Lactiplantibacillus plantarum and Pediococcus acidilactici strains isolated from cattle gut show probiotic potential

Abstract

Background

Lactic acid bacteria (LAB), particularly Lactiplantibacillus plantarum and Pediococcus acidilactici, are well-characterized probiotics, known for their beneficial role in promoting gut health in animals.

Results

In this study, 48 presumptive LAB were obtained from the cattle gastrointestinal tract (feces and saliva). Among these, two strains (CS-23 and BC-14) exhibited significant probiotic properties, including acid tolerance (67.67–96.33%), bile salt tolerance (49.33–83.60%) auto-aggregation (74.24–76.21%), co-aggregation (10.35–23.93%), cell surface hydrophobicity (56.10–69.35%), antioxidant (DPPH scavenging: 27.15 and 34.66%), and antimicrobial activity (inhibition zone: 8.3–14.0 mm). Both strains were susceptible to key antibiotic classes (macrolides and beta-lactams) and exhibited neither gelatinase nor hemolytic activities, indicating their safety for use. Based on 16 S rRNA gene sequence analysis, the strains were identified as Lactiplantibacillus plantarum CS-23 and Pediococcus acidilactici BC-14, exhibiting close phylogenetic relatedness to indigenous probiotic strains of animal origin.

Conclusion

The indigenous strains Lactiplantibacillus plantarum CS-23 and Pediococcus acidilactici BC-14 exhibit promising probiotic potential and could be used as effective candidates for improving livestock health.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12866-026-05266-3.

Introduction

The gastrointestinal tract (GIT) of cattle harbors a diverse and balanced microbiota that maintains gut balance. Certain factors like improper diet, environmental changes, and farming conditions can disrupt the gut microbiota, which leads to the potential risk of infections [1]. The imbalanced gut microbiota favors the growth of pathogens like Salmonella, Staphylococcus, E. coli, Clostridium, etc., leading to severe disease outbreaks. It is the major cause of animal mortality and economic losses [2]. Numerous antibiotics are used to treat animal infections, but their irrational use has led to the destruction of intestinal microbiota and the emergence of antibiotic-resistant microbes [3]. Antibiotic replacement with other biocontrol agents seems to be the general trend in livestock [4]. Probiotics are a suitable alternative to replace antibiotics and improve animal health [5]. Probiotics can be considered part of the native microbiome, which is involved in regulating gut homeostasis [6]. Probiotic microorganisms constitute a highly complex biological ecosystem, which has a symbiotic association with their host and are extensively used in several industries, including animal nutrition and medicine. A probiotic strain must possess certain traits, especially the ability to tolerate gastrointestinal tract (GIT) conditions, adherence to the mucosal cell surface, and self-compatibility [7]. Likewise, they must meet the safety criteria, i.e., antibiotic susceptibility and inability to degrade blood cells, DNA, etc. [8]. Lactic acid bacteria (LAB), particularly members of Lactiplantibacillus plantarum and Pediococcus acidilactici are Gram-positive, rod-shaped, beneficial microbes that are generally recognized as safe (GRAS). These bacteria mainly reside in the gastrointestinal tract of ruminants, where they contribute to gut homeostasis, enhance nutrient absorption, regulate immune responses, produce antimicrobial metabolites, and help reduce serum glucose and cholesterol [9]. Numerous commercial probiotic strains have been reported and utilized to date. Despite the commercial availability of various probiotic strains, the isolation and characterization of native LAB from the cattle GIT remain underexplored [10]. The increasing demand for probiotics and their prophylactic and therapeutic attributes constantly encourages researchers to screen the indigenous strains as they are more effective and adaptive to their native host [11]. Thus, screening of indigenous potential probiotic strains from the animal’s gut is highly desirable to improve livestock [12]. This current study aims to isolate and characterize indigenous LAB strains from cattle’s gut, evaluate their probiotic potential and safety attributes, and identify them using 16 S rRNA gene analysis.

Materials and methods

Sample collection

This study was approved by the Ethical Review Board of COMSATS University (CUI), Islamabad, under reference # CUI/Bio/ERB/11-2023/3. Fecal and saliva samples were collected aseptically from the dairy farms of Punjab, Pakistan, including Chakwal, Chiniot, Gujarat, Gujranwala, Jhang, Jhelum, and Rawalpindi. Details of different farms are provided in Table S1. Fecal samples were collected directly from rectum with sterile gloves, transferred into the individual sterile conical tubes (50 mL), which were sealed to avoid cross-contamination among samples [13]. Saliva samples were collected from the mouth using a sterile swab [14]. Each sample was preserved in 20% glycerol, labeled, and stored in dry ice bags.

Isolation of bacteria

The samples (fecal = 1 g and saliva = 1 ml) were homogenized in sterile saline (0.85%), diluted (10^− 1 to 10^− 9), serially and plated onto De Man-Rogosa-Sharpe (MRS) agar [13, 14]. The agar plates were incubated at 37 ± 2℃ for 48 h. After incubation, the appearing bacterial colonies were marked based on variable colony morphology and counted. Distinct colony types were sub-cultured under the same conditions (MRS, 37 ± 2℃, 48 h) to purify them. The pure colonies were preserved in 20% (v/v) glycerol prepared in sterile distilled water at -80℃ for further analysis. The colony-forming units per gram (CFU/g or ml) were computed using the formula 1 [15].

1

Presumptive identification and probiotic characterization

The LAB isolates were presumptively identified phenotypically (colony morphology, Gram reaction) and biochemically (catalase activity) [10, 16]. All the isolates were characterized for different probiotic traits, including acid and bile tolerance, autoaggregation, co-aggregation, cell surface hydrophobicity, antioxidant potential, and antimicrobial activity, as described below.

Acid tolerance assay

The acid tolerance ability of LAB isolates was tested by following the method [10, 17]. Briefly, the isolates were cultivated in MRS broth at 37 ± 2℃ for 16–18 h. The bacterial cells were harvested by centrifuging at 10,000 xg for 10 min, washed twice with phosphate-buffered saline (PBS), and suspended in MRS broth at their respective pH (2, 3, 6.5). The cell suspension in MRS broth at pH 6.5 was used as the control. The cell concentration in each treatment was normalized based on their absorbance at 600 nm and incubated at 37℃ for 3 h. The viable cells in each treatment were counted using the serial dilution method and expressed as log CFU/mL. The acid tolerance ability of each isolate was evaluated based on its survival rate using the formula 2 at different pH levels.

2

Where: control (pH 6.5); treatment (pH 2, 3).

Bile salt tolerance assay

The bile salt tolerance of LAB was assessed as described in previous studies [10, 16] with some modifications. The isolates were cultivated in MRS broth ammended with different concentrations of bile salt (0.3% and 0.5%) at 37 ± 2℃ for 16-18 h. The LAB isolate grown in MRS broth without bile salt was used as a control. The growth was observed by measuring the optical density (OD 600). The tolerance ability of each isolate was calculated as a survival rate using the formula (2) as described in Sect. (acid tolerance assay).

Auto-aggregation assay

The auto-aggregation ability of the LAB isolates was tested as previously described [10, 18]. Briefly, isolates were grown on MRS broth at 37℃ for 18 h. The culture was harvested by centrifuging at 18,000 xg for 10 min. The cell pellet was washed (2x) with PBS and redissolved in the same buffer to attain a final concentration of 10⁸ CFU/mL. The cell suspension was incubated at 37℃ for 4 h. Phosphate-buffered saline without treatment was used as a control. The OD was measured at 600 nm at the beginning (t = 0 h) and after the incubation (t = 4 h). The auto-aggregation percent (%) was calculated using the formula 3.

3

Where A _t = Final absorbance (after 4 h) ; A₀ = Initial absorbance (at 0th h) t.

Co- aggregation assay

The co-aggregation ability of LAB isolates with two pathogenic bacterial strains, i.e., Staphylococcus aureus (ATCC25923) and Salmonella typhimurium (ATCC14028), was assessed following the method in previous studies [10, 18]. The cell suspension of LAB and pathogenic strains was prepared as described in the previous section (auto-aggregation). The LAB and pathogenic isolate were mixed in equal volume. The individual strains of LAB and pathogenic bacteria were utilised as control. After 5 h of incubation at 37℃, the OD was measured at 600 nm. The co-aggregation percent (%) was determined using the following equation (4).

4

Whereas A_{LAB =} Absorbance of LAB; A_PAT = Absorbance of pathogenic isolates, and A_MIX = Absorbance of the LAB and pathogenic bacteria mixed in equal ratio.

Cell surface hydrophobicity (CSH) assay

The cell surface hydrophobicity of LAB isolates was determined according to the method [19]. Briefly, LAB isolates were cultured in MRS broth at 37℃ for 18 h. The cell pellet was obtained by centrifugation at 12,000 rpm for 10 min and washed twice with PBS. The pellet was resuspended in the same buffer, and OD of cells (x) was measured at 600 nm. This OD was taken as a control Suspension of each cell was mixed with respective solvents (n-hexane / xylene). To separate the two phases, the suspension was vortexed and allowed to stand for 1 min at 37℃. The OD of aqueous phase (y) was measured at 600 nm. Cell surface hydrophobicity (%) was calculated using the equation (5).

5

Antioxidant activity

The antioxidant activity of LAB isolates was evaluated based on a 2,2-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging assay [10, 20]. Briefly, a culture of each strain (1 mL) was mixed with 2.0 mL of DPPH radical solution and incubated in the dark at 37℃ for 30 min. The mixture was then centrifuged at 10000 xg for 10 min to separate cells. The absorbance of the supernatant was measured at 517 nm. The DPPH radical scavenging ability was calculated by using the equation (6).

6

Antimicrobial activity

The pathogenic bacteria Staphylococcus aureus (ATCC25923), and Salmonella typhimurium (ATCC14028) were obtained from the National Institute of Health (NIH) in Islamabad, Pakistan. The antimicrobial activity of LAB isolates was evaluated using the agar spot method as described by Schillinger and Cke [21]. Briefly, the LAB isolates were cultured in MRS broth at 37℃, spotted (5 uL) on the MRS agar and incubated at 37℃ for 24 h. The pathogens were cultured in LB broth (1 × 10⁶ cells/mL), overlayed onto the previously spotted MRS agar plates and incubated at 37℃ for 24 h. The diameter of the inhibition zone was measured in millimeters (mm).

Scoring of potential probiotics

The potential LAB isolates were selected based on their probiotic traits using a standardized scoring system as described previously [22] with certain modifications. Each trait was scored on a 0–5 scale based on its activity (Table S2). Final score of each isolate was determined by taking the average of scores of all traits. The isolates with the highest score were selected for further studies.

Self-compatibility assay

The self-compatibility of potential probiotic strains was detected using an antibacterial assay. Each strain was used as an indicator as well as the test strain. The strains with no antimicrobial activity against each other were considered compatible.

Antibiotic susceptibility assay

The probiotic strains were evaluated for their susceptibility or resistance using the disk diffusion method [10, 16]. The antibiotics tested were selected based on the Enterococcus spp. criteria provided by the Clinical Laboratory Standard Institute (CLSI) [23]. Purified LAB strains were lawned onto MRS agar and allowed to dry for 2–3 min. Antibiotic discs were placed on each plate inoculated with LAB and incubated at 30 ± 2℃ for 18-24 h. The zone of inhibition was recorded and interpreted based on CLSI and EUCAST criteria to designate strains as susceptible and resistant [24]. A plate without an antibiotic disc was used as a control. The multiple antimicrobial resistance (MAR) index of each strain was calculated using the equation (7).

7

Hemolytic activity and gelatinase activity

For hemolytic activity, the strains were inoculated on MRS agar ammended with 5% blood and incubated at 37℃ for 48-72 h [25]. Staphylococcus aureus (ATCC14028) was used as a positive control [26]. The appearance of a discolouration zone (colourless to greenish) around each colony. Gelatinase activity was evaluated as described earlier [27] with minor modifications. Briefly, LAB isolates were grown on MRS agar containing gelatin (3% w/v) and incubated at 37℃ for 48-72 h. The appearance of clear zone around each colony indicated gelitinaseactivity.

Molecular identification of potential probiotics

The potential probiotic strains were identified through 16 S rRNA gene analysis. The genomic DNA was extracted using the CTAB method, including a blank extraction control [28]. The 16 S rRNA gene was amplified using universal primers (P1 and P6) [10, 29]. The PCR reactions were performed with the LAB strains and a negative control (non-template control). The PCR reaction mixture consisted of DNA (10 ng), MgCl₂ (10 mM), dNTPs (0.5–1 mM), primers (10 mM of each), Taq polymerase (1.5 U), and PCR buffer (1X). The cycling condition consisted of initial denaturation at 95℃ for 5 min; 25 cycles of denaturation at 95℃ for 1 min, annealing (1 min) at 58℃, and extension for 1 min at 72℃, followed by a single cycle of final extension for 10 min at 72℃. The PCR product was electrophoresed and visualized under a gel documentation system (Biometra, Germany). A DNA ladder (1Kb, Thermo Scientific) was utilised to compare the size of the gene. Amplicons were purified by using a PCR purification kit (Thermo) and sequenced by Macrogen Inc., Korea. Raw sequences were trimmed, annotated, and analyzed for homology at EZ taxon database. Each 16 S rRNA gene sequence was submitted to the gene bank. The 16 S rRNA gene-based phylogenetic analysis was done by assessing the sequences of respective type strains from the EZ taxon. These sequences were aligned, trimmed, and converted to MEGA X format using MEGA X software. The phylogenetic tree was constructed using the Neighbor Joining Method. A bootstrap test was done with 1000 replicates.

Statistical analysis

All experiments were conducted using a completely randomized design (CRD) with n = 3 biological replicates, pretreatment. Statistics 8.0 was used to analyze the data. Data normality was checked, confirming that most groups followed normal distribution. One-way analysis of variance (ANOVA) was used to determine significant differences among isolates (p ≤ 0.05). Fisher’s least significant difference (LSD) test was used for pairwise comparisons at a significance level of p ≤ 0.05 [18]. Pearson correlation and Principal component analysis (PCA) were constructed by using Origin 2022.

Results and discussion

Isolation of presumptive lactic acid bacteria (LAB)

A total of 48 presumptive LAB isolates were obtained from the gut (fecal and saliva/ruminal) of cows and buffaloes (Table S3). Out of 48, 21 isolates were obtained from cow feces, 8 from cow’s saliva, and 19 from buffalo´s feces. Prevelance of variable isolates in the cattle´ s gut had been reported earlier [30, 31]. Our results depicting variability of LAB isolates among sample types and host spp. are align with findings in various ecological habitats. For example, a recent study on artisanal Turkish cheeses reported 500 LAB isolates from 16 genera and 30 spp., showing that LAB diversity is mainly influenced by nutrient availability, environmental conditions and ecological niche [32]. Such distribution also highlights how host physiology, habitat, and diet affect the gut microbial populations. Such distribution reflects the presence of diverse microbes in various host spp., and highlights the influence of host physiology, diet, and habitat on gut microbial population. The population of isolates was variable, ranging from 3.4 to 7.2 log CFU/g or mL as shown in Table S3. The isolate CF-13 showed the highest number of cells (7.2 log CFU/g) while BC-9 exhibited the least number of cells (3.4 log CFU/g). Our findings align with previous studies [33], in which the commercial probiotic Lactobacillus rhamnosus GG exhibited a comparable cell count. Similar variability has been reported in pigs and wild boars, where LAB counts ranged from 4.46 to 7.39 CFU/mg [34]. The variability in cell population of LAB isolates usually depends on different factors like the host (animal) type/ habitat, feeding regime and environment [35]. These findings highlight the significance of host-associated factors in maintaining the probiotic LAB reservoir.

Morphological and biochemical traits of presumptive lactic acid bacteria

The LAB isolates in this study exhibited diverse colony morphologies ranging from white, milky white, creamy white to yellowish white. The colony shape of all bacterial isolates was round/circular except for four strains (BC-1, BC-2, BC-13, and BC-17), which showed irregular shapes. Similarly, the size of all bacterial colonies was different, i.e., small (1–2 mm), moderate (3–4 mm), and large (> 5 mm), as shown in Table S3. All the presumptive LAB isolates were Gram-positive and catalase-negative. The presumptive identification based on these traits had been reported in earlier studies [36]. Similar observation has been reported in recent studies, where LAB typically appear as Gram-positive, catalase–negative and varying colony colors and textures based on the isolation source and medium [37, 32]. The overall pattern of LAB isolates presumptive features is the same, although the isolates of the current study showed variation among colony colors and sizes. These morphological variations among LAB isolates may reflect strain-level differences influenced by the nature of the host, sample type, and environmental adaptations [7]. Morphological variation has been reported in LAB isolated from different ecological habitat, such as gastrointestinal samples, fermented food and animal sources [38]. Such findings highlight that, although colony-level variations exist, the core features remain same for presumptive identification. Moreover, the similarity in Gram–staining and catalase reaction further supports the precise presumptive identification of LAB.

Probiotics attributes of lactic acid bacteria (LAB)

Acid tolerance

Acid tolerance determines the ability of strain to adopt the gastrointestinal tract. The tested isolates tolerated the acidic conditions (pH 2 and 3) with a survival rate of 30–97.63% (Table S4). At pH 2, 52% of LAB isolates maintained viability compared to 85% at pH 3. The highest acid tolerance at pH 2 (survival rate ≥ 83%) was shown by eight isolates, followed by twenty-six isolates at pH 3. The potential strain BC-14 showed maximum tolerance at pH 3 (96.33%), and isolates CS-29 exhibited the highest tolerance (72.67%) at pH 2. The acid tolerance ability of potential LAB isolates is listed in Table 1. Similar findings were reported by earlier studies [37, 39] where LAB typically exhibits maximum viability at pH 3 compared to pH 2. Our findings are further strengthened by other studies [10, 40, 41], in which probiotic strain Pediococcus pentosaceus E3 and the widely used commercial strain Lactobacillus rhamnosus GG exhibited survival rates of approximately ≤ 58% at pH 2.5. In comparison, our potential strains showed better survival at both pH 2 and pH 3, indicating greater strength under acidic conditions. The variation in acid tolerance at different pH levels is dependent on the strain type, habitat, and host [19]. Mechanistically, the decreased survival at pH 2 may be due to inhibition of the F₀F₁-ATPase pump [42]. Additional mechanism including membrane fatty acid modifications, upregulation of acid-shock and stress response proteins, have been reported to contribute to LAB acid resistance [43]. These findings depict that LAB isolates possess a promising acid tolerance capability, which helps them to survive in harsh GIT conditions to perform their function effectively.

Table 1.

Acid and bile salt tolerance of potential lactic acid bacteria (LAB)

Isolates	Acid tolerance		Bile salt tolerance
	pH 3	pH2	0.3%	0.5%
CS-23	93.33 ± 0.88b	67.67 ± 1.20c	83.60 ± 0.88b	58.57 ± 0.29b
CS-25	72.40 ± 0.38e	-	89.33 ± 1.76a	74.00 ± 1.53a
CS-29	90.24 ± 0.69c	86.96 ± 0.77a	66.00 ± 0.58c	56.30 ± 1.45b
BC-5	58.33 ± 0.33f	-	61.00 ± 1.73d	15.00 ± 1.15d
BC-14	96.33 ± 0.88a	72.67 ± 1.45b	83.00 ± 0.80b	49.33 ± 0.73c
BC-15	78.30 ± 0.35d	67.67 ± 1.20c	83.00 ± 1.16b	48.33 ± 1.20c

The values are mean of three replications and values within the same column bearing different superscript letters (a-f) are significantly different at p ≤ 0.05 according to Fisher’s Least Significant Difference (LSD); ± indicates standards errors, (-) indicates no activity

Bile salt tolerance

The ability of LAB isolates to survive and grow in high concentrations of bile salt certifies their adherence to the epithelium of gut. LAB isolates exhibited resistance against various bile salt concentrations, i.e., 0. 3% and 0.5% with a survival rate of 1.5% to 89.3% (Table S4). All LAB isolates tolerated 0.3% bile salt, whereas 81% exhibited tolerance at 0.5%. At 0.3% bile, most isolates showed moderate to high tolerance, indicating a strong capability to withstand physiological bile levels. However, a notable reduction in viability was observed at 0.5%, where only one isolate showed maximum tolerance (survival rate ≥ 74%), and 34 isolates exhibited the least survival (survival rate ≤ 25). The potential strain CS-25 exhibited maximum bile salt tolerance at both 0.3% and 0.5% with a high survival (89% and 74%) as shown in (Table 1). Similar findings were reported in earlier studies [6], where most LAB isolates showed good resistance at 0.3% bile salt compared to 0.5%. Likewise, both Pediococcus pentosaceus E3 and L. rhamnosus GG exhibit bile tolerance of ≤ 66% at 0.3% [40, 41], which is comparatively lower than that of most strains in our study, emphasizing the effectiveness of the potential strain to withstand gut bile tolerance. Bile salt hydrolase activity (BSH) is a well-known mechanism used by LAB isolates to resist bile salt toxicity through the deconjugation of bile salts. The difference in the trend of bile salt tolerance ability may also be due to different expressions of bile resistance-linked proteins, nature/ type of probiotic strains, and geographical distribution of the host [19]. Recent studies further suggest that deregulation of various efflux transporters and stree response genes contributes to LAB survival under bile stress [44]. Isolates depicting significant bile tolerance, mainly at 0.5%, represent potential candidates for further characterization.

Auto-aggregation ability

The LAB isolates exhibited variable auto-aggregation, i.e., 22.4–96.8% (Table S5). Six isolates showed maximum auto-aggregation ability (≥ 80%), 35 isolates showed moderate auto-aggregation (≥ 45% ≤ 79%), and 7 isolates showed the least auto-aggregation (≥ 22% ≤ 44%). Isolate BC-15 showed the highest auto-aggregation potential (94.75%) compared to other potential strains (Table 2). Our results were aligned with earlier studies demonstrating that many LAB strains showed strong autoaggregation ability [45]. Notably, our potential isolates showed even higher autoaggregation potential compared to widely used commercial probiotic L. rhamnosus GG (35%) and well-characterized P. acidilactici CLP03 (81.44%) [10, 46, 47]. High auto-aggregation capacity was correlated with strong adhesion. Previous studies assure that potential autoaggregation depends on multiple factors such as cell surface proteins, carbohydrates, lipoteichoic acid, and environment of intestine [48]. Current research highlights the role of surface layer (S-layer) components in strengthening cell to cell interactions and supporting probiotic fitness [49]. Therefore, LAB isolates with strong autoaggregation potential may thus possess better colonization ability in the host gut, making them a desirable candidate for probiotic applications.

Table 2.

Auto-aggregation, co-aggregation, cell surface hydrophobicity and 2,2 diphenyl-1-picrylhydrazyl (DPPH) scavenging ability of potential lactic acid bacterial isolates

No of isolates	Auto-aggregation percentage	Co-aggregation ability		Hydrophobicity				DPPH scavenging ability
		S. aureus	S. typhimurium		n-Hexane	Xylene
CS-23	76.21 ± 0.6c	23.93 ± 0.09a	10.35 ± 0.01c	56.10 ± 0.83b		68.20 ± 0.92b	34.66 ± 0.39d
CS-25	80.32 ± 0.57b	6.41 ± 0.05e	23.01 ± 0.04a	49.70 ± 0.89c		59.43 ± 0.81c	48.43 ± 0.90b
BC-5	47.90 ± 0.83d	10.75 ± 0.16c	2.15 ± 0.15e	68.38 ± 0.33a		75.43 ± 0.81a	73.36 ± 0.38a
BC-14	74.27 ± 0.68c	15.52 ± 0.09b	22.42 ± 0.08b	69.35 ± 0.60a		68.50 ± 0.91b	27.15 ± 0.81e
BC-15	94.75 ± 0.58a	7.90 ± 0.07d	7.41 ± 0.03d	68.90 ± 0.61a		53.84 ± 0.67d	41.23 ± 0.42c

The values are mean of three replications and values within the same column bearing different superscript letters (a-e) are significantly different at p ≤ 0.05 according to Fisher’s Least Significant Difference (LSD); ± indicates standards errors, (-) indicates no activity

Co-aggregation ability

The LAB isolates showed variable co-aggregation with S. aureus, ranging from 1.15% − 35.3% and 1.8% − 40.9% with S. typhimurium (Table S5). The highest co-aggregation was shown by potential isolate CS-23 with S. aureus (23.93%), while CS-29 with S. typhimurium (40.9%) (Table 2). Similar findings were reported in earlier studies [45, 50], where the strains showed higher co-aggregation with S. typhimurium than that of S. aureus. In another study [17], reported that LAB isolates showed strong co-aggregation ability with S. aureus as compared to that of S. typhimurium. Overall, our tested strains showed good co-aggregation potential, the same as commercial strain L. rhamnosus GG, which exhibited co-aggregation rates of 35% with S. typhimurium and 38% with S. aureus [46]. However, a slight increase was noted with P. acidilactici CS-29 with S. typhimurium. These differences may be due to the strain-specific trait and the surface proteins of cell i.,e exopolysaccrides, S-layer proteins and adhesins, which modulate LAB-pathogen interaction [51]. Such ability of LAB suggests a protective role in the host GIT tract by inhibiting pathogens. The tendency of LAB strains to co-aggregate with enteric pathogens provide a protective role in maintain gut health, through competitive exclusion of pathogen colonization. Isolates showing higher co-aggregation potential in this study could be served as candidates for further probiotic evaluation.

Cell surface hydrophobicity (CSH)

Another important trait of probiotics is hydrophobicity, which is the measure of the capability of a bacterial strain towards hydrophobic solvents. The cell surface hydrophobicity potential of LAB was variable with different solvents. In this study, LAB isolates exhibited a wide range of hydrophobicity, from 2.74% − 81.8% with n-hexane and 25.3% − 79.9% with xylene (Table S5). The potential isolate BC-14 showed the highest cell surface hydrophobicity (69.35%) with n-hexane, while BC-5 showed the highest hydrophobicity potential (75.43%) with xylene (Table 2). Our potential strains showed higher cell surface hydrophobicity compared to reference strain L. rhamnosus GG (55%) [46]. Our findings are strengthened by earlier studies [16, 17] in which LAB isolates showed more affinity for n-hexane as compared to xylene. In contrast, other researchers reported that LAB isolates showed more affinity towards xylene solvent as compared to n-hexane [52], indicating that CSH vary depending on the strain and experimental conditions. Such variability in CSH may depend upon numerous factors, i.e., the chemical composition of hydrophobic solvents, the intrinsic mechanism of LAB isolate, and bacterial cell surface proteins [53]. The LAB isolates with high hydrophobicity may possess significant mucosal adhesion capacity, making them promising probiotic candidates. Hydrophobicity is not only associated to adhesion but may also reflect adaptive mechanisms influencing interactions with epithelial cells and the gut environment [54]. Overall, the good hydrophobicity observed in our tested isolates suggest a potential for mucosal adhesion, meanwhile further in vitro and in vivo trails are essential to confirm this trait and its contribution to probiotic functionality.

Antioxidant activity

Antioxidant activity is an essential trait of probiotic LAB, as it helps to reduce oxidative stress, supporting intestinal barrier integrity, and enhancing overall host health. In this study, LAB isolates showed variable antioxidant activity based on their DPPH scavenging ability (16.9% to 73.4%) (Table S5). Isolate BC-5 exhibited the strongest scavenging ability (73.4%) compared to the others. The DPPH scavenging ability of potential LAB isolates is given in Table 2. According to previous reports [55], L. plantarum LP9010 demonstrated good DPPH scavenging potential (75%) like our potential strain BC-5. DPPH scavenging activity of probiotics helps them to decrease the oxidative stress, stabilize intestinal microbiota, and improve animal health [11]. Such variation in DPPH scavenging ability may be due to the nature of probiotic strains and the type of stress conditions. The observed diversity in DPPH scavenging activity among LAB isolates suggests that only a few potential strains harbor this functional attribute, which may enhance their ability to withstand oxidative stress in the host’s GIT. Antioxidant potential may be linked to production of various enzymes (glutathione peroxidase and superoxide dismutase), cell surface proteins and metabolites i.e., short chain fatty acids and exopolysaccharides [56]. Overall, the variation observed in antioxidant potential among our isolates indicates that only few potential strains harbor this functional attribute, which may enhance their ability to withstand oxidative stress in the host’s GIT. However, expression analysis and in vivo trails are essential to confirm these effects.

Antimicrobial activity of LAB isolates

Antimicrobial activity is a significant property of LAB that eradicates pathogenic microbes, a major cause of animal morbidity and mortality. About 73% of isolates exhibited antimicrobial activity against S. typhimurium, while 56% of isolates were effective against S. aureus (Table 3). Based on the zone of inhibition, 2 isolates exhibited maximum inhibition (17.7 mm) against S. aureus, followed by 3 isolates that showed the highest inhibition (15–18 mm) against S. typhimurium. Similarly, both the commercial probiotic L. ramhnosus GG and the well-characterized strain L. plantarum KLDS 1.0344 exhibited antimicrobial patterns comparable to those observed in our tested isolates [57]. Our findings are further supported by earlier studies [17], where LAB isolates showed higher inhibitory activity against S. typhimurium compared to S. aureus. The observed variations may be due to the intrinsic nature of probiotics, mainly their ability to produce various antimicrobial compounds, including bacteriocins, siderophores, organic acids, hydrogen peroxide, and antibiotics [31]. Current researchers have highlighted the strain specific production of bacteriocins and other antimicrobial metabolites play an important role in regulating competitive interactions with pathogens and maintaining gut microbial balance [58].

Table 3.

Antimicrobial activity of LAB isolates against. S. typhimurium and S. aureus

Isolates	ZOI against S. typhimurium	Isolates	ZOI against S. aureus
CF-1	8.30 ± 0.67lmn	CF-1	8.32 ± 0.33h
CF-3	12.23 ± 0.58efgh	CF-6	12.70 ± 0.33bcde
CF-4	10.00 ± 0.58ijkl	CF-8	13.32 ± 0.58bcd
CF-6	10.32 ± 0.88hijk	CF-9	12.00 ± 0.58bcdef
CF-8	13.00 ± 0.58cdef	CF-11	11.70 ± 0.88cdefgh
CF-9	13.70 ± 0.67cde	CF-12	11.00 ± 0.58efg
CF-10	13.70 ± 0.33cde	CF-14	11.10 ± 0.57efg
CF-11	12.00 ± 0.58efgh	CF-15	10.33 ± 0.88fg
CF-12	11.00 ± 0.58ghijk	CF-16	13.00 ± 0.58bcd
CF-13	11.30 ± 0.55ghijk	CF-21	12.00 ± 0.58bcdef
CF-14	8.30 ± 0.88lmn	CF-22	13.30 ± 0.88bc
CF-15	7.30 ± 0.67m	CS-23	12.14 ± 0.58bcdef
CF-16	10.3 ± 0.87hijk	CS-24	12.75 ± 0.88bcde
CF-17	7.20 ± 0.58no	CS-25	12.35 ± 0.88bcde
CF-18	5.30 ± 0.33o	CS-26	12.70 ± 0.33bcde
CF-19	12.30 ± 0.88defg	CS-27	13.10 ± 0.58bcd
CF-20	11.32 ± 0.88fghij	CS-28	13.70 ± 0.88b
CF-21	13.10 ± 0.57cdef	BC-1	8.23 ± 0.58gh
CF-22	9.72 ± 0.67jkl	BC-4	10.34 ± 0.88fg
CS-23	14.00 ± 0.57cd	BC-5	13.33 ± 0.58bc
CS-24	11.70 ± 0.33fghi	BC-6	11.70 ± 0.67cdefg
CS-25	16.10 ± 0.58ab	BC-8	11.31 ± 0.33defg
CS-26	13.70 ± 0.33cde	BC-9	17.70 ± 0.88a
CS-27	14.32 ± 0.67bc	BC-10	17.00 ± 0.58a
CS-28	11.00 ± 0.58ghijk	BC-13	13.20 ± 0.58bcd
BC-1	12.32 ± 0.58efgh	BC-14	11.30 ± 0.88defg
BC-4	14.75 ± 0.88bc	BC-15	8.30 ± 0.58h
BC-5	18.00 ± 0.88a
BC-6	14.00 ± 0.58cd
BC-8	7.70 ± 0.33mn
BC-9	14.20 ± 0.58cd
BC-14	8.30 ± 0.32lmn
BC-15	8.30 ± 0.29lmn
BC-18	9.30 ± 0.89lmn
BF-20	10.00 ± 0.33hijk

The values are mean of three replications and values bearing different superscript letters (a-o) in the same columns are significantly different from each other at p ≤ 0.05 according to fisher’s least significant test (LSD); ± indicates standards errors

These findings highlight the selective antimicrobial activity of LAB strains, which support their application as targeted probiotics or natural biocontrol agents in animal health management.

Selection of potential probiotics and their compatibility

Six LAB isolates, i.e., CS-23, CS-25, CS-29, BC-5, BC-14, and BC-15, were selected as potential probiotics based on their highest probiotic trait scoring approach. The score distribution of probiotic traits of each isolate is shown in Fig. 1, and their Pearson correlation is shown in Fig. 2. Probiotic isolates having good traits, i.e., host survival and pathogen inhibition, were assigned greater weight in the scoring system. All the potential probiotic isolates showed no antimicrobial activity against each other, which ensured their safety for the related class of bacteria as well as their suitability to be used as a consortium. The potential probiotic isolates mainly exhibited self-compatibility with each other. Similar findings were earlier reported [59], in which all LAB strains were compatible among each other and showed no inhibitory activity. However [60], stated that only the L. murinus strain was inhibited by L. mucosae, L. johnsonii, and L. slalivarius in the compatibility assay. This change in compatibility pattern mainly depends on the type/nature and different inhibitory substances produced by LAB isolates. Recent studies also depicted that inter-and intra-spp compatibility among LAB strain is significant for their synergetic function in multi strain formulations [61]. Moreover, compatibility supports the stability of probiotic consortia and also increase their efficacy in pathogen exclusion and gut colonization. Overall, the potential LAB isolates in this study demonstrated broad-spectrum probiotic attributes along with functional compatibility, and cross-interaction, making them effective candidates for further in vivo evaluation and formulation development in feed supplements.

Fig. 1.

Scores distribution of probiotic traits among LAB isolates. Each cell represent the score assigned to a specific trait for individual isolate. A = S. aureus, B = S. typhimurium, C = n-Hexane, D= xylene. The various colors in heatmap indicates trait intensity i.e., red= higher/stronger values, white= intermediate values, and blue= lower/weaker values

Fig. 2.

Pearson Correlation matrix among the different probiotic traits of lactic acid bacteria. (LAB). The values in the cell represents the Pearson correlation coefficient (r), at p ≤ 0.05. Positive correlations are shown in red, negative in blue, with color intensity proportional to the strength of the correlation. DPPH = 2, 2 diphenyl-1-picrylhydrazyl, A = S. aureus, B = S typhimurium, C = n-Hexane, D= xylene

Despite the comprehensive in vitro characterization of probiotic attributes, certain limitations must be acknowledged. The study lacks gene analysis of probiotic attributes (acid, bile, cell colonization, antioxidant, antimicrobial analysis), which are essential to understand the functional efficacy of a strain within the host’s GIT. The PCA analysis was done to find the relationship between probiotic traits and to identify the combinations that highlight the most favorable isolates. PC1 had a 43.5% variance and showed better acid tolerance, co-aggregation, hydrophobicity, and DPPH scavenging traits. These traits are responsible for gut survival and epithelial adhesion [16, 41]. PC2 showed a 28.9% variance and it showed better bile tolerance, autoaggregation, and antimicrobial activity, which are important for controlling pathogens from the intestinal tract. The distribution of LAB isolates along PC1 and PC2 dimensions shows the functional variation among strains (Fig. 3). The potential strain CS-23 and BC-15 group closely with PC2, representing better bile tolerance and colonization capability. While BC-14 and CS-29 group closely in PC1, indicating strong epithelial interaction and optimal oxidative stress resistance. However, BC-5 is associated with both PC1 and PC2, indicating strong antioxidant, hydrophobicity, and moderate antimicrobial activity, and CS-25 is only associated with high antibacterial activity. The PCA highlights attributes and functions that help distinguish strains with multi-functional properties from those with dominant traits [19]. Similar findings were reported in earlier studies [7, 33], where the multivariate analysis was conducted using PCA in order to select the best probiotic strain by taking all probiotic properties by strain clustering and identifying isolates with complementary probiotic functions. Recent studies highlight the importance of multivariate tools i.e., PCA, in probiotic evaluation, as they facilitate integration of multiple in vitro attributes for an unbiased and comprehensive strain selection [62].

Fig. 3.

Principal component analyses (PCA) biplot showing the distribution of potential LAB isolates based on probiotic traits. The first two principal component analysis i.e., PC1 and PC2 explains the total variance. Red dots indicate LAB isolate codes, while vectors represent probiotic traits. % = Percentage. A = S. aureus, B = S. typhimurium, C = n-Hexane, D= xylene

Phenotypic resistance and MAR indices

Antibiotic resistance is a major safety concern in newly discovered probiotic strains. In this study, six potential probiotic strains were tested for susceptibility/ resistance to 3 classes of antibiotics (n = 6) (Fig. 4A). Five isolates were susceptible (83%) to the macrolide class of antibiotics (erythromycin and rifamycin). In contrast, one isolate (CS-25) showed intermediate sensitivity (16.7%) (Fig. 4a-1). For beta-lactam antibiotics (ampicillin and penicillin), 2 isolates (33.3%) were sensitive, whereas 4 (66.7%) isolates were resistant. All LAB isolates (100%) were resistant to tetracycline and streptomycin, with varying MAR indices of 0.32 to 0.66 (Fig. 4a-2). Two isolates (CS-23 and BC-15) were susceptible to both macrolide and beta-lactam antibiotics, with the least MAR index (0.32), depicting reduced antibiotic resistance stress. Antibiotic resistance has been reported in numerous LAB strains and commercial formulations [59, 60, 63], where L. rhamnosus LGG showed susceptibility towards 41 antibiotics and resistance to 18 (beta-lactam and aminoglycoside antibiotics). It is an unlikely concern if there is intrinsic resistance that is non-transferable to other strains [64–66]. The variation of LAB in antibiotic sensitivity/ resistance pattern is a strain-dependent property, which may be due to their intrinsic nature to adapt to the particular habitat. Their genomic determinants and escape rate should be further studied to assess the risk based on antimicrobial resistance. Similarly, hemolytic and gelatinase activities are considered as virulence traits. In the current study, the potential probiotic isolates showed no hemolytic (Fig. 4B) and gelatinase activity. The absence of hemolytic and gelatinase activity is considered a prerequisite in the selection of potential probiotic strains [66–68]. The selected isolates fulfill most of the safety criteria and could be considered and utilized as potential probiotics for use in the dairy industry. However, for a more in-depth safety evaluation, genomic analysis of antibiotic resistance and virulence-associated factors is required.

Fig. 4.

Safety assessment of probiotic strains A= Phenotypic representation of antibiotic susceptibility/resistant, a-1 = Antimicrobial resistance profile, a-2 = Multiple antimicrobial resistance index, B = Hematolytic activity of probiotic strains. The double line around the colony indicates the halo zone (beta hemolysis), whereas the single line around colonies indicates no halo zone (no hemolysis)

Molecular identification of potential probiotics

Six potential LAB isolates with the highest probiotic traits (score = 2.58–3.17) were identified by 16 S rRNA gene analysis (Fig S1). The potential probiotic strains were identified as Lactiplantibacillus plantarum (CS-23 and BC-15) and Pediococcus acidilactici (CS-25, CS-29, BC-5, and BC-14). The 16 S rRNA nucleotide sequences have been submitted to GenBank under accession numbers OP723337, OP721097, OP721061, OP721065, OP720973, and OP721029. The 16 S rRNA gene sequencing is a universal and well-established genotypic approach for the identification of bacteria. The potential probiotics screened in this study were identified as P. acidilactici and L. plantarum, are generally recognized as safe, and they are being widely used as probiotics worldwide [9, 48, 67]. Furthermore, the indigenous strains of P. acidilactici and L. plantarum showed phylogenetic lineage with the strains of the same species, which have strong probiotic potential and animal origin. These strains were distantly related to the weak probiotics and pathogenic strains (Fig. 5). These findings were supported by the earlier studies, where the phylogenetic analysis depicted that LAB strains were closely related to the isolates obtained from the same ecological niche as compared to isolates obtained from other sources [69, 70, 71]. The molecular identification of these indigenous probiotic strains, P. acidilactici and L. plantarum, correlates with their safety attributes like antimicrobial drug resistance, virulence, and gelatinase activity [72]. Although 16 S rRNA sequencing is a reliable approach for bacterial identification at the species level, however, it may lack the accuracy needed for strain-level identification. For more accurate identification methods, like multiple locus sequence typing (MLST) and whole genome sequencing, are essential.

Fig. 5.

The phylogenetic tree of probiotics with reference strains based on 16 S rRNA gene. The evolutionary history was inferred using the Neighbor-Joining method. The percentage of replicate trees in which the associated taxa clustered together in the bootstrap test (1000 replicates) are shown next to the branches. The evolutionary distances were computed using the Maximum Composite Likelihood method and are in the units of the number of base substitutions per site. This analysis involved 37 nucleotide sequences. Evolutionary analyses were conducted in MEGA11. Scale bar represent 0.05 substitution per nucleotide position. Test LAB strains isolated from cattle feces and saliva are indicated in grey-colored branch lengths. Branches are color-coded based on various source type of each isolate. Probiotic potential is presented using star symbols: green (strong), pink (moderate), yellow (weak), red (virulent), and blue (uncharacterized)

Conclusion

The gut of cattle is a good reservoir of indigenous LAB, but very few have strong probiotic potential. Among the isolates, L. plantarum CS-23 and P. acidilactici BC-14 exhibited the most remarkable probiotic attributes. Both strains showed susceptibility to a broad range of antibiotics and lacked hemolytic as well as gelatinase activity, indicating a favorable preliminary safety profile. Phylogenetic lineage revealed a close evolutionary relationship with probiotic strains of the same species and animal origin, while exhibiting clear divergence from pathogenic and less effective strains. These findings suggest that the CS-23 and BC-14 hold significant probiotic potential. Despite in vitro characterization providing valuable preliminary insights. Therefore, comprehensive whole genome analysis and development of formulation followed by in vivo trials in livestock, are necessary to fully elucidate their safety and efficacy for potential application in the livestock sector. Overall, these findings could help develop an effective feed additive to improve gut health and growth performance in livestock.

Authors' contributions

Asiya Batool : Conceptualization, Methodology, Writing – original draft, Analysis, Data Curation, Review and editing. Humaira Yasmin: Conceptualization, Analysis, Investigation, Writing& Editing. Muhammad Abdul Farah: Writing& Editing, Analysis, Investigation. Naeem Khan: Writing & Editing. Muhammad Nadeem Hassan: Project administration, investigation Conceptualization, Resources, Supervision, Validation, Writing-review and editing.

Funding

This work was supported by the Pakistan Science Foundation (PSF) under research grant number NSLP/PSF/CUI/C # 820.

Data availability

All data generated during this study is included in this article and its supplementary files. The nucleotide sequences of the bacterial strains used in this study has been deposited in the National Center for Biotechnology Information (NCBI; [https://www.ncbi.nlm.nih.gov](https:/www.ncbi.nlm.nih.gov) ) GenBank database under the accession numbers OP723337, OP721097, OP721061, OP721065, OP720973, and OP721029.

Declarations

Ethics approval and consent to participate

All animal procedures in this study were conducted in accordance with institutional and national guidelines for the care and use of animals in research. Ethical approval was obtained from the Ethical Review Board of COMSATS University Islamabad (CUI) under reference number (CUI/Bio/ERB/1111–2023/3) and the Institutional Ethical Committee of the Livestock Research Station, National Agriculture Research Center (NARC), Islamabad, Pakistan, via letter number LRS/NARC (1/12/2024). Both institutes provided their Informed Consent from the owners.

Consent for publication

N/A.

Competing interests

The authors declare no competing interests.