# Oral indomethacin modifies small intestine biofilms and host-microbe interaction mediators

Arezoo Haghighi, András Tóth, Zsuzsanna Demeter, Barbara Hutka, Anna Zsidai, Lili Lengyel, Samaneh Haghighi, Melissa Pannier, Guillaume Le Cosquer, Elodie Meunier, et al.

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## To cite this version:

Arezoo Haghighi, András Tóth, Zsuzsanna Demeter, Barbara Hutka, Anna Zsidai, et al. Oral indomethacin modifies small intestine biofilms and host-microbe interaction mediators. Life Sciences, 2026, 384, pp.124114. 10.1016/j.lfs.2025.124114. inserm-05392928

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Oral indomethacin modifies small intestine biofilms and host-microbe interaction mediators

Arezoo Haghighi<sup>a,b</sup>, András S. Tóth<sup>a,b</sup>, Zsuzsanna O. Demeter<sup>a,b</sup>, Barbara Hutka<sup>a,c</sup>,
Anna Zsidai<sup>a,b</sup>, Lili Lengyel<sup>a,b</sup>, Samaneh Haghighi<sup>d</sup>, Melissa Pannier<sup>e</sup>, Guillaume Le Cosquer<sup>e,f</sup>,
Elodie S. Meunier<sup>e</sup>, Bence Ágg<sup>b,g,h</sup>, Nóra Makra<sup>i</sup>, Eszter Ostorházi<sup>i</sup>, Balázs Ligeti<sup>j</sup>,
Krisztián Kovács<sup>k</sup>, Ágnes Kelemen<sup>l</sup>, Anna Jakab<sup>m</sup>, Gerda Wachtl<sup>a,b</sup>, Gábor Kökény<sup>d,n</sup>,
Dóra Szabó<sup>i,o</sup>, Péter Ferdinandy<sup>b,g,h</sup>, Jean-Paul Motta<sup>e</sup>, Nathalie Vergnolle<sup>e</sup>, Klára Gyires<sup>a,b</sup>,
Zoltán S. Zádori<sup>a,b,*</sup>

ARTICLE INFO

Keywords:

Nonsteroidal anti-inflammatory drugs

Indomethacin

Enteropathy

Microbiota

Toll-like receptor

Biofilm

Antimicrobial peptides

A B S T R A C T

**Background and aims:** Nonsteroidal anti-inflammatory drugs (NSAIDs) can cause small intestinal injury and dysbiosis. Although NSAID-induced dysbiosis is well-characterized and contributes to enteropathy, the changes in host-bacterial interactions during enteropathy remain largely unexplored. Here we assessed the expression pattern of six toll-like receptors (TLRs) and three antimicrobial peptides (AMPs) over the course of indomethacin (IND)-induced enteropathy in rats, and evaluated their correlations with inflammation and dysbiosis. In addition, we assessed for the first time the effect of IND on small intestinal mucosal biofilm structure.

**Materials and methods:** Mucosal injury, inflammation and expression of TLR and AMP genes were evaluated at five time points following IND administration. Gut microbiota composition was determined by 16S rRNA gene sequencing. Small intestinal mucosal biofilms were visualized using fluorescent in situ hybridisation.

**Key findings:** We found that TLR1, TLR2 and cathelicidin were upregulated, TLR5 was downregulated, whereas TLR6 and TLR9 were not altered in enteropathy. TLR4 expression showed only subtle differences, but correlated with α-defensin 5 and β-defensin 2 levels. We found several correlations between TLRs, AMPs, inflammation and gut bacteria in severe enteropathy, but in early disease stage TLR1, TLR2, TLR5 and cathelicidin expression were more strongly associated with inflammation, whereas TLR4 and defensins were more dependent on gut dysbiosis. IND treatment also caused mild damage to the mucosal microbiota biofilm.

**Significance:** This is the first comprehensive characterization of the time-dependent changes in TLRs, AMPs and mucosal biofilm in NSAID-treated rats, which may help to identify new strategies for the treatment of enteropathy.

<footnote><sup>a</sup> Department of Pharmacology and Pharmacotherapy, Semmelweis University, Budapest, Hungary</footnote>

<footnote><sup>b</sup> Center for Pharmacology and Drug Research & Development, Semmelweis University, Budapest, Hungary</footnote>

<footnote><sup>c</sup> Pharmacological and Drug Safety Research, Gedeon Richter Plc, Budapest, Hungary</footnote>

<footnote><sup>d</sup> Institute of Translational Medicine, Semmelweis University, Budapest, Hungary</footnote>

<footnote><sup>e</sup> Toulouse University, IRSD, INSERM, INRAe, ENVT, Toulouse, France</footnote>

<footnote><sup>f</sup> Department of Gastroenterology and Pancreatology, Toulouse University Hospital (CHU Toulouse), Toulouse, France</footnote>

<footnote><sup>g</sup> Cardiometabolic and HUN-REN-SU System Pharmacology Research Group, Department of Pharmacology and Pharmacotherapy, Semmelweis University, Budapest, Hungary</footnote>

<footnote><sup>h</sup> Pharmahungary Group, Szeged, Hungary</footnote>

<footnote><sup>i</sup> Institute of Medical Microbiology, Semmelweis University, Budapest, Hungary</footnote>

<footnote><sup>j</sup> Faculty of Information Technology and Bionics, Pázmány Péter Catholic University, Budapest, Hungary</footnote>

<footnote><sup>k</sup> Department of Laboratory Medicine, Semmelweis University, Budapest, Hungary</footnote>

<footnote><sup>l</sup> Department of Histopathology, Central Hospital of Northern Pest - Military Hospital, Budapest, Hungary</footnote>

<footnote><sup>m</sup> Department of Pathology, Forensic and Insurance Medicine, Semmelweis University, Budapest, Hungary</footnote>

<footnote><sup>n</sup> International Nephrology Research and Training Center, Semmelweis University, Budapest, Hungary</footnote>

<footnote><sup>o</sup> HUN-REN Human Microbiota Study Group, Institute of Medical Microbiology, Semmelweis University, Budapest, Hungary</footnote>

<footnote>* Corresponding author at: Department of Pharmacology and Pharmacotherapy, Semmelweis University, Nagyvárad tér 4., 1089, Budapest, Hungary.
E-mail address: zadori.zoltan@semmelweis.hu (Z.S. Zádori).</footnote>
# 1. Introduction

Nonsteroidal anti-inflammatory drugs (NSAIDs) are among the most commonly used prescription and over-the-counter drugs. Around 30 million people in the United States take NSAIDs regularly [1], and their use is steadily increasing worldwide [2]. However, NSAID use is associated with an increased risk of gastrointestinal (GI) damage, which is not limited to the gastroduodenal region but can also occur in the mid-distal small bowel. Capsule enteroscopy studies have revealed that up to 70% of long-term NSAID users may develop NSAID enteropathy [3,4]. A notable aspect of NSAID enteropathy is its complex and not yet fully elucidated pathogenesis, which makes it challenging to develop effective treatment strategies [5,6].

Over the past decades, numerous studies have demonstrated the pivotal role of intestinal bacteria in the development of enteropathy [4,7]. Current estimates suggest that the distal parts of the small intestine, where enteropathy typically develops, harbour approximately 10<sup>11</sup> bacteria [8], belonging mainly to the *Firmicutes* phylum [9,10]. In enteropathy, these bacteria are gradually replaced by Gram-negatives, such as members of the *Proteobacteria* and *Bacteroidetes* phyla [4,11], and these changes exhibit a strong correlation with the severity of intestinal inflammation and damage [12,13]. The significance of bacteria in enteropathy is underscored by the observations that certain antibiotics, probiotics, or fecal microbiota transplantation may offer protection against NSAID-induced intestinal injury [4,7,14,15]. Nevertheless, the clinical benefit of these strategies is still uncertain [16], and the prolonged use of even "safe" antibiotics, such as rifaximin, can be associated with the development of multi-drug resistant pathogens [17].

Instead of directly modulating gut microbiota composition, targeting the receptors involved in bacterial recognition may offer a more subtle way to control host-bacterial interactions. The intestinal epithelial and immune cells express a wide array of pattern recognition receptors (PRRs) capable of detecting diverse microbial components and initiating various immune responses [18]. Toll-like receptors (TLRs) represent one of the best characterized families of PRRs [19], comprising 10 members (TLR1-TLR10) in humans and 13 in rodents [20]. Of these, TLR4, TLR5, TLR9, and TLR2 and its heterodimerizing partners, TLR1 and TLR6 are primarily involved in bacterial detection [19]. The activation of these receptors by specific bacterial components leads to the induction of inflammation and the secretion of various antimicrobial peptides, including defensins [21,22] and cathelicidin [23], which play a crucial role in the suppression of bacterial invasion and the eradication of pathogens. TLRs, however, do not only influence but are also influenced by inflammatory cytokines and bacteria [24–27]. Changes in intestinal TLR expression and dysregulated immune responses are believed to play a fundamental role in the perpetuation of inflammation in diseases such as inflammatory bowel disease (IBD) [28–31], and TLR ligands were shown to either reduce [32–35] or aggravate [9,36] intestinal inflammation and injury depending on the model used.

Interestingly, despite the long-recognized importance of intestinal bacteria in enteropathy, NSAID-induced changes in host-bacterial interactions remain largely unexplored. Studies so far have focused mainly on TLR4 and TLR2 in this context, demonstrating that the activation of TLR4 is critical in enteropathy development [37], whereas TLR2 ligands are rather protective, although it also depended on the timing of ligand administration [38]. In addition, both TLR4 and TLR2 have been shown to be upregulated in enteropathy [39–42], which was linked to dysbiosis and increased mucosal adherence of *Enterobacteriaceae* and *Bacteroides* [41]. Nevertheless, the finding that NSAIDs increase TLR4 and TLR2 expression is not universal [37,40,43], suggesting that changes in TLRs depend on several factors such as disease severity and the actual composition of gut microbiota. With regard to other TLRs, one study has reported no change in intestinal TLR5 expression following fecal transplantation from mice exposed to the NSAID indomethacin (IND) [44], whereas, to the best of our knowledge, no studies have been conducted to assess the NSAID-induced changes in TLR1, TLR6 and TLR9

expression in the gut. Similarly, little is known about the changes in AMPs and their associations with TLRs, inflammation and dysbiosis in enteropathy.

In addition, intestinal bacteria not only exist in planktonic form, but also form larger mucin-embedded aggregates that adhere to the mucosa and food particles, called biofilms [45]. Mucosal microbiota biofilms play a crucial role in bacterial-host interactions, and alterations in biofilm biogeography are associated with several intestinal diseases such as IBD [46–48]. Although NSAID-induced changes in gut microbiota composition have been extensively studied [4,11], and several studies have shown that certain NSAIDs can interfere with bacterial biofilm formation in vitro [49], the effect of NSAIDs on small intestinal mucosal biofilm integrity and its role in enteropathy is not known.

In order to better understand NSAID-induced changes in bacterial-host interactions and identify potential targets for therapeutic intervention, we have assessed the time course of changes in the small intestinal gene expression of TLR1, TLR2, TLR4, TLR5, TLR6, TLR9, and of three major AMPs (α-defensin 5, β-defensin 2 and cathelicidin /CAMP/) in rats with acute enteropathy, and have comprehensively evaluated their correlations with intestinal inflammation and dysbiosis. Changes in TLR and AMP expressions were also determined in rats receiving low-dose IND repeatedly over a prolonged time, which is more similar to the clinical scenario of long-term NSAID use. In addition, we have also assessed, for the first time, the effect on IND on small intestinal bacterial biofilm structure both in vivo and ex vivo.

# 2. Materials and methods

## 2.1. Animals

Experiments were carried out on non-fasted male Wistar rats weighing 220–240 g (Toxi-Coop Ltd., Budapest, Hungary). Animals were housed in a temperature (22 ± 2 °C)- and humidity-controlled room at a 12-h light/dark cycle. Food and water were available ad libitum. All efforts were made to minimize animal suffering and to reduce the number of animals used in the experiments. All procedures conformed to the Directive 2010/63/EU of European Convention for the protection of animals used for scientific purposes. The experiments were approved by the National Scientific Ethical Committee on Animal Experimentation and permitted by the government (Food Chain Safety and Animal Health Directorate of the Government Office for Pest County (PE/EA/1118–6/2020)).

## 2.2. Study design

Indomethacin-induced changes in intestinal expression of TLRs and AMPs were assessed in two consecutive, separate studies. Before each study, rats were co-housed to homogenize their gut microbiota, and then were randomly allocated into vehicle- (1 % hydroxyethylcellulose) or IND-treated groups.

First, in order to assess the time course of changes in intestinal TLRs' and AMPs' expression in acute enteropathy, rats were treated with a single, large dose (20 mg/kg) of IND (Sigma, St. Louis, MO, USA) via gastric gavage, as previously described [13], and then were euthanized at 6, 12, 24, 48 or 72 h after the IND treatment, with 8–8 animals in each group. Because our preliminary experiments showed that the microbiota composition of vehicle (1 % hydroxyethylcellulose)-treated rats is stable over time, provided that there are no changes in their environment, control animals were euthanized only 72 h after the administration of vehicle.

In the second study, we aimed to treat rats with lower doses of IND, 2 and 4 mg/kg twice daily for 14 and 7 days, respectively. We chose this protocol based on the results of pilot studies in which prolonged administration of IND at a dose of 4 mg/kg/day for 4 weeks caused moderately severe enteropathy in rats. However, in the present study the 4 mg/kg dose of IND given twice a day caused severe enteropathy, and
<table><caption>**Table 1**
List of primer sequences.</caption><thead><tr><th>Gene</th><th>Primer sequence (5'-3')</th><th>Accession number</th></tr></thead><tbody><tr><td>Tlr1</td><td>forward CTG ATC TCT TGC CAC GCA AC<br>reverse TCT GAG AAC CGC TCA ATC CC</td><td>NM_001172120.2</td></tr><tr><td>Tlr2</td><td>forward TTG GGT GGA GAA CCT CAT GG<br>reverse GGA ACA AAG TCC CGC TTG TG</td><td>NM_198769.2</td></tr><tr><td>Tlr4</td><td>forward AAC CTC AAC GAC CTC ACA GG<br>reverse GTT CTC ACT GGG CCT TAG CC</td><td>NM_019178.2</td></tr><tr><td>Tlr5</td><td>forward GCC AGA GCC AGA TTG AAG TCT TGA<br>reverse GAG AGG CTG GAG TTC ATC TTC ACA</td><td>NM_001145828.1</td></tr><tr><td>Tlr6</td><td>forward CCT CTG GGA TAG CCT CTG CAA C<br>reverse CAG AGA ACT GGG TCA TGC TTC C</td><td>NM_207604.1</td></tr><tr><td>Tlr9</td><td>forward TGC TGG ACC TAA GCG AGA AC<br>reverse AAG GAA CTT GCC AGG TGG AG</td><td>NM_198131.1</td></tr><tr><td>Defa5</td><td>forward GCA CTC CGA CCA AGA AGA ATC C<br>reverse CAT CTG ACA AAT AGC AGT CTC GTT C</td><td>NM_173329.2</td></tr><tr><td>Defb2</td><td>forward AGG GTT CTG TCT CAA TGC TG<br>reverse TGC AAC AGT AGA ACT TGT ATG GG</td><td>NM_001037507.2</td></tr><tr><td>Camp</td><td>forward GGC ATC ACA GCA GTT ACC TG<br>reverse GCT GTA TAC CAG GCG CAT C</td><td>NM_001100724.1</td></tr><tr><td>Il1b</td><td>forward TGG CAA CTG TCC CTG AAC TC<br>reverse GGG CTT GGA AGC AAT CCT TAA TC</td><td>NM_031512.2</td></tr><tr><td>Il10</td><td>forward GAA CCA CCC GGC ATC TAC TG<br>reverse AGG AGT TGC TCC CGT TAGC</td><td>NM_012854.2</td></tr><tr><td>Tnf</td><td>forward ACT GAA CTT CGG GGT GAT TG<br>reverse GCT TGG TGG TTT GCT ACG AC</td><td>NM_012675.3</td></tr><tr><td>Rpl13a</td><td>forward GGA TCC CTC CAC CCT ATG ACA<br>reverse CTG GTA CTT CCA CCC GAC CTC</td><td>NM_173340.2</td></tr></tbody></table>

animals were euthanized one day earlier than originally scheduled due to their poor general condition. As in the first study, control animals were euthanized only at one time point, after a two-week vehicle treatment.

At the time of sacrifice, blood samples were collected and the small intestines were excised. The length of the whole small intestine was measured, as one parameter to assess intestinal inflammation. Then, full-thickness pieces of the distal small intestine and luminal contents of the same sites were snap-frozen and stored at -80 °C for further assays. In addition, distal jejunal segments were fixed in either 10 % formalin for standard histologic analysis, or in Methacarn fixative (methanol 60 %/chloroform 30 %/acetic acid 10 %) for visualizing the spatial organization of microbiota biofilm [48].

### 2.3. Western blot analysis

Tissue samples were homogenized and Western blot was performed as described previously [13]. The primary antibodies used were anti-COX-2 (D5H5) (12282S, 1:1000, Cell Signaling Technology, Danvers, MA, USA), anti-interleukin-1β (ab9722, 1:1000) and anti-pentraxin-3 (ab125007, 1:1000) (Abcam, Cambridge, UK) and anti-myeloperoxidase (AF3667, 1:1000, R&D Systems, Minneapolis, MN, USA). As secondary antibodies HRP-linked anti-rabbit IgG (7074, 1:10000, Cell Signaling Technology, Danvers, MA, USA) and anti-goat IgG (ab97120, 1:10000, Abcam, Cambridge, UK) were used. Anti-GAPDH (D16H11) (5174S, 1:1000, Cell Signaling Technology, Danvers, MA, USA) was used to control for sample loading and protein transfer and to normalize the content of target protein. Membranes were trimmed before the antibody treatment if the bands of interest were far apart. At least two repetitions were performed for each experiment. Signals were detected with a chemiluminescence kit (BioRad, Hercules, CA, USA) by Chemidoc XRS+ (BioRad, Hercules, CA, USA).

### 2.4. qRT-PCR measurements

Total RNA was obtained from 10 to 30 mg of small intestine tissue using the QIAzol extraction method (Qiagen, Hilden, Germany). RNA concentration was measured with Nanophotometer (Implen GmbH,

Munich, Germany). Reverse transcription was performed from 1 µg of total RNA with a SensiFAST cDNA synthesis kit (Bioline, London, UK) according to the manufacturer's protocol. Target genes were amplified on a LightCycler® 480 II instrument (Roche, Germany) using the SensiFAST SYBR Green master mix (Bioline, London, UK). Expression levels were calculated with the $2^{-\Delta\Delta CT}$ evaluation method and Rpl13a was used as reference gene. The sequences of primers used are listed in Table 1.

### 2.5. Histology

Distal small intestinal samples were excised and prepared using the Swiss-roll technique, then were fixed in 10 % formalin, embedded in paraffin, sectioned (4 µm), and stained with haematoxylin and eosin. Histologic damage was graded blindly by two histopathologists. Histological changes were scored in six categories: villus morphology, the presence of erosions, the presence and extent of ulcers, the severity of mucin depletion, inflammatory cell infiltration and peritoneal reaction (Supplementary Table 1). The sub-scores were then added together to produce a total score of 17. Five grades of damage severity were then determined from these scores: grade 0 (0), grade 1 (1-3), grade 2 (4-8), grade 3 (9-13) and grade 4 (14-17).

### 2.6. Hematological analysis

Whole blood samples were collected in Vacuette K-3 EDTA tubes (Greiner Bio-One, Kremsmünster, Austria). Hematological analysis was performed using a Sysmex XN 1000 (Sysmex America, Lincolnshire, IL, USA) hematology analyzer according to routine laboratory protocols. The results were calculated automatically using the Sysmex XN 1000 hematology software.

### 2.7. Gut microbiota analysis

Bacterial DNA was extracted from 80 mg of small intestinal sample using the QIAamp PowerFecal DNA Kit (Qiagen, Hilden, Germany) and was further purified using AMPure XP beads (Beckman Coulter, Brea, CA, USA) according to the manufacturer's protocols. The concentration of genomic DNA was measured using a Qubit 2.0 Fluorometer with Qubit dsDNA HS Assay Kit (Thermo Fisher Scientific, Waltham, MA, USA). Bacterial DNA was amplified with tagged primers (5'-TCG TCG GCA GCG TCA GAT GTG TAT AAG AGA CAG CCT ACG GGN GGC WGC AG and 5'-GTC TCG TGG GCT CGG AGA TGT GTA TAA GAG ACA GGA CTA CHV GGG TAT CTA ATC C), covering the V3-V4 region of the bacterial 16S rRNA gene. PCR and DNA purifications were performed according to Illumina's demonstrated protocol (Part # 15044223 Rev. B). The PCR product libraries were quantified and qualified by using DNA 1000 Kit on Agilent 2100 Bioanalyzer instrument (Agilent Technologies, Waldbronn, Germany). Equimolar concentrations of libraries were pooled and sequenced on an Illumina MiSeq platform (Illumina, San Diego, CA, USA) using MiSeq Reagent Kit v3 (600 cycles PE). Bioinformatic analysis was carried out as described previously [50].

In a separate analysis we assessed the changes in the *Lachnospiraceae* family in response to repeated IND treatments. To this end, we designed specific primers targeting all bacteria and bacteria belonging to the *Lachnospiraceae*. Bacterial 16S rRNA gene sequences were obtained from the European Nucleotide Archive and aligned in UGENE [51] to identify potential specific primers. Final primers were selected based on testing them in SILVA [52] and Ribosomal Database Project (RDP) [53] rRNA gene databases (Table 2). Primers were synthesized commercially (Integrated DNA Technologies, IA, USA). Bacterial DNA was extracted from 100 mg luminal samples by ZymoBIOMICS DNA Miniprep Kit (Zymo Research, USA), then qPCR was performed as described in section 2.4. Expression levels were calculated with the $2^{-\Delta\Delta CT}$ evaluation method and normalized for the universal bacterial primer [54].
<table><caption>**Table 2**

Primer sequences for bacterial analysis.</caption><thead><tr><th>Target</th><th>Primer sequence (5'-3')</th><th>Amplicon size (bp)</th><th>Applied annealing temperature (°C)</th></tr></thead><tbody><tr><td rowspan="2">Lachnospiraceae</td><td>forward</td><td>GGG AAG<br>AWA ATG<br>ACG GTA CCT<br>GAC TAA</td><td rowspan="2">140</td><td rowspan="2">62</td></tr><tr><td>reverse</td><td>RCG CTC CCT<br>TTA CAC CCA<br>GTA AA</td></tr><tr><td rowspan="2">All bacteria</td><td>forward</td><td>CCA GCA GCC<br>GCG GTA ATA<br>CG</td><td rowspan="2">260</td><td rowspan="2">62</td></tr><tr><td>reverse</td><td>TGG ACT ACC<br>AGG GTA TCT<br>AAT CCT GTT</td></tr></tbody></table>

## 2.8. Fluorescent in situ hybridization

Distal small intestinal tissues were Methacarn-fixed and paraffin-embedded. Slides were hybridized with 1 µM of a universal bacterial 16S fluorescent rRNA probe (EUB338-Cy3, 5'-GCT GCC TCC CGT AGG AGT-3' Cy5, Eurogentec, Seraing, Belgium) and counterstained for DNA by 4',6-diamidino-2-phenylindole (DAPI, 1:1000, Sigma, France) and polysaccharides content (wheat germ agglutinin labeled with fluorescein, L4895, 1:1000, Sigma, France) [48]. Representative images were obtained from blind acquisition of 4 different fields per animal on a Leica LSM 710 confocal microscope. FIJI freeware was used for final image mounting (v.1.51). Biofilm damage was assessed in blinded fashion. While we initially adopted the scoring system from Motta et al. (2019), we found it necessary to adapt it for the small intestine (Table 3). Given the distinct organization of the small intestinal microbiota compared to the colon, we focused primarily on the “overall density” of the mucosal biofilm, as this metric appears more relevant to small intestinal microbiota organization.

## 2.9. Ex vivo analysis of gut microbiota biofilms

Ex vivo analysis of intestinal microbiota biofilms was performed as described previously with slight modifications [48]. Briefly, rat small intestinal sections (ileum) from 5 different animals (Wistar rats, conventionally housed in CREFRE animal facility, Toulouse, France) were immediately transferred in sterile tubes containing anaerobic media (Brain Heart Infusion, BHI broth, Sigma-Aldrich, St. Quentin Fallavier, France) supplemented with hemin/menadione (1/0.01 mg, Sigma) and L-cysteine (0.5 mg/ml, Sigma, France). Tissue was immediately homogenized using a microtube pestle, then mucosa-associated microbiota was cultured overnight in BHI broth anaerobic media. Overnight anaerobic cultures were diluted to 0.1 OD 600 nm density and biofilms were generated for 72 h in the Calgary Biofilm Device (Innovotech, Edmonton, Canada). All steps described herein were performed under anaerobic conditions (Don Whitley D20) and incubated in hermetic jars with an anaerobic catalyser (Anaeropack, Thermo Fisher, France). Mature biofilms were transferred onto a new plate of fresh microbial broth (BHI) containing different concentration of IND

(0.01–10 µM from a stock solution of 20 mM in DMSO, I7378 from Sigma, France) or vehicle (microbial broth). After 24 h exposure to IND, the biofilm biomass density and bacterial metabolic activity (indicative of viable bacteria) were determined respectively by safranin-O staining (Gram's solution 10%, Sigma, France) and rezasurin (1 mg/ml in water, Sigma, France). Briefly, biofilms were transferred into new well plate containing stains and incubated for 2 h (resazurin) or 20 min (safranin-O). Safranin was recovered by sonication at 45 kHz (USC-500 T, VWR) in a plate containing acetic acid (30%, safranin-O). Specific absorbance (safranin-O) and fluorescence (resazurin/resofurin) were measured using a fluorescent spectrophotometer (Ensight, Perkin Elmer, Villepinte, France). Biofilm rate of dispersal was assessed by measuring viable bacteria released in wells after 24 h and normalized by the biofilm biomass. All results were expressed as a percentage change relative to the mean of the vehicle-treated group, which was set as 100%. An average of at least 5 biofilm replicates were performed for each concentration.

## 2.10. Cell culture experiments

Human colorectal adenocarcinoma DLD-1 cells were maintained in RPMI 1640 (21875–034, Gibco, Waltham, USA) medium supplemented with 10% fetal bovine serum (FBS) (35–079-CV, Corning, USA) and 1% Antibiotic-Antimycotic Solution (30–004-CI, Corning) at a 37 °C in a standard cell culture incubator (5% CO₂). Cells were seeded on stripwell 96 well plates (CLS2592, Corning) at a density of 5000 cells/well and the effect of IND (0.1–100 µM) on cell viability was measured with CellTiter-Glo Luminescent Viability Assay (G7571, Promega, Madison, USA) according to the manufacturer's protocol and luminescence intensity was measured using VarioscanLUX (Thermo Fischer Scientific) plate reader. To investigate the effect of IND on TLR expression, DLD-1 cells were seeded in 6-well plates at a density of 1.5*10⁵ cells/well and grown for 72 h. After incubation, cells were treated with 1 µM and 10 µM IND or its vehicle (0.1% DMSO), and cells were harvested 0 and 24 h after IND or vehicle treatment. The sequences of primers for human TLRs and GAPDH are listed in Supplementary Table 2. All experiments were performed in duplicate and repeated three times independently.

## 2.11. Data analysis

The bioinformatic analysis was performed essentially as described by Mansour et al. (2020) [55]. Briefly, raw reads were first evaluated for quality using FastQC and MultiQC. Low-quality sequences were then filtered and trimmed with Trimmomatic [56], retaining only sequences with a minimum length of 36 nucleotides and discarding base calls with a Phred score below 20. Taxonomic classification was performed using the SILVA SSU Ref NR 99 database (release 132), which had been pre-processed and indexed by Kraken2 [57] using a k-mer length of 31. The final microbiota composition was estimated using Bracken [58]. Subsequently, a Bayesian-multiplicative zero replacement was applied via the zCompositions package in R [59], followed by a Centered Log-Ratio (CLR) transformation performed with the scikit-bio Python library [60]. Taxa with a median CLR value below 2 were excluded from downstream statistical analyses. Alpha diversity was quantified using the Shannon index, and only taxa supported by at least 50 reads were considered for

<table><caption>**Table 3**

Biofilm damage score of the small intestinal microbiota.</caption><thead><tr><th rowspan="2"></th><th colspan="4">Biofilm damage score</th></tr><tr><th>0</th><th>1</th><th>2</th><th>3</th></tr></thead><tbody><tr><td>Biofilm distance to epithelia</td><td>distant</td><td>close</td><td>contact</td><td>numerous contacts</td></tr><tr><td>Biofilm density</td><td>mild</td><td>scattered</td><td>low</td><td>few</td></tr><tr><td>Bacterial translocation into lamina propria</td><td>none</td><td>few</td><td>numerous</td><td>extensive</td></tr><tr><td>Immune cell transmigration into the biofilm</td><td>none</td><td>mild</td><td>dense</td><td></td></tr><tr><td>Bacteria morphology</td><td>more than 5 morphotypes</td><td>&lt;4 morphotypes</td><td>&lt;3 morphotypes</td><td>Predominance of one morphotype</td></tr></tbody></table>
<figure><img src="image_4.png" /><figcaption>**Fig. 1.** High-dose indomethacin (IND, 20 mg/kg) induces severe inflammation and tissue damage in the small intestine of rats. **A)** The experimental protocol. Rats received vehicle (VEH) or IND and were euthanized at different time points. **B-E)** The effect of IND on animal weight, small intestinal length, histologic injury and expression of different inflammatory mediators and cytokines. The results are expressed as the mean ± SEM, circles represent individual data points. For statistical analysis two-way repeated measures ANOVA (weight changes), one-way ANOVA (intestine length, inflammatory genes and proteins) and Kruskal-Wallis test (histologic score) were used, followed by Fisher's LSD and uncorrected Dunn's tests, respectively. *p* < 0.05, **p** < 0.01, ***p** < 0.001 compared to vehicle (VEH)-treated group.</figcaption></figure>

further analysis. The overall microbiota composition was visualized by Principal Component Analysis (PCA), and differences in community structure were statistically assessed using Permutational Multivariate Analysis of Variance (Permanova). Differential abundance testing was conducted with the Wilcoxon rank-sum test, applying the Benjamini-Hochberg method [61] to control the false discovery rate (FDR).

Statistical analysis of the data was performed with one-way ANOVA or two-way ANOVA, followed by Fisher's LSD post hoc test, or in case of nonparametric values Kruskal-Wallis test followed by uncorrected Dunn's post hoc test. Outliers detected by Grubb's test were excluded from the analyses. A probability of *p* < 0.05 was considered statistically significant.

Associations between TLRs, AMPs, bacterial abundances and inflammatory markers were investigated as described in previous works [13,50,62]. Briefly, two correlation analyses were performed across all the samples of the first (acute enteropathy induced by a single high dose of IND) and the second (enteropathy induced by repeated administration of lower doses of IND) study separately by calculating Spearman's rank correlation coefficients (Rho-values) and corresponding *p*-values for pairs of the investigated parameters (first study: TLRs vs.

inflammatory markers, TLRs vs. AMPs, inflammatory markers vs. AMPs, TLRs and AMPs vs. bacterial abundances, inflammatory markers vs. bacterial abundances; second study: TLRs vs AMPs). Rho-values were visualized on separate heat maps for each comparison marking correlations considered to be significant according to either the uncorrected *p*-value or the *p*-value corrected by the Benjamini-Hochberg procedure (q-value) [60]. The correlation analyses were conducted using version 4.3.3 of the R software package [63].

## 3. Results

### 3.1. IND-induced acute enteropathy differentially regulates intestinal TLR and AMP expressions

First, we aimed to assess the changes in the gene expression of TLRs and AMPs and their correlation with inflammation in severe acute enteropathy. To this end, we treated rats with a single high dose of IND (20 mg/kg), and collected the small intestine at five different time points, from 6 to 72 h post-dose (Fig. 1A). The severity of IND-induced enteropathy gradually increased over time and was associated with weight
<figure><img src="image_5.png" /><figcaption>A</figcaption></figure>

<figure><img src="image_6.png" /><figcaption>C</figcaption></figure>

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<figure><img src="image_13.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_14.png" /><figcaption>D</figcaption></figure>

<figure><img src="image_15.png" /><figcaption>E</figcaption></figure>

<figure><img src="image_16.png" /><figcaption>**Fig. 2.** High-dose indomethacin (IND, 20 mg/kg) induces characteristic changes in the intestinal gene expression of TLRs and AMPs, which are differently associated with inflammation. **A** and **C**) The results are expressed as the mean + SEM, circles represent individual data points. For statistical analysis one-way ANOVA was used, followed by Fisher's LSD test. *p* = 0.05, **p** = 0.01, ***p** = 0.001 compared to vehicle (VEH)-treated group. **B**, **D** and **E**) Heat maps of Spearman's correlation coefficients between TLRs, AMPs and tissue injury markers in animals treated with VEH or IND 20. *p*-values were corrected by the Benjamini–Hochberg procedure yielding *q*-values. White asterisk: uncorrected *p* < 0.05, black asterisk: *q* < 0.05.</figcaption></figure>

loss (Fig. 1B, left), shortening of the intestine (Fig. 1B, right), and various histological changes such as erosions, ulcers and inflammatory infiltrates, which could be detected as early as 6 h in some animals (Fig. 1C). Mucosal inflammation was also indicated by the elevation of several inflammatory mediators and cytokines, such as MPO, PTX3, COX2, IL1β and TNFα, all of which peaked at 48 h (Fig. 1D, E). In contrast, the gene expression of the anti-inflammatory IL10 gradually decreased.

Next, we assessed the gene expression of the six bacterial TLRs in the small intestine of enteropathic animals. We found that IND-induced acute enteropathy was associated with different changes in TLR transcript levels. The gene expression of TLR2 increased gradually and showed a significant increase at 24 and 48 h (Fig. 2A), similar to that of several inflammatory mediators. TLR1, a co-receptor of TLR2, increased similarly to TLR2 and peaked at 48 h. The strong association between inflammation and TLR1/2 expression was also confirmed by Spearman's correlation analysis (Fig. 2B). In contrast to TLR1, the expression of TLR6, the other heterodimerizing partner of TLR2, remained unchanged during the observation period. Although TLR4 and TLR9 mRNA expressions tended to increase at some time points, they showed high inter-individual variability and were not significantly different between control and enteropathic animals (Fig. 2A). There was no clear association between inflammation and the expression of TLR4, TLR6 or TLR9, except that COX-2 correlated positively with TLR4 (Fig. 2B). Finally, TLR5 was downregulated in enteropathic animals and showed a strong inverse correlation with inflammation (Fig. 2A, B).

With regard to AMPs, the expression of genes encoding rat α-defensin 5 (Defa5) and β-defensin 2 (Defb2) was highly variable and did not increase significantly compared to the control group (Fig. 2C), but they both showed a strong positive correlation with TLR4 expression (*q* < 0.001, Fig. 2D). In contrast, the gene encoding rat cathelicidin (Camp) increased significantly as early as 12 h after IND administration and

correlated positively with inflammation, TLR1, TLR2 and TLR4, and negatively with TLR5 (Fig. 2D, E).

## 3.2. Severe but not mild enteropathy caused by repeated IND exposure is associated with similar changes in TLRs and AMPs as acute enteropathy induced by a single high-dose of IND

We next sought to determine whether the observed changes in TLRs and AMPs could be recapitulated by treating animals repeatedly with lower doses of IND over a longer period of time. To this end, rats received 2 or 4 mg/kg IND twice a day for 14 or 6 days, respectively (Fig. 3A). 4 mg/kg IND given repeatedly caused severe intestinal damage and inflammation similar to 20 mg/kg IND given once in the first study, whereas repeated administration of 2 mg/kg IND caused only mild inflammation (Fig. 3B), and enteropathy was mainly reflected by changes in hematological parameters, such as reduced hemoglobin levels and increased neutrophil to lymphocyte ratio (Fig. 3C and Supplementary Table 3).

Severe enteropathy induced by repeated administration of 4 mg/kg IND caused changes in TLR and AMP expression similar to 20 mg/kg IND given once. TLR1, TLR2 and CAMP genes were upregulated and TLR5 was downregulated, whereas the expressions of TLR4, TLR6 and TLR9 were not significantly different between control and enteropathic animals (Fig. 3D, E). Interestingly, in contrast to acute enteropathy, prolonged exposure to IND also resulted in increased α-defensin 5 expression. Correlation analysis confirmed the positive and negative correlation of CAMP with TLR2 and TLR5, respectively, and the association between TLR4 and β-defensin 2 expressions (Fig. 3F). In addition, α-defensin 5 correlated with both TLR1 and TLR2 in this cohort.

In contrast to severe enteropathy, chronic mild enteropathy had only a mild and rather inhibitory effect on TLR expressions, in particular on those of TLR1 and TLR5 (Fig. 3D). As for the AMPs, CAMP tended to
<figure><img src="image_17.png" /><figcaption>Fig. 3. Repeated administration of indomethacin (IND) at doses of 2 and 4 mg/kg induces mild and severe enteropathy, respectively, with different changes in TLR and AMP expression. A) The experimental protocol. Rats received vehicle (VEH) or IND and were euthanized after 6 or 14 days. B-E) The effect of chronic IND treatment on small intestinal length, expression of different inflammatory mediators and cytokines, hematological parameters, and TLR and AMP expression. The results are expressed as the mean ± SEM, circles represent individual data points. For statistical analysis one-way ANOVA was used, followed by Fisher's LSD test. n = 5-8/group. \*p < 0.05, \*\*p < 0.01, \*\*\*p < 0.001 compared to vehicle (VEH)-treated group. F) Heat map of Spearman's correlation coefficients between TLRs and AMPs in animals treated repeatedly with VEH or lower doses of IND (2 and 4 mg/kg). p-values were corrected by the Benjamini-Hochberg procedure yielding q-values. White asterisk: uncorrected p < 0.05, black asterisk: q < 0.05.</figcaption></figure>

increase, while defensins did not change in mild enteropathy.

Taken together, severe enteropathy caused an upregulation of TLR2, TLR1 and CAMP, and also of α-defensin 5 by repeated drug exposure, whereas TLR5 decreased in both mild and severe enteropathy. TLR4, TLR6 and TLR9 did not change significantly in either case, but TLR4 expression correlated with that of the defensins.

### 3.3. Changes in distinct TLRs and AMPs are differently associated with inflammation and dysbiosis

As TLRs, AMPs and gut bacteria are closely interrelated, we next characterized the small intestinal luminal microbiota of control animals and those treated with a single 20 mg/kg dose of IND by 16S rRNA gene sequencing. In line with literature data [11], IND had only minor effect on the Shannon index, an indicator of α-diversity (Fig. 4A), but caused marked changes in the overall microbiota composition. As shown by the PCA score plot (Fig. 4B), IND-induced dysbiosis developed gradually, being detectable as early as 6 h post-dose, and was characterized by a marked shift from Gram-positive to Gram-negative organisms (Figs. 4C-4E) (Permanova p value <0.001 between VEH and IND-treated groups). More specifically, the proportion of the Gram-positive *Firmicutes* decreased from 96.7 % to 57.1 % within 72 h, mainly due to a decrease in genera such as *Lactobacillus*, *Limosilactobacillus*, *Ligilactobacillus*, *Romboutsia* and *Turicibacter*. In parallel, the *Enterococcus* genus, as well as several genera belonging to *Proteobacteria* and *Bacteroidota*, such as *Escherichia-Shigella* and *Bacteroides* increased significantly in enteropathy. Several Gram-negatives, in particular, those belonging to the *Gammaproteobacteria*, showed a strong positive association with inflammation, whereas several Gram-positives correlated inversely with

that (Supplementary Figs. 1 A, 1B).

Next, we sought to determine whether IND-induced changes in small intestinal TLRs and AMPs were associated with specific microbiota signatures. To this end, we have comprehensively evaluated the correlation between the expression of TLRs, AMPs and the proportion of all identified bacterial families and genera in rats with acute enteropathy induced by 20 mg/kg IND. Changes in TLR1, TLR2, TLR5 and CAMP expression correlated with the luminal abundance of several bacterial families and genera, although only the positive correlation between TLR2 and the genus *Vibrio* remained significant when individual p-values were corrected for multiple testing (Fig. 5 and Supplementary Fig. 2). Nevertheless, the observed associations between TLRs, AMPs and bacteria were likely driven mainly by inflammation rather than direct TLR-bacterial interactions. Namely, inflammation-associated bacteria, such as members of the *Gammaproteobacteria* or *Enterococcus*, showed positive correlation with TLR1, TLR2 and CAMP, and negative correlation with TLR5, whereas the opposite was true for several Gram-positives (e.g. *Lactobacillaceae*, *Erysipelotrichaceae*), which decreased in inflammation. Conversely, TLRs and AMPs whose expression was less or not dependent on acute intestinal inflammation, such as TLR4, TLR6, TLR9 and defensins, showed only little or no correlation with bacteria. More specifically, TLR6 and TLR9 showed only negative correlation with *Pectobacteriaceae* and *Bifidobacteriaceae*, respectively, whereas α-defensin 5 correlated positively with the *Lachnospiraceae* family.

In an attempt to identify direct associations between TLRs, AMPs and bacteria without the confounding influence of inflammation, we performed a subgroup analysis on animals euthanized at the earliest time point, 6 h after treatment with 20 mg/kg IND. This allowed us to differentiate animals with uninflamed intestinal mucosa from those
<figure><img src="image_18.png" /><figcaption>**Fig. 4.** High-dose indomethacin (IND, 20 mg/kg) causes marked changes in small intestinal microbiota composition. A) The effect of IND on small intestinal bacterial diversity, estimated by the Shannon index (Kruskal-Wallis test, followed by uncorrected Dunn's tests, *n* = 8/group). B) Principal component analysis (PCA) score plot comparing the small intestinal microbiota composition of rats treated with either vehicle (VEH) or IND. C-E) IND-induced changes in the relative abundance of bacterial phyla and major families and genera. The differential abundance tests were performed using ANCOM with Holm-Bonferroni correction for multiple comparison, *n* = 8/group. \*q < 0.05, \*\*q < 0.01 compared to VEH-treated group.</figcaption></figure>

having already mild inflammation, as indicated by higher tissue levels of inflammatory markers such as IL-1β (Fig. 6A). In addition, some animals that only partially matched those with detectable inflammation, showed small intestinal dysbiosis with a shift towards Gram-negative bacteria (*Verrucomicrobiota*, *Bacteroidota*, *Proteobacteria*) and *Actinobacteriota* (Fig. 6B). These changes were attributed to changes in the *Akkermansiaceae*, *Muribaculaceae*, *Enterobacteriaceae* and *Bifidobacteriaceae* families, respectively. When we compared IND-treated animals with and without dysbiosis, we found mild inflammation in non-dysbiotic animals, but more severe inflammation and also higher CAMP levels in animals with dysbiosis (Fig. 6C), suggesting that inflammation and CAMP upregulation were exacerbated, but not primarily caused by changes in the microbiota. When we compared TLR and defensin expression in animals with and without inflammation, and also in animals with and without dysbiosis (Fig. 6D, E), we found that the changes in TLR1, TLR2 and TLR5 expression did not depend on the presence of dysbiosis, but showed a strong association with inflammation (Fig. 6D). In contrast, the expression of TLR4, α-defensin 5 and β-defensin 2 showed a strong association with dysbiosis in early enteropathy, in particular with the expansion of the *Enterobacteriaceae* family (Fig. 6E). There was no difference in TLR6 and TLR9 expression between animals with and without dysbiosis or inflammation (not shown).

To investigate the potential direct effects of IND on epithelial TLR expression, experiments were conducted using the human colon cancer cell line DLD-1. This cell line expresses both TLR2 and TLR5 [64,65] and, to a lesser extent, TLR4 [66]. We found that 24 h of treatment with IND at 1 µM and 10 µM concentrations had no significant time- or concentration-dependent effect on the expression of any of the tested TLRs (Supplementary Fig. 3). These results suggest that IND does not directly affect the expression of TLRs in epithelial cells and that changes

in TLRs in enteropathy predominantly occur in non-epithelial lamina propria cells and/or as part of the complex immune response associated with enteropathy and dysbiosis. Finally, based on our finding that α-defensin 5 expression was more dependent on dysbiosis than inflammation in early enteropathy, we hypothesized that the increased α-defensin 5 expression after repeated IND administration but not in acute enteropathy was due to differences in the microbiota profile of the animals. When we compared IND-induced changes in the ileal microbiota in the two models, we found a marked difference in the levels of *Lachnospiraceae*. It showed a modest, transient decrease in acute enteropathy caused by 20 mg/kg IND at 48 h, but tended to increase at 72 h, and showed a significant increase at 6 days of 4 mg/kg IND treatment (Fig. 7A and B). As mentioned above, this was the only bacterial family that correlated positively with α-defensin 5 expression in acute severe enteropathy, and we found a similar positive correlation between α-defensin 5 and *Lachnospiraceae* in animals treated with IND for 6 days (*p* = 0.02, Fig. 7B).

### 3.4. IND induced enteropathy causes mild damage to the mucosal microbiota biofilm structure

Although the importance of mucosa-adherent microbiota biofilms in bacterial-host interactions is well recognized [45], and certain NSAIDs have been shown to have anti-biofilm activity in vitro [49], little is known about the effect of NSAIDs on intestinal biofilms in the small intestine, and their role in enteropathy. Therefore, we assessed biofilm integrity in Methacarn fixed ileal samples of VEH- and IND-exposed rats by fluorescent in situ hybridization. As IND enteropathy is characterized by multiple transmural ulcers which are sharply delineated from non-ulcerated areas of the mucosa (Fig. 8A), we focused on areas with
<figure><img src="image_19.png" /><figcaption>**Fig. 5.** Changes in TLRs and AMPs caused by high-dose indomethacin (IND) are differently associated with changes in intestinal bacteria. Heat map of Spearman's correlation coefficients between TLRs, AMPs and the small intestinal luminal abundance of identified bacterial families in animals treated with VEH or IND 20. p-values were corrected by the Benjamini-Hochberg procedure yielding q-values. \*uncorrected *p* < 0.05.</figcaption></figure>

maintained mucosal structure to assess IND-induced changes in biofilm integrity. We found that the biofilm of IND-treated animals was significantly different compared to control animals, mainly due to a reduced biofilm density at 6 and 12 h post-dose (Fig. 8B, C). However, we did not observe bacterial adhesion to epithelium, bacteria clustering into inner mucus layer, or even bacterial translocation into lamina propria, indicating that IND had only a mild damaging effect on biofilm organization.

Because of the early onset of damage and its partial recovery at later time points we hypothesized that IND has a direct damaging effect on the microbiota biofilm. Therefore, we reconstituted mucosa-associated rat ileal microbiota and cultured it under its natural biofilm phenotype under anaerobic conditions for 72 h. Mature biofilms were then exposed to different concentrations of IND for 24 h. IND up to 10 µM had no effect on biofilm biomass (cells and total matrix-associated content), on bacterial viability within the biofilm, or on the release of live biofilm-dispersed bacteria (Fig. 8D), suggesting that the mild damaging effect of IND on microbiota biofilm structure observed in vivo is secondary to

enteropathy-induced changes in the host. Although there was no clear association between mucosal injury and changes in biofilm integrity, interestingly the biofilm density damage score showed a negative correlation with TLR5 expression (*p* = 0.04) (Supplementary Fig. 4) and also tended to correlate positively with the levels of PTX3 (*p* = 0.07), IL-1β and CAMP (*p* = 0.09 for both).

## 4. Discussion

Here we show that NSAID enteropathy has differential effects on the expression of six bacterial TLRs and three major intestinal AMPs. TLR2, TLR1 and CAMP expression increased in the inflamed gut, TLR5 expression decreased, whereas TLR6 and TLR9 did not show significant changes in enteropathy. Although TLR4 expression showed only subtle differences limited to a subset of animals, it correlated positively with defensin expression, and also with the abundance of the *Enterobacteriaceae* family in the early phase of enteropathy. We also show that, in
<figure><img src="image_20.png" /><figcaption>**Fig. 6.** The expression of TLRs and AMPs is differently associated with inflammation and dysbiosis in the early stage of acute enteropathy. A and B) 6 h after treatment with indomethacin (IND, 20 mg/kg) a part of the animals had mild inflammation (marked with green data points and green circles in the PCA score plot) and/or significant small intestinal dysbiosis (black data points). C-E) The expression of inflammatory markers, TLRs and AMPs in VEH-treated and different sub-groups of IND-treated animals. The results are expressed as the mean ± SEM, circles represent individual data points. For statistical analysis one-way ANOVA was used, followed by Fisher's LSD test. *p* < 0.05, compared to VEH-treated group, \*p* < 0.05 compared to IND-treated animals with low IL-1β levels.</figcaption></figure>

<figure><img src="image_21.png" /><figcaption>**Fig. 7.** The level of the *Lachnospiraceae* family shows time-dependent changes in IND enteropathy and correlates with the expression of α-defensin 5. A) The relative abundance of the *Lachnospiraceae* family in animals with acute enteropathy, derived from the data of Illumina sequencing. The results are expressed as the mean ± SEM, circles represent individual data points. *p* < 0.05 compared to VEH-treated group. B) Relative changes in *Lachnospiraceae* levels (determined by qPCR and normalized for the universal bacterial primer) and their correlation with α-defensin 5 expression. \*p* < 0.05 compared to VEH-treated group.</figcaption></figure>

addition to marked changes in luminal microbiota composition, IND in vivo treatment caused mild damage to the mucosal microbiota biofilm structure, which does not seem to be due to a direct effect of IND on microbial biofilms, but rather is a consequence of changes in host mucosal mediator expression.

Although the pivotal role of TLR-bacterial interactions in maintaining gut homeostasis is well recognized [33,67] and dysregulated TLR signalling has been linked to inflammatory gut disorders such as IBD [68], there are limited and inconclusive data on whether and how the expression of TLRs is altered in NSAID enteropathy. Therefore, the primary aim of the present study was to determine the gene expression pattern of all six TLRs mainly responsible for bacterial detection in both acute and chronic IND enteropathy, and to assess whether these changes are related to inflammation and dysbiosis. In addition, as increased secretion of AMPs is a pivotal part of the complex defense response initiated by TLRs against intestinal pathogens [21,69], we also assessed the expression profile of three major intestinal AMPs, α-defensin 5, β-defensin 2 and cathelicidin [70], in parallel with that of the TLRs.

We found that severe intestinal inflammation caused by a single high

dose of IND and also by repeated administration of lower doses of IND was associated with a modest upregulation of the ileal TLR2 gene. A similar 1.5-fold increase in the ileal TLR2 expression in IND-treated rats was also reported by Teran-Ventura et al. [41], although not by other groups [40,43]. These discrepancies are likely due to differences in the protocols used, resulting in different intestinal inflammation, on which TLR2 upregulation is strongly dependent. In acute enteropathy, TLR2 increased mainly at time points when inflammation peaked, whereas in chronic mild enteropathy it did not increase but rather tended to decrease, similar to what others have reported in mild gut inflammation caused by other triggers [26]. Nevertheless, the overexpression of TLR2 in severe IND enteropathy is in harmony with the increased TLR2 expression reported in diclofenac-treated rats [42], colitic mice [71], as well as in patients with IBD [29,31] and diverticulitis [29], suggesting that increased TLR2 signalling contributes to gut inflammation induced by various triggers, including NSAIDs.

An important characteristic of TLR2 is that it can form a heterodimer with either TLR1 or TLR6, which also determines its ligand specificity [72,73]. Our finding that TLR1 increased in parallel with TLR2, whereas
A

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C

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D

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<figure><img src="image_27.png" /><figcaption>**Fig. 8.** Indomethacin (IND) damages the small intestinal mucosal biofilm in vivo, but not ex vivo. A) A representative histological image of severe IND enteropathy, characterized by multiple transmural ulcers (denoted by arrows) separated by non-ulcerous areas (scale bar: 1 mM, haematoxylin and eosin staining). B) Changes in gut microbiota biofilm organization were recorded by an observer unaware of treatments. The results are expressed as the mean ± SEM, circles represent individual data points. n = 4–5/group. For statistical analysis Kruskal-Wallis test was used, followed by uncorrected Dunn's tests. C) Representative images of Methacarn fixed small intestinal sections. All bacterial cells were labeled with the universal probe Eub338 for fluorescent in situ hybridization (red) and wheat germ agglutinin was used to stain the polysaccharides-rich mucus layer (green). All images were counterstained with the nuclear stain, 4',6-diamidino-2-phenylindole DAPI (blue). D) Multispecies anaerobic biofilms were generated by culturing microbiota of small intestinal tissues collected from five rats. Biofilms were then exposed to various concentrations of IND (0.01–10 µM). IND had no effect on total biofilm biomass (Safranin-O assay), on biofilm bacteria survival (rezasurin metabolic assay) and had no significant impact on the rate of biofilm dispersal. The results are expressed as the mean ± SEM, n = 5–8/group.</figcaption></figure>

TLR6 did not show significant changes, suggests that IND-induced inflammation is associated with the overexpression of the TLR2/TLR1, but not the TLR2/TLR6 receptor complex. This is consistent with previous findings linking the TLR2/TLR1, but not the TLR2/TLR6 dimer to pro-inflammatory responses in the intestinal mucosa [74]. It also resonates with the ligand recognition profile of TLR2/TLR1, which mainly binds triacyl lipopeptides, a typical component of Gram-negative bacteria [75]. Although we found indeed several significant correlations between TLR2/TLR1 expression and the luminal abundance of Gram-negatives in enteropathic animals, these are not necessarily caused by direct interactions between TLR2/TLR1 and bacteria, given the strong association of both with inflammation. In fact, our finding that TLR2/TLR1 expression was increased to similar levels in animals with and without dysbiosis in the early phase of enteropathy, but was significantly higher in animals with higher IL-1β levels regardless of their gut microbiota composition, suggests that the upregulation of TLR2/TLR1 was not primarily caused by an increase in Gram-negative bacteria.

A part of the TLR2-induced innate immune response against bacteria is the increased secretion of several AMPs, such as α-defensins [22], β-defensin 2 [21] and CAMP [76,77]. Of these, the expression pattern of CAMP was most similar to that of TLR2. It increased gradually with

disease progression and correlated positively with most of the inflammatory mediators measured. This finding is in agreement with the inflammation-dependent overexpression of intestinal CAMP in IND- and dextran sulfate sodium-treated mice [23,78], as well as in IBD patients [24,79], and highlights the importance of this peptide in the control of various inflammatory intestinal disorders. As both CAMP and TLR2 were increased in the inflamed gut, there was also a positive correlation between their expression in all animals with severe enteropathy. Although we cannot exclude the possibility that CAMP expression was directly regulated by TLR2, the findings that CAMP but not TLR2 levels were influenced by dysbiosis in early enteropathy and also tended to increase in chronic mild enteropathy suggest that CAMP expression was not dependent on TLR2 signalling.

Although a characteristic feature of NSAID-induced intestinal dysbiosis is the marked expansion of *Gammaproteobacteria* [11], and several members of this class are flagellated and recognized by TLR5 [80], little is known about the role of TLR5 in enteropathy. Our study shows that TLR5 expression changed in the opposite direction to that of TLR2 and TLR1, and was significantly reduced in enteropathic animals. This was consistently found in all IND-treated groups throughout the study, including animals with both severe and mild enteropathy, resulting in a
strong negative correlation between TLR5 expression and inflammation. Notably, a similar downregulation of TLR5 was found in the mucosa of colitic mice [81] and also in IBD patients [82], although there are also data showing that TLR5 remains unchanged or even increases in IBD [28,29]. Since both gut bacteria [26] and inflammatory cytokines [81,83] can control TLR5 expression, and since *Gammaproteobacteria* increased in parallel with inflammation, it is difficult to dissect the contribution of each to the downregulation of TLR5. However, the finding that TLR5 expression decreased in early enteropathy in both dysbiotic and non-dysbiotic animals suggests that this was caused by inflammation rather than changes in gut bacteria. In fact, if we consider that TLR5 deficiency has been linked to dysbiosis and reduced ability to manage *Proteobacteria* [35,84], our result that TLR5 downregulation preceded the overgrowth of *Proteobacteria* in some animals suggests that it was a cause rather than a consequence of dysbiosis. Based on these results we propose that an early decrease in TLR5 signalling contributes to NSAID-induced enteropathy and dysbiosis, and that administration of a TLR5 ligand at an early stage of enteropathy may provide protection against it, similar to gut injury of other origins [35].

Based on the existing literature, both lipopolysaccharide (LPS), a major component of the outer membrane of Gram-negatives, and its receptor TLR4 play a crucial role in the pathogenesis of enteropathy. NSAID-induced small intestinal injury is greatly attenuated in TLR4-mutant and TLR4-/- mice [37,40], whereas LPS treatment increases injury in antibiotic-treated animals [85]. Interestingly, it is less clear whether enteropathy is also associated with changes in TLR4 expression. There is some evidence that intestinal TLR4 is overexpressed in NSAID enteropathy [39,40], but there are also studies in which small intestinal TLR4 gene and protein expression remained unchanged after IND treatment [37,43], or showed only mild changes compared to the large intestine [41]. In the present study, TLR4 expression did not change significantly over the course of enteropathy, but there was a clear upregulation of TLR4 in a subset of IND-treated animals, independent of disease severity, and this was associated with higher α-defensin 5 and β-defensin 2 levels. We could not identify a specific reason for the differential expression of TLR4 in animals with moderate to severe enteropathy, as it was not related to the severity of inflammation, the proportion of Gram-negative bacteria or the degree of mucosal biofilm damage. However, in the early stage of enteropathy, increased TLR4 and defensin expression was highly dependent on the luminal abundance of Gram-negatives, particularly the *Enterobacteriaceae* family. These results suggest that TLR4 has a more complex role in enteropathy than previously appreciated, and in addition to its well-documented role in mediating intestinal inflammation, it also promotes mucosal defense by increasing the expression of defensins. This is consistent with previous reports that TLR4 can modulate α- and β-defensin expression in the intestine [21,22] and also with the dual role of TLR4 in colitis, where it can both contribute to intestinal homeostasis and epithelial repair [33,86] and exacerbate inflammation [87,88]. However, it is not yet clear why TLR4 expression did not increase in parallel with the increasing abundance of *Enterobacteriaceae* in later stages of the disease. We speculate that this may be related to the concomitant activation of TLR2, which can suppress TLR4 expression in enteropathy [38], but this requires further investigation.

Although α-defensin 5 expression showed high inter-individual variability in acute enteropathy and increased in only a subset of animals regardless of the degree of inflammation, it increased significantly in severe chronic enteropathy. This finding contrasts with previous studies linking acute IND-induced enteropathy to Paneth cell deficiency and reduced α-defensin 5 expression in mice [89,90]. Although the reason for this contradiction is not clear and may also depend on species differences, we propose that it may be caused, at least in part, by differences in gut microbiota composition, in particular *Lachnospiraceae* levels. The influence of NSAID enteropathy on *Lachnospiraceae* is inconsistent [11] and, as our present data show, is also likely to depend on the duration of enteropathy. We found that it decreased during the

first two days of acute enteropathy, but increased at 72 h and also after a 6-day treatment, and showed a positive correlation with α-defensin 5 expression in both acute and chronic enteropathy. Given that several bacteria belonging to the *Lachnospiraceae* produce butyrate [91], a short-chain fatty acid that has been shown to induce α-defensin 5 gene expression and secretion [92,93], it can be assumed that changes in the *Lachnospiraceae* family are among the factors that regulate α-defensin 5 expression in enteropathy.

Finally, although the widespread use of sequencing technologies has greatly increased our knowledge of NSAID-induced changes in gut microbiota composition, very little is known about the effect of these drugs on mucosal microbiota biofilms and their role in the pathogenesis of enteropathy. The effect of NSAIDs on bacterial biofilms has mainly been investigated in vitro, with results suggesting that certain NSAIDs, such as acetylsalicylic acid, diclofenac or ibuprofen, may have anti-biofilm activity via inhibition of quorum sensing or production of extracellular polysaccharides required for biofilm formation [49]. Here, we investigated for the first time, whether IND treatment had any effect on the integrity of the mucosal biofilm in the rat small intestine and importantly, we found that it was compromised in non-ulcerous areas of the mucosa in enteropathy, particularly in the early stages of the disease. The damage was mild and transient, but did not seem to allow luminal bacteria to come into direct contact with the epithelium at this stage. This finding suggests that changes in overall commensal biofilm density were not sufficient to trigger mucosal bacterial invasion and disease development, and were a consequence rather than a cause of enteropathy. This is supported by the findings that ex vivo cultured ileal biofilms exposed to IND did not show any changes in biomass, bacterial viability or bacterial dispersion. That said, pathobiont taxa may still undergo spatial repositioning within the small intestinal biofilm in the context of NSAID enteropathy. Although there was no clear association between the extent of biofilm damage and the expression of any of the individual host-related factors measured, the finding that biofilm density damage scores correlated negatively with TLR5 expression and tended to correlate positively with PTX3, IL-1β and CAMP suggests that biofilm alterations were likely caused by factors released from the inflamed mucosa that remain to be identified. These findings suggest that changes in microbial density may have a localized and biologically significant impact related to the gut transcriptome. One could hypothesize that this transient response could, in turn, shape the outcome of subsequent events, including infectious or inflammatory challenges.

Although our study identified characteristic changes in intestinal expression of TLRs and AMPs in response to IND treatment, we should emphasize that we only used male animals, as in most studies focusing on enteropathy [12,13,37-43,50]. However, there is some evidence that enteropathy-associated mucosal injury, immune responses, and dysbiosis may differ to some extent between males and females [94,95], therefore, validation of our findings in females is warranted. Additionally, we employed 16S sequencing, a cost-effective and simple method for profiling gut microbiota composition, but it has limitations compared to shotgun metagenomics. For instance, it provides limited resolution at the species or strain level and offers little direct information about microbial functional potential or gene content [96]. Therefore, while our findings on indomethacin-induced intestinal dysbiosis align with existing literature [4,11], further research combining shotgun metagenomics and functional gene profiling is required to determine the specific microbial taxa or functions involved in enteropathy and establish the link between microbial changes and host TLR/AMP expression and biofilm responses. Finally, our association analysis incorporated only tissue inflammation, a key feature of enteropathy. Mild inflammation is evident in the early stages of enteropathy, but it would be worthwhile measuring even earlier events in enteropathy development, such as increased permeability [6], and assessing their associations with TLRs and AMPs. Similarly, given that a large proportion of the population uses NSAIDs chronically [1,2], it is particularly important to assess changes in TLRs and AMPs after long-term NSAID use and during tissue
<figure><img src="image_28.png" /><figcaption>**Fig. 9.** Associations between toll-like receptors (TLRs), antimicrobial peptides (AMPs), inflammation and intestinal bacteria in indomethacin (IND)-induced early enteropathy. Changes in TLR1, TLR2, TLR5 and CAMP expression were mainly associated with inflammation, whereas changes in TLR4 expression and defensins were mainly dependent on gut dysbiosis. In addition, there was an unexpected association between mucosal microbiota biofilm changes in density and inflammatory gene expression.</figcaption></figure>

regeneration, and the potential benefits of interventional strategies
targeting TLRs.

In conclusion, we show that IND enteropathy is associated with
characteristic changes in the intestinal TLR and AMP expression pattern.
TLR1, TLR2 and CAMP are upregulated, TLR5 is downregulated, TLR4
and defensins show only modest and highly variable changes, whereas
TLR6 and TLR9 are not significantly altered in enteropathic rats.
Although these changes are associated with both tissue inflammation
and gut dysbiosis, two closely related factors, dissection of these factors
in animals with early enteropathy suggests that changes in TLRs and
AMPs are differentially dependent on them (Fig. 9). Changes in TLR1,
TLR2, TLR5 and CAMP expression were strongly associated with
inflammation, whereas changes in TLR4 expression and defensins were
more dependent on gut dysbiosis in early disease. In addition, we show
for the first time that IND enteropathy is associated with mild changes in
the integrity of the mucosal microbiota biofilm, likely as a consequence
of inflammation-induced changes in the intestinal mucosa. Our findings
provide new insights into the complex interactions between TLRs,
AMPs, intestinal inflammation and bacteria, and may help to identify
new strategies for the treatment of NSAID enteropathy.

CRediT authorship contribution statement

**Arezoo Haghighi:** Writing – original draft, Visualization, Investi-
gation, Formal analysis, Conceptualization. **András S. Tóth:** Writing –
review & editing, Investigation. **Zsuzsanna O. Demeter:** Investigation.
**Barbara Hutka:** Investigation. **Anna Zsidai:** Investigation. **Lili Len-**
**gyel:** Investigation. **Samaneh Haghighi:** Investigation. **Melissa**
**Pannier:** Investigation. **Guillaume Le Cosquer:** Investigation. **Elodie**
**S. Meunier:** Investigation. **Bence Ágg:** Writing – review & editing,
Formal analysis. **Nóra Makra:** Formal analysis. **Eszter Ostorházi:**
Formal analysis. **Balázs Ligeti:** Writing – review & editing, Formal
analysis. **Krisztián Kovács:** Formal analysis. **Agnes Kelemen:** Formal
analysis. **Anna Jakab:** Formal analysis. **Gerda Wachtl:** Supervision,
Investigation. **Gábor Kökény:** Writing – review & editing. **Dóra Szabó:**

Writing – review & editing. **Péter Ferdinandy:** Writing – review &
editing, Funding acquisition. **Jean-Paul Motta:** Writing – review &
editing, Supervision, Funding acquisition, Formal analysis, Conceptu-
alization. **Nathalie Vergnolle:** Writing – review & editing, Supervision,
Funding acquisition, Conceptualization. **Klára Gyires:** Writing – review
& editing, Conceptualization. **Zoltán S. Zádori:** Writing – review &
editing, Writing – original draft, Visualization, Supervision, Funding
acquisition, Conceptualization.

**Declaration of competing interest**

PF is the founder and CEO of Pharmahungary Group, a group of R&D companies.

Acknowledgements

The work was supported by the National Research, Development and
Innovation Office of Hungary (NKFI FK 138842 to ZSZ). Project no. RRF-
2.3.1-21-2022-00003 has been implemented with the support provided
by the European Union (to BA and PF). The research was financed by the
Thematic Excellence Programme (2020-4.1.1.-TKP2020) of the Ministry
for Innovation and Technology in Hungary, within the framework of the
Therapeutic Development and Bioimaging thematic programmes of the
Semmelweis University. This project has received funding from the
HUN-REN Hungarian Research Network and from the Semmelweis
Lendület 2024 grant (to PF). Authors thank the INSERM dotation (to
NV), ANR dysBIOFILM ANR-22-CE14-0041-01 (to JPM) and the na-
tional program "Microbiota" (PTM) from INSERM (to NV). The authors
are grateful for the technical assistance of Sámuel Jenei. Fig. 9. was
created with BioRender.com.

**Appendix A. Supplementary data**

Supplementary data to this article can be found online at https://doi.
org/10.1016/j.lfs.2025.124114.
## Data availability

The Manuscript includes the Data availability statement: Raw sequencing data is deposited in Zenodo at https://zenodo.org/records/ 15012367.

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