CD28 co-stimulation restores T cell responsiveness in NOD mice by overcoming deficiencies in Rac-1/p38 mitogen-activated protein kinase signaling and IL-2 and IL-4 gene transcription

Jian Zhang1,4, Konstantin V. Salojin1 and Terry L. Delovitch1,2,3

1 Autoimmunity/Diabetes Group, The John P. Robarts Research Institute, London, Ontario N6G 2V4, Canada
2 Departments of Microbiology and Immunology, and
3 Medicine, University of Western Ontario, London, Ontario N6A 5C1, Canada

Correspondence to: T. L. Delovitch, Autoimmunity/Diabetes Group, The John P. Robarts Research Institute, 1400 Western Road, London, Ontario N6G 2V4, Canada


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results and discussion
 References
 
Previously, we reported that T cell hyporesponsiveness induced by TCR ligation is causal to autoimmune diabetes in NOD mice. Neonatal CD28 co-stimulation reverses T cell hyporesponsiveness and protects NOD mice from diabetes by an IL-4-mediated mechanism, indicating that a deficiency in TCR signaling may be overcome by CD28/B7-2 co-stimulation in NOD T cells. To investigate which co-stimulation-induced signaling events mediate this protection, we analyzed the activity of Ras, Rac-1, mitogen-activated protein kinases (MAPK) and several transcription factors in TCR-activated NOD T cells in the presence or absence of CD28 co-stimulation. We show that CD28 co-stimulation restores normal TCR-induced activation of Rac-1 and p38 MAPK in NOD T cells. Deficiencies in TCR-induced nuclear expression of activating protein (AP)-1 binding proteins as well as activation of AP-1 and NF-AT in the IL-2 and IL-4 P1 promoters are also corrected by CD28 co-stimulation. Thus, CD28 co-stimulation reverses NOD T cell hyporesponsiveness by restoring TCR signaling leading to the activation of AP-1 and NF-AT during IL-2 and IL-4 gene transcription. Our findings provide additional evidence that CD28 co-stimulation amplifies signals delivered by the TCR and further explain the mechanism by which CD28 co-stimulation may protect against autoimmune diabetes.

Keywords: NOD mice, signal transduction, T cell hyporesponsiveness


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results and discussion
 References
 
During the spontaneous development of autoimmune type I diabetes (TID) in NOD mice, effector T cells cell infiltrate the pancreatic islets of Langerhans (insulitis), and progressively and selectively destroy the insulin-producing islet ß cells (1). The reactivity of these T cells with islet ß cell-specific autoantigens suggests that a loss of T cell tolerance to self-antigens elicits this autoimmmune disease. The precise mechanism(s) of loss of tolerance is incompletely understood, although it is clear that a failure in T cell-mediated immune regulation may be a principal factor (1).

Previously, we reported that beginning at 3–5 weeks of age, NOD T cells exhibit a proliferative hyporesponsiveness (anergy) upon TCR stimulation that is mediated by reduced IL-2 and IL-4 production (24). Addition of IL-4, a Th2-type cytokine, completely restores NOD T cell responsiveness, whereas addition of IL-2, a Th1-type cytokine, only partially restores NOD T cell responsiveness even at high doses (3,4). Treatment of NOD mice with low doses of IL-4 (50 ng/dose) protects them from insulitis and TID (3,4). These findings suggested that Th2 cells are anergic in NOD mice and that Th2 cell hyporesponsiveness mediates the onset of TID. We also showed that NOD T cell hyporesponsiveness is associated with defective TCR-mediated signaling along the protein kinase C (PKC)/Ras/mitogen-activated protein kinase (MAPK) pathway of T cell activation (5). Interestingly, we observed that a block in Ras activation mediates hyporesponsiveness in NOD T cells (5) and similar results were reported for anergic murine CD4+ Th1 clones (69). More recently, we demonstrated that TCR-stimulated NOD T cells exhibit a constitutive down-regulation of Ras-associated GDP-releasing activity as a result of the inability of the murine `son of sevenless' guanine nucleotide-releasing factor to be translocated from the cytoplasm to the plasma membrane in association with Grb-2 (10). These defects might be attributed in part to the sequestration of CD4-associated Lck from the TCR–CD3 complex and the differential activation of the Fyn–TCR{zeta}–Cbl pathway in NOD T cells (10,11).

Optimal T cell activation requires signaling through the TCR and CD28 co-stimulatory receptor (1215). In vivo studies, including those conducted using CTLA-4–Ig transgenic and CD28-knockout mice, have suggested that CD28–B7 interaction plays an important role in the generation of Th2 cells (1619). This notion was further substantiated by our observation that neonatal CD28 co-stimulation with an anti-CD28 mAb can prevent TID in NOD mice by an IL-4-dependent mechanism that promotes a shift from a Th1- to a Th2-biased environment (20). CD28 co-stimulation also completely restores NOD T cell hyporesponsiveness in vitro, indicating that T cell hyporesponsiveness (anergy) in vivo is different from the classical T cell anergy in which T cell proliferation is not reversed by CD28 co-stimulation (15). Furthermore, B7–CD28 co-stimulation has been shown to be essential for the generation of CD4+CD25+ regulatory T cells that control TID in NOD mice (21).

To further understand how CD28 co-stimulation reverses NOD T cell hyporesponsiveness and protects from TID, the activities of Ras, Rac-1, MAPK and several transcription factors involved in IL-2 and IL-4 production were analyzed in TCR-stimulated NOD T cells in the presence or absence of CD28 co-ligation. We demonstrate that the TCR-induced activation of Rac-1, p38 MAPK and the transcription factors activating protein (AP)-1 and NF-AT are deficient in NOD T cells compared to T cells from diabetes-resistant strains, and that these deficiencies may be reversed by TCR–CD28 co-stimulation. Our findings provide additional support for the notions that CD28 co-stimulation amplifies signals transmitted along the TCR pathway and as a result may prove therapeutic for protection against autoimmune diabetes.


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results and discussion
 References
 
Mice
NOD/Del, NOR/Lt and NOD.B10H-2b mice were bred and maintained in a specific pathogen-free facility at The John P. Robarts Research Institute (London, Ontario, Canada). C57BL/6J (B6) mice were purchased from Taconic (Germantown, NY) and maintained in the Animal Care facility at the University of Western Ontario (London, Ontario, Canada). Female mice were used at 6–10 weeks of age.

Antibodies and reagents
The following reagents were purchased from Santa Cruz Biotechnology (Santa Cruz, CA): rabbit polyclonal antibodies against mouse extracellular signal regulated kinase (ERK)-1, c-Jun N-terminal kinase (JNK)-1 and p38 MAPK; and mouse mAb against c-Jun/AP-1, JunB and c-Fos; GST–c-Jun (1–79), GST–activating transcription factor (ATF)-2 (1–505) and glutathione–agarose. An anti-actin mAb and myelin basic protein (MBP) were obtained from Sigma (St Louis, MO). The 145-2C11 anti-CD3 and 37.51 anti-CD28 mAb were purified by Protein G-affinity chromatography (Pharmacia Biotech, Uppsala, Sweden) of the supernatants of the B cell hybridomas kindly supplied by Dr J. Bluestone (University of Chicago, Chicago, IL) and Dr J. Allison (University of California, Berkeley, CA) respectively. Anti-v-H-Ras mAb and anti-Rac-1 mAb were purchased from Oncogene Science (Manhasset, NY) and Upstate Biotechnology (Lake Placid, NY) respectively. An anti-NF-ATc mAb (7AG) and an anti-NF-ATp (R-59) polyclonal antibody were kindly supplied by Drs G. Crabtree (Stanford University, CA) and A. Rao (Harvard University, Boston, MA) respectively.

Cell isolation and stimulation
Splenic T cells were isolated on T cell enrichment columns (R & D Systems, Minneapolis, MN) to a purity of >95% as detected by flow cytometric analysis of their CD3 surface expression. The percent distribution of CD4+ and CD8+ T cells and surface expression of TCR and CD28 was found to be very similar in NOD and control B6 mice. This ensures that the differences observed do not reflect an unappreciated change in T cell composition in these mouse strains. To decrease high basal levels of p38 MAPK activity, T cells were cultured for 5 h at 37°C in RPMI 1640 medium supplemented with 10% heat-inactivated FCS, 10 mM HEPES, 0.1 mg/ml streptomycin, 100 U/ml penicillin, 0.05 mM 2-mercaptoethanol and 2 mM glutamine (all purchased from Gibco/BRL, Burlington, Ontario, Canada). Cells were then incubated with the biotinylated anti-CD3 (1 µg/ml), anti-CD28 (1 µg/ml) or anti-CD3 + anti-CD28 mAb for 3 min at 37°C and then cross-linked with streptavidin (5 µg/ml) for 15 min. For assays of transcription factor activity, purified T cells were stimulated with biotinylated anti-CD3 in the presence or absence of anti-CD28 for 4 h at 37°C.

Ras and Rac-1 assays
Splenic T cells (5x107/ml) from NOD and B6 mice were incubated in phosphate-free RPMI 1640 medium for 1h at 37°C and labeled with [32P]orthophosphate (1 mCi/ml) for 2 h at 37°C in phosphate-free RPMI 1640 containing 10% dialyzed heat-inactivated FCS. Cells were either left unstimulated or stimulated with anti-CD3 or anti-CD3 + anti-CD28 for 15 min at 37°C as described above. Ras and Rac-1 assays were performed by polyethyleneimine–cellulose thin layer chromatography, as described (22,23). Positions of the 32P-labeled guanine nucleotides were determined according to the mobility of unlabeled GDP and GTP markers visualized under UV light.

In vitro kinase assays
After stimulation, T cells were lysed in ice-cold lysis buffer containing 1% Triton X-100, 10 mM Tris, pH 7.5, 150 mM NaCl, 2 mM EGTA, 50 mM ß-glycerophosphate, 2 mM Na3VO4, 10 mM NaF, 1 mM DTT, 1 mM PMSF, 10 µg/ml leupeptin and 10 µg/ml aprotinin. In vitro kinase assays were performed using MBP, GST–c-Jun and GST–ATF-2 fusion proteins as substrates for ERK-1, JNK-1 and p38 MAPK respectively, as described (24). Following 30 min at 30°C, reactions were terminated by the addition of SDS sample buffer, samples were boiled and kinase reaction products were resolved by SDS–PAGE. For the solid JNK assay, cell lysates were incubated for 4 h at 4°C with 10 µg GST–c-Jun pre-coupled to glutathione–agarose beads followed by an in vitro kinase reaction. Immunoblots revealed that quiescent NOD and control T cells contain equivalent amounts of the ERK-1, JNK and p38 MAPK proteins.

Nuclear expression of AP-1 proteins and nuclear translocation of NF-AT proteins
Purified splenic T cells (107/ml) were incubated with biotinylated anti-CD3 or biotinylated anti-CD3 + anti-CD28 and then cross-linked with streptavidin for either 5 min, 15 min or 4 h, as indicated. The cells were harvested, washed with ice-cold PBS and lysed during 30 min at 4°C in lysis buffer (20 mM HEPES, pH 7.5, 5 mM NaCl, 3 mM MgCl2, 1 mM DTT, 5% glycerol and 0.4% NP-40) containing protease (2.5 mM PMSF, 40 µg/ml aprotinin, 40 µg/ml leupeptin and 2 mM EDTA) and phosphatase (1 mM Na3VO4 and 10 mM NaF) inhibitors. Nuclei were separated by centrifugation at 600 g at 3°C and cytoplasmic extracts (supernatant) were added to boiling SDS–PAGE reducing sample buffer. Nuclei were rinsed in the hypotonic buffer (20 mM HEPES, pH 7.5, 5 mM NaCl and 3 mM MgCl2) containing the inhibitors and lysed in boiling sample buffer. Cytosolic and nuclear proteins were separated on 10% SDS–PAGE, transferred to nitrocellulose membranes, probed with anti-c-Fos, anti-JunB, anti-c-Jun, anti-ATF-2, anti-NF-ATc or anti-NF-ATp antibodies and identified by ECL chemiluminescence (Amersham, Piscataway, NJ), as described (25). Equivalent protein loading was verified by immunoblotting for actin. Relative amounts of proteins detected were quantitated using a Molecular Imager System and Molecular Analyst imaging software (BioRad, Hercules, CA).

Electrophoretic mobility shift assays (EMSA)
EMSA were performed as described (26). Briefly, double-stranded oligonucleotides were labeled with [{gamma}-32P]ATP using T4 polynucleotide kinase (NEB). Aliquots of these oligonuccleotides (2x104 c.p.m.) were incubated with 5 µg of nuclear protein and 2 µg of poly(dI–dC) for 30 min at 23°C in a binding reaction buffer containing 10 mM HEPES, pH 7.4, 1 mM EDTA, 50 mM NaCl, 8% glycerol, 1 mM DTT and 0.5 mM PMSF. The following IL-2 gene-derived oligonucleotides were used: mouse NF-AT, 5'-TCGAAAGAGGAAAATTTGTTTCATACAGAAGGCG-3; and mouse AP-1, 5'-TCGAGAAATTCCAGAGAGTCATCAGAAGA-3'. The oligonucleotide containing the IL-4 promoter P1 element was 5'-GACAATCTGGTGTAATAAAATTTTCCAATG-3'. Protein–DNA complexes were electrophoresed on a 5% polyacrylamide gel using 0.5xTBE buffer. In antibody supershift assays, polyclonal antibodies (1 µg/ml) to c-Jun/AP-1, JunB, NF-ATp and NF-ATc were pre-incubated with nuclear extracts (5 µg) for 20 min before addition of the oligonucleotides (26).


    Results and discussion
 Top
 Abstract
 Introduction
 Methods
 Results and discussion
 References
 
CD28 co-stimulation restores Rac-1 but not Ras activity in NOD T cells
Previously, we (5) and others (7) showed that Ras activity is significantly reduced following TCR ligation in anergic T cells. Although Ras may be activated by CD28 ligation (22), CD28 co-stimulation can also activate small GTPases, such as Rac-1 (23,27). More recently, we observed that TCR and CD28 stimulation accelerate Rac-1 GDP/GTP exchange in Jurkat T cells and primary mouse T cells (23). We therefore examined whether Rac-1 activity is also deficient in anergic NOD T cells, and whether CD28 co-stimulation restores NOD T cell Rac-1 and Ras activities to normal levels. NOD and B6 splenic T cells, which express similar levels of CD3 and CD28 on their surface as previously described (10), were labeled with [32P]orthophosphate, and 32P-labeled guanine nucleotides bound to Ras and Rac-1 were analyzed after TCR or TCR–CD28 stimulation. Small GTPases are activated as a result of the conversion of an inactive GDP-bound GTPase to an active GTP-bound state. Basal Ras activity, as assayed by the relative amount of detectable GTP-bound Ras, was found to be equivalent in unstimulated NOD and B6 T cells (Fig. 1Go, upper panel). In contrast, Ras activity was significantly increased in B6 but not NOD T cells and the levels of Ras activity were not further enhanced upon TCR–CD28 co-ligation in either NOD or B6 T cells. Thus, although CD28 stimulation alone elicits a low level of Ras activation (data not shown), Ras does not appear to play a major role in CD28-mediated co-stimulatory signaling, consistent with our observations in Jurkat T cells (23).



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Fig. 1. Effects of CD28 co-stimulation on Ras and Rac-1 GDP/GTP loading in NOD and B6 T cells. NOD and B6 splenic T cells (2x107/ml) were labeled for 2 h with [32P]orthophosphate (1 mCi/ml), incubated for 3 min at 37°C with anti-CD3 (1 µg/ml) or anti-CD3 + anti-CD28 (1 µg/ml) mAb and then cross-linked with streptavidin (5 µg/ml) for 15 min. 32P-labeled guanine nucleotides bound to Ras (A) and Rac-1 (B) were fractionated by polyethyleneimine thin layer chromatography as indicated in Methods. The relative amounts of GTP loading on Ras in (A) and GDP loading on Rac-1 in (B) are indicated below each respective panel. Data shown in (A) and (B) are each from one of two reproducible experiments.

 
Rac-1 activity was determined by assaying the level of GDPrather than GTP-bound Rac-1 (23), since the high intrinsic GTPase activity of Rho family proteins precludes the detection of GTP incorporated by these small GTPases (28). After activation, Rac-1 exists in an active GTP-bound state only for a short period of time. As a result of the high intrinsic GTPase activity, this GTP-bound state is short-lived and GTP-bound activated Rac-1 undergoes a rapid GTP to GDP nucleotide exchange after stimulation. Thus, the increased level of 32P-labeled GDP bound to Rac-1 is the only indicator of Rac-1 nucleotide exchange. We found that basal Rac-1 activities were similar in unstimulated NOD and B6 T cells (Fig. 1Go, lower panel). Whereas Rac-1 was increased only modestly after TCR stimulation in NOD T cells, Rac-1 activity was significantly enhanced in activated B6 T cells. Rac-1 activity in both NOD and B6 T cells was further elevated after TCR–CD28 co-ligation. Previously, we found that CD28 ligation alone also induces modest Rac-1 activation in B6 T cells but not in T cells from NOD control NOR or NOD.B10H-2b mice (our unpublished observations). Although NOR mice are MHC matched and congenic with NOD for 88% of their genome, they are diabetes resistant (29). Like NOD mice, NOR mice develop insulitis and their T cells are hyporesponsive to TCR stimulation (11). NOD.B10H-2b mice are congenic with NOD, MHC matched to B6 mice and their T cells are also hyporesponsive to TCR stimulation (our unpublished observations). Taken together, our data demonstrate that CD28 co-stimulation restores Rac-1 but not Ras activity in NOD T cells.

CD28 co-stimulation corrects defective activation of p38 MAPK in NOD T cells
TCR ligation activates the Ras/ERK signaling pathway, while CD28 co-ligation induces a Rac-1-mediated pathway (23,27). Since CD28 co-stimulation restores Rac-1 but not Ras activity in NOD T cells, we next analyzed whether downstream effectors of Ras and Rac, e.g. ERK and JNK/p38 MAPK respectively, are activated after TCR or TCR–CD28 stimulation. NOD and B6 splenic T cells were stimulated with anti-CD3 or anti-CD3 + anti-CD28 for 15 min at 37°C, and kinase activities associated with ERK-1 and p38 MAPK immunoprecipitates were assayed. In B6 T cells, ERK-1 was fully activated by TCR but not by CD28 stimulation and its activity was not further enhanced by TCR–CD28 co-ligation (Fig. 2Go). In contrast, ERK-1 was not activated by TCR stimulation in NOD T cells. p38 MAPK was activated only modestly in response to either TCR or CD28 ligation (24 and data not shown) in NOD and B6 T cells, and the level of p38 MAPK activation observed was appreciably lower in NOD T cells (Fig. 2Go). However, TCR–CD28 co-ligation elicited high levels of p38 MAPK activity in both NOD and B6 T cells. JNK was activated by phorbol myristate acetate stimulation but not by TCR or TCR–CD28 ligation, in agreement with our report that p38 MAPK but not JNK mediates signal integration during TCR–CD28 co-stimulation in primary murine T cells (24). It is noteworthy that the defects observed in MAPK activation are not mouse strain dependent, since T cells from another non-autoimmune mouse strain, BALB/c, responded normally to anti-TCR stimulation. In addition, T cells from NOR mice, which are congenic with NOD, displayed similar deficiencies in the activation of ERK-1 and p38 MAPK (our unpublished observations). These observations suggest that CD28 ligation activates p38 MAPK possibly via a Rac-1 signaling pathway.



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Fig. 2. CD28 co-stimulation corrects defective activation of p38 MAPK in NOD T cells. NOD (N) and B6 splenic T cells (5x106) were stimulated for 15 min at 37°C with anti-CD3 or anti-CD3 + anti-CD28 as in Fig. 1Go, lysed and immunoprecipitated with either anti-ERK-1 or anti-p38 MAPK mAb. The kinase activities associated with ERK-1 and p38 MAPK immunoprecipitates were assayed using MBP and ATF-2 as substrates. For the JNK assay, cell lysates were incubated for 4 h at 4°C with 10 µg GST–c-Jun pre-coupled to glutathione–agarose beads followed by an in vitro kinase reaction. The relative amounts of each of the proteins detected are indicated below each gel lane. Equal loading of ERK-1 and p38 MAPK was confirmed by anti-ERK-1 and anti-p38 MAPK immunoblotting. Data shown are from one of two reproducible experiments.

 
Defective activation of c-Fos, c-Jun, JunB and NF-AT in NOD T cells is restored by CD28 co-stimulation
The AP-1 transcription complex, which consists of a Jun–Jun homodimer or Jun–Fos heterodimer, is critical for regulated IL-2 expression (30). Decreased binding of AP-1 to the IL-2 promoter has been implicated in the molecular basis of T cell anergy (31). Defective transcription of the IL-2 gene is associated with impaired expression of c-Fos, FosB and JunB in anergic Th1 cells (26). We have observed that the production of IL-2 and IL-4, and mainly IL-4, is significantly increased in anti-CD28-treated NOD T cells, suggesting that CD28 signaling prevents type I diabetes in NOD mice by an IL-4-dependent mechanism (21). In view of the latter findings and those described above for AP-1, we initially investigated the role of AP-1 factors in the control of NOD T cell hyporesponsiveness by examining their nuclear expression induced following TCR and TCR–CD28 stimulation in NOD and control B6 T cells. The relative amounts of ATF-2, c-Jun, c-Fos and JunB in the nucleus were found to be reduced in NOD T cells compared to B6 T cells in response to TCR ligation (Fig. 3AGo). However, these amounts of ATF-2, c-Jun and JunB were elevated appreciably and to about equivalent levels in the nucleus of NOD and B6 T cells after TCR–CD28 co-ligation. While c-Fos nuclear expression was considerably lower in TCR-stimulated NOD T cells compared to B6 T cells, these levels were normalized in NOD and B6 T cells upon TCR–CD28 co-stimulation. In contrast, c-Fos nuclear expression was fully induced by TCR engagement and did not increase further upon TCR–CD28 co-ligation in B6 T cells. These results are consistent with the notion that c-Fos activation is mainly dependent on the canonical MAPK pathway.



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Fig. 3. Impaired TCR-induced nuclear expression of c-Fos, c-Jun, JunB and ATF-2 in NOD T cells is restored by CD28 co-stimulation. (A) NOD and B6 splenic T cells were stimulated for 4 h at 37°C with anti-CD3 or anti-CD3 + anti-CD28. Nuclear proteins were extracted as indicated in Methods, and immunoblotted with anti-c-Fos, anti-c-Jun, anti-JunB and anti-ATF-2 mAb. The relative amounts of each of the proteins detected are indicated below each gel lane. (B) B6 splenic T cells were pretreated for 15 min at 37°C with 10 µM SB203580 and were then stimulated for with anti-CD3 + anti-CD28 for 15 min. In vitro kinase assay of p38 MAPK was performed as in Fig. 2Go. Data from one of three reproducible experiments are shown.

 
Recently, we found that JNK is not activated in primary murine T cells in response to TCR and/or CD28 ligation and that p38 MAPK may be involved in the induction of c-Jun protein expression (24). To determine whether p38 MAPK is also involved in the induction of JunB expression, JunB nuclear expression was evaluated in B6 splenic T cells that were pre-treated for 15 min with 10 µM SB203580, a specific inhibitor of p38 MAPK activity (32), prior to TCR–CD28 co-stimulation for 4 h. JunB expression was completely inhibited by SB203580 (Fig. 3BGo), demonstrating that p38 MAPK activity is indeed required for the induction of JunB expression in primary murine T cells. This result is compatible with the recent observation that JunB is phosphorylated by p38 MAPK in an in vitro kinase assay (33). It is noteworthy that 10 µM SB203580 did not inhibit the activity of ERK (24 and data not shown), which can also phosphorylate JunB (33). The results demonstrates the specificity of inhibition by SB203580.

NF-ATc and NF-ATp, two NF-AT family members, regulate the transcription of both IL-2 and IL-4, particularly IL-4 (3437). Studies of NF-AT-deficient mice indicate that both NF-ATc and NF-ATp are important transcription factors that control IL-4 gene expression (36,37), and that NF-ATp may be responsible for the early expression of IL-4 (37). Whereas NF-AT proteins are phosphorylated and reside in the cytoplasm in resting cells, these proteins are dephosphorylated and translocate rapidly to the nucleus upon stimulation of cells with calcium-mobilizing agents (34). NF-AT proteins may control the transcription of AP-1 factors c-Fos and c-Jun in activated T cells (38). Accordingly, we examined the role of activation of NF-AT factors in NOD T cell hyporesponsiveness by determining the ability of NF-ATc and NF-ATp to be translocated from the cytoplasm to the nucleus in NOD and B6 T cells after TCR or TCR–CD28 stimulation. TCR-induced activation of both NF-ATc (Fig. 4AGo) and NF-ATp (Fig. 4BGo) proteins was first detectable by 5 min after stimulation and was further increased by 15 min in both NOD and B6 T cells. Note that the levels of NF-ATc and NF-ATp activation were considerably higher in B6 than NOD T cells. Interestingly, by 15 min after TCR–CD28 co-stimulation, NF-ATc and NF-ATp activities were enhanced ~2- to 3-fold respectively in both NOD and B6 T cells (Fig. 4A and BGo). These data demonstrate that a deficiency in NF-AT activation in NOD T cells can be overcome by CD28 co-stimulation. It is also interesting that TCR–CD28 co-stimulation did not trigger an increase in Ca2+ flux (our unpublished observations), consistent with the report that CD28 co-stimulation induces NF-AT activation via a Ca2+-independent mechanism (39).



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Fig. 4. Defective TCR-induced translocation of NF-AT from the cytoplasm to the nucleus in NOD T cells is restored by CD28 co-stimulation. NOD and B6 splenic T cells were either not stimulated (Unstim) or were stimulated for 5 and 15 min at 37°C with either anti-CD3 or anti-CD3 + anti-CD28 mAb. Cytosolic (Cyto) and nuclear (Nuc) proteins were separated and immunoblotted with anti-NF-ATc (A) or anti-NF-ATp (B) respectively. Note that NF-ATc translocates rapidly from the cytoplasm to the nucleus. NF-ATc in its phosphorylated state is present as several bands migrating with an apparent Mr of 160–190 kDa, while nuclear NF-ATc is dephosphorylated and migrates more rapidly (apparent Mr of 80–150 kDa) (34). Data from one of two reproducible experiments are shown.

 
CD28 co-stimulation restores DNA-binding activity of AP-1 and NF-AT in the promoter regions of the IL-2 and IL-4 genes in NOD T cells
Having shown that CD28 co-stimulation augments the expression of c-Fos, c-Jun and JunB, as well as the nuclear translocation of NF-AT, it was of interest to compare the activity of these transcription factors in NOD versus control B6 splenic T cells. Specific oligonucleotides encompassing the promoter regions of the IL-2 and IL-4 genes were used in EMSA to determine the DNA-binding capacity of these proteins to these promoter regions. TCR ligation induced only a modest activation of AP-1 and NF-AT in the IL-2 promoter, and the extent of binding observed in B6 T cells exceeded that in NOD T cells (Fig. 5A and BGo, left panels). However, a high and essentially equivalent degree of activation of AP-1 and NF-AT was noted in NOD and B6 T cells after TCR–CD28 co-ligation. The specificities of AP-1 and NF-AT binding were determined by the addition of unlabeled competitor DNA containing the AP-1 and NF-AT binding sites in the IL-2 promoter (our unpublished data). To further analyze the binding specificity of transcription factors in the AP-1 and NF-AT complexes present in TCR–CD28 co-stimulated NOD T cells, antibody supershift assays were performed. These assays showed that c-Fos, JunB and c-Jun, but not Jun D, are the major components of the AP-1 complex that bind to the IL-2 promoter after TCR–CD28 co-ligation of NOD T cells (Fig. 5A and BGo, right panels). These results also indicate that both NF-ATp and NF-ATc proteins bind to the NF-AT site of the IL-2 promoter in these activated cells.



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Fig. 5. CD28 co-stimulation restores the activities of the NF-AT and AP-1 elements in the IL-2 promoter and P1 element in the IL-4 promoter in NOD T cells. (A and B) Left panels. NOD and B6 splenic T cells were stimulated for 4 h at 37°C with anti-CD3 or anti-CD3 + anti-CD28. Nuclear proteins were reacted with specific oligonucleotides for AP-1 and NF-AT within the IL-2 promoter. Right panels. The specificity of DNA–protein complexes was confirmed by antibody-induced supershift assays of nuclear proteins from anti-CD3 + anti-CD28 co-stimulated NOD T cells. (C) Nuclear proteins from anti-CD3 + anti-CD28-activated NOD and B6 splenic T cells were subjected to EMSA using the IL-4 P1 element, and the specificity was confirmed by antibody-induced supershifts of nuclear proteins from anti-CD3 + anti-CD28 co-stimulated NOD T cells. Data shown in (A)–(C) are derived from one of three reproducible experiments.

 
Analysis of human and murine IL-4 promoters has revealed multiple regulatory sequences in the first 310 bp of the promoter (4043). Five DNA sequences (designated P0, P1, P2, P3 and P4 sites) homologous to the DNA binding sequence of NF-AT have been identified (3941). JunB regulates IL-4 expression during T cell differentiation (33) and a multimerized P1 region (AP-1 + NF-AT) confers Th2-restricted expression of transcriptional activity during T cell differentiation (44). However, AP-1 (c-Jun and JunD) family members may also bind to IL-4 P1 upon CD28 ligation (45). To further analyze how CD28 co-stimulation regulates increased IL-4 production in NOD T cells, we examined the activation of IL-4 P1 binding proteins in nuclear extracts of NOD and B6 splenic T cells after TCR and TCR–CD28 stimulation for 4 h at 37°C. EMSA conducted using murine IL-4 P1 oligonucleotides showed that only modest levels of binding to the IL-4 P1 complex occurred in TCR-stimulated NOD and B6 T cells, and the level of binding was less in activated NOD than B6 T cells (Fig. 5CGo, left panel). CD28 co-stimulation elicited similar high levels of binding to the IL-4 P1 complex in NOD and B6 T cells. Antibody supershift assays indicated that NF-ATc, NF-ATp and JunB were present in the IL-4 P1 complex of CD28 co-stimulated NOD T cells (Fig. 5CGo, right panel).

Thus, CD28 co-stimulation appears to increase IL-2 and IL-4 production in anergic NOD T cells by the restoration of activities of Rac-1 and p38 MAPK that lead to the activation of both AP-1 factors and NF-AT. Although many transcription factors are involved in the regulation of IL-4 gene transcription, we analyzed the activities of only NF-ATc and NF-ATp since they are the only ones that control transcription of both the IL-2 and IL-4 genes. Moreover, NF-AT may control AP-1 transcription (38). Our data suggest that the activities of AP-1 and NF-AT coordinately regulate the transcription of the IL-2 and IL-4 genes. Further experimentation is required to identify the factors that regulate IL-4 gene transcription induced by CD28 co-stimulation in NOD T cells.

In summary, our data demonstrate that CD28 co-stimulation restores the defect in TCR-mediated signaling and proliferative hyporesponsiveness in peripheral T cells from NOD mice by correcting the deficiencies in the nuclear expression of c-Fos, c-Jun, JunB and ATF-2 as well as NF-ATc and NF-ATp. The differences noted in kinase activities and activities of transcriptional factors between NOD and control mice were not absolute. Rather, these differences likely reflect the significantly higher threshold of TCR-mediated activation of NOD T cells, as we previously reported (46). Thus, our studies provide further insight into how CD28 co-stimulation protects NOD mice from autoimmune diabetes by an IL-4 dependent mechanism.


    Acknowledgments
 
We thank Drs J. Allison, J. Bluestone, G. R. Crabtree and A. Rao for generously providing reagents, Mrs C. Richardson for maintaining our mouse colony, and all members of our laboratory for their valuable advice and encouragement. We also thank A. Leaist for her cheerful assistance with the preparation of this manuscript. This work was supported by grants from the Canadian Diabetes Association and Juvenile Diabetes Foundation International to T. L. D. J. Z. and K. S. are recipients of Juvenile Diabetes Foundation International postdoctoral fellowships.


    Abbreviations
 
AP activating protein
ATF activating transcription factor
B6 C57BL/6J
EMSA electrophoretic mobility shift assay
ERK extracellular-regulated signal kinase
JNK c-Jun N-terminal kinase
MBP myelin basic protein
MAPK mitogen-activated protein kinases
TID type I diabetes
PKC protein kinase C

    Notes
 
4 Present address: Department of Orthopedic Surgery, Rush-Presbyterian-St Luke's Medical Center, 1653 W. Congress Parkway, Chicago, IL 60612, USA Back

Transmitting editor: A. Cooke

Received 7 August 2000, accepted 8 December 2000.


    References
 Top
 Abstract
 Introduction
 Methods
 Results and discussion
 References
 

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