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Mature dendritic cells (DCs) are crucial for the induction of adaptive immune responses and perturbed DC homeostasis can result in autoimmune disease. Either uncontrolled expansion1 or enhanced survival2,3 of DCs can result in a variety of autoimmune diseases in mouse models. In addition, increased maturation signals, through overexpression of surface Toll-like receptors (TLRs)4 or stimulation by type I interferon (IFN)5, has been associated with systemic autoimmunity. Whereas recent studies have focused on identifying factors required for initiating the maturation process, the possibility that resting DCs also express molecules that ‘hold’ them in an immature state has generally not been considered. Here we show that nuclear factor– kB1 (NF-kB1) is crucial for maintaining the resting state of DCs. Self-antigen–pulsed unstimulated DCs that do not express NF-kB1 were able to activate CD8+ T lymphocytes and induce autoimmunity. We further show that NF-kB1 negatively regulates the spontaneous production of tumor necrosis factor-a (TNF-a), which is associated with increased granzyme B expression in cytotoxic T lymphocytes (CTLs). These findings provide a new perspective on functional DC maturation and a potential mechanism that may account for pathologic T cell activation.

It is presently unclear whether DCs are tolerogenic by default or whether intrinsic factors are required to maintain the resting state and prevent induction of autoimmunity. The role of NF-κB1 is of particular interest, as it may both positively6–8 and negatively9,10 regulate gene transcription. Here we present a system for evaluat-ing the contribution of defined genes or pathways to DC function, where the induction of CD8+ T cell–mediated autoimmune responses is dependent upon self-antigen presentation by DCs that have been matured by TLR stimulation. Notably, NF-κB1–deficient DCs do not require TLR-induced maturation to initiate self-reactive CTL activity. The absence of NF-κB1 results in dysregulated production of TNF-α,

which affects the intracellular abundance of the serine protease granzyme B in CD8+ T cells and has an impact on the induction of diabetes. These findings represent a major shift in our understanding of DC biology, as they are the first indication that resting DCs possess intrinsic factors that inhibit the activation of the adaptive immune system. This work demonstrates that proinflammatory functions of DCs, including TNF-α production, are regulated even in the absence of external stimulation and that the removal of these repressive mech-anisms can result in the activation of autoreactive T lymphocytes.

We have developed a model based on transgenic mice (RIP-gp) expressing the lymphocytic choriomeningitis virus (LCMV) glycopro-tein (gp) on pancreatic islet beta cells under the control of the rat insulin promoter (RIP). Previous work has shown infection of RIP-gp mice with LCMV resulted in activation of gp-specific T lymphocytes and destruc-tion of beta cells11. In this study, we describe a new method to induce diabetes by transferring bone marrow–derived DCs (BMDCs) pulsed with gp-derived epitopes into RIP-gp mice. The induction of diabetes was dependent upon DC maturation, as wild-type (WT) gp peptide–pulsed DCs must be stimulated with CpG oligodeoxynucleotides to initiate autoimmune diabetes (Fig. 1a,b). Furthermore, the induction of diabetes was antigen specific, as non–peptide-pulsed mature DCs were unable to induce diabetes (Supplementary Fig. 1a). Additionally, this response depended on CD8+ T cell activation, as depletion of CD8+ cells and not depletion of CD4+ cells before DC transfer completely prevented diabetes induction (Supplementary Fig. 1b). Histological examination of pancreas samples showed that mature, peptide-pulsed DCs promoted increased CD8+ islet infiltration (Fig. 1c,d). Therefore, the induction of autoimmunity was dependent on the transfer of mature, gp peptide–pulsed DCs, which led to the activation of tissue-specific CTLs.

To evaluate the role of NF-κB1, we generated BMDCs from Nfkb1−/− mice. Flow cytometric analysis of surface markers confirmed that con-ventional DCs could be generated in vitro in the absence of NF-κB1 (Supplementary Fig. 2). The transfer of TLR-stimulated NF-κB1–deficient DCs resulted in the induction of diabetes in RIP-gp mice, indicating that NF-κB1 expression is not required for CTL activation

Nuclear factor-κB1 controls the functional maturation of dendritic cells and prevents the activation of autoreactive T cellsDilan Dissanayake1,2, Håkan Hall1, Nancy Berg-Brown1, Alisha R Elford1, Sara R Hamilton1, Kiichi Murakami1, Leslie Summers Deluca2, Jennifer L Gommerman2 & Pamela S Ohashi1–3

1Campbell Family Institute for Breast Cancer Research, Ontario Cancer Institute, Toronto, Ontario, Canada. 2Department of Immunology, University of Toronto, Toronto, Ontario, Canada. 3Department of Medical Biophysics, University of Toronto, Toronto, Ontario, Canada. Corresposndence should be addressed to P.S.O. (pohashi@uhnresearch.ca).

Received 25 July; accepted 12 October; published online 13 November 2011; doi:10.1038/nm.2556

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(Supplementary Fig. 3). Notably, unlike WT DCs, NF-κB1–deficient DCs could also trigger diabetes in the absence of TLR stimulation (Fig. 2a,b). The induction of diabetes was associated with increased in vivo CTL activity in mice given peptide-pulsed unstimulated NF-κB1-deficient or CpG-matured WT DCs (Fig. 2c). Furthermore, histological analysis revealed an increased degree of CD8+ infiltrate in islets of mice receiving unstimulated NF-κB1-deficient DCs compared with those receiving unstimulated WT DCs (Fig. 2d,e). Therefore, NF-κB1–deficient DCs do not require maturation stimuli to promote robust autoimmune responses in vivo.

Given that prolonged antigen presentation may be associated with T cell activation2,3, we examined whether DC survival was enhanced in the absence of NF-κB1. We found no significant difference in viability between WT and NF-κB1–deficient DCs after granulocyte-monocyte

colony–stimulating factor (GM-CSF) withdrawal in vitro (Supplementary Fig. 4a). To evaluate the duration of antigen presentation in vivo, we trans-ferred peptide-pulsed DCs to mice, followed by carboxyfluorescein suc-cinimidyl ester (CFSE)-labeled gp-specific T cells at different time points. The proportion of T cells that proliferated decreased with time after DC transfer, with little proliferation taking place when T cells were trans-ferred 72 h after either WT or NF-κB1–deficient DCs (Supplementary Fig. 4b). Together, these results suggest that the induction of diabetes by unstimulated NF-κB1–deficient DCs was not a function of prolonged antigen presentation.

To confirm that these findings were not limited to BMDCs, we assessed the ability of T cells to become tolerized in NF-κB1–deficient hosts. Previous work has shown that repeated administration of peptide in the absence of maturation stimuli results in the expansion

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Figure 1 TLR-stimulated, antigen-pulsed DCs induce diabetes in RIP-gp mice. (a–d) DCs were cultured overnight with or without CpG ODN 1826 (10 µM) as indicated before being pulsed with peptides and transferred to RIP-gp mice. (a) Blood glucose concentration after DC transfer. Each line represents an individual mouse with five representative mice depicted per group. (b) Diabetes incidence after transfer of unstimulated (squares) and CpG-stimulated (triangles) DCs for 15–20 mice per group. *P < 0.0001. (c) Pancreas sections from RIP-gp mice 6 d after receiving unstimulated or CpG-stimulated DCs, stained with antibody to CD8. (d) Quantification of the degree of CD8+ T cell infiltration in the pancreas (n = 3 mice per group with 30 or more islets examined per mouse). All results are representative of more than three independent experiments.

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unstimulated WT or NF-κB1–deficient DCs, stained with antibody to CD8. (e) Quantification of the degree of CD8+ T cell pancreatic infiltration seen by histology (n = 3 mice per group with 30 or more islets examined per mouse). (f) Proportion of Thy1.1+CD8+ P14 T cells in the blood of WT and Nfkb1−/− hosts that were infused with 5 µg gp33-41 peptide on days 0, 3 and 6 (shown by arrows). (g) The proportion and absolute number of Thy1.1+ cells in the spleens of WT and Nfkb1−/− mice on day 8 of treatment as in f. *P < 0.05. **P < 0.02. (h) CFSE dilution of Thy1.1+CD8+ splenocytes collected from WT (red) and Nfkb1−/− (blue) mice 8 d after the first peptide infusion and re-stimulated with peptide ex vivo. (i) Quantification of divided cells as described in h cultured with or without 10 U ml−1 IL-2. *P < 0.05. NS, not significant. Results are representative of three independent experiments. Error bars indicate mean ± s.d.

Figure 2 NF-κB1–deficient DCs do not require TLR maturation to induce diabetes. (a–e) BMDCs were generated from WT or Nfkb1−/− mice as indicated, pulsed with peptides and transferred to RIP-gp mice. (a) Blood glucose concentration after DC transfer. Each line represents an individual mouse with five representative mice depicted per group. (b) Diabetes incidence after transfer of unstimulated WT (squares) and unstimulated Nfkb1−/− (triangles) DCs for more than ten mice per group. *P < 0.001. (c) Quantification of gp33-41–specific in vivo CTL activity 8 d after transfer of peptide-pulsed DCs. *P = 0.0001. (d) RIP-gp pancreas sections from mice 6 d after receiving

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of antigen-specific T cells, followed by their deletion12,13. The remain-ing T cells are anergic, as indicated by their limited proliferative capac-ity and responsiveness to the addition of interleukin-2 (IL-2)12,13. We therefore transferred Thy1.1+ gp33–41–specific T cells to WT or NF-κB1–deficient hosts, followed by repeated infusions of peptide, and measured the expansion of the transferred cells. In the blood, we found a higher proportion of Thy1.1+ CD8+ cells in Nfkb1−/− hosts compared to WT mice (Fig. 2f). Additionally, 8 d after transfer, the spleens of Nfkb1−/− mice had a significantly higher proportion and number of Thy1.1+ cells, compared to WT mice (Fig. 2g), which is consistent with hyperactivity of resting NF-κB1–deficient antigen-presenting cells. To assess tolerance of the transferred cells, we iso-lated CD8+ splenocytes from WT and Nfkb1−/− mice 2 d after the last peptide transfer and re-stimulated them with peptide ex vivo. T cells that had encountered antigen on WT antigen-presenting cells had a lower proliferative capacity compared to T cells isolated from NF-κB1–deficient hosts (Fig. 2h). Furthermore, the proliferative defect in T cells that had been transferred to WT mice could be rescued by adding IL-2, suggesting anergy was induced in T cells transferred to WT mice but not Nfkb1−/− mice (Fig. 2i). Taken together, these results suggest that the absence of NF-κB1 in resting antigen-presenting cells is associated with impaired induction of T cell tolerance.

Spontaneous autoimmunity has not been described in previous stud-ies using NF-κB1–deficient mice14. Accordingly, Nfkb1−/− RIP-gp mice did not develop spontaneous hyperglycemia (data not shown). It is possible that the lack of diabetes in Nfkb1−/− RIP-gp mice is due to an inability to expand self-reactive T cells, as previous work has shown defective T cell proliferation in the absence of NF-κB1 (ref. 7). To increase the frequency of self-reactive T cells, we bred Nfkb1−/− RIP-gp mice that possess the P14 gp-specific T cell receptor transgene. It has previously been shown that, despite the high frequency of islet-reactive T cells, spontaneous diabetes does not occur in P14-expressing RIP-gp mice11. However, we found that approximately 55% of NF-κB1– deficient P14-expressing RIP-gp mice developed spontaneous hyperglycemia starting at 7 weeks of age (P < 0.0001) (Supplementary Fig. 5). This demonstrates that in the presence of self-reactive T cells, NF-κB1 has a role in preventing spontaneous autoimmunity.

To understand how NF-κB1 was influencing DC function, we examined the expression of various markers associated with DC maturation. Unexpectedly, expression of major histocompatibility complex (MHC) classes I and II, CD40, CD80 and CD86 were not elevated in NF-κB1–deficient DCs (Fig. 3a). In fact, MHC class II and CD80 expression were significantly lower in NF-κB1–deficient DCs (Fig. 3a), perhaps reflecting the requirement for NF-κB1 to maintain expression of these molecules, as has been suggested previously15. We next examined secretion of proinflammatory cytokines.

Unstimulated NF-κB1–deficient DCs produced higher amounts of TNF-α and IL-6 but not IL-1, IL-12, IFN-α or IFN-β than unstimu-lated WT DCs (Fig. 3b). There was also no detectable production of IL-10 by either unstimulated WT or unstimulated NF-κB1–deficient DCs (Fig. 3b). Furthermore, using DCs deficient for both NF-κB1 and TNF-α, we found that spontaneous IL-6 production was dependent on TNF-α production (data not shown), possibly through feedback pathways that have previously been described16. These data support previous observations that upregulation of co-stimulatory molecules and secretion of IL-12 by DCs does not necessarily correlate with the ability to effectively initiate CTL responses17.

To test for the specific requirement of TNF-α in the induction of diabetes, we generated Nfkb1−/−;Tnf−/− mice. Unlike Nfkb1−/− DCs, Nfkb1−/−;Tnf−/− DCs were unable to induce diabetes (Fig. 4a,b). Similarly, whereas CpG-stimulated WT DCs induced diabetes upon transfer to RIP-GP mice, CpG-stimulated TNF-α–deficient DCs were unable to induce diabetes (Supplementary Fig. 6a). Conversely, CpG-stimulated IL-6–deficient DCs and Pycard−/− DCs, which are defective in IL-1 production18, were capable of inducing diabetes, suggesting that IL-6 and IL-1 are dispensable for the induction of diabetes (Supplementary Fig. 6a). WT DCs that were cultured in medium containing recombinant TNF-α were unable to promote diabetes, indicating that the paracrine effects of TNF-α on DCs was not sufficient for autoimmunity (Supplementary Fig. 6b). To fur-ther confirm that TNF-α was essential upon transfer to the host, we generated TNFR1- or TNFR2-deficient RIP-gp mice and found that unstimulated NF-κB1–deficient DCs were unable to induce diabetes in these mice (Fig. 4c,d). Therefore, the spontaneous production of TNF-α by NF-κB1–deficient DCs was crucial for the induction of CD8+ T cell mediated autoimmunity.

Although TNF-α is a well-characterized cytokine that is associ-ated with a number of autoimmune diseases19–21, it is unclear which aspects of CD8+ T cell activation are affected by its overproduction. Following transfer of unstimulated Nfkb1−/−;Tnf−/− DCs, we found that neither the proportion of antigen-specific T cells (Fig. 4e) nor antigen-specific production of IFN-γ was decreased (Fig. 4f), suggesting that DC-derived TNF-α does not contribute to these aspects of T cell activation. Furthermore, pancreatic infiltration by gp-specific CD8+ T cells was not impaired upon transfer of Nfkb1−/−;Tnf−/− DCs (Fig. 4g). However, pancreas-infiltrating lym-phocytes from RIP-gp mice that had received Nfkb1−/−;Tnf−/− DCs showed markedly lower amounts of granzyme B (Fig. 4h), which is a key mediator of beta cell destruction in diabetes22. We confirmed these results using in vitro co-culture of antigen-specific T cells and peptide-pulsed DCs. Cultures of CD8+ T cells with unstimu-lated NF-κB1–deficient DCs or CpG-stimulated WT DCs resulted

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in higher intracellular granzyme B expression in the T cells com-pared with cocultures containing unstimulated WT DCs (Fig. 4i). Furthermore, the addition of TNF-α to cocultures with unstimu-lated WT DCs resulted in increased CD8+ T cell granzyme B expres-sion (Fig. 4i). Collectively, these studies show that the absence of NF-κB1 leads to the production of TNF-α by unstimulated DCs, which can have an impact on CTL effector function by promoting granzyme B upregulation.

Previous studies assessing the contribution of a specific factor to autoimmune disease have typically used mice that lack the molecule in all tissues, which could potentially obscure varying functions that the molecule has in different cell types. For example, NF-κB1– deficient mice are protected from experimental autoimmune encepha-litis because of their impaired T cell proliferation and differentiation7, whereas studies with macrophages in the setting of lipopolysaccha-ride tolerance have revealed a regulatory role for NF-κB1 (ref. 9). Additionally, expression of NF-κB in pancreatic beta cells can influence their susceptibility to apoptosis, potentially confounding diabetes studies in knockout mice23. The model presented here allows for the study of molecules specifically within DCs for their contribu-tion during interactions with autoreactive T cells.

The occurrence of autoimmunity has previously been linked to NF-κB hyperactivity24–28. It is worthy to note, however, that these studies typically examined NF-κB activity as a whole, often after mat-uration by an external ligand such as lipopolysaccharide. Although NF-κB1 p50 heterodimerization with p65 has a proinflammatory role upon TLR stimulation, p50 is found in transcriptionally repres-sive homodimers in many unstimulated cells, including DCs29–31. The absence of NF-κB1 may therefore have different functional

consequences in stimulated and unstimulated cells. Correspondingly, in addition to heightened NF-κB responses after stimulation, nonobese diabetic (NOD) mice show markedly lower nuclear abun-dance of p50 homodimers in unstimulated DCs compared with the diabetes-resistant NOR/Lt strain25. Bcl-3–deficient mice, which show defective maintenance of p50 homodimers, also showed increased susceptibility to diabetes in the NOD model10,32. These studies are consistent with a regulatory function for NF-κB1 p50 in limiting autoimmune diabetes.

In this report, we have demonstrated a role for NF-κB1 in negatively regulating TNF-α production in resting DCs and preventing the sub-sequent induction of CD8+ effector activity. Notably, human studies have identified a polymorphism in the TNF-α promoter region that results in reduced binding by p50 homodimers33. This polymorphism is associated with a variety of autoimmune diseases, including type 1 diabetes34–36. Our work describes a mechanism that may explain the increased incidence of autoimmunity in this population.

Recent studies have shown that transcriptional regulators may actively maintain quiescence in lymphocytes37,38. The work presented here indicates that, in addition to previously identified extrin-sic mechanisms of DC maturation via pattern recognition receptor ligation, factors within DCs exist to limit CTL induction in the steady state. Further research into such DC-intrinsic regulatory factors may provide key insights into the initiation of autoimmune pathology by aberrant DC activity.

METHoDSMethods and any associated references are available in the online version of the paper at http://www.nature.com/naturemedicine/.

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Figure 4 The induction of diabetes in RIP-gp mice is dependent upon TNF-α production by transferred DCs. (a–d) BMDCs were generated from Nfkb1−/− (a) and Nfkb1−/−;Tnf−/− (b) mice, pulsed with peptides and transferred to RIP-gp mice or BMDCs were generated from Nfkb1−/− mice and pulsed with peptides before transfer to TNFR1-deficient RIP-gp (c) or TNFR2-deficient RIP-gp mice (d). Blood glucose concentrations were followed after DC transfer. Each line represents an individual mouse with four representative mice depicted per group. (e–h) Unstimulated WT, unstimulated Nfkb1−/−, unstimulated Nfkb1−/− ;Tnf−/− or CpG-stimulated WT DCs were transferred to C57BL/6 mice. (e) Proportion of tetramer-positive (Tet+) CD8+ T cells in blood 8 d after DC transfer. (f) Quantification of IFN-γ–producing CD8+ splenocytes 8 d after DC transfer. (g) Number of tetramer-positive pancreas-infiltrating lymphocytes (PILs) 6 d after DC transfer to RIP-gp mice. (h) Representative histogram and median fluorescence intensity of granzyme B in PILs 6 d after transfer of unstimulated WT (red), unstimulated Nfkb1−/− (blue), unstimulated Nfkb1−/−;Tnf−/− (green) or CpG-stimulated WT (orange) DCs to RIP-gp mice. (i) Representative histogram and median fluorescence intensity of granzyme B in CD8-sorted P14 splenocytes cocultured with unstimulated WT (red), unstimulated Nfkb1−/− (blue), CpG-stimulated WT (orange) or WT DCs with 10 ng ml−1 exogenous TNF-α (green). Histograms are gated on CD8+CD44+ cells. *P < 0.05; **P < 0.001; ***P < 0.0001. Results are representative of three independent experiments. Error bars indicate mean ± s.d.

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Note: Supplementary information is available on the Nature Medicine website.

AcKNOwLEDGMENtSWe would like to thank S. Mariathasan (Genentech) for kindly providing Pycard-deficient mice. We would also like to thank L.T. Nguyen for critical evaluation of the manuscript. This work was supported by a Canadian Institutes for Health Research (CIHR) grant to P.S.O. and CIHR studentship to D.D. P.S.O. holds a Canada Research Chair in Autoimmunity and Tumor Immunity. This research was funded in part by the Ontario Ministry of Health and Long Term Care (OMHLTC). The views expressed do not necessarily reflect those of the OMHLTC.

AUtHOR cONtRIBUtIONSD.D. and P.S.O. designed and supervised the project, analyzed the data and wrote the manuscript. H.H., A.R.E., S.R.H. and K.M. assisted with experiments. N.B.-B., L.S.D. and J.L.G. performed supporting experiments.

cOMPEtING FINANcIAL INtEREStSThe authors declare no competing financial interests.

Published online at http://www.nature.com/naturemedicine/. Reprints and permissions information is available online at http://www.nature.com/reprints/index.html.

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nature medicine doi:10.1038/nm.2556

oNLINE METHoDSMice and blood glucose monitoring. We purchased C57BL/6 mice and gene targeted NF-κB1-deficient (Nfkb1−/−), TNF-α–deficient (Tnf−/−), TNFRI-deficient (Tnfrsf1a−/−), TNFRII-deficient (Tnfrsf1b−/−), IL-6–deficient (Il6−/−) and Thy1.1 congenic mice from the Jackson Laboratory (B6.Cg-Nfkb1tm1bal/J, B6.129S-Tnf tmGkl/J, B6.129-Tnfrsf1atm1Mak/J, B6.129S2-Tnfrsf1btm1Mwm/J, B6.129S2-Il6-tm1Kopf/J and B6.PL-Thy1a/CyJ, respectively). We obtained Pycard-deficient (Pycard−/−) mice from Genentech. Generation of RIP-GP and P14 mice was previously described11,39. We measured blood glucose con-centrations using Accu-chek III Glucometers and Chemstrips (Roche) and considered mice diabetic after two consecutive measurements of >15 mM. All mice were maintained and mouse experiments were performed at the Ontario Cancer Institute animal facility according to institutional guidelines and with approval of the Ontario Cancer Institute Animal Ethics Committee.

DC generation and transfer. BMDCs were generated as previously described40. Briefly, we collected bone marrow from mouse femurs and tibiae and cultured it in complete RPMI containing 40 ng ml−1 GM-CSF (Peprotech) with medium changes on days 3, 6 and 8 of culture. On day 10, we collected and cultured nonadherent DCs with CpG ODN 1826 (ACGT), lipopolysaccharide (Sigma) or TNF-α (Peprotech) for 16 h. For transfer, we pulsed DCs for 2 h with 1 µg ml−1 each of gp33–41 (KAVYNFATM), gp276–286 (SGVENPGGYCL) and gp61–80 (GLNGPDIYKGVYQFKSVEFD). We then washed cells three times in Hank’s Buffered Saline Solution (HBSS) and administered them intra-venously at 2 × 106 cells per mouse through tail vein injection. For CD8+ or CD4+ T cell depletion in RIP-GP mice, we injected YTS 169 or YTS 191 antibodies from in-house hybridoma cultures respectively 3 d and 1 d before DC transfer.

Flow cytometry and antibodies. We purchased monoclonal antibodies specific for CD11c, B220, MHC class II, CD80 and CD86 from eBioscience, and CD11b, CD8, CD4, MHC class I and CD40 from BD Pharmingen. CFSE was purchased from Molecular Probes. We performed intracellular staining by incubating cells with GolgiPlug for 6 h followed by staining using Cytofix/Cytoperm (BD Pharmingen) and monoclonal antibody specific for IFN-γ purchased from eBioscience or granzyme B purchased from Invitrogen. We conducted tetramer staining using streptavidin (Invitrogen) and monomers for H-2Db:KAVYNFATM and H-2Kb:AVYNFATC. We acquired flow cytometry data on a FACSCalibur (BD) and analyzed using FlowJo software (Tree Star).

In vivo cytotoxic T lymphocyte assay. We vaccinated C57BL/6 mice with 2 × 106 peptide-pulsed DCs by tail vein injection as described above. Eight days later, we pulsed C57BL/6 splenocytes with gp33–41 peptide or a control adenovirus-derived (AV) peptide (SGPSNTPPEI) and labeled with CFSE (Molecular Probes) at 10 µM and 1 µM, respectively, for 10 min at 37 °C. After two washes, we administered the target cells intravenously via tail vein injection to the DC-vaccinated mice at 1.5 ×1 07 target cells of each peptide specificity per mouse. We also infused naive mice with targets to assess ratio of gp33-41 pulsed and AV-pulsed targets. We collected splenocytes 4 h later and detected CFSE-labeled cells by flow cytometry on a FACSCalibur (BD). We calculated specific lysis as (percentage AV – percentage gp33–41) / percentage AV × 100 and normalized on the basis of values seen in naive controls.

Immunohistochemistry and antibodies. We used liquid nitrogen to snap-freeze pancreata in optimal cutting temperature medium and

prepared sections for CD8 staining with primary rat mAb (YTS 169). We performed quantification using light microscopy.

Dendritic cell persistence experiments. For assessment of in vitro viability, we generated BMDCs as described above and cultured them in 24-well plates in complete RPMI without GM-CSF. We isolated cells 24, 48 and 72 h after start of culture and stained them with annexin V (BD Pharmingen) according to the manufacturer’s instructions.

For assessment of DC-antigen persistence in vivo, we generated BMDCs and transferred them intravenously to C57BL/6 mice at 2 × 106 cells per mouse. One, two or three days after BMDC transfer, we CD8-enriched splenocytes from Thy1.1+ P14 mice using CD8a+ T cell Isolation Kit II (Miltenyi Biotec) and labeled them with CFSE before intravenous infusion. Three days after each T cell transfer, we recovered splenocytes and conducted flow cytometry to determine dilution of CFSE-labeled CD8+Thy1.1+ cells. We also transferred T cells to naive mice on each day to determine fluorescence intensity of undivided cells.

Cytokine detection. We collected supernatants from DCs cultured for 16 h at 2 × 106 per ml. We used OptEIA ELISA kits from BD Biosciences to measure TNF-α, IL-6, IL-12, IL-1β and IL-10 concentrations in supernatants follow-ing the manufacturer’s instructions. We used Verikine ELISA kits purchased from PBL InterferonSource to measure IFN-α and IFN-β concentrations in supernatants following the manufacturer’s instructions.

Granzyme B expression. We perfused RIP-gp mice using PBS with 75 U ml−1 heparin (Sigma). We then removed pancreases and digested them using RPMI containing collagenase D and DNase I (Roche). We purified lymphocytes from the digest using Percoll (Amersham Biosciences) before intracellular staining for granzyme B.

For in vitro coculture experiments, we enriched CD8+ splenocytes by mag-netic separation using a CD8a+ T cell Isolation Kit II (Miltenyi Biotec). We cocultured purified CD8+ cells with BMDCs at a 5:1 ratio for 48 h before intra-cellular staining for granzyme B.

In vivo tolerance experiment. We collected and negatively sorted Thy1.1+ P14 splenocytes to enrich for CD8+ cells using magnetic separation using a CD8a+ T cell Isolation Kit II (Miltenyi Biotec). We transferred 3 × 106 purified cells intravenously to C57BL/6 mice on day –1. On days 0, 3 and 6, mice received 5 µg of gp33–41 peptide. On days 2, 4, 6 and 8, we drew blood from the tail vein, and on day 8 we removed spleens to assess Thy1.1+ population by flow cytometry. We labeled day-8 splenocytes with CFSE and cocultured them in vitro with gp33–41–pulsed C57BL/6 stimulator cells, with or without 10 U ml−1 IL-2 (Novartis) for 3 d before flow cytometry for CFSE dilution.

Statistical analyses. We analyzed differences in diabetes incidence by log-rank test. We analyzed differences in in vivo CTL activity, cell numbers and supernatant cytokine measurements by unpaired t tests. We compared median fluorescence intensities of granzyme B expression in independent experiments using paired t tests.

39. Pircher, H., Burki, K., Lang, R., Hengartner, H. & Zinkernagel, R.M. Tolerance induction in double specific T cell receptor transgenic mice varies with antigen. Nature 342, 559–561 (1989).

40. Lutz, M.B. et al. An advanced culture method for generating large quantities of highly pure dendritic cells from mouse bone marrow. J. Immunol. Methods 223, 77–92 (1999).

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