego, CA); proteasome inhibitors (PI), 50 µM MG-132 (z-Leu-Leu-Leu-CHO) and 50 µM PSI (z-Ile-Glu(OtBu)-Ala-Leu-CHO) (CalBiochem); 400 µM chloroquine (Sigma, St Louis, MO); 50 µg/mL Brefeldin A (Sigma); 2 mM EGTA (ethyleneglycoltetraacetic acid; Sigma); and 50 µg/mL cycloheximide (Sigma). Cells were stained and analyzed for tetramer, CD8, and perforin or GzmA as above. Modifying agents were added to the FACS blocking buffer and were maintained throughout the staining procedure.Immunomagnetic enrichment of tetramer-positive population PBMCs, stained with PE-labeled HLA-A2 or -B8 tetramers in sterile phosphate-buffered saline (PBS) with 2% fetal calf serum (FCS) for 40 minutes in the cold, were washed and incubated with -PE
Miltenyi (Miltenyi Biotech, Bergisch Gladbach, Germany) beads for
another 15 minutes. Tetramer-PE cells that bound the beads were
immunomagnetically selected on a Miltenyi column following the
manufacturer's instructions. An aliquot of selected cells was
costained with -CD8-Cy5 to ascertain the levels of enrichment. Usually, more than 100-fold enrichment of the tetramer-positive population could be obtained.
Cytotoxicity assay Log-phase autologous B lymphoblastoid cell line (BLCL) target cells were labeled with 100 µCi (3.7 MBq) chromium Cr 51 for 1 hour, washed 3 times in RPMI 1640 medium with 10% FCS, and resuspended at 105/mL as described.33 Labeled target cells (104) were added to triplicate U-bottom microtiter wells in the presence or absence of relevant antigenic peptides. Effector cells were prepared from freshly isolated, density-separated PBMCs used directly or after enrichment for tetramer-positive cells as above. After incubating target cells with peptides (1 µg/mL) for 1 hour at 37°C, effector cells suspended at indicated E/T ratios in 100 µL were added to the wells, and the plates were incubated at 37°C over CO2 for 6 hours. Supernatants (35 µL) were counted on a Top Count (Packard, Meriden, CT) microplate reader, and the percentage of specific cytotoxicity was calculated from the average cpm as [(average cpm spontaneous
release)/(total release spontaneous release) × 100]. The
spontaneous release for all experiments was less than 20%.
Peptide-specific cytotoxicity was calculated as the difference between
percentage specific cytotoxicity against peptide-loaded targets and
targets incubated with medium.
Statistical analysis Comparisons between groups were analyzed by 2-sided Wilcoxon rank sum test. Results were compared for percentage tetramer-positive cells expressing the indicated markers analyzed for each HIV tetramer and for HIV tetramer-positive cells versus EBV and/or CMV tetramer-positive cells in HIV-infected donors and for each virus between HIV-seropositive samples and healthy donor samples. Data were expressed as medians and ranges. Differences of expression in the HIV-seropositive samples were also tested for correlation with CDC stage, CD4 count (fewer than 500 cells/mm3, 500 or more cells/mm3) and for differences depending on whether plasma viremia was suppressed (fewer than 100 copies/mL, 100 or more copies/mL). Given the exploratory nature of the analysis, P values were not adjusted for multiple comparisons.
Despite persistent infection, HIV-specific tetramer-positive CD8 T cells have the phenotype of memory cells and lack perforin required for cytolysis It has been hypothesized that antigen-primed CD8 T cells segregate into a memory population that expresses the RO isoform of CD45 and is CD27+ and an effector CTL population that expresses the RA isoform of CD45 and is CD27 .14 However, this
simple phenotypic picture may not be true in patients with persistent
infection.15-18 A few studies have shown that HIV
tetramer-binding cells from most HIV-infected subjects are of the
memory subtype because they are CD27+ and
CD45RA and express high levels of
bcl-2.13,19 Other properties (GzmA+,
CD28, CCR7, and CD62L),
however, may not be typical of either naive or memory
cells.12,13,19,34 Although the tetramer-positive cells
express GzmA, they generally do not stain for perforin13
and thus may be incapable of lysing HIV-infected target cells by
granule exocytosis.9 These results were confirmed in the
present analysis of HIV tetramer-positive CD8 T cells from 35 HIV-infected subjects stained with 3 HLA A2.1 tetramers
produced with gag or RT peptides and 1 HLA B8 env tetramer (Table 2).
The subjects represent a cross-section of disease stages (Table 1), but
nearly half (16 of 35 or 46%) are stage A subjects with no history of
HIV-related symptoms. The CD4 counts of the group were also relatively
well preserved (median, 437 cells/mm3), but 7 patients had
CD4 counts lower than 200/mm3, and the range of CD4 counts
was wide (range, 38-1005/mm3). Approximately one third of
these patients were receiving highly active antiretroviral therapy
(HAART), but one third of the samples were obtained before HAART was
available, and approximately one third of the subjects were not
receiving any antiretroviral drugs because of stable disease without
drugs, drug intolerance, or individual preference. Among the latter
group were 6 long-term asymptomatic subjects (CW5, CW7, CW14, CW16,
CW18, CW20) who had been infected for more than 5 years and maintained
CD4 counts greater than 450/mm3 without antiviral drug
therapy. Approximately one third of all subjects had plasma viremia
below the levels of detection.
The frequencies of CD8 T cells reacting with each of the 3 HLA A2.1-restricted gag and RT tetramers were similar, but the HLA B8 env epitope was less commonly recognized. The mean proportion of CD8 T cells that recognized gag SLYNTVATL was 0.67% (range, 0.18%-1.85%; n = 11); that recognized RT epitope ILKEPVHGV was 0.53% (range, 0.27%-0.95%; n = 6); and that recognized RT epitope KYTAFTIPSI was 0.64% (range, 0.60%-0.81%; n = 5). Only an average of 0.09% CD8 T cells in 5 B8+ donors recognized the env epitope YLKDQQLL (range, 0.05%-0.13%). Despite the fact that most of the subjects had uncontrolled viral
production, HIV tetramer-positive cells did not have the phenotype of
effector CTLs. Although a median of 85% (range, 26%-95%) HIV
tetramer-positive cells stained for GzmA, only a median of 10% stained
for perforin. However, perforin expression in HIV-reactive cells was
heterogeneous in HIV-infected persons (range, less than 1%-70%).
Surprisingly fewer HIV tetramer-positive cells stained for perforin
than did CD8 T cells as a whole (median, 27%; range, 8%-85%).
Moreover, few HIV tetramer-positive cells had other characteristics attributed to effector CTLs: only 12% (range, 0%-23%) down-modulated CD27, and 14% (range, 0%-59%) expressed CD45RA. Representative flow
cytometry dot plots are shown in Figure
1A for a long-term asymptomatic donor.
Figure 2 and Table
3 show the phenotypic profile for all
subjects in aggregate. The properties of the tetramer-positive cells
recognizing each of the HIV epitopes were statistically indistinguishable from each other.
The phenotypic properties of the HIV tetramer-positive cells did
not correlate significantly with clinical disease parameters (Table
4). The 20 HIV tetramer-positive
samples were grouped by Centers for Disease Control (CDC)
stage, CD4 count (fewer than 500 cells/mm3 or 500 or
more cells/mm3), or plasma viral load (fewer than
100 copies/mL or 100 or more copies/mL). HIV-specific cells did not
vary significantly with any of these parameters and were mostly
CD27+CD45RA
Lack of perforin staining is not attributed to degranulation in culture or low sensitivity of detection Because of the discrepancy between staining for GzmA and perforin in HIV-specific CD8 T cells, we performed experiments to verify that low perforin staining was not caused by degranulation during freezing, thawing, or staining or by low sensitivity from a short perforin half-life within CD8 T cells. Parallel analysis of fresh samples and thawed samples gave comparable results (data not shown). Perforin is stored in acidic cytotoxic granules, which are specialized secretory lysosomes. We determined whether agents that block the acidification of the granules or that inhibit proteolysis by granule cathepsins, which process granzymes into their active form, enhance perforin protein levels in cells. The FACS buffer used for tetramer staining contains 5 mM EDTA, which chelates Ca++ and blocks perforin polymerization and granule exocytosis. Therefore, decreased perforin staining because of Ca++-dependent degranulation is unlikely. Because the composition of the Caltag fixation and permeabilization reagents are proprietary, however, we also compared staining using these reagents in the presence of an additional 2 mM EGTA (Figure 3). For these experiments, tetramer-positive cell lines were generated through the stimulation of PBMCs from an HLA B8-expressing donor with the EBV B8-restricted peptide (Table 2). Cells were preincubated for 2 hours at 37°C and were costained for the B8 EBV tetramer and for perforin and GzmA in the presence of agents that block granule exocytosis (2 mM EGTA), protein secretion (25 µg/mL Brefeldin A), lysosomal acidification (400 µM chloroquine), or proteolysis. Protease inhibitors that were tested included a cocktail of cathepsin inhibitors (z-FA-fmk and CA-074me) and a cocktail of proteosome inhibitors (MG-132 and PSI). These inhibitors were tested individually and in combination. Data from 4 experiments are shown in Figure 3. Although incubation with chloroquine, which blocks endosome and lysosome acidification, enhanced GzmA mean fluorescence intensity, it had no effect on perforin staining. In fact none of the reagents enhanced perforin mean fluorescence intensity or percentage of perforin-staining cells (not shown). Therefore, the lack of perforin detection seems unlikely to have been secondary to artifacts caused by degranulation or by rapid degradation or secretion of perforin during processing and staining.
In addition, cycloheximide, an inhibitor of protein synthesis, had little effect on perforin levels but did reduce GzmA somewhat. These results taken together suggest that GzmA may have a relatively short half-life in cells because of lysosomal degradation and that it is constantly synthesized to maintain the cellular pool. However, de novo synthesis of perforin does not occur in these cells. These preliminary results must be verified by more formal studies of perforin and GzmA protein synthesis and degradation, which are outside the scope of this study. CD8 T cells specific for EBV and CMV in HIV-infected donors and healthy donors are also mostly perforin negative To determine whether the properties of HIV-specific CD8 T cells are similar to those of CD8 T cells directed at other chronic infections, we costained EBV- and CMV-specific CD8 T cells with HLA A2.1 and HLA B8 tetramers and with perforin and GzmA in the same donors and in healthy donors. In HIV-infected patients, the proportions of EBV and CMV tetramer-positive cells that stained for perforin were similar to the staining pattern of HIV tetramer-positive cells. Most cells specific for these chronic infections are also perforin negative (Figures 1, 2). Only 14% (range, 1.4%-75%) of CMV tetramer-positive cells (n = 10) and 10% (range, 0%-76%) of EBV tetramer-positive cells (n = 23) were perforin positive (P = .92 and P = .77, respectively) compared with HIV tetramer-positive cells. These results differ from those reported by Appay et al,13 who found that perforin was preferentially down-modulated in HIV-specific CD8 T cells compared with CMV-specific CD8 T cells. As for HIV-specific cells, in an occasional donor, a high proportion of EBV or CMV tetramer-positive CD8 T cells was also perforin positive. However, donors with high proportions of perforin staining cells recognizing one virus did not have a high proportion of perforin-positive cells recognizing other viruses. These rare instances may correspond to recent infections or flares in viral production to which an effective CD8 T-cell response is generated. Plasma samples were unavailable to examine this. We also found no significant difference by 2-sided Wilcoxon rank sum test in perforin expression in EBV and CMV tetramer-positive cells in healthy donors compared with HIV-infected donors: 13% (0%-43%) of EBV-specific cells (n = 7) and 13% (range, 6%-48%) of CMV-specific cells (n = 4) from healthy donors were perforin positive (P = .87 and P = .68, respectively, compared with HIV-specific cells) (Figure 2; Table 3).EBV and CMV tetramer-positive cells in HIV-infected donors are more
often GzmA+, CD27 In HIV-infected donors, EBV- and CMV-specific cells were more uniformly GzmA+ than were HIV-specific cells (Figure 2; Table 3). GzmA was expressed by 85% (range, 26%-95%) of HIV tetramer-positive cells compared with 90% (range, 70%-100%) of EBV tetramer-positive cells and 96% (range, 78%-99%) of CMV tetramer-positive cells in HIV-infected donors (P = .005 and P = .002, respectively). GzmA expression in EBV- and CMV-specific cells from healthy donors was indistinguishable from that in EBV- and CMV-specific cells in HIV-infected donors (P = .60 and P = .50, respectively), with most of these circulating cells expressing GzmA. Although 88% (range, 77%-100%) of HIV tetramer-positive cells express CD27, 80% (range, 21%-100%) of EBV tetramer-positive cells and 35% (range, 19%-68%) of CMV tetramer-positive cells express CD27 in HIV-infected donors (P = .02 and P < .001, respectively, compared with HIV). The difference in CD27 expression between EBV- and CMV-specific cells in the HIV-infected donors was also highly significant (P < .001). Similarly, though few HIV-specific cells express CD45RA (14%; range, 0%-59%), in the same infected donors more EBV-specific cells (32%; range, 0%-79%, P = .006) and still more CMV-specific cells (54%; range, 14%-86%; P = .001) express this marker. Again, the difference between EBV- and CMV-specific cell expression of CD45RA was significant (P = .030). By CD27 expression, EBV- and CMV-specific CD8 T cells in healthy donors
looked more like memory cells than they did in HIV-infected donors.
Although 91% (range, 70%-100%) of healthy donor EBV
tetramer-positive cells are CD27+, only 80% (range,
21%-100%) of HIV-seropositive donor EBV-specific cells are
CD27+ (P = .050). Similarly, for CMV
tetramer-positive cells, 76% (range, 61%-86%) of healthy donor and
35% (range, 19%-68%) of HIV-infected donor cells expressed CD27
(P = .021). However, CD45RA expression was statistically
indistinguishable on EBV- and CMV-specific cells in healthy and
HIV-infected donors (P = .31 and P = .13,
respectively). The lack of clear differences in CD45RA expression may
reflect the fact that CD45RA does not correlate as well as CD27
down-modulation with CTL effector status (see "Most
perforin+ CD8 T cells are also CD27 Most perforin+ CD8 T cells are also
CD27  a median of
65% of perforin+ cells is CD45RA+ in healthy
donor or HIV-seropositive samples. CD27 down-modulation correlated
better with perforin expression than did CD45RA, but neither was a
perfect indicator of potential CTL function in HIV-infected donors.
Neither marker was useful in identifying the GzmA+ subset
of CD8 T cells.
Perforin expression is required for cytotoxicity by freshly tested virus-specific cells in healthy donors and HIV-infected donors To verify that the low levels of perforin expression in CMV and EBV tetramer-positive cells signified impaired cytotoxicity, we tested cytotoxicity by using freshly isolated healthy donor (n = 3) and HIV-seropositive donor (n = 4) PBMCs against autologous BLCLs incubated with EBV or CMV peptides (Tables 5, 6). Although cell lines in which the frequency of perforin expression on tetramer-positive cells was greater than 25% demonstrated peptide-specific cytotoxicity (range, 12%-23%) above background at E/T ratios of 100:1 to 200:1, PBMCs with low numbers of perforin+ tetramer-positive cells did not. However, EBV-specific cells from one donor (CW16) did not have cytotoxic activity above background, even though the frequency of tetramer-positive cells was reasonably high (0.76%) and most tetramer-positive cells stained for perforin.
Because the frequency of tetramer-positive cells was low (0.05%-4.1%
of total PBMCs), we also verified these results by testing cytotoxicity
after immunomagnetic enrichment of tetramer-positive cells in 5 samples
(Tables 5, 6). Representative flow cytometry plots and
cytotoxicity assays for one sample with perforin-staining tetramer
cells and one sample without are shown in Figure
5. After enrichment, 16% to 80% of the
cells were tetramer positive. Results for total PBMCs were verified in
the samples enriched for tetramer-positive cells tested at lower E/T
ratios. Only samples with at least 25% of tetramer cells expressing
perforin were capable of significant levels of antigen-specific
cytotoxicity. The sample from CW16 was again unable to lyse specific
targets. Therefore, as expected, perforin is required for cytotoxic
function. Results for sample CW16, however, suggest that perforin
staining may not be sufficient for cytotoxicity.
In this study most cells specific for the chronic viruses HIV, EBV, and CMV in nearly all HIV-infected donors and healthy donors do not express high levels of perforin, the key determinant of immediate cytotoxic function. The low frequency of high perforin expression by chronic virus-specific CD8 T cells is especially striking because approximately one quarter of CD8 T cells are perforin positive in HIV-infected patients across the disease spectrum.12 We investigated whether low perforin expression might be an artifact of protein secretion or cytotoxic granule exocytosis during cell processing and staining or whether high protein turnover caused by proteolytic degradation might account for the low levels of perforin staining. We did not find any of these to be the case. Further, ex vivo cytotoxicity correlates with high perforin expression. When more than 25% of tetramer-positive cells express perforin, specific cytotoxicity is readily detectable; when the frequency is lower, as it is in most samples, specific cytotoxicity is not much above background, even when assays are performed on populations enriched for tetramer reactivity. Thus, low perforin expression by EBV- and CMV-specific CD8 T cells from most healthy and HIV-seropositive persons suggests that, as has been shown for HIV-specific CD8 T cells, the cytotoxic function of EBV- and CMV-specific cells tested directly without in vitro culture is also limited in many donors. These results imply that in patients with chronic antigen exposure, CD8 T-cell cytotoxicity, and possibly other functions, is tightly regulated so that most cells that have seen antigen repeatedly are not cytotoxic. This may protect from damage that could arise by CTL recognition and destruction of uninfected host cells displaying weakly reacting self-peptides. Therefore, lack of specific cytotoxicity is not peculiar to HIV-specific cells and is not restricted to HIV-infected donors. Our results differ from those of Appay et al,13 who found that HIV-specific CD8 T cells were specifically impaired in perforin expression and cytotoxic function compared with CMV-specific cells. However, a more recent paper17 by this group, published after this submission, suggests that perforin expression among CD8 T cells responding to 4 persistent infections (EBV, CMV, HCV, and HIV) may not be that dissimilar. Apparent differences between studies might be explained by the inclusion of a few patients with recent infection, which might be missed because the primary infection with these viruses can be subclinical. This might explain the occasional donor with a high frequency of perforin-staining antiviral cells. Future studies of viral antibody status may determine whether this is the case. Alternatively, occasional patients with exceptional control of viral production may respond to a burst in viral replication, as they do to a cleared infection. Understanding the reasons for this wide variation in perforin staining will be important for understanding what regulates cytotoxic function. These results suggest that the best chance for controlling an infection
is early, presumably when CD8 T cells highly express perforin
and can eliminate infected cells. During that time, CD8 T cells
contribute substantially to controlling the total body viral burden, as
has been demonstrated conclusively in SIV-infected macaques.6,7 Our data support the notion that later,
during the chronic phase, their ability to control viral production is more limited. This is also supported by a study of perforin expression in lymphoid tissues of HIV-infected patients in which the proportion of
CD8 T cells expressing perforin within 4 to 5 months of symptomatic primary HIV infection was significantly higher than in chronic infection (0.3%-1.5% of all lymphoid cells in recent infection vs
mean of less than 0.1% in chronic infection).10 In
another study in macaques infected with pathogenic SIV, most SIV
tetramer-positive cells produced IFN- The low frequency of perforin expression is unlikely to be secondary to recent degranulation because one would then expect both perforin and GzmA to be depleted. However, GzmA expression is high. In fact our studies of perforin and GzmA staining after blocking protein synthesis suggest that GzmA is continuously synthesized and degraded in CTLs. Nonetheless, it remains possible that perforin expression might be so tightly regulated, in a manner analogous to cytokines,40 that constitutive expression is low and cells require antigenic stimulation for it to be readily detectable. When interpreting these data, it is also important to bear in mind that, at least in murine models, low levels of perforin expression are adequate for CTL cytotoxicity.41 Although in this study lack of cytotoxicity correlates with low expression of perforin, sufficient perforin for cytotoxic function may still be expressed in tetramer-positive cells but may be below the sensitivity level of flow cytometry detection. In that case, the lack of cytotoxic function in perforin low cells might be caused by other factors, such as defective signaling of cytotoxicity by the T cell.20,42 Because EBV- and CMV-specific CD8 T cells in healthy donors also
generally lack perforin and are not cytotoxic, CD4 T-cell depletion may
not be the underlying reason for CD8 T-cell dysfunction during chronic
HIV infection. However, this does not mean that CD4 T cells are not
important. Although CD4 cell proliferative responses to CMV can be
measured in many chronically infected healthy donors, many
antigen-specific CD4 T cells may still be partially anergized. Many
CMV-reactive CD4 cells in HIV infection are activated to produce
IFN- CD4 lymphoproliferative responses to HIV p24 (which correlate with IL-2 production) have been suggested to be associated with improved disease prognosis primarily because they lead to an increase in CD8 T-cell function.47-50 Although this is an attractive hypothesis, data presented here suggest that this must be examined more closely. Because most HAART-treated patients and healthy donors have lymphoproliferative responses to CMV antigens, one would anticipate that their CMV-specific CD8 T cells would have higher levels of perforin. However, this is not the case. Moreover, the CD8 T cells from the long-term asymptomatic HIV-infected subjects in this study did not have higher proportions of perforin expression (data not shown). Looking at perforin expression and CTL function in long-term nonprogressors (LTNPs) and patients with CD4 proliferative responses to p24 merits further study. Perforin expression is the defining feature of terminally
differentiated effector CD8 T cells. Although other phenotypic changes, such as CD27 down-modulation and expression of GzmA and CD45RA, may
often accompany differentiation into effector CTLs, our
findings here suggest that antigenic stimulation conditions for
different viral infections may alter the differentiation program. EBV-
and CMV-specific CD8 T cells in HIV-infected donors are more likely than HIV-specific cells to be CD27 In this study, CD27 down-modulation was the phenotypic change that correlated most closely with perforin expression. However, in HIV-infected donors, one fifth of perforin-positive cells were CD27+. CD45RA expression was not a good indicator of perforin expression in healthy donors or HIV-seropositive donors. Phenotypic properties of effector CTLs in healthy donors or in patients with cleared infection cannot be extrapolated to the more complex state of persistent infection. Differences in expression of CD8 differentiation molecules in cells responding to different viruses or between HIV-infected and healthy donors (Figure 2; Table 3) may also reflect differences in the likelihood of recent encounters with cells actively replicating virus. For the markers we studied, the EBV-specific cells in HIV-infected donors are intermediate in activation phenotype between the HIV- and CMV-tetramer populations. This suggests that CMV replication may be more active than EBV replication in HIV-infected donors. In healthy donors, most EBV- and CMV-specific cells have a memory phenotype, suggesting that they have not recently encountered antigen. In HIV-infected donors, higher proportions of EBV- and CMV-specific cells have down-modulated CD27, and there is also a trend toward more expression of GzmA and CD45RA. Therefore, the cells specific for these viruses appear to have been more recently activated in HIV-infected patients than in healthy donors. This is probably because they have more recently encountered antigen, as is implied by higher frequencies of cells infected with these herpesviruses in immunosuppressed persons.51,52 The decrease in CD27 down-modulation and CD45RA re-expression on HIV tetramer-positive CD8 T cells compared with EBV- and CMV-specific cells, even in patients with uncontrolled viral replication, could indicate that HIV antigen presentation or T-cell recognition may be particularly impaired in HIV infection.13 HIV-specific CD8 T cells may look like memory cells because they have not recently recognized, or been stimulated by, an infected cell. This may be because of viral mutation or nef-mediated down-modulation of major histocompatibility complex class 1 or because of other unknown viral effects on antigen presentation or of impaired signaling by the T cell itself.20,42,53,54
Submitted March 13, 2002; accepted August 7, 2002.
Prepublished online as Blood First Edition Paper, August 22, 2002; DOI 10.1182/blood-2002-03-0791.
Supported by National Institutes of Health (NIH) grants AI-42519 and AI-45406 (J.L.); and the NIH-funded Dana Farber Cancer Institute-Beth Israel Deaconess Medical Center-Children's Hospital Center for AIDS Research (AI28691) and Case Western Reserve University Center for AIDS Research (AI36219).
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Judy Lieberman, Center for Blood Research, Harvard Medical School, 800 Huntington Ave, Boston, MA 02115; e-mail: lieberman{at}cbr.med.harvard.edu.
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A. Iannello, O. Debbeche, S. Samarani, and A. Ahmad Antiviral NK cell responses in HIV infection: II. viral strategies for evasion and lessons for immunotherapy and vaccination J. Leukoc. Biol., July 1, 2008; 84(1): 27 - 49. [Abstract] [Full Text] [PDF]
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B. E. Palmer, E. Boritz, and C. C. Wilson Effects of Sustained HIV-1 Plasma Viremia on HIV-1 Gag-Specific CD4+ T Cell Maturation and Function J. Immunol., March 1, 2004; 172(5): 3337 - 3347. [Abstract] [Full Text] [PDF]
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H. Katano, M. A. Ali, A. C. Patera, M. Catalfamo, E. S. Jaffe, H. Kimura, J. K. Dale, S. E. Straus, and J. I. Cohen Chronic active Epstein-Barr virus infection associated with mutations in perforin that impair its maturation Blood, February 15, 2004; 103(4): 1244 - 1252. [Abstract] [Full Text] [PDF]
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R. Draenert, C. L. Verrill, Y. Tang, T. M. Allen, A. G. Wurcel, M. Boczanowski, A. Lechner, A. Y. Kim, T. Suscovich, N. V. Brown, et al. Persistent Recognition of Autologous Virus by High-Avidity CD8 T Cells in Chronic, Progressive Human Immunodeficiency Virus Type 1 Infection J. Virol., January 15, 2004; 78(2): 630 - 641. [Abstract] [Full Text] [PDF]
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X. Wang, D. M. Hone, A. Haddad, M. T. Shata, and D. W. Pascual M Cell DNA Vaccination for CTL Immunity to HIV J. Immunol., November 1, 2003; 171(9): 4717 - 4725. [Abstract] [Full Text] [PDF]
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Top
Abstract
Introduction
(IFN-
) compared with control cells
(
).