The Probiotic Effects of Lactobacillus intestinalis Y10 Isolated From Feces of Wild Brown Rat (Rattus norvegicus)

TL;DR

This study identifies a probiotic strain from wild rat feces that survives harsh conditions and inhibits harmful bacteria, showing potential for rodent-specific probiotic use.

Contribution

The study characterizes Lactobacillus intestinalis Y10 as a novel probiotic candidate with strong survival and antimicrobial properties from wild rat feces.

Findings

L. intestinalis Y10 showed 72.4% viability after 2 hours in acidic conditions (pH 2.5).

The strain exhibited high hydrophobicity (93.80%) and auto-aggregation (81.68%), indicating strong mucosal adhesion.

It demonstrated concentration-dependent antimicrobial activity against E. coli, P. aeruginosa, and S. aureus.

Abstract

The gut microbiota of wild mice exhibits a significant correlation with their environmental adaptability, particularly highlighted by the dominance of Lactobacillus spp. This study evaluated the probiotic traits of Lactobacillus intestinalis Y10, a fecal isolate from Rattus norvegicus, through in vitro assays. The strain demonstrated two‐hour survival in acidic conditions (pH 2.5; 72.4% viability) and maintained viability under 0.3% bile salts for 24 h. It also showed high hydrophobicity index (93.80%) and auto‐aggregation percentage (81.68%), indicative of superior mucosal adhesion potential. Organic acid–mediated antimicrobial and antibiofilm activity of strain Y10 against Escherichia coli, Pseudomonas aeruginosa, and Staphylococcus aureus subsp. aureus exhibited concentration dependence. Antibiotic susceptibility profiling identified sensitivity to cell wall–targeting agents (penicillin and cephalosporin) but resistant to antibiotics that inhibit nucleic acid synthesis and cytoplasmic membrane function, suggesting the possibility of clinical compatibility with fluoroquinolones or diaminopyrimidines for treatment of specific infections to minimize gut microbiota disruption. These findings position L. intestinalis Y10 as a promising candidate for rodent‐specific probiotic applications, combining environmental resilience with pathogen exclusion capabilities. Further in vivo validation and mechanistic studies are required to translate these traits into therapeutic interventions for gastrointestinal dysbiosis.

Figures

Maximum likelihood (ML) phylogenetic tree based on 16S rRNA gene sequences. Bootstrap values (> 70%) calculated for 1000 replicates are shown at branch nodes. Pediococcus damnosus JCM 5886 was used as an outgroup. Strains isolated in this study are designated by dots. Bar, 0.03 substitutions per nucleotide position.

Biofilm inhibition effects of L. intestinalis Y10 CFCS. The continuously twofold diluted CFCS was cocultured with Escherichia coli ATCC 25922, Pseudomonas aeruginosa ATCC 27853, and Staphylococcus aureus subsp. aureus ATCC 25923, respectively, for 24 h.

Tables

TABLE 1. Antibiotic susceptibility profiles of Lactobacillus intestinalis Y10.

Antibiotics and concentrations	Zone diameters (mm) a	Antibiotics and concentrations	Zone diameters (mm) a
Amoxicillin 25 μg	+++	Tylosin 30 μg	+++
Amoxicillin–clavulanic acid 30 μg	+	Clindamycin 2 μg	++
Penicillin G 10 U	+++	Lincomycin 2 μg	+
Ceftiofur 30 μg	+++	Florfenicol 30 μg	+++
Cephalothin 30 μg	+++	Trimethoprim 5 μg	—
Bacitracin 10 IU	+++	Trimethoprim‐sulfamethoxazole 25 μg	—
Amikacin 30 μg	—	Ciprofloxacin 5 μg	—
Gentamicin 30 μg	+	Enrofloxacin 5 μg	—
Kanamycin 30 μg	—	Norfloxacin 10 μg	—
Streptomycin 10 μg	—	Ofloxacin 5 μg	—
Doxycycline 30 μg	+	Colistin sulfate 10 μg	—
Minocycline 30 μg	+	Polymyxin B 300 IU	—

TABLE 2. MIC and MBC values of Lactobacillus intestinalis Y10.

	E. coli ATCC 25922	P. aeruginosa ATCC 27853	S. aureus ATCC 25923
MIC	1/4	1/8	1/4
MBC	1/2	1/4	1 a

Full text

1. Introduction

Probiotics are live microorganisms that, when administered in sufficient quantities, provide a health benefit to the host [1]. They encompass various microorganisms, such as Lactobacillales, Bacillus, Bifidobacterium, and certain yeasts. Lactobacillus, the earliest discovered probiotic genus, has been proven to play crucial roles in pathogenic bacterium antagonism, intestinal disease treatment, immune regulation, anticancer activity, and metabolic regulation [2, 3].

Lactobacillus intestinalis is a native commensal in the gastrointestinal tract of rodents, where it contributes to vital host–microbe interactions [4]. It modulates immune responses by metabolizing dietary vitamin A to retinoic acid, thereby suppressing colitis‐associated Th17 inflammation [5, 6]. Additionally, its mercury‐binding capacity alleviates mercury toxicity on the intestinal epithelium [7]. Emerging evidence further links L. intestinalis abundance to lipid metabolism, indicating its potential therapeutic role in improving serum lipid profiles and attenuating atherosclerotic progression [8, 9]. However, despite these established roles, the probiotic properties of L. intestinalis remain critically underexplored, especially wild‐derived strains. Only one study has focused on laboratory‐adapted strains, characterized through conventional assays, such as tolerance test, aggregation assay, and antibiotic susceptibility [10].

Notably, gut microbiota derived from wild mouse populations drives distinctive composition compared to laboratory‐reared counterparts [11, 12]. Wild rodents, inhabiting ecosystems with fluctuating nutrient availability and pathogen exposures, are likely to harbor wild‐derived strains with superior environmental resilience and antimicrobial activity. Laboratory mice reconstituted with the wild mouse gut microbiota demonstrate enhanced infection resistance and reduced inflammation, underscoring the ecological advantage of undomesticated microbial lineages [12, 13]. It is therefore speculated that the evolutionary adaptation of Lactobacillus spp. to natural ecosystems confers enhanced probiotic properties relative to laboratory strains. Moreover, the remarkable stability of lyophilized Lactobacillus formulations and their compatibility with oral administration routes significantly facilitate clinical translation and practical applications. However, systematic evaluation of core probiotic characteristics in wild rodent–derived L. intestinalis remains absent, leaving a significant knowledge gap in host‐adapted probiotic development.

To qualify as probiotics, microorganisms must undergo a systematic selection process [14]. Firstly, the candidate strains must survive gastrointestinal stressors, including gastric acidity, pepsin, and bile salts [14–16]. Secondly, they need to possess adhesion abilities in order to colonize the gastrointestinal tract. This adherence capability is closely linked to the auto‐aggregation capacity and hydrophobic properties of the potential probiotic strains [14, 15, 17]. Additionally, effective probiotics demonstrate strong antipathogenic activities, facilitated by the production of extracellular antimicrobial components, including bacteriocins, hydroperoxides, organic acids, and peptides [3, 18]. Lastly, probiotics must meet safety criteria, ensuring the absence of toxicity, infectivity, and transferable antibiotic resistance genes among the candidates [19].

Thus, this study aimed to investigate the probiotic potential of L. intestinalis Y10, a strain isolated from the feces of a wild brown rat (Rattus norvegicus), through comprehensive in vitro evaluation of its survival traits, adhesion ability, antimicrobial activity, and safety profile. Our findings will provide foundational data on the probiotic properties of wild‐derived L. intestinalis strains, supporting its development as a biosafe therapeutic candidate for treating dysbiosis in rodent models.

2. Materials and Methods

2.1. Bacteria Strain and Culture Condition

Escherichia coli ATCC 25922, Pseudomonas aeruginosa ATCC 27853, and Staphylococcus aureus subsp. aureus ATCC 25923 were provided by the Bioresource Collection and Research Center (Hsinchu, Taiwan) and utilized as target pathogens. All the three strains were cultured in tryptic soy broth (Difco, Le Pont de Claix, France) and stored in 50% glycerol at −20°C.

2.2. Isolation of Lactobacillus

L. intestinalis Y10 was isolated from a wild female brown rat (Rattus norvegicus) trapped at the National Chung Hsing University in Taichung City, Taiwan. The rat was released immediately after noninvasive fecal collection. Individual cases did not require ethical approval. The collected fecal sample was mixed with sterile saline, followed by vortexing. The mixture was then streaked on an MRS agar plate (Difco, Le Pont de Claix, France) and cultured aerobically and anaerobically at 37°C for 48 h. Subsequently, colonies were picked up for pure culture and subjected to Gram staining and microscopic examination. Finally, isolates were stored at −20°C in MRS broth supplemented with 50% glycerol for further analysis.

2.3. Bacterial Identification

To remove the MRS broth, the overnight culture of isolates was centrifuged at 12,000 rpm for five minutes. The extracted DNA was obtained using the phenol–chloroform method with the GenoMaker reagent kit (GenePure Technology, Taichung, Taiwan). For the amplification of the 16S rRNA gene, a polymerase chain reaction (PCR) was performed using universal primers: forward (5′‐ AGA GTT TGA TCC TGG CTC AG‐3′) and reverse (5′‐GGT TAC CTT GTT ACG ACT T‐3′) [20]. Each 20 μL PCR was prepared with the following components: 1.6 μL of dNTP Mixture, 2 μL of 10x Reaction Buffer, 0.2 μL of 5 U/μL APP Taq DNA polymerase (GeneTeks BioScience, Taipei, Taiwan), 0.2 μL of each primer (50 μM/μL), 14.8 μL of Nuclease‐Free Water (Thermo Fisher Scientific, Waltham, USA), and 1 μL of sample DNA. Amplification was performed under the following conditions: initial denaturation at 95°C for 10 min; 35 cycles of denaturation (95°C, 30 s), annealing (55°C, 30 s), and extension (72°C, 90 s); followed by a final extension at 72°C for 10 min. The PCR product was then sent to a commercial sequencing company (Genomics BioSci & Tech. Co., Ltd, New Taipei, Taiwan) for sequencing. The resulting nucleotide sequence was compared to the GenBank database using the Geneious Prime software (Version 2023.2.1).

For phylogenetic analyses, 16S rRNA gene sequences of type strains were downloaded from the GenBank database. Sequences were aligned using the plugin MAFFT v7.490 [21] in Geneious Prime. ModelFinder [22] implemented in PhyloSuite v1.2.3 [23, 24] was used to test the best‐fit evolutionary model. The maximum likelihood (ML) phylogenies were inferred under the GTR + I + G4+F model for 1000 ultrafast [25] bootstraps using IQ‐TREE v2.2.0 [26].

2.4. Preliminary Screening of Lactobacillus Strains With Antimicrobial Activity

Isolated strains were cultured in MRS broth for three days at 37°C. The cell‐free culture supernatant (CFCS) was collected by centrifugation at 3500 rpm for five minutes, followed by filtration using a sterile Millex‐GS syringe filter unit (0.22 μm, Merck KGaA, Darmstadt, Germany). To assess the activity, E. coli ATCC 25922, P. aeruginosa ATCC 27853, and S. aureus subsp. aureus ATCC 25923 were utilized as indicators.

The antimicrobial activity of the isolated strains was initially evaluated using a microplate‐based method. Metabolites from the isolated strains and pathogens were mixed in a 96‐well plate (BIOFIL, Guangzhou, China). Each well contained 90 μL of the pathogen suspension at a concentration of 10^5^ colony‐forming units per milliliter (CFU/mL) and 10 μL of the CFCS. The growth control was 90 μL of the pathogen suspension and 10 μL of MRS broth. Each well was replicated three times. The microplate was then incubated at 37°C, and the optical density at 600 nm (OD_600 nm_) was measured at 0, 8, and 16 h for P. aeruginosa and S. aureus, as well as at 0, 6, and 12 h for E. coli, using a microplate reader (Tecan Sunrise, Australia).

2.5. Acid and Bile Salt Tolerance

The acid and bile salt tolerance testing was conducted according to previous studies with minimal modifications [27, 28]. The overnight culture of L. intestinalis Y10 was centrifuged at 3500 rpm for 10 min and washed three times with sterile phosphate‐buffered saline (PBS, pH 7.4). It was then resuspended, and the optical density (OD_600 nm_) was adjusted to 1.0 ± 0.05 (10^7^ CFU/mL) using the same buffer solution with a cell density meter (Ultraspec 10, Amersham Biosciences).

For the acid tolerance test, 600 μL of the washed cell suspension was inoculated, respectively, into 6 mL of MRS broth which was buffered with PBS and preadjusted to pH 2.5 and pH 7.4 by hydrogen chloride and sodium hydroxide. The mixture was incubated at 37°C. At each interval (0, 1, 2, 4, and 6 h), 100 μL of the mixture was serially diluted 10‐fold using sterile saline, and 25 μL of each dilution was spotted onto MRS agar in duplicate. Colony count was performed after anaerobic incubation at 37°C for 48 h and expressed as log CFU/mL. The survivability of L. intestinalis in acidic PBS was also tested using the same method, and the survival rate (%) was calculated as viable cell counts in the test group/control group × 100 at the same time points, with both counts expressed in CFU/mL.

For the bile salt tolerance test, two groups containing 6 mL of MRS broth were established with bile salt (cholic acid–deoxycholic acid sodium salt mixture, Sigma‐Aldrich, Cat. No. B8756) concentrations of 0% (control) and 0.3% (w/v). Each group was inoculated with 600 μL of the washed cell suspension followed by incubation at 37°C. Viable cell counts were performed at 0, 1, 2, 4, 6, and 24 h using the same methods in the acid tolerance test. About 100 μL aliquots underwent serial 10‐fold dilution with sterile saline, and 25 μL of each dilution was spotted onto MRS agar plates in duplicate. All the plates were incubated anaerobically at 37°C for 48 h.

2.6. Cell Surface Hydrophobicity Assay

The in vitro cell surface hydrophobicity was determined using the method described by Ekmekci et al. [29]. The cell suspension was prepared as described in the acid and bile salt tolerance test, with the initial optical density at 600 nm (OD_600 nm_) measured as A 0. Thereafter, 3 mL of the cell suspension was mixed with 1 mL of toluene (Chung Shing Chemicals Co., Ltd., Taiwan) in a glass test tube and vortexed for 90 s. The mixture was then incubated at 37°C for 30 min. After incubation, the aqueous phase was carefully extracted into a new tube, and the OD_600 nm_ was measured immediately as A_1_ for three replicates. The cell surface hydrophobicity (%) was calculated as the following equation:

$$[eqn]$$

2.7. Auto‐ and Co‐Aggregation Assays

The auto‐ and co‐aggregation assays were conducted following the method described by Reuben et al. [30]. For the auto‐aggregation assay, an overnight culture of L. intestinalis Y10 was pelleted and washed with sterile PBS (pH 7.4). The OD_600 nm_ of the washed cell suspension was adjusted to 0.5 ± 0.05 and recorded as A0. The bacteria suspension was then incubated stationarily at 37°C for 24 h. At 2, 4, 6, 12, and 24 h, 100 μL of the upper phase was carefully collected without disturbing the settled aggregates and transferred to a 96‐well plate for three replicates, and the OD_600 nm_ (A _ t _) was measured. The auto‐aggregation percentage was calculated according to the following equation:

$$[eqn]$$

For the co‐aggregation assay, E. coli ATCC 25922, P. aeruginosa ATCC 27853, and S. aureus subsp. aureus ATCC 25923 were utilized. The cell suspensions of the pathogens were prepared in the same manner. For each co‐aggregation sample, 3 mL of the L. intestinalis suspension was mixed with 3 mL of the suspension of each pathogen and vortexed. The absorbance (OD_600 nm_) of each mixture was recorded as A mix0. These mixtures were then incubated stationarily at 37°C for 24 h. At 2, 4, 6, 12, and 24 h, the OD_600 nm_ of the supernatant was measured for three replicates and recorded as A _mixt _. The co‐aggregation percentage was calculated as the following equation:

$$[eqn]$$

2.8. Antibiotic Susceptibility Test

To determine the antibiotic susceptibility of L. intestinalis Y10, the Kirby–Bauer disk diffusion method [31] was employed. L. intestinalis suspension was inoculated on MRS agar at a concentration of 10^8^ CFU/mL using a sterile swab. Antibiotic disks, including amoxicillin 25 μg, bacitracin 10 IU, ceftiofur 30 μg, cephalothin 30 μg, ciprofloxacin 5 μg, clindamycin 2 μg, colistin sulfate 10 μg, doxycycline 30 μg, enrofloxacin 5 μg, florfenicol 30 μg, gentamicin 30 μg, lincomycin 2 μg, trimethoprim–sulfamethoxazole 25 μg, tylosin 30 μg (Liofilchem, Roseto degli Abruzzi, Italy), amikacin 30 μg, amoxicillin–clavulanic acid 30 μg, kanamycin 30 μg, minocycline 30 μg, norfloxacin 10 μg, ofloxacin 5 μg, penicillin G 10 U, polymyxin B 300 IU, streptomycin 10 μg, and trimethoprim 5 μg (Oxoid, Basingstoke, UK), were used in the experiment. Disks were placed on the surface of the medium using sterile forceps. The plates were then incubated anaerobically at 37°C for 24 h. The diameter (mm) of the inhibition zone around each disk was measured and compared with the zone diameters provided by Charteris et al. [32].

2.9. Antagonistic Activity by Agar Well Diffusion Method

The agar well diffusion method [33, 34] was used to assess the antagonistic activity of L. intestinalis Y10 against P. aeruginosa, S. aureus, and E. coli, with minimal modifications. Following 20‐h incubation in MRS broth, the CFCS of strain Y10 was obtained by centrifugation at 3500 rpm for 10 min and subsequent membrane filtration sterilization through 0.22‐μm pore‐size filters. The CFCS, initially exhibiting a pH of 3.89, was adjusted to pH 6.5–7.0 using 1 mM sodium hydroxide. The neutral CFCS underwent filtration sterilization before use. About 100 μL of pathogen suspensions at densities of 10^5^ and 10^8^ CFU/mL was homogenously added into 25 mL of Mueller–Hinton agar (MHA, Difco, Le Pont de Claix, France) before solidification. Six wells with a diameter of 7 mm were created, and 100 μL of the neutral CFCS was added to their respective wells. After 20 h of incubation at 37°C under aerobic conditions, the diameter (mm) of the inhibition zone was measured.

2.10. Determination of Minimal Inhibitory Concentration (MIC) and Minimum Bactericidal Concentration (MBC)

The determination of the MIC and MBC was performed using the broth microdilution method [35, 36]. The CFCS of L. intestinalis Y10 was subjected to twofold serial dilution in MRS broth, resulting in a final volume of 50 μL in each well. Simultaneously, 50 μL of 10^5^ CFU/mL pathogen suspended in Mueller–Hinton broth (MHB) was inoculated into each well. The growth control consisted of MRS broth with the bacterial inoculum, while the negative control contained equal volumes of MRS broth and MHB. The microtiter plates were incubated at 37°C for 20 h. Subsequently, 50 μL of the mixture from each well was dropped onto tryptic soy agar (TSA, Difco, Le Pont de Claix, France), and the colonies were counted after incubating at 37°C for 24 h. The MIC was defined as the lowest concentration of CFCS that inhibited the visible growth of microorganisms after overnight incubation. The MBC was determined as the lowest concentration of CFCS that prevented bacterial growth after subculture.

2.11. Minimum Biofilm Inhibitory Concentration (MBIC) Test

The MBIC of L. intestinalis Y10 was analyzed following the protocol published by Stepanović et al. [37]. The microtiter plates were prepared using the same manner as for determining MIC. Each well contained 50 μL of pathogen suspension (10^5^ CFU/mL) and 50 μL of twofold serially diluted CFCS. The plate was then incubated aerobically for 24 h at 37°C. After incubation, the plates were washed with a saline solution, fixed with pure methanol, stained with crystal violet, and dried. The dye bound to the adherent cells was dissolved by adding 33% (v/v) acetic acid. Subsequently, the absorbance at 570 nm was measured using a microplate reader. The biofilm inhibition rate was calculated as the following equation:

$$[eqn]$$

In the equation, A _ t _ represents OD_570 nm_ of the tested well, in which the CFCS of L. intestinalis Y10 was cocultured with each pathogen. A _ n c _ represents OD_570 nm_ of the negative control. A _ g c _ represents OD_570 nm_ of the growth control. Therefore, the MBIC_50_ and MBIC_90_ were determined as the lowest concentration of CFCS that inhibited biofilm formation by 50% and 90%.

2.12. Statistical Analysis

The data were analyzed using a two‐way analysis of variance, and multiple comparisons were performed using Tukey’s test. Statistical significance was determined at a threshold of p < 0.05, and the results are presented as mean ± standard deviation (SD).

3. Results

3.1. Isolation and Identification of Lactic Acid Bacteria

A total of 12 candidate strains, exhibiting gram‐positive characteristics, were isolated from the feces of wild rats. Based on the obtained 16S rRNA gene ranging from 1415 bp to 1445 bp, three strains were identified as Limosilactobacillus reuteri (99.9%–100%), three as Limosilactobacillus oris (99.9%), two as Lactobacillus crispatus (99.9%–100%), two as Lactobacillus johnsonii (99.9%–100%), one as Ligilactobacillus murinus (100%), and one as L. intestinalis (99.8%). A ML phylogenetic tree was constructed based on the 16S rRNA gene sequences of the type strains belonging to Lactobacillus, Limosilactobacillus, and Ligilactobacillus (Figure 1). The tree revealed three major clades, with Pediococcus damnosus JCM 5886 serving as the outgroup. Only branches with bootstrap support values > 70% are shown. All strains isolated in this study clustered within these three clades, and their phylogenetic positions were consistent with the results obtained from BLASTn sequence analysis.

Maximum likelihood (ML) phylogenetic tree based on 16S rRNA gene sequences. Bootstrap values (> 70%) calculated for 1000 replicates are shown at branch nodes. Pediococcus damnosus JCM 5886 was used as an outgroup. Strains isolated in this study are designated by dots. Bar, 0.03 substitutions per nucleotide position.

3.2. Preliminary Screening and Acid and Bile Salt Tolerance

During the preliminary screening, L. intestinalis Y10 exhibited a high antibacterial activity against three pathogens, as the optical density at 600 nm showed no increase after incubation (Figure 2). Based on these results, and considering the limited data on L. intestinalis probiotic properties, L. intestinalis Y10 was selected for further experiments.

FIGURE 2Preliminary screening results of antimicrobial activity. (a) CFCS of the isolated strains was incubated with Escherichia coli ATCC 25922. (b) CFCS of the isolated strains was incubated with Pseudomonas aeruginosa ATCC 27853. (c) CFCS of the isolated strains was incubated with Staphylococcus aureus subsp. aureus ATCC 25923. GC, growth control.(a)(b)(c)

After 6 h of cultivation in acidic MRS broth (pH = 2.5), strain Y10 exhibited a concentration of 10^6.41^ CFU/mL, whereas the control group reached 10^8.41^ CFU/mL (p < 0.01, Figure 3(a)). It demonstrated resistance to low pH conditions for a period of 2 hours, with a survival rate of 72.4% (Figure 3(b)). In terms of bile salt tolerance, the viable cell counts in the group treated with 0.3% bile salts were significantly lower than the control group after 4 h (p < 0.05) (Figure 3(c)). However, L. intestinalis Y10 was able to survive exposure to 0.3% bile salts for 24 h, although there was no proliferation observed.

FIGURE 3Acid and bile salt tolerance test of Lactobacillus intestinalis Y10. (a) Growth curve in MRS broth at pH 2.5 and 7.4 within 6 h. (b) Survival rate in PBS at pH 2.5 and 7.4 within 6 h. The survival rate (%) was calculated as (viable cells in experimental group [CFU/mL]/viable cells in control group [CFU/mL]) × 100. (c) Growth curve in MRS broth with 0.3% bile salt.(a)(b)(c)

3.3. Cell Surface Hydrophobicity Assay

In the in vitro cell surface hydrophobicity assay, L. intestinalis Y10 demonstrated a high hydrophobicity index of 93.80% ± 0.60% in toluene. This result indicates a high adhesion property of L. intestinalis Y10.

3.4. Auto‐Aggregation Assay and Co‐Aggregation Assay

L. intestinalis Y10 exhibited a strong auto‐aggregation ability, with an auto‐aggregation percentage of 81.68% after 24 h (Figure 4(a)), indicating that L. intestinalis Y10 has a strong capability to adhere to epithelial cells and form a protective barrier. The co‐aggregation percentages of L. intestinalis Y10 with three pathogens were also assessed, and the results are presented in Figure 4(b). After 24 h, L. intestinalis Y10 displayed significant co‐aggregation with S. aureus ATCC 25923 (73.53%), followed by P. aeruginosa ATCC 27853 (64.25%) and E. coli ATCC 25922 (64.23%). These results confirm that L. intestinalis Y10 has the ability to competitively inhibit the adhesion of pathogenic strains, further supporting its potential as a beneficial microorganism.

FIGURE 4Auto‐aggregation and co‐aggregation assay measured at 2, 4, 6, 12, and 24 h. (a) Auto‐aggregation percentage of Lactobacillus intestinalis Y10. (b) Co‐aggregation percentages of Lactobacillus intestinalis Y10 with Escherichia coli ATCC 25922, Pseudomonas aeruginosa ATCC 27853, and Staphylococcus aureus subsp. aureus ATCC 25923, respectively.(a)(b)

3.5. Antibiotic Susceptibility Test

The susceptibility of L. intestinalis Y10 to 24 antibiotics was assessed, and the results are presented in Table 1 and Figure S1. L. intestinalis Y10 exhibited higher susceptibility to antibiotics targeting cell wall synthesis, including penicillins, cephalosporins, and bacitracin. Conversely, it displayed resistance to antibiotics that inhibit nucleic acid synthesis and cytoplasmic membrane function, such as diaminopyrimidines, fluoroquinolones, and polymyxins. These findings indicate the specific antibiotic sensitivities of L. intestinalis Y10, highlighting its potential utility and limitations in therapeutic applications.

3.6. Antimicrobial Activity, MIC, and MBC

In the agar well diffusion test, it was observed that neutral CFCS did not exhibit any inhibition zone. This suggests that the antibacterial activity displayed by L. intestinalis Y10 is primarily attributed to its organic acid production. The MIC and MBC values of the original CFCS against the three tested pathogens are presented in Table 2 and expressed as dilutions. Among the pathogens, the CFCS of L. intestinalis Y10 demonstrated the strongest antibacterial activity against P. aeruginosa ATCC 27853, while exhibiting the weakest antibacterial activity against S. aureus ATCC 25923.

3.7. Biofilm Inhibitory Test

As depicted in Figure 5, the inhibitory effects of L. intestinalis Y10 on biofilm formation were found to be concentration‐dependent. Among the tested pathogens, L. intestinalis Y10 exhibited the strongest inhibitory effect on the biofilm formation of S. aureus, with an inhibition rate as high as 99.48% ± 0.68%. This was followed by P. aeruginosa (97.51% ± 0.69%) and E. coli (96.83% ± 0.65%). The MBIC_90_ of L. intestinalis Y10 corresponded to its MIC for the respective pathogens. Specifically, for E. coli and S. aureus, the MBIC_90_ was achieved with a fourfold diluted CFCS of L. intestinalis Y10. Meanwhile, for P. aeruginosa, the MBIC_90_ was obtained with an eightfold diluted CFCS. In the experiment, only the MBIC_50_ for E. coli was determined, which required an eightfold diluted CFCS of L. intestinalis Y10.

Biofilm inhibition effects of L. intestinalis Y10 CFCS. The continuously twofold diluted CFCS was cocultured with Escherichia coli ATCC 25922, Pseudomonas aeruginosa ATCC 27853, and Staphylococcus aureus subsp. aureus ATCC 25923, respectively, for 24 h.

4. Discussion

Both captive and wild mice exhibited a relatively high abundance of Lactobacillaceae within their gut microbial communities, accompanied by pronounced seasonal dynamics [38]. Given the documented multifunctional health‐promoting effects of lactic acid bacteria, the exploration of wild rodent–derived Lactobacillus species holds significance due to their potential evolutionary adaptations to complex ecological niches, which may confer unique probiotic properties compared to domesticated counterparts. Previous studies on wild mouse gut microbiota have demonstrated that environmentally adapted gut microbiota from wild mice often exhibit enhanced functional traits, such as anti‐inflammatory effects and pathogen inhibition [12, 13]. Building upon these findings, this study aimed to systematically evaluate the probiotic potential of Lactobacillus strains isolated from wild brown rats (Rattus norvegicus). Through systematic screening and characterization, 12 strains were successfully isolated and taxonomically identified. Among these, L. intestinalis Y10 was prioritized for subsequent in‐depth experimental investigations due to its pronounced antimicrobial activity against pathogens and its status as one of the least‐studied candidates in the identified collection.

Before intestinal colonization, probiotic microorganisms must demonstrate resistance to gastric acidity and intestinal bile salts during transit through the gastrointestinal tract. Acid and bile salt tolerance represents a fundamental prerequisite for probiotic functionality, requiring maintenance of viability for 3 hours under simulated gastric conditions and three to 8 hours at physiological bile salt concentrations [39]. Assessment revealed that L. intestinalis Y10 exhibited moderate resistance to acidic environments, achieving 72.4% survival after 2 h at pH 2.5, though viability sharply declined beyond 4 h under acidic stress. Compared to the laboratory‐adapted L. intestinalis strain which declined from 10^6^·^72^ CFU/mL to 10^6^·^13^ CFU/mL, this strain demonstrated superior acid tolerance after 6 h of exposure to acidic conditions, declining from 10^6^·^63^ CFU/mL to 10^6^·^41^ CFU/mL [10]. Under bile salt stress (0.3% w/v), strain Y10 exhibited growth within 4 h and maintained viability for 24 h without proliferation, contrasting with domesticated strains showing active growth under similar conditions [10]. Results suggested that both acidic environment and bile salt exert inhibitory effect on the growth of strain Y10. This limitation can be mitigated by employing strategies, such as the addition of protective compounds (e.g., amino acids, trehalose, and inulin) or encapsulation, which significantly improve probiotic resilience against gastrointestinal stressors, thereby improving its practical application [40–44].

The adhesion properties of probiotics, characterized by hydrophobicity and auto‐aggregation, are critical for their initial contact with mucosal or epithelial cells [45, 46]. Hydrophobicity mediates initial attachment via nonspecific interactions with mucosal surfaces, facilitating subsequent specific binding mediated by specific cell wall components [47, 48]. In this study, L. intestinalis Y10 exhibited significantly higher cell surface hydrophobicity (93.80% ± 0.60% in toluene) than laboratory‐adapted strains (44.74 ± 2.6%), consistent with established strain‐specific variation in this trait [10, 45]. The hydrophobicity in toluene of strain Y10 surpassed values reported for other Lactobacillus isolates which range from 0% to 83% [49, 50]. Furthermore, hydrophobicity has been established as one of the species‐specific influencing factors of auto‐aggregation capacity [46, 51]. After 24‐h incubation at 37°C, the wild‐derived strain Y10 in this study demonstrated superior auto‐aggregation (81.68%) compared to both laboratory‐adapted L. intestinalis strain (< 30%) and other lactobacilli (29.32%–59.51%) [10, 52]. These findings indicate that the wild‐derived strain of L. intestinalis exhibits enhanced adhesion properties, potentially extending gut residency time while inhibiting pathogens and protecting intestinal epithelium [45]. These properties may confer a competitive advantage for its successful gastrointestinal colonization and persistence.

The strong adhesion capacity of probiotics enables competitive occupation of host cell surface receptors and promotes co‐aggregation with pathogens, thereby inhibiting biofilm formation [48]. Co‐aggregation establishes a protective barrier that impedes pathogenic colonization and facilitates the proximity of antimicrobial substances produced by probiotic strains to pathogens, enhancing pathogen suppression in the gastrointestinal tract [53, 54]. In this study, L. intestinalis Y10 displayed notably high co‐aggregation with S. aureus (73.53%), followed by P. aeruginosa (64.25%) and E. coli (64.23%) after 24‐h incubation, exceeding values reported for laboratory‐derived strains [10]. Various factors can influence the co‐aggregation property, including the auto‐aggregation ability between Lactobacillus and pathogen strains [55], the incubation time [56], the source of pathogenic strains [57], and the origin of probiotic strains [47]. The observed disparities likely arise from divergent L. intestinalis strain origins and inherent auto‐aggregation properties. The antipathogen activity of probiotics is multifactorial and also involves the production of antimicrobial compounds. Based on the MIC and MBC, the CFCS of L. intestinalis Y10 exhibited potent inhibitory and bactericidal activity against P. aeruginosa, followed by E. coli and S. aureus. However, the neutral CFCS did not exhibit any inhibition zone toward the three pathogens, suggesting that the antibacterial activity of L. intestinalis Y10 was primarily due to organic acids. Organic acids, such as lactic acid and acetic acid, are recognized as key antibacterial substances produced by Lactobacillus [3]. These organic acids lower both the extracellular and intracellular pH, destabilizing bacterial cell membranes and thereby inhibiting bacterial growth [3, 58]. In addition, Gram‐negative bacteria are more susceptible to the pH‐dependent antibacterial effect compared to Gram‐positive bacteria and yeasts [59], consistent with the findings in this study. The pH values of the L. intestinalis Y10 CFCS were 4.48, 5.16, and 3.78 at the concentrations corresponding to the MBC of E. coli, P. aeruginosa, and S. aureus, respectively, aligning with their growth‐permissible pH ranges. These results further corroborate the antibacterial effect of organic acids. Furthermore, the MBIC_90_ values of the L. intestinalis Y10 CFCS against three pathogens were similar to their MIC values, indicating that the antibiofilm mechanism may also be attributed to the action of organic acids. The equivalence between the MBIC_90_ and MIC further suggests that organic acids can effectively kill most bacteria, thereby reducing the production of biofilms from viable cells [60].

Probiotic safety assessments necessitate rigorous antimicrobial resistance screening. This process identifies strains harboring transferable resistance determinants that could disseminate to pathogens, thereby mitigating the emergence of resistant strains [61]. In this study, L. intestinalis Y10 exhibited susceptibility to antibiotics that target cell wall synthesis, including penicillins, cephalosporins, and bacitracin, but showed resistance to antibiotics that inhibit nucleic acid synthesis and cytoplasmic membrane function, such as diaminopyrimidines, fluoroquinolones, and polymyxins. This profile diverges from laboratory‐rat‐derived strains [10], but aligns with documented intrinsic resistance in most Lactobacillus species to aminoglycosides (including gentamicin, kanamycin, streptomycin, and neomycin), and many nucleic acid inhibitors [61–64]. Critically, the intrinsic resistance of lactobacilli poses minimal horizontal transfer risk according to the European Food Safety Authority (EFSA) [65]. Consequently, the resistance pattern of L. intestinalis Y10 may theoretically support co‐administration with fluoroquinolones or diaminopyrimidines in targeted anti‐infective regimens to reduce collateral damage to commensal microbiota. Human studies have shown that the concomitant administration of probiotics with antibiotics reduces the risk of antibiotic‐associated diarrhea (AAD) in adults [66]. However, comprehensive safety validation for new strains, including transfer risk assessment and therapeutic efficacy evaluation, remains essential before clinical implementation.

This study investigated the probiotic potential of L. intestinalis Y10 through comprehensive in vitro evaluations, yet several methodological limitations merit acknowledgment. Firstly, this study exclusively evaluated a single bacterial strain, with resultant data serving as a strain‐specific reference. Such single‐strain analysis provides limited generalizability for comparative assessment of Lactobacillus functional attributes between wild and laboratory rats. Secondly, the experimental design omitted in vivo assessments; consequently, the physiological efficacy of this wild‐type strain in animal models remains experimentally undefined. Finally, although established protocols were rigorously followed, quantitative discrepancies may arise from reagent source variations, dosage inconsistencies, and technical execution heterogeneities. For instance, aggregation measurements demonstrate sensitivity to multiple confounding variables, including culture media composition, temporal assessment windows, incubation temperatures, and pre‐analytical bacterial treatments (e.g., thermal, sonication, or enzymatic interventions) [67]. Consequently, integrative genomic characterization would significantly enhance phenotypic interpretation accuracy. The acid tolerance phenotype correlates mechanistically with amino acid decarboxylase and F1‐F0‐ATPase gene expression profiles, while comparative genomics has identified 112 bile salt tolerance–associated genes in lactobacilli, with malate dehydrogenase (sfcA) validated as a molecular biomarker via functional overexpression [68–70]. Aggregation mechanisms involve greater complexity, with experimental evidence identifying key proteins and their genes, such as S‐layer proteins, adhesins, exopolysaccharides, and aggregation‐promoting factors [67]. Additionally, specific gene determinants of antibiotic resistance have been experimentally documented, including blaZ for β‐lactams, Cat for chloramphenicol, mef(A) for macrolide, and aph(3′)‐III for aminoglycoside. [61].

5. Conclusion

In conclusion, this study characterized a strain of L. intestinalis Y10 isolated from the fecal microbiota of wild brown rats (Rattus norvegicus), demonstrating significant probiotic potential through comprehensive in vitro evaluations. The strain showed resistance to acid and bile salt, exhibited high adhesion property based on hydrophobicity and auto‐aggregation tests, and displayed antipathogenic effects through co‐aggregation, as well as the determination of MIC and MBC of its CFCS. It also demonstrated the capability to inhibit the formation of biofilms by pathogens. Moreover, antibiotic susceptibility profiling revealed sensitivity to antibiotics that target cell wall synthesis, such as penicillins, cephalosporins, and bacitracin. These findings collectively suggest that L. intestinalis Y10 possesses multifunctional probiotic properties and has great potential to treat gastrointestinal disorders. However, further in vivo studies utilizing animal models and molecular investigations are warranted to elucidate the precise mechanisms underlying its probiotic functions and to validate its therapeutic potential for clinical applications in gastrointestinal disorders.

Author Contributions

Sijia Yu: investigation, formal analysis, data curation, and writing–original draft. Peihang Hong: formal analysis, data curation, and software. Chao‐Min Wang: conceptualization, methodology, visualization, and writing–review and editing. Shyun Chou: software, formal analysis, and writing–original draft. Cheng‐Hung Lai: conceptualization, visualization, supervision, and writing–review and editing.

Funding

This research did not receive any external funding. This study was conducted with the support of surplus funds from previous industry‐academia collaboration projects (Project No.: 111D511 and 111D512).

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting Information

Supporting Figure S1. Antibiotic susceptibility test results of Lactobacillus intestinalis Y10. The plates were incubated anaerobically at 37°C for 24 h.

Supporting information

Supporting Information Additional supporting information can be found online in the Supporting Information section.