Targeting PPARγ–NF-κB: (+)-catechin reverses Clostridioides difficile toxin A-induced intestinal barrier dysfunction

ABSTRACT

Background

Clostridioides difficile toxin A (TcdA) is a key factor in antibiotic-associated intestinal epithelial injury. (+)-Catechin (CC) exhibits anti-inflammatory and antioxidant properties, but its role in protecting intestinal epithelial barrier function during TcdA exposure remains unclear.

Methods

Human colon epithelial Caco-2 cells were treated with TcdA to induce barrier dysfunction, followed by CC at varying concentrations. Barrier integrity was assessed using trans-epithelial electrical resistance (TEER), Lucifer yellow paracellular flux, and lactate dehydrogenase (LDH) release. The involvement of peroxisome proliferator-activated receptor gamma (PPARγ) and nuclear factor kappa B (NF-κB) signaling was evaluated using short hairpin RNA (shRNA)-mediated PPARγ knockdown.

Results

TcdA treatment disrupted Caco-2 barrier function, indicated by decreased TEER, increased Lucifer yellow flux, and elevated LDH release. TcdA treatment also reduced PPARγ expression and was associated with increased activation of the NF-κB signaling pathway. CC treatment significantly alleviated these effects in a concentration-dependent manner, with 100 μg/mL showing optimal protection. PPARγ knockdown partially attenuated the protective effects of CC and diminished its inhibitory effect on NF-κB activation, suggesting that PPARγ contributes to the barrier-protective role of CC.

Conclusion

CC attenuates TcdA-induced intestinal epithelial barrier dysfunction and is associated with modulation of the PPARγ–NF-κB signaling pathway.

1. Introduction

Long-term or repeated antibiotic use disrupts the gut microbial balance and can directly injure colonic epithelial cells, compromising the integrity of the intestinal epithelial barrier and increasing the risk of intestinal inflammation [1,2]. Among antibiotic‑associated complications, Clostridioides difficile (C. difficile) is a highly transmissible, Gram‑positive anaerobic bacterium that accounts for 15–25% of antibiotic‑associated diarrhea cases [3]. The development of intestinal disease caused by C. difficile is closely linked to bacterial colonization, secretion of the C. difficile toxins A (TcdA) and C. difficile toxins B (TcdB), and direct interactions with the intestinal epithelium [4]. These toxins activate inflammatory signaling pathways in epithelial cells and disrupt tight-junction proteins, leading to increased barrier permeability and intestinal inflammation [5–7]. Therefore, interventions targeting the interaction between C. difficile toxins and the intestinal epithelium are regarded as promising approaches for both prevention and therapy.

In recent years, traditional Chinese medicine and natural bioactive compounds have attracted considerable attention for their ability to modulate intestinal barrier function and inflammatory responses [8]. For example, the classical herbal formula Wulingsan has demonstrated favorable efficacy in regulating gastrointestinal disorders [9]. Network pharmacology analyses suggest that several flavonoids in Wulingsan, including (+)-catechin (CC) [10]. CC, a major flavonoid component of tea, is a natural polyphenolic compound widely present in green tea, black tea, and cocoa. Other tea-derived flavonoids have also been reported to reduce epithelial barrier damage and inflammation caused by this pathogen, highlighting their potential to protect intestinal epithelial integrity [11]. CC may play a key role in alleviating intestinal injury induced by C. difficile. Bioinformatics Analysis Tool for Molecular Mechanism of Traditional Chinese Medicine (BATMAN-TCM) network pharmacology analysis identifies peroxisome proliferator-activated receptor gamma (PPARγ) as both a potential target of CC and a key regulator in C. difficile-related disease [12]. PPARγ has been shown to suppress activation of the nuclear factor kappa B (NF-κB) signaling pathway, whose excessive activation is a major contributor to epithelial barrier dysfunction [13,14].

Based on this background, the present study aims to investigate the protective effects and molecular mechanisms of CC against TcdA-induced intestinal epithelial barrier disruption. We focus on the PPARγ–NF-κB signaling pathway to evaluate whether CC can restore epithelial barrier function by modulating this pathway, thereby providing experimental evidence and theoretical support for the application of natural bioactive compounds in the prevention and treatment of intestinal disease caused by C. difficile.

2. Materials and methods

2.1. Cell culture

Dulbecco’s Modified Eagle Medium (DMEM; C0891, Beyotime, China) was utilized to culture human colonic epithelial Caco-2 cells (AW-CH0049, AnWei-sci, China). It was supplemented with 10% fetal bovine serum (FBS; C0235, Beyotime, China) and 1% penicillin–streptomycin (CA005-010, GenDEPOT, USA). In a humidified environment with 5% CO₂ at 37°C (Forma Steri-Cult, Thermo Fisher, USA), the cells were incubated.

2.2. Preparation of C. difficile toxins

TcdA/B were purified using a known process [15]. In brief, C. difficile (HZB354278, Huizaobio, China) was cultured anaerobically (HYQX, Hengzi, China) in brain-heart infusion (BHI) medium (FD3192, Hangzhou Fude Biological Technology, China) for 5 days. The culture supernatant was concentrated using a 100 kDa ultrafiltration tube (FUF058, Beyotime, China). TcdA was extracted from the concentrated mixture by DEAE-Sepharose anion-exchange chromatography (1.0 × 10 cm, ~8 mL bed volume; 420401-1010, DWK Life Science, China) [16]. Eluted fractions containing TcdA were pooled, desalted, and quantified by BCA assay (P0009, Beyotime, China).

2.3. Experimental design

PPARγ-targeting short hairpin RNA (shRNA) (shPPARγ; pRP[shRNA]-U6 >hPPARγ[shRNA#1], VectorBuilder, China) or a negative control shRNA (sh-NC) was transfected into Caco-2 cells using the Lipo6000 reagent (C0526, Beyotime, China). The transfection effectiveness was verified by Western blot during a 48-h incubation period at 37°C. Cells were allocated to the following groups: Control, TcdA, TcdA + CC (25 µg/mL), TcdA + CC (50 µg/mL), TcdA + CC (100 µg/mL), TcdA+CC, TcdA+CC+shNC, and TcdA+CC+shPPARγ. Cells in the TcdA group were treated with TcdA at a final concentration of 10 µg/mL for 16 h [15]. After being treated with 10 μg/mL TcdA for 16 h, Caco-2 cells in the TcdA, TcdA + CC (25 µg/mL), TcdA + CC (50 µg/mL), and TcdA + CC (100 µg/mL) groups were incubated with 0, 25, 50, or 100 μg/mL CC (HY-N0898, MCE, USA) for 6 h [17]. The TcdA+CC group received 100 µg/mL CC after TcdA exposure. The corresponding shRNA was transfected into Caco-2 cells in the TcdA+CC+shNC and TcdA+CC+shPPARγ groups, which were subsequently grown for 48 h, treated with 10 μg/mL TcdA for 16 h, and then incubated with 100 μg/mL CC for 6 h. Using a light microscope with a digital camera (200×, Zeiss, Germany), cell morphology was investigated. Thus, in all experiments, CC was administered after TcdA exposure, representing a therapeutic (post-treatment) model to assess reversal of established barrier injury.

2.4. Lactate dehydrogenase (LDH) assay

A CytoTox 96 non-radioactive kit (YZ-G1780, Nobleryder, China) was implemented to evaluate cytotoxicity. 96-well plates (FCP962, Beyotime, China) were planted with 100 µL of Caco-2 cells per well. Following the recommended procedures, 50 µL of CytoTox 96 reagent was added, incubated for 30 min, and then, 50 µL of stop solution was added. Utilizing a microplate reader (Bio-Rad, USA), absorbance was determined at 490 nm.

2.5. Transepithelial electrical resistance (TEER)

Caco-2 cells (2 × 10⁴ cells per well) were seeded in 24-well Transwell inserts (3413, Corning, USA) and cultured until a confluent monolayer was formed prior to TEER measurement. An epithelial volt-ohm meter (EVM-MT-03–01, YX-Intl, China) was deployed to quantify TEER. From each measurement, the resistance of the medium and blank filters was deducted [18].

2.6. Paracellular tracer flux

Lucifer yellow, a fluorescent tracer, was applied to gauge paracellular permeability. Cells were cultivated for 24 h after being seeded on 24-well Transwell inserts. The apical chamber (200 µL) was filled with Lucifer yellow (20 µg/mL; HY-D0208, MCE, USA) and incubated for 5 h. Fluorescence was quantified using a plate reader (Molecular Devices, China) at 405 nm excitation and 500–550 nm emission after the medium from the basolateral chamber was collected.

2.7. Western blot

50 µL of radioimmunoprecipitation assay (RIPA) buffer (P0013, Beyotime, China) containing protease inhibitors (P1006, Beyotime, China) was utilized to lyse the cells in each well. A BCA kit was utilized to calculate the protein concentrations after centrifugation was done to clarify the lysates. The protein was separated using 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (P0690, Beyotime, China) and then transferred onto polyvinylidene difluoride membranes (FFP24, Beyotime, China). After blocking the membranes for 2 h (room temperature) using 5% nonfat milk in TBST (20 mM Tris-HCl, 150 mM NaCl, 0.05% Tween-20, pH 7.4), the primary antibodies (Table 1) were incubated for the entire night at 4°C. Following washing, the membranes were incubated for 1 h (room temperature) with secondary antibodies coupled with horseradish peroxidase. A ChemiDocTM XRS imaging system (Bio-Rad, USA) and an enhanced chemiluminescence kit (KGC4603, KeyGene, China) served to view the bands. After normalizing to glyceraldehyde 3-phosphate dehydrogenase (GAPDH), band intensities were acquired using ImageJ software and expressed as relative protein levels.

Table 1.

Antibodies used in Western blot analysis.

Antibody name	Molecular weight	Host species	Catalog number	Supplier	Type	Dilution
PPARG	58 kDa	Rabbit	ab59256	Abcam	Primary antibody	1:1000
P65	65 kDa	Rabbit	ab32536	Abcam	Primary antibody	1:1000
p-P65	65 kDa	Rabbit	ab76302	Abcam	Primary antibody	1:1000
ZO-1	195kDa	Rabbit	ab307799	Abcam	Primary antibody	1:1000
Occludin	59kDa	Rabbit	ab216327	Abcam	Primary antibody	1:1000
Claudin1	22kDa	Rabbit	ab211737	Abcam	Primary antibody	1:2000
GAPDH	36 kDa	Rabbit	ab181602	Abcam	Primary antibody	1:4000
Goat Anti-Rabbit IgG H&L (HRP)	—	Goat	ab6721	Abcam	Secondary antibody	1:4000

2.8. Enzyme-linked immunosorbent assay (ELISA)

Concentrations of IL-8 and TNF‑α in cell culture supernatants were measured using commercially available ELISA kits according to the manufacturers’ protocols. Briefly, Caco‑2 cells were seeded in 24‑well plates at 2.5 × 10⁵ cells/well and cultured to confluence. After the indicated treatments (as described above), cell culture supernatants were collected, centrifuged at 1000 × g for 10 min at 4°C to remove cellular debris, and stored at −80°C until analysis. The levels of IL‑8 and TNF‑α were determined using human IL‑8 (E-EL-H6008, Elabscience, China) and TNF‑α (E-EL-H0109, Elabscience, China) ELISA kits. The absorbance was read at 450 nm using a microplate reader, and cytokine concentrations were calculated from standard curves generated with recombinant human IL‑8 and TNF‑α standards provided with the kits. All measurements were performed in triplicate.

2.9. Statistical analysis

The program GraphPad Prism 8.0 was applied to examine all of the data. All data are presented as mean ± standard deviation (SD) from three independent biological replicates (n = 3 per group per experiment, with each replicate representing independently cultured Caco‑2 cells). Prior to one‑way analysis of variance (ANOVA), normality (Shapiro–Wilk test) and homogeneity of variances (Levene’s test) were verified; assumptions were met for all datasets. Comparisons among multiple groups were conducted using one‑way ANOVA followed by Tukey’s post hoc test. A value of p < 0.05 was considered statistically significant.

3. Results

3.1. CC reverses TcdA-induced intestinal epithelial barrier dysfunction

Caco-2 intestinal epithelial cells were killed by TcdA exposure, and this impact was lowered by CC therapy (Figure 1(A)). At 100 µg/mL CC, the best protection was seen (Figure 1(A)), suggesting that CC effectively counteracts TcdA-induced epithelial injury. Cells in the TcdA group displayed higher Lucifer yellow flux and greater LDH release in comparison to the Control group, but TEER diminished (p < 0.001, Figure 1(B-D)). The increase in paracellular permeability together with elevated LDH release further confirmed that TcdA disrupted membrane integrity and epithelial tight junction function. Relative to the TcdA group, CC at 50 µg/mL and 100 µg/mL greatly lowered LDH release and raised TEER in a concentration-dependent manner (p < 0.01, Figure 1(B,C)). Additionally, 25, 50, and 100 µg/mL CC suppressed Lucifer yellow flux, with 100 µg/mL CC exhibiting the highest inhibition (p < 0.001, Figure 1(D)). These findings demonstrate that CC preserves intestinal epithelial barrier function by reducing cytotoxicity and paracellular permeability under TcdA stimulation. Exposure to TcdA raised phosphorylated P65/total P65 (p-P65/P65) and inhibited PPARγ expression (p < 0.001, Figure 2(A-C)). This indicates activation of the NF-κB inflammatory signaling pathway and concurrent inhibition of PPARγ-mediated regulatory signaling following toxin exposure. These alterations were reversed by treatment with 25, 50, or 100 µg/mL CC; the largest effect was shown at 100 µg/mL (p < 0.001, Figure 2(A-C)). The simultaneous suppression of NF-κB activation and restoration of PPARγ expression suggest that the protective effect of CC may be associated with modulation of the PPARγ–NF-κB signaling axis. To further evaluate the functional relevance of NF‑κB modulation, we measured the levels of pro‑inflammatory cytokines IL‑8 and TNF‑α by ELISA. As shown in Figure 2(D,E), TcdA significantly increased IL‑8 and TNF‑α production compared to the Control group (p < 0.001). CC treatment reduced these cytokines in a concentration‑dependent manner, with 100 μg/mL CC achieving the greatest inhibition (p < 0.01). These data indicate that CC suppresses TcdA‑induced inflammatory responses.

Figure 1.

(+)-Catechin (CC) reverses Clostridioides difficile toxins A (TcdA)-induced intestinal epithelial barrier dysfunction.

(A) Morphology of Caco-2 cells in Control, TcdA, TcdA + CC (25 µg/mL), TcdA + CC (50 µg/mL), and TcdA + CC (100 µg/mL) groups observed by optical microscopy. Magnification: 200×; scale bar = 100 μm. (B) Lactate dehydrogenase (LDH) levels in Caco-2 cells were measured using the CytoTox 96 non-radioactive cytotoxicity assay. (C) Transepithelial electrical resistance (TEER) measurements of Caco-2 cell monolayers. (D) Paracellular flux of Lucifer yellow in Caco-2 cells. Catechin was added after TcdA exposure (therapeutic protocol). Data are presented as mean ± standard deviation (n = 3 independent biological replicates). ***p < 0.001 vs. Control; ^{^^}p < 0.01, ^{^^^}p < 0.001 vs. TcdA.

Figure 2.

CC regulates peroxisome proliferator-activated receptor gamma (PPARγ)-nuclear factor kappa B (NF-κB) signaling in TcdA-treated Caco-2 cells.

(A–C) Western blot analysis of PPARγ and phosphorylated P65/total P65 (p-P65/P65) levels in Control, TcdA, TcdA + CC (25 µg/mL), TcdA + CC (50 µg/mL), and TcdA + CC (100 µg/mL) groups. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as a loading control. (D–E) ELISA measurement of IL‑8 (D) and TNF‑α (E) levels in Control, TcdA, TcdA + CC (25 µg/mL), TcdA + CC (50 µg/mL), and TcdA + CC (100 µg/mL) groups. (F–I) Western blot analysis of tight junction proteins ZO-1 (G), Occludin (H), Claudin1 (I) levels in Control, TcdA, TcdA + CC (25 µg/mL), TcdA + CC (50 µg/mL), and TcdA + CC (100 µg/mL) groups. GAPDH was used as a loading control. Catechin was added after TcdA exposure (therapeutic protocol). Data are mean ± standard deviation (n = 3 independent biological replicates). ***p < 0.001 vs. Control; ^{^^}p < 0.01, ^{^^^}p < 0.001 vs. TcdA.

We further examined the expression of tight junction proteins ZO‑1, Occludin, and Claudin-1. As shown in Figure 2(F–I), TcdA markedly reduced all three proteins (p < 0.001). The expression of ZO‑1, Occludin, and Claudin-1 was restored by CC in a concentration‑dependent manner (p < 0.01). These data provide molecular evidence that CC protects epithelial barrier integrity by rescuing tight junction proteins. Based on these results, 100 µg/mL CC was selected for subsequent mechanistic experiments to further investigate its regulatory role in intestinal barrier protection.

3.2. CC protects the intestinal barrier through the PPARγ–NF-κB pathway

PPARγ expression was dramatically lowered by TcdA therapy, and this drop was reversed by CC administration (p < 0.001, Figure 3(A,B)). This suggests that CC directly modulates PPARγ expression under inflammatory stress conditions. PPARγ levels were lower in the TcdA+CC+shPPARγ group than in the TcdA+CC+shNC group (p < 0.001, Figure 3(A,B)). Effective silencing of PPARγ allowed us to further determine whether the protective effects of CC depend on PPARγ signaling. Additionally, TcdA-induced boosts in LDH release and Lucifer yellow flow, as well as reduction in TEER, were reversed by CC treatment (p < 0.001, Figure 3(C-E)). By knockdown of PPARγ, the effects of CC were partially reversed (p < 0.01, Figure 3(C-E)). These findings suggest that PPARγ is a critical mediator of CC-induced barrier protection. The TcdA group’s p-P65/P65 rose in comparison to the Control group (p < 0.001, Figure 4(A,B)). The TcdA+CC group showed a decline in p-P65/P65 when compared to the TcdA group (p < 0.001, Figure 4(A,B)). Furthermore, compared with the TcdA+CC+shNC group, the p-P65/P65 rose in the TcdA+CC+shPPARγ group (p < 0.001, Figure 4(A,B)). This suggests that PPARγ knockdown reactivates NF-κB signaling even in the presence of CC, suggesting that PPARγ may function upstream of NF-κB in mediating the protective effect of CC. We also examined IL‑8 and TNF‑α levels under PPARγ knockdown conditions. As shown in Figure 4(C,D), CC treatment markedly reduced TcdA-induced IL-8 and TNF-α production (p < 0.001). PPARγ knockdown (shPPARγ) significantly reversed this inhibitory effect (p < 0.01), indicating that CC suppresses inflammation at least in part through PPARγ.

Figure 3.

CC reverses TcdA-induced intestinal epithelial barrier dysfunction through PPARγ.

(A–B) Western blot analysis of PPARγ levels in Control, TcdA, TcdA+CC, TcdA+CC+short hairpin RNA negative control (shNC), and TcdA+CC+short hairpin RNA targeting PPARγ (shPPARγ) groups. GAPDH served as a loading control. (C) LDH levels measured by CytoTox 96 non-radioactive cytotoxicity assay. (D) TEER measurements of Caco-2 monolayers. (E) Paracellular Lucifer yellow flux in Caco-2 cells. Catechin was added after TcdA exposure (therapeutic protocol). Data are mean ± standard deviation (n = 3 independent biological replicates). ***p < 0.001 vs. Control; ^{^^^}p < 0.001 vs. TcdA; ^##p < 0.01, ^###p < 0.001 vs. TcdA+CC+shNC.

Figure 4.

CC inhibits NF-κB activation in TcdA-treated Caco-2 cells through PPARγ.

(A–B) Western blot analysis of phosphorylated P65/total P65 (p-P65/P65) levels in Control, TcdA, TcdA+CC, TcdA+CC+shNC, and TcdA+CC+shPPARγ groups. GAPDH was used as a loading control. (C–D) ELISA measurement of IL-8 (C) and TNF-α (D) levels in Control, TcdA, TcdA+CC, TcdA+CC+shNC, and TcdA+CC+shPPARγ groups. (E–H) Western blot analysis of tight junction proteins ZO‑1 (F), Occludin (G), Claudin1 (H) in Control, TcdA, TcdA+CC, TcdA+CC+shNC, and TcdA+CC+shPPARγ groups. GAPDH was used as a loading control. Catechin was added after TcdA exposure (therapeutic protocol). Data are mean ± standard deviation (n = 3 independent biological replicates). ***p < 0.001 vs. Control; ^{^^^}p < 0.001 vs. TcdA; ^###p < 0.001 vs. TcdA+CC+shNC.

We further examined the expression of tight junction proteins ZO-1, Occludin, and Claudin-1 under PPARγ knockdown. As shown in Figure 4(E-H), TcdA reduced all three proteins (p < 0.001). CC treatment restored their expression to near-control levels (p < 0.001). PPARγ knockdown (shPPARγ) significantly reversed the restorative effect of CC, reducing ZO‑1, Occludin, and Claudin-1 back to levels comparable to the TcdA group (p < 0.001). These results confirm that CC protects intestinal epithelial barrier integrity by restoring tight junction proteins in a PPARγ‑dependent manner.

4. Discussion

This study investigated the protective effects of CC on intestinal epithelial barrier dysfunction induced by TcdA in Caco-2 cells. CC exhibits potent anti-inflammatory, antioxidant, and cytoprotective properties, helping to mitigate cellular damage caused by oxidative stress and inflammatory stimuli [19]. These effects contribute to the maintenance of cellular homeostasis and the protection of tissue integrity under pathological conditions. A previous study has reported that CC alleviates diabetic nephropathy by inhibiting endoplasmic reticulum stress-related nucleotide-binding oligomerization domain-like receptor family pyrin domain containing 3 (NLRP3) inflammasome signaling [20]. CC also reduces porphyromonas gingivalis-induced inflammation by regulating Toll-like receptor 2/4 (TLR2/4) and inflammasome pathways [21]. And CC protects against doxorubicin-induced oxidative stress and inflammatory cardiotoxicity in rats [22]. These reports emphasize the role of CC in modulating inflammation and oxidative stress, but few studies have explored its direct effects on intestinal epithelial barrier function during C. difficile toxin exposure. In the present study, treatment with TcdA markedly disrupted the intestinal epithelial barrier. CC significantly ameliorated all of these injury markers, indicating a strong barrier-protective effect. The integrity of the intestinal epithelial barrier depends critically on tight junction proteins such as ZO‑1, occludin, and claudins [23]. Our data showed that TcdA substantially reduced the expression of these proteins, and CC treatment restored them in a concentration‑dependent manner. This provides direct molecular evidence that CC protects barrier function by preserving tight junction integrity, which is consistent with the observed recovery of TEER and reduction in paracellular flux. This indicates, for the first time, that this study systematically demonstrates the direct protective effect of CC against TcdA-induced intestinal epithelial barrier dysfunction, extending its known anti-inflammatory and antioxidant properties.

Further analysis demonstrated that TcdA downregulated PPARγ expression but activated NF-κB signaling, whereas CC reversed these changes and restored barrier homeostasis. Consistent with previous reports in C. difficile toxin models [15,24]. Our findings show that TcdA activates NF-κB signaling, leading to barrier dysfunction in Caco-2 cells. Activation of NF‑κB typically leads to production of pro‑inflammatory cytokines such as IL‑8 and TNF‑α, which amplify inflammation [25]. In line with this, we observed that TcdA significantly increased IL‑8 and TNF-α levels, and CC treatment suppressed their production. Moreover, PPARγ knockdown not only reversed the protective effect of CC on tight junction proteins but also abrogated the suppressive effect on IL-8 and TNF‑α, further confirming that CC exerts its anti-inflammatory and barrier-protective effects through the PPARγ–NF-κB axis. Moreover, PPARγ serves as an important regulator in this process. Importantly, unlike previous work focused only on NF-κB inhibition, the present study provides evidence that CC counteracts TcdA-mediated injury by regulating the PPARγ–NF-κB axis. Specifically, CC restored PPARγ expression, which in turn inhibited NF-κB activation. This study suggests the pivotal role of the upstream regulator PPARγ in barrier restoration, suggesting that activating or stabilizing PPARγ may represent a novel strategy to suppress NF-κB-mediated barrier injury. Moreover, the PPARγ–NF-κB axis appears to be critical in TcdA-induced intestinal epithelial barrier dysfunction, and the protective effect of CC relies at least in part on this pathway. Although our PPARγ knockdown experiments support a functional role for PPARγ in the protective effect of CC against TcdA, they do not definitively establish causality. It remains possible that CC regulates NF‑κB activity through other PPARγ‑independent mechanisms, such as direct antioxidant effects or modulation of other transcription factors [26,27]. Further studies using PPARγ agonists or genetic rescue experiments are needed to establish a direct causal relationship.

This study has several limitations that should be considered. First, all experiments were performed exclusively in Caco-2 cell monolayers, a model that lacks both immune cells and microbiota key components of the intestinal microenvironment. Therefore, our findings cannot fully replicate the complexity of the gut and should be extrapolated to physiological conditions with caution. Therefore, our findings should be extrapolated to physiological conditions with caution, and future in vivo studies using appropriate animal models (e.g., TcdA‑induced murine colitis) are required to confirm the physiological relevance and pharmacokinetic properties of CC. Such models would allow evaluation of CC’s oral bioavailability, intestinal tissue distribution, and its ability to suppress NF‑κB activation and restore barrier function in a complex physiological environment with intact immune and microbiota components. Second, only TcdA was examined; TcdB is also a key virulence factor of C. difficile and may cause epithelial damage. TcdA was selected as it is the primary toxin mediating early barrier injury, but future studies should test CC against TcdB or the combination of both toxins. Third, the catechin concentrations used (25–100 μg/mL) are higher than typical plasma levels but consistent with previous Caco‑2 studies [17]; LDH data at 100 μg/mL showed no cytotoxicity, although non‑specific effects at high doses cannot be completely excluded. Fourth, the precise molecular mechanism by which PPARγ regulates NF‑κB remains unclear and warrants further investigation.

5. Conclusion

In conclusion, this study indicates that CC attenuates TcdA-induced intestinal epithelial barrier dysfunction and is associated with modulation of the PPARγ–NF-κB signaling pathway. CC treatment restores PPARγ expression and suppresses NF-κB activation, thereby contributing to the maintenance of epithelial barrier integrity. These findings suggest a regulatory role of PPARγ in barrier protection and provide preliminary experimental evidence supporting further investigation of natural polyphenols as potential therapeutic candidates for intestinal barrier disorders. While these results are limited to an in vitro model, they offer a rational basis for future in vivo investigations and, ultimately, for exploring CC‑based dietary or pharmacological strategies to mitigate toxin‑mediated intestinal injury in clinical settings.

Funding Statement

This work was supported by the Zhejiang Provincial Traditional Chinese Medicine Science and Technology Project [No. 2025ZL559], “Shuangqing” Talent Development Program of Jiaxing Traditional Chinese Medicine Hospital, Zhejiang Province, Clinical ”Small but Strong” Innovation Team of Jiaxing Traditional Chinese Medicine Hospital, Zhejiang Province, Zhejiang Traditional Chinese Medicine Science and Technology Project [No. 2026ZL0873], Jiaxing University Horizontal Research Project “Open Competition Mechanism” [No. KY20260105399]. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Article highlights

• (+)-Catechin attenuates Clostridioides difficile toxin A (TcdA)-induced intestinal epithelial barrier dysfunction in Caco-2 cells.

• (+)-Catechin restores tight junction proteins (ZO-1, occludin, claudin-1) and suppresses pro-inflammatory cytokines (IL-8, TNF-α).

• The barrier-protective effect of (+)-catechin is mediated through the PPARγ–NF-κB signaling axis, with PPARγ serving as a key upstream regulator.

Disclosure statement

The authors report that there are no competing interests to declare. The authors have no relevant affiliations or financial involvement with any organization or entity with a financial interest in or financial conflict with the subject matter or materials discussed in the manuscript. This includes employment, consultancies, honoraria, stock ownership or options, expert testimony, grants or patents received or pending, or royalties.

Data availability statement

The data that support the findings of this study are available from the corresponding author KC upon reasonable request.