Goat-derived Lactobacillus amylovorus improved floppy kid syndrome via regulating gut microflora

Abstract

Background

Floppy Kid Syndrome (FKS) severely restricts goat farming due to high mortality from metabolic disturbances and gut dysbiosis. Here, we aimed to isolate a goat-derived probiotic and evaluate its capacity to restore gut homeostasis in FKSaffected goats.

Results

Lactobacillus amylovorus isolates was obtained via selective colony morphology, Gramstain and catalase testing, and confirmed by 16 S rRNA sequencing (≥ 98% identity). The isolate survived at pH 2.0 (72%) and pH 3.0 (85%), tolerated 0.3% bile salts (survival > 10%), and retained > 40% viability after 60 s at 60 °C. In disk diffusion assays, inhibition zones averaged 16 mm against Escherichia coli and 12 mm against Staphylococcus aureus. In FKSaffected goats receiving 1 × 10⁸ CFU/day for 3 days, Shannon diversity increased by 25% (p < 0.05), Simpson index by 18% (p < 0.05), and shared species with healthy controls rose from 8 to 17. Relative abundance of EscherichiaShigella declined by 40%, while Lactobacillus and Ruminococcus increased by 30% and 22%, respectively (p < 0.05).

Conclusions

Goat-derived L. amylovorus displays robust acid, bile and heat tolerance, inhibits key pathogens, and effectively regulates gut microbiota in FKSaffected goats. These findings support its promise as a novel probiotic intervention for preventing and treating Floppy Kid Syndrome.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12917-025-04967-7.

Introduction

The gut microbiota refers to the microbial community residing in the digestive tract of animals, comprising bacteria, fungi, viruses, and other microorganisms [1]. These microbes play essential roles in food digestion, vitamin synthesis, and immune regulation, as well as in maintaining the stability of the gut environment. Generally, the gut microbiota can be categorized into three groups: beneficial bacteria, pathogenic bacteria, and conditional pathogenic bacteria [2, 3]. Under a balanced microbial state, the host can sustain normal physiological functions. However, when dysbacteriosis occurs (characterized by a significant reduction in beneficial bacteria and an overgrowth of pathogenic and conditional pathogenic bacteria) the gut barrier may be compromised, allowing pathogens to invade the host and disrupt the homeostasis between the microbiota and internal environment, thereby leading to various diseases [4]. The microbial composition varies along different sections of the gut tract, with Firmicutes, Bacteroidetes, and Proteobacteria generally dominating the gut microbiota. Furthermore, studies have demonstrated that anaerobes account for over 98% of the total gut microbiota, whereas aerobes and facultative anaerobes constitute only a minor fraction [5, 6]. Due to environmental constraints, the microbial diversity and abundance in the stomach and proximal intestine are relatively low. However, as the pH decreases in the distal intestine, microbial diversity and richness increase significantly, with studies showing that bacterial abundance in the cecum can be up to ten times higher than in the proximal gut [7–10].

The balance of gut microbiota is closely linked to host health. Recent studies have indicated that gut microbiota diversity is strongly associated with the occurrence of cardiovascular diseases, certain cancers, inflammatory bowel disease, type 1 diabetes, and allergic disorders [11–23]. Notably, probiotics exhibit significant protective effects by inhibiting pathogenic bacteria, modulating immune responses, and maintaining gut barrier integrity. Their metabolic products, such as short-chain fatty acids, have even been shown to possess anticancer properties [15, 24]. In light of the growing concern over antibiotic overuse, probiotics have emerged as promising alternatives due to their safety profile and lack of antibiotic resistance.

Lactic acid bacteria (LAB) are widely recognized for their probiotic properties. Since Pasteur first discovered and named them in 1857, LAB have been classified into various genera, including Pediococcus, Streptococcus, Lactobacillus, Bifidobacterium, and Leuconostoc [25]. LAB contribute to gut homeostasis, facilitate digestion and nutrient absorption, enhance immune responses, and inhibit the growth of pathogenic bacteria through mechanisms such as competitive exclusion, secretion of antimicrobial compounds (e.g., organic acids, bacteriocins, and hydrogen peroxide), and interference with pathogen colonization [26–35]. Moreover, LAB can regulate innate and adaptive immune responses, reduce inflammatory cytokine levels, reinforce gut barrier function, and even exhibit antiviral properties [36–42]. Given these beneficial attributes, the International Scientific Association for Probiotics and Prebiotics (ISAPP) stated in 2014 that appropriate probiotic supplementation can significantly improve host health [43]. In animal husbandry, LAB have been utilized as feed additives to enhance calf immunity, prevent diarrhea, improve piglet gut microbiota composition, and increase the quality of silage, demonstrating broad application prospects [44–50].

However, the global overuse of antibiotics has driven the emergence of virulent multidrug-resistant (MDR) bacterial pathogens, severely limited treatment options and raising morbidity and mortality worldwide [51]. This public health crisis has spurred intense interest in effective non‐antibiotic strategies, with probiotics emerging as promising alternatives to counter MDR infections and safeguard both animal and human health [52].

L. amylovorus, a key member of the Lactobacillus genus, was first isolated by Nakamura from cattle farm waste and fermented corn mixtures [53]. As a Gram-positive bacterium, L. amylovorus forms convex, smooth, and round colonies with a white, opaque appearance and regular edges. Under the microscope, L. amylovorus appears as single or in chains, lacks spores. L. amylovorus thrives optimally at pH 5.5–6.0 and can survive in highly acidic conditions (pH 3.0–4.5), demonstrating exceptional acid tolerance among non-spore-forming bacteria. L. amylovorus adheres robustly to gut epithelial surfaces, produces digestive enzymes that enhance starch breakdown, and secretes bacteriocins to inhibit pathogens including Salmonella, Escherichia coli and Staphylococcus aureus. It also upregulates host defense peptides and tight-junction proteins, thereby reinforcing barrier integrity and promoting nutrient absorption, highlighting its potential role in promoting overall host well-being [54–56].

Floppy kid syndrome (FKS) is a metabolic disorder disease in newborn goats caused by gut dysbiosis, characterized by an increased abundance of pathogenic bacteria and a decline in probiotic bacteria, leading to hypoglycemia and acidosis. Metagenomic sequencing has revealed a significantly higher relative abundance of L. amylovorus in the gut tract of healthy goats compared to FKS-affected goats, suggesting that L. amylovorus may play a crucial role in maintaining gut microbial balance, facilitating glucose absorption, and preventing disease onset [57]. Therefore, this study aims to isolate and identify L. amylovorus from healthy goat feces, characterize its growth, stress-tolerance and antimicrobial properties in vitro, and evaluate its capacity to regulate gut microbiota balance in FKS-affected goats, providing a theoretical and practical basis for its probiotic application.

Materials and methods

Sample collection

Contents from the duodenum of healthy goats were collected aseptically, stored in sterile centrifuge tubes, and frozen at − 80 °C. Upon processing, samples were thawed, serially diluted in sterile PBS, and plated on MRS (DeMan, Rogosa and Sharpe) solid medium. Plates were cultured anaerobically at 37 °C for 48 h. Colonies with morphology characteristic of L. amylovorus (white, convex, smooth edges, ~ 2–3 mm diameter) were picked and purified by continuous streaking procedure on MRS solid medium, selected a single colony and inoculated it onto MRS liquid medium for cultured. Pure isolates were confirmed as Gram-positive before further identification. Per liter, MRS liquid medium comprises peptone 10.0 g, beef extract 10.0 g, yeast extract 4.0 g, glucose 20.0 g, dipotassium phosphate 2.0 g, ammonium citrate 2.0 g, sodium acetate 5.0 g, magnesium sulfate 0.2 g, manganese sulfate 0.04 g, and Tween 80 1.0 g; and MRS solid medium comprises peptone 10.0 g, beef extract 5.0 g, yeast extract 4.0 g, glucose 20.0 g, dipotassium phosphate 2.0 g, ammonium citrate 2.0 g, sodium acetate 5.0 g, magnesium sulfate 0.2 g, manganese sulfate 0.05 g, agar 15.0 g, and Tween 80 1.0 g.

Isolate identification

Gram staining was performed to observe bacterial morphology. Among the biochemical tests, assays such as fermentation of cellobiose, mannitol, maltose, lactose, inulin, sorbitol, sucrose, salicin, and arbutin, as well as for arginine decarboxylase, lysine decarboxylase, ornithine decarboxylase, and urease activities following biochemical tube test protocols (Hope Bio-Technology Co.,Ltd, Qingdao, China) [56]. Molecular identification was performed via PCR amplification with the universal primers F (5’-AGAGTTTGATCCTGGCCTCA-3’) and R (5’-GGTTACCTTGTTTGTTACGACTT − 3’), the 50 µL PCR reaction mix contained 2 µL each of forward and reverse primers (10 µmol/L), 25 µL Premix Taq, 2 µL DNA template, and 19 µL ddH₂O; amplification was carried out with an initial denaturation at 95 °C for 5 min, followed by 35 cycles of 95 °C for 30 s, 54 °C for 30 s, and 72 °C for 45 s, then a final extension at 72 °C for 10 min. The products were then detected via agarose gel electrophoresis. Samples with the target bands were sent for sequencing (Sangon Biotech Co.,Ltd, Shanghai, China). The sequencing results were compared to the NCBI database via BLAST.

Growth characteristics of L. amylovorus

Growth curve determination

The purified bacterial was grown to the stationary phase by MRS liquid medium. The bacterial was then inoculated into MRS liquid medium at a 1% inoculum ratio and cultured under anaerobic conditions at 37 °C with shaking at 225 r/min. Every 2 h, 100 µL of the bacterial suspension was taken, and the optical density at 600 nm (OD_600) was measured using a microplate reader. A growth curve was plotted with time as the x-axis and OD_600 values as the y-axis.

Simulated artificial gastric fluid and artificial intestinal fluid tolerance test

Prepared artificial gastric fluid by dissolving 10 g pepsin in 800 mL deionized water, adding 16.4 mL concentrated HCl, mixing, and bringing to 1 L; adjust pH to 2.0 or 3.0 and sterilize by 0.22 μm filtration. For the test, 10 µL of log‑phase L. amylovorus culture was inoculated into 990 µL MRS liquid medium and cultured 16 h, then 100 µL of this culture (10% v/v) was mixed with 900 µL artificial gastric fluid and cultured at 37 °C for 2 h. After cultured, samples were serially diluted (10^–4–10^–5) and plated on MRS solid medium; colonies were counted after 24 h anaerobic incubation, and survival rate calculated.

Prepared artificial intestinal fluid by dissolving 6.8 g KH₂PO₄ in 500 mL deionized water, adjusting pH to 6.8, diluting to 1 L, adding 1.0 g trypsin per 100 mL, and sterilizing by 0.22 μm filtration. AIF was prewarmed to 37 °C. Following the same protocol as for artificial gastric fluid, 100 µL of log‑phase culture (10% v/v) was cultured in 900 µL AIF at 37 °C for 2 h, then diluted, plated, and counted to determine survival.

Bile salt tolerance test

A 10% bile salt stock solution was prepared and added to MRS liquid medium to achieve final bile salt concentrations of 0.2%, and 0.3%. L. amylovorus in the logarithmic phase was inoculated into the medium at a 1% ratio and cultured at 37 °C for 2 h. After culture, the bacterial suspension was serially diluted, and 10⁻⁴–10⁻⁵ dilutions were inoculated into MRS solid medium. After 24 h of culture, viable bacterial counts were determined, and survival rates were calculated.

Heat tolerance test

MRS liquid medium was prepared and aliquoted into test tubes, which were preheated to 80 °C, 70 °C, 60 °C, and 37 °C, respectively. L. amylovorus in the logarithmic phase was added to each tube at a 1% inoculum ratio, cultured at the designated temperatures for 60 s, and then rapidly cooled in an ice-water bath. After culture, the bacterial suspension was serially diluted, and 10⁻⁴–10⁻⁵ dilutions were inoculated into MRS solid medium. After 24 h of culture, viable bacterial counts were determined, and survival rates were calculated.

Antibiotic sensitivity test

Antibiotic sensitivity testing of L. amylovorus was performed using the disk diffusion method according to the standards of the Clinical and Laboratory Standards Institute (CLSI). The specific procedures were as follows: The purified L. amylovorus was inoculated into BHI liquid medium and cultured under anaerobic conditions at 37 °C with shaking at 220 r/min for 16 h. The bacterial suspension was then adjusted to a concentration of 0.5 McFarland standard (1.5 × 10⁸ CFU/mL). Under a clean bench, 100 µL of the diluted bacterial suspension was pipetted and evenly spread onto the surface of a 90 mm MH agar (Hope Bio-Technology Co.,Ltd, Qingdao, China) plate using a disposable spreader. The plate was left at room temperature for 10–15 min until the bacterial inoculum layer dried. Bacterial susceptibility paper was then placed onto the agar surface, ensuring a minimum distance of 24 mm between bacterial susceptibility papers, the diameter of bacterial susceptibility papers is 6.35 mm. The plates were cultured in an inverted position under anaerobic conditions at 37 °C for 18–24 h. After culture, the diameter of the bacteriostatic circle was measured using a vernier caliper, and the results were recorded according to CLSI guidelines (Supplementary Material 1 A).

In vitro antibacterial test

This study evaluated the antibacterial activity of L. amylovorus against Escherichia coli and Staphylococcus aureus using the Oxford cup diffusion method (E. Coli and S. aureus isolated from the FKS-affected goats previously [58]). E. coli and S. aureus suspensions (10⁵–10⁶ CFU/mL) were prepared and 100 µL was spread evenly on LB agar plates. Sterile Oxford cups (inner diameter 6 mm) were gently pressed into the agar to form wells. Each well was filled with 200 µL of 18 h anaerobic culture of L. amylovorus. Plates were pre-cultured at 4 °C for 8–10 h for diffusion, then cultured anaerobically at 37 °C for 24 h. After cultured, the total diameter of the inhibition zone (including the 6 mm well) was measured with a vernier caliper and recorded.

Probiotic effects of L. amylovorus on the gut microbiota of goats

Clinical observation of symptoms

The condition of each goat was observed before and after L. amylovorus gavaged, to determine whether the goats were affected by FKS. The specific symptoms of the FKS-affected goats are as follows: mental depression, slow heart rate, cool ears and nose, weakness in the limbs (sometimes with the forelimbs knelt or in a “V” shape), difficulty suckling, and abdominal bloating.

Preparation of bacterial inoculum

L. amylovorus was cultured anaerobically in MRS liquid medium at 37 °C. A logarithmic-phase bacterial suspension was inoculated into fresh MRS liquid medium at a 1% inoculation ratio. Based on the bacterial growth curve, the culture was incubated for 14 h to reach the maximum bacterial density. The bacterial suspension was stored at 4 °C until use. Only viable bacteria were administered during the animal experiments.

Animal grouping and treatment

Nine newborn goats (5–6 days old) were randomly selected from the same farm, including three goats affected with FKS and six healthy goats of similar age. The animals were divided into three experimental groups: FKS-affected goats + L. amylovorus treatment group (T Group): Three FKS-affected goats received 10 mL of L. amylovorus suspension (1 × 10⁸ CFU/mL) daily for three consecutive days; Healthy goat + L. amylovorus treatment group (H Group): Three healthy goats received 10 mL of L. amylovorus suspension (1 × 10⁸ CFU/mL) daily for three consecutive days; Healthy control group (J Group): Three healthy goats received no treatment.

Sample collection and processing

Rectal swabs were collected from all goats on days 1, 3, and 5 of the trial. Sterile cotton swabs were used to collect fecal samples from the rectum and transferred to sterile centrifuge tubes. To ensure sufficient material for analyses, at least 1 g of fecal sample was collected per tube. Each group had three replicates per time point. Samples were designated based on treatment and time points as follows: FKS-affected goats + L. amylovorus treatment group: T1D, T3D, T5D; Healthy goat + L. amylovorus treatment group: H1D, H3D, H5D; Healthy control group: J1D, J3D, J5D.

16 S rRNA sequencing analysis

Extracting DNA from the rectal swab samples using a DNA extraction kit (TianGen, China). The V3–V4 hypervariable region of the bacterial 16 S rRNA gene was amplified using the universal primers F (5′-CCTAYGGGRBGCASCAG-3′) and R (5′-GGACTACNNGGGTATCTAAT-3′). After PCR amplification, the target bands were purified and recovered using the Universal DNA purification kit (TianGen, China). Library construction was performed using the NEBNext^® Ultra DNA Library Prep Kit, followed by high-throughput sequencing on the Illumina platform. The raw sequencing output (“Raw Data”) inevitably contains a proportion of lowquality or contaminant reads (“Dirty Data”). To ensure that downstream analyses are accurate and reliable, we first merge pairedend reads and apply quality filtering to remove these contaminants, yielding “Clean Data.” Next, we denoise the Clean Data using DADA2 or deblur to generate the final set of Amplicon Sequence Variants (ASVs). For each ASV, its representative sequence is taxonomically annotated to provide specieslevel identification and corresponding abundance profiles. In parallel, we calculate ASV abundances, compute αdiversity metrics, and construct Venn diagrams to quantify withinsample diversity and to identify shared versus unique ASVs across different samples or treatment groups. Finally, we perform Principal Coordinate Analysis (PCoA) to assess and visualize differences in microbial community structure between groups.

2.4. Statistical analysis

Statistical analysis was performed using IBM SPSS Statistics 26 (IBM Corp., Armonk, NY, USA) and GraphPad Prism 8 (GraphPad Software, San Diego, CA, USA). Differences between multiple groups were evaluated by one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test. T-test and Wilcoxon rank-sum test were applied for comparisons between two groups, α-diversity comparisons among three groups used Kruskal–Wallis. β-diversity significance was assessed by PERMANOVA, p < 0.05 was considered statistically significant.

Results

In this study, we aimed to evaluate the probiotic potential of goat-derived Lactobacillus amylovorus and its capacity to restore gut microbial balance in Floppy Kid Syndrome (FKS). We hypothesized that isolates of L. amylovorus would exhibit strong stress tolerance (acid, bile, heat), possess antimicrobial activity against key pathogens, and, when administered to FKSaffected goats, shift their gut microbiota toward a healthier composition. To test this, we first isolated and characterized the growth and stresstolerance traits of L. amylovorus in vitro, conducted antibiotic and antibacterial assays, then performed a controlled gavage trial in FKS goats with subsequent 16 S rRNA community profiling to assess changes in diversity and taxonomic composition.

Isolation and identification results of L. amylovorus

The isolates colonies appeared as circular, slightly raised, milky white colonies with a diameter of approximately 2–3 mm on MRS plates (Fig. 1A). Isolates exhibited gram- positive staining. Positive results were observed for fiber disaccharides, mannitol, maltose, lactose, inulin, sorbitol, sucrose, salicin, and arginine decarboxylase, whereas negative results were recorded for urease, lysine decarboxylase, and ornithine decarboxylase. PCR amplification of the bacteria was performed, and the products were analyzed via 1.5% agarose gel electrophoresis. The amplified bands were approximately 1500 bp long, which is consistent with the expected length of the primers used. BLAST analysis of GenBank revealed over 98% homology with known sequences of L. amylovorus. These results confirmed the identity of the isolates as L. amylovorus.

Fig. 1.

A Growth characteristics of L. amylovorus on MRS solid medium; B Growth curves of L. amylovorus

Growth characteristics of L. amylovorus

L. amylovorus entered the logarithmic growth phase after 6 h of culture, reached the stationary phase at 14 h, and gradually entered the death phase after 22 h (Fig. 1B).

L. amylovorus exhibited a survival rate of 84.6% at pH 3.0 and 73.07% at pH 2.0, indicating a degree of acid tolerance, with survival rates above 70%. After 2 h in artificial intestinal fluid, over 95% survival was observed, suggesting minimal impact of the artificial intestinal fluid on the bacteria. The survival rates of L. amylovorus decreased with increasing temperature. At 60 °C, the survival rate was 42.14%, but it decreased sharply to 3.07% at 70 °C, and no survival was observed at 80 °C. The survival rates of L. amylovorus decreased with increasing bile salt concentration, but greater than 10% survival was still observed at 0.3% bile salt after 2 h, indicating some degree of bile salt tolerance. The L. amylovorus were resistant to norfloxacin, gentamicin, and amikacin but sensitive to amoxicillin, cefixime, doxycycline, florfenicol, clarithromycin, and erythromycin. L. amylovorus exhibited antimicrobial activity against both E. coli and S. aureus, with a stronger inhibitory effect on E. coli than on S. aureus (Supplementary Material 1B-G).

Probiotic effects of L. amylovorus on gut microbiota

All FKS-affected goats in T Group did not die and their symptoms improved.

Species rarefaction curves were plotted on the basis of OUT numbers at different sequencing depths, revealing a decreasing slope with increasing depth, indicating that no new species would be discovered even with further sequencing, thus confirming the adequacy of sequencing depth and quantity (Fig. 2A).

Fig. 2.

A Species rarefaction curve; B Alpha diversity indices across different treatment groups; C Alpha diversity indices across same treatment group; (D) PCoA plots based on genus level; E Changes in species abundance; F Species abundance venn diagram; G The relative abundance of phylum level; H The relative abundance of genus level; I Genus-Level abundance heatmap across different treatment groups; J Genus-Level abundance heatmap across same treatment groups. T: FKS-affected goats + L. amylovorus treatment group; H: Healthy goat + L. amylovorus treatment group; J: Healthy control group; 1D: Day 1 of the trial; 3D: Day 3 of the trial; 5D: Day 5 of the trial

No significant differences were observed in the alpha indices among the three groups between day 1 and day 5 (p > 0.05) (Fig. 2B). However, the overall trend on day 1 indicated that the T group had lower diversity and richness than the healthy control group. On day 5, while there were still no significant differences (p > 0.05), the H group presented greater diversity and richness than did the healthy control group, although the richness was still lower than that of the healthy control group. Comparing the alpha diversity indices between days 1 and 5 in the same treatment groups revealed no significant differences in the healthy control group. The Shannon index for the H group significantly increased (p < 0.05), and the Simpson, Chao1, and ACE indices also increased, although not significantly (p > 0.05). The T group presented significant increases in the Shannon and Simpson indices (p < 0.05), with increases in the Chao1 and ACE indices that were not significant (p > 0.05) (Fig. 2C).

Beta diversity was analyzed via nonmetric multidimensional scaling (NMDS), which provides a method for assessing differences in microbial community structure. For the same treatment group, bacterial community structures were compared over different days. In the healthy control group, there was significant overlap between days 1, 3, and 5 of the experiment, indicating little change in the microbial community structure during the trial. For the H group, partial overlap occurred between day 3 and days 1 and 5, but no overlap was observed between days 1 and 5, suggesting changes in the community structure. Similarly, in T group, partial overlap was observed between day 3 and the other days, whereas there was no overlap between days 1 and 5, indicating differences in community structure between these two days (Fig. 2D). When bacterial community structures across different treatment groups were compared on the same day, on day 1, the healthy control group and the H group were close in distance with some overlap, whereas the T group was greater in distance from the other two groups with no overlap, indicating significant differences in community structure. By day 5, there was some overlap between all groups, with the T group having a greater overlap with the healthy control group, suggesting that the microbial community structure in the FKS group had become more similar to that in the other groups.

The line graph shows that the overall number of species remained stable in the healthy control group. In H group, the number of species first decreased but then increased, with the final species count higher than day 1, totaling 204 species, indicating an overall increasing trend (Fig. 2E). The T group showed a gradual increase in species count from 121 species on day 1 to 151 species by the end of the experiment, but the total species count remained lower than that in the healthy control group. The Venn diagram shows that on day 1, only 8 species were shared between the T group and the healthy control group, with 68 endemic species. By day 5, the number of shared species had increased to 17, whereas the number of endemic species had decreased to 57. In H group, there were 109 endemic species on day 1, which increased to 116 by day 5. The number of shared species among the three groups increased from 36 species on day 1 to 53 species on day 5. These results suggest that gavaged with L. amylovorus promotes an increase in the richness of the digestive tract microbiota in FKS-affected goats (Fig. 2F).

At the phylum level, the microbial structure in the healthy control group remained stable, with the dominant phyla being Proteobacteria, Bacteroidetes, and Firmicutes. In H group, the dominant phyla were Firmicutes and Proteobacteria on day 1, but Bacteroidetes increased sharply on day 3, making Firmicutes, Bacteroidetes, and Proteobacteria the dominant phyla. On day 5, the relative abundance of Bacteroidetes slightly decreased, whereas that of Firmicutes increased. In T group, the dominant phyla were Proteobacteria, Firmicutes, and Bacteroidetes, with the relative abundance of Proteobacteria decreasing over time, whereas Firmicutes and Bacteroidetes increased (Fig. 2G).

At the genus level, a bar chart of the top ten genera according to relative abundance was generated. In the healthy control group, the relative abundance of each genus remained stable, with no significant changes; these genera were dominated by Escherichia-Shigella, Bacteroides, Butyricicoccus, and Lactobacillus, most of which are probiotic genera (Fig. 2H). In H group, the relative abundance of Butyricicoccus significantly decreased on day 3, whereas the relative abundances of Bacteroides and Lactobacillus increased. By day 5, the relative abundance of Butyricicoccus slightly increased, whereas the relative abundances of Bacteroides and Lactobacillus slightly decreased. During the experiment, the relative abundance of Escherichia-Shigella gradually decreased but remained higher than that in the healthy control group, whereas the relative abundances of Bacteroides and Lactobacillus were greater than those in the control group, with the abundance of Escherichia-Shigella being lower. In T group, Escherichia-Shigella remained the most dominant genus, but its relative abundance decreased over time, whereas Butyricicoccus and Lactobacillus slightly increased. Compared with the healthy control group, the T group presented a greater relative abundance of Escherichia-Shigella, whereas the relative abundances of Bacteroides and Lactobacillus were lower in T group than in the healthy control group.

On day 1 of the experiment, the genera with the greatest differences among the three groups were Escherichia-Shigella, Butyricicoccus, Lactobacillus, and Bacteroides. On day 5, the genera with the greatest differences were Escherichia-Shigella, Lactobacillus, and Bacteroides (Fig. 2I). When comparing different time points within the same treatment group, significant microbial differences were observed. In the H group, the most variable taxa were Escherichia-Shigella, Butyricicoccus, Lactobacillus, and Bacteroides. Escherichia-Shigella and Butyricicoccus showed an overall decreasing trend, while Bacteroides and Lactobacillus exhibited an increasing trend. In the T group, the most significantly altered taxa included Escherichia-Shigella, Bacteroides, Butyricicoccus, Clostridium sensu stricto 1, Lactobacillus, [Ruminococcus] gnavus group, and Fusobacterium. Among these, Escherichia-Shigella, Bacteroides, and Fusobacterium showed a decreasing trend, whereas Butyricicoccus, Lactobacillus, and [Ruminococcus] gnavus group exhibited an increasing trend (Fig. 2J).

Based on the differentially abundant taxa identified, individual species–level abundance variation charts were plotted. On Day 1, Escherichia–Shigella reached its highest relative abundance—over 60%—in the T group, whereas its abundance remained below 40% in both the J group and the H group. The T group exhibited a significantly higher Escherichia–Shigella abundance than the J and H groups (p < 0.05). Butyricicoccus peaked in the H group, was intermediate in the J group, and fell below 10% in the T group; however, these differences were not statistically significant (p > 0.05). Lactobacillus was most abundant in the H group—only marginally (≤ 1%) higher than in the J group—but was nearly undetectable in the T group, with both J and H groups showing significantly greater Lactobacillus abundance than the T group (p < 0.05). Bacteroides was most abundant in the J group, followed by the T group, and lowest in the H group, though the differences did not reach significance (p > 0.05). The low Bacteroides abundance in the H group may reflect competitive inhibition by Butyricicoccus, which dominated that group at roughly 30% relative abundance and may suppress Bacteroides growth (Fig. 3A).

Fig. 3.

A Major changes in bacterial species in different treatment groups (Day 1); B Major changes in bacterial species in different treatment groups (Day 5); C Major changes in bacterial species in H groups; D Major changes in bacterial species in T Groups. T: FKS-affected goats + L. amylovorus treatment group; H: Healthy goat + L. amylovorus treatment group; J: Healthy control group; 1D: Day 1 of the trial; 3D: Day 3 of the trial; 5D: Day 5 of the trial; * stands for p<0.05

By Day 5, Escherichia–Shigella remained highest in the T group, intermediate in the J group, and lowest in the H group. Compared with Day 1, these betweengroup differences in Escherichia–Shigella abundance were no longer significant (p > 0.05), and its abundance in the T group had declined to around 40%, suggesting that oral L. amylovorus treatment inhibited Escherichia–Shigella growth. Bacteroides abundance was highest in the H group, followed by the J group, and lowest in the T group; the rapid increase in Bacteroides in the H group may also reflect a stimulatory effect of L. amylovorus on Bacteroides. Butyricicoccus declined in all groups compared with Day 1, converging at roughly 10% across treatments. Lactobacillus remained most abundant in the H group, next in the J group, and lowest in the T group; moreover, the gap between the H and J groups widened from Day 1, indicating that L. amylovorus treatment further promoted Lactobacillus proliferation (Fig. 3B).

When comparing Days 1, 3, and 5 within the H group, Escherichia–Shigella showed a slight, nonsignificant decline over time (p > 0.05). Butyricicoccus dropped sharply between Days 1 and 3 (p < 0.05), then rose modestly by Day 5 but remained significantly below its Day 1 level (p < 0.05). Bacteroides increased from nearly 0% on Day 1 to a significant peak on Day 3 (p < 0.05), then declined slightly by Day 5 yet stayed significantly above the Day 1 baseline (p < 0.05). Lactobacillus rose significantly on Day 3 (p < 0.05), then dipped slightly by Day 5, returning to a level statistically similar to Day 1 (p > 0.05) (Fig. 3C). These trends suggest that L. amylovorus promotes Bacteroides and Lactobacillus growth but competes with the native dominant Butyricicoccus, thereby suppressing its abundance—hence the opposing temporal patterns between Butyricicoccus and the other two genera.

In the T group, Escherichia–Shigella steadily declined over time and reached its lowest level on Day 5, significantly lower than on Day 1 (p < 0.05). Clostridium also trended downward, falling below 1% by Day 5 (p < 0.05). Bacteroides followed the same initial risethenfall pattern observed in the H group. Butyricicoccus, in contrast, rose over time—opposite the H group trend—but its Day 5 abundance remained around 10%, with no significant change (p > 0.05). Ruminococcus exhibited an overall increasing trend, peaking on Day 5 with a significant rise from Day 1 (p < 0.05). Lactobacillus likewise rose steadily, with the most pronounced increase by Day 5 (p < 0.05) (Fig. 3D). In summary, L. amylovorus treatment in FKSaffected goats (T group) significantly reduced the relative abundance of pathogenic Escherichia–Shigella and Clostridium, while markedly increasing beneficial Lactobacillus and Ruminococcus abundances, thereby reshaping the gut microbiota toward a healthier profile.

Discussion

In this study, L. amylovorus was isolated from the duodenum samples of healthy goats using traditional culturing methods. The accuracy of the experiment was enhanced by amplifying and sequencing the isolates via PCR. Previous research indicated that the FKS-affected goats exhibited significant gut dysbacteriosis, with a significant increase in pathogenic bacteria (E. coli, S. aureus) and a significant decrease in probiotic bacteria (L. amylovorus) [57]. In contrast, the relative abundance of L. amylovorus was higher in healthy goats, suggesting its potential role in maintaining gut microbiota homeostasis. As a member of the Lactobacillus, L. amylovorus has been shown to contribute to the establishment of the digestive ecosystem in ruminants [59]. Lactobacillus species can promote gut health by inhibiting pathogenic bacteria and stabilizing the gut microbiota [60]. Moreover, studies have shown that L. amylovorus can suppress the inflammation caused by enterotoxic E. coli by inhibiting the TLR4 signaling pathway [61]. These findings support the potential of L. amylovorus as a probiotic for mitigating dysbiosis and inhibiting pathogenic bacteria in FKS-affected goats, suggesting that this probiotic could serve as a promising therapeutic tool for treating FKS.

Regarding its growth characteristics, L. amylovorus formed circular, slightly raised, milkywhite colonies of approximately 2–3 mm on MRS solid medium and stained Grampositive. Biochemical profiling showed positive fermentation of cellobiose, mannitol, maltose, lactose, inulin, sorbitol, sucrose, salicin and arginine decarboxylase, with negative results for urease, lysine decarboxylase and ornithine decarboxylase, consistent with its known probiotic characterization [53]. In liquid culture, L. amylovorus entered the logarithmic phase after 6 h of cultivation, reached the stationary phase at 14 h, and entered the decline phase after 22 h. Compared to the findings of Liu et al. [62], the growth rate of L. amylovorus in this study was relatively slower, which may be related to differences in culture conditions and genetic variation among strains. Additionally, the probiotic function of L. amylovorus in ruminants depends on its tolerance to gastric acid, intestinal fluids, and bile salts. The results showed that L. amylovorus retained over 70% viability in artificial gastric juice at pH 2.0, and its survival rate in artificial intestinal fluid was nearly unaffected, confirming its ability to persist and function in the gut tract. Furthermore, L. amylovorus exhibited good survivability at 0.1% bile salt concentration, further supporting its potential as a probiotic in the digestive system of animals. Temperature also influences the survival of probiotics. In this study, L. amylovorus maintained over 40% viability at temperatures exceeding 60 °C, indicating strong heat resistance, which is essential for its survival and functionality within goats.

Since probiotics often need to be co-administered with antibiotics during treatment, to ensure its safety in clinical applications, the antibiotic sensitivity of L. amylovorus was assessed. The isolate was resistant to norfloxacin, gentamicin, and amikacin but was susceptible to amoxicillin, cefoxitin, doxycycline, fleroxacin, clarithromycin, and erythromycin. These findings provide valuable reference points for combination therapy strategies, highlighting the need to avoid concurrent use with antibiotics to which the strain is resistant. In vitro inhibitory tests demonstrated that L. amylovorus exerted inhibitory effects against both E. coli and S. aureus, with a stronger inhibition observed against E. coli. This selective inhibition may be related to the specific expression of the bacteriocin amylovorin L, which has been reported to exert antibacterial effects by disrupting the outer membrane structure of Gram-negative bacteria [63, 64].

Further animal trials analyzed changes in the gut microbiota before and after intervention using 16 S rDNA sequencing. Alpha diversity analysis revealed that, compared to the healthy control group, L. amylovorus did not significantly affect the diversity of the gut microbiota in healthy goats, although a slight increase in species diversity was observed, which aligns with the findings of Wang et al. [65]. Comparing the microbiota of FKS-affected goats on day 1 and day 5 of the trial showed that the species diversity of the gut microbiota significantly increased by day 5. This suggests that L. amylovorus can enhance the species diversity in the gut microbiota of FKS-affected goats. Principal Coordinate Analysis (PCoA) showed that the gut microbiota structure of FKS-affected goats, which was distinctly different from that of the healthy group on day 1, partially overlapped with the healthy group by day 5. This indicates that the gut microbiota structure of the FKS-affected goats gradually became more similar to that of healthy goats, suggesting that L. amylovorus has a regulatory effect on the gut microbiota of FKS-affected goats. When comparing the species composition and abundance between the T group, the H group, and the healthy control group, an upward trend in species abundance was observed in both the T group and the H group. This indicates that L. amylovorus can promote an increase in the gut microbiota in both healthy goats and FKS-affected goats.

Comparing the relative abundance of gut microbiota across the three groups revealed notable differences. At the genus level, the relative abundance of different genera in the healthy control group remained relatively stable, with no significant fluctuations. In H group, the relative abundance of Escherichia-Shigella showed a downward trend over time, though this decrease was not statistically significant. However, the Butyricicoccus genus exhibited a significant decline, while Bacteroides showed a significant upward trend. The relative abundance of Lactobacillus spiked significantly on the third day of the trial, but after a slight drop on the fifth day, the difference compared to the first day was no longer significant. By day 5 of the trial, the relative abundance of Escherichia-Shigella in H group was lower than that in the healthy control group, while the relative abundance of both Lactobacillus and Bacteroides was higher compared to the healthy control group. This indicates that L. amylovorus supplementation may help suppress harmful bacteria such as Escherichia-Shigella and promote probiotics, contributing to a healthier gut microbiota balance. In T group, the relative abundance of pathogenic bacteria like Escherichia-Shigella and Fusobacterium decreased significantly over time, while the relative abundance of probiotics such as Lactobacillus and Ruminococcus significantly increased. There was also a trend of increased relative abundance for Bacteroides and Butyricicoccus, although the difference was not statistically significant. Compared to the healthy control group, on the first day of the experiment, the Escherichia-Shigella levels in T group were significantly higher, while Lactobacillus levels were significantly lower than in H group. By the fifth day of the experiment, the differences in Escherichia-Shigella and Lactobacillus levels between the supplemented group and the healthy control group were no longer significant, indicating that the gut microbiota of the FKS goat group was progressively resembling that of healthy goats. Overall, supplementing L. amylovorus in healthy goats promotes the growth of probiotics like Lactobacillus and Bacteroides, though its inhibitory effect on Escherichia-Shigella is not significant. In FKS-affected goats, L. amylovorus significantly reduces the relative abundance of Escherichia-Shigella and promotes the growth of probiotics like Lactobacillus and Ruminococcus. Bacteroides produce short-chain fatty acids like acetate, facilitating carbohydrate metabolism, while Ruminococcus breaks down cellulose, supplying energy to the host [66–68]. The opposite trends in Butyricicoccus between the T group and H group may be due to differences in the relative abundance of Butyricicoccus. L. amylovorus may compete with Butyricicoccus when it is present in higher relative abundance, inhibiting its growth. However, when Butyricicoccus is present in lower abundance, the suppression of Escherichia-Shigella by L. amylovorus indirectly aids the growth of Butyricicoccus. The metabolic product butyrate from Butyricicoccus plays a crucial role in maintaining gut health [36, 69, 70]. Additionally, Butyricicoccus can metabolize various polysaccharides, supporting gut fermentation [71]. Research suggests that a reduction in the relative abundance of butyrate-producing bacteria may trigger inflammation [72].

In summary, L. amylovorus has the ability to exert probiotic effects on the digestive tracts of animals. It can increase the diversity and species richness of the gut microbiota in FKS-affected goats and help modulate their gut microbiome. It significantly reduces the relative abundance of pathogenic bacteria such as Escherichia-Shigella in FKS-affected goats while promoting probiotics such as Lactobacillus and Ruminococcus, which can help prevent and treat FKS. Additionally, in healthy goats, L. amylovorus promotes the growth of probiotics such as Lactobacillus and Bacteroides without significantly altering the existing gut microbiome structure of goats.

L. amylovorus shows probiotic potential for preventing and treating FKS. However, further clinical trials are necessary to determine whether its combination with other drugs or antibiotics provides better therapeutic outcomes and to evaluate its ability to enhance immunity and productivity in healthy goats.

Conclusion

Goat-derived Lactobacillus amylovorus demonstrated robust probiotic properties, including high acid (pH 2.0–3.0), bile (0.3%), and heat (60 °C) tolerance, and potent inhibition of Escherichia coli and Staphylococcus aureus. In FKSaffected goats, daily gavage with 1 × 10⁸ CFU for 3 days significantly increased gut microbial diversity (Shannon + 25%, Simpson + 18%), boosted beneficial genera (Lactobacillus, Ruminococcus), and reduced pathogenic Escherichia–Shigella by 40% (p < 0.05). These findings underscore L. amylovorus ability to restore gut homeostasis, making it a promising probiotic intervention for preventing and treating Floppy Kid Syndrome in goat production.

Authors’ contributions

Conceptualization, D. C.; methodology, D.C.; software, C.C.; validation, D.C., C.C. and Y.Z.; formal analysis, C.C. and Y.Z.; investigation, C.C., Y.Z. and X.W.; resources, D.C.; data curation, C.C.; writing—original draft preparation, C.C.; writing—review and editing, C. C., J.T. and D.C.; visualization, C.C.; supervision, D.C. and J.T.; project administration, D.C..All authors reviewed the manuscript.

Funding

This study was funded by the earmarked fund for Jiangsu Agricultural Industry Technology System (JATS [2023]449), partially funded by the Priority Academic Program Development of Jiangsu Higher Education Institutions (PAPD) (2018).

Data availability

The datasets used and/or analysed during the current study are available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

All experimental protocols, including the sample collection and clinical trials of goats, were carried out under the permission and guidelines of the Animal Care and Use Committee of the College of Veterinary Medicine, Yangzhou University (Approval ID: 202108926). In addition, as the study involved client-owned animals, written informed consent was obtained from the owners prior to sample collection. All methods and protocols in this study comply with the ARRIVE guidelines.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.