A Bacteroides fragilis protease activates host PAR₂ to induce intestinal pain and inflammation

SUMMARY

Protease-activated receptor 2 (PAR₂) is a central regulator of intestinal barrier function, inflammation and pain. Upregulated intestinal proteolysis and PAR₂-signaling are implicated in inflammatory bowel diseases (IBDs) and irritable bowel syndrome (IBS), conditions often associated with gut microbiome alterations. To identify potential bacterial regulators of PAR₂ activity, we developed a functional assay for PAR₂ processing to screen a library of diverse gut microbes. We identify multiple bacteria that secrete proteases capable of cleaving host PAR₂. Using chemoproteomic profiling with a covalent irreversible inhibitor, we uncover a previously uncharacterized Bacteroides fragilis protease Bfp1 and show that it cleaves and activates PAR₂ in multicellular and murine models. PAR₂ cleavage by Bfp1 disrupts the intestinal barrier, sensitizes nociceptors, and triggers colonic inflammation and abdominal pain. Collectively, our findings uncover Bfp1-mediated PAR₂-processing as an axis of host-commensal-interaction in the gut that has the potential to be targeted for therapeutic intervention in IBD or IBS.

eTOC Blurb

Lakemeyer and colleagues analyze the secretome of gut bacteria to identify factors that target host PAR₂. They find that the Bacteroides fragilis protease Bfp1 cleaves and activates PAR₂, disrupting intestinal barrier function and triggering inflammation and pain. This suggests microbial proteolysis may function through host signaling pathways to regulate pathogenesis.

Graphical Abstract

INTRODUCTION

An imbalance of the gut microbiota, known as dysbiosis, has been linked to gastrointestinal diseases such as colorectal cancer,¹ inflammatory bowel disease (IBD)² and irritable bowel syndrome (IBS)³, as well as seemingly unrelated conditions including Parkinson’s disease⁴ and response to chemotherapy.⁵ Despite the clinical relevance of these observations, the underlying mechanisms that define specific bacterial contributions to pathogenesis remain largely unclear. While individual bacterial metabolites have been shown to be regulators of bacterial-host signaling,^6,7 the role of secreted proteins, especially bacterial enzymes, has been mostly overlooked.⁸ Intriguingly, the gut is the organ most exposed to proteases, both from endogenous and exogenous sources.⁹ These proteases not only mediate the digestion of dietary proteins but also regulate inflammation, pain, cell migration, apoptosis and intestinal permeability.¹⁰ Excessive proteolysis in the gut caused by host or microbial proteases has been identified as a major contributor to IBD^2,11 and IBS.¹² Moreover, proteases play pivotal roles in microbial homeostasis and mediate microbe-microbe interactions.^8,9 Therefore, identifying and modulating bacterial protease activities are promising strategies for the development of therapies for microbiome-related diseases.^13,14

Protease-activated receptor 2 (PAR₂) is a central regulator of epithelial barrier function, intestinal inflammation and pain. PAR₂ is a G-protein coupled receptor (GPCR) that mediates a number of downstream signaling pathways. In contrast to most GPCRs, PAR₂ is not activated by small molecule binding. Rather, proteolysis within the extracellular N-terminal domain (NTD) reveals a neo-N-terminus that acts as a tethered ligand for the cleaved receptor (Figure 1A) or induces conformational changes that activate the cleaved receptor.¹⁵ Excessive proteolysis in the gut and consequent PAR₂-signaling have been linked to IBD and IBS severity and intestinal pain.^11,12 While the modulation of PAR₂ signaling is understood as a virulence strategy for pathogens such as Enterococcus faecalis,¹⁵ it remains unclear which extracellular proteases from gut bacteria modulate host PAR₂ signaling and thereby affect gut physiology and pathology.

Figure 1. Identification of bacterial proteases that cleave the PAR₂-NTD.

(A) Mechanism of activation of PAR₂ through host or microbial proteases.

(B) Schematic representation of the dual-labeled substrate probe and workflow for the screening of bacterial cultures for PAR₂-NTD cleaving proteases. After incubation with culture supernatants, intact or cleaved substrate is affinity-enriched on magnetic Ni-NTA beads and fluorescence in solution measured to quantify proteolysis.

(C) Results from the PAR₂-NTD proteolysis screening mapped onto a phylogenetic tree of all bacterial strains tested. Bar graph depicts PAR₂-NTD proteolysis relative to no-protease control (0% cleavage) and treatment with excess of trypsin (100% cleavage).

(D) PAR₂-processing activity of most active supernatants upon pre-incubation DMSO (control) or serine hydrolase inhibitor fluorophosphonate-alkyne (FP-alkyne, 20 μM). Data are presented as mean ± SEM.

Here, we describe a screening platform for functional profiling of the secretomes of gut bacteria that process human PAR₂, which identified members of the Bacteroidota phylum with high PAR₂-cleaving activity. By applying covalent inhibitors and chemical proteomics, we identified a previously uncharacterized and unique serine protease that we have named Bfp1 from Bacteroides fragilis, which efficiently processes PAR₂, resulting in receptor activation, internalization, and increased transepithelial permeability in intestinal organoids and in ex vivo Ussing chamber experiments. Intracolonic administration of B. fragilis culture supernatants as well as engraftment of mice with wild-type (WT) B. fragilis bacteria resulted in strong PAR₂-mediated colonic inflammation and nociception that was absent in a Δbfp1-knockout (KO) strain. Colonic administration of Bfp1 in vitro led to excitation of sensory nerves and this sensitization was lost in mice lacking PAR₂ in NaV1.8-expressing nociceptors. Overall, our work has uncovered Bfp1-mediated PAR₂-processing as a mechanism of host-commensal interactions in the gastrointestinal tract that has profound impact on host physiology.

RESULTS

Development of an affinity-pulldown platform to profile microbial PAR2-proteolysis

To identify bacterial strains that process the human PAR₂-NTD via secreted proteases, we developed a sensitive in vitro screening platform for directly testing non-concentrated bacterial culture supernatants in a multiplexed manner. Using expressed protein ligation and bioorthogonal chemistry (Figure S1), we generated a synthetic dual-labeled reporter substrate representing the 46 amino acid-spanning PAR₂-NTD with N-terminal hexahistidine (His₆) affinity tag and C-terminal Cy5 fluorophore (Figure 1B). Upon cleavage by microbial proteases, the fluorophore is separated from the affinity tag. When the His₆-tagged peptides are removed from the solution via Ni-NTA-based affinity pulldown, the extent of PAR₂-NTD cleavage can be quantified by the amount of released fluorescent fragment remaining in the soluble fraction (Figure 1B). Optimization using recombinant trypsin, the canonical PAR₂ activator, and Enterococcus faecalis culture supernatants that contain the known PAR₂-cleaving protease gelatinase¹⁶ resulted in a robust and sensitive assay that requires minimal volumes (10 μL) of non-concentrated culture supernatants (Figure S2).

We applied our screening platform to a collection of 140 human gut microorganisms, representing 104 species and spanning 5 phyla, that has previously been applied to profile the production of microbial metabolites.¹⁷ To create comparable datasets, we cultivated all strains in a rich, undefined medium (Mega Medium, MM), which supports the growth of diverse bacteria.¹⁷ Bacteria were grown in 96-well microtiter plates, harvested by centrifugation and sterile-filtration, and subsequently tested in the PAR₂-proteolysis assay. As an outcome of this global screening, we identified culture supernatants from 45 bacterial strains that processed more than 50% of the PAR₂-substrate in our assay (Figure 1C, Table S1). While it has previously only been hypothesized that commensals might interact with the human host via PAR-modulation, these results highlight that a remarkable number of bacterial strains secrete potential PAR₂-processing proteases. In addition to select Bacillota species, such as two Enterococcus faecalis strains, we observed very strong proteolytic activities within the Bacteroidota phylum. In particular, five Bacteroides fragilis strains showed consistently high PAR₂-processing, ranging from 78% – 86% of total substrate. For the validation of their proteolytic activity, all “hit” strains were individually cultivated on a large scale, harvested at early stationary phase (to exclude release of intracellular proteases through cell lysis) and subjected to repeated testing with a shortened proteolysis time (to select for the most active supernatants). Out of the 45 initial hits, 27 passed these more stringent selection criteria (Figure 1D [blue bars] and S3, see Supporting Information). It is noteworthy that apart from the two E. faecalis strains, Clostridium sporogenes ATCC 15579 remained the only non-Bacteroidota PAR₂-processor identified.

Identification of uncharacterized B. fragilis serine proteases by chemical proteomics

Proteases can be classified based on their active sites into serine-, cysteine, metallo-, aspartic-/glutamic-, asparagine- and threonine-proteases. Previous work has linked excessive serine protease activities with intestinal diseases such as IBD and IBS.^8,18,19 Multiple serine proteases from the host or exogenous sources are known to cleave and activate PAR₂.²⁰ As serine proteases are additionally predicted to be the largest group of proteases to be secreted by bacteria,²¹ and can be efficiently inhibited by broad-spectrum inhibitors, we focused our protein identification efforts specifically on serine proteases.

Bacterial culture supernatants were pre-treated with a covalent, broad-spectrum serine hydrolase probe, fluorophosphonate (FP)-alkyne (Figure 2A)²², prior to the PAR₂-proteolysis assay (Figure 1D). FP-alkyne partially inhibited PAR₂ cleavage by select strains, including Bacteroides finegoldii BEI HM-727 and Bacteroides thetaiotaomicron wh302. The strongest inhibitory effect was observed for five B. fragilis strains, resulting in 58 – 87 % inhibition compared to vehicle (DMSO)-treated samples (Figure 1D). We performed activity-based protein profiling (ABPP, schematic workflow in Figure 2B)²³ on culture supernatants of the reference strain B. fragilis NCTC 9343 to identify the PAR₂-processing protease(s). We first optimized labeling conditions via gel-based ABPP using a fluorescent FP-tetramethylrhodamine (FP-TMR) probe (Figure S4), prior to quantitative ABPP using an FP-biotin probe, streptavidin-based affinity-enrichment of labeled proteins and LC-MS/MS-based analysis. This identified 20 proteins, 10 of which are potential peptidases or proteases, that were strongly (> 4-fold) and significantly (p < 0.05) enriched compared to the vehicle (DMSO) control (Figure 2C, Table S2). For the unambiguous identification of the PAR₂-processing protease, we screened a small set of 11 covalent protease inhibitors (Figure S5A) and identified the peptide chloromethyl ketone MLP7 (Figure 2A) as an inhibitor of the PAR₂-processing activity of B. fragilis (Figure S5B). Pretreatment of culture supernatants with MLP7 prior to labeling with FP-biotin blocked the protease-labeling by FP-probes in a competitive and concentration-dependent manner (Figure 2B and S4). Importantly, the MS-based competitive ABPP experiment resulted in only two protease candidates with the UniProt²⁴ identifiers Q5LDF9 (gene name BF9343_2070) and Q5LIA5 (gene name BF9343_0342) (Figure 2D, Table S2). Both enzymes are uncharacterized putative lipoproteins that, according to InterPro analysis²⁵, belong to the poorly characterized families of S41-proteases or tail-specific proteases (Figure 2E).

Figure 2. Identification of two uncharacterized B. fragilis proteases.

(A) Chemical structures of inhibitors FP-alkyne and MLP7.

(B) Schematic representation of competitive ABPP workflow.

(C) Volcano plot of FP-biotin protein targets in the secretome of B. fragilis NCTC 9343. Significantly enriched peptidase/protease candidates are colored in blue.

(D) Volcano plot for competitive ABPP experiment with pretreatment by MLP7 uncovers two PAR₂-processing candidates, TspA and Bfp1.

(E) InterPro domain analysis of the proteases Q5LIA5 (TspA) and Q5LDF9 (Bfp1).

(F) Phylogenetic distribution of bfp1 and tspA based a database of 1,520 reference genomes of culturable bacteria from human gut microbiota.

(G) Growth curves for WT B. fragilis and the B. fragilis Δbfp1 and ΔtspA protease knockouts.

(H) Relative PAR₂-proteolysis of the WT B. fragilis and respective knockout strains confirms that Bfp1 is the responsible PAR₂-processing protease.

(I) Structural prediction of Bfp1 (AlphaFold model) with a typical S41 serine protease domain and an extended helical domain of unknown function.

BLAST analysis against 1,520 reference genomes from cultivated human gut bacteria²⁶ resulted in 381 Bacteroidota spp. with homologs of Q5LIA5 (Figure 2F, Table S3). In contrast, Q5LDF9 was only identified in the genomes of the 31 unambiguously annotated B. fragilis strains in the database (Figure 2F, Table S4). We therefore propose to name the conserved Q5LIA5 protease tail-specific protease A (TspA) and the Q5LDF9 enzyme Bacteroides fragilis serine protease 1 (Bfp1). The unique distribution of Bfp1 in B. fragilis and lack of Bfp1-negative B. fragilis strains suggests a specific physiological role for this serine protease.

Genetic deletion mutants uncover Bfp1 to be the PAR2-processing protease

To determine which enzyme candidate was responsible for our observed PAR₂-processing activity, we generated two B. fragilis NCTC9343 strains in which either the tspA or the bfp1 gene was genetically deleted by heterologous recombination (Supporting Information).²⁷ Both deletion mutants showed only a slightly delayed growth compared to the WT strain, highlighting that these enzymes are not essential for bacterial survival in vitro under our conditions (Figure 2G). Next, bacterial culture supernatants were subjected to the PAR₂-proteolysis assay. While knockout of tspA did not alter PAR₂-processing compared to WT B. fragilis, the supernatants from the B. fragilis Δbfp1 strain exhibited only residual PAR₂-processing activity, which could not be further reduced by pretreatment with FP-alkyne (Figure 2H). Consequently, we concluded that Bfp1 is the serine protease that mediates PAR₂-NTD proteolysis in B. fragilis culture supernatants. The predicted structure of Bfp1²⁸ contains a proteolytic core with its active site serine residue and an extended helical domain which is annotated as “domain of unknown function” (Figure 2I).²⁵

Bfp1-mediated cleavage of full-length PAR2 at the cell-surface

To determine whether bacterial Bfp1 can cleave the extracellular NTD of intact PAR₂ at the cell surface and stimulate endocytosis of the cleaved receptor, consistent with activation, we incubated B. fragilis culture supernatants with HEK293T cells expressing human PAR₂ with an extracellular HA-epitope and an intracellular fluorescent mApple tag (Figure 3A). Cells were exposed to bacterial supernatants, and HA epitope and mApple were localized by immuno-fluorescence microscopy. In unstimulated cells (vehicle) and in cells exposed to Mega Medium only (control), HA and mApple colocalized at the plasma membrane (Figure 3B). Trypsin (10 nM, 30 min, positive control) removed the HA-tag and caused redistribution of mApple to endosomes, consistent with PAR₂ cleavage and activation. A similarly strong phenotype was observed upon treatment with culture supernatants from WT B. fragilis and this altered localization was partially restored for B. fragilis Δbfp1, highlighting that Bfp1 indeed processes the intact, membrane-bound PAR₂ (Figure 3B). Concomitantly, relative quantification of the internalized mApple-PAR₂ fluorescence signal compared to membrane-associated signal showed significantly increased endocytosis upon administration of trypsin (p < 0.05) and supernatant from WT B. fragilis (p < 0.001) but not B. fragilis Δbfp1 (Figure S6A).

Figure 3. Bfp1 processes PAR₂ at the cell-surface, induces endocytosis and calcium influx.

(A). Cartoon showing the proteolytic processing of N-terminal HA-epitope and C-terminal mApple fluorescent tagged PAR₂ to release the HA tag.

(B). Cells (HEK293) expressing dual tagged PAR2 (as shown in (A)) were exposed to trypsin (10 nM) or 10-fold concentrated supernatants from WT B. fragilis or Δbfp1-knockout cultures, equally concentrated Mega Medium (MM), or purified Bfp1 (500 nM), and analyzed by immuno-fluorescence microscopy to detect HA (green), mAppple (red) or DAPI (blue). Trypsin (positive control) removes Flag and stimulates redistribution of mApple to endosomes (arrows), denoting PAR₂ cleavage, activation and internalization. WT B. fragilis supernatant has similar effects. B. fragilis Δbfp1-knockout supernatant does not completely remove Flag, denoting slower cleavage of PAR₂. Recombinantly expressed and purified Bfp1 also induces PAR₂ cleavage and internalization. Scale bar: 10 μm, arrowheads denote surface PAR₂, arrows show internalized receptor.

(C, D). Intracellular calcium levels [iCa²⁺] of HEK293-Par₂^+/+ or HEK293-Par₂^−/− cells upon addition of purified Bfp1 (500 nM), as measured by Fluo-4 fluorescence. Pretreatment with the PAR₂ antagonist AZ3451 (1 μM) blocked Bfp1-mediated calcium influx. Data are presented as mean ± SEM, n = 3. Statistical analysis: one-way ANOVA * p<0.05.

To further validate that the observed effects are specifically mediated by Bfp1, we aimed to perform recombinant production and purification of soluble Bfp1 without its signal peptide sequence using Escherichia coli. Despite extensive efforts on optimization, only minimal amounts of catalytically active Bfp1 could be purified and validated for its catalytic activity (Figure S6B–E), indicating a potential importance protein maturation under native conditions in B. fragilis. We used the limited supply of active purified Bfp1 to validate the cleavage of the full-length PAR₂ (Figure 3B, last panel), which resembled the phenotype of WT B. fragilis supernatant, highlighting that supernatant is a valid substitute for the purified protein.

Canonical activation of PAR₂ induces calcium influx, increasing the intracellular calcium levels [iCa²⁺].¹⁵ Similarly, we observed that purified Bfp1 (500 nM) induced a significantly prolonged increased [iCa²⁺] in HEK293 PAR₂^+/+ cells (AUC: 196 ± 49), but not in PAR2^−/− cells (AUC: 10 ± 9, p = 0.019) or PAR2^+/+ cells treated with the PAR₂-antagonist AZ3451²⁹ (AUC: 2 ± 2, p = 0.014 to PAR₂^+/+) (Figure 3C, D). Taken together, these data demonstrate that Bfp1 is able to cleave membrane-bound PAR₂ and induce intracellular signaling.

Bfp1 increases epithelial permeability in human intestinal organoids

PAR₂ is a central modulator of the intestinal epithelial barrier and excessive activation of PAR₂ has been linked to increased paracellular permeability and “leaky gut” syndrome.^2,9,30 To assess whether Bfp1 modulates intestinal permeability in a physiologically relevant human model, we subjected organoids from healthy ileal adult tissue to bacterial culture supernatants and analyzed paracellular permeability in organoid-derived monolayers and apical-out organoids (Figure 4A). Apical-out organoids facilitate permeability testing as effector molecules that are predicted to interact with the apical side can be added directly to the culture medium, circumventing the need for laborious intra-organoid injection.³¹

Figure 4. Bfp1 increases epithelial permeability in human intestinal organoids and murine colons.

(A) Experimental workflow for testing epithelial permeability.

(B) Time-dependent permeability analysis of organoid-derived monolayers from healthy ileal adult tissue upon exposure to bacterial culture supernatants (4-fold concentrated) or controls, as determined by transepithelial electrical resistance (TEER). Trypsin: 4 nM, PAR2-agonist peptide 2-furoyl-LIGRLO-amide (2F): 1 μM, mean + SD, n = 3.

(C, D) Permeability analysis of apical-out organoids, as determined by FITC-dextran permeability. Quantitative analysis (C) and representative microscopy images of organoids (D). Trypsin: 8 nM, 2F: 25 μM, violin plot with mean ± median. Statistical testing via two-way ANOVA. * p<0.05, ** p<0.01, **** p<0.0001. Nonsignificant (ns) test differences are not presented. Scale bar: 50 μm.

(E) Ex vivo assessment of colonic permeability using Ussing chamber methodology, as determined by FITC-dextran permeability across the colonic epithelium. Mean + SD, n = 6 or 7. Statistical testing for WT B. fragilis vs. MM via two-way ANOVA. ** p < 0.01, *** p < 0.001.

Treatment of monolayers with bacterial supernatants resulted in a significantly decreased transepithelial electrical resistance (TEER) for WT B. fragilis, but not for B. fragilis Δbfp1, which was not significantly different from the MM-treated control (Figure 4B). Upon incubation for 2 hours, the residual resistance for WT B. fragilis-treated monolayers was 378 ± 30 Ω/cm², which corresponds to 49% of the resistance for B. fragilis Δbfp1 (736 ± 10 Ω/cm², p < 0.001). In accordance with these results, culture supernatants from WT B. fragilis, but not from B. fragilis Δbfp1 induced a high percentage of organoids (71.6 ± 7.3 compared to 39.3 ± 10.7, p < 0.01) that were permeable to FITC-dextran (Figure 4C), as determined by confocal microscopy (Figure 4D).

To validate increased permeability in a mouse model, we performed Ussing-chamber experiments using ex vivo murine colons. Exposure to WT B. fragilis supernatants significantly increased colonic permeability, as indicated by elevated FITC-dextran transmission across the colonic epithelium at 2- (p < 0.01), 2.5- (p < 0.001), and 3-hours (p < 0.001) post-application compared to MM application (baseline) (Figure 4E). In contrast, tissues treated with the B. fragilis Δbfp1 supernatant did not show increased FITC-dextran permeability at any time point. These findings highlight that Bfp1-containing culture supernatants induce an overall impairment of the epithelial barrier that, during homeostatic conditions, protects the mucosa from inflammatory factors in the intestinal lumen.

Bfp1 causes PAR2-dependent excitation of nociceptors and enhances mechanosensitivity

PAR₂ is expressed by a subpopulation of dorsal root ganglia (DRG) nociceptors, where activation results in sensitization that leads to pain.^3,32,33 To determine whether B. fragilis proteases also cause PAR₂-dependent sensitization of nociceptors, we recorded patch clamp measurements of DRG neurons from WT Par₂^+/+ or global Par₂^−/− knockout (KO) mice. The rheobase (minimum input current required to fire an action potential) of DRG nociceptors was measured to assess sensitization. The rheobase of nociceptors from Par₂^+/+ mice incubated with culture MM (control) for 10 min at RT was 37.6 ± 4.3 pA (n = 29 neurons) (Figure 5A). Incubation with WT B. fragilis supernatant reduced the rheobase to 15.5 ±1.3 pA (n = 31 neurons, p < 0.001 to control), denoting heightened excitability. In sharp contrast, supernatant from B. fragilis Δbfp1 did not affect rheobase (Figure 5A, B). Moreover, application of WT B. fragilis supernatant resulted in an increased number of action potentials (WT: 5.9 ± 0.9, Δbfp1: 2.6 ± 0.5, p < 0.01) (Figure 5C) and more depolarized resting membrane potential (WT: −43.5 ± 1.1 mV, Δbfp1: −48.0 ± 1.2 mV, p < 0.05) compared to B. fragilis Δbfp1 (Figure 5D). Preincubation of supernatant from WT B. fragilis with the serine protease inhibitor FP-alkyne fully abrogated the reduction in rheobase (38.6 ± 2.5 pA, n = 28 neurons, p < 0.001 to WT supernatant) (Figure 5A). Similarly, preincubation of neurons with the PAR₂ antagonist AZ3451²⁹ prevented the reduction in rheobase (42.9 ± 5.1 pA, n = 24 neurons, p < 0.001 to WT supernatant). Corroborating these results, WT B. fragilis did not reduce the rheobase of DRG neurons isolated from Par₂^−/− global KO mice (44.7 ± 9 pA, n = 19 neurons, p<0.001 to WT supernatant in WT mice) (Figure 5A). The results show that WT B. fragilis supernatants cause hyperexcitability of mouse nociceptors and that effects are mediated by Bfp1-catalyzed cleavage of PAR₂.

Figure 5. Bfp1 induces nociception and enhances mechanosensitivity in ex vivo samples.

(A) Excitation of dorsal root ganglia (DRG) nociceptors by 0.5-fold concentrated B. fragilis culture supernatants or Mega Medium (MM) control. To validate Bfp1- and PAR₂-mediated response, supernatant was pre-treated with serine protease inhibitor FP-alkyne (100 μM, 60 min, RT), DRG were preincubated with PAR₂-antagonist AZ3451 (1 μM, 60 min, RT), or neurons from Par₂^−/− global KO mice were studied. L1-L5 DRG neurons were used for patch clamp recordings and the data was collected exclusively from small diameter neurons (≤ 25 μm). Data show a summary of all rheobase activity measurements. Statistical analysis: one-way ANOVA * p < 0.05, ** p < 0.01, *** p < 0.001. Sample numbers are depicted in the plot. (B–D) Detailed analysis of DRG nociceptor excitation. Data for rheobase (B), action potential (C) and resting membrane potential (D) of DRG neurons treated with 1-fold WT or Δbfp1 B. fragilis culture supernatants. Data are presented as mean ± SEM, Statistical testing via unpaired t-test. *p < 0.05, **p < 0.01, **** p < 0.0001.

(E–L) Effect of supernatants from WT B. fragilis and Δbfp1 cultures on murine colonic afferent neuron activity in Par₂^+/+ WT, Par₂^−/−-Nav1.8 KO as compared to Krebs buffer (vehicle). Spontaneous baseline activities (E, G, I, K) and overall mechanosensitive response of colonic afferent nerves to distension (F, H, J, L). Data are presented as mean ± SEM. Statistical testing via Wilcoxon test (baseline activity) and two-way ANOVA (mechanosensitive response), ns = not significant, *p < 0.05, **p < 0.01, **** p < 0.0001.

To determine whether B. fragilis proteases can signal from the intestinal lumen to sensitize nociceptor terminals in the colon wall to mechanical stimuli, we performed extracellular recordings for afferent nerves innervating isolated segments of the mouse colon.³³ Samples were perfused through the murine colonic lumen while action potential firing under baseline conditions and in response to noxious mechanical distension was measured. In WT Par2^+/+ mice, perfusion of MM alone had no effect on either spontaneous baseline firing (Figure 5E, p = 0.1594, n = 6 mice, 19 neuronal units) or the response to mechanical distension (Figure 5F, p = 0.4217, n = 6) compared to buffer as a vehicle control. Culture supernatants from WT B. fragilis significantly increased both spontaneous firing (Figure 5G and S7A, p = 0.0022, n = 6 mice, 23 neuronal units) and neuronal activation in response to colonic distension (Figure 5H, p = 0.0406, n = 6). Single-unit analysis revealed that 58% of the afferents were activated upon perfusion (Figure S7B) and that the activation in both distension response and spontaneous activity was driven by the wide-dynamic range units, not the high-threshold units (Figure S7C–G). In sharp contrast, perfusion of B. fragilis Δbfp1 supernatants in Par₂^+/+ colons (Figure 5I, J) did not affect spontaneous firing (p = 0.4304, n = 6 mice, 22 neuronal units) or the response to colonic distension (p = 0.9383, n = 6). Activation of colonic nociceptors and their mechanical sensitization by WT B. fragilis supernatants were not observed in colons from Nav1.8 Par₂^−/− knockout mice, with targeted deletion of PAR₂ from NaV1.8+ve nociceptors³³ (Figure 5K,L, p = 0.0972, n = 6 mice, 24 neuronal units for baseline activity; p = 0.9595, n = 6 for distension response). Colons from Par₂^+/+ Cre-mice (control) showed the same significantly increased responses to culture supernatants from WT B. fragilis as observed for WT Par₂^+/+ mice (Figure S7H,I, baseline firing: p < 0.0001, n = 7 mice, 39 neuronal units, colonic distension response: p = 0.0375, n = 7). Taken together, these results demonstrate that WT B. fragilis culture supernatants induce excitation of colonic afferent nerves, which is mediated by Bfp1-triggered activation of PAR₂ on NaV1.8+ve nociceptors.

Intracolonic injection of Bfp1 activates PAR2 and causes PAR2-dependent inflammation and nociception

Mice expressing PAR₂ C-terminally fused to monomeric ultrastable green fluorescent protein (Par₂-mugfp) allow specific localization of PAR₂ and analysis of its redistribution during disease.³⁰ In the normal colon, PAR₂ is localized to the basolateral and apical membrane of colonocytes. In the inflamed colon, PAR₂ redistributes to endosomes of colonocytes, as a result of its proteolytic activation.³⁰ It is well established that PAR₂ signals from endosomes of colonocytes, causing increased paracellular permeability, influx of luminal contents and inflammation, while signaling from endosomes of nociceptors leads to hyperexcitability and pain.^30,33 To assess whether bacterial proteases can activate PAR₂ in the colon, MM (control) and bacterial supernatants were injected into the colon of Par₂-mugfp mice. After 3 h, the colon was excised and PAR₂-muGFP was localized by immunofluorescence using GFP antibodies. In mice treated with MM, PAR₂-muGFP was confined to the basolateral and apical membranes of colonocytes (Figure 6A). In mice injected with supernatant from WT B. fragilis, PAR₂-muGFP was depleted from the plasma membrane and detected in endosomes (Figure 6B). Supernatant from B. fragilis Δbfp1 did not cause endocytosis, with PAR₂-muGFP confined to the plasma membrane (Figure 6C). The quantification of receptor internalization revealed two distinct populations in WT B. fragilis samples. We hypothesize that this variation might be due to the technical challenges encountered in collecting and processing the portion of the colon that was in contact with the supernatant, which might explain no overall statistically significant difference compared to controls (Figure S8). Overall, these results are consistent with the hypothesis that Bfp1 from B. fragilis cleaves and activates PAR₂ in the mouse colon, stimulating endocytosis of the receptor.

Figure 6. Intracolonic injection of Bfp1 evokes endocytosis of PAR₂-muGFP, increases proinflammatory cytokines/chemokines and induces colonic nociception through PAR₂ activation.

(A–C) Localization of GFP immunoreactivity (green) in isolated segments of colon from PAR₂-muGFP mice incubated with 10x concentrated (A) Mega Medium (MM), or 10x culture supernatants from (B) WT B. fragilis and (C) B. fragilis Δbfp1. Arrowheads showing PAR₂-muGFP localization on the plasma membrane of colonocytes (A, C). Arrows showing PAR₂-muGFP trafficking in intracellular compartments (B). Nuclei are visualized by DAPI-stain (blue). Representative images, independent experiments, n = 3 mice. Scale bar, 20 μm.

(D–I) Colonic cytokine mRNA levels at 3 h after intracolonic administration of Vehicle (Veh.), Mega Medium (MM) and supernatants of WT B. fragilis or B. fragilis Δbfp1 in Par₂^+/+ WT (D–F) and Par₂^−/− global KO (G–H) mice. Mean ± SEM, One-Way ANOVA, Tukey’s multiple comparison test n = 5 or 6 mice.

(J–N) Abdominal mechanical allodynia measurement following intracolonic administration of supernatant

(J). Mega medium (MM), B. fragilis Δbfp1, and WT B. fragilis evoked colonic nociception and area under curve (AUC) in Par₂-mugfp (K), Par₂^−/− global KO (L) and Par₂-Nav1.8 (M). (N) Graphical representation of WT B. fragilis-evoked colonic nociception and AUC in Par₂-mugfp, Par₂^−/− global KO, and Par₂-Nav1.8. Mean ± SEM, n = 5 or 6 mice. Two-way ANOVA, Tukey’s multiple comparison test. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

(O, P) Visceral motor response (VMR) to colorectal distension (CRD) of mice receiving two consecutive B. fragilis supernatant enemas (S1 and S2). Administration of supernatants from WT B. fragilis (O) but not B. fragilis Δbfp1 (P) resulted in significantly increased VMR compared to the initial response following B. fragilis Δbfp1 supernatant. Mean ± SEM, n = 5 or 7 mice. Two-way ANOVA, *p < 0.05.

Notably, intracolonic injection of supernatant from WT B. fragilis but not B. fragilis Δbfp1 stimulated a marked increase in the expression of tumor necrosis factor α (TNFα) (>4-fold higher, p = 0.0245), interleukin (IL)-1β (>20-fold higher, p = 0.0075) and CXCL1 (>45-fold higher, p = 0.0056) mRNA compared to vehicle or MM in the colon of Par₂^+/+-mugfp mice, consistent with an inflammatory response (Figure 6D–F). In contrast, injection of the same samples into the colon of global Par₂^−/− KO mice did not stimulate any significant cytokine or chemokine transcription (TNFα, 0-fold change, p = 0.3184; IL1β, 0-fold change, p = 0.3470 and CXCL1, 1-fold higher, p = 0.2235; Figure 6G–I). Additionally, we tested a panel of 40 cytokines and chemokines in Par₂^+/+-mugfp mice and identified several (e.g. CXCL5, CXCL12, I-CAM1 and IL16) to be significantly elevated upon intracolonic administration of supernatants from WT B. fragilis but not B. fragilis Δbfp1 or MM control (Figure S9). Thus, Bfp1 induces PAR₂-dependent inflammation in the colon.

After intracolonic injection, PAR₂ agonists are known to cause mechanical allodynia in the colon.^3,30,33 To determine whether Bfp1 causes mechanical allodynia, we injected MM (control) or bacterial supernatants into the mouse colon. At various time points, we measured withdrawal responses to stimulation of the abdomen with calibrated von Frey filaments (VFF) to assess mechanical allodynia (Figure 6J). In WT Par₂^+/+ mice, MM decreased the withdrawal threshold from 1.2 ± 0.0 g (baseline) to 0.5 ± 0.0 g, after 1 h, indicative of mechanical allodynia (Figure 6K). WT B. fragilis supernatant caused a significantly larger decrease in threshold from 1.3 ± 0.1 g (baseline) to 0.1 ± 0.0 g after 1 h, which was maintained for at least 6 h (0.3 ± 0.1 g), consistent with long-lasting mechanical allodynia of the colon. Supernatant from B. fragilis Δbfp1 induced a significantly lower response compared to WT supernatant with a baseline threshold of 1.3 ± 0.0 g and a 1 h injection time point mechanical threshold of 0.3 ± 0.1 g (Figure 6K). In Par₂^−/− global KO mice (Figure 6L) or in mice with selective PAR₂ deletion from Nav1.8+ve nociceptors (Par₂^−/−Nav1.8) (Figure 6M), the withdrawal thresholds to MM, WT B. fragilis and B. fragilis Δbfp1 supernatant were comparable. Analysis of the area-under-the-curve reveals that the nociceptive threshold changes following the injection of WT B. fragilis supernatant in the colon are significantly reduced in Par₂^+/+ mice, showing a twofold decrease compared to both Par₂^−/− global knockout and Par₂^−/−Nav1.8 mice (p < 0.0001). Thus, the nociceptive response is largely mediated by nociceptor PAR₂ because it is absent from mice with global or targeted PAR₂ deletion on Nav1.8+ve nociceptors (Figure 6N).

Noxious colorectal distension (CRD) triggers the visceral motor response (VMR), a nociceptive brainstem reflex consisting of abdominal muscle contractions, which can be monitored by electromyography.³³ This approach allows assessment of visceral sensitivity in conscious mice and further strengthens our mechanical sensitivity data. We administered colorectal enemas containing supernatants from WT B. fragilis or B. fragilis Δbfp1 to mice. At 90 min from the injection, we recorded VMR to CRD at 40, 60 and 80 μL. We observed increased visceral sensitivity to CRD in mice treated with supernatant from WT B. fragilis (Figure 6O), but not B. fragilis Δbfp1 (Figure 6P). Together, these results support the hypothesis that B. fragilis Bfp1 activates PAR₂ in the colon to cause an increase in colonic pain.

Engraftment of the mouse colon with B. fragilis triggers abdominal nociception

To determine the impact of secreted Bfp1 on colonic pain in a mouse model, we repopulated antibiotic-treated mice with either WT B. fragilis or B. fragilis Δbfp1 knockout strains (Figure 7A). Mice were treated with an antibiotic cocktail (ampicillin, vancomycin, neomycin, metronidazole) in drinking water for 7 days. We analyzed fecal pellets by qRT-PCR on days 0 (pre-treatment), 3, and 7 of the antibiotic treatment to assess the extent of elimination of colonic bacteria. On day 3, all mice treated with antibiotic cocktail had no detectable bacterial DNA (Figure S10A). Mice then received vehicle, WT B. fragilis or B. fragilis Δbfp1 by gavage for 8–19 days to attain engraftment, which was verified by qRT-PCR analysis of fecal pellets for B. fragilis DNA (Figure S10B), and via metagenomic analysis (Figure S10C–F).

Figure 7. Engraftment of the mouse colon with B. fragilis results in Bfp1- and PAR₂-dependent nociception and behavioral changes.

(A–G) Experimental design for the repopulation and engraftment with B. fragilis strains following antibiotic depletion of the natural intestinal microbiota of Par₂^+/+ mugfp (B–D) and Par₂^−/− global KO (E–G) mice. (B, E) Abdominal mechanical threshold throughout the whole antibiotics/repopulation experiment. Area under curve (AUC) during repopulation (C, F) and Engraftment (D, G) periods.

(H–M) Assessment of spontaneous non-evoked behavior of engrafted mice at day 19 of the experimental protocol. (H, K) Visits to the center of the Arena. (I, L) Body Grooming events, (J, M) Number of still events. Mean ± SEM, n = 5 mice, One-Way ANOVA, and Two-Way ANOVA with Tukey’s multiple comparison test. *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001.

Abdominal nociception was evaluated by measuring withdrawal responses to abdominal stimulation with VFFs. Antibiotic treatment of Par₂-mugfp mice (equivalent to Par₂^+/+ WT)³⁰ decreased the withdrawal threshold to VFF stimulation compared to mice that had not received antibiotics, indicating mechanical allodynia (Figure 7B–D). The administration of vehicle to antibiotic-treated mice resulted in a normalization of the withdrawal threshold to pre-experiment baseline by 15 days. In marked contrast, in mice repopulated with WT B. fragilis, mechanical allodynia was fully sustained during the periods of repopulation and engraftment for at least 19 days (Figure 7B). In comparison, repopulation with B. fragilis Δbfp1 resulted in a partial normalization of mechanical allodynia by day 19. At day 19, the von Frey withdrawal threshold for antibiotic-treated mice was 1.3 ± 0.0 g, 1.2 ± 0.0 g for vehicle-treated mice, 0.3 ± 0.0 g for WT B. fragilis repopulated mice, and 0.9 ± 0.1 g for B. fragilis Δbfp1 repopulated mice (Figure 7B). Area-under-the-curve analysis for the period from day 17 to 19 revealed a highly significant (p<0.001) difference between mice engrafted with WT B. fragilis (0.3 ± 0.1) compared to mice engrafted with the Δbfp1 strain (0.8 ± 0.1, Figure 7D). In stark contrast, engraftment of mice with whole-body deletion of PAR₂ (Par₂^−/− global, Figure 7E–G) resulted in a drastically reduced colonic nociception upon repopulation with WT B. fragilis at day 19 (0.7 ± 0.1 g) that was not significantly different compared to repopulation with B. fragilis Δbfp1 (0.8 ± 0.1 g, Figure 7E). Accordingly, area-under-the-curve analysis for the period from day 8 to 18 (Figure 7F) and day 17 to 19 (Figure 7G) also did not result in a significant difference between the WT and Δbfp1 knockout strains. In contrast to the heightened mechanical allodynia in mice repopulated with WT B. fragilis, the mRNA expression of TNFα, IL1β and CXCL1 in the colon at 19 days was not significantly different between antibiotic-treated Par₂^+/+ or Par₂^−/− mice receiving vehicle, WT B. fragilis or B. fragilis Δbfp1 (Figure S11).

At day 19 of the engraftment period, we evaluated spontaneous pain using exploratory, grooming and locomotor behaviors of the mice in a behavioral spectrometer, which allows objective quantification of pathological and pain-like behavior in preclinical mouse models of disease (Figure 7 H–M and S12).³⁴ For Par₂₋mugfp mice, there were no detectable differences in any exploratory, grooming or locomotor behavior between non-antibiotic-treated mice and antibiotic-treated mice receiving vehicle or B. fragilis Δbfp1 (Figure 7H–J and S12A–C). In contrast, antibiotic-treated mice receiving WT B. fragilis showed significantly decreased activity (Figure S12A) and number of visits to the central area compared to mice receiving B. fragilis Δbfp1 (Figure 7H, p < 0.0001 and Figure S13, Video 1). Moreover, antibiotic-treated mice receiving WT B. fragilis showed a significant increase in the number of total body grooming events (Figure 7I, p<0.01) and still events (Figure 7J, p < 0.05) compared to mice receiving B. fragilis Δbfp1. Notably, none of the described exploratory, grooming or locomotor behaviors were significantly different for Par₂^−/− global KO mice (Figure 7K–M and Figure S12E–F). Taken together, these results are consistent with the hypothesis that B. fragilis Bfp1 mediates not only the induction of intestinal permeability and initiation of the proinflammatory signaling, but also colonic pain and pain-like behavior in mice via the PAR₂-axis.

DISCUSSION

Diseases of gastrointestinal pathology, including IBD and IBS, are thought to arise from dysregulation across three interrelated systems: the gut microbiota, the epithelial barrier, and immune responses.^35–37 Previous studies have linked excessive protease activities and elevated PAR₂-signaling to disrupted barrier function, inflammation, abdominal pain and IBD severity,^11,19,38 but the role of specific gut bacterial proteases has yet to be fully elucidated.^9–12,19,39 Abdominal pain is also a unifying symptom of IBS, and has been related to increased proteolysis and activation of PAR₂ in the colon.^3,33,40 To identify bacterial proteases capable of cleaving PAR₂, we developed an assay that enables direct screening of bacteria found in the human gut for their ability to produce and secrete proteases that cleave PAR₂. By using the full-length N-terminal domain of PAR₂, it was possible to screen for proteases that cleave the receptor at any location within the N-terminus accessible to an extracellular enzyme. Using this approach, we found that a substantial number of intestinal bacteria (19% of our 140-member library), particularly Bacteroides spp., secrete PAR₂-processing proteases (Figure 1C, D). Subsequent functional studies using broad-spectrum inhibitors and activity-based probes showed that a previously uncharacterized serine protease Bfp1, produced by B. fragilis, cleaves the N-terminus of PAR₂, resulting in impaired barrier function, pain and intestinal inflammation.

While the onset and progression of IBD and IBS are multifaceted and likely cannot be attributed to a single bacterial species or enzyme, our findings identify a potentially valuable new target for therapeutics that could suppress some of the key pathological symptoms of IBD and IBS.^3,41,42 Our work highlights the complex set of interactions taking place between gut bacteria and the host mediated by secreted microbial proteases. Furthermore, because diverse bacterial species produce protease activities that are capable of cleaving PAR₂, this receptor could serve as a possible nexus for competition between bacteria in a community.

Cleavage of PAR₂ by Bfp1 results in receptor endocytosis (Figure 3), which is consistent with its canonical activation.^30,33 However, PARs are GPCRs capable of triggering diverse downstream processes through ‘biased signaling’, wherein the location of proteolytic cleavage determines coupling to specific G-proteins and distinct signaling pathways.^20,43 Since our assay detects cleavage at any location along the full N-terminal domain, our top hits from the screen have the potential to mediate a diverse range of downstream effects, either through (i) activation by canonical or biased signaling or (ii) receptor inactivation via removal of the tethered ligand. The ability to modulate multiple activities of PAR₂ may markedly affect gut physiology, considering the central role of PAR₂ in controlling epithelial barrier integrity, colonic inflammation and visceral pain.^33,44–46 Given that we identified ten putative serine proteases/peptidases in the supernatant of B. fragilis, it was striking that genetic deletion of Bfp1 alone was sufficient to disrupt PAR₂-activation. This suggests that the cohort of proteases secreted by B. fragilis and other Bacteroidota spp. may play different roles in modulating host or microbial processes and that we have only identified one such node which involves PAR₂ as a substrate.

Previous studies found that antibiotic treatment reduces bacterial serine protease activity in the mouse colon, leading to diminished PAR₂ processing and expression by colonocytes, but the source of this activity has remained unclear.⁴⁷ Beyond the previously identified PAR₂ activator from E. faecalis,¹⁶ our work expands the limited list of PAR₂-NTD-processing bacteria in the human intestine^18,48 to include Clostridium sporogenes and 14 diverse Bacteroidota species (representing 24 strains). Notably, Bfp1 is one of the few bacterial proteases identified and validated as a PAR₂-activating enzyme. Although the strong proteolytic activity of B. fragilis and other Bacteroidota spp. was first characterized more than 35 years ago,^49–51 these bacteria have historically been studied primarily for their ability to degrade complex carbohydrates and produce bioactive metabolites. Despite these advances, the functional roles of Bacteroidota proteases remain underexplored,^52,53 with few proteases mechanistically characterized.^54–59 Notably, the human gut metagenome contains over 285 putative bacterial serine proteases,⁸ making targeted enzyme identification a significant challenge. Using our competitive ABPP approach (Figure 2), we efficiently pinpointed Bfp1 as the protease responsible for PAR₂ proteolysis, demonstrating the power of chemical proteomics to functionally annotate uncharacterized microbial enzymes.

Our finding that Bacteroidota spp. secrete PAR₂-processing proteases aligns well with recent reports that demonstrate a link between bacteria-associated proteolysis and IBD/IBS.^60–62 A recent multi-omics study revealed uncharacterized proteases secreted from B. vulgatus that induce colitis in mouse models.⁶³ Notably, as B. vulgatus is also a top hit in our screen, it is intriguing to hypothesize that the reported proinflammatory phenotypes are mediated by proteolytic processing of PAR₂.

While enterotoxigenic B. fragilis (ETBF), producing the metalloprotease fragilysin (BFT), contributes to acute diarrheal, secretory diarrhea and colonic epithelial damage, non-toxigenic B. fragilis is generally regarded as a commensal in the human intestine.⁶⁴ However, studies focused on the role of B. fragilis in IBD and IBS have yielded diverse outcomes, with evidence for both protective^65–67 and pathogenic^68–72 effects. The majority of these studies focus on metagenomic analyses which are often correlative. In the current study, we identify a specific bacterial effector, Bfp1, its human receptor, PAR₂, and the physiological phenotypes (loss of barrier function, inflammation and pain) that result from their interaction, thus providing a mechanistic understanding of this specific axis of host-microbe interactions. The seemingly conflicting results of prior studies and our findings may, in part, reflect differences in the regulation and expression of secreted factors like Bfp1, which could be influenced by presently unknown molecular cues or environmental stimuli. Furthermore, B. fragilis may have evolved Bfp1 to activate PAR₂-signaling and create an ecological niche conducive to its survival. Conversely, other PAR₂-processing activities identified in our proteolysis screen might exert opposing effects to those of Bfp1, potentially deactivating PAR₂-signaling as a strategy to suppress inflammatory conditions or to counteract dominance of a species such as B. fragilis. PAR₂ activation can induce both pathologic and protective effects, which may also account for the detrimental and beneficial actions of B. fragilis.²⁰

While there are a number of animal models of IBD, current advances in the use of intestinal organoids enable establishment of highly controllable cell models that mimic the interactions between microbes and human gut tissue. Previous work using classical organoids in which B. fragilis supernatants were added basolaterally suggested that B. fragilis does not impact barrier integrity.⁷⁰ In contrast, we find that physiologically-relevant apical treatment of organoids with supernatant containing Bfp1 disrupts barrier function to a similar extent as an optimized PAR₂ agonist or trypsin, a protease known to activate PAR₂ (Figure 4). This result demonstrates the value of the apical-out organoid model³¹ and suggests that the prior inability to detect an impact of B. fragilis on barrier function may be attributed to the inability of secreted proteases to reach the epithelial apical side. A loss of barrier function triggered by Bfp1 and PAR₂ would be expected to induce an influx of bacteria and macromolecules from the colon lumen, leading to colonic inflammation and pain.

Our findings that Bfp1 increases excitability of isolated DRG nociceptors, sensitizes colonic nociceptors to mechanical stimuli, and evokes abdominal mechanical allodynia and pain-like behavior in mice through mechanisms largely dependent on PAR₂ are in line with the known actions of PAR₂ in the colon. PAR₂ activation modulates the function of several ion channels involved in DRG neuronal activation. Excitatory voltage-gated Na⁺ currents⁴² and transient receptor potential V1 and V4 currents^73,74 are augmented by PAR₂ activation in DRG neurons whereas inhibitory voltage-gated K+ currents are suppressed.⁷⁵ PAR₂ agonists, including proteases and peptide analogs of the tethered ligand, sensitize transient receptor potential ion channels⁷⁴ and induce hyperexcitability³³ of nociceptors. Moreover, proteases from macrophages, mast cells, and from colon biopsies of IBS patients induce visceral nociception upon intracolonic injection by a PAR₂-mediated mechanism.^{3,30,33,46,76} PAR₂ agonists also cause a loss of barrier function in the colon^30,45 and stimulate the release of neuropeptides from colonic nociceptors,^44,77 which would lead to inflammation and ensuing pain. Activation of PAR₂ in nociceptors and colonocytes would be expected to cause pain in mice. However, our results that selective deletion of PAR₂ from NaV1.8 nociceptors diminishes the pronociceptive actions of Bfp1 to a similar extent as global PAR₂ deletion, suggests that Bfp1 mostly evokes pain through nociceptor-PAR₂.

Collectively, our findings that Bfp1-positive culture supernatants consistently induce PAR₂-dependent phenotypes across organoid, neuronal, and mouse models, closely reflecting the established effects of PAR₂ signaling, provide strong evidence for a physiologically relevant axis of host-microbe interaction. By identifying Bfp1 as a molecular effector of intestinal pain, barrier function and inflammation, this study establishes a crucial framework to investigate how microbial proteases drive pathogenesis. Future research will be required to build upon these findings to further elucidate the complex roles of microbial proteases and explore their potential as therapeutic targets for restoring intestinal homeostasis and mitigating inflammatory and painful diseases.

RESOURCE AVAILABILITY

Lead contact

Further information and requests for resources should be directed to and will be fulfilled by the lead contact, Matthew Bogyo (mbogyo@stanford.edu).

Materials availability

All cell lines, chemical probes and reagents generated in-house for this study will be made available subject to a materials transfer agreement with Stanford University or New York University.

Data and code availability

Proteomics data have been deposited at https://www.ebi.ac.uk/pride/ under the identifier PXD059166 and are publicly available as of the date of publication. Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

STAR★METHODS

EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

Animals.

The following strains of mice were used: C57BL/6J wild-type mice (#000664 JAX^®); knock-in mice expressing PAR₂ fused to monomeric ultrastable GFP (Par₂-mugfp); Par₂^−/− global knockout mice (#0004993 JAX^®); mice with PAR₂ deleted in Nav-1.8+ve nociceptors (Par₂^−/−-Nav1.8) and PAR₂-Cre control mice (Par₂^+/+-Cre). Par₂^−/−-Nav1.8 mice were previously obtained by breeding mice expressing Par₂ flanked by LoxP sites (Par₂ lox/lox) with mice expressing Cre recombinase targeted to nociceptors using the NaV1.8 promoter (Scn10a).³³ Par₂^−/−-NaV1.8 mice lacked immunoreactive PAR₂ in NaV1.8+ neurons of the dorsal root ganglia (DRG). Male and female mice (8–12 weeks old; 20–25 g) were used. Data were analyzed for sex-based differences. No significant differences were observed between male and female, and data were therefore pooled for final presentation, or male mice only were used. Mice were group housed (5 × cage), with the exception for the one that underwent telemetry implantation; mice were single-housed. Mouse randomization for the different experimental groups was done using Research Randomizer (https://matheusararipe.github.io/mouse-randomization-js/mouse_ultima_versao.html). All animals were maintained in a light-controlled (12-h light/dark cycle) and temperature-controlled (22 ± 4 °C) environment with ad libitum access to food and water. The New York University and Queen’s University Animal Ethics Committees approved all experimental protocols, which conformed to The American and Canadian Council of Animal Care guidelines.

Primary cells and cell lines.

Human embryonic kidney (HEK) 293 cells (female-derived) were maintained in culture Dulbecco’s Modified Eagle Medium (DMEM) + 10% fetal bovine serum (FBS) + penicillin/streptomycin, 50 IU/mL at 37°C in 5% CO₂. The de-identified human ileal organoids from healthy tissue used in this study were received from an organoid biobank established by Dr. Calvin Kuo and grown as specified in Methods details.

Bacterial strain library.

Bacterial strain library was received from Dr. Justin Sonnenburg as frozen glycerol stocks in 96-well plates and was stored at −80 °C before use. Individual bacterial stocks were prepared as glycerol stocks under anaerobic conditions in sealed glass tubes with a septum and stored at −80 °C.

Bacterial culture media and growth conditions.

Liquid handling and cultivations for all strains (except for E. coli strains for molecular cloning and recombinant protein expression) were performed inside an anaerobic chamber (Coy Lab Products) with an atmosphere of 5% H₂, 5% CO_2, and 90% N₂. Unless stated otherwise, bacteria were grown in Mega Medium (see Supplementary Table 8) at 37°C.¹⁷ Identity of the strains was verified by 16S rRNA gene sequencing, as previously described.¹⁷

METHOD DETAILS

Chemical synthesis of Cys-PEG₄-Cy5.

Synthesis of I-1:

N-(tert butoxycarbonyl)-S-trityl-L-cysteine (160 mg, 0.35 mmol, 1.0 eq) and HBTU (144.0 mg, 0.38 mmol, 1.1 eq) were dissolved in DMF (3 mL). (2-[2-[2-[2-(2-Azidoethoxy)ethoxy]ethoxy]ethoxy]ethanamine (99.6 mg, 0.38 mmol, 1.1 eq) and DIPEA (89.2 mg, 0.69 mmol, 2.0 eq) were added and the solution was stirred at rt for 15 h. The solvent was removed under reduced pressure and the residue was redissolved in EtOAc. The organic layer was washed with 1 M citric acid, 5% NaHCO₃ and brine and the solvent was removed under reduced pressure. The residue was dissolved in MeOH (4mL) and Pd/C (10%, 30 mg, 0.28 mmol) was added. Reduction was performed for 17 h using a hydrogen-filled balloon. The MeOH was removed under reduced pressure, the residue resuspended in EtOAc, filtered over celite and the solvent was removed under reduced pressure. Purification by preparative HPLC (A: H₂O + 0.1% TFA, B: MeCN + 0.1% TFA, gradient: 5 to 95% B in 17 min) and lyophilization yielded the product as a white solid (205 mg, 0.26 mmol, 74%).

Synthesis of Cys-PEG₄-Cy5:

Sulfo-Cyanine5 (“Cy5”, 15 mg, 23 μmol, 1.0 eq) and HBTU (10 mg, 27 μmol, 1.2 eq) were dissolved in DMF (1 mL) and I-1 (18 mg, 23 μmol, 1.0 eq) and DIPEA (8.7 mg, 68 μmol, 3 eq) were added. The reaction was stirred overnight. Next, DCM (0.8 mL), TIPS (12 μL) and TFA (0.2 mL) were added and the reaction was stirred at rt for 60 min. The solvents were removed under reduced pressure and the residue was purified by preparative HPLC (A: H₂O + 0.1% TFA, B: MeCN + 0.1% TFA, gradient: 5 to 95% B in 17 min), yielding the product as a blue powder after lyophilization (10 mg, 4.8 μmol, 20%).

Generation of His-PAR₂-NTD-Cy5 peptide by expressed protein ligation.

Design:

The PAR₂-NTD-peptide, consisting of the 46 amino acids spanning human PAR₂-NTD and a N-terminal hexahistidine (His₆)-sequence, was fused to the intein Mxe GyrA. To improve efficiency of native chemical ligation and protein quantification, a two amino acid linker sequence (WG) was introduced before the GyrA sequence. Nucleotide and protein sequences can be found in Supplementary Table S5.

Recombinant protein expression and purification of His-PAR₂NTD-GyrA:

Plasmid pET28a_His-PAR₂NTD-GyrA was transformed into chemically competent E. coli Rosetta2(DE3) cells with selection using 50 μg ml⁻¹ kanamycin and 34 μg ml⁻¹ chloramphenicol for expression. For recombinant protein expression, an overnight culture of transformed cells was inoculated into 1 L of LB supplemented with 50 μg ml⁻¹ kanamycin and 34 μg ml⁻¹ chloramphenicol and grown with shaking until the optical density at 600 nm (OD₆₀₀) = 0.6. After the culture was cold-shocked on ice for 20 min, 0.5 mM isopropyl β-D-1-thiogalactopyranoside (IPTG) was added to the culture to induce protein expression, and the culture was incubated for 20 h at 18 °C. The bacterial pellet was flash-frozen in liquid nitrogen and stored at −80 °C. The pellet was lysed in 30 ml of lysis buffer (50 mM N-2-hydroxyethylpiperazine-N-2-ethane sulfonic acid (HEPES), 10 mM imidazole, 150 mM NaCl, pH 8) supplemented with 750 mM trehalose by probe sonication on ice (3 min at 30% power in 1 s bursts; 1 min at 60% power in 1 s bursts; 3 min at 30% power in 1 s bursts and 1 min at 60% power in 1 s bursts). Cell debris was pelleted at 38,400 × g for 20 min at 4 °C, and the lysate was transferred to a new tube and centrifuged at 12,000 × g for 15 min at 4 °C. The lysate was then clarified sequentially through a 5-μm and a 1-μm filter before purification. The clarified lysate was injected into an ÄKTA purifier FPLC system (GE Healthcare) and separated with a 5 mL HisTrap FF crude column (GE Healthcare). The column was subsequently washed with lysis buffer supplemented with 10 mM imidazole, 20 mM imidazole, and 40 mM imidazole. The protein was eluted with lysis buffer supplemented with 300 mM imidazole, and fractions containing protein (as determined by the ultraviolet (UV) trace and SDS–PAGE) were combined. The imidazole-concentration of the final protein sample was reduced to ca. 30 mM by consecutive buffer-exchange with buffer (200 mM HEPES, 150 mM NaCl, pH 8).

Native chemical ligation:

On-resin native chemical ligation was used to C-terminally conjugate the His₆-PAR₂-peptide with Cys-PEG₄-Cy5. To do so, 1.2 mL of 300 μM His₆-PAR₂-GyrA (360 nmol) in thioesterification buffer (200 mM HEPS, 150 mM NaCl, pH 8) was added to 2.4 mL of 150 mM sodium 2-mercaptoethanesulfonate (Mesna) in freshly degassed thioesterification buffer and incubated at 4°C for 18 h while gently rocking. The sample was diluted with 1.8 mL of pulldown-buffer (200 mM HEPES, 20 mM NaCl, 20 mM imidazole, pH 7.2) and 300 μL of pre-washed Ni-NTA agarose beads (Qiagen) was added. Pulldown of His₆-PAR₂-Mesna was performed at 4°C for 2 h while gently rocking. The supernatant containing cleaved GyrA was removed, and the beads were washed six times with 500 μL of pulldown-buffer. For native chemical ligation, beads were resuspended in freshly degassed NCL-Buffer (200 mM HEPES, 20 mM NaCl, 20 mM Tris(2-carboxyethyl)phosphine (TCEP), 100 mM 4-Mercaptophenylacetic acid (MPAA), pH 7.4), 18 μL of 50 mM Cys-PEG₄-Cy5 (2.5 eq. based on original His₆-PAR₂-Mesna) was added. The reaction was incubated under an argon atmosphere at 4 °C for 16 h while gently rocking. The supernatant was removed and beads were washed six times with 300 μL of wash-buffer-1 (200 mM HEPES, 20 mM NaCl, 1 mM TCEP, pH 7.2). For alkylation, beads were resuspended in 300 μL of freshly made alkylation-buffer (200 mM HEPES, 20 mM NaCl, 1 mM TCEP, 5 μM iodoacetamide) and incubated at rt in the dark for 90 min. The supernatant was removed and beads were washed three times with 500 μL of wash-buffer-2 (200 mM HEPES, 20 mM NaCl, pH = 7.2) and three times with 500 μL of ultrapure water. His₆-PAR₂NTD-Cy5 peptide was eluted from the Ni-NTA beads by treatment with four times 150 μL of 0.5% TFA in ultrapure water. The peptide was purified by preparative HPLC (column: Aeris 3.6 μm Widepore C4 200, 250 × 4.6 mm; solvents: A: 0.1% TFA in water, B: 0.1% TFA in MeCN; gradient: 5% B to 80% B in 30 min) and lyophilized. The final peptide-TFA salt (1.13 mg; 75 nmol, 21% based on Cy5-absorbance) was dissolved in 0.05% formic acid in ultrapure water and its concentration was determined by Cy5 absorbance (e = 271,000 M⁻¹cm⁻¹ at 649 nm). Identity and purity of the His-PAR₂-NTD-Cy5 peptide was verified by HPLC-HRMS (EM_calcd.: = 6950.39 Da, EM_measured: = 6950.38 Da).

Preparation of Mega Medium.

The preparation of Mega Medium was performed as previously published¹⁷ and the recipe is outlined in Supplementary Table 8. Prior to preparation of the medium itself, we prepared the following stock solutions:

1 M potassium phosphate buffer, pH 7.2:

We first prepared 1 M KH₂PO₄ (monobasic) - 68.045 g KH₂PO₄ (anhydrous) in Milli-Q water to 500 mL. We then proceeded to prepare 1 M K₂HPO₄ (dibasic) - 174.18 g K₂HPO₄ (anhydrous) in Milli-Q water to 1 L. Both potassium phosphate powders should be added to actively stirred water to dissolve them well. Finally, added monobasic to dibasic to achieve pH 7.2 (typically this required ~430 mL monobasic added to 1 L dibasic).

Vitamin K solution:

We dissolved 40 mg menadione (Vitamin K3, Sigma M5625) in 40 mL 100% EtOH.

TYG salts solution:

0.5 g of MgSO₄·7H₂O (Sigma 230391), 10.0 g of NaHCO₃ (Sigma S5761) and 2.0 g of NaCl (Sigma S7653) were dissolved in Milli-Q water to 1 L final volume.

FeSO₄·7H2O (0.4 mg/mL):

40 mg FeSO₄·7H₂O (Sigma F8633) was dissolved in 100 mL Milli-Q water.

0.8% (w/v) CaCl₂:

0.4 g CaCl₂·2H2O (Sigma C7902) was dissolved in 50 mL Milli-Q water.

Resazurin anaerobic indicator (0.25 mg/mL):

25 mg resazurin (Sigma R2127) was dissolved in 100 mL distilled H2O and stored protected from light at 4 °C.

Histidine-Hematin solution:

0.2 M histidine, pH 8.0, was prepared by mixing 4.2 g histidine-HCl monohydrate (Sigma H7875) in 80 mL Milli-Q water. pH was adjusted to 8 with 10 N NaOH (the histidine dissolved as the pH went up) and volume was adjusted to 100 mL with Milli-Q water. In a second step, 12 mg hematin (Sigma H3281) was dissolved in 10 mL of 0.2 M histidine, pH 8.0 through vigorous shaking for several hours and filter-sterilized using a 0.2 μm filter.

SCFA supplement:

Prepared by mixing 17 mL of glacial acetic acid (Sigma A6283), 6 mL of propionic acid (Sigma P5561), 4 mL butyric acid (Sigma B103500) and 1 mL of isovaleric acid (Sigma 129542).

Final preparation of Mega Medium:

Briefly, we added all ingredients of the Base solution (Supplementary Table 8) and mixed well. pH was adjusted to ~7.0, in practice, this took slightly less than 2.5 mL 10 M KOH per 500 mL media. After pH measurement, we added 3.6 – X mL water where X is mL of KOH added. Mega Medium was autoclaved (121°C for 25 minutes at 20 PSI, 20 minutes dry). After autoclaving we added vitamins and supplements (Supplement Solution) into the cooled medium as specified in Supplementary Table 8.

PAR₂-proteolysis screening.

Bacterial cultivation and supernatant generation:

Microbes were cultivated in 96-well plates with every well corresponding to an individual bacterial strain, as previously described.¹⁷ Starter cultures (1 mL scale) were grown for 24 h at 37 °C without shaking. Growth was verified by OD₆₀₀ measurement. For main cultures, Mega Medium (1 mL) was inoculated from overnight cultures (1:50) and cultivated for 24 h without shaking. Growth was monitored in parallel in a 96-well plate format via OD₆₀₀-measurements using a Cerillo^® Stratus plate-reader mounted on a plate shaker (200 rpm). Culture plates were taken outside the anaerobic chamber, centrifuged (4500 × g, 4°C, 15 min) and supernatants were sterile filtered using 0.22 μm filter plates. The cell-free supernatants were aliquoted, snap-frozen and stored at −80 °C until used for activity assays.

Proteolysis and affinity-pulldown:

Supernatants (9 μL) and His-PAR₂-NTD-Cy5 substrate (1 μL of 20 μM in 0.01% FA) were incubated for 3 h at 37 °C. Next, samples were cooled in a cold-block and centrifuged. For affinity-pulldown, Ni-NTA magnetic beads (PureCube^®) were washed in pulldown-buffer (200 mM HEPES, 0.025 % Tween-80, pH = 8.8) and 1 μL of original bead suspension in 16 μL of pulldown-buffer were added to each sample. Pulldown was performed for 2 h at 4 °C while shaking. Samples were centrifuged, beads were sedimented using a magnetic rack and 10 μL of the supernatant was transferred to a small-volume 384 well-plate (Greiner). Fluorescence of the proteolytically cleaved peptide fragments was determined using a Cytation3 plate-reader (excitation: 640 nm, emission: 670 nm). For each sample, PAR₂-processing activity was normalized to trypsin (100% activity) and Mega Medium-background (0% activity). The screening results were plotted onto a phylogenetic tree (based on 16S rRNA sequences) of the strain tested via iTOL (https://itol.embl.de).

Validation of PAR₂-processing activity and inhibition by covalent inhibitors.

To validate the PAR₂-processing activity, verified strains (via 16S rRNA sequencing) with more than 50% PAR₂-processing activity in the initial screen were cultivated individually. To do so, 10 mL Mega Medium was inoculated with 0.1 mL of an overnight-culture and cultivated at 37 °C without shaking. Growth was monitored in parallel in a 96-well plate format via OD₆₀₀-measurements using a Cerillo^® Stratus plate-reader mounted on a plate shaker (200 rpm). When a culture reached early stationary phase, the culture tube was centrifuged (4500 × g, 4°C, 15 min) and supernatants were sterile filtered using a 0.22 μm PES syringe filter. Supernatants were then subjected to activity-assays. For each assay, 9 μL of each supernatant was pre-incubated with 0.2 μL DMSO (control) or covalent inhibitor (50x stock concentration) at 37 °C for 1h. Samples were centrifuged and 1 μL of His-PAR₂-NTD-Cy5 substrate (20 μM in 0.01% FA) was added to each well. Proteolysis was performed for 1.5 h (instead of 3 h for the initial screen) at 37°C. Affinity-pulldown and fluorescence measurements were performed as described above. For each sample, PAR₂-processing activity was normalized to trypsin (100% activity) and Mega Medium-background (0% activity). Experiments were performed in duplicates (n = 2).

Gel-based ABPP.

B. fragilis NCTC 9343 culture supernatants were harvested at early stationary phase (ca. 9h cultivation) via centrifugation (5000 × g, 15 min, 4 °C) and sterile-filtration (0.22 μm PES sterile filter). Supernatants were then concentrated 5-fold using 10 kDa MW-cutoff-filters. For labeling, 60 μL of 5-fold supernatants were incubated with 0.6 μL of DMSO or MLP7 (0.5 mM, 2 mM, or 10 mM) for 1 h at 37 °C. Subsequently, samples were treated with 0.6 μL of DMSO or FP-TMR (0.1 mM or 1 mM) for 1 h at 37 °C. Samples were precipitated by chloroform-methanol precipitation. In brief, each sample was treated with methanol (240 μL), chloroform (60 μL), and water (180 μL) with mixing after each addition step. After centrifugation (18000 × g, 5 min, rt) the upper layer was carefully removed. Methanol (180 μL) was added and the sample was mixed and centrifuged (18000 × g, 5 min, rt) again. The full supernatant was removed and the remaining pellet dissolved in 1x Laemmli-Buffer. Labeled proteins were separated via SDS–PAGE. In-gel fluorescence was visualized using the Cy3 channel on Typhoon 9410 Imager (Amersham Biosciences).

MS-based ABPP sample preparation and analysis.

B. fragilis NCTC 9343 cultures were grown in 110 mL of Mega Medium in triplicates for 9 h until early-stationary phase. Cultures were centrifuged (5000 × g, 15 min, 4 °C) to pellet bacterial cells and supernatants were filtered using a 0.22 μm PES sterile filter. 100 mL of the supernatants were concentrated to a final volume of 20 mL using a 10 kDa MW-cutoff concentrator. For each biological replicate, the supernatant was aliquoted into three separate tubes (6 mL each), representing the three labeling conditions “DMSO”, “Probe” and “Competition”. Supernatants were incubated with 60 μL DMSO (for “DMSO” and “Probe”) or 60 μL of 10 mM MLP7 (for “Competition”) at 37 °C for 1 h. Subsequently, samples were treated with 60 μL of DMSO (for “DMSO”) or 60 μL of 1 mM FP-Biotin (for “Probe” and “Competition”) and incubated at 37 °C for 1 h. For precipitation, 606 μL of 100% trichloroacetic acid (TCA) solution was added to each sample, incubated on ice for 30 min, and centrifuged (12,000 × g, 4 °C for 15 min). The supernatants were removed, the pellets were resuspended in a mix of 0.5 mL PBS and 4.5 mL acetone and centrifuged (12,000 × g, 4 °C for 15 min). The supernatants were removed, and the pellets washed again with a mix of 0.375 mL PBS and 3.4 mL acetone. Protein pellets were solubilized in 0.5 ml PBS containing 0.4% (w/v) SDS and transferred to Lo-bind Eppendorf tubes containing 50 μL of avidin-agarose bead slurry (Sigma Aldrich) pre-equilibrated with 0.4% SDS (w/v) in PBS (3 × 1 mL, 400 × g, 5 min, rt), and rotated for 1 h at room temperature. The beads were then washed with 0.4% (w/v) SDS in PBS (3 × 1 mL), 6 M urea (2 × 1 mL) and finally PBS (3 × 1 mL). Beads were resuspended in 200 μL X-buffer (7 M urea, 2 M thiourea in 20mM HEPES buffer, pH 7.5). Upon reduction with 5 mM TCEP (2 μL of 500 mM stock in ddH₂O) for 1 h at 37°C, proteins were alkylated using 10 mM IAA (4 μL of 500 mM stock in ddH₂O) for 30 min at 25°C and samples were quenched with 10 mM DTT (4 μL of 500 mM stock in ddH₂O) for 30 min at 25°C. Enzymatic digestion using LysC (1 μL of 0.5 μg/μL, Wako, MS-grade) was first carried out for 2 h at 25°C, upon which samples were diluted with triethylammonium bicarbonate (TEAB) buffer (600 μL of 50 mM stock in ddH₂O) and digested with trypsin (1.5 μL of 0.5 μg/μL in 50 mM acetic acid, Promega, sequencing grade) for a further 16 h at 37°C. Samples were acidified to 1% (v/v) FA and desalted using SepPak^® C18 cartridges (50 mg, Waters) with a vacuum manifold. The cartridges were first washed with ACN (2 × 1 ml) and equilibrated with 0.1% (v/v) TFA (3 × 1 mL) prior to loading the samples. After washing with 0.1% (v/v) TFA (3 × 1 mL) and 0.5% (v/v) FA (1 × 0.5 mL), peptides were eluted in 80% (v/v) ACN containing 0.5% FA (3 × 0.25 mL) and freeze-dried using a speedvac centrifuge. For tandem-mass-tag (TMT)-labeling, samples were redissolved in 23 μL of 100 mM HEPES-Buffer (pH = 8.5) and treated with 7.5 μL of respective TMT-reagent (Thermo Scientific 10-plex kit, each label was dissolved to 13 μg/μL in anhydrous acetonitrile) and incubated (25 °C, 500 rpm) for 1 h. The reactions were stopped by adding 0.3 μL of 50% hydroxylamine and incubating it (25 °C, 500 rpm) for 15 min. Samples were freeze-dried using a speedvac centrifuge, desalted using C18-tips (Pierce^® C18 tips, 100 μL, Thermo Scientific) according to the manufacturer’s protocol and freeze-dried using a speedvac centrifuge. For measurement, samples were dissolved in 20 μL of 1% formic acid in water. MS analysis was performed using an Orbitrap Eclipse Tribrid mass spectrometer (Thermo Scientific, San Jose, CA, USA) with liquid chromatography using an Acquity M-Class UPLC (Waters Corporation, Milford, MA, USA). A flow rate of 300 nL/min was used, where mobile phase A was 0.2% formic acid in water and mobile phase B was 0.2% formic acid in acetonitrile. Analytical columns were prepared in-house with an I.D. of 100 microns pulled to a nanospray emitter using a P2000 laser puller (Sutter Instrument, Novato, CA, USA). The column was packed using C18 reprosil Pur 1.8-micron stationary phase (Dr. Maisch) to a length of ~25 cm. Peptides were directly injected onto the analytical column using a gradient (2%–45% B, followed by a high-B wash) of 180 min. MS data were acquired using an MS3 data-dependent acquisition method. MS1 profile scans were acquired in the Orbitrap (resolution, 120,000; scan range, 400–1600 m/z ; AGC target, 4 × 10⁵ and maximum injection time, 50 ms). Monoisotopic peak determination was set to ‘peptide’. Only charge states 2 – 6 were included. Dynamic exclusion was enabled (repeat count, n = 1; exclusion duration, 30 s; mass tolerance: low, 10 ppm and high, 10 ppm, excluding isotopes). An intensity threshold of 5 × 10³ was set. Data-dependent MS2 spectra were acquired in centroid mode across a mass range of 400–1,600 m/z. Precursor ions were isolated using the quadrupole (isolation window, 0.7 m/z), fragmented using collision-induced dissociation (CID) (collision energy, 35%; activation time, 10 ms and activation Q, 0.25) and detected in the ion trap (scan range mode, auto m/z normal; scan rate, turbo; AGC target, 1 × 10⁴ and maximum injection time, 35 ms). Data-dependent MS3 spectra were acquired in the Orbitrap (resolution, 50,000; scan range, 100–500 m/z; AGC target, 1 × 10⁵and maximum injection time, 200 ms) following higher-energy collisional dissociation (HCD) activation (collision energy, 55%) using Synchronous Precursor Selection from up to 10 precursors. Peptide and protein identifications were performed using MaxQuant (version 2.3.0.3)⁷⁹ with Andromeda as the search engine. Group-specific parameters were set to ‘Reporter ion MS3’ with 10plex TMT isobaric labels for N-terminal and lysine residue modification selected. Reporter mass tolerance was set to 0.003 Da. The following parameters were used: carbamidomethylation of cysteines as fixed modifications, oxidation of methionine and acetylation of N-terminus as dynamic modifications, and trypsin/P as the proteolytic enzyme. Default settings were used for all other parameters. Searches were performed against the UniProt database for B. fragilis NCTC 9343 (proteome ID: UP000006731, downloaded on 11.02.2022.) with the following parameters: minimum peptide length, 6; maximum peptide mass, 6,000 Da; minimum peptide length for unspecific search, 6 and maximum peptide length for unspecific search, 40. Identification was performed with at least two unique peptides and quantification only with unique peptides. Statistical analyses were performed with Perseus v.2.0.3.0.⁸⁰ Putative contaminants, reverse hits and proteins identified by side only were removed. Label-free quantitation intensities were log₂-transformed. Missing values were imputed using a normal distribution (width, 0.3; down-shift, 1.8). P values were calculated using a two-sided, two-sample t-test. Data are available under: https://www.ebi.ac.uk/pride/ (Project accession: PXD059166, Token: ceCR6vYkusLh).

BLAST-N analysis.

BLAST-N analysis (Version BLASTN 2.16.0+ ) against 1,520 reference genomes from cultivated human gut bacteria (Bioproject ID 482748)²⁶ was performed at https://blast.ncbi.nlm.nih.gov/Blast.cgi⁸¹ using the nucleotide sequences of Bfp1 (BF9343_2070) and TspA (BF9343_0342) as input with the following parameters: Program selection: discontiguous megablast, Max. target sequences: 500. All other parameters were set to default. The phylogenetic tree for the Blast results was generated with phyloT software (https://phylot.biobyte.de), using the full genomes of 216 species in which the full 1520-member database was clustered, and plotted via iTol (https://itol.embl.de).

Clean genetic deletion of the bfp1 and tspA genes.

The two clean deletion strains B. fragilis NCTC 9343 Δbfp1 and ΔtspA were generated via two-step allelic exchange based on the vector pLGB30 according to the protocol from Garcia-Bayona et al.²⁷ In brief, the plasmids pLGB30-bfp1 and pLGB30-tspA were generated by PCR according to Supplementary Table S6. Each plasmid was conjugated in B. fragilis NCTC 9343 using E. coli S17–1 λpir as the conjugative donor strain. Exconjugants with chromosomally integrated plasmids were recovered on BHIS plates containing 200 μg/mL gentamycin and 6 μg/mL tetracycline. Second crossover events were selected using BHIS plates containing 10 mM L-rhamnose. Deletion of the target genes was confirmed by PCR using suitable primers (see Supplementary Table S7).

Human intestinal organoids.

Human organoid line HC921 established in the Calvin Kuo lab (Stanford University) from healthy ileal tissue was cultured in IntestiCult Organoid Growth Media (Stemcell) supplemented with 10 μM Y27632 (MedChem Express, HY-10583) and 2.5 μM CHIR 99021 (Tocris, 102875–390). TrypLE Express Enzyme (Thermo Fisher Scientific, 12604013) was used to passage organoids before embedding the cells in Matrigel (Corning, 354234) and growing organoids in a 24-well plate with 400 μl of growth medium usually with 1:5 to 1:8 ratio every 7–10 days.

Polarity reversal and FITC-dextran assay.

To achieve polarity reversal and obtain apical-out organoids, we followed a protocol by Co et al.³¹ and confirmed the polarity reversal using Phalloidin-647 (Cell Signaling Technologies, 8940S). Briefly, organoids 5–7 days after passage were harvested and Matrigel was removed using 5 mM EDTA, 4 °C, 30 minutes. Organoids were washed with AdMEM/F-12 (Thermo Fisher Scientific, 12634010) and then incubated in IntestiCult growth medium for 2 days in ultra-low attachment plates (Corning, 3473) pre-treated with Anti-Adherence Rinsing Solution (Stemcell Technologies, 07010).

Flipped organoids were transferred into 50 μl of phenol-red free DMEM/F-12 (Thermo Fisher Scientific, 21041025) and 10 μl of concentrated bacterial supernatants from WT B. fragilis, B. fragilis Δbfp1-knockout, Mega Medium or 5 mM EDTA were added. Organoids were incubated for 30 minutes at 37 °C before adding 40 μl of 2 mg/ml FITC-dextran (Sigma-Aldrich, 46944–100MG-F) and incubated for 15 minutes. Organoids were then centrifuged for 2 min, 250xg, the supernatant was removed, and organoids were resuspended in the remaining liquid. 5 μl of the sample was transferred onto a microscopy slide and a chamber was created using vacuum grease and a coverslip to prevent mechanical stress. Organoids were then imaged on confocal microscopy. Samples were processed sequentially to ensure precise timing. MFIs were calculated using Zen software (Zeiss). MFI for each organoid was normalized to background for the particular image and values were plotted using GraphPad Prism 10.

Organoid-derived monolayers and transepithelial resistance measurement.

Organoid-derived monolayers were prepared according to Stemcell’s protocol with minor alterations.⁸² Briefly, Transwells (Corning, 3413) were pre-coated with Matrigel. Organoids were harvested with TrypLE and then trypsin to ensure dissociation into single cells. Cells were seeded at 75,000 cells/well in 100 μl of the recommended medium. Once monolayers were formed (3–4 days), cells were treated apically with the following solutions: 50 μl phenol-red free DMEM/F-12, 10 μl 40x concentrated bacterial supernatants or Mega Medium (final 4x concentrated), PAR₂-agonistic peptide 2-furoyl-LIGRLO-amide in Mega Medium (final 1 μM), or purified Trypsin in Mega Medium (final 4 nM) and 40 μl of FITC-dextran (2mg/ml). At selected time points, transepithelial resistance was measured using EVOM2. Measured values were normalized by subtracting the background (coated well without cells treated apically with Mega Medium) and calculating the resistance per cm².

Cell lines.

Human embryonic kidney (HEK) 293 cells (female-derived), which endogenously express PAR₂, were maintained in culture medium (DMEM + 10% fetal bovine serum (FBS) + penicillin/streptomycin, 50 IU/ml) at 37°C in 5% CO₂. To make a cell line expressing fluorescently tagged PAR₂, cDNA encoding human PAR₂ with N-terminal HA tag and C-terminal mApple fluorescent tag was designed and purchased from Twist Bioscience. HEK293 cells were transfected with 1 μg of the pTwist_CMV_Hygro-HA-PAR₂-mApple plasmid using PEI (Invitrogen) with a DNA:PEI ratio of 1:6 and cultured for 48 h. Single cells were suspended, plated in 96 well plates, and HA-PAR₂-mApple expressing cells were selected in culture medium + hygromycin (200 μg/ml). mApple fluorescence was confirmed by microscopy. A single colony with high and uniform expression of HA-PAR₂-mApple was selected, expanded for further experimentation, and maintained in culture medium with hygromycin (100 μg/ml). HEK293 cells in which PAR₂ was deleted via CRISPR/Cas genome editing (HEK-PAR₂-KO) have been described previously.⁸³

Cellular PAR₂ cleavage assays.

HEK-HA-PAR₂-mApple cells (30,000 cells) were plated on poly-D-lysine-coated 12 mm round glass coverslips in a 24 well plate and incubated overnight. Cells were washed and incubated in HBSS-H for 30 min. Cells were untreated or incubated with Mega Medium (10x diluted, control), trypsin (10 nM, positive control), WT B. fragilis or B. fragilis Δbfp1 supernatant (10x diluted), or purified Bfp1 (500 nM) for 30 min at 37°C. Cells were washed 1x in ice-cold HBSS-H, fixed in 4% paraformaldehyde on ice for 20 min and washed with PBS. Cells were incubated in blocking buffer (PBS + 3% normal horse serum + 0.3% saponin, pH 7.4) for 1 h at room temperature (RT), followed by rat anti-HA (1:1000) in PBS at 4°C overnight. Cells were washed 3x in PBS and incubated with donkey anti-rat-AlexaFlour488 (1:1000, ThermoFisher) for 1 h at RT. Cells were then washed, incubated with DAPI (1 μM, 5 min), re-washed, and mounted with ProLong^® Gold Antifade Mountant (ThermoFisher). Cells were imaged on an inverted Leica SP8 confocal microscope with a 63x objective (1.4 NA). Images were processed with ImageJ (NIH) and figures were made with Adobe Illustrator.

In vitro quantification of HA-PAR₂-mApple localization.

In ImageJ, the mApple channel was saturated and the researcher designated the ROI by tracing the outer edge of each cell. The ROIs were converted to a mask which was then eroded by 5 pixels (0.9 μm) to create the cytosolic mask. The cytosolic mask was subtracted from the original mask to get a band that was 5 pixels (0.9 μm) wide corresponding to the outer edge of the cell. That band was designated the membrane mask. The membrane mask and the cytosolic mask were then applied individually to the original PAR₂-mApple channel and PAR₂-mApple fluorescence was quantified along the plasma membrane (RFU_mem) and cytosolic (RFU_cyto) regions of the cell. PAR₂-mApple internalization was presented as a ratio of RFU_cyto / RFU_mem.

Cellular PAR2 activation assays.

Fluo-4 fluorescence was imaged using a Leica DMi8 wide field microscope (excitation 470±20 nm, emission 527±15 nm) with a 10x objective (0.3 NA). Baseline fluorescence was measured (F0), the cells were challenged with bacterial supernatants (10x), trypsin (10 nM), Mega Medium (control), or the purified protease Bfp1 (500 nM). Changes in fluorescence, which are proportional to [Ca²⁺]_i were recorded (F). [Ca²⁺]_i responses were quantified using ImageJ (NIH). To account for the high levels of fluorescence of the Mega Medium, a cell free region of interest for each experiment was also quantified (Background fluorescence, FB), and [Ca²⁺]_i was normalized to baseline and background fluorescence (ΔF= (F - F0) - FB).

Patch clamp electrophysiology.

Dorsal root ganglia (DRG) (L1-L5) from C57BL/6J wild-type mice were incubated in collagenase (1 mg/ml, Sigma-Aldrich) and dispase (1 mg/ml, Sigma-Aldrich) in minimal essential medium (Invitrogen) with 10% fetal bovine serum (FBS) for 30 min at 37℃ in a humidified atmosphere (95% air, 5% CO₂). Neurons were mechanically dispersed by trituration through a fire-polished Pasteur pipette. Neurons were resuspended in Neurobasal Plus Medium with B27 supplement (Invitrogen), 10% FBS and penicillin-streptomycin. Neurons were plated onto poly-D-lysine (0.05 mg/ml, Invitrogen) coated glass coverslips and maintained at 37℃ in a humidified atmosphere (95% air, 5% CO₂). The culture medium was replaced with serum-free Neurobasal Plus medium 2 h after plating. Patch clamp recordings were made from small diameter (≤ 25 μm) DRG neurons within 30 h of plating. Perforated patch clamp recordings were made using a pipette solution containing amphotericin B (240 μg/ml, Thermo Scientific) in current clamp mode at RT. Recordings were made using an amplifier (Axopatch 200B, Molecular Devices) and an analog-digital converter (Digidata 1440A, Molecular Devices) controlled with a PC running pCLAMP 10 software. After a giga-ohm seal was established, neurons were equilibrated for 15 min to achieve membrane perforation and electrical access. Changes in excitability were quantified by measuring rheobase (minimum current required to elicit an action potential), determined by applying depolarizing current steps in 10 pA increments. A ramp current protocol (100 pA, 1 sec) was used to determine the frequency of action potentials firing. The recording chamber was continuously perfused with an external solution at 2 ml/min. Solutions had the following composition (mM): pipette solution: K-gluconate 110, KCl 30, HEPES 10, MgCl₂ 1, CaCl₂ 2 (pH 7.2, adjusted with KOH; 290 mOsm); external solution: NaCl 143, KCl 5, HEPES 10, glucose 10, MgCl₂ 1, CaCl₂ 2 (pH 7.4, adjusted with NaOH; 305 mOsm). Neurons were pre-incubated with bacterial supernatants, diluted to a final concentration of 0.5x or 1x in external solution, for 10 min at RT before measuring changes in excitability. The following bacterial samples were tested: Mega Medium, WT B. fragilis supernatant, B. fragilis Δbfp1 supernatant. To determine the requirement for protease activity, supernatants were preincubated with the serine protease inhibitor FP-alkyne (100 μM, 60 min, RT) or vehicle before incubation with neurons. To determine the contribution of PAR₂, neurons were pre-incubated with the PAR₂ antagonist AZ3451 (1 μM, 60 min, RT) or vehicle before exposure to supernatant, or neurons from Par₂^−/− global KO mice were studied.

Ex vivo assessment of alterations to colonic permeability.

Dissection and Ussing chamber experiments were conducted as previously described.⁸⁴ Briefly, mice were euthanized, and the colon was immediately excised and placed in ice-cold, oxygenated Krebs buffer. The mid-colon (2–4cm from the pelvic brim) was isolated, opened along the mesenteric border, and 0.2cm² of whole tissue was mounted onto cassettes (Physiologic Instruments, San Diego, CA) for Ussing chamber studies. Each cassette was exposed to 4 mL of oxygenated Krebs buffer (115 mM NaCl, 1.25 mM CaCl₂, 1.2 mM MgCl₂, 2.0 mM KH₂PO₄, 25 mM NaHCO₃, and 10 mM glucose, pH 7.4) maintained at 37°C. After a 30-minute equilibration period, 4-kDa FITC-labeled dextran (Sigma-Aldrich, 46944–100MG-F) was added to the apical compartment at a concentration of 2 mg/mL to assess macromolecular permeability. Simultaneously, 50 μL of 40x B. fragilis WT, Δbfp1, or MM was added to the apical side, with an equal volume of Krebs buffer removed prior to addition. Serosal samples were collected every 30 minutes over a 3-hour period, and each was replaced with fresh oxygenated Krebs buffer. FITC-dextran concentrations were measured using a fluorescence plate reader (excitation: 485 nm; emission: 525 nm; SpectraMax iD3 Multi-Mode Microplate Reader) and quantified against a standard curve. Data was then normalized to the surface area of the tissue, and results were expressed as μg/mL/cm².

Extracellular afferent nerve recording.

The distal colon of C57BL/6J wild-type mice, along with the intact neurovascular bundle of the inferior mesenteric artery, was removed and placed in a Sylgard-lined organ bath that was continuously superfused (10 mL/min) with oxygenated (5 % CO₂-95 % O₂) Krebs buffer solution (mM): 118.4 NaCl, 24.9 NaHCO₃, 1.2 MgSO₄, 1.2 KH₂PO₄, 11.7 glucose, and 1.9 CaCl₂, pH 7.4 at 34 °C. The Krebs solution also contained the l-type calcium channel blocker nifedipine (3 μM), the muscarinic acetylcholine receptor antagonist atropine (5 μM), and the cyclooxygenase inhibitor indomethacin (3 μM). The colonic tissue preparation was then cannulated at both ends. The proximal end was attached to an infusion pump to allow continuous perfusion of the Krebs buffer solution (0.2 mL/min), while the distal end was linked to a pressure transducer that measured intracolonic pressure. The lumbar splanchnic nerve bundles emanating from the colon were separated into small strands and then drawn into a glass suction recording electrode attached to a Neurolog headstage (NL100; Digitimer). The signal was filtered through a 10 Hz low-cut filter to remove low-frequency noise and a 5000 Hz high-cut filter to remove high-frequency noise. The analogue signal was digitized with a Micro 1401 MKII interface and recorded using CED Spike2 6.1 software. A ramp distension was applied by closing the outflow drain of the preparation until the intracolonic pressure reached 60 mmHg to ensure the selected nerve bundle was mechanically sensitive. Nerve strands that did not display an increase in action potential (AP) frequency in response to distension were discarded. The mechanical distensions were performed every 15 min until the magnitude of the distension response was reproducible (defined as ≤ 20% change in maximal AP discharge frequency between three distensions). Following confirmation of nerve stability, Mega Medium or WT B. fragilis or B. fragilis Δbfp1 supernatant was applied intraluminally by mixing 20μL of the 40x supernatant with 780 μL Krebs solution and perfusing for 10 min. At the end of the 10 min, but with supernatant still present in the colon, the afferent nerve response to ramp distension was reassessed. The intraluminal solution was then switched back to Krebs, and the colon was washed out for 30 min, distending every 15 min. Single-unit analysis was used to discriminate individual afferent fibers.⁸⁵ The analysis was conducted using the Spike2 sorting function. The basal firing rate was defined as the average spontaneous AP firing frequency (Hz) of individual axons (discriminated nerve units) in the absence of distensions. The basal AP discharge rate was measured for 120 s before the control distension and for 120 s just before the distension in the presence of supernatant. The distension response immediately before the supernatant was used as the vehicle distension. Nerve excitation in response to distensions performed in the presence of supernatant was compared with the vehicle distension response to determine its impact on the magnitude and pressuresensitivity of distension responses.

In vivo Visceromotor Response (VMR) to Colorectal Distension (CRD).

Telemetry implantation and colorectal distensions were performed as previously described.⁸⁶ Briefly, mice were anesthetized with isoflurane (2.5%), placed on a heating pad, and given bupivacaine (2 mg/kg, i.d.) and meloxicam (20 mg/kg, s.c.) as analgesics before surgery. A PhysioTel ETA-F10 telemetric transmitter (Data Science International, Saint Paul, MN) was inserted into the abdominal cavity. Electrode tips were sutured onto the external oblique muscle (~5–10 mm apart) to measure electromyographic (EMG) activity. Post-operative analgesia included meloxicam (2 μL/g s.c.) for three days. Mice were acclimatized in a restrainer (Kent Scientific Corporation) for 30 minutes daily over two days prior to visceromotor response recording.

On day 12 post-surgery, all mice received an enema containing B. fragilis Δbfp1 supernatant (37.5 μL supernatant + 112.5 μL sterile 0.9% saline, total 150 μL). Four days later (day 16), mice were randomized into two groups: one group was re-exposed to the Δbfp1 supernatant enema to control for sensitization effects from repeated colorectal distensions, while the other group received an enema of the B. fragilis WT supernatant (same volume and preparation). Prior to each enema, mice were fasted for 12 hours, gently anesthetized under isoflurane (2.5%), and given a 100 μL enema of sterile saline to empty the colon. After 15 minutes, the experimental enema was administered. Mice were held vertically suspended for 60 seconds to minimize leakage, returned to their cages without food but with water access and left undisturbed for 90 minutes. Following these 90 minutes, mice were sedated with isoflurane, placed in a restrainer, and a 4F arterial embolectomy catheter (Fogarty 120804FF, Edwards Lifesciences) was inserted in the distal 3 cm of the colorectum and secured to the tail. After 15 minutes of recovery from anesthesia, colorectal distensions were performed stepwise at volumes of 40, 60, and 80 μL (two 10-second distensions per volume, with 3-minute intervals). EMG activity was recorded with Ponemah v6.5 software (Data Science International). The mean basal EMG activity recorded 10 seconds prior to distension was subtracted from the mean activity during distension. Results are expressed as mV/s.

Intracolonic administration of bacterial supernatants.

Investigators were blinded to treatments and genotypes. Mice were lightly sedated (3–5% isoflurane). The following samples were administered into the colon lumen by enema (150 μl, 3 cm from anus): vehicle (sterile sodium chloride 0.9%); Mega Medium (10-fold concentrated); WT B. fragilis supernatant (10-fold concentrated); or B. fragilis Δbfp1 supernatant (10fold concentrated).

Colonic pain.

Mice were acclimatized to the room, apparatus, and investigator for 2 h per day for 2 days before the study. The abdomen was divided into 9 equal quadrants. von Frey filaments of increasing force were applied to the central quadrant, corresponding to the colonic region. Responses to von Frey filament stimulation included arching of the back, jumping, and raising the rear legs. Responses were measured as a baseline and hourly for 6 h after intracolonic administration of samples. Results are expressed as a threshold in grams. To determine the contribution of PAR₂ to pain, bacterial supernatants were similarly administered to Par₂^−/− global KO or Par₂^−/−-Nav1.8 mice.

Colonic inflammation.

The colon was removed 3 h after intracolonic administration of samples and was snap-frozen. RNA was extracted using a Direct-zol RNA Kit (Zymo Research, #R2050). cDNA was synthesized from 50 ng of DNase-treated RNA using a High-Capacity cDNA Reverse Transcriptase Kit (Applied Biosystems, #4374966). The cDNA (50 or 100 ng) was subjected to 40 cycles of qRT-PCR amplification using the QuantStudio 3 Real-Time PCR System. The mRNA expression of TNFα (#Mm00443258_m1), IL-1β (#Mm00434228_m1) and CXCL1 (#Mm04207460_m1) was measured by qPCR using TaqMan^® gene expression and TaqMan^® Fast Advanced Master Mix (#4444557). The relative amounts of target genes were calculated by normalizing the expression with the reference gene GAPDH (#Mm9999915_g1). The 2-ΔΔCT method was used to compare the reference and target gene levels.

Cytokine profiling was performed using the Mouse Cytokine Array, Panel A (R&D Systems, Catalog # ARY006, USA) according to the manufacturer’s instructions. This membrane-based antibody array simultaneously detects the relative expression levels of 40 mouse cytokines and chemokines. In brief, colonic tissue collected from mice subjected to intracolonic injection of WT B. fragilis, B. fragilis Δbfp1 and MM supernatants was homogenized and clarified by centrifugation at 10,000 × g for 10 minutes at 4°C to remove debris. Protein concentrations were determined using the BCA protein assay (Thermo Fisher Scientific), and all samples were normalized to the same total protein concentration (typically 300 μg per membrane). Nitrocellulose membranes pre-spotted with capture antibodies were blocked for 1 hour at room temperature in the array buffer provided. Samples were then mixed with a cocktail of biotinylated detection antibodies and incubated with the membranes overnight at 4°C on a rocking platform. Following incubation, membranes were washed thoroughly and incubated with streptavidin–HRP for 30 minutes at room temperature. After a final series of washes, the membranes were developed using the chemiluminescent detection reagent provided in the kit. Signal was captured following 300 seconds exposure, using a digital imaging system (ChemiDoc MP, Bio-Rad), and densitometry analysis was performed using ImageJ (NIH). The relative expression of each cytokine was calculated by quantifying the mean pixel density of duplicate antibody spots after background subtraction and normalized with the cytokine expression in a non-treated mouse colonic tissue (used as a control). Cytokines were considered differentially expressed if they showed a consistent change in signal intensity relative to control conditions across at least 2 biological replicates.

PAR₂ endocytosis.

Samples were administered into the colon of Par₂-mugfp mice as described above. The colon was removed and fixed (4% paraformaldehyde, PFA in PBS, 2 h, 4°C), cryoprotected (30% sucrose, PBS, 48 h, 4°C), and embedded in tissue freezing medium (TFM, #TFM-5, General Data). Frozen sections (8 μm) were prepared. Sections were blocked in 10% normal donkey serum (NDS), 0.05% Triton X-100 in PBS (1 h, RT). Sections were incubated with rabbit anti-GFP (1:400, overnight, 4°C; #600–401-215L, Rockland Immunochemicals). Slides were washed and incubated with donkey anti-rabbit Alexa Fluor^® 488 (1:1000, 45 min, RT; Invitrogen). Slides were incubated with DAPI (1 μg/ml, 5 min) and mounted in ProLong^® Gold Antifade (ThermoFisher). Sections were observed using a Leica SP8 confocal microscope with HCX PL APO 40x (NA 1.30) or 63x (NA 1.40) oil objectives (Leica-Microsystems). Images were processed using Adobe Photoshop and Illustrator.

Colon segmentation and quantification of internalization.

To segment cells and quantify PAR₂-muGFP internalization, 2-channel images from the mouse colon containing PAR₂-muGFP and DAPI staining were processed using Cellpose (Cellpose-SAM model, Cell diameter 50 μm).^87,88 The cell segmentation by Cellpose was converted to ROIs which were saved and imported into ImageJ. The ROIs of the segmented cells were used to create two masks. The cytosolic mask was generated by eroding 2 pixels (0.8–0.9 μm) along the outer edge of the ROI generated by Cellpose. The plasma membrane mask was generated by subtracting the cytosolic mask from the original mask, resulting in a band, 2 pixels thick along the outer edge of the cell. These cytosolic and membrane masks were used to generate new ROIs that were used to quantify the relative fluorescence units (RFU) of muGFP fluorescence along the plasma membrane (RFU_mem) and from the cytosol (RFU_cyto) of the colonocytes. The PAR₂-muGFP internalization was presented as a ratio of RFU_cyto / RFU_mem.

Antibiotic treatment and repopulation.

To deplete intestinal bacteria, mice received a mixture of antibiotics in drinking water / 10% sucrose for 7 d (days 0–7): ampicillin (1 g/L), vancomycin (0.5 g/L), neomycin (1 g/L) and metronidazole (1 g/L). During the repopulation phase (days 8–18), mice received vehicle (control), WT B. fragilis or B. fragilis Δbfp1 (1×10⁸ cells per 100 μL) every other day by gavage (100 μL). The engraftment phase continued from day 18. Fecal pellets were collected every other day and were snap-frozen for assessing the depletion of gut bacteria and repopulation with B. fragilis by qRT-PCR. Fecal pellet bacterial DNA was extracted using the QIAmp^® Power Fecal^® Pro DNA Kit, (Qiagen, USA). Bacterial DNA was quantified by qRT-PCR using 16S primers (100 nmol/L) for total bacterial DNA (Universal_forward: AAACTCAAAKGAATTGACGG, Universal_reverse: CTCACRRCACGAGCTGAC) or for B. fragilis DNA (Bf_DNA_fwd: TGATTCCGCATGGTTTCATT, Bf_DNA_rev: CGACCCATAGAGCCTTCATC) with 2x Fast SYBR Green master mix (#43–856-12, Applied Biosystem, USA) and 15 ng bacterial DNA. qRT-PCR conditions were: denaturation 94° for 30 s; 40 cycles of 94° for 5 s, 60° for 30 s. B. fragilis DNA extracted from culture was used to plot a standard curve for both B. fragilis and universal primer sets, which was used to calculate bacterial DNA concentration. Abdominal withdrawal responses to stimulation with von Frey filaments were measured on day 0 before antibiotic treatment, day 7 after antibiotic treatment, and alternate days during the repopulation and engraftment phases.

Metagenomic analysis.

Microbial DNA was extracted and purified from mouse fecal pellets using the QIAamp PowerFecal Pro DNA kit according to the manufacturer’s instructions. Shotgun metagenomic sample preparation was performed at the Genomic Technology Center at NYU Langone Medical Center, using the Nextera Flex Prep HT library prep. kit and paired-end sequenced (2×150 bp) on a NovaSeq X+ 10B 300 Cycle Flowcell occupying one lane. A read depth of 22 million reads per sample was targeted. Paired-end raw sequencing reads in FASTQ format were subjected to quality control using KneadData (https://huttenhower.sph.harvard.edu/kneaddata/). The cleaned reads were analyzed using MetaPhlAn 4.0.4⁸⁹ to characterize the species-level taxonomic composition of each sample. The resulting taxonomic profiles were integrated into Phyloseq⁹⁰ objects in R for downstream analysis and visualization. Alpha diversity was calculated at the species level using the microbiome package (version 1.20.0), and a Wilcoxon rank-sum test was used to identify any statistically significant differences. Beta diversity analyses were performed at the species level using Bray–Curtis dissimilarity and Weighted UniFrac distance metrics, and the results were visualized by principal coordinate analysis (PCoA). Statistical differences were determined by permutational multivariate analysis of variance (PERMANOVA).^91–93

Spontaneous non-evoked behavior.

On day 19 of the engraftment, non-evoked behavior was assessed using the behavioral spectrometer (Behavior Sequencer, Behavioral Instruments). Mice were individually placed in the center of the behavioral spectrometer and their ambulatory, exploratory and grooming behaviors were recorded, tracked, evaluated, and analyzed using a computerized video tracking system (Viewer3, BiObserve) for 30 min. Total distance traveled in the open field, average velocity of locomotion, visit to the center of the arena, ambulation, and grooming were recorded and analyzed.

Cloning, expression and purification of recombinant Bfp1.

The expression plasmid for recombinant Bfp1 was purchased from GenScript. In brief, the gene of Bfp1 without its signal peptide sequence but including an N-terminal Strep-tag and TEV-cleavage site was cloned into a pET28a vector and transformed into Escherichia coli Rosetta pLys cells (for the amino acid sequence see Supplementary Table S5). For overexpression, LB-media supplemented with kanamycin (50 μg/mL) and chloramphenicol (34 μg/mL) was inoculated 1:100 with an overnight culture and incubated at 37 °C and 200 rpm. Upon reaching an OD₆₀₀ = 0.6, the expression was induced by adding 1 mM Isopropyl β-D-1-thiogalactopyranoside (IPTG), and cells cultivated for at 37 °C, 200 rpm for 5 h. The cells were harvested (4700 rpm, 4 °C, 20 min) and the pellet was washed with PBS once. Cell lysis was performed using B-PER Bacterial Protein Extraction Reagent (Thermo Scientific) according to the manufacturer’s protocol. Cell debris was removed from the soluble cell lysate by centrifugation (11000 rpm, 4 °C, 30 min). Protein purification was performed on an FPLC-System (Äkta FPLC Explorer-900 by Amersham Biosciences) using a Strep-Tactin^®XT 4Flow^® high capacity FPLC column 5mL (IBA Lifesciences). The buffers utilized during the method were buffer W (50 mM HEPES, 150 mM NaCl, 10 mM CaCl₂,1 mM DTT pH 8.0) and buffer BXT (Buffer W + 50 mM Biotin). The column was prepared by 3 column volumes (CV) wash with Mili Q followed by equilibration with 3 CV of buffer W before the lysate was applied on the column with a flow rate of 0.5 mL/min. The column was washed with buffer W untill the UV reached baseline. Following this, the bound Bfp1 was eluted using buffer BXT with isocratic elution. Buffer exchange to buffer X (50 mM HEPES, 150mM NaCl pH 8.0) was performed thrice using a 10 kDa molecular weight cutoff filter (6000 × g at 4 °C, 10 min). The sample was subsequently incubated at 20 °C until proteolytic activity was observed via enzyme assay. Precipitated protein was removed by centrifugation (6000 × g at 4 °C, 10 min) and supernatants were concentrated using a 10 kDa molecular weight cutoff filter (6000 × g at 4 °C, 10 min). An SDS-PAGE gel analysis of the purification is presented in Supplementary Figure S6A.

Protein purity and identity were verified by intact-protein LC-MS using a Dionex Ultimate 3000 UHPLC system coupled to a Bruker Maxis Q-Tof mass spectrometer. Chromatography was performed using a ZORBAX 300SB-C3 column (Agilent Technologies) at 65 °C with a 15 min gradient (5 % B to 95% B over 12 min; A: 0.1% formic acid in water, B: 0.1 % formic acid in acetonitrile). The raw MS spectra were processed using Compass DataAnalysis 4.3 (Bruker) and then deconvoluted using UniDec (6.03).⁹⁴ Raw data and deconvoluted spectra are presented in Supplementary Figure S6B,C. Purified Bfp1 was detected as the full protein without its Start-Methionine (MW_calculated: 63557 Da, MW_measured: 63599 Da) with a mass difference of 42 Da which could potentially correspond to N-terminal acetylation of the protein.

Enzyme activity of purified Bfp1 was determined using a fluorogenic substrate assay. In a black flatbottom 384-well plate, 50 μL of Bfp1 (1 μM in buffer X) was incubated with 100 μM FP-alkyne (inhibition control) or DMSO (positive control) for 1 h at 37°C. Next, 100 μM of fluorogenic substrate Boc-Val-Leu-Lys-AMC was added and peptidolytic release of AMC was monitored by fluorescence (excitation: 355 nm, emission 466 nm) at 38.5 °C using a microplate-reader every 1 min for 60 min. Activity data for purified Bfp1 are presented in Supplementary Figure S6D.

QUANTIFICATION AND STATISTICAL ANALYSIS

Statistical analysis.

Data are presented as mean ± SEM. For studies of cells, n>5 experiments were performed. For experiments with mice, 5–6 mice were used per treatment. Differences were assessed using Student’s t-test for two comparisons and 1 or 2 way-ANOVA followed by Tukey for multiple comparisons test. For 3D organoid experiments 30–130 organoids were assessed per treatment. A nonparametric Kruskal-Wallis test followed by a Dunn’s multiple comparisons test was performed to assess differences. p<0.05 was considered significant at the 95% confidence level in all cases. All tests were performed in GraphPad Prism (v. 10.4.1).

Supplementary Material

1

Document S1. Figures S1–S13 and Supplementary References.

2

Supplementary Table S1. Results of the initial proteolysis screen, Related to Figure 1.

3

Supplementary Table S2. Proteomics results of direct and competitive ABPP, Related to Figure 2.

4

Supplementary Table S3. BLAST-N Analysis Results for tspA, Related to Figure 2.

5

Supplementary Table S4. BLAST-N Analysis Results for bfp1, Related to Figure 2.

6

Supplementary Table S5. Nucleotide and protein sequences, Related to STAR-Methods.

7

Supplementary Table S6. Plasmids used in this study, Related to STAR-Methods.

8

Supplementary Table S7. Primers used in this study, Related to STAR-Methods.

9

Supplementary Table S8. Preparation of Mega Medium, Related to STAR-Methods.

10

Video 1. Spontaneous behavior of mice after antibiotic treatment, repopulation and engraftment with B. fragilis, Related to Figure S13. The videos show the spontaneous behavior of mice over 90 s (speeded-up three times) on day 19 of the antibiotic treatment, repopulation and engraftment protocol. Par₂^+/+ mugfp mice received water and vehicle (control) or antibiotic cocktail followed by vehicle, WT B. fragilis, or B. fragilis Δbfp1. Recordings are from individual mice that are representative of n = 5 mice.

Spontaneous behavior of mice after antibiotic treatment, repopulation and engraftment with B. fragilis, Related to Figure 7.

KEY RESOURCES TABLE

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
Roche Rat anti-HA 3F10 monoclonal	Millipore	Cat# 11867423001 RRID: AB_390918
DYKDDDDK Tag (9A3) mouse mAb	Cell Signaling	Cat# 8146 RRID: AB_2687462
Donkey anti-mouse-Alexa488	ThermoFisher	Cat# A-21202 RRID: AB_141607
Donkey anti-rat-Alexa568	ThermoFisher	Cat# A78946 RRID: AB_2910653
Phalloidin-647	Cell Signaling Technologies	Cat# 8940S
Bacterial and virus strains
B. fragilis NCTC 9343 Δbfp1	This work	N/A
B. fragilis NCTC 9343 ΔtspA	This work	N/A
Intestinal strain collection	Table S1	N/A
Biological Samples
Murine colons – C57Bl/6 mice -Ussing chamber, afferent nerve electrophysiology	Queen’s University	N/A
Bacterial culture supernatants	This work	N/A
Chemicals, peptides, and recombinant proteins
Fluo-4 AM	Cayman	Cat# 31143
HBSS	Invitrogen	Cat# 14025092
Trypsin	Sigma-Aldrich	Cat# T6424
AZ3451	Sigma-Aldrich	Cat# SML2050
PEI – MW 25000	Polysciences Inc.	Cat# 23966
His-PAR2−NTD-Cy5	This work	N/A
IntestiCult Organoid Growth Medium	Stemcell	Cat# 06010
Y27632	MedChem Express	Cat# HY-10583
CHIR 99021	Tocris	Cat# 102875-390
Matrigel	Corning	Cat# 354234
FITC-dextran	Sigma-Aldrich	Cat# 46944-100MG-F
Collagenase I, Dispase II	Sigma-Aldrich	Cat# 1148089
Neurobasal Plus Medium	Invitrogen-Gibco	Cat# A3582901
Critical commercial assays
Proteome Profiler Mouse cytokine Arrey Kit, Panel A	R&D System	Cat# ARY006
QIAamp PowerFecal Pro DNA Kit	QIAGEN	#51804
Deposited data
Reference genomes from cultivated human gut bacteria	Zou et. al.26	NCBI Bioproject ID: 482748
Proteomics Data	This work	https://www.ebi.ac.uk/pride/ Identifier: PXD059166
Experimental models: Cell lines
Human Embryonic Kidney HEK293 cell line	ATCC	Cat# CRL-1573 RRID: CVCL_0045
HEK293-PAR2−/− KO cell line	University of Calgary – Dr Morley D. Hollenberg	N/A
HEK293(HA-PAR2-mApple)	This study	N/A
Human ileal organoid line	Calvin Kuo lab	HC921
Experimental models: Organisms/strains
Afferent nerve/Ussing chamber/CRDVMR data – C57Bl/6 mice	Queen’s University	N/A
Par2−/− KO (B6.Cg-F2rl1tm1Mslb/J) mice	JAX	Cat# IMSR_JAX:004993
Par2−/−-Nav1.8 mice	Bunnett Lab	N/A
Par2+/+ mugfp mice	Bunnett Lab/ Monash University	N/A
Par2−/− Nav1.8/ Par2+/+-Cre mice	Queen’s University	N/A
Oligonucleotides
Primers used in this study	Table S6	N/A
Recombinant DNA
pTwist_CMV_Hygro-HA-PAR2-mApple	Twist Biosciences	N/A
pET28a_His-PAR2NTD-GyrA	Genscript	N/A
pET28a_Strep-trBfp1	Genscript	N/A
pLGB30	Comstock lab27	Addgene #126620
Software and Algorithms
BLAST-N 2.16.0	NCBI	RRID:SCR_004870
iTOL	EMBL	RRID:SCR_018174
Spike2 6.1	Cambridge Electronic Design	RRID:SCR_000903
Ponemah v6.5	Data Science International	RRID:SCR_017107
MaxQuant 2.3.0.3	MPI for Biochemistry	RRID:SCR_014485
Perseus v.2.0.3.0	MPI for Biochemistry	RRID:SCR_015753
ImageJ	NIH	RRID:SCR_003070
GraphPad Prism	GraphPad	RRID:SCR_002798
Affinity Designer	Serif	RRID:SCR_016952
Research Randomizer	Laboratory of Cellular Communication, Oswaldo Cruz Institute 78	N/A
Other
Mega Medium	Han et al.17 , see STAR-Methods and Table S8	N/A

Highlights.

• Intestinal bacteria secrete proteases that affect host PAR₂ signaling

• Chemoproteomics identifies Bfp1, a serine protease released by Bacteroides fragilis

• Bfp1 activates PAR₂ to disrupt epithelial integrity, induce nociception and inflammation

• Bfp1-PAR₂-axis links the microbiota to pain and inflammation in the gut