International Immunology

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International Immunology, Vol. 12, No. 7, 1005-1013, July 2000
© 2000 Japanese Society for Immunology

Loss of CD28 expression on CD8⁺ T cells is induced by IL-2 receptor chain signalling cytokines and type I IFN, and increases susceptibility to activation-induced apoptosis

Nicola J. Borthwick^1,4, Mark Lowdell², Mike Salmon³ and Arne N. Akbar¹

¹ Departments of Clinical Immunology and
² Haematology, Royal Free and University College Hospital Medical Schools, London NW3 2PF, UK
³ Department of Rheumatology, Birmingham University, Birmingham B15 2TJ, UK
⁴ Immunology Unit, Department of Infectious and Tropical Diseases, London School of Hygiene and Tropical Medicine, London WC1E 7HT, UK

Correspondence to: N. J. Borthwick, Immunology Unit, Department of Infectious and Tropical Diseases, London School of Hygiene and Tropical Medicine, Keppel Street, London WC1E 7HT, UK

	Abstract

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CD8⁺CD28^– T cells are selectively expanded during viral infections, indicating their importance in anti-viral immune responses. Since little is known about the differentiation of CD8⁺CD28^– cells, we investigated the generation, function and survival characteristics of this subset. In healthy individuals CD8⁺CD28^– T cells contained more elevated levels of perforin and IFN- than the CD8⁺CD28⁺ subset, indicating that they can have an effector function. CD8⁺CD28^– cells were selectively expanded when activated CD8⁺CD28⁺ T cells were cultured in IL-2, IL-7 or IL-15. Moreover, the generation of CD8⁺CD28^– cells was accelerated by type I IFN suggesting that these cytokines which are released during viral infections influence CD8⁺ T cell differentiation. We did not observe re-expression of CD28 by CD8⁺CD28^– T cells in any of the experiments performed. Activated T cells are susceptible to activation-induced cell death (AICD) if re-stimulated in the absence of co-stimuli. AICD was induced in both CD28⁺ and CD28^– subsets of activated T cells when stimulated with anti-CD3 antibody in the absence of co-stimuli but the magnitude of death was greater in the CD28^– subset. While co-stimulation through LFA-1 (CD11a and CD18) significantly reduced AICD in the CD8⁺CD28⁺ subset, death was not prevented in CD8⁺CD28^– cells. These results suggest that CD8⁺CD28^– T cells are more functionally differentiated than the CD8⁺CD28⁺ subset and indicate they may represent a terminally differentiated effector population which is destined for clearance by apoptosis at the end of the immune response.

Keywords: activation-induced apoptosis, CD8, CD28, cytotoxic T lymphocytes, type 1 interferon

	Introduction

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Immune responses to viral infections involve the proliferation and differentiation of CD8⁺ T lymphocytes into an effector population that controls viral replication. The expansion of effector cells is accompanied by an increased sensitivity to apoptosis, which regulates proliferation and maintains lymphocyte homeostasis (1,2). Nevertheless, a small proportion of activated CD8⁺ T cells survive, forming the memory population. Although the precise relationship between CD8⁺ effector and memory T lymphocytes is not known, determining the signals that result in effector cell generation and the mechanisms that control clearance of these cells are important for understanding CD8 memory.

The T cell antigen CD28 is known to provide important co-stimulatory signals that facilitate activation through the TCR, regulating proliferation and preventing the induction of anergy or apoptosis (3). In humans, the majority of resting CD4⁺ T lymphocytes express this marker; however, a small proportion, ~25%, of CD3⁺CD8⁺ T cells do not (4). Viral infections are associated with an expansion of CD8⁺CD28^– cells. In particular, this subset is increased during HIV-1 infection (5,6) where high proportions are associated with disease progression (7). Increased proportions of CD8⁺CD28^– T cells are also found during Epstein–Barr virus (EBV)-induced infectious mononucleosis (EBVIM) (4), and have similarly been reported in acute graft versus host disease (8), in patients with common variable immunodeficiency (9) and in Chagas disease induced by the Leishmania parasite (10).

The function of CD8⁺CD28^– T cells remains controversial, but their association with disease suggests that they are an effector population. In HIV-1-infected individuals CD8⁺CD28^– T cells were shown to mediate HIV-1-specific cytotoxicity (11), although cytotoxic T lymphocyte (CTL) activity has also been demonstrated in the CD8⁺CD28⁺ subset (12). The CD8⁺CD28^– subset can also have a suppressor function as clones with a CD8⁺CD28^– phenotype generated from the gingiva were found to suppress proliferation (13) and, similarly, allospecific suppressor cells can be generated from CD8⁺CD28^– cells following multiple mixed lymphocyte reaction stimulations (14).

The factors controlling the generation of CD8⁺CD28^– T cells are not clear. It is known that long-term maintenance of T cell lines in IL-2 results in an accumulation of this subset (15); however, it is unclear whether this is due to the preferential expansion of an existing subset or to differentiation from CD28⁺ cells. Similarly, it is not known if other cytokines can induce this phenotypic change and whether this is reversible. The role of cell proliferation on the loss of CD28 expression is unknown. In particular type I IFN have been shown to augment the induction and differentiation of virus-specific CTL (16,17) but their precise role in CD8⁺ T cell differentiation and the loss of the CD28 antigen expression has not been determined.

In this study we have investigated the in vitro generation, reactivation and apoptosis of CD8⁺CD28^– T cells. Our observations indicate there is a lineage relationship between CD8⁺CD28⁺ and CD8⁺CD28^– lymphocytes, and that loss of the CD28 antigen identifies more differentiated cells that are likely to be removed by apoptosis.

	Methods

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Lymphocyte purification and activation
Venous blood from healthy laboratory personnel was taken into preservative-free heparin and peripheral blood mononuclear cells (PBMC) isolated by density gradient centrifugation. Plastic adherent cells were prepared by incubating PBMC in medium containing 20% FCS at 37°C for 2 h. Purified T lymphocytes were prepared by depletion of monocytes, NK cells and B cells using CD14 (RM052, IgG1; Immunotech, Marseilles, France), CD16 (RG8, IgG1; Immunotech) and CD19 (RFB9, IgG1; Royal Free Hospital) respectively, together with goat anti-mouse (GAM)-coated Dynabeads (Dynal, Bromborough, UK). CD8⁺ T cells were prepared by removal of CD4⁺ T cells using RFT4 (IgG1; Royal Free Hospital) plus Dynabeads. In some cases, purified CD28⁺ and CD28^– subsets were prepared from activated lines by cell sorting after staining with a CD28–phycoerythrin (PE) conjugate (L293, IgG1; Becton Dickinson, Oxford, UK) using a Vantage flow cytometer (Becton Dickinson). PBMC and purified T cells were cultured in RPMI 1640 medium supplemented with 10% FCS, 100 IU/ml penicillin, 100 µg/ml streptomycin and 2 mM L-glutamine (Gibco/BRL, Paisley, UK).

IL-2-dependent T cell lines were prepared from PBMC after activation with phytohaemagglutinin (PHA-P; 0.5 µg/ml; Murex Biotech, Dartford, UK) or anti-CD3 (OKT3, IgG2a, 1 µg/ml; ATCC, Rockville, MD) for 5 days. The cells were then washed and re-cultured at 10⁶/ml in medium containing 5 ng/ml recombinant IL-2 (R & D Systems, LOCATION??) and could be maintained in this way for 4–6 weeks. Only lines that had been in IL-2 for >1 week were used in apoptosis studies. CD8⁺ T cell lines were prepared by activation of purified CD8⁺ T cells in the presence of 10% autologous plastic adherent cells followed by maintenance in IL-2 as before. In some cases, PBMC were stimulated with soluble anti-CD3 in the presence or absence of IFN- and/or IFN-ß (10 ng/ml; PeproTech, London, UK). Before the start of all experiments involving T cell lines, the cell cultures were Ficolled to remove dead cells and debris, and were >95% viable prior to culture under experimental conditions.

Cytotoxicity and proliferation assays
CTL were measured in a redirected killing assay using the JAM test (18) which measures DNA fragmentation and cell death. Target cells were the mouse line P815 which were labelled overnight with [³H]thymidine (5x10⁶ P815/ml, 5 µCi [³H]thymidine/ml). Target cells were washed and incubated with effectors at a range of ratios for 5 h in the presence of 5 µg/ml anti-CD3 mAb (OKT3; IgG2a; ATTC). Plates were harvested using a Filtermate 196 cell harvester (Canberra Packard, Pangbourne, UK) and counted in a Topcount scintillation counter (Canberra Packard). Killing was calculated using the following formula: percent specific lysis = (S – E)/Sx100, where S = counts from targets cultured without effectors for 5 h and E = counts from experimental wells. Lymphocyte proliferation was assayed in CD28⁺ and CD28^– subsets after stimulation with plate-bound anti-CD3 (10 µg/ml; UCHT1; IgG1; Professor P. Beverley Edward Jenner Institute for Vaccine Research, Newbury, UK) in the presence or absence of anti-CD18 (10 µg/ml, L130, IgG1; Becton Dickinson). Cells were stimulated for 3 days and harvested and counted as for the JAM assay.

Activation-induced cell death (AICD)
IL-2-dependent T cell lines were washed, Ficolled to remove dead cells and then re-cultured at 1x10⁶/ml in the presence or absence of plate bound anti-CD3 (UCHT1, IgG1, 10 µg/ml). Antibodies against the co-stimulatory molecules CD28 (Kolt2, IgG1; Dr Kimitaka Sagawa, Department of Immunology, Kurume University School of Medicine, Kurume, Japan), CD5 (UCHT2, IgG1; Professor P. Beverley), CD11a (MHM24, IgG1; Professor A. McMichael, John Radcliffe Hospital, Oxford) and CD18 (L130, IgG1; Becton Dickinson) were added at pre-titrated optimal concentrations. Cultures were assayed for apoptosis as indicated below.

Enumeration of viable cells and cells in apoptosis
Viable cells were distinguished by their forward angle scatter and 90° side scatter profiles, and were counted using a Cytoron Absolute flow cytometer (Ortho Diagnostics, High Wycombe, UK). Cells in apoptosis were identified by the characteristic sub-G₀/G₁ DNA peak after staining with propidium iodide (PI; Sigma, Poole, UK). Briefly, cells were permeabilized with 70% ice-cold ethanol and stored at –20°C for up to 1 week prior to analysis. The ethanol was washed off and the cells stained with 5 µg/ml PI in PBS containing 50 ng/ml RNase (Sigma) for 30 min at room temperature prior to analysis. Additionally, apoptotic cells were identified by their morphological appearance under light microscopy following staining of cytospin preparations with May–Grunwald Giemsa.

Phenotypic analyses
The following mAb were used in double and triple combinations for flow cytometric analyses. CD3 (OKT3, IgG2a), CD4 (RFT4, IgG1) and CD8 (RFT8 IgG1 and IgM) recognize T lymphocytes, helper and suppressor/cytotoxic subsets respectively. Lymphocytes were also investigated for their expression of CD28 (Becton Dickinson). Antibodies were either directly conjugated to FITC, PE or PE–Cy5 or were used indirectly with isotype-specific second layers conjugated to either FITC or PE (Southern Biotechnology Associates, Birmingham, AL). The expression of perforin in CD8⁺CD28⁺ and CD8⁺CD28^– subsets was measured by flow cytometry. PBMC and T cell lines were permeabilized using Leucoperm (Serotec, Oxford, UK), following the manufacturers instructions and stained using the combination CD8–PE–Cy5 (Dako, Cambridge, UK), CD28–PE (Becton Dickinson) and anti-perforin (clone G9, IgG2b; PharMingen, Oxford, UK) plus goat anti-mouse IgG2b–FITC (Southern Biotechnology Associates). Fluorescence was measured using a FACScan flow cytometer (Becton Dickinson) and data analysed using CellQuest software. IFN- production was measured in lymphocytes activated with phorbol myristate acetate (PMA, 5 ng/ml; Sigma) and ionomycin (1 µM; Sigma) for 4 h in the presence of monensin (3 µM; Calbiochem-Novabiochem, Nottingham, UK). Cell cultures were harvested, permeabilized with Leucoperm (Serotec), and stained using the combination CD8–PE–Cy5 (Dako), CD28–PE (Becton Dickinson) and IFN-–FITC (Cambridge-Bioscience, Cambridge, UK).

	Results

Top Abstract Introduction Methods Results Discussion References

 
Functional characteristics of CD8⁺CD28⁺ and CD8⁺CD28^– T cells
Healthy individuals show only a small proportion (25%) of CD28^– T cells within the CD8⁺ subset in peripheral blood. To investigate if these resting populations had defined functional repertoires, peripheral CD8⁺ T cells from healthy adults were examined for their expression of perforin, indicative of CTL function and the ability to produce IFN- upon in vitro stimulation. In the individuals examined (n = 6) 20.1 ± 6.3% of resting CD8⁺ T cells contained perforin. Interestingly, a greater proportion of CD8⁺CD28^– T cells contained this protein although it could also be detected within the reciprocal CD8⁺CD28⁺ subset (CD8⁺CD28⁺ 11.8 ± 3.4%; CD8⁺CD28^– 33.5 ± 8.6%; P < 0.02; Fig. 1a). This supports previous findings in which CTL activity of PBMC was found to be higher in the CD8⁺CD28^– subset (19). A similar pattern was observed when IFN- production was examined (CD8⁺CD28⁺ 13.3% ± 2.0%; CD8⁺CD28^– 23.5 ± 3.7%; P < 0.007; Fig. 1b). Both perforin and IFN- expression were also investigated in polyclonally generated CD8⁺ T cell lines. In these activated cells it was again clear that perforin expression was more restricted to the CD28^– subset, while IFN- was produced by both CD28⁺ and CD28^– cells (Fig. 1c and d).

View larger version (46K): [in this window] [in a new window]   Fig. 1. FACScan profiles showing examples of perforin and IFN- staining in CD8+CD28+ and CD8+CD28– subsets. Resting PBMC (a and b) and a T cell line were triple stained with CD8, CD28 and either perforin (a and c) or IFN- (b and d) as outlined in Methods. Profiles show expression within cells brightly staining with anti-CD8 that fell within the viable cell FSC and SSC scatter profiles. Numbers refer to the percentage of cells that fell within each quadrant.

 
Interestingly, while a greater proportion of activated CD28^– cells expressed perforin, this was not reflected in their ability to kill targets in a redirected killing assay. Instead, when cell lines were sorted for CD28 expression, both CD8⁺CD28⁺ and CD8⁺CD28^– subsets showed similar levels of killing (donor 1 cytotoxicity at E:T = 25:1 CD28⁺ 19.7 ± 0.5%, CD28^– 24.4 ± 2.03; donor 2 CD28⁺ 47.9 ± 2.5, CD28^– 50.1 ± 4.0; donor 3 CD28⁺ 56.4 ± 5.8, CD28^– 50.3 ± 2.4). The cytotoxic activity in the CD28^– subset may occur as a result of perforin-independent mechanisms of CTL killing, but nevertheless this data suggests that after activation in vitro both CD28⁺ and CD28^– subsets have effector function both in terms of CTL killing and IFN- production.

Differentiation of CD8⁺CD28^– T cells
As the proportion of CD8⁺CD28^– cells is increased in T cell lines we next investigated factors influencing the differentiation of this subset. We observed that polyclonal stimulation of CD8⁺ T cells with either PHA or anti-CD3 induced an early up-regulation of CD28 antigen expression on day 3 that subsequently declined during culture in IL-2. Using PHA as the mitogen (Fig. 2) this switch had occurred in 66.9 ± 4.5% of CD8⁺ cells by day 14, but was slightly slower using anti-CD3 as the initial stimulus (% CD8⁺CD28^– cells on day 14; 40.6 ± 1.5%). After 4 weeks in culture, however, the majority of these anti-CD3-stimulated cells had lost CD28 expression (% CD8⁺CD28^– cells on day 28; 89.3 ± 3.3%). It is apparent that the long-term culture of CD8⁺ T cells in IL-2 results in an accumulation of CD28^– cells; however, it was not clear whether these cells differentiated from the CD8⁺CD28⁺ subset or arose due to the expansion of existing CD8⁺CD28^– cells. To examine this, cell lines cultured in IL-2 for 7 days were sorted into CD8⁺CD28⁺ and CD8⁺CD28^– subsets. These were then re-cultured for up to 3 weeks in IL-2 or other cytokines that signal through the common chain of the IL-2 receptor to determine the effect on CD8⁺ T cell expansion and CD28 expression. Cell separations using a FACS cell sorter were necessary as cross-linking CD28 with antibody on magnetic beads down-regulates CD28 antigen expression. Both the intensity of CD28 expression and the percentage of CD28⁺ cells gradually declined with time when the sorted CD28⁺ cells were cultured in the presence of either IL-2, IL-7 or IL-15 (Fig. 3a, c and d). In contrast, cells cultured in IL-4 retained CD28 expression (Fig. 3b). None of these cytokines induced the re-expression of CD28 in the CD8⁺CD28^– sorted population (data not shown). Both CD8⁺CD28⁺ and CD8⁺CD28^– subsets were observed to expand when cultured in the presence of IL-2, IL-7 or IL-15 and survived without significant expansion in the presence of IL-4, suggesting that proliferation might be important in inducing the loss of the CD28 antigen. To examine this further, day 3 PHA blasts were cultured long-term in IL-2, IL-4 or IL-2 + IL-4 and CD28 expression determined over time. Cells cultured in IL-4 alone expanded less well and by day 14 virtually all the cells were dead. In contrast, cells maintained in IL-2 or IL-2 + IL-4 expanded and survived beyond day 27. Both of these cultures demonstrated a similar loss of CD28 expression (% CD8⁺CD28^– cells on day 14; IL-2 70.6%, IL-2 + IL-4 67.4%). It appears, therefore, that the maintenance of CD28 expression in the presence of IL-4 was not due to the effects of this cytokine per se. Instead, cell division induced by IL-2 (or IL-15 or IL-7) is a pre-requisite for the acquisition of the CD8⁺CD28^– phenotype, supporting the hypothesis that CD28^– cells differentiate from CD28⁺ in response to these cytokines.

View larger version (12K): [in this window] [in a new window]   Fig. 2. Loss of CD28 expression during maintenance of T cell lines in IL-2. PBMC from healthy adults were isolated and the CD3+CD8+ population investigated for CD28 expression using the triple combination CD3–FITC, CD8–PE/Cy5 and CD28–PE. The bars represent the mean ± SEM percentage of CD8+CD28– cells within the CD3+, lymphocyte gate at the time points shown (n = 6).

 

View larger version (33K): [in this window] [in a new window]   Fig. 3. Effect of chain signaling cytokines on CD28 expression. CD8+CD28+ T cells were prepared by cell sorting from a T cell line that had been maintained in IL-2 for 7 days (open histogram). The cells were then cultured in either IL-2 (a), IL-4 (b), IL-7 (c) or IL-15 (d) for a further 7 days and re-investigated for CD28 expression (shaded histogram). The cells sorted for CD28 expression were 8.7% CD28– at the start of the experiment. The numbers refer to the percentage of CD28– after culture in the presence of each of the cytokines. Log10 fluorescence is shown. Representative experiment illustrated (n = 3).

 
The increase in CD8⁺CD28^– T cells that occurs in vivo during viral infections is likely to be influenced by a number of factors, however; one important feature of viral infections is the release of type I IFN by the infected cells (20). We therefore investigated the effect of IFN- and IFN-ß on T lymphocyte proliferation and CD8 phenotype during stimulation with anti-CD3 and subsequent maintenance in IL-2. We found both IFN- and IFN-ß accelerated the appearance of the CD8⁺CD28^– population compared to anti-CD3 alone. This was apparent after anti-CD3 activation for only 3 days when the initial up-regulation of CD28 was diminished if IFN was included. Thereafter, the proportions of CD8⁺CD28^– T cells were increased in cultures supplemented with IFN- or IFN-ß compared to IL-2 alone (Fig. 4). Thus the presence of type I IFN may accelerate the generation of CD8⁺CD28^– effector cells.

View larger version (17K): [in this window] [in a new window]   Fig. 4. Accelerated loss of CD28 expression by type I IFN. The loss of CD28 expression was investigated in CD8+ T cells activated and maintained in IL-2 in the presence of IFN- or -ß. (a) Time course showing the percentage CD8+CD28– T cells after 3 days stimulation with anti-CD3 (start) and after subsequent culture in either IL-2 alone (open bar), IL-2 plus IFN- (shaded bar) or IL-2 plus IFN-ß (filled bar) for the time points indicated. Representative experiment illustrated (n = 4). FACScan profiles of CD28 expression in CD3+CD8+ T cells after (b) IL-2 alone, (c) IL-2 plus IFN- and (c) IL-2 plus IFN-. Percentages are the proportion of cells within the marker region, M1, set using negative control reagents. Cells on day 14 are shown.

 
Differential survival of CD8⁺CD28⁺ and CD8⁺CD28^– T lymphocytes
Both the CD8⁺ T cells in the peripheral blood of EBVIM patients and in vitro generated T cell lines are highly susceptible to spontaneous apoptosis when cultured in the absence of appropriate growth factors (21). We have previously shown that this spontaneous apoptosis can be prevented by the cytokines IL-2, IL-7 and IL-15 (21,22). To determine the extent of spontaneous apoptosis in CD8⁺CD28⁺ and CD8⁺CD28^– subsets, we measured cell recovery after withdrawal of IL-2 in CD8⁺ T cell lines immunophenotyped for CD28 expression. Both subsets were equally susceptible to spontaneous apoptosis as reflected by the decrease in cell recovery after 48 h (CD8⁺CD28⁺ 21.9 ± 7.7%; CD8⁺CD28^– 29.0 ± 7.5%; n = 3). The addition of IL-2 prevented apoptosis in both CD28⁺ and CD28^– subsets.

The CD8⁺ T cells from both EBVIM patients and in vitro generated T cell lines are also susceptible to AICD following stimulation through the TCR (22). We therefore examined the ability of CD8⁺CD28⁺ and CD28^– T cells to be re-stimulated in vitro with anti-CD3 antibody. Activation of CD8⁺ T cell lines for 24 h with anti-CD3 alone induced a marked decrease in viable cell recovery to levels below those seen after IL-2 withdrawal (IL-2 withdrawal 66.8 ±9.2%; anti-CD3 47.4 ± 6.2%; n = 8; P < 0.004). Death was blocked when a CD95 neutralizing antibody was added, indicating the involvement of this molecule in the death process (% apoptosis after 24 h; CD3 58.3%, CD3 + blocking anti-CD95 antibody 16.2%, CD3 + control antibody 52.6%).

The outcome of stimulation through the TCR is in part regulated by co-stimulatory signals given by accessory molecule–ligand interactions. To determine if co-stimulatory signals could prevent AICD, CD8⁺ T lines were activated in the presence of a number of antibodies known to be co-stimulatory for anti-CD3-induced activation of resting T lymphocytes. When used in the absence of anti-CD3, none of these antibodies had any effect on cell survival after IL-2 withdrawal or during maintenance in IL-2. Co-stimulation through CD2, CD5 and CD28 did not prevent AICD as measured after 48 h; however, using antibodies directed against either the or ß chains of LFA-1 did increase the number of viable cells recovered (Fig. 5a). A time course of cell recovery after anti-CD3 activation with or without co-stimuli (Fig. 5b), showed that the initial decrease in cell recovery due to apoptosis occurred within the first 2 days after activation. Provided IL-2 was added to the cultures, this was followed by increased cell numbers presumably due to expansion of the residual cells (Fig. 5b). Cultures in which cells were co-stimulated with either anti-CD11a or anti-CD18 showed increased cell expansion at later time points compared to cells stimulated with anti-CD3 alone (Fig. 5b).

View larger version (21K): [in this window] [in a new window]   Fig. 5. Effect of co-stimulation on anti-CD3-induced cell death of CD8+ T cell lines. (a) T cell lines were stimulated with immobilized anti-CD3 in the presence of antibodies directed against the molecules indicated. The proportions of viable cells relative to initial input were determined by flow cytometry after 48 h. The survival above background (48.1 ± 9.54%, mean ± SEM) is illustrated. The effect of co-stimulation on cell survival was compared to survival after anti-CD3 alone using Student's t-test; n = 6, *P < 0.02, **P < 0.003. (b) Time course of lymphocyte recovery following stimulation of T cell lines with anti-CD3 alone (A), anti-CD3 + anti-CD28 (B), anti-CD3 + anti-CD11a (C) or anti-CD3 + anti-CD18 (D). After 3 days cells were washed and re-cultured at 106/ml in medium plus IL-2. Data shown are the percentages of cells recovered relative to initial input (100%). Data points are the mean of duplicate wells.

 
In order to examine AICD in CD28⁺ and CD28^– subsets these cells were sorted from CD8⁺ T cell lines that had been maintained in IL-2 for 7 days. As shown in Fig. 6(a) in a representative experiment, stimulation with anti-CD3 induced AICD in both subsets; however, this was more marked in cells lacking CD28. As seen previously in unsorted CD8⁺ T cells, the addition of IL-2 had little effect on AICD in either subset compared to anti-CD3 alone. Co-stimulation through CD11a or CD18 enhanced cell recovery, particularly in the CD8⁺CD28⁺ subset where the cell survival was similar to cultures maintained in IL-2. Although it is possible that the presence of CD28 antibody on the positively sorted cells might enhance survival, the inability of anti-CD28 to prevent AICD (Fig. 5) suggests this is unlikely. The CD8⁺CD28^– T cells could also be re-stimulated in the presence of anti-CD11a or anti-CD18 antibodies but survival was not increased to levels observed in CD8⁺CD28⁺ cultures or in cells cultured with IL-2 alone. The increased cell recovery seen after co-stimulation through LFA-1 was again due to a decrease in the proportion of cells in apoptosis (Fig. 6b). In Fig. 6(b) apoptosis was measured by morphological criteria and numbers appear small as they do not include the cells that have undergone apoptosis and then gone into secondary necrosis and fragmented. The patterns of cell survival and apoptosis are nevertheless largely reciprocal. The observed increase in cell survival within the CD28⁺ subset following co-stimulation through LFA-1 was reflected when the proliferative responses of the subsets were investigated. Stimulation of sorted CD28⁺ cells with anti-CD3 induced low-level proliferation which was increased if anti-CD18 antibodies were included in the cultures (CD8⁺CD28⁺, c.p.m. day 3; IL-2 2700, CD3 26,500, CD3 + CD18 68,800). The reciprocal CD28^– subset showed a lower response to anti-CD3 but co-stimulation did enhance the response (CD8⁺CD28^–, c.p.m. day 3; IL-2 2000, CD3 8300, CD3 + CD18 33,800). No re-expression of CD28 was observed on CD28^– cells even upon co-stimulation; however, maintenance of CD8⁺CD28⁺ cells in IL-2 after stimulation once again led to down-regulation of CD28 expression. Thus, re-stimulation causes marked apoptosis particularly in CD8⁺CD28^– T cells, while CD8⁺CD28⁺ T cells can more easily be re-activated particularly in the presence of appropriate co-stimuli.

View larger version (20K): [in this window] [in a new window]   Fig. 6. AICD in CD8+CD28+ and CD8+CD28– subsets. CD8+ T cell lines were sorted into CD28+ (open bar) and CD28– (filled bar) subsets and re-cultured as indicated. (a) The proportions of viable cells relative to initial input were determined by flow cytometry after 24 h. (b) Cells in apoptosis were detected by their characteristic morphological appearance after May–Grunwald Giemsa staining of cytospins. A minimum of 500 cells per sample was counted by two independent investigators. Representative experiment illustrated (n = 3).

 

	Discussion

Top Abstract Introduction Methods Results Discussion References

 
The differentiation of CD8⁺ effector and memory cells remains unclear. In this study we investigated CD8⁺CD28^– T cells, a subset that is particularly associated with acute and chronic viral infections. This subset has an effector function as determined by perforin expression, IFN- production and cytotoxic activity. We have shown that CD8⁺CD28^– T cells differentiate from the CD28⁺ subset—a phenomenon that could not be reversed. The CD8⁺CD28^– subset was prone to apoptosis unless maintained in either IL-2, IL-15 or IL-7 and was poorly re-stimulated through the TCR. This suggests that loss of CD28 marks end-stage effectors and indicates that less-differentiated cells, including early effectors and naive cells, retain CD28 expression.

We have previously observed in EBVIM patients that CD8⁺CD28^– T cells have other phenotypic features indicative of activation in vivo such as expression of CD45RO and loss of CD45RB, and high levels of HLA-DR, LFA-1 (CD11a and CD18), CD38 and CD44 (4,23 and N. Borthwick unpublished observations). Some of these markers are also seen on resting CD8⁺CD28^– T cells from healthy individuals; however, these resting cells additionally express CD57, a marker not increased on CD8⁺ T cells during acute EBVIM (N. Borthwick, unpublished observations). We found that the majority of CD8⁺CD28^– T cells from healthy people contained perforin and, in addition, some of these cells could also synthesize IFN-, indicating that this population contains cells with effector function. This agrees with other studies on CD8⁺ cells from healthy subjects in which CTL activity in a redirected killing assay was found mainly in the CD28^– subset (19). This indicates that CD8⁺CD28^– T cells are effector cells perhaps retained from a previous infection(s) and indeed the acquisition of CD57 is generally associated with more chronic infections such as HIV-1 (24). Additionally, CD8⁺CD28^–CD57⁺ T cells show evidence of oligoclonal expansion (25,26) accompanied by a decrease in telomere length (27), indicating the cells have previously encountered antigen and undergone clonal expansion.

Until recently, the lineage relationship of CD8⁺CD28⁺ and CD8⁺CD28^– subsets were not clear. We have demonstrated in vitro that loss of CD28 antigen expression on CD8⁺ T cells occurs when TCR-stimulated cells are maintained in the chain signalling cytokines IL-2, IL-7 or IL-15, each of which induced proliferation. In addition cell-sorting experiments confirmed that CD28^– cells differentiate from the CD28⁺ in response to these stimuli. Studies in HIV-1⁺ patients looking at the complementary-determining region 3 of the TCR have also suggested that CD8⁺CD28⁺ and CD8⁺CD28^– cells are phenotypic variants of the same lineage (28). These observations strongly suggest that CD8⁺CD28^– cells differentiate from the CD8⁺CD28⁺ subset following activation and cell division.

We showed that type I IFN accelerated the loss of CD28 on CD8⁺ T cells. These cytokines are required for the induction and differentiation of both virus- and tumour-specific CTL (17,29,30). In most cases IFN is thought to act indirectly via a secondary mediator, e.g. in mice the proliferation of CD8⁺ memory cells in vivo induced by type I IFN is thought to occur indirectly through the release of IL-15 (31). Our studies show that the presence of type I IFN promotes the acquisition of an end-stage effector phenotype. Furthermore, recent data suggests that type I IFN can also provide an anti-apoptotic signal to T cells in vivo (32), and can prevent apoptosis following IL-2 withdrawal and anti-CD95-induced death in vitro (33,34). This indicates that type I IFN released during viral infections may promote both the generation and long-term persistence of effector cells following initial antigen-specific stimulation.

The CD8⁺ effector cells that arise during acute viral infections are remarkably sensitive to apoptosis (21). Nevertheless, following acute infections a memory response is retained, suggesting that signals preventing the death of effectors are important in the generation of memory. Both CD8⁺CD28⁺ and CD8⁺CD28^– subsets were equally dependent on cytokines to prevent spontaneous apoptosis. This is perhaps not unexpected as we have previously shown in EBVIM and HIV-1⁺ patients that both subsets show a similar decrease in Bcl-2, an anti-apoptotic molecule that is important in the regulation of T cell apoptosis due to growth factor withdrawal (4). Both subsets were also susceptible to AICD, a phenomenon that is observed when activated cells in cell cycle are re-stimulated through the TCR in the absence of appropriate co-stimuli (35). The important differences between the subsets were that CD8⁺CD28^– cells showed an increased AICD compared to CD8⁺CD28⁺ T cells and furthermore were only poorly co-stimulated through CD18. These observations suggest that while both subsets will survive and can manifest their effector function if supplied with sufficient cytokine, the CD8⁺CD28^– subset responds less well to antigen re-encounter.

The reason for the particular sensitivity of the CD8⁺CD28^– subset to AICD is not clear. It may in part be due to the absence of the CD28 molecule itself which prevents the cells from being triggered via classical CD28/B7-1 or B7-2 co-stimulatory pathways (36). In other systems co-stimulation through this pathway has been shown to prevent apoptosis (37,38); however, we were unable to prevent AICD of T cell lines using anti-CD28, even when the cells retained CD28 expression. In contrast, antibodies directed against LFA-1 (CD11a and CD18) were able to prevent AICD particularly within the CD8⁺CD28⁺ subset. Signals through LFA-1 have been shown to co-stimulate resting T cell activation and to promote conjugate formation (39), facilitating other molecular interactions (40–42). Signals through LFA-1 have also been shown to provide anti-apoptotic signals to germinal centre B cells (43), to prevent anti-CD3-induced apoptosis of cortical thymocytes (44) and are involved in the delay of T cell apoptosis following transendothelial migration (N. Borthwick, submitted).

Direct infection of cells with a number of viruses including the herpes virus CMV causes up-regulation of ICAM-1, the ligand for LFA-1 (45). This may have a number of important consequences. Firstly, it is known that viruses such as CMV can infect endothelial cells and the up-regulation of ICAM-1 may enable effector CD8⁺ T cells to be selectively recruited to sites of viral infection. Additionally, LFA-1/ICAM-1 interactions play an important role in conjugate formation between CTL and infected target cells (46). The mechanisms whereby the re-encounter of antigen by CTL induces lysis of target rather than effector cells are poorly understood and it is possible that LFA-1 interactions may have a role to play.

Using tetrameric MHC–peptide complexes it is possible to visualize antigen-specific CD8⁺ T cells. One study investigating the phenotype of EBV-specific CD8⁺ T cells during primary infection and 3 years afterwards found that the frequency of CD8⁺CD28⁺ peptide-specific cells was unchanged during this time, while peptide-specific CD8⁺CD28^– cells were significantly decreased (47). Additionally, it has recently been shown in HIV-1⁺ patients that responses to HIV-1, the persistent viral infection, are found within the CD8⁺CD28^– subset, while recall responses in the same patients to EBV or influenza virus are found mainly in the CD8⁺CD28⁺ subset (48). This indicates that the continued presence of HIV-1 antigens drives the CD8 subset to the more differentiated end-stage effector phenotype, while true memory responses following complete resolution of infection, such as influenza, are found within the CD8⁺CD28⁺ subset. From these and our own observations we suggest that CD28 expression on CD8⁺ cells is a useful marker to distinguish late, more differentiated effectors (CD28^–) from naive, unstimulated cells and early effectors (CD28⁺). We would also hypothesize that CD8 memory cells would be found within the CD28⁺ compartment because cells that retain CD28 expression can be re-stimulated if given the appropriate co-stimuli, while CD28^– cells cannot.

	Acknowledgments

 
This work was funded by the Medical Research Council grant no. G9218555MA, the Nuffield Foundation Oliver Bird Fund grant no. RHE/96/263/G and Action Research.

	Abbreviations

 

AICD activation-induced cell death

CTL cytotoxic T lymphocyte

EBV Epstein–Barr virus

EBVIM Epstein–Barr virus-induced infectious mononucleosis

PBMC peripheral blood mononuclear cell

PE phycoerythrin

PHA phytohaemagglutinin

PI propidium iodide

	Notes

 
Transmitting editor: P. L. C. Beverley

Received 21 July 1999, accepted 13 March 2000.

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