2.1. In vitro characterization of lactobacillus bacteria

For a collection of healthy uterus probiotic cultures, a total of 34 cows and 17 buffaloes were selected for this experiment from the Ambulatory Clinic of Kamdhenu University, Sanoda, Taluka Dehgam, Gandhinagar, Gujarat, India. The cows and buffaloes reported for artificial insemination (AI) with apparently healthy and with optimum body condition score (2.5–3), without any reproductive deformities or infection (white slide test negative) were enrolled for the study. The age of the selected animals ranged from 4 to 8 years (4.8 ± 3.5 years; median ± SD years) with bodyweight from 310 to 510 kg (359 ± 110 kg; median ± SD kg). As the animals were reported for AI, no approval for IAEC is needed for the experiment. Our (Gohil et al., 2022) research outlines the process of collecting vaginal swab samples from the fornix of the vagina close to the external orifice of the uterus. Indeed, samples from each cow and buffalo were collected from the fornix of vagina using the uterine swab (part no. 17214/2951/2950; Minitub GmBH, Hauptstrasse 41, 84184, Tiefenbach, Germany). Before taking the vaginal samples, external genitalia were washed with 4% chlorhexidine (Excelle, DRE Veterinary, India). The samples were transported maintaining a 4°C temperature and subsequently subjected to isolation of probiotic cultures.

Microorganisms isolated from vaginal swab samples were suspended in 4 ml of 1× PBS buffer and incubated at 37°C with intermittent mixing using a vortex. Dilutions were prepared and then isolation was carried out using the spread plate and pour plate methods on microbiological media (MRS; recommended for the growth of the Lactobacilli group of bacteria). Pure bacterial cultures were obtained through sub-culturing and stored at −80°C in 25% glycerol for future experiments. Primary identification was performed based on morphological characteristics, Gram staining, and catalase reaction results. The microorganisms were identified based on the guidelines provided by Bergey’s Manual of Systematic Bacteriology (Kandler, 1986). To evaluate and compare the probiotic properties of our bacterial isolates, we chose a well-established and widely recognized probiotic strain, Lacticaseibacillus rhamnosus GG (LRGG).

Bacterial cultures were grown overnight in MRS broth incubated at 37°C with shaking. Bacterial cells were harvested by centrifugation (2,000 × g, 15 min, 4°C), washed twice in PBS buffer (NaCl = 8 g/L; KCl = 200 mg/L; Na₂HPO₄ = 1.44 g/L; KH₂PO₄ = 245 mg/L pH 7.4 ± 0.2) and finally, OD₆₀₀ was adjusted to 0.55–0.60 in the same buffer. Auto-aggregation was determined at initial time (0 h) and after 2 h of incubation of these isolated bacterial cultures by measuring absorbance at 600 nm. The percentage of the auto-aggregation assay is expressed as:

A0, the optical density at 0 h; At, the optical density after 2 h.

To assess the co-aggregation, an equal volume of each pathogen (Staphylococcus aureus and E. coli) was mixed and incubated at room temperature. Absorbance was measured at 600 nm at 0 h and after 2 h of incubation and the percentage of co-aggregation determined as:

Ax and Ay represent absorbance of probiotic isolates and pathogenic bacteria individually in the control tube and A(x + y) represents the absorbance of the mixture of probiotic isolate with pathogenic bacteria.

The cell surface hydrophobicity percentage of bacterial isolates toward hydrocarbons such as chloroform, ethyl acetate, and xylene was evaluated following the protocol outlined by Chen et al. (2014). Bacterial cell pellets were harvested as described previously which was followed by centrifugation at 2,000 × g, 15 min and 4°C, and then resuspended in a PBS buffer. Each hydrocarbon was added to the cell suspension in the proportion of 1:3 respectively, vortexed for 5 min, and incubated for 20 min at room temperature. Further, the OD was measured using the aqueous phase at 600 nm (At). The percentage of the cell surface hydrophobicity (% H) was calculated as follows:

A0, the optical density at 0 h; At, the optical density after 20 min.

In order to investigate the mechanism of inhibition of pathogenic bacterial growth, an agar well diffusion assay was performed. Firstly, LAB strains were cultured in MRS broth for 48 h at 37°C and their cell-free supernatant (CFS) was obtained by centrifugation at 5,000 × g for 10 min at 4°C. The obtained CFS was then concentrated at 65°C for 1.5 h using a vacuum concentrator. Overnight cultures of indicator microorganisms, E. coli and S. aureus, were washed twice in PBS solution and resuspended in fresh PBS solution to obtain a concentration of approximately 10⁷ cells/ml (OD₆₀₀ ∼ 0.25). 100 μl cell suspension was spread onto nutrient agar plates, and wells with 8 mm diameter were created in the agar. A total of 100 μl of concentrated CFS was added to each well, and fresh MRS broth was used as a negative control. The plates were then incubated aerobically at 37°C for 24 h, and the inhibitory activities were calculated by measuring the diameter of the zones of inhibition around the wells (Akabanda et al., 2014).

Quantification was conducted by doing broth dilution once activity was observed in the CFS agar well diffusion assay. To determine the minimum inhibitory concentration (MIC) of probiotic cells against pathogens, the broth dilution procedure was employed, using CFS obtained from probiotic cells. The concentrated CFS was tested against the pathogens to evaluate its effectiveness in inhibiting their growth. The pathogen inoculum was prepared by growing the pathogen in a nutrient broth media until it reached the logarithmic phase. The cells were collected by centrifugation and resuspended in saline to obtain a cell suspension with a turbidity equivalent to a 0.5 McFarland standard. The CFS was prepared by growing the probiotic cells in MRS medium for 48 h, and the cells were removed by centrifugation. The supernatant was filtered through a 0.22-μm filter to obtain the cell-free supernatant. The CFS was diluted with the growth medium to obtain a series of twofold dilutions, ranging from 1:2 to 1:32. Inoculation in the microtiter plate involved adding equal volumes of the diluted CFS and pathogen suspension to each well in the dilution series. The microtiter plate was then incubated at 37°C for 24 h, and the MIC was determined by visually inspecting the wells for growth of the pathogen by measuring OD₆₀₀. The lowest concentration of the CFS at which no visible growth of the pathogen was observed was defined as the MIC (Lim et al., 2018).

The antimicrobial activity by partially purified bacteriocin, bacterial culture sample A and B, were grown in MRS broth for 24 h under shaking conditions at 37°C for 48 h. After incubation, CFS was collected by centrifugation (10,000 rpm, 15 min, 4°C). 500 ml of CFS was precipitated by 80% saturated ammonium sulfate. After overnight stirring at 4°C, the resulting precipitate was collected by centrifugation at 12,000 × g for 15 min and dissolved in a minimal volume of 0.1 M sodium acetate buffer (pH 6.5), which was further centrifuged (12,000 × g for 15 min) and the supernatant was collected. The collected supernatant was used to study the antibacterial activities of the samples that were tested against pathogenic Gram-positive and Gram-negative bacteria. The indicator strains were inoculated in the appropriate soft agar media, and the antibacterial activities were determined by measuring the zone as previously described. CFS with neutralized pH 7.0 was used to study antimicrobial activity against pathogenic bacteria. All experiments were conducted in triplicate. In order to investigate the acid production properties of the Lactobacilli strains, MRS medium containing 0.017% W/V bromocresol purple was utilized. Bromocresol purple functions as a pH indicator by changing its color from purple to yellow under acidic conditions.

The hemolytic activities were assessed using the cells streaked on Columbia agar plates supplemented with 5% defibrinated buffalo blood and cultured for 48 h at 37°C to determine their hemolytic activity. The development of a visible zone of hemolysis encircling the colonies (++hemolysis), partial hemolysis, as well as a greenish-brown zone (+hemolysis) or no reaction (+hemolysis) was used to evaluate the hemolytic reaction after incubation. S. aureus ATCC 25923 cells were used as positive controls. Furthermore, potential probiotic bacterial strains were tested for their susceptibility to antibiotics using the disc diffusion method according to the criteria of the National Committee for Clinical Laboratory Standards with some modifications. HiMedia 20 Icosa Universal-1, Icosa Universal-2, and Icosa G-I-Plus antibiotic discs containing a total of 20 different antibiotics were placed on the surface of the solid nutrient agar, and after incubation for 24 h at 37°C, the diameters of the inhibition zones were noted.

The fatty acid methyl esterase (FAME) analysis was conducted using a comprehensive strategy that involved several steps ranging from sample preparation to GC-MS operation and analysis. The objective of the analysis was to determine the change in fatty acid components of two different bacterial cells. To achieve this, two sets of samples were prepared: the first set involved harvesting cells from MRS agar plates, while the second set involved collecting cell pellets after centrifugation of young active cultures. The samples were then subjected to the FAME analysis using the aforementioned strategy. The peaks were compared with standard esters using gas chromatography fatty acid methyl esters (GC-FAME) method as described and reported by Sasser (2006) and Das et al. (2019). From the supernatant of the culture, metabolites extraction and derivatization were performed. Here protein precipitation was done using ACN (acetonitrile) and allowed to dry overnight at 65–70°C, followed by derivatization using N,O-Bis(trimethylsilyl)trifluoroacetamide (BSTFA) was performed at 70°C. Afterward, GC-MS run was performed to separate and analyze the samples (Kinani et al., 2008). Exo-polysaccharide (EPS) secretion was detected by growing culture on MRS medium and also supplementing MRS media with 2% sugar concentrations of monosaccharides such as glucose, fructose, and galactose (Nambiar et al., 2018; Butorac et al., 2021). Colonies which were obtained after streaking followed by incubation for 24 h, were observed and analyzed using stereo microscopy (Magnüs MSZ-Bi, Olympus Pvt. Ltd., Noida, India).

To verify the cell adhesion assay, the probiotic cells were labeled with CFDA-SE and analyzed by flow cytometry after adhering to epithelial cells. Entire protocol was performed following previously described protocol in Wang et al. (2005) and Zhang et al. (2022), with slight modifications. Probiotic cells were harvested from overnight grown culture at 3,000 × g for 10 min. To remove excess media, the cell pellet was washed twice with sterile PBS. Here, the probiotic cells were labeled with CFDA-SE (50 μM) at 37°C for 20 min. The labeling reaction was terminated once the incubation period was accomplished by pelleting the cells followed by washing with sterile PBS twice, to remove excess dye molecules as described in Wang et al. (2005). As described in Zhang et al. (2022), by doing certain modifications, goat endometrial cells were seeded in the 24 well plate at a concentration of 50,000 cells/well and incubated at 37°C in 5% CO₂. This was followed by 100 μl aliquot of labeled probiotic cells (10⁸ cells/ml in RPMI-1640 medium) were then added to a 24-well plate in RPMI medium and incubated at 37°C in 5% CO₂ for 2 h. Supernatant was aspirated and cells were washed twice with 500 μl of sterile PBS. Later, the cells were treated with 0.05% Triton X-100 and incubation of the system was carried out for 10 min. After which the sample was analyzed in Flow Cytometer (BD FACSAria Fusion™).