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Memory Functions and Death Proneness in Three CD4⁺CD45RO⁺ Human T Cell Subsets¹

Laboratory of Immunology, Department of Radiobiology, Radiation Effects Research Foundation, Hiroshima, Japan

	Abstract

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We propose a classification of human CD4⁺CD45RO⁺ memory T cells into three new subsets based on cell surface expression levels of CD43. The first subset consists of cells whose CD43 expression is relatively high; this subset also contains the highest proportion of recall Ag-reactive precursors, and its constituent cells respond far more strongly than cells in either of the other subsets to immobilized CD3 Ab in addition to secreting substantially more IFN- and IL-4. Cells of the second subset express similar levels of CD43 to naive cells, and they also respond weakly to TCR-mediated stimuli as judged by either their ability to proliferate or capacity for cytokine production. The third subsets consists of cells whose CD43 expression levels are clearly down-regulated; its cells appear to be anergic to TCR-mediated stimuli, and when examined ex vivo many of them appear to be undergoing either spontaneous apoptosis via a caspase-independent pathway or Fas-mediated apoptosis via a caspase-dependent pathway, even in the resting state. An analysis of telomere lengths revealed that the typical telomere of a cell in the second subset was significantly longer than the typical telomere in the first or third subset. Taken together, these results appear to indicate that CD4⁺CD45RO⁺ T cells fall into three functionally differing subsets, one being a subset of cells with fully matured memory phenotype, a second being a less mature subset of cells that retain longer telomeres and whose memory functionality is marginal, and a third consisting of anergic cells that give every appearance of being death-prone and/or in the process of dying.

	Introduction

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Memory T cells generated during a primary response appear to survive for long periods and are therefore available to provide immediate protection as well as to assist in the provision of rapid and effective responses if and when a relevant Ag is re-encountered (1, 2, 3). However, we know very little about the mechanisms involved in the generation and persistence of memory T cells, or about the factors that control their activities in vivo. Murine CD4 and CD8 memory T cells appear to have much less stringent requirements for TCR-MHC interactions for their long-term homeostatic proliferation and survival than do naive T cells, insofar as can be judged by kinetic analyses of T cells expressing transgenic TCRs following their transfer to MHC-deficient mice (4, 5). Unfortunately, attempts to study the process involved in T cell memory and homeostasis in humans have been hampered by the absence of such well-defined in vivo experimental systems.

Studies of the in vivo kinetics of mature T cell pools in humans have been heavily reliant upon following the kinetics of restoration of the human T cell compartment after its partial elimination by disease (6, 7), or following its eradication by radiotherapy or chemotherapy (8, 9). In most cases, such studies have involved studying the differentiation, long-term survival, and cell division of human T cells using such cellular markers as chromosome aberrations (10), mutations (11, 12), telomere lengths (9, 13, 14), and TCR excision circles (7, 15, 16); in addition, investigators have tended to use the surface expression of CD45 isoforms to distinguish between naive and memory T cells (2, 17). One problem of the latter approach, though, is that human memory T cells that express the memory type CD45 isoform (RO⁺ or RA^-) are by no means uniform in their functional and maturational properties. Recent papers indicate that human CD4 T cells expressing the memory type CD45 isoform can be divided into two populations that differ with respect to their expression of the homing receptors associated with nonidentical memory functions (18, 19), i.e., CCR7^- memory cells, so called "effector memory cells" that preferentially enter inflamed tissues and perform immediate effector functions, and CCR7⁺ memory cells, "central memory cells" that express lymph-node homing receptors and lack immediate effector functions. It has also been suggested that human peripheral T cells displaying the memory surface phenotype can be anergic, and also may be capable of acting in a regulatory capacity in much the same manner as the better-understood regulatory T cells of mice (20, 21). Our knowledge of the number and nature of possible human memory T cell subsets is therefore somewhat limited and in obvious need of clarification. In this study, we sought to develop a new classification system by using leucosialin (CD43) as a surface marker with which to divide human CD4⁺CD45RO⁺ memory T cells into three distinct subsets; we also sought to provide a detailed characterization of each subset on the basis of its phenotypic and/or functional attributes.

We chose to use CD43, a highly glycosylated transmembrane protein that is expressed in all hematopoietic cells except mature B cells and erythrocytes (22), as a marker in our study primarily because previous workers have suggested that it may have a role in the regulation of several T cell functions (23) including adhesion (24, 25), activation (26), and proliferation (27), in addition to an involvement in cell survival and apoptosis (28, 29). However, we are well aware that there are still many conflicting views about the precise function of CD43 (23, 30, 31). Of particular interest, though, is a recent study in which it is indicated that the up-regulation of CD43 expression in activated CD4⁺ T cells could have a negative effect on activation-induced cell death, and that this could make high levels of expression of CD43 of value as a defining marker for CD4⁺ memory T cells in the mouse (32). However, the evidence cited in support of this idea was not very strong, and we feel that much further experimentation will be required before it is likely to gain widespread acceptance.

In this paper we suggest dividing human CD4⁺CD45RO⁺ T cells into three subsets on the basis of CD43 expression levels. Although we have not yet identified all of the effects of the CD43 protein on the generation and/or longevity of memory T cells, our results to date do seem to indicate that CD43 will be a useful marker in any future nomenclature that may be developed with a view to describing the functional heterogeneity of human memory T cells. Implications of our work for an improved understanding of the events involved in human memory T cell maturation and termination are also discussed.

	Materials and Methods

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Antibodies

A novel CD43 mAb, HSCA-2 (IgG1 subclass), was prepared by immunizing BALB/c mice with KG-1 cells and fusing their spleen cells with NS1 myeloma cells. HSCA-2 mAbs purified from mouse ascites were then labeled with FITC (Sigma-Aldrich, St. Louis, MO). Binding of this labeled Ab to KG-1 cells was completely blocked by other known CD43 mAbs such as DFT-1 (33) (Coulter-Immunotech, Marseille, France) and 1G10 (BD PharMingen, San Diego, CA). HSCA-2 mAb bound to HeLa cell transfectants expressing human CD43 but not to mock-transfected HeLa cells; it also recognized the sialocarbohydrate moiety of a novel CD43 isoform expressed in T, NK, and activated B cells, but not in monocytes or granulocytes (S. Kyoizumi and T. Ohara, manuscript in preparation). This mAb was filed for participation in the 8th International Workshop on Human Leukocyte Differentiation Ags (to be held in Adelaide, Australia). Unconjugated CCR7 mAb, PE-conjugated CD4, CD25, and CD62L mAbs and streptavidin were purchased from BD PharMingen. PerCP-labeled CD4 and CD8 mAbs were from BD Biosciences (San Jose, CA). PE-conjugated CD27, CD28, CD54, and CD95 mAbs were from Coulter-Immunotech. PE-conjugated TCR, CD2, CD29, CD43, CD44, and CD45RO mAbs and Tricolor-labeled CD45RO mAb were from Caltag Laboratories (Burlingame, CA). PE-labeled CXCR3 mAb for a Th1 surface marker (34) was obtained from R&D Systems (Minneapolis, MN) and biotinylated CRTH2 mAb for a Th2 surface marker (35) was kindly provided by Dr. K. Nagata (BML, Kawagoe, Japan). Apoptosis-inducible Fas (CD95) mAb was purchased from Medical and Biochemical Laboratories (Nagoya, Japan).

Cell preparations and flow cytometry

PBMCs from healthy adult volunteers (n = 17) and cord blood mononuclear cells (CBMCs)⁵ (n = 7) were isolated by density centrifugation in Ficoll-Hypaque (ICN Biomedical, Aurora, OH). For triple-color analysis PBMCs and CBMCs were simultaneously stained with either PerCP-labeled CD4 or CD8 in combination with FITC-labeled CD43 and PE-conjugated CD45RO mAbs. CD4⁺ or CD8⁺ lymphocytes were gated on forward/side scatter and PerCP fluorescence. The proportions of CD4⁺CD45RO^- cells (RO^- subset) and CD4⁺CD45RO⁺ cells expressing high (M1 subset), intermediate (M2 subset), and low (M3 subset) levels of CD43 were measured by flow cytometry with FACScan (BD Biosciences) (see Fig. 1A). To analyze the surface phenotype of each subset, CD4⁺ cells were purified by positive enrichment using MACS (Miltenyi Biotec, Bergish Gladbach, Germany). In brief, PBMCs were incubated with magnetic beads conjugated with CD4 mAbs, followed by enrichment of positive cells using an autoMACS device (Miltenyi Biotec) according to the manufacturer’s instructions. Purified CD4⁺ T cells were stained with FITC-labeled CD43 (HSCA-2), Tricolor-labeled CD45RO, and PE directly labeled TCR, CD2, CD4, CD25, CD27, CD28, CD29, CD44, CD54, CD62L, CD95, or CXCR3 mAbs, or PE indirectly labeled CRTH2 (with biotinylation followed by PE-streptavidin), or CCR-7 (followed by biotinylated anti-mouse IgM and PE-streptavidin) mAbs.

View larger version (31K): [in this window] [in a new window]   FIGURE 1. Flow cytometric analyses of CD43 and CD45RO expression in CD4+ (A and C) and CD8+ (B and D) lymphocytes from adults (A and B) and cord blood (C and D) by triple-color immunofluorescence. Four different subsets can be defined within CD4+ cell populations from adult blood: CD45RO+ cells expressing higher (M1), intermediate (M2), and lower (M3) levels of CD43, and CD45RO- (RO-) cells. In each donor a window for the M2 subset was set in a region where the CD43 level was from approximately one-half to 2-fold of the mean CD43 intensity for RO- cells. These four CD4+ T cell subsets in adult lymphocytes differ in forward light scatter (E). Results are representative of 10 different blood samples.

Cell proliferation assays

For T cell responses to recall Ags in bulk culture, 96-well flat-bottom plastic plates were used. Sorted T cells (5 x 10⁴ cells/well) were stimulated with tuberculosis purified protein derivative (PPD; Connaught Laboratories, Willowdale, Ontario, Canada) at 5 µg/ml, tetanus toxoid (TT; Calbiochem, La Jolla, CA) at 5 µg/ml, or inactivated influenza virus (IV) H1N1 (Advanced Immuno Chemical, Long Beach, CA) at 2.5 µg/ml in the presence of autologous monocytes (2.5 x 10⁴ cells/well), which were previously isolated using magnetic beads coated with anti-CD14 Ab (Miltenyi Biotec) and irradiated with x-ray at 30 Gy. The culture medium used for this assay was RPMI 1640 supplemented with 10% human serum. Proliferation was measured on day 5 for PPD and day 7 for TT and IV by adding [³H]thymidine (NEN Life Science Products, Boston, MA) at 1 µCi/well during the last 16 h of culture. All cultures were set up in triplicate. Immobilized CD3 mAb was prepared by binding OKT3 mAb (10 µg/ml in sodium bicarbonate buffer, pH 9.6) in flat-bottom 96-well plates at room temperature for 2 h and then washing the plates with RPMI 1640 supplemented with 10% FCS. For proliferative responses to anti-CD3 mAb, sorted T cells were stimulated with either immobilized CD3 (OKT-3) mAb or soluble CD3 mAb (2 µg/ml) in the presence of autologous monocytes. The culture medium used for this assay was RPMI 1640 medium supplemented with 10% FCS. The effects of soluble CD28 mAb (1 µg/ml; Coulter-Immunotech), human rIL-2 (10 ng/ml; PeproTech, Rocky Hill, NJ), or PMA (10 ng/ml; Sigma-Aldrich) on immobilized CD3 mAb-induced cell proliferation were also evaluated. Incorporation of [³H]thymidine was measured on day 3 during the last 16 h of culture.

Limiting dilution assays

To set up a limiting dilution assay (LDA) of PPD-reactive CD4 T cells, graded numbers of total CD4⁺ T cells or of the four sorted CD4⁺ T cell subsets were seeded into 96-well flat-bottom plates along with 2.5 x 10⁴ x-irradiated (30 Gy) autologous CD14⁺ monocytes as APC. Twenty-four well replicate series per subset were set up for each dilution. The cells were cultured in RPMI supplemented with 10% human serum. After 3 days in culture, human rIL-2 was added to each well at a final concentration of 0.1 ng/ml. After an additional 11 days of culture, cell proliferation was measured by adding [³H]thymidine to each well at a concentration of 1 µCi/well. The response of each well was scored positive when radioactivity exceeded the mean + 3 SD in a set of 24 control wells containing APC in the absence of responder cells. The frequencies of reactive cells were calculated by fitting a generalized linear model with a complementary log-log link using L-Calc limiting dilution analysis software (StemSoft Software, Vancouver, Canada).

Cytokine measurement

T cells were stimulated with immobilized CD3 mAb (OKT-3 mAb) in the presence of CD28 mAb (1 µg/ml), IL-2 (0.1–100 ng/ml), or PMA (10 ng/ml) for 24 or 48 h. Production of cytokines by T cell subsets was measured in the culture supernatants by ELISA using matched pairs of Abs specific for IL-4 and IFN- (BD PharMingen). Briefly, anti-cytokine capture Abs (1 µg/ml) were coated to the wells of an enhanced protein-binding ELISA plate overnight. Nonspecific binding was blocked by incubation with blocking buffer (PBS containing 4% BSA). Culture supernatants and standards were added and incubated for 2 h at room temperature. After washing a plate with blocking buffer, biotinylated anti-cytokine detection Ab (1 µg/ml) was added and incubated for 1 h at room temperature. After washing with blocking buffer, avidin-HRP conjugate was added and incubated for 30 min at room temperature. For color development, 3,3',5,5'-tetramethylbenzidine substrate (Kirkegaard & Perry Laboratories, Gaithersburg, MD) and H₂O₂ (0.01%) were added and incubated for 30 min before absorbance at 405 nm was recorded.

Induction and detection of cell death

For the induction of cell death in culture, PI-unstained viable T cells from the M1, M2, M3, and RO^- subsets were separated in a cell sorter and then cultured for 16–72 h with RPMI supplemented with 10% FCS in the presence and absence of Fas mAb. The involvement of caspases in spontaneous and Fas-mediated cell death was evaluated by adding the peptide inhibitor Z-VAD-fmk (Sigma-Aldrich) to the cultures at various concentrations. Possible inhibitory effects of IL-2 on cell death were also examined. Cell death was detected by staining cultures with FITC-labeled annexin V (Coulter-Immunotech) according to the manufacturer’s instructions. Briefly, fresh or cultured T cells were washed with PBS and resuspended in a binding buffer (140 mM NaCl, 2.5 mM CaCl₂, 10 mM HEPES (pH 7.4)). The cell suspensions were then mixed with 0.5 µg/ml FITC-labeled annexin V and 1 µg/ml PI and incubated at room temperature for 15 min in the dark. To detect intracellular activated caspases, the cell suspensions were incubated with an FITC conjugate of Z-VAD-fmk (Promega, Madison, WI) at 10 µM for 20 min (36). The fluorescence signals from FITC and PI were measured by flow cytometry using a single laser FACScan. Apoptotic cell death was confirmed by detecting the cleavage of DNA into oligonucleosomal fragments using an Apoptosis Ladder Detection kit (Wako Pure Chemical, Osaka, Japan).

Telomere length measurement

We measured the telomere lengths of CD4 T cells using fluorescence in situ hybridization as previously described (14, 37). In situ hybridization was performed using 1 x 10⁵ cells from each CD4 T cell subset in a hybridization mixture containing 70% formamide solution, 20 mM Tris (pH 7), 0.1% BSA, and either 0.3 µg/ml telomere-specific FITC-labeled peptide nucleic acid probe (C₃T A₂; Sawady Technology, Tokyo, Japan) or peptide nucleic acid probe specific for the alphoid sequences of the X chromosome as a control for background fluorescence. Samples were subjected to heat denaturation of DNA for 10 min at 80°C followed by hybridization overnight at 20°C. The cells were then washed with 70% formamide solution, 20 mM Tris (pH 7), 0.1% BSA, and 0.1% Tween 20. Following a final wash without formamide, the cells were suspended in PBS/0.1% BSA containing 10 µg/ml RNase A and incubated for 2 h at room temperature. Before flow cytometry, 0.06 µg/ml 7-aminoactinomycin-D (7-AAD) was added to each cell suspension and fluorescence from the FITC-probe and 7-AAD was analyzed by FACScan. Interphase cells were gated on 7-AAD fluorescence and forward scatter to obtain the fluorescence histograms derived from FITC-labeled telomere or control probes. The specific telomere fluorescence of cells was calculated by subtracting the mean background fluorescence from the mean fluorescence obtained with the telomere probe. We confirmed that intraindividual variation of telomere fluorescence is very low (coefficient of variations is <5% for triplicate samples). Also, we obtained constant data for each individual when tests were done at different experimental times (coefficient of variations was 5% for three different samplings). To estimate telomere length from telomere fluorescence we used a calibration curve obtained by plotting telomere fluorescence against the values for telomere length obtained by Southern blot analysis for PBMC (n = 16) and CBMC (n = 2). The lengths of the mean terminal restriction enzyme fragments were determined by hybridization with the appropriate oligonucleotide telomere probe on a Southern blot. The slope of the calibration curve (Y = 39.0X, R² = 0.79) was used for the estimation of telomere length (Y in base pairs) from telomere fluorescence (X in arbitrary units).

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
CD4⁺CD45RO⁺ T cell subsets differentially express CD43

To study possible variations in CD43 expression levels in human memory T cells, we analyzed human adult peripheral blood and cord blood T cells by three-color flow cytometry using mAbs against CD43 (HSCA-2), CD45RO, and CD4 or CD8 (Fig. 1, A–D). We found that adult CD4⁺ T cells could be divided into four subsets; one was CD45RO^- and expressed an intermediate level of CD43 (designated the RO^- subset). There were also three CD45RO⁺ subsets: an M1 subset consisting of cells that expressed high levels of CD43; an M2 subset consisting of cells that expressed intermediate levels of CD43; and an M3 subset consisting of cells that expressed low levels of CD43 (Fig. 1A). HSCA-2 mAb turned out to be the most useful of the CD43 mAbs for separating these three CD45RO⁺ subsets in flow cytograms (data not shown). In contrast to our findings with adult blood, almost all of the cord blood CD4⁺ T cells proved to be CD45RO^- cells expressing intermediate levels of CD43 (Fig. 1C). Adult CD8⁺ T cells were also found to contain cells in which CD43 expression was either up- or down-regulated (Fig. 1B). We now describe the more detailed results of our analyses of adult CD4⁺ T cells.

The proportions of the three adult CD4⁺CD45RO⁺ T cell subsets (see above) that we could detect in a group of 10 normal donors (mean age, 43 years; age range, 27–52 years) were as follows: M1, 14.3 ± 6.5% (mean ± SD); M2, 17.5 ± 4.5%; M3, 3.9 ± 1.5%. The forward scatter histograms indicated that the M1 subset cells were the largest of the CD4⁺ T cells, while the M3 subset cells were larger than RO^- cells but smaller than either M1 or M2 cells in all 10 donors (Fig. 1E). By making use of three-color flow cytometry of MACS-purified CD4 T cells (Table I), we found that M1 cells express significantly higher levels of coreceptor and costimulatory molecules (such as CD2, CD4, and CD28) than either M2 or M3 cells. Expression of adhesion molecules such as CD29, CD44, and CD54 also appeared to be expressed at higher levels in M1 subset cells than in M2 or M3 cells. The proportion of cells with an effector memory (CD27^- or CCR7^-) phenotype was higher in the M1 subset than in the M2 or M3 subset. Interestingly, the expression levels of CD25 and CD62L cells were somewhat higher in the M3 subset than in the other subsets. Also, CD95 expression levels were slightly higher in the M3 subset than in the M1 or M2 subsets. The M1 subset appeared to contain proportionally more Th1 cells than the others, while the M3 subset appeared to contain proportionally more Th2 cells than the others.

To analyze memory functions in the three CD4⁺CD45RO⁺ T cell subsets, we examined the proliferative responses of each subset to appropriate recall Ags (PPD, TT, and IV presented by autologous CD14⁺ APC) in bulk culture (Fig. 2). M1 subset cells responded more strongly than M2 and M3 subset cells to the test Ags. M2 cells did respond, albeit very weakly, while M3 cells did not appear to respond at all. To estimate precursor frequencies (PF), we performed LDA of detectably PPD-reactive cells in the various CD4⁺ T cell subsets from two healthy donors (Fig. 3). We found that the M1 subset contained the highest frequency of precursors in both donors and that the M2 subset contained less than one-eighth as many; the M3 subset had an even lower PF than the M2 subset.

View larger version (15K): [in this window] [in a new window]   FIGURE 2. Proliferative responses to recall Ags in CD4+ T cell subsets. MACS-purified CD4+ T cells were sorted by FACS into the indicated subsets and thereafter stimulated with PPD, TT, or IV in the presence of autologous CD14+ APC. Proliferation was measured on day 5 (PPD) or 7 (TT and IV) by adding [3H]thymidine during the last 16 h of culture. Results are expressed as mean cpm ± SD and are representative of three donors.

View larger version (31K): [in this window] [in a new window]   FIGURE 3. LDAs of proliferative responses to PPD Ag in the four CD4+ T cell subsets from two donors whose PFs of the total CD4 T cells were the lowest (upper panel) and the highest (lower panel) of six laboratory controls. The PF as calculated by regression analysis is in parentheses.

View larger version (31K): [in this window] [in a new window]   FIGURE 4. Proliferative responses to CD3 mAb in CD4+ T cell subsets. Purified CD4+ T cell subsets were stimulated with immobilized CD3 mAb (iCD3; A and B), soluble CD3 mAb (sCD3; 2 µg/ml) in the presence of CD14+ APC (A), or immobilized CD3 in the presence of soluble CD28 mAb (1 µg/ml) or IL-2 (10 ng/ml) (B). Proliferation was measured on day 3 following addition of [3H]thymidine for the last 16 h of culture. Results are expressed as mean cpm ± SD and are representative of three donors.

The cytokine production profile of each CD4⁺ T cell subset was measured following stimulation with immobilized CD3 mAb in the presence or absence of IL-2 (Figs. 5, B and C, and 6), CD28 mAb, or PMA (Fig. 6). Of the four subsets, M1 cells secreted most IFN- and IL-4 in response to stimulation with CD3 mAb plus IL-2, CD28 mAb, or PMA, almost certainly reflecting the fact that the M1 subset contains many mature Th1 and Th2 cells as judged by their expression of Th1 or Th2 surface markers (Table I). M2 subset cells produced lower levels of IFN- or IL-4 than M1 subset cells in both the presence and absence of secondary signals. Similarly, M3 subset cells produced only extremely low levels of IFN- and IL-4 in response to CD3 stimulation both with and without CD28 mAb but appeared to produce substantial amounts of IL-4 in the presence of IL-2 or PMA.

View larger version (18K): [in this window] [in a new window]   FIGURE 5. Effects of IL-2 on immobilized CD3 mAb-induced proliferation and cytokine production in CD4+ T cell subsets. Purified CD4+ T cell subsets were stimulated by immobilized CD3 mAb in the presence of various concentrations of human IL-2. Proliferation was measured on day 3 following the addition of [3H]thymidine for the last 16 h of the culture (A). Production of IFN- (B) and IL-4 (C) was measured in supernatants of the 2-day cultures by ELISA. Results are representative of three donors.

View larger version (23K): [in this window] [in a new window]   FIGURE 6. Capacity to produce IFN- and IL-4 in the CD4+ T cell subsets. Purified CD4+ T cell subsets were stimulated with immobilized CD3 mAb, either alone or in combination with soluble CD28 mAb (1 µg/ml), IL-2 (10 ng/ml), or PMA (10 ng/ml). Supernatants were collected on day 1 after initiation of the cultures. Cytokine levels of the supernatants were determined by ELISA. Results are expressed as mean ± SD and are representative of three donors.

We considered the possibility that M3 subset cells might not survive very well in culture, as an explanation for our inability to detect a response to CD3 mAb. To test this possibility, we isolated and cultured viable PI^- cells in each CD4⁺ T cell subset and followed their fate, paying particular attention to death-associated events, by flow cytometry (Fig. 7). Only a small percentage of the PI^- cells displayed typical death-associated phenotypes with respect to changes in cell size (Fig. 7A) or annexin V binding (Fig. 7B) in the immediate aftermath of cell isolation. The percentages of dying cells were significantly greater in the M3 subset than in the other subsets after 16 h in culture but increased slowly in all subsets if culture was prolonged beyond 24 h (Fig. 8A). After 16 h of culture, isolated annexin V⁺ M3 cells exhibited typical apoptotic features, including the nuclear shrinkage and fragmentation associated with chromatin condensation (Fig. 9A). Apoptosis of M3 subset cells was confirmed by DNA electrophoresis, with oligonucleosomal DNA fragmentation becoming obvious in the M3 population as a whole but even more obvious in isolated annexin V⁺ M3 cells (Fig. 9B). The addition of anti-Fas mAb to the culture medium significantly enhanced apoptosis in the M3 cell population but had little or no effect on the cells of the other subsets (Fig. 8). These findings indicate that M3 subset cells are especially prone to both spontaneous and Fas-mediated apoptosis.

View larger version (26K): [in this window] [in a new window]   FIGURE 7. Flow cytometric analysis of spontaneous apoptosis in CD4+ T cell subsets. Purified CD4+ T cell subsets were cultured in RPMI supplemented with 10% FCS for 16 h. Apoptosis was followed by light scattering of the RO- and M3 subsets (A) and by binding of FITC-annexin V or FITC-Z-VAD-fmk to the M3 subset (B). A region for positive staining was determined using Fas mAb-treated Jurkat cells. Values are the percentages of positive cells. Results are representative of five donors

View larger version (20K): [in this window] [in a new window]   FIGURE 8. Spontaneous and Fas mAb-induced apoptosis in the CD4+ T cell subsets. Purified CD4+ T cell subsets were cultured in the presence and absence of anti-Fas mAb for various times. Time courses of spontaneous cell death in the RO-, M1, M2, and M3 subsets, Fas mAb-induced cell death in the M3 subset (A), and dose dependency of Fas mAb-induced cell death at 16 h (B) were analyzed by flow cytometry using FITC-annexin V. Results on the dose dependency (B) are expressed as the mean ± SD for three donors. *, The percentage of annexin V+ cells in the M3 subset was significantly larger in the presence of Fas mAb than in the absence of Fas mAb by t test (n = 3, p < 0.01).

View larger version (91K): [in this window] [in a new window]   FIGURE 9. Analysis of spontaneous cell death in the M3 subset cells as judged by cellular morphology (A) and DNA ladder formation (B). Purified CD4+ T cell subsets were cultured for 16 h and both annexin V+ and annexin V- cells in cultured M3 subset were sorted. Cytospin preparations of annexin V+ (A, upper panel) and annexin V- (A, lower panel) M3 cells were then stained with May-Grunwald-Giemsa. A bar indicates 10 µm. B, Total DNA extracted from RO-, M1, total M3, annexin V+ M3, and annexin V- M3 cells were electrophoresed with markers (HaeIII-digested 174 DNA).

View larger version (18K): [in this window] [in a new window]   FIGURE 10. Effects of Z-VAD-fmk and IL-2 on spontaneous and Fas mAb-induced apoptosis in the M3 subset. Purified M3 subset cells were cultured in the presence and absence of anti-Fas mAb for 16 h. Various concentrations of Z-VAD-fmk (A) and IL-2 (B) were added to the cultures at the time of initiation. Cell death was measured by flow cytometry using FITC-annexin V. The concentration of DMSO in the mock experiment (A) was 0.05%, equivalent to that used in the experiment with 25 µM Z-VAD-fmk. Results are representative of five different blood samples.

To obtain some insight into the in vivo growth kinetics of the various CD4⁺ T cell subsets, we decided to determine the average telomere lengths of representative cells of the various CD4⁺ T cell subsets by flow cytometry following hybridization with a FITC-labeled telomere probe (Fig. 11A). Analysis of the relevant cell populations from five healthy donors revealed that the cells of all three CD45RO⁺ subsets had telomeres that were significantly shorter than those of RO^- cells, although interindividual variation was relatively large (Fig. 11B). In addition, we found that the telomeres of M1 and M3 subset cells were of approximately equal length but were shorter than a typical M2 cell telomere. These findings could well imply that the M1 and M3 subset cells are descendants of M2 subset cells.

View larger version (28K): [in this window] [in a new window]   FIGURE 11. Flow cytometric measurement of telomere lengths in the CD4+ T cell subset cells. A, Representative flow histogram of telomere and background fluorescence was obtained by hybridization with telomere and control probes, respectively, in the CD4+ T subset cells from a healthy 37-year-old donor (B, •). B, The means of specific telomere fluorescence were compared among the CD4+ T cell subsets and the CBMC from five healthy donors (age range, 34–56 years) and seven neonates, respectively. The specific telomere fluorescence of cells was calculated by subtracting the mean background fluorescence from the mean fluorescence obtained with the telomere probe. Statistical significance was determined by Wilcoxon signed-rank test (*, p < 0.05). N.S., Not significant.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

We consider that the cells that make up the largest subset of CD4⁺CD45RO⁺ T cells (the M2 subset) are intermediate between naive cells and M1 memory T cells and are best described as premature memory T cells. Support for this proposition comes from the following observations. First, the telomeres of cells in M2 subset cells appear to be considerably longer than those of fully matured memory M1 cells. Thus, M2 cells may actually mature into M1 cells after an additional series of cell divisions. We now have obtained preliminary evidence that indicates that the addition of CD3 mAb can stimulate M2 cells to proliferate while at the same time causing a marked increase in their CD43 expression levels (our unpublished observation). Second, there appear to be fewer recall Ag-reactive precursor cells in M2 populations than in M1 populations. Third, M2 cells appeared to respond only very weakly—as assessed by both their proliferative response and their cytokine outputs—when exposed to immobilized CD3 mAb in the absence of secondary signals. Additional signals in the form of APC, CD28, or IL-2R enhanced the CD3-stimulated proliferative activities of M2 cells to a level more typical of M1 cells, perhaps indicating that their TCR-mediated cell proliferative capacities are much more reliant on secondary signals than those of M1 cells. Interestingly, though, it became evident that M2 cells were able to produce only a relatively small amount of IFN- and IL-4, regardless of the availability or otherwise of secondary signals.

The M3 subset is the smallest of the three CD45RO⁺ T subsets, and its cells are noteworthy mainly because they seem to be both anergic and unusually prone to cell death. They display no discernible reactivity to immobilized CD3 mAb in the absence of costimulatory signals, even though their TCR/CD3 expression levels are similar to those of M1 or M2 subset cells. This seeming unresponsiveness of M3 cells to TCR stimuli could be a function of their death proneness in culture, but because 50% of the cells in an M3 cell population appear to be capable of surviving for 3 days in culture in either the presence or absence of immobilized CD3 mAb (data not shown), it seems reasonable to assume that the surviving cells are themselves anergic. Moreover, IL-2 is capable of causing at least partial reversion of anergy in the M3 cell population. Stimulation of M3 subset cells with IL-2 or PMA together with immobilized CD3 mAb also appeared to stimulate IL-4 production but not IFN- production. These results may indicate that the M3 subset contains proportionately more Th2 cells than other subsets. This is consistent with our findings from Th1/Th2 surface marker analysis. With regard to IL-2 effects, it should be noted that more than half of the M3 subset cells express CD25. The surface phenotypes and anergy associated with M3 cells indicate that they have a resemblance to previously described murine (21, 43) and human (20, 44, 45, 46, 47, 48) CD4⁺CD25⁺ T cell populations that display regulatory activity. However, the M3 subset cells we have been studying seem to be functionally different from these so-called regulatory T cells, if only because they did not appear to be capable of suppressing the CD3-stimulated proliferation of M1 and naive cells in either the presence or the absence of APC (data not shown).

We suspect that the spontaneous apoptosis of M3 cells that we observed is mainly mediated by a caspase-independent signal transduction pathway, if only because only 20% of the total apoptotic activity appeared to be sensitive to Z-VAD-fmk. Also, only almost one-third of the annexin V⁺ M3 cells in a 16-h culture could be stained with FITC-Z-VAD-fmk. Thus, it can be assumed that spontaneous apoptosis of M3 cells is different from conventional lymphokine withdrawal cell death, which involves activation of a caspase (49, 50). There is already some good evidence that both caspase-independent and nonclassical forms of apoptosis occur in mature lymphocytes (51, 52, 53, 54) and thymocytes (55). Interestingly, we found that spontaneous apoptosis of M3 cells could be at least partially prevented by the addition of IL-2. This finding is consistent with previous reports showing that IL-2 can prevent the spontaneous cell death of resting human CD4⁺CD45RO⁺ (56) and CD4⁺CD25⁺ (47) T cells, but unfortunately the caspase dependency of cell death was not determined in either case. Therefore, we conclude that IL-2 could be acting as a survival factor in human resting T cells that are anergic and prone to apoptosis, probably by interfering with the signaling pathway that leads to spontaneous apoptosis.

Treatment with Fas mAb appeared to significantly enhance killing of M3 subset T cells. Unlike spontaneous apoptosis, Fas mAb-induced apoptosis can be completely blocked by Z-VAD-fmk and hence appears to be a caspase-activated pathway. This may mean that a Fas-mediated caspase activation pathway operates in M3 subset cells even in resting state, although it is unknown whether this pathway is similar to that observed in activation-induced cell death (50). By contrast, M1 and M2 subset cells showed no sensitivity to Fas-mediated death despite their ability to express Fas on their cell surfaces. A detailed comparative study of the signaling pathways involved in spontaneous and Fas-mediated apoptosis is necessary if we are to learn more about the differential sensitivities of resting T cell population to cell death signals (50).

There is some experimental evidence that indicates that CD43 takes part in cell signaling pathways in T cells (22, 23), possibly by helping to costimulate T cells (26, 27). Therefore, it seems reasonable to assume that CD43 play a part in certain of the cell signaling events that are likely to be involved in memory T cell activation. However, recent reports suggest that CD43 molecules of T cells may be actively excluded from the immunological synapse and that this event is mediated by relocalization of cytoskeletal adapter proteins that bind to CD43 (57). These suggestions led to a proposal that movement of CD43 could modulate T cell activation by sequestering negative regulatory proteins away from the site of TCR signaling (58). This could mean that the up-regulated expression of CD43 in M1 subset cells causes an increase in activation signaling, perhaps in concert with other up-regulated costimulatory and/or adhesion molecules. Because high-level expression of CD43 appears to protect T cell hybridoma cells against activation-induced cell death, at least in the mouse (32), it may be that one net effect of down-regulating CD43 expression is the sensitization of M3 T cells to apoptotic cell death. Thus, although the precise molecular mechanisms underlying the costimulatory and antiapoptotic effects of CD43 remain to be determined, it seems reasonable to assume that CD43 expression levels will prove to be important in the control of both cell activation and cell survival processes in memory T cells.

Based on the findings reported in this work, we will outline some of the key events and processes that we believe are likely to be involved in the maturation and eventual termination stages of the life span of human CD4 memory T cells. Soon after Ag exposure, naive T cells may undergo repeated cycles of cell division associated with the shortening of their telomeres. Calculations based on estimated telomere length shortening per mitosis (75 bp) (14), together with observed differences in average telomere length between RO^- and M2 cells (3700 bp), combine to suggest that 50 population doublings are involved in the transformation of stimulated T cells into cells at the premature memory stage herein designated the M2 stage. Approximately 15 further population doublings then appear to be required for the conversion of M2 cells into M1-type cells, by which time they are behaving as typical fully functioning mature memory T cells. M3 cells, by contrast, behave more like cells that are approaching the end of their ability to function as normal CD4 T cells. Such cells may well arise from fully mature M1 cells, possibly when M1-type cells begin to lose some of their key properties as they approach senescence, although it is unclear as yet whether this type of senescence process is associated with activation-induced or cytokine withdrawal cell death following M1 cell activation. We can also envisage circumstances in which activated premature M2 cells with recall Ags are directly transformed into death-prone M3-type subset cells without first having been converted into fully mature memory M1-type cells as in the pathway outlined above. Such a balance between the two putative pathways (M2M1M3 and M2M3) could be very important element in the maintenance of CD4 memory T cell pools.

	Acknowledgments

 
We are grateful to Mika Yamaoka for her excellent assistance with FACS analysis, Mika Yonezawa for manuscript preparation, and Dr. Donald MacPhee for his valuable suggestions.

	Footnotes

 
¹ This publication is based on research performed at the Radiation Effects Research Foundation (Hiroshima and Nagasaki, Japan). Radiation Effects Research Foundation is a private nonprofit foundation funded equally by the Japanese Ministry of Health, Labor and Welfare and the U.S. Department of Energy through the National Academy of Sciences. A part of this study was supported by funds for Research Promotion on AIDS Control from the Japanese Ministry of Health and Welfare.

² Current address: Life Science RD Center, Life Science Laboratories, Kaneka Corporation, Takasago-shi, Hyogo, Japan.

³ Current address: Cellular Signal Analysis, Department of Bio-Signal Analysis, Applied Medical Engineering Science, Yamaguchi University Graduate School of Medicine, Ube City, Yamaguchi, Japan.

⁴ Address correspondence and reprint requests to Dr. Seishi Kyoizumi, Laboratory of Immunology, Department of Radiobiology, Radiation Effects Research Foundation, 5-2 Hijiyama Park, Minami Ward, Hiroshima, 732-0815 Japan. E-mail address: kyoizumi{at}rerf.or.jp

⁵ Abbreviations used in this paper: CBMC, cord blood mononuclear cell; PPD, purified protein derivative; TT, tetanus toxoid; IV, influenza virus; 7-AAD, 7-aminoactinomycin-D; PF, precursor frequency; LDA, limiting dilution assay; PI, propidium iodide.

Received for publication January 28, 2002. Accepted for publication April 23, 2002.

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