Regulating colonic dendritic cells by commensal glycosylated large surface layer protein a to sustain gut homeostasis against pathogenic inflammation

ABSTRACT Microbial interaction with the host through sensing receptors, including SIGNR1, sustains intestinal homeostasis against pathogenic inflammation. The newly discovered commensal

_Propionibacterium_ strain, P. UF1, regulates the intestinal immunity against pathogen challenge. However, the molecular events driving intestinal phagocytic cell response, including colonic

dendritic cells (DCs), by this bacterium are still elusive. Here, we demonstrate that the glycosylation of bacterial large surface layer protein A (LspA) by protein _O_-mannosyltransferase

1 (Pmt1) regulates the interaction with SIGNR1, resulting in the control of DC transcriptomic and metabolomic machineries. Programmed DCs promote protective T cell response to intestinal

_Listeria_ infection and resist chemically induced colitis in mice. Thus, our findings may highlight a novel molecular mechanism by which commensal surface glycosylation interacting with

SIGNR1 directs the intestinal homeostasis to potentially protect the host against proinflammatory signals inducing colonic tissue damage. You have full access to this article via your

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SUGAR UTILIZERS REDUCES _CLOSTRIDIODES DIFFICILE_ COLONIZATION Article Open access 09 October 2020 INTRODUCTION Commensal bacteria, via their surface layer (S-layer) gene products, and the

gastrointestinal phagocytic cells expressing sensing receptors (e.g., SIGNR1) synergistically interact to fine-tune the T cell signaling that is critical for protecting the host against

pathogenic inflammation exerted by intestinal infections.1,2 In this process, bacterial S-layer macromolecules along with induced metabolites transduce critical signals via cognate receptors

into these cells that profoundly control the host homeostasis to protect against tissue damage.3,4 Although the bacterial S-layer proteins display a similar architecture composed of a

peptidoglycan layer decorated with proteins and polysaccharides, various modifications, particularly glycosylation, exhibit strain-specific properties that differentially modify the host

immune physiology.5 Disruption of mutualistic interactions of the commensal’s S-layer with the host triggers deleterious signals that may manifest in pathogenic inflammation potentially

impairing the intestinal barrier function.6 Thus, understanding how host intestinal immunity is regulated through the recognition of these well-structured bacterial gene products by their

cognate receptors7 to coordinate protective immune responses is currently of particular therapeutic significance and requires further mechanistic investigations.8 _Propionibacterium_ strain,

P. UF1, is a newly discovered commensal bacterium isolated from the gut microbiota of premature infants fed human breast milk.9 This bacterium increases the frequency of colonic Th17 and

Treg cells9 involved in mucosal barrier repair and regulation of the intestinal inflammation.10 Induced bacteria-specific Th17 cell differentiation requires the bacterial dihydrolipoamide

acetyltransferase (DlaT), an enzymatic component of the pyruvate dehydrogenase complex.9 Chromosomal deletion of _dlaT_ gene impairs the regulation of protective Th17 cell response to

intestinal and systemic _Listeria monocytogenes (L. m)_ infection.9,11 Furthermore, P. UF1 regulates the neonatal T cells against necrotizing enterocolitis (NEC)-like injury in mice9 and

enhances the neonatal protective T cells against intestinal pathogen infection over time.12 However, the bacterial effector mechanisms potentially instructing the function of colonic DCs to

possibly control protective T cell immunity remain largely unknown. Here, we demonstrate that the glycosylation of bacterial LspA interacting with SIGNR1 is a pivotal factor, which

transcriptionally and metabolically programs colonic DCs, leading to protective T cell activation in steady state and during intestinal infection. Further, glycosylated LspA-SIGNR1

interaction critically protects mice against colitis-induced intestinal barrier injury. Errors in the bacterial glycosylation significantly disrupt the intestinal homeostasis, manifesting in

an inflammatory condition resulting in pathogen persistence and colonic tissue damage. Thus, this finding highlights the critical relevance of the glycosylated LspA in programming DC

immunophysiology to mitigate pathogenic inflammation and the induced colitogenic potential in mice. RESULTS GLYCOSYLATION OF LSPA BY PMT1 Knowing the significance of bacterial S-layer

complexes in communicating with host cells,13 we sought to investigate the functional relevance of P. UF1 S-layer proteins potentially involved in the regulation of colonic DC function. One

of the S-layer proteins of P. UF1 is LspA, which contains six N-terminal LGFP repeats [L-G-X-P-X(7-8)-D/N-G] involved in cell membrane anchoring and a C-terminal N-acetylglucosaminidase-like

domain, potentially implicated in bacterial cell wall metabolism (Supplementary Fig. 1a). Phylogenetic analysis demonstrated that LspA was highly conserved in P. UF1 and closely related

_Propionibacterium_ strains. Moreover, LspA homologs were also found in evolutionarily distantly related bacterial species, including _Bifidobacterium_ and _Geodermatophilus_ (Supplementary

Fig. 1b). Thus, to elucidate the functional significance of LspA within P. UF1 molecular machinery, the _lspA_ gene was deleted from the bacterial chromosome, resulting in Δ_lspA_ P. UF1

(Fig. 1a, b). Δ_lspA_ P. UF1 demonstrated enhanced bacterial clusters and autoagglutination (Fig. 1c), suggesting the critical involvement of this protein in bacterial S-layer structures.

Further, deletion of LspA significantly affected the bacterial transcriptomic and metabolomic signaling, including differential metabolic pathways involved in peptidoglycan biosynthesis,

amino and nucleotide sugar metabolism, fructose and mannose metabolism (Supplementary Fig. 2a). The analyzed metabolites involved in protein glycosylation (e.g., GDP-mannose and mannose

1-phosphate), along with those important for cell wall metabolism (e.g., GlcNAc-6-phosphate and UDP-GlcNAc), were significantly deregulated within Δ_lspA_ P. UF1 compared to P. UF1

(Supplementary Fig. 2b). RNA-Seq analysis further documented differentially expressed genes implicated in bacterial mannosylation and nucleotide sugar metabolism, including

phosphatidylinositol mannosyltransferase _pimA_14 and GDP-mannose-dependent alpha-mannosyltransferase _mgtA_15 (Supplementary Fig. 2c). Thus, these data emphasize the importance of LspA in

the regulation of glycan metabolism that may fundamentally impact the bacterial S-layer glycosylation. The bacterial S-layer proteins are generally glycosylated for their noncovalent

anchoring to the cell surface and interactions with environmental factors and host immune cells.5 Data demonstrated that the S-layer of P. UF1 reacted with concanavalin A (ConA), a

mannose/glucose-binding lectin, while LspA deficiency resulted in the loss of ConA binding (Fig. 1d), suggesting that LspA may be glycosylated. Therefore, we investigated the

glycosyltransferases responsible for adding glycol moieties to the bacterial S-layer using genome-wide bioinformatic analysis. Pmt1, a potential member of protein _O_-mannosyltransferase

family responsible for mannose transfer to serine and threonine residues of proteins in yeast16, was identified in P. UF1 genome. Further analysis demonstrated that Pmt1 homologs fell into

separate and loosely related groups of bacteria, including Actinobacteria (Fig. 1e). The _pmt1_ gene was then deleted in P. UF1 to assess the status of LspA glycosylation (Fig. 1f). Although

Δ_pmt1_ P. UF1 showed similar S-layer protein patterns when compared to P. UF1, no binding to ConA was observed for S-layer proteins isolated from Δ_pmt1_ P. UF1 (Fig. 1g). To underscore

the role of Pmt1 in the glycosylation of LspA, this protein was overproduced by Δ_lspA_ P. UF1 and Δ_pmt1_ P. UF1 strains to isolate and purify the glycosylated LspA (G-LspA) and

non-glycosylated LspA (NG-LspA), respectively (Fig. 1h). While both G-LspA and NG-LspA were recognized by anti-LspA serum antibodies, only purified G-LspA bound to ConA and illuminated

staining for glycoprotein (Fig. 1i). Thus, Pmt1 is critically required for the glycosylation of LspA. _O_-MANNOSYLATED LSPA INTERACTION WITH SIGNR1 To elaborate on the nature of LspA

glycosylation, the purified G-LspA and NG-LspA proteins were treated with PNGase F to release any N-glycans, permethylated and analyzed by MALDI-MS. Here, no _N_-linked glycans were detected

in either of the LspA proteins (Supplementary Fig. 1c). The _O_-linked glycans were released by β-elimination procedure and permethylated prior to MALDI-MS analysis. Signals corresponding

to Hex1-Hex6 were observed in the G-LspA protein (Fig. 2a), but not in the NG-LspA (Supplementary Fig. 1d). Furthermore, glycan compositional analysis demonstrated that mannose (Man) was the

major monosaccharide of G-LspA, with a retention time of 10.9 min (Fig. 2b). In contrast, NG-LspA showed no traces of Man (Supplementary Fig. 1e). Note that a minor peak of glucose was also

detected in both samples. However, glucose, as a very common contaminant, could be derived from reagents and detected as a free and minor glucose peak in HPAEC analysis. Moreover, glycomic

analysis of released oligosaccharides demonstrated that Man3 was the major glycan in the G-LspA, comprising 77% of the total glycans. While Man2 and Man4 oligosaccharides were minor glycans,

only traces of Man1, Man5 and Man6 were detected in the G-LspA (Fig. 2c). The purified G-LspA protein was then digested with trypsin and elastase, resulting in peptides with >75%

coverage (Supplementary Fig. 1a). LC-MS/MS analysis of the enriched peptides revealed seven _O_-glycopeptides at the N-terminus of LspA (Fig. 2d). In addition to the 41 threonine/serine

residues involved in Man attachment, adjacent proline and alanine residues that may facilitate local conformational changes for protein _O_-glycosylation17 were also found in all the

glycopeptides (Fig. 2d). Further, GC-MS analysis was performed to investigate the glycosyl linkages and positions of released _O_-glycans. Data demonstrated that Man oligosaccharides of LspA

were short linear chains interconnected via (1 → 6)-linkage and (1 → 2)-linkage (Fig. 2e). Man(1 → 6)Man(1 → 2)Man was the major trisaccharide component in the G-LspA, while disaccharide

Man(1 → 6)Man and tetrasaccharide Man(1 → 6)Man(1 → 2)Man(1 → 2)Man comprised a small percentage (Fig. 2e). These data indicate that LspA is a mannosylated S-layer glycoprotein with linear

short-chain _O_-glycans. SIGNR1 expressed by myeloid DCs recognizes characteristic molecular patterns with complex mannose and fucose structures in bacteria and fungi.18 Recently, we

observed the binding of P. UF1 to SIGNR1, but not SIGNR3.9 To precisely delineate the role of G-LspA binding to SIGNR1 in regulating DCs to subsequently initiate T cell commitment, the

interaction of G-LspA with SIGNR1 was biochemically investigated. Here, G-LspA and NG-LspA proteins were first separated by SDS-PAGE, transferred to PVDF membrane, and then incubated with

SIGNR1-hFc fusion protein. This protein binding complex was analyzed by subsequent incubation with anti-human Fc secondary antibody. Data demonstrated that the purified G-LspA bound

specifically to SIGNR1-hFc, and this binding was abolished in the presence of EDTA (Fig. 2f). Further, SIGNR1 interaction with G-LspA was assessed by ELISA showing G-LspA binding to

SIGNR1-hFc (Fig. 2g), but not to Dectin-1-hFc used as a control fusion protein (Fig. 2h). This binding was completely blocked by pre-incubation of SIGNR1-hFc with anti-SIGNR1 antibody, or

with the competitive ligand zymosan that is composed of β-glucan, α-mannan and mannosyl proteins (Fig. 2g). In contrast, no binding was observed for NG-LspA using similar assays (Fig. 2g).

Furthermore, the G-LspA exhibited a dose-dependent binding with SIGNR1-hFc using protein concentrations ranging from 0.08 μg/ml to 10 μg/ml with a _K__d_ value of 2.617 μg/ml G-LspA (Fig. 

2i). In contrast, NG-LspA did not react with SIGNR1-hFc, even with higher protein concentrations up to 20 μg/ml (Fig. 2i), highlighting the specificity of SIGNR1 binding to glycosylated

LspA. ACTIVATION OF COLONIC DCS BY GLYCOSYLATED LSPA IN STEADY STATE SIGNR1 was majorly expressed by colonic CD11chi MHCIIhi DCs (Supplementary 3a, b). To shed light on the relevance of LspA

glycosylation interacting with colonic DCs, CD11chi MHCIIhi CD11b+ F4/80_−_ DCs were FACS sorted (Supplementary Fig. 4a, b) from mice gavaged with P. UF1 or Δ_lspA_ P. UF1 to analyze their

transcriptome by RNA-Seq. Data demonstrated the modulation of costimulatory molecules (_Cd40_, _Cd80_, _Cd86_, and _Tnfsf4_) in colonic DCs by P. UF1 (Fig. 3a). NF-κB signaling (_Casp4_,

_Traf1_, _Tnfrsf1b_, _Mapk6_, _Nfkbie_, and _Nfkbiz_), cytokine/chemokine transcripts (e.g., _Il1b, Il12b, Cxcl1_, and _Cxcl2_), and antigen presentation-related genes (e.g., _Serpinb9_,

_Rab8b_) were also significantly augmented in DCs derived from mice gavaged with P. UF1 compared to Δ_lspA_ P. UF1. In contrast, DCs derived from Δ_lspA_ P. UF1-gavaged mice showed

activation of _Sod3_, _Rhoh_ and _Klf2_, which may instruct functional suppression in these cells. Migrating DCs constitutively express genes with regulatory functions.19 Accordingly, DCs

derived from mice gavaged with P. UF1 had elevated expression of genes, such as _Cd274_, _Spred1_, _Etv3_, _Tnfnip2_, _Stat3_, _Stat4_, and _Stat5a_. These cells were also enriched with

genes implicated in DC development (_Edn1, Cish_), migration (_Nrp2_, _Ccr10_, _Eps8_), and differentiation (_Pdk1_, _Hilpda_), while DCs of Δ_lspA_ P. UF1-gavaged mice exhibited increased

quantities of genes suppressing cellular regulatory functions (e.g., _Cyr61_, _Sdc1_). In addition, transcription factor _Irf4_ controlling Th17 cell cytokine machinery,20 cell cycle

inhibitor _Cdkn1a_ involved in Treg cell formation,21 and T cell-attracting chemokines _Ccl17_ and _Ccl22_, were all significantly activated in DCs of P.UF1-gavaged mice. In contrast,

Δ_lspA_ P. UF1 enhanced the DC expression of _Cd55_ and _Gilz_ genes associated with suppression of T cell function.22,23 Gene set enrichment analysis (GSEA) demonstrated that gene sets for

metabolic activities [e.g., glycolysis and oxidative phosphorylation (OXPHOS)] and DC activation, including activation of innate responses, regulation of I-κB/NF-κB signaling, cytokine

receptor activity and cell migration, were enriched in DCs derived from mice gavaged with P. UF1 (Fig. 3b). In contrast, glycosylated LspA deficiency abrogated cellular protein responses in

DCs, resulting in enhanced endoplasmic reticulum (ER) stress-associated activities (Fig. 3b), which were associated with increased transcripts of unfolded protein response (UPR)-related

genes _Xbp1_, Derl3, and _Edem2_ in these cells (Fig. 3a). Further, P. UF1-induced DC activation (e.g., _Cd80_, _Cd86_, _Il1b_, and _Il12b_) was abrogated due to _Signr1_ deficiency (Fig. 

3c). Moreover, loss of LspA glycosylation (Δ_lspA_ P. UF1-gavaged _Signr1__+/+_ mice) mirrored the deficiency of SIGNR1 (P. UF1-gavaged _Signr1__−_/_−_ mice), suggesting a role of

glycosylated LspA-SIGNR1 interaction in DC activation. Having shown that LspA deficiency impacted the transcriptome of DCs, we asked whether LspA deficiency would also influence the

regulation of T cell polarization in steady state. Thus, C57BL/6 mice were gavaged with P. UF1 or Δ_lspA_ P. UF1 to analyze colonic DC and T cell responses. Data demonstrated that Δ_lspA_ P.

UF1 compared to P. UF1 significantly decreased DC and T cell responses, including IL-10+ Th17 cells and 10+ Tregs (Supplementary Fig. 5a, b). Notably, no difference in intestinal bacterial

colonization was observed in mice gavaged with P. UF1 or Δ_lspA_ P. UF1, as both strains transiently colonized conventional mice (Supplementary Fig. 5c), suggesting that glycosylated LspA is

not involved in bacterial colonization, but instructs DC activation to polarize T cells. PROGRAMMING TRANSCRIPTOMIC AND METABOLOMIC DC-MACHINERIES DURING INTESTINAL INFECTION Intestinal DCs

rapidly respond to invading pathogens.24 However, these cells may be functionally conditioned by pathological surrounding signals induced by intestinal infections.25 To elaborate on the

bacterial glycosylated LspA influencing DC transcriptome, colonic MHCIIhi CD11chi CD11b+ F4/80_−_ DCs were FACS sorted from mice gavaged with P. UF1 or Δ_lspA_ P. UF1 and then orally

infected with Δ_actA L. m_. RNA-Seq analysis demonstrated that P. UF1 controlled the expression of genes associated with DC activation (e.g., _Cstl_, _Tsc1_) and suppression of TGF-β

signaling (e.g., _Tgbfi_, _Eaf2_, and _Cited2_) (Fig. 4a). Further, transcripts of _Tnip3_ regulating NF-κB activation and _Dok-2_ suppressing Ras-Erk signaling were stimulated in colonic

DCs isolated from mice gavaged with P. UF1. Importantly, _Cd209b_ (_Signr1_), whose activation highly depends on the mannosylation of glycans, was upregulated in DCs derived from mice

gavaged with P. UF1. In contrast, Toll-like receptors (_Tlr3_, _Tlr4_, and _Tlr12_) associated with proinflammatory response were upregulated in DCs of mice gavaged with Δ_lspA_ P. UF1 (Fig.

 4a, b). _Mx1_ and _Irf1_, selectively activated during TLR-induced DC stimulation,26 were also enriched in DCs derived from Δ_lspA_ P. UF1-gavaged mice. Consistently, a proinflammatory

status was readily observed in DCs of mice gavaged with Δ_lspA_ P. UF1. Accordingly, a set of proinflammatory genes (e.g., _Ifng_, _Tnfsf15_, and _Ifi204_), several genes associated with

cell apoptosis (e.g., _Daxx_, _Nab2_, and _Fbxw7_), and stress-associated activities were activated in DCs derived from mice gavaged with Δ_lspA_ P. UF1 (Fig. 4a, b). In addition, GSEA

demonstrated enhanced mitochondrial biogenesis in DCs isolated from P. UF1-gavaged mice, as indicated by enriched gene sets for OXPHOS, TCA cycle and respiratory chain, and NADH

dehydrogenase complex (Fig. 4c). Moreover, pathways regulating apoptosis and protein localization to ER were also enhanced in these cells. In contrast, DCs from Δ_lspA_ P. UF1-gavaged mice

exhibited dysregulated protein folding and ER stress-associated activities (Fig. 4c), consistent with increased transcripts of the UPR-related genes _Xbp1_ and _Derl3_ in these cells (Fig. 

4a, b). Inflammatory DC response is associated with declined mitochondrial activity promoted by TLR signaling.27 Having demonstrated that glycosylated LspA controlled TLR-activation and

enriched gene sets for OXPHOS (Fig. 4), we further assessed whether P. UF1 expressing glycosylated LspA would  maintain mitochondrial respiration during intestinal infection. Thus, colonic

DCs were enriched by magnetic beads from mice gavaged with P. UF1 or Δ_lspA_ P. UF1 and orally infected with Δ_actA L.m_ to analyze the real-time changes in the rate of extracellular

acidification (ECAR), a measurement of glycolysis, and the mitochondrial rate of oxygen consumption (OCR). While no difference in the ECAR was observed, OCR was significantly decreased in

enriched DCs derived from mice gavaged with Δ_lspA_ P. UF1 compared to DCs derived from P. UF1-gavaged mice (Fig. 5a). These data support the notion that glycosylated LspA may regulate DC

metabolic function during intestinal infection. It was recently demonstrated that DC function can be impacted by cellular metabolic factors that facilitate the biosynthetic and bioenergetic

needs of these cells.28 To elaborate on this notion, colonic MHCIIhi CD11chi CD11b+ F4/80_−_ DCs were FACS sorted from mice gavaged with P. UF1 or Δ_lspA_ P. UF1 and orally infected with

Δ_actA L. m_ to analyze the metabolomic activity of these cells. Here, a distinct metabolome was identified in DCs derived from P. UF1-gavaged mice compared to those derived from Δ_lspA_ P.

UF1-gavaged mice (Fig. 5b). Metabolic pathways, including arginine and proline metabolism, purine metabolism and _de novo_ fatty acid biosynthesis, were differentially activated in DCs

isolated from the aforementioned groups of mice (Fig. 5c). Putatively annotated eicosatrienoic acid, known as an anti-inflammatory metabolite, 4-aminobutanal indicative of anti-inflammatory

putrescine metabolism, and proline involved in suppressing reactive oxygen species (ROS),29 were markedly accumulated in DCs from P. UF1-gavaged mice compared to the other group (Fig. 5d).

In contrast, putative metabolites associated with proinflammation and energy starvation  [e.g., methylimidazoleacetic acid and 5-aminoimidazole-4-carboxamide ribonucleotide (AICAR)] and cell

apoptosis-inducing deoxyadenosine were significantly enriched in DCs derived from Δ_lspA_ P. UF1-gavaged mice (Fig. 5d). Thus, these data specifically demonstrate the requirement of

glycosylated LspA expressed by P. UF1 that programs the regulation of DC function during _Listeria_-induced inflammation. REGULATING PROTECTIVE T CELL RESPONSE BY P. UF1 EXPRESSING

GLYCOSYLATED LSPA To further elucidate the functional relevance of glycosylated LspA in modulating colonic homeostasis during intestinal infection, C57BL/6 mice were gavaged with P. UF1,

Δ_lspA_ P. UF1 or PBS, and then orally infected with Δ_actA L. m_. Notably, P. UF1, compared to Δ_lspA_ P. UF1, regulated proinflammatory responses by controlling the frequencies and cell

counts of IL-1β+, IL-6+, and IL-12/23p40+ DCs (Fig. 6a). However, such induced DC regulation was abolished in _Signr1__−/−_ mice gavaged with P. UF1, Δ_lspA_ P. UF1 or PBS and subsequently

infected with Δ_actA L. m_ (Fig. 6a), denoting a potential role of glycosylated LspA-SIGNR1 interaction in controlling inflammatory DC response. Further, tuned DC response was associated

with T cell regulation, leading to an increased percentage and number of Th17 and IL-10+ Treg cells in mice gavaged with P. UF1 compared to Δ_lspA_ P. UF1 or PBS (Fig. 6b). Loss of

protective immunity in _Signr1__+/+_ mice gavaged with Δ_lspA_ P. UF1 resulted in delayed Δ_actA L. m_ clearance compared to mice gavaged with P. UF1 (Fig. 6c). In addition, transcripts of

proinflammatory molecules such as _Il6_, _Il12b_, _Ifng_, _Ccl5_, and _Cxcl2_ were significantly decreased in colonic tissues of _Signr1__+/+_ mice gavaged with P. UF1 compared to those

gavaged with Δ_lspA_ P. UF1 or PBS (Fig. 6d). In contrast, the protective T cell immunity was blunted in _Signr1__−/−_ mice gavaged with P. UF1, Δ_lspA_ P. UF1 or PBS and then infected with

Δ_actA L. m_ (Fig. 6b), and no difference in pathogen clearance was observed in these groups of mice (Fig. 6c). Furthermore, data demonstrated that deficiency in bacterial glycosylated LspA

did not induce any changes in the gut microbial composition of mice gavaged with Δ_lspA_ P. UF1 compared to P. UF1 during Δ_actA L. m_ infection (Supplementary Fig. 6a, b). Together, these

results indicate the crucial role of glycosylated LspA-SIGNR1 interaction in regulating colonic DCs to orchestrate protective T cell response to intestinal _Listeria_ infection. AMELIORATING

DEXTRAN SULFATE SODIUM (DSS)-INDUCED COLITIS BY P. UF1 EXPRESSING GLYCOSYLATED LSPA IL-17A and IL-10+ Tregs contribute to protection against acute intestinal colitis.30,31,32 To further

investigate the protective role of glycosylated LspA expressed by P. UF1 in chemically induced colitis, groups of mice were gavaged with P. UF1, Δ_lspA_ P. UF1 or PBS, and the experimental

colonic tissue damage was induced by 3% DSS. Data demonstrated that P. UF1, compared to other groups of mice, significantly reduced the disease severity, as indicated by the reduction of

weight loss, diarrhea and fecal blood scores (Fig. 7a, b), and increased colon length (Fig. 7c). Furthermore, severe signs of inflammation-induced thickening of the bowel wall and loss of

regular blood vessel structures were observed in the colons of PBS- and Δ_lspA_ P. UF1-gavaged mice but not in P. UF1-gavaged mice (Fig. 7d). Histological analysis demonstrated severe tissue

destruction, characterized by extensive segmental to diffuse mucosal epithelium and crypt loss with associated mucosal and submucosal inflammation in the colonic tissues of mice gavaged

with PBS or Δ_lspA_ P. UF1, while DSS-induced colitis was substantially mitigated in mice gavaged with P. UF1 (Fig. 7e, f). Furthermore, obtained data also demonstrated that DSS-treated mice

gavaged with P. UF1 were markedly protected from leaky gut when compared with other groups, as documented by FITC-dextran assay (Fig. 7g). Consistently, enhanced transcripts of tight

junction proteins (_Cldn2_, _Cldn3_, _Cldn7_, _Cldn8_, and _ZO-3_) in colonic tissues of P. UF1-gavaged mice were observed compared to the other groups (Fig. 6h). Thus, glycosylated LspA

expressed by P. UF1 contributes to the maintenance of intestinal barrier integrity. To further elucidate the protective role of glycosylated LspA-SIGNR1 signaling in DSS-induced colitis,

_Signr1__−/−_ mice were gavaged with P. UF1, Δ_lspA_ P. UF1 or PBS and treated with DSS. Once again, lack of SIGNR1 interaction with glycosylated LspA resulted in no improvement of

colitogenic disease progression in any of the groups of mice (Fig. 7a–f). Here, the disruption of glycosylated LspA-SIGNR1 interaction abolished the regulation of intestinal permeability

(Fig. 7g) and blunted the expression of tight junction proteins in _Signr1__−/−_ mice (Fig. 7i). Collectively, these data suggest that glycosylated LspA interacting with SIGNR1 sustains gut

homeostasis to protect against detrimental signals inducing tissue damage. DISCUSSION The maintenance of gut homeostasis requires a rigidly balanced dialog between the resident microbes and

the host.1 This can be established through the interaction of a variety of bacterial gene products with their cognate receptors expressed by phagocytic cells.6 Disruption of such elegantly

induced communication may result in pathogenic inflammation and intestinal tissue damage.33 In this study, we demonstrate a protective mechanism at the intestinal interface whereupon

glycosylated LspA interacting with SIGNR1 dictates DC response that in turn regulates protective T cells against intestinal pathogen infection and chemically induced colitis. There has

recently been a growing interest in the bacterial S-layer proteins with multiple regulatory properties.34,35 Accordingly, the composition of the S-layer proteins of _Propionibacterium_

species is remarkably variable, and the functions of bacterial S-layer proteins are strain-dependent.36 For instance, surface layer protein B (SlpB), highly expressed by some of _P.

freudenreichii_ strains, facilitates bacterial adhesion to epithelial cells.37 While no SlpB homolog is found in P. UF1, LspA constitutes the major extractable S-layer protein in this

bacterium. The high protein expression may also suggest that LspA, particularly in P. UF1, is a valuable factor for not only supporting the S-layer structure of this bacterium but could also

be important for fine-tuning intestinal immunity and may serve as an elegant vehicle for mucosal vaccine and therapeutic approaches.38 Although LspA is conserved, the levels of protein

expression vary dramatically in different _Propionibacterium_ strains,35 indicating potentially differential gene regulation in these bacterial strains. With this notion in mind and the

unique expression pattern of LspA by P. UF1, we were prompted to further elaborate on its physiological characteristics and its potential implication in regulating intestinal immunity. Here,

we demonstrate that deletion of _lspA_ leads to fundamental changes in pathways associated with S-layer carbohydrate metabolisms, including mannosylation, suggesting the critical role of

LspA in maintaining the S-layer glycosylation profile. Indeed, ConA binding assays comparing S-layer proteins isolated from P. UF1 and Δ_lspA_ P. UF1 show that glycosylation of other

potential glycoproteins is also likely impaired by LspA deficiency. Further, changes in S-layer glycosylation may directly impact the bacterial cell-cell interactions, leading to enhanced

autoagglutination due to LspA deficiency in P. UF1, possibly as a result of altered cell surface hydrophobicity. This is consistent with the observations that surface glycosylation, such as

flagella glycosylation, is highly associated with autoagglutination, which is an important step for microcolony formation on the intestinal epithelial cells that contributes to intestinal

colonization.39,40 In this study, we also clearly demonstrate that LspA is an _O_-mannosylated glycoprotein, uniquely representing the first glycoprotein characterized in _Propionibacteria_.

Furthermore, the glycosylation of LspA critically requires Pmt1 activity. This feature is reminiscent of protein glycosylation in _Streptomyces_ and _Mycobacterium_, whereupon the

_O_-glycosylation of surface lipoproteins or phosphate-binding protein PstS, are highly dependent on the membrane-associated lipoprotein Pmt.41 Thus, the Pmt-mediated protein

_O_-mannosylation seems to be a general pathway in actinomycetes. In fungi and yeasts, up to seven Pmt family members have been identified thus far, and the homomeric and/or heteromeric

interactions among Pmt members cooperatively initiate the protein glycosylation, resulting in the biosynthesis of diversified glycan structures.42 However, Pmt1 seems to be the only protein

glycosyltransferase in P. UF1, and no other protein glycosylation is observed in the S-layer proteins isolated from Δ_pmt1_ P. UF1. Furthermore, the linear short-chain mannose is found to be

the major glycan structure of bacterial LspA; thus, concluding that Pmt1 is the crucial enzyme responsible for modifying S-layer proteins, particularly LspA, with simple mannoses in P. UF1.

Recognition of microbial gene products by sensing receptors, including C-type lectins, is essential for translating the nature of microbes into gene-transcriptional and metabolic programs

that may initiate the regulation of DC function to prime T cell polarization.43 SIGNR1, a murine homolog of human DC-SIGN, conditions intestinal DCs for the induction of oral tolerance44 and

plays a key role in host defense against pathogen infection.45 However, further rigid molecular studies are still required to mechanistically elucidate the gene and metabolic programs that

are modulated by the interactions of this sensing receptor with the bacterial surface glycosylation. Here, we demonstrate that LspA glycosylation is required to be recognized by its

receptor, SIGNR1, leading to the regulation of DC activation in steady state and during pathogen-induced inflammation. Accordingly, in steady state P. UF1, via its glycosylated LspA, seems

to optimally induce the activation of colonic DCs when compared with its counterpart, Δ_lspA_ P. UF1. Mainly, P. UF1 regulates the expression of costimulatory, cytokine and antigen

presentation molecules (e.g., _Cd40_, _Il1b, Serpinb9_, _Rab8b,_ and _Bcl3_) in colonic DCs while glycosylated LspA deficiency results in the activation of suppressive molecules (e.g.,

_Sod3_, _Rhoh_ and _Klf2, Cyr61_, _Sdc1_) that may dysfunction colonic DCs to properly induce T cell differentiation. More importantly, P. UF1 decorated with glycosylated LspA regulates DCs

that control T cell response to pathogen infections. Here, regulatory signaling appears to be activated by LspA glycosylation, as _Tgfbi_, _Eaf2_, and _Cited2_ genes associated with

suppression of TGF-β signaling are tuned down in DCs derived from mice gavaged with P. UF1 compared to Δ_lspA_ P. UF1. Further, proinflammatory genes (e.g., _Ifng_ and _Ifi204_) are also

downregulated in DCs derived from mice gavaged with P. UF1. Consistently, anti-inflammatory polyunsaturated fatty acid eicosatrienoic acid is highly enriched in DCs of P. UF1-gavaged mice,

while proinflammatory metabolites, such as methylimidazoleacetic acid and AICAR, are significantly elevated in DCs of Δ_lspA_ P. UF1-gavaged mice during _Listeria_ infection. In addition,

DCs derived from Δ_lspA_ P. UF1-gavaged mice exhibit significant quantities of genes associated with cell apoptosis and stress response, consistent with decreased cellular proline levels

that are important for protecting against various cellular stresses.29 This may emphasize the impaired DC metabolism induced by glycosylated LspA deficiency, which is further supported by

reduced mitochondrial respiration and enriched metabolite AICAR that serves as a metabolic activator responding to energy starvation of the cells. Collectively, these data suggest that LspA

glycosylation may be an important factor that modulates colonic DC response via transcriptomic and metabolic reprogramming. Proinflammatory DC regulation may impact T cell polarization.7

Indeed, dysregulated cellular pathways in DCs induced by Δ_lspA_ P. UF1 significantly reduce the protective T cell immunity and correspondingly manifest in uncontrolled intestinal pathogen

infection, resulting in delayed pathogen clearance and enhanced intestinal inflammation. In addition, no protection is seen in mice when either SIGNR1 or its ligand, glycosylated LspA, is

deficient, indicating the critical role of the glycosylated LspA-SIGNR1 axis in regulating colonic DC response that dictates protective T cell response against pathogen infection. We have

previously shown that P. UF1 induces DlaT-specific Th17 cells, which are indispensable for protection against _Listeria_ infection.9,11,12 While DlaT expression is not impacted by _lspA_

deletion (Supplementary Fig. 2c), induced Th17 cells notably require a regulated cytokine environment initiated by DCs through glycosylated LspA interacting with SIGNR1. Interestingly,

glycosylated LspA deficiency does not impact the phylum of gut microbiota in bacterially-gavaged groups of mice and then infected with _Listeria_, suggesting that glycosylated LspA may not

contribute to any changes in the intestinal bacterial community. Yet, do changes in bacterial metabolome due to glycosylated LspA deficiency contribute to reduced regulation of DCs during

intestinal infection? Although critical metabolic changes are observed within Δ_lspA_ P. UF1, we posit that these metabolites may be mostly restricted to intracellular bacterial metabolic

networks in response to altered S-layer glycosylation and may not directly impact the immune cells. Nonetheless, this important notion still requires further rigid investigations. Ample data

demonstrate that IL-17A serves as a protective factor against DSS-induced colitis by maintaining intestinal tight junctions and promoting epithelial repair.31,46 Here, Th17 cells, together

with IL-10+ Tregs, regulated by colonic SIGNR1+ DCs interacting with glycosylated LspA, may be required for ameliorating DSS-induced colitis, all of which potentially result in the

maintenance of tight junction expression that controls the intestinal permeability and the mitigation of chemically induced inflammation and intestinal tissue damage. In summary, our data

demonstrate the molecular mechanism through which DC functions can be fine-tuned via bacterial glycosylated LspA interacting with SIGNR1. Such a regulated DC response is pivotal in priming

protective T cell response to intestinal infection and plays a critical role in mitigating DSS-induced colitis in mice. Induced immune regulatory processes involving the fine-tuned

receptor-ligand interaction, mainly glycosylated LspA-SIGNR1, elicits functional gut homeostasis during the intestinal inflammatory condition. Thus, consistent with this finding,

glycosylation of a bacterial gene product such as LspA may be an important feature for a future formulated prebiotic, conjugated vaccines and therapeutic targets that not only induce the

regulation of innate and protective T cells against intestinal pathogen challenges, but may also prevent induced pathogenic inflammation that triggers tissue damage and the progression of

intestinal proinflammatory diseases (e.g., colitis) in affected patients.47,48 Thus, a deeper understanding of beneficial bacterial gene products influencing the regulation of gut

homeostasis may pave the way for the development of new pre or postbiotic therapeutic strategies to potentially treat inflammatory bowel disease (IBD) or colon cancer.49,50 Finally, our work

uniquely illustrates the significance of the mannosylated ligand, bacterial LspA, and its critical binding to SIGNR1 that sustains the optimal activation and regulation of colonic DCs in

intestinal steady state and during inflammatory condition. Conclusively, shedding light on the relevance of a sensing receptor, SIGNR1 and its human homolog, DC-SIGN, may reveal the critical

innate factor in resisting detrimental signals inducing tissue damage that manifest in IBD. Thus, activating critical signals involving SIGNR1 (DC-SIGN) in health and human diseases may

advance our vision to improve and develop therapeutic platforms mitigating IBD, which increasingly affects more than 3.5 million worldwide.51 MATERIALS AND METHODS MICE C57BL/6 mice (6–9

weeks old) were obtained from Jackson Laboratory and maintained under specific pathogen-free, _Helicobacter_-free conditions. _Signr1__−/−_ mice were provided by Huang Shau-Ku (Johns Hopkins

University School of Medicine, Baltimore, Maryland, USA). All animal studies were approved by the Animal Care and Use Committee of the University of Florida under the protocol number

201708484. Mice were maintained in accordance with the Animal Welfare Act and the Public Health Policy on Humane Care. BACTERIAL ADMINISTRATION In steady state, _Signr1__+/+_ and

_Signr1__−/−_ mice were gavaged with P. UF1 or Δ_lspA_ P. UF1 (109 CFU/mouse/100 μl) every 3 days over the course of 12 days, and mice were euthanized on day 14 to isolate colonic immune

cells. During Δ_actA L. m_ infection, _Signr1__+/+_ and _Signr1__−/−_ mice were gavaged with P. UF1, Δ_lspA_ P. UF1 (109 CFU/mouse/100 μl) or PBS on days −7, −4, −1, and 2. Mice were denied

food for 4 h on day 0 and then orally infected with 100 μl PBS containing 50 mg/ml CaCO3 and approximately 109 CFU of Δ_actA L. m_. Fecal samples were collected on days 1–4, and mice were

sacrificed on day 7. Pathogen loads were determined by plating serial dilutions of fecal samples on BHI agar supplemented with 200 μg/ml streptomycin. CELL ISOLATION AND FLOW CYTOMETRY

Density gradient centrifugation using Percoll was performed to isolate lamina propria cells from mouse colon. Briefly, tissues were collected and fecal contents in the colon were carefully

removed. Colons were opened longitudinally and cleaned with cold PBS and then shaken in PBS containing 20 mM Hepes and 10 mM EDTA for 30 min at 37 °C. Tissues were cut into small pieces and

incubated with digestion solution [RPMI 1640 containing 10% FBS (Thermo Fisher Scientific, Waltham, MA), 0.4% β-mercaptoethanol, 400 U/ml collagenase VIII (Sigma Aldrich, St. Louis, MO) and

100 μg/ml DNase I (Sigma Aldrich, St. Louis, MO)] for 1.5 h at 37 °C in a 5% CO2 incubator. Digested tissues were filtered through 100 μm cell strainer (Genesee Scientific, San Diego, CA),

and cells were resuspended in 5 ml of 40% Percoll (Sigma Aldrich, St. Louis, MO) and overlaid on 5 ml of 80% Percoll. Cells in the interphase were collected after gradient centrifugation

(1258 g, 25 min, 25 °C). Flow cytometry was performed as described previously9 with some modifications. Isolated cells were stimulated with 50 ng/ml PMA (Sigma Aldrich, St. Louis, MO) and

500 ng/ml ionomycin calcium salt (Sigma Aldrich, St. Louis, MO) for 4 h and 5 μg/ml brefeldin A (BioLegend, San Diego, CA) was added 2 h before cells were harvested. Stimulation was

performed in IMDM medium (Sigma Aldrich, St. Louis, MO) containing 10% FBS, 1% penicillin/streptomycin, 0.4% β-mercaptoethanol, 10 mM Hepes and 2 mM L-Glutamine. Dead cells were detected

using LIVE/DEAD® Fixable Blue Dead Cell Stain Kit (Thermo Fisher Scientific, Waltham, MA), followed by incubation with Mouse Fc Blocking Reagent (Miltenyi Biotec, Auburn, CA). Cells were

first stained for cell-surface markers and then resuspended in fixation/permeabilization solution [Cytofix/Cytoperm kit (BD Biosciences, San Diego, CA) for DC cytokine analysis, or Foxp3

Transcription Factor Staining Buffer Set (eBiosciences, San Diego, CA) for T cell analysis]. Cells were stained with following fluorescent antibodies: eVolve 655-CD45 (catalog 86-0451-42),

eFlour506-CD11c (catalog 69-0114-80), PE-MHCII (catalog 12-5321-82), PE/Cy5-F4/80 (catalog 15-4801-80), APC-SIGNR1 (catalog 17-2093-82), PE/Cy5-CD8 (catalog 15-0081-83), PE/Cy7-Pro-IL-1β

(catalog 25-7114-82)/rat IgG1 κ, FITC-IL-6 (catalog 11-7061-82)/rat IgG1 κ, eFlour 450-IL-12/23p40 (catalog 48-7123-82)/mouse IgG1 κ, and eFlour450-FoxP3 (catalog 48-5773-82)/rat IgG2a κ

from Thermo Fisher Scientific; APC/Cy7-CD11b (catalog 101226), PerCP/Cy5.5-CD64 (catalog 139308), APC/Cy7-CD3 (catalog 100330), Brilliant Violet 605-CD4 (catalog 100548), PE-IL-17A (catalog

506904)/rat IgG1 κ, and FITC-IL-10 (catalog 505006)/rat IgG2b κ from BioLegend. Data were collected by an LSR II Fortessa (BD Biosciences, San Jose, CA) and analyzed with FlowJo software

(version 10) (TreeStar, Ashland, OR). After dead and doublet cell exclusion, and the subsequent CD45+ selection, DCs were defined as CD11chi MHCIIhi CD11b+ F4/80_−_ (Supplementary Fig. 7a)

and T cells as CD3+ CD8_−_ CD4+ (Supplementary Fig. 7b). DC SORTING Colonic cell suspensions were prepared from mice gavaged with P. UF1 or Δ_lspA_ P. UF1 in steady state or during

intestinal Δ_actA L. m_ infection. Cells were labeled with a cocktail of fluorescent antibodies specific for: PE/Cy7-CD11c (BioLegend, catalog 117318), PE-MHCII (Thermo Fisher Scientific,

catalog 12-5321-82), APC-CD11b (BioLegend, catalog 101212), FITC-F4/80 (Thermo Fisher Scientific, catalog 11-4801-82). Dead cells were identified and excluded using LIVE/DEAD Fixable Violet

Dead Cell Stain (Thermo Fisher Scientific, Waltham, MA). CD11chi MHCIIhi CD11b+ F4/80_−_ DCs (Supplementary Fig. 4a) were isolated using a SONY SH800S Cell Sorter (Sony, Tokyo, Japan). The

purity of sorted cells analyzed by flow cytometry was determined to be >98% (Supplementary 4b). Subsequently, cell pellets were resuspended in RLT plus buffer (Qiagen, Germantown, MD) for

RNA extraction or snap-frozen in liquid nitrogen for metabolomic analysis. RNA-SEQ ANALYSIS Total RNA was extracted from about 1 × 104 CD11chi MHCIIhi CD11b+ F4/80_−_ DCs isolated from each

individual mouse using an RNeasy Plus Micro Kit (Qiagen, Germantown, MD). cDNA was generated using a SMART-Seq HT kit (Takara Bio Inc., Mountain View, CA) and RNA-Seq libraries were

constructed using a Nextera XT DNA Library Preparation Kit (Illumina, Inc., San Diego, CA). Barcoded samples were sequenced on an Illumina HiSeq instrument (Illumina, Inc., San Diego, CA) at

the University of Florida ICBR NextGen DNA Sequencing Core Facility. The sequencing reads were mapped to the _Mus musculus_ genome (NCBI GRCm38/mm10) using STAR aligner (v2.6.0), and count

table was generated using SubReads featureCounts (v1.6.0). Significantly altered genes (RPKM ≥ 1, FDR _P_ < 0.05, fold change ≥ 1.5) were identified by DESeq2.

Regularized-log-transformation of count data was performed for heatmap plotting. Gene set enrichment analysis (GSEA) was performed in the javaGSEA (v3.0) using GO, Hallmark, KEGG and

REACTOME database. EXTRACELLULAR FLUX ANALYSIS Colonic DCs were isolated from mice gavaged with P. UF or Δ_lspA_ P. UF1 and orally infected with Δ_actA L. m_ using a Pan-DC Enrichment Kit

(StemCell Technologies, Vancouver, Canada). For real-time analysis of extracellular acidification rate (ECAR) and oxygen consumption rate (OCR), DCs were analyzed using an XFe96

Extracellular Flux Analyzer (Seahorse Bioscience, North Billerica, MA) as described.52 Briefly, enriched colonic DCs (3 × 105 cells/well, pooled from 3 mice) were analyzed in non-buffered

RPMI medium supplemented with 2.5 μM dextrose, 2 nM glutamine, and 1 μM sodium pyruvate. ECAR and OCR were analyzed in response to 1 μM oligomycin, 1.25 μM fluoro-carbonyl cyanide

phenylhydrazone (FCCP), 1 μM rotenone and 1 μM antimycin A. HIGH-RESOLUTION METABOLOMICS ANALYSIS FACS sorted colonic CD11chi MHCIIhi CD11b+ F4/80_−_ DCs were isolated from mice gavaged with

P. UF1 or Δ_lspA_ P. UF1 and orally infected with Δ_actA L_. _m_. Note: colonic cells of 2 mice were combined to obtain 60,000–80,000 DCs/sample. After vortex and incubation with

acetonitrile-water (2:1) at 4 °C for 30 min, DC samples were centrifuged and the supernatants were analyzed by LC-MS. Each sample was run in triplicates on an Orbitrap Fusion Tribrid Mass

spectrometer with the resolution of 120,000 (Thermo Fisher, San Diego, CA), with dual chromatography using a 5 min C18 reversed-phase chromatography in negative electrospray ionization (ESI)

mode and a 5 min HILIC chromatography in positive ESI mode over a mass-to-charge ratio (_m_/_z_) range of 85–1250. Student’s _t_-test was performed between treatment groups. Subsequently,

metabolic pathway analysis was performed by _Mummichog_ software (v2.0) with default parameters. 786 significant metabolite features in negative mode and 1234 metabolite features in positive

mode were used as input to _Mummichog_. The pathways represented by at least two significant metabolites and enriched at _P_ < 0.05 in positive mode (HILIC column) are presented.

DSS-INDUCED COLITIS _Signr1__+/+_ and _Signr__1−/−_ mice were treated with 3% DSS in drinking water for 5 consecutive days (made fresh every 2–3 days) to induce colitis. Mice were monitored

for disease progression through day 12 (_Signr1__+/+_ mice) or day 10 (_Signr1__−/−_ mice as they are more susceptible to induced colitis) after DSS treatment. For bacterial administration,

mice were orally gavaged with P. UF1, Δ_lspA_ P. UF1 or PBS on days −7, −4, −1, 2, 5, and 8 for a total of 6 gavages. Colitis severity was determined by histopathology. Tissues were fixed,

sectioned, and stained with hematoxylin and eosin (Histology Tech Services, Gainesville, FL). Stained sections were evaluated by a boarded veterinary pathologist. Macroscopic damage in the

colons of DSS-treated mice gavaged with P. UF1, Δ_lspA_ P. UF1 or PBS was visualized with a Multi-Purpose Rigid Telescope attached to a TELE PACK X (Karl Storz-Endoscope, Germany), as

described previously.13 FITC-DEXTRAN GUT PERMEABILITY ASSAY DSS-treated mice were orally gavaged with FITC-dextran 4000 (Sigma-Aldrich, St. Louis, MO), a nonmetabolizable macromolecule that

is used as a permeability probe. All mice were orally gavaged with FITC-dextran (0.6 mg/g mouse weight) and sacrificed 4 h later for serum harvest. Fluorescent intensity in the serum was

measured using a microplate reader (BioTek, Winooski, VT) with an excitation wavelength of 485 nm and an emission wavelength of 519 nm. FITC-dextran concentrations in the mouse sera were

determined from standard curves generated by serial dilution of FITC-dextran. Serum from mice that were not gavaged with the permeability tracer was used as a control and subtracted from the

tested samples. STATISTICAL ANALYSIS Statistical analyses were performed using GraphPad Prism v7.0. Prior to statistical analysis, normality was tested using the Shapiro-Wilk normality

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glucose transporters. _Front Immunol._ 10, 833 (2019). Article  CAS  PubMed  PubMed Central  Google Scholar  Download references ACKNOWLEDGEMENTS This work was supported by NIH R01 DK109560

(to M.M.). Glycomic and glycoproteomic analyses were supported in part by NIH 1S10OD018530 and P41GM10349010 grants (to P.A.) at the Complex Carbohydrate Research Center. AUTHOR INFORMATION

AUTHORS AND AFFILIATIONS * Department of Infectious Diseases and Immunology, University of Florida, Gainesville, FL, 32611, USA Yong Ge, Minghao Gong, Mojgan Zadeh, Jing Li & Mansour

Mohamadzadeh * Division of Gastroenterology, Hepatology and Nutrition, Department of Medicine, University of Florida, Gainesville, FL, 32611, USA Yong Ge, Minghao Gong, Mojgan Zadeh & 

Mansour Mohamadzadeh * Department of Comparative, Diagnostic and Population Medicine, University of Florida, Gainesville, FL, 32611, USA Jeffrey R. Abbott * Department of Pathology,

Immunology and Laboratory Medicine, University of Florida, Gainesville, FL, 32611, USA Wei Li & Laurence Morel * Complex Carbohydrate Research Center, University of Georgia, Athens, GA,

30602, USA Roberto Sonon, Nitin T. Supekar & Parastoo Azadi * Division of Pulmonary, Allergy, Critical Care and Sleep Medicine, Department of Medicine, Emory University School of

Medicine, Atlanta, GA, 30322, USA Yating Wang, Dean P. Jones & Shuzhao Li Authors * Yong Ge View author publications You can also search for this author inPubMed Google Scholar * Minghao

Gong View author publications You can also search for this author inPubMed Google Scholar * Mojgan Zadeh View author publications You can also search for this author inPubMed Google Scholar

* Jing Li View author publications You can also search for this author inPubMed Google Scholar * Jeffrey R. Abbott View author publications You can also search for this author inPubMed 

Google Scholar * Wei Li View author publications You can also search for this author inPubMed Google Scholar * Laurence Morel View author publications You can also search for this author

inPubMed Google Scholar * Roberto Sonon View author publications You can also search for this author inPubMed Google Scholar * Nitin T. Supekar View author publications You can also search

for this author inPubMed Google Scholar * Parastoo Azadi View author publications You can also search for this author inPubMed Google Scholar * Yating Wang View author publications You can

also search for this author inPubMed Google Scholar * Dean P. Jones View author publications You can also search for this author inPubMed Google Scholar * Shuzhao Li View author publications

You can also search for this author inPubMed Google Scholar * Mansour Mohamadzadeh View author publications You can also search for this author inPubMed Google Scholar CONTRIBUTIONS M.M.

directed the experiments, which were executed by Y.G., M.G., M.Z., and J.L. Y.G. and M.Z. performed animal experiments; Y.G. performed flow cytometry analysis, genetic and biochemical

assays. Y.G. and J.L. constructed RNA-Seq libraries. M.G. analyzed RNA-Seq data. J.R.A. evaluated and scored all colonic tissue sections. W.L. and L.M. performed extracellular flux analysis.

R.S., N.T.S., and P.A. analyzed glycomics and glycoproteomics. M.G., Y.W., D.P.J. and S.L. performed, analyzed, and directed metabolomic studies. Y.G. and M.M. wrote the paper.

CORRESPONDING AUTHOR Correspondence to Mansour Mohamadzadeh. ETHICS DECLARATIONS COMPETING INTERESTS The authors declare no competing interests. ADDITIONAL INFORMATION PUBLISHER’S NOTE

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PERMISSIONS Reprints and permissions ABOUT THIS ARTICLE CITE THIS ARTICLE Ge, Y., Gong, M., Zadeh, M. _et al._ Regulating colonic dendritic cells by commensal glycosylated large surface

layer protein A to sustain gut homeostasis against pathogenic inflammation. _Mucosal Immunol_ 13, 34–46 (2020). https://doi.org/10.1038/s41385-019-0210-0 Download citation * Received: 08

July 2019 * Revised: 30 August 2019 * Accepted: 23 September 2019 * Published: 16 October 2019 * Issue Date: January 2020 * DOI: https://doi.org/10.1038/s41385-019-0210-0 SHARE THIS ARTICLE

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