Myeloid Dendritic Cells in Non-Obese Diabetic Mice have Elevated Costimulatory and T Helper-1-Inducing Abilities

doi:10.1006/jaut.2002.0597, available online at http://www.idealibrary.com on

Journal of Autoimmunity (2002) 19, 23–35

Myeloid Dendritic Cells in Non-Obese Diabetic Mice have Elevated Costimulatory and T Helper-1-Inducing Abilities Annette M. Marleau and Bhagirath Singh Department of Microbiology and Immunology and John P. Robarts Research Institute, University of Western Ontario, London, Ontario, N6A 5C1, Canada

Received 27 December 2001 Accepted 10 June 2002 Key words: dendritic cells, NOD mice, T cells, type 1 diabetes

Type 1 diabetes (T1D) in the non-obese diabetic (NOD) mouse begins with activation of islet-reactive T helper-1 (Th1) cells by dendritic cells (DCs). Since multiple genetic loci contribute to T1D, we evaluated the hypothesis that NOD DCs possess inherent characteristics that contribute to the autoimmune phenotype. When compared to a representative Th1 (C57BL/6) and Th2 (BALB/C) control strain, in vitro generated NOD myeloid DCs matured normally. Functionally, NOD DCs exhibited higher expression of CD80/86 and IL-12 production during stimulation of naı¨ve T cells, even in comparison to C57BL/6 DCs, the prototype strain for vigorous, Th1-biased immunity. These features of NOD DCs translated into aberrantly elevated IFN-
synthesis, enhanced T-cell proliferation, and heightened CD69 expression. Further, NOR DCs, from an NOD-related, autoimmune-resistant strain, did not display this hyper-responsiveness, suggesting that these abnormalities are genetic features of NOD DCs that are related to disease pathogenesis. Cumulatively, these results indicate that NOD DCs are inherently biased towards abnormally high costimulation and Th1-induction, two features that would be expected to confer activation and persistence of autoreactive T cells. © 2002 Elsevier Science Ltd. All rights reserved.

Introduction

hypo-responsiveness of thymocytes to T-cell receptor (TCR) ligation [9, 10]. However, despite emerging insight into the functions of NOD T cells, putative APC-related defects have been less extensively investigated. In NOD mice, particular attention has been focused on the unique MHC class II molecule on APCs, I-Ag7, which demonstrates poor binding of certain peptides, a feature which has been suggested to confer inefficient thymic selection [11–14]. Homozygous I-Ag7 expression is essential but by itself, not sufficient, for development of disease [15], indicating the important contribution of non-MHC loci. NOD bone marrow-derived DCs and macrophages reportedly exhibit defective phenotypic and functional maturation [16–21]. Analogous DC immaturity has been documented in humans with islet autoimmunity [22, 23] as well as in the Biobreeding rat model of T1D [24, 25]. However, not all existing data supports the idea of generalized APC immaturity in diabetes. Both myeloid DCs and macrophages in NOD produce markedly elevated levels of IL-12 in response to maturation stimuli [26–28]. This finding contradicts the supposed immaturity of NOD APCs, as high IL-12 production is a hallmark feature of mature DCs [reviewed in 29]. Recently, a locus associated with human T1D, Idd18, was identified in the region encoding IL-12 p40 [30], highlighting an apparent parallel between DC defects in human diabetes and those in NOD mice. Further,

Type 1 diabetes (T1D) is characterized by emergence of activated, self-reactive T cells that progressively destroy the pancreatic insulin-producing  cells [1]. Disease is initiated by the presentation of self- and/or pathogen-derived antigens by antigen-presenting cells (APCs), in particular, dendritic cells (DCs), which are the first cells to circumscribe the islets [2–4]. A preferential skewing of CD4 + T cells towards an IFN-
and TNF--producing T-helper-1 (Th1) phenotype has been demonstrated to drive the chronic inflammatory process [5–6]. Dysregulation of the Th1/Th2 equilibrium is evident in the murine model of spontaneous T1D, the non-obese diabetic (NOD) mouse, in which a high ratio of IFN-
/IL-4 in the islets is predictive of disease development [7]. In T1D, defective regulation of autoreactive T cells is under polygenic control with multiple genetic loci interacting to produce the autoimmune phenotype [1]. Several of the known lymphocyte defects are manifested at the T-cell level, including impaired development of protective Th2 cells [1], reduced numbers of regulatory CD4 + CD25 + T cells in circulation [8], and Correspondence to: Dr Bhagirath Singh, Department of Microbiology and Immunology, University of Western Ontario, London, Ontario, Canada, N6A 5C1. Tel.: (519) 663-3427; Fax: (519) 661-3499; E-mail: [email protected] 23 0896–8411/02/$-see front matter

© 2002 Elsevier Science Ltd. All rights reserved.

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the immunostimulatory nature of NOD DCs was exemplified by the recent observation that these cells exhibit dysregulated NF-B activation [28], a transcription factor known to regulate IL-12 gene expression [31]. Importantly, information is lacking concerning the activities of NOD DCs in a functional context. The role of DCs in propagation of autoimmunity is of particular significance, since DCs are responsible for naı¨ve T-cell activation [32]. At an immature stage, DCs reside at non-lymphoid locations, where they are specialized for antigen uptake but have low costimulatory molecules [reviewed in 29]. Upon activation by pathogens or inflammatory cytokines, DCs progressively acquire characteristics of mature cells and concomitantly migrate to local lymph nodes. Such maturing DCs are phenotypically and functionally distinct from their precursors, displaying upregulated surface expression of costimulatory molecules, an increased propensity for cytokine synthesis and enhanced antigen presentation capabilities [33]. In vitro, immature DCs can be derived from myeloid progenitors in the presence of GM-CSF and IL-4 [34–36]. Further supplementation of cultures with LPS and/or proinflammatory cytokines leads to the differentiation of phenotypically and functionally mature DCs [37]. Thus, in vitro DC maturation is a useful system for analysis of molecular and functional aspects of DC biology. DC maturation is also elaborated over the course of interactions with T cells [reviewed in 29]. APC functions are enhanced via engagement of CD40 molecules on DCs with CD40 ligand (CD40L) on CD4 + T cells, leading to IL-12 production [38–41]. Additionally, CD8 + T cells provide IFN-
for coinduction of IL-12 [42]. In turn, DCs provide costimulatory signals to T cells and dictate the cytokine microenvironment, thereby influencing Th lineage commitment [reviewed in 29]. CD80 and CD86 molecules on APCs interact with CD28 on T cells, serving to sustain T cell clonal expansion and survival [43], in addition to dictating the magnitude of T-cell activation [44]. In autoimmunity, costimulation via B7/CD28 is essential for productive IL-12 responses and the subsequent differentiation of pathogenic Th1 effector cells [45]. Based on the role of DCs as initiators of islet inflammation, combined with putative abnormalities of this lineage, we evaluated NOD myeloid DC function during interactions with naive T cells by comparison to DCs of non-autoimmune control strains. First, normalcy of NOD DC differentiation was demonstrated in both the immature and LPS/ IFN-
-activated states. In a functional context, two parameters of NOD DC maturation were assessed; the ability to prime naı¨ve T-cell activation/proliferation and Th-skewing capabilities. We herein report that NOD DCs possess elevated costimulatory and IFN-
inducing proclivities in comparison to DCs from a panel of control strains, BALB/C, C57BL/6 and NOR, an NOD-related recombinant strain. Moreover, these exaggerated immune responses were inherently DC-related, rather than representing T-cell-dependent differences. Hyper-responsiveness of

A. M. Marleau and B. Singh

NOD DCs points towards a contribution of DC abnormalities to the persistent and Th1-skewed responses in T1D.

Materials and Methods Mice Female NOD/Lt mice were bred in the animal facility at the John P. Robarts Research Institute (London, Ontario, Canada). C57BL/6, BALB/C and NOR mice were purchased from The Jackson Laboratory (Bar Harbor, ME, USA). For generation of (NOD×C57BL/6) F1 offspring, NOD males were mated with C57BL/6 females and the first generation (F1) females were used for experiments. Mice were maintained under specific pathogen-free conditions in the animal facility at the University of Western Ontario, London, Ontario, Canada. All experiments were performed using female mice between 4–7 weeks of age. For NOD mice, this age period represents the pre-diabetic stage.

DC preparation Bone marrow-derived DCs were prepared using a modification of a previously described protocol [34]. Briefly, bone marrow precursors were harvested by flushing the femurs and tibiae of C57BL/6, BALB/C, NOD and NOR mice using 23-gauge needles. Cells were counted using a haemocytometer and trypan blue exclusion. Cells were suspended in complete medium composed of RPMI (Life Technologies, Gaithersburg, MD, USA) supplemented with 5×10 −5 M 2-ME, 10 mM HEPES, 2 mM glutamine, 5 IU/ml penicillin-streptomycin, and 10% heatinactivated foetal calf serum (FCS) (HyClone Laboratories, Logan, UT, USA). Media was also supplemented with murine recombinant (r)GM-CSF (20 ng/ml; Cedarlane Laboratories, Hornby, Ontario) and murine rIL-4 (10 ng/ml; Cedarlane) for stimulating DC growth. Cells were plated in six-well plates at concentrations of 1–5×106 cells/ml and incubated at 37°C in humidified air with 5% CO2. On the second day of culture, non-adherent cells were discarded cytokineand cultures were supplemented with supplemented media and supernatant. Subsequently, cells were fed every two days by splitting the contents of each well into two and adding volume fresh, cytokine-supplemented media per well. The nonadherent and loosely adherent DC-containing fraction was harvested at days 7–9 of culture. To purify DCs, MACS was used (Miltenyi Biotec, Auburn, CA, USA). Briefly, DCs were positively selected by first staining with either FITC (fluorescein isothiocyanate)-, PE (phycoerythrin)-, or biotin-conjugated anti-CD11c (HL3) antibody (Ab) (BD Pharmingen, San Diego, CA, USA). Cells were then stained with anti-FITC, anti-PE, or anti-biotin Microbeads and run through an LS+ MACS column in a MidiMacs magnet (Miltenyi 3 4

1 4

1 2

T-cell activation by NOD dendritic cells

Biotec). DC purity was typically greater than 96%, as verified by FACS. In some experiments, DCs were activated for 24 h in GM-CSF-containing media (20 ng/ml) supplemented with lipopolysaccharide (LPS; Sigma, St. Louis, MO, USA) at 100 ng/ml and murine rIFN-
(BD Pharmingen) at 100 U/ml. T-cell purification Spleens were aseptically dissected, dissociated, and passaged through a 40 m pore size nylon cell strainer in complete RPMI. Red blood cells were lysed using ACK lysis buffer (Life Technologies). T cells (including both CD4 + and CD8 + cells) were subsequently purified using nylon wool columns. T-cell purity was typically 90%, as verified by FACS using FITC-conjugated anti-TCR  chain Ab (H57-597; BD Pharmingen). CFSE labelling of T cells Purified T cells that were suspended in complete RPMI were washed once in sterile PBS to remove any residual FCS and were subsequently re-suspended in PBS. T cells were then labelled with 5-(and 6-)carboxyfluorescein diacetate succinimidyl ester (CFSE; Molecular Probes, Eugene, OR, USA), as previously described [46]. Briefly, 5×106 T cells/ml in PBS were co-incubated with an equal volume of a 1:250 dilution of 5 mM CFSE stock solution for 10 min at 37°C in the dark. An equal volume of FCS (HyClone Laboratories) was then added to quench excess CFSE. Cells were washed once in RPMI and re-suspended at the desired concentration. Uniform labelling of cells was verified by flow cytometry. DC-T cell co-cultures MACS-purified DCs were cultured with nylon woolpurified T cells and soluble anti-mouse CD3 at 0.5 g/ ml (Cedarlane). Cells were plated in 24-well plates in final volumes of 2 ml RPMI. In various experiments, between 1.5–3 million of each cell type was plated per well. Exogenous cytokines were not added to DC/T cultures. Cells and/or supernatants were then harvested at the appropriate time post-stimulation for flow cytometry and ELISA, respectively. To harvest DCs, non-adherent cells were first collected and trypsin-EDTA (Life Technologies) was then added to adherent cells for 5 min at 37°C followed by gentle pipetting to fully detach cells. For harvesting T cells, non-adherent cells were collected. For collection of supernatant samples, supernatant was separated from cell suspensions by centrifugation and samples were then stored at −80°C before analysis. Flow cytometric analysis Before staining, cells were suspended in PBS and Fc receptors were blocked with 4 l of anti-CD16/32 Ab

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(2.4G2; BD Pharmingen) per 106 cells. Cells were subsequently incubated with 0.5 g per 106 cells of the relevant Ab for 40 min at 4°C. The following antimouse Abs from BD Pharmingen were used: FITC-, PE-, and biotin-conjugated anti-CD11c (HL3), PEconjugated anti-CD80 (16-10A1), PE-conjugated antiCD86 (GL1), PE-conjugated anti-CD40 (HM40-3), FITC-conjugated anti-TCR  chain (H57-597), PE-conjugated anti-CD25 (3C7), and FITC- or PEconjugated anti-CD69 (H1.2F3). Isotype-specific control Abs were also purchased from BD Pharmingen. After staining, cells were washed once in PBS and analysed on a FACScan (Becton Dickinson, Mountain View, CA, USA). Data were analysed with CellQuest software (Becton Dickinson). Cytokine ELISA assays Cytokines in supernatant of DC or DC/T cell co-cultures were measured using commercially available OptEIA kits for IL-12 p70 and IFN-
(BD Biosciences). Assays were performed according to the manufacturer’s instructions. Plates were read using a Bio-Rad ELISA plate reader (Richmond, CA, USA). Results are expressed as picograms per millilitre (pg/ml). Proliferation assays To measure DC and/or T-cell proliferation, the appropriate cell combinations were plated in triplicate in 96-well round-bottom plates in final volumes of 200 l per well containing 200,000 total cells. Plates were incubated for 30 or 72 h at 37°C and cells were pulsed with 1 Ci/well of [3H]-thymidine (NEN-DuPont, Boston, MA, USA) for the last 18 h of incubation. Incorporation of [3H]-thymidine was measured using a liquid scintillation counter (LKB Instruments, Gaithersburg, MD, USA). Statistical analysis Comparison between two means was performed using the Student t-test. P<0.05 was considered significant. This test was used to compare cytokine release and T-cell proliferation by [3H]-thymidine incorporation.

Results Characterization of bone marrow-derived DCs from NOD and non-autoimmune control strains As significant effort has been invested into investigation of T-cell signalling and development in NOD, we assessed the contribution of putative DC anomalies to the aberrant immune responses found in these mice. In initial studies, we generated myeloid

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Figure 1. Comparable phenotype of NOD and control bone marrow-derived DCs. Immature bone marrow-derived DCs were prepared from BALB/C, C57BL/6, and NOD mice and MACS-purified based on CD11c expression (using anti-FITC Microbeads). DCs were then re-plated for an additional 24 h in GM-CSF (20 ng/ml) to maintain immature cells or, alternatively, matured with LPS (100 ng/ml) and IFN-
(100 U/ml). For FACS analysis, FITC-labeled CD11c + cells were gated and analysed for co-expression of CD80, CD86 and CD40 using PE-conjugated Abs. Gated regions were analysed for the percentage of CD11c + cells that co-expressed CD80, CD86 and CD40 (A immature DCs; C mature DCs) and mean fluorescence intensity of staining on the positively stained cells (B immature DCs; D mature DCs). All data are representative of three independent experiments with similar results acquired in each repetition.

DCs from their bone marrow precursors using GM-CSF (20 ng/ml) and IL-4 (10 ng/ml) and evaluated DC maturation based on phenotype. Application of in vitro generated cells allowed the inherent genetic propensities of a pure myeloid DC population to be characterized. NOD DCs were evaluated in parallel with DCs from non-autoimmune control mice, C57BL/6, a prototypical Th1-type strain, and BALB/C, a Th2-biased strain. NOD DCs were found to be comparable to those of controls with respect to expression of CD80, CD86 and CD40 (Figure 1A, percentage of positive cells; Figure 1B, mean fluorescence intensity of staining). On all immature DCs, CD80 expression predominated, whereas CD86 and CD40 expression was minimal, particularly for BALB/C DCs. Costimulatory molecule expression was also characterized on immature DCs that were activated with LPS 100 ng/ml) and IFN-
(100 U/ml) for 24 h. Treatment of DCs with LPS/IFN-
confers the ability for high-level IL-12 production and augments their T-cell activation capabilities [48, 49]. LPS specifically programs DCs for terminal maturation and definitive growth arrest [50]. As anticipated,

maturation was evident by an increased percentage of DCs expressing CD80, CD86 and CD40 (Figure 1C), as well as increased levels of expression of these molecules (Figure 1D). Overall, costimulatory molecule expression by NOD DCs was within the range of controls. Lastly, we compared proliferation of NOD and C57BL/6 DCs. Maturation has been associated with declining proliferative capacity as cells differentiate from CD11c − CD86 − precursors to CD11c + CD86 − immature DCs and finally, to CD11c + CD86 + mature DCs [51]. Bone marrow-derived NOD and C57BL/6 DCs were re-plated for 24 h in the presence of GM-CSF/IL-4 for immature DCs or GM-CSF/IL-4 in combination with LPS/IFN-
for mature DCs (Figure 2). Proliferation of both adherent and nonadherent/loosely adherent populations was assessed. Overall, levels of [3H]-thymidine uptake were low for all DC fractions, suggesting that the cell turnover was low and/or that only a small population of cells proliferated. For immature DCs of both strains, more proliferation was evident in adherent vs non-/semiadherent fractions. Further, LPS/IFN-
-activated,

T-cell activation by NOD dendritic cells

Figure 2. Comparable proliferation of NOD and control DCs. Immature DCs (iDC) from NOD and C57BL/6 were seeded in 96-well round-bottomed plates (200,000 cells/ well) in media supplemented with GM-CSF (20 ng/ml). Alternatively, immature DCs were plated with GM-CSF in combination with LPS (100 ng/ml) and IFN-
(100 U/ml) for assessment of mature DC (mDC). To assess proliferation, cultures were pulsed with [3H]-thymidine for the last 18 h of culture. After a total of 30 h of culture, non-adherent cells were transferred to separate wells and the remaining adherent cells were detached by treatment with 50 l trypsinEDTA per well. Adherent and non-adherent fractions were then assayed separately for proliferation. Results are represented as the mean counts per minute (cpm) of triplicate cultures±SD.

mature cells were virtually non-proliferating. These results conform to the established ontogeny of DCs from murine bone marrow in vitro, which begins with adherent, [3H]-thymidine-incorporating clusters of growing DC and culminates with the release of more mature, non-proliferating and nonadherent cells [34]. No significant differences in proliferative capacity were noted between NOD and C57BL/6 for any of the DC fractions. Thus, it was verified that the CD11c + cells obtained from each strain were of equal purity with respect to contaminating progenitors and/or were at a comparable differentiation stage. Based on these findings, differences in DC potency could then be monitored in T-cell-dependent assays without the confounding influence of variations in DC proliferation/numbers. In summary, this evaluation of myeloid DC maturation stands in dispute of the idea that NOD DCs are developmentally impaired.

Enhanced maturation of NOD DCs during T-cell stimulation To evaluate NOD DC functions, we selected a T-celldependent model system. During cognate DC/T-cell interactions, bi-directional costimulatory signals activate the APC [52–54]. Importantly, T-cell derived signals are both quantitatively and qualitatively different from those conferred by LPS and proinflammatory cytokines alone, with T cells having an enhanced ability to boost DC activation [42]. In the present study, naı¨ve, splenic T cells were nylon woolpurified to include both CD4 + and CD8 + subsets since this affords two distinct sets of signals for optimal DC activation. T cells were co-incubated with syngeneic myeloid DCs and anti-CD3 was added as an antigen

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Figure 3. NOD DCs have elevated expression of CD80 and CD86 during T-cell stimulation. Myeloid DCs (MACSpurified using FITC-conjugated anti-CD11c Ab and antiFITC Microbeads) from NOD, C57BL/6 and BALB/C were cultured with their respective syngeneic, splenic T cells plus anti-CD3 (0.5 g/ml) (1.5×106 DCs and 3×106 T cells per well). After 72 h, cells were harvested for FACS analysis. DCs were gated based on positive staining with FITCconjugated anti-CD11c Ab. DCs were also clearly distinguishable from T cells based on their high forward (FSC-H) and side (SSC-H) scatter properties (refer to representative dot plots showing DC gates). Assessment of positive staining for CD80 and CD86 was done using PE-conjugated Abs. Marker lines on each histogram delineate the positively stained cells with reference to their respective isotype control Abs. The numbers on each histogram represent the percentage of positive cells and mean fluorescence intensity values (below).

surrogate for stimulation of the TCR complex. In this model system, initial T-cell activation would be reliant upon DC-derived costimulation and further elaborated by DC-activating properties of the T cells. In comparison to DC/T-cell combinations from BALB/C and C57BL/6, NOD DCs exhibited substantially elevated CD80 and CD86, as evidenced by both higher percentages of positive cells and mean fluorescence intensity values (Figure 3). Notably, BALB/C DCs exhibited the lowest costimulatory molecule expression during DC/T-cell communication and their viability was also comparatively reduced during the culture period (data not shown). Overall, the profound elevations of costimulatory molecule expression on NOD DCs were suggestive of a functionally hyper-responsive cell. It remained possible that inherent aberrances of NOD T cells contributed to differences in reciprocal activation observed during NOD DC-T-cell interactions. An intrinsically increased propensity towards IFN-
production has been documented in NOD T cells [55], a cytokine that acts as a potent DC activator [56]. The influence of T-cell abnormalities

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Enhanced capacity for Th1 polarization by NOD DCs

Figure 4. Elevated costimulatory molecule expression is an inherent feature of maturing NOD DCs. Immature DCs (MACS-purified using FITC-conjugated anti-CD11c and anti-FITC Microbeads) from NOD and C57BL/6 were cultured with (NOD×C57BL/6) F1 T cells and anti-CD3 (0.5 g/ml) in 24-well plates (1.5×106 DCs and 3×106 T cells per well). After 72 h, cells were harvested and stained for FACS. DCs were gated based on positive staining with FITC-conjugated anti-CD11c and evaluated for expression of CD80 and CD86 using PE-conjugated Abs. Histogram marker lines on each graph delineate the positive cells with reference to isotype controls. Percentage of positive cells and mean fluorescence intensity values are indicated. This experiment was repeated two additional times with comparable results.

was addressed by using a genetically common T cell in co-cultures with NOD and C57BL/6 DCs. Notably, BALB/C DCs were excluded from further experiments based on two premises. First, based on their dissimilarity from NOD and C57BL/6 with respect to costimulatory molecule expression during both LPS/IFN-
and T-cell-dependent activation (Figures 1 and 3). Moreover, it could be extrapolated that these features of BALB/C DCs might contribute to the inherent Th2 predisposition of this animal. On the other hand, NOD and C57BL/6 DCs were both presumed to have inherently elevated APC functions, thus C57BL/6 could serve as a standard for non-autoimmune, Th1-type responses. Splenic T cells from F1 progeny of mated C57BL/6 and NOD mice were cultured with NOD or C57BL/6 DCs (plus anti-CD3), thereby ensuring that T cells would be selected by and thus compatible with both MHC backgrounds. Using this model system, we confirmed the elevated CD80 and CD86 expression by NOD DCs (Figure 4). This finding established that NOD DCs inherently drove their elevated maturation status in a manner independent of NOD T-cell defects.

It was of interest to determine whether elevated maturation status of NOD DCs was correlated with increased Th1 polarization. DC-derived IL-12 p70, produced via CD40 signalling pathways, is essential for development of IFN-
-producing T cells [57], and thus, these cytokines are coordinately upregulated [49]. In the aforementioned experiments using GM-CSF/IL-4 induced DCs co-cultured with T cells and anti-CD3, CD40 expression and IL-12 p70 production were decipherable, but only at suboptimal levels (data not shown). Since these DCs were immature at the establishment of DC/T co-culture, having barely detectable surface CD40 (Figure 1A, B), it is likely that initial CD40-CD40L interactions that are required for IL-12 p70 induction were insufficient. The prevailing view is that CD40-CD40L interactions are required for Th1 induction by DCs that have already been preactivated in a classical manner by a pro-inflammatory Th1 mediator. Thus, to properly evaluate the Th1 cytokine profile induced by NOD DCs, immature DCs from NOD and C57BL/6 were pre-activated with LPS and IFN-
for 24 h and subsequently co-cultured with (NOD× C57BL/6) F1 T cells and anti-CD3. Supernatant samples from activated DC/T-cell cultures were analysed for IL-12 p70 and IFN-
. Both IL-12 p70 (Figure 5A) and IFN-
(Figure 5B) were significantly elevated in co-cultures that contained NOD DCs and T cells. Importantly, LPS/IFN-
-activated NOD DCs in the absence of T cells also exhibited higher IL-12 p70 production than C57BL/6 DCs (Figure 5A). For DCs of both strains, IL-12 p70 production was further augmented by DC/T cell mutual activation. Although both NOD and C57BL/6 DCs uniformly expressed CD40 during communications with T cells, expression levels were higher on NOD DCs (Figure 5C). These experiments demonstrate that several qualitatively distinct signals are altered in NOD DCs, including their T cell stimulatory activity (CD80/86-CD28 pathways) and their Th1-driving ability (CD40-mediated signalling). Increased T-cell proliferation is induced by NOD DCs To demonstrate the functional outcome of enhanced NOD DC maturation, (NOD×C57BL/6) F1 T-cell proliferation induced by NOD and C57BL/6 DCs was assessed by [3H]-thymidine incorporation. In preliminary experiments, the kinetics of proliferation was established (data not shown). For all strains, proliferation was still minimal at 48 h of co-culture with both DC types, but was maximal by 72–96 h. Using NOD DCs, maximal T-cell proliferation was detected using both 1:1 and 1:10 DC/T-cell ratios, with no statistically significant difference between these two DC concentrations (Figure 6A). Moreover, although 1:10 DC/Tcell ratios were sufficient using NOD DCs, 1:1 ratios were required to maximize proliferation in the

T-cell activation by NOD dendritic cells

Figure 5. Increased Th1 cytokine production is induced by NOD DCs. Immature C57BL/6 and NOD DCs were preactivated with GM-CSF (10 ng/ml) in combination with LPS (100 ng/ml) and IFN-
(100 U/ml) to induce DC maturation. After 24 h, DCs were washed twice in RPMI and co-cultured with (NOD×C57BL/6) F1 T cells and anti-CD3 (0.5 g/ml) (1.5×106 DCs and 3×106 T cells per well). Additionally, untreated naı¨ve T cells (3×106 per well) and immature DCs (1.5×106 per well) in GM-CSF and LPS/ IFN-
-treated media were plated. Supernatant samples were collected after 16 h for T cells and DCs plated individually and after 72 h for DC/T-cell co-cultures. Assays were done for the presence IL-12 p70 (A) and IFN-
(B) using ELISA. Results are represented as picograms per millilitre (pg/ml) of cytokine in triplicate cultures±SD. In A: *P<0.01, in B: *P<0.05. In C, cells from DC/T-cell co-cultures were double stained for FACS analysis using FITC- and PE-conjugated isotype control Abs (upper panels) or FITC-conjugated anti-CD11c plus PE-conjugated anti-CD40 (lower panels). Numbers indicate the percentage of positive cells in each quadrant and, in brackets of upper right quadrants, the mean fluorescence intensity values for CD40 expression on CD11c + CD40 + cells.

presence of C57BL/6 DCs, and their potency was significantly reduced at 1:10 ratios. As expected, reducing the DC number per well from a 1:10 to 1:100 DC/T-cell ratio resulted in lower overall proliferation due to a reduced availability of costimulation on a per

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Figure 6. Enhanced maturation status of NOD DCs is associated with higher T-cell proliferation. In A, immature NOD and C57BL/6 DCs were cultured with (NOD×C57BL/ 6) F1 T cells (plus anti-CD3; 0.5 g/ml). T cells were plated at 1×105 per well in 96-well round-bottomed plates and DCs were added at 1×105, 1×104 or 1×103 per well to make 1:1, 1:10 or 1:100 DC/T-cell ratios, respectively. As controls, DCs or unstimulated T cells were plated individually. Proliferation was assayed by [3H]-thymidine uptake after an 18 h pulse and a total culture time of 96 h. Results represent the mean counts per minute (cpm) of triplicate cultures±SD. *P>0.05, **P<0.01, ***P<0.05. In B, NOD and C57BL/6 DCs were co-cultured with (NOD×C57BL/6) F1 T cells (plus anti-CD3; 0.5 g/ml) that had been pre-labeled with CFSE. After 96 h, the division profiles were monitored by FACS analysis. Open histogram curves represent the non-divided profile of naı¨ve unstimulated F1 T cells alone (negative controls). The filled histograms (black) depict the division profile of gated T cells (TCR  chain + cells) that were cultured with anti-CD3 (0.5 g/ml) and either NOD or C57BL/6 DCs (activated T cells). Histogram marker lines demarcate the non-dividing T-cell populations and percentages indicate the proportion of DC/anti-CD3-stimulated T cells that remained undivided. The mean fluorescence intensity (MFI) for all T cells is also displayed. Each of these experiments was repeated twice, with similar results obtained in each trial.

T cell basis. At 1:100, NOD DCs were still the more effective stimulators of proliferation. Elevated proliferation measured using [3H]thymidine incorporation may be attributable to either an increased number of cycling cells and/or more successive rounds of division per stimulated cell. To distinguish between these alternatives, F1 T cells were stained with CFSE prior to co-culture with DCs. There were no differences in the number of divisions that

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F1 T cells underwent in the presence of NOD vs C57BL/6 DCs, as evidenced by the similarities in fluorescence intensity at the advanced division peaks (Figure 6B). However, a higher percentage of F1 T cells divided under the influence of NOD vs C57BL/6 DCs, as demonstrated by a lower overall mean fluorescence intensity of the CFSE label and an increased percentage of cycling T cells. This finding could only be attributed to DC differences, since the F1 T cells were purified from the same batch of pooled spleens. Overall, the proliferation experiments illustrated that elevated costimulatory molecule expression by NOD DCs translates into enhanced T-cell activation.

Phenotypic differences of T cells in the presence of NOD DCs To further evaluate T-cell activation by NOD DCs, co-cultured (NOD×C57BL/6) F1 T cells were monitored for expression of two early activation markers, CD25 ( chain of the high affinity IL-2 receptor) and CD69 (Figure 7). Expression of CD25 on TCR  chain + T cells was comparable in the presence of NOD and C57BL/6 DCs, suggesting that the increased proliferation afforded by NOD DCs was not due to differences in T-cell responsiveness to IL-2. However, CD69 expression levels were markedly elevated on TCR  chain + cells stimulated by NOD vs C57BL/6 DCs. CD69, a molecule with costimulatory properties, is important in early T-cell activation and proliferation [58]. The finding that select costimulatory capabilities of T cells are elevated in the presence of NOD DCs provides a likely explanation for enhanced DC maturation that ensues.

Hyper-responsiveness of NOD DCs can be correlated with progression to autoimmunity To correlate the previous findings with the development of autoimmunity, we compared NOD DCs to those of the MHC-syngeneic strain, NOR. This recombinant strain has a proportion of NOD alleles but is diabetes-resistant [59, 60]. Maturing bone marrowderived DCs from NOD and NOR were compared phenotypically during interactions with T cells of (NOD×C57BL/6) F1 origin. Elevated CD80 and CD86 expression levels were confirmed to be features specific to NOD DCs that was not shared by NOR (Figure 8A; refer to mean fluorescence intensity values). Furthermore, the elevated maturation status of NOD DCs was correlated with an enhanced activation phenotype of (NOD×C57BL/6) F1 T cells (Figure 8B). Although CD25 expression was comparable using NOD and NOR DCs, expression levels of CD69 were substantially elevated in the presence of NOD DCs. Based on phenotype, it was concluded that expression levels of activation markers on NOR DCs and their respective T cells were comparable to those found in the control C57BL/6 DC/T-cell cultures (Figures 4 and 7).

Figure 7. Phenotype of T cells activated by NOD vs C57BL/6 DCs. DCs were purified from bone marrow cultures using biotin-conjugated anti-CD11c and anti-biotin Microbeads. NOD and C57BL/6 DCs (1.5×106 cells/well) were cultured with (NOD×C57BL/6) F1 T cells (3×106 cells/well) and anti-CD3 (0.5 g/ml). Cells were harvested for FACS analysis after 36 h of culture to monitor expression levels of early activation markers on T cells. For FACS, instrument settings were optimized for T cells, which were gated based on positive staining with FITC-conjugated anti-TCR  chain Ab. These gated regions (circled on dot plots) had slightly different forward and side scatter properties where NOD and C57BL/6 DCs were used. T cells were analysed for expression of activation markers, CD25 and CD69, using FITC-conjugated Abs (histograms with dark outlines) or isotype control (hatched histograms). Marker lines on histograms indicate the positively stained population relative to isotype control staining. The numbers provided represent the percentage of positive cells and mean fluorescence intensities (below). Two additional repetitions of these experiments gave comparable results.

To characterize NOR DCs with respect to cytokine production, ELISAs for IL-12 p70 were done. In comparison to NOD DCs, NOR cells exhibited a 50% lower capacity for IL-12 p70 production in 72-h culture with (NOD×C57BL/6) F1 T cells (Figure 9A). At a functional level, elevated IL-12 p70 translated into a requirement for fewer NOD DCs in order to attain a maximal IFN-
response (Figure 9B). In titration experiments, wherein DC/T ratios between 10:1 and 1:100 were prepared, ten-fold fewer NOD than NOR DCs were required for maximal IFN-
production. Further, the maximum attainable IFN-
concentrations were reduced with NOR DCs. Overall, these experiments demonstrate that the NOD genotype is specifically affected by an innate hyper-responsiveness of DCs with respect to both costimulatory molecules and Th1 polarization. Additionally, the correlation

T-cell activation by NOD dendritic cells

Figure 8. Elevated costimulatory molecule expression by NOD DCs is not a feature of DCs from autoimmuneresistant NOR mice. Immature bone marrow-derived DCs were generated from NOR and NOD mice and MACSpurified using biotin-conjugated anti-CD11c Ab and antibiotin Microbeads. To assess DC activation, DCs from each strain (2×106 cells/well) were cultured with (NOD×C57BL/ 6) F1 T cells (4×106 cells/well) and anti-CD3. After 72 h, cells were stained with PE-conjugated Abs against CD11c, CD80, and CD86. For FACS analysis, the DC population was gated based on CD11c expression and high forward/side scatter properties. Histogram marker lines on each graph delineate the positively stained cells with reference to isotype controls. Numbers on histograms denote the percentage of positive cells and mean log fluorescence intensity values (A). To assess T-cell phenotype, co-cultures were established, as described above, with the exception that cells were harvested at 36 h for FACS analysis. Cells were double stained to measure markers of T-cell activation (B). Upper panels: FITC- and PE-conjugated isotype control Abs; Center panels: FITC-conjugated anti-TCR  chain plus PE-conjugated anti-CD25; Lower panels: FITC-conjugated anti-TCR  chain plus PE-conjugated anti-CD69. Numbers in quadrants represent the percentage of positive cells and mean fluorescence intensity values for CD25 or CD69 expression on double positive cells (in brackets of upper right quadrants). Data for each of A and B are representative of at least two independent experiments with similar results obtained in each repetition.

between control levels of DC function and diabetesresistance in NOR suggests that these NOD-specific aberrances are relevant to T1D pathogenesis.

31

Figure 9. Elevated Th1 cytokine production is a feature of NOD, but not NOR, DCs. Immature DCs were generated from NOR and NOD mice. To induce maturation, cells were re-plated for 24 h with GM-CSF (20 ng/ml), LPS (100 ng/ ml), and IFN-
(100 U/ml). To assay DC cytokine production in the presence of T cells, NOR and NOD DCs (1.5×106 cells/well) were cultured with (NOD×C57BL/6) F1 T cells (3×106) and anti-CD3 (0.5 g/ml). After 72 h, supernatant samples were collected to measure IL-12 p70 production using ELISA (A). *P<0.05. To measure T-cell cytokine responses, titrated numbers of NOD and NOR DCs were cultured with a fixed number of (NOD×C57BL/6) F1 T cells (2×106 cells/well) and anti-CD3 (0.5 g/ml). The final DC/T ratios used were 10:1, 1:1, 1:10, and 1:100. After 72 h, supernatant samples were harvested for ELISA assays for IFN-
(B). *P<0.05, **P>0.05. All ELISA results are expressed as picograms per millilitre (pg/ml) of cytokine in triplicate wells±SD.

Discussion Given the importance of cognate DC-T-cell interactions in autoimmunity, this study has specifically addressed both the costimulatory capabilities of NOD DCs as well as their Th1 polarizing abilities. A common consensus has been that NOD DCs are phenotypically and functionally immature [18–21]. In contrast, we found NOD DCs to be phenotypically comparable to those of control strains in both the immature and LPS/IFN-
-activated state (Figure 1). Importantly, however, maturation of DCs in the presence of T cells produced quite different results. At the more immature stage (GM-CSF and IL-4 generated cells), substantially elevated CD80 and CD86 expression was noted on NOD DCs during bi-directional activation with T cells and anti-CD3 (Figures 3 and 4). T-cell activation was coordinately upregulated, including proliferation (Figure 6) and

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CD69 expression (Figure 7). Although CD80 and CD86 are reliable indicators of DC maturation status, these findings do not exclude the possibility that other costimulatory interactions between ligands and their receptors are differentially regulated by NOD DCs. Moreover, when NOD DCs were primed with LPS/IFN-
prior to culture with T cells, their CD40 expression and IL-12 p70 production were exaggerated compared to cultures containing DCs from nonautoimmune control strains (Figures 5 and 9). The apparent discordance between LPS/IFN-
vs T-celldriven maturation likely reflects the divergent gene expression patterns that can be induced in maturing DCs by different stimuli [50]. Maturation of DCs using LPS promotes abrupt and terminal maturation. In contrast, DC maturation in vivo is a continuous and coordinated process mediated by DC-T-cell interactions [reviewed in 29]. This study demonstrates that NOD DCs have an inherent genetic ability to stimulate elevated immune responses. At the level of DC/T interactions, these findings do not concur with the prevailing view that APC immaturity, specifically, in the non-B cell compartment, accounts for the blunted T-cell proliferative responses that have been described in NOD [46]. In vivo, this defect is believed to translate into a failure of T-cell susceptibility to activation-induced cell death and ultimately, long-term persistence of autoreactive cells. Importantly, Delovitch and colleagues have demonstrated unresponsiveness of NOD T lineage cells to TCR ligation [9]. Further, NOD T cells have inherent apoptosis resistance [61–64]. These T-cell abnormalities are themselves consistent with the contention that T-cell responses are dysregulated. Taken in combination with the present data, we propose that DC and T-cell defects work in parallel to produce an autoimmune phenotype. Conceivably, DC-mediated hyper-stimulation of a T-cell population that is AICD-resistant leads to the long-term persistence of Th1-type, autoreactive cells. Evidence has accumulated to support the notion that genetically disparate immune responses between Th1 and Th2-predisposed mice are related to inherent differences in APCs. Macrophages from C57BL/6 and BALB/C have distinct metabolic programs that differentially regulate the bias towards Th1 and Th2 responses, respectively [65]. In animal models of infection with intracellular pathogens, DCs of Th2biased BALB/C mice are suggested to be inherently inferior producers of IL-12 and IL-15 compared to those of C57BL/6, conferring lower IFN-
responses and disease susceptibility [66, 67]. DC maturation, in particular, elevated costimulatory molecule expression, has been associated with both prolonged T-cell responses and induction of more strongly Th1-biased immunity [reviewed in 29]. Agents that inhibit maturation of myeloid DCs, such as IL-10, corticosteroids and prostaglandins, convert DCs from Th1- to Th2skewing cells. In the present study, C57BL/6 DCs, from a strain that is Th1-biased, displayed high costimulatory abilities, whereas DCs of BALB/C were comparatively immature and also underwent apoptosis prematurely in vitro (data not shown).

A. M. Marleau and B. Singh

Based on comparison to these strains, NOD DCs were clearly at the extreme Th1 pole of this spectrum. However, it is not yet established whether interactions between NOD DCs and T cells are of prolonged duration. Protracted DC-driven responses would allow a greater proportion of T cells to be engaged and activated. Indeed, the present results are consistent with an increased number of dividing T cells in the presence of NOD DCs (Figure 6B) as well as elevated expression of the CD69 costimulatory molecule on T cells (Figures 7 and 8). Arrest of NOD DCs at a highly immunostimulatory stage could also account for the elevated IL-12 p70 and IFN-
production in DC/T co-cultures containing NOD DCs (Figures 5 and 8). IL-12 production by DCs is normally downregulated at late stages of maturation, accompanied by a transition to IL-10 production and preferential priming of Th2 and non-polarized T cells [68]. In turn, autocrine IL-10 also modulates surface expression of MHC and costimulatory molecules, thereby limiting both T-cell activation and Th1 polarization [69]. Development of destructive autoimmunity is associated with prolonged antigen presentation [70]. Both a prolonged half-life of MHC/peptide complexes in addition to a minimal antigen dose are critical parameters for progression to disease. In autoimmune-prone mice, the presence of ongoing immune responses is exemplified by the formation of spontaneous germinal centers [71]. In the context of the present study, these events are likely a sequel to hyperactivity of the NOD APC compartment. The relevance of the present findings to development of overt disease are exemplified by the observation that NOR DCs functionally resemble those of other nonautoimmune controls, rather than NOD. Despite the fact that NOR mice develop a mild inflammatory infiltrate at the periphery of the islets, minimal T-cell infiltration ensues [60]. As the potentially deleterious responses in NOR are regulated at the level of antigen presentation, prior to involvement of T cells, this suggests that the susceptibility loci that differ between NOR and NOD genomes are related to APC functions. Activation of transcription by NF-B is responsible for coordinated upregulation of MHC expression, costimulation, and cytokine synthesis [72]. Recently, Tisch’s group demonstrated an elevation of IL-12 production by NOD DCs that was dependent upon persistent hyperactivation of NF-B [28, 73]. Although the specifics of the affected intracellular signalling pathways have not been dissected, the present data suggests that several upstream, maturation-related signalling pathways that converge on NF-B might be involved. At a functional level, it is clear that aberrances of NOD DCs have several possible contributions to autoimmuity: (1) elevated costimulation which is accompanied by the potential to drive more potent T-cell activation and (2) exaggerated Th1 polarization that confers an increased propensity for autoreactivity. Cumulatively, these features of NOD DCs could be responsible for the hyperactivity of self-reactive T cells in T1D.

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Acknowledgements The authors would like to thank Thomas Ichim for helpful suggestions and review of this manuscript. This work was supported by research grants from the Juvenile Diabetes Foundation (JDRF) and the Canadian Institutes of Health Research (CIHR). A.M. is the recipient of graduate studentship awards from CIHR and the Natural Sciences and Engineering Research Council of Canada (NSERC).

References 1. Delovitch T.L., Singh B. 1997. The nonobese diabetic mouse as a model of autoimmune diabetes: immune dysregulation gets the NOD. Immunity 7: 727–738 2. Jasen A., Homo-Delarche F., Hooijkas H., Leenen P.J., Dardenne M., Drexhage H.A. 1994. Immunohistochemical characterization of monocyte-macrophages and dendritic cells involved in the initiation of insulitis and beta cell destruction in NOD mice. Diabetes 43: 667–675 3. Rosmalen J.G., Homo-Delarche F., Durant D., Kap M., Leenen P.J., Drexhage H.A. 2000. Islet abnormalities associated with an early influx of dendritic cells and macrophages in NOD and NODscid mice. Lab. Invest. 80: 769–777 4. Shinomiya M., Nadano D., Shinomiya H., Onji M. 2000. In situ characterization of dendritic cells occurring in the islets of nonobese diabetic mice during the development of insulitis. Pancreas 20: 290–296 5. Rabinovitch A. 1994. Immunoregulatory and cytokine imbalances in the pathogenesis of IDDM. Therapeutic intervention by immunostimulation? Diabetes 43: 613–621 6. Liblau R.S., Singer S.M., McDevitt H.O. 1995. Th1 and Th2 CD4 + T cells in the pathogenesis of organ-specific autoimmune diseases. Immunol. Today 16: 34–38 7. Fox C.J., Danska J.S. 1997. IL-4 expression at the onset of islet inflammation predicts nondestructive insulitis in nonobese diabetic mice. J. Immunol. 158: 2414–2424 8. Salomon B., Lenschow D.J., Rhee L., Ashourian N., Singh B., Sharpe A., Bluestone J.A. 2000. B7/CD28 costimulation is essential for the homeostasis of the CD4 + CD25 + immunoregulatory T cells that control autoimmune diabetes. Immunity 12: 431–440 9. Zipris D., Lazarus A.H., Crow A.R., Hadzija M., Delovitch T.L. 1991. Defective thymic T cell activation by concanavalin A and anti-CD3 in autoimmune nonobese diabetic mice: evidence for thymic T cell anergy that correlates with the onset of insulitis. J. Immunol. 146: 3763–3771 10. Bergman M.L., Penha-Goncalves C., Lejon K., Holmberg D. 2001. Low rate of proliferation in immature thymocytes of the non-obese diabetic mouse maps to the Idd6 diabetes susceptibility region. Diabetologia 44: 1054–1061 11. Stratmann T., Apostolopoulos V., Mallet-Designe V., Corper A.L., Scott C.A., Wilson I.A., Kang A.S., Teyton L. 2000. The I-Ag7 MHC class II molecule linked to murine diabetes is a promiscuous peptide binder. J. Immunol. 165: 3214–3225 12. Latek R.R., Suri A., Petzold S.J., Nelson C.A., Kanagawa O., Unanue E.R., Fremont D. 2000.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

Structural basis of peptide binding and presentation by the type I diabetes-associated MHC class II molecule of NOD mice. Immunity 12: 699–710 Moustakas A.K., Routsias J., Papadopoulos G.K. 2000. Modelling of the MHC II allele I-A(g7) of NOD mouse: pH-dependent changes in specificity at pockets 9 and 6 explain several of the unique properties of this molecule. Diabetologia 43: 609–624 Ridgway W.M., Ito H., Fasso M., Yu C., Fathman C.G. 1998. Analysis of the role of variation of major histocompatibility complex class II expression on nonobese diabetic (NOD) peripheral T cell response. J. Exp. Med. 188: 2267–2275 Wicker L.S., Todd J.A., Peterson L. 1995. Genetic control of autoimmune diabetes in the NOD mouse. Annu. Rev. Immunol. 13: 179–200 Serreze D.V., Gaedeke J.W., Leiter E.H. 1993. Hematopoietic stem-cell defects underlying abnormal macrophage development and maturation in NOD/Lt mice: defective regulation of cytokine receptors and protein kinase C. Proc. Natl Acad. Sci. 90: 9625–9629 Serreze D.V., Gaskins H.R., Leiter E.H. 1993. Defects in the differentiation and function of antigen presenting cells in NOD/Lt mice. J. Immunol. 150: 2534–2543 Morel P.A., Vasquez A.C., Feili-Hariri M. 1999. Immunobiology of DC in NOD mice. J. Leukoc. Biol. 66: 276–280 Strid J., Lopes L., Marcinkiewicz J., Petrovska L., Nowak B., Chain B.M., Lund T. 2001. A defect in bone marrow derived dendritic cell maturation in the nonobesediabetic mouse. Clin. Exp. Immunol. 123: 375–381 Feili-Hariri M., Morel P.A. 2001. Phenotypic and functional characteristics of BM-derived DC from NOD and non-diabetes-prone strains. Clin. Immunol. 98: 133–142 Dahlen E., Hedlund G., Dawe K. 2000. Low CD86 expression in the nonobese diabetic mouse results in the impairment of both T cell activation and CTLA-4 up-regulation. J. Immunol. 164: 2444–2456 Jansen A., van Hagen M., Drexhage H.A. 1995. Defective maturation and function of antigen-presenting cells in type 1 diabetes. Lancet 345: 491–492 Takahashi K., Honeyman M.C., Harrison L.C. 1998. Impaired yield, phenotype, and function of monocyte-derived dendritic cells in humans at risk for insulin-dependent diabetes. J. Immunol. 161: 2629–2635 Delemarre F.G., Simons P.J., de Heer H.J., Drexhage H.A. 1999. Signs of immaturity of splenic dendritic cells from the autoimmune prone biobreeding rat: consequences for the in vitro expansion of regulator and effector T cells. J. Immunol. 162: 1795–1801 Delemarre F.G., Hoogeveen P.G., De Haan-Meulman M., Simons P.J., Drexhage H.A. 2001. Homotypic cluster formation of dendritic cells, a close correlate of their state of maturation. Defects in the biobreeding diabetes-prone rat. J. Leukoc. Biol. 69: 373–380 Alleva D.G., Pavlovich R.P., Grant C., Kaser S.B., Beller D.I. 2000. Aberrant macrophage cytokine production is a conserved feature among autoimmune-prone mouse strains: elevated interleukin (IL)-12 and an imbalance in tumor necrosis factor-alpha and IL-10 define a unique cytokine profile in macrophages from young nonobese diabetic mice. Diabetes 49: 1106–1115

34

27. Alleva D.G., Johnson E.B., Wilson J., Beller D.I., Conlon P.J. 2001. SJL and NOD macrophages are uniquely characterized by genetically programmed, elevated expression of the IL-12(p40) gene, suggesting a conserved pathway for the induction of organ-specific autoimmunity. J. Leukoc. Biol. 69: 440–448 28. Weaver D.J., Poligone B., Bui T., Abdel-Motal U.M., Baldwin A.S., Tisch R. 2001. Dendritic cells from nonobese diabetic mice exhibit a defect in NF-kappab regulation due to a hyperactive I kappa B kinase. J. Immunol. 167: 1461–1468 29. Banchereau J., Briere F., Caux C., Davoust J., Lebecque S., Liu Y.J., Pulendran B., Palucka K. 2000. Immunobiology of dendritic cells. Annu. Rev. Immunol. 18: 767–811 30. Morahan G., Huang D., Ymer S.I., Cancilla M.R., Stephen K., Dabadghao P., Werther G., Tait B.D., Harrison L.C., Colman P.G. 2001. Linkage disequilibrium of a type 1 diabetes susceptibility locus with a regulatory IL12B allele. Nat. Genet. 27: 218–221 31. Yoshimura S., Bondeson J., Brennan F.M., Foxwell B.M., Feldmann M. 2001. Role of NFkappaB in antigen presentation and development of regulatory T cells elucidated by treatment of dendritic cells with the proteasome inhibitor PSI. Eur. J. Immunol. 31: 1883–1893 32. Banchereau J., Steinman R.M. 1998. Dendritic cells and the control of immunity. Nature 392: 245–252 33. Sallusto F., Lanzavecchia A. 1999. Mobilizing dendritic cells for tolerance, priming, and chronic inflammation. J. Exp. Med. 189: 611–614 34. Inaba K., Inaba M., Romani N., Aya H., Deguchi M., Ikehara S., Muramatsu S., Steinman R.M. 1992. Generation of large numbers of dendritic cells from mouse bone marrow cultures supplemented with granulocyte/macrophage colony-stimulating factor. J. Exp. Med. 176: 1693–1702 35. Scheicher C., Mehlig M., Zecher R., Reske K. 1992. Dendritic cells from mouse bone marrow: in vitro differentiation using low doses of recombinant granulocyte-macrophage colony-stimulating factor. J. Immunol. Methods 154: 253–264 36. Sallusto F., Lanzavecchia A. 1994. Efficient presentation of soluble antigen by cultured human dendritic cells is maintained by granulocyte/macrophage colony-stimulating factor plus interleukin 4 and downregulated by tumor necrosis factor alpha. J. Exp. Med. 179: 1109–1118 37. Winzler C., Rovere P., Rescigno M., Granucci F., Penna G., Adorini L., Zimmermann V.S., Davoust J., Ricciardi-Castagnoli P. 1997. Maturation stages of mouse dendritic cells in growth factor-dependent long-term cultures. J. Exp. Med. 185: 317–328 38. Caux C., Massacrier C., Vanbervliet B., Dubois B., Van Kooten C., Durand I., Banchereau J. 1994. Activation of human dendritic cells through CD40 cross-linking. J. Exp. Med. 180: 1263–1272 39. Schoenberger S.P., Toes R.E., van der Voort E.I., Offringa R., Melief C.J. 1998. T-cell help for cytotoxic T lymphocytes is mediated by CD40-CD40L interactions. Nature 393(6684): 480–483 40. Bennett S.R., Carbone F.R., Karamalis F., Flavell R.A., Miller J.F., Heath W.R. 1998. Help for cytotoxic-T-cell responses is mediated by CD40 signaling. Nature 393(6684): 478–480

A. M. Marleau and B. Singh

41. Ridge J.P., Di Rosa F., Matzinger P. 1998. A conditioned dendritic cell can be a temporal bridge between a CD4 + T-helper and a T-killer cell. Nature 393(6684): 474–478 42. Mailliard R.B., Egawa S., Cai Q., Kalinska A., Bykovskaya S.N., Lotze M.T., Kapsenberg M.L., Storkus W.J., Kalinski P. 2002. Complementary dendritic cell-activating function of CD8+ and CD4+ T cells: helper role of CD8+ T cells in the development of T helper type 1 responses. J. Exp. Med. 195: 473–483 43. Salomon B., Bluestone J.A. 2001. Complexities of CD28/B7: CTLA-4 costimulatory pathways in autoimmunity and transplantation. Annu. Rev. Immunol. 19: 225–252 44. Schweitzer A.N., Borriello F., Wong R.C., Abbas A.K., Sharpe A.H. 1997. Role of costimulators in T cell differentiation: studies using antigen-presenting cells lacking expression of CD80 or CD86. J. Immunol. 158: 2713–2722 45. Chang J.T., Segal B.M., Shevach E.M. 2000. Role of costimulation in the induction of the IL-12/IL-12 receptor pathway and the development of autoimmunity. J. Immunol. 164: 100–106 46. Noorchashm H., Moore D.J., Noto L.E., Noorchashm N., Reed A.J., Reed A.L., Song H.K., Mozaffari R., Jevnikar A.M., Barker C.F., Naji A. 2000. Impaired CD4 T cell activation due to reliance upon B cell-mediated costimulation in nonobese diabetic (NOD) mice. J. Immunol. 165: 4685–4696 47. Mueller D.L., Jenkins M.K., Schwartz R.H. 1989. Clonal expansion versus functional clonal inactivation: a costimulatory signaling pathway determines the outcome of T cell antigen receptor occupancy. Annu. Rev. Immunol. 7: 445–480 48. Snijders A., Kalinski P., Hilkens C.M., Kapsenberg M.L. 1998. High-level IL-12 production by human dendritic cells requires two signals. Int. Immunol. 10: 1593–1598 49. Hilkens C.M., Kalinski P., de Boer M., Kapsenberg M.L. 1997. Human dendritic cells require exogenous interleukin-12-inducing factors to direct the development of naive T-helper cells toward the Th1 phenotype. Blood 90: 1920–1926 50. Granucci F., Vizzardelli C., Virzi E., Rescigno M., Ricciardi-Castagnoli P. 2001. Transcriptional reprogramming of dendritic cells by differentiation stimuli. Eur. J. Immunol. 31: 2539–2546 51. Morelli A.E., Zahorchak A.F., Larregina A.T., Colvin B.L., Logar A.J., Takayama T., Falo L.D., Thomson A.W. 2001. Cytokine production by mouse myeloid dendritic cells in relation to differentiation and terminal maturation induced by lipopolysaccharide or CD40 ligation. Blood 98: 1512–1523 52. Kitajima T., Caceres-Dittmar G., Tapia F.J., Jester J., Bergstresser P.R., Takashima A. 1996. T cell-mediated terminal maturation of dendritic cells: loss of adhesive and phagocytotic capacities. J. Immunol. 157: 2340–2347 53. Peng X., Kasran A., Warmerdam P.A., de Boer M., Ceuppens J.L. 1996. Accessory signaling by CD40 for T cell activation: induction of Th1 and Th2 cytokines and synergy with interleukin-12 for interferon-gamma production. Eur. J. Immunol. 26: 1621–1627 54. Rescigno M., Piguet V., Valzasina B., Lens S., Zubler R., French L., Kindler V., Tschopp J., Ricciardi-Castagnoli P. 2000. Fas engagement induces the maturation of dendritic cells (DCs), the release of

T-cell activation by NOD dendritic cells

55.

56.

57.

58. 59.

60.

61.

62.

63.

interleukin (IL)-1beta, and the production of interferon gamma in the absence of IL-12 during DC-T cell cognate interaction: a new role for Fas ligand in inflammatory responses. J. Exp. Med. 192: 1661–1668 Koarada S., Wu Y., Ridgway W.M. 2001. Increased entry into the IFN-gamma effector pathway by CD4+ T cells selected by I-Ag7 on a nonobese diabetic versus C57BL/6 genetic background. J. Immunol. 167: 1693–1702 Ito T., Amakawa R., Inaba M., Ikehara S., Inaba K., Fukuhara S. 2001. Differential regulation of human blood dendritic cell subsets by IFNs. J. Immunol. 166: 2961–2969 Heufler C., Koch F., Stanzl U., Topar G., Wysocka M., Trinchieri G., Enk A., Steinman R.M., Romani N., Schuler G. 1996. Interleukin-12 is produced by dendritic cells and mediates T helper 1 development as well as interferon-gamma production by T helper 1 cells. Eur. J. Immunol. 26: 659–668 Ziegler S.F., Ramsdell F., Alderson M.R. 1994. The activation antigen CD69. Stem Cells 12: 456–465 Serreze D.V., Prochazka M., Reifsnyder P.C., Bridgett M.M., Leiter E.H. 1994. Use of recombinant congenic and congenic strains of NOD mice to identify a new insulin-dependent diabetes resistance gene. J. Exp. Med. 180: 1553–1558 Fox C.J., Danska J.S. 1998. Independent genetic regulation of T-cell and antigen-presenting cell participation in autoimmune islet inflammation. Diabetes 47: 331–338 Lamhamedi-Cherradi S.E., Luan J.J., Eloy L., Fluteau G., Bach J.F., Garchon H.J. 1998. Resistance of T-cells to apoptosis in autoimmune diabetic (NOD) mice is increased early in life and is associated with dysregulated expression of Bcl-x. Diabetologia 41: 178–184 Leijon K., Hammarstrom B., Holmberg D. 1994. Non-obese diabetic (NOD) mice display enhanced immune responses and prolonged survival of lymphoid cells. Int. Immunol. 6: 339–345 Colucci F., Cilio C.M., Lejon K., Goncalves C.P., Bergman M.L., Holmberg D. 1996. Programmed cell death in the pathogenesis of murine IDDM: resistance to apoptosis induced in lymphocytes by cyclophosphamide. J. Autoimmun. 9: 271–276

35

64. Colucci F., Bergman M.L., Penha-Goncalves C., Cilio C.M., Holmberg D. 1997. Apoptosis resistance of nonobese diabetic peripheral lymphocytes linked to the Idd5 diabetes susceptibility region. Proc. Natl. Acad. Sci. 94: 8670–8674 65. Mills C.D., Kincaid K., Alt J.M., Heilman M.J., Hill A.M. 2000. M-1/M-2 macrophages and the Th1/Th2 paradigm. J. Immunol. 164: 6166–6173 66. Liu T., Nishimura H., Matsuguchi T., Yoshikai Y. 2000. Differences in interleukin-12 and -15 production by dendritic cells at the early stage of Listeria monocytogenes infection between BALB/C and C57BL/6 mice. Cell Immunol. 202: 31–40 67. Wakeham J., Wang J., Xing Z. 2000. Genetically determined disparate innate and adaptive cell-mediated immune responses to pulmonary Mycobacterium bovis BCG infection in C57BL/6 and BALB/C mice. Infect. Immun. 68: 6946–6953 68. Langenkamp A., Messi M., Lanzavecchia A., Sallusto F. 2000. Kinetics of dendritic cell activation: impact on priming of TH1, TH2 and nonpolarized T cells. Nat. Immunol. 1: 311–316 69. Corinti S., Albanesi C., la Sala A., Pastore S., Girolomoni G. 2001. Regulatory activity of autocrine IL-10 on dendritic cell functions. J. Immunol. 166: 4312–4318 70. Ludewig B., Odermatt B., Ochsenbein A.F., Zinkernagel R.M., Hengartner H. 1999. Role of dendritic cells in the induction and maintenance of autoimmune diseases. Immunol. Rev. 169: 45–54 71. Luzina I.G., Atamas S.P., Storrer C.E., daSilva L.C., Kelsoe G., Papadimitriou J.C., Handwerger B.S. 2001. Spontaneous formation of germinal centers in autoimmune mice. J. Leukoc. Biol. 70: 578–584 72. Yoshimura S., Bondeson J., Foxwell B.M., Brennan F.M., Feldmann M. 2001. Effective antigen presentation by dendritic cells is NF-kappaB dependent: coordinate regulation of MHC, co-stimulatory molecules and cytokines. Int. Immunol. 13: 675–683 73. Poligone B., Weaver D.J. Jr, Sen P., Baldwin A.S. Jr, Tisch R. 2002. Elevated NF-kappaB activation in nonobese diabetic mouse dendritic cells results in enhanced APC function. J. Immunol. 168: 188–196