Abstract
The intestinal immune system must be able to respond to a wide variety of infectious organisms while maintaining tolerance to non-pathogenic microbes and food antigens. The Vitamin A metabolite retinoic acid (RA) has been implicated in the regulation of this balance, partially by regulating innate lymphoid cell (ILC) responses in the intestine. However, the molecular mechanisms of RA-dependent intestinal immunity and homeostasis remain elusive. Here we define a role for the transcriptional repressor Hypermethylated in cancer 1 (HIC1, ZBTB29) in the regulation of ILC responses in the intestine. Intestinal ILCs express HIC1 in a vitamin A-dependent manner. In the absence of HIC1, group 3 ILCs (ILC3s) are lost, resulting in increased susceptibility to infection with the bacterial pathogen Citrobacter rodentium. In addition, the loss of ILC3s leads to a local and systemic increase in IFN-γ-producing T cells that prevents the development of protective immunity against infection with the parasitic helminth Trichuris muris. Thus, RA-dependent expression of HIC1 in ILC3s regulates intestinal homeostasis and protective immunity.
Author Summary Innate lymphoid cells (ILCs) are emerging as important regulators of immune responses at barrier sites such as the intestine. However, the molecular mechanisms that control this are not well described. In the intestine, the Vitamin A metabolite retinoic acid (RA) has been shown to be an important component of the homeostatic mechanisms. In this manuscript, we show that the RA-dependent transcription factor Hypermethylated in cancer 1 (HIC1, ZBTB29) is required for ILC homeostasis and function in the steady state as well as following infection with the bacterial pathogen Citrobacter rodentium or the helminth parasite Trichuris muris. Thus, HIC1 links RA signalling to intestinal immune responses. Further, our results identify HIC1 as a potential target to modulate ILC responses in vivo in health and disease.
INTRODUCTION
The intestinal immune system is held in a tightly regulated balance between immune activation in response to potential pathogens and the maintenance of tolerance to innocuous antigens, such as food and commensal flora. Disruption of this balance can lead to the development of serious inflammatory disorders, such as food allergy or inflammatory bowel disease (IBD). A complex network of different immune cell types including dendritic cells (DCs), macrophages, innate lymphoid cells (ILCs), and T cells, are essential for both the induction of active immunity and the maintenance of intestinal homeostasis.
The vitamin A metabolite all-trans-retinoic acid (atRA) plays an important role in shaping intestinal immunity by regulating both the innate and adaptive immune systems. atRA that is generated by the metabolism of Vitamin A by intestinal epithelial cells (IECs) and a subset of CD103-expressing intestinal dendritic cells (CD103+ DCs) has been shown to directly affect the localization and function of lymphocytes. For example, atRA has been shown to induce expression of chemokine receptors (CCR9) and integrins (α4 and β7) that are associated with homing to, and retention in, the intestinal microenvironment (Iwata et al., 2004; Kim et al., 2015; Mora and von Andrian, 2009). In addition, atRA has been shown to control the balance of regulatory T (Treg) cells and CD4+ T helper 17 (TH17) cells in the intestine by promoting Treg cell differentiation and inhibiting TH17 cell development (Sun et al., 2007; Mucida et al., 2007; Elias et al., 2008; Hill et al., 2008; Xiao et al., 2008; Takaki et al., 2008). Similarly, atRA controls the development of ILC subsets in the intestine, as mice raised on a Vitamin A-deficient (VAD) diet display reduced numbers of ILC3s (Spencer et al., 2014; Wilhelm et al., 2016), with one study showing a concomitant increase in ILC2 numbers and enhanced type 2 immunity within the intestine (Spencer et al., 2014). In addition, intestinal DC differentiation is influenced by atRA as mice raised on a VAD diet display reduced numbers of CD103+ CD11b+ DCs (Zeng et al., 2016). Thus, atRA-dependent processes are central to the function of intestinal TH cells, ILCs, and DCs in vivo. However, the molecular mechanisms downstream of atRA signaling that control immune cell function and homeostasis remain unknown.
Hypermethylated in cancer 1 (HIC1, ZBTB29) is a transcriptional factor that was first identified as a gene that is epigenetically silenced in a variety of human cancers (Chen et al., 2003; Wales et al., 1995). HIC1 has been shown to regulate cellular proliferation, survival and quiescence in multiple normal and tumour cell lines (Lin et al., 2013; Valenta et al., 2006; Van Rechem et al., 2010; Chen et al., 2005). HIC1 is a member of the POZ and Kruppel/Zinc Finger and BTB (POK/ZBTB) family of transcription factors that consists of regulators of gene expression that are critical in a variety of biological processes (Kelly and Daniel, 2006). Importantly, several members of the POK/ZBTB family are key regulators in immune cell differentiation and function, including: BCL6, PLZF and ThPOK (Lee and Maeda, 2012; Nurieva et al., 2009; Savage et al., 2008; Wang et al., 2008; Muroi et al., 2008). Recently, we identified HIC1 as an atRA responsive gene in intestinal TH cells and demonstrated a T cell-intrinsic role for HIC1 in the regulation of intestinal homeostasis as well as in development of several models of intestinal inflammation (Burrows, et al. 2017).
In this study, we show that HIC1 is critical for intestinal immune homeostasis by regulating ILC3s. Deletion of HIC1 in hemoatopoietic cells or RORγt-expressing cells results in a significant reduction in the number of ILC3s in the intestine. Further, we identify an ILC3-intrinic role for HIC1 in regulating intestinal TH cell responses to commensal bacteria and immunity to infection with the intestinal pathogens Citrobacter rodentium and Trichuris muris. These results identify a central role for atRA-dependent expression of HIC1 in ILC3s in the regulation of intestinal immune responses.
RESULTS
HIC1 is expressed by intestinal ILCs and is critical for intestinal immune homeostasis
We have previously shown that HIC1 was highly expressed by T cells, DCs and macrophages within the intestinal lamina propria (LP) and intraepithelial compartments. (Burrows et al., 2017). Using mice with a fluorescent reporter gene inserted in the Hic1 locus (Hic1Citrine mice) (Pospichalova et al., 2011) we were able to determine that in addition to the previously identified populations, lineage-negative (linneg) CD90.2+ CD127+ ILCs isolated from the intestinal lamina propria express HIC1 (Fig 1A). Similar to our previous results, HIC1 expression in ILCs was dependent on the availability of atRA, as Hic1Citrine mice reared on a VAD diet did not express HIC1 in ILCs within the LP (Fig 1B). To determine the role of HIC1 in ILCs, we first crossed mice with loxP sites flanking the Hic1 gene (Hic1fl/fl mice) with mice that express the Cre recombinase under control of the Vav promoter (Vav-Cre mice) to generate mice with a hematopoietic specific deletion of Hic1 (Hic1Vav mice). Upon hematopoietic cell-specific deletion of HIC1 we observed a significant change in ILC populations in the LP. In the steady state, we observed significantly fewer RORγt+ ILCs (ILC3s) in the LP of Hic1Vav mice, with a significant reduction in the number of RORγt+ TBET+ ILC3s (Fig 1C,D). We also detected a small but significant increase in the number of CD4+ ILC3s (also known as lymphoid tissue inducer (LTi) cells) but no change in numbers of the canonical Gata3+ ILC (ILC2) population (Fig 1C,D). Thus HIC1 expression in hematopoietic cells is critical for regulation of ILC populations in the LP.
HIC1 does not regulate ILC precursors in the bone marrow
As we observed a significant reduction of ILC3s in the LP in the absence of HIC1, we next tested directly whether the lack of HIC1 affected the upstream development of ILC precursors in the bone marrow. ILCs develop in the bone marrow through a lineage pathway that begins with a common lymphoid progenitor (CLP) and progresses through an α4β7-expressing lymphoid progenitor (αLP), a common progenitor to all helper-like ILCs (ChILP) and, in the case of ILC2s, an ILC2 precursor (ILC2p) (Zook and Kee, 2016). Analysis of surface marker expression on lineage-negative, CD45+ bone marrow cells showed that HIC1 was not required for the development of CLP, αLP, ChILP or ILC2p populations (Fig 2A,B). Thus, the reduced number of ILC3s in the LP is not due to a reduced frequency of ILC precursors and suggests that HIC1 is required for ILC3 homeostasis in the periphery.
Hematopoietic specific deletion of HIC1 results in susceptibility to intestinal bacterial infection
ILC3s have been shown to play a significant role in resistance to infection with the attaching and effacing bacterial pathogen Citrobacter rodentium (Satoh-Takayama et al., 2008; Sonnenberg et al., 2011). Following infection with C. rodentium, Hic1Vav mice exhibited enhanced weight loss and significantly higher bacterial burdens in the feces compared to Hic1fl/fl controls (Fig 3A,B). Furthermore, infected Hic1Vav mice but not Hic1fl/fl mice had dissemination of bacteria to the liver (Fig 3C), demonstrating a significant impairment in the intestinal barrier following infection. Associated with impaired bacterial containment and clearance were reduced levels of transcripts for the cytokines Il17a and Il22, as well as the intestinal antimicrobial peptide Reg3g (Fig 3D). Thus, expression of HIC1 within hematopoietic cells is critical to mount a proper immune response against C. rodentium.
ILC3-intrinsic HIC1 expression is critical for defence against intestinal bacterial infection
As T cells, CD103+ CD11b+ DCs and ILC3s are all important in initiating and propagating ILC3/TH17 responses in the intestine (Persson et al., 2013; Kinnebrew et al., 2012; Collins et al., 2014; Klose and Artis, 2016) and these population are perturbed in Hic1Vav mice, we next sought to determine the effect of HIC1 deficiency in these specific cell populations during infection C. rodentium. We crossed Hic1fl/fl mice with mice expressing Cre under the control of either the Cd4 promoter or Itgax promoter to generate T cell-specific (Hic1CD4 mice) and dendritic cell-specific (Hic1CD11c mice) HIC1-deficient mice. Both Hic1CD4 mice (S1A-C Fig) and Hic1CD11c mice (S1D-F Fig) were as resistant to infection with C. rodentium as control Hic1fl/fl mice, with equivalent weight loss, fecal bacterial burdens and expression of cytokines and antimicrobial peptide mRNA in the intestine. Thus, these results demonstrate that expression of HIC1 in T cells or CD11c-expressing cells is not required for immunity to bacterial infection and suggests loss of HIC1 in another cell population is responsible for the phenotype observed in Hic1Vav mice.
To determine the role of HIC1 expression in ILC3s during infection with C. rodentium, we crossed Hic1fl/fl mice with mice expressing Cre recombinase under the control of the Rorc promoter (Hic1Rorc mice). Following infection with C. rodentium, and similar to what we observed in the Hic1Vav mice, Hic1Rorc mice displayed increased weight loss, higher fecal bacterial burdens and increased bacterial dissemination than control Hic1fl/fl mice (Fig 4A-C). Associated with increased susceptibility was reduced expression of IL17a, IL22 and Reg3g in intestinal tissues (Fig 4D). Based on our results, we hypothesized that HIC1 is an important regulator of ILC3 function during C. rodentium infection. To better examine the effect of HIC1 deletion on ILC3s, we examined the intestinal LP of Hic1fl/fl mice and Hic1Rorc mice at day 4 post C. rodentium infection. Enhanced susceptibility to infection with C. rodentium that we observed in Hic1Rorc mice correlated with the loss of ILC3s and reduced numbers IL-22-producing ILC3s in (Fig 4E,F). Taken together, these results suggest that expression of HIC1 in RORγt+ ILC3s is critical for resistance to intestinal bacterial infection.
Hematopoietic deficiency of HIC1 results in susceptibility to intestinal helminth infection
Our results show that HIC1 is an atRA-responsive factor that is critical for regulation of ILC3 homeostasis and function. As a recent study demonstrated that blockade of RA signalling results in dysfunctional ILC3 responses along with a compensatory increase in ILC2 responses and enhanced immunity to infection with the intestinal helminth parasite Trichuris muris (Spencer et al., 2014), we hypothesized that in addition to the defective ILC3 response observed in both Hic1Vav mice and Hic1Rorc mice, we would find enhanced ILC2 responses in the absence of HIC1. To test this, we infected Hic1fl/fl and Hic1Vav mice with T. muris. In contrast to our expectations, we did not observe a heightened protective ILC2/Th2 cell response, but found that Hic1Vav mice were susceptible to infection. Hic1Vav mice maintained a significant worm burden and enlarged mesenteric lymph node 21 days after infection (Fig 5A,B). Histological analysis revealed that T. muris-infected Hic1Vav mice displayed a reduced frequency of goblet cells, with parasites embedded within the caecal epithelium (Fig 5C,D). Associated with the increased susceptibility, Hic1Vav mice mounted a non-protective type 1 response following infection, as restimulation of the draining mesenteric lymph node (mLN) revealed a significant increase in secreted IFN-γ (Fig 5E) and expression of the Ifng in the intestinal tissues (Fig 5F), concomitant with a reduced type 2 response. We also observed a switch in the T. muris-specific antibody response from IL-4-dependent IgG1 to IFN-γ-dependent IgG2a in the serum of infected Hic1Vav mice (Fig 5G). Thus, loss of HIC1 in hematopoietic cells results in increased susceptibility to infection with T. muris.
HIC1 expression in T cells and dendritic cells is dispensable for immunity to T. muris
To determine which cell type required HIC1 expression to promote immunity to T. muris, we infected both Hic1CD4 mice and Hic1CD11c mice. Strikingly, loss of HIC1 in T cells or CD11c-expressing cells had no effect on the development of protective immunity against T. muris. We observed equivalent expulsion of worms and expression of Ifng and Il13 in the intestinal tissues between control Hic1fl/fl mice, Hic1CD4 mice (S2A,B Fig) and Hic1CD11c mice (S2C,D Fig). Thus, we conclude that expression of HIC1 in T cells or CD11c-expressing cells is not required for immunity to T. muris.
ILC3-intrinsic expression of HIC1 is required for immunity to T. muris
As there was no observable role for HIC1 in T cell or DC function during T. muris infection, we next sought to determine if there was a defect in HIC1 deficient ILC3s that prevent clearance of the parasite. Surprisingly, Hic1Rorc mice infected with T. muris were susceptible to infection, maintaining a significant parasite burden at day 21 post-infection (Fig 6A). Similar to Hic1Vav mice, we observed enlarged mesenteric lymph nodes (Fig 6B), as well as increased inflammatory cell infiltration, reduced number of goblet cells, submucosal edema, and parasites embedded within the caecal epithelium (Fig 6C,D). Consistent with the lack of protective immunity, we found that Hic1Rorc mice displayed high levels of secreted IFN-γ from restimulated mLN cells as well as increased expression of Ifng in intestinal tissues (Fig 6E,F), which is associated with increased levels of IFN-γ-dependent IgG2a antibodies in the serum (Fig 6G). However, we failed to observe reduced expression of type 2 cytokines in the mLN or intestine (Fig 6E,F), suggesting that the heightened levels of IFN-γ was promoting susceptibility in the context of a protective type 2 immune response. Thus, ILC3-intrinsic expression of HIC1 is critically required for the development of protective type 2 immune responses against T. muris.
Neutralization of IFN-γ in T. muris-infected Hic1Rorc mice promotes resistance
Based on our results showing heightened levels of IFN-γ, we next asked whether antibody blockade of IFN-γ would render Hic1Rorc mice resistant to infection. We found that treatment of T. muris-infected Hic1Rorc mice with α-IFN-γ antibody promoted resistance, with a complete clearance of parasites in antibody-treated mice by day 21 post infection (Fig 7A). Increased resistance was associated with increased numbers of goblet cells (Fig 7B,C) along with reduced levels of IFN-γ and heightened levels of IL-4 and IL-13 (Fig 7D,E). Antibody treatment also resulted in reduced levels of T. muris-specific IgG2a and increased levels of IgG1 in the serum (Figure 7F). Thus, these results suggest that ILC3-specific expression of HIC1 is not required for resistance to T. muris infection when IFN-γ responses are neutralized.
ILC3-intrinsic HIC1 is required to limit commensal bacteria specific TH cell responses
ILC3s play a central role in intestinal immune homeostasis by limiting T cell responses against commensal bacteria (Hepworth et al., 2013; Hepworth et al., 2015). A subset of intestinal ILC3s can present commensal bacterial antigen to CD4+ T cells through MHCII but lack any co-stimulatory molecules and thus induce anergy in commensal-specific T cells (Hepworth et al., 2013). As we observed reduced numbers of ILC3s as well as enlarged mLNs and heightened levels of IFN-γ production from restimulated T cells from the mLN of naïve and T. muris-infected Hic1Rorc mice, we hypothesized that the increased IFN-γ production was due to dysregulated T cell responses to bacteria. Consistent with this, we observe a significant reduction in the number of regulatory MHCII+ ILC3s in the intestinal LP of Hic1Rorc mice at steady state (Fig 8A,B). Interestingly, analysis of CD4+ T cells from the mLN of Hic1Rorc mice at steady state reveals that in comparison to control mice, Hic1Rorc mice exhibited significantly increased frequencies of proliferating Ki67+ CD4+ T cells (Fig 8D), effector/effector memory CD44high CD62Llow CD4+ T cells (Fig 8E) as well as IFN-γ+ CD4+ T cells (Fig 8F), indicative of disrupted immune cell homeostasis. Consistent with these responses being driven by commensal bacteria, oral administration of a cocktail of antibiotics to Hic1Rorc mice was associated with significantly reduced peripheral IFN-γ+ CD4+ T cells and CD44high CD62Llow CD4+ T cells and mLN size (Fig 8C-F). Taken together, these results suggest that ILC3-intrinsic expression of HIC1 is required to limit commensal specific TH cell responses in the steady state.
Discussion
Our results demonstrate that in the steady state, HIC1 is expressed by intestinal ILCs in a Vitamin A-dependent manner. In the absence of HIC1, we observed a dramatic decrease in intestinal ILC3 numbers, which was associated with a failure to clear C. rodentium infection. In addition, the reduction of regulatory MHCII+ ILC3s resulted in an increased frequency IFN-γ-producing T cells locally and systemically. The heightened levels of IFN-γ in the absence of HIC1 inhibited the ability of Hic1Vav mice and Hic1Rorc mice to mount a protective TH2 cell-associated immune response against T. muris infection. Together, these results highlight an important role for HIC1 not only in regulating intestinal immune homeostasis but also in mounting proper immune responses to diverse intestinal infections.
In the absence of HIC1, we found a significant reduction in the number of ILC3s with no effect on ILC2s in the intestine. Specifically, there were reduced numbers of RORγt+ TBET+ ILC3s and an increase in the number of CD4+ LTi cells. This is consistent with studies that have demonstrated that these two lineages have distinct developmental pathways; LTi cells would develop in the fetus while TBET+ ILC3s develop postnatally and rely on environmental signals (Klose et al., 2013; Sanos et al., 2009; Spencer et al., 2014). Interestingly, it has been shown that atRA signalling is also important for generation of LTi cells in the fetus (van de Pavert et al., 2014). However, our results suggest that HIC1 is not involved in fetal LTi formation, as we find no differences in LTi numbers or lymphoid structures in the absence of HIC1. Further, the development of ILC progenitor cells in the bone marrow is not perturbed by loss of HIC1, suggesting that the primary role of HIC1 is to regulate the development and function of adult cells in the periphery.
Resistance to intestinal infection with C. rodentium is mediated by IL-22, and ILC3s are the predominant IL-22-producing cell population during the first week of infection (Zheng et al., 2008; Sonnenberg et al., 2011). There are contradictory studies on which ILC3 populations are key for resistance to C. rodentium with both CD4+ LTis and natural cytotoxicity receptor (NCR)+ ILC3s each being described as either individually critical or redundant (Rankin et al., 2016; Sonnenberg et al., 2011; Song et al., 2015). Another study looking at TBET+ ILC3s (which include NCR+ ILC3s) demonstrated that TBET expression in a subset of ILC3s is critical for resistance to C. rodentium infection (Rankin et al., 2013). Our results are consistent with a role for NCR+ or TBET+ ILC3s in immunity to C. rodentium as Hic1CD4 mice (deficient for HIC1 in T cells and LTi) are resistant to infection while Hic1Rorc mice (deficient for HIC1 in T cells and all ILC3s) are susceptible. Thus, HIC1 expression in ILC3s is critical for immunity to C. rodentium.
In addition to a reduction in ILC3s, a recent study identified that mice raised on a VAD diet displayed increased ILC2 numbers and heightened type 2 immunity to helminth infection in the intestine (Spencer et al., 2014). However, in the absence of HIC1, we did not observe an increase in ILC2 numbers nor increased resistance to infection with T. muris. Instead, we detected increased production of IFN-γ and an inability to mount a protective TH2 cell response to T. muris infection. Further, treatment of T. muris-infected Hic1Rorc mice with a neutralizing antibody against IFN-γ rendered the mice resistant to infection, demonstrating that HIC1-dependent responses are dispensable in the absence of IFN-γ and that the effects of RA on ILC2s are likely independent of HIC1.
Taken together, these results establish a role for the transcriptional repressor HIC1 as an atRA-responsive cell-intrinsic regulator of ILC3 cell function in the intestine, and identify a potential regulatory pathway that could be targeted to modulate ILC3 responses in the intestine.
METHODS
Ethics statement
Experiments were approved by the University of British Columbia Animal Care Committee (Protocol number A13-0010) and were in accordance with the Canadian Guidelines for Animal Research.
Mice
The generation of Hic1Citrine mice has been described (Pospichalova et al., 2011) and Hic1fl/fl mice will be described elsewhere (manuscript in preparation). Cd4-Cre mice were obtained from Taconic, Vav-Cre mice were obtained from T. Graf (Centre for Genomic Regulation, Barcelona, Spain) and CD11c-Cre (B6.Cg-Tg(Itgax-cre)1-1Reiz/J) and RORc-Cre (B6.FVB-Tg(RORc-cre)1Litt/J) mice were obtained from the Jackson Laboratory (Bar Harbor, ME, USA). Animals were maintained in a specific pathogen-free environment at the UBC Biomedical Research Centre animal facility.
Diet Studies
Vitamin A-deficient (TD.09838) diet was purchased from Harlan Teklad Diets. At day 14.5 of gestation, pregnant females were administered the vitamin A-deficient diet and maintained on diet until weaning of litter. Upon weaning, females were returned to standard chow, whereas weanlings were maintained on special diet until use.
Isolation of Lamina Propria Lymphocytes
Peyer’s patches were removed from the small intestine, which was cut open longitudinally, briefly washed with ice-cold PBS and cut into 1.5 cm pieces. Epithelium was stripped by incubated in 2mM EDTA PBS for 15 minutes at 37°C and extensively vortexed. Remaining tissue was digested with Collagenase/Dispase (Roche) (0.5 mg/mL) on a shaker at 250 rpm, 37°C, for 60 minutes, extensively vortexed and filtered through a 70µm cell strainer. The flow-through cell suspension was centrifuged at 1500rpm for 5 min. The cell pellet was re-suspend in 30% Percoll solution and centrifuged for 10 minutes at 1200 rpm. The pellet was collected and used as lamina propria lymphocytes.
Antibodies and flow cytometry
Absolute numbers of cells were determined via hemocytometer or with latex beads for LP samples. Intracellular cytokine (IC) staining was performed by stimulating cells with phorbol 12-myristate 13-acetate (PMA), ionomycin, and Brefeldin-A (Sigma) for 4 hours and fixing/permeabilizing cells using the eBioscience IC buffer kit. All antibody dilutions and cell staining were done with PBS containing 2% FCS, 1 mM EDTA, and 0.05% sodium azide. Fixable Viability Dye eFluor 506 was purchased from eBioscience to exclude dead cells from analyses. Prior to staining, samples were Fc-blocked with buffer containing anti-CD16/32 (93, eBioscience) and 1% rat serum to prevent non-specific antibody binding. Cells were stained with fluorescent conjugated anti-CD11b (M1/70), anti-CD11c (N418), anti-CD19 (ID3), anti-CD5 (53-7.3), anti-CD8 (53.67), anti-CD3 (KT3)(2C11), anti-NK1.1 (PK136), anti-B220 (atRA-6B2), anti-Ter119 (Ter119), anti-Gr1 (RB6-8C5) produced in house, anti-CD4 (GK1.5), anti-CD25 (PC61.5), anti-CD45 (30-F11), anti-CD90.2 (53-2.1), anti-Gata3 (TWAJ), anti-RORgt (B2D), anti-Tbet (eBio4B10), anti-Flt3 (A2F10), anti-ckit (ACK2), anti-TCRβ (H57-597), anti-MHCII (I-A/I-E) (M5/114.15.2), anti-F4/80 (BM8), anti-α4β7 (DATK32), anti-IFN-γ (XMG1.2), anti-IL-22 (IL22JOP), anti-IL-13 (eBio13A), anti-Ki67 (SolA15) purchased from eBioscience, anti-CD127 (5B/199), anti-CD62L (MEL-14), anti-CD44 (IM7), anti-CD64 (X54.5/7.1.1) purchased from BD Biosciences. Data were acquired on an LSR II flow cytometer (BD Biosciences) and analysed with FlowJo software (TreeStar).
Citrobacter rodentium infection
Mice were infected by oral gavage with 0.1 ml of an overnight culture of Luria-Bertani (LB) broth grown at 37°C with shaking (200 rpm) containing 2.5 × 108 cfu of C. rodentium (strain DBS100) (provided by B. Vallance, University of British Columbia, Vancouver, British Columbia, Canada). Mice were monitored and weighed daily throughout the experiment and sacrificed at various time points. For enumeration of C. rodentium, fecal pellets or livers were collected in pre-weighed 2.0 ml microtubes containing 1.0 ml of PBS and a 5.0 mm steel bead (Qiagen). Tubes containing pellets or livers were weighed, and then homogenized in a TissueLyser (Retche) for a total of 6 mins at 20 Hz at room temperature. Homogenates were serially diluted in PBS and plated onto LB agar plates containing 100 mg/ml streptomycin, incubated overnight at 37°C, and bacterial colonies were enumerated the following day, normalizing them to the tissue or fecal pellet weight (per gram).
Trichuris muris infection
Propagation of Trichuris muris eggs and infections were performed as previously described (Antignano et al., 2011). Mice were infected with approximately 150 – 200 embryonated T. muris eggs by oral gavage and monitered over a period of 21 days. Sacrificed mice were assessed for worm burdens by manually counting worms in the ceca using a dissecting microscope. Cecal tissues were fixed overnight in 10% buffered formalin and paraffin-embedded. A total of 5-µm-thick tissue sections were stained with periodic acid–Schiff (PAS) for histological analysis. mLNs were excised and passed through a 70 µm cell strainer to generate a single-cell suspension. mLN cells (4 × 106/mL) were cultured for 72 h in media containing 1 µg/mL each of antibodies against CD3 (145-2C11) and CD28 (37.51; eBioscience, San Diego, CA). Cytokine production from cell-free supernatant was quantified by ELISA using commercially available antibodies (eBioscience).
In vivo neutralization of IFN-γ
Mice were infected with T. muris as described above. On day 4 post infection, mice were injected i.p. with 500 µg of either control IgG or anti-IFN-γ (XMG1.2) (produced in-house by AbLabBiologics, UBC (Vancouver, BC)), constituted in sterile PBS. Mice were repeatedly injected thereafter on days 8, 12, and 16 prior to sacrifice on day 21.
RNA isolation and quantitative real-time PCR
Tissues were mechanically homogenized and RNA was extracted using the TRIzol method according to the manufacturer's instructions (Ambion). cDNA was generated using High Capacity cDNA reverse transcription kits (Applied Biosystems). Quantitative PCR was performed using SYBR FAST (Kapa Biosystems) and SYBR green-optimized primer sets run on an ABI 7900 real-time PCR machine (Applied Biosystems). Cycle threshold (CT) values were normalized relative to beta-actin (Actb) gene expression. The primers used were synthesized de novo: Il4 forward 5’- TCGGCATTTTGAACGAGGTC - 3’ and reverse 5’- CAAGCATGGAGTTTTCCCATG-3’; Il13 forward 5’- CCTGGCTCTTGCTTGCCTT-3’ and reverse 5’- GGTCTTGTGTGATGTTGCTCA-3’; Il17a forward 5’-AGCAGCGATCATCCCTCAAAG-3’ and reverse 5’-TCACAGAGGGATATCTATCAGGGTC-3’; Il22 forward 5’-ATGAGTTTTTCCCTTATGGGGAC-3’ and reverse 5’-GCTGGAAGTTGGACACCTCAA-3’; Ifng forward 5’-GGATGCATTCATGAGTATTGCC-3’ and reverse 5’-CCTTTTCCGCTTCCTGAGG-3’; Reg3g forward 5’-CCGTGCCTATGGCTCCTATTG- 3’ and reverse 5’-GCACAGACACAAGATGTCCTG −3’ Actb forward 5’-GGCTGTATTCCCCTCCATCG-3’ and reverse 5’-CCAGTTGGTAACAATGCCATGT-3’.
Serum ELISA
Serum was collected from mice 21 days post-infection with T.muris. Immulon plates (Thermo Fischer Scientific, NY) were coated with 5 µg/mL of dialyzed T. muris antigen overnight at 4°C. Wash buffer was PBS containing 0.05% Tween 20. Plates were blocked and serum samples were diluted in 3% bovine serum albumin in PBS/0.05% Tween 20. Serum samples were incubated on plates for 1 hour at room temperature. Plates were then incubated with rat anti-mouse IgG1 or IgG2a conjugated to horseradishperoxidase (BD Pharmingen, CA) for 1 hour at room temperature. Plates were developed using 3,3’,5,5’- tetramethylbenzidine (TMB) substrate (Mandel Scientific, ON) and stopped with 1N HCl. Plates were read at 450 nm on a Spectramax 384 (Molecular Devices, CA).
Antibiotic treatment
Mice received antibiotics (0.5 g/l of each ampicillin, gentamicin, neomycin and metronidazole, 0.25g/l vancomycin, with 4 g/l Splenda for taste) in their drinking water from weaning until euthanization (~10 week of age).
Statistics
Data are presented as mean ± S.E.M. A two-tailed Student’s t-test using GraphPad Prism 5 software determined statistical significance. Results were considered statistically significant with P < 0.05.
AUTHOR CONTRIBUTIONS
K.B., F.A., A.C., and C.Z. designed and performed the experiments. T.M.U. and V.K. provided assistance and contributed reagents and materials. K.B. and C.Z. wrote the manuscript.
ACKNOWLEDGMENTS
We would like to thank R. Dhesi, L. Rollins (BRC core), A. Johnson (UBCFlow), M. Williams (UBC AbLab), T. Murakami (BRC Genotyping), I. Barta (BRC Histology), and all members of BRC mouse facility for excellent technical assistance. This work was supported by the Canadian Institutes of Health Research's (CIHR) Canadian Epigenetics, Environment and Health Research Consortium (grant 128090 to C. Zaph) and operating grants (MOP-89773 and MOP-106623 to C. Zaph) and an Australian National Health and Medical Research Council (NHMRC) project grants (APP 1104433 and APP1104466 to C. Zaph). F. Antignano is the recipient of a CIHR/Canadian Association of Gastroenterology/Crohn’s and Colitis Foundation of Canada postdoctoral fellowship. C. Zaph is a Michael Smith Foundation for Health Research Career Investigator and a Veski Innovation Fellow.