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Differential Contributions of APC Subsets to T Cell Activation in Nonobese Diabetic Mice¹

Department of Microbiology and Immunology and Robarts Research Institute, University of Western Ontario, London, Ontario, Canada

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 
Despite the pivotal role of dendritic cells (DC) in shaping immunity, little is known about their functionality in type 1 diabetes. Moreover, due to the paucity of DC in vivo, functional studies have relied largely upon in vitro-expanded cells to elucidate type 1 diabetes-associated functional abnormalities. In this study, we provide a comprehensive analysis of the functional capabilities of in vivo-derived DC subsets from NOD mice by comparing DC to other NOD APC types and to DC from autoimmune-resistant strains. NOD DC closely resemble those from nonautoimmune strains with respect to costimulation and cytokine production. The exception is the CD8⁺CD11b^–DC subset which is numerically reduced in NOD spleens, but not in the pancreatic lymph nodes, while DC from both tissues produce little IL-12 in this strain. This defect results in unusual deferral toward macrophage-derived IL-12 in NOD mice; NOD macrophages produce aberrantly high IL-12 levels that can overcompensate for the DC defect in Th1 polarization. APC subset use for autoantigen presentation also differs in NOD mice. NOD B cells overshadow DC at activating islet-reactive T cells, whereas DC and B cells in NOD-resistant mice are functionally comparable. Differential involvement of APC subsets in T cell activation and tolerance induction may prove to be a crucial factor in the selection and expansion of autoreactive T cells.

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 
The NOD mouse recapitulates many aspects of the pathogenesis of type 1 diabetes (T1D)⁵ in humans and is therefore widely applied in studies addressing the cellular mechanisms of autoimmunity. APC, mainly dendritic cells (DC) and macrophages, first appear at the islet periphery at 4 wk of age and their appearance correlates with transient hyperinsulinemia and islet neogenesis (1, 2). Islet-infiltrated DC and macrophages produce TNF- which drives the proinflammatory response during insulitis (3), leading to CD4⁺ and CD8⁺ T cell recruitment and activation (4).

Numerous studies have investigated the role of APC, particularly B cells and macrophages, in T1D pathogenesis in NOD mice. Studies involving ablated B cell development or manipulated Ag-presentation function have shown that B cells are essential for disease (5, 6, 7, 8, 9). NOD B cells demonstrate efficient presentation of islet Ag to T cells, which is attributable to Ag capture via Ig receptors (10). NOD B cells also exhibit a plethora of aberrant activation characteristics, including resistance to tolerance induction (11), increased NF-B activity (12), hyperproliferation, resistance to apoptosis and enhanced costimulation (13, 14). Similarly, we and others have demonstrated that NOD macrophages and bone marrow (BM)-derived DC exhibit elevated IL-12 production and NF-B hyperactivation (15, 16, 17, 18, 19, 20, 21). However, surprisingly little information is available concerning the functional capabilities of in vivo-derived DC in NOD mice. Studies of NOD DC have mainly used in vitro-generated cells, which are now recognized to be phenotypically and functionally distinct from the heterogeneous DC subsets that exist in vivo (22). Specifically, the 7-day GM-CSF plus IL-4 culture protocols for generating DC primarily expand a myeloid population whose in vivo equivalent is not known.

In this study, we present a comprehensive functional analysis of splenic CD8⁺ and CD8^– DC, the DC subsets which are prevalent throughout mouse lymphoid tissues, in NOD and autoimmune-resistant mice. In the spleen, CD8⁺ DC reside in the T cell areas while CD8^– DC are localized in the marginal zones (23, 24). Both DC subsets are capable of priming naive T cells, although this is accomplished using different cytokine pathways and in response to distinct endogenous signals and microbial products (25). This report identifies specific abnormalities in the numbers and IL-12-producing capability of CD8⁺ DC in the NOD periphery. These data also challenge several long-standing presumptions, namely that in vitro- and in vivo-derived DC are functionally comparable and that an increased proclivity toward promoting type 1 cytokine responses is a generalized trait of NOD APC. Moreover, by placing DC in the spectrum of NOD APC activity, we show that macrophage-derived IL-12 and Ag presentation by B cells override the functional capabilities of DC. Therefore, in terms of mediating inappropriate T cell activation, DC appear to harbor minimal functional impairment in comparison to the other NOD professional APC.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 
Mice

Female NOD/Lt mice were bred in the animal facility at the Robarts Research Institute (London, Canada). Female C57BL/6, BALB/c, and NOD-resistant (NOR) mice were purchased from The Jackson Laboratory. Mice were maintained in the specific pathogen-free facility at the University of Western Ontario (London, Canada). All experiments were performed in accordance with institutional guidelines for animal care. Female NOD mice were compared with age- and sex-matched diabetes-resistant control strains. Unless otherwise specified, mice were used between 4 and 6 wk of age and were therefore nondiabetic. Diabetes incidence was monitored by weekly measurement of venous blood glucose concentrations in nonfasting mice using Glucometer Elite strips (Bayer). Mice with two consecutive blood glucose concentrations >300 mg/dl were considered diabetic, which typically occurred between 15 and 20 wk of age in our colony.

Culture medium, cytokines, TLR agonists

RPMI 1640 medium was supplemented with 2 mM L-glutamine, 0.5% HEPES, 5 µg/ml penicillin, 100 U/ml streptomycin (Invitrogen Life Technologies) and 10% (v/v) FCS (HyClone Laboratories). Murine cytokines (GM-CSF, IL-4, IFN-, and TNF-) were purchased from Cedarlane Laboratories and reconstituted in sterile water. LPS (Escherichia coli serotype 055:B5) was obtained from Sigma-Aldrich. Polyinosine-polycytidylic acid (poly(I:C)) was purchased from Sigma-Aldrich and reconstituted in sterile PBS.

Isolation of splenic DC, macrophages, and B cells from spleens

Mice were euthanized by CO₂ narcosis and spleens were harvested. Pooled spleens were cut into small fragments and digested for 30 min at room temperature with gentle and continuous agitation in RPMI 1640 containing 1 mg/ml collagenase A (Roche Diagnostics) and 40 µg/ml DNaseI (Roche Diagnostics). Spleen fragments were intermittently resuspended by gentle pipetting. Cell suspensions were filtered through a sterile nylon mesh to remove undigested material and were subsequently treated for 5 min with PBS supplemented with 5% FCS and 5 mM EDTA (pH 7.2) to disrupt DC/T cell complexes. Following another washing step in medium, RBC were removed using ACK lysing buffer (BioWhittaker). For labeling and purification of DC, the medium consisted of PBS containing 2 mM EDTA and 0.5% BSA. FcR were blocked for 15 min with anti-mouse CD16/CD32 (2.4G2; BD Pharmingen) at 4°C. Cells were labeled with MACS CD11c (N418) Microbeads (Miltenyi Biotec) for 15 min at 4°C. Positive selection for CD11c⁺ cells was done using a MidiMACS Separator and MACS columns (Miltenyi Biotec) according to the manufacturer’s instructions. The resulting purity of DC was consistently >90% as verified by flow cytometry.

To purify DC subsets, a two-step method was used. DC were first pre-enriched from bulk splenocytes by negative selection using a dendritic cell isolation kit (Miltenyi Biotec). Briefly, non-DC were labeled with an Ab mixture and removed by MACS separation. CD11c^highCD8⁺ and CD11c^highCD8^– DC were sorted using a FACSVantage (BD Biosciences) and cell purity was verified to be at least 97%.

To purify splenic macrophages, CD4 (GK1.5), CD8 (HO2.2), CD11c (HL3), and B220 (RA3-3A1/6.1)-expressing cells were depleted from the cell suspensions by complement-mediated lysis. Macrophages were then isolated by adherence to plastic. Flow cytometry was done to verify the purity of the cells, which were routinely >92% CD11b⁺ and >65% F4/80⁺.

To purify B cells, splenocytes were labeled with mouse CD19 microbeads or CD45R/B220 microbeads and positive selection was performed using MACS separation columns according to the manufacturer’s instructions (Miltenyi Biotec), which routinely yielded a population that was >80% positive for B cell surface markers.

Maturation of DC and macrophages in vitro

Splenocytes or purified APC populations were matured in vitro with combinations of proinflammatory cytokines and TLR agonists. For DC maturation, cells were treated with LPS (100 ng/ml) plus TNF- (10 ng/ml), GM-CSF (10 ng/ml) plus IL-4 (10 ng/ml) plus IFN- (20 ng/ml) ± poly(I:C) (100 ng/ml, unless otherwise indicated) or LPS (100 ng/ml) plus irradiated CD40L-transfected J558 cells (a gift from P. Lane, University of Birmingham, Birmingham, U.K.). J558 cells were gamma-irradiated before use with 2500 rad from a Cobalt 60 source. Preliminary experiments were conducted to determine that 1 DC:1 J558 cell was an optimal ratio for induction of IL-12p70 synthesis by DC. Macrophages were matured by treatment with LPS (1 µg/ml). After 24 h of culture, unless otherwise indicated, cells were pelleted and prepared for flow cytometry. Supernatant samples were stored at –70°C to test cytokine concentrations by ELISA.

In vivo DC activation

Four-week-old NOD and NOR mice (n = 3/group) were injected i.v. with PBS (vehicle control), LPS (30 µg/mouse), or poly(I:C) (100 µg/mouse). At 6 h postinjection, splenocytes were pooled from each treatment group and CD11c⁺ cells were selected using the MACS separation technique. DC (5 x 10⁵/ml) were plated in medium without exogenous cytokines for 30 h and supernatant samples were collected for analysis of the IL-12p70 content by ELISA.

Flow cytometry

All staining steps were performed at 4°C in PBS. FcR were blocked with anti-CD16/32 Ab (clone 2.4G2; BD Pharmingen) and cells were subsequently incubated with 0.5 µg/10⁶ cells of the relevant Ab for 45 min. The following anti-mouse Abs were purchased from BD Pharmingen: anti-CD11c (HL3), anti-CD80 (16-10A1), anti-CD86 (GL1), anti-CD40 (HM40-3), anti-CD4 (L3T4), anti-CD8 (53-6.7), anti-CD11b (M1/70), anti-B220 (RA3-6B2), and anti-I-A^k (10-3.6). Anti-mouse F4/80 Ab was purchased from Serotec (CI:A3-1). Isotype-matched control Abs were purchased from BD Pharmingen and Cedarlane Laboratories. Samples were analyzed on a FACSCalibur flow cytometer (BD Biosciences) using CellQuest software (BD Biosciences). Live cells were selected by forward/side scatter gating.

Flow cytometry to identify dead and apoptotic cells was performed by staining splenocytes with Abs against DC subset markers (CD11c and CD8) in combination with annexin V and propidium iodide (PI) using a kit from BD Pharmingen. Apoptotic DC were identified as annexin V-positive, PI-negative cells.

BrdU labeling and analysis

Groups of three or four mice (NOD and C57BL/6) were given BrdU (0.8 mg/ml; Sigma-Aldrich) in sterile drinking water that was changed daily. After 2 or 5 days, splenocytes were stained for cell surface molecules and were subsequently stained to detect BrdU incorporation by flow cytometry using a kit from BD Pharmingen.

Intracellular cytokine staining

Pancreatic lymph nodes (PLN) were excised from 6-wk-old NOD and NOR mice and were pressed through a sterile nylon steel mesh to obtain cell suspensions. Flow cytometry was performed using freshly isolated PLN-derived cells to identify CD11c^highCD11b^high and CD11c^highCD11b^low/– populations. Cells obtained were cultured (2 x 10⁶/ml) with GM-CSF (10 ng/ml) plus IL-4 (10 ng/ml) plus IFN- (20 ng/ml) plus poly(I:C) (100 ng/ml) at 37°C for 10 h. Brefeldin A (5 µg/ml) was added for the last 5 h of culture to prevent cytokine secretion by blocking intracellular transport processes. Surface staining for CD11c and CD11b was conducted followed by fixation and permeabilization using a commercially available kit (Cedarlane Laboratories) and staining for IL-12p40/p70 (BD Pharmingen) for flow cytometry.

APC/T cell cocultures and T cell proliferation assays

Spleens were mechanically dissociated by passage through a nylon steel mesh and RBC were lysed. T cell purification was performed using mouse CD4 subset mini-column kits from R&D Systems according to the manufacturer’s instructions. MACS-purified (CD11c⁺) or sorted DC subsets (CD11c⁺CD8⁺ and CD11c⁺CD8^– cells) were cultured with syngeneic T cells plus soluble anti-mouse CD3 (0.5 µg/ml; Cedarlane Laboratories) or with allogeneic C57BL/6 T cells. In other assays, macrophages and DC were either plated separately or together in varying ratios (total APC equaling 2 x 10⁵/ml) in 96-well round-bottom plates with fixed numbers of syngeneic T cells (10⁶/ml) and soluble anti-mouse CD3 (0.5 µg/ml). The APC were freshly purified or preactivated for 18 h with LPS (1 µg/ml) for macrophages and with GM-CSF (10 ng/ml) plus IL-4 (10 ng/ml) plus IFN- (20 ng/ml) plus poly(I:C) (100 ng/ml) for DC. Mature APC were washed three times to remove residual cytokines before coculture with T cells. For measurement of proliferation, APC/T cell combinations were plated in triplicate in 96-well round-bottom plates in final volumes of 200 µl/well. Titrations of APC were plated with fixed numbers of T cells as indicated in the graph legends. Plates were incubated in 5% CO₂ at 37°C for 3 or 4 days for syngeneic and allogeneic assays, respectively. Cells were pulsed with 1 µCi/well [³H]thymidine (NEN-DuPont) for the last 18 h of culture. [³H]Thymidine uptake was measured using a cell harvester (Tomtec) and a liquid scintillation counter (Wallac). The results were expressed as the mean cpm ± SD from triplicate wells. To measure cytokine concentrations, APC/T cell combinations were plated in 24-well plates at the indicated concentrations and supernatant samples were taken from 60 h cultures for ELISA.

Assessment of glutamic acid decarboxylase (GAD) p524-reactive T cell activation

GAD65 peptide comprised of amino acids 524–543 (GADp524) was synthesized in our laboratory using the Merrifield solid-phase technique using an ABI Peptide Synthesizer (Applied Biosystems) as before (26). Peptides were purified using HPLC on a C18 reverse-phase semipreparative Synchropak RP-P column (Synchrom), lyophilized, and stored at –20°C until use. Purity of the peptides was verified using mass spectrometry. Peptides were reconstituted in double-distilled H₂O and sterilized by passage through a 22-µm filter before incorporation into cell cultures.

APC subsets were used to stimulate induced and spontaneous proliferation responses to GADp524. For induced responses, 6-wk-old NOD or NOR mice were immunized s.c. in the hind footpads with 100 µg of GADp524 emulsified in CFA. Popliteal lymph nodes were removed 10 days later and teased through nylon meshes to obtain single-cell suspensions. CD4⁺ T cells were then isolated from pooled lymph node samples using mouse CD4 subset mini column kits (R&D Systems). Splenocytes from separate, nonimmunized mice (4–5 wk of age) were harvested for purification of DC (CD11c⁺ cells) by MACS separation. CD4⁺ T cells (2 x 10⁵/well) together with titrations of DC were plated in 96-well microtiter plates with GADp524 for 4 days. GADp524 was added at the optimal concentration of 10 µM.

For assaying spontaneous T cell responses, CD4⁺ T cells were purified from the spleens of nonimmunized 8- to 10-wk-old NOD mice as described and were cocultured with dilutions of MACS-purified B cells (B220⁺ cells), MACS-purified DC (CD11c⁺ cells), or FACS-sorted CD8⁺ or CD8^– DC from NOD and NOR mice. Before incorporation into T cell cocultures, B cell proliferation was inhibited by treating the cells with 50 µg/ml mitomycin C for 30 min at 37°C, followed by thorough washing, counting, and resuspension in medium. Cultures were established in triplicate wells in 96-well flat-bottom plates with GADp524 at the indicated concentrations. [³H]Thymidine incorporation assays were used to measure T cell proliferation after 5 days. In the spontaneous assays, IFN- was also measured by ELISA using 48-h supernatant samples and cells were taken at 24 h for flow cytometry.

Cytokine measurements by ELISA

Cytokines in supernatant samples were measured using OptEIA ELISA sets for IL-12p70, IL-12p40, IL-10, IFN-, IL-4, TNF-, and IL-18 (BD Pharmingen) in accordance with the manufacturer’s instructions. All samples were analyzed in duplicate wells. Plates were read using a Bio-Rad ELISA plate reader. All results are expressed as the mean picograms per milliliter of cytokine ± SD from duplicate or triplicate wells.

Statistics

Statistical comparisons of two groups were performed using the Student t test and multiple comparisons were conducted using one-way ANOVA tests and Bonferroni multiple comparison tests for subsequent pairwise comparisons where appropriate. A p value <0.05 was considered significant.

	Results

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 
NOD spleens have an altered DC subset composition

To assess the frequency of DC in NOD mice, comparisons were drawn to splenic DC from autoimmune-resistant NOR, C57BL/6, and BALB/c mice. The NOR strain shares the NOD MHC haplotype as well as a proportion of the diabetes susceptibility genes. Although NOR islets exhibit APC infiltration, the subsequent recruitment of T cells is minimal, suggesting that differences in APC activity in NOD mice are responsible for the transition to overt disease (27). Flow cytometry was performed to identify expression of CD11c, a surface molecule which phenotypically identifies DC in murine lymphoid organs (24) (Fig. 1A, upper panels, depicting the lymphocyte gate from representative mice). We observed that the percentages of CD11c^high cells in NOD spleens were within a normal range based on comparisons to age-matched control mice, representing 3% of gated lymphocytes (Fig. 1B). The absolute numbers of CD11c^high cells were not significantly different in NOD and age-matched control spleens (Fig. 1C). Moreover, diabetic NOD mice did not display altered percentages or numbers of CD11c^high cells compared with age-matched normoglycemic NOD mice or control strains (Fig. 1, B and C). In all of the strains examined, the numbers of CD11c^high cells increased between 4 and 8 wk of age but did not change between 8 and 16 wk of age.

View larger version (43K): [in this window] [in a new window]   FIGURE 1. Percentages and numbers of splenic DC subsets in NOD and autoimmune-resistant control strains. A, Flow cytometry was performed to analyze DC subsets in spleens from 8-wk-old NOD and autoimmune-resistant mice. Upper panels, Viable lymphocytes were gated based on forward and side scatter properties and the percentages of CD11chigh cells were analyzed. Lower panels, CD11chigh cells were gated to identify CD8+ populations. B, Summary of the percentages of CD11chigh cells in spleens from the indicated strains/ages of mice, as determined by flow cytometry. Diabetic mice were between 15 and 19 wk of age, whereas NOD mice in all other age categories were normoglycemic. C, The absolute numbers of CD11chigh cells (x106) were calculated by multiplying total splenocyte counts by the percentages of CD11chigh cells in B. D, Summary of the percentages of CD8+ DC among gated CD11chigh cells from NOD and control strain spleens. *, A statistically significant difference between NOD and age-matched autoimmune resistant mice. The results are from six to eight mice per group and are expressed as the mean ± SD from individual animals.

DC subsets were also gated on the basis of CD8, CD11b, and CD4 expression; the splenic CD11c^high population consists of CD8⁺CD11b^low/–CD4^–, CD8^–CD11b^highCD4⁺, and CD8^– CD11b^highCD4^– populations in autoimmune-resistant mice (28). We observed that the NOD splenic DC compartment contained a pronounced reduction in the percentages of CD8⁺CD11b^low/– cells and a coordinate increase in the proportions of CD8^–CD11b^high cells at 8 wk of age. CD8⁺CD11b^low/– cells represented 18.0 ± 2.4, 25.2 ± 1.8, 27.8 ± 3.4, and 24.4 ± 2.7% of gated CD11c^high cells in NOD, NOR, C57BL/6, and BALB/c mice, respectively. The CD8^–CD11b^high population comprised 81.6 ± 3.5, 74.0 ± 2.3, 69.8 ± 0.9, and 71.9 ± 3.2% of NOD, NOR, C57BL/6, and BALB/c CD11c^high cells, respectively (n = 6–8 mice/strain, p < 0.05 for NOD vs control strain populations). We also observed that the CD8^–CD11b^high subset of NOD DC contained increased percentages of CD4⁺ cells compared with control strains; CD4⁺ cells comprised 70.4 ± 3.2, 58.5 ± 1.2, and 54.2 ± 1.8% of gated CD11c^highCD8^–CD11b^high splenocytes in 8-wk-old NOD, NOR, and BALB/c mice, respectively. However, because few functional distinctions between CD8^–CD4⁺ and CD8^–CD4^– DC subsets have been characterized to date, we chose to categorize DC on the basis of the broader CD8⁺ and CD8^– subsets (as depicted in Fig. 1A) in the subsequent experiments.

Unlike the other DC subsets, the frequency of plasmacytoid DC is highly variable between mouse strains (29). Our data revealed that the percentages and numbers of CD11c^lowB220⁺ cells in spleens from 8-wk-old NOD mice were within the broad range found in autoimmune-resistant mice (data not shown). Therefore, for the purpose of this report, we have focused on the CD8⁺ and CD8^– DC subsets in NOD mice, excluding the plasmacytoid DC population.

Limited differentiation of CD8⁺ DC and skewing toward the CD8^– DC subset in NOD mice

Murine DC have a short lifespan in the spleen; the CD8⁺ subset has an 3-day lifespan, whereas CD8^– DC are longer-lived (31, 32). NOD and C57BL/6 mice (6 wk of age) were given the DNA precursor BrdU in their drinking water for 2 or 5 consecutive days to measure the turnover of DC populations using previously described methods (31, 32). Briefly, splenocytes were surface-stained for DC subset molecules, fixed/permeabilized, and subsequently stained with fluorescent anti-BrdU or isotype control Ab for flow cytometry. Because DC are largely nonproliferating, the BrdU-labeled population represents the cells that have entered the splenic DC compartment either through expansion of a splenic precursor or through replenishment by precursors from the blood or BM (31). Representative histograms depicting BrdU-stained CD11c^high cells (total DC) and the CD8⁺ and CD8^– DC subsets in NOD and C57BL/6 spleens on day 2 of labeling are provided in Fig. 2A and are summarized for all mice on days 2 and 5 of labeling (Fig. 2A, bar graph). The data revealed that lower percentages of BrdU⁺ cells were found within the NOD DC compartment on days 2 and 5. Importantly, however, evaluation of the BrdU labeling kinetics of individual DC subtypes showed no differences between NOD and C57BL/6 mice with respect to the percentages of CD8⁺ or CD8^– DC that had incorporated BrdU (Fig. 2A). This finding suggested that the difference in BrdU labeling kinetics of total CD11c^high cells was attributable to the differing DC subset composition in NOD and C57BL/6 mice, rather than due to actual differences in DC half-life. In support of this notion, the absolute numbers of BrdU⁺CD11c^highCD8⁺ cells in NOD spleens were reduced and the numbers of BrdU⁺CD11c^highCD8^– cells were increased in comparison to C57BL/6 mice after 5 days of labeling (Fig. 2B). Lastly, we also examined the steady-state survival of NOD, NOR, and C57BL/6 DC by staining splenocytes with annexin V and PI. The percentages of annexin V⁺PI^– cells, which represent the cells undergoing apoptosis, did not differ in the CD8⁺ or CD8^– DC populations from NOD, NOR, or C57BL/6 spleens (Fig. 2C). Collectively, these data suggest that the increased frequency of CD8^– DC and the reduced differentiation of CD8⁺ DC in NOD mice are attributable to differences in the availability of DC precursors rather than due to altered DC turnover. These data provide valuable insight that the activity of DC precursors is altered in NOD mice although further studies will be required to decipher whether differences in precursor migration and/or lineage commitment account for these DC subset abnormalities.

View larger version (34K): [in this window] [in a new window]   FIGURE 2. Turnover and survival of DC subsets in NOD and C57BL/6 spleens. A and B, To measure the incorporation of BrdU in newly generated DC, 6-wk-old NOD and C57BL/6 mice were given BrdU in their drinking water for 2 or 5 consecutive days. Splenocytes were harvested and stained for CD11c and CD8, fixed/permeabilized, and stained with anti-BrdU Ab to identify proliferating cells. A, Representative histograms depict the percentages of BrdU+ cells among gated CD11Chigh, CD11chighCD8+, and CD11chighCD8– populations on day 2 of labeling. The bar graph depicts the mean percentages ± SD (n = 4 mice/group) on days 2 and 5 of BrdU labeling. B, The absolute numbers (x105) of BrdU+ CD11chighCD8+ and BrdU+CD11chighCD8– cells were calculated after 5 days of BrdU labeling. C, Apoptosis of DC was assessed by staining freshly isolated NOD, NOR, and C57BL/6 splenocytes with Abs against DC subset-specific markers, annexin V, and PI and tabulating the percentages of annexin V-positive PI-negative cells among gated CD11chighCD8+ and CD11chighCD8– populations (n = 4 mice/strain). *, A statistically significant difference between NOD and control strain DC populations.

Because functional defects of APC have been proposed to play a role in diabetes pathogenesis, we asked whether the immunostimulatory capabilities of NOD DC subsets were altered in comparison to autoimmune-resistant strains. Resting DC subsets from NOD mice did not display differences in expression of CD80, CD86, or CD40 relative to DC from NOR and BALB/c spleens (Table I, geo mean fluorescence intensity (MFI) values). To assess the phenotype of NOD DC during maturation, several treatments were tested: LPS (100 ng/ml) plus irradiated J558 cells (a CD40L-transfected cell line), LPS (100 ng/ml) plus TNF- (10 ng/ml), and GM-CSF (10 ng/ml) plus IL-4 (10 ng/ml) plus IFN- (20 ng/ml) plus poly(I:C) (100 ng/ml). Splenocytes were incubated with the indicated stimuli for 18 h and flow cytometry was done to assess costimulatory molecule expression on DC subsets. Comparable expression levels of CD80 and CD40 were found on CD8⁺ and CD8^– DC from NOD and autoimmune-resistant mice under the various maturation conditions (Table I), although NOD and NOR CD8⁺ DC exhibited reduced CD86 expression compared with BALB/c DC in response to proinflammatory stimuli. However, in allogeneic MLR assays with naive C57BL/6 T cells, NOD and NOR total DC (Fig. 3A), CD8⁺ DC (Fig. 3B), and CD8^– DC (Fig. 3C) at varying concentrations demonstrated a similar allostimulatory ability, proving that NOD DC are competent costimulators.

View larger version (19K): [in this window] [in a new window]   FIGURE 3. Equivalent allostimulation by NOD and NOR DC populations. MACS-purified CD11c+ cells (A) or FACS-sorted CD11chighCD8+ (B) and CD11chighCD8– DC (C) from NOD and NOR spleens were cultured at the indicated dilutions with allogeneic C57BL/6 T cells (105 cells/well). T cells alone (T) did not proliferate. Proliferation was determined by addition of [3H]thymidine for the last 18 h of the 96-h culture. Results are presented as mean cpm ± SD of triplicate wells. T cell proliferation elicited by NOD vs NOR DC was not statistically different.

NOD CD8⁺ DC demonstrate abnormally low IL-12 production and increased IL-10 production in response to innate and T cell-derived signals

Numerous studies have reported that NOD macrophages and in vitro-differentiated DC produce increased quantities of IL-12 in response to maturation stimuli (15, 16, 18, 20, 21); therefore, we explored whether in vivo-derived DC from NOD mice possess similar characteristics. IL-12p70 is the biologically active heterodimer comprised of p40 and p35 subunits, whereas the p40 form is produced in excess as free monomers or dimers (33). IL-12p70 synthesis by murine splenic DC requires multiple signals which are transmitted through TLR and CD40 and/or proinflammatory cytokines (34, 35). To assess IL-12 production, we cultured NOD and NOR DC (MACS-purified CD11c⁺ cells) with a cytokine mixture consisting of GM-CSF (10 ng/ml), IL-4 (10 ng/ml), and IFN- (20 ng/ml) in combination with poly(I:C) (100 ng/ml), a treatment which is known to induce IL-12p70 production by CD8⁺ DC (36). Significantly, analysis of cytokine secretion showed that NOD DC produced reduced quantities of IL-12p70 and IL-12p40 in comparison to NOR DC in response to the cytokine mixture plus titrated doses of poly(I:C) (Fig. 4A). Expectedly, DC that were treated with cytokines in the absence of poly(I:C), and are therefore partially mature, did not produce bioactive IL-12p70. However, production of IL-12p40 by NOD DC was reduced in comparison to NOR DC (Fig. 4A). NOD and NOR DC produced equivalent quantities of two other proinflammatory cytokines, TNF- and IL-18 (Fig. 4A). As a second method of stimulating IL-12 secretion, NOD and NOR DC were treated with LPS (100 ng/ml) plus titrations of irradiated CD40L-transfected J558 cells, which also revealed significantly impaired IL-12p70 production by NOD DC (Fig. 4B), thereby demonstrating that this feature of NOD DC was not stimulus specific.

View larger version (43K): [in this window] [in a new window]   FIGURE 4. Selective reduction of IL-12 synthesis by NOD DC. A, MACS-purified splenic DC (8 x 105 cells/ml) from NOD and NOR mice were stimulated with GM-CSF (10 ng/ml) + IL-4 (10 ng/ml) + IFN- (20 ng/ml) (designated as mature DC) ± poly(I:C) at the indicated doses. Immature DC were treated with GM-CSF (10 ng/ml) alone. Supernatants were sampled after 24 h for IL-12 p70, IL-12 p40, TNF-, and IL-18 measurement by ELISA. B, Splenic DC (1 x 106/ml) were cultured with LPS (100 ng/ml) plus titrations of irradiated CD40L-transfected J558 cells (ratios ranging from 1 DC:5 J558 cells to 10 DC:1 J558 cell) and IL-12p70 was measured in 24-h supernatant samples. C, Assessment of IL-12p70 production by DC following in vivo activation. Four-week-old NOD and NOR mice (n = 3/strain) were injected i.v. with LPS (30 µg/mouse), poly(I:C) (100 µg/mouse), or PBS (vehicle control). Pooled splenocytes from each strain were taken at 6-h postinjection and CD11c+ cells were selected using MACS columns. DC were plated (8 x 105 cells/ml) in nonsupplemented RPMI 1640 medium for a collection of 30-h supernatant samples for measurement of IL-12p70 by ELISA. D, Comparison of IL-12 production by APC in NOD and NOR PLN. Flow cytometry was performed using PLN-derived lymphocytes that were treated with GM-CSF (10 ng/ml) + IL-4 (10 ng/ml) + IFN- (20 ng/ml) + poly(I:C) (100 ng/ml) for 10 h with inclusion of brefeldin A for the last 5 h of culture. Cells were gated according to CD11chighCD11b+ and CD11chighCD11blow/– populations (dot plots) and analyzed to measure intracellular IL-12p40/70 expression (histograms delineating the percentages of positive cells). IL-12p40/p70 expression was not detected in nonstimulated control DC cultures (data not shown). *, p < 0.05 for NOD vs NOR or C57BL/6 DC for a particular treatment.

The PLN are the primary site of autoreactive T cell priming and tolerance induction to islet Ags in NOD mice (37). Comparison of the DC subset composition in the PLN from 6 wk-old NOD and NOR mice revealed that the populations of CD11c^highCD11b^high and CD11c^highCD11b^low/– cells were equivalent between the strains. The percentages of CD11c^highCD11b^high cells were 0.35 ± 0.10 and 0.36 ± 0.13%, and the CD11c^highCD11b^low/– cells comprised 0.25 ± 0.07 and 0.29 ± 0.08% of NOD and NOR PLN, respectively. Analysis of intracellular IL-12 expression was performed by stimulating total PLN-derived cells with GM-CSF plus IL-4 plus IFN- plus poly(I:C) and analyzing the DC populations by flow cytometry. First, we checked that overnight culture of lymphocytes did not affect the detection of DC subsets; we observed no differences in the percentages of DC subsets among total lymphocytes before vs after the 10-h culture (data not shown), nor between poly(I:C)/cytokine-stimulated cultures from NOD and NOR mice (Fig. 4D, dot plots). The CD11c^highCD11b^low/– population from NOD PLN contained reduced percentages of IL-12-expressing cells upon stimulation, and also displayed a reduced intensity of IL-12 staining compared with the corresponding NOR population (Fig. 4D, histograms). In contrast, the NOD/NOR CD11c^highCD11b^high populations did not express IL-12p40/p70. Hence, unlike the splenic DC compartment, the composition of DC subsets was similar in NOD and NOR PLN; however, the reduced IL-12 production by NOD DC was observed in both lymphoid organs.

To assess cytokine production by DC subsets, CD8⁺ and CD8^– DC were FACS-sorted from NOD, NOR, and BALB/c spleens and cultured for 24 h with GM-CSF plus IL-4 plus IFN- plus poly(I:C) or LPS plus CD40L-transfected J558 cells. Significantly, CD8⁺ DC from NOD mice produced reduced quantities of IL-12p70 in comparison to NOR and BALB/c DC in both culture conditions (Fig. 5A). CD8⁺ DC were the major IL-12p70-producing subset, whereas IL-12p70 secretion by CD8^– DC was negligible in response to poly(I:C)/cytokine stimulation. Interestingly, in response to LPS/CD40L stimulation, the CD8^– DC subset produced small quantities of IL-12p70 that were consistently reduced in NOD cultures; however, the difference from control strain DC did not reach statistical significance (Fig. 5A). TNF- was observed to be a major cytokine product of activated CD8^– DC and was produced in comparable quantities by NOD and control strain DC during maturation with poly(I:C)/cytokines (Fig. 5B). Therefore, the cytokine production defect of the NOD DC compartment results from a numerical deficiency of the major IL-12-producing CD8⁺ DC subset combined with a reduced ability of CD8⁺ DC to produce IL-12p70.

View larger version (43K): [in this window] [in a new window]   FIGURE 5. NOD CD8+ DC exhibit reduced IL-12p70 production and increased IL-10 synthesis during maturation. A–C, Comparison of cytokine production by NOD, NOR, and BALB/c DC subsets. FACS-sorted CD8+ and CD8– DC (5 x 105 cells/ml) were cultured for 24 h in GM-CSF (10 ng/ml) + IL-4 (10 ng/ml) + IFN- (20 ng/ml) + poly(I:C) (100 ng/ml) or LPS (100 ng/ml) + CD40L transfected J558 cells (1 DC: 1 J558 cell). IL-12p70 (B), TNF- (C), and IL-10 (D) concentrations were measured by ELISA. D and E, Analysis of the kinetics of cytokine production by MACS-purified CD11c+ cells (5 x 105 cells/ml) from NOD, NOR, and BALB/c spleens. Cells were cultured in GM-CSF + IL-4 + IFN- + poly(I:C) and supernatant samples were acquired after 14, 24, or 40 h of culture for measurement of the quantities of IL-12p70 (D) and IL-10 (E). *, A significant difference between NOD DC vs NOR or BALB/c DC for a particular treatment.

We performed a time-course analysis of IL-12p70 and IL-10 production by MACS-purified CD11c⁺ cells in response to poly(I:C)/cytokine stimulation. Interestingly, there was an inverse correlation between the quantities of IL-12p70 (Fig. 5D) and IL-10 (Fig. 5E) in mature DC cultures; the NOR DC compartment produced increased IL-12p70 and reduced IL-10 levels whereas NOD DC displayed the opposite cytokine bias. The deficit in IL-12p70 production by NOD DC was observed after 14, 24, and 40 h of poly(I:C)/cytokine treatment, indicating that the reduced quantities of IL-12p70 did not result from delayed kinetics of DC maturation in NOD cultures (Fig. 5D). Once again, IL-10 synthesis by NOD DC was noticeably increased in poly(I:C)/cytokine-stimulated cultures; however, a statistically significant difference from control strain cells was only evident after 40 h of culture (Fig. 5E). Taken together, these data reveal an unusual cytokine profile of NOD CD8⁺ DC in response to proinflammatory stimuli, marked by increased IL-10 and reduced IL-12 production.

A limited ability to elicit Th1-polarized responses is associated with reduced IL-12p70 synthesis by NOD CD8⁺ DC

To assess the role of NOD DC in Th1 cytokine polarization, CD8⁺ and CD8^– DC from NOD and NOR spleens were pretreated with poly(I:C)/cytokines for 18 h, washed thoroughly to remove residual cytokines, and were subsequently cultured with syngeneic CD4⁺ T cells and anti-CD3 to stimulate the TCRs. In preliminary experiments, a 1:10 DC-T cell ratio induced optimal IFN- production by T cells in 48 h cultures with 0.5 µg/ml anti-CD3 Ab (in comparison to 1:1, 1:5, or 1:20 ratios; data not shown); therefore, these cellular proportions were used in the subsequent experiments. The cytokine and proliferation data from comparisons of NOD and NOR DC/T cocultures are summarized in Table II. Control cultures containing immature (i.e., freshly purified) CD8⁺ or CD8^– DC as stimulators revealed robust T cell proliferation but minimal IFN- production, verifying that microbial/cytokine stimulation is required for Th1 polarization. Notably, T cell proliferation elicited by immature DC subsets from NOD and NOR mice was equivalent and it was also evident that CD8^– DC were more proficient than CD8⁺ DC at inducing CD4⁺ T cell responses, in agreement with prior studies of T cell stimulation in vitro by DC from nonautoimmune strains (40, 41). In cultures containing poly(I:C)/cytokine-matured CD8^– DC, NOD and NOR cells demonstrated comparable T cell proliferation and IFN- and IL-4 production. Poly(I:C)/cytokine-stimulated CD8^– DC from NOD and NOR mice did not produce IL-12p70, thereby implicating alternate DC-derived cytokines in Th1 induction by this DC subset. Significantly, CD8⁺ DC from NOD mice incited considerably lower levels of IFN- secretion and IL-12p70 production was coordinately reduced (2.5-fold) whereas T cell proliferation and IL-4 concentrations were equivalent in NOD and NOR cultures. Addition of 2.5 ng/ml recombinant murine IL-12p70 to NOD CD8⁺ DC/T cell cultures not only equalized the IL-12p70 concentrations but also augmented the IFN- concentrations to the approximate levels found in NOR CD8⁺ DC/T cell cultures. These findings provide evidence for the adjuvant effect of IL-12p70 on Th1 induction in this assay. In contrast, supplementation of NOR CD8⁺ or NOD/NOR CD8^– DC/T cell cultures with 2.5 ng/ml rIL-12p70 had comparatively modest effects on IFN- production, suggesting that IFN- production in these cultures was already maximized. Altogether, these results delineate a reduced IL-12-producing capability of NOD CD8⁺ DC which correlates with an impaired ability to polarize T cells toward IFN- production.

DC and macrophages are believed to act cooperatively in promoting proinflammatory T cell responses in NOD mice (3). Given their contrasting defects of IL-12 production in this strain, we compared the contributions of these APC types to Th1 priming in NOD and NOR mice. We isolated splenic CD11c⁺ cells using MACS separation and splenic macrophages by depletion of nonmacrophage lineage cells followed by selection based on plastic adherence (refer to Materials and Methods). DC were first matured with poly(I:C)/cytokines (as described in the previous experiments) and macrophages were treated with LPS (1 µg/ml). These distinct stimuli were used for DC and macrophage maturation to elicit maximal IL-12p70 production by each APC type, as determined in preliminary experiments (data not shown). Mature APC were subsequently washed and cultured with syngeneic T cells and soluble anti-CD3. A constant ratio of APC to T cells was maintained (1 APC:10 T cells) but the composition of macrophages and DC within the APC population was varied to assess their relative contributions to T cell activation.

We first verified the cytokine production capabilities of NOD and NOR splenic macrophages. LPS-treated NOD macrophages produced increased quantities of IL-12p40 (Fig. 6A) whereas IFN- production by mature NOD and NOR macrophages was low and equivalent (Fig. 6B). As anticipated, poly(I:C)/cytokine-stimulated DC demonstrated the reverse trend in IL-12p40 production between the strains (Fig. 6A) whereas IFN- was not a product of stimulated DC (Fig. 6B). IL-12p40 synthesis by APC was also considerably augmented in cocultures with T cells (Fig. 6A). The quantities of IL-12p40 and IFN- were increased in cultures of LPS-pretreated NOD macrophages with T cells/anti-CD3, whereas cocultures containing poly(I:C)/cytokine-stimulated NOD DC demonstrated the opposite trend in comparison to NOR cultures (Fig. 6, A and B). When mature DC and macrophages (1 DC: 1 macrophage) were admixed with syngeneic T cells/anti-CD3, the quantities of IL-12p40 and IFN- were increased in NOD vs NOR cultures (Fig. 6, A and B). In contrast, at 10:1 ratios of DC to macrophages, the opposite trend was observed: NOR cells produced increased quantities of IL-12p40 and IFN-. Expectedly, cultures of immature APC (1 DC:1 macrophage) with T cells/anti-CD3 contained low concentrations of IL-12p40 and IFN- which did not differ in NOD vs NOR cultures. T cell proliferation did not differ between NOD and NOR cultures during T cell activation by immature (Fig. 6C) or mature APC types (Fig. 6D) admixed at varying ratios. This finding implies that Th1 biases rather than T cell numbers confer the cytokine production differences between NOD and NOR cultures. Collectively, these experiments demonstrate that activated NOD macrophages are endowed with an enhanced Th1-promoting proclivity whereas the NOD DC compartment is impaired in its capacity to condition CD4⁺ T cells toward IFN- production. NOD macrophages can thus overcompensate for the DC defect in Th1 activation depending on the relative abundance of APC types. This pattern differs for the NOR APC compartment, where the contributions of DC to Th1 differentiation outweigh those of macrophages.

View larger version (29K): [in this window] [in a new window]   FIGURE 6. NOD macrophages have an augmented Th1-priming ability and can compensate for the IL-12 production defect of the NOD DC compartment. NOD and NOR splenic DC and macrophages (MAC) were freshly purified (immature) or were treated with maturation stimuli individually for 18 h before coculture with T cells. DC were pretreated with GM-CSF (10 ng/ml) + IL-4 (10 ng/ml) + IFN- (20 ng/ml) + poly(I:C) (100 ng/ml) and macrophages were matured with LPS (1 µg/ml) for 18 h and were washed thoroughly to remove residual cytokines. Dilutions of immature or mature APC (2.0 x 105/ml) were plated alone or with syngeneic T cells (2.0 x 106/ml) ± anti-CD3 (0.5 µg/ml) in 24-well plates. Supernatants were sampled after 60 h to measure the quantities of IL-12p40 (A) and IFN- (B) by ELISA. C and D, To assess T cell proliferation, dilutions of immature (C) or mature (D) APC (admixed DC and macrophages totaling 1.2 x 105/ml) were plated in the indicated combinations with their respective syngeneic T cells (1.2 x 106/ml) and anti-CD3. Control wells were plated lacking anti-CD3. Thymidine uptake was measured after 72 h of culture and the results were expressed as the mean cpm from triplicate wells ± SD. *, A significant difference between NOD and NOR cultures.

Ag presentation is another hallmark of immunostimulatory DC; however, in NOD mice, the functions of DC in autoantigen presentation remain undefined. T cells reactive against a peptide of GAD65, the immunodominant 524–543 epitope (GADp524), are predominant at 3–4 wk of age during disease initiation in NOD mice (42, 43); therefore, this peptide was used for Ag-presentation studies to assess the activation of islet-reactive T cells by NOD DC in vitro. Ag-specific T cells from GADp524/CFA-immunized NOD and NOR mice were used in crossover combinations with varying numbers of NOD/NOR splenic DC (MACS-purified CD11c⁺ cells) for measurement of T cell proliferation in thymidine incorporation assays. The results revealed that GADp524-specific T cells from NOD mice responded more vigorously than NOR T cells to peptide restimulation, irrespective of the DC genotype and at various DC dilutions (Fig. 7A), suggesting that the differences between the strains were T cell dependent.

View larger version (42K): [in this window] [in a new window]   FIGURE 7. Autoreactive T cell activation by NOD and NOR APC. A, Activation of in vivo-expanded GADp524-reactive T cells by NOD vs NOR DC. GAD65 p524-specific CD4+ T cells were purified from pooled popliteal lymph nodes of NOD and NOR mice that had been immunized with peptide (100 µg/footpad) emulsified in CFA 10 days earlier. CD4+ T cells (2 x 105/well) were cultured in crossover combinations with titrations of MACS-purified DC from the spleens of separate, nonimmunized NOD and NOR mice and 10 µM GADp524 for 96 h. Control wells containing CD4+ T cells plus peptide without DC were included to measure the residual T cell expansion resulting from the in vivo priming. B–D, Activation of spontaneously primed GADp524-reactive T cells by NOD vs NOR APC. B cells (B220) and DC (CD11c+) were MACS-purified from the spleens of 8-wk-old NOD and NOR mice. Dilutions of DC and B cells from NOD and NOR spleens were plated with splenic CD4+ T cells from unprimed NOD mice (6 x 105/well) and GADp524. Control wells contained APC and T cells without peptide. In B, two concentrations of APC (2.5 x 105 or 5.0 x 105 cells/well) were plated with NOD CD4+ T cells ± 10 µM GADp524. In C and D, titrations of GADp524 were plated with DC or B cells (5.0 x 105/well) and splenic CD4+ T cells (7 x 105/well) for measurement of proliferation (B and C) and IFN- production (D) after 5 days and 72 h, respectively. E, Analysis of IFN- production by GAD65-reactive T cells stimulated by DC subsets. FACS-sorted DC subsets (CD11chighCD8+ and CD11chighCD8– cells; 5 x 104 cells/well) from NOD and NOR spleens were cultured with unprimed CD4+ T cells (2 x 105/well) from NOD spleens plus GADp524 (5 µM). Supernatant samples were collected after 48 h of culture. F, MACS-purified NOD/NOR DC or B cells (2 x 105/well) from spleens were plated with NOD CD4+ T cells (2 x 106/well) and GADp524 (5 µM) and cells were harvested after 24 h to analyze expression of I-Ag7 (MHC class II) on APC populations by flow cytometry. [3H]Thymidine incorporation assays (mean cpm ± SD) and ELISA (mean picograms per milliliter ± SD) were performed to analyze proliferation and cytokine concentrations, respectively. *, A significant difference for a particular APC subset from NOD vs NOR mice.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 
Our investigations have uncovered numerical, phenotypic, and functional abnormalities in the APC compartment of NOD mice. This analysis has identified several key distinguishing features of CD8⁺ and CD8^– DC in NOD mice in comparison to autoimmune-resistant strains. First, the composition of DC subsets differs in NOD spleens, marked by a bias toward the CD8^– subset and a deficit of CD8⁺ DC, a difference which was not observed in the PLN of NOD vs NOR mice and therefore suggests that DC numbers are differentially regulated in the steady-state but not during autoreactive T cell activation. BrdU labeling and apoptosis studies indicate that DC turnover and survival do not differ in NOD and control strain spleens, but instead implicate altered differentiation or recruitment of DC precursors in the bias toward the CD8^– DC subset in NOD mice. Second, this study demonstrates that CD8⁺ and CD8^– DC in NOD mice do not harbor gross defects in maturation, as have been ascribed to NOD B cells, macrophages, and in vitro-generated DC. CD8⁺ DC from NOD mice have a diminished ability to synthesize IL-12 but are otherwise functionally normal. In the context of the abundant literature documenting the functional abnormalities of other NOD professional APC types, the present work suggests that the NOD DC compartment exhibits the least functional impairment. To accentuate this point, our comparisons between NOD professional APC types revealed that B cells are the most efficient presenters of autoantigen and that macrophages mediate aberrantly elevated Th1 priming in this strain.

This is one of only a few reports which address the functional capabilities of naturally occurring DC in NOD mice and represents the most comprehensive study to date. Previous studies of NOD DC defects have used in vitro-generated cells owing to their relative homogeneity and availability in large numbers. Studies of clinical T1D have also relied mainly upon in vitro-generated cells and have not yet reached a consensus concerning their functional capabilities. A recent study from our laboratory has addressed this issue by analyzing unmanipulated DC subsets in whole blood samples from patients with T1D (44). Intriguingly, DC from patients with diabetes exhibit impaired IFN- production and modestly reduced 12p70 secretion, although their costimulation and T cell activation abilities are intact. Unlike NOD macrophages, which reproducibly exhibit cytokine and activation abnormalities irrespective of their source (15, 16, 17, 20), we have observed that DC in NOD mice are functionally heterogeneous. This raises the issue of whether studies of in vitro-generated DC can be extrapolated as having functional relevance in vivo. An equivalent population in vivo may be represented by the small subset of blood-derived monocytes that are recruited to lymphoid tissue and undergo DC differentiation during inflammation (45). These monocyte-derived DC codifferentiate with macrophages, possess a high phagocytic ability, and express high levels of MHC class II and costimulatory molecules; therefore, this population closely resembles in vitro-generated DC, which are also monocyte-derived and exhibit GM-CSF-dependent differentiation. In contrast, CD8^– and CD8⁺ DC in the spleen are derived from a resident precursor that is distinct from monocytes (46). Hence, the fact that in vitro-generated DC appear to be developmentally and functionally related to macrophages may underlie their parallel defects in NOD mice.

Our data reveal that IL-12 production by NOD DC is impaired as a result of a numerical deficiency of CD8⁺ DC coupled with a diminished capacity of this subset to synthesize IL-12, as we have demonstrated in vitro in response to two distinct stimulation conditions, during DC-T cell interactions, and in vivo. Moreover, the abnormally low IL-12 production by both spleen and PLN-derived CD8⁺ DC supports the idea that this is an inherent feature of NOD CD8⁺ DC rather than an environmentally programmed one. Our data also suggest that this NOD defect is cytokine specific, although it is presently unclear which cytokine pathways are predominantly used by the CD8^– DC subset to mediate type 1 cytokine responses. For example, a recent study has demonstrated that LPS-activated CD8^– DC can direct IL-12-independent Th1 differentiation through up-regulation of Delta 4, which signals through the Notch receptor on T cells (47). There is presently little information available concerning the unique molecular signatures of CD8⁺ vs CD8^– DC in autoimmune-resistant mice. Interestingly however, CD8⁺ DC have been shown to mediate CD4⁺ T cell apoptosis in vitro, leading to reduced T cell survival and restricted proliferation in comparison to the CD8^– subset (40, 41). We also observed that NOD and control strain CD8^– DC were more effective than CD8⁺ DC at eliciting T cell proliferation during anti-CD3 and GADp524-mediated stimulation, possibly due to reduced T cell survival during activation by the latter population. The diverse activation pathways used by DC subsets to stimulate T cells will therefore be an important area of further study.

The fact that IL-12 up-regulation is uncoupled from the costimulatory capabilities of mature NOD CD8⁺ DC suggests that select branches of the DC maturation program are altered in NOD mice. A signaling mediator which could be responsible for the variable IL-12 production by NOD and control strain DC is MyD88, an adaptor molecule upstream of NF-B which regulates the synthesis of IL-12 family members (48). MyD88 is essential for cytokine production by activated DC but is not required for MHC class II or costimulatory molecule up-regulation (49). IL-10 overproduction may also contribute to the poor IL-12 responses of NOD CD8⁺ DC. IL-10 regulates proinflammatory cytokine production by DC (38) and is a hallmark of tolerogenic DC which activate regulatory T cells and are associated with diabetes protection (50). However, the putative role of DC-derived IL-10 in immune regulation in the NOD mouse requires further clarification because the reduced IL-12 production was not always accompanied by a statistically significant increase in IL-10 production during NOD DC maturation. Aside from their immunogenic roles, studies of the functional capabilities of NOD DC subsets in tolerance induction are also warranted. CD8⁺ DC are highly specialized for tolerance induction, having the ability to restrict IL-2 production by CD8⁺ T cells (51), induce apoptosis of CD4⁺ T cells (40, 41), mediate cross-tolerance to cell-associated Ags (52) and catabolize the amino acid tryptophan, which mediates immune suppression and diabetes protection (53). The reduced CD8⁺ DC numbers in the steady-state could contribute to the loss of self-tolerance in NOD mice. Indeed, T1D can be prevented by adoptive transfer of DC into NOD mice (54, 55, 56) or by therapeutic interventions which mediate increased activity of immature or semimature DC with tolerogenic properties (50, 57, 58, 59). Alternatively, the abnormal cytokine bias of activated CD8⁺ DC from NOD mice may be involved in the diabetes-protective effects conferred by immune stimulation following treatment with microbial products, such as CFA and bacillus Calmette-Guérin (60, 61), or TLR agonists, such as LPS and poly(I:C) (62, 63). In this scenario, DC maturation in NOD mice would lead to a propensity toward tolerogenic, IL-10-driven T cell responses and protection from T1D. Whether immunity or tolerance prevails would therefore depend on the balance of tolerogenic DC vs proinflammatory APC during autoreactive T cell activation.

Although this study does not define the involvement of DC subsets in vivo during the course of disease, our findings are directly relevant to the understanding of the inherent cellular defects that are associated with T1D susceptibility. The Idd4 locus on mouse chromosome 11 has been associated with increased IL-12 synthesis (64), differential expression of genes involved in IFN response pathways (65), and elevated GM-CSF production in NOD mice (66). All of these functions were associated with an increased proclivity toward proinflammatory responses by NOD APC. Identification of these autoimmune-associated phenotypes was based on characterization of macrophages and BM-derived DC; hence, our study illuminates the fact that these abnormalities cannot be extrapolated to all NOD APC. Moreover, because the presentation of NOD DC defects is stimulus specific, our findings argue that NOD DC are fully functional in terms of mediating efficient T cell immunity, with the exception of signals that elicit IL-12-dependent type 1 cytokine polarization. For example, NOD DC are highly effective at inducing expansion and IFN- production by spontaneously primed, GADp524-reactive T cells, an effect which does not require an IL-12-inducing stimulus. It is also noteworthy that neither IL-12 nor its IFN--inducing effects are required for diabetes pathogenesis (67, 68, 69); therefore, the reduced IL-12-producing capability of DC may have little impact on their involvement in autoreactive T cell activation in NOD mice.

Interestingly, although NOD mice with functionally incompetent B cells are diabetes-resistant, they still harbor pathogenic T cells (6, 70). This observation indirectly implicates DC in the initial selection of diabetogenic T cells, but suggests that the subsequent expansion and/or maintenance of autoreactive T cells relies heavily on the B cell compartment. Macrophages are not capable of naive T cell priming and most likely have a role in the amplification of autoreactive T cell responses, particularly given their high IL-12 production. One possible interpretation of our findings is that B cells and macrophages possess a functional advantage among NOD professional APC owing to their aberrantly activated state in the periphery (71) and their hyperactivity during maturation (13, 14), whereas DC are resting in the steady-state and possess a largely normal ability to activate T cells. Importantly, however, the fact that the presentation of NOD DC defects is context-dependent suggests that the relevance of APC functional defects to autoimmunity will depend upon the maturation signals and the local APC composition. Hence, although the importance of one particular APC subset over another is difficult to predict in vivo, DC likely possess complex and subset-specific roles in T cell-mediated autoimmunity.

	Acknowledgments

 
We express gratitude to Thomas Ichim for valuable input and to Edwin Lee-Chan for assistance in preparation of the manuscript.

	Disclosures

Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 
The authors have no financial conflict of interest.

	Footnotes

 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

¹ This work was supported by grants from the Canadian Institutes of Health Research. A.M.M. was a recipient of a doctoral award from the Canadian Institutes of Health Research.

² Current address: Department of Immunology, The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, CA 92037.

³ Current address: Lawson Health Research Institute, London, Ontario, Canada, N6A 4V2.

⁴ Address correspondence and reprint requests to Dr. Bhagirath Singh, Department of Microbiology and Immunology, University of Western Ontario, London, Ontario, Canada, N6A 5C1. E-mail address: bsingh{at}uwo.ca

⁵ Abbreviations used in this paper: T1D, type 1 diabetes; DC, dendritic cell; BM, bone marrow; NOR, NOD resistant; PI, propidium iodide; PLN, pancreatic lymph node; GAD, glutamic acid decarboxylase.

Received for publication July 20, 2007. Accepted for publication February 8, 2008.

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Top Abstract Introduction Materials and Methods Results Discussion Disclosures References

 

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