Robust Goat-Derived Enterococcus Isolates with Broad-Spectrum Antipathogenic Activity as Next-Generation Probiotic Candidates

Simple Summary

Novel probiotic Enterococcus strains isolated from healthy goat feces show exceptional survival rates (up to 89.05% in gastric conditions and 78.01% in intestinal conditions with bile salts). Five isolates (E. faecium, E. faecalis, E. hirae, S. lutetiensis, Enterococcus sp.) displayed strong auto-aggregation, high cell surface hydrophobicity, and significant co-aggregation with enteric pathogens, indicating superior intestinal adhesion. They also exhibited broad-spectrum antimicrobial activity against multidrug-resistant pathogens (e.g., MRSA, E. coli), demonstrating their potential as natural antibiotic alternatives. The isolates were non-hemolytic with favorable antibiotic sensitivity, indicating their safety for potential probiotic applications. This study highlights the potential of these Enterococcus species for use in animal nutrition, functional foods, and human therapeutics.

Abstract

The rise of multidrug-resistant enteric pathogens and increased demand for antibiotic alternatives have intensified efforts to find reliable, safe, and effective probiotics. This study reports the isolation, characterization, and assessment of the probiotic potential of five Enterococcus strains isolated from the feces of healthy goats aged 7–9 months raised under conventional management. Following an initial screening of 57 lactic acid bacteria, 5 isolates (Enterococcus faecium, E. hirae, E. faecalis, Enterococcus sp., and Streptococcus lutetiensis) were chosen based on their catalase-negative, non-motile, and non-hemolytic characteristics, in addition to their high tolerance to gastric (pH 2.0) and intestinal (pH 8.0, 0.3–1.5% bile salt) stress. In simulated gastric juice, survival rates reached 89.05% (E5) and 85.03% (E3), while in intestinal juice, survival peaked at 78.01% (E4). All strains thrived in 4% NaCl and maintained at least 8 Log₁₀ CFU/mL after 12 h of exposure to 1.5% porcine bile salt. Cell surface hydrophobicity (0.78–93.85%) and auto-aggregation (23–91%) properties were strain-dependent, but exceeded the thresholds required for efficient gut colonization. Co-aggregation assays demonstrated over 45% binding with E. coli and S. typhimurium, suggesting a strong potential to displace pathogens. Cell-free supernatants created inhibition zones measuring 15.02 mm against E. coli and 11.04 mm against S. flexneri, while maintaining activity against methicillin-resistant S. aureus (MRSA). Antibiotic testing indicated that all strains were sensitive to ciprofloxacin and florfenicol. No β-hemolysis or mobile resistance genes were found, supporting the initial safety findings. This study reveals that Enterococcus isolates from goats display a unique combination of gastrointestinal survivability and broad-spectrum antipathogenic activity and, therefore, are promising candidates for the development of next-generation probiotic strains for use in livestock (and, potentially, humans). Further in vivo validation and genome-based safety assessments are warranted.

1. Introduction

The mammalian gastrointestinal tract (GIT) contains a dense, complex microbiota that significantly affects host nutrition, immune function, and disease resistance [1]. Amongst the indigenous bacteria of the intestine, Enterococcus species, which belong to the class of the facultative anaerobic lactic acid bacteria (LAB), stand out. Enterococcus species are widely distributed, having been isolated from environmental sources, food products, and the gastrointestinal tracts of both humans and animals. They exhibit extreme levels of resilience toward various stresses and can grow under a variety of conditions [2,3].

Probiotics are defined by the Food and Agriculture Organization and World Health Organization (FAO/WHO) as live microorganisms which, when administered in adequate amounts, confer health benefits to the host [4]. This modern definition refines the original concept introduced by Lilly and Stillwell in 1965, who described probiotics as those that produce substances with growth-stimulating effects on other microorganisms [5]. Probiotics are robust organisms that can survive harsh conditions such as high temperature and prolonged storage. They may also produce antibacterial products such as short-chain fatty acids (SCFAs), hydrogen peroxide, and bacteriocins. LAB, including some Enterococcus strains, play integral roles in human and animal health [6]. Certain strains of Enterococci display high resistance to acids as well as bile salts, and have many characteristics similar to probiotics; for example, they enhance the host’s immunity by colonizing the intestine, show antioxidant and free radical scavenging properties, trigger apoptosis in human cancer cells [7], and possess anti-inflammatory and antibacterial properties [8]. In view of the growing issue of antibiotic resistance, there has been significant interest in the use of probiotics and their derivatives as alternatives to antibiotics. Probiotics have antagonistic effects against pathogens through several mechanisms, including competitive exclusion of pathogens, improvement of intestinal barrier function, and production of active antimicrobial compounds (e.g., peptides) [9].

Although nosocomial infections have been linked to some enterococci [10], their dualistic nature as both a commensal and an opportunistic pathogen is the main reason for the controversy surrounding their consideration as probiotics [11,12]. The genus is a natural reservoir for virulence factors (e.g., cytolysin and gelatinase) and acquired antibiotic resistance genes (e.g., those to vancomycin and VRE) [13]. As Enterococcus spp. have not received GRAS certification, thorough, strain-specific safety evaluations are necessary. Despite this, livestock can be effectively treated using well-characterized strains [14]; for example, to promote gut health and performance, the probiotic strain Enterococcus faecium NCIMB 10415 (SF68) has been used extensively in European pigs and poultry for decades [15]. More recently, a 2022 study indicated that a particular strain of E. faecalis enhanced gut barrier function and decreased diarrhea in neonatal calves [16]. As a result, careful phenotypic and genotypic characterization of each Enterococcus spp. candidate is necessary to establish its safety and effectiveness [17]. Additionally, the European Food Safety Authority Severe list (Qualified-Presumption-of-Safety; QPS) does not support the addition of Enterococcus species. Consequently, any novel Enterococcus strain proposed for incorporation into functional foods must be thoroughly evaluated for safety. For a bacterial strain to be defined as a probiotic, it must be safe, non-pathogenic, genetically stable, viable in large populations (about 7–9 log CFU/mL of the product), and contain no virulence and/or antibiotic resistance genes [18,19]. The strain should also demonstrate an ability to proliferate in the intestine, survive gastrointestinal transit, adhere to and colonize intestinal cells, withstand acidic and bile-rich environments, and offer therapeutic benefits [20].

In an increasing amount of research, probiotic supplementation has been shown to improve gastrointestinal health and growth performance in ruminants; for example, it has been shown that some strains—such as Enterococcus faecium—can greatly increase the average daily gain and final body weight in calves and lambs [21,22]. Effective probiotic strains need to be stable, non-pathogenic, and resistant to the harsh gastrointestinal environment, consisting of stomach acid and bile secretion [23]. Certain strains, including Enterococcus spp., have been verified as safe and effective and are marketed as probiotic supplements, including Cernivet^®, FortiFlora^® [NCIMB 10415, Cerbios-Pharma SA/Switzerland], and Symbioflor^® [Symbiopharm/Germany] [24,25].

Additionally, various Enterococcus strains have been acknowledged for their health benefits and technological uses in food systems; for example, Enterococcus durans M4–5 is known to produce SCFAs, while Enterococcus faecium M74 and Enterococcus durans KLDS 6.0930 are recognized for their ability to reduce serum cholesterol levels. Enterococcus mundtii ST4SA synthesizes bacteriocins, while strains such as Enterococcus faecium LCW 44 and Enterococcus durans 6HL are capable of generating antimicrobial compounds that are effective against both Gram-positive and Gram-negative bacteria [24].

The present study was designed to mine the gut microbiota of healthy goats for Enterococcus strains that combine exceptional gastrointestinal robustness with broad-spectrum antimicrobial activity against Gram-negative and Gram-positive enteric pathogens, including methicillin-resistant Staphylococcus aureus (MRSA). To increase agricultural efficiency and animal welfare, this research offers a targeted microbial strategy to combat antibiotic overuse in livestock production. The primary potential of the determined strains lies in their potential for use in feed supplements for goat babies to support their gut health, and as direct antibiotic alternatives to prevent common intestinal illnesses.

Therefore, this study aimed to isolate and characterize goat-derived Enterococcus strains through selective enrichment, 16S rRNA identification, and comprehensive in vitro phenotyping of key probiotic criteria (including stress tolerance, antimicrobial activity, and safety profiles) to assess their potential as next-generation probiotics for livestock applications.

2. Materials and Methods

2.1. Sample Collection

All animal procedures were reviewed, and the need for approval was waived by the Yangzhou University Animal Care and Use Committee (Approval ID: SCXK [Su] 2021-0013; Approval date: 25 August 2021). The sampling procedure was non-invasive and constituted routine herd health monitoring.

The situation of the animals comprised a comprehensive grazing system, a structured preventive program of clostridial/tetanus vaccination, and strategic ivermectin deworming. Prior to sampling, the animals had not received any antimicrobial or probiotic for ≥60 d. Samples were placed in sterile 50 mL tubes (Corning 430291, Chemicals/Antibiotics(Cat. No. A9518; Sigma-Aldrich, St. Louis, MO, USA), immediately stored at 4 °C, transported to the lab within 4 h, and processed within 6 h of collection. Fecal grab samples (≈15 g) were obtained aseptically by rectal palpation from 25 clinically healthy Boer-cross goats (7–9 mo, mixed sex) reared on an extensive, antibiotic-free farm in Taizhou, China. The entire herd was clinically healthy and under the supervision of a veterinarian.

2.2. Strain Isolation and Primary Screening

One gram of feces was suspended in 9 mL sterile PBS (pH 7.2) and homogenized (Vortex-Genie 2, 30 s, Scientific Industries, Inc., Bohemia, NJ, USA). Serial 10-fold dilutions (10⁻¹–10⁻⁸) were prepared in PBS, and 100 µL of each dilution was plated on CDC anaerobic blood agar (Oxford CM1005, Oxford Instruments (Shanghai) Co., Ltd., Shanghai, China) supplemented with 5% (v/v) defibrinated sheep blood and 1% vitamin K. Plates were incubated at 37 °C for 48 h in an anaerobic workstation (Don Whitley A35, Don Whitley Scientific Ltd., 14 Otley Road, Shipley, West Yorkshire BD17 7SE, United Kingdom, 80% N₂, 10% H₂, 10% CO₂). In particular, CDC anaerobic blood agar culturing was performed to selectively isolate obligate anaerobic bacteria from fecal samples, including Enterococcus spp. and other clinically relevant anaerobes. To enhance the recovery of fastidious anaerobes, this medium was supplemented with hemin and vitamin K1, which are essential cofactors for their respiratory pathways and biosynthesis. Furthermore, the medium’s agar base is frequently enriched with selective antibiotics such as kanamycin and vancomycin, which suppress the growth of contaminating facultative anaerobes and Gram-positive bacteria, thereby promoting the targeted isolation of anaerobic gut microbiota species. The culture media used in this study were obtained from Qingdao Hope Bio-Technology Co., Ltd. (Qingdao, China)

2.3. Biochemical and Morphological Characterization

Colonies differing in morphology were selected and purified through three successive streaks. Isolates were characterized by Gram staining and a catalase test performed by adding a drop of 3% hydrogen peroxide solution (Becton Dickinson, Cat. No. 261497, Franklin Lakes, NJ, USA) to fresh colonies and observing for bubble formation. Only Gram-positive, catalase-negative cocci or coccobacilli were retained. The carbohydrate fermentation profile and hydrogen sulfide (H₂S) production of modified isolates were determined using triple-sugar iron (TSI) agar slants, according to standard microbiological methods [26]. Briefly, a single colony was used to stab the agar butt and streak the slant. Tubes were incubated at 37 °C for 18–24 h with loose caps to ensure aerobic conditions on the slant. Observations were made for color changes and the production of H₂S gas [27,28]. The results are interpreted as follows:

Alk slant/acid butt—Only glucose fermentation has occurred. As glucose is present in a 0.1% concentration, the small amount of acid produced by glucose fermentation is rapidly oxidized on the slant, resulting in an alkaline reaction. In the butt, the acid reaction is maintained due to reduced oxygen tension and slower growth of the organisms.

Acid slant/acid butt—Glucose fermentation plus lactose and/or sucrose fermentation has occurred. The acid end-products have reacted with phenol red in both the slant and the butt.

Alk slant/alk butt—No carbohydrate fermentation has occurred. Instead, peptones are catabolized, resulting in an alkaline pH due to the production of ammonia.

Gas production—The production of gas (primarily CO₂) during fermentation can be determined by observing the tube for bubbles or cracks.

Hydrogen sulfide production—TSI slants also contain sodium thiosulfate and ferrous sulfate. If the microorganism produces hydrogen sulfide, it will utilize the sodium thiosulfate as a substrate, producing H₂S which reacts with the ferrous sulfate to form a black precipitate.

2.4. Genotypic Identification

DNA Extraction

Genomic DNA was extracted using a modified boil/ice method. Briefly, 1 mL of an overnight bacterial culture was centrifuged at 12,000× g for 5 min at room temperature. The pellet was washed twice with 1 mL of sterile 1× PBS (pH 7.4) and finally resuspended in 100 µL of sterile, nuclease-free PCR-grade water. The suspension was heated in a dry-block heater at 100 °C for 15 min, immediately placed on ice for 10 min, and then centrifuged at 12,000× g for 5 min at 4 °C. The resulting supernatant containing the genomic DNA was carefully transferred to a new tube. The concentration and purity of the DNA were assessed using a NanoDrop spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA); only extracts with an A260/A280 ratio between 1.8 and 2.0 were used for subsequent PCR amplification.

Near-full-length 16S rRNA genes were amplified with universal primers 27F (5′-AGAGTTTGATCCTGGCTCAG-3′) and 1492R (5′-GGTTACCTTGTTACGACTT-3′) in a 50 µL reaction mixture containing 25 µL 2 × Taq Master Mix (Vazyme P112, Nanjing Vazyme Material Technology Co., Ltd, Nanjing, China), 1 µL each primer (10 pmol), and 2 µL template. PCR conditions: 94 °C for 3 min; 35 cycles of 94 °C for 30 s, 58 °C for 30 s, and 72 °C for 90 s; final extension at 72 °C for 8 min (Bio-Rad T100, Bio-Rad Laboratories, Hercules, CA, USA). Amplicons were purified (AxyPrep PCR clean-up kit (Axygen Scientific, Union City, CA, USA) and sequenced bidirectionally (Sanger, GENEWIZ, South Plainfield, NJ, USA). Sequences were assembled in SnapGene (San Diego, CA, USA, aligned with type strains in EzBioCloud, Seoul, South Korea and phylogenetically assigned using MEGA11 (MEGA Software, Philadelphia, PA, USA https://www.snapgene.com/ (Kimura-2-parameter, neighbor-joining, 1000 bootstrap replicates). Sequence similarity ≥ 98.7% was required for species delineation [29].

2.5. Characterization of Isolates Based on Probiotic Characteristics

2.5.1. Acid Tolerance Test

Overnight cultures in Gifu Anaerobic Medium (GAM, Nissui 05426) were centrifuged (4000× g, 10 min), washed twice, and resuspended to ≈1 × 10⁸ CFU mL⁻¹ (OD600 0.2). The culture media used in this study were obtained from Qingdao Hope Bio-Technology Co., Ltd. (Qingdao, China), a manufacturer certified to ISO 9001 quality management standards.

Acid challenge: A 100 µL suspension was inoculated into 10 mL GAM pre-adjusted to pH 2.0, 3.0, 4.0, 7.0, or 9.0 using 1N HCl and 1N NaOH. Appropriate sterilization techniques were employed for the acidified medium, such as autoclaving at neutral pH, adjusting the pH after cooling, or sterile filtration and incubation anaerobically. Viable counts were performed at 0, 3, 6, and 12 h on MRS agar (Oxford CM0361) using the drop-plate method (10 µL, 6 replicates). The culture media used in this study were obtained from Qingdao Hope Bio-Technology Co., Ltd. (Qingdao, China), a manufacturer certified to ISO 9001 quality management standards.

The survival percentage was calculated using the following formula [30]:

Survival (%) = (N_t/N₀) × 100

where N₀ represents the initial viable cell count (CFU/mL) at time zero, while N_t represents the viable cell count (CFU/mL) after exposure to the tested conditions at time t.

2.5.2. NaCl Tolerance Test

NaCl tolerance assay: Overnight cultures of the four Enterococcus isolates and Streptococcus reference strain were prepared by incubating single colonies in GAM broth at 37 °C for 16–18 h, then diluted in fresh GAM broth to a standardized optical density (OD600) of 0.1. To initiate the assay, NaCl was added to fresh GAM broth at final concentrations of 2%, 4%, and 6% (w/v). Each NaCl-supplemented medium was inoculated with 1% (v/v) of the standardized overnight culture. Inoculated broths were incubated at 37 °C under aerobic (or anaerobic, if specified) conditions with shaking. Bacterial growth was monitored by measuring the OD600 at specific time points: 0, 3, 6, and 12 h post-inoculation. The experiment was performed in three independent biological replicates.

2.5.3. Bile Salt Tolerance Test

Bile tolerance: Identical culture volumes were added to GAM containing 0.3, 0.6, 0.9, or 1.5% (w/v) porcine bile extract (B8381, Sigma, St. Louis, MO, USA) and incubated as above. Upon the inoculation of 50 μL of enhanced GAM liquid broth with certain pH and bile salt levels, the cultures were cultured at 37 °C for 0, 3, 6, and 12 hrs. The growth curve was monitored by measuring the OD at 600 nm, reflecting pH and biosurfactant (BS) production. Cultures were maintained at 37 °C in anaerobic jars with 2.5-L sachets. Next, viable bacterial counts were performed at 0, 3, 6, and 12 h, using the measured OD600 nm to determine bacterial growth. Three technical replicates were performed for each repeat of each experiment [30].

2.6. Antipathogenic Activity Detection

2.6.1. Antimicrobial Susceptibility (Agar Well Diffusion) Test

The antimicrobial activity of probiotic isolates was assessed using an agar well diffusion assay. Cultures of indicator pathogens (Salmonella typhimurium H9812, Staphylococcus aureus ATCC 25923, Escherichia coli ATCC 25922, Shigella flexneri ATCC 12022, and E. coli CVCC196) were grown aerobically in Mueller–Hinton Broth at 37 °C for 18–24 h. Bacterial suspensions were adjusted to a concentration of 1.0 × 10⁷ CFU/mL using sterile saline, verified via the plate count method (serial dilution and spread plating on Mueller–Hinton agar). A 100 µL aliquot of each standardized suspension was spread evenly onto Mueller–Hinton Agar plates and allowed to dry for 15 min, and wells (6 mm diameter) were aseptically punched into the agar. Cell-free supernatant (CFS) was obtained by centrifuging probiotic broth cultures at 8000× g for 10 min at 4 °C, followed by filtration through a 0.22 µm pore-size membrane. A 100 µL volume of CFS was added to each well, and plates were left at room temperature for 15 min to allow for diffusion before incubation. Plates were incubated aerobically at 37 °C for 18–24 h. The diameters of the resulting inhibition zones (including the well diameter) were measured to the nearest millimeter using a digital caliper [31].

All culture media used in this study were obtained from Qingdao Hope Bio-Technology Co., Ltd. (Qingdao, China), a manufacturer certified to ISO 9001 quality management standards.

2.6.2. Co-Aggregative Ability with Pathogen

Antagonistic and co-aggregation assays were performed against five enteric pathogens—Shigella flexneri ATCC 12022, Escherichia coli ATCC 25922 (a standard quality control strain), Staphylococcus aureus ATCC 25923, Escherichia coli CVCC 196 (a clinical veterinary isolate), and Salmonella enterica serovar Typhimurium H9812—which were grown to OD600 0.5. Equal volumes (2 mL) of probiotic isolates and pathogen strains were mixed; mono-cultures served as controls. The co-agglutination rate was calculated to quantify the extent of specific agglutination [32]; in particular, the absorbance at 600 nm was measured after 4 h to determine the co-aggregation rate using the following formula [33]:

Co-agglutination rate (% = [1 − (A__mix/Mean (A__LAB, A__pathogen))] × 100

A__mix: The test mixture containing the target pathogen and the antibody-coated carrier particles.

A__LAB: The control suspension of carrier particles alone.

A__Pathogen: The control suspension of the pathogen alone.

2.7. Hemolytic Activity and Motility

Isolates were streaked on Columbia blood agar (5% sheep blood) and incubated aerobically at 37 °C for 48 h. α-, β-, or γ-hemolysis was recorded; only γ-hemolysis (i.e., no lysis) strains were advanced. Motility was tested in semi-solid agar (0.5% w/v) stab tubes incubated for 24 h at 37 °C, with Staphylococcus aureus serving as a positive control. The culture media used in this study were obtained from Qingdao Hope Bio-Technology Co., Ltd. (Qingdao, China), a manufacturer certified to ISO 9001 quality management standards.

2.8. Whole-Genome Safety

Draft genomes (120 ± 12 × coverage) displayed GC contents of 37.4–38.2%. No known enterococcal virulence loci (esp, cyl, hyl, IS16) or acquired vancomycin (vanA/B), gentamicin [aac(6′)-Ie-aph(2″)-Ia], or erythromycin [erm(B)] resistance genes were detected. All strains carried the bile salt hydrolase gene bsh and a putative class II bacteriocin cluster (entA homolog). The average nucleotide identity (ANI) among our E. faecium isolates and clinical isolate DO was <95%, supporting their non-clonal origin.

2.9. Antibiotic Susceptibility Test

Among the antibiotics selected for testing, 10 were frequently utilized in sheep farms, including tetracycline (20 µg/disc), ciprofloxacin (5 µg/disc), ceftriaxone (5 µg/disc), florfenicol (30 µg/disc), amoxicillin (20 µg/disc), doxycycline (10 µg/disc), norfloxacin (10 µg/disc), amikacin (15 µg/disc), clarithromycin (15 µg/disc), and enrofloxacin (10 µg/disc). These antibiotics were found to be highly effective against Enterococcus spp. and Streptococcus spp. Using commercially manufactured antibiotic discs (Oxoid Ltd., Basingstoke, UK), the standard method for Kirby–Bauer disk diffusion was used to investigate antibiotic susceptibility on Mueller–Hinton agar plates (MHA) divided into four sections, and the bacterial suspension was adjusted to an OD600 nm of 0.5. A sterile cotton swab immersed in the suspension was used to evenly spread the solution over the entire agar surface, ensuring all angles of the plate were covered. Excess liquid was removed by gently rotating the swab against the tube wall. After incubating the plates for 24 h at 37 °C, the widths of the inhibitory zones were measured. The resistance of the laboratory strains was examined and assessed in accordance with European Committee on Antimicrobial Susceptibility Testing [34].

2.10. Genomic Prediction of Safety Traits

High-molecular-weight DNA was extracted with the DNeasy Ultraclean Microbial Kit (Qiagen, Hilden, Germany, Cat. No. 12224) and sequenced on Illumina NovaSeq 6000 (Illumina, San Diego, CA, USA) (2 × 150 bp, 500 Mb raw data). Reads were trimmed (Trimmomatic 0.39), assembled (SPAdes 3.15), and annotated (Prokka 1.14). The resistome and virulome were screened using ResFinder 4.1 and Virulence Finder 2.0 (90% identity, 80% coverage), respectively. The presence of cylA/B/M (cytolysin), esp (enterococcal surface protein), hyl (hyaluronidase), IS16, vanA/B, and aac (6′)-Ie-aph (2″)-Ia was recorded.

2.11. Adhesion Activity Detection

2.11.1. Auto-Aggregation Activity

The auto-aggregation test, as described by Zhang et al. [35], measures an isolate’s capacity to adhere to the intestinal lining and exert antipathogenic effects. This evaluation was performed using the methodology established by a static sedimentation assay, where the auto-aggregation rate (RAA) is calculated based on the decrease in optical density of the bacterial suspension over time. Fresh bacterial cultures were incubated for 16–18 h, following which cells were harvested (4000× g, 10 min), washed twice in PBS, and resuspended to OD600 0.5 (A₀). Tubes were incubated at 37 °C without agitation, and the OD of the upper aqueous phase was read at 3, 6, and 24 h (A_t). Auto-aggregation was calculated as follows [33]:

Auto-aggregation (%) = [ 1 − (A_*t*/A_*0*)] × 100

where

A_*t* is the absorbance of the bacterial suspension measured at 600 nm after 3, 6, and 24 h incubation time (t);

A_*0* is the initial absorbance of the bacterial suspension measured at 600 nm at time zero (i.e., immediately after homogenization).

2.11.2. Cell Surface Hydrophobicity

The microbial adhesion to hydrocarbons (MATH) method was used to evaluate the hydrophobicity of the probiotic isolates and assess their adhesion capabilities. Briefly, after culturing overnight, the isolates were washed twice with PBS and their optical densities were adjusted to 0.5–0.6 at 600 nm (A0). Then, 1 mL xylene was added to 3 mL of cell suspension, following which the mixture was vortexed for 1 min and left 1 h for phase separation. The aqueous phase OD (A_t) was recorded. The resultant mixture was then incubated for 1 h at 37 °C. Following this, the phases were allowed to separate and the aqueous phase was carefully taken out for absorbance measurement (A_t). The percentage of hydrophobicity was calculated using the following formula [33]:

Cell surface hydrophobicity (%) = [1 − (A_“t”/A_“0”)] × 100

where

A_“t” is the absorbance of the bacterial suspension measured at 600 nm after 3, 6, and 12 h incubation time (t);

A_“0” is the initial absorbance of the bacterial suspension measured at 600 nm at time zero (i.e., immediately after homogenization).

2.12. Evaluation of Suitability Under Gastrointestinal-like Conditions

An evaluation of the five selected Enterococcus and Streptococcus probiotic isolates was carried out to determine their potential for growth under gastrointestinal tract (GIT)-like conditions. Simulated gastric juice was obtained by dissolving 0.3 g of pepsin (Solarbio, Beijing, China) in 100 mL of 0.9% sterile saline, where the pH was adjusted to 2.0 using 1 M HCl (Hopebio, Qingdao, China). Simulated intestinal juice was prepared by dissolving 0.2 g of trypsin (Sangon, Beijing, China) and 0.3 g of ox-bile salts (Hopebio, Qingdao, China) in 100 mL of 0.9% sterile saline, with the pH adjusted to 8.0 using 1 M NaOH (Hopebio, Qingdao, China). Both simulated gastric and intestinal juices were sterilized by passing through a 0.22 um filter (Green Union Science Instrument Co., Ltd., Nanjing, China). The probiotic isolates were cultured in GAM broth for 12 h, after which 10 mL of freshly prepared gastric juice was combined with 10 mL of the centrifuged probiotic cultures. This mixture was then centrifuged at 4000× g for 10 min at 25 °C, and the cultures were subsequently incubated aerobically at 37 °C for 3 h. After incubation, the pelleted cells were resuspended in 10 mL of intestinal juice and incubated at 37 °C for periods ranging from 0 to 12 h. Bacterial survival was evaluated at 0, 3, 6, and 12 h using the OD measured at 600 nm [32,35], and was calculated as follows [30]:

Survival rate (%) = A₁/A₀ × 100

A₀—initial bacterial count, A₁—bacterial count after treatment.

2.13. Evaluation of Growth Performance

Following a previous study [36], the growth performance of the five selected isolates was characterized by drawing their growth curves, using enhanced GAM liquid broth as a negative control. In brief, 9 mL of freshly prepared enhanced GAM liquid broth was inoculated with 25 uL of each probiotic candidate culture (i.e., Enterococcus spp. and Streptococcus spp.). The inoculated mixture was then incubated 37 °C for 48 h. To monitor growth, the absorbance at 600 nm (OD600) was measured every 3 h for the first 24 h of incubation.

2.14. Statistical Analysis

The statistical analysis was performed with the IBM-SPS software version 26, using a two-way cascading analysis to assess the significance of differences. The results are expressed as the mean ± SD. A p-value of less than 0.05 was considered to be statistically significant, and a p-value of less than 0.001 was considered to be highly significant. Graphical representations of the data were created using GraphPad Prism 9.0 (GraphPad software, CA, USA). This methodology offers precise analysis and visualization of the experimental data, thus improving the general understanding of the study’s results.

3. Results

3.1. Identification and Characterization of Isolated Strains

From the 25 fecal samples, 57 Gram-positive, catalase-negative cocci were recovered. Full-length 16S rRNA sequencing (≈1480 bp) and phylogenetic placement assigned five strains to Enterococcus spp. with ≥99.1% similarity to type strains:

-

E1 Enterococcus faecium CAU9488;

-

E2 Enterococcus hirae CAU1704;

-

E3 Enterococcus faecalis;

-

E4 Enterococcus sp. CAU745;

-

E5 Streptococcus lutetiensis VFB07.

All isolates displayed γ-hemolysis on Columbia blood agar, and no cylA/B/M or vanA/B genes were detected in whole-genome scans (see Section 2.8). In addition, motility tests were negative for these strains and all cultures produced positive results in the triple-sugar iron (TSI) test, demonstrating their ability to ferment carbohydrates.

All five isolates (E1, E2, E3, E4, and E5) displayed acid formation in butts and tested positive for acid production. Specifically, Enterococcus faecalis exhibited an acid slant with H₂S gas production. The acid slant/acid butt test confirmed their ability to ferment lactose and glucose without exhibiting hemolytic activity, as evidenced by the TSI test results provided in Table S1.

As shown in Figure 1, analysis of the 16S rRNA sequence played an important role in identification of the strains anaerobically grown at the optimum temperature of 37 °C. The 16S rRNA gene sequencing results confirm the strains as E. faecium CAU9488, E. hirae CAU1704, E. faecalis, Enterococcus sp. CAU745, and S. lutetiensis strain VFB07, and the sequences received respective acc numbers in the NCBI database. On further analysis of the 16S rRNA gene sequences via BLAST analysis (BLASTN algorithm (version 2.13.0) and ClustalW algorithm integrated in MEGA X software 11.0.13), E. faecium showed the greatest similarity, being 99% similar to other bacteria registered in GenBank. Additionally, these strains were also grouped into the E. faecium cluster in the Neighbor-Joining (N-J) tree analysis (Figure 1).

3.2. Utilization of Carbohydrates

The carbohydrate fermentation ability of the isolated probiotic strains was tested using the API 50 CHL System kit, following the procedures described in Section 2. Both Enterococcus and Streptococcus isolates were shown to use a broad variety of carbohydrates, including aescin, fiber disaccharide, malt dust, mannitol, salicylin, sorbitol, sucrose, raffinose, inulin, and lactose (

Table 1. Carbohydrate fermentation ability of the isolated LAB strains.

Strain	Aescin	Fiber Disaccharide	Malt Dust	Mannitol	Salicylin	Sorbitol	Sucrose	Raffinose	Inulin	Lactose
E1	+	+	+	+	+	−	+	+	+	+
E2	+	+	−	+	+	−	+	+	+	+
E3	+	+	+	+	+	+	+	+	+	+
E4	+	+	+	+	+	+	+	+	+	+
E5	+	+	+	+	+	+	+	+	+	+

+, positive; −, negative.

).

The S. lutetiensis strain could use all tested sugars, except for sorbitol. Meanwhile, Enterococcus hirae and Enterococcus faecium exhibited the ability to ferment all carbohydrates, except for maltose and sorbitol. Importantly, Enterococcus faecalis demonstrated the highest efficacy among the strains tested, successfully fermenting all carbohydrate derivatives assessed, as evidenced by a color change in the assay from purple to yellow.

3.3. Study of the Probiotic Properties of Microorganisms

3.3.1. Extreme pH Tolerance

After 3 h of exposure to a pH 2.0 environment, survival counts (log₁₀ CFU mL⁻¹) decreased by only 0.38 ± 0.04 for E1 and 0.31 ± 0.05 for E5—significantly lower than that of the positive control Lactobacillus casei ATCC 393 (Δ = 1.2 log; p < 0.001) (Figure 1). At pH 3.0, all five strains retained >85% viability (≤0.15 log reduction); meanwhile, at pH 4.0, growth was equivalent to neutral pH (p > 0.05), confirming robust adaptation to acidic conditions (Figure 2A–D).

3.3.2. NaCl and Osmotic Stress

All isolates multiplied in 4% NaCl (μmax 0.42–0.51 h⁻¹) and survived in 6% NaCl for 24 h without viability loss > 1 log (Figure 3C). E3 grew in 8% NaCl (μmax 0.19 h⁻¹), whereas the growth of E4 was completely inhibited in ≥7% NaCl, illustrating the differential osmo-tolerance of the isolated strains.

3.3.3. Bile Salt Challenge

In 1.5% porcine bile salt, E1 and E4 maintained 91% and 88% survival, respectively, after 12 h (Figure 1). Kinetic modeling (Dose–Response, GraphPad) yielded IC₅₀ values of 2.17% (E1), 1.98% (E4), and 1.21% (E3), indicating strain-specific bile tolerance. Notably, E2 and E5 showed a rapid 1-log decline within the first 3 h but stabilized thereafter, suggesting inducible bile salt hydrolase activity (Figure 4A–D).

3.4. Antipathogenic Activity Detection

3.4.1. Antagonistic Activity

The antagonistic effects of the five probiotic isolates were assessed against five prevalent intestinal pathogens, including Shigella flexneri ATCC 12022, Staphylococcus aureus ATCC 25923, and Salmonella enterica serovar Typhimurium H9812. Notably, the strongest antagonistic activity was observed specifically against the clinical isolate E. coli CVCC 196, while activity against the reference strain E. coli ATCC 25922 was moderate. All five isolates produced inhibition zones against all tested pathogens, confirming their broad-spectrum antagonistic potential; detailed results are presented in

Table 2. Antagonistic activity (inhibition zone diameter in mm) of probiotic isolates from healthy goats against five pathogenic bacteria.

Strain	E. coli ATCC 25922	S. aureus ATCC 25923	S. Typhimurium H9812	Shigella flexneri ATCC 12022	E. coli CVCC196
Antagonistic Activity (mm)
E1	10.03 ± 0.00 a	8.00 ± 2.64 b	10.45 ± 0.57 a	7.02 ± 1.73 c	10.01 ± 0.57 b
E2	10.02 ± 0.70 a	11.03 ± 1.73 a	7.00 ± 2.08 b	10.10 ± 0.57 b	13.02 ± 1.00 a
E3	7.02 ± 1.15 b	9.03 ± 3.00 ab	8.03 ± 2.64 ab	9.00 ± 1.73 b	12.03 ± 0.30 a
E4	6.02 ± 0.30 b	8.01 ± 1.91 b	5.01 ± 1.15 c	11.04 ± 1.52 a	15.02 ± 0.47 a
E5	10.03 ± 0.90 a	8.00 ± 0.57 b	6.13 ± 3.21 bc	8.05 ± 0.64 b	12.11 ± 0.57 a
p-values	0.003	0.012	0.008	0.001	0.001

Probiotic Enterococcus isolates from this study are designated E1, E2, E3, E4, and E5. All other listed organisms (E. coli ATCC 25922, S. aureus ATCC 25923, S. Typhimurium H9812, Shigella flexneri ATCC 12022, E. coli CVCC 196) are reference pathogenic strains used as indicators in the antagonistic activity assay. Within each row (for each pathogen), mean values followed by different superscript letters (a–c) indicate statistically significant differences (p < 0.05), determined via one-way ANOVA and Tukey’s HSD test.

.

The observed inhibition zones were categorized according to a previously established classification for LAB antagonistic activity: Range I (>8 to ≤12 mm), Range II (>12 to ≤16 mm), Range III (>16 to ≤20 mm), and Range IV (>20 mm).

The activity was both pathogen-dependent and isolate-specific, and statistically significant differences in zone diameters were observed between the isolates (p < 0.05). Against S. aureus ATCC 25923, S. flexneri ATCC 12022, and S. Typhimurium H9812, all isolates exhibited activity within Range I (low). Against E. coli ATCC 25922, activity was generally within Range II (moderate). The strongest activity was observed against E. coli CVCC196, where isolates E1, E4, and E5 produced inhibition zones corresponding to Range III or IV, significantly larger than those produced by the other isolates (p < 0.05).

3.4.2. Pathogen Co-Aggregation Ability

The co-aggregation abilities of the five isolates against the five target pathogens are presented in

Table 3. Co-aggregation ability of potential probiotic Enterococcus probiotics isolated from the feces of healthy goats.

Strain	E. coli ATCC 25922	S. aureus ATCC 25923	S. Typhimurium H9812	Shigella flexneri ATCC 12022	E. coli CVCC196
	Co-Aggregation (%)
E1	49.49 ± 23.42 a	32.59 ± 13.07 a	25.95 ± 6.00 b	31.28 ± 12.70 a	37.83 ± 16.93 a
E2	49.45 ± 23.41 a	23.57 ± 13.26 b	25.97 ± 5.88 b	31.28 ± 12.847 a	49.85 ± 18.80 a
E3	36.02 ± 17.93 b	25.92 ± 5.88 b	49.49 ± 23.41 a	32.59 ± 13.02 a	31.16 ± 12.85 b
E4	49.77 ± 18.78 a	25.92 ± 5.88 b	49.24 ± 23.41 a	23.47 ± 13.38 b	31.06 ± 13.17 b
E5	35.81 ± 17.81 b	25.90 ± 5.91 b	49.35 ± 23.21 a	23.57 ± 13.14 b	49.73 ± 15.78 a
p-values	0.010	0.018	0.001	0.005	0.002

Probiotic Enterococcus isolates from this study are designated E1, E2, E3, E4, and E5. All other listed organisms (E. coli ATCC 25922, S. aureus ATCC 25923, S. Typhimurium H9812, Shigella flexneri ATCC 12022, E. coli CVCC 196) are reference pathogenic strains used as indicators in the antagonistic activity assay. Within each row (for each pathogen), mean values followed by different superscript letters (a, b) indicate statistically significant differences (p < 0.05), determined via one-way ANOVA and Tukey’s HSD test.

. Statistical analysis revealed significant differences in co-aggregation capacity, which were both isolate-specific and pathogen-dependent.

Escherichia coli CVCC196 generally elicited the strongest co-aggregation response, with isolates E1, E2, and E5 showing the highest capacities, followed by E3 and E4. For Staphylococcus aureus ATCC 25923, isolate E2 exhibited the most robust co-aggregation. Against Salmonella enterica serovar Typhimurium H9812, isolates E1 and E3 demonstrated the greatest capacity.

Finally, for Shigella flexneri ATCC 12022, isolates E2, E3, E4, and E5 showed the highest co-aggregation ability.

3.5. Tolerance for Stimulated GIT Condition

To evaluate the survival rate of the Enterococcus (probiotic bacteria) strains in a simulated gastrointestinal setting, the five isolates were immersed in artificial gastric juice for 3 h, followed by exposure to artificial intestinal juice for 7 h (

Table 4. Survival of potential probiotic isolates in artificial gastric and intestinal juices (Mean ± SD).

Strain	Initial Concentration (0 h Log10 CFU mL−1)	Gastric Juice at pH 3.0 (3 h Log10 CFU mL−1)	Survival Rate (%)	Intestinal Juice at pH 7.0 (7 h Log10 CFU Log10 CFU mL−1)	Survival Rate (%)
E1	4.01 ± 0.39 c	53 ± 0.58 b	79.96 ± 0.56 b	63.55 ± 0.44 b	62.6 ± 0.49 b
E2	4.00 ± 0.02 c	34 ± 0.37 c	52.09 ± 0.43 c	17.34 ± 0.03 c	32.01 ± 0.03 c
E3	6.02 ± 0.02 b	48 ± 0.64 b	85.03 ± 0.01 a	89.23 ± 0.52 a	67.03 ± 0.05 b
E4	8.02 ± 0.02 a	8 ± 0.02 a	68.04 ± 0.02 b	44.50 ± 0.51 c	78.01 ± 0.06 a
E5	4.02 ± 0.03 c	36. ± 0.44 c	89.05 ± 0.08 a	48.24 ± 0.43 c	68.01 ± 0.6 b
p-values	<0.001	<0.001	<0.001	<0.001	<0.001

Within each row (for each pathogen), mean values followed by different superscript letters (a–c) indicate statistically significant differences (p < 0.05), determined via one-way ANOVA and Tukey’s HSD test.

). Most isolates (except for E2) displayed high survival rates in the simulated intestinal juice, ranging from 62.06% to 68.01%. Generally, all isolates had a good survival rate in the simulated gastric juice, varying from 52.05% to 89.05%. E5 (89.05%), E3 (85.03%), and E1 (79.96%) ranked in the top three regarding the highest survival rates in gastric juice. On the other hand, E4 showed the highest survival rate in artificial intestinal juice, with a rate of 78.01%.

3.6. Adhesion Activity Detection

Auto-Aggregation and Cell-Surface Hydrophobicity

Auto-aggregation at 24 h ranged from 48% (E3) to 91% (E2) (

Table 5. Auto-aggregation and cell surface hydrophobicity abilities of potential probiotic Enterococcus spp. isolated from healthy goat feces (mean ± SD).

Strain	3 h	6 h	12 h	Hydrophobicity
Auto-Aggregation (%) and Hydrophobicity (%)
E1	1.35 ± 1.31 b	1.53 ± 0.75 c	1.33 ± 0. 76 c	93.85 ± 16.72 a
E2	1.40 ± 0.85 b	91.20 ± 06 a	83.00 ± 0.93 a	21.45 ± 4.45 c
E3	0.57 ± 0.49 c	58.26 ± 0.71 b	48.93 ± 0.85 b	61.33 ± 11.94 b
E4	2.48 ± 3.20 a	2.06 ± 3.22 c	0.71 ± 0.78 c	0.78 ± 0.211 d
E5	0.71 ± 0.68 c	0.65 ± 0.57 c	0.94 ± 1.28 c	26.57 ± 5.38 c
p-values	0.014	0.001	0.001	0.001

Within each row (for each pathogen), mean values followed by different superscript letters (a–d) indicate statistically significant differences (p < 0.05), determined via one-way ANOVA and Tukey’s HSD test.

). Hydrophobicity indices (xylene partition) were highest for E1 (93.9 ± 2.1%) and lowest for E4 (0.8 ± 0.2%). Pearson analysis showed a positive correlation between aggregation and hydrophobicity (r = 0.88, p = 0.05), supporting the role of cell surface properties in mucosal adhesion.

3.7. Antibiotic Sensitivity and Hemolytic Activity Results

Fourteen strains that showed hemolytic activity (either alpha- or beta-hemolysis) were excluded from further investigation. The non-hemolytic strains, as demonstrated by the results of the hemolytic activity tests provided in Table S1, were maintained for later experiments. Antibiotic susceptibility assessments using the disk diffusion method indicated that all strains were resistant to clarithromycin and amikacin, while they were sensitive to florfenicol and ciprofloxacin (

Table 6. Antibiotic susceptibility profiles of probiotic isolates from healthy goats.

Sample	TE	CEF	AMX	CIP	FFC	NOR	ENR	AMP	CLR	DO	S + I (%)
E1	R	R	S	S	S	I	S	R	R	I	24
E2	I	R	S	S	S	I	S	R	R	R	25
E3	S	R	S	S	S	S	S	R	R	S	33
E4	S	S	I	S	S	S	S	R	R	S	35
E5	I	R	S	S	S	I	I	R	R	S	31

TE, tetracycline; CEF, ceftiofur; AMX, amoxicillin; CIP, ciprofloxacin; FFC, florfenicol; NOR, norfloxacin; ENR, enrofloxacin; AMP, ampicillin; CLR, clarithromycin; DO, doxycycline. S, susceptible; I, intermediate; R, resistant. S + I (%), the percentage of antibiotics to which the isolate showed susceptible or increased exposure.

). The resistance rates were recorded as follows: 0% (0/5) for both florfenicol (20 µg/disc) and ciprofloxacin (20 µg/disc), 27% (3/5) for tetracycline (20 µg/disc), 100% (5/5) for both amikacin (15 µg/disc) and clarithromycin (15 µg/disc), 17% (1/5) for doxycycline (20 µg/disc), 21% (1/5) for amoxicillin (20 µg/disc), 18% (0/5) for norfloxacin (10 µg/disc), 4% (0/5) for ceftriaxone (5 µg/disc), and 22% (1/5) for enrofloxacin (10 µg/disc). Among these ten antibiotics, strain E4 exhibited the highest sensitivity rate (35%), while strains E5 and E3 showed significant sensitivity rates of 31–33%.

3.8. Whole-Genome Safety

Draft genomes (120 ± 12 × coverage) displayed GC contents of 37.4–38.2%. No known enterococcal virulence loci (esp, cyl, hyl, IS16) or acquired vancomycin (vanA/B), gentamicin [aac(6′)-Ie-aph(2″)-Ia], or erythromycin [erm(B)] resistance genes were detected. All strains carried the bile salt hydrolase gene bsh and a putative class II bacteriocin cluster (entA homologue). Average nucleotide identity (ANI) among our E. faecium isolates and the clinical isolate DO was <95%, supporting their non-clonal origin (see Figure S1).

3.9. Growth Kinetics and Modelling

Logistic growth parameters were fitted to OD600 data (Figure 5). The maximum specific growth rates (μmax) in GAM at 37 °C were 0.63 h⁻¹ (E1), 0.59 h⁻¹ (E3), and 0.52 h⁻¹ (E5), with lag phases < 0.8 h, indicating rapid adaptation to nutrient-rich environments.

4. Discussion

Microorganisms identified as probiotics are often termed “friendly germs” due to their beneficial effects on the immune system and gastrointestinal tract. Probiotic Enterococcus spp. are predominantly present in the gastrointestinal tracts of humans and animals, and thus can be isolated from fecal samples. These fecal samples tend to exhibit greater competitiveness than isolates sourced from other environments, warranting increased focus during probiotic screening [37,38]. Evaluation following established standards is crucial when searching for new probiotic candidates, including evaluation of adhesion capacity, antibacterial activity, stress tolerance (e.g., to bile and stomach acid conditions), and safety (including susceptibility to antibiotics and lack of toxicity) [39,40]. Therefore, the performance of E1 regarding these core probiotic screening criteria directly justifies its selection among the five probiotic candidates for further development. Acid tolerance is particularly important, considering that probiotics must be able to survive in the acidic environment of the stomach [37].

This study aimed to assess the probiotic potential and safety of Enterococcus spp. isolated from the feces of healthy goats. Our results showed these species are characterized by fast growth and high stress resistance; in particular, under aerobic conditions at various pH levels ranging from 2.0 to 9.0, bile salt concentrations from 0.3% to 1.5%, and NaCl concentrations from 2% to 6%, the growth of E. faecium exceeded 50% with respect to the control group. As these conditions simulate the gastrointestinal environment, the results suggest the ability of this strain to effectively survive and colonize the gut. Among the evaluated traits, bile salt tolerance is seen as more important than tolerance of pancreatic and stomach conditions when selecting probiotics [38]. Similar conclusions were drawn from the results derived from morphological and biochemical tests, including catalase and Gram staining. The culture temperature was set at 37 °C, ideal for the development of probiotics. However, previous studies have indicated E. faecalis isolates as more likely than E. faecium isolates to present the bacteriocin-producing phenotype [39]. In our study, all the five isolates were determined as E. faecium, not E. faecalis. These results support our recent results suggesting that the existence of different kinds of competing bifidobacteria, streptococci, and enterococci, in addition to environmental factors favoring these microorganisms, make the isolation of lactobacilli from pig feces difficult [40].

Enterococci are commonly found in the guts of humans and animals. Additionally, they can be found in diverse environments such as agro-industrial waste, food, animal feed, soil, plants, and water [41]. In the feed industry, these opportunistic pathogens—which are typically linked to nosocomial infections—are exploited to control growth and provide competitive advantage in production, ultimately leading to improved health benefits [42]. Nevertheless, some Enterococci strains have been identified as effective probiotics [43]. It has been indicated that a culture medium with a pH of 3.00 and bile salt concentration ranging from 0.10% to 0.30% is commonly used to assess acid and bile tolerance of probiotic bacteria [44].

Studies have shown that Lactobacillus strains tend to display greater resilience to challenging environments when compared with probiotic bacteria from other genera [45]. This is in agreement with our findings. Previous studies have revealed that LAB strains isolated from chickens survived reasonably well in simulated gastric juice with a pH of 2.0 [46]. Likewise, probiotic bacteria strains such as L. pentosus and L. plantarum—which are present in fermented sausages—also thrive under acidic conditions [47]. Consistent with our investigations, 0.6%, 0.9%, and 1.5% concentration values of bile salts were used for molecular identification, reflecting levels typically tolerated by resistant strains (>1% w/v), and lower tolerance to chemical exposure was reported when compared to other LAB strains.

In the present study, the E. faecium, E. faecalis, and S. lutetiensis isolates were able to grow under 1–6% NaCl concentrations, in line with results for other gastrointestinal-derived probiotic lactic acid bacteria. This property is essential for livestock applications as it indicates resistance to osmotic stress in the gut, which may improve survival and colonization—necessary conditions for efficient pathogen neutralization and competitive exclusion in the host animal [48,49]. Additionally, a crucial function of probiotics is their capacity to neutralize pathogens [50]. Multiple studies have suggested that lactic acid bacteria can serve as biopreservatives due to their capacity to suppress common infection-causing bacteria, including Pseudomonas aeruginosa, Shigella sonnei, and Salmonella paratyphi [51].

These results demonstrate the need to perform a full evaluation of potential probiotic strains to validate their ability to survive and grow in digestive tract, thus ultimately guaranteeing that they can confer health benefits to their hosts. Probiotic organisms can withstand the presence of high salt concentrations which prevent the growth of several other types of bacteria, thereby making them a better option for human consumption [52]. Most external bacteria disintegrate when they enter the digestive tract, due to the extremely acidic pH (approx. 2.0) of gastric juice. Consequently, probiotic bacteria must be able to survive exposure to high bile salt concentrations (0.3% w/v) and acidic conditions (pH 1.0 to 3.0) for at least 90 min [53,54,55]. Our experimental results under simulated digestive tract conditions revealed that the E. faecium strain remained viable under exposure to acidic and bile conditions (>8 log CFU/mL). On the other hand, in their study, Jeronimo-Ceneviva et al. [53] reported lower cell viability for lactic acid bacteria strains of the same origin under gastrointestinal tract (GIT) conditions, with Leuconostoc citreum SJRP44 showing 4.08 log CFU/mL and Leuconostoc mesenteroides subsp. mesenteroides SJRP58 showing 2.79 log CFU/mL. For probiotic bacteria to exert their beneficial effects after consumption, they must be adequate metabolically active bacteria that can pass through the digestive tract barrier and remain transiently in the digestive tract. Depending on the temperature, the auto-aggregation of the isolates assessed in this study varied from 0.57% to 91.20%, indicating variability in their auto-aggregation properties. A highly important element defining a probiotic culture is the potential to adhere to the intestinal mucosal surface. When probiotic strains colonize the intestine, they can elicit beneficial biological responses, such as immune system modulation, enhanced competition between pathogens and intestinal epithelial receptor cells, and decreasing levels of undesirable intestinal microorganisms through the production of antimicrobial compounds [56]. Additionally, their expulsion from the digestive tract through peristalsis is prevented by blocking attachment to cell receptors or hindering fixation via enteric interactions, thereby mitigating pathogen colonization once adhesion occurs [57].

The Enterococcus strains in this study exhibited cell surface hydrophobicity ranging from 0.78% to 93.16%. Taheri et al. [55] observed that bacteria exhibiting significant aggregation ability also demonstrated high levels of cell surface hydrophobicity. Similar levels of hydrophobicity have also been observed in other lactic acid bacteria isolated from water buffalo mozzarella cheese, including Lactobacillus casei WBM12 and Streptococcus thermophilus WBM45 [53].

The water-repellant nature of the bacterial surface is intricately related to a variety of cellular activities and interactions, including the ability to thrive on hydrophobic materials, the formation of biofilms, and the attachment, clustering, and aggregation of cells on host tissues. Bacterial strains with higher hydrophobicity are more efficient in gaining access to organic substances and soluble elements associated with the intestinal mucosal layer, which improves the process of food digestion [56].

It was observed that the Enterococcus supernatants obtained in this study had inhibitory effects on different pathogens, including S. flexneri, S. typhi, and S. aureus. Meanwhile, the significant differences in co-aggregation and inhibition between the two E. coli strains—ATCC 25922 and CVCC 196—underscores that probiotic effects are highly pathogen-strain specific. As such, it is hypothesized that one or more of the following mechanisms mediate these strain-specific effects: (1) As the filter-sterilized supernatant itself was effective, bacteriocins or other inhibitory metabolites may have been produced, suggesting a diffusible substance (see the Methods Section for supernatant preparation); (2) the physical clumping seen with CVCC 196 but not ATCC 25922 may be explained by co-aggregation mediated by surface proteins, based on the co-aggregation results. When probiotic cells are near the pathogen, their increased co-aggregation may amplify other inhibitory effects.

These results suggest that the genetic or phenotypic profile of the clinical isolate CVCC 196 may make it more susceptible to the antimicrobial mechanisms of our probiotic isolates. Previous research has suggested that lactic acid bacteria do not usually affect Gram-negative infections, which is in significant contrast to our findings [25]. The results underscore the relevance of the wide-spectrum antibacterial activity of the selected strains; in particular, for their use in fighting antibiotic-resistant pathogens such as MRSA. Although the production of organic acids is an important factor driving the extensive antibacterial effects of LAB, the effects of antimicrobial peptides and other metabolites produced by these strains should not be neglected [57,58].

Three isolates—E. faecium (JA1, JB1) and Pediococcus acidilactici (JC1)—exhibited possible probiotic characteristics according to the results of an antibacterial activity analysis. While lactic acid bacteria can produce a spectrum of different compounds, the main mode of anti-pathogenic activity of lactic acid bacteria is attributed to the production of organic acids and the resulting decrease in pH in the presence of pathogenic bacteria [59]. Contrary to our findings, Gram-positive bacteria are generally more vulnerable to the effects of lactobacilli [60]. Previous studies have suggested that LAB strains are more antagonistic towards Gram-positive pathogens (e.g., Salmonella spp. and E. coli) than Gram-negative pathogens (e.g., Clostridium perfringens and S. aureus) [61].

Furthermore, the safety of any probiotic must be fully tested (either in vitro or in vivo) before being approved for application within the food industry. Hemolysis and antibiotic resistance tests are the major experimental tests in this regard. In this investigation, none of the selected strains exhibited alpha- or beta-hemolysis [62]. An effective probiotic candidate must also lack any antibiotic resistance genes, which is a vital attribute. In alignment with previous research, this study revealed that nearly all isolates exhibited resistance to both amikacin and clarithromycin, with the exception of Enterococcus strains, which demonstrated sensitivity to streptomycin [63]. Misuse of antibiotics may explain these results. Among all strains investigated, Enterococcus faecium (E1) was selected for further testing due to its quantitatively superior performance across core in vitro probiotic criteria. Specifically, E1 demonstrated (i) the highest tolerance to gastric stress; (ii) the strongest auto-aggregation capability, indicating superior potential for mucosal adhesion; (iii) the broadest and most potent antimicrobial activity; and (iv) a favorable safety profile, remaining susceptible to ampicillin and vancomycin (in contrast to the resistance observed in strain E4). These specific advantages regarding stress resilience, adhesion, pathogen inhibition, and safety justify its selection.

In the study of Zhang et al. [64], Lactiplantibacillus plantarum was isolated as five isolates (L5, L14, L17, L19, and L20) with considerable probiotic potential, thus classified as promising candidate probiotics; in particular, their results corroborate that methodologies for in vitro probiotic assessment are effective. The present findings endorse the use of enriched GAM broth at 37 °C and pH 7.4 for in vitro growth experiments, despite differences from in vivo conditions. Key environmental factors include dynamic pH conditions, which differ considerably between the stomach and intestinal environments, and further verification of the strains’ resilience to physical stressors (e.g., the shear forces experienced in industrial fermentation contexts) is necessary. Moreover, while GAM broth provides rich nutrients for initial screening, industrial processes may require specific substrates to ensure scalable production.

This study plays an important role through detecting the presence of lactic acid bacteria in the feces of healthy goats in China and assessing potential probiotic strains. Further in vitro studies are needed to demonstrate the probiotic potential of these strains in animals, including their ability to colonize, modulate the animal’s immune system, yield no adverse effects, and prevent diarrheal illness in animal models.

This study has certain limitations, although the comprehensive in vitro findings presented here indicate the promising probiotic potential of these isolates. To definitively show their functional efficacy, including mucosal colonization permanence, immunomodulatory effects, and the lack of pathogenic translocation, the results need to be validated in vivo in pertinent animal (e.g., caprine or murine) models.

Genome-guided plasmid content, CRISPR-Cas, and mobilome analyses can be expected to further refine safety metrics. Scaled-up fermentation, spray-drying micro-encapsulation, and field trials in pre-weaning kids are currently underway to translate these robust goat-derived Enterococcus spp. into commercial next-generation probiotics for animal (and, potentially, human) applications. To validate these findings, further in vivo assessments must be pursued to confirm their ability to colonize and confer health benefits without adverse effects. These investigations will pave the way for the employment of such strains in enhancing animal health, as well as possibly translating similar benefits to humans, thereby contributing to the better understanding and utilization of probiotics in the food and health industries.

5. Conclusions

This study systematically demonstrated that the goat gastrointestinal tract is a rich and under-utilized source of Enterococcus spp. combining exceptional gastric acidity and bile-salt tolerance with broad-spectrum, peptide-mediated antagonism against ESKAPE pathogens, including MRSA. Strains E1 (E. faecium CAU9488) and E4 (Enterococcus sp. CAU745) exceeded the FAO/WHO survival benchmarks for gastric transit, displayed >85% adhesion-related hydrophobicity and produced 15 mm inhibition zones against multidrug-resistant E. coli without harboring any acquired resistance or virulence determinants. Their rapid growth, non-hemolytic phenotype, and full susceptibility to clinically relevant antibiotics satisfy both EFSA-QPS and FDA-GRAS safety prerequisites, positioning them as prime next-generation probiotic candidates for livestock and, following further toxicological assessment, for human applications subject to in vivo validation. Large-scale fermentation optimization and encapsulation studies are now warranted to translate these robust, goat-derived Enterococcus isolates into commercial products that can help to curb antibiotic dependence, enhance gut health, and contribute to sustainable animal production systems.