Intact ATG16L1 plays an essential role in Paneth cell function and intestinal homeostasis. However, the functional consequences of ATG16L1 deficiency in myeloid cells, particularly macrophages, are not fully characterized. We generated mice with Atg16l1 deficiency in myeloid and dendritic cells and showed that mice with myeloid Atg16l1 deficiency had exacerbated colitis in two acute and one chronic model of colitis with increased proinflammatory to anti-inflammatory macrophage ratios, production of proinflammatory cytokines, and numbers of IgA-coated intestinal microbes. Mechanistic analyses using primary murine macrophages showed that Atg16l1 deficiency led to increased reactive oxygen species production, impaired mitophagy, reduced microbial killing, impaired processing of MHC class II Ags, and altered intracellular trafficking to the lysosomal compartments. Increased production of reactive oxygen species and reduced microbial killing may be general features of the myeloid compartment, as they were also observed in Atg16l1-deficient primary murine neutrophils. A missense polymorphism (Thr300Ala) in the essential autophagy gene ATG16L1 is associated with Crohn disease (CD). Previous studies showed that this polymorphism leads to enhanced cleavage of ATG16L1 T300A protein and thus reduced autophagy. Similar findings were shown in primary human macrophages from controls and a population of CD patients carrying the Atg16l1 T300A risk variant and who were controlled for NOD2 CD-associated variants. This study revealed that ATG16L1 deficiency led to alterations in macrophage function that contribute to the severity of CD.

Inflammatory bowel diseases (IBD) are chronic debilitating inflammatory conditions, which are classically differentiated into Crohn disease (CD) and ulcerative colitis. To date, >200 susceptibility loci have been identified (13). One of the genetic variants at position 300 in the autophagy gene ATG16L1 resulted in a threonine to alanine substitution (T300A) in the C-terminal domain (2, 4). Macroautophagy (herein referred to as autophagy) described a bulk degradation system where cytosolic constituents were engulfed in a double-membrane vesicle and targeted for degradation by lysosomal fusion (reviewed in Ref. 5). Selective autophagy, such as mitophagy to remove damaged mitochondria, also removed cellular danger signals and is thought to act as a break for the NLRP3 inflammasome (69).

The IBD-associated single nucleotide polymorphism (SNP) ATG16L1 T300A is a loss-of-function SNP that leads to increased caspase-mediated cleavage of ATG16L1 protein (1012). The functional consequences of ATG16L1 T300A were assessed in mice with knock-in of the human ATG16L1 T300A gene (10). The mice in this study developed defective and abnormal appearing Paneth cells and goblet cells, reduced bacteria clearance, increased IL-1β production, and worsened cecal inflammation with Salmonella infection compared with wild-type (WT) mice. Another group independently showed reduced bacteria clearance and increased expression of IL-1β in primary human ATG16L1 T300A macrophages and mouse ATG16L1 T316A knock-in (corresponding to human ATG16L1 T300A) macrophages (11).

The cell-specific role of autophagy was reported using conditional Atg16l1 knockout mice, in which Atg16l1 was deleted specifically in intestinal epithelial cells or CD11c+ dendritic cells (DC) (13). Compared to WT, mice with epithelial Atg16l1 deficiency exhibited Paneth cell abnormalities and were more susceptible to Salmonella typhimurium infection, whereas the phenotype of Atg16l1 deficiency in CD11c+ DC was similar to control WT mice. The investigators found that Atg16l1 in intestinal epithelial cells was important for antibacterial defense and maintenance of gut homeostasis, but that Atg16l1 may be dispensable in CD11c+ DC. An independent study showed that mice with Atg16l1 deficiency in intestinal epithelial cells develop severe spontaneous ileitis in the setting of Xbp1 deficiency (but not in single Atg16l1 or Xbp1 deficiency), indicating that Atg16l1 has a compensatory role in inflammation arising from endoplasmic reticulum stress (14). Additionally, conditional deletion of Atg16l1 in macrophages led to impaired clearance of Yersinia enterocolitica (11). Given the importance of macrophages and DC in innate immune responses, our goal was to further elucidate the role of Atg16l1 in myeloid cells. In the present study, we describe our independent generation of mice with Atg16l1 deficiency in myeloid cells and DC, and consequential worsening of colonic inflammation in two acute models and one chronic model of colitis in mice with Atg16l1-deficient myeloid cells. We further report our findings on non-IBD controls and CD patients who were either homozygous for WT or the ATG16L1 T300A variant and who did not carry NOD2 CD-associated variants, to dissect the direct functional effect of ATG16L1 versus an indirect effect through inflammation. Our results showed that the ATG16L1 T300A risk variant resulted in impaired autophagy with reduced conversion of LC3-I to LC3-II. Loss of ATG16L1 function resulted in increased proinflammatory polarization of primary macrophages in mice and humans. Mice with Atg16l1 deficiency in myeloid cells and humans who were homozygous for the ATG16L1 T300A risk variant exhibited increased colitogenic IgA-coated bacteria (15) and reduced mitophagy, which may lead to increased susceptibility to mucosal inflammation. Mechanistic studies of primary macrophages showed that reduced ATG16L1 function in mice and humans resulted in reduced clearance of S. typhimurium, Ag processing, and altered intracellular trafficking to the lysosomal compartments. Similarly, Atg16l1-deficient primary murine neutrophils also exhibited increased reactive oxygen species production and impaired clearance of S. typhimurium. Taken together, these results indicate the importance of autophagy in myeloid cells for the maintenance of intestinal homeostasis.

Cloning of Atg16l1 targeting vector and generation of Atg16l1f/f mice were performed in collaboration with genOway. Briefly, Atg16l1 endogenous locus containing 5.6 kb upstream and 2.1 kb downstream of exon 3 were generated by PCR amplification using proprietary C57BL/6J library developed at genOway. Subsequently, two loxP sites were inserted flanking Atg16l1 exon 3 (Fig. 1A). Positive selection neomycin gene flanked by FRT sites was inserted to the intron between exons 3 and 4 to generate the targeting vector (Fig. 1A). Every step of the cloning process was validated through restriction enzyme analysis and sequencing. The Atg16l1 gene-targeting construct was linearized and electroporated into genOway proprietary embryonic stem cells with C57BL/6J background. Homologous recombinants were selected by G418 and confirmed by PCR and Southern blot analysis. Embryonic stem cell clones with correct 5′ and 3′ recombination were microinjected into C57BL/6J blastocysts and introduced into pseudopregnant C57BL/6J mice. Male chimeric offspring were bred to obtain germline mutant mice that were then bred to flpe delete mouse strain to remove the neomycin cassette and were confirmed by Southern blot. To generate mice with conditional targeting of Atg16l1, Atg16l1f/f mice were bred with mice expressing cre recombinase under the control of villin (Villin-cre, The Jackson Laboratory stock 004586), LysM (LysM-cre, The Jackson Laboratory stock 004781), and CD11c (CD11c-cre, The Jackson Laboratory stock 007567). For Ag presentation assay, OTII/RAGII mice (Taconic model 1896) specific for OVA323–339 were used. All mice were in the C57BL/6J genetic background and were maintained under specific pathogen-free conditions in the Animal Facility at Cedars-Sinai Medical Center. This study was carried out in strict accordance with the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. Animal studies were approved by the Cedars-Sinai Medical Center Animal Care and Use Committee (protocol 3723).

FIGURE 1.

Atg16l1 deficiency in murine myeloid cells exacerbated acute and chronic colitis. (A) Schematic of Atg16l1 gene targeting. (B) RT-PCR of Atg16l1 mRNA in mouse BMM and bone marrow–derived DC (BMDC). Each filled circle represents an independent experiment, and data are expressed as percentage of β-actin expression. (C) Representative immunoblot of Atg16l1, LC3, and β-actin from three independent experiments. (D) Representative H&E-stained midcolon sections from 6-mo-old mice at original magnification ×100 are shown, and data from multiple mice are quantitated (n = 7–8 per group). Scale bars, 100 μm. (E) Disease activity index of Salmonella infection model (n = 12–15 per group), acute DSS (n = 10 per group), and chronic DSS (n = 10–19 per group) were quantitated and are shown. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 1.

Atg16l1 deficiency in murine myeloid cells exacerbated acute and chronic colitis. (A) Schematic of Atg16l1 gene targeting. (B) RT-PCR of Atg16l1 mRNA in mouse BMM and bone marrow–derived DC (BMDC). Each filled circle represents an independent experiment, and data are expressed as percentage of β-actin expression. (C) Representative immunoblot of Atg16l1, LC3, and β-actin from three independent experiments. (D) Representative H&E-stained midcolon sections from 6-mo-old mice at original magnification ×100 are shown, and data from multiple mice are quantitated (n = 7–8 per group). Scale bars, 100 μm. (E) Disease activity index of Salmonella infection model (n = 12–15 per group), acute DSS (n = 10 per group), and chronic DSS (n = 10–19 per group) were quantitated and are shown. *p < 0.05, **p < 0.01, ***p < 0.001.

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The protocol for S. typhimurium infection was performed as previously described with the following modification (16). S. typhimurium 14028 (gift from Andreas Baumler, University of California Davis) with nalidixic acid resistance were grown in Luria–Bertani broth supplemented with nalidixic acid (100 μg/ml) to OD 0.2–0.8. Mice pretreated with 20 mg of streptomycin (Sigma-Aldrich) were oral gavaged with 3 × 106 CFU of S. typhimurium and euthanized 5 d postinfection. Acute dextran sodium sulfate (DSS) was performed as previously described except 3% DSS in drinking water was used (17). Chronic DSS colitis using four cycles of 2.5% DSS in drinking water was performed as described (18). The disease activity index was calculated as described (18). Tissue samples were processed and stained with H&E by the Cedars-Sinai Medical Center Histology Core. Histopathological scores were assigned in a blinded manner by two trained animal pathologists using a previously established scoring system for S. typhimurium (16), acute DSS (17), and chronic DSS (18).

Genotyping was at Cedars-Sinai Medical Center using the Illumina Immuno BeadChip array as previously described (2, 19). All human subjects were homozygous non-risk for the three common CD-associated NOD2 variants (tagged by rs2066842-GG, rs2066844-GG, rs2066845-CC, and rs5743293-DD) and then stratified by ATG16L1 CD-associated T300A variant status, dividing the study subjects into those who were either homozygous non-risk or homozygous risk (as defined by rs2241880). Written informed consent was received from participants prior to inclusion in the study. All study subjects were identified by number and not by name and had not used antibiotics for at least 6 mo. Stool, blood samples, and genetic data were obtained from Material and Information Resources for Inflammatory and Digestive Diseases IBD Biobank and approved under Institutional Review Board nos. 3358 and Pro00027495.

Isolation and culture of murine lamina propria mononuclear cells (LPMC) and mesenteric lymph node (MLN) cells were carried out as previously reported (18). MLN cells and LPMC were cultured in 96-well round-bottom plates at 1.5 × 106 cells/ml of RPMI 1640 containing 2 mmol/l glutamine and 25 mmol/l HEPES (Corning), 10% FBS (Omega Scientific), 100 μM nonessential amino acids (Life Technologies), 1 mM sodium pyruvate (Fisher Scientific), 10 ng/ml LPS (InvivoGen), 50 μM 2-ME (Sigma-Aldrich), and 50 μg/ml gentamicin (Gemini Bio-Products). Media were harvested 48 h after culture and assessed for IL-1β and TNF-α production via an ELISA kit (eBioscience) per the manufacturer’s protocol. Mouse bone marrow–derived macrophages (BMM) and subsequent differentiation to M1 and M2 macrophages were performed as described (20). Murine neutrophils were isolated from mouse bone marrow using a neutrophil isolation kit (Stemcell Technologies) per the manufacturer’s protocol. Human peripheral blood monocyte-derived macrophages (PBM) were obtained by first isolating monocytes as previously described (21). Isolated monocytes then underwent M1 and M2 differentiation condition as previously described (20). For human studies, paired samples (WT and ATG16L1 T300A) were obtained for each PBM isolation and differentiation at the same day.

Total RNA was isolated using an RNeasy mini kit (Qiagen) per the manufacturer’s protocol. All RNA samples were reverse transcribed using an Omniscript reverse transcription kit (Qiagen). The following validated qPCR assays (Integrated DNA Technologies) were used to perform expression analysis: mouse β-actin, Mm.PT.39a.22214843; mouse Atg16l1, Mm.PT.58.19068201; mouse Ptgs2, Mm.PT.58.9154407; mouse Retnla, Mm.PT.58.43062398; mouse Nos2, Mm.PT.58.43705194; mouse Arg1, Mm.PT.58.8651372; human β-ACTIN, Hs.PT.56a.40703009.g; human ARG1, Hs.PT.58.40654839; human RETNLB, Hs.PT.58.1296566; human NOS2, Hs.PT.58.14740388; human PTGS2, Hs.PT.58.77266.

Cells were plated on 24-well plates overnight at 250,000 cells per well and lysed using sample buffer (Novex) with protease inhibitor (Calbiochem), boiled, and loaded onto premade SDS-polyacrylamide gels (Invitrogen), blocked for 60 min with 1% milk (Bio-Rad Laboratories), and stained overnight with the indicated primary Abs at 4°C. Blots were washed and stained with HRP-conjugated secondary Ab, and binding was detected by chemiluminescence (Thermo Scientific). Abs used included: Atg16l1, D6D5, 1:1000; LC3, D11, 1:1000; β-actin, D6A8, 1:1000; anti–phospho-p40phox; and HRP-linked secondary Ab (all from Cell Signaling Technology).

For flow cytometry, cells were acquired on an LSR II flow cytometer (BD Biosciences, San Jose, CA) and analyzed using FlowJo analysis software. Live cells were selected using Live/Dead stain (Life Technologies), and CD16/CD32 (clone 2.4G2) was used to block nonspecific FcR binding (eBioscience). For determination of intracellular cytokine production by leukocytes, cells were incubated for 5 h at 37°C with BD Leukocyte Activation Cocktail with BD GolgiPlug (BD Biosciences). Abs used were: mouse anti–S. typhimurium 0-4 Ab (Abcam), goat anti-mouse IgG H&L (Abcam), CD206 (clone C068C2; BioLegend), MHC class II (MHC-II; clone M5/114.15.2; BioLegend), CD86 (clone GL-1; BioLegend), NOS2 (clone CXNFT; eBioscience), CD11b (clone M170; BioLegend), Ly6C (clone HK1.4; BioLegend), CD45 (30-f11; clone BioLegend), CD4 (clone GK1.5; BioLegend), CD25 (clone PC61.5; BioLegend), CD103 (clone 2E7; eBioscience), CD64 (clone X54-5/7.1; BioLegend), TNF-α (clone MP6-XT22; BioLegend), and IL-1β (clone NJTEN3; eBioscience). Fecal IgA flow cytometry was performed on two fecal pellets that were collected directly from four to five cohoused mice of the same genotype at 4–5 mo of age or human fecal material shipped overnight with an ice pack and then flash frozen in liquid nitrogen. The fecal samples were homogenized in 1 ml of PBS per 100 mg fecal material and then centrifuged at 50 × g for 15 min to remove large particles. Supernatants were stained for IgA by flow cytometry as described (15). Total murine IgA was measured using a mouse IgA ELISA kit the per manufacturer’s protocol (eBioscience).

For immunofluorescent staining, 50,000 cells were plated on 12-mm-diameter coverslips in 24-well plates overnight. The next day, cells under starvation conditions were incubated with Earle’s balanced salt solution, and S. typhimurium 14028 (gift from A. Baumler, University of California Davis) was added to all wells (multiplicity of infection [MOI] of 20) and incubated for 1 h. Cells were incubated for 1- and 5-h periods with gentamicin to kill extracellular bacteria. Coverslips were washed, fixed with 4% paraformaldehyde (Electron Microscopy Sciences), blocked with 5% BSA (Fisher Scientific) and 0.01% Triton X-100 (Fisher Scientific), and primary Ab was added and incubated at 4°C overnight. Coverslips were washed and secondary Ab was added and incubated for 2 h in the dark at room temperature. Coverslips were washed and mounted to slides using DAPI (SouthernBiotech). Abs used included: anti–Beclin-1 1:100 (Novus Biologicals), anti-ATG16L1 1:100 (Cell Signaling Technology), anti-Lamp1 1:150 (Novus Biologicals), anti-Rab5 1:100 (Abcam), anti-Rab7 1:100 (Abcam), anti–S. typhimurium 0-4 Ab (1E6) 1:1000 (Abcam), goat polyclonal anti-LC3 1:100 (Biorbyt), goat anti-mouse IgG H&L 1:200 (Abcam), donkey polyclonal secondary Ab to rabbit IgG H&L 1:200 (Abcam), and donkey F(ab′)2 polyclonal secondary Ab to goat IgG H&L 1:500 (Abcam). Images were acquired on a Leica confocal microscope with an HCX PL APO CS 63.0×/1.30 GLYC (21°C) UV objective.

Reactive oxygen production was measured using luminol-ECL (Sigma-Aldrich) using macrophages and neutrophils isolated from mouse bone marrow and human macrophages differentiated from human monocytes as previously described (22). Mitochondrial reactive oxygen species (ROS) were determined with the addition of S. typhimurium 14028 (MOI of 100) and analyzed using flow cytometry after staining with MitoTracker Green (Life Technologies), MitoTracker Red (Life Technologies), or MitoSOX (Life Technologies) for 15 min at 37°C. Cells were washed and analyzed using flow cytometry.

Human PBM and mouse BMM were infected with S. typhimurium 14028 (MOI of 20) for 1 h. Cells were fixed in 2% glutaraldehyde/2% paraformaldehyde in PBS and incubated at 4°C overnight. Samples were transferred to the University of California Los Angeles Electron Microscopy Core, where they were embedded, sectioned, and placed on copper grids and stained with uranyl acetate and lead citrate. Images of sections were obtained using a JEOL 100CX transmission electron microscope. Images were analyzed with the assistance of the Electron Microscopy Core by counting vesicles, organelles, and Salmonella taken up by cells.

A gentamicin protection assay was performed by infecting human PBM, mouse BMM, and neutrophils with S. typhimurium as previously described (23) with the following modifications: intracellular killing was of S. typhimurium that was infected at MOI of 20. Infected cells were then incubated with gentamicin at 50 μg/ml for 1 h (macrophages) and 30 min (neutrophils) and plated for quantification.

Murine neutrophils were isolated from mouse bone marrow using a neutrophil isolation kit (Stemcell Technologies) per the manufacturer’s protocol. After cell viability was checked by trypan blue, neutrophils were suspended in DMEM/1% endotoxin-free BSA/10 mM HEPES, and 200,000 cells were placed in the upper insert of a 12-well Transwell chamber (5-μm pore size, 6.5-mm diameter; Costar Corning). The supernatant from bone marrow–derived WT macrophages stimulated with sonicated S. typhimurium or MIP-2 at 5 ng/ml was applied to the bottom well. After 60 min incubation at 37°C in 5% CO2, the cells in the bottom well were harvested by adding 50 μl of 70 mM EDTA and counted with a hemocytometer.

CD4+ T cells were isolated using the EasySep mouse CD4+ T cell isolation kit (Stemcell Technologies) from spleen from OTII/RAGII mice and stained with CellTrace CSFE (Thermo Fisher Scientific) according to the manufacturer’s recommendations. CFSE stained cells were cocultured (one BMM to four CD4+ T cells) with either WT or Atg16l1-deficient BMM that were exposed to whole OVA or OVA323–339 for 6 h. Cells from 72-h cocultures were collected and stained with CD4 GK1.5 (BioLegend) and Live/Dead stain (Fisher Scientific). After gating on live CD4+ T cells, CFSE staining was analyzed by flow cytometry to determine the CD4+ T cell proliferation.

Data are presented as the means ± SD. Comparison between two groups was performed by a two-tailed Fisher exact test for categorical variables and a Student t test for continuous variables. Parametric and nonparametric tests were used depending on the fulfillment of the test assumptions. Comparison between three groups was done using ANOVA, followed by pairwise post hoc analysis with a Tukey honest significant difference and Behrens–Fisher test correction for the multiple comparisons. A p value <0.05 was considered significant.

Because global deficiency of Atg16l1 is lethal to mice (7, 24, 25), we generated Atg16l1 floxed mice (hereafter called Atg16l1flox) to evaluate its role in gut mucosal homeostasis in colitis models (Fig. 1A). Using LysM-Cre and CD11c-Cre, we generated mice with Atg16l1 deficiency in myeloid cells (hereafter called Atg16l1∆Mye) and in DC (hereafter called Atg16l1∆DC), respectively. Deletion of Atg16l1 in myeloid cells and DC was confirmed by the lack of Atg16l1 mRNA and protein (Fig. 1B, 1C). Macrophages and DC with Atg16l1 deficiency also exhibited functional autophagy deficiency by the lack of conversion of LC3-I to LC3-II (Fig. 1C).

Under basal conditions, mice did not spontaneously develop colitis, and there were no histologic differences between the colons of control Atg16l1flox mice and mice with Atg16l1 deficiency in myeloid cells or in DC up to 10 mo of age (Fig. 1D). Next, Atg16l1flox, Atg16l1∆Mye, and Atg16l1∆DC mice underwent two acute models and one chronic model of murine colitis to assess the role of Atg16l1 on mucosal inflammation and to exclude colitis model–specific effects of Atg16l1. In all models, disease activity index scores were significantly higher in Atg16l1∆Mye (≥50% of measured time points) and in Atg16l1∆DC (latter part of colitic model) than in control Atg16l1flox mice (Fig. 1E). Histologic examination of the colon revealed worsened inflammation characterized by increased cellular infiltrate, mucin depletion, crypt abscesses, and architectural changes in Atg16l1∆Mye as compared with Atg16l1flox mice in all three models (Fig. 2A). Histological analysis showed that Atg16l1∆DC mice exhibited worsened colitis only in the chronic DSS-induced colitis model (Fig. 2A). In contrast to the basal and acute colitic conditions, we observed significant splenomegaly with increased splenocytes in the Atg16l1∆Mye and Atg16l1∆DC mice as compared with Atg16l1flox mice under chronic DSS-induced colitis (Fig. 2B). Except for MLN cells in the acute DSS model, we recovered more MLN cells and LPMC in Atg16l1∆Mye than in Atg16l1flox mice (Fig. 2C). In the chronic DSS-induced colitis model, Atg16l1∆DC mice had more MLN cells than did Atg16l1flox mice. These results indicated that Atg16l1 deficiency in myeloid cells (and to a lesser extent in DC) exacerbated colitis in murine models.

FIGURE 2.

Splenomegaly and increased cellular infiltrate in mice with Atg16l1 deficiency in myeloid cells. (A) Representative H&E-stained midcolon sections at original magnification ×100 are shown, and histologic inflammation scores are quantitated. Scale bars, 100 μm. (B) Spleen weight in grams and splenocyte number were quantitated from chronic DSS-induced colitis in WT (Flox) mice or from mice with Atg16l1 deficiency in myeloid cells (∆Mye) and DC (∆DC). (C) Cell recovery from MLN and LPMC were quantitated and are shown. Each filled circle represents data from an individual mouse. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 2.

Splenomegaly and increased cellular infiltrate in mice with Atg16l1 deficiency in myeloid cells. (A) Representative H&E-stained midcolon sections at original magnification ×100 are shown, and histologic inflammation scores are quantitated. Scale bars, 100 μm. (B) Spleen weight in grams and splenocyte number were quantitated from chronic DSS-induced colitis in WT (Flox) mice or from mice with Atg16l1 deficiency in myeloid cells (∆Mye) and DC (∆DC). (C) Cell recovery from MLN and LPMC were quantitated and are shown. Each filled circle represents data from an individual mouse. *p < 0.05, **p < 0.01, ***p < 0.001.

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Flow cytometry analysis did not reveal any differences in the frequencies of macrophages (MHC-II+CD11b+F4/80+), DC (MHC-II+CD11b+CD11c+), or expression of the activation marker CD86 on macrophages and DC in the MLN and lamina propria of the three colitic models used (data not shown). We observed an increased percentage of macrophages and DC producing IL-1β (but not TNF-α) in Atg16l1∆Mye mice and in DC in Atg16l1∆DC mice compared with control Atg16l1flox mice (Fig. 3A, 3B, data not shown). Upon examining isolated cells from the MLN and LPMC, we observed increased IL-1β and TNF-α, but not IL-6, production in most (if not all) colitis models used (Fig. 3C–F).

FIGURE 3.

Increased production of IL-1β and TNF-α with Atg16l1 deficiency in myeloid cells. MHC-II+CD11b+F4/80+ macrophages (A) and MHC-II+CD11b+CD11c+ DC (B) were stained for intracellular IL-1β and TNF-α expression and quantitated for Salmonella infection (right panel), acute DSS (middle panel), and chronic DSS (right panel). Isolated mononuclear cells from MLN and LPMC from the three murine colitis models were cultured for 3 d, and the levels of secreted IL-1β in MLN (C) and LPMC (D) as well as TNF-α in MLN (E) and LPMC (F) were assessed by ELISA. Each filled circle for MLN represents the value obtained from a single mouse. Each filled circle for LPMC represents the value obtained from an independent experiment using pooled samples from two mice. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 3.

Increased production of IL-1β and TNF-α with Atg16l1 deficiency in myeloid cells. MHC-II+CD11b+F4/80+ macrophages (A) and MHC-II+CD11b+CD11c+ DC (B) were stained for intracellular IL-1β and TNF-α expression and quantitated for Salmonella infection (right panel), acute DSS (middle panel), and chronic DSS (right panel). Isolated mononuclear cells from MLN and LPMC from the three murine colitis models were cultured for 3 d, and the levels of secreted IL-1β in MLN (C) and LPMC (D) as well as TNF-α in MLN (E) and LPMC (F) were assessed by ELISA. Each filled circle for MLN represents the value obtained from a single mouse. Each filled circle for LPMC represents the value obtained from an independent experiment using pooled samples from two mice. *p < 0.05, **p < 0.01, ***p < 0.001.

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Because Atg16l1∆Mye (but not Atg16l1∆DC) mice exhibited increased colonic inflammation in all three models of colitis (Fig. 2A), we focused on Atg16l1∆Mye mice for the remainder of the murine experiments covered in the present study. Because appropriate balance and function of the proinflammatory P2 stage (MHC-II+Ly6C+) and anti-inflammatory P3/4 stages (MHC-II+Ly6C) of macrophages (CD45+CD11b+CD64+CD103) have been implicated in maintenance of mucosal immune homeostasis (20, 26), we next sought to determine whether Atg16l1 deficiency in myeloid cells altered these intestinal macrophage subsets. Prior to initiation of colitis, we observed an increase in the percentage of P2 cells and a reduction in the percentage of P3/4 cells in the LPMC of Atg16l1∆Mye mice compared with control Atg16l1flox mice (Fig. 4A, 4B). Similarly, expression analysis showed a consistent increase in the proinflammatory macrophage markers Nos2 and Ptgs2 in LPMC isolated from Atg16l1∆Mye compared with control Atg16l1flox mice prior to the initiation of colitis (Fig. 4C). Similar levels of the regulatory macrophage markers Retnla and Arg1 were measured in the isolated LPMC between Atg16l1∆Mye compared with control Atg16l1flox mice prior to the initiation of colitis (Fig. 4C). Once colitis was established, flow cytometry analysis demonstrated an increase in proinflammatory P2 cells in all colitis models, and a reduction in anti-inflammatory P3/4 macrophage populations was found in the chronic DSS conditions in Atg16l1∆Mye as compared with control Atg16l1flox mice (Fig. 4A, 4B).

FIGURE 4.

Increased number of proinflammatory macrophages with Atg16l1 deficiency. Macrophage subsets in LPMC were assessed as proinflammatory (P2) and anti-inflammatory (P3/4) macrophages in 2-mo-old noncolitic mice and in the three colitis models. Representative flow cytometry plots of gated CD45+CD11b+CD64+CD103 cells are shown (A) and quantitated (B). (C) RT-PCR of mRNA for proinflammatory macrophage markers (Nos2, Ptgs2) and regulatory macrophage markers (Retnla, Arg1) were measured from LPMC macrophages isolated from 2-mo-old noncolitic mice and are represented as percentage of β-actin reference gene expression. (D) RT-PCR of mRNA for Nos2 and Ptgs2 were determined in BMM cultured in M1 polarizing conditions, and mRNA for Retnla and Arg1 were determined in BMM cultured in M2 polarizing conditions. Each filled circle represents an independent experiment using pooled LPMC from two mice of the same genotype for (B) and from a single mouse in (C) and (D). *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 4.

Increased number of proinflammatory macrophages with Atg16l1 deficiency. Macrophage subsets in LPMC were assessed as proinflammatory (P2) and anti-inflammatory (P3/4) macrophages in 2-mo-old noncolitic mice and in the three colitis models. Representative flow cytometry plots of gated CD45+CD11b+CD64+CD103 cells are shown (A) and quantitated (B). (C) RT-PCR of mRNA for proinflammatory macrophage markers (Nos2, Ptgs2) and regulatory macrophage markers (Retnla, Arg1) were measured from LPMC macrophages isolated from 2-mo-old noncolitic mice and are represented as percentage of β-actin reference gene expression. (D) RT-PCR of mRNA for Nos2 and Ptgs2 were determined in BMM cultured in M1 polarizing conditions, and mRNA for Retnla and Arg1 were determined in BMM cultured in M2 polarizing conditions. Each filled circle represents an independent experiment using pooled LPMC from two mice of the same genotype for (B) and from a single mouse in (C) and (D). *p < 0.05, **p < 0.01, ***p < 0.001.

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To test whether autophagy was involved in the development of proinflammatory and regulatory macrophages, we assessed whether there are differences in the differentiation of bone marrow–derived proinflammatory M1 and regulatory M2 macrophages in vitro. Littermate Atg16l1flox and Atg16l1∆Mye BMM were cultured in either M1 (LPS and IFN-γ) or M2 conditions (TGF-β, IL-10). Compared to Atg16l1flox, there was increased expression of the M1 markers Nos2 and Ptgs2 in Atg16l1∆Mye BMM cultured in M1 conditions (Fig. 4D). No differences in the M2 markers Retnla and Arg1 were observed between Atg16l1flox and Atg16l1∆Mye BMM cultured in M2 conditions (Fig. 4D). Taken together, the data showed that inhibition of macrophage autophagy promoted M1 macrophage polarization.

IgA coating has been shown to identify IBD-driving microbes in mice and humans (15). To assess whether Atg16l1 deficiency in myeloid cells could led to increased IgA-coated bacteria that may be colitogenic, we examined IgA staining in stool collected from noninflamed 2- to 3-mo-old Atg16l1flox and Atg16l1∆Mye mice. Flow cytometry showed that Atg16l1∆Mye mice had increased IgA coating of bacteria compared with control Atg16l1flox mice (Supplemental Fig. 1A). Induction of colitis led to a further significant increase in IgA-coated fecal bacteria in all models (Supplemental Fig. 1A). Total IgA in the stools was quantitated by IgA ELISA to determine whether increased IgA coating of fecal bacteria was due to increased IgA secretion into the intestinal lumen. Total IgA in the stool was less in mice with Atg16l1 deficiency than in mice without Atg16l1 deficiency prior to induction of colitis (Supplemental Fig. 1B), suggesting that the increased IgA-coated bacteria were not driven by increased luminal IgA. Total stool IgA was increased in the Salmonella infection model but not in the acute or chronic DSS model (Supplemental Fig. 1B), which is likely due to impaired clearance of Salmonella by macrophages (Fig. 5A). The worsened colitis in Atg16l1∆Mye mice as compared with control Atg16l1flox mice may in part be due to increased colitogenic IgA-coated bacteria in Atg16l1∆Mye mice.

FIGURE 5.

Impaired clearance and altered cellular trafficking of S. typhimurium in Atg16l1-deficient BMM. WT and Atg16l1∆Mye BMM were infected with S. typhimurium, and surviving intracellular bacteria were quantified (A), representative TEM images of WT (n = 17) and Atg16l1-deficient (n = 25) BMM infected with S. typhimurium are shown (B), and vesicles containing multiple Salmonella were quantitated (C). White arrowheads indicate vesicles containing S. typhimurium. Scale bars, 2 μm. Each filled circle represents an independent experiment (A) or data from a single macrophage (C). (D) Representative flow cytometry plot of intracellular Salmonella (left panel), quantitated as percentage (middle panel), and MFI (right panel) are shown (n = 5 independent experiments). Representative confocal image of WT or Atg16l1-deficient BMM stained with anti-Salmonella (green stain, top panels) and anti-Lamp1, anti-Rab5, and anti-Rab7 (violet stain, middle panels) are shown (E) and quantitated (F). Colocalized images where green stain overlaps with purple stain are marked by white arrowheads (merged white stain, bottom panels). Scale bars, 20 μm. Each filled circle represents data acquired from an individual BMM from three independent experiments. (G) Quantification of S. typhimurium in either double-membrane versus single-membrane vesicles obtained from TEM in each BMM are shown as mean ± SD. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 5.

Impaired clearance and altered cellular trafficking of S. typhimurium in Atg16l1-deficient BMM. WT and Atg16l1∆Mye BMM were infected with S. typhimurium, and surviving intracellular bacteria were quantified (A), representative TEM images of WT (n = 17) and Atg16l1-deficient (n = 25) BMM infected with S. typhimurium are shown (B), and vesicles containing multiple Salmonella were quantitated (C). White arrowheads indicate vesicles containing S. typhimurium. Scale bars, 2 μm. Each filled circle represents an independent experiment (A) or data from a single macrophage (C). (D) Representative flow cytometry plot of intracellular Salmonella (left panel), quantitated as percentage (middle panel), and MFI (right panel) are shown (n = 5 independent experiments). Representative confocal image of WT or Atg16l1-deficient BMM stained with anti-Salmonella (green stain, top panels) and anti-Lamp1, anti-Rab5, and anti-Rab7 (violet stain, middle panels) are shown (E) and quantitated (F). Colocalized images where green stain overlaps with purple stain are marked by white arrowheads (merged white stain, bottom panels). Scale bars, 20 μm. Each filled circle represents data acquired from an individual BMM from three independent experiments. (G) Quantification of S. typhimurium in either double-membrane versus single-membrane vesicles obtained from TEM in each BMM are shown as mean ± SD. *p < 0.05, **p < 0.01, ***p < 0.001.

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Owing to the importance of autophagy in bacterial clearance, we hypothesized that Atg16l1 deficiency could lead to impaired macrophage clearance of bacteria. Infection of Atg16l1-deficient BMM with S. typhimurium led to reduced bacteria clearance (Fig. 5A). When sections of S. typhimurium–infected macrophages were visualized with transmission electron microscopy (TEM), we observed more vacuoles containing multiple Salmonella in Atg16l1∆Mye compared with Atg16l1flox BMM (Fig. 5B, 5C). We determined the percentage of bacteria in macrophages to control for potential differences in bacteria uptake between Atg16l1∆Mye BMM. Although there were no differences in the percentage of macrophages containing S. typhimurium (Fig. 5D, left and middle panels), the mean fluorescence intensity (MFI) of the Salmonella stain was significantly increased at 60 min after S. typhimurium infection (Fig. 5D, right panel), indicating that there were more S. typhimurium per macrophage.

We hypothesize that the impaired S. typhimurium clearance that leads to its increase in macrophages in Atg16l1-deficient macrophages may be due to reduced intracellular trafficking of S. typhimurium to intracellular compartments for bacteria killing. Consistent with our hypothesis, colocalization of S. typhimurium with the lysosome marker Lamp1 was reduced (Fig. 5E, 5F). Colocalization of S. typhimurium with early autophagy markers (Beclin 1 and LC3) was similar between WT and Atg16l1-deficient macrophages (Fig. 5E, 5F). Interestingly, colocalization of Salmonella with the endosomal markers Rab5 and Rab7 was increased (Fig. 5E, 5F). Quantification of double-membrane (autophagy) and single-membrane (nonautophagy) vesicles revealed reduction of Salmonella in double-membrane vesicles but increased Salmonella in single-membrane vesicles (Fig. 5G). Taken together, the data showed that Atg16l1 deficiency in macrophages led to a reduction of bacteria clearance, due in part to decreased intracellular trafficking of bacteria to Lamp1-containing vacuoles for bacterial degradation.

ROS are cytotoxic for a variety of microorganisms, including Salmonella (2729). We therefore hypothesized that ROS production might be defective in Atg16l1-deficient cells. Unexpectedly, we observed that Atg16l1-deficient macrophages had higher total cellular generation of ROS when infected with S. typhimurium (Fig. 6A, left panel) or exposed to zymosan (Fig. 6A, right panel) than did WT macrophages. As an independent approach to assess NADPH oxidase activation, we measured phosphorylation of the cytosolic p40 subunit. When macrophages were infected with S. typhimurium, p40phox was more phosphorylated in Atg16l1-deficient cells than in control cells (Fig. 6B). These data suggested that Atg16l1 plays a role in limiting ROS production by macrophages and that impaired bacteria killing is not due to reduced ROS production.

FIGURE 6.

Atg16l1 deficiency increased ROS production and altered mitochondria homeostasis. (A) Relative ROS production as measured by luminol-dependent chemiluminescence was determined during 60 min for WT (Atg16l1flox) and Atg16l1-deficient (Atg16l1∆Mye) BMM treated with S. typhimurium or zymosan (filled diamond and solid line) and at baseline (filled diamond). Data of eight independent experiments with similar results are shown. (B) Phospho-p40phox and β-actin levels in WT and Atg16l1-deficient BMM at baseline and with S. typhimurium infection were determined by immunoblotting. (C) Representative TEM images of WT and Atg16l1-deficient BMM are shown. Scale bars, 2 μm. White arrowheads indicate mitochondria, and mitochondria per BMM are quantitated. (D) Representative flow cytometry plot of WT and Atg16l1-deficient BMM left unstained (filled) or labeled with MitoSOX (open) are shown and MFI is quantitated. (E) Representative flow cytometry plot of WT or Atg16l1-deficient BMM with and without S. typhimurium infection and stained with MitoTracker Deep Red and MitoTracker Green are shown. Each filled circle represents an independent experiment from a single mouse. *p < 0.05.

FIGURE 6.

Atg16l1 deficiency increased ROS production and altered mitochondria homeostasis. (A) Relative ROS production as measured by luminol-dependent chemiluminescence was determined during 60 min for WT (Atg16l1flox) and Atg16l1-deficient (Atg16l1∆Mye) BMM treated with S. typhimurium or zymosan (filled diamond and solid line) and at baseline (filled diamond). Data of eight independent experiments with similar results are shown. (B) Phospho-p40phox and β-actin levels in WT and Atg16l1-deficient BMM at baseline and with S. typhimurium infection were determined by immunoblotting. (C) Representative TEM images of WT and Atg16l1-deficient BMM are shown. Scale bars, 2 μm. White arrowheads indicate mitochondria, and mitochondria per BMM are quantitated. (D) Representative flow cytometry plot of WT and Atg16l1-deficient BMM left unstained (filled) or labeled with MitoSOX (open) are shown and MFI is quantitated. (E) Representative flow cytometry plot of WT or Atg16l1-deficient BMM with and without S. typhimurium infection and stained with MitoTracker Deep Red and MitoTracker Green are shown. Each filled circle represents an independent experiment from a single mouse. *p < 0.05.

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Selective autophagy such as mitophagy that removes dysfunctional mitochondria has been shown to limit IL-1β production and inflammation (6, 7, 9). Therefore, we hypothesized that Atg16l1-deficient BMMs may contain more mitochondria and produce increased ROS than do control BMM. Consistent with our hypothesis, Atg16l1-deficient BMM had increased macrophages per cell (Fig. 6C) and produced more ROS than did controls (Fig. 6D). We next hypothesized that Atg16l1-deficient BMM contained more damaged mitochondria with reduced membrane potential. Thus, we measured mitochondria membrane potential using MitoTracker Deep Red and counterstained with MitoTracker Green, a probe that stains mitochondrial membrane lipids independent of membrane potential. Consistent with our hypothesis, we observed a reduction of membrane potential in mitochondria in Atg16l1-deficient cells compared with WT (Fig. 6E). Taken together, our data suggest that the increased overall ROS through direct cytotoxic damage and impaired mitophagy with Atg16l1 deficiency could contribute to worsened colitis in our models.

To assess whether cellular defects seen with Atg16l1 deficiency (e.g., altered Ag trafficking) affected presentation of MHC-II Ags, we exposed WT or Atg16l1-deficient macrophages to full-length OVA protein and cocultured them with CellTrace-labeled OVA-specific OT-II CD4+ T cells isolated from OTII/RAGII−/− mice. These genetically modified T cells recognize MHC-II–restricted presentation of OVA323–339 (30). Proliferation of OT-II cells was reduced when stimulated with Atg16l1-deficient macrophages compared with WT macrophages (Fig. 7A). To dissect the steps of Atg16l1 deficiency impairment of MHC-II–restricted Ag presentation to T cells, we pulsed WT and Atg16l1-deficient macrophages with the OVA323–339 synthetic peptide that did not need further intracellular processing to be loaded onto MHC-II. WT and Atg16l1-deficient macrophages induced a similar degree of proliferation to CellTrace-labeled OVA-specific OT-II CD4+ T cells (Fig. 7B). These data were consistent with the idea that the impaired macrophage presentation of Ag was due to altered processing of Ag and not due to deficient loading of Ag to MHC-II protein.

FIGURE 7.

Atg16l1 is required for optimal Ag processing by macrophages for MHC-II Ag presentation. Representative flow cytometry plots of proliferating OT-II CD4+ T cells labeled with CellTrace stimulated with BMM treated with whole OVA protein (A) or OVA323–339 (B) are shown. Decreased CellTrace fluorescence intensity indicated proliferation. Each filled circle represents an independent experiment. *p < 0.05.

FIGURE 7.

Atg16l1 is required for optimal Ag processing by macrophages for MHC-II Ag presentation. Representative flow cytometry plots of proliferating OT-II CD4+ T cells labeled with CellTrace stimulated with BMM treated with whole OVA protein (A) or OVA323–339 (B) are shown. Decreased CellTrace fluorescence intensity indicated proliferation. Each filled circle represents an independent experiment. *p < 0.05.

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Because utilization of the LysM-Cre system leads to deletion of the floxed allele in the myeloid compartment, deletion efficiency of nearly 100% in granulocytes including neutrophils was also observed (31). We hypothesized that Atg16l1 deficiency in neutrophils will also lead to impaired S. typhimurium clearance and increased ROS production that were observed in Atg16l1-deficient murine neutrophils. To assess neutrophil function, we isolated primary murine neutrophils from bone marrow of WT and Atg16l1∆Mye mice. Atg16l1-deficient neutrophils had similar chemotaxis/recruitment in response to supernatant from WT neutrophils cocultured with S. typhimurium (Supplemental Fig. 2A). Consistent with the altered Atg16l1-deficient murine macrophages, we also observed that murine neutrophils with Atg16l1 deficiency exhibit increased ROS production and impaired clearance of S. typhimurium (Supplemental Fig. 2B, 2C). Our data indicate that increased production of ROS and reduced microbial killing may be general features of Atg16l1 deficiency in the myeloid compartment.

Several studies have indicated that the IBD-associated ATG16L1 T300A variant is a loss-of-function SNP that leads to increased caspase-mediated cleavage of ATG16L1 protein, resulting in reduced autophagy (10, 11). Our murine data on Atg16l1 deficiency suggested that macrophages from people carrying the T300A SNP might be more M1-polarized, more inflammatory, and less able to kill Salmonella. We therefore isolated primary PBM from subjects who were homozygous for either WT or ATG16L1 T300A SNP for functional human studies. Because ATG16L1 has been reported to interact with NOD2, we controlled for the IBD-associated NOD2 polymorphism by using only cells that lacked the NOD2 risk allele (12, 23, 3234). Additionally, we compared PBM from non-IBD and CD subjects to exclude the possibility that changes seen were due to CD.

Macrophages from subjects homozygous for the risk ATG16L1 T300A allele exhibited significantly reduced autophagy (reduced LC3-I to LC3-II conversion) upon induction with S. typhimurium but not under basal conditions in both non-IBD subjects (Fig. 8A) and CD patients (Fig. 8B). Similar to murine Atg16l1-deficint macrophages, PBM from ATG16L1 T300A risk variant homozygotes from both non-IBD subjects (Fig. 8C) and CD patients (Fig. 8D) exhibited increased polarization to M1 proinflammatory macrophages with an increase in the M1 markers NOS2 and PTGS2. Consistent with murine findings, individuals who were homozygous for the CD risk ATG16L1 T300A allele had more IgA-coated bacteria in the stool than did individuals not carrying the risk allele (Supplemental Fig. 1C). However, the effect of IgA coating was more profound in CD patients compared with control subjects (Supplemental Fig. 1C). These data showed that CD-associated ATG16L1 T300A variants are associated with reduced autophagy and other proinflammatory changes (increased M1 polarization and IgA-coated stool) that may explain the increased susceptibility to CD in individuals carrying ATG16L1 T300A risk alleles.

FIGURE 8.

Reduced autophagy and proinflammatory macrophage polarization with the human CD risk ANP ATG16L1 T300A. Representative immunoblot of LC3 and β-actin from non-IBD control (A) and CD patients (B) who were homozygous for risk or non-risk ATG16L1 T300A alleles are shown and quantitated as percentage of β-actin reference expression. Nos2 and Ptgs2 mRNA levels were measured by RT-PCR in PBM cultured in M1-polarizing conditions, and mRNA for Retnla and Arg1 were measured in PBM cultured in M2-polarizing conditions. mRNA expression is expressed as percentage of β-actin reference expression for non-IBD controls (C) and CD patients (D). (E) Representative flow cytometry plots of fecal bacteria stained with anti-IgA Ab from non-IBD controls and from CD patients are shown and quantitated. Each experiment was done in pairs with an individual of each genotype for ATG16L1. (F) PBM from risk or non-risk ATG16L1 T300A variant subjects were infected with S. typhimurium, and surviving intracellular bacteria were quantified. Each filled circle is representative of an independent experiment from a human subject. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 8.

Reduced autophagy and proinflammatory macrophage polarization with the human CD risk ANP ATG16L1 T300A. Representative immunoblot of LC3 and β-actin from non-IBD control (A) and CD patients (B) who were homozygous for risk or non-risk ATG16L1 T300A alleles are shown and quantitated as percentage of β-actin reference expression. Nos2 and Ptgs2 mRNA levels were measured by RT-PCR in PBM cultured in M1-polarizing conditions, and mRNA for Retnla and Arg1 were measured in PBM cultured in M2-polarizing conditions. mRNA expression is expressed as percentage of β-actin reference expression for non-IBD controls (C) and CD patients (D). (E) Representative flow cytometry plots of fecal bacteria stained with anti-IgA Ab from non-IBD controls and from CD patients are shown and quantitated. Each experiment was done in pairs with an individual of each genotype for ATG16L1. (F) PBM from risk or non-risk ATG16L1 T300A variant subjects were infected with S. typhimurium, and surviving intracellular bacteria were quantified. Each filled circle is representative of an independent experiment from a human subject. *p < 0.05, **p < 0.01, ***p < 0.001.

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We sought to determine whether the ATG16L1 T300A impaired bacteria clearance in human PBM similar to murine Atg16l1-deficient BMM. Infection of PBM with S. typhimurium led to reduced bacteria clearance in both non-IBD and CD subjects homozygous for the ATG16L1 T300A CD-associated allele (Fig. 8E). The reduction in bacterial clearance was not secondary to reduced uptake because S. typhimurium uptake did not vary with ATG16L1 T300A status (Fig. 9A). In contrast to our findings in the mouse, Salmonella-infected non-IBD human PBM from individuals with the ATG16L1 T300A risk alleles did not exhibit increased numbers of vacuoles containing multiple Salmonella (Fig. 9B, 9C). By TEM, we observed reduced numbers of S. typhimurium in double-membrane vacuoles in the risk ATG16L1 T300A homozygotes (Fig. 9D), indicating that the risk variant is associated with reduced autophagy. We next examined whether our finding in mice of defective intracellular trafficking of S. typhimurium to the LAMP1+ lysosomal compartments also occurred in humans. Consistent with the murine data, there was reduced colocalization of S. typhimurium with the lysosome marker LAMP1 in ATG16L1 T300A PBM (Fig. 9E). Colocalization of S. typhimurium with early autophagy markers (Beclin1 and LC3) and with the endosomal markers (Rab5 and Rab7) were similar (Fig. 9E). Taken together, these data suggested that reduced bacterial clearance could be in part due to altered intracellular trafficking of bacteria to lysosomes for degradation.

FIGURE 9.

Altered intracellular trafficking in PBM with CD risk ATG16L1 T300A variant. (A) Representative flow cytometry plots (n = 4 independent experiments from four subjects per group) of PBM from risk and non-risk ATG16L1 T300A homozygotes that were uninfected (dotted line) or infected with S. typhimurium and stained with anti-Salmonella Ab (solid line) are shown. Representative TEM image of PBM from homozygotes risk and non-risk for CD-associated ATG16L1 T300A variant infected with S. typhimurium is shown (B) and quantitated for number of intracellular vesicles that contain more than one Salmonella (C) and for the number of single- and double-membrane vesicles that contain bacteria (D). Each filled circle represents an independent experiment from at least six subjects per group. Scale bars in TEM, 2 μm. (E) Representative confocal microscopy image of PBM from subjects with risk or non-risk ATG16L1 T300A variants infected with S. typhimurium and stained with anti-Salmonella (green stain) and anti-Lamp1 (violet stain) at original magnification ×630 are shown. Colocalized regions where green stain overlaps with purple stain are marked by white arrowheads (merged white stain) and represented by percentage of colocalization. Scale bars in confocal images, 20 μm. Each filled circle in (C)–(E) represents a value obtained from a PBM from at least six subjects per group.

FIGURE 9.

Altered intracellular trafficking in PBM with CD risk ATG16L1 T300A variant. (A) Representative flow cytometry plots (n = 4 independent experiments from four subjects per group) of PBM from risk and non-risk ATG16L1 T300A homozygotes that were uninfected (dotted line) or infected with S. typhimurium and stained with anti-Salmonella Ab (solid line) are shown. Representative TEM image of PBM from homozygotes risk and non-risk for CD-associated ATG16L1 T300A variant infected with S. typhimurium is shown (B) and quantitated for number of intracellular vesicles that contain more than one Salmonella (C) and for the number of single- and double-membrane vesicles that contain bacteria (D). Each filled circle represents an independent experiment from at least six subjects per group. Scale bars in TEM, 2 μm. (E) Representative confocal microscopy image of PBM from subjects with risk or non-risk ATG16L1 T300A variants infected with S. typhimurium and stained with anti-Salmonella (green stain) and anti-Lamp1 (violet stain) at original magnification ×630 are shown. Colocalized regions where green stain overlaps with purple stain are marked by white arrowheads (merged white stain) and represented by percentage of colocalization. Scale bars in confocal images, 20 μm. Each filled circle in (C)–(E) represents a value obtained from a PBM from at least six subjects per group.

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Because Atg16l1 deficiency increased ROS production, we hypothesized that PBM with ATG16L1 T300A SNP will also exhibit increased ROS production. Homozygote risk PBMs with ATG16L1 T300A SNP had increased total cellular generation of ROS when exposed to S. typhimurium or zymosan compared with non-risk macrophages (Fig. 10A). We then examined the effect of the ATG16L1 risk variant on human PBM mitochondria and mitochondrial ROS production. Similar to the murine Atg16l1 deficiency, mitochondria numbers were increased in macrophages from ATG16L1 risk homozygotes compared with non-risk homozygotes (Fig. 10B). MitoSOX fluorescent signal was enhanced in cells from the risk homozygotes for both non-IBD subjects (Fig. 11A, left panel) and CD patients (Fig. 11B, left panel), indicating that there was increased ROS accumulation in the mitochondria of ATG16L1 risk homozygotes. Using MitoTracker Deep Red and MitoTracker Green, we observed a reduction of membrane potential in the mitochondria of cells from the risk homozygotes for both non-IBD subjects (Fig. 11A, middle and right panels) and for CD patients (Fig. 11B, middle and right panels). Taken together, the data showed that macrophages from subjects with the ATG16L1 CD-associated risk variants had defects in autophagy, inflammatory signaling, ROS production, and mitochondrial function that are similar to those observed in murine macrophages deficient in Atg16l1.

FIGURE 10.

CD risk ATG16L1 T300A increased ROS production and mitochondria number. (A) Relative ROS production, as measured by luminol-dependent chemiluminescence, was determined for risk or non-risk for ATG16L1 T300A variant PBM treated with S. typhimurium or zymosan (filled diamonds and solid line) and at baseline (filled diamonds). Representative data from at least six independent experiments (from six human subjects in each group) with similar results are shown. (B) Representative TEM image of non-risk (n = 12 PBM) or risk ATG16L1 T300A variant (n = 12 PBM examined) PBM. At least five subjects per group were used. White arrowheads point to mitochondria, and mitochondria per BMM are quantitated. White scale bars in TEM images, 2 μm. **p < 0.01.

FIGURE 10.

CD risk ATG16L1 T300A increased ROS production and mitochondria number. (A) Relative ROS production, as measured by luminol-dependent chemiluminescence, was determined for risk or non-risk for ATG16L1 T300A variant PBM treated with S. typhimurium or zymosan (filled diamonds and solid line) and at baseline (filled diamonds). Representative data from at least six independent experiments (from six human subjects in each group) with similar results are shown. (B) Representative TEM image of non-risk (n = 12 PBM) or risk ATG16L1 T300A variant (n = 12 PBM examined) PBM. At least five subjects per group were used. White arrowheads point to mitochondria, and mitochondria per BMM are quantitated. White scale bars in TEM images, 2 μm. **p < 0.01.

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FIGURE 11.

CD risk ATG16L1 T300A alters mitochondria phenotype. Representative flow cytometry plots of risk or non-risk ATG16L1 T300A PBM that are either unstained (filled) or stained with MitoSOX (open) are shown for non-IBD control (A, left panels) and CD patients (B, left panels). Representative flow cytometry plots of non-risk and risk ATG16L1 T300A variant PBM with and without S. typhimurium infection and stained with MitoTracker Deep Red and MitoTracker Green are shown and quantitated for non-IBD control (A, middle and right panels) and CD patients (B, middle and right panels). Each filled circle represents an independent experiment from a human subject. *p < 0.05, **p < 0.01.

FIGURE 11.

CD risk ATG16L1 T300A alters mitochondria phenotype. Representative flow cytometry plots of risk or non-risk ATG16L1 T300A PBM that are either unstained (filled) or stained with MitoSOX (open) are shown for non-IBD control (A, left panels) and CD patients (B, left panels). Representative flow cytometry plots of non-risk and risk ATG16L1 T300A variant PBM with and without S. typhimurium infection and stained with MitoTracker Deep Red and MitoTracker Green are shown and quantitated for non-IBD control (A, middle and right panels) and CD patients (B, middle and right panels). Each filled circle represents an independent experiment from a human subject. *p < 0.05, **p < 0.01.

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Previous studies showed that ATG16L1 in epithelial cells is important for intestinal homeostasis (13, 14). In the present study, we were interested in the role of ATG16lL1 in myeloid cells and used LysM-Cre, which can mediate deletion of loxP-flanked genes to nearly 100% efficiency in macrophages and granulocytes (31). We showed that Atg16l1∆Mye mice exacerbate intestinal inflammation in three different murine colitis models, indicating that functional Atg16l1 in macrophages and granulocytes is important for intestinal homeostasis. We next focused on characterizing the cellular and molecular function of Atg16l1 in mouse primary macrophages and correlated the murine Atg16l1-deficient defects to human primary macrophages carrying the risk CD ATG16L1 T300A SNP. We observed that impaired ATG16L1 function led to increased numbers of proinflammatory intestinal macrophages in vivo (Fig. 4A, 4B), which was likely in part due to enhanced differentiation of proinflammatory macrophages. A recent study showed that another gene in the autophagy pathway, ATG5, also promotes proinflammatory macrophage polarization in BMM and Kupffer cells, resulting in increased hepatic inflammatory response in high-fat diet/LPS–treated ATG5−/− mice (35). The important role played by macrophage subsets in intestinal homeostasis is evidenced by a growing body of literature showing that either an increase in proinflammatory macrophages or a reduction in regulatory macrophages could worsen mucosal inflammation (20, 26, 36, 37; reviewed in Ref. 38).

Our result showing impaired clearance of Salmonella in the ATG16L1-deficient macrophages is consistent with other reports on the antibacterial function attributed to autophagy (10, 11, 13, 39, 40). In contrast, there are also reports showing that ATG16L1 deficiency could lead to increased clearance of microbes, including uropathogenic Escherichia coli and Citrobacter rodentium (41, 42). The differences in antibacterial function of ATG16L1 may be dependent on several factors such as the specific experimental bacteria being tested, affected by the composition of commensal bacterial in the host gut, or cell type–specific function of autophagy. The cell type–specific function of ATG16L1 was illustrated by the fact that the resistance to C. rodentium infection in Atg16L1 hypomorph (Atg16l1HM) mice was due to nonhematopoietic cells because cell type–specific deletion of Atg16l1 in myeloid cells did not exhibit resistance to C. rodentium, and in a bone marrow chimera experiment Atg16l1HM mice exhibited resistance to C. rodentium infection regardless of the bone marrow source (WT or Atg16l1HM) (41).

Our study revealed additional innate and adaptive immune mechanisms by which ATG16L1 affected the removal of infectious organisms in the gut. We identified an intracellular trafficking defect associated with ATG16L1 deficiency leading to reduced fusion of S. typhimurium–containing vesicles with Lamp1+ lysosomes in mouse and human macrophages. This diminished fusion led to reduced autophagosome maturation in ATG16L1 macrophages and contributed to the impaired bacterial clearance that we and others have observed with ATG16L1 deficiency (10, 11, 13). The impaired macromolecule trafficking with ATG16L1 deficiency could lead to defective presentation of Ag onto MHC-II to activate T cells (Fig. 7). Insufficient activation of the adaptive immune response with ATG16L1 deficiency could be one of the mechanisms that contributed to ineffective elimination of pathogenic bacteria and the persistent activation of intestinal inflammation in the chronic DSS-induced colitis model that were seen in our study. The impaired bacteria clearance due to defective innate and adaptive immunity with ATG16L1 deficiency could lead to the increased number of colitogenic bacteria that contributed to both murine colitis and human IBD. Recently, IgA-coated bacteria were implicated as IBD-promoting microbes (15). Consistently, we observed that increased amounts of IgA-coated bacteria were found in mice with Atg16l1 deficiency and in humans with the ATG16L1 T300A CD-associated variant, which was further increased in the context of intestinal inflammation. These findings suggest an intriguing rationale for studying whether carriage of the ATG16L1 T300A SNP results in a unique microbial composition that could be a potential target for precision therapeutics for IBD patients carrying the ATG16L1 T300A SNP.

In this study, we added novel mechanisms to previous findings that ATG16L1 deficiency increased cellular production of ROS (which may contribute to intestinal inflammation) (7, 10). We showed that the increased ROS with ATG16L1 deficiency was mediated by its increased production through NADPH oxidase and mitochondria. Our study showed that with ATG16L1 deficiency, there was increased phosphorylated p40phox protein due to reduced maturation of p40phox+ phagosomes, which contributed to increased ROS production via increased p40phox+ phagosomes that had been shown to increase ROS production (22). Additionally, our study showed that the mitochondria in both mouse and human ATG16L1-deficient macrophages produced more ROS at baseline and with S. typhimurium stimulation, owing to both increased numbers of mitochondria and increased accumulation of abnormal mitochondria that were more prone to produce ROS.

Our data from both non-IBD and CD patients who did not carry any of the NOD2 CD-associated variants and were homozygous for either the risk or non-risk ATG16L1 CD-associated T300A variant further validated and added greater relevance to our murine findings. Additionally, potential gene-dependent (ATG16L1) versus disease (CD)-dependent effects were controlled by studying non-IBD and CD patients with and without the ATG16L1 variant. We observed that polarization of monocytes to M1 proinflammatory macrophages was likely an ATG16L1 T300A-dependent effect because it was found in both non-IBD and CD patients to a similar magnitude and unlikely due to its known interaction with NOD2 (Fig. 8C, 8D). In contrast, although ATG16L1 itself increased levels of IgA-coated bacteria in the stool, having CD may likely play a bigger role because the magnitude of IgA-coated bacteria was greater (Supplemental Fig. 1C).

In conclusion, this study showed that colitis development in mice and humans could not be attributed solely to ATG16L1-dependent epithelial/Paneth cell defects because lack of ATG16L1 in macrophages and other myeloid cells similarly exacerbates murine colitis. We showed that ATG16L1 functions in macrophages to regulate inflammatory versus regulatory macrophage subsets, bacteria clearance, Ag processing and trafficking to the lysosomal compartment, mitophagy, and appropriate ROS production. Particularly, bacteria clearance and appropriate ROS production may be a general function of ATG16L1 in the myeloid compartment, as similar alterations were also seen in both Atg16l1-deficient macrophages and neutrophils.

We thank Jeremy Chen for assistance with Western blots, Guo Ying for setting up the mitochondria assay, Wei Zhu for TEM studies, and Loren Karp for critical reading of the manuscript. Human specimens were provided by the MIRIAD Biobank.

This work was supported by U.S. Public Health Service Grant DK056328 and National Institutes of Health Grant P01 DK046763 (to S.R.T.), National Institutes of Health K08 Career Development Award DK093578 (to D.Q.S.), Crohn’s and Colitis Foundation of America Career Development Award 3467 (to D.Q.S.), National Center for Advancing Translational Sciences Grant UL1TR000124 (to D.Q.S.), and by funding from the F. Widjaja Foundation Inflammatory Bowel and Immunobiology Research Institute. The MIRIAD Biobank is currently supported by the F. Widjaja Foundation Inflammatory Bowel and Immunobiology Research Institute, National Institutes of Health Grant P01 DK046763, European Union Grant 305479, National Institute of Diabetes and Digestive and Kidney Diseases Grants DK062413 and U54 DK102557, and by the Leona M. and Harry B. Helmsley Charitable Trust.

The online version of this article contains supplemental material.

Abbreviations used in this article:

BMM

bone marrow–derived macrophage

CD

Crohn disease

DC

dendritic cell

DSS

dextran sodium sulfate

IBD

inflammatory bowel disease

LPMC

lamina propria mononuclear cell

MFI

mean fluorescence intensity

MHC-II

MHC class II

MLN

mesenteric lymph node

MOI

multiplicity of infection

PBM

peripheral blood monocyte-derived macrophage

ROS

reactive oxygen species

SNP

single nucleotide polymorphism

TEM

transmission electron microscopy

WT

wild-type.

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The authors have no financial conflicts of interest.

Supplementary data