NOD/ShiLtJ (NOD), NOD.Cg-Rag1^tm1Mom (NOD-Rag1^−/−), NOD.Cg-Rag1^tm1MomTg (TcraAI4)^1Dvs/DvsJ (NOD.AI4α-Rag1^−/−), and NOD.Cg-Rag1^tm1MomTg (TcrbAI4)^1Dvs/DvsJ (NOD.AI4β-Rag1^−/−) mice were purchased from The Jackson Laboratory (Bar Harbor, ME). F1 hybrid progeny developed from outcrosses of NOD.AI4α-Rag1^−/− to NOD.AI4β-Rag1^−/− (hereafter referred to as NOD.AI4-Rag1^−/−) developed diabetes between 3 and 5 weeks of age. NADPH oxidase 2 deficient NOD-Ncf1^m1J mice were generated as previously described (23, 24). NOD.Ncf1^m1J.Cg-Rag1^tm1Mom (NOD-Ncf1-Rag1^−/−) mice were generated through F2 mating of NOD-Rag1^−/− to NOD-Ncf1^m1J. Mice were genotyped as described (25) to ensure homozygosity of Rag1^−/− as well as D5Mit30 and D5Mit31 to test for inheritance of the Ncf1^m1J allele. Mice at the F2 generation that were homozygous for the targeted deletion of Rag1 and the mutant allele of Ncf1 were used as founders for this mouse strain. Female mice were used for all experiments. All mice used in this study were housed in specific pathogen free facilities, and all studies herein were approved by the institutional animal care and use committee at the University of Florida.

Fluorescently labeled antibodies including: Phycoerythrin (PE)-labeled α-CXCR4 (2B11), Brilliant violet 421-labeled α-CD8 (53-6.7), allophycocyanin (APC)-labeled α-CD3ε (BM8), and APC-labeled α-T-bet (4B10), PE-labeled α-granzyme B (NGZB), PE-labeled α-interferon gamma (IFNγ) (XMG1.2), APC labeled α-TNFα (MP6-XT22) [eBioscience (San Diego, CA)] as well as PE-labeled α-H2K^d [Biolegend (San Diego, CA)] were used. Recombinant mouse granulocyte-macrophage colony stimulating factor (rmGM-CSF) and rmIL-4 were purchased from R&D systems (Minneapolis, MN). Pam3CysSerLys4 (Pam3CSK4) and Polyinosinic-polycytidylic acid (Poly(I:C)) were purchased from Invivogen (San Diego, CA). Lipopolysachharide (LPS) was purchased from Sigma (St. Louis, MO). Polybead amino 3.0 μm microspheres were purchased from Polysciences (Warrington, PA). Horse cytochrome c was purchased from Sigma. Alexa Fluor 647 (AF647) and DQ ovalbumin (DQ-OVA) were purchased from Life technologies (Grand Island, NY). Fluorescein isothiocyanate (FITC) conjugated ovalbumin (OVA) was purchase from Sigma.

Mouse spleens or lymph nodes were collected, homogenized to a single cell suspension, and subjected to hemolysis with Gey's solution. Negative selection of CD8⁺ T cells from was performed using magnetic beads [mouse CD8⁺ T cell isolation kit (Miltenyi Biotec)], according to the manufacturer's protocol. CD4⁺ T cells from NOD as well as CD8⁺ T cells from NOD and NOD-Ncf1^m1J were purified by negative selection with magnetic beads according to the manufacturer's protocol using a CD4⁺ T cell isolation kit or a CD8⁺ T cell isolation kit (Miltenyi Biotec), respectively. Purity, >96%, was confirmed by flow cytometric analysis on a BD LSR Fortessa.

Pre-diabetic (8 weeks old) NOD and NOD-Ncf1^m1J T cell donors were used for adoptive transfer experiments. Splenocytes were purified as described above. CD4⁺ and CD8⁺ T cells were mixed at a ratio of 3:1 and 10⁷ cells were transferred intraperitoneally (i.p.) to 8 week old NOD-Rag1^−/− or 8 week old NOD-Ncf1^m1J-Rag1^−/− recipients. Transfers were divided into 4 groups: NOD-Rag1^−/- mice received either (1) NOD CD4⁺ + NOD CD8⁺ or (2) NOD CD4⁺ + NOD-Ncf1^m1J CD8⁺, while the remaining two groups were NOD-Ncf1^m1J-Rag1^−/− recipients of (3) NOD CD4⁺ + NOD CD8⁺ or (4) NOD CD4⁺ + NOD-Ncf1^m1J CD8⁺. Mice were monitored weekly for diabetes onset as described previously (23). Engraftment of cells was confirmed by flow cytometry.

Bone marrow derived DCs (BMDCs) were generated by 8 days of culture in complete RPMI 1,640 media with 10% FBS (26). The culture media was supplemented with 1,000 U/mL rmGM-CSF and 500 U/mL rmIL4. Maturation was induced by 24-h treatment with 100 ng/mL Pam3CSK4, 25 μg/mL poly (I: C), 100ng/mL LPS, 1ug/mL R848, or 5 μg/mL CpG2336 respectively.

Real time quantitative PCR was performed as previously reported (27–31). In general, total RNA from DCs was isolated with TRIzol (Invitrogen, Carlsbad, CA) and cDNA was prepared using the Superscript III First-Strand Synthesis System (Invitrogen) according to the manufacturer's protocol. SYBR Green I (Bio-Rad) analysis was performed on a LightCycler 480 II (Roche, Basel, Switzerland). The amplification program utilized the following steps for all primer sets: 95°C for 10 min, then 45 cycles of 95, 60, and 72°C for 30 s. Melting-curves were performed for each PCR reaction to ensure specificity. Primers were used according to qPrimerDepot (NIH) and previous reports and listed as follows (28, 32):

Flow cytometry was performed to detect the surface proteins on and phagocytosis by BMDC. BMDC were counted and re-suspended in PBS at 2 × 10⁷ cells/mL. Approximately 10⁶ cells were labeled with antibodies at the proper dilution. Fluorescence was measured using a LSR Fortessa (BD Bioscience, San Jose, CA). Data were collected and analyzed using Flowjo 7.6.1 software.

Phagocytosis by BMDC was determined using beads coupled with the fluorescent pH insensitive indicator dye AF647. BMDC were incubated at 37°C for 20 min in the presence of fluorescent beads, washed extensively with cold PBS, and then immediately analyzed by flow cytometry. BMDCs with different numbers of bead phagocytized were gated by assessing fluorescence intensity of AF647. The percentages of cells that had taken up increasing numbers of beads were calculated based on the cell count of each gate to the total CD11c positive cells (Figure 3).

Phagosomal pH was measured by fluorescence quenching of the pH sensitive dye Fluorescein 5-isothiocyanate (FITC) after phagocytosis (15, 22). A detailed experimental process was described by Savina et al. (22). Briefly, 3 μm Polybead® Amino Microspheres activated by 8% glutaraldehyde were incubated with Ovalbumin (OVA) conjugated with FITC (pH sensitive) and AF647 (pH insensitive) in PBS at 4°C overnight. After 30 min of aldehyde blocking with glycine (1M), labeled beads were washed three times and suspended in PBS at a concentration of 1.7 × 10⁷ beads per microliter. BMDCs were stained with α-CD11c antibody. After washing with PBS, BMDCs were incubated with the FITC/AF647 coupled beads for 15 min at 37°C, at a BMDC to bead ratio of 1:3 and washed three times with ice cold PBS to remove the excess beads. Bead pulsed BMDCs were re-suspended in warm RPMI media, incubated for the indicated time points. At each time point, BMDCs were immediately placed on ice and subjected to flow cytometry analysis. The mean fluorescence intensity (MFI) emission for both dyes was determined for BMDCs containing a single phagocytosed bead. The ratio of MFI between FITC and AF647 was employed to indicate the phaghosomal pH value. The phagosomal pH values were determined by establishing a standard curve. To develop the standard curve, BMDC were incubated with coupled beads for 30 min and then re-suspended in HBSS containing 0.1% Triton X-100 adjusted to a pH ranging from 5.5 to 8.0. After an 8-min incubation, these cells are immediately analyzed by flow cytometry and the emission ratio of the two fluorescent probes of BMDC incubated in HBSS at each pH was collected.

Ovalbumin with a self-quenching conjugate that exhibits green fluorescence upon proteolytic degradation (DQ-OVA) and AF647 were covalently conjugated to 3 μm Polybead® Amino Microspheres as described above. Anti-CD11c labeled BMDC were pulsed with DQ-OVA/AF647 coupled beads for 15 min in 37°C media. Extracellular beads were washed away with cold PBS and BMDC were chased for a series of time periods. At each time point fluorescence was measured by flow cytometry and OVA degradation was calculated as the MFI ratio between DQ-OVA and AF647.

Horse cytochrome c was employed to measure antigen translocation from the phagosome into the cytoplasm for cross presentation as previously described (33). BMDC were seeded in 96-well plates and incubated with cytochrome c (0–4 mg/mL) for 24 or 48 h. BMDC apoptosis induced by cytoplasmic cytochrome c serves as an indicator of exogenous antigen being translocated into the cytoplasm (33, 34). BMDC not treated with cytochrome c were used as a negative control and cell viability assessed via MTT assay as previously reported (35).

Cell proliferation was measured by incorporation of radiolabeled tritiated thymidine ([³H]TdR). [³H]TdR was added to BMDC-T cell culture at 1 μCi per well for 16 h before harvesting. After harvest the radioactivity was measured as described previously (24).

The ability of BMDC to internalize whole protein antigen and then induce CD8⁺ T cell activation was assessed using an in vitro cross-presentation assay. To measure the cross-presentation of autoantigen, insulin reactive G9C8 and AI4 T cells were purified and incubated with BMDCs from NOD or NOD-Ncf1^m1J at a ratio of 5:1. Antigens (beads conjugated with Insulin, freeze thawed necrotic NIT-1 cells [a NOD derived β cell line (36)], heat-inactivated insulin as whole antigen, or insulin B15-23 peptide antigen) were added to the cell mixture. CD8⁺ T cell prolifeation was measured by [³H]TdR incorporation an assays (23)].

Statistical analysis was performed using GraphPad Prism (GraphPad Software, Inc., La Jolla, CA) or SAS 9.2 (SAS Institute Inc., Cary, NC). Statistical significance between mean values was determined using the Student's t-test with a significance level at P = 0.05. Time to diabetes onset after adoptive transfer was analyzed by Kaplan-Meier curve. The log-rank test was employed to compare the survival distributions of different groups and correct for multiple comparisons. Except for the survival analysis, all data reported here are representative of at least three independent experiments performed in triplicate.

To assess defects in the diabetogenicity of CTLs and antigen presenting cells (APCs) lacking intracellular superoxide production, purified naïve CD8⁺ T cells collected from prediabetic NOD or NOD-Ncf1^m1J donor mice were adoptively co-transferred with CD4⁺ T cells from prediabetic NOD into either NOD-Rag1^−/− or NOD-Rag1^−/−.Ncf1^m1J recipient animals. Recipients were followed for onset of T1D. As previously noted (23), transfer of NOD.Ncf1^m1J CD8⁺ T cells into immune deficient NOD-Rag1^−/- mice resulted in a significant delay and reduction in disease onset when compared to NOD CD8⁺ T cells (Figure 1 and Table 1. Group 1 vs. Group 2: p = 0.05). Protection was also was also observed when transferring NADPH oxidase 2-competent immune cells (23) into NOD-Rag1^−/−.Ncf1^m1J (Figure 1). Specifically, transfer of NOD CD4⁺ and CD8⁺ T cells into NOD-Rag1^−/−.Ncf1^m1J recipients led to a reduction and delay in T1D onset compared to NOD-Rag1^−/− animals (Figure 1 and Table 1. Group 1 vs. Group 3: p = 0.03). Group 2 and Group 3 were statistically indistinguishable suggesting that cells within the NADPH oxidase 2 deficient recipient mice, likely DC, were providing protection against T1D similar to that observed with lack of NADPH oxidase 2 activity in CD8⁺ T cells (23). The group exhibiting the lowest frequency of diabetes development involved NOD-Rag1^−/−.Ncf1^m1J recipients of NOD.Ncf1^m1J CD8⁺ T cells (Figure 1 and Table 1, Group 4). Indeed, groups 1 and 2 exhibited significantly more T1D when compared to Group 4 (Table 1). While 5 of 9 mice in Group 3 developed T1D compared to only 1 of 10 in Group 4 (Figure 1), statistical comparison of these two groups did not reach significance (Table 1: p = 0.09). Hence, there is a need for p47^phox within both CD8⁺ T cells and DCs for full T1D pathogenesis to be observed.

Maturation of DCs is one of the earliest events in the initiation of T1D (2, 37). In the process of maturation, DCs up-regulate surface co-stimulatory markers and MHC molecules in conjunction with the production of pro-inflammatory cytokines to prime antigen specific T cells. DCs can be activated by an array of Toll-Like Receptor (TLR) or Pattern Recognition Receptor (PRR) signaling pathways and in doing so, respond to infection and damage. Accordingly, many of the TLR signaling pathways have been reported to be associated with T1D (38, 39). To test if is required in TLR signal transduction and the ensuing up-regulation of co-stimulatory molecules on DC, bone marrow-derived DCs (BMDC) from NOD or NOD-Ncf1^m1J were treated for 24 h with either Poly(I:C), LPS, R848, or CpG2336 to activate TLR3, TLR4, TLR7/8, or TLR9, respectively. The activation markers CD80, CD86, and 4-1BBL were measured by surface staining and analyzed by multicolor flow cytometry (Figures 2A–C). All four ligands significantly boosted the expression of CD80 and CD86 (Figures 2A,B), while 4-1BBL was up-regulated by Poly(I:C) and LPS but not R848 nor CpG2336 (Figure 2C). When comparing BMDC from NOD and NOD-Ncf1^m1J, DCs from both strains showed indistinguishable increases in surface expression (Figures 2A–C) suggesting the p47^phox was not required for co-stimulatory molecule up-regulation during DC maturation.

T_H1 responses are the dominant path for pathogenic T cell differentiation in T1D. As professional APCs, DCs control CD4⁺ T cell differentiation though mechanisms of antigen presentation and co-stimulation as well as secretion of cytokines and ROS, which provide the “third signal” for activation (40, 41). IL-12 is the primary cytokine produced by DCs to drive the T_H1 response (42, 43), and IL-12 is required for the priming of diabetogenic CD8⁺ T cells. We have previously demonstrated that macrophages from NOD-Ncf^m1J mice have compromised IL-12 production (24). Similarly, in the context of the B6 or B10 genetic background, Ncf1 has been proposed to play a suppressive role on IL-12 p35 production (44, 45). To investigate the role of Ncf1 in IL-12 expression by DCs and the impact on NOD T cell differentiation, IL-12p35 mRNA (Figure 2D) and production of the pro-inflammatory cytokine TNFα (Figure 2E) were examined 12 h after DC activation with the ligands for TLR1/2, TLR2/6, TLR4, TLR7, or TLR9, Activation of the series of TLRs significantly increased the level of IL-12p35 and TNFα mRNA. Interestingly, we found BMDCs from NOD-Ncf^m1J showed no significant deficiency in the transcription of these cytokines (Figures 2D,E). Overall, DC maturation in NOD-Ncf^m1J mice did not appear deficient in terms of the up-regulation of co-stimulatory molecules or pro-inflammatory cytokines upon targeted stimulations when compared to DCs from NADPH oxidase 2-intact NOD mice.

Type 1 interferons (T1-IFN) are required for the initiation of T1D (46). These cytokines also promote DC cross-presentation by modulating antigen survival within the phagosome, endocytic routing and processing, and hence enhance the cross-priming of islet specific CTLs (47, 48). To determine if Ncf1 regulates the expression and responses of T1-IFN by DCs, BMDC from NOD and NOD-Ncf^m1J were activated by Poly(I:C), R848, or CpG2336, and cell surface levels of CCR7 (Figure 2F) and stem cell antigen-1 (Sca-1, Figure 2G) were assessed by flow cytometry. CCR7 is a T1-IFN-induced chemokine receptor that functions to recruit DCs from peripheral tissue into lymphoid organs (49, 50). Sca-1 is a phosphatidylinositol-linked membrane protein that is also T1-IFN-induced on DCs. After 24 h of stimulation with TLR ligands, CCR7 was mildly upregulated (Figure 2F), while Sca-1 was significantly increased (Figure 2G) to an equal extent on DCs from both the NOD and NOD-Ncf^m1J strains.

To confirm that Ncf1 in DCs is not necessary for production and response of T1-IFN, transcription of the T1-IFN-inducible genes interferon regulatory factor-7 (IRF7), interferon stimulated gene 15 (ISG15), and myxoma response protein-1 (Mx1) in BMDCs from NOD or NOD-Ncf1^m1J were measured after 12-h stimulation by Poly(I:C), R848, or CpG2336 for respective activation of TLR3, TLR7/8, or TLR9. Significant elevations in the expression of IRF7 (Figure 2H), ISG15 (Figure 2I), and Mx1 (Figure 2J) were seen after stimulation with Poly(I:C) or R848, while small increases in transcription were observed following TLR9 activation with CpG2336 (Figures 2H,J). However, no differences were observed in NOD-Ncf1^m1J vs. NADPH oxidase 2 expressing DCs in terms of response to T1-IFN (Figures 2F–J). Taken together, Ncf1 in DCs does not play a role in the inducible T1-IFN production or response after activation of an array of TLRs.

Since Ncf1 is not required for DC maturation measured by upregulation of surface markers and production of pro-inflammatory cytokines following stimulation with various TLRs, we asked if ROS is necessary for the presentation of antigen by DCs. It has been reported that ROS produced by NADPH oxidase 2 is critical for the cross-presentation of antigen in MHC Class I by DCs to CD8⁺ T cells. Cross-presentation can be roughly divided into four steps: (1) phagocytosis of antigens; (2) limiting antigen degradation; (3) translocation of exogenous antigen into cytoplasm; and (4) presentation of antigens onto MHC Class I molecules. The proposed mechanism whereby NADPH oxidase 2 promotes an increase in antigen cross-presentation is through attenuating the phagosomal acidification following phagocytosis, thus facilitating antigen translocation into the cytoplasm (15, 21, 22). To examine if ROS is required for DC phagocytosis, we employed AlexaFluor 647 (AF647) labeled latex beads as a probe. Antibody-labeled (α-CD11c) BMDC from NOD and NOD-Ncf^m1J were incubated with AF647-labeled beads for 15 min at 37°C and subjected to flow cytometry (Figure 3A). Quantitative analysis of the CD11c positive population was conducted by gating on DCs that had taken up different numbers of beads. The percentage of DCs having phagocytized from one to five beads showed a declining trend (Figure 3B). However, bead uptake was comparable between NOD and NOD-Ncf1^m1J (Figure 3B). Thus, we concluded that Ncf1 is dispensable for antigen uptake by DCs.

Although it has been reported that ROS are necessary for regulation of phagosomal acidification, some studies have argued that ROS are dispensable in phagosomal pH regulation (15, 17). These previous conflicting studies were conducted using the C57BL/6 (B6) background rather than the autoimmune-prone NOD mouse. One significant difference regarding antigen presentation between NOD and B6 mice involves the Slc11a1 gene within the Idd5 interval on Chromosome 1. Nramp1, the gene product of Slc11a1, promotes acidification of phagosomes in DCs and this protein is deficient in B6 mice (51) leading to compromised phagosomal acidification following phagocytosis. Here we used cells derived from NOD mice with a functional Nramp1 (52). To examine phagosomal acidification we utilized FITC, a widely used pH sensitive fluorescence dye. FITC conjugated ovalbumin (DQ-OVA) and the pH insensitive dye AF647 were linked to latex beads. BMDC were then incubated with these FITC/AF647-fluorescent beads. Notably, even with functional Slc11a1 alleles, phagosomal pH in NOD DC was almost neutral through the initial 60 min after phagocytosis. In stark contrast, we observed rapid and significant decreases in phagosomal pH for NOD-Ncf1^m1J DC. Indeed, pH quickly dropped to around 5 within the first 15 min of the observation period (Figure 4), suggesting a rapid DC phagosomal acidification when Ncf1 was non-functional. The final pH value measured in NOD-Ncf1^m1J DCs was 1 unit lower than the previously reported values using gp91^phox-defective B6 DCs (15), which could be rescued to approximately pH 7 by adding the V-ATPase inhibitor concanamycin A (Figure 4B). Overall, Ncf1 indispensable for NOD DCs to maintain a neutral pH shortly after phagocytosis.

In DCs, for efficient cross-presentation of exogenous antigen, protein degradation is restricted immediately following phagocytosis. As shown in the previous reports and above, DCs constrain phagosomal acidification to inhibit antigen degradation (15, 21, 22). Due to the strong decrease of phagosome pH values observed in NOD-Ncf1^m1J DC, we expected to observe accelerated antigen digestion. To test this, we employed DQ-conjugated ovalbumin (DQ-OVA), which is designed to have self-quenched fluorescence group pairs when the three-dimensional conformation is intact. Upon degradation of this protein, the fluorescence and quenching groups detach from one another, and green fluorescence is emitted upon excitation (Figure 5A). Here, latex beads were linked with DQ-OVA and AF647 and incubated with BMDCs from NOD or NOD-Ncf1^m1J mice. DCs were analyzed, as described in the Materials and Methods by flow cytometry at the indicated time points post-phagocytosis. In NOD DCs, only a mild increase in fluorescence (~5%) was observed as late as 60 min after pulsing (Figure 5). This lack of antigen degradation is associated with maintenance of neutral pH (Figure 4). However, in NOD-Ncf1^m1J DC there was a rapid increase in FITC fluorescence intensity. Indeed, only 15 min post-pulsing, the fluorescence intensity of DQ-OVA increased by 50% (Figure 5B). Therefore, NOD-Ncf1^m1J DC rapidly exhibited a reduction in pH (Figure 4) and degradation of ovalbumin (Figure 5) indicating that without Ncf1, there is accelerated antigen degradation in the phagosome.

In cross-presentation, exogenous protein is exported into the cytoplasm by an unknown mechanism and processed by the proteasome for peptide antigen loading into MHC Class I molecules (14). To measure protein translocation into the cytosol, we employed cytochrome c as a probe. Cytochrome c is a protein that is sequestered in the inner mitochondrial membrane and functions as part of the electron transport chain. During apoptosis, cytochrome c is released into the cytoplasm and complexes into the apoptosome to induce apoptotic cell death. When cross-presenting cells such as BMDCs are incubated with recombinant cytochrome c, they will internalize cytochrome c and transport the intact protein from endosomes into cytoplasm, which triggers apoptosome activation and caspase-3 dependent apoptosis; in contrast, in cells that are incapable of translocating exogenous cytochrome c to cytoplasm, apoptosis would not be triggered (33, 34). Thus, we employed cellular apoptosis to indicate antigen translocation by incubating BMDC with cytochrome c. NOD and NOD-Ncf1^m1J DC were treated with increasing concentrations of cytochrome c [1 to 4 mg/mL] for 24 or 48 h, and apoptosis was measured by MTT assay. In both 24 and 48-h groups, cytochrome c induced a dose dependent killing of NOD DCs (Figure 6). However, cytochrome c did not induce apoptosis of NOD-Ncf1^m1J DCs indicating that cytochrome c was not transported into the cytoplasm in p47^phox deficient DCs (Figure 6), consistent with accelerated antigen degradation in NOD-Ncf1^m1J DC.

To determine if excessive antigen degradation and inhibition of antigen translocation have an impact on CTL activation, an in vitro cross-presentation assay was performed. BMDC from NOD-Ncf1^m1J or NOD were pulsed with antigen and incubated with G9C8 cells, an insulin specific CD8⁺ T cell clone (53). T cell proliferation was then measured by [³H]TdR incorporation. When DCs were pulsed with the specific insulin B15-23 peptide antigen, T cell proliferation was comparable in both groups. This suggests that when antigen processing is not required, NOD-Ncf1^m1J DCs have no deficiency in priming diabetogenic CD8⁺ T cells, consistent with the data above showing equal upregulation of co-stimulatory markers and pro-inflammatory cytokine expression by Ncf1-intact and -deficient DCs (Figure 2). In contrast, when DCs were primed with heat-inactivated insulin, G9C8 CD8⁺ T cell proliferation did not occur when the DCs were from NOD-Ncf1^m1J mice (Figure 7B). Strong G9C8 proliferation was induced when Ncf1-intact NOD DCs processed and presented heat-inactivated insulin (Figure 7B). Together, these data indicate that antigen cross-presentation is severely affected by the absence of Ncf1.

To mimic the scenario in T1D, necrotic NIT-1 cells (pancreatic β cell line derived from a NOD mouse) were added to DC-G9C8 co-cultures. As expected, NOD-Ncf1^m1J DCs were incapable of inducing diabetogenic T cell activation compared with the Ncf1-intact control DC (Figure 7C). This highlights the indispensable role of NADPH oxidase 2 in the DC to activate diabetogenic T cells in T1D. In summary, processing of exogenous autoantigen by NOD-Ncf1^m1J DCs is deficient, and without a functional Ncf1, DCs are not able to cross-present autoantigens to diabetogenic CD8⁺ T cells and drive the activation of these CTL.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.