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DRAK2 is a member of the death-associated protein (DAP)-like family of serine/threonine kinases. Members of this family induce apoptosis in various cell types. DRAK2, in particular, is specifically expressed in T cells and B cells, and it is differentially regulated during T cell development. To determine whether DRAK2 regulates lymphocyte apoptosis, we produced Drak2−/− mice. Contrary to our expectations, Drak2−/− T cells did not demonstrate any defects in apoptosis or negative selection; however, T cells from Drak2−/− mice exhibited enhanced sensitivity to T cell receptor-mediated stimulation with a reduced requirement for costimulation. These results provide evidence that DRAK2 raises the threshold for T cell activation by negatively regulating signals through the TCR. In contrast to other models of T cell hypersensitivity, Drak2−/− mice were remarkably resistant to experimental autoimmune encephalomyelitis (EAE). These results expose a new pathway regulating T cell activation and highlight the intricacies of induced autoimmune disease.
The acquired immune system of vertebrates evolved in response to relentless pressure applied by coevolving infectious agents (Hedrick, 2004). It is unique in exhibiting unlimited potential for molecular recognition as well as a diverse means for inducing cellular destruction. In fact, it is the only physiological system in animals capable of actively executing the loss of organismal integrity. Consequently, numerous mechanisms have evolved that regulate the activation of an immune response and limit its extent and duration. This is particularly important at the level of T cell activation. Regulatory molecules expressed in T cells participate in maintaining cellular quiescence, setting the threshold for activation and limiting the magnitude of a response through negative feedback control (Veillette et al., 2002). Defects in these molecules result in T cell hypersensitivity and often contribute to an increased propensity for autoimmune disease. Examples include phosphatases such as SHP-1, PEP, PTP-Pest, CD45, SHIP-1, and PTEN; the E3 ubiquitin ligases Cbl-b, c-Cbl, Itch, and GRAIL; inhibitory cell surface receptors CTLA-4 and PD-1; and transcription factors Foxj1, Foxp3, and LKLF (Anandasabapathy et al., 2003; Bachmaier et al., 2000; Brunkow et al., 2001; Chiang et al., 2000; Clark et al., 1999; Coyle and Gutierrez-Ramos, 2001; Fang et al., 2002; Kuo et al., 1997; Lin et al., 2004).
T cell apoptosis is another critical component of tolerance and plays an essential role at two stages in the natural life of a T cell. Developing thymocytes are induced to die as a consequence of high reactivity for self-MHC and peptide complexes, thus reducing the number of potentially autoreactive T cells that progress through development. Apoptosis is also an essential feature of the waning phase of the immune response to an infectious agent.
The signal transduction events leading to apoptosis are well studied and, unlike most other pathways, rarely involve protein phosphorylation. An exception may be apoptosis promoted by the DAP-like family of kinases. Members of this family include DAP kinase, DRP-1, ZIP kinase, DRAK1, and DRAK2 (Kogel et al., 2001). These serine/threonine kinases share homology within their N-terminal kinase domains but vary extensively in their C-terminal domains. All members of this family induced apoptosis after ectopic expression in cell lines (Cohen et al., 1997; Kawai et al., 1999; Sanjo et al., 1998). DAP kinase-related apoptosis-inducing protein kinase 1 (DRAK1) and DRAK2 were identified by EST homology searches for genes containing kinase domains similar to DAP kinase. In addition to its putative role in apoptosis, DRAK is of particular interest because it is expressed most abundantly in lymphoid organs. In fact, an analysis of gene expression patterns in 46 different human tissues revealed that DRAK (also referred to as STK17) is one of only 31 genes expressed exclusively in lymphoid tissue (Su et al., 2002).
Given that apoptosis is so critical for maintaining a well-regulated immune system and that members of the DAP-like kinase family induce apoptosis, we investigated the role of DRAK2 in lymphoid cells. In extensive studies of Drak2−/− mice, we found no evidence for an alteration in the sensitivity of thymocytes or mature T cells to apoptosis. Instead, we found that T cells deficient in Drak2 exhibited increased sensitivity to TCR signals with a reduced requirement for costimulation. From these studies we conclude that DRAK2 functions as yet another negative regulator of T cell activation. Surprisingly, despite this T cell hypersensitivity, Drak2−/− mice were highly resistant to EAE, an unexpected result in light of other models of T cell hypersensitivity.
A query of the Gene Expression Atlas created by the Genomics Institute of the Novartis Research Foundation (Su et al., 2002) for Drak2 expression in mouse tissues demonstrated that like the pattern in human tissues, Drak2 mRNA expression is highest in lymphoid organs (Figure 1A). To confirm that DRAK2 protein retained the same expression pattern as Drak2 mRNA and to further identify the subsets of cells within the lymphoid tissue that expressed DRAK2, we examined expression of DRAK2 protein by Western blot. In the thymus, DRAK2 expression is low during the double negative (DN) stage, then is increased during the double positive (DP) stage, and continues to increase in the single positive (SP) stage (Figure 1B). In peripheral lymphoid tissues, DRAK2 expression is highest in B cells but is also expressed at high levels in both CD4+ and CD8+ T cells. DRAK2 is not expressed at significant levels in NK cells, macrophages, or dendritic cells. Due to the lymphoid-specific nature of DRAK2 and the ability of its family members to induce apoptosis, we examined the function of DRAK2 by generating Drak2−/− mice. We note that analysis of the mouse genome revealed that there is no mouse homolog to human DRAK1.
Drak2-deficient mice were generated by homologous recombination in 129SVJ embryonic stem cells targeted with a construct that replaced a 400 bp fragment encompassing the intron 2/exon 3 junction and most of exon 3 (which contains the lysine residue required for ATP binding) with a neomycin resistance cassette (Supplemental Figure S1A available online at http://www.immunity.com/cgi/content/full/21/6/781/DC1/). Drak2−/− mice were recovered at the expected Mendelian frequencies and did not exhibit any gross abnormalities when compared to littermate controls.
Given the reported ability of DRAK2 to induce apoptosis after ectopic expression in cell lines (Matsumoto et al., 2001; Sanjo et al., 1998), we tested whether negative selection in Drak2−/− mice was altered in two in vivo models. First, Drak2−/− mice were crossed to H-Y TCR transgenic mice. This TCR is specific for a peptide from the male antigen bound to H-2Db, and T cells expressing this receptor are deleted before the DP stage of development in male mice (Kisielow et al., 1988). Second, the role of DRAK2 was evaluated in late-stage negative selection in AND TCR transgenic mice that expressed H-2b/s (Page et al., 1998). In this model of negative selection, thymocytes are deleted after the DP stage, but before the CD4+CD8− (CD4SP) stage. A deficiency in Drak2 did not affect the loss of self-reactive T cells in either model (data not shown). In addition, the role of DRAK2 in thymocyte apoptosis was tested in vitro with several apoptosis-inducing agents. Surprisingly, there were no significant differences in thymocyte apoptosis induced by anti-CD3, peptide-pulsed antigen-presenting cells (APC), anti-Fas, dexamethasone, staurosporine, etoposide, or υ-irradiation (Supplemental Figure S2 and data not shown). Together, these data suggest that DRAK2 does not play an essential role in induction of thymocyte apoptosis or negative selection.
Although there was no defect in thymocyte negative selection or differences in total thymic cellularity, Drak2−/− thymuses had a slight, yet statistically significant, increase in the percent of CD4SP thymocytes compared to wild-type (wt) mice (Supplemental Figure S3A). This increase coincided with a concomitant decrease in the percent of DP thymocytes and was enhanced when Drak2−/− mice were bred to mice expressing MHC class II-restricted TCR transgenes, either OT-II or AND (Supplemental Figure S3A and data not shown). However, there was no effect on subset distribution when Drak2−/− mice were bred to OT-I or P14 mice, MHC class I-restricted TCR transgenic mice (Supplemental Figure S3A). Thus, the absence of Drak2 enhanced development of CD4+ thymocytes but had little effect on CD8+ thymocytes. In addition, the levels of CD5 and CD69 were slightly increased on OT-II;Drak2−/− and OT-I; Drak2−/− DP thymocytes compared to wt DP thymocytes (Supplemental Figure S3B). This increase in CD5 and CD69 expression in the absence of Drak2 suggests that the intensity of the signal through the TCR was enhanced in Drak2−/− thymocytes. We conclude that DRAK2 negatively regulates signals initiated through the TCR.
An analysis of the lymphocytes present in the spleens of Drak2-deficient mice revealed an increase in the number of cells with characteristics of memory T cells. The percent of CD62Llo cells was consistently increased, but the difference in other memory markers, such as CD44 and CD45RB was more variable (Supplemental Figure S4A). There was no difference between Drak2−/− and wt T cells in the expression of transient markers of activation such as CD69, CD25, or LY6C (Supplemental Figure S4B). The percent of CD62Llo cells was the largest and most consistent difference between wt and Drak2−/− T cells, whereas this increase was not observed in Drak2−/− mice expressing the OT-I or OT-II TCR transgenes (data not shown). This suggests that DRAK2 does not directly affect CD62L expression and that the appearance of these cells may arise as a result of an encounter with environmental or self-antigens. This further suggests that in the absence of Drak2, the T cells responded to antigens that are otherwise ignored by the immune system.
If DRAK2 acts as a negative regulator of T cell signaling, then one would predict that Drak2−/− T cells would be hypersensitive to antigenic stimulation. To test this, we stimulated OT-I;Drak2−/− T cells in vitro with the agonist peptide OVAp or an altered form of this peptide G4. The G4 peptide varies from the agonist peptide by one amino acid residue but has a 8-fold reduced affinity for the OT-I TCR and therefore induces a weak T cell response (Rosette et al., 2001). Consistent with a deficiency in a negative regulator, Drak2−/− T cells proliferated more in response to antigenic stimulation, and this difference was greater in response to the weak agonist (Figure 2A).
The observed phenotype may be the result of hypersensitive T cells; however, it may also be caused by the APC because DRAK2 is also expressed in B cells (Figure 1B). To determine whether the effect of Drak2 ablation was T cell autonomous, the APC were exhaustively depleted by magnetic sorting, and the purified T cells were stimulated in vitro with anti-CD3 alone. Wt T cells were completely unresponsive to low doses of plate bound anti-CD3, while Drak2−/− T cells exhibited a robust proliferative response even in the absence of any source of costimulation (Figure 2B). Addition of anti-CD28 antibody evoked a response in Drak2−/− and Drak2−/− T cells that was virtually indistinguishable (Figure 2B). These data demonstrate that in the absence of Drak2, T cells responded to suboptimal stimulation that did not evoke a response in wt T cells, and this effect was independent of the APC population.
Given the fact that there were more CD62Llo cells in the spleen of Drak2−/− mice, it was possible that the hypersensitivity of Drak2−/− T cells was a consequence of an increase in the number of CD62Llo cells. To address this possibility, the CD62Llo cells were removed from the purified T cells by magnetic sorting, and the remaining T cells were stimulated in vitro with anti-CD3. Even in the absence of CD62Llo cells, Drak2−/− T cells proliferated to doses of anti-CD3 that did not induce proliferation of wt T cells, and again both proliferated equally well when costimulation was present (Figure 2C).
Because DRAK2 functions as a negative regulator of T cell activation, we compared the response of Drak2−/− T cells to T cells lacking another regulatory molecule, Cbl-b. Cbl-b reduces the amount of signaling through the TCR, and in its absence, T cells are hypersensitive to antigenic stimulation (Bachmaier et al., 2000; Chiang et al., 2000; Naramura et al., 2002). As shown previously, Cbl-b−/− T cells were hypersensitive to suboptimal doses of anti-CD3 similar to Drak2−/− T cells (Figure 2C). However, the sensitivity of the Cbl-b−/− T cells was greater than that of the Drak2−/− T cells, suggesting that DRAK2 and Cbl-b may inhibit separate signaling pathways.
In order to understand the mechanism in which DRAK2 regulates T cell activation, we examined other consequences of TCR stimulation such as cytokine production and expression of activation-induced molecules. First, we measured the amount of IL-2 produced by naive T cells in response to anti-CD3 alone. Analogous to the proliferation results, Drak2−/− T cells produced more IL-2 than wt T cells, but not as much IL-2 as Cbl-b−/− T cells (Figure 3A). This provides at least one explanation for the increased sensitivity of Drak2−/− T cells to suboptimal stimuli. We also measured the amount of IL-4 and IFNγ in unpolarized and Th1/Th2 polarized T cells. Both wt and Drak2−/− T cells produced the appropriate cytokines under Th1 and Th2 skewing conditions (Figure 3B). In addition, under nonskewing conditions, Drak2−/− T cells produced more IFN-γ, IL4, and IL-2 compared to controls.
In addition to cytokine production, we analyzed the expression level of various costimulatory molecules on purified, naive T cells before and after activation. Prior to T cell activation, the levels of the cytokine receptors, costimulatory molecules, and TNFR family members were equivalent between Drak2−/− and wt T cells (Figure 3C). However, after suboptimal stimulation with anti-CD3, the levels of various costimulatory markers varied between wt and Drak2−/− T cells. The greatest difference was in the level of CD25 (Figure 3C). The increased expression of CD25 was highest at 24 hr and maintained for at least 72 hr after stimulation (Figure 3C). Likewise, the expression levels of other costimulators including IL-7R, ICOS, CD27, OX40, and 41BB were also increased on Drak2−/− T cells in response to suboptimal stimulation (Figure 3C). Consistent with previous experiments, the levels of these costimulatory molecules were similar between wt and Drak2−/− T cells when stimulated by anti-CD3 and anti-CD28. Although several positive costimulatory molecules increased in the absence of Drak2, there were also increases in two negative regulators, PD-1 and CTLA-4. These results exemplify the robust regulation of the immune response whereby enhanced stimulation is self-limiting.
As T cells are stimulated and respond to cytokines, there is ongoing cell death. Although there was no defect in apoptosis of thymocytes, we considered the possibility that the hyperproliferative phenotype of Drak2−/− peripheral T cells resulted from diminished cell death after activation. In order to analyze the level of apoptosis associated with T cell activation, purified T cells were labeled with CFSE, stimulated with anti-CD3, and stained with Annexin V to detect the percentage of apoptotic cells at each round of cell division. T cells from wt mice did not undergo proliferation without costimulation (Figure 4A), and two effects became apparent as anti-CD28 was titrated into culture. At low concentrations of anti-CD28, a large proportion of the wt T cells progressed through the cell cycle, but a large percentage of those cells died at each round of division. As costimulation was increased, the same proportion of cells proliferated, but the apoptosis was dramatically reduced. This illustrates the two activities mediated by costimulation, proliferation associated with cytokine production and survival. The question was whether Drak2−/− T cells would divide without the concomitant cell death. Drak2−/− T cells were capable of dividing in response to anti-CD3 stimulation with minimal anti-CD28 antibody, but the Drak2−/− cells still exhibited a proportional cell death (Figure 4A). In fact, the response to anti-CD28 was virtually identical to that elicited in wt cells with the exception that the dose response was shifted to lower concentrations of agonististic anti anti-CD28 antibody.
The net recovery of cells from these experiments was plotted as a function of the time to reflect both proliferation and apoptosis (Figure 4B). Wt CD8+ T cells expanded at low concentrations of anti-CD28, whereas wt CD4+ T cells required 10100-fold more costimulation to achieve the same cell production. This was similar for Drak2−/− T cells with the exception that both of the responses required even lower concentrations of anti-CD28. The loss of Drak2 would therefore not appear to obviate the need for costimulation, but rather T cells are more sensitive to costimulation as measured by either proliferation or survival. Together these data demonstrate that the enhanced proliferation exhibited by Drak2−/− T cells did not arise from a defect in cell death associated with activation.
Although there was no defect in apoptosis of Drak2−/− T cells in vitro, we wanted to confirm that apoptosis after an in vivo immune response was also intact in the absence of Drak2. Thus, mice were infected with lymphocytic choriomeningitis virus (LCMV). At various days postinfection, the number of CD8+ T cells specific for the gp33 epitope of LCMV and the number of CD4+ T cells specific for gp61 epitope were calculated. As shown in Figure 4C, there was a robust response to LCMV in both wt and Drak2−/− mice. Furthermore, the absence of Drak2 did not affect the contraction phase of the response, supporting the in vitro data measuring cell death (Figure 4C). To be sure that there was not an alteration in the migration of T cells to nonlymphoid tissue, we also counted the number of LCMV-specific T cells in the lung, and the response in this organ was similar to that in the spleen (data not shown). The difference between the number of wt and Drak2−/− T cells that responded to LCMV was not dramatic, which would be predicted for a response in which costimulation is likely to be maximal. In addition to counting the number of LCMV-specific T cells, we also measured the amount of IFN-γ in the serum during the response. Interestingly, the level of IFN-γ present in the serum of Drak2−/− mice was greater than that of wt mice before the peak of the response and during the contraction phase (Figure 4D). This is consistent with the fact that Drak2−/− T cells make more IL-2, IL-4, and IFN-γ than wt T cells in vitro (Figure 3B), suggesting that Drak2−/− T cells retain some aspects of hypersensitivity in vivo.
In order to understand how DRAK2 influences T cell activation, we looked for differences in signaling molecules downstream of TCR activation. To narrow down the number of possible candidates, we first established whether DRAK2 was localized in the cytoplasm or the nucleus of naive T cells and if this localization changed after activation. Cytoplasmic and nuclear fractions were prepared from purified, wt T cells that were stimulated with anti-CD3 and anti-CD28. In naive, unstimulated T cells, DRAK2 was located in both the cytoplasm and nucleus (Figure 5A). Interestingly, 2 min after activation, the level of DRAK2 in the nucleus decreased and remained low for up to 48 hr (Figure 5A). The level of DRAK2 in the cytoplasm also decreased after activation, but this was not evident until 24 hr after stimulation (Figure 5A). At this time, the level of Drak2 mRNA was also lower than naive T cells (Figure 5B). These experiments demonstrate that DRAK2 could interact with both cytoplasmic and nuclear proteins and that TCR stimulation directly affects DRAK2 localization. Furthermore, this expression pattern is similar to other negative regulators of T cell activation such as Foxj1, Tob, and LKLF (Kuo et al., 1997; Lin et al., 2004; Tzachanis et al., 2001).
Next, we tested whether well-characterized signaling pathways downstream of the TCR were altered in the absence of Drak2. First, we measured calcium flux in response to TCR stimulation. In Drak2−/− CD4+ T cells, the amount of calcium fluxed in response to anti-CD3 stimulation was increased relative to wt T cells (Figure 5C); however, there was no difference when the T cells were stimulated with ionomycin, which bypasses the signal through the TCR. This provides evidence that the absence of Drak2 affects early events in TCR signaling. To test if other signaling molecules were altered, we stimulated OT-I;Drak2−/− and Drak2+/+ T cells for 5 min with peptide-pulsed APC. In these experiments, the APC were pulsed with the agonist peptide OVAp or the suboptimal peptide G4. Consistent with an increase in calcium flux, there was also an increase in the phosphorylation of ERK1/2 and c-Jun in Drak2−/− T cells compared to wt cells (Figure 5D). Like the proliferation data, the difference was greater in response to the suboptimal stimulus. Interestingly, not all TCR signaling pathways were affected as there was no difference in phosphorylation of JNK, p38, or IκB-α, a molecule in the NF-κB pathway. In addition, we did not observe a difference in the phosphorylation of molecules proximal to the TCR including TCRζ, LCK, and ZAP70 (data not shown). Although we have not identified a molecule with which DRAK2 directly interacts, in its absence, critical pathways downstream of TCR signaling are overstimulated.
The data presented thus far establish that DRAK2 is a negative regulator of T cell activation, and in its absence, T cells are hypersensitive to suboptimal stimuli. Based on other examples of T cell hypersensitivity, we predicted that Drak2−/− mice would be more susceptible to autoimmune disease. To test for spontaneous autoimmunity, we aged 9 Drak2+/+ and 15 Drak2−/− littermates for at least 15 months and analyzed the major organs of these mice for cellular infiltrate. In addition, we measured the amount of anti-nuclear and anti-dsDNA antibodies in the serum of the aged mice. Of the 15 Drak2−/− mice, only one mouse had a very mild infiltrate in the pancreas. In the remaining wt and Drak2−/− mice, there were no signs of cellular infiltration (data not shown). In addition, none of the Drak2−/− mice had higher levels of autoantibodies than the wt littermates (data not shown). Contrary to expectations, there was no obvious increase in these indicators of autoimmune disease.
In order to test if Drak2 had an impact on induced autoimmune disease, we compared the responses of wt, Drak2−/−, and Cbl-b−/− mice in EAE, a model of multiple sclerosis. In this model, C57BL/6 mice are immunized with a peptide from myelin oligodendrocyte glycoprotein (MOG) emulsified in complete Freund’s adjuvant. Induction of disease is dependent on MOG-reactive CD4+ T cells that cross the blood-brain barrier and induce inflammation in the spinal cord and the brain. The onset of disease in wt mice occurred close to day 14 after the initial immunization and reached a maximum around day 26 (Figure 6A and Table 1). As previously reported (Chiang et al., 2000), Cbl-b−/− mice were more susceptible to EAE in that a greater number of mice exhibited signs of disease, and the severity of this disease was elevated compared to wt mice (Table 1). Contrary to our expectations, Drak2−/− mice were not more susceptible to EAE, but rather they were extremely resistant to disease (Figure 6A). Both the incidence and the severity of disease were dramatically reduced in Drak2−/− mice (Table 1). In accord with symptoms, Drak2−/− mice exhibited a reduced number of infiltrating cells in the nervous system (Figure 6B), although there were similar numbers of MOG-specific cells in the spleen and the draining lymph nodes 20 days after immunization (data not shown). Therefore, both Drak2−/− and Cbl-b−/− T cells were hypersensitive to suboptimal stimuli; however, the impact this hypersensitivity had on autoimmunity was completely contradictory. In fact, thus far, a deficiency in Drak2 is the only model of T cell hypersensitivity that results in a resistance to autoimmunity.
DRAK2 is unique in that it is one of the few proteins that is specifically expressed in lymphoid tissue. Based on its similarity to DAP-like family members and direct experimentation (Sanjo et al., 1998), we predicted that DRAK2 would play a role in T cell apoptosis. However, we demonstrate here that the absence of Drak2 does not affect apoptosis of T cells in a number of different assays. This includes apoptosis of thymocytes in vitro and two models of in vivo negative selection. In addition, there was no defect in death of mature T cells after in vitro stimulation or after an in vivo immune response. Furthermore, overexpression of Drak2 in Jurkat T cells did not result in an increase of apoptosis (data not shown). Therefore, we conclude that DRAK2 does not play a critical role in T cell apoptosis.
An analysis of Drak2−/− mice revealed that it encodes a negative regulator of T cell activation, and this is supported by several distinct phenotypic differences in T cell development and activation. First, positive selection of CD4+ T cells was enhanced as illustrated by the increase in the number of CD4+ thymocytes and an increase in the levels of CD5 and CD69 expression. There was an increase in memory-like T cells that was antigen specific. The requirement for costimulation was dramatically reduced although Drak2−/− T cells retained sensitivity to costimulation (Figure 2B). In fact, the effects of costimulation, including both proliferation and survival, were identical in Drak2−/− and wt T cells, with the exception that Drak2−/− T cells responded to lower doses of anti-CD28 antibodies (Figures 4A and 4B). Control experiments showed this hypersensitivity was not due to differences in APC or due to the fact that there were more CD62Llo T cells in Drak2−/− mice. Together, these data reveal that DRAK2 functions to negatively regulate signals involved in T cell activation.
The most surprising characteristic observed in Drak2−/− mice was the resistance to EAE despite the T cell hypersensitivity. In other models of negative T cell regulation that have been tested for autoimmunity, the mice were either equally susceptible or more susceptible to disease when compared with wt mice. In fact, deficiencies in Cbl-b (Bachmaier et al., 2000; Chiang et al., 2000), Itch (Fang et al., 2002), Sts-1 and Sts-2 (Carpino et al., 2004), Mgat5 (Demetriou et al., 2001), SHP-1 (Kozlowski et al., 1993; Shultz et al., 1993; Tsui et al., 1993), Foxj1 (Lin et al., 2004), CTLA-4 (Tivol et al., 1995; Waterhouse et al., 1995), PD-1(Nishimura et al., 1999), SHIP (Helgason et al., 1998), and Pten (Suzuki et al., 2001) all resulted in an increased susceptibility to autoimmunity. Thus far, DRAK2 appears to be the only negative regulatory molecule that nonetheless conveys resistance to autoimmunity.
Due to the complexity of EAE, there are many opportunities for a regulatory molecule to affect disease. It is possible that there is a defect in CD4+ T cell expansion and survival in response to MOG immunization; however, this does not seem likely given the results of experiments measuring the T cell response in culture or in response to an infectious agent in vivo. In addition, at day 20 after immunization with MOG, there were similar numbers of antigen-reactive cells in the spleen and draining lymph nodes of Drak2−/− and wt mice (data not shown). Rather, the loss of disease appears to originate from a loss of CNS infiltration. The number of MOG-specific cells in the spinal cord and brain of the Drak2−/− mice was reduced compared to wt and Cbl-b−/− mice, and this number correlated strongly with disease. We are currently investigating the possible reasons for the difference in migration to the central nervous system (CNS), and obvious possibilities include a disregulation of adhesion molecules, chemokines, or chemokine receptors. Consistent with this possibility, there were more CD62Llo cells in the spleen of Drak2−/− mice compared to wt mice (Supplemental Figure S4).
Another way in which the absence of Drak2 could affect autoimmunity is by altering the cytokine expression of T cells. The amount and timing of cytokine expression is critical in EAE. An increase in Th2 cytokines can impart resistance to EAE and, paradoxically, IFN-γ has also been suggested to play an inhibitory role (Pedotti et al., 2003; Willenborg et al., 1996). In addition, T cells deficient in MAP kinase phosphatase 5 (Mkp5) have elevated levels of activated JNK and produce more cytokines than wt T cells; however, Mkp5-deficient mice are resistant to EAE (Zhang et al., 2004). Clearly, the role of cytokines in EAE is not completely understood, but it is conceivable that an increase in either IFN-γ or IL-4 by Drak2−/− T cells could result in decreased EAE (Figure 3B).
Drak2-deficiency could affect EAE as a consequence of altered B cell regulation. Previous experiments showed that B cells can play a protective role in EAE through the production of IL-10 (Cross et al., 2001; Dittel et al., 2000; Wolf et al., 1996). Because DRAK2 is highly expressed in B cells, it is possible that Drak2−/− B cells are also hypersensitive and produce more disease-inhibiting IL-10. Consistent with this, Drak2−/− B cells were hyperproliferative to suboptimal stimuli in vitro (data not shown).
Finally, the effect of Drak2 on autoimmunity may be through regulatory T cells. Given that Drak2−/− conventional T cells are hypersensitive to antigenic stimulation, it is possible that Drak2−/− regulatory T cells are also hypersensitive to stimulation, and the response of the regulatory T cells could dominate over that of the conventional CD4+ T cells that induce disease. However, in vitro, Drak2−/− regulatory T cells did not suppress T cell activation more effectively than wt T cells (data not shown). In addition, we did not detect an increase in the number of CD4+CD25+CD69− or CD4+GITR+T cells in the CNS or draining lymph nodes at day 20 after MOG immunization in Drak2−/− mice compared to wt mice (data not shown). However, this does not eliminate the possibility that there are more regulatory T cells present early in the response, and this is sufficient to inhibit disease.
In conclusion, the absence of Drak2 does not appear to negatively impact the health or fecundity of mice. Yet, a Drak2 deletion renders mice resistant to autoimmune disease though fully capable of mounting a normal immune response to an infectious virus. It thus presents an evolutionary enigma–what was the selective pressure to evolve a regulatory molecule that inhibits the effectiveness of an immune response and increases the propensity for autoimmunity? Whatever the answer, we note that DRAK2 constitutes a cell type-specific target that could be inhibited to treat autoimmune disease without the risks normally associated with immune inhibition.
Drak2−/− mice were generated by homologous recombination in 129SVJ embryonic stem cells. The genomic clone corresponding to Drak2 was obtained by screening a 129SVJ mouse genomic lambda phage library (Stratagene, La Jolla, CA) by using full-length Drak2 cDNA as a probe. Most Drak2−/− mice analyzed were backcrossed to C57Bl/6 at least six generations, unless indicated in the figure legend. In all cases, littermates were used as controls. Cbl-b−/− mice were a generous gift from Hua Gu and were backcrossed 12 generations to C57BL/6.
DN, CD8+, CD4+, and postselection DP thymic subsets were purified from C57BL/6 thymuses by sorting with a FACSVantage (Becton Dickinson, San Jose, CA) after magnetic depletion with MACS beads (Miltenyi Biotech, Auburn, CA). Preselection DP thymocytes were obtained from a MHC I/II−/− (β2M−/−;I-Ab−/−) thymus and sorted for expression of CD4 and CD8 with the FACSVantage. Peripheral T cells were purified from the lymph nodes by negative selection with biotin-CD4 or CD8 and B220, CD11b, DX5, and MHC class II (I-Ab) antibodies (eBioscience, San Diego, CA), followed by separation with streptavidin-conjugated magnetic beads (Miltenyi Biotech, Auburn, CA). B cells were purified from the spleen by positive selection with B220-conjugated magnetic beads (Miltenyi Biotech, Auburn, CA). NK cells and macrophages were obtained from splenocytes that were first depleted of T and B cells followed by sorting based on CD11b and NK1.1 expression. Macrophages were CD11b+ NK1.1− and NK cells were NK1.1+ with about half also expressing CD11b. Dendritic cells were obtained from collagenase-digested spleens that were then spun on a BSA gradient. CD19+ and Thy1+ cells were removed by magnetic sorting, and the remaining cells were sorted for expression of CD11c with a FACSVantage. All subsets were greater than 94% pure.
Single cells suspensions of thymus, lymph node, and spleen were stained with FITC-, PE-, TC-, PerCP-, and APC-conjugated antibodies against CD4, CD8, CD3, CD5, CD25, CD69, CD44, TCRβ, CD62L, PD-1, ICOS, 41BB, CD27, CD127 (IL-7R) Ly6C, CTLA4, and OX40 (Pharmingen, San Diego, CA; Beckman Coulter, Fullerton, CA; and eBioscience, San Diego, CA). Cells were analyzed by flow cytometry on a FACSCalibur (Becton Dickinson, San Jose, CA). Analysis was performed with FlowJo software (TreeStar, Inc., Ashland, OR).
Purified T cells or whole spleen were labeled with 5,6-carboxyfluorescein diacetate succinimidyl ester (CFSE, Molecular Probes, Eugene OR) at 2.5 μM in PBS for 10 min at 37°C, washed with RPMI containing 10% FCS, and incubated with plate bound anti-CD3 or peptide as described below.
Anti-CD3 was plate bound by first binding goat anti-hamster (Vector Laboratories, Burlingame, CA) followed by the appropriate concentration of anti-CD3 ascites (145-2C11). Anti-CD28 ascites (N37) were added at varying concentrations. After 24–72 hr, T cells were harvested and stained with Annexin V (Pharmingen, San Diego, CA).
CD4+ T cells, purified as described above, were stimulated in Th1 or Th2 conditions as described previously (Rengarajan et al., 2002). After 4 days, the cells were restimulated for 5 hr in the presence of Brefeldin A, fixed, and permeablized with Cytofix/Cytoperm Plus staining kit (Pharmingen, San Diego, CA). The treated cells were then stained with PE-anti-IL-2, PE-anti-IFN-g, or PE-anti-IL4 (eBioscience, San Diego, CA).
LCMV virus was a generous gift from Raymond Welsh. Mice were infected with 2 × 105 PFU of LCMV via intraperitoneal injection. T cells were isolated from the spleen at various days postinfection and stimulated in vitro for 5 hr in the presence of gp33 or gp61 peptides, IL-2, and Brefeldin A, then stained for intracellular IFN-γ with the cytofix/cytoperm plus kit to detect LCMV-specific T cells. In addition, blood was collected via cardiac puncture in the presence of 15 μl of 0.5 M EDTA. The blood was spun down and plasma was frozen at −80°C and used to determine levels of IFN-γ by ELISA.
T cells were purified as described above and stimulated with anti-CD3 and anti-CD28. After the various time points, the cells were harvested into ice-cold PBS to stop the reaction. The cells were lysed in a way to separate the cytoplasmic fraction from the nuclear fraction as described previously (Frasca et al., 2002).
Purified T cells were stimulated with plate-bound anti-CD3 for 24 hr, harvested, washed with PBS, and frozen at −20°C. RNA was isolated from frozen cell pellets with RNeasy mini kits (Qiagen, Valencia, CA) and labeled and hybridized as described (Wodicka et al., 1997) to mouse GeneChip arrays (Affymetrix, Santa Clara, CA). Duplicate chips were used for each condition. Data was analyzed with Resolver Gene Expression Data Analysis software (Rosetta Inpharmatics, Seattle, WA).
Calcium flux was measured by labeling cells with 2 μM Fura red, 1 μM Fluo-4, and 0.2% Pluronic (Molecular Probes, Eugene, OR) for 30 min at room temperature in serum-free media. Cells were washed twice and rested for 20 min in the dark. For stimulation, biotinylated anti-CD3 (eBioscience, San Diego, CA) was added to labeled cells on ice for 15 min, washed, then resuspended with prewarmed streptavidin, and analyzed by flow cytometry. Calcium mobilization was determined by measuring the ratio of Fluo-4:Fura red with FlowJo software (Treestar, Inc., Ashland, OR).
The APC were peritoneal macrophages that were harvested by peritoneal lavage 5 days after injection with 1 ml thioglycollate. The macrophages were labeled with CD11b-conjugated magnetic beads (Miltenyi Biotech, Auburn, CA) and positively selected on a magnetic column. The magnetically labeled APC were then incubated with 5 μM OVAp or 50 μM G4 peptides for 3 hr at 37°C. Purified CD8+ T cells were added to the APC, spun at high speed for 5 sec, and incubated for an additional 5 min at 37°C. Ice-cold PBS was added to stop the stimulation, and tubes were incubated on ice and centrifuged at 4°C. Each sample was then run over the magnet again to separate the APC and T cells. The T cell fraction was washed in PBS and lysed with the cytoplasmic lysis buffer described above. Equal amounts of total protein were loaded on a 10% SDS gel for Western blot analysis with antiphosphorylated ERK (Rabbit polyclonal), phosphorylated p38 (28B10), phosphorylated IκB-α (Ser 32, rabbit polyclonal) all purchased from Cell signaling, (Beverly, MA), ZAP-70 (Transduction Labs, Lexington, KY), and phosphorylated c-Jun (KM-1, Santa Cruz Biotechnology, Inc., Santa Cruz, CA).
Mice were immunized with 125 μg of MOG35-55 (MEVGWYRSPFSR VVHLYRNGK, Genemed Synthesis, San Francisco, CA) emulsified in complete Freund’s adjuvant containing 0.4 mg of H37Ra mycobacterium tuberculosis (Fisher Scientific, Tustin, CA) in each hind flank. The mice also received 200 ng of Bordetella pertussis toxin (List Biological, Campbell, CA) intraperitoneally immediately after immunization and again on day 2. On day 7, the mice were boosted with another 125 μg of MOG35-55 emulsified in complete Freund’s adjuvant, followed by one injection of 200 ng Bordetella pertussis toxin. Each day, the mice were scored for disease using the following scale: 0, no signs of disease; 0.5, altered gait and/or hunched appearance; 1, limp tail; 2, partial hind limb paralysis; 3, complete hind limb paralysis; 4, complete hind limb paralysis and partial forelimb paralysis. Mouse cages were coded and individual mice were scored without reference to genotype. Mice were euthanized when they reached a score of 4.
The authors thank Vincent Liuew for excellent technical assistance and John Walker for expression profiling analysis. This work was supported by National Institutes of Health grants 5T32DK07233 and AI53091-01 and by the Giannini Family Foundation.