Inhibitory effects of Levilactobacillus brevis IBRC-M10790 on apoptosis and inflammation induced by Clostridioides difficile culture supernatant in vitro

Abstract

Clostridioides difficile infection (CDI) is a major cause of healthcare-associated diarrhea that contributes significantly to global morbidity and mortality. Bacterial virulence factors, mostly toxins, play key roles in CDI pathogenesis. Probiotic supplementation is a potential strategy to reduce the adverse effects of C. difficile and support intestinal homeostasis. This study aimed to investigate the inhibitory effects of live Levilactobacillus brevis IBRC-M10790 (LLB) and its membrane vesicles (LBMVs) on apoptosis and inflammation induced by released C. difficile virulence factors in vitro. We employed human colorectal adenocarcinoma Caco-2 and HT-29 cell lines, which are widely used as in vitro models against CDI. Viability and apoptosis of both cell lines were assessed using MTT and Annexin V/PI flow cytometry assays. Anti-inflammatory and anti-apoptotic effects of LLB and LBMVs were investigated following treatment with cell-free supernatants of toxigenic C. difficile RT001 (Tox-S), as well as the culture filtrates of non-toxigenic C. difficile RT084 and ATCC 700057 strains. The expression of apoptosis-related genes (BAX, BCL-2, Caspase-3, Caspase-9) and inflammatory markers (IL-6, IL-8, IL-1β, TNF-α) was measured by RT-qPCR, and cytokine production was analyzed by ELISA. C. difficile Tox-S and culture filtrate significantly reduced cell viability and increased the expression of apoptotic and proinflammatory markers in Caco-2 and HT-29 cells. LLB and LBMVs effectively modulated cell viability, reduced apoptosis, and downregulated the expression and production of inflammatory cytokines in both cell lines after exposure to C. difficile culture supernatants. These findings suggest that LLB and LBMVs could be exploited as potential supplement to the current treatment strategies against C. difficile-induced cellular injury and inflammation.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-025-04975-5.

Introduction

Clostridioides difficile infection (CDI) is the most common cause of healthcare facility-associated diarrhea, which can lead to substantial morbidity, mortality, and economic burden worldwide¹. C. difficile is an obligate anaerobic, sporogenic, toxin-producing bacillus, with clinical manifestations ranging from asymptomatic carriage and mild diarrhea to life-threatening pseudomembranous colitis (PMC), toxic megacolon, and death^2,3. The multifactorial pathogenicity of C. difficile is mediated by several virulence factors. The major exotoxins, namely toxin A (TcdA) and toxin B (TcdB), disrupt the cytoskeleton and tight junctions of intestinal epithelial cells and cause inflammation and tissue damage. Some strains also produce a binary toxin (CDT) that enhances the bacterium colonization and disease severity⁴. In addition, surface layer proteins (SLPs) and cell wall proteins (CWPs) improve adhesion, immune evasion, and biofilm formation, which promote persistence and resistance to host defenses^5–7. Flagellar structures are another components that contribute to bacterial motility and host epithelial interactions, therefore, contribute to CDI pathogenesis⁸.

Currently, antibiotic therapy with vancomycin or fidaxomicin is recommended for the treatment of initial mild to moderate CDI episodes in adults, with metronidazole considered an alternative when first-line options are unavailable⁹. However, treatment failure occurs in approximately 20% of cases, which leads to recurrent CDI (rCDI) that requires other therapeutic strategies^10,11. Notwithstanding the antibiotic effectiveness in treating microbial infections, it is clear that antibiotics by virtue of their suppressive effects on the native gut microbiota, often are the cause of dense colonization of C. difficile in the intestine¹². Moreover, with the emergence of highly virulent types and antibiotic-resistant C. difficile strains associated with severe infections, the successful treatment of this recalcitrant infection has become a major global challenge for human health¹³.

Microbiome-oriented therapeutics, such as probiotics, prebiotics, and fecal microbiota transplantation (FMT), have emerged as promising milestones to restore gut microbiota balance and prevent CDI¹⁴. Probiotics are live microorganisms, which in adequate dosage can normalize the composition and function of the gut microbiome and reduce the risk of pathogen colonization¹⁵. These health-promoting microbes can generate various bioactive compounds with anti-inflammatory and antimicrobial properties¹⁶. Although clinical evidence remains limited regarding the role of probiotics in preventing rCDI, several lactic acid bacteria have shown the potential to inhibit C. difficile virulence by modulating toxin production and inflammatory responses^17–20. Levilactobacillus brevis, which is commonly isolated from fermented foods, plants, and the human intestinal tract, has exhibited several probiotic properties, such as acid and bile resistance, adhesion to intestinal cells, and antimicrobial activities^21,22. By virtue of these features, L. brevis has become enable to maintain homeostasis, improve the integrity of epithelial barriers, and attenuate inflammation²³.

Mounting evidence show that Gram-positive organisms, such as lactic acid bacteria, release membrane vesicles (MVs), which are nanosized lipid bilayer structures that contain proteins, lipids, and nucleic acids²⁴. Recent investigations have demonstrated the therapeutic potential of MVs derived from probiotic bacteria in various disease contexts, such as wound healing²⁵ gastrointestinal inflammation²⁶ and vaccine development against encapsulated pathogens²⁷. These findings provide a scientific rationale for using probiotic-derived MVs as therapeutic agents for intestinal infections like CDI.

In vitro studies using human intestinal epithelial cell lines, such as Caco-2 and HT-29, have demonstrated that exposure to C. difficile toxins triggers apoptotic pathways, which are characterized by caspase activation, mitochondrial dysfunction, and epithelial barrier disruption^28,29. Given the imperative requirement of new anti-clostridial treatments, we investigated the inhibitory effects of L. brevis IBRC-M10790 (LLB) and its derived MVs (LBMVs) on the expression of apoptosis-related genes (BAX, Caspase-3, Caspase-9, and BCL-2), as well as proinflammatory cytokines (IL-6, IL-8, IL-1β, and TNF-α) in human colorectal adenocarcinoma Caco-2 and HT-29 cells treated with cell-free supernatant (Tox-S) of toxigenic C. difficile RT001, as well as the culture filtrate of non-toxigenic C. difficile RT084 and ATCC 700057 strains. RT084 represents a non-toxigenic clinical isolate, whereas ATCC 700057 serves as a well-characterized non-toxigenic reference strain. This experimental design provides a better understanding of the protective potential of LLB and LBMVs against the cytotoxicity of C. difficile-released virulence factors under in vitro conditions.

Materials and methods

C. difficile culture and growth conditions

The three C. difficile strains used in this study, including toxigenic C. difficile RT001 (A⁺B⁺) and the non-toxigenic RT084 (A⁻B⁻) and ATCC 700057 (A⁻B⁻), were obtained from the Department of Anaerobic Bacteriology in the Research Institute for Gastroenterology and Liver Diseases in Tehran, Iran. RT001 was selected as it is among the most prevalent toxigenic strains circulating in Iranian healthcare settings and produces both TcdA and TcdB, which makes it a clinically relevant strain for studying CDI pathogenesis. Additionally, RT084 is a locally isolated, non-toxigenic strain, included to evaluate cellular responses to C. difficile filtrates in the absence of toxins that allow us to distinguish background effects from toxin-mediated clinical strains. All the strains were incubated in cycloserine-cefoxitin-fructose agar (CCFA, Mast Group Ltd., Merseyside, UK) supplemented with 5% sheep blood for 48–72 h under anaerobic conditions (85% N₂, 10% CO₂ and 5% H₂) generated by the Anoxomat^® Gas Exchange System (Mart Microbiology BV, Holland)³⁰.

Preparation of Tox-S and culture filtrate

Tox-S and culture filtrates of C. difficile were obtained according to the previously described procedure with modifications³¹. C. difficile strains were cultured on CCFA medium for 48 h under anaerobic conditions. Suspensions equivalent in turbidity to a 2 McFarland standard were adjusted in 0.85% sterile saline. A volume of 100 µl of each suspension was inoculated into a 10 ml pre-reduced brain heart infusion (BHI) broth and incubated for 5 h under anaerobic conditions. Using a shaking incubator (120 rpm), the broth cultures, which had been aseptically sealed, were incubated for 5 days at 37 °C (Daihan Labtech CO., Ltd, South Korea). To remove cells and debris, cultures were centrifuged at 4,000 × g for 5 min and the supernatants were filtrated using 0.22 μm-pore size filters (Sigma Aldrich, MO, USA) to obtain cell-free filtrates, which contain a complex mixture of released bacterial components, including toxins, surface-associated proteins, enzymes, and other virulence factors. The presence of C. difficile toxin A and B in the supernatant of the toxigenic strain (RT001) was evaluated by enzyme-linked immunosorbent assay (ELISA, Thermo scientific, MA, USA) according to the manufacturer’s instructions. Non-toxigenic strains (RT084 and ATCC 700057) were similarly processed to obtain culture filtrates and examined by ELISA.

L. brevis culture and growth conditions

L. brevis strain IBRC-M10790, which has been previously isolated from turmeric, was gifted from the culture collection of Takgene Zist Company (Tehran, Iran)³². The bacteria were cultured on de Man-Rogosa-Sharpe agar (MRS, Merck, Darmstadt, Germany), and incubated at 37 °C under the abovementioned anaerobic conditions for 48–72 h.

Safety assessment

The safety of L. brevis was investigated by assessing their hemolytic, DNase, and gelatinase activities according to the protocols as described previously^33–35. Staphylococcus aureus ATCC 25923 was served as the positive control for each test.

Isolation of LBMVs

The isolation of LBMVs was carried out based on the previously described method with some modifications^36,37. Briefly, A loopful of grown colonies on MRS agar was cultured in 1 L of basal medium (BHI broth) supplemented with 0.5% yeast extract, 0.05% (w/v) L-cysteine (Sigma Aldrich, MO, USA), 5 µg/ml hemin, 1 µg/ml vitamin K1 under anaerobic conditions for 24 h to yield OD600 of 0.5, an equivalent of 1 × 10⁸ ~ 1.5 × 10⁸ cfu/ml. The bacterial suspension was centrifuged for 20 min at 4 °C at 10,000 g typically 4–6 cycles until there was no visible cell debris. The supernatant was filtrated through a 0.22 μm filter (Sigma Aldrich, MO, USA) to remove residual bacteria. Subsequently, 36 ml volumes of the supernatant were ultracentrifuged at 150,000 g for 5 h, at 4 °C. The obtained pellet was resuspended in 300 µl sterile phosphate-buffered saline (PBS, pH = 7). The extracted MVs were immediately stored at -80 °C for further experiments.

Characterization of LBMVs

The protein concentrations of the isolated MVs were measured using bicinchoninic acid (BCA) protein assay kit (DNAbiotech, Tehran, Iran). The presence of lipopolysaccharide (LPS) in purified MVs was evaluated by LAL Chromogenic Endotoxin Quantitation Kit (Thermo Fisher Scientific, MA, USA) according to the manufacturer’s instructions.

Transmission electron microscopy (TEM)

To evaluate the shape and size of MVs, briefly, 10 µl of MVs were pipetted onto a 400-mesh copper grid with carbon-coated formvar film. After 2 min incubation and removing the extra liquid by blotting, the grid was placed on 10 µl of 2% (w/v) uranyl acetate. After blotting and repeating the last step, the grid became air-dried³⁸. A Philips EM208 TEM system with an accelerating voltage of up to 100Kv and an image magnification of 89 kX was used to examine the structure of the isolated MVs.

Cell culture and growth conditions

Caco-2 and HT-29 cells were obtained from the Research Institute for Gastroenterology and Liver Diseases, Shahid Beheshti University of Medical Sciences, Tehran, Iran. Cells were maintained in the high-glucose Dulbecco’s modified Eagle’s medium (DMEM, Gibco-Invitrogen, CA, USA) with 100 U/ml of penicillin, 100 µg/ml of streptomycin, and 10% heat-inactivated fetal bovine serum (FBS, Gibco-Invitrogen, CA, USA) at 37 °C and a humidified atmosphere with 5% CO₂ until they reached 80% confluency. The cells were routinely sub-cultured by trypsinization upon reaching confluence^39,40. For all experiments, Caco-2 and HT-29 cells between passages 20 and 30 were used to ensure consistency and reproducibility.

Cell viability assay

Cell viability was measured by utilizing the Cell Proliferation Kit I (Sigma Aldrich, USA) according to the manufacturer’s guidelines. Briefly, 5 × 10⁵ cells/well were seeded in 96-well plates and allowed to adhere overnight. Subsequently, cells were then treated for 24 h with the following preparations: culture filtrates from the non-toxigenic C. difficile RT084 (100 µg/ml and 500 µg/ml) and ATCC 700057 strains (500 µg/ml); Tox-S derived from the toxigenic C. difficile RT001 (100 µg/ml, 500 µg/ml), LLB at various MOIs (10, 50, and 100), and different concentrations of LBMVs (10, 100, 1000 ng/ml). These concentrations were selected based on our previous dose-finding studies^40,41. The MTT solution (5 mg/ml in PBS; Sigma-Aldrich, USA) was added to each well and incubated for 4 h at 37 °C. The supernatant was carefully removed, and the resulting formazan crystals were solubilized by adding 100 µl of dimethyl sulfoxide (DMSO; Sigma-Aldrich, USA) to each well. Absorbance was measured at 570 nm using a microplate reader (BioTek Instruments, USA), and a reference wavelength of 630 nm. Untreated cells were included as a negative control (representing maximum cell viability), and cells treated with hydrogen peroxide (H₂O₂) at a concentration of 250 µM were used as a positive control for cell death. The assay was performed in triplicate. The percentage of cell viability of treated cells was measured through the following formula: cell viability (%) = (X × 100%)/Y, where “X” is the absorbance of treated cells and “Y” is the absorbance of untreated cells.

Treatment of Caco-2 and HT-29 cells with culture filtrates, Tox-S, LLB, and LBMVs

For gene expression and cytokine production analysis, Caco-2 and HT-29 cells were pre-treated with culture filtrate of RT084 (100 µg/ml), ATCC 700057 (500 µg/ml) strains, and Tox-S (100 µg/ml) of RT001, and incubated for 24 h at 37 °C in a CO₂ incubator. After incubation, cells were treated with LLB (MOI 100), and LBMVs (100 ng/ml) for 24 h at 37 C in 5% CO₂. Untreated cells and cells only treated with C. difficile supernatants were used as controls. Experiments were carried out in triplicate.

Cytokine measurement

To assess the production of inflammation-related cytokines, culture supernatants from treated Caco-2 and HT-29 cells were collected and analyzed using commercial ELISA kits for IL-8 (ZB-10089 C-H96-48), IL-6 (ZB-10090 C-H9648), IL-1β (ZB-10143 C-H9648), and TNF-α (ZB-10082 C-H9648) (ZellBio, Lonsee, Germany), in accordance with the manufacturer’s instructions. Standard curves were generated for each cytokine, with standard concentration ranges as follows: IL-8 (40-1280 ng/ml), IL-6 (30–960 pg/ml), IL-1β (250-80000 pg/ml), and TNF-α (7.5–240 ng/ml). Absorbance was measured at 450 nm using a microplate reader (ELx808, BioTek Instruments, VT, USA). The cytokine assays were performed in triplicate.

Flow cytometry by Annexin V-FITC/PI staining

Apoptotic cells were quantified by Annexin V-FITC/PI staining (BD Biosciences, San Jose, CA, USA) using a FACSCalibur™ flow cytometer (Becton, Dickinson and Company, Franklin Lakes, NJ, USA). In brief, Caco-2 cells (4 × 10⁵ cells/well) were treated with culture filtrate of RT084 (100 µg/ml) and ATCC 700057 (500 µg/ml) strains, Tox-S (100 µg/ml) of C. difficile, LLB (MOI 100), and LBMVs (100 ng/ml) and maintained for 24 h at 37 °C in a CO₂ incubator. Afterward, cells were harvested by trypsinization and were rinsed with PBS. Cells were then collected and resuspended in a binding buffer [10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (pH 7.5), 2.5 mM CaCl₂ and 140 mM NaCl). Finally, cells were mixed with 5 µl of Annexin V-fluorescein isothiocyanate and 10 µl of propidium iodide and incubated for 15 min in the dark room, prior to flow cytometric analysis^42,43. FlowJo 8.8.6 software was applied to analyze the flow cytometry data (Tree Star Inc., Ashland, OR, USA).

Total RNA extraction and RT-qPCR

Total RNA was extracted from treated Caco-2 and HT-29 cells by RNeasy Plus Mini Kit (Qiagen, GmbH, Germany) based on the manufacturer’s instructions. The quantity and purity of the obtained RNA samples were measured via ultraviolet spectroscopy (NanoDrop spectrophotometer, ND-1000, Thermo Scientific, MA, USA). The RNA was reverse-transcribed to cDNA by using BioFACT™ RT-Kit (BIOFACT CO., Ltd. Daejeon, South Korea) according to the manufacturer’s instructions. The RT-qPCR analysis was performed with Rotor-Gene^® Q (Qiagen, GmbH, Germany) real-time PCR system using BioFACT™ 2X Real-Time PCR Master Mix (BIOFACT CO., Ltd. Daejeon, South Korea). β-actin served as the housekeeping gene. Relative expression was calculated by the 2^−ΔΔCt method, and the RNA input was normalized against the housekeeping gene β-actin. The expression levels were presented as the fold change relative to the control samples. The oligonucleotide primers and amplification conditions used for gene expression analysis of BCL-2, Caspase-3, Caspase-9, BAX, TNF-α, IL-1β, IL-8, and IL-6 are listed in Table 1.

Table 1.

List of oligonucleotide primers used for RT-qPCR assay in this study.

Target gene	Oligonucleotide sequences (5′–3′)	Product size (bp)	Amplification procedures	References
TNF-α	F: CCCAGGGACCTCTCTCTAATC R: ATGGGCTACAGGCTTGTCACT	84	95 °C 15 min, 40 cycles (95 °C 20 s; 55 °C 40 s; 72 °C 35 s), 72 °C 5 min	44
Il-1β	F: GCACGATGCACCTGTACGAT R: CACCAAGCTTTTTTGCTGTGAGT	64	95 °C 15 min, 40 cycles (95 °C 20 s; 55 °C 40 s; 72 °C 35 s), 72 °C 5 min	45
IL-6	F: GCACTGGCAGAAAACAACCT R: TCAAACTCCAAAAGACCAGTGA	119	95 °C 15 min, 40 cycles (95 °C 20 s; 56 °C 40 s; 72 °C 35 s), 72 °C 5 min	44
IL-8	F: CTCTTGGCAGCCTTCCTGATT R: ACTCTCAATCACTCTCAGTTCT	147	95 °C 15 min, 40 cycles (95 °C 20 s; 56 °C 40 s; 72 °C 35 s), 72 °C 5 min	44
BCL-2	F: GAGCTGGTGGTTGACTTTCTC R: TCCATCTCCGATTCAGTCCCT	119	95 °C 10 min, 40 cycles (95 °C 10 s; 56 °C 35 s; 72 °C 20 s), 72 °C 5 min	46
Caspase-3	F: AGGACTCTAGACGGCATCCA R: CAGGAGACTTGGTGCAGTGA	106	95 °C 10 min, 40 cycles (95 °C 10 s; 56 °C 35 s; 72 °C 20 s), 72 °C 5 min	47
Caspase-9	F: GTCCTCAAACCTTCCTGGAAC R: GCCCAAGCTCTTTTTCATCC	141	95 °C 10 min, 40 cycles (95 °C 10 s; 56 °C 35 s; 72 °C 20 s), 72 °C 5 min	48
BAX	F: TTCATCCAGGATCGAGCAGG R: TGAGACACTCGCCTCAGCTTC	104	95 °C 10 min, 40 cycles (95 °C 10 s; 56 °C 35 s; 72 °C 20 s), 72 °C 5 min	47
β-actin	F: ATGTGGCCGAGGACTTTGATT R: AGTGGGGTGGCTTTTAGGATG	107	95 °C 10 min, 40 cycles (95 °C 10 s; 55–56 °C 35 s; 72 °C 20 s), 72 °C 5 min	49

Statistical analysis

All experiments were performed in triplicate, and results are presented as mean ± standard deviation (SD). Statistical analyses were conducted with GraphPad Prism 5 software version 5.04 (GraphPad Software, Inc., San Diego, CA, USA). Comparisons between multiple groups were performed using one-way analysis of variance (ANOVA), followed by Tukey’s post hoc test for multiple comparisons. Comparisons were made between each treatment group and the untreated control group, as well as between groups treated with only C. difficile culture filtrates or Tox-S and groups co-cultured with LLB or LBMVs. The differences were considered statistically significant when P < 0.05; *P < 0.05, **P < 0.01, ***P < 0.001 and ****P < 0.0001.

Results

Safety assessment

In order to ensure the biosafety of the L. brevis strain IBRC M10790 for potential use as a probiotic strain, its hemolytic and gelatinase activity, and DNase production were examined. The strain exerted no hemolysis and gelatinase activity. Also, it was unable to produce DNase enzyme.

Characterization of LBMVs

The extraction of LBMVs is summarized in Fig. S1A. The protein profile of the isolated MV was in the range of 10–200 kDa as observed by SDS-PAGE (Fig. S1B). The morphological characteristics of the isolated MVs were evaluated by TEM analysis (Fig. S1C). These findings revealed that L. brevis was able to produce nanosized, spherical, membrane-bound vesicles. Th uncropped gel of protein profile of MVs is shown in Fig. S1D.

Cell viability of Caco-2 and HT-29 cells treated with culture filtrates, Tox-S, LLB, and LBMVs

H₂O₂, used as a positive control, markedly decreased viability in both Caco-2 and HT-29 cells (P < 0.0001), confirming the sensitivity of the assay. Exposure to Tox-S derived from the toxigenic C. difficile RT001 significantly reduced Caco-2 cell viability in a dose-dependent manner. Notably, RT001 Tox-S at 100 µg/ml and 500 µg/ml significantly decreased viability compared to the untreated control (P < 0.05 and P < 0.01, respectively). In addition, culture filtrate from the non-toxigenic RT084 at 500 µg/ml induced a significant reduction in Caco-2 cell viability (P < 0.05). Similarly, in HT-29 cells, RT001 Tox-S at 100 µg/ml and 500 µg/ml resulted in a significant reduction in cell viability (P < 0.05 and P < 0.01, respectively). However, culture filtrates from RT084 and ATCC 700057 did not significantly affect HT-29 cell viability (Fig. S2-S3).

Effects of Tox-S, LLB, and LBMVs on the expression level of apoptotic-related genes in Caco-2 and HT-29 cells

Overnight exposure of Caco-2 and HT-29 cells with 100 µg/ml Tox-S of RT001 (P < 0.0001, P < 0.0001), and 100 µg/ml culture filtrate of RT084 (P < 0.0001, P < 0.0001), and 500 µg/ml culture filtrate of 700,057 (P < 0.01, P < 0.01), resulted in marked decrease in BCL-2 gene expression. Inversely, LLB and LBMVs upregulated the expression level of BCL-2 in both treated cell lines. However, this elevation was not statistically significant (Fig. 1A).

Fig. 1.

The effects of Tox-S and culture filtrates of C. difficile strains, LLB and LBMVs on apoptosis-related gene expression in Caco-2 and HT-29 cells. Data represents relative gene expression of BCL-2 (A), BAX (B), Caspase-3 (C), and Caspase-9 (D) measured by RT-qPCR in Caco-2 and HT-29 cells after 24 h treatment with Tox-S derived from the toxigenic C. difficile RT001 (100 µg/ml), culture filtrates of the non-toxigenic RT084 (100 µg/ml) and ATCC 700057 (500 µg/ml) strains, LLB (MOI 100), or LBMVs (100 ng/ml). Gene expression data were normalized to β-actin as the reference gene. Data are presented as mean ± SD from three independent experiments. Statistical comparisons were performed using one-way ANOVA followed by Tukey’s post hoc test. Comparisons were made between each treatment group and the untreated control group, and between Tox-S or culture filtrate-treated groups and co-treatment groups with LLB or LBMVs. A P value of < 0.05 was considered significant (*P < 0.05; **P < 0.01; ***P < 0.001; ****P < 0.0001) by one-way ANOVA statistical analysis. LLB, live Levilactobacillus brevis; LBMVs, Levilactobacillus brevis membrane vesicles.

Tox-S of RT001 (P < 0.0001; P < 0.0001) and culture filtrate of RT084 (P < 0.001; P < 0.001) significantly upregulated the expression of BAX after 24 h in Caco-2 and HT-29 cells, whereas culture filtrate of ATCC 700057 caused a limited effect. In contrast, LLB effectively reduced the expression of BAX (P < 0.01; P < 0.05) in Caco-2 and HT-29 cells in comparison to control, as depicted in Fig. 1B. As shown in Fig. 1C, Tox-S of RT001 (P < 0.001; P < 0.01) and culture filtrate of RT084 (P < 0.05; P < 0.05) significantly upregulated Caspase-3 expression after 24 h in Caco-2 and HT-29 cells, while LLB and LBMVs notably decreased its expression level compared to control. This reduction was significant for LLB (P < 0.05; P < 0.01) in Caco-2 and HT-29 cells, and LBMVs (P < 0.01) in HT-29 cells. (Fig. 1C). Moreover, Tox-S of RT001 (P < 0.0001; P < 0.0001) and culture filtrate of RT084 (P < 0.01; P < 0.01) significantly elevated the expression level of Caspase-9 in Caco-2 and HT-29 cells as shown in Fig. 1D. Conversely, LLB (P < 0.05) significantly decreased the Caspase-9 gene expression in HT-29 cells compared to control (Fig. 1D).

LLB and LBMVs decreased C. difficile-induced apoptosis in Caco-2 and HT-29 cells

LLB and LBMVs upregulated the expression level of BCL-2 in Caco-2 and HT-29 cells treated with Tox-S of RT001 and culture filtrate of RT084, however, this reduction was not statistically significant (Fig. 2A). LLB could substantially reduce the expression of BAX in Caco-2 and HT-29 cells stimulated with Tox-S of RT001 (P < 0.01; P < 0.05) and culture filtrate of RT084 (P < 0.01; P < 0.05) (Fig. 2B). LLB also lowered the expression of Caspase-3 (P < 0.05; P < 0.05, P < 0.05; P < 0.05, respectively), as well as Caspase-9 (P < 0.05; P < 0.05, P < 0.05; P < 0.05, respectively) in Caco-2 and HT-29 cells treated with Tox-S of RT001 and culture filtrate of RT084 (Fig. 2C-D). Additionally, the BAX/BCL-2 ratio was significantly elevated in Caco-2 and HT-29 cells exposed to Tox-S of RT001 (P < 0.0001; P < 0.0001) and culture filtrate of RT084 (P < 0.001; P < 0.001) compared to the control (Fig. S4A). However, treatment with LLB (P < 0.05; P < 0.05) and LBMVs (P < 0.05; P < 0.05) led to a significant reduction in the BAX/BCL-2 ratio in Caco-2 and HT-29 cells treated with RT001 or RT084 supernatants (Fig. S4B).

Fig. 2.

The protective effects of LLB and LBMVs on C. difficile Tox-S and culture filtrates induced apoptotic gene dysregulation in Caco-2 and HT-29 cells. Relative expression of BCL-2 (A), BAX (B), Caspase-3 (C), and Caspase-9 (D) in Caco-2 and HT-29 treated with Tox-S of C. difficile RT001 (100 µg/ml), culture filtrates of RT084 (100 µg/ml) and ATCC 700057 (500 µg/ml) strains, LLB, and LBMVs. Gene expression data were normalized to β-actin as the reference gene. Data are presented as mean ± SD from three independent experiments. Statistical comparisons were made as described in Fig. 1. LLB, live Levilactobacillus brevis; LBMVs, Levilactobacillus brevis membrane vesicles.

The detection of apoptosis in Caco-2 cells was further assessed through flow cytometry analysis using annexin V-FITC/PI staining (Fig. 3). The results obtained from the control group indicated that 87.5%, 2.75%, 7.48%, and 2.29% of cells were alive during early apoptosis, late apoptosis, and necrotic stage (Fig. 3A). Flow cytometry demonstrated an increase in late apoptotic cells following treatment with Tox-S of RT001, culture filtrate of RT084, and culture filtrate of ATCC 700057 (14.3%, 13.5%, and 12%, respectively) in comparison to the control group (Fig. 3B-D). In contrast, the number of late apoptotic cells decreased after exposure to LLB and LBMVs (4.93% and 5.93%, respectively) as compared to control cells (Fig. 3E-F). Furthermore, LLB and LBMVs diminished the rate of apoptosis in stimulated cells with Tox-S of RT001, culture filtrate of RT084, and culture filtrate of ATCC 700057 (LLB 5.95%, 3.05% and, 2.98%, respectively; LBMVs: 8.43%, 5.56%, and 4.61%, respectively) (Fig. 3G-L).

Fig. 3.

The inhibitory effects of LLB and LBMVs on C. difficile Tox-S and culture filtrates induced apoptosis in Caco-2 cells determined by Flow cytometry. Flow cytometry by Annexin V-FITC/PI staining was used to determine the proportion of apoptosis in Caco-2 cells after 24 h treatment with the indicated groups. Density plots show necrotic (Q1), late apoptotic (Q2), early apoptotic (Q3), and viable cells (Q4). Treatment of Caco-2 cells with C. difficile Tox-S or culture filtrates exposed to LLB and LBMVs resulted in reduced apoptosis (A-L). The bar graphs represent percentages of early (M), late (N), and total (O) apoptotic cells. Data are presented as mean ± SD from three independent experiments. Statistical comparisons were made as described in Fig. 1. LLB, live Levilactobacillus brevis; LBMVs, Levilactobacillus brevis membrane vesicles.

Similar to flow cytometry histograms, an elevation in the rate of total apoptosis in Caco-2 cells was observed after treatment with Tox-S of RT001, culture filtrate of RT084, and culture filtrate of ATCC 700057 (P < 0.0001) (Fig. 3M-O). Whereas LLB and its derived MVs significantly (P < 0.0001) diminished the rate of total apoptosis in stimulated cells with supernatants of RT001, RT084, and ATCC 700057 when compared to their corresponding controls (Fig. 3M-O).

Effects of C. difficile Tox-S and culture filtrate, and LLB and LBMVs on the expression of inflammation-related mediators in Caco-2 and HT-29 cells

As shown in Fig. 4, Tox-S of RT001 and culture filtrate of RT084 enhanced the expression level of IL-8, IL-6, IL-1β, and TNF-α cytokines in Caco-2 and HT-29 cells. In more detail, Tox-S of RT001 significantly upregulated the expression of IL-8 (P < 0.001, P < 0.001), IL-6 (P < 0.0001, P < 0.001), IL-1β (P < 0.0001, P < 0.001), and TNF-α (P < 0.001, P < 0.0001), compared to control in Caco-2 and HT-29 cells, respectively. Similarly, culture filtrate of RT084 also markedly increase IL-6 (P < 0.05, P < 0.05), IL-1β (P < 0.01, P < 0.05), and TNF-α (P < 0.001, P < 0.0001), compared to control in Caco-2 and HT-29 cells, respectively. The culture filtrate of ATCC 700057 variably changed the expression level of inflammation-related genes, with a significant reduction observed only for TNF-α (P < 0.05), as shown in Fig. 4A-D. Treatment with LLB significantly reduced the expression level of IL-8 (P < 0.001, P < 0.01, respectively), IL-6 (P < 0.05, P < 0.05, respectively), and TNF-α (P < 0.001, P < 0.0001, respectively) genes in Caco-2 and HT-29 cells after 24 h. (Figure 4A-B and D). Treatment with LBMVs also significantly reduced the expression level of IL-6 (P < 0.05, P < 0.05, respectively) and TNF-α (P < 0.001, P < 0.001, respectively) in Caco-2 and HT-29 cells after 24 h. Additionally, LLB (P < 0.01) and LBMVs (P < 0.01) significantly decreased the IL-1β gene expression in HT-29 cells compared to the control (Fig. 4A and C).

Fig. 4.

The effects of Tox-S and culture filtrates of C. difficile strains, LLB and LBMVs on proinflammatory cytokine expression and production in Caco-2 and HT-29 cells. Relative gene expression of IL-8 (A), IL-6 (B), IL-1β (C), and TNF-α (D) in Caco-2 and HT-29 cells was measured by RT-qPCR after treatment with Tox-S of C. difficile RT001 (100 µg/ml), culture filtrates of RT084 (100 µg/ml) and ATCC 700057 (500 µg/ml) strains, LLB (MOI 100), and LBMVs (100 ng/ml) overnight. The production of proinflammatory cytokines IL-8 (E), IL-6 (F), IL-1β (G), and TNF-α (H) in culture supernatants was assessed by ELISA. Data are presented as mean ± SD from three independent experiments. Statistical comparisons were made as described in Fig. 1.

As represented in Fig. 4E-H, the production of IL-8, IL-6, IL-1β, and TNF-α in Caco-2 and HT-29 cells treated with Tox-S, LLB, and LBMVs was further quantified using ELISA. Consistent with the results of gene expression, stimulation of Caco-2 calls with Tox-S of RT001 led to a significant elevation in the production of IL-8 (P < 0.0001, P < 0.001), IL-6 (P < 0.001, P < 0.001), IL-1β (P < 0.001, P < 0.001), and TNF-α (P < 0.01, P < 0.01) in Caco-2 and HT-29 cells. Similarly, culture filtrate of RT084 significantly increased the production of IL-8 (P < 0.01) in HT-29 cells, IL-6 (P < 0.05; P < 0.01) IL-1β (P < 0.01; P < 0.05), and TNF-α (P < 0.01; P < 0.05) in Caco-2 and HT-29 cells, respectively. On the other hand, treatment with LLB significantly reduced the secretion of IL-8 (P < 0.05, P < 0.01), IL-6 (P < 0.001, P < 0.001), and TNF-α (P < 0.0001, P < 0.0001) in Caco-2 and HT-29 cells, respectively (Fig. 4E-F and H). Also, Treatment with LBMVs notably decreased the production of IL-6 (P < 0.01, P < 0.01, respectively) and TNF-α (P < 0.01, P < 0.01, respectively) in Caco-2 and HT-29 cells (Fig. 4F and H).

LLB and LBMVs mitigated C. difficile supernatant-induced inflammation in Caco-2 and HT-29 cells

As represented in Fig. 5A-B and D, treatment with LLB and LBMVs significantly decreased IL-8 (P < 0.0001, P < 0.001; P < 0.0001, P < 0.001, respectively), IL-6 (P < 0.0001, P < 0.0001; P < 0.0001, P < 0.0001, respectively), and TNF-α (P < 0.01, P < 0.001; P < 0.05, P < 0.05, respectively) in Caco-2 and HT-29 cells stimulated with Tox-S of RT001 after 24 h. Similarly, Treatment with LLB and LBMVs significantly decreased IL-8 (P < 0.0001, P < 0.0001; P < 0.0001, P < 0.0001, respectively), IL-6 (P < 0.0001, P < 0.0001; P < 0.0001, P < 0.0001, respectively), and TNF-α (P < 0.01, P < 0.001; P < 0.01, P < 0.01, respectively) in Caco-2 and HT-29 cells stimulated with Tox-S of RT084 after 24 h. Furthermore, LLB and LBMVs significantly downregulated IL-8 (P < 0.0001, P < 0.0001; P < 0.0001, P < 0.0001, respectively), and TNF-α (P < 0.01, P < 0.001; P < 0.01, P < 0.001, respectively) in Caco-2 and HT-29 cells stimulated with culture filtrate of ATCC 700057. LLB and LBMVs also significantly reduced IL-6 (P < 0.05, P < 0.05, respectively) in Caco-2 cells stimulated with culture filtrate of ATCC 700057. Furthermore, treatment with LLB and LBMVs decreased the gene expression of IL-1β in Caco-2 and HT-29 cells stimulated with Tox-S of RT001 and culture filtrate of RT084 and ATCC 700057 strains. However, this reduction was not statistically significant (Fig. 5C).

Fig. 5.

The protective effects of LLB and LBMVs on C. difficile Tox-S and culture filtrates induced proinflammatory response in Caco-2 and HT-29 cells. Relative gene expression of IL-8 (A), IL-6 (B), IL-1β (C), and TNF-α (D) assessed by RT-qPCR, and cytokine secretion levels of IL-8 (E), IL-6 (F), IL-1β (G), and TNF-α (H) determined by ELISA after treatment with Tox-S of C. difficile RT001 (100 µg/ml), and culture filtrates of RT084 (100 µg/ml) and ATCC 700057 (500 µg/ml) strains. Data are presented as mean ± SD from three independent experiments. Statistical comparisons were made as described in Fig. 1. LLB, live Levilactobacillus brevis; LBMVs, Levilactobacillus brevis membrane vesicles.

Further analysis using ELISA showed that treatment with LLB and LBMVs significantly reduced the secretion of IL-8 (P < 0.0001, P < 0.01; P < 0.001, P < 0.05, respectively) and IL-6 (P < 0.0001, P < 0.01; P < 0.0001, P < 0.05, respectively) in Caco-2 and HT-29 cells stimulated with Tox-S of RT001. In addition, LLB and LBMVs treatment notably decreased IL-8 (P < 0.01, P < 0.01; P < 0.05, P < 0.05, respectively), and IL-6 (P < 0.01, P < 0.01; P < 0.05, P < 0.05, respectively) in Caco-2 and HT-29 cells stimulated with culture filtrate of RT084 as represented in Fig. 5E-F. LLB and LBMVs reduced the production of IL-1β in Caco-2 and HT-29 cells stimulated with Tox-S of RT001 and culture filtrate of RT084 and ATCC 700057 strains. However, this reduction was not statistically significant (Fig. 5G). LLB also significantly decreased the production of TNF-α in Caco-2 and HT-29 cells treated with Tox-S of RT001 (P < 0.05, P < 0.05, respectively), RT084 (P < 0.001, P < 0.001, respectively) and culture filtrate of ATCC 700057 (P < 0.001, P < 0.001, respectively). Moreover, LBMVs effectively diminished TNF-α production (P < 0.05; P < 0.05, respectively) in Caco-2 and HT-29 cells exposed to culture filtrate of ATCC 700057 (Fig. 5H).

Discussion

Recent steadfast microbial-based therapeutic developments have paved the way for designing certain strategies to defeat CDI^22,50. FMT is currently regarded as the most effective therapeutic option for treating rCDI in patients who fail to respond to standard antibiotic therapies⁵¹. However, FMT is generally reserved for patients with multiple recurrences rather than initial or mild cases of CDI, and challenges such as donor variability, potential pathogen transmission, and limited public acceptance have prompted the search for safer, more standardized alternatives⁵². Notably, it can be suggested that part of FMT therapeutic efficacy may be mediated by bacterial MVs present in the donor material, which can deliver bioactive components and modulate host immune responses. Thus, the application of probiotic consortia and bacterial-derived vesicle therapies can be promising adjunctive or prophylactic strategies that regulate gut microbiome dysbiosis and potentially reduce CDI and limit reinfections⁵³. By modulating inflammatory pathways and supporting microbiota resilience, these microbes have the potential to serve either as preventive agents in patients at risk of CDI or as complementary therapies to reduce the risk of relapse after antibiotic treatment⁵⁴.

Induction of apoptosis in microbial infections is generally a host defense mechanism⁵⁵. Some pathogens have evolved virulence mechanisms that manipulate apoptotic pathways to promote their replication and spread in a hostile host environment³⁵. According to previous studies, TcdA and TcdB have shown the potential to induce both apoptosis and necrosis of epithelial cells in human intestinal xenografts in a chimeric mouse model. Therefore, these toxins could gain access to the underlying lamina propria and submucosa to affect other cell types⁵⁶. In another study, Saavedra et al.⁵⁷ indicated that C. difficile enterotoxins induce activation of executioner Caspase-3 and Caspase-7 via the intrinsic apoptosis pathway, and demonstrated that this type of apoptosis is critical for in vivo host defense during the early stages of CDI in a mouse model of pseudomembranous colitis. In vitro studies have also shown that TcdA exposure affected the activation of the caspase-cascade system as evidenced by alterations in the mitochondrial outer membrane permeabilization, which triggers apoptosis through the mitochondrial pathway, dependent on Rho inactivation and mitochondrial damage⁵⁸.

Mounting evidence has elucidated that TcdA and TcdB can activate Caspase-3 via initiator caspases, including Caspase-8 and Caspase-9^59–61. Following stimulation by TcdA or TcdB, BCL-2 coordinates the permeability of the mitochondrial outer membrane and triggers the secretion of cytochrome C. Furthermore, the formation of apoptosome complexes actuates the secretion of Caspase-9, which represents the intrinsic pathway of apoptosis⁶¹. Cleaved Caspase-9 can trigger Caspase-3 afterward, which induces apoptosis in the long run⁶². However, Fettucciari et al.⁶¹ reported that TcdB exerted its apoptotic effect on enteric glial cells through caspase-dependent and mitochondria-independent pathway, in which the expression level of BAX or B-cell lymphoma-x long isoform (BCL-XL) were not significantly changed. In this regard, Matte et al.⁶³ demonstrated that TcdA has the potential to stimulate the mitochondrial pathway of apoptosis in both ovarian and colonic cancer cells by a caspase-independent mechanism. The authors discussed that this process was regulated by antiapoptotic members of the BCL-2 family. It has been reported by Zhang et al.⁶⁴ that infecting breast cancer mouse models with TcdB noticeably suppressed the expression of BCL-2, which is an inhibitor of apoptosis and another biomarker for the mitochondrial cell death cascade. On the other hand, while examining the transduction pathways involved in controlling the expression of BCL-2 in neuroblastoma SK-N-SH cells, Lombet et al.⁶⁵ reported that TcdB decreased the basal expression of BCL-2 to 25% after overnight treatment. They also suggested that RhoA and PtdIns3 kinase might be part of the transduction pathway involved in protein kinase C activation, leading to BCL-2 expression and resistance towards apoptosis. An increasing body of evidence suggests that a greater ratio of BAX/BCL-2 could be an indicator of the intrinsic apoptosis pathway, which can result in the activation of Caspase-3. Using flow cytometry and RT-qPCR, we showed that the overnight exposure of Caco-2 and HT-29 cells to supernatants of RT001 and RT084 resulted in the overexpression of apoptotic BAX, Caspase-9 and Caspase-3 and downregulation of antiapoptotic BCL-2. Furthermore, changes in the expression level of Caspase-9 and Caspase-3 were consistent with the BAX/BCL-2 ratio. The apoptotic response induced by RT084 supernatants may be mediated by non-toxin virulence factors produced by these strains, which can play critical roles in the severity of CDI and may contribute to differences in the expression of apoptotic genes^66,67.

Several studies have demonstrated that certain probiotic Lactobacillus strains can promote apoptotic activities against colorectal cancer cells to inhibit their viability and proliferation significantly^68–70. In a study conducted by Saber et al.⁷¹ while assessing the anticancer effect of the potential probiotic Pichia kudriavzevii AS-12 isolated cell-free supernatant on HT-29 and Caco-2 cell lines, the authors noticed a reduction in the expression level of BCL-2 and increase in the expression level of pro-apoptotic genes including BAD, Caspase-3, Caspase-8, Caspase-9, and Fas-R. In contrast, in this work, we observed that after treating the cells with LLB or LBMVs for 24 h, the expression of BCL-2 enhanced significantly (Fig. 6).

Fig. 6.

Proposed mechanisms by which LLB and LBMVs alleviate apoptosis and inflammation in epithelial cells exposed to C. difficile released virulence factors. Treatment with LLB and LBMVs modulates apoptosis by upregulating anti-apoptotic gene (BCL-2) and downregulating pro-apoptotic genes (BAX, Caspase-3, Caspase-9), and attenuates inflammation by reducing IL-8, IL-6, and TNF-α expression. LLB, live Levilactobacillus brevis; LBMVs, Levilactobacillus brevis membrane vesicles.

Although there are limited data on the impact of probiotic administration on C. difficile-induced apoptosis, in a study performed by Yarmohammadi et al.³³it has been observed that the potentially probiotic strain Lactobacillus gasseri ATCC 33323 decreased BCL-2 expression after infecting with H. pylori strains. El-Khadragy et al.⁷² demonstrated that oral administration of probiotics or yogurt before or after infection with Schistosoma mansoni remarkably upregulated BCL-2 expression level and downregulated the expression of the BAX gene. In the present study, we observed that LBMVs could not entirely normalize the apoptosis-related genes in Caco-2 and HT-29 cells and made no significant changes in the expression level of apoptosis-related genes in cells treated with released C. difficile virulence factors. However, LLB efficiently downregulated the expression level of apoptosis-related genes in Caco-2 and HT-29 cells stimulated with Tox-S from RT001 or culture filtrate from RT084. In addition, the diminished BAX/BCL-2 ratio induced by LLB was accompanied by a decrease in apoptosis-related gene expression levels.

Both TcdA and TcdB possess potent proinflammatory activities and are capable of stimulating gut epithelial and immune cells to produce different cytokines and chemokines⁷³. In animal models, TcdA has induced neutrophil infiltration, as well as the production of cyclooxygenase-2, prostaglandin E2, and inflammatory cytokines, which mediate tissue inflammation^58,74. Once the protective epithelial barrier has been breached, TcdA and TcdB come into contact with submucosal macrophages, monocytes, and dendritic cells to trigger the dissemination of the inflammatory cascade via further release of proinflammatory cytokines IL-1α, IL-1β, IL-6, IL-8 and TNF-α, which can predominantly exert their effects on neutrophils^75,76. Furthermore, it has been shown that non-toxigenic strains may still contribute to host cell inflammation through other virulence factors such as SLPs and structural components, which can interact with Toll-like receptor 4 (TLR4) and trigger an inflammatory response independent of toxin production. Working on CCL-241 and HISM cell lines, Ng et al.⁷⁷ reported that in HISM cells, but not in CCL-241 cells, treatment with C. difficile culture extract containing TcdA, TcdB, and LPS slightly increased IL-6 expression level. Yu et al.⁷⁸ identified a tremendous increase in IL-8 (80-fold), IL-6 (42.3-fold), and IL-1β (143-fold) in serum samples of CDI patients, which proposes that these cytokines play a vital role in CDI pathogenesis. Nicholas et al.⁷⁹ also described ATCC 43255-derived MVs as important nano complexes, which stimulated the expression of IL-6, IL-8, and IL-1β, therefore eliciting a proinflammatory response and cytotoxicity in Caco-2 cells. In accordance with the abovementioned studies, we noticed that when Caco-2 and HT-29 cells were treated with supernatant of C. difficile strains, the production of IL-6, TNF-α, and IL-1β increased significantly. However, changes related to IL-8 production were only significant when Tox-S of RT001 was applied. Accordingly, it can be suggested that the major proinflammatory effects are primarily mediated by the classical toxins TcdA and TcdB in our experiment.

In the present work, when administered solely, LLB as well as LBMVs noticeably downregulated IL-6, IL-8, and TNF-α expression and had no significant impact on the expression level of IL-1β. However, data obtained from a meta-analysis performed by Milajerdi et al.⁸⁰ indicated that probiotic supplementation significantly reduced serum concentrations of proinflammatory cytokines, including hs-CRP, TNF-α, IL-6, IL-12, and IL-4, but did not influence IL-1β, IL-8, IFN-γ, and IL-17 concentrations. In a study carried out by Boonma et al.⁸¹ assessing 34 Lactobacillus spp. isolated from infant feces revealed that L. rhamnosus L34 and L. casei L39 were able to produce heat-stable factors capable of suppressing inflammation by downregulating C. difficile-induced IL-8 production in colonic epithelial cells. Moreover, it was suggested that L. rhamnosus L34 and L. casei L39 can produce factors that are able to modulate inflammation stimulated by C. difficile. Hence, they described these strains as potential probiotics against C. difficile. Also, in another study by Wei et al.⁸² the anti-TcdA and -TcdB capability of Bifidobacterium longum JDM301 was investigated. Using cytokine quantification by cytometric bead array, researchers found that B. longum JDM301 reduced IL-6 and TNF-α production in the colon tissues of mice with CDI. Notably, we indicated that LLB and LBMVs almost had a similar effect on downregulating the expression level of IL-6, IL-8, and TNF-α in Caco-2 and HT-29 cells stimulated with supernatants of RT001 and RT084. On the other hand, LLB and LBMVs decreased the production of TNF-α in cells treated with the culture filtrate of ATCC 700057. The difference between the two non-toxigenic strains may be attributed to distinct structural components or secreted virulence factors in RT084, which would be capable of inducing a stronger basal inflammatory response compared to the reference strain ATCC 700057. Accordingly, our findings suggest that L. brevis IBRC-M10790 and its MVs may protect intestinal epithelial cells from C. difficile Tox-S-induced injury by upregulating the anti-apoptotic gene BCL-2 and suppressing the expression of pro-apoptotic mediators such as BAX, Caspase-3, and Caspase-9. These protective effects are accompanied by a reduction in the production of inflammatory cytokines, including IL-6, IL-8, TNF-α, and IL-1β, as schematically illustrated in Fig. 6.

Conclusion

In summary, this study demonstrates that LLB and LBMVs can modulate cell death cascade in Caco-2 and HT-29 cells stimulated by released C. difficile virulence factors through modulating the expression of apoptosis-related genes. Moreover, treatment with LLB and LBMVs mitigated the expression and production of proinflammatory markers in both cell lines after exposure to C. difficile culture supernatants, which suggests a potential to inhibit CDI-associated inflammation. Based on these findings, L. brevis IBRC-M10790 and its derived MVs show promise in ameliorating apoptotic and inflammatory responses induced by C. difficile virulence factors. This probiotic could be exploited as a potential adjunct and supplement to the current treatment strategies against CDI. However, our experiments were performed in vitro using human epithelial cell lines and relied on bacterial supernatants rather than purified toxins. Further in vitro and in vivo investigations are warranted to identify the specific metabolites and by-products generated by this probiotic strain and decipher their putative mechanisms of action against C. difficile pathogenesis.

Electronic supplementary material

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Author contributions

Masoumeh Azimirad, Maryam Noori, Samira Alipour, and Tannaz Salehi performed cell culture and molecular assays. Maryam Noori contributed to figure illustration. Masoumeh Azimirad, Armitasadat Emami Meibodi, and Abbas Yadegar participated in data analysis and interpretation, reviewed the literature, and wrote the manuscript draft. Abbas Yadegar designed the study and participated in data analysis, conceptualization, writing and editing, and project administration. Abbas Yadegar and Mohammad Reza Zali critically edited the manuscript. All authors read the final version of the manuscript and approved the list of authors.

Funding

This study was supported financially by a grant [no. RIGLD 992] from the Foodborne and Waterborne Diseases Research Center, Research Institute for Gastroenterology and Liver Diseases, Shahid Beheshti University of Medical Sciences, Tehran, Iran.

Data availability

All data generated or analyzed during this study are included in this published article [and its supplementary information files].

Competing interests

The authors declare no competing interests.

Ethics approval

This work does not contain any studies related to human participants or animals. The study was approved by the Institutional Review Board of the Research Institute for Gastroenterology and Liver Diseases, Shahid Beheshti University of Medical Sciences, Tehran, Iran (Project No. IR.SBMU.RIGLD.REC.1395.211).