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* Autoimmunity Branch, National Institute of Arthritis and Musculoskeletal and Skin Diseases (NIAMS),
Experimental Immunology Branch, National Cancer Institute, and
National Institute on Aging, National Institutes of Health, Bethesda, MD 20892
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Abstract |
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was produced by CCR7–CD27+ and to a slightly lesser extent by CCR7–CD27– T cells. IL-4 secretion was predominantly conducted by CCR7–CD27– memory CD4 T cells. Thus, by using both CCR7 and CD27, distinct maturational stages of CD4 memory T cells with different functional activities were defined. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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Notably, however, the use of CCR7 as a marker distinguishing recirculating TCM from cytokine-secreting TEM has been challenged. Thus, it has been reported that the majority of cytokine-secreting human T cells reside in the CCR7-expressing subset and as a result may be capable of lymphoid organ homing (7). Nevertheless, these authors did find a higher frequency of polarized Th1 or Th2 cells in the CCR7-negative TEM subset. Furthermore, new data indicate the presence of "pre-Th1" and "pre-Th2" cells in the TCM compartment, which secrete low amounts of IFN- or IL-4, and can be induced to differentiate into Th1 and Th2 effector memory T cells by antigenic stimulation or in response to homeostatic cytokines (8).
In addition to CCR7, phenotypic expression of other cell surface proteins, especially costimulatory molecules, has been used to study the differentiation of memory T cells. In this respect, CD27, a member of the TNFR family, has also been used to delineate stages of T cell differentiation (9, 10, 11). CD27 is expressed by naive CD4 and CD8 T cells as well as by most memory T cells and has been assigned a costimulatory function. Initial up-regulation of CD27 expression upon TCR engagement is followed by irreversible loss after repeated antigenic stimulation (12, 13). CD27 expression in CD4 memory T cells distinguishes two populations whose distinct properties have been characterized (11). The loss of CD27 correlates with an increase of IL-4 production. Accumulation of these cells has been reported in patients with chronic allergic diseases, rheumatoid arthritis, and in aged donors (11, 14).
Although the combination of CCR7 and CD27 has been used in an attempt to define a maturational pathway of CD8 T cells (15, 16), the differentiation steps of CD4 memory T cells characterized by expression of CCR7 and CD27 have not been defined.
Therefore, the purpose of this study was to characterize the maturational pattern of CD4 memory T cells by expression of CCR7 and CD27.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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The investigated population consisted of 42 healthy individuals (50% women, age: 40.6 ± 2.5 years), whose blood was provided by the Department of Transfusion Medicine, Clinical Center, National Institutes of Health. Informed consent was obtained from all subjects.
PBMC were obtained by centrifugation of heparinized blood over Ficoll-Hypaque (Amersham Biosciences). T cells or CD4+ T cells were negatively selected on a magnetic column using the AutoMACS (Miltenyi Biotec) and the respective isolation kits for Pan T cell or CD4 isolation (Miltenyi Biotec). In some cases, CD4+ T cells were further subjected to positive selection for CD45RO-expressing cells by the CD45RO kit and AutoMACS. All procedures were conducted according to the manufacturer’s instruction. The purity of the resultant population was analyzed by flow cytometry and was generally >95%.
Flow cytometry
For phenotypic analysis, T cells were incubated for 30 min at 4°C with a FITC-coupled mAb against CCR7 (clone no. 150503; R&D Systems), PE-conjugated anti-CD27 mAb (clone no. M-T271; BD Pharmingen), PerCPCy5.5-coupled anti-CD4 mAb (clone no. SK3; BD Biosciences), and allophycocyanin-conjugated anti-CD45RO mAb (clone no. UCHL-1; BD Pharmingen). Abs of the appropriate IgG isotypes were used as negative controls. Naive T cells served as positive control for staining with anti-CCR7 and anti-CD27. For chemokine receptor analysis, CD4+CD45RO+ T cells were isolated by AutoMACS and stained with the above-mentioned FITC-anti-CCR7, PE-anti-CD27, and PerCPCy5.5-anti-CD4, as well as with allophycocyanin-conjugated mAb against CXCR4 (clone no. 12G5; BD Biosciences) or a biotinylated mAb against CXCR5 (clone no. RF8B2; BD Pharmingen), followed by streptavidin-coupled allophycocyanin. Cells were analyzed using a FACSCalibur flow cytometer (BD Biosciences). The acquired data were analyzed using Flow Jo software (Tree Star).
Cell separation by flow cytometry
For cytokine analysis, proliferation assays, assessment of telomere length, and telomerase activity, cells were sorted using a MoFlo flow cytometer (DakoCytomation). Anti-CCR7-FITC, anti-CD27-PE, anti-CD4-Per-CPCy5.5, and anti-CD45RO-allophycocyanin were in some cases supplemented with anti-CD3-PE-Cy7 to avoid purification steps on the AutoMACS and the inherent cell loss. Postsort controls of the respective sorted populations routinely exhibited >95% purity.
Cell culture and cell counting assays
After an initial overnight activation with 10 ng/ml PMA (Sigma-Aldrich) and 1.34 µM ionomycin (Calbiochem), sorted memory T cell subsets were washed and maintained in 50 U/ml IL-2 (Biological Research Branch, Division of Cancer Treatment and Diagnosis, National Cancer Institute) in U-bottom 96-well-plates (Costar). Culture medium consisted of Ultra Culture serum-free medium (BioWhittaker). After 1 wk, cells were stimulated with anti-CD3 (64.1: 1 µg/ml) and cultured for another week, after which they were activated again with PMA/ionomycin as described above. Phenotypic characterization was performed immediately after the last stimulation and after an additional 9 days of incubation in IL-2. Alternatively, cells were stained 3 days after each respective stimulation and after an additional resting period of 4 days.
For analysis of actual cell numbers, 5 x 104 cells from each population tested were stimulated with PMA/ionomycin overnight and then cultured with 50 U/ml IL-2. Cell numbers were assessed by flow cytometry using SPHERO beads (Spherotech) after an additional 72 h of culture. Dead cells were excluded by staining with 7-aminoactinomycin D (Molecular Probes). The proliferative capacity is expressed as the ratio of cells recovered from the various subsets to the cells recovered from the CCR7+/CD27+ subset. Alternatively, proliferation of T cell subsets was measured after stimulation with anti-CD3 mAb. A total of 3 x 104 cells of each respective subset were incubated in triplicate with plate-bound anti-CD3 (64.1: see above), and proliferation was measured after 72 h by [3H]thymidine incorporation.
Cell cycle progression of T cell subsets after stimulation was examined by DNA staining with propidium iodide (PI).
Telomere length measurement
Telomere length of five individuals (age range: 43–72 years; mean: 55.0 ± 4.9) was measured by flow-fluorescence in situ hybridization (FISH) (17). Isolated T cells were incubated in PBS + 1% BSA. Cells were then incubated in the dark for 15 min in a hybridization mixture containing either 0.3 µg/ml FITC-labeled peptide nucleic acid-telomere FISH probe (Applied Biosystems) or without probe for autofluorescence background measurement. Cells were then heated at 86°C for 15 min and hybridized in the dark at room temperature for 1.5 h. Cell bodies were then gently washed and resuspended. LDS 751 (Exciton), a DNA binding dye, was added to the resuspended cells (10 ng/ml), and fluorescence intensities of the telomere probe and LDS were measured by flow cytometry using a FACSCalibur (BD Biosciences). Single lymphocytes were identified by forward scatter and by total DNA content, and these cells were gated for telomere binding fluorescence. Quantitation of telomere length was performed by interpolating the fluorescence signal from the peptide nucleic acid telomere probe to a standard curve generated by beads with known fluorescence units (Bangs Laboratories). Variations in hybridization conditions between experiments were normalized by comparison to an aliquot of a control cell preparation run with each experiment.
Telomerase activity
Naive CD4 T cells and sorted CD4 memory T cell subsets were stimulated overnight with 10 ng/ml PMA plus 1.34 µM ionomycin followed by a 3-day culture period in medium supplemented with IL2. Telomerase activity was assayed three days later using the TRAPeze Telomerase Detection kit (Chemicon/Serologicals) following the manufacturer’s instructions. Telomerase activity in serial dilutions of cell lysates was detected by its ability to extend telomeric repeats on a substrate, the template strand primer. These telomerase products were then amplified by a two-step PCR for 31 cycles and run on a 12% PAGE gel. DNA bands were visualized using SYBR-green (Molecular Probes) and a PhosphorImager (Molecular Dynamics). A competitive internal standard was used to control for the efficiency of the PCR amplification as well as to detect the presence of Taq polymerase inhibitors. Telomerase activity was calculated as the intensity of telomerase products generated divided by the intensity of the internal standard. A heat-inactivated lysate and a buffer-only lysate were used as negative controls, and TSR8 control template and EL-4 lysate were used as a positive control. To control for day-to-day variation, one sample from each experimental group was repeated in a single assay. These values were then used as a reference to normalize the other samples from the same experimental day.
Cytokine analysis
FACS-sorted CD4 memory T cell subsets (1 x 104/well) were activated with PMA/ionomycin overnight, and supernatants were collected after 20 h. Cytokines in supernatants were measured by Cytometric bead array (CBAkit-II; BD Biosciences) according to the manufacturer’s manual using a FACSCalibur (BD Biosciences). To normalize individual variation, the cytokine secretion of CCR7–/CD27+ and CCR7–/CD27– was expressed as a percentage of the secretion observed in the CCR7+/CD27+ population.
Statistical methods
Values are expressed as mean ± SEM, and for statistical analysis Student’s t test was applied.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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Using naive CD45RO– CD4 T cells that uniformly express CCR7 and CD27 as a positive control and isotype-matched control mAb as a negative control, we could identify three major subsets of memory CD4+T cell: the CCR7+/CD27+, the CCR7–/CD27+, and the CCR7–/CD27–. Within CD4+ T cells of healthy individuals, the majority of cells expressed both CCR7 and CD27 (mean ± SEM: 69.8 ± 2.1%). Memory T cells that did not express CCR7 but were CD27 positive accounted for 21.5 ± 1.8%, whereas only 5.6 ± 0.6% of memory CD4+ T cells were CCR7–/CD27– (Fig. 1). The observed patterns of CCR7 and CD27 expression in healthy individuals did not correlate with the proportion of CD4+CD45RO+ total memory cells and did not correlate significantly with the age of the donors (age range: 8–72 years; mean age ± SEM: 40.6 ± 2.5).
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To examine the relationship between the CD4 memory T cell subsets, the effect of in vitro stimulation on cell surface phenotype was assessed. Repeated stimulation altered the phenotype of CCR7+/CD27+ and CCR7–/CD27+ CD4 memory T cell subsets (Fig. 2A). A portion of CCR7+/CD27+ memory CD4 T cells (mean ± SEM: 30.6 ± 4.3%) became CCR7–/CD27+, whereas approximately one-half (mean ± SEM: 46.7 ± 5.0%) became negative for both surface markers. Furthermore, the majority (mean ± SEM: 80.5 ± 6.9%) of the CCR7–/CD27+ cell population lost expression of CD27 and became CCR7–/CD27– after repeated stimulation (Fig. 2A). To examine the stability of the phenotypic change, cells were also assessed after an additional resting period. As can be seen in Fig. 2A, the altered phenotype remained stable during the resting period of 9 days. CD4+ T cells appeared to alter their phenotype in a stepwise manner following activation (Fig. 2B). After one round of stimulation, the majority of the CCR7–/CD27+ converted to a CCR7–/CD27– phenotype (mean: 77.0 ± 1.8%). In contrast, >75% of CCR7+/CD27+ cells lost CCR7 after the first stimulation and then progressively converted to CCR7–/CD27–. The bulk of naive cells retained their CCR7+/CD27+ phenotype after one stimulation (68.4 ± 9.6%), after which they increasingly became CCR7–/CD27– with additional stimulation (Fig. 2B). Finally, the CCR7–/CD27– subset retained its phenotype despite repeated stimulation and did not (re)-express either CCR7 or CD27 (Fig. 2B). These results clearly identify a differentiation pathway of CD4+ memory T cells in which CCR7+/CD27+ cells mature to CCR7–/CD27+ cells and finally to CCR7–/CD27– terminally differentiated memory T cells.
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The subsets were assessed for the capacity to proliferate after in vitro stimulation. Initially, cells were stimulated with PMA/ionomycin so that intrinsic differences in proliferative capacity could be assessed separately from potential changes in TCR signaling. As shown in Fig. 3, A and B, 3 days after in vitro stimulation with PMA/ionomycin, the number of cells detected increased in all populations, except for the CCR7–/CD27– subset. Proliferative capacity decreased from naive cells to CCR7+/CD27+ (Fig. 3A), further declining in the CCR7–/CD27+ T cells. The lowest cell recovery was noted in CCR7–/CD27– memory T cells (Fig. 3B).
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To gain more insight into the balance of cell death and proliferation manifested by the subsets, cell cycle analysis was conducted. A representative example is shown in Fig. 4A. Immediately ex vivo, most cells from each subset were in G0-G1, and at most, 2% (CCR7–/CD27–) were in G2-M (Fig. 4B). Upon stimulation, each of the subsets progressed into the cell cycle to varying degrees as noted in Fig. 4, C and D. Naive and CCR7+/CD27+ memory T cells showed a similar pattern of cell cycle progression with 
30% of the cells in S-G2-M and only a small percentage becoming apoptotic. In contrast, CCR7–/CD27+ and CCR7–/CD27– subsets exhibited similar cell progression patterns that were different from those in naive and CCR7+/CD27+ cells. A rather small percentage of these cells was stimulated to enter S-G2-M, whereas a sizeable portion of the cells become apoptotic (Fig. 4D).
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Previous studies have indicated that telomeres shorten significantly as T cells divide during in vivo differentiation from naive to memory cells. To test the lineage relationship between memory CD4 subsets suggested by the in vitro experiments, we examined the ex vivo telomere length of the respective subsets by flow-FISH. As shown in Fig. 5A, consistent differences in telomere length were observed among these populations. Naive CD4 T cells of five healthy individuals (age range: 43–72 years; mean: 55.0 ± 4.9; 60% female) had significantly longer telomeres than any of the memory CD4 subsets. Moreover, subsets within the memory population had significant differences in telomere length, with longest telomeres in the CCR7+/CD27+ subset, followed by the CCR7–/CD27+ subset, whereas CCR7–/CD27– cells had the shortest telomeres.
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Inducible telomerase activity in memory CD4 subsets
As telomere length is influenced by the activity of telomerase, inducible telomerase activity was examined in CD4 T cell subsets. The populations derived from each normal donor showed very consistent patterns of inducible telomerase activity, which was higher in the naive population than any of the memory subsets (p < 0.002; Fig. 6A). Within the CD4 memory subsets, the CCR7+/CD27+ subset had higher activity than either the CCR7–/CD27+ or CCR7–/CD27– subsets (both p < 0.002), both of which had similarly low telomerase activity levels (Fig. 6B).
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The functional properties of the memory T cell subsets were examined by measuring cytokine secretion after stimulation (Fig. 7). IFN- and IL-4, the main cytokines associated with mature Th1 and Th2 cells, respectively, showed a differential secretion pattern in CD4 memory subsets. The greatest IFN- secretion was observed in CCR7–/CD27+ and less in the CCR7+/CD27+ and CCR7–/CD27– populations. In contrast, as already reported for CD27– cells (13), CCR7–/CD27– cells showed the highest IL-4 production of the three populations, whereas the IL-4 secretion of the CCR7–/CD27+ subset was comparable to the CCR7+/CD27+. Secretion of the anti-inflammatory cytokine, IL-10, was greatest in the CCR7+/CD27+ population, decreased in CCR7–/CD27+ cells, and was even less in the CCR7–/CD27–. IL-2 production showed an expected decrease from CCR7+/CD27+ to CCR7–/CD27+ and CCR7–/CD27–.
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To investigate the trafficking potential of the CD4 memory T cell subsets, the expression of CXCR4 and CXCR5 was examined. As shown in Fig. 8, the expression of CXCR4 decreased from CCR7+/CD27+ > CCR7–/CD27+ > CCR7–/CD27–. As with CXCR4, a decrease in CXCR5 expression was observed from CCR7+/CD27+ to CCR7–/CD27+ and CCR7–/CD27–. The expression of CXCR4 and CXCR5 in the CCR7+/CD27+ was significantly (p < 0.05) higher than in the CCR7–/CD27+ subset.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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CCR7+/CD27+ memory CD4 T cells CCR7–/CD27+ to the terminally differentiated CCR7–/CD27– CD4 memory T cells.
In normal individuals, the majority (70%) of memory T cells found in blood bear both CCR7 and CD27 on their surfaces, 20% express only CCR7, and only a very small proportion (5%) of memory cells do not express either markers. An even lower percentage (3.0 ± 0.3%, data not shown) exhibited the CCR7+/CD27– phenotype. We did not investigate this subset and its function is currently unknown.
In vitro culture experiments demonstrated a stepwise conversion of CCR7+/CD27+ into a CCR7–/CD27+ population and the progression of the CCR7–/CD27+ into a CCR7–/CD27– subset. Because the induced phenotypes were maintained after a resting period, a transient change of expression of either marker by recent stimulation (10, 18) seems unlikely. The conclusion that there is a stepwise down-regulation of CCR7 and then CD27 during CD4 memory T cells maturation is consistent with the finding that stimulated naive CD4 T cells rapidly down-regulated CCR7, whereas CD27 only declined at a later round of proliferation (19). A similar finding was recently observed in bulk cultures of alloreactive CD45RO+ memory T cells (20).
As the down-regulation of CD27 exhibited similarities to the loss of CD28 with replicative senescence (21), we examined the expression of CD28 by the various CD4+ memory T cell subsets. In accordance with previous findings (22), the CD28- cells were confined to the CCR7-/CD27- subset, constituting about half of this population (data not shown). Thus, the CD28- memory cells represent a portion of the CCR7-/CD27- subset. Notably, the marked decrease in proliferative capacity appeared to be characteristic of the entire CCR7-/CD27- subset, and not a unique property of the CD28- fraction.
Progressive decrease in telomere length from naive CD4 T cells to the CCR7+/CD27+ memory CD4 T cell population and further to the CCR7–/CD27+ subset and the CCR7–/CD27– CD4 memory cells substantiated the model of stepwise CD4 T cell maturation and expanded upon the known correlation of telomere shortening and maturation from the naive to memory stage (23) in CD4 T cells. These results have clearly delineated the CCR7– population into two subsets based on the expression of CD27 and also established the lineal relationship between the previously designated CCR7+ TCM and the previously designated CCR7– TEM. Previous data have not clearly demonstrated whether TCM and TEM are independent subsets or stages of maturation along a continuum of differentiation. The current data clearly support the latter conclusion.
The intrinsic in vitro proliferative capacity of the memory subsets differed. This was demonstrated initially with PMA/ionomycin stimulation to bypass possible variations in TCR signaling and confirmed by stimulation with anti-CD3. In previous reports, memory CD45RO T cells have been shown to exhibit a higher turnover rate and a relatively short lifespan compared with naive T cells (24, 25, 26); furthermore, recently TEM of healthy individuals were found to manifest a higher in vivo turnover rate compared with TCM and naive CD4 T cells (27). These results are consistent with the telomere length analysis conducted here. If TEM had an increased in vivo turnover than TCM, it would be anticipated that they would have shorter telomeres, as we found in the CCR7– subsets, indicating that the TEM had undergone more rounds of in vivo proliferation than TCM. However, the high turnover of TEM shown in the aforementioned study differed from the diminished in vitro proliferative capacity of both CCR7– subsets shown here. These results suggest that there might be in vivo constraints on proliferation by CCR7+ TCM, perhaps related to increased requirements for productive activation. Alternatively, TEM may have greater in vivo exposure to Ag presented by the appropriate APC. Importantly, the in vivo experiments did not examine the potential proliferative capacity as was examined here by cell cycle analysis, cell counting by FACS beads and thymidine incorporation. Together, these data are most consistent with the conclusion that proliferating TCM eventually give rise to TEM with diminished in vivo and in vitro proliferative potential. The endogenous signals that may contribute to the somewhat increased in vivo proliferation of TEM remain to be delineated.
The shortening of telomere length in dividing cells is counteracted by telomerase, an enzyme that is able to extend telomeres by synthesis of terminal telomeric repeats. Although regulation of telomerase activity in lymphocytes is not completely understood, decreased induction of telomerase activity has been observed with both repeated in vitro stimulation, as well as in vivo differentiation from naive to memory cells, and in parallel with telomere length shortening (28, 29). In line with these findings, we observed decreases in induced telomerase activity in the CCR7–/CD27+ and CCR7–/CD27– subsets with shorter telomeres. However, as telomerase activity might be influenced by stimulation and subsequent entry into the cell cycle (27), the lower proportion of in vitro-stimulated cycling cells in the CCR7–/CD27+ and CCR7–/CD27– compared with naive or CCR7+/CD27+ memory cell subsets could account at least in part for the reduced telomerase activity of the CCR7– subsets.
Another feature of maturation and differentiation of cells is the acquisition of certain functional properties such as cytokine secretion. In this respect, a difference in cytokine secretion between TCM and TEM has been postulated (1) but also disputed in human as well as murine models (5, 7), whereas the loss of CD27 is commonly associated with higher IL-4 production (13, 30). We found a clear pattern of cytokine secretion of the CD4+ T cell subsets. We observed increased IFN- and decreased IL-2 secretion by CCR7–/CD27+ and CCR7–/CD27– cells compared with the CCR7+/CD27+ population. This result is consistent with the conclusion that there is an association of "effector" cytokine secretion with the loss of CCR7 expression. However, the TEM population, as defined by CCR7– only, was not homogeneous, and the subdivision into CCR7–/CD27+ and CCR7–/CD27– cells revealed different cytokine secretion patterns. Whereas the production of IL-2 and IFN- was somewhat greater in the CCR7–/CD27+ compared with the CCR7–/CD27– cells, IL-4 production was predominantly conducted only by CCR7–/CD27– memory T cells.
These findings are consistent with previous results that IFN- secretion is initiated early after activation and then increases with successive cell cycles, whereas IL-4 production only appears after several cell divisions (31). The current data indicate that the CCR7–/CD27– population constitutes the most terminally differentiated state of memory cells, enriched in IL-4-producing cells and IFN- secretors. The CCR7–/CD27+ subset represents an earlier maturational stage consisting of cells that have already been biased toward Th1 cells but also of cells that are not yet irreversibly committed and might become Th2 cells in the following maturational step to CCR7–/CD27– cells. Interestingly, the pattern of IL-10 secretion was dissociated from IL-4 production, as production of this anti-inflammatory cytokine was predominantly confined to the CCR7+/CD27+ cells.
The enhanced IL-10 production by CCR7+/CD27+ cells might indicate that a pool of T regulatory 1 (Tr1) cells resides within the CCR7+/CD27+ subset. Tr1 cells suppress immune responses and are known to express CCR7 (reviewed in Ref.32). Because Tr1 cells display a low proliferative capacity, it is somewhat surprising that they were prominently found in the CCR7+/CD27+ subset that exhibited the greatest proliferative capacity.
The loss of IL-10-producing cells in the CCR7–/CD27+ and CCR7–/CD27– T cell populations is consistent with the conclusion that TEM, which enter tissue, are devoid of this population of regulatory cells so as to ensure maximal capacity to respond to pathogens peripherally. The current results are compatible with the possibility that regulation of immune responses by Tr1 cells may largely occur in secondary lymphoid organs.
Another characteristic of cell maturation and differentiation is the change in trafficking potential. The central role of CXCR4 in the trafficking and maintenance of hemopoietic cells has been described previously (33, 34). It is likely that the tendency to recirculate to lymphoid organs through the interaction of this receptor with its ligand, CXCL12 (stromal cell-derived factor 1), declines with maturational state. In this respect, the unique homing potential of the terminally differentiated CCR7–/CD27– subset has been addressed previously (6) and the predominance of CCR7–/CD27– T cells in gut, lung, and liver has been demonstrated. Similar to CXCR4, a reduction of CXCR5+ cells was observed in the more differentiated T cell subsets. The ligand for CXCR5, CXCL13, is a chemokine expressed in B cell follicles and serves to attract follicular helper T cells that have an enhanced capacity to provide help to B cells (35) It is possible that a fraction of the CXCR5+CCR7+/CD27+ cells represents follicular helper cells, although previous studies have shown that CD27– memory T cells have an enhanced capability of providing B cell help (11). This makes it more likely that the follicular helper cells may reside within the CCR7–CD27– population. The decrease in CXCR5 expression in the more mature stages of CD4 memory T cells might again represent a decreased inclination of the majority of these cells to circulate through lymphoid tissue as these cells differentiate and is in accordance with recently published findings (8).
Altogether, the data presented in this study lead to a more detailed view of human CD4 memory T cell maturation. We propose a model of differentiation, in which the sequential stages can be distinguished by the phenotypic expression of CCR7 and CD27. The results clearly indicate a stepwise conversion of CCR7+/CD27+ T cells via the stage of CCR7–/CD27+ to a terminally differentiated CCR7–/CD27– state. The relationship between these subsets was demonstrated by the stepwise unidirectional differentiation in vitro, as well as differences in telomere length, cell cycle analysis, and proliferative capacity. Furthermore, as the cells mature, they acquire different functional properties illustrated by a change in cytokine secretion and chemokine receptor expression. Thus, the defined subsets represent different maturational stages of CD4 memory T cells with unique functional properties
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Acknowledgments |
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Disclosures |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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Footnotes |
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1 This work was supported by a Max Kade Foundation Postdoctoral Research Exchange Grant (to R.D.F.). 
2 R.D.F. and X.S. contributed equally to this work.
3 Address correspondence and reprint requests to Dr. Peter E. Lipsky, National Institute of Arthritis and Musculoskeletal and Skin Diseases, National Institutes of Health, Building 10, Room 6D47C, Bethesda, MD 20892. E-mail address: lipskyp{at}mail.nih.gov
4 Abbreviations used in this paper: TCM, central memory T cell; TEM, effector memory T cell; HEV, high endothelial venule; PI, propidium iodide; FISH, fluorescence in situ hybridization; Tr1, T regulatory 1.
Received for publication May 17, 2005. Accepted for publication September 1, 2005.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionDisclosuresReferences |
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-chemokine, thymus-derived chemotactic agent 4, with activity on T lymphocytes and mesangial cells. J. Immunol. 159:5671.-5679. [Abstract]
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D. Verhoeven, S. Sankaran, M. Silvey, and S. Dandekar Antiviral Therapy during Primary Simian Immunodeficiency Virus Infection Fails To Prevent Acute Loss of CD4+ T Cells in Gut Mucosa but Enhances Their Rapid Restoration through Central Memory T Cells J. Virol., April 15, 2008; 82(8): 4016 - 4027. [Abstract] [Full Text] [PDF]
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B. Santner-Nanan, N. Seddiki, E. Zhu, V. Quent, A. Kelleher, B. F. de St Groth, and R. Nanan Accelerated age-dependent transition of human regulatory T cells to effector memory phenotype Int. Immunol., March 1, 2008; 20(3): 375 - 383. [Abstract] [Full Text] [PDF]
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A. P. Soares, T. J. Scriba, S. Joseph, R. Harbacheuski, R. A. Murray, S. J. Gelderbloem, A. Hawkridge, G. D. Hussey, H. Maecker, G. Kaplan, et al. Bacillus Calmette-Guerin Vaccination of Human Newborns Induces T Cells with Complex Cytokine and Phenotypic Profiles J. Immunol., March 1, 2008; 180(5): 3569 - 3577. [Abstract] [Full Text] [PDF]
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T. J. Scriba, B. Kalsdorf, D.-A. Abrahams, F. Isaacs, J. Hofmeister, G. Black, H. Y. Hassan, R. J. Wilkinson, G. Walzl, S. J. Gelderbloem, et al. Distinct, Specific IL-17- and IL-22-Producing CD4+ T Cell Subsets Contribute to the Human Anti-Mycobacterial Immune Response J. Immunol., February 1, 2008; 180(3): 1962 - 1970. [Abstract] [Full Text] [PDF]
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S. L. Shiao, N. C. Kirkiles-Smith, B. R. Shepherd, J. M. McNiff, E. J. Carr, and J. S. Pober Human Effector Memory CD4+ T Cells Directly Recognize Allogeneic Endothelial Cells In Vitro and In Vivo J. Immunol., October 1, 2007; 179(7): 4397 - 4404. [Abstract] [Full Text] [PDF]
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A. N. Akbar and M. Vukmanovic-Stejic Telomerase in T Lymphocytes: Use It and Lose It? J. Immunol., June 1, 2007; 178(11): 6689 - 6694. [Abstract] [Full Text] [PDF]
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C. Romagnani, K. Juelke, M. Falco, B. Morandi, A. D'Agostino, R. Costa, G. Ratto, G. Forte, P. Carrega, G. Lui, et al. CD56brightCD16- Killer Ig-Like Receptor- NK Cells Display Longer Telomeres and Acquire Features of CD56dim NK Cells upon Activation J. Immunol., April 15, 2007; 178(8): 4947 - 4955. [Abstract] [Full Text] [PDF]
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C. Riou, B. Yassine-Diab, J. Van grevenynghe, R. Somogyi, L. D. Greller, D. Gagnon, S. Gimmig, P. Wilkinson, Y. Shi, M. J. Cameron, et al. Convergence of TCR and cytokine signaling leads to FOXO3a phosphorylation and drives the survival of CD4+ central memory T cells J. Exp. Med., January 22, 2007; 204(1): 79 - 91. [Abstract] [Full Text] [PDF]
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M. Stubbe, N. Vanderheyde, M. Goldman, and A. Marchant Antigen-Specific Central Memory CD4+ T Lymphocytes Produce Multiple Cytokines and Proliferate In Vivo in Humans J. Immunol., December 1, 2006; 177(11): 8185 - 8190. [Abstract] [Full Text] [PDF]
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