Glucocorticoid-induced TNF receptor family-related protein (GITR) is expressed on activated and regulatory T cells, but its role on these functionally opposing cell types is not fully understood. Here we describe that transgenic expression of GITR’s unique ligand (GITRL) induces a prominent increase of both effector and regulatory CD4+ T cells, but not CD8+ T cells. Regulatory T cells from GITRL transgenic mice are phenotypically activated and retain their suppressive capacity. The accumulation of effector and regulatory T cells is not due to enhanced differentiation of naive T cells, but is a direct result of increased proliferation. Functional consequences of increased numbers of both regulatory and effector T cells were tested in an autoimmune model and show that GITR stimulation is protective, as it significantly delays disease induction. These data indicate that GITR regulates the balance between regulatory and effector CD4+ T cells by enhancing proliferation of both populations in parallel.

Members of the TNF receptor superfamily are able to directly and indirectly affect the course of an immune response (1). The glucocorticoid-induced TNF receptor family-related protein (GITR)4 is a member of this family and has been implicated in regulating both innate and adaptive immune responses (2, 3). Regulatory T cells are well known for their expression of GITR, although this receptor is also expressed on activated nonregulatory T cells, as well as B cells, monocytes and macrophages, dendritic cells, and mast cells (3, 4, 5, 6, 7). Its ligand (GITRL) can be found on a variety of cells, including dendritic cells, macrophages, and B cells (7, 8, 9). GITRL is transiently up-regulated on these APCs upon stimulation via the transcription factor NF-1 (8), and it most likely exerts its main function during inflammatory responses. Indeed, agonistic Abs and (cells expressing) recombinant GITRL enhance T cell proliferation in vitro upon TCR triggering (2, 5, 6, 8, 10), which suggests that GITR acts as a costimulatory factor for T cells. GITR stimulation on regulatory T cells in coculture with effector T cells has been suggested to neutralize the suppressive capacity of regulatory T cells (5, 6, 8). However, it was later shown that GITR ligation on regulatory T cells does not directly affect their suppressive capacity, but that GITR stimulation on nonregulatory T cells allows them to escape suppression by regulatory T cells (11). GITR is not essential for regulatory T cell function, as regulatory T cells from GITR−/− mice display a normal capacity to suppress T cell proliferation in vitro (2, 12). This leaves unanswered what the function of GITR is on regulatory T cells.

Studies using GITR−/− mice showed that the absence of GITR was protective in several disease models, which was attributed to an impaired effector function of T cells (13, 14, 15). Correspondingly, studies that have directly addressed the function of GITR on T cells in vivo by deliberate stimulation of the receptor with agonistic Abs concluded that GITR has a proinflammatory role within the immune system through its costimulatory effects on T cells (6, 7, 16, 17). However, these Abs have their limitations when studying the impact of GITR stimulation in complex disease models, in particular since anti-GITR Abs have been reported to cause depletion of regulatory T cells (7). Moreover, recent studies on the crystal structure of GITRL have revealed that this ligand can exist in multiple oligomerization states that depend on binding to the receptor (18, 19), and it therefore remains to be addressed whether cross-linking GITR with agonistic Abs exerts the same downstream effects as signaling induced by membrane-bound GITRL. Thus, to properly address the consequence of direct GITR stimulation on T cell function in vivo, we generated transgenic (TG) mice in which GITRL is constitutively expressed on B cells. Our findings demonstrate that GITR stimulation in vivo very effectively increases the absolute number of both effector and regulatory CD4+ T cells through enhanced proliferation of both cell types. In agreement with increased regulatory T cell numbers, GITRL TG mice showed a marked delay in disease onset upon induction of experimental autoimmune encephalomyelitis (EAE), an experimental model for multiple sclerosis. We propose that GITR plays an important role in the regulation of both regulatory and effector CD4+ T cell numbers in vivo by enhancing their turnover.

cDNA encoding murine GITRL was obtained via PCR on total splenic cDNA and cloned into the pGEM-T plasmid (Promega). This construct was digested with NotI and XhoI to obtain a 600-bp fragment containing the mGITRL cDNA, which was cloned into the NotI-XhoI site of a CD19-pC3 plasmid (kindly provided by Patrick Derksen, Academic Medical Center, The Netherlands), resulting in the GITRL expression construct under control of the human CD19 promoter. This construct was linearized via AatII digestion (see Fig. 1 A) and microinjected into pronuclei of C57BL/6 fertilized oocytes and implanted into pseudopregnant female C57BL/6 mice. Transgenic founders were identified by PCR analysis of tail or ear DNA, using the following PCR primers: pC3s1 (5′-GCAGTGACTCTCTTAAGGTAGCC-3′) and mGITRL4a (5′-CTTGAGTGAAGTATAGATCAGTGTA-3′). Three GITRL TG founder lines (RW14, RW18, RW20) were propagated by mating with wild-type (WT) C57BL/6 mice, and offspring were tested for the presence of the transgene by PCR analysis of tail or ear DNA with the same primers.

FIGURE 1.

Generation of B cell-specific GITRL TG mice. A, Schematic representation of the hCD19-mGITRL DNA construct. The human CD19 promoter (hatched bar) is followed by a chimeric intron (white bar), mGITRL cDNA (dotted bar), and a poly(A) tail (black bar). B, PCR analysis of genomic tail DNA from WT or GITRL TG mice (founder lines RW14, RW18, and RW20). C, Representative staining for GITRL on splenic B220+ cells from WT, RW14, RW18, and RW20 mice and (D) the expression of GITRL on splenic B220+ B cells as the average geometric mean fluorescence intensity (GeoMFI) ± SD for three mice per group. E, Representative staining for GITR expression on splenic CD3+ cells from WT, RW14, RW18, and RW20 mice and (F) the expression of GITR on CD3+ T cells as the average geoMFI ± SD. **, p < 0.005.

FIGURE 1.

Generation of B cell-specific GITRL TG mice. A, Schematic representation of the hCD19-mGITRL DNA construct. The human CD19 promoter (hatched bar) is followed by a chimeric intron (white bar), mGITRL cDNA (dotted bar), and a poly(A) tail (black bar). B, PCR analysis of genomic tail DNA from WT or GITRL TG mice (founder lines RW14, RW18, and RW20). C, Representative staining for GITRL on splenic B220+ cells from WT, RW14, RW18, and RW20 mice and (D) the expression of GITRL on splenic B220+ B cells as the average geometric mean fluorescence intensity (GeoMFI) ± SD for three mice per group. E, Representative staining for GITR expression on splenic CD3+ cells from WT, RW14, RW18, and RW20 mice and (F) the expression of GITR on CD3+ T cells as the average geoMFI ± SD. **, p < 0.005.

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GITRL TG mice were maintained on a C57BL/6 background and bred in the animal department of the Academic Medical Center (Amsterdam, The Netherlands) under specific pathogen-free conditions. Mice were used at 6–24 wk of age, age- and sex-matched within experiments, and were handled in accordance with institutional and national guidelines. All experiments have been reviewed and approved by the Academic Medical Center Animal Ethics Committee. For measurement of in vivo T cell proliferation, mice were injected i.p. with 1 mg of BrdU (Sigma-Aldrich) and sacrificed for analysis 16 h later.

Single-cell suspensions were obtained by mincing the specified organs through 40-μm cell strainers (BD Biosciences). Erythrocytes were lysed with an ammonium chloride solution and cells were subsequently counted using an automated cell counter (Casy; Schärfe System). Cells (5 × 105 to 5 × 106) were collected in staining buffer (PBS with 0.5% BSA; Sigma-Aldrich) and stained for 30 min at 4°C with Abs in the presence of anti-CD16-CD32 (clone 2.4G2). The following fluorescently or biotin-labeled mAbs (and clone names) were obtained from BD Pharmingen: anti-B220 (RA3-6B2), anti-CD3ε (145-2C11), anti-CD4 (L3T4), anti-CD8 (Ly-2), anti-CD62L (clone MEL-14), and anti-CD69 (H1.2F3); or from eBioscience: anti-GITRL (ebioYGL386), anti-FoxP3 (NRRF-30), anti-GITR (DTA-1), anti-CD44 (IM7), anti-CTLA4 (4C10-4B9), anti-PD1 (RMP1-30), anti-CD134 (OX86), anti-CD27 (LG.7F9), anti-CD103 (M290), and anti-CD45.1 (104 or A20). PE-conjugated anti-CD25 was obtained from Miltenyi Biotec. For the detection of biotinylated Abs, streptavidin-PE (Caltag Laboratories), streptavidin-allophycocyanin (BD Pharmingen), or streptavidin-conjugated PerCP-Cy5.5 (BD Pharmingen) was used. Intracellular stainings for FoxP3 and/or BrdU were performed using Foxp3 fixation/permeabilization concentrate and diluent (eBioscience), according to the manufacturer’s protocol. For BrdU staining, FITC-conjugated anti-BrdU/DNase (BD Pharmingen) was added during the FoxP3 staining step and stained for 25 min at room temperature. Data were collected on a FACSCalibur or FACSCanto (BD Biosciences) and were analyzed with using FlowJo software (Tree Star).

To determine direct ex vivo cytokine production, splenocytes were plated at 1 × 106 cells/well in a 96-well round-bottom plate and stimulated with 1 ng/ml PMA and 1 μM ionomycin. After 2 h of incubation at 37°C, 1 μg/ml of the protein-secretion inhibitor brefeldin A was added (Sigma-Aldrich) and cells were cultured for another 4 h. Afterward, cells were washed and stained for CD3 and CD4 or CD8, followed by fixation and permeabilization (BD Biosciences). Cells were then incubated for 30 min with fluorescently labeled Abs against IFN-γ, IL-2, IL-10, or IL-17 (eBioscience), thoroughly washed, and analyzed by flow cytometry.

To analyze the effect of GITR engagement on the proliferative capacity of WT responder cells, T cells were enriched from spleens of WT mice by negative selection using CD19+ beads (Miltenyi Biotec). T cell-enriched splenocytes were labeled with 0.25 μM CFSE in PBS at 37°C for 10 min and stimulated with 100 ng/ml anti-CD3 (clone 145-2C11) for 3 days in the presence of irradiated (10 Gy) WT or GITRL TG B cells with or without 200 U/ml IL-2. B cells were isolated by positive selection using CD19+ beads (Miltenyi Biotec). For the analysis of the expression of CD25 and CD69, non-CFSE-labeled enriched WT T cells were used, stimulated similarly, and analyzed after 1 day. To determine the effects of GITR ligation on IL-2 production in vitro, T cell-enriched splenocytes were stimulated as described above, in the presence of 10 μg/ml blocking anti-CD25 Ab (clone PC61) to prevent IL-2 consumption. Culture supernatant was harvested after 1 day of stimulation and frozen at −20°C. The IL-2 ELISA (BD Biosciences) was performed according to instructions from the manufacturer.

Splenic CD4+CD25 (responder cells) and CD4+CD25high (regulatory cells) T cells were isolated by cell sorting using a FACSAria (BD Biosciences) and purity of sorted populations was consistently >96%. Responder cells were mixed with regulatory T cells at different ratios in 96-well tissue culture plates. The cells were stimulated with 10 μg/ml soluble anti-CD3 (clone 145-2C11) plus irradiated (10 Gy) WT splenocytes (APCs) at 37°C for 72 h. Hereafter, cells were pulsed for 16 h with 1 μCi of 3H-TdR ([methyl-3H]thymidine; Amersham Pharmacia)/well, and incorporation of 3H-TdR was determined using a beta plate scintillation counter (Wallac; 1450 Microbeta Plus liquid scintillation counter). Data are presented as percentage proliferation compared with maximum responder cell proliferation of triplicate assays.

For adoptive transfers, naive (CD25) CD4+ T cells were purified from spleens and peripheral lymph nodes of Ly5.1 mice by negative selection using the CD4+CD25+ regulatory T cell isolation kit (Miltenyi Biotec). Purified CD4+CD25 cells (purity >90%) were labeled with 0.25 μM CFSE in PBS at 37°C for 10 min and injected after washing (with or without 1 × 106 in 200 μl of PBS) i.v. into WT and GITRL TG recipient mice. Distribution and phenotype of transferred cells was analyzed 3 days later by flow cytometry.

EAE was induced by s.c. immunization of mice in the hind flanks using 50 μg of MOG35–55 peptide in CFA containing 1 mg/ml heat-inactivated Mycobacterium tuberculosis (Difco) on day 0. Mice also received 200 ng of pertussis toxin (Sigma-Aldrich) i.v. on days 0 and 2. Disease severity was assessed according to the following scale: 0, no disease; 1, flaccid tail; 2, loss of hind leg spreading reflex; 3, hind limb weakness; 4, unilateral hind limb paralysis; 5, bilateral hind limb paralysis; 6, abdominal paralysis; 7, moribund; 8, dead. All mice were sacrificed 14 days following EAE induction, after which brain and spinal cord was frozen in Tissue-Tek (Sakura Finetek) at −80°C for immunohistochemical analysis.

Cryostat sections (8 μm) of spinal cord and brain of 4 WT and 4 TG mice were fixed in acetone, containing 1% H2O2 for 10 min. Then, sections were incubated with monoclonal rat anti-mouse Abs to CD68 (a kind gift from Siamon Gordon, Oxford, U.K., clone FE-11), CD4, and FoxP3 (eBioscience), diluted in PBS with 8% BSA, 10% normal mouse serum, and 0.05% NaN3 for 1 h at 4°C. After washing in PBS, the sections were incubated with anti-rat HRP diluted in PBS, 8% BSA, 10% normal mouse serum, 0.05% NaN3, and 350 mM NaCl. Staining was visualized with diaminobenzidine (Sigma-Aldrich) applied for 10 min. Sections were counterstained with heamatoxylin for 30 s, dehydrated, and mounted in Entallan (Merck). As negative controls, primary Abs were either left out or substituted with an isotype control Ab. No immunoreactivity was seen for all negative controls.

For the visualization of cellular infiltrates in organs obtained from WT and GITRL TG mice, cryostat sections (7 μm) of the thyroid, kidney, liver, stomach, and small and large intestine were stained by Diff-Quick (Dade Behring) according to the manufacturer’s instructions and analyzed by light microscopy.

Statistical analysis of the data was performed using the unpaired Student’s t test or Wilcoxon rank-sum test where mentioned. For the EAE experiments, the effect on mean clinical score was assessed by calculating the area under the curve using the trapezoidal rule, followed by the Wilcoxon rank-sum test. Differences in cumulative incidence were analyzed on a per day basis using a χ2 test of a contingency table.

To study the function of GITR on T cells in vivo, we generated B cell-specific GITRL TG mice by expressing GITRL cDNA under control of the human CD19 promoter (Fig. 1,A). Through microinjection of fertilized oocytes, we acquired three founder lines (RW14, RW18, and RW20), which were identified by genomic PCR analysis (Fig. 1,B). GITRL TG mice were fertile, born at expected Mendelian frequencies and appeared as healthy as their littermate controls. Flow cytometry showed that GITRL was indeed significantly expressed on B cells in all founder lines, with highest expression on the RW18 line (Fig. 1, C and D). As receptor shedding upon ligand stimulation is a hallmark of various TNF receptor superfamily members (20, 21, 22, 23, 24), we determined the expression of GITR on T cells. T cells from GITRL TG mice showed decreased GITR expression compared with WT mice, which correlated with the level of GITRL expression (Fig. 1, E and F). This indicates that GITR is indeed functionally engaged by its ligand in these mice. The data shown below are obtained from experiments with the GITRL TG RW18 line, although a similar, but less pronounced phenotype was also found in the other founder lines (data not shown).

To establish the effects of GITR triggering in vivo, we analyzed the primary and secondary lymphoid organs of GITRL TG mice. T cell differentiation in the thymus of these mice was comparable to WT littermates (data not shown), as was cellularity of bone marrow, thymus, and peripheral and mesenteric lymph nodes (Fig. 2,A). However, spleens of GITRL TG mice contained significantly more leukocytes than WT mice (Fig. 2,A). This increase was due to elevated numbers of CD4+ T cells (Fig. 2, B and C), whereas numbers of B cells and CD8+ T cells were not significantly altered (Fig. 2, B and C). Analysis of FoxP3 expression indicated that a substantial part of this increase in CD4+ T cells could be attributed to an enlarged regulatory T cell compartment, as up to three times more CD4+FoxP3+ regulatory T cells were present in the spleens of GITRL TG mice (Fig. 2, D and E). Phenotypic analysis of FoxP3CD4+ T cells indicated that the nonregulatory fraction was also affected in GITRL TG mice, as significantly more CD4+ T cells with an effector memory-like (CD44+CD62L) and central memory-like (CD44+CD62L+) phenotype were identified (Fig. 2, D and E). This increase of CD4+ T cells with either a regulatory or a memory-like phenotype in GITRL TG mice apparently did not develop at the cost of the naive CD4+ population, since absolute numbers of naive CD4+ T cells were comparable with WT littermates (Fig. 2,E). No differences were found for CD8+ T cells with respect to their naive, effector memory-like and central memory phenotype (Fig. 2, F and G). Corroborating the specific increase in CD4+ effector T cells in GITRL TG mice, splenocyte stimulation with PMA-ionomycin showed increased production of the effector cytokines IL-2 (Fig. 2,H) and IFN-γ (Fig. 2, I and J) by CD4+, but not CD8+ T cells. Consistent with the increase in regulatory T cell numbers, we observed a trend toward more IL-10-producing CD4 T cells, but this difference was not significant (Fig. 2, K and L). These data thus indicate that GITR triggering in vivo enhanced the number of both regulatory and effector CD4+ T cells.

FIGURE 2.

GITRL TG mice have more effector and regulatory type CD4+ T cells. A, Absolute number of cells in spleen, bone marrow (BM), thymus, peripheral (pLN), and mesenteric (mLN) lymph nodes in 4- to 8-wk-old WT (open bar) and GITRL TG (filled bar) mice. Percentage and absolute number of (B) T and B cells or (C) CD4+ and CD8+ T cells in the spleen of WT and GITRL TG mice. D and E, Percentage and absolute number of splenic regulatory (FoxP3+) and nonregulatory (FoxP3) CD4+ T cells in WT and GITRL TG mice. Nonregulatory CD4+ T cells were subdivided in naive (CD44CD62L+), effector memory (EM) (CD44+CD62L) and central memory (CM) (CD44+CD62L+) cells. F and G, Percentage and absolute number of naive, EM, and CM cells of splenic CD8+ T cells. Production of (H) IL-2, (I and J) IFN-γ, and (K and L) IL-10 by CD4+ or CD8+ T cells in WT and GITRL TG mice after stimulation with PMA-ionomycin. *, p < 0.05; **, p < 0.005. Data represent the average value ± SD of three to five mice and are representative of two to four independent experiments.

FIGURE 2.

GITRL TG mice have more effector and regulatory type CD4+ T cells. A, Absolute number of cells in spleen, bone marrow (BM), thymus, peripheral (pLN), and mesenteric (mLN) lymph nodes in 4- to 8-wk-old WT (open bar) and GITRL TG (filled bar) mice. Percentage and absolute number of (B) T and B cells or (C) CD4+ and CD8+ T cells in the spleen of WT and GITRL TG mice. D and E, Percentage and absolute number of splenic regulatory (FoxP3+) and nonregulatory (FoxP3) CD4+ T cells in WT and GITRL TG mice. Nonregulatory CD4+ T cells were subdivided in naive (CD44CD62L+), effector memory (EM) (CD44+CD62L) and central memory (CM) (CD44+CD62L+) cells. F and G, Percentage and absolute number of naive, EM, and CM cells of splenic CD8+ T cells. Production of (H) IL-2, (I and J) IFN-γ, and (K and L) IL-10 by CD4+ or CD8+ T cells in WT and GITRL TG mice after stimulation with PMA-ionomycin. *, p < 0.05; **, p < 0.005. Data represent the average value ± SD of three to five mice and are representative of two to four independent experiments.

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To determine whether the strong increase of regulatory T cells in GITRL TG mice was restricted to the spleen, we analyzed the presence of these cells in bone marrow, thymus, peripheral and mesenteric lymph nodes, and liver in these mice. We found that GITRL TG mice have a systemic increase in regulatory T cell numbers, as all analyzed compartments, except for the bone marrow, showed a significantly higher fraction of FoxP3+CD4+ cells compared with WT mice (Fig. 3 A).

FIGURE 3.

Systemic increase of regulatory T cells via GITR signaling. A, The percentage of FoxP3+ cells within the CD4+ T cell compartment in spleen, bone marrow (BM), thymus, peripheral (pLN) and mesenteric (mLN) lymph nodes, and liver in WT and GITRL TG mice. B, Phenotypic analysis of splenic FoxP3 (filled graph) and FoxP3+ (open graph) CD4+ T cells for surface expression of CD25, CTLA4, PD1, OX40, CD27, CD62L, CD69, and CD103 in WT and GITRL TG mice. C, Average geometric mean fluorescence intensity (GeoMFI ± SD) or percentage positive cells (% ± SD) of these molecules on FoxP3+ CD4+ T cells. *, p < 0.05; **, p < 0.005. Data are representative of two independent experiments with each at least three mice per group.

FIGURE 3.

Systemic increase of regulatory T cells via GITR signaling. A, The percentage of FoxP3+ cells within the CD4+ T cell compartment in spleen, bone marrow (BM), thymus, peripheral (pLN) and mesenteric (mLN) lymph nodes, and liver in WT and GITRL TG mice. B, Phenotypic analysis of splenic FoxP3 (filled graph) and FoxP3+ (open graph) CD4+ T cells for surface expression of CD25, CTLA4, PD1, OX40, CD27, CD62L, CD69, and CD103 in WT and GITRL TG mice. C, Average geometric mean fluorescence intensity (GeoMFI ± SD) or percentage positive cells (% ± SD) of these molecules on FoxP3+ CD4+ T cells. *, p < 0.05; **, p < 0.005. Data are representative of two independent experiments with each at least three mice per group.

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Next, we analyzed the activation status of splenic regulatory and nonregulatory T cells. Apart from the described changes in CD44 and CD62L expression (see Fig. 2, D and E), nonregulatory CD4+ T cells in GITRL TG mice were comparable with their WT counterparts on the basis of several other costimulatory and activation molecules (Fig. 3,B). On the other hand, we found that CD4+FoxP3+ regulatory T cells from GITRL TG mice consistently expressed lower levels of CD25, CD62L, and CTLA4 compared with their counterparts in WT mice (Fig. 3, B and C). Additionally, the expression of PD1 was increased in GITRL TG mice, while a large fraction of regulatory T cells from GITRL TG mice expressed the adhesion molecule CD103 (αE integrin) on their surface (Fig. 3, B and C). The expression of OX40, CD27, and CD69 was comparable to WT mice (Fig. 3, B and C). As FoxP3+ regulatory T cells can be divided in two functionally distinct subsets, namely naive (CD62L+CD103) and effector regulatory T cells (CD62LCD103+) (25), we conclude that constitutive GITR stimulation not only leads to more regulatory T cells, but specifically stimulates the formation of regulatory T cells with an effector phenotype.

To determine whether the altered phenotype of regulatory T cells in GITRL TG mice mirrored a change in their function, we performed in vitro proliferation assays, in which WT responder T cells (CD4+CD25) were stimulated with anti-CD3/CD28 in the presence of increasing numbers of regulatory T cells (CD4+CD25+) from WT or GITRL TG mice. From these experiments it can be concluded that regulatory T cells from GITRL TG mice are fully capable of suppressing responder T cell proliferation and were equally anergic as regulatory T cells from WT mice (Fig. 4 A). This conclusion challenges previous reports, which have suggested that GITR stimulation on regulatory T cells is sufficient to abrogate their suppressive capacity (5, 6).

FIGURE 4.

Regulatory T cell function of WT and GITRL TG mice. A, Ability of purified WT and GITRL TG regulatory (CD4+CD25+) T cells to suppress WT responder (CD4+CD25) T cells. B, Ability of WT regulatory T cells to suppress proliferation of WT and GITRL TG responder T cells. Cells were cultured at different ratios for 4 days with 10 μg/ml soluble anti-CD3 mAb in the presence of irradiated WT splenocytes as APCs; for the final 16 h [3H]thymidine was added and incorporation was measured. Data are depicted as the percentage proliferation compared with responder T cells alone (average of triplicate wells ± SD) and are representative of two independent experiments. C, Wright-Giemsa staining of cryosections from thyroid gland, kidney, liver, stomach, and small and large intestine of 12-mo-old WT and GITRL TG mice. Data are representative of two mice per group.

FIGURE 4.

Regulatory T cell function of WT and GITRL TG mice. A, Ability of purified WT and GITRL TG regulatory (CD4+CD25+) T cells to suppress WT responder (CD4+CD25) T cells. B, Ability of WT regulatory T cells to suppress proliferation of WT and GITRL TG responder T cells. Cells were cultured at different ratios for 4 days with 10 μg/ml soluble anti-CD3 mAb in the presence of irradiated WT splenocytes as APCs; for the final 16 h [3H]thymidine was added and incorporation was measured. Data are depicted as the percentage proliferation compared with responder T cells alone (average of triplicate wells ± SD) and are representative of two independent experiments. C, Wright-Giemsa staining of cryosections from thyroid gland, kidney, liver, stomach, and small and large intestine of 12-mo-old WT and GITRL TG mice. Data are representative of two mice per group.

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We also analyzed the susceptibility of GITRL TG- vs WT-derived responder T cells to the suppressive capacity of WT regulatory T cells, as it has been reported that GITR stimulation allows T cells to escape suppression by regulatory T cells (11). These experiments revealed that responder T cells from GITRL TG mice could still be adequately suppressed by regulatory T cells (Fig. 4 B), thereby indicating that chronic GITR stimulation in vivo is not sufficient to induce an enduring state of insensitivity to regulatory T cell activity. Instead, it rather suggests that this previously described capacity of GITR is only effective when given together with TCR stimulation (11).

To establish whether regulatory T cells in GITRL TG mice are indeed functionally active in vivo, we examined several organs of 12-mo-old GITRL TG mice for cellular infiltrates as a sign of organ inflammation and autoimmunity. Loss of function of regulatory T cells has been associated with increased numbers of autoreactive T cells, which can induce several forms of autoimmunity, including thyroiditis, glomerulonephritis, gastritis, and inflammatory bowel disease (26, 27, 28). However, despite the strong increase of effector CD4 T cells observed in lymphoid organs of GITRL TG mice (Fig. 2), we did not find any sign of inflammation or cellular infiltration in either the thyroid gland, kidney, liver, stomach, or intestines of these mice (Fig. 4 C). This indicates that GITR stimulation does not impair the function of regulatory T cells in vivo, and considering the increase of effector T cells, this suggests that the regulatory T cells in GITRL TG mice are rather competent in preserving the homeostasis within this more active immune system.

To investigate whether the increase in effector and regulatory type T cells could be explained by an increased survival potential mediated through GITR signaling, we examined the expression profile of ∼40 pro- and antiapoptotic proteins using an advanced PCR approach called multiplex ligation-dependent amplification (29). However, this comprehensive analysis did not reveal any significant differences in the apoptotic gene expression profile of naive, effector, and regulatory CD4 T cells isolated from GITRL TG mice compared with WT mice (data not shown).

Next, we set out to determine the effects of GITR triggering on T cell proliferation. CFSE-labeled T cells from WT mice were stimulated with suboptimal anti-CD3 in a 1:1 ratio with WT or GITRL TG irradiated B cells for a period of 3 days. We found that increased GITRL availability enhanced CD4+ T cell proliferation, but did not affect CD8+ T cell proliferation (Fig. 5,A). The enhanced proliferation of CD4+ T cells via GITR engagement was no longer apparent when extra IL-2 was added to these cultures, indicating that GITR engagement affects the early proliferative capacity of CD4+ T cells. We found that increased GITR ligation raised the percentage of CD4+ T cells entering cell division (Fig. 5,B), as well as the number of divisions that these cells underwent (Fig. 5,C). Since the addition of IL-2 enhanced T cell proliferation to a similar extent as the addition of GITRL TG B cells, we questioned whether GITR stimulation induced IL-2 production. Indeed, when WT T cells were stimulated with anti-CD3, the addition of GITRL TG B cells induced almost 3-fold more IL-2 than WT B cells (Fig. 5,D). Moreover, GITR ligation increased the expression of CD25 and CD69 on CD4+ T cells, confirming the enhanced IL-2 production and increased activation induced by GITR stimulation (Fig. 5, E and F).

FIGURE 5.

GITR costimulation in vitro enhances CD4+ T cell proliferation and IL-2 production. A, Anti-CD3-induced proliferation of CFSE-labeled WT T cells cultured for 3 days in the presence of WT (filled graph) or GITRL TG (open graph) B cells with or without IL-2. Triplicate wells analyzed for (B) the average precursor frequency and (C) the average division index (i.e., the number of divisions that the dividing population underwent). D, IL-2 concentration in supernatants (average of triplicate wells ± SD) after stimulating WT T cells as in A for 24 h in the presence of a blocking Ab against CD25 to prevent IL-2 consumption. Expression of CD25 (E) and CD69 (F) on CD4+ T cells stimulated as in A after 24 h. G, Naive WT CD4+CD25 T cells from Ly5.1 mice were CFSE-labeled and injected i.v. in WT (filled graph) or GITRL TG (open graph) mice. Donor cells, gated on CD45.1+CD4+ T cells, were analyzed 3 days after transfer for expression of CFSE, CD103, and CD62L expression. A representative staining from three mice per group is shown. *, p < 0.05; **, p < 0.005.

FIGURE 5.

GITR costimulation in vitro enhances CD4+ T cell proliferation and IL-2 production. A, Anti-CD3-induced proliferation of CFSE-labeled WT T cells cultured for 3 days in the presence of WT (filled graph) or GITRL TG (open graph) B cells with or without IL-2. Triplicate wells analyzed for (B) the average precursor frequency and (C) the average division index (i.e., the number of divisions that the dividing population underwent). D, IL-2 concentration in supernatants (average of triplicate wells ± SD) after stimulating WT T cells as in A for 24 h in the presence of a blocking Ab against CD25 to prevent IL-2 consumption. Expression of CD25 (E) and CD69 (F) on CD4+ T cells stimulated as in A after 24 h. G, Naive WT CD4+CD25 T cells from Ly5.1 mice were CFSE-labeled and injected i.v. in WT (filled graph) or GITRL TG (open graph) mice. Donor cells, gated on CD45.1+CD4+ T cells, were analyzed 3 days after transfer for expression of CFSE, CD103, and CD62L expression. A representative staining from three mice per group is shown. *, p < 0.05; **, p < 0.005.

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To determine whether constitutive GITR triggering alone was sufficient to induce activation and/or proliferation of naive T cells in vivo, we isolated naive nonregulatory CD4+CD25 T cells from Ly5.1+ WT donor mice, labeled them with CFSE, and transferred them into WT or GITRL TG (Ly5.2+) recipients. Three days after transfer, naive T cells transferred to both WT and GITRL TG mice showed no CFSE dilution and did not alter their expression levels of CD62L or CD103 (Fig. 5 G). Thus, despite the fact that GITRL TG mice contained more T cells with an effector memory-like phenotype, these data imply that stimulation through GITR alone is not sufficient to induce activation or proliferation of naive T cells. However, when TCR triggering is provided, GITR stimulation enhanced the production of IL-2 and increased proliferation of CD4+ T cells in vitro.

As GITR engagement could directly and specifically enhance CD4+ T cell proliferation, we investigated the proliferative capacity of CD4+ T cells in WT and GITRL TG mice in vivo. As measured by Ki-67 expression, GITRL TG mice had more nonregulatory T cells (CD4+FoxP3) in cell cycle in the spleen than did WT mice (24 ± 2.2% in GITRL TG mice vs 14 ± 1.2% in WT mice) (Fig. 6, A and B). The fraction of regulatory T cells (CD4+FoxP3+) that stained positive for Ki-67 was not significantly different between WT and GITRL TG mice, but the fraction of Ki-67+ cells within the regulatory T cell compartment is already high (∼30%) in WT mice (Fig. 6, A and B).

FIGURE 6.

GITR triggering enhances proliferation of effector and regulatory T cells in vivo. A, Representative intracellular staining for FoxP3 and Ki-67 on splenic CD4+ T cells from WT and GITRL TG mice and (B) the percentage Ki-67+ cells of FoxP3 and FoxP3+CD4+ T cells (average ± SD). C, Representative intracellular staining for BrdU and FoxP3 on splenic CD4+ T cells from WT and GITRL TG mice 16 h after i.p. injection of 1 mg BrdU. D, The percentage BrdU+ cells of FoxP3 and FoxP3+ CD4+ T cells (average ± SD). Characterization of proliferating FoxP3 and FoxP3+CD4+ T cells based on CD62L (E) or CD103 (F) expression. The percentage of BrdU+ cells in each fraction is depicted for WT (open bar) and GITRL TG (filled bar) mice (average ± SD). Data are representative for two independent experiments with each at least three mice per group. *, p < 0.05; **, p < 0.005.

FIGURE 6.

GITR triggering enhances proliferation of effector and regulatory T cells in vivo. A, Representative intracellular staining for FoxP3 and Ki-67 on splenic CD4+ T cells from WT and GITRL TG mice and (B) the percentage Ki-67+ cells of FoxP3 and FoxP3+CD4+ T cells (average ± SD). C, Representative intracellular staining for BrdU and FoxP3 on splenic CD4+ T cells from WT and GITRL TG mice 16 h after i.p. injection of 1 mg BrdU. D, The percentage BrdU+ cells of FoxP3 and FoxP3+ CD4+ T cells (average ± SD). Characterization of proliferating FoxP3 and FoxP3+CD4+ T cells based on CD62L (E) or CD103 (F) expression. The percentage of BrdU+ cells in each fraction is depicted for WT (open bar) and GITRL TG (filled bar) mice (average ± SD). Data are representative for two independent experiments with each at least three mice per group. *, p < 0.05; **, p < 0.005.

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To directly assess T cell proliferation in vivo, WT and GITRL TG mice were injected i.p. with BrdU and incorporation of this compound was analyzed 16 h later (Fig. 6, C and D). Within the nonregulatory CD4+ T cell compartment, we found that GITRL TG mice contained more BrdU+ cells than did WT mice and this increase was most pronounced in the CD62L effector fraction (Fig. 6,C–E). A GITR-mediated increase in proliferation was even more profound for regulatory T cells, as the percentage of CD4+FoxP3+ T cells that had incorporated BrdU had more than doubled compared with WT littermates (Fig. 6, C and D). In this case, it was the CD62L+CD103 fraction of regulatory T cells that showed the most profound increase in BrdU incorporation (Fig. 6, E and F). No effect of GITRL overexpression on BrdU incorporation was found in CD8+ T cells (data not shown). Overall, these data indicate that GITR affects the numbers of regulatory T cells as well as the memory/effector pool of nonregulatory CD4+ T cells in vivo by regulating their proliferation.

To examine the significance of the expansion of both regulatory and effector CD4+ T cells on a complex immune response in vivo, GITRL TG mice were subjected to EAE, an experimental model for multiple sclerosis. We selected this model, as it induces autoimmunity by selective depletion and loss of function of regulatory T cells through treatment of immunized mice with pertussis toxin, which probably facilitates autoreactive T cells to develop and cause nerve damage in the CNS (30, 31). It is therefore conceivable that GITRL TG mice are protected from disease (i.e., limb paralysis) because they have more regulatory T cells, but it could also be that they display enhanced susceptibility to EAE compared with WT mice, because of their increased effector T cell formation. We found that regulatory mechanisms dominated the EAE response in GITRL TG mice, as they had a significant delay in disease onset compared with WT mice, which correlated with a lower clinical score (Fig. 7, A and B). Although disease was delayed in GITRL TG mice, it was not inhibited, as the cumulative incidence and clinical score were similar between both groups at the experimental endpoint (Fig. 7, A and B), suggesting that autoreactive cells did develop in these mice.

FIGURE 7.

Delayed EAE induction though GITR ligation. A, The cumulative incidence and (B) the average clinical score following EAE induction in WT (□) and GITRL TG mice (▪). Experiment depicted contains 10 animals per group and is representative of two independent experiments. Significant differences (p < 0.05) of the area under the curve were determined by Wilcoxon rank sum test (highlighted area). The absolute numbers of (C) CD4+ T cells and (D) IFN-γ- and IL-17-producing CD4+ T cells on day 5 following EAE induction in the draining (inguinal and lumbar) and nondraining (axillary and brachial) lymph nodes from WT and GITRL TG mice are shown. E, Percentage regulatory (FoxP3+) and (F) nonregulatory effector (FoxP3CD62L) T cells within the CD4+ compartment was determined in peripheral blood following immunization (day 0) and pertussis toxin injection (days 0 and 2) of WT (□) and GITRL TG (▪) mice. Data represent the average ± SD of four mice. Asterisks denote significant differences between WT and GITRL TG mice on a particular day (*, p < 0.05; **, p < 0.005); black dots denote significant differences between consecutive days for either group (•, p < 0.05; ••, p < 0.005. Cellular infiltrates in brain and spinal cord were analyzed for mice with a clinical score of 5 and in which disease onset occurred 4–5 days before experimental endpoint. G, Size of cellular infiltrates in the spinal cord of WT and GITRL TG mice. Representative immunohistochemical staining and quantification of the spinal cord of WT and GITRL TG mice shows the presence of (H) CD68+ macrophages, (I) CD4+ T cells, and (J) FoxP3+ regulatory T cells on a heamatoxylin background staining (×20 magnification). Arrow indicates a single FoxP3+ cell. For quantification purposes, at least six infiltrates were analyzed per staining per mouse.

FIGURE 7.

Delayed EAE induction though GITR ligation. A, The cumulative incidence and (B) the average clinical score following EAE induction in WT (□) and GITRL TG mice (▪). Experiment depicted contains 10 animals per group and is representative of two independent experiments. Significant differences (p < 0.05) of the area under the curve were determined by Wilcoxon rank sum test (highlighted area). The absolute numbers of (C) CD4+ T cells and (D) IFN-γ- and IL-17-producing CD4+ T cells on day 5 following EAE induction in the draining (inguinal and lumbar) and nondraining (axillary and brachial) lymph nodes from WT and GITRL TG mice are shown. E, Percentage regulatory (FoxP3+) and (F) nonregulatory effector (FoxP3CD62L) T cells within the CD4+ compartment was determined in peripheral blood following immunization (day 0) and pertussis toxin injection (days 0 and 2) of WT (□) and GITRL TG (▪) mice. Data represent the average ± SD of four mice. Asterisks denote significant differences between WT and GITRL TG mice on a particular day (*, p < 0.05; **, p < 0.005); black dots denote significant differences between consecutive days for either group (•, p < 0.05; ••, p < 0.005. Cellular infiltrates in brain and spinal cord were analyzed for mice with a clinical score of 5 and in which disease onset occurred 4–5 days before experimental endpoint. G, Size of cellular infiltrates in the spinal cord of WT and GITRL TG mice. Representative immunohistochemical staining and quantification of the spinal cord of WT and GITRL TG mice shows the presence of (H) CD68+ macrophages, (I) CD4+ T cells, and (J) FoxP3+ regulatory T cells on a heamatoxylin background staining (×20 magnification). Arrow indicates a single FoxP3+ cell. For quantification purposes, at least six infiltrates were analyzed per staining per mouse.

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To obtain more insight in the mechanism of this difference in disease progression, we examined the CD4+ T cell response in draining lymph nodes from WT and GITRL TG mice early after immunization. After 5 days, the impact of the immunization was readily visible in both mouse strains, as the draining (inguinal and lumbar), but not nondraining (axillary and brachial), lymph nodes had considerably increased in size (data not shown). In contrast to what might be expected from the delay in disease progression, we found that the development of effector CD4+ T cells was not perturbed in GITRL TG mice, as they contained even more effector CD4+ T cells than did WT mice (Fig. 7,C). Absolute numbers of regulatory CD4+ T cells were also increased in GITRL TG mice (Fig. 7,C). Restimulation of lymph node cells with PMA-ionomycin corroborated an increase in effector T cells, as GITRL TG mice contained more IFN-γ-producing cells, whereas IL-17 production was not affected (Fig. 7 D). Thus, GITR stimulation does not negatively affect the formation of autoreactive T cells following EAE induction, but, if anything, rather enhances it.

To further investigate the relative contribution of regulatory and effector CD4+ T cells in disease development in these mice, we examined the levels of both cell types in circulation following EAE induction. During the first 9 days, the fraction of regulatory CD4+ T cells decreased in both groups of mice, but GITRL TG mice continuously displayed more regulatory T cells in circulation than did WT mice (Fig. 7,E). Both GITRL TG and WT mice showed a disease-related increase of effector type CD4+ T cells in peripheral blood, but this occurred later in GITRL TG than in WT mice, which is interesting, as it correlates with the observed delay in disease development (Fig. 7 F). At 14 days after immunization, when both mouse strains had a comparable clinical score, there were no significant differences in effector or regulatory CD4+ T cells in the blood of GITRL mice compared with WT mice.

Finally, to investigate whether the observed differences in EAE signified merely a difference in time or also in quality of the immune response, we analyzed the cellular infiltrates in the CNS of mice with a similar clinical score. The infiltrated areas were comparable in size between GITRL TG and WT mice (Fig. 7,G). Moreover, these mice showed a comparable influx of macrophages (Fig. 7,H) and CD4+ T cells (Fig. 7,I) in the CNS, and the influx of FoxP3+ cells was very low in both groups (Fig. 7 J). Therefore, these data suggest that GITR engagement does not change the quality of this autoimmune response, but rather delays disease development. This is not due to a decrease in the formation of disease-related effector type CD4+ T cells, but might be related to an altered recirculation of these cells.

Since its discovery in 1997, GITR has been the focus of many studies that address its biological function in cellular immunology (2, 4, 5, 6, 8). These studies have indicated that GITR has costimulatory effects during T cell activation, but it is still not fully understood at what level GITR triggering affects both T cell activation and regulatory T cell function and how this influences immune responses in vivo. Here we describe that in vivo GITR stimulation through its natural ligand increased absolute numbers of both effector and regulatory type CD4+ T cells. Detailed analysis revealed that this accumulation was a direct consequence of enhanced proliferation of both cell types in GITRL TG mice. The increase of effector and regulatory CD4+ T cells was not at the expense of the naive CD4+ T cell pool. Together with the finding that transferred naive WT T cells do not get activated in GITRL TG mice (Fig. 5,G), this indicates that enhanced GITR triggering is not sufficient to activate naive T cells, but that this process is still fully dependent on TCR activation. When TCR triggering is provided, GITR stimulation does enhance the expansion of newly activated CD4+ T cells, as can be concluded from the in vitro stimulation (Fig. 5,A–C) and EAE immunization experiments (Fig. 7,C). However, the BrdU incorporation experiments indicate that constitutive GITR triggering also enhances proliferation of effector and regulatory CD4+ T cells during the steady-state situation (Fig. 6 E). To what extent the TCR is also required for this increased level of homeostatic proliferation in GITRL TG mice is not yet clear, as this expansion might also be driven by increased availability of cytokines like IL-2. Transfer experiments with TCR-transgenic T cells could shed further light on this issue.

GITR ligation in vivo does not affect the anergic state of regulatory T cells in vitro, nor does it influence the suppressive function of regulatory T cells (Fig. 4,A). The fact that GITRL TG mice do not develop any sign of organ inflammation or autoimmunity (Fig. 4,C), despite the expansion of effector T cells, supports this notion and indicates that regulatory T cells are fully functional in vivo in these mice and maintain homeostasis. The hypothesis that GITR regulates the size, but not the function, of the regulatory T cell pool is supported by the observation that GITR−/− mice have normally functioning regulatory T cells, but fewer absolute numbers (2, 11). In vitro studies have shown that agonistic anti-GITR Abs can induce proliferation of regulatory T cells in vitro in an IL-2-dependent manner, also without affecting their suppressive activity (5, 11). We found that GITRL expression on B cells increased IL-2 production by CD4+ T cells in vivo (Fig. 2,H) and in vitro (Fig. 5 A), which is most likely a direct effect, as GITR cross-linking with Abs can induce IL-2 production through TRAF-5 (TNFR-associated factor-5)-mediated NF-κB activation (32). These results have two important implications for our understanding of the biological function of GITR on T cells. First, since regulatory T cells depend on exogenous IL-2 for their proliferation (33), these findings indicate that GITR drives proliferation of both regulatory and effector T cells through the induction of IL-2 from the latter. Second, it explains why GITR stimulation on nonregulatory T cells allows them to escape suppression by regulatory T cells (11), since it was recently shown that regulatory CD4+ T cells exert their suppressive function through consumption of IL-2 produced by activated T cells, leading to apoptosis of the latter (34). Since GITR triggering increases the production of IL-2, nonregulatory T cells can thereby escape from or delay cytokine deprivation-induced apoptosis. These implications fit in a previously postulated model for GITR function (7), in which it was also suggested that when GITRL expression decreases at the end of an immune response, this would render effector T cells susceptible to suppression by an expanded, activated regulatory T cell pool. Transgenic GITRL expression does not allow us to test this hypothesis in our system, but it is worth following up on this idea, as it implies that GITR is indirectly involved in termination of a T cell response.

Detailed analysis revealed that GITR ligation in vivo modified the expression of several key proteins expressed by regulatory T cells (Fig. 3). We found that the IL-2 receptor is down-regulated on regulatory T cells of GITRL TG mice, which is most likely a direct consequence of increased IL-2 consumption driving enhanced proliferation (33). This is in agreement with recent findings that homeostatically proliferating regulatory CD4+ T cells in vivo express lower levels of the IL-2 receptor than do nonproliferating cells (35). Furthermore, GITRL TG mice contained more regulatory T cells with an activated phenotype, expressing low levels of CD62L and high levels of CD103 (Fig. 3) (25). This is interesting, because we found that BrdU predominantly incorporated in the CD62L+ and CD103 population of regulatory T cells in GITRL TG mice (Fig. 6). This would thus indicate that GITR ligation induces proliferation of CD62L+CD103 regulatory T cells and that during this proliferation they become activated and accumulate as CD62LCD103+ regulatory T cells. This would be in agreement with an earlier study, which described that regulatory T cells with a high turnover down-modulate CD62L after several cell divisions (36). Since CD62L is required for HEV-dependent lymphocyte entry into lymph nodes and CD103 is an integrin necessary for the homing and retention of cells at inflammatory sites, these data suggest that regulatory T cells in GITRL TG mice are more prone to enter (inflamed) peripheral tissues than secondary lymphoid organs compared with their WT counterparts. Indeed, we found that liver and bone marrow of GITRL TG mice accumulate more CD62LCD103+ regulatory T cells than do WT mice (data not shown), but since the supply of regulatory T cells is also increased in these mice, it requires more specific migration experiments to adequately address this issue.

An intriguing finding from our analysis of GITRL TG mice is that the functional consequences of GITR engagement were restricted to CD4+ T cells, as no effects on the proliferation or effector cell formation of CD8+ T cells could be detected, neither in vitro nor in vivo (Figs. 2 and 5,A–C and data not shown). This is in contrast with other studies in which a role for GITR on CD8+ T cell responses was demonstrated, using agonistic GITR Abs or GITR−/− mice (17, 37, 38). We found that both CD4+ and CD8+ T cells in GITRL TG mice had down-modulated surface expression of GITR compared with WT mice (Fig. 1 and data not shown), which indicates that GITR was functionally engaged by its ligand on both cell types. In WT mice, GITR expression is higher on CD4+ nonregulatory T cells than on CD8+ T cells (2) (and data not shown), which could be the reason why GITRL expression has a stronger effect on CD4+ T cells than on CD8+ T cells. This might also relate to the finding that the costimulatory effect of GITR cross-linking with an anti-GITR Ab is apparent at a lower anti-CD3 concentration in CD4+ T cells than in CD8+ T cells (11). Moreover, GITR up-regulation following T cell activation is dependent on CD28 engagement in CD4+ cells but not CD8+ T cells (11, 38, 39). Thus, although GITR functions on both CD4+ and CD8+ T cells, it is differently regulated in these subsets. In our hands, deliberate triggering of GITR on CD8+ T cells in vivo by its natural ligand clearly does not translate into functional consequences, or at least not as strong as the effects found on CD4+ T cells.

The synchronized expansion of regulatory and effector CD4+ T cells that is induced upon GITR stimulation might seem contradictory for protective immunity, as these cell types obviously have opposite functions. However, recent in vivo studies have shown that regulatory T cells expand with similar kinetics as effector CD4+ T cells upon HSV-2 infection (40) or immunization with Freund’s complete adjuvants (41), so that their ratio remains relatively constant. Coincident expansion of regulatory and effector T cells could be a direct consequence of responsiveness of regulatory T cells to IL-2 produced by effector T cells (42), and our data suggest that GITR could play a role in this process. It is most likely that the simultaneous increase of regulatory and effector T cells is the reason why GITRL overexpression induces a mild phenotype compared with transgenic overexpression of other members of the TNF superfamily, such as CD70, OX40L, 4-1BBL, and LIGHT, which leads to severe immunopathology induced by effector T cells (20, 43, 44, 45, 46, 47). The clinical consequence of an immune response might even depend on this ratio of effector vs regulatory T cells, as the experimental induction of both adjuvant arthritis and type 1 diabetes correlates with an increase of this ratio (48, 49, 50). The same might apply for the EAE model, as depletion of regulatory T cells resulted in enhanced disease progression and severity (51). We found that the delay in disease induction observed in GITRL TG mice was not due to a inhibition in the formation of effector CD4+ T cells in the draining lymph nodes (Fig. 7,C), but rather correlated with a delay in the increase of effector CD4+ T cells in circulation (Fig. 7, E and F). As no differences were observed in final disease severity nor cellular infiltrates in the brain parenchyma, these observations suggest that GITR stimulation enhances both formation of effector and regulatory CD4+ T cells in lymph nodes and might delay autoimmunity by regulating emigration of effector CD4+ T cells from the lymph nodes. Whether GITR triggering has a direct effect on the egress of activated T cells from lymph nodes or that this is an indirect effect mediated by regulatory T cells awaits further investigation.

In conclusion, we have shown that GITR serves as a costimulatory molecule in that it induces proliferation of regulatory as well as effector CD4+ T cells in vivo. We suggest that up-regulation of GITRL on APCs during the initiation of an immune response, through the increase of proinflammatory stimuli, enhances IL-2 production and thereby the proliferation of cognate CD4+ T cells, which also makes them less susceptible to suppression by regulatory T cells. At the same time, GITRL expression during this early phase induces the expansion of regulatory T cells, aided by the presence of exogenous IL-2 from proliferating nonregulatory T cells. These regulatory T cells might be important to re-establish the status quo of the immune system at later stages of the response.

We thank Alex de Bruin, Sten Libregts, and Dr. Koen van der Sluijs for technical assistance, as well as the staff of the animal facility of the Academic Medical Center for excellent animal care. We appreciate the dedicated assistance from Dr. Marian van Roon and coworkers in generating the GITRL TG mice. We thank Hella Aberson and Dr. Eric Eldering for the development of the murine apoptosis multiplex ligation-dependent amplification probe set and Dr. Janneke Samsom for providing us with reagents for the IL-2 ELISA. Finally, we thank Drs. Janneke Samsom, Esther Nolte-'t Hoen, Eric Eldering, and Kris Reedquist for critical reading of the manuscript and helpful discussions.

The authors have no financial conflicts of interest.

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 study was supported by a VICI Grant (to R.A.W.v.L.) and a VIDI Grant (to M.A.N.) from The Netherlands Organization of Scientific Research.

4

Abbreviations used in this paper: GITR, Glucocorticoid-induced tumor necrosis factor receptor family-related protein; GITRL, GITR ligand; EAE, experimental autoimmune encephalomyelitis; TG, transgenic; WT, wild type.

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