Inflammatory bowel disease (IBD), clinically comprising Crohn’s disease (CD), and ulcerative colitis (UC) are chronic inflammatory disorders of the gut with complex etiologies (Kaser et al., 2010). The understanding of pathogenic mechanisms underlying the development of IBD has aided the development of novel biological therapies. Agents that block the TNF pathways have been particularly successful in IBD therapy (Melmed and Targan, 2010). However, a large proportion of patients either fail to respond or develop tolerance to TNF therapy, highlighting the need for new therapeutic targets (Baumgart and Sandborn, 2007). IL-1β is a proinflammatory cytokine with a wide range of systemic and local effects. Primarily produced by innate leukocytes, IL-1β can modulate the function of both immune and nonimmune cells. Stimulation with IL-1β promotes the activation and effector functions of dendritic cells, macrophages, and neutrophils (Dinarello, 1996). Moreover, IL-1β can induce neutrophilia and promote neutrophil migration (Dinarello, 2009). IL-1β promotes T cell activation and survival (Ben-Sasson et al., 2009) and has recently been shown to act in concert with other proinflammatory cytokines to promote the differentiation of CD4+ Th17 cells (Sutton et al., 2006, 2009; Acosta-Rodriguez et al., 2007; Chung et al., 2009). The potent inflammatory activity of IL-1β is reflected by the tight mechanisms in place to regulate its secretion. IL-1β is translated as an inactive 31 kD precursor (pro-IL-1β) after TLR stimulation, which is cleaved into its activated 17 kD form by caspase-1, also known as interleukin-1β converting enzyme (ICE; Thornberry et al., 1992). Activation of caspase-1 relies on the formation of a multimolecular scaffold known as the inflammasome, which is triggered by the activation of intracellular NOD-like receptors (NLR) by endogenous or exogenous danger signals (Martinon et al., 2009). Upon secretion into the extracellular space, IL-1β binds and signals via the IL-1 receptor 1 (IL-1R1), which is expressed on a wide range of cell types, including epithelial and endothelial cells, hepatocytes, and innate and adaptive leukocytes (Sims and Smith, 2010). Regulatory mechanisms downstream of IL-1β secretion are also in place to limit the action of IL-1β, including the production of a natural antagonist for its receptor, IL-1RA, and the expression of a decoy receptor, IL-1R2 (Dinarello, 1996). Several clinical studies have reported high levels of IL-1β secretion by colon lamina propria monocytes from patients with active IBD (Satsangi et al., 1987; Mahida et al., 1989; Ligumsky et al., 1990; Reinecker et al., 1993; McAlindon et al., 1998). IL-1β levels in the colon correlated with disease activity and high levels of IL-1β were associated with active lesions (Casini-Raggi et al., 1995; Ludwiczek et al., 2004), suggesting an important role of this cytokine in promoting localized inflammation. High levels of colonic IL-1β are also a feature of many animal models of colitis (Cominelli et al., 1990; Okayasu et al., 1990; McCall et al., 1994), and treatment with IL-1–blocking agents has been successful in ameliorating acute models of intestinal injury and inflammation (Thomas et al., 1991; Cominelli et al., 1992; Siegmund et al., 2001). Furthermore, different genetic lesions associated with IBD development in animal models are associated with increased IL-1β. For example, macrophages from knock-in mice bearing the most common CD-associated variant of the Nod2 gene produced high levels of IL-1β after stimulation with muramyl dipeptide (MDP; Maeda et al., 2005). These NOD2 mutant mice also developed exacerbated disease in response to acute dextran sulfate sodium (DSS)–induced intestinal injury, which was significantly ameliorated by the administration of recombinant IL-1RA (Maeda et al., 2005). In addition, conditional deletion of the CD-linked autophagy gene Atg16l1 in the hematopoietic system of mice resulted in increased IL-1β production after LPS stimulation and increased susceptibility to DSS-mediated intestinal injury, a phenotype reversed by co-treatment with αIL-18 and αIL-1β antibodies (Saitoh et al., 2008). The importance of IL-1β in modulating intestinal inflammation has been confirmed by infection studies, as blocking IL-1β ameliorated inflammatory pathology in both Clostridium difficile–associated colitis and Salmonella typhimurium–induced enteritis (Müller et al., 2009; Ng et al., 2010). Although IL-1β has been linked to Th17 cell responses, the role of IL-17A in the development of intestinal inflammation is controversial. In acute models of intestinal injury, IL-17A plays a disease protective role, with Il17a−/− mice showing increased pathology and leukocyte infiltration after DSS administration (Ogawa et al., 2004). However, studies in chronic inflammatory models have highlighted a more complex role for IL-17A. Studies from our laboratory demonstrated a pathogenic role for IL-17A in Helicobacter hepaticus (H. hepaticus)–driven innate immune IBD (Buonocore et al., 2010). Similarly, administration of an αIL-17A antibody ameliorated the spontaneous colitis developed by mice bearing a conditional deletion of the transcription factor Stat3 in regulatory Foxp3+ T cells (Chaudhry et al., 2009). Th17 cells are enriched in the inflamed gut both in animal models and in humans (Fujino et al., 2003; Nielsen et al., 2003; Hue et al., 2006; Kullberg et al., 2006) and T cells lacking the transcription factor RORγ, that directs Th17 differentiation, could not transfer colitis to C57BL/6 Rag1−/− mice (Leppkes et al., 2009). However, T cell–derived IL-17A is not absolutely required for the development of intestinal pathology in T cell transfer models of colitis and it has been proposed that T cell–derived IL-17A and IL-17F might play a redundant role in driving intestinal inflammation (Izcue et al., 2008; Leppkes et al., 2009; O’Connor et al., 2009). These conflicting results might be explained by an as yet undiscovered additional pathogenic function of Th17 cells. Alternatively, a complex network of proinflammatory cells may contribute to IL-17A–mediated pathology in vivo (Littman and Rudensky, 2010). In this study, we aimed to assess the role of IL-1β in chronic intestinal inflammation. As a result of the pluripotent activity of IL-1β, we used complementary animal models of chronic colitis to selectively analyze the effects of IL-1β on adaptive and innate immune-mediated intestinal inflammation. Our results show that IL-1β signals are required for the development of severe inflammation in both T cell–independent and T cell–mediated colitis. Moreover, we identified key mechanisms underlying the pathogenic function of IL-1β, including a central role for this cytokine in promoting the accumulation of IL-17A–producing innate and adaptive immune cells. RESULTS IL-1β plays a key role in innate intestinal inflammation To specifically analyze the role of IL-1β in modulating innate inflammatory responses in the intestine, we infected T cell– and B cell–deficient 129SvEv Rag2−/− mice with H. hepaticus, which results in the development of colonic and cecal inflammation that is entirely dependent on innate immunity (Erdman et al., 2003; Maloy et al., 2003). We first examined whether H. hepaticus–induced intestinal inflammation was associated with increased levels of active IL-1β by analyzing the levels of IL-1β secretion by organ explants from the gastrointestinal tract of H. hepaticus–infected 129SvEv Rag2−/− mice. Intestinal inflammation in the colon and cecum of H. hepaticus–infected 129SvEv Rag2−/− mice was associated with high levels of secreted IL-1β (Fig. 1 A). In contrast, no increase in IL-1β levels was observed in the ileum of H. hepaticus–infected 129SvEv Rag2−/− mice (Fig. 1 A). Given that both colonization and H. hepaticus–induced inflammation occur primarily in the colon and cecum, this result suggests that IL-1β secretion is directly linked to local intestinal inflammation. Innate leukocytes appear to be the major source of IL-1β secretion in the lamina propria in IBD patients (Mahida et al., 1989; McAlindon et al., 1998). We observed high levels of IL-1β secretion by purified colonic lamina propria leukocytes (cLPLs) of H. hepaticus–infected 129SvEv Rag2−/− mice (Fig. 1 B), confirming that chronic intestinal inflammation correlates with increased local secretion of IL-1β by innate leukocytes. Figure 1. H. hepaticus–induced innate immune driven typhlocolitis is associated with IL-1β secretion. 129SvEv Rag2−/− mice were infected with H. hepaticus and sacrificed >8 wk after infection. (A) IL-1β secretion from organ explants from colon, cecum, and ileum incubated overnight in complete medium. Results are shown as mean ± SEM (n = 3 for uninfected control and n = 14 for H. hepaticus–infected mice, pooled from 2 independent experiments). (B) IL-1β levels in the supernatants of cLPLs cultured overnight in complete media. Results are shown as mean ± SEM (n = 7 for uninfected controls and n = 22 for H. hepaticus-infected mice, pooled from 3 separate experiments). **, P 8 wk of infection. LPLs were isolated from the large intestine, Sca1+Thy1.2Hi ILCs were FACS sorted, and Il1r1 expression was evaluated by qRT-PCR (mean ± SEM, n = 2 from 2 independent experiments; 8–10 mice were pooled in each experiment). (B–E) 129SvEv Rag2−/− mice were infected with H. hepaticus and treated weekly with 1 mg of αIL-1β antibody or isotype control (i.p.). After 8 wk, mice were sacrificed and cLPLs were isolated. (B) Total numbers of Sca1+ Thy1.2Hi ILCs in the colon lamina propria as evaluated by FACS analysis. (C) Total numbers of IL-17A– or IFN-γ–producing ILCs from the colon of indicated mice groups. (D) Cytokine production by cLPLs after overnight culture in complete medium alone (n/a) or in the presence of 10 ng/ml IL-23. Data are represented as mean ± SEM from 2 pooled independent experiments (n = 6–11). (E) Il23r expression by cLPLs as evaluated by qRT-PCR. Data are normalized on Hprt expression. (F and G) 129SvEv Rag2−/− mice were infected with H. hepaticus for 8 wk and CD45+lin−Sca1+Thy1.2Hi ILCs were FACS sorted from the colon. (F) ILCs were cultured overnight in complete medium alone (n/a) or in the presence of 10 ng/ml IL-1β. Il23r and Rorc expression levels were evaluated by qRT-PCR and normalized on Hprt expression. Data are shown as mean ± SEM from 2 pooled independent experiments. In each experiment, 10–18 mice were pooled. (G) ILCs were cultured overnight in complete medium alone (n/a) or in the presence of 10 ng/ml IL-23. Il1r1 expression was evaluated by qRT-PCR and normalized on Hprt expression. Data are shown as mean ± SEM from two pooled independent experiments. In each experiment, 10–18 mice were pooled. *, P 6 wk of age when used. Bacteria. H. hepaticus NCI-Frederick isolate 1A, isolated from the same mouse colony as isolate Hh-1 (strain 51449; American Type Culture Collection), was grown on blood agar plates containing Trimethoprim, vancomycin, and polymixin B (Oxoid) under microaerophilic conditions, as previously described (Maloy et al., 2003; Young et al., 2004). H. hepaticus viability was confirmed using fluorescent microscopy with a bacterial live/dead kit (BacLight; Invitrogen). For induction of innate immune colitis, 129SvEv Rag2−/− mice were infected with H. hepaticus (∼108 CFU) by oral gavage three times on alternate days. H. hepaticus–infected mice were sacrificed 8–10 wk after infection. In vivo antibody treatment. To block IL-1β activity in vivo, H. hepaticus–infected 129SvEv Rag2−/− mice were injected i.p. with 1 mg αIL-1β–blocking mAb (αIL-1β) or 1 mg isotype control mAb from the day of the first inoculation with H. hepaticus, and treatment repeated weekly for the duration of the experiment. Induction of colitis with naive CD4+CD45RBHi T Cells. Naive CD4+CD45RBHi T cells were purified (>98%) from spleens of C57BL/6, C57BL/6 Il1r1−/− , IL23r −/−, or Csf2−/− mice via FACS sorting, as previously described (Izcue et al., 2008). In brief, single-cell suspensions were depleted of CD8+, MHC class II+, Mac-1+, and B220+ cells by negative selection using a panel of rat monoclonal antibodies, followed by sheep anti–rat–coated Dynabeads (Invitrogen). After staining with APC-conjugated αCD4, PE-conjugated αCD25, and FITC–αCD45RB, naive CD4+CD45RBHi T cells were purified by cell sorting with a cell sorter (MoFlo; Dako). Naive T cell suspensions were washed in sterile PBS, and age- and sex-matched C57BL/6 Rag1−/− recipient mice received 4 × 105 CD4+CD45RBHi T cells by i.p. injection. For co-transfer experiments, 1:1 mixtures of CD45.2+ Il23r−/− and CD45.1+ WT CD4+CD45RBHi T cells were injected i.p. (total cell number = 4 × 105). Mice were sacrificed when symptoms of clinical disease (weight loss and/or diarrhea) developed in control groups, ∼5–8 wk after transfer of naive CD4+ T cells, unless otherwise indicated. For mice sacrificed 2 wk after transfer, 2 × 106 CD4+CD45RBHi T cells were transferred per mouse. Assessment of intestinal inflammation. Samples of cecum and proximal, mid, and distal colon were prepared as previously described (Izcue et al., 2008), and inflammation was graded according to the following scoring system. Each sample was graded semiquantitatively from zero to three for four criteria: (1) degree of epithelial hyperplasia and goblet cell depletion; (2) leukocyte infiltration in the lamina propria; (3) area of tissue affected; and (4) the presence of markers of severe inflammation such as crypt abscesses, submucosal inflammation, and ulcers. Scores for each criterion were added to give an overall inflammation score for each sample of 0–12. Scores from proximal, mid, and distal colon were averaged to obtain inflammation scores for the colon. Sections were scored in a blinded fashion to avoid bias. Isolation of leukocyte subpopulations and FACS. Cell suspensions from spleen, liver, MLN, and the lamina propria were prepared as described previously (Buonocore et al., 2010). Cells were washed, incubated with anti-Fc receptor (αCD16/32), and stained for flow cytometric analysis using combinations of the following antibodies: PeCy7-conjugated αCD4 and PerCP-Cy5.5-conjugated αTCR-β (T cells) and FITC-conjugated αCD45.2, αCD11b conjugated to PE, and Gr1 conjugated to PerCP-Cy5.5 (innate cells). Intracellular staining was performed as previously described (Ahern et al., 2010): cells were restimulated for 4 h with 0.1 µg/ml PMA (Sigma-Aldrich), 1 µg/ml ionomycin (Sigma-Aldrich), and 10 µg/ml Brefeldin A (Insight Biotechnology), washed, and stained for surface markers. Cells were then fixed overnight in eBioscience Fix/Perm buffer at 4°C. Cells were washed and permeabilized in permeabilization buffer (eBioscience) with 2% rat serum for 1 h at 4°C. Cells were then stained with αIL-17A conjugated to PE, αIFN-γ conjugated to APC or e450, and αKi-67 conjugated to FITC (BD) or appropriate isotype controls (BD) for 30 min at 4°C. ILCs were identified as previously described (Buonocore et al., 2010) using αThy1.2 PE-conjugated and FITC-conjugated αLy6A/E (Sca1) (both from BD Biosciences). Lineage+ cells were excluded using PerCP-Cy5.5–conjugated αCD11b, αGr1, αB220, and αCD3e. For intracellular staining of ILCs, 10 µg/ml brefeldin A (Insight Biotechnology) was added for the last 4 h of overnight cultures in complete RPMI media. Cells were then processed similarly to T cells. For analysis of colonic T cells, CD4+ TCRβ+ cells were sorted from cLPL preparations according to the expression of the surface markers CD4 and TCRβ with a cell sorter (MoFlo; Dako) and restimulated overnight with 0.1 µg/ml PMA and 1 µg/ml ionomycin. Cell culture. FACS sorted ILCs were cultured overnight in RPMI, 10% FCS, 100 U/ml penicillin/streptomycin, 2 mM l-glutamine, 0.05 mM β-mercaptoethanol (complete RPMI), and 10 ng/ml IL-1β. For Th17 differentiation from total CD4+ T cells, splenic CD4+ T cells were isolated from WT or Il1r1−/− mice and cultured in IMDM, 10% FCS, 100 U/ml penicillin/streptomycin, 2 mM l-glutamine, 0.05 mM β-mercaptoethanol (complete IMDM) in the presence of 2.5 ng/ml TGF-β, and 50 ng/ml IL-6 5, plus plate-bound αCD3 (2 µg/ml) and αCD28 (1 µg/ml). For Th17 differentiation from naive CD4+ T cells, naive CD4+ CD62LHi CD44low CD25− were FACS sorted from the spleen and MLN of WT C57BL/6 or Il23r−/− mice and cultured for the indicated time in complete IMDM in the presence of 200 pg/ml TGF-β, 20 ng/ml IL-6, 20 ng/ml IL-1β, 10 ng/ml IL-21, 20 ng/ml IL-23, 10 µg/ml αIFN-γ and αIL-4, and plate-bound αCD3 and αCD28 (both at 5 µg/ml). Quantification of gene expression by qRT-PCR. Quantitative real time PCR (qRT-PCR) was performed as previously described (Hue et al., 2006) with homogenization of frozen tissue samples performed using a FastPrep 24 homogenizer (MP Biomedicals) with lysing matrix D beads (MP Biomedicals). Primers for Hprt (Mm01545399_m1), cxcl1 (Mm04207460_m1), cxcl2 (Mm0436450_m1), cxcl5 (Mm0436451_g1), bcl2 (Mm0477631_m1), bclxl (Mm0437783_m1), Il1r1 (Mm00434237_m1), and rorc (Mm01261022_m1) were obtained from Applied Biosystems. Levels of gene expression were normalized on Hprt expression, unless otherwise indicated. In-house primer sequences are as follows: CD3, forward, 5′-TTACAGAATGTGTGAAAACTGCATTG-3′, reverse, 5′-CACCAAGAGCAAGGAAGAAGATG-3′, and probe, 5′-ACATAGGCACCATATCCGGCTTTATCTTCG-3′; CCR3 forward, 5′-TGTTTACCTCAGTTCATCCACGG-3′, reverse, 5′-CAGAATGGTAATGTGAGCAGGAA-3′, and probe, 5′-TCTGCTCAACTTGGCCATCTCTGACC-3′; CCR5 forward, 5′-CATCGATTATGGTATGTCAGCACC-3′, reverse, 5′-CAGAATGGTAGTGTGAGCAGGA-3′, and probe 5′-TACCTGCTCAACCTGGCCATCTCTGA-3′; CCR6 forward, 5′-ACTCTTTGTCCTCACCCTACCG-3′, reverse, 5′-ATCCTGCAGCTCGTATTTCTTG-3′, and probe, 5′-ACGCTCCAGAACACTGACGCACAGTA-3′; CXCR6 forward, 5′-AAGCTGAGGACTCTGACAGATGTGT-3′, reverse, 5′-CCAAAAGGGCAGAGTACAGACAA-3′, and probe, 5′-CTGCTGAACTTGCCCCTGGCTGAC-3′; IL-23R forward, 5′-CCATCTGGATGATATAGTGATACCTTCT-3′, reverse, 5′-ATGGTCTTGGGTACAGTATCGTTTG-3′, and probe, 5′-CGTCCATCATTTCCAGGGCTCACACT-3′ (Ahern et al., 2010). Amplification was performed using TaqMan Fast Universal PCR Master Mix (Applied Biosystems). Apoptosis assay. For in vitro assessment of apoptosis, total splenocytes from C57BL/6 or Il1r1−/− mice were depleted of CD8+, MHC class II+, Mac-1+, and B220+ cells by negative selection using sheep anti–rat–coated Dynabeads (Invitrogen; purity ≥82%). Enriched CD4+ cells were cultured for 3 d with 2.5 µg/ml or 10 µg/ml each of plate-bound αCD3 and αCD28 (eBioscience). APC anti–Annexin-V antibody and 7-Aminoactinomycin (7AAD; eBioscience) were used for apoptosis assessment. All cells were acquired on a FACSCalibur (BD) or on a CyAn ADP flow cytometer (Beckman Coulter), and analysis performed using FlowJo (Tree Star) software. Cytokine detection. For cytokine detection, colonic lamina propria cells we cultured overnight in complete RPMI media. IL-6, IFN-γ, IL-17, TNF, and IL-1β levels in culture supernatants were measured using FlowCytomix Bead-based assay (eBioscience) and cytokine concentration was normalized to cell number. To assess innate lymphoid cell function, colonic lamina propria cells were cultured with or without recombinant IL-23 (10 ng/ml; R&D) overnight, and IL-17 and IFN-γ were measured in the supernatants as above. Organ explants were prepared as previously described (Hue et al., 2006) and cultured overnight in complete RPMI media. IL-1β levels in the supernatants were determined by Bio-Plex Cytokine assay (Bio-Rad Laboratories), and concentrations were normalized to the weight of the explants. Statistics. The nonparametric Mann-Whitney test was used for assessment of statistical significance of in vivo experiments. The Student’s t test was used to assess the significance of in vitro experiments. Weight curves were compared using two-way ANOVA. Data were considered significant when P < 0.05.