|
|
|
|
|
|
|
||
ARTICLE |
CORRESPONDENCE David Voehringer: david.voehringer{at}med.uni-muenchen.de
 Top ABSTRACT RESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
DC) mice showed increased frequencies of CD4 single-positive thymocytes and infiltration of CD4 T cells into peripheral tissues. They developed spontaneous autoimmunity characterized by reduced body weight, splenomegaly, autoantibody formation, neutrophilia, high numbers of Th1 and Th17 cells, and inflammatory bowel disease. Pathology could be induced by reconstitution of wild-type (WT) mice with bone marrow (BM) from DC mice, whereas mixed BM chimeras that received BM from DC and WT mice remained healthy. This demonstrates that DCs play an essential role to protect against fatal autoimmunity under steady-state conditions.
© 2009 Ohnmacht et al.
The adaptive immune system can respond to a huge variety of pathogens as a result of a broad repertoire of antigen receptors on T and B cells generated by genomic recombination during development of these cells. To avoid autoimmune reactions, self-reactive lymphocytes have to be deleted or rendered tolerant. Normal polyclonal and self-tolerant T cell repertoires depend on positive and negative selection of developing T cells in the thymus. Positive selection is mediated by thymic cortical epithelial cells, whereas negative selection can occur in the cortex or in the medulla and is induced by both BM–derived cells and medullary thymic epithelial cells (1–5). It has been demonstrated that thymic DCs are very efficient in mediating negative selection of developing thymocytes (5–9). Furthermore, peripheral DCs can migrate to the thymus and contribute to negative selection (9, 10). However, because B cells (11), and perhaps other cells of hematopoietic origin, could also be involved in negative selection, it remains unclear whether a selective lack of DCs would result in impaired clonal deletion and release of self-reactive T cells into the periphery. Self-reactive T cells that escaped clonal deletion in the thymus need to be further controlled by peripheral tolerance mechanisms to prevent tissue damage (12). Under steady-state conditions, DCs are thought to play an important role in peripheral tolerance induction by various mechanisms, including production of soluble factors like IL-10, TGF-β or indoleamine 2,3-dioxygenase (13–15), induction of T reg cells (16–18), and initiation of abortive T cell proliferation resulting in clonal deletion of autoreactive T cells (19, 20). However, it remains unclear whether DCs are required to protect from spontaneous onset of autoimmunity. To address this important question, we generated constitutively DC-depleted mice. These mice rapidly developed spontaneous autoimmunity, which demonstrates for the first time that DCs are essential to maintain a self-tolerant immune system.
CD11c-Cre/R-DTA mice (
RESULTS
Top
ABSTRACT
RESULTS
DISCUSSION
MATERIALS AND METHODS
REFERENCES
Efficient ablation of DCs in CD11c-Cre/R–diptheria toxin A (DTA) mice
To determine the role of DCs for maintenance of self-tolerance, we bred mice that selectively express the Cre recombinase in DCs (CD11c-Cre mice) (21) with a strain carrying the diphtheria toxin 
chain (DTA) under control of a loxP-flanked stop cassette in the ubiquitously expressed ROSA26 locus (R-DTA mice) (22). As a consequence, DTA is expressed directly in DCs causing their constitutive elimination. DC mice, for short) lack >90% of DCs in thymus, spleen, and LNs (Fig. 1 A). Ablation affected all major DC subsets, including myeloid, lymphoid, and plasmacytoid DCs, whereas the recently described interferon-producing killer DC population (IKDC; CD11clo NK1.1+B220+) (23, 24) was not affected (Fig. 1 B). Furthermore, only few remaining Langerhans cells were detectable in epidermal sheaths of the ear from DC mice (Fig. 1 C). DCs can efficiently present foreign antigens and prime naive T cells. To determine whether DC mice are impaired in generating a primary immune response, we analyzed the efficiency of CD4 T cell priming in DC mice by adoptive transfer of OVA-specific TCR transgenic CD4 T cells (OT-II) followed by vaccination with MVA-OVA (25), a modified vaccinia virus Ankara which encodes chicken ovalbumin complementary DNA. At the peak of T cell expansion, 4 d after vaccination, total cell counts of transferred OT-II cells in the spleen of DC mice were fourfold lower as compared with control mice (Fig. 2 A). In addition, OT-II cells in DC mice were only partially activated, which is indicated by inefficient down-regulation of the surface marker CD62L (Fig. 2 B). Low expansion of OT-II cells was also observed when DC-depleted OT-II cells were transferred, indicating that remaining DCs or other APCs were able to induce a weak CD4 T cell proliferation (Fig. S1, available at http://www.jem.org/cgi/content/full/jem.20082394/DC1). Indeed, purified B cells, macrophages, or deletion-resistent DCs were all able to stimulate OT-II cell proliferation in vitro (Fig. S2). We further observed only a weak OVA-specific response of CD8 T cells upon MVA-OVA immunization of DC mice (Fig. 2, C and D). To determine the requirement of DCs for generation of an efficient immune response against gastrointestinal nematodes, we infected DC mice with the helminth Nippostrongylus brasiliensis. Adult worms were efficiently cleared from WT mice, whereas DC mice could not eliminate the parasites (Fig. 2 E). Furthermore, N. brasiliensis-infected DC mice showed impaired recruitment of eosinophils to the lung (Fig. 2 F). These immunization and infection experiments clearly demonstrate that DCs are required for efficient priming of naive T cells and execution of protective immune responses, which is consistent with their role as major APCs of the immune system. Next, we addressed the question of whether DCs are required for maintenance of immunological tolerance against self-antigens.
|
|
DC miceDC mice results in impaired negative selection, we analyzed the frequency of different thymocyte populations in DC and control mice. DC mice showed 30% higher frequencies and total numbers of CD4 single-positive (SP) thymocytes, whereas CD8 SP cells were not increased (Fig. 3, A and B; and Fig. S3, available at http://www.jem.org/cgi/content/full/jem.20082394/DC1). The TCR expression level on SP thymocytes was comparable between DC and control mice (Fig. S3). Furthermore, the expression levels of CD5, CD24, and CD69 on CD4 or CD8 SP thymocytes were comparable between DC and control mice, indicating normal thymocyte maturation (unpublished data). The increase of CD4 SP cells was not a result of recirculation of peripheral CD4 T cells because DC mice showed no significant increase of Qa2+CD24lo cells (Fig. S4) (26, 27). We generated BM chimeras to determine whether the increased frequency of CD4 SP cells is linked to hematopoietic cells and whether it can be prevented by coadministration of BM from WT mice. Lethally irradiated WT mice received either BM from DC mice or WT mice (single chimeras) or a 1:1 mixture of BM from DC and WT donors (mixed chimeras). 9 wk after reconstitution, the frequency of CD4 SP cells in DC
WT chimeras was 21%, as compared with 11% in mixed DC + WTWT chimeras or WTWT chimeras (Fig. 3 C). The ratio of CD4 SP/CD8 SP thymocytes was significantly increased in DC mice and DCWT chimeras as compared with WT mice or mixed chimeras (Fig. S5). This supports the concept that DCs contribute mainly to clonal deletion of CD4 but not CD8 T cells (28), although thymic epithelial cells can also mediate negative selection of both T cell subsets (29, 30).
|
DC mice. To this end, CD11c-Cre mice on C57BL/6 background were crossed once with R-DTA mice on a BALB/c background carrying the endogenous retroviruses Mtv-6, 8, and 9, which encode sAgs that cause deletion of Vβ3+ (Mtv-6) or Vβ11+/Vβ12+ (Mtv-8/9) thymocytes. Because only Mtv-6 is expressed by thymic DCs (31), we expected impaired deletion of Vβ3+ CD4 SP thymocytes in DC mice, whereas deletion of Vβ11+ and Vβ12+ thymocytes should not be affected. Indeed, 
60% of Vβ3+ CD4 SP cells was deleted in DC-sufficient mice whereas only 25% was deleted in DC mice (Fig. 3 D). In contrast, negative selection of Mtv-8/9–responsive Vβ11+ and Vβ12+ cells appeared to be DC independent. It has been estimated that 50–60% of initially positively selected CD4 T cells are subsequently deleted by hematopoietic cells (29). As we detected only a 30% increase of CD4 SP cells in DC mice, it remains possible that other cell types, including thymic B cells, also participate in negative selection (11). We found no difference in the frequency of natural T reg cells in the thymus of DC mice as compared with control mice (Fig. 3 E), which supports previous observations demonstrating that intrathymic natural T reg cell development occurs independently of MHC class II expression on BM-derived cells (32, 33).
DC mice develop severe pathology
DC mice were born at the expected Mendelian ratio but they appeared smaller in size, had a hunched posture, and their body weight was reduced by 30% as compared with controls at 6 wk of age (Fig. 4 A). 40% of the mice died within 8 wk of age (Fig. 4 B). Macroscopic analysis of peripheral lymphoid organs of 6–8-wk-old DC mice revealed that spleen and LNs were enlarged (Fig. 4 C). Small intestine and colon were inflamed and the fat pads were missing (Fig. 4 D). Histological analysis of colon, small intestine, kidney, and liver revealed the presence of cellular infiltrates (Fig. 4 E).
|
DC and control mice (Fig. 5 A). However, the number of F4/80+ macrophages was increased twofold and Gr-1+ cells increased more than 20-fold (Fig. 5 A). Further analysis revealed that the expanded Gr-1+ population consisted mainly of Gr-1hi cells, which represent neutrophils with segmented nuclei (Fig. S6, A and B, available at http://www.jem.org/cgi/content/full/jem.20082394/DC1). These cells could also be found in peripheral organs like kidney, liver, small intestine, and colon of DC mice (Fig. S6 C). To exclude the possibility that unspecific expression of DTA in nonhematopoietic cells was responsible for these phenotypic alterations and to determine whether pathology can be prevented in the presence of DCs, we generated single DCWT and mixed DC + WTWT BM chimeras. Importantly, reduced body weight and neutrophilia was only observed in single chimeras and not in mixed chimeras, suggesting that DCs originating from WT BM provide protection from spontaneous development of pathology (Fig. 5 C).
|
–producing CD4 T cells in DC miceDC mice seem to be impaired in negative selection of CD4 T cells, we considered it likely that pathology was caused by autoreactive CD4 T cells that left the thymus and could not be controlled by peripheral tolerance mechanisms. Flow cytometric analysis of peripheral T cells from 6–8-wk-old DC mice revealed that CD4 T cells (but not CD8 T cells) showed an increased frequency of activated cells (CD62LloCD44hi) (Fig. 6 A). The frequency of IFN-– and IL-17A–producing cells was increased 10-fold in DC as compared with control mice, which indicates that CD4 T cells had differentiated toward a Th1 or Th17 effector phenotype, respectively (Fig. 6, B and C). It has been shown that Th17 cells are associated with autoimmune disorders (34) and promote neutrophilia (35), which is consistent with the high number of neutrophils in DC mice. Interestingly, the spontaneous increase of Th1 and Th17 cells occurred despite an almost normal frequency of peripheral T reg cells (Fig. 6 B). Analysis of the TCR-Vβ repertoire revealed no major differences between DC and control mice, which demonstrates that the effector T cell populations in DC mice did not result from oligoclonal T cell expansion (Fig. S7, available at http://www.jem.org/cgi/content/full/jem.20082394/DC1).
|
DC as compared with control mice, which suggests that pathology might be caused by tissue infiltration of autoreactive CD4 T cells (Fig. 7). The frequency of T reg cells among total lymphocytes in the small intestine was increased in DC as compared with control mice and, therefore, the inflammatory immune response does not seem to be caused by a local lack of T reg cells in DC mice (Fig. S8, available at http://www.jem.org/cgi/content/full/jem.20082394/DC1). We also observed a significant reduction of CD8 T cells in the small intestine of DC mice, which could at least partially be explained by infiltration of large numbers of CD4 T cells resulting in a relative decrease of other lymphocyte populations. In addition, DCs in the small intestine might be required for survival or recruitment of CD8 T cells. Increased CD4 T cell infiltration was also observed in single DCWT BM chimeras as compared with mixed DC + WTWT BM chimeras, providing further evidence that DCs are required to prevent spontaneous CD4 T cell activation and infiltration of peripheral tissues (Fig. S9).
|
DC miceDC mice despite the near complete ablation of DCs, we measured the concentration of immunoglobulin isotypes in the serum. All isotypes were elevated with the largest increase observed for IgM and IgG2a (Fig. 8 A). To determine whether DC mice generated autoantibodies, we performed Western blot analysis where protein extracts from kidney, liver, and spleen of Rag-deficient mice was probed with serum from four individual DC mice and one negative littermate as control. Several distinct bands could be observed that varied between different tissues (Fig. 8 B). Individual mice showed only partially overlapping patterns, indicating that autoantigens from individual mice recognized different self-antigens. Staining of HEp-2 cells with serum from DC and control mice revealed the presence of antinuclear antibodies (ANAs) in DC mice (Fig. 8 C). To further define whether ANAs from individual mice recognize different nuclear antigens, we stained tissue sections from the liver of Rag-deficient mice with sera from DC mice. We observed a variety of staining patterns of subnuclear structures with sera from individual mice, which complements the result of the Western blot analysis (Fig. 8 D). Because intestinal inflammation was observed in most DC mice, we further analyzed the staining pattern of autoantibodies directed against intestinal antigens. Tissue sections from the small intestine of Rag-deficient mice were stained with serum from three individual DC mice. The sera stained either nuclei (mouse #1), the lamina propria (mouse #2), or the epithelial layer (mouse #3), whereas no staining was observed with serum from negative littermates (Fig. 8 E). Collectively, DC mice generated a variety of autoantibodies directed against nuclear and tissue-specific antigens with specificities that differed between individual mice.
|
|
DISCUSSION |
|---|
|
TopABSTRACTRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
DC mice has also been crossed to another strain of conditional ROSA26-DTA mice that encodes DTA and the neomycin resistance gene behind a loxP-flanked LacZ cassette (37). These mice developed a myeloid proliferative syndrome indicated by an increased frequency of Gr-1+ cells in a manner similar to that in DC mice (38). However, they remained healthy and no signs of autoimmunity were reported. This apparent discrepancy may be explained by the fact that plasmacytoid DCs and Langerhans cells were not ablated in these mice so that immune tolerance may still be functional. Both plasmacytoid DCs and Langerhans cells have been shown to provide powerful tolerogenic signals to prevent activation of alloreactive or allergen-specific T cells, so it seems likely that they also control self-reactive T cells (39–42).
Negative selection by DCs in the thymus is a very efficient process. Using reaggregate cultures, it has been demonstrated that central tolerance against alloantigens can be achieved by only one DC per 200 thymocytes (6). Furthermore, deletion of peptide MHC-specific TCR transgenic cells was still efficient when the frequency of thymic APCs was lowered to 1% of normal numbers (8). Therefore, the potential requirement of DCs for negative selection can only be observed in mice with almost complete depletion of thymic DCs. The efficient ablation of thymic DCs in DC mice may result in release of autoreactive T cells into the periphery where they could be primed by the few remaining DCs or other APCs. Importantly, autoimmunity was not caused by unspecific DTA expression in nonhematopoietic cells or by bystander toxicity of DTA released from dieing DCs because autoimmunity could be induced by reconstitution of WT mice with BM from DC mice but not by cotransfer of BM from WT and DC mice. The role of DCs in regulation of autoimmunity remains controversial. It has been shown that extending the lifespan of DCs can also result in autoimmunity in some (43, 44) but not all transgenic models (45).
Why does autoimmunity develop when the life span of DCs is shortened or prolonged? When DC life span is prolonged, they might increase their expression of costimulatory molecules which could lead to priming of self-reactive T cells (43). Our results indicate that if DC life span is too short so that they cannot build up a normal-sized pool of DCs, tolerance induction of CD4 T cells is impaired. This can occur at two levels: in the thymus (impaired negative selection) and in the periphery (lack of tolerogenic DCs). We observed that DC mice have significantly more CD4 SP thymocytes as compared with control mice. In addition, DC mice show less efficient deletion by DC-restricted endogenous retroviral sAgs. Both observations are consistent with the view that DCs play an important role in negative selection of CD4 T cells. Mice that express MHC class II only on thymic cortical epithelial cells generate autoreactive CD4 T cells, yet these mice do not develop spontaneous autoimmunity as a result of lack of MHC class II expression on APCs (3). Transfer of these autoreactive T cells into nonirradiated WT mice results in only mild autoimmunity, which is different from the severe phenotype of DC mice we describe in this paper (46). Therefore, it seems likely that in addition to their important role in negative selection, DCs might be required to maintain peripheral tolerance. This assumption is further supported by our recent observation that autoreactive T cells accumulated in mice with DCs that were defective for uptake of apoptotic cells (47).
It remains unclear at the moment whether the activated CD4 T cells were primed by the few remaining DCs or by other hematopoietic MHC class II+ APCs, such as B cells or macrophages, or whether they recognize MHC class II directly on tissue cells such as enterocytes (48–50). This latter possibility might explain why the intestine is particularly vulnerable to fatal autoimmune attack. However, it also remains possible that the barrier function of the intestines is compromised in DC mice, which might result in a local inflammatory response by commensal gut flora and subsequently initiate autoimmunity. Marguerat et al. (50) have shown that ulcerative colitis develops in MHC class II KOWT BM chimeras at 6–8 wk after reconstitution with heavy T cell infiltration into the lamina propria, which supports the possibility that antigen presentation by nonhematopoietic cells might be sufficient for priming of autoreactive T cells. In addition, conditional ablation of MHC class II in hematopoietic and endothelial cells resulted in a similar increase of CD4 SP thymocytes to that described in this paper for DC mice (51). However, peripheral T cells were not spontaneously activated, suggesting that priming of autoreactive T cells requires MHC class II expression on hematopoietic or endothelial cells.
It has been demonstrated that mice lacking integrin vβ8 on DCs have reduced frequencies of T reg cells in the colon and develop late onset of autoimmune colitis (15). Furthermore, depletion of T reg cells results in increased numbers and spontaneous activation of DCs under steady-state conditions and fatal autoimmunity (52). These results suggested that T reg cells prevent autoimmunity by modulation of DC activation. This model does not apply to the situation described in this paper because peripheral T reg cells were reduced only by 20–30% in spleen and LNs of DC mice and autoimmunity developed despite (and as a result of) the absence of DCs. However, it remains possible that the repertoire of T reg cells is altered or that a critical subpopulation of T reg cells is missing in DC mice so that some autoreactive T cells may escape the control by T reg cells. The spontaneously autoreactive T cells observed in DC mice illustrate that DCs play an important role for tolerance induction at multiple levels throughout T cell development and homeostasis. Thymic DCs remove thymocytes with a high TCR avidity for self-antigens. As a result, peripheral T cells express TCRs with relatively low avidity for self-antigens but they probably still need to be kept under control by peripheral DCs to avoid autoimmunity.
Collectively, our results demonstrate for the first time that DCs play a central role in preventing spontaneous autoimmunity under steady-state conditions and suggest a functional coevolution between the T cell compartment and DCs. The result of this development is the efficient establishment of immune tolerance against self-antigens by DCs. In the absence of DCs, self-reactive CD4 T cells can become prevalent and induce devastating autoimmunity.
|
MATERIALS AND METHODS |
|---|
|
TopABSTRACTRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
subunit (DTA) into the genomic ROSA26 locus behind a loxP-flanked STOP cassette by homologous recombination in ES cells (129/Sv background) (22). Therefore, cells were immediately killed when they expressed the Cre recombinase. R-DTA mice were backcrossed four generations to C57BL/6 background. CD11c-Cre/R-DTA mice (DC mice) were generated by crossing R-DTA mice (22) to CD11c-Cre mice (provided by B. Reizis, Columbia University, New York, NY) (21). Unless indicated otherwise, these mice were maintained on a mixed 129/Sv x C57BL/6 background. Control mice were negative littermates from this breeding. Rag-deficient mice on a C57BL/6 background were originally obtained from The Jackson Laboratory. Mice were housed according to institutional guidelines and used at 6–8 wk of age unless otherwise indicated. The animal experiments were approved by the Regierung von Oberbayern.
BM chimeras.
2 x 106 BM cells from indicated mice were injected into lethally (1,200 rad) irradiated recipient mice. Chimeras were kept with antibiotica containing drinking water (2 g/liter neomycin sulfate and 100 mg/liter polymyxin B). Mice were analyzed 9–10 wk after reconstitution.
OT-II transfer and MVA-OVA vaccination.
Total splenocytes from OT-II/Thy1.1 mice containing 1.5 x 106 TCR transgenic cells were transferred into DC or control mice that were injected 1 d later i.v. with 107 infectious units of MVA-OVA (25). 4 d later, splenocytes were analyzed by staining with anti-CD4, anti-Thy1.1, and anti-CD62L to determine the frequency and activation status of transferred OT-II cells. To monitor the expansion of endogenous OVA-specific CD8 T cells, peripheral blood samples of a separate set of mice were stained with anti-CD8 and Kb-OVA257-264 pentamers (ProImmune) at different days after immunization with 107 infectious units of MVA-OVA.
N. brasiliensis infection.
Third-stage larvae (L3) of N. brasiliensis were washed extensively in sterile 0.9% saline at 37°C, and 500 organisms were injected s.c. into mice. Mice were provided with antibiotics-containing water (2 g/liter neomycin sulfate and 100 mg/liter polymyxin B sulfate; Sigma-Aldrich) for the first 5 d after infection.
Flow cytometry.
Single cell suspensions were prepared by collagenase digestion (small intestine, colon, and kidney), by liberase CI and DNase I digestion (Roche; Fig. 1), or by mechanical dispersion, incubated with anti-CD16/CD32 blocking antibody (2.4G2) for 5 min at room temperature, and stained with the corresponding antibody mixtures on ice. The following monoclonal antibodies were purchased from Invitrogen, unless otherwise indicated: PE–Alexa Fluor 700–labeled anti-CD4, APC- or FITC-labeled anti-CD8, biotinylated anti-CD11b (eBioscience), PE- or APC-labeled anti-CD11c (BD), APC-labeled anti-F4/80 (eBioscience), PE-labeled anti–Siglec-F (BD), PE-labeled anti-CD25 (eBioscience), PE-labeled anti-CD44, FITC-labeled anti-CD62L, Alexa Fluor 647–labeled anti-B220, biotinylated anti–Gr-1, biotinylated anti–MHC class II (clone M5/114.15.2; eBioscience), biotinylated anti-NK1.1 (clone PK136; eBioscience), FITC-labeled anti-TCR screening panel (BD), PE-Cy5.5–labeled streptavidin, and PE- or APC-labeled streptavidin (SouthernBiotech). CFSE staining was performed as previously described (53). Intracellular staining for Foxp3 was performed with the anti–mouse/rat Foxp3 staining set (eBioscience). Intracellular cytokine staining was performed with FITC-labeled anti–IFN- (XMG1.2; eBioscience), PE-labeled anti-IL-4 (BVD6-24G2; Invitrogen), and Alexa Fluor 647–labeled anti–IL-17A (eBioTC11-18H10.1; eBioscience) after cells had been stimulated for 4 h with 1 µg/ml ionomycin and 40 ng/ml PMA with Brefeldin A added at 5 µg/ml for the last 2 h. Cells were analyzed on a FACSCalibur instrument (BD).
Histology.
Cryosections of tissues fixed in 4% paraformaldehyde were stained with a Hemacolor staining kit for microscopy (Merck) according to manufacturer's instructions. Staining for Langerhans cells in the ear was performed as previously described (54), except that biotinylated anti–I-A/I-E (M5/114) and Alexa Fluor 555–labeled streptavidin were used for detection. CD4+ T cells in the small intestine were detected by staining 5-µm cryosections with biotinylated anti-CD4 (RM4-5; Invitrogen) followed by Cy3-conjugated streptavidin (Jackson ImmunoResearch Laboratories) for detection. ANAs were detected using HEp-2 cells fixed on glass slides (EUROIMMUN AG). The staining pattern of autoantibodies was determined by staining of tissue sections of the liver or small intestine from Rag-deficient mice with serum from DC or control mice followed by goat anti–mouse Cy3. Pictures were acquired with a 10x/0.40 U Plan SApo or a 60x/1.35 U Plan SApo objective on a microscope (BX41; Olympus) equipped with a camera (F-View II; Olympus) and cell^F software (Olympus). Original magnification was 80 or 480x, respectively.
Western blot.
Tissue samples were homogenized on ice in lysis buffer containing 50 mM Tris, pH 8.0, 150 mM NaCl, 1% CHAP, and a protease inhibitor cocktail (Complete mini tablets). The lysates were centrifuged for 10 min at 18,600 g. The supernatants were mixed with sample buffer (62.5 mM Tris-HCL, pH 6.8, 25% glycerol, 2% SDS, and 5% β-mercaptoethanol), separated by a 10% SDS-PAGE, and 0.2 µm were electrophoretically transferred onto an Immun-Blot PVDF membrane (Bio-Rad Laboratories). The membranes were blocked with 5% nonfat dried milk in PBS for 1 h, washed with PBS/0.05% Tween20, and incubated for 1 h in a 1:100 dilution of sera from individual mice. After washing with PBS/Tween20, the bound antibodies were reacted with donkey anti–mouse IgG HRP (1:3,000; Jackson ImmunoResearch Laboratories) for 1 h and revealed with a chemiluminescence reagent (Western Lightning; PerkinElmer).
ELISA.
Immunoglobulin isotypes were determined using a commercial ELISA kit (SouthernBiotech). Serum IgE levels were analyzed using the mAb B1E3 for coating and the biotinylated mAb EM95 for detection.
Statistical analysis.
P-values were calculated with Student's t test using SigmaPlot software (SPSS Inc.).
Online supplemental material.
Fig. S1 shows the proliferation of transferred OT-II cells in DC mice. Fig. S2 shows in vitro stimulation of OT-II cells with different APCs from DC mice. Fig. S3 shows total number and TCR expression level on SP thymocytes. Fig. S4 shows the frequency of mature and immature CD4 SP thymocytes. Fig. S5 shows the ratio of CD4/CD8 SP thymocytes in single and mixed chimeras. Fig. S6 describes the phenotype of expanded Gr-1+ cells. Fig. S7 shows the TCR-Vβ repertoire in DC and control mice. Fig. S8 shows the frequency of Foxp3+ cells in the small intestine of DC and control mice. Fig. S9 shows T cell infiltration in peripheral organs of single and mixed chimeras. Online supplemental material is available at http://www.jem.org/cgi/content/full/jem.20082394/DC1.
|
Acknowledgments |
|---|
This work was supported by the Deutsche Forschungsgemeinschaft with an Emmy Noether grant to D. Voehringer (Vo944/2-2), with an SFB 455 grant to T. Brocker, and with an SFB 456 grant to I. Drexler.
The authors have no conflicting financial interests.
Submitted: 23 October 2008
Accepted: 3 February 2009
|
REFERENCES |
|---|
|
TopABSTRACTRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
|---|
Anderson, G., N.C. Moore, J.J. Owen, and E.J. Jenkinson. 1996. Cellular interactions in thymocyte development. Annu. Rev. Immunol. 14:73–99.[CrossRef][Medline]
Gallegos, A.M., and M.J. Bevan. 2004. Central tolerance to tissue-specific antigens mediated by direct and indirect antigen presentation. J. Exp. Med. 200:1039–1049.
Laufer, T.M., J. DeKoning, J.S. Markowitz, D. Lo, and L.H. Glimcher. 1996. Unopposed positive selection and autoreactivity in mice expressing class II MHC only on thymic cortex. Nature. 383:81–85.[CrossRef][Medline]
Volkmann, A., T. Zal, and B. Stockinger. 1997. Antigen-presenting cells in the thymus that can negatively select MHC class II-restricted T cells recognizing a circulating self antigen. J. Immunol. 158:693–706.[Abstract]
McCaughtry, T.M., T.A. Baldwin, M.S. Wilken, and K.A. Hogquist. 2008. Clonal deletion of thymocytes can occur in the cortex with no involvement of the medulla. J. Exp. Med. 205:2575–2584.
Matzinger, P., and S. Guerder. 1989. Does T-cell tolerance require a dedicated antigen-presenting cell? Nature. 338:74–76.[CrossRef][Medline]
Brocker, T., M. Riedinger, and K. Karjalainen. 1997. Targeted expression of major histocompatibility complex (MHC) class II molecules demonstrates that dendritic cells can induce negative but not positive selection of thymocytes in vivo. J. Exp. Med. 185:541–550.
Merkenschlager, M., M.O. Power, H. Pircher, and A.G. Fisher. 1999. Intrathymic deletion of MHC class I-restricted cytotoxic T cell precursors by constitutive cross-presentation of exogenous antigen. Eur. J. Immunol. 29:1477–1486.[CrossRef][Medline]
Proietto, A.I., S. van Dommelen, P. Zhou, A. Rizzitelli, A. D'Amico, R.J. Steptoe, S.H. Naik, M.H. Lahoud, Y. Liu, P. Zheng, et al. 2008. Dendritic cells in the thymus contribute to T-regulatory cell induction. Proc. Natl. Acad. Sci. USA. 105:19869–19874.
Bonasio, R., M.L. Scimone, P. Schaerli, N. Grabie, A.H. Lichtman, and U.H. von Andrian. 2006. Clonal deletion of thymocytes by circulating dendritic cells homing to the thymus. Nat. Immunol. 7:1092–1100.[CrossRef][Medline]
Kleindienst, P., I. Chretien, T. Winkler, and T. Brocker. 2000. Functional comparison of thymic B cells and dendritic cells in vivo. Blood. 95:2610–2616.
Steinman, R.M., D. Hawiger, and M.C. Nussenzweig. 2003. Tolerogenic dendritic cells. Annu. Rev. Immunol. 21:685–711.[CrossRef][Medline]
Akbari, O., R.H. DeKruyff, and D.T. Umetsu. 2001. Pulmonary dendritic cells producing IL-10 mediate tolerance induced by respiratory exposure to antigen. Nat. Immunol. 2:725–731.[CrossRef][Medline]
Munn, D.H., M.D. Sharma, J.R. Lee, K.G. Jhaver, T.S. Johnson, D.B. Keskin, B. Marshall, P. Chandler, S.J. Antonia, R. Burgess, et al. 2002. Potential regulatory function of human dendritic cells expressing indoleamine 2,3-dioxygenase. Science. 297:1867–1870.
Travis, M.A., B. Reizis, A.C. Melton, E. Masteller, Q. Tang, J.M. Proctor, Y. Wang, X. Bernstein, X. Huang, L.F. Reichardt, et al. 2007. Loss of integrin alpha(v)beta8 on dendritic cells causes autoimmunity and colitis in mice. Nature. 449:361–365.[CrossRef][Medline]
Dhodapkar, M.V., R.M. Steinman, J. Krasovsky, C. Munz, and N. Bhardwaj. 2001. Antigen-specific inhibition of effector T cell function in humans after injection of immature dendritic cells. J. Exp. Med. 193:233–238.
Jonuleit, H., E. Schmitt, G. Schuler, J. Knop, and A.H. Enk. 2000. Induction of interleukin 10–producing, nonproliferating CD4+ T cells with regulatory properties by repetitive stimulation with allogeneic immature human dendritic cells. J. Exp. Med. 192:1213–1222.
Mahnke, K., Y. Qian, J. Knop, and A.H. Enk. 2003. Induction of CD4+/CD25+ regulatory T cells by targeting of antigens to immature dendritic cells. Blood. 101:4862–4869.
Steinman, R.M., and M.C. Nussenzweig. 2002. Avoiding horror autotoxicus: the importance of dendritic cells in peripheral T cell tolerance. Proc. Natl. Acad. Sci. USA. 99:351–358.
Hawiger, D., K. Inaba, Y. Dorsett, M. Guo, K. Mahnke, M. Rivera, J.V. Ravetch, R.M. Steinman, and M.C. Nussenzweig. 2001. Dendritic cells induce peripheral T cell unresponsiveness under steady state conditions in vivo. J. Exp. Med. 194:769–779.
Caton, M.L., M.R. Smith-Raska, and B. Reizis. 2007. Notch–RBP-J signaling controls the homeostasis of CD8– dendritic cells in the spleen. J. Exp. Med. 204:1653–1664.
Voehringer, D., H.E. Liang, and R.M. Locksley. 2008. Homeostasis and effector function of lymphopenia-induced "memory-like" T cells in constitutively T cell-depleted mice. J. Immunol. 180:4742–4753.
Taieb, J., N. Chaput, C. Menard, L. Apetoh, E. Ullrich, M. Bonmort, M. Pequignot, N. Casares, M. Terme, C. Flament, et al. 2006. A novel dendritic cell subset involved in tumor immunosurveillance. Nat. Med. 12:214–219.[CrossRef][Medline]
Chan, C.W., E. Crafton, H.N. Fan, J. Flook, K. Yoshimura, M. Skarica, D. Brockstedt, T.W. Dubensky, M.F. Stins, L.L. Lanier, et al. 2006. Interferon-producing killer dendritic cells provide a link between innate and adaptive immunity. Nat. Med. 12:207–213.[CrossRef][Medline]
Kastenmuller, W., G. Gasteiger, J.H. Gronau, R. Baier, R. Ljapoci, D.H. Busch, and I. Drexler. 2007. Cross-competition of CD8+ T cells shapes the immunodominance hierarchy during boost vaccination. J. Exp. Med. 204:2187–2198.
Ramsdell, F., M. Jenkins, Q. Dinh, and B.J. Fowlkes. 1991. The majority of CD4+8- thymocytes are functionally immature. J. Immunol. 147:1779–1785.[Abstract]
Sprent, J., and H. Kishimoto. 2002. The thymus and negative selection. Immunol. Rev. 185:126–135.[CrossRef][Medline]
Gallegos, A.M., and M.J. Bevan. 2006. Central tolerance: good but imperfect. Immunol. Rev. 209:290–296.[CrossRef][Medline]
van Meerwijk, J.P., S. Marguerat, R.K. Lees, R.N. Germain, B.J. Fowlkes, and H.R. MacDonald. 1997. Quantitative impact of thymic clonal deletion on the T cell repertoire. J. Exp. Med. 185:377–383.
Sprent, J., H. Kosaka, and E.K. Gao. 1992. T cell tolerance after bone marrow transplantation in mice. Bone Marrow Transplant. 10:5–9.[Medline]
Moore, N.C., G. Anderson, D.E. McLoughlin, J.J. Owen, and E.J. Jenkinson. 1994. Differential expression of Mtv loci in MHC class II-positive thymic stromal cells. J. Immunol. 152:4826–4831.[Abstract]
Aschenbrenner, K., L.M. D'Cruz, E.H. Vollmann, M. Hinterberger, J. Emmerich, L.K. Swee, A. Rolink, and L. Klein. 2007. Selection of Foxp3+ regulatory T cells specific for self antigen expressed and presented by Aire+ medullary thymic epithelial cells. Nat. Immunol. 8:351–358.[CrossRef][Medline]
Ribot, J., G. Enault, S. Pilipenko, A. Huchenq, M. Calise, D. Hudrisier, P. Romagnoli, and J.P. van Meerwijk. 2007. Shaping of the autoreactive regulatory T cell repertoire by thymic cortical positive selection. J. Immunol. 179:6741–6748.
Bettelli, E., M. Oukka, and V.K. Kuchroo. 2007. T(H)-17 cells in the circle of immunity and autoimmunity. Nat. Immunol. 8:345–350.[CrossRef][Medline]
Smith, E., A. Zarbock, M.A. Stark, T.L. Burcin, A.C. Bruce, P. Foley, and K. Ley. 2007. IL-23 is required for neutrophil homeostasis in normal and neutrophilic mice. J. Immunol. 179:8274–8279.
Bennett, C.L., and B.E. Clausen. 2007. DC ablation in mice: promises, pitfalls, and challenges. Trends Immunol. 28:519–525.[CrossRef][Medline]
Brockschnieder, D., Y. Pechmann, E. Sonnenberg-Riethmacher, and D. Riethmacher. 2006. An improved mouse line for Cre-induced cell ablation due to diphtheria toxin A, expressed from the Rosa26 locus. Genesis. 44:322–327.[CrossRef][Medline]
Birnberg, T., L. Bar-On, A. Sapoznikov, M.L. Caton, L. Cervantes-Barragan, D. Makia, R. Krauthgamer, O. Brenner, B. Ludewig, D. Brockschnieder, et al. 2008. Lack of conventional dendritic cells is compatible with normal development and T cell homeostasis, but causes myeloid proliferative syndrome. Immunity. 29:986–997.[CrossRef][Medline]
de Heer, H.J., H. Hammad, T. Soullie, D. Hijdra, N. Vos, M.A. Willart, H.C. Hoogsteden, and B.N. Lambrecht. 2004. Essential role of lung plasmacytoid dendritic cells in preventing asthmatic reactions to harmless inhaled antigen. J. Exp. Med. 200:89–98.
Ochando, J.C., C. Homma, Y. Yang, A. Hidalgo, A. Garin, F. Tacke, V. Angeli, Y. Li, P. Boros, Y. Ding, et al. 2006. Alloantigen-presenting plasmacytoid dendritic cells mediate tolerance to vascularized grafts. Nat. Immunol. 7:652–662.[CrossRef][Medline]
Hadeiba, H., T. Sato, A. Habtezion, C. Oderup, J. Pan, and E.C. Butcher. 2008. CCR9 expression defines tolerogenic plasmacytoid dendritic cells able to suppress acute graft-versus-host disease. Nat. Immunol. 9:1253–1260.[CrossRef][Medline]
Merad, M., F. Ginhoux, and M. Collin. 2008. Origin, homeostasis and function of Langerhans cells and other langerin-expressing dendritic cells. Nat. Rev. Immunol. 8:935–947.[CrossRef][Medline]
Stranges, P.B., J. Watson, C.J. Cooper, C.M. Choisy-Rossi, A.C. Stonebraker, R.A. Beighton, H. Hartig, J.P. Sundberg, S. Servick, G. Kaufmann, et al. 2007. Elimination of antigen-presenting cells and autoreactive T cells by Fas contributes to prevention of autoimmunity. Immunity. 26:629–641.[CrossRef][Medline]
Chen, M., Y.H. Wang, Y. Wang, L. Huang, H. Sandoval, Y.J. Liu, and J. Wang. 2006. Dendritic cell apoptosis in the maintenance of immune tolerance. Science. 311:1160–1164.
Nopora, A., and T. Brocker. 2002. Bcl-2 controls dendritic cell longevity in vivo. J. Immunol. 169:3006–3014.
Laufer, T.M., L. Fan, and L.H. Glimcher. 1999. Self-reactive T cells selected on thymic cortical epithelium are polyclonal and are pathogenic in vivo. J. Immunol. 162:5078–5084.
Luckashenak, N., S. Schroeder, K. Endt, D. Schmidt, K. Mahnke, M.F. Bachmann, P. Marconi, C.A. Deeg, and T. Brocker. 2008. Constitutive crosspresentation of tissue antigens by dendritic cells controls CD8+ T cell tolerance in vivo. Immunity. 28:521–532.[CrossRef][Medline]
Li, X.C., W. Almawi, A. Jevnikar, J. Tucker, R. Zhong, and D. Grant. 1995. Allogeneic lymphocyte proliferation stimulated by small intestine-derived epithelial cells. Transplantation. 60:82–89.[CrossRef][Medline]
Westendorf, A.M., M. Templin, R. Geffers, S. Deppenmeier, A.D. Gruber, M. Probst-Kepper, W. Hansen, R.S. Liblau, F. Gunzer, D. Bruder, and J. Buer. 2005. CD4+ T cell mediated intestinal immunity: chronic inflammation versus immune regulation. Gut. 54:60–69.
Marguerat, S., H.R. MacDonald, J.P. Kraehenbuhl, and J.P. van Meerwijk. 1999. Protection from radiation-induced colitis requires MHC class II antigen expression by cells of hemopoietic origin. J. Immunol. 163:4033–4040.
Shimoda, M., F. Mmanywa, S.K. Joshi, T. Li, K. Miyake, J. Pihkala, J.A. Abbas, and P.A. Koni. 2006. Conditional ablation of MHC-II suggests an indirect role for MHC-II in regulatory CD4 T cell maintenance. J. Immunol. 176:6503–6511.
Kim, J.M., J.P. Rasmussen, and A.Y. Rudensky. 2007. Regulatory T cells prevent catastrophic autoimmunity throughout the lifespan of mice. Nat. Immunol. 8:191–197.[CrossRef][Medline]
Weston, S.A., and C.R. Parish. 1990. New fluorescent dyes for lymphocyte migration studies. Analysis by flow cytometry and fluorescence microscopy. J. Immunol. Methods. 133:87–97.[CrossRef][Medline]
Larsen, C.P., R.M. Steinman, M. Witmer-Pack, D.F. Hankins, P.J. Morris, and J.M. Austyn. 1990. Migration and maturation of Langerhans cells in skin transplants and explants. J. Exp. Med. 172:1483–1493.
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Facebook 
Reddit 
Technorati 
Twitter What's this?
Related Article
This article has been cited by other articles:
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| TABLE OF CONTENTS |
|
