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A Subset of CD8 Memory T Cells from Old Mice Have High Levels of CD28 and Produce IFN-γ
Clinical Immunology Vol. 104, No. 3, September, pp. 282–292, 2002 doi:10.1006/clim.2002.5221
A Subset of CD8 Memory T Cells from Old Mice Have High Levels of CD28 and Produce IFN-␥ Anavelys Ortiz-Sua´rez* and Richard A. Miller* ,† ,‡ ,㛳 ,§ ,1 *Cellular and Molecular Biology Graduate Program and †Department of Pathology, University of Michigan School of Medicine; Ann Arbor, Michigan 48109-0940; and ‡Institute of Gerontology, 㛳Geriatrics Center, and §VA Medical Center, Ann Arbor, Michigan 48105
Using carboxyfluorescein diacetate succinimidyl ester (CFSE)-tagged cells to measure proliferation in vivo, we found that only memory CD8 ⴙ cells from mice older than 18 months gave measurable levels of proliferation and that the proportion of memory CD8 ⴙ T cells able to proliferate in a nonirradiated recipient increased with age. CD8 cells that had proliferated in vivo contained higher levels of CD28 when compared to CD8 cells that had not divided. Cells with high levels of CD28 were preferentially able to divide in nonirradiated recipients. Using ex vivo intracellular staining analysis, we determined that most of the CD8 ⴙ T cells that were capable of dividing in vivo produced IFN-␥ after isolation from recipient mice or their original host. These studies thus document the presence in aged mice of a population of CD28 hi CD8 ⴙ cells whose ability to proliferate in vivo without antigenic stimulation and to produce IFN-␥ may be involved in immune regulation. © 2002 Elsevier Science (USA) Key Words: CTL; T lymphocytes; cytokines; IFN-␥ memory; spleen; lymph nodes. INTRODUCTION
Age in humans and rodents leads to immunologic defects which include deficits in the formation of highaffinity antibodies (1), generation of long-lasting immune responses after vaccination (2, 3), and expression of delayed-type hypersensitivity reactions to antigens initially encountered earlier in life (1). Immunodeficiencies associated with age appear to be due to some extent to the fact that fewer T cells from aged individuals are activated in response to mitogens as measured by proliferative capacity, calcium flux, or IL-2 receptor up-regulation (1, 4, 5). An age-dependent shift from naı¨ve T cells (predominant in young individuals) to memory T cells has also been observed (1). This could help to explain the decline in the number of functionally reactive T cells with age to mitogens in vitro and 1
To whom correspondence and reprint requests should be addressed at 5316 CCGC, 1500 E. Medical Center Dr., Ann Arbor, MI 48109-0940. E-mail: [email protected]. 1521-6616/02 $35.00 © 2002 Elsevier Science (USA) All rights reserved.
could possibly have an effect on the parallel decrease with age in the in vivo responsiveness to newly encountered antigens. Another possibility is that anergized T cells accumulate with age (6 –9). Several groups have reported that aging leads to changes in the repertoire of T cells in both mice and humans (10 –22). Older human beings and mice often develop large clones of CD8 ⫹ T cells (10 –18). These clones can represent as much as 80% of the CD8 ⫹ T cells in an individual (17). Large clones of this kind are not typically seen among CD4 ⫹ T cells, although smaller clones are often detected in the CD4 population (19 –22). CD8 ⫹ T-cell clones are detectable in almost every person over the age of 40 (16 –17). Similarly, expanded CD8 ⫹ T cells clones appear in middle-aged mice, and longitudinal studies have shown that their frequency increases rapidly after the age of 18 months (17, 18, 22, 28). It has been reported (28) that most of these mouse CD8 ⫹ T-cell clones are long lasting but nevertheless transient. Hence, the T-cell repertoire of aged mice seems highly unstable, rather than terminally altered, although infrequent stable alteration can sometimes be seen (28). These CD8 ⫹ T-cell clones show no signs of transformation to malignancy. Human and mice containing these clones have normal life spans with only rare evidence of T-cell neoplasia (11). Nevertheless, the percentage of CD8 ⫹ T-cell clones bearing particular Vs in old animals increased in comparison with other CD8 ⫹ T cells (17, 18, 25). Consequently, one can speculate that even though the clones are not overtly malignant, they have at some earlier stage made a transition to a state that allows them to proliferate in vivo more readily than other cells because of changes in cell-cycle kinetics, susceptibility to apoptotic stimuli, or some other alteration. The cause of this age-dependent skewing is not yet known. Recently, it has been shown (25) that the expanded CD8 ⫹ clones are very similar to nonclonal memory CD8 ⫹ cells in that both express similar surface molecules. Both bear high levels of CD44 and similar amounts of IL-2R. Like memory phenotype
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CD8 ⫹ T cells (23, 24), cells of the expanded clones divide slowly in nonirradiated mice by a process that is probably antigen independent, driven by IL-15, and inhibited by IL-2 (25). However, the proliferating progeny of the large CD8 ⫹ T-cell clones accumulated, in vivo, more rapidly than the progeny of nonclonal CD8 ⫹ T cells (25). Marrack and her colleagues have suggested that increased sensitivity to the stimulatory effects of IL-15 or lowered sensitive to the inhibitory effects of IL-2 contributes to the accumulation of clonally expanded CD8 T cells in older mice. The fact that old mice and humans accumulate a population of clonal T cells suggests that this population might contribute to the immunological deficits seen in elderly individuals. The experiments described in this article were designed to determine whether age alters the ability of mouse T cells to proliferate in nonirradiated hosts and to further characterize the cells that are able to divide in vivo without intentional antigenic stimulation. MATERIALS AND METHODS
Mice Specific-pathogen-free male (BALB/c ⫻ C57BL/6)F 1 (CB6F1) mice were purchased from the National Institute on Aging’s contract colonies at Harlan (Indianapolis, IN). Unless otherwise stated, “young” mice were used at 4 – 8 months of age, and “old” mice were 18 –25 months old. Mice were housed for at least 2 weeks after shipment in a specific-pathogen-free holding colony at the University of Michigan and given free access to food and water. Sentinel animals from this colony were examined quarterly for serological evidence of viral infection; all such tests were negative during the experimental period. Mice that were found to have splenomegaly or macroscopically visible tumors at the time of sacrifice were not used for any experiments. Abs and Cell Staining All phycoerythrin (PE), CyChrome (CyC), and biotinylated mAbs were purchased from PharMingen (San Diego, CA). These included PE-labeled antibodies to mouse CD4, CD8, CD28, CD44, Bcl-2, Fas, IFN-␥, IL-4, IL-10, and IgG; biotinylated antibodies to mouse CD28, CD44, CD69, CD25 (IL-2R␣), CD45RA, and IgG; and CyChrome (CyC)-labeled antibodies to mouse CD8 and CD4, as well as streptavidin-CyC and streptavidinfluorescein isothiocyanate (FITC). Rabbit antimouse IgG and goat antirat IgG were purchased from ICN (Costa Mesa, CA). Purified ascites to murine CD4 (clone GK1.5) were produced in our laboratory from cell lines purchased from the American Type Culture Collection (ATCC, Manassas, VA).
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For flow-cytometric analysis of cell-surface markers, cells were stained with the appropriate antibodies for 30 min on ice. Cells were then washed with phosphatebuffered saline (PBS)/1% bovine serum albumin (BSA) and fixed with 4% paraformaldehyde for 30 on ice. Cells were resuspended in PBS/1% BSA for analysis. For intracellular cytokine staining, cells were first activated with 100 ng/ml of phorbol myristate acetate (PMA) plus 500 ng/ml of ionomycin for 4 h. In the last 2 h of treatment monensin (Sigma, St. Louis, MO) was added to the media at a final concentration of 3 M. Cells were harvested and stained with the appropriate antisurface protein mAb and washed as described above. Cells were fixed and permeabilized simultaneously with 4% paraformaldehyde, 0.5% Saponin (Sigma), and 1% BSA for 30 min on ice. Cells were washed once with PBS/1% BSA and 0.5% Saponin (intracellular staining buffer) and stained with the appropriate anticytokine mAb for 30 min on ice. Cells were washed once with intracellular staining buffer and PBS/1% BSA. Cells were then resuspended in PBS/1% BSA for flow cytometry analysis. For Bcl-2 staining we followed the protocol suggested by the manufacturer of the antibody kit (PharMingen). T Cell Purification To prepare T cells, spleens were gently rubbed between frosted glass slides to obtain single cell suspensions in Hank’s balanced salt solution (HBSS) containing 0.2% bovine serum albumin (HBSS-BSA). Erythrocytes were removed by centrifugation over Lympholyte-M (Cedarlane Laboratory, Ontario, Canada), and adherent cells and B cells were depleted by panning over rabbit antimouse IgG Ab-coated plates. To prepare purified CD8 ⫹ cells, erythrocyte-depleted spleen cells were counted and incubated with rat antiCD4 for 30 min on ice. Adherent cells, B cells, and CD4 ⫹ cells were depleted by panning with goat antirat IgG Ab-coated plates. For purification of memory and naı¨ve or CD28 hi and CD28 lo CD8 ⫹ T cells, cells were stained with antimouse CD8 and either antimouse CD44 or CD28 for 30 min on ice. Cells were then sorted, using a FACSVantage instrument from Becton–Dickinson, into CD8 ⫹ populations bearing low or high amounts of either CD44 or CD28. Typical preparations showed the recovered cells in each population to be 97–99% pure. T Cell Transfer and CFSE Staining Purified cells were labeled with 5 mM Carbofluorescein diacetate succinimidyl ester (CFSE) at 1 ⫻ 10 7 cells/ml in PBS/0.1% BSA for 10 min at 37°C. Cells were then washed once with cold PBS/0.1% BSA and
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cold HBSS-BSA. CFSE-labeled cells were injected intravenously into the lateral tail vein of nonirradiated, syngeneic young mice in 200 l of PBS/0.1% BSA. At appropriate time points, cells from spleens and pooled lymph nodes (brachial, axillary, and inguinal) of recipient mice were isolated and stained for cytometric analysis as described above. Statistical Analyses To measure percentage of proliferation, we used a quantitative analysis that has been described before (29). It relies on the fact that the size of each subset of proliferating T cells is related to the size of the corresponding precursor subset by a function of 2 n, where n is the number of division cycles achieved in the recipient mice. Therefore if a specific fluorescence peak corresponding to n mitotic divisions contains E cells, then the number of precursor T cells (P) which must have divided n times to generate them can be estimated as E divided by 2 n. The total number of original precursor cells needed to account for the observed progeny can be calculated as the sum of the P values for each of the peaks, one for mitotic cycle, in the histogram. The percentage of cells in the recovered population that has given rise to a clone of proliferating progeny in vivo can then be calculated as the ratio of the sum of the P values to the total number of CFSE cells recovered (the sum of P plus the number of CFSE-positive nonproliferating cells). Data are presented in text and figures as the mean ⫾ SEM, and n indicates the number of individual experiments. Comparisons among different varieties of donor cells were made using the paired Student’s t test. RESULTS ⫹
CD8 T Cells from Old Mice Proliferate in Nonirradiated Recipients To determine whether age alters the ability of mouse T cells to proliferate in nonirradiated hosts, T cells from young (6 months) and old (18 months) CB6F1 mice were isolated and labeled with CFSE, a cell-tracking reagent. This dye is retained in cells for several weeks, and the label is inherited by daughter cells after cell division, making it possible to distinguish cells that have not divided from those that have divided one or more times after labeled cells are injected into host animals. CFSE-labeled T cells were injected iv into 4-month-old CB6F1 mice. Two to 12 days later, T cells were isolated from the lymph nodes (LN) and spleens of the recipient mice and aliquots stained with either PE-CD4 or PE-CD8. Figure 1 shows representative results. At two days after injection CD8 low and CD8 hi CFSE-labeled cells were detectable in both LN (Fig. 1,
FIG. 1. CD8 ⫹ T cells from old mice are able to proliferate in nonirradiated recipient mice. T cells were isolated from young and old CB6F1 mice, labeled with CFSE, and then injected into 4-monthold CB6F1 recipient mice. T cells from recipients’ spleen and pooled lymph nodes were isolated 2 or 12 days later and stained with ␣-CD8-PE or ␣-CD4-PE. Representative CD8 or CD4 vs CFSE dot plots of T cells isolated at days 2 and 12 from the lymph nodes of recipient mice injected with CFSE-labeled T cells from either young or old mice.
top panels) and spleen (not shown). As expected, these cells all had equally high levels of CFSE, indicating an absence of proliferation at this early time point. By day 12, however, many of the CD8 hi CFSE-labeled cells recovered from LN (Fig. 1) or spleen (not shown) had CFSE levels that were 50 or 25% of that shown by the brightest CFSE-positive cells, indicating that these cells had divided once or twice in the previous 12-day period. These cells appeared at high frequency only in hosts that had received CFSE-labeled cells from old donors. In contrast, very few CD4 hi T cells from old donors divide in nonirradiated recipient mice (Fig. 1, lower panels), and neither CD4 nor CD8 T cells derived from young donors showed evidence of more than minimal proliferation. A series of experiments was conducted to determine the kinetics of T-cell expansion in vivo, and the data are summarized in Fig. 2, which shows the percentage of CD8 ⫹ and CD4 ⫹ cells from young and old donors, isolated from the lymph nodes and spleens of the recipient mice, that had divided at least once at varying time points after inoculation. The proportion of recovered CFSE-positive CD8 cells that had undergone at least one in vivo mitosis increased steadily over at least 12 days for recipients of cells from old donors, but
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the recovered CD8 cells from 18-month-old donors had divided at least once in vivo, and this value increased significantly (P ⫽ 0.03), i.e., to 35%, in CD8 cells derived from 24-month-old donors. Proliferation of recovered CD4 ⫹ T cells remained low regardless of the age of the donor mice. Phenotype of CD8 ⫹ T Cells That Have Proliferated in Vivo Resembles That of Nonactivated Memory T Cells
FIG. 2. Time course of in vivo proliferation of CD4 and CD8 T cells from young and old donors injected into nonirradiated recipient mice. The bars represent the proportion of injected cells (⫾ SEM) capable of at least one round of proliferation estimated from the number of recovered cells and their inferred number of cell cycles; n represents the number of independent experiments.
remained low for young CD8 cells and for CD4 cells from donors of any age. By day 12 after injection on average 16% of the injected CD8 cells from old donors had been able to divide, in comparison with 3% from CD8 from young donors (P ⫽ 0.003). At day 12 the proportion of CD4 cells from old donors that had divided was only 3%, significantly (P ⫽ 0.004) lower than the proportion of proliferating CD8 cells from aged donors. The number of CFSE-labeled cells recovered at day 12 was similar in recipients of CD8 cells from young donors and CD4 cells from young and old mice (Table 1), but slightly higher in recipients of old CD8 cells, presumably because these cells are undergoing proliferation. Age-Dependent Increase in CD8 Cells That Proliferate in Vivo In order to determine at what age T cells from individual mice acquire the ability to proliferate in nonirradiated recipients, we isolated T cells from donors of different ages and analyzed their CFSE levels after 12 days of in vivo expansion. Figure 3 summarizes these results. We saw no significant proliferation of CD8 ⫹ T cells from mice 12 months or younger; fewer than 3% of the recovered CFSE-labeled cells from these younger donors had diminished CFSE levels. Sixteen percent of
Multiparameter flow cytometry was used to determine the cell-surface phenotype of CFSE-labeled CD8 cells recovered at day 12 from mice injected with T cells from aged donors. Figure 4 shows a representative result. Most of the CD8 ⫹ T cells that have divided in vivo have high levels of CD44 and CD45RA and low levels of CD25 and CD69. In contrast, antigen-driven T-cell proliferation typically leads to increases in cellsurface levels of CD25 and CD69 (30). Thus our results suggest that in vivo proliferation of CD8 cells from old donors is antigen independent. This is in agreement with previous reports where either CD8 ⫹ T-cell clones or nonclonal memory CD8 ⫹ T cells were observed to proliferate after injection into  2 microglobulin knockout mice (25). The phenotype we obtain in these experiments correlates with the phenotype seen in cells undergoing homeostatic proliferation after transfer into irradiated mice (30). After isolation from recipient mice, CD8 cells that proliferated in vivo appear to have a memory phenotype, as indicated by high-level expression of CD44. To see if high levels of CD44 were also present, prior to transplantation, on the precursors of the proliferating population, we purified naı¨ve and memory CD8 ⫹ T cells from old mice and injected them into nonirradiated recipients (data not shown). We found that only memory CD8 ⫹ T cells from old donor mice, i.e., cells with high levels of CD44, were able to divide in nonirradiated recipient mice. CD8 Cells That Are Able to Proliferate in Nonirradiated Recipients Have High Levels of CD28 Although some initial reports (17) suggested that the clonal CD8 cells found in aged mice were CD28 negative, reevaluation using a different anti-CD28 antibody showed that these cells did indeed express CD28 (25). Our own data, shown in Fig. 5, suggest that the levels of CD28 on cells which have proliferated in vivo are typically higher than those that did not divide after injection. All the CFSE-positive CD8 T cells recovered from the 6-month-old donor expressed CD28, but at modest levels (CD28 lo) (Fig. 5, left panel). The middle
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TABLE 1 Percentage of CFSE-Labeled Cells Recovered at Day 12 Old donors (%)
Lymph nodes Spleen
Young donors (%)
CD8 ⫹ T cells
CD4 ⫹ T cells
CD8 ⫹ T cells
CD4 ⫹ T cells
1.16 ⫾ 0.07 0.62 ⫾ 0.03
0.68 ⫾ 0.11 0.45 ⫾ 0.05
0.75 ⫾ 0.07 0.38 ⫾ 0.04
0.98 ⫾ 0.07 0.54 ⫾ 0.04
Note. Table 1 shows the percentages (mean ⫾ SEM) of CFSE-labeled cells in the isolated spleen and lymph node cells from recipient mice 12 days after injection and illustrates the distribution into CD4 and CD8 subsets. CD4 ⫹ and CD8 ⫹ T-cell percentages were determined by quantifying cells from the samples stained with ␣-PE-CD4 and ␣-PE-CD8 (Fig. 2) (n ⫽ 6).
panel shows cells from an 18-month-old donor: in this case 65% of the proliferating cells, and only 12% of the nonproliferating cells, express higher than normal levels of CD28 (CD28 hi cells). The effect was more dramatic in the 24-month-old donor, for which almost all (90%) of the proliferating cells, but only 33% of the nonproliferating cells, were CD28 hi (Fig. 5, right panel). The results shown in Fig. 5 suggested that elevated CD28 levels might help to identify CD8 T cells able to proliferate in vivo. Figure 6 shows results of an experiment to determine if aging alters the proportion of memory CD8 T cells with high levels of CD28. In a series of two experiments, the average number of CD28 hi cells among the memory CD8 population in the
FIG. 3. Effect of donor age on in vivo proliferation of CD4 and CD8 T cells. The upper and lower panels show the percentages of CD8 ⫹ (solid) and CD4 ⫹ (open) cells from the different donors that had divided by day 12, isolated from the lymph nodes and spleens of recipient mice respectively; n represents the number independent experiments; values are means ⫾ SE.
lymph nodes of aged mice was 50.6 ⫾ 13.8% (mean ⫾ SEM); in contrast 18.6 ⫾ 5.6% of CD8 cells from young donors were CD28 hi. We obtained similar numbers in cells from spleen. To understand if the CD28 hi subset of CD8 ⫹ cells was indeed responsible for the proliferation seen in previous experiments, we purified CD28 hi and CD28 lo CD8 ⫹ T cells from old mice and injected them separately into nonirradiated recipients. Figure 7 shows the results of a representative experiment and summary statistics from a series of four such experiments. The data show that only CD28 hi CD8 ⫹ T cells from old donor mice were able to divide in nonirradiated recipient mice. These
FIG. 4. Expression of surface markers on CD8 ⫹ T cells from old mice. T cells were stained after isolation from recipient mice with one of the following biotinylated antibodies for flow cytometry analysis: ␣-CD44, ␣-CD25, ␣-CD45RA, or ␣-CD69. Biotinylated antibodies were detected with streptavidin-CyC. The figure shows dot plots of the various surface markers vs CFSE on the gated CD8 ⫹ cells. The numbers represent the percentage of cells in each quadrant. Similar results were obtained in a second experiment.
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FIG. 5. CD8 cells that are able to divide in vivo have higher levels of CD28. T cells were stained after isolation from recipient mice with biotinylated ␣-CD28. Biotin was detected with SAv-CyC. T cells were gated on the CFSE ⫹ CD8 ⫹ cells. The figure shows CD28 vs CFSE on the gated cells from different-aged donor mice. The numbers represent the percentage of cells in each quadrant. Similar results were obtained in a second experiment.
results suggest that exceptionally high levels of CD28 may help to identify precursors of CD8 cells with abnormal responses to homeostatic regulation for further functional and biochemical study. CD8 ⫹ T Cells That Have Proliferated in Vivo Produce IFN-␥ Accumulation with age of CD8 ⫹ T cells with altered growth control could potentially contribute to the decline in immune responses seen in elderly individuals. In order to understand the possible role of these clones in regulation of the immune system, we measured using intracellular staining their production of some important immunoregulatory cytokines that are known to be produced by at least some CD8 ⫹ T cells, including IFN-␥, IL-4, and IL-10. Figure 8 shows results from
analysis of lymph node cells from mice 12 days after injection of CFSE-labeled T cells from 23-month-old mice. We found that approximately 80% of the CD8 cells that had been able to divide in nonirradiated recipients were able to produce IFN-␥ after brief culture (4 h) with PMA and ionomycin; about 30% of the recovered CFSE-positive but nonproliferating cells were also IFN-␥ producers (Fig. 8, upper panels). Only a small number of the cells (12%) that proliferated in vivo were IL-4 producers (Fig. 8, lower panels). There was no difference in the percentage of IL-4 producing cells between the population that proliferated in vivo and the population that did not proliferate (Fig. 8, lower right panel). We saw no significant production of IL-10 by proliferating or nonproliferating cells (data not shown). To determine if these cells were producing IFN-␥ in their original host prior to injection into recipient mice, we measured this cytokine by intracellular staining on CD8 ⫹ CD28 high T cells from young and old mice. Figure 9 shows results of a representative experiment and summary statistics from a series of three such experiments. The data show that only CD8 ⫹ CD28 high T cells from old mice are able to produce IFN-␥ in their original host. We found that approximately 68% of the CD8 ⫹ CD28 high T cells in old mice were able to produce IFN-␥, as compared to 9% for the CD8 ⫹ CD28 high T cells in young mice. CD8 ⫹ T Cells That Have Proliferated in Vivo Have High Levels of Bcl-2
FIG. 6. A population of CD28 hi in memory CD8 cells from old mice. The figure shows CD28 vs CD44 on cells that have been gated as CD8 ⫹ from a young and an old mouse prior to injection into recipient host. Numbers represent the percentage of cells in each quadrant. Similar results were obtained in a total of two independent experiments.
Accumulation in aged mice of CD8 cells with the ability to expand in nonirradiated hosts could in principle reflect alterations in levels of antiapoptotic or proapoptotic molecules. To test this hypothesis we
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FIG. 7. Only CD8 T cells with higher levels of CD28 are able to divide in vivo. CD28 hi and CD28 lo CD8 ⫹ T cells from old CB6F1 mice (20 –23 months old) were isolated, labeled with CFSE, and injected into nonirradiated syngeneic recipient mice. Twelve days after injection recovered cells from spleens and lymph nodes were stained for CD8 and CD28. T cells from the old mice were gated on the CFSE ⫹ CD8 ⫹ cells. Representative CD28 vs CFSE dot plots of gated cells isolated from lymph nodes of recipient mice injected with either CD28 lo or CD28 hi CFSE-labeled CD8 ⫹ T cells are shown. The numbers represent the percentage of cells in each quadrant. The number of CD28 lo or CD28 hi CD8 ⫹ T cells that divided in vivo was calculated. The right panel shows the mean ⫾ SEM for the percentage of CD28 lo and CD28 hi CD8 ⫹ T cells isolated from the lymph nodes and the spleens of recipient mice that were able to divide in vivo (n ⫽ 4).
measured levels of Bcl-2 and Fas in cells isolated from the lymph nodes and spleens of recipient mice that had been injected with CFSE-labeled T cells from young or
FIG. 8. Cytokine production by CD8 T cells in old mice. T cells from 23-month-old CB6F1 mice were labeled with CFSE and injected into young recipients. Cells from recipients’ spleens and pooled lymph nodes were isolated at day 12 and cultured for 4 h with 100 ng/ml PMA and 500 ng/ml ionomycin. In the last 2 h of culture, 3 M monensin was added. Cells were harvested and stained with ␣-CD8CyC. After fixation and permeabilization, cells were stained with ␣-IFN␥-PE, ␣-IL-4-PE, ␣-IL-10-PE (not shown), or ␣-IgG-PE. T cells from the old mice were gated on the CFSE ⫹ CD8 ⫹ cells. Left panels represent dot plots of cytokine production (IFN-␥ or IL-4) vs CFSE on the gated cells. Right panels show the percentages of CD8 ⫹ isolated from the lymph nodes and spleens of recipient mice that were able to produce the indicated cytokine, both for cells that had not divided in vivo (“undivided”) and for cells that had divided at least once in vivo, as means ⫾ SEM for n ⫽ 2 experiments. The thresholds shown represent the 97% cutoff points for the ␣-IgG-stained controls.
old donors. Figure 10 shows representative results from one of two independent experiments. The levels of Bcl-2 on the CD8 ⫹ T cells from old donors that were able to proliferate in vivo are higher in comparison to Bcl-2 levels of CD8 ⫹ T cells from young donors (Fig. 10). The left panel shows cells from a 6-month-old donor. Consistent with our previous results, most cells showed no evidence of in vivo mitosis. Among these, 75% expressed low, but easily detectable, levels of Bcl-2, and only 25% expressed the higher levels indicated by the threshold shown in the figure. The right panel shows cells from a 23-month-old donor: In this case 81% of the proliferating cells, and only 45% of the nonproliferating cells, expressed levels of Bcl-2 that exceeded the arbitrary threshold. In a series of experiments using four old donors, the mean proportion of Bcl-2 hi cells was found to be 81 ⫾ 3.3% for proliferating cells, and only 46 ⫾ 5% for the nonproliferating cells, as compared to 30 ⫾ 4.3% for young donors (N ⫽ 2). In contrast, we found no difference among proliferating and nonproliferating cells in levels of Fas (data not shown). The fact that CD8 ⫹ T cells that proliferate and survive in vivo have higher levels of Bcl-2 than T cells that did not divide suggests that Bcl-2-mediated resistance to cell death may contribute to their unusual potential for in vivo replication. DISCUSSION
The accumulation, in aging mice, of large clones of CD8 ⫹ T cells (17, 18, 25) is likely to reflect alterations in the ability of some CD8 cells to control division and/or cell death in the face of homeostatic controls that are still poorly understood. The proportion of aged
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FIG. 9. IFN-␥ production of CD8 T cells in old mice. Cells from the spleens and pooled lymph nodes of 7- and 22-month-old mice were isolated and cultured for 4 h with 100 ng/ml PMA and 500 ng/ml ionomycin. In the last 2 h of culture, 3 M monensin was added. Cells were harvested and stained with ␣-CD8-CyC and ␣-CD28-FITC. After fixation and permeabilization, cells were stained with ␣-IFN-␥-PE or ␣-IgG-PE (not shown). T cells from the old mice were gated on the CD8 ⫹ cells. Left and middle panels represent dot plots of cytokine production vs CD28 on the gated cells. Right panes show the percentages of CD8 ⫹ CD28 high T cells isolated from the lymph nodes of young and old mice that were able to produce IFN-␥ as means ⫾ SEM for n ⫽ 2 experiments. The thresholds shown represent the 98% cutoff points for the ␣-IgG-stained controls. Similar results were obtained in spleen samples.
T cells for which these homeostatic mechanisms have gone awry cannot be estimated simply by counting the number of cells within large in vivo clones because smaller clones, if they exist, will not be recognizable by methods based on either TCR V distributions or V spectratyping. It has recently been reported that CD8 ⫹ T-cell clones, as well as memory CD8 ⫹ T cells not demonstrably members of any obvious clone, are able to divide in vivo after transfer into normal animals (25). To explore this phenomenon further, we have measured the proliferative capacity of T cells from young and old mice by flow cytometry using the fluorescent dye CFSE. We found that CD8 ⫹ T cells from old
FIG. 10. CD8 T cells that are able to divide in vivo have high levels of Bcl-2 expression. T cells recovered from recipients’ pooled lymph nodes were isolated at day 12 and stained with ␣-CD8-CyC. After fixation and permeabilization, cells were stained with ␣-Bcl-2PE. T cells from mice injected with T cells from young and old donors were gated on the CFSE ⫹ CD8 ⫹ cells (both panels). Representative Bcl-2 vs CFSE dot plots of gated cells isolated from lymph nodes of recipient mice injected with CFSE-labeled T cells from either young or old donor are shown above. We obtained the same results in a second independent experiment.
donors, but not young donors, were able to proliferate in nonirradiated hosts. In contrast, CD4 ⫹ T cells from mice of any age did not divide in vivo. CD8 ⫹ T cells from old donors continued to proliferate in vivo for at least 12 days. When recovered from recipient mice, CD8 ⫹ T cells that had undergone at least one round of mitosis exhibit high expression of CD44, similar to resting memory cells. By isolating naı¨ve and memory CD8 T cells from old mice, we showed that only the CD44 hi subset of CD8 ⫹ T cells contains cells able to divide in nonirradiated recipient mice. The CD44 lo subset of CD8 cells, which contains most of the naı¨ve CD8 cells, contained no cells able to divide in vivo (31). Donor mice aged 12 months or younger contained no detectable numbers of CD8 ⫹ T cells able to proliferate in vivo, but in mice older than 18 months the CD8 ⫹ T-cell expansions increased with the age of the donor mice. This timing is consistent with previous information about appearance of T-cell clones in aging mice; such clones are typically not detected before the age of 12 months and then increase in frequency with age (11, 17, 18, 28). Using a three-color flow cytometric system that allowed us to study the surface markers of the CFSEtagged cells recovered from recipient mice, we found that the proliferating CD8 cells bear CD25 low and CD69 low phenotypes. Antigen-driven proliferation leads to increases in cell surface CD25 and CD69 expression (30). Thus the low level of CD25 and CD69 on CFSEpositive proliferating cells suggests that the in vivo proliferation is not dependent on activation through the T-cell receptor. Several experiments have shown that, unlike naı¨ve or actively responding CD8 T cells, the persistent population of CD8 memory T cells is not
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dependent on Ag or even MHC class I for its survival (40 – 42). The phenotype we obtain in these experiments correlates with the phenotype seen in cells undergoing homeostatic proliferation after transfer into irradiated mice (30). Interestingly most of the CD8 cells that are able to divide in nonirradiated hosts are CD45RA ⫹. CD8 ⫹ CD45RA ⫹ T cells, compared to CD8 ⫹, CD45RA ⫺ cells, are relatively nonresponsive to TCR stimulation in tests for cytokine production (32–35). It is not yet clear whether the CD8 cells whose proliferative abnormalities cause them to accumulate in aged mice are or are not normally responsive to antigenic stimulation. These cells clearly do produce IFN-␥ after injection into young hosts, but whether they produce this or other cytokines after TCR or antigenic stimulation is still to be determined. It is noteworthy, however, that anergic CD8 ⫹ T cells have been reported to persist and function in vivo (43, 44). Our data show that the CD8 ⫹ T cells that are able to divide in vivo have higher levels of CD28 when compared either with CD8 ⫹ T cells from old donors that did not divide in vivo or with recovered CFSE-positive cells from a young donor. In addition, we noted a population of memory CD8 cells, higher levels of CD28, present only in aged mice, from which emerged nearly all of the CD8 cells that were able to divide in vivo. Our data suggest that exceptionally high levels of CD28 may help to identify precursors of CD8 cells with abnormal responses to homeostatic regulation for further functional and biochemical study. We do not know at this point whether CD28 per se contributes to the altered growth control of these cells or if it only serves as a marker for cells in the subset. We note, though, that CD28 is expressed almost universally on CD8 T cells of both young and old mice (see Figs. 5 and 6 and also Ref. 45) and thus that expression of CD28 is not sufficient to confer abnormal in vivo proliferation. The high levels of Bcl-2 (see Fig. 10) suggest that clonal accumulation of abnormal CD8 cells in old mice might reflect an altered balance among proapoptotic and antiapoptotic factors in these cells. IL-15, previously implicated in the expansion of CD8 cells in aged mice (25), may well help to induce Bcl-2 and thereby diminish susceptibility to programmed cell death. The effects of aging on CD28 expression in the CD8 ⫹ subset are apparently different in humans, in which aging leads to a characteristic loss of CD28, predominantly among CD8 ⫹ T cells (11, 14 –16). This CD28 ⫺ population consists of a subset of memory T cells, and in many respects their phenotype and function are highly suggestive of differentiated armed effector T cells in that they produce IFN-␥ and TNF-␣ and express high levels of granzyme A and perforin. They also contain Ag-specific memory CTL (46, 47) and exert a potent cytolytic activity without requiring deliberate in vitro activation (48, 49). In humans, the CD28 ⫺ T cells
are clonally expanded (11, 14 –16), terminally differentiated lymphocytes with little proliferative response to mitogenic stimulation in vitro (49, 50). Recent data suggest that they derive from their CD28 ⫹ counterpart (50, 51). The etiology of most expansions in healthy individuals is not known. However, studies of T-cell receptor usage suggested that these cells arose in response to antigenic stimulus (14, 52). These clonal expansions are maintained in vivo at a high frequency for long periods of time (11, 14, 16) and are sometimes noted in association with infections (53, 54), autoimmune conditions (55, 56), or neoplasia (57, 58). It is not clear whether aging humans have a specific CD8 ⫹ subset, like the one we document in aged mice, from which clonal expansions are derived, nor whether such a precursor population has unusually high or low levels of CD28 expression. It is also possible that differences in source of cells—spleen and lymph nodes for mouse and blood for humans— contribute to the apparent differences in age effect on CD28 levels. Measurement of cytokine production provides a first step toward understanding the possible role of these expanded CD8 cells in late-life immunodeficiency. The cells do produce IFN-␥ after stimulation with PMA and ionomycin, but produce only low levels of IL-4 and no detectable IL-10. Although IFN-␥ has been shown to be immunosuppressive in some systems (59), a good deal of additional work, including tests for other cytokines and chemokines, will be needed to test the idea that the increased frequency of unregulated, clonal CD8 cells in old age has functional consequences. ACKNOWLEDGMENTS We wish to thank Dr. Charles D. Surh for advice and technical assistance. This work was supported by NIH Grant R01-AG19619. A.O.S. was supported by the Rackham Merit Fellowship and NIH Training Grants GM07315-21 and T32-AI07413. REFERENCES 1. Miller, R. A., The aging immune system: Primer and prospectus. Science 273, 70, 1996. 2. Nordin, A. A., and Collins, G. D., Limiting dilution analysis of alloreactive cytotoxic precursor cells in aging mice. J. Immunol. 131, 2215, 1983. 3. De Paoli, P., Battistin, S., and Santini, G. F., Age-related changes in human lymphocyte subsets: Progressive reduction of the CD4 CD45R (suppressor inducer) population. Clin. Immunol. Immunopathol. 48, 290, 1988. 4. Thoman, M. L., and Weigle, W. O., Partial restoration of Con A-induced proliferation, IL-2 receptor expression, and IL-2 synthesis in aged murine lymphocytes by phorbol myristate acetate and ionomycin. Cell Immonol. 114, 1, 1988. 5. Vie, H., and Miller R. A., Decline with age in the proportion of mouse T cells that express IL-2 receptor after mitogen stimulation. Clin. Mech. Ageing Dev. 33, 313, 1986. 6. Miller, R. A., Jacobson, B., Weil, G., and Simons, E. R., Diminished calcium influx in lectin-stimulated T cells from old mice. J. Cell Physiol. 132, 337, 1986.
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24. Zhang, X., Sun, S., Hwang, I., Tough, D. F., and Sprent, I., Potent and selective stimulation of memory-phenotype CD8⫹ T cells in vivo by IL-15. Immunity 8, 591, 1998. 25. Ku, C. C., Kappler, J., and Marrack, P., The growth of the very large CD8⫹ T cell clones in older mice is controlled by cytokines. J. Immunol. 166(4), 2186 –2193, 2001. 26. Tough, D. F., and Sprent, J., Turnover of naive- and memoryphenotype T cells. J. Exp. Med. 179, 1127, 1994. 27. Murali-Krishna, K., Lau, L. L., Sambhara, S., Lemonnier, F., Altman, J., and Ahmed, R., Persistence of memory CD8 T cells in MHC class I-deficient mice. Science 286, 1377, 1999. 28. LeMaoult, J., Messaoudi, I., Manavalan, J. S., Potvin, H., Nikolich-Zugich, D., Dyall, R., Szabo, P., Weksler, M. E., and Nikolich-Zugich, J., Age-related dysregulation in CD8 T cell homeostasis: Kinetics of a diversity loss. J. Immunol. 165(5), 2367– 2373, 2000. 29. Gudmundsdottir, H., Wells, A. D., and Turka, L. A., Dynamics and requirements of T cell clonal expansion in vivo at the singlecell level: Effector function is linked to proliferative capacity. J. Immunol. 162(9), 5212, 1999. 30. Murali-Krishna, K., and Ahmed, R., Cutting edge: Naı¨ve T cells masquerading as memory cells. J. Immunol. 165, 1733–1737, 2000. 31. Bruno, L., von Boehmer, H., and Kirberg, J., Cell division in the compartment of naive and memory T lymphocytes. Eur. J. Immunol. 26, 3179, 1996. 32. Bell, E. B., Spartshott, S. M., and Bunce, C., CD4 ⫹ T-cell memory, CD45R subsets and the persistence of antigen—a unifying concept. Immunology Today 19(2), 60, 1998. 33. Woodward, B., Hillyer, L., and Hunt, K., T cells with a quiescent phenotype (CD45RA⫹) are overabundant in the blood and involuted lymphoid tissues in wasting protein and energy deficiencies. Immunology 96, 246, 1999. 34. Lightstone, L., and Marvel, J., CD45RA⫹ T cells: Not simple virgins. Clin. Sci. (Colch). 85(5), 515, 1993. 35. Pihlgren, M., Lightstone, L., Mamalaki, C., Rimon, G., Kioussis, D., and Marvel, J., Expression in vivo of CD45RA, CD45RB and CD44 on T cell receptor-transgenic CD8⫹ T cells following immunization. Eur. J. Immunol. 25(6), 1755, 1995. 36. Miyazaki, T., Liu, Z. J., Kawahara, A., Minami, Y., Yamada, K., Tsujimoto, Y., Barsoumian, E. L., Permutter, R. M., and Taniguchi, T., Three distinct IL-2 signaling pathways mediated by bcl-2, c-myc, and lck cooperate in hematopoietic cell proliferation. Cell 81, 223, 1995. 37. Mor, F., and Cohen, L. R., IL-2 rescues antigen-specific T cells from radiation or dexamethasone-induced apoptosis: Correlation with induction of Bcl-2. J. Immunol. 156, 515, 1996. 38. Akbar, A. N., Borthwick, N. J., Wickremasinghe, R. G., Panayoitidis, P., Pilling, D., Bofill, M., Krajewski, S., Reed, J. C., and Salmon, M., Interleukin-2 receptor common-chain signaling cytokines regulate activated T cell apoptosis in response to growth factor withdrawal: Selective induction of anti-apoptotic (bcl-2, bcl-xL) but not pro-apoptotic (bax, bcl-xS) gene expression. Eur. J. Immunol. 26, 294, 1996. 39. Grayson, J. M., Zajac, A. J., Altman, J. D., and Ahmed, R., Cutting edge: Increased expression of Bcl-2 in antigen-specific memory CD8 ⫹ T cells. J. Immunol. 164, 3950 –3954, 2000. 40. Hou, S., Hyland, L., Ryan, K. W., Portner, A., and Doherty, P. C., Virus-specific CD8⫹ T-cell memory determined by clonal burst size. Nature 369, 652, 1994. 41. Lau, L. L., Jamieson, B. D., Somasundaram, T., and Ahmed, R., Cytotoxic T-cell memory without antigen. Nature. 369, 648, 1994.
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42. Xiang, R., Lode, H. N., Gillies, S. D., and Reisfeld, R. A., T cell memory against colon carcinoma is long-lived in the absence of antigen. J. Immunol. 163, 3676, 1999. 43. Blish, C. A., Dillon, S. R., Farr, A. G., and Fink, P. J., Anergic CD8⫹ T cells can persist and function in vivo. J. Immunol. 163(1), 155–164, 1999. 44. Deeths, M. J., Kedl, R. M., and Mescher, M. F., CD8⫹ T cells become nonresponsive (anergic) following activation in the presence of costimulation. J. Immunol. 163(1), 102–110, 1999. 45. Kirk, C. J., Freilich, A. M., and Miller, R. A., Age-related decline in activation of JNK by TCR- and CD28-mediated signals in murine T-lymphocytes. Cell Immunol. 197(2), 75– 82, 1999. 46. Fiorentino, S., Dalod, M., Olive, D., Guillet, J. G., and Gomard, E., Predominant involvement of CD8⫹CD28⫺ lymphocytes in human immunodeficiency virus-specific cytotoxic activity. J. Virol. 70, 2022, 1996. 47. Ho, H. N., Hultin, L. E., Mitsuyasu, R. T., Matud, J. L., Hausner, M. A., Bockstoce, D., Chou, C. C., O’Rourke, S., Taylor, J. M., and Giorgi, J. V., Circulating HIV-specific CD8⫹ cytotoxic T cells express CD38 and HLA-DR antigens. J. Immunol. 150, 3070, 1993. 48. Fagnoni, F. F., Vescovini, R., Mazzola, M., Bologna, G., Nigro, E., Lavagetto, G., Franceschi, C., Passeri, M., and Sansoni, P. Expansion of cytotoxic CD8⫹ CD28⫺ T cells in healthy agein people, including centenarians. Immunology 88, 501, 1996. 49. Azuma, M., Phillips, J. H., and Lanier, L. L., CD28⫺ T lymphocytes: Antigenic and functional properties. J. Immunol. 150, 1147, 1993. 50. Chamberlain, W., Falta, M., and Kotzin, B., Functional subsets within clonally expanded CD8⫹ memory T cells in elderly humans. Clin. Immunol. 94, 160 –172, 2000. 51. Fiorentini, S., Malacarne, F., Ricotta, D., Licenziati, S., Armenta-Solis, A., Ausenda, S., De Francesco, M. A., Garrafa, E., Simonini, A., Imberti, L. et al. Long term stimulation of CD8⫹CD28⫺ T cells induces stable disappearance of CD28 molecules: A possible mechanism accounting for the increased numReceived February 7, 2002; accepted with revision April 17, 2002
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A Subset of CD8 Memory T Cells from Old Mice Have High Levels of CD28 and Produce IFN-␥ Anavelys Ortiz-Sua´rez* and Richard A. Miller* ,† ,‡ ,㛳 ,§ ,1 *Cellular and Molecular Biology Graduate Program and †Department of Pathology, University of Michigan School of Medicine; Ann Arbor, Michigan 48109-0940; and ‡Institute of Gerontology, 㛳Geriatrics Center, and §VA Medical Center, Ann Arbor, Michigan 48105
Using carboxyfluorescein diacetate succinimidyl ester (CFSE)-tagged cells to measure proliferation in vivo, we found that only memory CD8 ⴙ cells from mice older than 18 months gave measurable levels of proliferation and that the proportion of memory CD8 ⴙ T cells able to proliferate in a nonirradiated recipient increased with age. CD8 cells that had proliferated in vivo contained higher levels of CD28 when compared to CD8 cells that had not divided. Cells with high levels of CD28 were preferentially able to divide in nonirradiated recipients. Using ex vivo intracellular staining analysis, we determined that most of the CD8 ⴙ T cells that were capable of dividing in vivo produced IFN-␥ after isolation from recipient mice or their original host. These studies thus document the presence in aged mice of a population of CD28 hi CD8 ⴙ cells whose ability to proliferate in vivo without antigenic stimulation and to produce IFN-␥ may be involved in immune regulation. © 2002 Elsevier Science (USA) Key Words: CTL; T lymphocytes; cytokines; IFN-␥ memory; spleen; lymph nodes. INTRODUCTION
Age in humans and rodents leads to immunologic defects which include deficits in the formation of highaffinity antibodies (1), generation of long-lasting immune responses after vaccination (2, 3), and expression of delayed-type hypersensitivity reactions to antigens initially encountered earlier in life (1). Immunodeficiencies associated with age appear to be due to some extent to the fact that fewer T cells from aged individuals are activated in response to mitogens as measured by proliferative capacity, calcium flux, or IL-2 receptor up-regulation (1, 4, 5). An age-dependent shift from naı¨ve T cells (predominant in young individuals) to memory T cells has also been observed (1). This could help to explain the decline in the number of functionally reactive T cells with age to mitogens in vitro and 1
To whom correspondence and reprint requests should be addressed at 5316 CCGC, 1500 E. Medical Center Dr., Ann Arbor, MI 48109-0940. E-mail: [email protected]. 1521-6616/02 $35.00 © 2002 Elsevier Science (USA) All rights reserved.
could possibly have an effect on the parallel decrease with age in the in vivo responsiveness to newly encountered antigens. Another possibility is that anergized T cells accumulate with age (6 –9). Several groups have reported that aging leads to changes in the repertoire of T cells in both mice and humans (10 –22). Older human beings and mice often develop large clones of CD8 ⫹ T cells (10 –18). These clones can represent as much as 80% of the CD8 ⫹ T cells in an individual (17). Large clones of this kind are not typically seen among CD4 ⫹ T cells, although smaller clones are often detected in the CD4 population (19 –22). CD8 ⫹ T-cell clones are detectable in almost every person over the age of 40 (16 –17). Similarly, expanded CD8 ⫹ T cells clones appear in middle-aged mice, and longitudinal studies have shown that their frequency increases rapidly after the age of 18 months (17, 18, 22, 28). It has been reported (28) that most of these mouse CD8 ⫹ T-cell clones are long lasting but nevertheless transient. Hence, the T-cell repertoire of aged mice seems highly unstable, rather than terminally altered, although infrequent stable alteration can sometimes be seen (28). These CD8 ⫹ T-cell clones show no signs of transformation to malignancy. Human and mice containing these clones have normal life spans with only rare evidence of T-cell neoplasia (11). Nevertheless, the percentage of CD8 ⫹ T-cell clones bearing particular Vs in old animals increased in comparison with other CD8 ⫹ T cells (17, 18, 25). Consequently, one can speculate that even though the clones are not overtly malignant, they have at some earlier stage made a transition to a state that allows them to proliferate in vivo more readily than other cells because of changes in cell-cycle kinetics, susceptibility to apoptotic stimuli, or some other alteration. The cause of this age-dependent skewing is not yet known. Recently, it has been shown (25) that the expanded CD8 ⫹ clones are very similar to nonclonal memory CD8 ⫹ cells in that both express similar surface molecules. Both bear high levels of CD44 and similar amounts of IL-2R. Like memory phenotype
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CD8 ⫹ T cells (23, 24), cells of the expanded clones divide slowly in nonirradiated mice by a process that is probably antigen independent, driven by IL-15, and inhibited by IL-2 (25). However, the proliferating progeny of the large CD8 ⫹ T-cell clones accumulated, in vivo, more rapidly than the progeny of nonclonal CD8 ⫹ T cells (25). Marrack and her colleagues have suggested that increased sensitivity to the stimulatory effects of IL-15 or lowered sensitive to the inhibitory effects of IL-2 contributes to the accumulation of clonally expanded CD8 T cells in older mice. The fact that old mice and humans accumulate a population of clonal T cells suggests that this population might contribute to the immunological deficits seen in elderly individuals. The experiments described in this article were designed to determine whether age alters the ability of mouse T cells to proliferate in nonirradiated hosts and to further characterize the cells that are able to divide in vivo without intentional antigenic stimulation. MATERIALS AND METHODS
Mice Specific-pathogen-free male (BALB/c ⫻ C57BL/6)F 1 (CB6F1) mice were purchased from the National Institute on Aging’s contract colonies at Harlan (Indianapolis, IN). Unless otherwise stated, “young” mice were used at 4 – 8 months of age, and “old” mice were 18 –25 months old. Mice were housed for at least 2 weeks after shipment in a specific-pathogen-free holding colony at the University of Michigan and given free access to food and water. Sentinel animals from this colony were examined quarterly for serological evidence of viral infection; all such tests were negative during the experimental period. Mice that were found to have splenomegaly or macroscopically visible tumors at the time of sacrifice were not used for any experiments. Abs and Cell Staining All phycoerythrin (PE), CyChrome (CyC), and biotinylated mAbs were purchased from PharMingen (San Diego, CA). These included PE-labeled antibodies to mouse CD4, CD8, CD28, CD44, Bcl-2, Fas, IFN-␥, IL-4, IL-10, and IgG; biotinylated antibodies to mouse CD28, CD44, CD69, CD25 (IL-2R␣), CD45RA, and IgG; and CyChrome (CyC)-labeled antibodies to mouse CD8 and CD4, as well as streptavidin-CyC and streptavidinfluorescein isothiocyanate (FITC). Rabbit antimouse IgG and goat antirat IgG were purchased from ICN (Costa Mesa, CA). Purified ascites to murine CD4 (clone GK1.5) were produced in our laboratory from cell lines purchased from the American Type Culture Collection (ATCC, Manassas, VA).
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For flow-cytometric analysis of cell-surface markers, cells were stained with the appropriate antibodies for 30 min on ice. Cells were then washed with phosphatebuffered saline (PBS)/1% bovine serum albumin (BSA) and fixed with 4% paraformaldehyde for 30 on ice. Cells were resuspended in PBS/1% BSA for analysis. For intracellular cytokine staining, cells were first activated with 100 ng/ml of phorbol myristate acetate (PMA) plus 500 ng/ml of ionomycin for 4 h. In the last 2 h of treatment monensin (Sigma, St. Louis, MO) was added to the media at a final concentration of 3 M. Cells were harvested and stained with the appropriate antisurface protein mAb and washed as described above. Cells were fixed and permeabilized simultaneously with 4% paraformaldehyde, 0.5% Saponin (Sigma), and 1% BSA for 30 min on ice. Cells were washed once with PBS/1% BSA and 0.5% Saponin (intracellular staining buffer) and stained with the appropriate anticytokine mAb for 30 min on ice. Cells were washed once with intracellular staining buffer and PBS/1% BSA. Cells were then resuspended in PBS/1% BSA for flow cytometry analysis. For Bcl-2 staining we followed the protocol suggested by the manufacturer of the antibody kit (PharMingen). T Cell Purification To prepare T cells, spleens were gently rubbed between frosted glass slides to obtain single cell suspensions in Hank’s balanced salt solution (HBSS) containing 0.2% bovine serum albumin (HBSS-BSA). Erythrocytes were removed by centrifugation over Lympholyte-M (Cedarlane Laboratory, Ontario, Canada), and adherent cells and B cells were depleted by panning over rabbit antimouse IgG Ab-coated plates. To prepare purified CD8 ⫹ cells, erythrocyte-depleted spleen cells were counted and incubated with rat antiCD4 for 30 min on ice. Adherent cells, B cells, and CD4 ⫹ cells were depleted by panning with goat antirat IgG Ab-coated plates. For purification of memory and naı¨ve or CD28 hi and CD28 lo CD8 ⫹ T cells, cells were stained with antimouse CD8 and either antimouse CD44 or CD28 for 30 min on ice. Cells were then sorted, using a FACSVantage instrument from Becton–Dickinson, into CD8 ⫹ populations bearing low or high amounts of either CD44 or CD28. Typical preparations showed the recovered cells in each population to be 97–99% pure. T Cell Transfer and CFSE Staining Purified cells were labeled with 5 mM Carbofluorescein diacetate succinimidyl ester (CFSE) at 1 ⫻ 10 7 cells/ml in PBS/0.1% BSA for 10 min at 37°C. Cells were then washed once with cold PBS/0.1% BSA and
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cold HBSS-BSA. CFSE-labeled cells were injected intravenously into the lateral tail vein of nonirradiated, syngeneic young mice in 200 l of PBS/0.1% BSA. At appropriate time points, cells from spleens and pooled lymph nodes (brachial, axillary, and inguinal) of recipient mice were isolated and stained for cytometric analysis as described above. Statistical Analyses To measure percentage of proliferation, we used a quantitative analysis that has been described before (29). It relies on the fact that the size of each subset of proliferating T cells is related to the size of the corresponding precursor subset by a function of 2 n, where n is the number of division cycles achieved in the recipient mice. Therefore if a specific fluorescence peak corresponding to n mitotic divisions contains E cells, then the number of precursor T cells (P) which must have divided n times to generate them can be estimated as E divided by 2 n. The total number of original precursor cells needed to account for the observed progeny can be calculated as the sum of the P values for each of the peaks, one for mitotic cycle, in the histogram. The percentage of cells in the recovered population that has given rise to a clone of proliferating progeny in vivo can then be calculated as the ratio of the sum of the P values to the total number of CFSE cells recovered (the sum of P plus the number of CFSE-positive nonproliferating cells). Data are presented in text and figures as the mean ⫾ SEM, and n indicates the number of individual experiments. Comparisons among different varieties of donor cells were made using the paired Student’s t test. RESULTS ⫹
CD8 T Cells from Old Mice Proliferate in Nonirradiated Recipients To determine whether age alters the ability of mouse T cells to proliferate in nonirradiated hosts, T cells from young (6 months) and old (18 months) CB6F1 mice were isolated and labeled with CFSE, a cell-tracking reagent. This dye is retained in cells for several weeks, and the label is inherited by daughter cells after cell division, making it possible to distinguish cells that have not divided from those that have divided one or more times after labeled cells are injected into host animals. CFSE-labeled T cells were injected iv into 4-month-old CB6F1 mice. Two to 12 days later, T cells were isolated from the lymph nodes (LN) and spleens of the recipient mice and aliquots stained with either PE-CD4 or PE-CD8. Figure 1 shows representative results. At two days after injection CD8 low and CD8 hi CFSE-labeled cells were detectable in both LN (Fig. 1,
FIG. 1. CD8 ⫹ T cells from old mice are able to proliferate in nonirradiated recipient mice. T cells were isolated from young and old CB6F1 mice, labeled with CFSE, and then injected into 4-monthold CB6F1 recipient mice. T cells from recipients’ spleen and pooled lymph nodes were isolated 2 or 12 days later and stained with ␣-CD8-PE or ␣-CD4-PE. Representative CD8 or CD4 vs CFSE dot plots of T cells isolated at days 2 and 12 from the lymph nodes of recipient mice injected with CFSE-labeled T cells from either young or old mice.
top panels) and spleen (not shown). As expected, these cells all had equally high levels of CFSE, indicating an absence of proliferation at this early time point. By day 12, however, many of the CD8 hi CFSE-labeled cells recovered from LN (Fig. 1) or spleen (not shown) had CFSE levels that were 50 or 25% of that shown by the brightest CFSE-positive cells, indicating that these cells had divided once or twice in the previous 12-day period. These cells appeared at high frequency only in hosts that had received CFSE-labeled cells from old donors. In contrast, very few CD4 hi T cells from old donors divide in nonirradiated recipient mice (Fig. 1, lower panels), and neither CD4 nor CD8 T cells derived from young donors showed evidence of more than minimal proliferation. A series of experiments was conducted to determine the kinetics of T-cell expansion in vivo, and the data are summarized in Fig. 2, which shows the percentage of CD8 ⫹ and CD4 ⫹ cells from young and old donors, isolated from the lymph nodes and spleens of the recipient mice, that had divided at least once at varying time points after inoculation. The proportion of recovered CFSE-positive CD8 cells that had undergone at least one in vivo mitosis increased steadily over at least 12 days for recipients of cells from old donors, but
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the recovered CD8 cells from 18-month-old donors had divided at least once in vivo, and this value increased significantly (P ⫽ 0.03), i.e., to 35%, in CD8 cells derived from 24-month-old donors. Proliferation of recovered CD4 ⫹ T cells remained low regardless of the age of the donor mice. Phenotype of CD8 ⫹ T Cells That Have Proliferated in Vivo Resembles That of Nonactivated Memory T Cells
FIG. 2. Time course of in vivo proliferation of CD4 and CD8 T cells from young and old donors injected into nonirradiated recipient mice. The bars represent the proportion of injected cells (⫾ SEM) capable of at least one round of proliferation estimated from the number of recovered cells and their inferred number of cell cycles; n represents the number of independent experiments.
remained low for young CD8 cells and for CD4 cells from donors of any age. By day 12 after injection on average 16% of the injected CD8 cells from old donors had been able to divide, in comparison with 3% from CD8 from young donors (P ⫽ 0.003). At day 12 the proportion of CD4 cells from old donors that had divided was only 3%, significantly (P ⫽ 0.004) lower than the proportion of proliferating CD8 cells from aged donors. The number of CFSE-labeled cells recovered at day 12 was similar in recipients of CD8 cells from young donors and CD4 cells from young and old mice (Table 1), but slightly higher in recipients of old CD8 cells, presumably because these cells are undergoing proliferation. Age-Dependent Increase in CD8 Cells That Proliferate in Vivo In order to determine at what age T cells from individual mice acquire the ability to proliferate in nonirradiated recipients, we isolated T cells from donors of different ages and analyzed their CFSE levels after 12 days of in vivo expansion. Figure 3 summarizes these results. We saw no significant proliferation of CD8 ⫹ T cells from mice 12 months or younger; fewer than 3% of the recovered CFSE-labeled cells from these younger donors had diminished CFSE levels. Sixteen percent of
Multiparameter flow cytometry was used to determine the cell-surface phenotype of CFSE-labeled CD8 cells recovered at day 12 from mice injected with T cells from aged donors. Figure 4 shows a representative result. Most of the CD8 ⫹ T cells that have divided in vivo have high levels of CD44 and CD45RA and low levels of CD25 and CD69. In contrast, antigen-driven T-cell proliferation typically leads to increases in cellsurface levels of CD25 and CD69 (30). Thus our results suggest that in vivo proliferation of CD8 cells from old donors is antigen independent. This is in agreement with previous reports where either CD8 ⫹ T-cell clones or nonclonal memory CD8 ⫹ T cells were observed to proliferate after injection into  2 microglobulin knockout mice (25). The phenotype we obtain in these experiments correlates with the phenotype seen in cells undergoing homeostatic proliferation after transfer into irradiated mice (30). After isolation from recipient mice, CD8 cells that proliferated in vivo appear to have a memory phenotype, as indicated by high-level expression of CD44. To see if high levels of CD44 were also present, prior to transplantation, on the precursors of the proliferating population, we purified naı¨ve and memory CD8 ⫹ T cells from old mice and injected them into nonirradiated recipients (data not shown). We found that only memory CD8 ⫹ T cells from old donor mice, i.e., cells with high levels of CD44, were able to divide in nonirradiated recipient mice. CD8 Cells That Are Able to Proliferate in Nonirradiated Recipients Have High Levels of CD28 Although some initial reports (17) suggested that the clonal CD8 cells found in aged mice were CD28 negative, reevaluation using a different anti-CD28 antibody showed that these cells did indeed express CD28 (25). Our own data, shown in Fig. 5, suggest that the levels of CD28 on cells which have proliferated in vivo are typically higher than those that did not divide after injection. All the CFSE-positive CD8 T cells recovered from the 6-month-old donor expressed CD28, but at modest levels (CD28 lo) (Fig. 5, left panel). The middle
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TABLE 1 Percentage of CFSE-Labeled Cells Recovered at Day 12 Old donors (%)
Lymph nodes Spleen
Young donors (%)
CD8 ⫹ T cells
CD4 ⫹ T cells
CD8 ⫹ T cells
CD4 ⫹ T cells
1.16 ⫾ 0.07 0.62 ⫾ 0.03
0.68 ⫾ 0.11 0.45 ⫾ 0.05
0.75 ⫾ 0.07 0.38 ⫾ 0.04
0.98 ⫾ 0.07 0.54 ⫾ 0.04
Note. Table 1 shows the percentages (mean ⫾ SEM) of CFSE-labeled cells in the isolated spleen and lymph node cells from recipient mice 12 days after injection and illustrates the distribution into CD4 and CD8 subsets. CD4 ⫹ and CD8 ⫹ T-cell percentages were determined by quantifying cells from the samples stained with ␣-PE-CD4 and ␣-PE-CD8 (Fig. 2) (n ⫽ 6).
panel shows cells from an 18-month-old donor: in this case 65% of the proliferating cells, and only 12% of the nonproliferating cells, express higher than normal levels of CD28 (CD28 hi cells). The effect was more dramatic in the 24-month-old donor, for which almost all (90%) of the proliferating cells, but only 33% of the nonproliferating cells, were CD28 hi (Fig. 5, right panel). The results shown in Fig. 5 suggested that elevated CD28 levels might help to identify CD8 T cells able to proliferate in vivo. Figure 6 shows results of an experiment to determine if aging alters the proportion of memory CD8 T cells with high levels of CD28. In a series of two experiments, the average number of CD28 hi cells among the memory CD8 population in the
FIG. 3. Effect of donor age on in vivo proliferation of CD4 and CD8 T cells. The upper and lower panels show the percentages of CD8 ⫹ (solid) and CD4 ⫹ (open) cells from the different donors that had divided by day 12, isolated from the lymph nodes and spleens of recipient mice respectively; n represents the number independent experiments; values are means ⫾ SE.
lymph nodes of aged mice was 50.6 ⫾ 13.8% (mean ⫾ SEM); in contrast 18.6 ⫾ 5.6% of CD8 cells from young donors were CD28 hi. We obtained similar numbers in cells from spleen. To understand if the CD28 hi subset of CD8 ⫹ cells was indeed responsible for the proliferation seen in previous experiments, we purified CD28 hi and CD28 lo CD8 ⫹ T cells from old mice and injected them separately into nonirradiated recipients. Figure 7 shows the results of a representative experiment and summary statistics from a series of four such experiments. The data show that only CD28 hi CD8 ⫹ T cells from old donor mice were able to divide in nonirradiated recipient mice. These
FIG. 4. Expression of surface markers on CD8 ⫹ T cells from old mice. T cells were stained after isolation from recipient mice with one of the following biotinylated antibodies for flow cytometry analysis: ␣-CD44, ␣-CD25, ␣-CD45RA, or ␣-CD69. Biotinylated antibodies were detected with streptavidin-CyC. The figure shows dot plots of the various surface markers vs CFSE on the gated CD8 ⫹ cells. The numbers represent the percentage of cells in each quadrant. Similar results were obtained in a second experiment.
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FIG. 5. CD8 cells that are able to divide in vivo have higher levels of CD28. T cells were stained after isolation from recipient mice with biotinylated ␣-CD28. Biotin was detected with SAv-CyC. T cells were gated on the CFSE ⫹ CD8 ⫹ cells. The figure shows CD28 vs CFSE on the gated cells from different-aged donor mice. The numbers represent the percentage of cells in each quadrant. Similar results were obtained in a second experiment.
results suggest that exceptionally high levels of CD28 may help to identify precursors of CD8 cells with abnormal responses to homeostatic regulation for further functional and biochemical study. CD8 ⫹ T Cells That Have Proliferated in Vivo Produce IFN-␥ Accumulation with age of CD8 ⫹ T cells with altered growth control could potentially contribute to the decline in immune responses seen in elderly individuals. In order to understand the possible role of these clones in regulation of the immune system, we measured using intracellular staining their production of some important immunoregulatory cytokines that are known to be produced by at least some CD8 ⫹ T cells, including IFN-␥, IL-4, and IL-10. Figure 8 shows results from
analysis of lymph node cells from mice 12 days after injection of CFSE-labeled T cells from 23-month-old mice. We found that approximately 80% of the CD8 cells that had been able to divide in nonirradiated recipients were able to produce IFN-␥ after brief culture (4 h) with PMA and ionomycin; about 30% of the recovered CFSE-positive but nonproliferating cells were also IFN-␥ producers (Fig. 8, upper panels). Only a small number of the cells (12%) that proliferated in vivo were IL-4 producers (Fig. 8, lower panels). There was no difference in the percentage of IL-4 producing cells between the population that proliferated in vivo and the population that did not proliferate (Fig. 8, lower right panel). We saw no significant production of IL-10 by proliferating or nonproliferating cells (data not shown). To determine if these cells were producing IFN-␥ in their original host prior to injection into recipient mice, we measured this cytokine by intracellular staining on CD8 ⫹ CD28 high T cells from young and old mice. Figure 9 shows results of a representative experiment and summary statistics from a series of three such experiments. The data show that only CD8 ⫹ CD28 high T cells from old mice are able to produce IFN-␥ in their original host. We found that approximately 68% of the CD8 ⫹ CD28 high T cells in old mice were able to produce IFN-␥, as compared to 9% for the CD8 ⫹ CD28 high T cells in young mice. CD8 ⫹ T Cells That Have Proliferated in Vivo Have High Levels of Bcl-2
FIG. 6. A population of CD28 hi in memory CD8 cells from old mice. The figure shows CD28 vs CD44 on cells that have been gated as CD8 ⫹ from a young and an old mouse prior to injection into recipient host. Numbers represent the percentage of cells in each quadrant. Similar results were obtained in a total of two independent experiments.
Accumulation in aged mice of CD8 cells with the ability to expand in nonirradiated hosts could in principle reflect alterations in levels of antiapoptotic or proapoptotic molecules. To test this hypothesis we
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FIG. 7. Only CD8 T cells with higher levels of CD28 are able to divide in vivo. CD28 hi and CD28 lo CD8 ⫹ T cells from old CB6F1 mice (20 –23 months old) were isolated, labeled with CFSE, and injected into nonirradiated syngeneic recipient mice. Twelve days after injection recovered cells from spleens and lymph nodes were stained for CD8 and CD28. T cells from the old mice were gated on the CFSE ⫹ CD8 ⫹ cells. Representative CD28 vs CFSE dot plots of gated cells isolated from lymph nodes of recipient mice injected with either CD28 lo or CD28 hi CFSE-labeled CD8 ⫹ T cells are shown. The numbers represent the percentage of cells in each quadrant. The number of CD28 lo or CD28 hi CD8 ⫹ T cells that divided in vivo was calculated. The right panel shows the mean ⫾ SEM for the percentage of CD28 lo and CD28 hi CD8 ⫹ T cells isolated from the lymph nodes and the spleens of recipient mice that were able to divide in vivo (n ⫽ 4).
measured levels of Bcl-2 and Fas in cells isolated from the lymph nodes and spleens of recipient mice that had been injected with CFSE-labeled T cells from young or
FIG. 8. Cytokine production by CD8 T cells in old mice. T cells from 23-month-old CB6F1 mice were labeled with CFSE and injected into young recipients. Cells from recipients’ spleens and pooled lymph nodes were isolated at day 12 and cultured for 4 h with 100 ng/ml PMA and 500 ng/ml ionomycin. In the last 2 h of culture, 3 M monensin was added. Cells were harvested and stained with ␣-CD8CyC. After fixation and permeabilization, cells were stained with ␣-IFN␥-PE, ␣-IL-4-PE, ␣-IL-10-PE (not shown), or ␣-IgG-PE. T cells from the old mice were gated on the CFSE ⫹ CD8 ⫹ cells. Left panels represent dot plots of cytokine production (IFN-␥ or IL-4) vs CFSE on the gated cells. Right panels show the percentages of CD8 ⫹ isolated from the lymph nodes and spleens of recipient mice that were able to produce the indicated cytokine, both for cells that had not divided in vivo (“undivided”) and for cells that had divided at least once in vivo, as means ⫾ SEM for n ⫽ 2 experiments. The thresholds shown represent the 97% cutoff points for the ␣-IgG-stained controls.
old donors. Figure 10 shows representative results from one of two independent experiments. The levels of Bcl-2 on the CD8 ⫹ T cells from old donors that were able to proliferate in vivo are higher in comparison to Bcl-2 levels of CD8 ⫹ T cells from young donors (Fig. 10). The left panel shows cells from a 6-month-old donor. Consistent with our previous results, most cells showed no evidence of in vivo mitosis. Among these, 75% expressed low, but easily detectable, levels of Bcl-2, and only 25% expressed the higher levels indicated by the threshold shown in the figure. The right panel shows cells from a 23-month-old donor: In this case 81% of the proliferating cells, and only 45% of the nonproliferating cells, expressed levels of Bcl-2 that exceeded the arbitrary threshold. In a series of experiments using four old donors, the mean proportion of Bcl-2 hi cells was found to be 81 ⫾ 3.3% for proliferating cells, and only 46 ⫾ 5% for the nonproliferating cells, as compared to 30 ⫾ 4.3% for young donors (N ⫽ 2). In contrast, we found no difference among proliferating and nonproliferating cells in levels of Fas (data not shown). The fact that CD8 ⫹ T cells that proliferate and survive in vivo have higher levels of Bcl-2 than T cells that did not divide suggests that Bcl-2-mediated resistance to cell death may contribute to their unusual potential for in vivo replication. DISCUSSION
The accumulation, in aging mice, of large clones of CD8 ⫹ T cells (17, 18, 25) is likely to reflect alterations in the ability of some CD8 cells to control division and/or cell death in the face of homeostatic controls that are still poorly understood. The proportion of aged
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FIG. 9. IFN-␥ production of CD8 T cells in old mice. Cells from the spleens and pooled lymph nodes of 7- and 22-month-old mice were isolated and cultured for 4 h with 100 ng/ml PMA and 500 ng/ml ionomycin. In the last 2 h of culture, 3 M monensin was added. Cells were harvested and stained with ␣-CD8-CyC and ␣-CD28-FITC. After fixation and permeabilization, cells were stained with ␣-IFN-␥-PE or ␣-IgG-PE (not shown). T cells from the old mice were gated on the CD8 ⫹ cells. Left and middle panels represent dot plots of cytokine production vs CD28 on the gated cells. Right panes show the percentages of CD8 ⫹ CD28 high T cells isolated from the lymph nodes of young and old mice that were able to produce IFN-␥ as means ⫾ SEM for n ⫽ 2 experiments. The thresholds shown represent the 98% cutoff points for the ␣-IgG-stained controls. Similar results were obtained in spleen samples.
T cells for which these homeostatic mechanisms have gone awry cannot be estimated simply by counting the number of cells within large in vivo clones because smaller clones, if they exist, will not be recognizable by methods based on either TCR V distributions or V spectratyping. It has recently been reported that CD8 ⫹ T-cell clones, as well as memory CD8 ⫹ T cells not demonstrably members of any obvious clone, are able to divide in vivo after transfer into normal animals (25). To explore this phenomenon further, we have measured the proliferative capacity of T cells from young and old mice by flow cytometry using the fluorescent dye CFSE. We found that CD8 ⫹ T cells from old
FIG. 10. CD8 T cells that are able to divide in vivo have high levels of Bcl-2 expression. T cells recovered from recipients’ pooled lymph nodes were isolated at day 12 and stained with ␣-CD8-CyC. After fixation and permeabilization, cells were stained with ␣-Bcl-2PE. T cells from mice injected with T cells from young and old donors were gated on the CFSE ⫹ CD8 ⫹ cells (both panels). Representative Bcl-2 vs CFSE dot plots of gated cells isolated from lymph nodes of recipient mice injected with CFSE-labeled T cells from either young or old donor are shown above. We obtained the same results in a second independent experiment.
donors, but not young donors, were able to proliferate in nonirradiated hosts. In contrast, CD4 ⫹ T cells from mice of any age did not divide in vivo. CD8 ⫹ T cells from old donors continued to proliferate in vivo for at least 12 days. When recovered from recipient mice, CD8 ⫹ T cells that had undergone at least one round of mitosis exhibit high expression of CD44, similar to resting memory cells. By isolating naı¨ve and memory CD8 T cells from old mice, we showed that only the CD44 hi subset of CD8 ⫹ T cells contains cells able to divide in nonirradiated recipient mice. The CD44 lo subset of CD8 cells, which contains most of the naı¨ve CD8 cells, contained no cells able to divide in vivo (31). Donor mice aged 12 months or younger contained no detectable numbers of CD8 ⫹ T cells able to proliferate in vivo, but in mice older than 18 months the CD8 ⫹ T-cell expansions increased with the age of the donor mice. This timing is consistent with previous information about appearance of T-cell clones in aging mice; such clones are typically not detected before the age of 12 months and then increase in frequency with age (11, 17, 18, 28). Using a three-color flow cytometric system that allowed us to study the surface markers of the CFSEtagged cells recovered from recipient mice, we found that the proliferating CD8 cells bear CD25 low and CD69 low phenotypes. Antigen-driven proliferation leads to increases in cell surface CD25 and CD69 expression (30). Thus the low level of CD25 and CD69 on CFSEpositive proliferating cells suggests that the in vivo proliferation is not dependent on activation through the T-cell receptor. Several experiments have shown that, unlike naı¨ve or actively responding CD8 T cells, the persistent population of CD8 memory T cells is not
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dependent on Ag or even MHC class I for its survival (40 – 42). The phenotype we obtain in these experiments correlates with the phenotype seen in cells undergoing homeostatic proliferation after transfer into irradiated mice (30). Interestingly most of the CD8 cells that are able to divide in nonirradiated hosts are CD45RA ⫹. CD8 ⫹ CD45RA ⫹ T cells, compared to CD8 ⫹, CD45RA ⫺ cells, are relatively nonresponsive to TCR stimulation in tests for cytokine production (32–35). It is not yet clear whether the CD8 cells whose proliferative abnormalities cause them to accumulate in aged mice are or are not normally responsive to antigenic stimulation. These cells clearly do produce IFN-␥ after injection into young hosts, but whether they produce this or other cytokines after TCR or antigenic stimulation is still to be determined. It is noteworthy, however, that anergic CD8 ⫹ T cells have been reported to persist and function in vivo (43, 44). Our data show that the CD8 ⫹ T cells that are able to divide in vivo have higher levels of CD28 when compared either with CD8 ⫹ T cells from old donors that did not divide in vivo or with recovered CFSE-positive cells from a young donor. In addition, we noted a population of memory CD8 cells, higher levels of CD28, present only in aged mice, from which emerged nearly all of the CD8 cells that were able to divide in vivo. Our data suggest that exceptionally high levels of CD28 may help to identify precursors of CD8 cells with abnormal responses to homeostatic regulation for further functional and biochemical study. We do not know at this point whether CD28 per se contributes to the altered growth control of these cells or if it only serves as a marker for cells in the subset. We note, though, that CD28 is expressed almost universally on CD8 T cells of both young and old mice (see Figs. 5 and 6 and also Ref. 45) and thus that expression of CD28 is not sufficient to confer abnormal in vivo proliferation. The high levels of Bcl-2 (see Fig. 10) suggest that clonal accumulation of abnormal CD8 cells in old mice might reflect an altered balance among proapoptotic and antiapoptotic factors in these cells. IL-15, previously implicated in the expansion of CD8 cells in aged mice (25), may well help to induce Bcl-2 and thereby diminish susceptibility to programmed cell death. The effects of aging on CD28 expression in the CD8 ⫹ subset are apparently different in humans, in which aging leads to a characteristic loss of CD28, predominantly among CD8 ⫹ T cells (11, 14 –16). This CD28 ⫺ population consists of a subset of memory T cells, and in many respects their phenotype and function are highly suggestive of differentiated armed effector T cells in that they produce IFN-␥ and TNF-␣ and express high levels of granzyme A and perforin. They also contain Ag-specific memory CTL (46, 47) and exert a potent cytolytic activity without requiring deliberate in vitro activation (48, 49). In humans, the CD28 ⫺ T cells
are clonally expanded (11, 14 –16), terminally differentiated lymphocytes with little proliferative response to mitogenic stimulation in vitro (49, 50). Recent data suggest that they derive from their CD28 ⫹ counterpart (50, 51). The etiology of most expansions in healthy individuals is not known. However, studies of T-cell receptor usage suggested that these cells arose in response to antigenic stimulus (14, 52). These clonal expansions are maintained in vivo at a high frequency for long periods of time (11, 14, 16) and are sometimes noted in association with infections (53, 54), autoimmune conditions (55, 56), or neoplasia (57, 58). It is not clear whether aging humans have a specific CD8 ⫹ subset, like the one we document in aged mice, from which clonal expansions are derived, nor whether such a precursor population has unusually high or low levels of CD28 expression. It is also possible that differences in source of cells—spleen and lymph nodes for mouse and blood for humans— contribute to the apparent differences in age effect on CD28 levels. Measurement of cytokine production provides a first step toward understanding the possible role of these expanded CD8 cells in late-life immunodeficiency. The cells do produce IFN-␥ after stimulation with PMA and ionomycin, but produce only low levels of IL-4 and no detectable IL-10. Although IFN-␥ has been shown to be immunosuppressive in some systems (59), a good deal of additional work, including tests for other cytokines and chemokines, will be needed to test the idea that the increased frequency of unregulated, clonal CD8 cells in old age has functional consequences. ACKNOWLEDGMENTS We wish to thank Dr. Charles D. Surh for advice and technical assistance. This work was supported by NIH Grant R01-AG19619. A.O.S. was supported by the Rackham Merit Fellowship and NIH Training Grants GM07315-21 and T32-AI07413. REFERENCES 1. Miller, R. A., The aging immune system: Primer and prospectus. Science 273, 70, 1996. 2. Nordin, A. A., and Collins, G. D., Limiting dilution analysis of alloreactive cytotoxic precursor cells in aging mice. J. Immunol. 131, 2215, 1983. 3. De Paoli, P., Battistin, S., and Santini, G. F., Age-related changes in human lymphocyte subsets: Progressive reduction of the CD4 CD45R (suppressor inducer) population. Clin. Immunol. Immunopathol. 48, 290, 1988. 4. Thoman, M. L., and Weigle, W. O., Partial restoration of Con A-induced proliferation, IL-2 receptor expression, and IL-2 synthesis in aged murine lymphocytes by phorbol myristate acetate and ionomycin. Cell Immonol. 114, 1, 1988. 5. Vie, H., and Miller R. A., Decline with age in the proportion of mouse T cells that express IL-2 receptor after mitogen stimulation. Clin. Mech. Ageing Dev. 33, 313, 1986. 6. Miller, R. A., Jacobson, B., Weil, G., and Simons, E. R., Diminished calcium influx in lectin-stimulated T cells from old mice. J. Cell Physiol. 132, 337, 1986.
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