Differentiation of human CD8 T cells: implications for in vivo persistence of CD8+CD28 cytotoxic effector clones
David N. Posnett,
James W. Edinger,
J. Sanil Manavalan,
Craig Irwin and
Gilles Marodon
Immunology Program, Graduate School of Medical Sciences, and the Department of Medicine, Cornell University, Weill Medical College, 1300 York Avenue, Box 56, New York, NY 10021
Correspondence to:
D. N. Posnett
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Abstract
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CD8 T cells contain a distinct subset of CD8+CD28 cells. These cells are not present at birth and their frequency increases with age. They frequently contain expanded clones using various TCR
ß receptors and these clones can represent >50% of all CD8 cells, specially in old subjects or patients with chronic viral infections such as HIV-1. Herein, it is shown that a large fraction of CD8+CD28 cells expresses intracellular perforin by three-color flow cytometry, in particular when this subset is expanded. Together with their known ability to exert potent re-directed cytotoxicity, this indicates that CD8+CD28 T cells comprise cytotoxic effector cells. With BrdU labeling, we show that CD8+CD28 cells derive from CD8+CD28+ precursors in vitro. In addition, sorted CD8+CD28+ cells gave rise to a population of CD8+CD28 cells after allo-stimulation. Moreover, ex vivo CD8+CD28+ cells contain the majority of CD8 blasts, supporting the notion that they contain the proliferative precursors of CD8+CD28 cells. CD95 (Fas) expression was lower in CD8+CD28 cells, and this subset was less prone to spontaneous apoptosis in ex vivo samples and more resistant to activation-induced cell death induced by a superantigen in vitro. Thus, the persistence of expanded clones in vivo in the CD8+CD28 subset may be explained by antigen-driven differentiation from CD8+CD28+ memory precursors, with relative resistance to apoptosis as the clones become perforin+ effector cells.
Keywords: age, apoptosis, CD8, CD28, human, oligoclonal, TCR
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Introduction
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In recent years a large number of TCR repertoire studies have uncovered oligoclonal T cell expansions in various diseases, including chronic viral infections such as HIV infection (1,2). These expansions also occur in apparently normal subjects. In a prior study we described clonal CD8 expansions in normal elderly human subjects (3). Clonal expansions were also found in normal young subjects and even children, among CD4 as well as CD8 cells, using more sensitive molecular assays (46). Several studies confirmed that CD8 clonal expansions were more prominent and more frequent in the elderly (3,5,7). Similar observations were made in old mice (8). In some human subjects individual clonal expansions have been followed for many years. Often, the expanded clones remain present for several years without much change in the percentage of cells (5,6,911). In these cases clonal `exhaustion' does not appear to be common.
Expanded CD8 clones can be dramatic and take over >50% of the CD8 compartment (5,9,10). Recent studies with tetrameric MHCantigen constructs have uncovered similarly large oligoclonal expansions of antigen-specific CD8 cells in response to viral infections (12). Usually these responses resolve, as after acute EpsteinBarr virus infection, but they may persist, as in the case of chronic HIV infection. The in vivo persistence of expanded CD8 clones in old subjects suggests chronic antigen exposure. The CD8 clones probably accumulate because of continuous production, perhaps combined with decreased elimination.
An early finding was that CD8 T cell clones that are expanded in vivo lack CD28 expression (3). Most CD28 T cells in the peripheral blood are CD8+ TCR
ß T cells, although they may include double-negative CD4CD8 T cells, CD4 single-positive T cells and TCR
T cells. CD8+CD28 TCR
ß T cells are not present at birth, but gradually increase in numbers with age, both in mice and in humans (3,5,7,13). Loss of CD28 expression is associated with a phenotype of antigen-experienced CD8+ T cells: CD57+CD11b+CD27CD49d(VLA4)+ (13,14) with variable expression of CD45RO/RA (3,14). Co-expression of chronic activation antigens like CD38 and HLA-DR can occur, particularly in HIV-infected subjects (1517). Acute activation antigens such as CD69, CD25 and CD71 are not usually expressed.
Stimulation of a T cell via the TCR and CD28 results in T cell responses such as proliferation, IL-2 secretion and activation of cytolytic effector function from memory precursor cytotoxic T lymphocytes (pCTL) (18). By contrast TCR stimulation without CD28 stimulation, for instance by antigen-presenting cells that lack CD80/86, can lead to anergy. CD8+CD28 T cells characteristically give weak proliferative responses (13), but are perfectly capable of exerting cytotoxic effector function (18). Some authors attribute antigen non-specific suppressor functions in co-culture assays to a similar, if not identical subset (13,14,1922).
The largely overlapping subset of CD8+CD27 cells has been shown to contain intracellular granzyme B and perforin, express Fas ligand, and produce IFN-
and tumor necrosis factor-
, and is thought to contain cytotoxic effector cells, while CD8+CD27+ cells comprise naive and memory CD8 subsets (14). Ex vivo CD8+CD28 T lymphocytes are enriched in cytotoxic effector function in a re-directed cytotoxicity assay using anti-CD3-coated P815 cells as target cells (13). In HIV-infected patients CD8+CD28 cells produce high levels of IFN-
(23) and may express intracellular TIA-1 (24), but results vary on the expression of perforin (24,25).
It is not yet clear whether all CD8+CD28 cells are cytotoxic effectors, whether they are functionally homogeneous or perhaps include a group of distinct maturational stages during differentiation of a CTL effector cell. Direct evidence of derivation from CD8+CD28+ precursors is still incomplete, especially since CD8+CD28 and CD8+CD28+ T cell clones were reported to maintain their phenotype during in vitro culture (13). It is also not clear why these cells sometimes accumulate as persistent clonal expansions and other times not. This report documents long-term persistence of clones within an expanded CD8+CD28 subset. As cytotoxic effector CD8+ cells mature they appear to lose CD28 expression, acquire intracellular perforin and become relatively resistant to apoptosis.
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Methods
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Human subjects and samples
Peripheral blood lymphocytes (PBL) were isolated by Ficoll-Hypaque density centrifugation from heparinized blood according to an IRB approved protocol. The TCR repertoire of normal subjects 15 and 16 was previously described (3). Subject 15 presented with an increased CD8+CD28 subset containing clonal expansions in 1993. In 1996 an asymptomatic renal adenocarcinoma was fortuitously discovered for which nephrectomy was performed. There was residual cancer in the contra-lateral kidney that has been followed by computed tomography scans since early 1997 without any change and without metastasis. Intravenous substance abusing (IVSA) patients, 2558 years old (mean = 39.4; n = 16), were recruited from a methadone clinic. Eight of these patients were HVB and two were HVC sero-positive. Seven HIV-1+ IVSA patients, 3345 years old (mean = 36.8), were all positive for HVB and two had HVC. HIV-1+ patients were all on HAART therapy, had 126641 CD4 cells/µl blood and low viral loads by the bDNA assay. Twenty normal subjects of various ages were recruited. `Old' subjects were 6579 years old (mean = 71.6). `Young' subjects were 2449 years old (mean = 36.3).
Antibodies and staining
The following antibodies were used for staining in either two- or three-color protocols. In all cases the antibodies were titrated for optimal results. TCR
ßFITC (PharMingen, San Diego, CA), CD57FITC (PharMingen), CD56FITC (Becton Dickinson, Mountain View, CA), CD94FITC (PharMingen), CD95FITC (Cal & Biological Labs), Annexin VFITC (Trevigen), anti-perforinFITC mAb
G9 (Ancell), CD4biotin FFB2.3, CD8biotin (PharMingen), CD28PE (PharMingen), goat anti-mouse IgFITC F(ab')2 (BioSource, Camarillo, CA), CD8PE (Pharmingen), streptavidinTriColor (Caltag), Vß22FITC (Coulter-Immunotech), Vß8FITC (Coulter-Immunotech), and Vß2 E2.2E7.2, Vß3 LE89, Vß5.1 IMMU157, Vß5.2/3 4H11, Vß6.7 OT145, Vß8 MX3, Vß9.1 FIN9, Vß11 C21, Vß12 SV11, Vß13.1 H131, Vß14 CAS1.1.3, Vß16 TAMAYA1.2, Vß17 C1, Vß20 ELL1.4 and Vß22 IMMU546, all from the last international workshop on TCR antibodies (26).
To evaluate TCR Vß subsets, a two-color staining protocol was used. About 105106 T cells were stained in microtiter wells with an anti-TCR Vß antibody for 30 min, then washed with staining buffer (HBSS, 5% FCS and 0.02% sodium azide), followed by staining with goat anti-mouse Ig F(ab')2 labeled with FITC for 30 min, another wash with staining buffer and staining with anti-CD8PE 1:50 dilution for 30 min. Washes were done by centrifuging at 2000 r.p.m. and flicking the plates to obtain dry pellets. Incubations were done at room temperature on a shaker in the dark.
For three-color staining a FITC-conjugated antibody (e.g. TCR
ßFITC) and the biotinylated antibody (e.g. CD8biotin) were used together for the first 30 min incubation, followed by a wash and then a 30 min incubation with the PE-conjugated antibody (e.g. CD28PE) together with streptavidinTriColor at a final dilution of 1:50. The cells were then washed and resuspended in wash buffer with 1% paraformaldehyde in a 0.5 ml volume for analysis on a Coulter Epics II cytofluorograph, calibrated with Coulter Standard-Brite beads and using Coulter XL software.
For cell sorting PBL were stained with TCR
ßFITC, CD8biotin and CD28PE as above, and sorted on a FACS Vantage (Becton Dickinson).
For intracellular perforin staining the cells were first permeabilized with wash buffer containing 4% paraformaldehyde, 0.1% saponin and 10 mM HEPES, and then stained using the regular three-color staining protocol.
For annexin VFITC staining, cells were first stained with CD8biotin, followed by CD28PE with streptavidinTriColor, a wash with staining buffer (10 mM HEPES, pH 7.4, 0.15 M NaCl, 5 mM KCl, 1 mM MgCl2 and 1.8 mM CaCl2) and then annexin VFITC for 15 min at room temperature at a final dilution of 1:50 (in 50 µl), followed by dilution in 450 µl staining buffer with 1% paraformaldehyde. For staining with annexin Vbiotin we stained as per the regular three-color protocol detailed above.
BrdU experiments
PBL (2x107) were cultured with 10 µg/ml phytohemagglutinin (PHA) and 40 U/ml IL-2 (Roche). After 3 days the cells were washed and resuspended at 106 cells/ml in fresh medium with PHA and IL-2, with or without 100 µM BrdU (Sigma, St Louis, MO). After 1 h the cells were washed and resuspended in media with IL-2. At the indicated time points thereafter cells were stained for CD8biotin, CD28PE and anti-BrdUFITC as per Penit et al. (27) with modifications. An aliquot of 5x106 cells was first washed with 5 ml wash buffer (HBSS, 10% FCS and 0.02% sodium azide), then stained with CD8biotinx30 min, then washed, stained with CD28PE and streptavidinTriColor for 30 min, washed, and fixed overnight in 1 ml 1% paraformaldehyde/0.5% Tween 20, at 4°C. The cells were then washed 4 times with 5 ml DNase I digestion buffer (20 mM TrisHCl, 10 mM Mg2+), centrifuged at 1000 r.p.m. for 10 min and resuspended in 100 µl buffer with 50 U DNase I for digestion at 37°C for 30 min. The cells were then washed with 5 ml stain buffer, resuspended in 100 µl, stained with 2 µl anti-BrdUFITC (Pharmingen) on a shaker for 30 min at room temperature, washed and resuspended in 0.5 ml stain buffer for analysis. These experiments could not be extended beyond 4872 h because of decreasing viability of the PHA-stimulated cells.
Cell cultures
PBL or CD4-depleted PBL were cultured with 200 ng/ml staphylococcal enterotoxin E (SEE), or with SEE pulsed BJAB lymphoblastoid cells (irradiated with 5000 rad), in RPMI with 10% FCS, supplemented with glutamine, penicillin/streptomycin and fungizone, for the indicated period of time. Where indicated, 40 U/ml of rhIL-2 (Boehringer Mannheim, Indianapolis, IN) was added on day 4 and 8 of culture. CD4 depletion was performed as follows. PBL were incubated with anti-CD4 at saturating concentrations for 30 min at room temperature, washed twice with sodium azide-free staining buffer, followed by a 30 min incubation with goat anti-mouse Ig F(ab')2-coated magnetic beads (Dynal, Oslo, Norway) at a 20:1 bead to cell ratio. Cells with bound magnetic beads were removed with a magnet. The remaining cells were washed and an aliquot was stained to assess the efficiency of CD4 cell removal which was usually >90%.
TCR ß chain CDR3 analysis
The CDR3 length assay is described elsewhere in detail (6). Briefly, first-step PCR reactions were set up with selected Vß-specific primers paired with a Cß primer. The products were then subjected to a second step run-off reaction using a nested fluorophore-labeled anti-sense Cß primer, run on a 6% acrylamide sequencing gel and visualized on a 373A Applied Systems DNA sequencer with fluorescent size markers in the 80380 bp range.
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Results
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TCR repertoire of CD8+CD28 T cells
The peripheral blood of adults contains a distinct population of CD4CD28 cells. CD28 TCR
ß+ cells were found nearly exclusively in the CD4 subset in three random normal subjects, aged 2734 (Fig. 1
). These TCR
ß+ CD4CD28 cells were single-positive CD8+ cells. Next we used the same three subjects to test CD28 expression in 12 Vß subsets gated on CD4 cells (Fig. 2
). Expanded CD8+ Vß subsets had previously been found to lack CD28 expression (3), but it was unclear to what extent non-expanded CD8+ Vß subsets also contained CD28 cells. The size of the Vß subset did not correlate well with the extent of CD28 loss. Thus, two large Vß subsets in subject 2, Vß17 and Vß2, contained 75 and 40% CD28 cells respectively. However, the minor Vß16 subset in subject 2 and the small Vß3 subset in subject 3 also contained a majority of CD28 cells (Fig. 2
). Therefore, Vß expansion was not a prerequisite for the CD28 phenotype. Overall, the prevalence of CD28 cells differed considerably from one Vß subset to the other.

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Fig. 2. Loss of CD28 expression differs among CD4 TCR Vß subsets. (A) The TCR Vß repertoire of CD4 cells from subjects 13 is shown. In subject 2, for example, there are two large subsets, Vß2 and Vß17. (B) The percentage of CD28 cells in each of the subsets shown in (A) is depicted. In subject 2, for example, Vß16 and Vß17, are particularly enriched in CD28 cells.
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Subject 15 (78 years old in 1997) was studied in detail because of a large TCR
ß+ CD8+CD28 subset. In subject 15, 58% of TCR
ß+ cells were CD8+ and 42% CD4+. Greater than 84% of the TCR+CD8+ cells lacked CD28 (Figs 6 and 7
). There were four Vß subset expansions that were followed over >4 years (Table 1
), comprising together between 46.9 and 31.1% of total CD8 cells. As an example of their phenotype, the Vß22 subset consisted predominantly of CD8+CD28 cells (Fig. 3B
). In 1993 the Vß5.2 cells contained a single dominant sequence, assessed by sequencing TCRß CDR3 regions (3). We used a PCR-based CDR3 length assay to detect restricted CDR3 length as a measure of oligoclonality. In each of the subsets examined the same peaks were present in 1993 and in 1997 (Fig. 3A
). In sum, the CD8+CD28 cells in subject 15 contain chronic oligoclonal expansions which could perhaps be driven by persistent antigens.

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Fig. 7. CD8 subsets in different groups of patients. The mean percentages of CD8+CD28 and CD8+CD28+ cells within PBL are given for the indicated groups with the SD of the mean and for three individual subjects. The fraction of perforin+ cells in each subset is indicated by the stippled section of the bar. Indicated P values for comparison of means were obtained with a t-test. Met43 is an IVSA patient with chronic HVC infection. Met43, LB and subject 15 were not included in the analysis of their respective groups.
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CD8+CD28 T cells derive from CD8+CD28+ precursors
It has been proposed that CD8+CD57+ differentiate from CD8+CD57 precursors (28,29) and that CD8+CD28 cells derive from CD8+CD28+ precursors, because the CD8+CD28 phenotype can be induced with in vitro stimulation of cord blood PBL (30) which typically contains only CD8+CD28+ T cells (13). To examine this question directly, BrdU-labeled T cells were used (Fig. 4
). Among gated BrdU+ cells, there was a gradual shift from a predominant CD8+CD28+ phenotype early in the culture to a CD8+CD28 phenotype (Fig. 4B
), demonstrating directly that CD8+CD28 cells were generated from cycling CD8+CD28+ precursors. As a control, the ratio of CD28+/CD28 cells in the total CD8 population remained constant, demonstrating that the switch from CD28+ to CD28 was specific to BrdU+ cells. Due to PHA stimulation, total cell numbers increased 2-fold during the observation period. Cell death was not observed by Trypan blue exclusion or forward scatter (FSC)/side scatter (SSC) light scatter characteristics. During the first 6 h of this experiment BrdU+ CD8+ cells contained on average 41.9% FSChi blast cells compared with 5.6% FSChi blasts in BrdU CD8+ cells. The BrdU+CD8+CD28+ subset contained 2-fold more FSChi blasts than the BrdU+ CD8+CD28 subset. Thus, it is unlikely that BrdU+ CD8+CD28 cells were overgrowing the cultures. Rather, a fraction of BrdU-labeled CD8+CD28+ precursors were converted to a CD8+CD28 phenotype over 48 h, providing clear evidence that CD8+CD28+ cells include precursors of CD8+CD28 cells. Similar results were obtained in three independent experiments.

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Fig. 4. CD8+CD28+ cells are direct precursors of CD8+CD28 cells. Cycling PHA-stimulated PBL were followed by three-color staining after a 1 h BrdU pulse. (A) Three-color FACS with anti-BrdUFITC: gated BrdU or BrdU+ cells were analyzed for CD8biotin and CD28PE staining 4 h after the BrdU pulse. (B) In a further experiment, CD8+CD28+ versus CD8+CD28 subset distribution was followed over 48 h among BrdU+ CD8 cells or among all CD8 cells. Similar results were obtained in two other experiments.
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Additional evidence was obtained by sorting TCR
ß+CD8+ CD28+ cells and culturing them to generate the TCR
ß+CD8+CD28 subset de novo (Fig. 5
). Spontaneous differentiation was not observed, e.g. in medium with exogenous IL-2 or in the presence of CD4 cells. However, when allogeneic stimulator non-T cells were added to the cultures a subset of the CD8 population lost expression of CD28 (Fig. 5
). This occured with or without the presence of added CD4 cells and required the presence of CD8+CD28+ cells as the latter could not be substituted by CD8CD28+ cells (not shown). The allogeneic stimulating T-depleted PBL could be irradiated (3000 rad), or not, with similar results (not shown).
Finally, we found that gated TCR
ß+ CD8+CD28+ cells obtained ex vivo from five normal subjects were enriched in distinct FSChi lymphoblasts when compared with TCR
ß+ CD8+CD28 cells (data not shown), suggesting that the CD8+CD28+ cells contain the proliferative precursors in vivo.
CD8+CD28 cells are perforin+ cytotoxic effectors
Stimulation of a T cell via the TCR and CD28 results in T cell responses such as proliferation, IL-2 secretion and activation of cytolytic effector function from memory pCTL (18). Given that CD8+CD28 cells were the progeny of CD8+CD28+ cells, we asked whether CD8+CD28 contained intracytoplasmic perforin granules. Cells permeabilized with saponin 0.1% were stained for CD8, CD28 and intracellular perforin. The CD28+ cells, which include TCR
ß+ CD4 and CD8 cells, were mostly negative for perforin staining (Fig. 6b and d
). In contrast the majority of CD28 cells were perforin+. Both CD8+CD28 and CD8CD28 subsets contained perforin+ cells. There were two intensities of perforin staining, a high-intensity cell type predominant in the CD8CD28 subset and an intermediate-intensity cell type predominant in the CD8+CD28 subset. The perforinint cells were TCR
ß+ but the perforinhi cells were TCR
ß TCR
, CD56+, CD16+ NK cells (Fig. 6ad
and data not shown).
Perforin+ CD8+CD28 TCR
ß+ were most prominent in subject 15. Most of the perforin+ CD8+ cells had lost expression of CD28 in this subject. By contrast, perforin+ CD8+CD28 were less prominent in other adults (Figs 6, 7 and 9

) and completely missing in cord blood (Fig. 6
), consistent with other results (13,31). The perforin+ cells in normal cord blood had a CD8CD28 or CD8loCD28 phenotype and were invariably TCR
ßTCR
NK cells (Fig. 6e and f
, and data not shown). CD8+CD28+ cells containing low levels of perforin were uncommon, but clearly present (Fig. 6d and 7
). It is possible that these are direct precursors of CD28 perforin+ cells.
The size of the various CD28, CD8 and perforin defined subsets was examined in IVSA patients, that were either HIV+ or HIV, and compared with normal young and old subjects, and with subject 15 (Fig. 7
). HIV+ IVSA patients had increased levels of both CD8+CD28 and CD8+CD28+ cells compared with normal subjects or with HIV IVSA patients. On average 55% of CD8+CD28 cells contained perforin in all groups, including HIV+ patients, but very few CD8+CD28+ cells were perforin+. Unlike the HIV+ patients, subjects 15, LB and Met43 had expansions of CD8+CD28, but not CD8+CD28+ cells. Moreover, a larger fraction of the CD8+CD28 cells was perforin+. In these subjects there is a specific accumulation of perforin+ CD8+CD28 cells.
Expanded CD8+CD28 T cells are relatively resistant to apoptosis
A frequently used mechanism to limit the duration of an immune response is activation-induced cell death (AICD). Since clonally restricted CD8+CD28 T cells persist in vivo for years, we asked whether CD8+CD28 cells might be inherently resistant to apoptosis. Annexin V staining was performed in cultures of T cells stimulated with the superantigen SEE, which targets Vß8 T cells. To check that efficient SEE stimulation had occurred, percentages of Vß8+ cells were measured. They increased from 25 to 1015% over the culture period. Both Vß8+ CD8+CD28+ and Vß8+ CD8+CD28 T cells expanded during these cultures in relative percentages and in absolute cell numbers (Fig. 8
). The percentage of annexin V+ cells was next determined in CD28+ and CD28 Vß8 T cells, and compared with bystander T cells not expressing Vß8 (Table 2
). Vß8+CD28+ showed the highest levels and Vß8+ CD28 showed background low levels of annexin V binding. Kinetic studies revealed that Vß8+ CD28 cells remained relatively resistant to apoptotic cell death over 13 days of culture. This was independent of the cell donor origin or the addition of exogenous IL-2. When using PBL most of the proliferative response to SEE occurs among CD4+ T cells (data not shown). CD8 cells were therefore isolated by negative sorting and stimulated with SEE. The same predominance of annexin V-binding cells among Vß8+CD28+ cells was observed in these CD8 enriched cultures (Table 2
, Experiments 3 and 4). The same results were obtained when SEE was pulsed on to BJAB antigen-presenting cells (Table 2
, Experiment 3), thus avoiding the presence of excess soluble SEE. Together, these results demonstrate that CD8+CD28 cells activated with a superantigen were less susceptible to apoptosis than their CD8+CD28+ counterparts. A likely explanation is that proliferative Vß8+ CD8+CD28+ precursor cells can have several different fates, including either apoptosis or differentiation in to Vß8+ CD8+CD28 cells that are more resistant to cell death. Similar results were obtained with another superantigen, Mycoplasma arthritidis mitogen (data not shown), which has a different specificity for Vß17+ T cells.

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Fig. 8. Expansion of Vß8 T cells in response to SEE. Percentages of Vß8 CD28+ and Vß8 CD28 cells at various time points after stimulation with SEE-pulsed, irradiated, BJAB lymphoblastoid cells, used at either a 1:10 or a 1:2 (B:T) ratio. Both bulk T cells and CD4-depleted T cells were stimulated and the cultures were supplemented with exogenous IL-2 at 40 U/ml on day 4. The efficiency of the CD4 depletion was >90%. During the 8 day observation period the total cell numbers increased ~3- to 5-fold, in particular after addition of IL-2. For annexin V staining on day 6 of this experiment, refer to Table 2 (Experiment 3).
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In these experiments with superantigens, AICD is the likely mechanism of cell death. AICD requires TCR-mediated stimulation and cell death is FasFas ligand mediated (32). To examine the spontaneous susceptibility of CD8 subsets to apoptosis, we first examined Fas (CD95) expression. In subject 15, CD8+CD28 cells express CD95, albeit at levels that are consistently 2- to 5-fold lower than those of CD8+CD28+CD95+ cells (mean channel fluorescence indicated with arrows in Fig. 9A
). The intensity of CD95 expression was also decreased in other subjects, when comparing CD8+CD28 and CD8+CD28+ cells (Fig. 9A
). A lower level of Fas expression is consistent with a relative decrease in susceptibility to apoptosis.
In subject 15 CD8 cells, but not CD4 cells, undergo spontaneous apoptosis. In freshly isolated ex vivo PBL stained immediately after isolation without in vitro culture, annexin V+ CD8+ cells were abundant in subject 15, in particular in the CD8+CD28+ subset (53%), but also to a lesser degree (24%) in the CD8+CD28 subset (Fig. 9A
). Spontaneously annexin V+ cells were seen in smaller numbers in subject 16 and were undetectable in a younger subject (Fig. 9A
). The same samples were examined for the relative presence of effector CD8 cells, demonstrating abundant perforin+ CD8+CD28 cells in subject 15 that were much more discrete in the other two subjects (Fig. 9B
). Therefore, abundant effector CD8 cells correlated with increased spontaneous annexin V-binding CD8 cells.
In vivo there is a steady state in subject 15, with maintenance of a relatively high number of CD8+ cells (~60% of all CD3+ T cells). If CD8+CD28+ cells represent the subset with proliferative self-renewal potential and >50% of this subset undergoes immediate spontaneous apoptosis ex vivo (Fig. 9A
), it follows that there must be a rapid rate of production of CD8 cells.
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Discussion
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Expansions of oligoclonal CD8+CD28 cells
It is striking that many clonal expansions of human T cells described in healthy or diseased subjects have a CD28 phenotype (2). Herein, we show variable expression of CD28 within the CD8 TCR repertoire of normal young subjects as evidence that even small Vß subsets that are not dramatically expanded may be dominated by CD8+CD28 cells and that the distribution of the CD8+CD28 phenotype is quite variable. This is consistent with the CD28 phenotype being a marker for effector T cells stimulated by random antigens in vivo. Some clones may expand dramatically leading to expanded Vß subsets (3). However, small Vß subsets may also contain many CD28 cells (Fig. 2
). In these cases, perhaps the clones undergo limited expansion, or clonal expansion is curtailed in vivo by AICD, or the clones may simply migrate in to tissues other than the blood.
In some elderly subjects there are chronic oligoclonal expansions within the CD8+CD28 subset. CD8+CD28 have shorter telomers than CD8+CD28+ cells, indicating a history of increased cell divisions (33,34). We found (i) that a large fraction of CD8+CD28+ cells is engaged in an apoptotic pathway, consistent with a high proliferation rate in this population, (ii) CD8+CD28 cells originate from CD8+CD28+ precursor cells and (iii) CD8+CD28 cells are relatively resistant to apoptosis compared with their CD8+CD28+ precursors. Therefore, CD8 cells that escape apoptosis may mature in to CD8+CD28 perforin+ effector T cells. Relative resistance to apoptosis among CD8+CD28 cells, combined with increased production of antigen-driven CD8 clones, provides the best explanation for the accumulation of CD8+CD28 clones in people like subject 15.
Which antigens might be at the origin of the CD8+CD28 clones? Non-cytopathic viruses that may be chronically or intermittently productive and yet asymptomatic in humans include the herpes viruses IVIII, HTLV-I and HTLV-II, HIV-1 and HIV-2, spumavirus, parvovirus B16, JC virus, and human endogenous retroviruses. Cytomegaloviruses in particular, but also other herpesvirus infections (35), have been shown to be associated with higher percentages of CD8+CD57+ (CD28) cells (28,36), which include CMV-specific T cells (19). HTLV-I and HTLV-II seropositive IVSA subjects are known to have CD8 cells that proliferate spontaneously ex vivo and co-express NK markers CD16/CD56 (37), a phenotype which may overlap with the CD8+CD28 subset.
Since there was no evidence of high antibody titers to the above listed viruses in subject 15 (data not shown), an alternative source for chronic antigenic stimulation was considered: an incidentally discovered renal cell carcinoma (RCC). A number of reports have described RCC-associated, serologically defined tumor antigens. Some of these appeared to be related to retroviral antigens. RCC are immunogenic and CTL may be effective in controlling tumor growth (38). In many patients in situ tumor-infiltrating lymphocytes contained oligoclonal expansions, but these were mostly not present in PBL (3941). As the CD8+CD28 expansion of subject 15 was present in the PBL several years prior to discovery of the RCC, it remains to be determined whether the CD8+CD28 cells are related to the tumor.
CD8+CD28 cells are cytotoxic effector cells with intracellular perforin
TCR
ß+CD8+CD28 cells were shown to include a population of cells with intracellular perforin, e.g. differentiated effector CTL. These cells are found in increased frequency in patients with acute and chronic viral infections, as exemplified by HIV-1 infected patients (Fig. 7
). However, in such patients we found a general predominance of all types of CD8 cells, and the normal ratios of perforin+ cells in the CD8+CD28 and CD8+CD28+ subsets were preserved. Elderly subjects without major CD8 clonal expansions had low levels of the CD8+CD28 and CD8+CD28+ subsets. However, three individuals (subject-15, LB and met-43) were identified with major expansions of CD8+CD28 cells that were predominantly perforin+ and yet the CD8+CD28+ subsets were not enlarged. This pattern clearly differs from what was seen in the chronic HIV-1-infected subjects. Thus, there are variations in differentiation patterns of CD8 cells. The data also suggest considerable heterogeneity among old subjects in the size of the CD8+CD28 subset.
CD280/0 mice can reject tumors (42), give an acute graft versus host response (43), a delayed type hypersensitivity response and a CTL response (44). Therefore it is likely that other co-stimulatory molecules compensate for the loss of CD28. CD2/LFA-3 (18) and CD27/CD70 (45) have both been shown to co-stimulate maturation of a CTL response. Interestingly, CD27 expression is lost in concert with CD28 expression when normal human CD8 cells differentiate in to effector CTL (14). Perhaps it is crucial that co-stimulation be avoided at a late stage of CD8 cell maturation.
CD8+CD28 T cells are relatively resistant to apoptosis
Loss of CD28 expression might serve to arrest a proliferative memory T cell response. AICD has been proposed as a mechanism to down-regulate a cellular immune response (32), and some authors claim that CD8 effector T cells are `apoptosis-prone' (46) and lose expression of Bcl-2 (35). However, Kaneko et al. found that CD8+CD28+, but not CD8+CD28 T cells, underwent apoptosis (47), using cells from patients with systemic lupus erythematosus and from normal controls. There were no differences in Fas and Bcl-2 expression, but CTLA-4 was only expressed in the CD8+CD28+ cells. Consistent with these data, we found CTLA-4 was not expressed on the CD8+CD28 cells of subject 15, but only on CD8+CD28+ cells (data not shown). Thus CD8+CD28+ cells appear to be the subset that is tightly regulated and subject to AICD or CTLA-4-mediated down-regulation. These results agree with our findings of decreased apoptosis in the CD8+CD28 relative to the CD8+CD28+ subset.
Zhang et al. (48) described murine antigen-specific CD8 T cells that escape AICD in vivo, in spite of the continued presence of antigen. These cells are anergic. They express activation antigens, including Fas and Fas ligand, and produce Tc2 cytokines (IL-4 and IL-10). The resistance of Tc2 cells to apoptosis has been studied in perforin0/0 mice, where IFN-
-producing Tc1, but not IL-5-secreting Tc2 CD8 cells, underwent Fas-mediated cytolysis (49). Moreover, Tc2 cells may express CD30 and produce soluble CD30 (49,50) which could interfere with CD30-mediated apoptosis. However, no expression of CD30 was found on the CD8+ T cells from subject 15 (data not shown).
A major requirement for apoptosis is a cycling cell (18,51). CD28 co-stimulation and the presence of IL-2 are important in advancing T cells through the cell cycle. Although CD28 co-stimulation is thought of as anti-apoptotic, due to stimulation of increased levels of Bcl-XL (52), this effect may not apply to AICD (53). Indeed, in some cells a CD28 co-stimulatory signal may facilitate cell death, as in double-positive thymocytes undergoing clonal deletion (54). It would seem important for a cytotoxic effector cell to avoid apoptosis. It undergoes repetitive antigen exposure when it serially kills one target cell after another (55) and repeated antigen exposures often result in AICD (32). Perhaps this is the reason for the loss of expression of CD28 and CD27 in cytotoxic effector cells.
 |
Acknowledgments
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We thank Mr M. Filomio for technical assistance, Dr Aaron Wells and the staff of the New York Hospital Adult Clinic, and the patients and normal volunteers for their support. Supported in part by PHS grant AI22333 and a NIDA supplement.
 |
Abbreviations
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AICD | activation-induced cell death |
CMV | cytomegalovirus |
CTL | cytotoxic T lymphocyte |
FSC | forward scatter |
HAART | highly active anti-retroviral therapy |
IVSA | i.v. substance abuse |
PBL | peripheral blood lymphocyte |
pCTL | precurosr CTL |
PE | phycoerythrin |
PHA | phytohemagglutinin |
RCC | renal cell carcinoma |
SEE | staphylococcal enterotoxin E |
SSC | side scatter |
 |
Notes
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Transmitting editor: A. Singer
Received 6 April 1998,
accepted 22 October 1998.
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