Male Wistar rats (8–10 weeks old; 220–250 g) were obtained from the Central Animal Facility. Animals were housed in polypropylene cages (three rats/cage) with autoclaved rice-husk bedding. Natural conditions were maintained at 22°C ± 2°C, 55%–60% relative humidity, and a 12-h light/dark cycle. Rats were acclimated for 7 days prior to experimentation. Standard laboratory chow (protein: 22%, fat: 5%; Provimi, India) and UV-sterilized drinking water were provided ad libitum.

All experimental procedures were approved by the Institutional Animal Ethics Committee (IAEC Approval No.: LL-2021056-K) and conducted in accordance with the Committee for the Purpose of Control and Supervision of Experiments on Animals (CPCSEA), Government of India, guidelines.

A total of 24 rats were randomly allocated using a computer-generated randomization list into three experimental groups (n = 8/group): control group, which received intraperitoneal saline and oral sterile water (vehicle); FLF group, which received GalN + LPS; and FLF + probiotic group, which received continuous probiotic supplementation for 14 days prior to FLF induction and continued until they were killed. Outcome assessments were performed by investigators blinded to group allocation.

A defined multistrain probiotic formulation comprising Lactobacillus rhamnosus GG (ATCC 53103), Lactobacillus plantarum (ATCC 14917), and Bifidobacterium longum (ATCC 15707) was used.

Each strain was cultured individually in de Man–Rogosa–Sharpe (MRS) broth (HiMedia, Mumbai, India) under anaerobic conditions at 37°C and harvested at the mid-logarithmic growth phase. Cultures were centrifuged, washed twice with sterile phosphate-buffered saline (PBS; pH 7.4), and resuspended in sterile PBS.

The final probiotic dose was 1 × 10⁹ CFU/rat/day (total dose), equally divided among the three strains. Fresh suspensions were prepared daily and administered via oral gavage (1 mL) using flexible feeding catheters. Viability was confirmed daily by serial dilution plating on MRS agar before administration. A complete list of reagents, antibodies, manufacturers, and catalog numbers is provided in Supplementary Table S2.

Fulminant liver failure was induced using a well-established GalN/LPS model. d-Galactosamine hydrochloride (700 mg/kg; Sigma-Aldrich, Burlington, USA) was dissolved in normal saline and administered intraperitoneally (10 mL/kg), immediately followed by lipopolysaccharide (10 µg/kg; Escherichia coli O111:B4; Sigma-Aldrich, i.p.). This dosing regimen was selected based on published benchmarks demonstrating reproducible induction of acute fulminant liver failure within 12–24 h. Animals were monitored continuously for clinical signs (lethargy, piloerection, reduced mobility), and humane endpoint criteria were strictly followed.

Body weight, food intake, rectal temperature, and clinical scores (activity, posture, grooming, respiratory rate) were recorded at 24-h intervals to minimize confounding variables.

At the humane experimental endpoint (12–24 h postinduction), rats were anesthetized with ketamine (75 mg/kg) and xylazine (10 mg/kg, i.p.). Blood was collected via cardiac puncture after a standardized fasting period. Serum was collected in clot-activator tubes, centrifuged at 3,500 rpm for 10 min at 4°C. Plasma was collected in EDTA-coated tubes for cytokine analysis. Liver and ileal tissues were harvested immediately. Samples were either snap-frozen in liquid nitrogen (protein/biochemical assays), fixed in 10% neutral-buffered formalin (histology), or stored in RNAlater, where applicable.

Rats were fasted for 4 h and administered FITC-dextran (4 kDa; 600 mg/kg; Sigma-Aldrich) orally. Serum was collected for 4 h postgavage, and fluorescence was measured using a microplate reader (excitation: 485 nm/emission: 528 nm). Concentrations were calculated using a standard curve (0–5,000 ng/mL). Gut microbiota composition was analyzed using 16S rRNA gene sequencing, with detailed methodology provided in Supplementary Methods S1.

Serum tumor necrosis factor alpha (TNF-α), interleukin (IL)-6, and IL-1β were evaluated using high-sensitivity rodent enzyme-linked immunosorbent assay (ELISA) kits (Elabscience, Wuhan, China). Absorbance was recorded at 450 nm with reference to 570 nm. All assays were performed in duplicates, with intra- and interassay CV < 10%.

Data were displayed as mean ± SD. Analyses were performed using GraphPad Prism 9.0. One-way ANOVA was used for group comparisons, followed by Tukey’s post hoc test for multiple comparisons. A p < 0.05 was considered statistically significant. Effect sizes (η²) and confidence intervals were reported where appropriate. Detailed statistical assumptions, post hoc testing, and effect size calculations are provided in Supplementary Table S1.

All animals in the control and probiotic groups survived throughout the study period. In the FLF group, three of eight rats died within 8–12 h following GalN/LPS administration, whereas no mortality occurred in the probiotic + FLF group. Surviving animals were included in biochemical and histological analyses, while deceased animals were included only in survival analysis. Baseline body weights and physiological parameters were comparable over bunches (p > 0.05), confirming consistency before experimental treatments. Routine health monitoring revealed no signs of distress in the probiotic or control groups during the supplementation period, indicating great tolerability of the probiotic formulation, as shown in Table 1.

Successful FLF induction was confirmed by the rapid onset of lethargy, piloerection, jaundice, and reduced mobility within 6–8 h of GalN/LPS injection. The FLF group showed a significant increase in serum ALT, AST, and bilirubin levels, along with a marked prolongation of Innate immune receptor (INR) (p < 0.001 vs. control), demonstrating severe hepatic damage, as shown in Table 2. Histological examination revealed mixed hepatocellular degeneration, sinusoidal collapse, and pronounced inflammatory infiltration, confirming the presence of serious hepatic injury.

Probiotic supplementation for 14 days was well tolerated, with no adverse effects observed. Rats treated with probiotics showed a modest but significant increase in body-weight gain compared with controls (p < 0.05), as shown in Table 3. Animals receiving probiotics displayed a modest yet noteworthy increase in body-weight gain compared with controls (p < 0.05), and fecal microbial analysis confirmed successful colonization, with elevated Lactobacillus and Bifidobacterium counts relative to the control group (p < 0.01). In the probiotic + FLF group, probiotic supplementation attenuated the clinical severity of FLF, with animals showing reduced clinical symptoms compared with the untreated FLF group.

GalN/LPS administration caused a profound increase in ALT, AST, total bilirubin, alkaline phosphatase (ALP), and INR (p < 0.001 vs. control). Probiotic supplementation significantly reduced ALT, AST, bilirubin, and ALP levels compared with the FLF group (p < 0.01), as shown in Table 4. Probiotics also improved coagulation status, as indicated by a decrease in INR. Exact numeric values (mean ± SD) for all hepatic biomarkers are provided in Table 4. Oxidative stress analysis revealed significantly elevated hepatic malondialdehyde (MDA) levels and depleted glutathione (GSH) and superoxide dismutase (SOD) activity in the FLF group. Probiotic pretreatment significantly reduced lipid peroxidation and restored antioxidant defenses.

FLF induction caused a pronounced disruption of intestinal barrier integrity, resulting in an approximately fourfold increase in plasma FITC-dextran levels compared with the control group (p < 0.001). Probiotic supplementation reduced FITC-dextran levels by approximately 40%–50% relative to the FLF group (p < 0.01), indicating partial restoration of epithelial barrier function (Figure 1).

Liver sections from FLF animals demonstrated severe necroinflammatory activity, characterized by widespread hepatocellular necrosis, inflammatory infiltration, sinusoidal congestion, and hemorrhage. In contrast, probiotic + FLF animals exhibited significantly preserved hepatic architecture, with fewer necrotic foci and reduced inflammatory cell infiltration (Figure 2). Intestinal histology revealed marked villus shortening, epithelial disorganization, and goblet-cell depletion in FLF rats. Probiotic pretreatment restored villus height, epithelial continuity, and goblet-cell density, consistent with improved mucosal barrier integrity. Hepatic MDA levels were significantly elevated in FLF rats, whereas GSH content and SOD activity were markedly reduced (p < 0.001). Probiotic supplementation significantly reduced oxidative damage and partially restored antioxidant enzyme activity. All oxidative stress parameters were normalized to total protein content.

Western blot densitometric analysis normalized to housekeeping proteins showed significant downregulation of ZO-1, occludin, and claudin-1 in the FLF group (p < 0.01; Figure 3). Probiotic supplementation restored tight junction protein expression by approximately 60%–80% relative to FLF animals (p < 0.01). Immunofluorescence analysis confirmed improved localization of tight junction proteins at the apical epithelial border in probiotic-treated animals. Z-stack imaging was not performed and is acknowledged as a limitation.

Microbiota profiling revealed significant dysbiosis in FLF rats, characterized by the depletion of beneficial Lactobacillus and Bifidobacterium genera and the overgrowth of pathogenic taxa, including Enterobacteriaceae and Clostridium spp. Probiotic supplementation effectively reversed these alterations, restoring beneficial microbial populations, suppressing pathogenic bacteria, and significantly improving alpha-diversity indices (p < 0.01; Figure 4).

Systemic inflammation was markedly elevated in FLF rats, as evidenced by significantly increased serum TNF-α, IL-6, and IL-1β levels (p < 0.001). Probiotic supplementation reduced circulating proinflammatory cytokines by approximately 40%–50% compared with FLF animals (p < 0.01) and significantly increased IL-10 levels (Figure 5). Only serum cytokines were measured; tissue cytokine levels were not assessed, which is acknowledged as a limitation.

Probiotics particularly diminished FITC-dextran porousness and reestablished the localization of ZO-1, occludin, and claudin-1, consistent with earlier work illustrating probiotic-induced stabilization of tight junction complexes (Babu and Mohanty, 2025; Plaza-Díaz et al., 2020). Reclamation of the epithelial barrier likely constrained the translocation of MAMPs, such as LPS, into the portal circulation, thereby decreasing downstream hepatic inflammation.

The critical diminishments in TNF-α, IL-1β, and IL-6 observed in the probiotic group reflect previous evidence that microbiota rectification constraints NF-κB-mediated inflammatory signaling and promotes safe homeostasis in models of intense liver damage (Babu and Mohanty, 2025; Beyaz Coşkun and Sağdiçoğlu Celep, 2022). These findings align with discoveries that probiotics and microbial metabolites can rebalance mucosal immunity and mitigate inflammatory intensification during hepatic stress (Chen et al., 2021; Liu et al., 2024).

Probiotic-treated rats displayed decreased lipid peroxidation and improved antioxidant status, findings congruent with reports that several Lactobacillus species activate Nrf2/HO-1 signaling to suppress oxidative damage (Zhou et al., 2022; Zhang J. et al., 2024). This antioxidant effect may synergize with the reduced cytokine burden to protect hepatocytes from apoptotic and necrotic damage.

Lower necroinflammatory changes, fewer TUNEL-positive cells, and improved hepatic architecture in treated animals emphasize the systemic effect of intestinal obstruction restoration on liver pathology. These enhancements parallel the defensive responses observed in FMT and probiotic interventions across various models of severe liver injury (Lamas-Paz et al., 2024; Yang et al., 2023).

A major quality of the study is the comprehensive assessment of epithelial integrity, inflammatory mediators, oxidative markers, and histological endpoints. However, this work does not fully characterize microbial composition or metabolite dynamics. Direct evaluation of NF-κB or Nrf2/HO-1 pathways was not performed. The study also reflects a pretreatment rather than a therapeutic administration approach. Future research should incorporate metagenomic and metabolomic profiling and assess safety and efficacy in large-animal or early clinical models.

Several limitations of the present study should be acknowledged. First, a single probiotic dose and fixed supplementation duration were used; therefore, dose–response relationships and optimal treatment windows could not be established. Second, although probiotic supplementation significantly improved survival, hepatic injury, intestinal permeability, and inflammatory markers, direct mechanistic pathways—including hepatic NF-κB activation, Nrf2/HO-1 signaling, and intestinal TLR pathways—were not directly assessed and are inferred from biochemical and cytokine profiles. Third, intestinal barrier integrity was evaluated functionally and structurally, but three-dimensional Z-stack imaging and electron microscopy were not performed, limiting ultrastructural resolution. Fourth, although serum cytokines and oxidative stress markers were measured, tissue cytokine levels, portal endotoxin concentrations, and direct bacterial translocation assays (e.g., mesenteric lymph node cultures) were not conducted. Fifth, although gut microbiota composition was analyzed, metagenomic or metabolomic profiling was not included, restricting functional interpretation of microbial changes. Finally, the sample size, while consistent with established FLF models, was not determined by an a priori power calculation, which may limit the detection of smaller effect sizes. Future studies incorporating dose–response analyses, mechanistic pathway interrogation, advanced imaging, and multiomics approaches will be essential to further define the probiotic-mediated gut–liver protective axis in fulminant liver failure.