E2F1 and E2F2 Are Differentially Required for Homeostasis-Driven and Antigen-Induced T Cell Proliferation In Vivo1
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免疫学杂志 2005年第14期
Abstract
Homeostasis-driven T cell proliferation occurs in response to a lymphopenic environment and is mediated by TCR and IL-7 signaling. In this report, we demonstrate a defect in the proliferation of murine naive and memory T cells lacking both E2F1 and E2F2 in response to lymphopenic conditions, suggesting that E2F1 and E2F2 function redundantly downstream of TCR and/or IL-7 signaling during homeostasis-driven proliferation. In contrast, T cell proliferation in response to antigenic stimulation is either unaffected (in vivo) or potentiated (ex vivo) by loss of E2F1 and E2F2, indicating divergent requirements for these E2F factors in T cell proliferation mediated by distinct stimuli. E2F1/E2F2 double knockout (DKO) T cells enter S phase in response to homeostatic signaling, but fail to divide, suggesting that S phase progression is either incomplete or defective. In addition, E2F1/E2F2 DKO mice do not recover normal T cell numbers following exposure to a sublethal dose of radiation, indicating that this defect in homeostasis-driven proliferation is physiologically relevant. Consistent with their failure in cell cycle progression, the differentiation of DKO T cells into memory T cells in response to homeostatic signals is significantly reduced. These observations support the idea that proliferation is required for memory T cell formation and also have implications for the development of clinical strategies to minimize the occurrence of lymphopenia-induced autoimmunity.
Introduction
T cell numbers are maintained at relatively constant levels in the peripheral immune system. Although environmental stimuli, including antigenic stimulation or exposure to lymphotoxic conditions, can result in a temporary increase or decrease, T cell numbers return to normal once the stimuli is removed. The total numbers of cells in both the naive and memory T cell compartments are determined by the relative contributions of lymphocyte generation, proliferation, differentiation, and death. Following exposure to an antigenic stimulus, T cell numbers are elevated. Once the Ag is cleared, many of the responding T cells undergo apoptosis or differentiate into memory T cells, thereby restoring naive T cell numbers to their normal level. In contrast, T cells undergo homeostasis-driven proliferation (HDP)3 in response to deficiencies in T cell numbers, or lymphopenia (reviewed in Ref. 1), and this response also results in the restoration of T cell numbers. HDP has both phenotypic and functional implications for the peripheral T cell compartment. Naive T cells that have undergone HDP express memory T cell surface markers and acquire increased effector function (reviewed in Ref. 2). Thus, both Ag-stimulated proliferation and HDP result in the formation of memory T cells.
Like Ag-stimulated proliferation, HDP of naive T cells requires TCR stimulation by MHC-peptide complexes (3, 4, 5, 6, 7). However, relatively weak TCR stimuli, often mediated by self-peptides, are sufficient for the induction of HDP and this feature may reflect decreased competition for access to APCs expressing the appropriate MHC-peptide complexes under lymphopenic conditions as well as the sensitization of T cells to TCR signaling. IL-7 is also required for HDP of naive T cells, and IL-7 levels are increased in lymphopenic hosts under a variety of conditions (reviewed in Ref. 8). In addition, exogenous IL-7 can stimulate T cell proliferation in vivo (9, 10, 11), consistent with the possibility that IL-7 functions to sensitize T cells to TCR stimuli and thereby promote HDP. These observations define TCR and cytokine signaling as minimal requirements for HDP of naive T cells. The requirements for HDP of memory T cells are both less stringent and less well defined (reviewed in Ref. 1). In the majority of cases, TCR stimulation by MHC-peptide complexes or MHC alone is not required for HDP of memory T cells. In addition, HDP of CD8 memory T cells can be supported by either IL-7 or IL-15, whereas no specific cytokine requirements for HDP of CD4 memory T cells have been defined.
We have been using mouse knockout models to study the roles of E2F1 and E2F2 in T cell proliferation. Members of the E2F family of transcription factors are key regulators of cell cycle progression. E2F1, E2F2, and E2F3 function to regulate expression of a relatively large set of genes that are induced at the G1 to S phase transition during the cell cycle. Many of these target genes encode proteins that are important for cell cycle progression, including cyclin A2, thymidylate synthase, DNA polymerase , and Cdc6 (12). Mutation of both E2F1 and E2F2 or E2F2 alone results in a number of hemopoietic phenotypes in mice. First, consistent with their known roles in the cell cycle, disruption of E2F1 and E2F2 results in defects in the development of a number of hemopoietic compartments due to impeded S phase progression in progenitors of the B cell, erythroid, and myeloid lineages as well as multipotent stem cells (13). These defects in the bone marrow are consistent with the known role for E2F members in promoting cell cycle progression. In contrast, T cells derived from mice mutant for both E2F1 and E2F2 or E2F2 alone not only enter S phase more rapidly than wild-type T cells following antigenic stimulation and proliferate much more extensively, but they also respond to lower levels of Ag (14, 15), indicating a role for E2F2 and, to a lesser extent, E2F1 in setting the TCR signaling threshold for Ag-stimulated proliferation. In addition, E2F2 mutant mice are predisposed to the development of autoimmunity (14). Taken together these observations reveal a somewhat paradoxical role for E2F1 and E2F2 in preventing inappropriate cell cycle progression. Given these results, we were interested in understanding how the loss of E2F1 and E2F2 might affect T cell proliferation in response to homeostatic stimuli. These studies have led to a further understanding of the events that occur downstream of TCR and cytokine signals during T cell HDP. In addition, the experiments we describe have important clinical and mechanistic implications with respect to memory T cell differentiation.
Materials and Methods
Mice
All experiments were performed using 4- to 8-wk-old E2F1/E2F2 double knockout (DKO) or control (E2f1+/+E2f2+/+, E2f1+/+E2f2+/–, E2f1+/–E2f2+/–, or E2f1+/–E2f2+/+) mice in either the B10.D2 (backcrossed for two to four generations) or BALB/c (backcrossed for three to five generations) backgrounds. No differences in HDP were observed between wild-type and E2F1 or E2F2 heterozygous T cells. DO11.10 TCR transgenic (DO TCR Tg) (16), E2F1 mutant (17), and E2F2 mutant (14) mice were obtained from Dr. P. Marrack (Howard Hughes Medical Institute, National Jewish Medical and Research Center, Denver, CO) or Dr. M. E. Greenberg (Children’s Hospital and Harvard Medical School, Boston, MA). Pure-bred recipient mice (B10.D2 or BALB/c) were obtained from The Jackson Laboratory. Mice were housed in the University of Colorado Health Sciences Center animal resource center and all animal procedures were performed according to Institutional Review Board approval protocols. Genotypes were determined by PCR analysis using genomic DNA isolated from tail biopsies. Where indicated mice were exposed to a single dose of gamma radiation from a cobalt 60 source at a distance of 25 cm. Complete blood counts were determined from a small sample of peripheral blood obtained from the tail and were measured using a Cell-Dyn 1700 hematology analyzer (Abbott Laboratories). For isolation of memory and naive T cells by single-cell sorting, DO TCR Tg, E2F1/E2F2 DKO and control mice were injected s.c. with 2.5 mg of chicken OVA protein (Sigma-Aldrich) in PBS plus 50 μl of CFA (Sigma-Aldrich) 32 days before sorting to induce the formation of DO TCR Tg memory T cells.
CFSE staining and lymphocyte transfers
Single-cell suspensions in PBS containing 1% FBS (FBS/PBS), 1 mM MgCl2, and 100 U/ml DNase I (Sigma-Aldrich) were obtained from lymph nodes and strained through nylon mesh. Where indicated, lymphocytes were stained in PBS containing 3 μM CFSE (Molecular Probes) for 15 min at 37°C. Both CFSE-labeled and unlabeled lymphocytes were washed in PBS for 30 min and were transferred to recipient mice by subocular or i.p. injection. Irradiated recipients were exposed to 450 rad of gamma radiation 1 day before transfer. Recipients received 2–8 x 106 lymphocytes/mouse. Transferred cells were recovered from the lymph nodes and spleens of recipient mice 14–21 days later or as indicated and analyzed.
Flow cytometry
Single-cell suspensions obtained from lymph nodes and/or spleen were strained through nylon mesh and washed once in PBS. In some cases, CD4+ T cells isolated from DO TCR Tg mice were purified by MACS using IMag CD4 MSC particles (BD Pharmingen) to facilitate efficient staining and analysis. Cells were stained in Ab solution (5 μg/ml fluorochrome-linked Abs, 1:50 2.4G2 tissue culture supernatant containing anti-FcR III/II Ab, and 1:100 goat serum (Invitrogen Life Technologies) in FBS/PBS) at a concentration of 108/ml and analyzed by flow cytometry. For single-cell sorting, RBC, macrophages, and B cells were depleted from combined lymph node and spleen suspensions by MACS using Ter119 microbeads (Miltenyi Biotec), CD11b microbeads (Miltenyi Biotec), and IMag anti-mouse CD45R/B220 particles (BD Pharmingen). The remaining cells were stained as described, and Thy1.2+ memory (CD44high, CD62Llow) and naive (CD44low, CD62Lhigh) T cells were purified by single-cell sorting. PE- and allophycocyanin-streptavidin conjugates and allophycocyanin anti-CD4, allophycocyanin anti-CD8, FITC anti-CD90.2 (Thy1.2), FITC anti-CD62L, PE anti-CD122, allophycocyanin anti-CD44, PE anti-CD127 (IL-7R), and PE anti-mouse IgG1 isotype control Abs were obtained from BD Pharmingen. Biotin- and fluorochrome-linked anti-DO TCR (KJ1.26) mAbs were obtained from Caltag Laboratories.
Stimulation of peripheral lymphocytes
For in vitro stimulation, single-cell suspensions obtained from lymph nodes of E2F1/E2F2 DKO and control mice or from the spleens of pure-bred mice were strained through nylon mesh, washed in PBS, resuspended at a final concentration of 1.0 x 107/ml in DMEM containing 10% FBS (HyClone Laboratories), 0.1 mM 2-ME, and 1% penicillin-streptomycin (Invitrogen Life Technologies), and plated in tissue culture dishes. Lymph node suspensions isolated from DO TCR Tg mice were combined 1:1 with nontransgenic splenocytes, which function as APCs, and stimulated with OVA protein (Sigma-Aldrich). Alternatively, lymph node suspensions were cultured alone and stimulated with 5 ng/ml PMA (Sigma-Aldrich) and 500 ng/ml ionomycin (Sigma-Aldrich). For in vivo stimulation, 107 CFSE-labeled lymphocytes were transferred to unirradiated recipients as described. Recipient mice received a single dose of OVA protein (Sigma-Aldrich) in PBS by i.p. injection 16–20 h after the lymphocyte transfer.
BrdU staining
Peripheral T cells were stimulated in vitro, as described, in the presence of 10 μM BrdU (Boehringer Mannheim). Alternatively, recipient mice were maintained on water containing 0.8 mg/ml BrdU and 10% sucrose, made fresh daily, to assess BrdU incorporation in vivo. In vitro cultures were harvested 48–52 h after stimulation. For in vivo experiments, single-cell suspensions were isolated from the lymph nodes and spleen of recipient mice at the indicated times and CD4+ T cells were isolated by MACS as described. Lymphocytes were exposed to UV-B for 10 min before staining with PE anti-DO TCR Ab (Caltag Laboratories), FITC anti-BrdU (DakoCytomation) Ab, and 7-aminoactinomycin D (7-AAD) as previously described (18). BrdU incorporation and DNA content in DO TCR Tg T cells was measured by flow cytometry.
Memory T cell surface marker and intracellular cytokine staining
Unlabeled DO TCR Tg lymphocytes were transferred to irradiated recipients. For memory and naive T cell controls, lymphocytes were isolated from a DO TCR Tg mouse and combined with B10.D2 lymphocytes at a ratio of 1:50. Aliquots of all samples were stained with Abs against memory cell surface markers and analyzed by flow cytometry or stimulated in vitro with PMA and ionomycin, as described. GolgiPlug (1 μl/ml; BD Pharmingen) was added to in vitro cultures to inhibit the export of cytokines. Cultures were harvested after 2 h, stained with PE anti-DO TCR and allophycocyanin anti-CD44 as described, fixed in 4% paraformaldehyde for 15 min at 4°C, and stored in PBS plus 1% BSA. The next day, fixed cells were incubated in 1x BD Pharmingen Perm/Wash buffer for 15 min at 4°C to permeabilize and stained with 5 μg/ml FITC anti-IL-2 (BD Pharmingen) plus 1:50 2.4G2 tissue culture supernatant containing anti-FcR III/II Ab in 1x BD Pharmingen Perm/Wash buffer at a concentration of 108/ml. Stained cells were washed two times for 20 min at 4°C in 1x BD Pharmingen Perm/Wash, resuspended in 1% FBS/PBS, and analyzed by flow cytometry.
Results
E2F1 and E2F2 are required for homeostasis-driven T cell proliferation
To determine whether E2F1 and E2F2 play a role in T cell HDP, we assessed the ability of T cells isolated from E2F1 mutant, E2F2 mutant E2F1/E2F2 DKO, or control mice to proliferate upon transfer to a lymphopenic host. Lymph nodes were harvested from DO TCR Tg E2F1/E2F2 DKO or control mice and stained with CFSE, a fluorescent dye that is taken up by live cells. CFSE-labeled cells were transferred to syngeneic wild-type recipients that had been rendered lymphopenic by exposure to a sublethal dose of gamma radiation. T cells were harvested from the lymph nodes and spleen of recipient mice 2–3 wk later, stained with fluorescent-linked Ab against the DO TCR (or other T cell markers, depending on the nature of the specific experiment), and the CFSE profiles of DO TCR+, CFSE+ donor-derived T cells were assessed by flow cytometry. With each round of division, the daughter cells inherit one-half of the CFSE of their parent and each generation of cells therefore has a characteristic fluorescence, which serves as an indicator of proliferation.
As shown in Fig. 1, E2F1/E2F2 DKO T cells exhibit defects in HDP. T cells isolated from control mice undergo significant proliferation in irradiated hosts (Fig. 1A). Cells that have undergone between 0 and 3 rounds of division are evident. In contrast, E2F1/E2F2 DKO T cells do not proliferate in irradiated recipient mice, indicating that E2F1, E2F2, or both together are required for this process. Consistent with this observation, the number of donor-derived T cells recovered from the lymph nodes and spleens of irradiated recipients of E2F1/E2F2 DKO T cells was 60% of the number recovered from recipients of control T cells, irrespective of whether the donor T cells were labeled with CFSE (Fig. 1B). Additional experiments revealed that T cells isolated from E2f1–/–E2f2+/– or E2f1+/–E2f2–/– mice do not exhibit defects in proliferation in irradiated hosts (Fig. 1C), suggesting that E2F1 and E2F2 function redundantly to promote proliferation under these conditions. As expected, when T cells isolated from mice of any of these genotypes are transferred into unirradiated recipients, they do not divide (Fig. 1A and data not shown), indicating that the observed proliferation requires lymphopenia and is therefore occurring in response to homeostatic signals. Thus, E2F1 and E2F2 function redundantly to promote T cell HDP in irradiated hosts. In addition, the lymphocyte compartment in DKO mice did not recover following exposure to a sublethal dose of gamma radiation, whereas lymphocyte numbers returned to near normal levels in wild-type mice (Fig. 1D and data not shown). At the last time point shown, significant reconstitution of the thymus in terms of cell number had not occurred in mice of either genotype and mature single positive thymocytes were not detected (data not shown), suggesting that reconstitution of the peripheral T cell compartment was predominantly a result of HDP, rather than thymopoiesis. Taken together, these observations indicate that the defect in HDP exhibited by E2F1/E2F2 DKO T cells results in a physiologically relevant deficit in recovery of the T cell compartment in response to lymphopenia.
Although E2F1/E2F2 DKO T cells exhibit significant defects in T cell HDP, this phenotype is not completely penetrant and in some cases, there is a small but distinct population of E2F1/E2F2 DKO T cells that do appear to proliferate in irradiated hosts (Figs. 1, A and B, and 2A). This observation raises the possibility that different T cell subsets may have different requirements for E2F1 and E2F2 for HDP. To investigate this possibility, we assessed the induction of HDP in several T cell subsets. The defect in HDP exhibited by E2F1/E2F2 DKO T cells is not restricted to a single T cell compartment, as both CD4+ and CD8+ T cells isolated from nontransgenic DKO mice exhibit this phenotype (Fig. 2A). In addition, both naive (CD44low, CD62Lhigh) and memory (CD44high, CD62Llow) T cells isolated from OVA-injected DO TCR Tg E2F1/E2F2 DKO mice exhibit defects in HDP (Fig. 2B). This observation also suggests that the defects in HDP that occur as a result of mutation of E2F1 and E2F2 are not due to abrogation of IL-7 signaling, as IL-7 is not required for HDP of CD4+ memory T cells (19). Consistent with this idea, DO TCR Tg T cells isolated from DKO and control mice exhibited similar cell surface expression of the IL-7R (Fig. 3A). Similarly, the defects in HDP exhibited by E2F1/E2F2 DKO T cells do not appear to occur as a result of differences in survival in irradiated hosts. Apparent precursor frequencies can be calculated for both DKO and control samples based on CFSE staining profiles and the absolute number of T cells recovered from recipient mice. For instance, in one experiment, 36.3% of the donor-derived control T cells recovered from irradiated recipients had not divided, 31.8% had divided once, 15.1% had divided twice, 10.3% had divided three times, and 6.6% had divided four times. For each generation, the minimum number of donor T cells that the recovered cells were derived from can be calculated by multiplying the total number of donor-derived T cells recovered by the percentage of donor-derived T cells in that generation and then dividing by 2x, where x is the number of rounds of proliferation that cells in that generation have gone through. In this example, 1.44 x 106 donor-derived T cells were recovered from the lymph nodes and spleen of an irradiated recipient of control cells and these cells were therefore derived from a total of 1.44 x 106 (0.363/20 + 0.318/21 + 0.151/22 + 0.103/23 + 0.066/24), or 8.31 x 105 cells. Similarly, 8.62 x 105 E2F1/E2F2 DKO T cells were recovered from an irradiated recipient and, because DKO T cells do not undergo proliferation in lymphopenic hosts, the apparent precursor frequency in this case is 8.62 x 105. Although this calculation does not provide information regarding the fraction of transferred cells that survive in irradiated recipients, it does indicate relative survival of E2F1/E2F2 DKO and control T cells in irradiated recipients. Using this method, apparent precursor frequencies were calculated and no significant difference between the precursor frequencies for DKO or control T cells were observed (Fig. 3B). Because similar numbers of T cells were transferred to recipients of both DKO and control T cells (the average ratio of donor-derived T cells recovered from E2F1/E2F2 DKO versus control recipients 1 day after transfer was 1.08 ± 0.06), apparent precursor frequency is a direct indicator of relative survival in these experiments. Thus, the observed differences in HDP of E2F1/E2F2 DKO and control T cells do not reflect a difference in their capacity for survival in irradiated hosts.
E2F1/E2F2 DKO T cells enter S phase in response to homeostatic signals
The phenotypes that have been observed in the bone marrow and the peripheral immune system suggest two distinct possibilities in terms of the underlying cause for the failure of E2F1/E2F2 DKO T cells to progress through the cell cycle in response to homeostatic signals. First, in the absence of E2F1 and E2F2, homeostatic signals may not stimulate sufficient accumulation of E2F targets to promote progression into S phase. In this case, E2F1/E2F2 DKO T cells would remain in G0 and G1 phases of the cell cycle following transfer into a lymphopenic host. Alternatively, E2F1/E2F2 DKO T cells may enter S phase but be impeded in S phase progression, similar to the phenotype exhibited by hemopoietic progenitors in the bone marrow (13). To address this issue, T cells were isolated from DO TCR Tg E2F1/E2F2 DKO or control mice and transferred to irradiated recipients. The recipients were maintained on BrdU-containing water for varying intervals of time after the transfers and BrdU incorporation and DNA content in donor-derived T cells were determined by flow cytometry. Parallel samples were stained with CFSE before transfer into irradiated recipients to monitor proliferation. Both E2F1/E2F2 DKO and control T cells exhibited significant incorporation of BrdU between 0 and 6 days after transfer into irradiated recipients indicating that DKO T cells do enter S phase (Fig. 5, A and B), despite the fact that they do not proliferate (Fig. 5D). Moreover, BrdU-positive cells with G1, S, and G2 DNA content were observed in both samples, indicating that E2F1/E2F2 DKO T cells not only initiate S phase, but undergo significant DNA synthesis in a lymphopenic environment (Fig. 5C), despite the fact that they do not divide (Fig. 5D). At later time points, an increasing percentage of donor-derived control cells exhibited BrdU incorporation. In contrast, the percentage of E2F1/E2F2 DKO T cells that incorporate BrdU decreases slightly between 6 and 15 days posttransfer. As shown in Fig. 3C, it is unlikely that E2F1/E2F2 DKO T cell survival in lymphopenic hosts is affected by loss of E2F1 and E2F2, and this decrease in the incidence of cells entering S phase is therefore consistent with the idea that E2F1/E2F2 DKO T cells are undergoing a cell cycle arrest, presumably in S phase. These data suggest that the defect in HDP due to mutation of E2F1 and E2F2 occurs by a similar mechanism to the failure in cell cycle progression that is manifested in hemopoietic precursors in the bone marrow.
E2F1 and E2F2 are required for homeostasis-driven memory T cell differentiation
Naive T cells that have undergone HDP develop a memory T cell phenotype. To determine whether E2F1/E2F2 DKO T cells acquire a memory phenotype following transfer to lymphopenic hosts, we examined the expression of memory T cell surface markers on E2F1/E2F2 DKO and control T cells after recovery from irradiated recipient mice. For control samples, DO TCR Tg lymphocytes were combined with nontransgenic lymphocytes at a ratio of 1:50, so that the density of both naive and memory phenotype DO TCR Tg T cells was similar in control samples and those derived from irradiated recipients. The comparison of populations with similar densities of Tg lymphocytes ensures that differences in gating on the DO TCR Tg fraction of the population do not contribute to observed differences in cell surface staining. Before transfer into recipient mice, E2F1/E2F2 DKO and control T cell populations exhibited staining patterns similar to naive T cell samples (data not shown). After recovery from lymphopenic recipients, control T cells exhibited a statistically significant increase in the proportion of CD44high memory T cells relative to the untransferred control sample (Fig. 6A). A substantial fraction of these CD44high cells exhibited altered expression of additional memory T cell markers relative to naive T cell samples, including IL-2R (CD122) and/or CD62L (Fig. 6B). Up-regulation of CD122 and down-regulation of CD62L were similar on endogenous memory T cells and memory T cells that had differentiated in response to homeostatic stimuli. In addition, CD44high memory T cells isolated from lymphopenic recipients of control T cells exhibited increased effector function as indicated by the rapid production of IL-2 in response to stimulation with PMA and ionomycin in vitro (Fig. 6B). Thus, control T cells that have undergone HDP exhibited an increase in the proportion of T cells expressing memory phenotypes, relative to populations derived from naive mice. In contrast, the expression of memory markers was not significantly different on E2F1/E2F2 DKO T cells rescued from lymphopenic hosts as compared with the untransferred control population (Fig. 6A). A small fraction of these DKO T cells do express memory markers and exhibit increased effector function (Fig. 6). However, it is not clear whether E2F1/E2F2 DKO T cells expressing a memory phenotype developed from naive T cells as a result of homeostatic signaling or were present in the original transferred population. In either case, memory T cell differentiation is not efficiently induced in response to irradiation-induced lymphopenia in the absence of E2F1 and E2F2.
Discussion
We have shown that the E2F1 and E2F2 transcription factors function redundantly to mediate proliferation of primary T cells in response to lymphopenic conditions in vivo, suggesting that E2F1 and E2F2 function downstream of TCR and/or IL-7R signaling during HDP. HDP is defective in multiple T cells subsets isolated from E2F1/E2F2 DKO mice, including CD4+ memory T cells and both CD4+ and CD8+ naive T cells. E2F1/E2F2 DKO T cells enter S phase in response to homeostatic signaling, but fail to divide, suggesting that S phase progression is either incomplete or defective. This phenotype is similar to the E2F1/E2F2-dependent defects in cell cycle progression exhibited by hemopoietic progenitors and may be due to a failure in the accumulation of E2F target genes required for S phase. Taken together, these observations indicate a role for E2F1 and E2F2 in promoting T cell proliferation in response to TCR stimuli in the context of homeostatic signaling in vivo. In contrast, E2F1/E2F2 DKO T cells proliferate in response to antigenic stimulation in vivo and hyperproliferate in response to suboptimal antigenic stimulation in vitro. Thus, E2F1 and E2F2 are differentially required for T cell proliferation in response to distinct TCR-mediated stimuli. Previous studies have shown that T cells undergoing proliferation in response to homeostatic signaling or antigenic stimulation exhibit similar gene expression profiles, suggesting that quantitative differences in TCR and/or cytokine signaling, rather than qualitative differences, distinguish HDP from Ag-mediated proliferation (20). In fact, only one gene was specifically regulated in response to homeostatic stimulation. Thus, the experiments we have described are particularly important in that they identify distinct molecular requirements for T cell proliferation in response to antigenic and homeostatic signaling and suggest a molecular mechanism by which quantitatively different signals can mediate similar effects on T cell proliferation. In addition, these observations reveal distinct aspects of E2F function that either promote or prevent cell cycle progression in response to TCR activation in peripheral T cells, depending on the nature of the stimulus. We are interested in understanding how the T cell activation signals defined in this study impinge on the cellular requirements for E2F function. One possibility is that the activity and/or abundance of E2F family members can be differentially regulated in a single-cell type in response to distinct stimuli or assay conditions. For instance, following TCR stimulation by Ag, E2F3 may be activated to perform functions that are dependent on E2F1 and E2F2 in the context of the weaker TCR signals that mediate HDP. Alternatively, it is possible that E2F-dependent target genes are similarly regulated in response to distinct stimuli, but regulation of these target genes may be differentially required in specific contexts.
What implications do these observations have for the T cell repertoire in E2F1/E2F2 DKO mice? Recent studies suggest that under normal circumstances, HDP is important during early development of the immune system for expansion of the relatively limited neonatal peripheral T cell population (21, 22, 23) and HDP may also function in older individuals to compensate for decreased T cell generation as a result of thymic involution. Thus, the defect in HDP exhibited by E2F1/E2F2 DKO T cells may contribute to the decrease in mature T cell numbers observed in DKO mice (13). This phenotype is expected to worsen with age as thymic output decreases and E2F1/E2F2 DKO T cells cannot compensate due to defects in HDP. Recent studies have also implicated lymphopenia and HDP in the development of autoimmunity (24, 25 and reviewed in Ref. 26). Older E2F1/E2F2 DKO and E2F2-deficient mice exhibit both lymphopenia and autoimmune features (13, 14, 15). Because E2F1/E2F2 DKO T cells exhibit defects in HDP, whereas E2F2-deficient T cells do not, a direct comparison of the timing and extent of the development of autoimmunity in these two genetic backgrounds should reveal the relative contributions of lymphopenia and HDP to this process.
These studies also have implications for the mechanism by which memory T cell differentiation occurs. Both homeostasis-driven and Ag-stimulated proliferation require TCR signaling and it has been previously shown that memory T cell differentiation during HDP is temporally correlated with proliferation such that only cells that have undergone multiple rounds of division express memory markers (27, 28, 29, 30). Based on these observations, it has been proposed that proliferation and TCR signaling are minimal requirements for the development of memory T cells. However, these experiments do not eliminate the possibility that it is the duration or quality of the TCR signal, rather than the extent of proliferation, which determines memory T cell differentiation. Mutation of E2F1 and E2F2 specifically abrogates T cell proliferation in response to homeostatic signals and has no apparent affect on TCR signaling, as indicated by the observation that E2F1/E2F2 DKO and control T cells enter S phase with similar kinetics and at a similar frequency in response to homeostatic signals. Thus, mutation of E2F1 and E2F2 allows for the separation of these two potentially important aspects of HDP-induced memory T cell differentiation, and DKO T cells thereby provide a unique opportunity to assess the effects of changes in proliferation in the absence of changes in TCR signaling. Despite the fact that E2F1/E2F2 DKO T cells receive a TCR signal that is sufficient to induce progression into S phase, they do not efficiently express memory phenotypes. Thus, the experiments presented in this study suggest that homeostatic signaling is not sufficient for the induction of memory T cell differentiation and are consistent with the idea that proliferation is required for efficient memory T cell differentiation.
A final consideration is the clinical relevance of these studies. HDP is important for repopulation of the peripheral immune system following the induction of lymphopenia either therapeutically, due to treatment with radiation or chemotherapies, or as a result of viral infection, particularly by HIV (31). In addition, HDP may contribute to the development of autoimmunity in several ways. Homeostatic TCR stimuli can be delivered by the same self Ags that mediate positive selection during T cell development, thus stimulating the clonal expansion of self-reactive T cells (5, 6, 7). HDP also induces memory T cell differentiation, thereby ensuring a persistent immune response. HDP-induced autoimmunity is thought to play an important role in organ transplant rejection, particularly if peripheral T cell depletion has been used as a method of immunosuppression (32). The studies described in this report suggest that therapeutic strategies using cell cycle inhibitors to suppress the development of graft-vs-host disease mediate protection by preventing both clonal expansion and memory T cell differentiation, thereby limiting the development of HDP-induced autoimmunity. Moreover, the development of strategies that result in specific inhibition of E2F1 and E2F2 may be useful in limiting autoimmunity with little or no effect on T cell activity toward bona fide Ags or proliferation of other cell types. This approach would be most useful in cases in which widespread immune suppression is not desirable, such as for the prevention of autoimmunity as a result of therapy- or virus-induced lymphopenia or due to a genetic predisposition. Further characterization of the molecular mechanism by which E2F1 and E2F2 mediate their role in HDP may reveal downstream effectors that can be more easily targeted for clinical intervention using currently available reagents.
Acknowledgments
We thank Jing Zhu and Feng Li for the initial observations that inspired this project, Dave Hildeman, Tom Mitchell, and Kent Teague for critical review of this manuscript, and the University of Colorado Health Sciences Center Cancer Center Flow Cytometry Core for technical assistance.
Disclosures
The authors have no financial conflict of interest.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 This work is supported by the National Institutes of Health Grant CA77314 and by a Scholar Award from the Leukemia and Lymphoma Society (to J.D.). Technical support and research facilities were provided by the University of Colorado Health Sciences Center Cancer Center and supported by National Institutes of Health Grant 2 P30 CA46934.
2 Address correspondence and reprint requests to Dr. James DeGregori, University of Colorado Health Sciences Center, PO Box 6511, Mail Stop 8101, Aurora, CO 80045. E-mail address: james.degregori@uchsc.edu
3 Abbreviations used in this paper: HDP, homeostasis-driven proliferation; Tg, transgenic; 7-AAD, 7-aminoactinomycin D; DKO, double knockout.
Received for publication January 19, 2005. Accepted for publication April 21, 2005.
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Homeostasis-driven T cell proliferation occurs in response to a lymphopenic environment and is mediated by TCR and IL-7 signaling. In this report, we demonstrate a defect in the proliferation of murine naive and memory T cells lacking both E2F1 and E2F2 in response to lymphopenic conditions, suggesting that E2F1 and E2F2 function redundantly downstream of TCR and/or IL-7 signaling during homeostasis-driven proliferation. In contrast, T cell proliferation in response to antigenic stimulation is either unaffected (in vivo) or potentiated (ex vivo) by loss of E2F1 and E2F2, indicating divergent requirements for these E2F factors in T cell proliferation mediated by distinct stimuli. E2F1/E2F2 double knockout (DKO) T cells enter S phase in response to homeostatic signaling, but fail to divide, suggesting that S phase progression is either incomplete or defective. In addition, E2F1/E2F2 DKO mice do not recover normal T cell numbers following exposure to a sublethal dose of radiation, indicating that this defect in homeostasis-driven proliferation is physiologically relevant. Consistent with their failure in cell cycle progression, the differentiation of DKO T cells into memory T cells in response to homeostatic signals is significantly reduced. These observations support the idea that proliferation is required for memory T cell formation and also have implications for the development of clinical strategies to minimize the occurrence of lymphopenia-induced autoimmunity.
Introduction
T cell numbers are maintained at relatively constant levels in the peripheral immune system. Although environmental stimuli, including antigenic stimulation or exposure to lymphotoxic conditions, can result in a temporary increase or decrease, T cell numbers return to normal once the stimuli is removed. The total numbers of cells in both the naive and memory T cell compartments are determined by the relative contributions of lymphocyte generation, proliferation, differentiation, and death. Following exposure to an antigenic stimulus, T cell numbers are elevated. Once the Ag is cleared, many of the responding T cells undergo apoptosis or differentiate into memory T cells, thereby restoring naive T cell numbers to their normal level. In contrast, T cells undergo homeostasis-driven proliferation (HDP)3 in response to deficiencies in T cell numbers, or lymphopenia (reviewed in Ref. 1), and this response also results in the restoration of T cell numbers. HDP has both phenotypic and functional implications for the peripheral T cell compartment. Naive T cells that have undergone HDP express memory T cell surface markers and acquire increased effector function (reviewed in Ref. 2). Thus, both Ag-stimulated proliferation and HDP result in the formation of memory T cells.
Like Ag-stimulated proliferation, HDP of naive T cells requires TCR stimulation by MHC-peptide complexes (3, 4, 5, 6, 7). However, relatively weak TCR stimuli, often mediated by self-peptides, are sufficient for the induction of HDP and this feature may reflect decreased competition for access to APCs expressing the appropriate MHC-peptide complexes under lymphopenic conditions as well as the sensitization of T cells to TCR signaling. IL-7 is also required for HDP of naive T cells, and IL-7 levels are increased in lymphopenic hosts under a variety of conditions (reviewed in Ref. 8). In addition, exogenous IL-7 can stimulate T cell proliferation in vivo (9, 10, 11), consistent with the possibility that IL-7 functions to sensitize T cells to TCR stimuli and thereby promote HDP. These observations define TCR and cytokine signaling as minimal requirements for HDP of naive T cells. The requirements for HDP of memory T cells are both less stringent and less well defined (reviewed in Ref. 1). In the majority of cases, TCR stimulation by MHC-peptide complexes or MHC alone is not required for HDP of memory T cells. In addition, HDP of CD8 memory T cells can be supported by either IL-7 or IL-15, whereas no specific cytokine requirements for HDP of CD4 memory T cells have been defined.
We have been using mouse knockout models to study the roles of E2F1 and E2F2 in T cell proliferation. Members of the E2F family of transcription factors are key regulators of cell cycle progression. E2F1, E2F2, and E2F3 function to regulate expression of a relatively large set of genes that are induced at the G1 to S phase transition during the cell cycle. Many of these target genes encode proteins that are important for cell cycle progression, including cyclin A2, thymidylate synthase, DNA polymerase , and Cdc6 (12). Mutation of both E2F1 and E2F2 or E2F2 alone results in a number of hemopoietic phenotypes in mice. First, consistent with their known roles in the cell cycle, disruption of E2F1 and E2F2 results in defects in the development of a number of hemopoietic compartments due to impeded S phase progression in progenitors of the B cell, erythroid, and myeloid lineages as well as multipotent stem cells (13). These defects in the bone marrow are consistent with the known role for E2F members in promoting cell cycle progression. In contrast, T cells derived from mice mutant for both E2F1 and E2F2 or E2F2 alone not only enter S phase more rapidly than wild-type T cells following antigenic stimulation and proliferate much more extensively, but they also respond to lower levels of Ag (14, 15), indicating a role for E2F2 and, to a lesser extent, E2F1 in setting the TCR signaling threshold for Ag-stimulated proliferation. In addition, E2F2 mutant mice are predisposed to the development of autoimmunity (14). Taken together these observations reveal a somewhat paradoxical role for E2F1 and E2F2 in preventing inappropriate cell cycle progression. Given these results, we were interested in understanding how the loss of E2F1 and E2F2 might affect T cell proliferation in response to homeostatic stimuli. These studies have led to a further understanding of the events that occur downstream of TCR and cytokine signals during T cell HDP. In addition, the experiments we describe have important clinical and mechanistic implications with respect to memory T cell differentiation.
Materials and Methods
Mice
All experiments were performed using 4- to 8-wk-old E2F1/E2F2 double knockout (DKO) or control (E2f1+/+E2f2+/+, E2f1+/+E2f2+/–, E2f1+/–E2f2+/–, or E2f1+/–E2f2+/+) mice in either the B10.D2 (backcrossed for two to four generations) or BALB/c (backcrossed for three to five generations) backgrounds. No differences in HDP were observed between wild-type and E2F1 or E2F2 heterozygous T cells. DO11.10 TCR transgenic (DO TCR Tg) (16), E2F1 mutant (17), and E2F2 mutant (14) mice were obtained from Dr. P. Marrack (Howard Hughes Medical Institute, National Jewish Medical and Research Center, Denver, CO) or Dr. M. E. Greenberg (Children’s Hospital and Harvard Medical School, Boston, MA). Pure-bred recipient mice (B10.D2 or BALB/c) were obtained from The Jackson Laboratory. Mice were housed in the University of Colorado Health Sciences Center animal resource center and all animal procedures were performed according to Institutional Review Board approval protocols. Genotypes were determined by PCR analysis using genomic DNA isolated from tail biopsies. Where indicated mice were exposed to a single dose of gamma radiation from a cobalt 60 source at a distance of 25 cm. Complete blood counts were determined from a small sample of peripheral blood obtained from the tail and were measured using a Cell-Dyn 1700 hematology analyzer (Abbott Laboratories). For isolation of memory and naive T cells by single-cell sorting, DO TCR Tg, E2F1/E2F2 DKO and control mice were injected s.c. with 2.5 mg of chicken OVA protein (Sigma-Aldrich) in PBS plus 50 μl of CFA (Sigma-Aldrich) 32 days before sorting to induce the formation of DO TCR Tg memory T cells.
CFSE staining and lymphocyte transfers
Single-cell suspensions in PBS containing 1% FBS (FBS/PBS), 1 mM MgCl2, and 100 U/ml DNase I (Sigma-Aldrich) were obtained from lymph nodes and strained through nylon mesh. Where indicated, lymphocytes were stained in PBS containing 3 μM CFSE (Molecular Probes) for 15 min at 37°C. Both CFSE-labeled and unlabeled lymphocytes were washed in PBS for 30 min and were transferred to recipient mice by subocular or i.p. injection. Irradiated recipients were exposed to 450 rad of gamma radiation 1 day before transfer. Recipients received 2–8 x 106 lymphocytes/mouse. Transferred cells were recovered from the lymph nodes and spleens of recipient mice 14–21 days later or as indicated and analyzed.
Flow cytometry
Single-cell suspensions obtained from lymph nodes and/or spleen were strained through nylon mesh and washed once in PBS. In some cases, CD4+ T cells isolated from DO TCR Tg mice were purified by MACS using IMag CD4 MSC particles (BD Pharmingen) to facilitate efficient staining and analysis. Cells were stained in Ab solution (5 μg/ml fluorochrome-linked Abs, 1:50 2.4G2 tissue culture supernatant containing anti-FcR III/II Ab, and 1:100 goat serum (Invitrogen Life Technologies) in FBS/PBS) at a concentration of 108/ml and analyzed by flow cytometry. For single-cell sorting, RBC, macrophages, and B cells were depleted from combined lymph node and spleen suspensions by MACS using Ter119 microbeads (Miltenyi Biotec), CD11b microbeads (Miltenyi Biotec), and IMag anti-mouse CD45R/B220 particles (BD Pharmingen). The remaining cells were stained as described, and Thy1.2+ memory (CD44high, CD62Llow) and naive (CD44low, CD62Lhigh) T cells were purified by single-cell sorting. PE- and allophycocyanin-streptavidin conjugates and allophycocyanin anti-CD4, allophycocyanin anti-CD8, FITC anti-CD90.2 (Thy1.2), FITC anti-CD62L, PE anti-CD122, allophycocyanin anti-CD44, PE anti-CD127 (IL-7R), and PE anti-mouse IgG1 isotype control Abs were obtained from BD Pharmingen. Biotin- and fluorochrome-linked anti-DO TCR (KJ1.26) mAbs were obtained from Caltag Laboratories.
Stimulation of peripheral lymphocytes
For in vitro stimulation, single-cell suspensions obtained from lymph nodes of E2F1/E2F2 DKO and control mice or from the spleens of pure-bred mice were strained through nylon mesh, washed in PBS, resuspended at a final concentration of 1.0 x 107/ml in DMEM containing 10% FBS (HyClone Laboratories), 0.1 mM 2-ME, and 1% penicillin-streptomycin (Invitrogen Life Technologies), and plated in tissue culture dishes. Lymph node suspensions isolated from DO TCR Tg mice were combined 1:1 with nontransgenic splenocytes, which function as APCs, and stimulated with OVA protein (Sigma-Aldrich). Alternatively, lymph node suspensions were cultured alone and stimulated with 5 ng/ml PMA (Sigma-Aldrich) and 500 ng/ml ionomycin (Sigma-Aldrich). For in vivo stimulation, 107 CFSE-labeled lymphocytes were transferred to unirradiated recipients as described. Recipient mice received a single dose of OVA protein (Sigma-Aldrich) in PBS by i.p. injection 16–20 h after the lymphocyte transfer.
BrdU staining
Peripheral T cells were stimulated in vitro, as described, in the presence of 10 μM BrdU (Boehringer Mannheim). Alternatively, recipient mice were maintained on water containing 0.8 mg/ml BrdU and 10% sucrose, made fresh daily, to assess BrdU incorporation in vivo. In vitro cultures were harvested 48–52 h after stimulation. For in vivo experiments, single-cell suspensions were isolated from the lymph nodes and spleen of recipient mice at the indicated times and CD4+ T cells were isolated by MACS as described. Lymphocytes were exposed to UV-B for 10 min before staining with PE anti-DO TCR Ab (Caltag Laboratories), FITC anti-BrdU (DakoCytomation) Ab, and 7-aminoactinomycin D (7-AAD) as previously described (18). BrdU incorporation and DNA content in DO TCR Tg T cells was measured by flow cytometry.
Memory T cell surface marker and intracellular cytokine staining
Unlabeled DO TCR Tg lymphocytes were transferred to irradiated recipients. For memory and naive T cell controls, lymphocytes were isolated from a DO TCR Tg mouse and combined with B10.D2 lymphocytes at a ratio of 1:50. Aliquots of all samples were stained with Abs against memory cell surface markers and analyzed by flow cytometry or stimulated in vitro with PMA and ionomycin, as described. GolgiPlug (1 μl/ml; BD Pharmingen) was added to in vitro cultures to inhibit the export of cytokines. Cultures were harvested after 2 h, stained with PE anti-DO TCR and allophycocyanin anti-CD44 as described, fixed in 4% paraformaldehyde for 15 min at 4°C, and stored in PBS plus 1% BSA. The next day, fixed cells were incubated in 1x BD Pharmingen Perm/Wash buffer for 15 min at 4°C to permeabilize and stained with 5 μg/ml FITC anti-IL-2 (BD Pharmingen) plus 1:50 2.4G2 tissue culture supernatant containing anti-FcR III/II Ab in 1x BD Pharmingen Perm/Wash buffer at a concentration of 108/ml. Stained cells were washed two times for 20 min at 4°C in 1x BD Pharmingen Perm/Wash, resuspended in 1% FBS/PBS, and analyzed by flow cytometry.
Results
E2F1 and E2F2 are required for homeostasis-driven T cell proliferation
To determine whether E2F1 and E2F2 play a role in T cell HDP, we assessed the ability of T cells isolated from E2F1 mutant, E2F2 mutant E2F1/E2F2 DKO, or control mice to proliferate upon transfer to a lymphopenic host. Lymph nodes were harvested from DO TCR Tg E2F1/E2F2 DKO or control mice and stained with CFSE, a fluorescent dye that is taken up by live cells. CFSE-labeled cells were transferred to syngeneic wild-type recipients that had been rendered lymphopenic by exposure to a sublethal dose of gamma radiation. T cells were harvested from the lymph nodes and spleen of recipient mice 2–3 wk later, stained with fluorescent-linked Ab against the DO TCR (or other T cell markers, depending on the nature of the specific experiment), and the CFSE profiles of DO TCR+, CFSE+ donor-derived T cells were assessed by flow cytometry. With each round of division, the daughter cells inherit one-half of the CFSE of their parent and each generation of cells therefore has a characteristic fluorescence, which serves as an indicator of proliferation.
As shown in Fig. 1, E2F1/E2F2 DKO T cells exhibit defects in HDP. T cells isolated from control mice undergo significant proliferation in irradiated hosts (Fig. 1A). Cells that have undergone between 0 and 3 rounds of division are evident. In contrast, E2F1/E2F2 DKO T cells do not proliferate in irradiated recipient mice, indicating that E2F1, E2F2, or both together are required for this process. Consistent with this observation, the number of donor-derived T cells recovered from the lymph nodes and spleens of irradiated recipients of E2F1/E2F2 DKO T cells was 60% of the number recovered from recipients of control T cells, irrespective of whether the donor T cells were labeled with CFSE (Fig. 1B). Additional experiments revealed that T cells isolated from E2f1–/–E2f2+/– or E2f1+/–E2f2–/– mice do not exhibit defects in proliferation in irradiated hosts (Fig. 1C), suggesting that E2F1 and E2F2 function redundantly to promote proliferation under these conditions. As expected, when T cells isolated from mice of any of these genotypes are transferred into unirradiated recipients, they do not divide (Fig. 1A and data not shown), indicating that the observed proliferation requires lymphopenia and is therefore occurring in response to homeostatic signals. Thus, E2F1 and E2F2 function redundantly to promote T cell HDP in irradiated hosts. In addition, the lymphocyte compartment in DKO mice did not recover following exposure to a sublethal dose of gamma radiation, whereas lymphocyte numbers returned to near normal levels in wild-type mice (Fig. 1D and data not shown). At the last time point shown, significant reconstitution of the thymus in terms of cell number had not occurred in mice of either genotype and mature single positive thymocytes were not detected (data not shown), suggesting that reconstitution of the peripheral T cell compartment was predominantly a result of HDP, rather than thymopoiesis. Taken together, these observations indicate that the defect in HDP exhibited by E2F1/E2F2 DKO T cells results in a physiologically relevant deficit in recovery of the T cell compartment in response to lymphopenia.
Although E2F1/E2F2 DKO T cells exhibit significant defects in T cell HDP, this phenotype is not completely penetrant and in some cases, there is a small but distinct population of E2F1/E2F2 DKO T cells that do appear to proliferate in irradiated hosts (Figs. 1, A and B, and 2A). This observation raises the possibility that different T cell subsets may have different requirements for E2F1 and E2F2 for HDP. To investigate this possibility, we assessed the induction of HDP in several T cell subsets. The defect in HDP exhibited by E2F1/E2F2 DKO T cells is not restricted to a single T cell compartment, as both CD4+ and CD8+ T cells isolated from nontransgenic DKO mice exhibit this phenotype (Fig. 2A). In addition, both naive (CD44low, CD62Lhigh) and memory (CD44high, CD62Llow) T cells isolated from OVA-injected DO TCR Tg E2F1/E2F2 DKO mice exhibit defects in HDP (Fig. 2B). This observation also suggests that the defects in HDP that occur as a result of mutation of E2F1 and E2F2 are not due to abrogation of IL-7 signaling, as IL-7 is not required for HDP of CD4+ memory T cells (19). Consistent with this idea, DO TCR Tg T cells isolated from DKO and control mice exhibited similar cell surface expression of the IL-7R (Fig. 3A). Similarly, the defects in HDP exhibited by E2F1/E2F2 DKO T cells do not appear to occur as a result of differences in survival in irradiated hosts. Apparent precursor frequencies can be calculated for both DKO and control samples based on CFSE staining profiles and the absolute number of T cells recovered from recipient mice. For instance, in one experiment, 36.3% of the donor-derived control T cells recovered from irradiated recipients had not divided, 31.8% had divided once, 15.1% had divided twice, 10.3% had divided three times, and 6.6% had divided four times. For each generation, the minimum number of donor T cells that the recovered cells were derived from can be calculated by multiplying the total number of donor-derived T cells recovered by the percentage of donor-derived T cells in that generation and then dividing by 2x, where x is the number of rounds of proliferation that cells in that generation have gone through. In this example, 1.44 x 106 donor-derived T cells were recovered from the lymph nodes and spleen of an irradiated recipient of control cells and these cells were therefore derived from a total of 1.44 x 106 (0.363/20 + 0.318/21 + 0.151/22 + 0.103/23 + 0.066/24), or 8.31 x 105 cells. Similarly, 8.62 x 105 E2F1/E2F2 DKO T cells were recovered from an irradiated recipient and, because DKO T cells do not undergo proliferation in lymphopenic hosts, the apparent precursor frequency in this case is 8.62 x 105. Although this calculation does not provide information regarding the fraction of transferred cells that survive in irradiated recipients, it does indicate relative survival of E2F1/E2F2 DKO and control T cells in irradiated recipients. Using this method, apparent precursor frequencies were calculated and no significant difference between the precursor frequencies for DKO or control T cells were observed (Fig. 3B). Because similar numbers of T cells were transferred to recipients of both DKO and control T cells (the average ratio of donor-derived T cells recovered from E2F1/E2F2 DKO versus control recipients 1 day after transfer was 1.08 ± 0.06), apparent precursor frequency is a direct indicator of relative survival in these experiments. Thus, the observed differences in HDP of E2F1/E2F2 DKO and control T cells do not reflect a difference in their capacity for survival in irradiated hosts.
E2F1/E2F2 DKO T cells enter S phase in response to homeostatic signals
The phenotypes that have been observed in the bone marrow and the peripheral immune system suggest two distinct possibilities in terms of the underlying cause for the failure of E2F1/E2F2 DKO T cells to progress through the cell cycle in response to homeostatic signals. First, in the absence of E2F1 and E2F2, homeostatic signals may not stimulate sufficient accumulation of E2F targets to promote progression into S phase. In this case, E2F1/E2F2 DKO T cells would remain in G0 and G1 phases of the cell cycle following transfer into a lymphopenic host. Alternatively, E2F1/E2F2 DKO T cells may enter S phase but be impeded in S phase progression, similar to the phenotype exhibited by hemopoietic progenitors in the bone marrow (13). To address this issue, T cells were isolated from DO TCR Tg E2F1/E2F2 DKO or control mice and transferred to irradiated recipients. The recipients were maintained on BrdU-containing water for varying intervals of time after the transfers and BrdU incorporation and DNA content in donor-derived T cells were determined by flow cytometry. Parallel samples were stained with CFSE before transfer into irradiated recipients to monitor proliferation. Both E2F1/E2F2 DKO and control T cells exhibited significant incorporation of BrdU between 0 and 6 days after transfer into irradiated recipients indicating that DKO T cells do enter S phase (Fig. 5, A and B), despite the fact that they do not proliferate (Fig. 5D). Moreover, BrdU-positive cells with G1, S, and G2 DNA content were observed in both samples, indicating that E2F1/E2F2 DKO T cells not only initiate S phase, but undergo significant DNA synthesis in a lymphopenic environment (Fig. 5C), despite the fact that they do not divide (Fig. 5D). At later time points, an increasing percentage of donor-derived control cells exhibited BrdU incorporation. In contrast, the percentage of E2F1/E2F2 DKO T cells that incorporate BrdU decreases slightly between 6 and 15 days posttransfer. As shown in Fig. 3C, it is unlikely that E2F1/E2F2 DKO T cell survival in lymphopenic hosts is affected by loss of E2F1 and E2F2, and this decrease in the incidence of cells entering S phase is therefore consistent with the idea that E2F1/E2F2 DKO T cells are undergoing a cell cycle arrest, presumably in S phase. These data suggest that the defect in HDP due to mutation of E2F1 and E2F2 occurs by a similar mechanism to the failure in cell cycle progression that is manifested in hemopoietic precursors in the bone marrow.
E2F1 and E2F2 are required for homeostasis-driven memory T cell differentiation
Naive T cells that have undergone HDP develop a memory T cell phenotype. To determine whether E2F1/E2F2 DKO T cells acquire a memory phenotype following transfer to lymphopenic hosts, we examined the expression of memory T cell surface markers on E2F1/E2F2 DKO and control T cells after recovery from irradiated recipient mice. For control samples, DO TCR Tg lymphocytes were combined with nontransgenic lymphocytes at a ratio of 1:50, so that the density of both naive and memory phenotype DO TCR Tg T cells was similar in control samples and those derived from irradiated recipients. The comparison of populations with similar densities of Tg lymphocytes ensures that differences in gating on the DO TCR Tg fraction of the population do not contribute to observed differences in cell surface staining. Before transfer into recipient mice, E2F1/E2F2 DKO and control T cell populations exhibited staining patterns similar to naive T cell samples (data not shown). After recovery from lymphopenic recipients, control T cells exhibited a statistically significant increase in the proportion of CD44high memory T cells relative to the untransferred control sample (Fig. 6A). A substantial fraction of these CD44high cells exhibited altered expression of additional memory T cell markers relative to naive T cell samples, including IL-2R (CD122) and/or CD62L (Fig. 6B). Up-regulation of CD122 and down-regulation of CD62L were similar on endogenous memory T cells and memory T cells that had differentiated in response to homeostatic stimuli. In addition, CD44high memory T cells isolated from lymphopenic recipients of control T cells exhibited increased effector function as indicated by the rapid production of IL-2 in response to stimulation with PMA and ionomycin in vitro (Fig. 6B). Thus, control T cells that have undergone HDP exhibited an increase in the proportion of T cells expressing memory phenotypes, relative to populations derived from naive mice. In contrast, the expression of memory markers was not significantly different on E2F1/E2F2 DKO T cells rescued from lymphopenic hosts as compared with the untransferred control population (Fig. 6A). A small fraction of these DKO T cells do express memory markers and exhibit increased effector function (Fig. 6). However, it is not clear whether E2F1/E2F2 DKO T cells expressing a memory phenotype developed from naive T cells as a result of homeostatic signaling or were present in the original transferred population. In either case, memory T cell differentiation is not efficiently induced in response to irradiation-induced lymphopenia in the absence of E2F1 and E2F2.
Discussion
We have shown that the E2F1 and E2F2 transcription factors function redundantly to mediate proliferation of primary T cells in response to lymphopenic conditions in vivo, suggesting that E2F1 and E2F2 function downstream of TCR and/or IL-7R signaling during HDP. HDP is defective in multiple T cells subsets isolated from E2F1/E2F2 DKO mice, including CD4+ memory T cells and both CD4+ and CD8+ naive T cells. E2F1/E2F2 DKO T cells enter S phase in response to homeostatic signaling, but fail to divide, suggesting that S phase progression is either incomplete or defective. This phenotype is similar to the E2F1/E2F2-dependent defects in cell cycle progression exhibited by hemopoietic progenitors and may be due to a failure in the accumulation of E2F target genes required for S phase. Taken together, these observations indicate a role for E2F1 and E2F2 in promoting T cell proliferation in response to TCR stimuli in the context of homeostatic signaling in vivo. In contrast, E2F1/E2F2 DKO T cells proliferate in response to antigenic stimulation in vivo and hyperproliferate in response to suboptimal antigenic stimulation in vitro. Thus, E2F1 and E2F2 are differentially required for T cell proliferation in response to distinct TCR-mediated stimuli. Previous studies have shown that T cells undergoing proliferation in response to homeostatic signaling or antigenic stimulation exhibit similar gene expression profiles, suggesting that quantitative differences in TCR and/or cytokine signaling, rather than qualitative differences, distinguish HDP from Ag-mediated proliferation (20). In fact, only one gene was specifically regulated in response to homeostatic stimulation. Thus, the experiments we have described are particularly important in that they identify distinct molecular requirements for T cell proliferation in response to antigenic and homeostatic signaling and suggest a molecular mechanism by which quantitatively different signals can mediate similar effects on T cell proliferation. In addition, these observations reveal distinct aspects of E2F function that either promote or prevent cell cycle progression in response to TCR activation in peripheral T cells, depending on the nature of the stimulus. We are interested in understanding how the T cell activation signals defined in this study impinge on the cellular requirements for E2F function. One possibility is that the activity and/or abundance of E2F family members can be differentially regulated in a single-cell type in response to distinct stimuli or assay conditions. For instance, following TCR stimulation by Ag, E2F3 may be activated to perform functions that are dependent on E2F1 and E2F2 in the context of the weaker TCR signals that mediate HDP. Alternatively, it is possible that E2F-dependent target genes are similarly regulated in response to distinct stimuli, but regulation of these target genes may be differentially required in specific contexts.
What implications do these observations have for the T cell repertoire in E2F1/E2F2 DKO mice? Recent studies suggest that under normal circumstances, HDP is important during early development of the immune system for expansion of the relatively limited neonatal peripheral T cell population (21, 22, 23) and HDP may also function in older individuals to compensate for decreased T cell generation as a result of thymic involution. Thus, the defect in HDP exhibited by E2F1/E2F2 DKO T cells may contribute to the decrease in mature T cell numbers observed in DKO mice (13). This phenotype is expected to worsen with age as thymic output decreases and E2F1/E2F2 DKO T cells cannot compensate due to defects in HDP. Recent studies have also implicated lymphopenia and HDP in the development of autoimmunity (24, 25 and reviewed in Ref. 26). Older E2F1/E2F2 DKO and E2F2-deficient mice exhibit both lymphopenia and autoimmune features (13, 14, 15). Because E2F1/E2F2 DKO T cells exhibit defects in HDP, whereas E2F2-deficient T cells do not, a direct comparison of the timing and extent of the development of autoimmunity in these two genetic backgrounds should reveal the relative contributions of lymphopenia and HDP to this process.
These studies also have implications for the mechanism by which memory T cell differentiation occurs. Both homeostasis-driven and Ag-stimulated proliferation require TCR signaling and it has been previously shown that memory T cell differentiation during HDP is temporally correlated with proliferation such that only cells that have undergone multiple rounds of division express memory markers (27, 28, 29, 30). Based on these observations, it has been proposed that proliferation and TCR signaling are minimal requirements for the development of memory T cells. However, these experiments do not eliminate the possibility that it is the duration or quality of the TCR signal, rather than the extent of proliferation, which determines memory T cell differentiation. Mutation of E2F1 and E2F2 specifically abrogates T cell proliferation in response to homeostatic signals and has no apparent affect on TCR signaling, as indicated by the observation that E2F1/E2F2 DKO and control T cells enter S phase with similar kinetics and at a similar frequency in response to homeostatic signals. Thus, mutation of E2F1 and E2F2 allows for the separation of these two potentially important aspects of HDP-induced memory T cell differentiation, and DKO T cells thereby provide a unique opportunity to assess the effects of changes in proliferation in the absence of changes in TCR signaling. Despite the fact that E2F1/E2F2 DKO T cells receive a TCR signal that is sufficient to induce progression into S phase, they do not efficiently express memory phenotypes. Thus, the experiments presented in this study suggest that homeostatic signaling is not sufficient for the induction of memory T cell differentiation and are consistent with the idea that proliferation is required for efficient memory T cell differentiation.
A final consideration is the clinical relevance of these studies. HDP is important for repopulation of the peripheral immune system following the induction of lymphopenia either therapeutically, due to treatment with radiation or chemotherapies, or as a result of viral infection, particularly by HIV (31). In addition, HDP may contribute to the development of autoimmunity in several ways. Homeostatic TCR stimuli can be delivered by the same self Ags that mediate positive selection during T cell development, thus stimulating the clonal expansion of self-reactive T cells (5, 6, 7). HDP also induces memory T cell differentiation, thereby ensuring a persistent immune response. HDP-induced autoimmunity is thought to play an important role in organ transplant rejection, particularly if peripheral T cell depletion has been used as a method of immunosuppression (32). The studies described in this report suggest that therapeutic strategies using cell cycle inhibitors to suppress the development of graft-vs-host disease mediate protection by preventing both clonal expansion and memory T cell differentiation, thereby limiting the development of HDP-induced autoimmunity. Moreover, the development of strategies that result in specific inhibition of E2F1 and E2F2 may be useful in limiting autoimmunity with little or no effect on T cell activity toward bona fide Ags or proliferation of other cell types. This approach would be most useful in cases in which widespread immune suppression is not desirable, such as for the prevention of autoimmunity as a result of therapy- or virus-induced lymphopenia or due to a genetic predisposition. Further characterization of the molecular mechanism by which E2F1 and E2F2 mediate their role in HDP may reveal downstream effectors that can be more easily targeted for clinical intervention using currently available reagents.
Acknowledgments
We thank Jing Zhu and Feng Li for the initial observations that inspired this project, Dave Hildeman, Tom Mitchell, and Kent Teague for critical review of this manuscript, and the University of Colorado Health Sciences Center Cancer Center Flow Cytometry Core for technical assistance.
Disclosures
The authors have no financial conflict of interest.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 This work is supported by the National Institutes of Health Grant CA77314 and by a Scholar Award from the Leukemia and Lymphoma Society (to J.D.). Technical support and research facilities were provided by the University of Colorado Health Sciences Center Cancer Center and supported by National Institutes of Health Grant 2 P30 CA46934.
2 Address correspondence and reprint requests to Dr. James DeGregori, University of Colorado Health Sciences Center, PO Box 6511, Mail Stop 8101, Aurora, CO 80045. E-mail address: james.degregori@uchsc.edu
3 Abbreviations used in this paper: HDP, homeostasis-driven proliferation; Tg, transgenic; 7-AAD, 7-aminoactinomycin D; DKO, double knockout.
Received for publication January 19, 2005. Accepted for publication April 21, 2005.
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