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. 2002 Dec;130(3):509–517. doi: 10.1046/j.1365-2249.2002.02005.x

Comparison between HIV- and CMV-specific T cell responses in long-term HIV infected donors

L PAPAGNO *, V APPAY *, J SUTTON *, T ROSTRON *, G M A GILLESPIE *, G S OGG *, A KING *, A T MAKADZANHGE *, A WATERS †, C BALOTTA ‡, A VYAKARNAM †, P J EASTERBROOK †, S L ROWLAND-JONES *
PMCID: PMC1906546  PMID: 12452843

Abstract

The mechanisms underlying non-progression in HIV-1 infection are not well understood; however, this state has been associated previously with strong HIV-1-specific CD8+ T cell responses and the preservation of proliferative CD4+ T cell responses to HIV-1 antigens. Using a combination of interferon-gamma (IFN-γ) ELISpot assays and tetramer staining, the HIV-1-specific CD8+ T cell populations were quantified and characterized in untreated long-term HIV-1-infected non-progressors and individuals with slowly progressive disease, both in relation to CD4+ T cell responses, and in comparison with responses to cytomegalovirus (CMV) antigens. High levels of CD8+ T cell responses specific for HIV-1 or CMV were observed, but neither their frequency nor their phenotype seemed to differ between the two patient groups. Moreover, while CMV-specific CD4+ T cell responses were preserved in these donors, IFN-γ release by HIV-1-specific CD4+ T cells was generally low. These data raise questions with regard to the role played by CD8+ T cells in the establishment and maintenance of long-term non-progression.

Keywords: cytotoxic T lymphocytes, HIV-1, T cell help, T cell maturation, long-term non-progressor

Introduction

The recent development of new technologies for the study of antigen-specific T cells has highlighted the importance of T lymphocytes in the control and the eradication of viral pathogens, and in particular HIV-1 [1]. More than any other virus, human immunodeficiency virus (HIV-1) represents a challenge to our immune defences, which are eroded progressively during HIV-1 infection. The presence of HIV-1-specific cytotoxic T lymphocytes (CTL) has been well described in subjects infected with HIV-1, and high levels of CTL were one of the first features to be documented in the small number of HIV-1-infected individuals who meet the definition of long-term non-progressors (LTNPs) [2]. However, despite the large numbers of HIV-specific CTL in asymptomatic individuals, their inability to eradicate HIV-1 and their eventual disappearance in late disease have suggested functional impairment of these cells [3]. Recent reports have also demonstrated that CD8+ T cell function is critically dependent on the support of CD4+ T cells [4]. Because one of the cardinal features of HIV-1 infection is the loss of CD4+ T cell proliferative responses from the earliest stages of infection [5–8], it has been proposed that lack of CD4+ T cell help could account for a defective CTL response which ultimately fails to contain viral replication [9]. We have reported previously that HIV-specific CD8+ T cells detected by tetramer-staining are distinguished by strikingly low perforin content, which is associated with less efficient target cell-killing than achieved by CMV-specific CTL from the same donor [10].

LTNPs may constitute a special case for CTL function, as preservation of HIV-1-specific CD4+ T cell proliferative responses have been demonstrated in some LTNPs, in contrast to the majority of infected people [8]. However, disease progression occurs ultimately in the majority of individuals, even among LTNPs showing clear evidence of long-standing viral control. The reason for this remains unclear. To address this question and to investigate the role played by CTL, a cross-sectional study was carried out in a well-characterized cohort of long-term infected patients that included both LTNPs and slow progressors (SP) who had not received any antiretroviral therapy (ART). We analysed the frequency and phenotype of HIV-1-specific CD8+ T cells in relation to CD4+ T cell responses to HIV antigens. HIV-1-specific CD8+ and CD4+ T cell responses were characterized in these subjects by means of ELISpot assays and FACS analysis, using peptide-HLA class I tetrameric complexes to examine cell surface markers and intracellular cytokines in HIV-specific populations. CMV-specific responses were also studied in the same subjects for comparison. We found large numbers of HIV-1-specific CD8+ T cells in the majority of patients regardless of clinical status; however, these cells exhibited a phenotype distinct from CMV-specific cells, as described previously [10], consistent with differences in cellular differentiation. Moreover, the magnitude of virus-specific IFN-γ-producing CD4+ T cell response was strikingly low for HIV-1, in contrast to CMV. Slow progressors and non-progressors were not distinguished by significant differences in the magnitude or phenotype of the HIV-specific CD8+ T cell response, whether or not specific IFN-γ-producing CD4+ responses were present.

Materials and methods

Study population

Samples were taken from members of a previously characterized cohort, established in 1995, of 165 long-term HIV-1-infected volunteers attending clinics in London, UK, who had been enrolled into a nested case–control study of the biological and behavioural correlates of non-progression in HIV-1 infection [11]. Of the original cohort, 46 subjects were defined as LTNPs based on a stable CD4+ T cell count>500 cells/µl at 8 years of infection, 92 as slow progressors based on a CD4+ T cell count <500 cells/µl at 8 years of infection and 27 rapid progressors who developed AIDS within 5 years of infection. The majority (95%) of patients were white homosexual or bisexual men. The study was approved by the local ethics committee and written, informed consent was obtained from all subjects. We identified a subgroup of 29 LTNP and SP patients who were all diagnosed HIV positive prior to 1988 and had therefore been infected for at least 15 years, but had not yet received ART at the time of sample evaluation [11,12]. We identified 17 patients who remained long-term non-progressors based on a stable CD4+ T cell count>500 cells/µl at 15 years; however, three of the LTNPs (001, 037 and 046) had progressed to a CD4+ T cell count below 500 cells shortly after sample evaluation and were therefore analysed as a separate group. Twelve individuals were defined as slow progressors based on a decline in their CD4+ T cell count to below 500 cells/µl at some time over the previous 10 years. All individuals were ART naïve at the time of sample evaluation. It was not possible to have a comparison group of individuals with more rapid HIV progression, as all the original rapid progressors had been commenced on therapy or had since died. HLA-typing was carried out by ARMS-PCR using sequence-specific primers as described previously [13]. Blood samples were either used fresh within 8 h or peripheral blood mononuclear cells (PBMCs) were separated from heparinized blood and cryopreserved for subsequent analysis.

Antigens and antibodies

Peptides were synthesized by FMOC chemistry, and corresponded to previously defined and optimized CTL epitopes [14] (Table 1). Whole HIV-1 p24 and gp120 proteins, nef and tat peptide pools were obtained from the National Institute for Biological Standards and Control (NIBSC) (Herts, UK). Human CMV lysate was purchased from Advanced Biotechnologies Inc. (Columbia, MD, USA). Anti-CD8 (Peridinin chlorophyll protein, PerCP or phycoerythrin, PE), anti-CD4 (PerCP), anti-CD69 (APC or PE), anti-CD27 (FITC or APC), anti-CD28 (FITC or APC) and antiperforin (FITC or PE) antibodies were purchased from Becton Dickinson or BD Pharmingen (San Diego, CA, USA). Monoclonal anti-IFN-γ (FITC) antibodies were purchased from R&D systems (Abingdon, UK) or Becton Dickinson.

Table 1.

Distribution of HIV-1 epitope-specific responses according to the class I HLA type of donors in the cohort

Frequency Frequency Frequency
HLA (no. patients) Protein Epitope no. patients % HLA (no. patients) Protein Epitope (no. patients) % HLA (no. patients) Protein Epitope (no. patients) %
A1 (4) p17 GSEELRSLY – – gp41 IPRRIRQGL 2 25 p24 KAFSPEVIPMF 5 63
nef ISERILSTY – – gp120 RPNNNTRKSI 2 25 B57 p24 ISPRTLNAW 4 50
B7 nef TPGPGVRYPL 3 37 (8) p24 TSTLQEQIAW 3 37
p17 SLYNTVATL 10 53 (8) nef FPVTPQVPLR – – pol IVLPEKDSW 2 25
p24 TLNAWVKVV 2 11 CMV TPRVTGGGAM 7 87
A2 (19) pol VIYQYMDDL 6 32 p24 KAFSPEVIPMF – –
pol ILKEPVHGV 8 47 p17 GGKKKYKL 1 14 B58 p24 ISPRTLNAW – –
nef VLEWRFDSRL – – p17 GGKKKYRL 2 29 (1) p24 TSTLQEQIAW – –
CMV NLVPMVATV 13 68 p24 DCKTILKAL 1 14
B8 p24 EIYKRWII 5 71
p17 KIRLRPGGK 4 36 (7) p24 DIYKRWII 3 43
gp41 RLRDLLLIVTR 1 9 gp41 YLKDQQLL – –
A3 (11) gp120 TVYYGVPVWK 1 9 pol GPKVKQWPL 3 43
pol AIFQSSMTK 4 36 nef FLKEKGGL 7 100 Cw3 p24 QAISPRTL – –
nef QVPLRPMTYK 5 45 (3)
p24 DRFFKTLRA 2 67
A11 (1) pol AIFQSSMTK – – B14 p24 DRFWKTLRA 1 33 p17 KYRLKHLVW – –
nef QVPLRPMTYK – – (3) p24 DLNTMLNTV 1 33 Cw4 p24 QASQEVKNW – –
p24 RAEQASQEV – – (3) gp120 SFNCGGEFF 1 33
p17 KYKLKHIVW 2 40
p24 RDYVDRFFKTL 1 20 B27 p24 KRWIILGLNK 3 60 Cw7 gp160 VYYGVPVWKEA – –
A24 (5) p24 IYKRWIIL 1 20 (5) p24 KRWIIMGLNK 3 60 (8)
gp41 YLKDQQLL 1 20
pol GYIEAEVI 1 20 p17 NSSKVSQNY – – p24 RAEQASQEV – –
p24 PPIPVGDIY 1 25 Cw8 gp120 NCSFNISTSI 1 33
A32 (3) gp120 RIKQIINMW – – B35 gp41 TAVPWNASW 1 25 (3) pol VTDSQYALGI 1 33
pol PIQKETWETW – – (4) pol NPDIVIYQY 1 25 nef KAAVDLSHFL 2 67
nef FPVRPQVPL 2 50
A33 (1) pol AIFQSSMTK – – nef VPLRPMTY – –
p24 RDYVDRFYKTL 1 17
A6801 (1) gp120 VTVYYGVPWK – – B44 gp120 AENLWVTVY 1 17
(6) RT EELRQHLLRW 2 33
A6802 (1) pol DTVLEDINL – –
pol ETAYFILKL – –
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Enzyme-linked immunospot (ELISpot) assays

Synthetic peptides (Table 1) were used at a concentration of 10 µm in IFN-γ enzyme-linked immunospot (ELISpot) assays to define CD8 T cell responses directly ex vivo, as described previously [15]. PHA was always included as a positive control. Recombinant HIV-1-derived proteins gp120, p24, p66 (10 µg/ml; NIBSC, Potters Bar, UK) and overlapping pooled peptides spanning HIV-1 Nef protein (5 µg/ml; NIBSC) with the addition of anti-CD28 MoAb co-stimulus (0·5 µg; Becton Dickinson, Oxford, UK) were used to determine HIV-1-specific CD4 T cell responses by IFN-γ ELISpot assays in PBMC depleted of CD8 T cells with anti-CD8-conjugated Dynabeads (Dynal, Merseyside, UK).

Preparation of HLA-peptide tetrameric complexes

Peptide-MHC tetramers were synthesized as described previously [16]. Briefly, the HLA molecule heavy chain cDNAs were modified by substitution of the transmembrane and cytosolic regions with a sequence encoding the BirA biotinylation enzyme recognition site. These modified HLA heavy chains, and β2-microglobulin, were synthesized in a prokaryotic expression system (pET, R&D Systems), purified from bacterial inclusion bodies, and allowed to refold with the relevant peptide by dilution. Refolded monomeric complexes were purified by FPLC and biotinylated using BirA (avidity), then combined with phycoerythrin (PE)-labelled streptavidin (Sigma) at a 4 : 1 molar ratio to form tetrameric HLA/peptide complexes (tetramers). The tetramers used in these studies were as follows: HLA-A*0201-SLYNTVATL (A2 gag p17), A*0201-ILKEPVHGV (A2 pol), A*0201-NLVPMVATV (A2 CMV), B7-TPGPGVRYPL (B7 nef), B7-IPRRIRQGL (B7 gp41), B7-RPNNNTRKSI (B7 gp120), B7-TPRVTGGGAM (B7 CMV), B8-FLKEKGGL (B8 nef), B8-DIYKRWII (B8 gag p24), B*2705-KRWIIMGLNK (B27 gag p24), B57-KAFSPEVIPMF (B57 gag p24) and Cw4-SFNCGGEFF (Cw4 gp120) complexes.

Flow cytometry

Cell surface and intracellular stainings were generally carried out directly on whole blood. Titrated tetramers (PE-conjugated) were added to 150 µl of heparinized blood for 15 min at 37°C, followed by addition of a panel of titrated antibodies (FITC, PerCP or APC-conjugated) and incubation for 15 min at room temperature (RT). The lymphocytes were then fixed and the red blood cells lysed using FACSTM lysis solution (Becton Dickinson). Cells were washed in PBS, 0·5 mm EDTA, 1% BSA, fixed and permeabilized in FACSTM Permeabilization buffer (Becton Dickinson). After washing, perforin staining was performed for 15 min at RT in the dark using a previously titrated concentration of antibodies. Cells were then washed and stored in Cell FixTM buffer (Becton Dickinson) at 4°C until analysing on a Becton Dickinson FACSCalibur flow cytometer appropriately compensated using freshly stained PBMCs. Four-colour stainings were performed and always included CD8 staining. The threshold for Perforin levels (high or low) was generally set up so that 35–50% of Perforin-positive cells were seen in the CD8+ T cell population.

Intracellular IFN-γ staining in CD4+ T cells

Intracellular cytokine staining assays were performed as described elsewhere [17]. In brief, 0·5 ml of whole blood were incubated with either streptococcal enterotoxin B (SEB, 2 µg/ml), CMV lysate or a pool of HIV-1 p24, HIV-1 gp120 recombinant proteins and nef or tat peptides at a final concentration of 10 µg/ml, and 1 µg/ml each of the co-stimulatory MoAbs anti-CD28 and anti-CD49d (Becton Dickinson). The blood was left at 37°C, 5% CO2 for 1 h, before the addition of 10 µg/ml of brefeldin A (Sigma-Aldrich). After an additional 5-h incubation at 37°C, 5% CO2, the red blood cells were lysed using FACSTM lysis solution. Cells were then permeabilized and stained as described above with anti-IFN-γ (FITC), anti-CD69 (PE), anti-CD4 (PerCP) and anti-CD8 (APC) antibodies.

Statistical analysis

The median viral load and CD4 cell count in the LTNPs and SPs at the time of sample evaluation were compared using Mann–Whitney U-test.

Results

Cohort characteristics

Based on the definition of long-term non-progressors [11,12], patients were categorized into two groups according to their current clinical status and CD4 cell count (Table 2). 14 patients were categorized as long-term non-progressors (LTNPs), as defined by absence of symptoms without antiretroviral therapy and stable CD4+ T cell counts>500 cells/µl at a median 15 years of infection (range 13–15 years). Three additional LTNPs (001, 037 and 046) had progressed with a decline in their CD4+ T cell count to below 500 cells/µl shortly after their immunological evaluation and were therefore analysed as a separate subgroup. Twelve patients were termed slow progressors (SPs), who had been infected for a median of 15 years but who showed evidence of disease progression (declining CD4+ T cell count below 500 cells/µl) at the time of study, but had not yet started antiviral therapy. The median CD4 cell count and viral load at the time of evaluation in LTNPs was 745 cells/µl and 18 473 copies/ml compared to 310 cells/µl (P = < 0·0001) and 68 266 copies/ml (P = 0·003) among the SPs. There were no other significant demographic differences between LTNPs and SPs. Our cohort showed the characteristic distribution of MHC class I haplotypes described in Caucasian populations (Table 2). Consistent with the previously described association of HLA-B57 with non-progression in HIV-1 infection [18–20], seven of the eight subjects expressing HLA-B*5701 were either LTNPs or switchers from LTNPs to SPs.

Table 2.

Clinical data of study patients at the time of sampling and numbers of HIV epitopes tested for each patient

No. of HIV epitopes tested
Group* Patient ID Gender Year of diagnosis CD4+ T cells (cells/µl) Viral load (copies/ml) HLA type ELISpot Tetramer
SP 012 M 1985 218 77 000 A2, A32, B7, B8, Cw7 19 6
018 M 1985 296 1 002 A3, A24, B14, B62, Cw3, Cw8 15 /
028 M 1987 272 9 600 A2, B27, B73, Cw1, Cw15 7 3
034 M 1986 221 263 000 A2, A33, B7, B35, Cw4, Cw7 17 6
041 M 1985 317 95 640 A2, A3, B8, B44, Cw3, Cw7 21 5
043 M 1985 489 18 266 A2, B15, B44, Cw4, Cw5 9 3
048 M 1987 387 51 361 A2, A68, B7, B62, Cw3, Cw12 12 5
114 M 1984 437 163 767 A2, A23, B8, B44, Cw7, Cw4 18 5
118 M 1985 129 4 556 A2, A24, B40, B62, Cw3 11 2
140 M 1985 378 44 367 A30, A74, B57, B15, Cw21, Cw7 5 /
147 M 1985 492 21 202 A3, B7, B62, Cw3, Cw7 11 3
156 M 1985 85 69 433 A2, A23, B44, Cw2, Cw5 7 2
LTNP to SP 001 M 1986 479 88 194 A1, A3, B14, B57, Cw8 17 1
037 M 1985 711 30 149 A3, A32, B27, B40, Cw2, Cw5 8 1
046 M 1985 663 104 000 A1, A2, B8, B57, Cw6, Cw12 15 6
LTNP 005 M 1985 695 64 458 A2, A24, B8, B57, Cw6, Cw7 15 6
011 M 1985 692 27 409 A3, A24, B27, B35, Cw1, Cw12 14 1
013 M 1986 840 733 A2, A3, B44, B58, Cw7 12 2
025 M 1985 650 17 300 A3, A31, B7, B35, Cw4, Cw7 12 2
039 M 1985 549 9 719 A2, A32, B27, B35, Cw1, Cw4 14 3
058 M 1987 589 3 050 A2, A3, B7, B51, Cw7, Cw14 13 5
065 M 1985 1993 23 521 A1, A2, B8, B57, Cw6, Cw7 21 5
085 F 1987 588 1 700 A2, A11, B7, B62, Cw3, Cw7 10 4
113 M 1984 588 <50 A2, A3, B27, B57, Cw2, Cw6 16 3
122 M 1984 736 166 A1, A24, B44, B51, Cw5, Cw15 8 /
134 M 1985 763 1 198 A1, B14, B52, Cw8, Cw12 9 /
200 F 1987 552 928 A2, A24, B8, B15, Cw7, Cw12 17 4
201 M 1988 815 60 A1, A2, B13, B57, Cw6 11 3
202 F 1988 685 <50 A1, A3, B7, B57, Cw6, Cw7 10 3
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*

Patients were classified into two groups according to clinical status, CD4 count and rate of CD4 decline, as described previously [11,12]. SP = slow processor; LTNP = long-term non-progressor.

HIV-1- and CMV-specific CD8+ T cell populations were studied using a panel of 70 previously optimized CTL epitopes in the ELISpot assays and by FACS analysis using 12 different peptide-HLA tetramers. The frequencies obtained with these methods were compared whenever the same epitope was tested by ELISpot and tetramer. In keeping with the results of other studies in chronic EBV [21] and HIV-1 infection [22], the mean value for tetramer staining was sixfold higher than the number of antigen-specific cells estimated using ELISpot assays, and no differences in this ratio were found between the two study groups (data not shown). However, there was a close correlation between ELISpot and tetramer frequencies (r = 0·757, P = < 0·0001), indicating that ELISpot assays provide a reliable relative measure of cell frequencies.

Large numbers of HIV-1- and CMV-specific CD8+ T cells are seen in both LTNPs and SPs

Large HIV-1-specific CD8+ T cell populations were found in the majority of study patients (Fig. 1a). This finding was not unique to long-term non-progressors, or LTNPs who had recently progressed, but was also observed in the slow progressors. Twelve of 14 LTNPs (86%) and 11 of 12 SPs (92%) (three of three in the LTNP to SP) displayed positive responses to at least one HIV-1 epitope in both ELISpot assay and FACS analysis. In one progressing donor (012), HIV-1-specific CD8+ T cells represented up to 24% of the CD8+ T cell population: 19·1% of circulating CD8+ T cells stained with a panel of tetramers (Fig. 1b), and at least a further 5% of responses (A2 p 17 SLY, A2 pol VIY, B8 pol GPK) were detected using ELISpot assays with peptides for which no tetramers were available. Large numbers of CD8+ T cells specific for CMV were also observed in the donors in this cohort (Fig. 2a). Seven of nine LTNPs (78%) and 10 of 10 SPs (100%) displayed a positive response for previously defined immunodominant CMV peptides restricted by either HLA-A2 or B7. The level of responses was similar to the frequencies described recently in healthy HIV-seronegative donors [23]. In one subject SP (114) 16·8% of the circulating CD8+ T cell population was found to be specific for a single CMV epitope by tetramer analysis (Fig. 2b).

Fig. 1.

Fig. 1

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HIV specific CD8+ T cells in SPs and LTNPs. (a) ELISpot assays were carried out using optimized CTL epitopes according to donor HLA types. Responses to CTL epitopes are added together, grouped by gene products, and shown for each donor. The values are expressed as sfu/106 PBMCs. The background IFN-γ production (usually not higher than 20–30 sfu/106 PBMCs) has been subtracted. (b) Ex vivo tetramer staining showing the largest populations specific for individual HIV epitopes found in any of our donors. The percentage is calculated after gating on the CD8+ population. Inline graphic, gag; Inline graphic, pol; Inline graphic, env; □, nef.

Fig. 2.

Fig. 2

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The number of CMV-specific CD8+ T cells in SPs and LTNPs. (a) ELISpot assays were carried out using optimized CTL epitopes in donors with either HLA-A2 or B7. The values are expressed as sfu/106 PBMCs, after subtraction of background IFN-γ production. (b) Ex vivo tetramer staining showing the largest population specific for a single CMV epitope detected in any of our cohort (114, SP). The percentage is calculated after gating on the CD8+ population.

The immunodominance of particular class I-restricted CD8+ T cell responses within our panel of 70 known CTL epitopes was evaluated (Table 1). Particularly high response rates were directed towards the following CTL epitopes: A2 p17 SLY, A2 pol ILK, A2 CMV, A3 nef QVP, B7 CMV, B8 p24 EIY, B57 p24 KAF and B57 p24 ISP. Interestingly, 100% of the seven HLA B8 subjects responded to an epitope in nef FLKEKGGL (Table 1), which has also been described as an immunodominant epitope in acute HIV-1 infection [24] and during viral rebound [25] in donors with HLA-B8. However, many other antigens triggered IFN-γ production (albeit at lower frequencies), emphasizing the breadth of CD8+ T cell responses specific for HIV-1.

HIV-1-specific CD8+ T cells exhibit a distinct phenotype from CMV-specific cells, regardless of clinical status

CD28 and CD27 are co-stimulatory receptors whose expression is associated with cellular differentiation and the intracellular expression of the cytotoxic factor perforin [10,26,27]. We have shown previously that in chronic HIV-1 infection the HIV-specific populations (mainly CD28−, CD27+, perforin low) differ in expression of these markers from CMV-specific CD8+ T cells (mainly CD28−, CD27−, perforin high) [10], which might be believed to represent better cytotoxic effector cells [26]. Here, we found that the phenotype of the HIV-specific tetramer-staining population in LTNPs did not differ from that seen in SPs (Fig. 3). Although a trend toward greater number of CD27− cells in the LTNPs may be observed, this one does not reach significance. No significant difference was seen in perforin expression between LTNPs (0% to 24% perforin high) and SPs (3% to 26% perforin high). A similar phenotype of HIV-specific CD8+ T cells appears very early following primary infection [28] and is also found in individuals with prolonged viral suppression [29]. Altogether these data suggest unexpectedly that the phenotype and the perforin level of the HIV-specific CD8+ T cells may be independent from the patient clinical status.

Fig. 3.

Fig. 3

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Expression of CD27, CD28 and perforin in HIV-specific CD8+ T cells in LTNPs and SPs. Expression on CMV-specific CD8+ T cells is presented as a comparison. The results are expressed as percentages of HIV-specific or CMV tetramer staining cells expressing CD27, CD28 or high level of perforin.

HIV-1-specific IFN-γ-producing CD4+ T cell responses in long-term HIV-1-infected individuals are infrequent and low in magnitude compared with CMV-specific CD4+ T cells

IFN-γ ELISpot assays and intracellular IFN-γ staining were used to assess HIV-1-specific T helper responses in our cohort of long-term HIV-1-infected individuals. Using ELISpot assays with a panel of HIV antigens we observed that eight of 13 LTNPs (61%) and four of eight SPs (50%) displayed HIV-1-specific CD4+ T cell responses (Fig. 4a): however, the magnitude of response was generally low (20–1650 sfu/106 CD8 depleted PBMCs, mean 178). A trend toward higher HIV specific IFN-γ-producing CD4+ T cell responses was observed in the LTNPs compared to individuals showing signs of progression; however, this did not reach statistical significance (P = 0·566), which may be a consequence of the limited sample size. Most donors had detectable CMV-specific CD4+ T cell responses: 10 of 11 LTNPs (91%) and eight of eight SPs (100%) showed IFN-γ production upon stimulation with CMV antigens, and the level of responses was significantly higher (58–4500 sfu/106 CD8 depleted PBMCs, mean 1538). The results from intracellular cytokine staining confirmed the ELISpot results: HIV-1-specific IFN-γ-producing cells were usually either not detected at all or were present at low levels in most subjects, contrasting with significant levels of CMV responses detected by FACS analysis (Fig. 4b).

Fig. 4.

Fig. 4

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Comparison between HIV-specific and CMV-specific CD4+ T cell responses. (a) IFN-γ ELISpot assays were carried out on CD8+ T cell-depleted PBMCs. The values are expressed as sfu/106 CD8+ T cell-depleted PBMCs. The background IFN-γ production was always subtracted. (b) Staining for intracellular IFN-γ and activation marker CD69 on CD4+ T cells upon stimulation with SEB, CMV or HIV antigens. The populations shown have been gated on CD4+ cells. The numbers show the percentages of CD4+/CD69+/IFN-γ+ cells. Inline graphic, HIV; ▪, CMV.

Discussion

Using a panel of 70 previously optimized CTL epitopes to screen for HIV-1-specific responses in the ELISpot assays, we found that many patients had responses directed towards well-characterized immunodominant antigens, such as A2 SLY, A2 ILK, A2 VIY, A3 QVP, B57 KAF, B57 ISP, B8 EIY and in particular B8 FLK, with usually high frequencies. It was not possible in this study to identify particular responses that are unique to non-progressors. However, it is intriguing that so many of the A2 donors responded to a conserved peptide in the active site of reverse transcriptase (VIYQYMMDL) which has been linked previously with non-progression [30] and which is an unusual response in most HIV-infected donors with HLA-A2 [31]. The mechanisms underlying the association of non-progression with the HLA allele B57, as described previously [18–20] and also observed in this study, remain unclear. However, it is worth noting that in this study B57-restricted CD8+ T cell populations did not differ in phenotype from CTL restricted by other less favourable HLA molecules. The breadth of the virus-specific CD8+ T cell-mediated response in these donors is striking and illustrates the complexity of detailed CTL studies in HIV-infected patients.

Long-term HIV-1-infected individuals displayed strong broadly directed CD8+ T cell responses against HIV-1 antigens, but the magnitude of the response did not distinguish between non-progressors and individuals with progressive disease. A potential limitation of our study is the use of optimized CTL epitopes to monitor HIV-specific CD8+ T cells which may result in an underestimation of the CD8 responses, compared with the use of overlapping peptides representing all the HIV proteins. This issue was emphasized recently in a study by Betts et al., where an extensive panel of overlapping peptides covering most of the HIV proteins was used [32]. Nevertheless, the authors concluded that overall frequencies of HIV-specific T cells are not the main determinant of immune-mediated protection in HIV-infection and disease progression, which is in keeping with our results as well as those of other groups [33]. A further possible explanation for our failure to discriminate between LTNPs and progressors on the basis of their CD8 responses or CD4 IFN-γ responses may be the fact that both groups represent patients with an unusually favourable outcome. Because only progressors who had not yet required antiretroviral therapy at 15 years were selected for the study these patients, despite a CD4+ T cell count below 500 cells, represent a relatively indolent disease course and may not be sufficiently distinct from LTNPs to show clear differences in immune responses.

The highest HIV-1-specific responses, accounting for almost 24% of circulating CD8+ T cells in this donor, were seen in a patient with clear evidence of disease progression. The class I HLA haplotype of this patient (which included HLA A2, B7 and B8) is one for which many CTL epitopes have been described, which facilitates substantially the detection of responses. It is possible that similarly high levels of HIV-1-specific CD8+ T cells could be present in other donors with HLA types for which fewer CTL epitopes have been mapped.

Large numbers of CMV-specific CD8+ T cells were found in our cohort, both in LTNPs and SPs (in one patient, up to 17% of the circulating CD8+ T cells specific for a single CMV epitope). In chronic infected patients, these cells differ from HIV-1-specific tetramer-staining populations, exhibiting a more differentiated phenotype (CD28−/CD27−) with higher perforin level [10]. However, we observed no obvious difference with regard to this phenotype between HIV-1-specific CD8+ T cells in LTNPs or SPs, indicating that the patient non-progression status does not result from a distinct HIV-1-specific CD8+ T cell phenotype.

We observed a trend towards a higher level of IFN-γ-producing CD4+ T cell responses to HIV-1 antigens in LTNPs compared to SPs, although this did not reach statistical significance. This is in contrast with the report of higher proliferative responses to HIV antigens in another study of LTNPs compared to progressors [8]. These discrepancies highlight the differences between IFN-γ production and proliferative capacity often observed in the study of antigen specific CD4+ T cells [34]. In contrast to HIV-specific responses, a vigorous CD4+ T cell activity was seen in many patients against CMV antigens, in keeping with a recent study by Pitcher et al. [17]. The different antigen preparations used (whole viral lysate for CMV versus single recombinant proteins or peptide pools representing p24, gp120, nef and tat for HIV-1) may differ in their ability to stimulate measurable CD4+ T cell responses but it seems unlikely that this fact would account for the extent of the difference in magnitude of HIV-1 and CMV-specific CD4+ T cell responses.

The precise role of CD4+ T cell help in supporting the activity of virus-specific cytotoxic T cells remains open to debate. The magnitude of the CMV-specific IFN-γ-producing CD4+ T cell activity and the more differentiated CD8+ T cells directed to CMV antigens that we observed, in contrast to low IFN-γ-producing CD4+ T cell activity and less differentiated CD8+ T cells directed to HIV antigens, might suggest an association between these two factors. However, these differences may also reflect specific immune responses varying in the context of two distinct virus infections. Interestingly, in a recent study of patients with detectable HIV-specific CD4+ T cell responses in primary HIV-1 infection, we did not find any differences in their HIV-specific CTL phenotype, which remained mainly CD28−/CD27+ [28].

In other studies of immune responses to HIV antigens in LTNPs, a consistent feature has been a strong cytotoxic T cell response to the virus [2,35]. However, it is not clear that the magnitude of the HIV-1-specific CD8+ T cell response correlates with disease outcome. Dalod et al. [36] found no correlation between the intensity of anti-HIV CD8+ T cell responses and either CD4+ count or viral load, although they noted that the polyclonality of the HIV-specific CD8+ T cell response was linked to CD4+ count. Gea-Banacloche and colleagues [37] noted similarly high levels of HIV-specific CD8+ T cells in both progressors and non-progressors, and did not find a correlation between the magnitude of the response and viral load.

Overall, neither the numbers nor the differentiation phenotype, including perforin levels, of HIV-1-specific CD8+ T cells appear to be correlated directly with the non-progressor state. These data therefore suggest that HIV-specific CTLs may not represent the determining factor responsible for non-progression in these HIV-infected patients.

Acknowledgments

We are very grateful to the staff and patients of the Caldecot Centre at King's College Hospital London which provided blood samples, and to Shahed Murad for the help on the statistical analysis. This work was supported by the Medical Research Council of the UK, the Elizabeth Glaser Paediatric AIDS Foundation and the NIH. SR-J holds an Elizabeth Glaser scientist award. The HIV antigens (gp120 and p24 recombinant proteins and nef and tat peptide pools) from Dr Harvey C Holmes (NIBSC) were obtained from the Centralized Facility for AIDS Reagents supported by EU Programme EVA (contract QLK2-CT-1999–00609) and the UK Medical Research Council.

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