Ligilactobacillus salivarius 8-2 attenuates intestinal mucosal injury in chicks via the mitochondrial fision-glycometabolism reprogramming axis

Abstract

The intestinal mucosa is crucial for nutrient absorption and barrier function. In this study, a newly identified strain of Ligilactobacillus salivarius (L. sa. 8-2) exhibited a mucosal protective effect in chick intestinal injury models. However, the underlying mechanisms need to be clarified. In vivo, chicks were pre-fed with L. sa. 8-2 supernatant or its precipitate and then challenged with lipopolysaccharide (LPS) or Salmonella Typhimurium (STm). The results showed that L. sa. 8-2 and its supernatant significantly reduced intestinal epithelial apoptosis, increased goblet cell density and Muc2 mRNA abundance, and upregulated Claudin1 protein levels. Specifically, compared with the LPS-challenged group, L. sa. 8-2 supernatant accelerated epithelial renewal, as shown by elevated protein levels of proliferating cell nuclear antigen (PCNA) and increased mRNA abundance of cell cycle regulators Ccnd1 and Cdk2. Moreover, the L. sa. 8-2 supernatant markedly increased protein levels of both the active intestinal stem cell (aISC) marker LGR5 and the reserve ISC (rISC) marker HOPX in the intestinal crypt. In vitro, the enteroids were pre-treated with L. sa. 8-2 supernatant and then challenged with LPS. It was shown that the L. sa. 8-2 supernatant elevated the protein levels of LGR5 and HOPX compared with the injured group. Moreover, co-staining of LGR5 and HOPX revealed that more HOPX⁺ cells were co-localized with LGR5⁺ cells in the L. sa. 8-2 supernatant pretreatment group, suggesting the activation of rISCs. Additionally, compared with the injured groups, L. sa. 8-2 supernatant preserved mitochondrial morphology and upregulated the fission-related protein FIS1. At both the mRNA and protein levels, L. sa. 8-2 supernatant upregulated the pyruvate metabolism-related enzyme LDHA and LDHB, while decreasing mRNA levels of Sdha (an oxidative phosphorylation-related enzyme), suggesting a metabolic shift from oxidative phosphorylation toward glycolysis and an enhancement of pyruvate utilization. Taken together, L. sa. 8-2 alleviates intestinal mucosal injury and promotes epithelial renewal by activating ISCs, which is mediated by promoting mitochondrial fission and subsequent enhancement of pyruvate utilization in the intestinal crypt.

Introduction

The intestinal mucosa functions as a selectively permeable barrier for nutrient absorption and also serves as a physical and immunological defense, relying on the structural integrity of the epithelial lining and intercellular junctional complexes such as tight junctions (Gehart and Clevers, 2019; Di Vincenzo et al., 2024). However, this barrier is highly vulnerable to inflammatory or infectious insults, which can compromise its integrity and physiological function. In the lipopolysaccharide (LPS)-induced septic mouse model, intestinal mucosal injury is characterized by elevated epithelial apoptosis and disruption of tight-junction proteins, including Occludin and Claudin-1 (Chen et al., 2023a). Similarly, in the Aflatoxin B1 (AFB1)-induced intestinal injury mouse model, mucosal damage manifests as significant downregulation of tight-junction proteins (zonula occludens-1 (ZO-1), Occludin, and Claudin-1) and mucin-2 (MUC2), accompanied by increased intestinal permeability and elevated pro-inflammatory cytokine levels (Li et al., 2024a). Trichinella spiralis infection increases the number and size of goblet cells in the small intestine (Luo et al., 2019). Accumulating evidence indicates that probiotics can protect the intestinal mucosal barrier against various insults. For instance, in DSS-induced colitis mouse models, Akkermansia muciniphila has been shown to suppress intestinal epithelial cell apoptosis, promote goblet cell function and MUC2 production, and enhance tight-junction integrity, collectively contributing to the restoration of the intestinal mucosal barrier (Martin-Gallausiaux et al., 2022). A multi-strain probiotic formulation containing L. rhamnosus LR 32, B. lactis BL 04, and B. longum BB 536 upregulates tight-junction protein ZO-1, Claudin-1, and Occludin while downregulating Claudin-2, thereby enhancing barrier integrity and protecting against inflammatory damage (di Vito et al., 2022). Using HRT18 monolayers and Caco-2 ALI/VIP mucus layer models, Bifidobacterium bifidum strains W23 and W28 were shown to enhance intestinal epithelial barrier integrity by desialylating the transmembrane mucin MUC13, thereby increasing transepithelial electrical resistance and promoting mucus layer colonization (Segui-Perez et al., 2025). Similarly, Lactobacillus plantarum T10 improves intestinal barrier integrity in DSS-induced colitis mice by producing exopolysaccharides that enhance tight-junction proteins and reduce inflammation (Tao et al., 2024). Lactobacillus plantarum BMCM12 secretes extracellular proteins that reduce the adhesion of Escherichia coli and Salmonella enterica, thereby reinforcing the intestinal barrier (Liu et al., 2020). Additionally, a mixture containing Akkermansia muciniphila and Clostridioides difficile can occupy specific ecological niches and function as microbial competitors, thereby crowding out pathogenic bacteria (Pereira et al., 2020). In this study, we aimed to examine the protective effects of a selected strain of Ligilactobacillus salivarius, isolated from Chinese native hens, on intestinal barrier function.

The intestinal mucosal homeostasis and repair depend on two distinct populations of intestinal stem cells (ISC), active Lgr5⁺ ISCs at the crypt base, and quiescent Hopx⁺/Bmi1⁺ ISCs located at the "+4" site of the crypt (Sangiorgi and Capecchi, 2008; Montgomery et al., 2011; Powell et al., 2012; Kim et al., 2024). Mitochondria in murine Lgr5⁺ ISCs display both high abundance and elevated activity (Rodríguez-Colman et al., 2017). Although quiescent under physiological conditions, +4 ISCs can rapidly proliferate and regenerate crypt-base columnar cells (CBC) following injury, making them reserve ISCs (rISC) (Smith et al., 2025). During the early period of intestinal inflammation in chickens, Krüppel-like factor 5 (KLF5) activates the rISCs by binding to the Hopx promoter directly, thus initiating mucosal repair after injury (Yu et al., 2023). Growing evidence suggests that probiotics exert regulatory effects on ISCs. For example, Lactobacillus salivarius promotes ISC activity by facilitating mitochondrial ATP synthesis (Luo et al., 2025). Lactobacillus rhamnosus GG promotes the repair process following intestinal mucosal injury by enhancing the regenerative capacity of ISCs in mice (Chen et al., 2023b). Limosilactobacillus reuteri D8 strain stimulates the regeneration of ISCs by increasing the number of Paneth cells in the crypts, promoting the repair of intestinal mucosa in DSS-induced colitis mice (Hou et al., 2018). Similarly, Akkermansia muciniphila accelerated the proliferation of Lgr5⁺ ISCs and promoted the differentiation of Paneth cells and goblet cells in the small intestine, as demonstrated in vivo in C57BL/6 mice and Lgr5-EGFP reporter mice (Kim et al., 2021). Additionally, the intestinal microbiota can regulate ISCs through pattern-recognition receptors (PRR) or by modulating the redox state and oxygen concentration in the intestine (Pral et al., 2021; Ye and Rawls, 2021). However, whether the newly isolated Ligilactobacillus salivarius in this study exerts regulatory effects on ISCs remains to be elucidated.

Mitochondria are highly dynamic organelles. Their fission generates smaller mitochondria, which facilitates efficient intracellular movement and organization; this process also ensures the inheritance of mitochondria during cell division (Yapa et al., 2021). Mitochondrial fusion maintains respiratory function and cellular homeostasis by facilitating the exchange of gene products and metabolites between mitochondria, particularly under metabolic and environmental stress (Adebayo et al., 2021). As a process sensitive to the intestinal environment, the dynamic balance between mitochondrial fission and fusion is crucial for energy generation, regulating mitochondrial function and ultimately affecting cellular fate. Specifically, oxidative stress in porcine enterocytes and experimental colitis in mice consistently trigger mitochondrial damage, characterized by structural deformation (e.g., cristae disruption) and bioenergetic failure, including impaired oxidative phosphorylation (OXPHOS) and ATP depletion (Li et al., 2022; Uddin et al., 2024). In addition, the mycotoxin deoxynivalenol promotes the expression of mitochondrial fission proteins (DRP1, FIS1, MIEF1, etc.) in porcine intestinal epithelial cells, leading to excessive mitochondrial fission and ROS accumulation, thus impairing the intestinal mucosal barrier function (Dai et al., 2020; Zhang et al., 2022). Escherichia coli can activate fission protein DRP1 in human colonic epithelial cells, inducing excessive mitochondrial fission that triggers cytochrome C release and disrupts intestinal epithelial barrier function (Mancini et al., 2021). Furthermore, mitochondrial fission promotes glycolysis and enhances differentiation activity. In a murine model of ulcerative colitis, excessive fission disrupts butyrate metabolism, thereby inhibiting stem cell proliferation and mucosal repair (Fu et al., 2024). In colorectal cancer cells, abnormally expressed genes induce phosphorylation of the mitochondrial DRP1 at Ser616 via ERK, leading to excessive mitochondrial fission, reduced OXPHOS levels, and increased glycolysis (Wu et al., 2024). FoxO and Notch signaling recruit DRP1 and induce mitochondrial fission, thereby activating glycolysis and promoting differentiation of mouse ISCs into secretory cells (Ludikhuize et al., 2020). However, it remains unclear whether the newly isolated Ligilactobacillus salivarius strain exerts its effects by influencing the mitochondrial fission-fusion balance and further modulating ISC activity.

Compared with OXPHOS, glycolysis has a limited capacity for ATP generation, which is partly compensated by its higher rate (Kern Coquillat et al., 2025). Cytoplasmic glycolysis rapidly supplies energy to maintain stem cell self-renewal capacity. Besides, glycolysis is the predominant bioenergetic pathway for endothelial cells (EC) (Fitzgerald et al., 2018). Hexokinase 2 (HK2), a key glycolytic enzyme, sustains self-renewal in mouse ISCs by regulating lactate levels (Li et al., 2023). Pyruvate dehydrogenase kinase 1/4 (PDK1/4) inhibits pyruvate entry into the mitochondrial TCA cycle, forcing mitochondrial OXPHOS to switch to cytoplasmic glycolysis. This enhances the function of senescent cells by producing lactate (Dou et al., 2023). mTORC2-AKT signaling can target activation of 6-phosphofructokinase 2 (PFKFB2), promoting glycolysis in intestinal epithelial cells and enhancing their "stemness" (Li et al., 2024b). In contrast, mitochondrial OXPHOS can alter stem cell differentiation. Dichloroacetic acid (DCA) inhibits cytoplasmic glycolysis, thereby enhancing the mitochondrial OXPHOS pathway and promoting the differentiation of mouse ISCs into secretory cells (Khaloian et al., 2020). Similarly, inosine activates mitochondrial function and the OXPHOS pathway in chronic CAR-T cells, thereby inducing their stem cell characteristics (Klysz et al., 2024). In dendritic cells (DC), metabolic reprogramming to a high-glycolytic phenotype is necessary for DC activation and regulates DC cytokine production and antigen-presenting capability (Kierans and Taylor, 2021). As one of the essential metabolic substances in the glycolytic pathway, pyruvate replenishes the cofactor NAD⁺ to maintain glycolysis through the action of lactate dehydrogenase (LDH), playing a central role in cellular energy production and biosynthesis (Rao et al., 2021). However, whether the newly isolated Ligilactobacillus salivarius regulates ISC activity through metabolic reprogramming remains to be explored.

In this study, a strain of Ligilactobacillus salivarius was isolated and screened from a Chinese native chicken breed. Based on its observed mucosal protective effect, this study will further elucidate its regulation of mucosal renewal and ISC activity, focusing on mitochondrial dynamics and subsequent glycometabolism reprogramming to uncover the mechanisms by which Ligilactobacillus salivarius regulating ISC activity.

Materials and methods

Isolation and purification of Ligilactobacillus

A 0.1 g fresh fecal sample was collected from free-range Zhenning (ZN) hens and homogenized in 1 mL of sterile normal saline. The mixture was fully blended and subjected to 10-fold serial dilution. Then, 100 μL of the 10⁻⁴ dilution was pipetted and evenly spread onto an MRS (de Man, Rogosa and Sharpe Medium) agar plate. After incubation at 37°C for 48 h, a well-isolated single colony was picked and cultured in MRS broth for 24 h. The resulting culture was streaked onto fresh MRS agar to obtain single colonies. This purification procedure was repeated three times to obtain a pure strain. A single purified colony was inoculated into MRS broth for large-scale cultivation. The bacterial suspension was mixed with 40% sterile glycerol to achieve a final glycerol concentration of 20%, and stored at −80°C for long-term preservation.

Gram staining

A drop of ultrapure water was placed at the center of a clean glass slide with a sterile inoculating loop. A single colony was picked and evenly smeared into the water drop. The smear was air-dried at room temperature before staining. The smear was stained with ammonium oxalate crystal violet for 1 min and gently rinsed with distilled water. Iodine solution was then applied for 1 min, followed by rinsing. Decolorization was performed using 95% ethanol for 45 s with gentle shaking, and the slide was rinsed again. Counterstaining with safranin was carried out for 1 min, followed by a final rinse with distilled water. After staining, bacterial morphology was observed under an optical microscope.

Antibacterial activity assay

Sterile LB (Luria-Bertani) agar (20 mL) was poured into each Petri dish. After the medium solidified, pre-placed sterile Oxford cups were kept in position until sample loading. Four indicator strains (Escherichia coli, Staphylococcus aureus, Salmonella Typhimurium, and Salmonella Enteritidis) were activated by subculturing 2 to 3 times on LB agar. A single colony of each indicator strain was inoculated into LB broth and incubated at 37°C for 10 h. Bacterial cells were harvested by centrifugation and resuspended in sterile phosphate-buffered saline (PBS) to adjust the bacterial concentration to 10⁸ CFU/mL. Subsequently, 200 μL of each bacterial suspension was evenly spread onto LB agar plates using sterile cotton swabs. L. sa. 8-2 was activated and cultured in MRS broth at 37°C for 13 h to achieve a concentration of 10⁸ CFU/mL. The cell-free fermentation supernatant was added to each Oxford cup at 200 μL per cup, with uninoculated sterile MRS broth defined as the blank control. All plates were incubated at 37°C for 24 h, and the diameters of the inhibition zones were measured. All experiments were performed in triplicate. The reference strain Lactobacillus salivarius CGMCC 1.1881 was used as the positive control throughout the assay.

Scanning and transmission electron microscopy

To analyze the morphology of the newly isolated Ligilactobacillus strain, scanning electron microscopy (SEM) was performed. The strain was fixed in 2.5% glutaraldehyde prepared in 0.1 M phosphate buffer (pH 7.4) at 4°C overnight. After fixation, the samples were washed three times with the same buffer and dehydrated using a graded ethanol series (50%, 70%, and 90%), with each step lasting 15 min. Critical-point drying was carried out with liquid CO₂. The dried samples were mounted on aluminum stubs, sputter-coated with a thin gold-palladium layer (∼10 nm in thickness), and observed using a field-emission scanning electron microscope (FEI Nova NanoSEM 450, USA) at an accelerating voltage of 5 kV.

To evaluate ultrastructural changes in intestinal crypts following mucosal injury, duodenal tissue samples were prepared for transmission electron microscopy (TEM). Samples were first fixed with 2.5% glutaraldehyde, post-fixed with 1% osmium tetroxide, and stained with 2% uranyl acetate. Subsequently, the samples were dehydrated in a graded ethanol series (50%, 70%, and 90% for 15 min each, followed by 100% for 20 min), then dehydrated twice with 100% acetone for 20 min each, The samples were infiltrated and embedded in Spurr resin. Ultrathin sections (70 to 90 nm) were prepared using a Leica UC 7 ultramicrotome (Wetzlar, Germany) and counterstained with lead citrate and saturated uranyl acetate. The ultrastructure of the crypt was visualized using a Hitachi H-7650 TEM (Tokyo, Japan).

Animals

A total of 120 Hy-Line White chicks (Gallus), 3 days old, were randomly divided into eight groups with 15 chicks per group (5 chicks per cage × three cages per group). The grouping strategy and treatments are summarized in Table 1. L. sa. 8-2 was prepared at the concentration of 10⁸ CFU/chick/day, and the fermentation supernatant (L. sa. 8-2 s) and bacterial cell precipitate (L. sa. 8-2 p) were obtained by centrifuging at 1250 × g for 5 min. The control group was fed a basal diet. Chicks in the MRS groups, the L. sa. 8-2 s groups and the L. sa. 8-2 p groups were fed a basal diet containing MRS, L. sa. 8-2 s or L. sa. 8-2 p at an equal volume for 14 consecutive days. At the end of the feeding trial, the chicks in the MRS+PBS group were injected intraperitoneally (i.p.) with PBS (pH 7.4). The chicks pre-fed with MRS, L. sa. 8-2 s, and L. sa. 8-2 p were divided into two subsets, one subset was injected i.p. with LPS (10 mg/kg BW, L2880, Sigma, MO, USA), the other subset was subjected to an oral challenge with Salmonella Typhimurium (STm) at 1 × 10⁹ CFU per chick. Based on Yu et al. (2021) and our preliminary experiments, at 4 h post-LPS injection or STm challenge, duodenal tissue and duodenal crypts were sampled from 6 individuals in each group. The present study was carried out following the Guiding Principles for the Care and Use of Laboratory Animals of Zhejiang University. The experimental protocols were approved by the Committee on the Ethics of Animal Experiments of Zhejiang University (No. 14933).

Table 1.

Animal experimental design.

Group	Pre-feeding for 14 days	Challenge/Injection
Control	Basal diet	/
MRS+PBS	Basal diet + MRS broth	PBS i.p.
MRS+LPS	Basal diet + MRS broth	LPS i.p.
MRS+STm	Basal diet + MRS broth	S. Typhimurium oral challenge
L. sa. 8-2 s+LPS	Basal diet + L. sa. 8-2 s	LPS i.p.
L. sa. 8-2 p+LPS	Basal diet + L. sa. 8-2 p	LPS i.p.
L. sa. 8-2 s+STm	Basal diet + L. sa. 8-2 s	S. Typhimurium oral challenge
L. sa. 8-2 p+STm	Basal diet + L. sa. 8-2 p	S. Typhimurium oral challenge

Note: n = 15 per group (5 chicks per cage × three cages per group). i.p., intraperitoneally; LPS, lipopolysaccharide; L. sa. 8-2 s, L. salivarius 8-2 supernatant; L. sa. 8-2 p, L. salivarius 8-2 precipitate; STm, S. Typhimurium.

Duodenal crypts isolation and enteroid culture

To investigate alterations in ISC activity, duodenal crypts were isolated and cultured in vitro following a previously described protocol (Li et al., 2018). Briefly, the duodenum was cut into 2 to 3 cm segments after making a longitudinal incision. The villi were gently scraped off, and the duodenal segments were then cleaned with PBS and gently shaken in 2 mM EDTA solution (pH 7.4). After passing through a 70 μm nylon cell strainer (352360, Corning, NY, USA), the suspensions were centrifuged to obtain crypts. The crypts were further used for mRNA abundance assay, protein level assay, and enteroid culture.

For enteroid culture, the isolated duodenal crypts were mixed with Matrigel (354231, Corning, NY, USA) and dropped into a flat-bottom plate as a hemispherical droplet. After the Matrigel solidified, a complete medium containing R-spondin 1, Noggin, N-acetylcysteine, N-2 supplement, and EGF was added. The enteroids were then cultured at 38.5°C in a 5% CO₂ atmosphere. During the initial stages of cultivation, L. sa. 8-2 supernatant was added, and the enteroids were cultured for 48 h. After 24 h of culture, LPS (2 μg/mL) was added to induce injury. For transcriptomic analysis, mRNA expression analysis, and protein expression analysis, the enteroids were harvested from Matrigel using Cell Recovery Solution (354253, Corning, NY, USA).

RNA sequencing

To screen for differentially expressed genes in the enteroids, organoid specimens were collected from three independent biological replicates per group. Following collection, samples were immediately snap-frozen on dry ice and transported to a commercial company (Novogene Co., Ltd., Beijing, China). Briefly, total RNA was extracted, and transcriptome sequencing was performed using the Illumina sequencing platform. Significantly differentially expressed genes were detected using the EdgeR algorithm implemented in the CLC Genomics Workbench (Qiagen, Hilden, Germany).

To ensure the quality and reliability of data analysis, reads containing adapters, ambiguous nucleotides (N), and low-quality reads were removed. The reference genome index was constructed, and clean paired-end reads were aligned to the chicken reference genome using HISAT2 v2.0.5. Differential expression analysis between groups was conducted using DESeq2 software (version 1.20.0), and the resulting P-values were adjusted using the Benjamini-Hochberg method to control the false discovery rate. Genes with an adjusted P-value < 0.05, as identified by DESeq2, were designated as differentially expressed. GO enrichment analysis and KEGG enrichment analysis of differentially expressed genes were conducted using the clusterProfiler R package.

Terminal deoxynucleotidyl transferase-mediated dUTP nick-end labelling (TUNEL) assay

Apoptosis in intestinal tissue was assessed qualitatively by TUNEL assay. Duodenal samples fixed in 4% paraformaldehyde (PFA) were embedded in paraffin and sectioned into 5 μm cross-sections. Staining was performed using the TUNEL BrightGreen Apoptosis Detection Kit (A112-01, Vazyme, Nanjing, China) according to the manufacturer’s instructions. TUNEL-positive cells exhibiting green fluorescence were visualized and captured using either a Nikon A1R or an Olympus IX81-FV1000 confocal laser scanning microscope (Tokyo, Japan). For each experimental group, a total of 30 well-preserved villi (derived from 15 sections from 5 individual chicks) were examined. The relative abundance and tissue distribution patterns of apoptotic signals were qualitatively assessed and described by comparing fluorescence intensity and the number of positive cells across different groups.

AB-PAS staining

For histological analysis, duodenal samples were fixed in 4% PFA. Following fixation, the tissues were embedded in paraffin and sectioned into 5 μm cross-sections. AB-PAS staining (G1285, Solarbio, Beijing, China) was conducted according to the manufacturer’s instructions to identify goblet cells. The number of goblet cells in each group was counted for a total of 30 well-preserved villi from 15 sections of 5 individual chicks, and the density of goblet cells was calculated as the number of positive cells per 100 μm of villus length.

Quantitative real-time PCR (qPCR)

Total RNA was extracted from intestinal tissue or crypts using FreeZol Reagent (R711-01, Vazyme, Nanjing, China) and reverse transcribed into cDNA using the HiScript III All-in-one RT SuperMix kit (R333-01, Vazyme, Nanjing, China). With cDNA as a template, real-time PCR reactions were performed on a Bio-Rad CFX96 Touch Real-Time PCR system (Hercules, CA, USA). The reaction mixture consisted of 1 μL of cDNA, 400 nM of each primer (Table 2), and 8.5 μL of 2 × SYBR qPCR Master Mix (Q711-02, Vazyme, Nanjing, China). The reaction conditions were as follows: initial denaturation at 95°C for 30 s, followed by 40 cycles of 95°C for 10 s and 60°C for 30 s. GAPDH was used for normalization. Data analysis was performed using the comparative Ct method (2^−△△Ct). All samples were run in triplicate, and all experiments were repeated at least three times.

Table 2.

Primers for qPCR analysis.

Genes	Accession No.	Primer sequences (5′−3′)	Product (bp)
Muc2	NM_001318434.1	F: TACTTCACCTTCAACCATTACAAC R: CATAGTCACCACCATCTTCTTCA	161
Ccnd1	NM_205381.1	F: CCTCAAGAAAAGCCGGTTGC R: CTGCGGTCAGAGGAATCGTT	86
Cdk2	NM_001199857.1	F: TCCGTATCTTCCGCACGTTG R: GCTTGTTGGGATCGTAGTGC	183
Hopx	NM_204556.2	F: AACAAGCATCCTGATCCCA R: TAGTCCCTCACAGACCCAC	149
Lgr5	XM_425441.7	F: CATACATTCCTAAGGGAGCAT R: GTGTCTAAGGGAGACCAAACC	191
Ldha	NM_205284.2	F:GAAGACGCCGGCAGTACA R: ACCAACCACGCTGATCTTGT	101
Ldhb	XM_046906327.1	F:GTGGTTTCCAACCCAGTGGA R: CTCTCAGCCATCAGGTAGCG	128
Sdha	NM_001277398.1	F: ACCATTTACCACCCCAGCAG R: ACCGTAGGCAAAACGGGAAT	112
HK2	NM_001396482.1	F:CCCTTAACCCTGGGAAGCAG R: CCTCGCTTGGTGAAGTCCAT	97
GAPDH	NM_204305.1	F: GATGGGTGTCAACCATGAGAAA R: CAATGCCAAAGTTGTCATGGA	116

Western blot (WB)

Total protein was extracted from intestinal tissue, crypts or cultured enteroids using RIPA buffer (P0013B, Beyotime, Shanghai, China). The protein concentration in the lysates was determined using a BCA protein assay kit (20201ES76, Yeasen, Shanghai, China). The extracted proteins were diluted with Protein Loading Buffer (FD006, Fdbio, Hangzhou, China) and boiled at 95°C for 15 min. Total protein (30 μg) was separated by SDS-PAGE and then transferred to a 0.22 μm PVDF membrane (Millipore, MA, USA). The membrane was blocked with 5% skimmed milk at room temperature for 1.5 h, followed by overnight incubation at 4°C with primary antibodies: anti-PCNA (ab29, Abcam, Cambridge, UK), anti-LGR5 (customized, HuaBio, Hangzhou, China), anti-HOPX (self-prepared), anti-OPA1 (self-prepared), anti-MFN1 (self-prepared), anti-FIS1 (self-prepared), anti-Villin1 (HA500509, HuaBio, Hangzhou, China), anti-Claudin1 (13050-1-AP, Proteintech, Wuhan, China), anti-LDHA (ER00702, HuaBio, Hangzhou, China), anti-LDHB (0807-1, HuaBio, Hangzhou, China) and anti-α-Tubulin (ER130905, HuaBio, Hangzhou, China). After rinsing in PBS, the membrane was incubated at room temperature for 1 h with HRP-conjugated anti-rabbit IgG (HA1001, HuaBio, Hangzhou, China) or anti-mouse IgG (HA1006, HuaBio, Hangzhou, China). The blots were detected using the SuperPico ECL chemiluminescence kit (E422-01, Vazyme, Nanjing, China) and visualized with a Bio-Rad ChemiDoc Touch Imaging System (Hercules, CA, USA). The densitometry of the blots was analyzed using ImageJ and normalized to α-Tubulin.

Immunofluorescence (IF)

To identify the distribution of proteins in intestinal tissue or enteroids, different sample preparation methods were applied. For paraffin sections, antigen retrieval was performed, followed by permeabilization with 0.3% Triton X-100 (Sigma, MO, USA). For enteroids, samples were fixed with 4% PFA for 15 min. Both types of samples were then blocked with normal goat serum (AR0009, Boster, Wuhan, China) for 30 min. Subsequently, the samples were incubated overnight at 4°C with the following primary antibodies: anti-LGR5 (customized, HuaBio, Hangzhou, China), anti-HOPX (self-prepared), and anti-Villin1 (HA500509, HuaBio, Hangzhou, China). After rinsing with PBS, the samples were incubated with TRITC-conjugated goat anti-mouse IgG (AS026, Abclonal, Wuhan, China) or ABflo 488-conjugated goat anti-rabbit IgG (AS053, Abclonal, Wuhan, China) at 37°C for 1 h, followed by DAPI staining at 37°C for 15 min. Images were captured using a Nikon A1R confocal laser scanning microscope (Tokyo, Japan) or an Olympus IX81-FV1000 confocal laser scanning microscope (Tokyo, Japan).

Statistical analysis

Statistical analysis was conducted using GraphPad Prism version 9.4.0. Outliers were identified and removed from the experimental raw data following the interquartile range principle. The Shapiro-Wilk test was used to assess the normality of the data. For comparisons between two groups, Student's t-test was applied to normally distributed data, while the Mann-Whitney U test was used for data with a non-normal distribution. In analyses involving multiple groups with normally distributed data, variance homogeneity was evaluated using Levene’s test. Based on this evaluation, either a one-way ANOVA with LSD post hoc test (assuming equal variances) or Welch’s ANOVA with Dunnett's T3 post hoc test (not assuming equal variances) was applied. For comparisons among multiple groups with non-normally distributed data, the Kruskal-Wallis H test was utilized. A P-value < 0.05 was considered statistically significant.

Results

A strain of Ligilactobacillus salivarius isolated from the gut microbiota

To isolate and identify superior Ligilactobacillus salivarius strains from the gut microbiota of ZN hens, a comprehensive screening process was conducted. A strain exhibiting optimal characteristics was selected for in-depth validation of its probiotic properties. On MRS agar, the strain showed vigorous growth, forming round, convex, milky-white colonies with smooth margins; colonies were plump, moist, and opaque (Fig. 1A). Gram staining confirmed it was Gram-positive, with short rod-shaped cells having rounded ends and no spore formation (Fig. 1B). Scanning electron microscopy (SEM) revealed a rod-shaped morphology, with cells approximately 2 to 3 μm in length and 0.5 to 1 μm in diameter (Fig. 1C). The sequencing identified the strain as Ligilactobacillus salivarius, designated L. salivarius 8-2 (L. sa. 8-2). In antibacterial assays, the strain significantly inhibited the growth of four common intestinal pathogens: Escherichia coli, Staphylococcus aureus, Salmonella Typhimurium, and Salmonella Enteritidis. Notably, its antibacterial activity was stronger than that of the commercial strain L. sa. CGMCC 1.1881, indicating superior probiotic potential (Fig. 1D, Table 3). These results demonstrate that L. sa. 8-2 possesses potent antibacterial properties against prevalent intestinal pathogens in the poultry industry, supporting its potential as a candidate for further probiotic development.

Fig. 1.

A newly isolated strain of Ligilactobacillus salivarius from Zhenning (ZN) hens. (A) Colony morphology of L. sa. 8-2. (B) Gram staining of L. sa. 8-2. (C) Scanning electron micrograph of L. sa. 8-2. (D) Antibacterial activity of L. sa. 8-2 against Escherichia coli, Staphylococcus aureus, Salmonella Typhimurium, and Salmonella Enteritidis.

Table 3.

Inhibition zone diameter.

Strain	Inhibition zone diameter (mm)
	Escherichia coli	Staphylococcus aureus	Salmonella Typhimurium	Salmonella Enteritidis
L. sa. 8-2	2.73±0.25	2.23±0.32	3.70±0.10	1.93±0.15
L. sa. CGMCC1.1881	2.37±0.06	1.90±0.20	2.30±0.10	1.57±0.15

L. sa. 8-2 alleviates intestinal injury and strengthens the intestinal physical barrier

To investigate whether L. sa. 8-2 exerts a protective effect against intestinal mucosal injury, two in vivo models, LPS challenge and STm infection, were utilized. As shown in Fig. 2A, challenge with either LPS or STm resulted in a significant increase in apoptotic cells as detected by TUNEL assay. In contrast, pre-feeding with L. sa. 8-2 markedly reduced the number of apoptotic cells (Fig. 2A). To assess alterations in epithelial barrier function, goblet cells and columnar cells were analyzed. AB-PAS staining showed that, in the LPS or STm group, goblet cell density increased by 62.50% to 93.75% compared with the MRS+PBS group. Interestingly, the L. sa. 8-2 p+LPS group and the L. sa. 8-2 s+STm group showed an even higher goblet cell density, with a further increase of 29.57% to 36.54% (P < 0.01) (Fig. 2B and C). Moreover, following STm challenge, Muc2 mRNA abundance was significantly increased by 137.74% and 63.21% in the L. sa. 8-2 s and L. sa. 8-2 p pretreatment groups, respectively, compared with the MRS+STm group (P < 0.01) (Fig. 2D). Additionally, the protein levels of Villin1 and Claudin1 were analyzed to evaluate the mechanical barrier of the intestinal mucosa. IF staining (Fig. 2E) confirmed that the intestinal epithelium was Villin1-positive. Furthermore, protein expression analysis showed that pretreatment with L. sa. 8-2 s dramatically elevated Claudin1 levels under either LPS or STm challenge (Fig. 2F–H). Collectively, L. sa. 8-2 and its supernatant mitigated intestinal injury and strengthened the intestinal physical barrier under infectious or inflammatory conditions.

Fig. 2.

L. sa. 8-2 reduced cell apoptosis and improved intestinal mucosal barrier function during intestinal injury. Chicks were fed the supernatant or precipitate of L. sa. 8-2 for 14 consecutive days, followed by challenge via i.p. injection with lipopolysaccharide (LPS) (10 mg/kg BW) or oral gavage with Salmonella Typhimurium (STm) (1 × 10⁹ CFU/chick). (A) TUNEL assay shows the alteration of apoptotic cells in the duodenum. Scale bar = 100 μm. (B-C) AB-PAS staining assay shows the alteration of goblet cell density in duodenal villi. Scale bar = 50  μm. (D) The alteration of Muc2 mRNA abundance in crypts, normalized to GAPDH. (E) Immunofluorescent staining of Villin1 protein in the duodenum. Scale bar = 50 μm. (F-H) The alteration of Villin1 (VLN1) and Claudin1 (CLDN1) protein levels in the duodenum. Control: chicks were fed a basal diet. MRS+PBS, MRS+LPS and MRS+STm: chicks were fed a basal diet containing MRS, then challenged with PBS, LPS or STm, respectively; L. sa. 8-2 s+LPS, L. sa. 8-2 s+STm: chicks were fed a basal diet containing L. sa. 8-2 supernatant, then challenged with LPS or STm, respectively; L. sa. 8-2 p+LPS, L. sa. 8-2 p+STm, chicks were fed a basal diet containing L. sa. 8-2 precipitate, then challenged with LPS or STm, respectively. Data are presented as mean ± standard deviation (n = 3). Columns without common letters indicate significant differences (P < 0.05) among groups.

L. sa. 8-2 enhanced the turnover of intestinal epithelium

To investigate the underlying mechanisms of barrier function enhancement, the cell cycle of the intestinal crypts was analyzed. Compared with the MRS+PBS group, STm challenge increased the expression levels of Ccnd1 and Cdk2 by 88.62% to 98.70%, respectively. Under LPS challenge, Ccnd1 expression decreased by 67.75% compared with the MRS+PBS group. However, pretreatment with L. sa. 8-2 s and L. sa. 8-2 p significantly elevated the abundance of Ccnd1 (by 94.98% to 293.48%) and Cdk2 (by 16.78% to 80.52%) (Fig. 3A and B). Moreover, higher levels of proliferating cell nuclear antigen (PCNA) were detected in the L. sa. 8-2 s+LPS group compared with the LPS or STm challenge groups, showing a 38.06% increase compared with the MRS+LPS group (P < 0.01) (Fig. 3C and D). These findings suggest that L. sa. 8-2 may accelerate the cell cycle and promote the proliferation of intestinal epithelial cells in crypts under injury.

Fig. 3.

L. sa. 8-2 enhanced the turnover of intestinal epithelium during intestinal injury. Chicks were fed the supernatant or precipitate of L. sa. 8-2 for 14 consecutive days, followed by challenge via i.p. injection with LPS (10 mg/kg BW) or oral gavage with STm (1 × 10⁹ CFU/chick). (A-B) The mRNA expression levels of cell cycle-related genes Ccnd1 and Cdk2 in the duodenum, normalized to GAPDH. (C-D) The alteration of PCNA protein level in the duodenum. Data are presented as mean ± standard deviation (n = 3). Columns without common letters indicate significant differences (P < 0.05) among groups.

L. sa. 8-2 promoted the activity of ISCs

To investigate the basis for the enhanced mucosal renewal rate, the activity of both active intestinal stem cells (aISC) and reserve intestinal stem cells (rISC) was assessed following mucosal damage. In vivo, compared with the MRS+PBS group, LPS and STm challenge decreased the protein levels of LGR5 and HOPX by 14.58% to 40.84% (Fig. 4A–C). However, pretreatment with L. sa. 8-2 s reversed this effect in the LPS-challenged group, significantly elevating LGR5 and HOPX protein levels by 59.09% and 173.00%, respectively, compared with the injured group. In enteroid models, protein level analysis showed elevated levels of LGR5 (by 74.96%, P < 0.01) and HOPX (by 43.11%, P < 0.01) in the L. sa. 8-2 s+LPS group compared with the MRS+LPS group (Fig. 4D–F). Co-staining for LGR5 and HOPX in enteroids confirmed that LPS treatment notably diminished the fluorescence intensity of both markers compared with the control group, whereas this effect was reversed in the L. sa. 8-2 s pretreatment group. Moreover, more HOPX⁺ cells were co-localized with LGR5⁺ cells in the pretreatment group (Fig. 4G). These findings suggest that L. sa. 8-2 may boost the activity of both active and reserve ISCs, thereby playing a crucial role in improving barrier function and facilitating mucosal repair.

Fig. 4.

L. sa. 8-2 promoted the activity of ISCs during intestinal injury. For in vivo, chicks were fed the supernatant or precipitate of L. sa. 8-2 for 14 consecutive days, followed by challenge via i.p. injection with LPS (10 mg/kg BW) or oral gavage with STm (1 × 10⁹ CFU/chick). For in vitro, enteroids were pretreated with the supernatant of L. sa. 8-2 for 24 h, followed by treatment with LPS (2  μg/mL) for 24 h. (A-C) The alteration of LGR5 and HOPX protein levels in the duodenum. (D-F) The alteration of LGR5 and HOPX protein levels in enteroids. (G) Immunofluorescent staining of HOPX and LGR5 protein in enteroids. Scale bar = 10 μm. Data are presented as mean ± standard deviation (n = 3). Columns without common letters indicate significant differences (P < 0.05) among groups.

L. sa. 8-2 protected the integrity of mitochondrion

To explore the mechanism underlying enhanced ISC activity, changes in mitochondrial morphology were investigated. Transmission electron microscopy (TEM) revealed distinct differences in mitochondrial morphology among the experimental groups (Fig. 5A). In both the LPS and STm groups, mitochondria exhibited pronounced vacuolation, discontinuous membrane structures, and blurred or even absent cristae, indicating severe mitochondrial dysfunction. In contrast, in the L. sa. 8-2 pretreatment groups, overall mitochondrial morphology remained regular, although cristae showed some degree of swelling, suggesting partial protection or restoration of mitochondrial function. Further analysis of mitochondrial dynamics-related proteins in the intestinal crypt demonstrated an increasing trend in the fission-related protein FIS1 in both the LPS and STm challenge groups. Moreover, FIS1 levels were further elevated in the L. sa. 8-2-pretreated groups, with levels 70.69% to 122.43% higher than those in the MRS groups (P < 0.01). In contrast, the expression levels of fusion-related proteins, including optic atrophy 1 (OPA1) and mitofusin 1 (MFN1), did not show significant changes compared with the MRS groups (Fig. 5B–E). These findings suggest that under mucosal damage conditions, L. sa. 8-2 may promote mitochondrial fission, thereby facilitating the removal of dysfunctional mitochondria and maintaining mitochondrial structural integrity.

Fig. 5.

L. sa. 8-2 alleviates mitochondrial damage during intestinal injury. Chicks were fed the supernatant or precipitate of L. sa. 8-2 for 14 consecutive days, followed by challenge via i.p. injection with LPS (10 mg/kg BW) or oral gavage with STm (1 × 10⁹ CFU/chick). (A) Transmission electron microscopy (TEM) of intestinal crypts. The upper row shows overall morphology of the crypt epithelium. Scale bar = 2 μm. The lower row shows enlarged images of mitochondria. Scale bar = 1 μm . (B-E) The alteration of FIS1, OPA1 and MFN1 protein levels in crypts. Data are presented as mean ± standard deviation (n = 3). Columns without common letters indicate significant differences (P < 0.05) among groups.

L. sa. 8-2 strengthens pyruvate utilization during intestinal mucosal repair

To uncover the mechanisms underlying the intestinal repair effects of L. sa. 8-2 supernatant, enteroids were pretreated with either MRS (control) or L. sa. 8-2 supernatant for 24 h, followed by LPS challenge for another 24 h. Transcriptome analysis of the enteroids was then conducted to explore the potential pathways involved in this repair process. Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis demonstrated that, compared with the MRS+LPS group, differentially expressed genes in the L. sa. 8-2 s+LPS group were significantly enriched in several metabolic pathways, including tyrosine metabolism, retinol metabolism, and pyruvate metabolism (Fig. 6A). The heat map of differentially expressed genes (Fig. 6B) revealed that, compared with LPS treatment alone, the L. sa. 8-2 s+LPS group exhibited downregulation of glycolytic genes (e.g., HK2, PFKP) and upregulation of pyruvate metabolism-related genes, particularly LDHA and LDHB, suggesting enhanced mitochondrial pyruvate utilization. qPCR analysis confirmed that, compared with LPS treatment alone, pretreatment with L. sa. 8-2 supernatant increased Ldha and Ldhb mRNA levels by 77.07% and 145.60% (P < 0.01), while it decreased mRNA levels of Sdha and the glycolysis-related gene HK2 by 47.31% and 35.87%, respectively (Fig. 6C–I). At the protein level, LDHA and LDHB in the L. sa. 8-2 s+LPS group were higher by 128.97% (P < 0.01) and 35.46% (P > 0.05) than those in the MRS+LPS group. Similarly, in vivo studies showed a consistent trend. Compared with the MRS+LPS group, pretreatment with L. sa. 8-2 supernatant increased the mRNA and protein levels of Ldha and Ldhb. However, in the L. sa. 8-2 s+LPS group, the mRNA abundance of Sdha and HK2 decreased by 50.45% and 55.44%, respectively, relative to the MRS+LPS group (Fig. 6J–P). These results suggest that L. sa. 8-2 strengthens local mitochondrial pyruvate utilization in the intestinal crypt during intestinal mucosal repair.

Fig. 6.

L. sa. 8-2 promotes glycolytic reprogramming during intestinal mucosal repair. For in vivo, chicks were fed the supernatant or precipitate of L. sa. 8-2 for 14 consecutive days, followed by challenge via i.p. injection with LPS (10 mg/kg BW) or oral gavage with STm (1 × 10⁹ CFU/chick). For in vitro, enteroids were pretreated with the supernatant of L. sa. 8-2 for 24 h, followed by treatment with LPS (2  μg/mL) for 24 h. (A) KEGG analysis revealed enriched pathways in the “L. sa. 8-2 s+LPS” group compared with the “MRS+LPS” group. (B) Hierarchical clustering analysis of differentially expressed genes between the “MRS+LPS” group and the “L. sa. 8-2 s+LPS” group. Red and blue blocks indicate upregulated and downregulated genes, respectively. (C-F) The alteration of Ldha, Ldhb, Sdha and HK2 mRNA abundance in enteroids, normalized to GAPDH. (G-I) The alteration of LDHA and LDHB protein levels in enteroids. (J-M) The alteration of Ldha, Ldhb, Sdha and HK2 mRNA abundance in the duodenum, normalized to GAPDH. (N-P) The alteration of LDHA and LDHB protein levels in crypts. Data are presented as mean ± standard deviation (n = 3). Columns without common letters indicate significant differences (P < 0.05) among groups.

Discussion

Our study demonstrated that feeding with L. sa. 8-2 prior to LPS or STm challenge in chicks reduced apoptotic cells and strengthened the intestinal mucosal barrier, as illustrated by an increased density of goblet cells and elevated protein levels of Villin1 and Claudin1 in the intestinal epithelium. These findings align with previous reports on the protective roles of various probiotics. For instance, Lactobacillus plantarum HNU082 (Lp082) was shown to protect the intestinal mucosal barrier in a DSS-induced colitis mouse model by increasing goblet cells and Muc2 expression (Wu et al., 2022). Similarly, a probiotic mixture (Lactobacillus acidophilus LA1, Lactobacillus reuteri LR92, Bifidobacterium breve Bbr8) enhanced the integrity of Occludin and ZO-1 proteins in Caco-2 cells (Tinazzi et al., 2024). Additionally, in broiler chicks, a combination of Lactobacillus salivarius and Pediococcus parvulus (Floramax® B11) reduced intestinal permeability and decreased the colonization of S. Enteritidis (Prado-Rebolledo et al., 2017). Furthermore, a combination of Lactobacillus crispatus, Lactobacillus salivarius, Lactobacillus gallinarum, Lactobacillus johnsonii, Enterococcus faecalis, and Bacillus amyloliquefaciens inhibited Salmonella colonization in the gastrointestinal tract of broilers (Neveling et al., 2020). Therefore, we propose that L. sa. 8-2 exerts a protective barrier effect against intestinal mucosal damage induced by LPS or STm infection.

It is well known that intestinal mucosal barrier function is closely linked to epithelial renewal. Our study further confirmed that L. sa. 8-2 promoted intestinal epithelial cell proliferation under impaired situation, as evidenced by upregulated PCNA expression and increased mRNA levels of Cdk2 and Ccnd1. Previously, it was demonstrated that L. salivarius enhances the activity of ISCs to promote epithelial turnover and improve the intestinal mucosal absorptive capacity (Liu et al., 2022). Similarly, succinate, produced by Lactobacillus metabolism, enhances the proliferation and differentiation of ISCs in laying hens (Zhou et al., 2022). Intestinal epithelial cell renewal is based on the activity of ISCs in the crypt. Furthermore, after intestinal injury, the activation of rISCs, which is characterized by upregulation of the Hopx, initiates mucosal repair (Yu et al., 2023). In our study, under injury conditions, L. sa. 8-2 promoted the protein expression of LGR5 and HOPX both in vivo and in vitro. Notably, co-staining of LGR5 and HOPX in enteroids revealed an increased number of HOPX⁺ cells in the L. sa. 8-2 s pretreatment group, and these cells were predominantly co-localized with LGR5⁺ cells, reflecting an accelerated transition of rISCs into aISCs during the repair phase. These findings indicate that L. sa. 8-2 promotes the initiation of mucosal repair. In addition, lactate derived from L. plantarum was shown to stimulate ISC proliferation in mice by enhancing mitochondrial respiration in the crypts and activating Wnt3/β-catenin signaling in ISCs (Lee et al., 2018). Collectively, we propose that L. sa. 8-2 strengthens ISC activity and plays a key role in intestinal mucosal renewal.

The repair of the intestinal mucosa rely on the activation and state transition of ISCs, a process highly dependent on the energy supply pathways. This study demonstrated that mitochondrial morphology was severely disrupted under mucosal injury conditions. Conversely, pretreatment with L. sa. 8-2 effectively preserved the overall mitochondrial architecture and attenuated ultrastructural damage. Furthermore, analysis of mitochondrial dynamics-related proteins confirmed that L. sa. 8-2 promoted mitochondrial fission under mucosal injury conditions. Fragmented mitochondria also enable the elimination of damaged mitochondria through mitophagy, thereby maintaining the quality of the bioenergetic machinery (Kawano et al., 2023). Moreover, enhancing mitochondrial fission promotes a metabolic shift toward glycolysis (Prieto et al., 2016). In our study, L. sa. 8-2 pretreatment led to a pronounced increase in the expression of LDHA over LDHB. The lactate dehydrogenase isoforms LDHA and LDHB differentially regulate pyruvate metabolism, and their functional balance is central to cellular energy production and biosynthesis (Gray et al., 2014). It is known that LDHA favors the reaction from pyruvate to lactate (Rahman et al., 2016; Park et al., 2018), which enables it to robustly sustain anaerobic glycolysis (Prochownik and Wang, 2021). Thereby, sustaining ATP production to supply energy rapidly for activated ISCs under injury conditions. In contrast, LDHB favors the reverse reaction (lactate to pyruvate). Concurrently, in the present study, a relative decrease in the expression of Sdha (a gene encoding a subunit of the OXPHOS complex II) was observed, reflecting a metabolic shift from OXPHOS toward glycolysis. Collectively, during the intestinal mucosal repair process, L. sa. 8-2 promotes ISC proliferation and differentiation by inducing mitochondrial fission, and enhances pyruvate utilization in the intestinal crypt.

In summary, L. sa. 8-2 alleviates intestinal mucosal injury and promotes epithelial renewal by activating ISCs, which is mediated by promoting mitochondrial fission and the subsequent enhancement of pyruvate utilization in the intestinal crypt.

Funding

This study was supported by the National Natural Science Foundation of China (No. 31972630), Hangzhou Chengxi Sci-tech innovation Corridor Management Committee, and the Scientific Research Fund of Zhejiang Provincial Education Department (Y202457021).

Availability of data and material

The authors confirm that the data supporting the findings of this study are available within the article. The datasets supporting the conclusions of this article are available in the NCBI GenBank no. PX314491.

CRediT authorship contribution statement

Yuping Hua: Writing – original draft, Investigation, Data curation. Minyao Zou: Investigation. Lingzhi Zhang: Investigation. Leixiao Chen: Investigation. Xi Rao: Investigation. Guozhen Wei: Investigation. Jiaxin Wei: Investigation. Mingping Wei: Investigation. Yihang Wei: Investigation. Jian Li: Writing – review & editing, Supervision, Funding acquisition, Data curation, Conceptualization.

Disclosures

The authors report there are no competing interests to declare.