DON standard was purchased from Qingdao Prubang Bioengineering Co., Ltd. Assay kits for superoxide dismutase (SOD), glutathione peroxidase (GSH-Px), MDA, and T-AOC were obtained from Nanjing Jiancheng Technology Co., Ltd. AG RNAex Pro RNA extraction reagent was sourced from Hunan Aikerui Bioengineering Co., Ltd. Hoechst 33342 staining solution, Annexin V-PE apoptosis detection kit, and CCK-8 reagent were supplied by Shanghai Biyuntian Biotechnology Co., Ltd. 2 × RealStar Fast dye qPCR premix and StarScript II genomic DNA removal reverse transcription premix reagents were purchased from Beijing Kangrun Chengye Biotechnology Co., Ltd. All qPCR primers were synthesized by Biotechnology (Shanghai) Co., Ltd. DMEM cell culture medium was obtained from Situofan Biotechnology Co., Ltd., and fetal bovine serum was procured from Shanghai Jiase Biotechnology Co., Ltd. Additional reagents including 0.25% trypsin solution and penicillin–streptomycin dual antibiotic solution were supplied by Wuhan Saiweier Biotechnology Co., Ltd.

The strain Lactobacillus rhamnosus MY-1 was isolated and preserved in our laboratory. The porcine intestinal epithelial cell line IPEC-J2 was also maintained in our laboratory. A total of 48 six-week-old male BALB/c mice with an initial body weight of 16 ± 2 g were purchased from the Animal Center of Zhengzhou University for use in the experiments.

Lactobacillus rhamnosus MY-1 stored at −80 °C was inoculated into MRS liquid medium for activation. A 1% (v/v) inoculum was then transferred into fresh MRS medium and incubated anaerobically at 37 °C for 24 h. The bacterial culture was collected and centrifuged at 12,000 r/min for 5 min, after which the supernatant was filtered through a 0.22 μm membrane to obtain the cell-free supernatant (CFS). The CFS was aliquoted and stored at −20 °C until use. For preparation of the degradation supernatant, MY-1 culture grown for 24 h was co-incubated with DON for 48 h. The mixture was then centrifuged and filtered following the same procedure, and the resulting supernatant was collected as the degradation supernatant and stored at −20 °C.

Cells were cultured in DMEM complete medium supplemented with 10% fetal bovine serum and 1% penicillin–streptomycin in a 37 °C incubator with 5% CO₂. When cells reached 70–80% confluence, they were digested and passaged using 0.25% trypsin. The cell suspension was adjusted to a density of 1 × 10⁵ cells/mL and seeded into 96-well plates (100 μL per well) or 6-well plates (1 mL per well). After 4 h of incubation to allow for full adhesion, the cells were used for treatments. The DON challenge concentration (0.25 μg/mL) and the MY-1 CFS intervention dose (5%, v/v) were selected based on preliminary dose–response experiments, which were conducted to optimize experimental conditions for subsequent mechanistic evaluation. The cells were then divided into four treatment groups: the control group received only complete culture medium; the MY-1 group was treated with 5% (v/v) of L. rhamnosus MY-1 CFS; the DON group was exposed to 0.25 μg/mL of DON toxin; and the MY-1 + DON group was treated with 5% (v/v) of the degradation supernatant obtained by co-culturing MY-1 with DON. All treatments were applied continuously for 36 h, after which cells were harvested for subsequent analysis.

Cell viability was assessed using the CCK-8 assay. After treatment, 10 μL of CCK-8 solution was added to each well, and the plates were further incubated for 1 h in the incubator. The absorbance of each well was then measured at a wavelength of 450 nm using a microplate reader. Cell viability was expressed as a percentage relative to the control group.

For observation of cellular morphology, samples were sequentially fixed with 2.5% glutaraldehyde (in 0.1 M PBS buffer) and 1% osmium tetroxide, thoroughly rinsed with PBS, and dehydrated using a graded acetone series (30 to 100%). Following dehydration, the samples were infiltrated stepwise with a 1:1 mixture of acetone and Epon-812 embedding resin, followed by pure resin, and then embedded and polymerized at 60 °C. The polymerized blocks were sectioned into 60–80 nm ultrathin slices using an ultramicrotome. After double-staining with uranyl acetate and lead citrate, the sections were observed and imaged under a transmission electron microscope at 20,000 × magnification. All cell experiments were performed in three independent replicates (n = 3).

Levels of MDA, SOD activity, and T-AOC were detected using commercially available assay kits (Jiancheng Bioengineering Institute, China), in accordance with the procedures described in a previous study (Tian et al., 2021). Each assay was conducted with three independent biological replicates (n = 3).

Cell apoptosis was detected using an Annexin V-PE apoptosis detection kit with in situ fluorescence microscopy. The specific procedure was as follows: after treatment under respective conditions, the culture medium from IPEC-J2 cells was removed, and 0.1 mL of fixative solution was added followed by incubation at room temperature for 10 min. The fixative was then discarded, and cells were washed twice with PBS (3 min per wash), with residual liquid completely aspirated. Next, 0.5 mL of Hoechst 33258 staining solution was added and allowed to stain for 5 min. After removal of the staining solution, cells were rinsed 2–3 times with PBS. Subsequently, 195 μL of Annexin V-PE binding buffer was added to resuspend the cells, followed by addition of 5 μL of Annexin V-PE and gentle mixing. The samples were incubated at room temperature (20–25 °C) in the dark for 10–20 min. Apoptosis was assessed by observing red fluorescence under a fluorescence microscope. Apoptosis experiments were performed in three independent replicates (n = 3).

All animal experiments were approved by the Institutional Animal Ethics and Welfare Committee. Prior to the experiment, the animals were acclimatized for one week under standard conditions (12-h light/dark cycle, with free access to food and water). To minimize bias, a completely randomized design was employed for group assignment. Specifically, each mouse was assigned a unique identification number, and group allocation (Control, MY-1, DON, MY-1 + DON) was determined using a computer-generated random number sequence. The mice were randomly assigned to four groups (n = 12) and treated as follows: the control group received 0.2 mL of sterile PBS daily by oral gavage; the MY-1 group was administered 0.2 mL of L. rhamnosus MY-1 bacterial suspension (1 × 10⁹ CFU/mL) daily; the DON group was fed a diet containing 3 mg/kg DON; and the MY-1 + DON group received both the 3 mg/kg DON-containing diet and daily gavage of an equivalent dose of MY-1 bacterial suspension. In this study, a DON dose of 3 mg/kg diet was selected, which corresponds to an estimated daily intake of approximately 0.45 mg DON/kg body weight (assuming a daily feed intake of 3 g and an average mouse body weight of 20 g). This dose is relevant to field exposure scenarios where contaminated feed may contain DON levels ranging from 1 to 5 mg/kg, thereby providing a realistic model for evaluating probiotic intervention under practical conditions. Wherever feasible, blinding was implemented during the experiment and outcome assessment. The personnel responsible for daily gavage, body weight/feed intake recording, and tissue collection were unaware of the group allocations (single-blind). Additionally, histopathological evaluation and quantitative PCR data analysis were performed by investigators blinded to the treatment groups. The experiment lasted 42 days. Body weight and feed intake were recorded weekly. At the end of the experiment, blood was collected from the orbital venous plexus, and serum was separated by centrifugation and stored at −20 °C. The mice were then euthanized by cervical dislocation under anesthesia, and segments of the duodenum, jejunum, ileum, and cecum were rapidly dissected. A portion of intestinal tissue was fixed in 4% paraformaldehyde for histological analysis, and the remainder was snap-frozen at −80 °C for subsequent molecular and microbiological assays. For all subsequent analyses, six mice per group (n = 6) were randomly selected.

Fixed mouse organ tissue samples stored at 4 °C were embedded in paraffin, and sections were prepared from all organs. These sections were stained with hematoxylin and eosin (H&E) and examined under a microscope. Similarly, fixed samples from different intestinal segments were embedded, sectioned, and H&E-stained for microscopic observation. Villus height (VH) and crypt depth (CD) were measured using ImageJ software, and the villus-to-crypt ratio (V/C) was calculated.

Frozen mouse organ tissues stored at −80 °C were homogenized, and total RNA was extracted using the AG SteadyPure Universal RNA Extraction Kit. RNA concentration and A260/A280 ratio were measured with a spectrophotometer. The extracted RNA was then reverse-transcribed into cDNA using the AG StarScript II Genomic DNA Removal Reverse Transcription Premix. Relative expression levels of target genes were determined by quantitative real-time PCR (qRT-PCR) using ChamQ Universal SYBR qPCR Master Mix (Novozymes, China) on a Roche LightCycler 480 system. With glyceraldehyde-3-phosphate dehydrogenase (GAPDH) as the housekeeping gene, relative gene expression was calculated using the 2-ΔΔCt method. Primer sequences for both cell and mouse samples are listed in Supplementary Table 1.

Following DNA extraction from cecal contents and quality verification via Nanodrop and agarose gel electrophoresis, the V3–V4 hypervariable regions of the bacterial 16S rRNA gene were amplified using barcoded primers 338F (5′-ACTCCTACGGGAGGCAGCA-3′) and 806R (5′-GGACTACHVGGGTWTCTAAT-3′). The resulting amplicons were purified, quantified, pooled in equimolar ratios, and used to construct sequencing libraries with the Illumina TruSeq Nano DNA LT Library Prep Kit. High-throughput paired-end sequencing was performed on the Illumina NovaSeq 6,000 platform. After demultiplexing based on barcode sequences, raw reads were processed using the QIIME2 pipeline, where denoising and amplicon sequence variant (ASV) clustering were conducted with DADA2. Taxonomic assignment was performed against the SILVA 138 database. All samples were rarefied to an even sequencing depth prior to downstream analysis. Alpha diversity was assessed using the Chao1, Shannon, and Simpson indices, and beta diversity was evaluated by Bray–Curtis distance and visualized via principal coordinates analysis (PCoA). Microbial composition was summarized at the phylum and genus levels. Differentially abundant taxa between groups were identified using linear discriminant analysis effect size (LEfSe) with a Kruskal–Wallis test (p < 0.05) for significance and a linear discriminant analysis (LDA) score threshold > 4.0; results were presented as LDA score histograms and cladograms to illustrate phylogenetic discriminative features.

All experimental data were analyzed using IBM SPSS Statistics 25 software. Normality was assessed by the Shapiro–Wilk test and homogeneity of variances by Levene’s test. Data meeting both assumptions were analyzed by one-way ANOVA followed by Tukey’s HSD test. For data violating the normality assumption (e.g., certain microbiota and gene expression data), appropriate transformations (log10, square-root, or arcsine-square-root) were applied. If transformed data still deviated from normality, non-parametric tests were used: the Kruskal–Wallis test followed by the Mann–Whitney U test with Bonferroni correction for pairwise comparisons. Microbiome compositional data were analyzed within QIIME2 using appropriate transformations (e.g., centered log-ratio). Differential abundance was assessed by LEfSe with the non-parametric Kruskal–Wallis test (p < 0.05) and LDA score > 4.0. Bar graphs were generated using GraphPad Prism 8. Data are presented as mean ± standard deviation (SD). Differences were considered statistically significant at p < 0.05 and highly significant at p < 0.01. The ‘n’ values represent independent biological replicates (animals or independent experiments).

To evaluate the protective effect of Lactobacillus rhamnosus MY-1 against DON-induced injury, we first assessed the viability and morphology of IPEC-J2 cells. DON exposure significantly reduced cell viability in a time- and concentration-dependent manner. Treatment with 0.25 μg/mL DON for 36 h decreased cell viability to approximately 50% (Figure 1A). A 5% concentration of L. rhamnosus MY-1 CFS maintained 100% cell viability over 48 h, whereas higher concentrations led to reduced viability (Figure 1B). Based on these pilot experiments, 0.25 μg/mL DON and 5% CFS were chosen as the optimal challenge and intervention doses, respectively, for all subsequent mechanistic investigations. Morphological analysis by transmission electron microscopy revealed that DON induced severe cellular damage, including nuclear condensation, mitochondrial swelling, and endoplasmic reticulum dilation. In contrast, the degradation supernatant group exhibited only mild abnormalities, similar to the control and MY-1 supernatant groups (Figure 1C). Moreover, cells treated with the degradation supernatant showed significantly restored viability compared to those treated with DON alone (Figure 1D). These results demonstrate that L. rhamnosus MY-1 effectively alleviates DON-induced cytotoxicity.

In IPEC-J2 cells, exposure to deoxynivalenol significantly induced oxidative stress, as evidenced by elevated MDA levels, whereas treatment with the CFS of Lactobacillus rhamnosus MY-1 alone enhanced antioxidant enzyme activity. Compared with the control group, the DON-treated group showed no significant changes in T-AOC or SOD activity, but exhibited a marked increase in MDA. In contrast, treatment with the MY-1 degradation supernatant significantly reduced MDA content and improved overall antioxidant capacity (Figure 2A). At the mRNA level, DON strongly induced an inflammatory response, significantly upregulating TNF-α and IL-1α while downregulating IL-4. The degradation supernatant notably reversed these changes, reducing pro-inflammatory cytokines and restoring expression of the anti-inflammatory cytokine IL-4 (Figure 2B). Furthermore, DON significantly downregulated the expression of the tight junction proteins ZO-1, Occludin, and Claudin-1, thereby compromising intestinal barrier integrity. Treatment with the CFS of L. rhamnosus MY-1 alone upregulated ZO-1 and Occludin expression, while the degradation supernatant significantly restored the expression levels of both proteins (Figure 2C). Collectively, these results demonstrate that L. rhamnosus MY-1 alleviates DON-induced oxidative stress, inflammatory response, and impairment of barrier function in IPEC-J2 cells.

Fluorescence microscopy analysis (Figure 3A) revealed that IPEC-J2 cells in the control group exhibited blue nuclei with no apparent signs of apoptosis. The supernatant group showed only minimal apoptotic red fluorescence. In contrast, the DON group displayed extensive necrotic and apoptotic features, whereas the degradation supernatant group exhibited markedly reduced apoptosis and improved cellular morphology compared to the DON group. Further analysis of apoptosis-related gene expression (Figure 3B) demonstrated that the mRNA levels of the pro-apoptotic genes BAX and Caspase-3 were upregulated in the DON group. However, the expression of both genes in the degradation supernatant group was lower than that in the DON group, with Caspase-3 expression even lower than that in the control group. These results collectively indicate that DON treatment was associated with upregulation of apoptosis-related genes and corresponding morphological changes, while the degradation supernatant was linked to an attenuation of these effects, suggesting its potential protective role in IPEC-J2 cells.

In vivo experiments showed that mice in the control and L. rhamnosus MY-1 groups remained healthy throughout the trial with no mortality. In contrast, one mouse in the DON group died on day 28 after exhibiting severe lethargy and anorexia; this animal was excluded from all subsequent analyses. The mortality in the DON group, although of low incidence, is consistent with the systemic toxic effects of DON exposure, likely resulting from severe intestinal barrier disruption, systemic inflammation, or metabolic dysregulation. The remaining mice in the DON group displayed persistent signs of toxicity, including mental exhaustion, reduced activity, and ruffled fur, whereas animals in the control, MY-1, and MY-1 + DON groups remained active and exhibited normal grooming behavior. Although no significant differences in initial body weight or feed intake were observed among the groups, the weight gain rate in the DON group during the 14–42 day challenge period was significantly lower than that of the control group. In contrast, the MY-1 + DON group showed a higher weight gain rate compared to the DON group, suggesting that DON inhibits growth and that MY-1 alleviates this effect (Supplementary Table 2; Supplementary Figure 1). To further investigate the impact of MY-1 on DON-induced intestinal injury, H&E staining was performed on the duodenum, jejunum, and ileum (Figure 4), and villus height, crypt depth, and the villus height-to-crypt depth ratio were measured. Histological analysis revealed intact and well-organized intestinal structures in the control, MY-1, and MY-1 + DON groups, whereas the DON group exhibited significant damage in the jejunal villi. Statistical analysis (Supplementary Table 3) indicated that ileal VH in the MY-1 group was significantly higher than in the control group. In the DON group, VH was significantly reduced in the duodenum and jejunum, CD was significantly increased in the jejunum and ileum, and V/C was significantly elevated in the duodenum and jejunum. By comparison, the MY-1 + DON group showed significantly higher VH and V/C values across all intestinal segments relative to the DON group, with ileal VH even exceeding that of the control group. These findings demonstrate that DON disrupts intestinal barrier integrity, while L. rhamnosus MY-1 effectively mitigates this damage and exerts a protective effect on the intestine.

The mRNA expression levels of tight junction proteins and apoptosis-related genes in the duodenum, jejunum, and ileum of mice were analyzed using quantitative real-time PCR. In the duodenum (Figure 5A), DON exposure significantly downregulated Claudin-1 and moderately reduced ZO-1 and Occludin expression. Administration of Lactobacillus rhamnosus MY-1 significantly reversed these changes, increasing the expression of all three tight junction proteins. In the jejunum (Figure 5B), DON markedly suppressed Occludin and reduced Claudin-1 expression, while MY-1 intervention significantly elevated Claudin-1 levels. In ileal tissue (Figure 5C), no significant differences in ZO-1 and Claudin-1 were observed among groups; however, DON significantly decreased Occludin expression, which was restored by MY-1 co-treatment. Regarding apoptosis-related genes, DON significantly upregulated BAX and Caspase-3 expression in the duodenum (Figure 5D) and ileum (Figure 5E), and increased Caspase-3 levels in the jejunum (Figure 5F). MY-1 intervention effectively counteracted these effects, significantly reducing BAX and Caspase-3 expression in the duodenum and ileum, and lowering Caspase-3 in the jejunum. Additionally, MY-1 significantly enhanced BCL-2 expression in the ileum compared to the DON group. These results demonstrate that Lactobacillus rhamnosus MY-1 effectively alleviates DON-induced intestinal barrier damage by modulating the expression of tight junction proteins and apoptosis-related genes.

Analysis of cecal microbiota alpha diversity revealed that, compared to the control group, the MY-1 group exhibited significant increases in the Chao1, Simpson, and Shannon indices, while the DON group showed significant decreases. All three indices were significantly higher in the MY-1 + DON group than in the DON group (Figure 6A). Principal coordinates analysis (PCoA) of beta diversity indicated good intra-group sample reproducibility and clear separation in microbial composition among the four groups (Figure 6B). At the phylum level, Bacteroidota, Firmicutes, Desulfobacterota, and Proteobacteria were the dominant phyla, with no significant differences observed among the four groups (Figure 6C). At the genus level, Bacteroides and Dubosiella were the predominant genera. The relative abundance of Bacteroides and Dubosiella was higher in the DON group than in the control group (Figure 6D). These results suggest that Lactobacillus rhamnosus MY-1 alleviates DON-induced gut microbiota dysbiosis by modulating its structural composition and diversity.

LEfSe analysis revealed that, compared with the control group, DON exposure significantly enriched several bacterial taxa in the mouse gut, including Erysipelotrichales, Erysipelotrichaceae, Dubosiella, Paramuribaculum, Firmicutes_D, Bacilli, and Desulfovibrio_R446353 (Figures 7A,B). Furthermore, supplementation with Lactobacillus rhamnosus MY-1 effectively reversed these DON-induced changes and significantly increased the relative abundance of beneficial taxa such as Clostridia_258483, Firmicutes_A, Lachnospiraceae, Lactobacillales, and Aerococcus (Figures 7C,D). These findings indicate that DON exposure induces gut microbiota dysbiosis, marked by the proliferation of pro-inflammatory and opportunistic pathogenic bacteria. In contrast, L. rhamnosus MY-1 counteracts DON-induced dysbiosis by promoting the colonization and growth of beneficial bacteria, thereby restoring a more balanced microbial community.