Multiomics Reveals Nonphagocytosable Microplastics Induce Colon Inflammatory Injury via Bile Acid-Gut Microbiota Interactions and Barrier Dysfunction

PS MPs are typical microplastics widely present in the environment and commonly used for toxicity studies. This study utilized two types of PS MPs (Zhichuan, Jiangsu, China). Conventional PS MPs, with a particle size of 10 μm, were used for toxicological research in animal and cell experiments. Additionally, fluorescent PS MPs (excitation wavelength: 520 nm; emission wavelength: 580 nm) with particle sizes of 100 nm, 1 μm, and 10 μm were employed to visualize ingestion, accumulation, and distribution in vivo and in vitro. The morphology and particle size of the PS MPs were confirmed using scanning electron microscopy (SEM, Sigma 300, ZEISS, Germany) and a laser particle size analyzer (Mastersizer 2000, Malvern, UK). The chemical composition of the PS MPs was verified through Fourier-transform infrared spectroscopy (FTIR; IRTracer 100, Shimadzu, Japan). Taurochenodeoxycholic acid (TCDCA) was obtained from Aladdin (CAS#: 516-35-8, Shanghai, China). The cell counting kit-8 (CCK-8), calcein/propidium iodide (PI) Live/Dead viability/cytotoxicity assay kit, reactive oxygen species (ROS) assay kit, and mitochondrial membrane potential assay kit with JC-1 were provided by Beyotime Biotechnology, Shanghai, China. The Annexin V-FITC/PI apoptosis detection kit was supplied by Jiangsu KeyGEN Biotechnology, China.

SPF BALB/c mice (4 weeks old, male) were obtained from the Anhui Experimental Animal Center (Hefei, Anhui, China). The mice were housed at 22–26 °C with controlled humidity and a 12 h light/dark cycle, with 1 week of acclimatization. All animal experiments utilized 10 μm PS MPs, diluted in PBS to a final concentration of 10 mg/mL. All experimental protocols were approved by the Animal Care and Use Committee of Anhui Medical University (ethics approval number: LLSC20242418).

The human normal colonic epithelial cell line (NCM460) was sourced from Warner Bio (Wuhan, China). NCM460 cells were cultured in RPMI1640 medium (Gibco, Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS; Gibco), 100 IU/mL penicillin, and 100 μg/mL streptomycin at 37 °C with 5% CO₂. Once the cell density exceeded 90%, they were seeded into six-well plates, 24-well plates, or 96-well plates for subsequent experiments.

Six healthy mice were randomly selected and fasted for 8 h. Fluorescent PS MPs at a concentration of 10 mg/mL were orally administered at 0, 2, 4, 8, and 12 h, with a gavage volume of 200 μL per mouse. The control group received an equivalent volume of PBS. Mice were anesthetized with pentobarbital sodium administered intraperitoneally. Using Small Animal In Vivo Optical Imaging (IVIS Luminz iii, PerkinElmer, USA), images were captured at each time point to monitor the ingestion, digestion, and accumulation of fluorescent PS MPs throughout the digestive tract, from the oral cavity to the anus, as well as their distribution in various organs.

Five healthy mice were randomly selected and orally administered fluorescent PS MPs at a concentration of 10 mg/mL, with a daily dosage of 1 mg per mouse for 6 weeks. After the exposure period, the mice were euthanized, and tissue samples, including the heart, lungs, liver, spleen, kidneys, stomach, and colon, were collected. The tissues were fixed in 4% paraformaldehyde, embedded in paraffin, and sectioned to a thickness of 4 μm. Hematoxylin and eosin (H&E) staining was performed for observation. To detect the presence of fluorescent PS MPs in tissue sections, a bright-field image was first obtained using a Leica upright fluorescence microscope, followed by fluorescence imaging in the appropriate channel. The two sets of images were then merged for analysis.

After 1 week of acclimatization, 10 healthy mice were randomly selected and weighed. They were subsequently divided into two groups: the control group (NC) and the model group (MP), each consisting of 5 mice. During gavage, the body weight of the mice was measured every other day. The MP group received a 10 mg/mL concentration of PS MPs at a dosage of 1 mg/day per mouse, while the control group was administered an equivalent volume of PBS. Gavage continued for 6 weeks. Body weights of both groups were measured before the final gavage, and mice were fasted for 8 h after the final gavage. Euthanasia was carried out by cervical dislocation following intraperitoneal anesthesia with pentobarbital sodium. Tissues, including serum, feces, heart, lungs, liver, spleen, kidneys, stomach, cecum, and colon, were collected.

An appropriate number of NCM460 cells were seeded in a 24-well plate and cultured for 24 h until stable growth and proliferation were achieved. Fluorescent PS MPs of varying sizes (100 nm, 1 μm, 10 μm) were then coincubated with the cells at a concentration of 1 mg/mL for another 24 h. After coincubation, the wells were washed several times with PBS to remove uninternalized MPs. Bright-field, red fluorescence, and DAPI fluorescence images were captured using a live cell workstation (Celldiscoverer 7, Zeiss, Germany). The images were exported and merged using the ZEN software provided by Zeiss as necessary.

Histopathological examinations were performed to assess inflammation in the colon and liver, as well as colonic mucus secretion. Colon and liver tissue samples were fixed in 4% paraformaldehyde, dehydrated, paraffin-embedded, and sectioned into 4 μm slices. The sections were stained with H&E and alcian blue periodic acid-Schiff (AB-PAS) reagents from Solarbio (Beijing, China). Tissue sections were observed and images were captured using a Leica upright microscope (DM6B, Leica, Germany).

Enzyme-linked immunosorbent assay (ELISA) kits were used to determine the expression levels of inflammatory factors in colon tissue and TBA levels in the liver, colon, and feces. The ELISA kits, provided by Meibiao Biotechnology (Jiangsu, China), included assays for mouse tumor necrosis factor-α (TNF-α), interleukin-1 β (IL-1β), interleukin-6 (IL-6), and interleukin-10 (IL-10). Mouse TBA ELISA kits were obtained from Zhongke Quality Inspection Biotechnology (Beijing, China). After supernatants were extracted from colon, liver, and fecal samples, procedures were strictly followed according to the manufacturer’s protocols for the respective commercial ELISA kits. Optical density (OD) values were measured at a wavelength of 450 nm by using a microplate reader (BioTek, USA). Data analysis involved constructing standard curves, and the expression levels of inflammatory factors and TBA were calculated based on these curves.

For the detection of oxidative stress biomarkers in the colon and liver, mouse colon and liver tissue homogenates were prepared according to the kit manufacturer’s instructions. The levels of reduced glutathione (GSH) and oxidized glutathione (GSSG) were assessed by using a colorimetric assay (GSH and GSSG assay kit, S0053, Beyotime, Shanghai, China). Glutathione peroxidase (GSH-Px) content was determined using the NADPH method (total glutathione peroxidase assay kit, S0058, Beyotime). Superoxide dismutase (SOD) content was measured using the WST-8 method (total SOD activity assay kit S0101S, Beyotime). Malondialdehyde (MDA) levels were determined using an MDA content assay kit (BC0025, Solarbio), and catalase (CAT) activity was measured using a catalase activity assay kit (BC0205, Solarbio). Protein concentrations were determined using a BCA protein assay kit (P0012, Beyotime).

Splenic lymphocytes were isolated from murine specimens, followed by erythrocyte depletion using hypotonic lysis. After centrifugation and washing with PBS, cellular suspensions were preincubated with an Fc receptor-blocking agent to prevent nonspecific antibody interactions. Surface epitopes were labeled for 30 min at 37 °C using the following fluorophore-conjugated monoclonal antibodies: PB450-anti-CD45, FITC-anti-CD4, APC-anti-CD8, PerCP/Cy5.5-anti-CD3, and PE-anti-CD25. Intracellular staining was performed after fixation and permeabilization with APC-conjugated Foxp3 and PE/Cy7-tagged IL-17A antibodies under identical thermal conditions. Processed samples were resuspended in a stabilization buffer and analyzed by using flow cytometry (Beckman Coulter CytoFLEX platform). Quantitative data interpretation was performed using FlowJo software (version 10.8.1, Tree Star Inc.).

Paraffin-embedded mouse colon tissue blocks were sectioned to a thickness of 3 μm and dewaxed in water. Endogenous peroxidase activity was blocked using an endogenous peroxidase blocking solution (AR1108, Boster, Wuhan, China) for 10 min at room temperature, followed by a wash with PBS. Antigen retrieval was performed using a microwave, and the tissue was then blocked with 5% BSA (BS114-5g, Biosharp, Hefei, Anhui, China) at room temperature for 2 h. Primary antibodies, including ZO-1 (1:200 dilution, 21773-1-AP, Proteintech, Wuhan, China), Occludin (1:200 dilution, 27260-1-AP, Proteintech), Claudin 1 (1:200 dilution, 13050-1-AP, Proteintech), and MUC1 (1:100 dilution, ET1611-14, Huabio, Hangzhou, China), were incubated overnight at 4 °C. The sections were then brought to room temperature, washed with PBS to remove unbound primary antibodies, and dried by gently wiping away excess water. Subsequently, the sections were incubated in the dark with FITC-labeled secondary antibodies (BL033A, Biosharp) at room temperature for 2 h. After incubation, the sections were washed with PBS on a shaker and mounted using an antifade mounting medium containing DAPI (P0131, Beyotime). Observations and imaging were performed with a Leica upright microscope.

Feces (20 mg) were ground and extracted with 200 μL of methanol/acetonitrile (v/v = 2:8), then stored at −20 °C for 10 min for quantification. The mixture was then centrifuged for 10 min at 12,000 rpm and 4 °C to obtain the supernatant. The supernatant was analyzed using an liquid chromatography-electrospray ionization-tandem mass spectrometry (LC-ESI-MS/MS) system equipped with a Waters ACQUITY UPLC HSS T3 C18 column (100 × 2.1 mm i.d., 1.8 μm), consisting of UHPLC (ExionLC AD) and MS (Applied Biosystems 6500 Triple Quadrupole). Mobile phase A consisted of water with 0.01% acetic acid and 5 mmol/L ammonium acetate, while mobile phase B consisted of acetonitrile with 0.01% acetic acid. The column temperature was set to 40 °C, and the injection volume was 3 μL. Detection was carried out using the QTRAP 6500+ LC–MS/MS mass spectrometer system (SCIEX, USA), equipped with an ESI Turbo ion–spray interface, operating in negative ion mode, and controlled by Analyst 1.6.3 software (Sciex). The ESI source parameters were as follows: ion source, ESI; source temperature, 550 °C; ion spray voltage (IS), −4500 V; curtain gas (CUR), 35 psi.

Sequencing was conducted by MetWare (http://www.metware.cn/↗) using the NovaSeq 6000 platform (Illumina, USA). Following the acquisition of raw sequencing data, filtering and merging were performed with Fastp (v0.22.0), FLASH (v1.2.11), and Vsearch (v2.22.1) to generate effective tags. Operational taxonomic unit (OTU) clustering and amplicon sequence variant (ASV) denoising were carried out based on these effective tags. Taxonomic annotation and multiple sequence alignment analyses were then performed on the sequences of the OTU/ASV, providing species identification and abundance distribution profiles. Differences in the community structure across various samples or groups were explored. The data were subsequently normalized, followed by analyses such as alpha diversity, beta diversity, differential species analysis with significance testing, network analysis, and functional prediction based on the normalized data.

For cell viability assays, NCM460 cells were seeded in a 96-well plate. After 24 h, the culture medium was removed, and various concentrations of PS MPs or TCDCA, diluted in RPMI1640 medium, were added. After 24 h of incubation, CCK-8 reagent was added, followed by a 2 h incubation. The OD values at 450 nm were measured by using a microplate reader, and cell viability was calculated.

NCM460 cells were also seeded into a 24-well plate and treated with specified concentrations of PS MPs or TCDCA for a designated period. Calcein AM/PI detection working solution was prepared according to the manufacturer’s instructions. After the culture medium was removed, cells were washed once with PBS, and residual liquid was removed. Then, 250 μL of the Calcein AM/PI working solution was added to each well, and the plate was incubated at 37 °C in the dark for 30 min. Following incubation, staining was observed under a fluorescence microscope and images were captured. Calcein AM emits green fluorescence (Ex/Em = 494/517 nm), while PI emits red fluorescence (Ex/Em = 535/617 nm).

For ROS detection, DCFH-DA was diluted in a serum-free culture medium at a 1:1000 ratio to achieve a final concentration of 10 μM. After removing the culture medium, an appropriate volume of diluted DCFH-DA was added to cover the cells, which were then incubated at 37 °C for 20 min. The cells were washed three times with a serum-free medium to remove any unincorporated DCFH-DA. Rosup was added to the positive control well. Finally, the cells were observed, and images were captured using a live-cell workstation at Ex/Em = 488/525 nm.

Following the manufacturer’s instructions, the JC-1 staining working solution was prepared. CCCP was diluted to 10 μM and used to treat the cells for 20 min as a positive control. The culture medium was removed from the 24-well plate, and the cells were washed once with PBS. Next, 1 mL of complete culture medium was added, followed by 1 mL of JC-1 staining working solution, which was mixed gently and thoroughly. The plate was incubated at 37 °C in a cell culture incubator for 20 min. During incubation, a 1× JC-1 staining buffer was prepared and kept on ice. After incubation, the supernatant was removed and the cells were washed twice with 1× JC-1 staining buffer. Finally, 1 mL of complete culture medium was added, and the cells were observed and imaged using a live-cell workstation. JC-1 aggregates emit red fluorescence (Ex/Em = 525/590 nm), while JC-1 monomers emit green fluorescence (Ex/Em = 490/530 nm).

For apoptosis detection, cells from each group were first collected after trypsin digestion without EDTA. The cells were washed twice with PBS, centrifuged at 800 rpm for 5 min, and the supernatant was discarded. Then, 500 μL of binding buffer from the kit was added to the cells, mixed gently to form a single-cell suspension, and transferred to flow cytometry tubes. Next, 5 μL of Annexin V-FITC was added to each tube and mixed thoroughly, followed by the addition of 5 μL of PI to each tube and gently mixed again. The tubes were incubated in the dark at room temperature for 10 min. Within 1 h, flow cytometry was performed to observe and detect the cells. The green fluorescence of Annexin V-FITC (Ex/Em = 488/530 nm) was detected through the FITC channel (FL1), and the red fluorescence of PI (Ex/Em = 488/630 nm) was detected through the PI channel (FL3).

For protein extraction, cells were lysed on ice using cell lysis buffer (P0013, Beyotime) containing the protease inhibitor PMSF (ST506, Beyotime, China). The protein concentration was determined using a BCA protein assay kit. Proteins were separated by 10% SDS-PAGE and transferred onto a PVDF membrane. The membrane was blocked in 5% skim milk for 1 h and then incubated overnight at 4 °C with appropriately diluted primary antibodies (Bax, Bcl-2, Caspase-3, Caspase-9, PARP, GAPDH). The next day, after washing three times with TBST, the membrane was incubated with corresponding secondary antibodies for 1 h. The membrane was washed three times with TBST, stained with enhanced chemiluminescence (ECL) reagents (Affinity, China), and imaged and analyzed using a chemiluminescence imaging system (Tanon, Shanghai, China).

All mice were acclimated for 1 week and weighed before being randomly assigned to four groups: the control group (NC), the experimental group (MP), the MP + ABX group, and the MP + TCDCA group, with five mice per group. PS MPs were administered via gavage to all experimental groups at a concentration of 10 mg/mL, with a dosage of 1 mg per mouse per day, while the control group received an equivalent volume of PBS. Gavage was performed continuously for 6 weeks. In the MP + ABX group, an antibiotic mixture (ABX) was provided via drinking water, consisting of 0.5 g/L vancomycin (CAS#: 1404-93-9, Aladdin), 1 g/L neomycin sulfate (CAS#: 1405-10-3, Aladdin), 1 g/L metronidazole (CAS#: 443-48-1, Aladdin), and 1 g/L ampicillin (CAS#: 69-52-3, Aladdin). The solution was replaced every 3 days for 6 weeks to deplete the intestinal microbiota. From the fifth week onward, mice in the MP + TCDCA group were orally administered with TCDCA dissolved in PBS at a dose of 200 mg/kg for 2 weeks. Prior to the final gavage, all mice were weighed, fasted for 8 h, and euthanized via cervical dislocation under pentobarbital sodium anesthesia. Samples including serum, feces, heart, lungs, liver, spleen, kidneys, stomach, cecum, and colon tissues were collected.

Data are presented as mean ± SEM. Statistical analyses were performed by using GraphPad Prism 10.1 software. One-way ANOVA followed by post hoc Tukey tests were used for statistical comparisons between groups. A P-value of <0.05 was considered statistically significant.

SEM was employed to capture the overall morphology of the PS MPs samples. Both low- and high-magnification images revealed that PS MPs of three different diameters displayed spherical structures with uniform sizes of 100 nm, 1 μm, and 10 μm, indicating excellent monodispersity (FigureA). Subsequently, a laser particle size analyzer was used to measure the size and distribution of the PS MPs, confirming that the average sizes were centered around 100 nm, 1 μm, and 10 μm, consistent with the SEM observations (FigureB). FTIR spectroscopy was then applied to analyze the chemical composition of the samples, and the characteristic absorption peaks of polystyrene were observed, confirming the material as polystyrene (FigureC).

The characterization of PS MPs demonstrated their excellent experimental properties. To evaluate the phagocytic capacity of colonic epithelial cells for PS MPs of varying sizes, NCM460 cells were cultured in vitro and coincubated with PS MPs of three different sizes for 24 h. The results indicated that 100 nm PS MPs were most readily engulfed by colonic epithelial cells, followed by 1 μm PS MPs, while 10 μm PS MPs could not be engulfed (FiguresA, S1↗).

To further investigate the uptake and distribution of 10 μm PS MPs following a single high-dose oral exposure (0–12 h) in vivo, small animal live imaging technology was used to simulate the digestion process of PS MPs in the gastrointestinal tract after ingestion by mice and explore their distribution across various tissues and organs at different time points (FigureB). The live imaging results showed that at 0 h, strong fluorescence signals from red fluorescent PS MPs were detected in the stomach immediately after gavage, with no notable signals in the intestines or other organs. At 2 h, PS MPs had reached the midlower abdomen, with significant fluorescence signals detected in the cecum and parts of the colon, while undigested PS MPs were still present in the stomach. By 4 h, the fluorescence signal in the stomach disappeared, suggesting the possible complete digestion of the PS MPs, while the MPs were primarily distributed in the left abdomen, with bright fluorescence signals observed in the left colon and rectum. At 8 h, as PS MPs were gradually excreted, the fluorescence signal in the abdomen diminished, accompanied by a decrease in fluorescence signals in the cecum and colon. A pronounced increase in fluorescence signal was observed around the anus. By 12 h, the overall fluorescence signals in the abdominal area significantly decreased, indicating that most PS MPs had been metabolized and excreted from the body. Furthermore, during this single high-dose oral exposure, no fluorescence signals were detected in organs beyond the gastrointestinal tract. In vivo imaging confirmed that single high-dose oral exposure to PS MPs primarily affected the gastrointestinal tract, with no evidence suggesting that distant organs were invaded by PS MPs.

However, long-term exposure to PS MPs in vivo led to accumulation in distant organs. After 6 weeks of gavage with red fluorescent PS MPs, histological tissue sections revealed red fluorescent particle accumulation in the colon, liver, and kidneys (FigureC). These results suggest that prolonged exposure to PS MPs may not only result in accumulation within the intestines but also lead to the accumulation of MPs in multiple organs.

Following prolonged exposure to PS MPs, changes in mouse body weight were first observed (FiguresB,C, S2A↗). The body weight growth curve indicated no significant difference between the MP and NC groups before the initiation of gavage (P > 0.05). However, during the gavage period, the body weight gain rate in the MP group was notably slower than that in the NC group. By the end of the gavage period, the body weight of mice in the MP group was significantly lower than that of the NC group (P < 0.05). After euthanizing the mice and collecting specimens, comparisons of colonic length and weight revealed that both were significantly reduced in the MP group compared to the NC group (P < 0.05) (FigureA,B). Furthermore, H&E staining of the MP group showed significant infiltration of inflammatory cells (FigureD). ELISA results showed that pro-inflammatory cytokines TNF-α, IL-1β, and IL-6 were significantly higher in the MP group compared to the NC group, while the anti-inflammatory cytokine IL-10 was significantly reduced (P < 0.05) (FigureE). These results suggest that prolonged exposure to PS MPs slowed weight gain in mice and induced colonic inflammation.

Analysis of oxidative stress markers in colonic tissues (FiguresF, S2B↗) showed that prolonged exposure to PS MPs significantly increased the levels of GSSG and MDA, while GSH levels were significantly decreased in the MP group (P < 0.05). Additionally, the expression levels of antioxidant enzymes GSH-Px, SOD, and CAT were significantly lower in the MP group compared to the NC group (P < 0.05). These results indicate that prolonged exposure to PS MPs caused an imbalance in the oxidative–reductive system of the mouse colon, leading to oxidative stress-induced damage to colonic tissues.

Th17/Treg cells contribute to the development of intestinal inflammation. To investigate whether PS MPs-induced colonic inflammation is linked to changes in the Th17/Treg ratio in mice, flow cytometry was employed to assess the ratio of Th17 and Treg cells in the spleen (FiguresA, S3↗). Quantitative analysis of CD4+ IL17A+ (Th17) and CD4+ CD25+ Foxp3+ (Treg) cells within spleen CD4+ T cells revealed that, compared to the control group, PS MPs significantly increased Th17 cell levels and decreased Treg cell levels, resulting in a notable elevation of the Th17/Treg ratio (P < 0.05). These results suggest that PS MPs exposure selectively increases Th17 levels while reducing Treg levels, leading to a disruption of intestinal immune homeostasis.

The intestinal mucosal epithelial barrier is a primary protective mechanism that shields the gut from damage by harmful stimuli. MPs may compromise this barrier, potentially triggering inflammation. AB-PAS staining showed that the MP group exhibited a reduction in goblet cells within the colonic mucosa, accompanied by decreased mucin secretion, leading to a reduced mucin coverage area in the colon compared to that in the control group (FigureB). Additionally, immunofluorescence (IF) staining of colon tissues was used to assess the expression of intestinal barrier proteins (FigureC). The results demonstrated a decrease in fluorescence intensity and lower expression levels of ZO-1, Occludin, Claudin 1, and MUC1 in the MP group.

In conclusion, long-term exposure to PS MPs results in colon oxidative stress, an imbalance in the Th17/Treg immune response, extensive infiltration of inflammatory cells in colon tissues, increased secretion of pro-inflammatory cytokines, and disruption of the intestinal epithelial barrier. These results suggest that PS MPs exposure contributes to dysregulation of intestinal homeostasis and induction of colonic inflammation in mice.

The long-term effects of PS MPs exposure on distant organs, such as the liver, were also investigated. FigureA presents specimens from the NC and MP groups, including the heart, liver, spleen, lungs, kidneys, and stomach. No significant differences in the appearance of these organs were observed between the two groups. However, upon weighing the organs, no significant differences were found in the masses of the heart, lungs, and stomach (P > 0.05). In contrast, the liver and spleen mass were significantly increased, while the kidney mass was significantly decreased in the MP group compared to the control group (P < 0.05) (FiguresB, S4↗). In addition to the intestine, the liver is one of the most commonly studied distant organs. H&E staining of liver tissue sections (FigureC) revealed inflammatory damage in the liver of the MP group. Furthermore, the activities of hepatic antioxidant enzymes, including GSH-Px, SOD, and CAT, were significantly reduced in the MP group (FigureE). A significant increase in TBA was also detected in the liver of the MP group (FigureD). These findings, combined with the results from FigureC, suggest that long-term PS MPs exposure leads to their accumulation in the liver, causing inflammatory damage, impairing antioxidant function, and disrupting BA metabolism.

Additionally, TBA levels in the colon and feces were significantly elevated in the MP group compared to the control group (P < 0.05) (FigureD). This suggests that the elevated hepatic TBA levels in the MP group result in increased secretion of TBA into the intestine, which, in turn, raises the fecal TBA levels. The mechanism underlying colonic inflammation and damage induced by PS MPs exposure may be related to elevated BA levels. Intestinal microbiota, which can convert primary BAs into secondary BAs, may play a pivotal role in this process. To explore this further, multiomics analysis of fecal BA metabolism and changes in intestinal microbiota were performed using BA metabolism metabolomics and 16S rRNA microbial sequencing technologies (FigureE).

BA metabolomic sequencing was conducted on fecal samples from the previously mentioned experiments. Principal component analysis (PCA) of fecal samples from both groups revealed that long-term exposure to PS MPs significantly altered the overall structure of fecal BAs, leading to notable differences in overall metabolism and increased variability (FigureA). Subsequently, cluster heatmap analysis was performed on all samples from both groups to observe changes in various BAs, as shown in FigureB. Using cluster analysis and the orthogonal partial least squares-discriminant analysis (OPLS-DA) model, variable importance in projection (VIP) scores were obtained. Differential BAs were selected based on P-values and fold change (FC) values from univariate analysis. FigureC highlights the top 20 differential BAs in terms of FC between the NC and MP groups, with the majority of BAs being elevated in the MP group. To further analyze the patterns of BA changes, the original levels of the selected differential BAs were normalized using unit variance scaling (UV scaling). A total of 17 differential BAs were identified, with 14 upregulated and 3 downregulated. Cluster heatmap analysis of these differential BAs is presented in FigureD. Additionally, violin plots were used to visually display the differences in levels and the overall distribution of differential BAs between the two groups (FiguresE, S5↗). These results clearly demonstrate the global disruption of BA metabolism induced by prolonged PS MPs exposure. Fecal BAs were predominantly elevated, with 14 out of 17 differential BAs (82.4%) showing significant increases, which aligns with enhanced hepatic synthesis and secretion (refer to liver and colon TBA data, FigureD). Among the 14 upregulated BAs, 11 were conjugated types (78.6%), with taurine-conjugated BAs predominating (8 out of 11, 72.7%). Furthermore, this abnormal BA profile may be linked to impaired gut microbiota functionality, suggesting the need for further investigation to elucidate the underlying mechanism.

In 16S rRNA sequencing, the OTUs were identified through clustering methods, and a Venn diagram was generated. The analysis revealed 724 OTUs shared between the NC and MP groups, with 372 unique to the NC group and 397 unique to the MP group (FigureA). Further investigation of species with high relative abundance at each taxonomic level and their proportional changes was conducted based on OTUs (FiguresB, S6A↗). The results demonstrated that PS MPs exposure induced multilevel shifts in the microbiota. At the phylum level, Firmicutes and Actinobacteriota were significantly enriched, while Bacteroidota, Proteobacteria, and Verrucomicrobiota were markedly depleted. These phylum-level alterations were driven by dynamic changes in the key families and genera. Firmicutes enrichment correlated with increased abundance of Lachnospiraceae and its associated genera and . In contrast, Bacteroidota depletion was linked to reduced levels of Bacteroidaceae and its dominant genus Bacteroides. At the species level, analysis revealed a bifurcation within the Lactobacillus genus: and were upregulated, while and the mucin-degrading species were suppressed. These taxon-specific responses suggest the selective regulation of microbial consortia involved in mucin metabolism and immunomodulation by PS MPs. To assess microbial diversity and community structure, a multiple sequence alignment of the top 100 genera was performed, and their phylogenetic relationships were inferred (Figure S6C↗). Principal component analysis (PCA) and principal coordinate analysis (PCoA) revealed significant alterations in the intestinal microbiota composition induced by PS MPs (FigureC). Simper analysis identified key taxa with intergroup differences, including Firmicutes, Bacteroidota, and Proteobacteria at the phylum level, as well as , , and at the genus level (Figure S6B↗). LEfSe analysis further highlighted 8 bacterial taxa significantly affected by PS MPs (Figure S6D↗).

To further elucidate the interaction between gut microbiota and BAs under PS MPs exposure, Spearman correlation analysis was performed on differential microbiota and BA metabolites (FiguresD, S7A,B↗). The results (P < 0.05) revealed significant positive correlations between elevated BAs and Actinobacteria at the phylum level, while negative correlations were observed with Proteobacteria. At the genus level, BAs positively correlated with potentially harmful bacteria such as and but negatively correlated with beneficial genera including , , and . Species-level analysis further showed that increased BA levels were positively associated with oxidative stress- and inflammation-inducing pathogens like , while negatively correlated with probiotics such as , , and , which play roles in maintaining barrier integrity, exerting anti-inflammatory effects, and regulating metabolism. These results suggest that PS MPs exposure may exacerbate colonic inflammation and injury by disrupting the correlation between specific microbiota and BAs, leading to the enrichment of potential pathogens and reduction of beneficial bacteria.

BA metabolism disorders are characterized by a significant elevation in conjugated BA levels, particularly tauro-conjugated species. To further investigate the role of BAs in PS MPs-induced colitis and intestinal injury, TCDCA, a representative tauro-conjugated BA, was selected for this study. CCK-8 assays were conducted to determine optimal treatment concentrations, revealing that PS MPs (0.5–2 mg/mL) and TCDCA (≥1200 μM) induced dose-dependent cytotoxicity in NCM460 cells (FiguresA, S8A↗). At 0.5 mg/mL, PS MPs reduced cell viability to 88.43%, while concentrations ≥2 mg/mL resulted in near-total cell death. TCDCA ≥1200 μM also exhibited significant toxicity. To minimize PS MPs-induced mortality while examining TCDCA’s role, 0.5 mg/mL PS MPs were combined with graded concentrations of TCDCA. The combination of 0.5 mg/mL PS MPs with 1000 μM TCDCA resulted in severe cytotoxicity (13.22% viability). The experimental groups included NC (control), MP (0.5 mg/mL PS MPs), TCDCA (1000 μM), and MP + TCDCA (combined treatment).

FigureB presents Calcein-AM/PI staining results where live (green) and dead (red) cells reflect the degree of damage. The NC group exhibited minimal red fluorescence, indicating normal cell viability. The MP group showed slight red signals, while TCDCA treatment increased red fluorescence moderately. Notably, the MP + TCDCA combination induced a dramatic increase in red signals, demonstrating synergistic cytotoxicity. ROS analysis (FigureA) revealed negligible green fluorescence in control cells, whereas intense signals were observed in Rosup-treated cells. MP exposure induced mild ROS generation, which was further enhanced by TCDCA. The combination group exhibited the strongest fluorescence, indicating significant ROS overproduction. JC-1 staining (FigureB) was used to assess mitochondrial membrane potential, with red/green fluorescence shifts indicating changes. NC cells showed predominant red signals (intact mitochondria), while CCCP-treated cells displayed a green dominance. Both MP and TCDCA treatments resulted in reduced red fluorescence compared with NC, with TCDCA showing slightly stronger green signals. The combination group showed near-complete conversion to green fluorescence, indicating severe mitochondrial dysfunction.

Subsequent flow cytometry and Western blot analyses provided further insights into apoptosis regulation. FigureC demonstrates an increase in apoptosis across groups: NC (1.33%), MP (5.48%), TCDCA (6.04%), and MP + TCDCA (29.95%), indicating synergistic pro-apoptotic effects. Western blot analysis of mitochondrial apoptosis markers (Figure S8B↗) showed consistent patterns when normalized to GAPDH. Pro-apoptotic proteins Bax, caspase-9, caspase-3, and PARP exhibited low expression in the NC group, increased in the MP and TCDCA groups, and reached the highest levels in the MP + TCDCA group, showing significant upregulation. Conversely, the antiapoptotic protein Bcl-2 exhibited the highest expression in the NC group, decreased in the MP and TCDCA groups, and was lowest in the MP + TCDCA group, demonstrating significant downregulation. These molecular changes align with the flow cytometry results, confirming that TCDCA significantly enhances PS MPs-induced apoptosis through activation of the mitochondrial pathway.

In vitro findings indicated that TCDCA exacerbates PS MPs-induced colonic epithelial damage, leading to subsequent in vivo validation. Broad-spectrum antibiotics (ABX) were used to deplete the gut microbiota, allowing for further investigation into how the microbiota–metabolite interaction influences colitis progression. Six-week murine experiments showed significant reductions in colonic length and weight in the MP group compared to NC controls (P < 0.05). Depletion of gut microbiota through ABX (MP + ABX group) mitigated these morphological changes, with significantly greater colonic length and weight observed compared to the MP group (P < 0.05). Conversely, TCDCA administration (MP + TCDCA group) exacerbated PS MPs-induced damage, with further reductions in colonic parameters compared to the MP group (P < 0.05, FiguresA,B, S9A↗).

Histopathological analysis (FigureC) revealed progressive mucosal alterations in the experimental groups. The NC group maintained intact mucosal architecture with well-organized intestinal villi and distinct crypt structures. MP exposure led to significant pathological changes, including villous atrophy, crypt disorganization, and substantial submucosal inflammatory infiltration, accompanied by vascular congestion and edema. Microbiota-depleted (MP + ABX) animals exhibited partial preservation of mucosal structures, with relatively ordered villi/crypt arrangements and attenuated inflammatory manifestations compared to the MP group. The MP + TCDCA group, however, exhibited severe mucosal destruction characterized by epithelial necrosis, total villous/crypt loss, and exacerbated submucosal vascular abnormalities with dense inflammatory infiltrates. ELISA analysis of inflammatory mediators (FiguresD, S9B↗) revealed distinct cytokine profiles. Pro-inflammatory cytokines (TNF-α, IL-1β, IL-6) were elevated in the MP, MP + ABX, and MP + TCDCA groups compared to NC (P < 0.05), with the MP + ABX group showing significant reduction and the MP + TCDCA group exhibiting maximal elevation compared to MP controls. Conversely, the anti-inflammatory cytokine IL-10 displayed an inverse pattern, with suppressed levels across all treatment groups (P < 0.05), partially restored in MP + ABX and further diminished in MP + TCDCA.

The integrated in vitro and in vivo findings demonstrate that TCDCA exacerbates PS MPs-induced colitis and intestinal injury in mice, with its interaction with the gut microbiota likely playing a critical role in the aggravated toxic effects of PS MPs.