All experiments were conducted in vitro and did not involve live animals or human participants; ethical approval was not required for this study. All laboratory procedures and data handling were followed. Institutional safety guidelines for the laboratory, according to Thailand’s Enhancing Safety Practices in Research Laboratories (ESPReL; number 2-0021-0052-7), were strictly observed.

The study was conducted between 2021–2024 at the Microbiology Laboratory, Faculty of Liberal Arts and Science, Kasetsart University, Kamphaeng Saen Campus, Nakhon Pathom, Thailand.

Thai ground herbs with particle sizes of approximately 0.3–0.5 mm, including turmeric (C. longa), galingale (B. rotunda), and gotu kola (C. asiatica), were purchased from Vejpongosot Yaowarat Co., Ltd., Bangkok, Thailand. The herbs were stored in a cold, dry chamber for approximately 2–3 days.

Black sesame seeds (S. indicum) were obtained from Thanya Farm Co., Ltd., Nonthaburi, Thailand. The seeds were stored in a cold, dry room for approximately 2–3 days and pulverized in a blender to obtain particle sizes of approximately 0.3–0.5 mm before being used in the studies.

The probiotic strains used in this study were Ligilactobacillus salivarius KUKPS6202, Lacticaseibacillus paracasei KUKPS6201, Lactobacillus acidophilus KUKPS6107, Lacticaseibacillus rhamnosus KUKPS6007, Limosilactobacillus reuteri KUKPS6103, Bacillus coagulans KPS-TF02, and Saccharomyces cerevisiae subsp. boulardii KUKPS6005 [21, 22]. These strains were previously identified for their high probiotic potential and are listed by the Ministry of Public Health, Thailand. All probiotic strains were isolated from fermented foods purchased from different Thai local markets. All probiotic microorganisms were nonhaemolytic [21] and did not exhibit any antimicrobial resistance genes.

The probiotic bacteria were cultivated in nutrient broth (NB; Merck, Darmstadt, Germany) and de Man Rogosa and Sharpe medium (MRS; Merck, Darmstadt, Germany), while the probiotic yeast was cultured in yeast extract–malt extract medium (YM; Himedia, Mumbai, India). Incubation was performed at 37°C for 24–48 h. The cultures were stored at 4°C for short-term use and −20°C for long-term storage until further tests were needed.

A patented mung bean–soybean medium consisting of 1% (w/v) soybean, 1% (w/v) mung bean, and 0.5% (w/v) coconut sugar was prepared as previously described [23]. All probiotic strains were previously tested for growth enhancement. The samples were incubated at 37°C with shaking at 150 rpm to assess the effects of L. salivarius, L. paracasei, L. acidophilus, L. rhamnosus, L. reuteri, B. coagulans, and S. boulardii on growth in mung bean–soybean medium. Growth was monitored at 0, 24, 48, and 72 h using the spread plate count method.

Turmeric, galingale, and gotu kola were fermented with probiotic bacteria. Seven probiotic strains were cultivated in mung bean–soybean medium at 37°C for 48 h. The probiotic strain was separately mixed with 10% (w/v) dried herbal powder in water, heated to 100°C for 5 min, and cooled to 45°C.

A 25% (w/v) mixture of the probiotic inoculum (approximately 10⁶ colony-forming units [CFU]/mL) and herbal water (initial pH approximately 6) was added to the inoculum and incubated for 24 h at 37°C without agitation. The mixtures (final pH approximately 5) were placed in separate glass bottles and stored at 4°C until further use.

The antimicrobial activity of aqueous herbal extracts fermented with probiotics was tested using the agar well diffusion method [24] with some modifications. Pathogenic bacteria including Aeromonas hydrophila, Bacillus cereus, Escherichia coli, Staphylococcus aureus, Staphylococcus epidermidis, Serratia marcescens, Salmonella Typhimurium, Proteus vulgaris, and Vibrio parahaemolyticus were used as indicator strains.

Aqueous herbal extracts fermented with each single probiotic and multi-strain probiotics (100 μL) were added to the pathogen agar plates. The herbal extract was used as a negative control, while streptomycin at 0.1 mg/mL served as a positive control. The agar plates were incubated at 37°C for 48 h, and then the zones of inhibition were measured. The experiment was replicated three times. A clear zone around each well was used to assess the antibacterial activity.

The aqueous herbal extract fermented with multi-strain probiotics was prepared as described above. A 1% (w/v) black sesame extract was added to the herbal extract before probiotic fermentation.

The growth of probiotic strains in 1% black sesame extract in aqueous herbal extracts was compared with that in aqueous herbal extracts without sesame extract. Probiotic microorganisms were cultivated in 10 mL of MRS, NB, and YM broth media. The probiotics were then inoculated into each prebiotic-based medium and incubated for 24 h at 37°C.

Culture samples (1 mL) were collected after 0, 6, 12, 24, and 48 h. The growth of probiotics was measured using the dilution plate method [21] on different agar media. Briefly, 1 mL of sample was suspended in 9 mL of 0.85% normal saline before being diluted about 10 times until the suitable dilution was achieved. One hundred microliters of successive decimal dilutions were spread on medium agar (MRS, NA, or YM agar). The experiment was replicated four times. Then, the product fermented with black sesame extract in aqueous herbal extracts fermented with multi-strain probiotics was used for further analysis.

The fermented product was evaluated for the inhibition of α-amylase activity based on the modified methods of Kusano et al. [25] and Luyen et al. [26]. Potato starch (100 mg) was boiled in 5 mL of phosphate buffer (pH 7.0), cooled to room temperature, and combined with 50 μL of the fermented sample and 30 μL of 0.1 M phosphate buffer. After 5 min of pre-incubation, 20 μL of a 2 mg/mL α-amylase solution was added and incubated at 37°C for 15 min.

The reaction was stopped by adding 50 μL of 1 M HCl, followed by 50 μL of iodine solution. The absorbance was measured at 650 nm using a microplate reader (SPECTROstar Nano, BMG LABTECH, Ortenberg, Germany). Acarbose was used as the positive control. The half-maximal inhibitory concentration (IC50) was calculated. All determinations were performed at least three times.

The percentage inhibition (IC50) of absorbance at OD650 was plotted as a function of the acarbose concentration and was used to calculate the acarbose equivalent antioxidant capacity. The experiment was replicated three times.

The α-glucosidase enzyme inhibition assay, based on Luyen et al. [26] and Ali et al. [27], involved combining the fermented sample solution and 40 μL of 0.5 U/mL α-glucosidase in 120 μL of 0.1 M phosphate buffer (pH 7.0). After pre-incubation for 5 min, 40 μL of a 5 mM p-nitrophenyl-α-D-glucopyranoside solution was added, and the mixture was incubated for 30 min at 37°C.

The absorbance of the released p-nitrophenol was measured at 405 nm. Acarbose was used as the positive control. The IC50 was determined. All determinations were performed at least three times.

The percentage inhibition (IC50) of absorbance at OD405 was plotted as a function of acarbose concentration and was used to calculate the antioxidant capacity of acarbose. The experiment was replicated three times.

Lipase activity was measured using a Lipase Assay Kit (Elabscience®, Houston, TX, USA) as described below. First, 10 μL of the control sample was added to the corresponding control well. Similarly, 10 μL of the fermented sample was added to each well. In the control well, 40 μL of the buffer solution was added, while in the sample wells, 40 μL of the substrate working solution was added.

The wells were thoroughly mixed for 5 s in a microplate reader (SPECTROstar Nano), followed by incubation at 37°C for 20 min. After incubation, 150 μL of the chromogenic working solution was added to each well, followed by thorough mixing for 5 s. Then, the wells were incubated in the dark for an additional 30 min at 37°C.

The optical density of each well was measured at 412 nm using a microplate reader. The experiment was replicated three times. The lipase activity was calculated using the following formula:

Lipase activity (mU/L) = (ΔA / ε × b) × f × T × 10⁶ × 1,000

where ΔA = ODsample − ODcontrol, ε = molar absorption coefficient (14,150 L/mol×cm), b = height of the reaction system (0.6 cm), T = incubation time of the reaction (20 min), and f = dilution factor of the sample before the test. One mol/L equals 10⁶ μmol/L.

The analytical sensitivity of the assay is 0.03 U/L. This was determined by adding two standard deviations to the mean optical density obtained when the zero standard was assayed 20 times, and the corresponding concentration was calculated (Elabscience®).

2,2-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging activity assay

The antioxidant activity of the fermented samples was assessed using the 2,2-diphenyl-1-picrylhydrazyl radical scavenging assay (DPPH; Sigma-Aldrich, Saint Louis, MO, USA) following a modified protocol by Brand-Williams et al. [28]. Ten microliters of the sample supernatant was added to a test tube containing 190 μL of 0.06 mM DPPH solution in 95% ethanol. The mixture was vortexed thoroughly and incubated in the dark for 30 min at room temperature.

The absorbance was measured at 515 nm using a microplate reader (SPECTROstar Nano). The antioxidant activity was compared with that of a standard Trolox solution, with 95% ethanol as the blank. All determinations were performed at least three times.

The percentage inhibition (IC50) of absorbance at OD515 was plotted as a function of Trolox concentration and was used to calculate the equivalent antioxidant capacity. The experiment was replicated three times.

The DPPH radical scavenging activity (%) was calculated as [(Acontrol − Asample) / Acontrol] × 100, where Acontrol is the absorbance of the control (DPPH solution without the sample) and Asample is the absorbance of the sample after reaction with DPPH. Antioxidant activity was expressed as IC50 of Trolox divided by IC50 of the sample.

Experiments were conducted following a statistical design [30]. Data analysis was performed using the Statistical Package for the Social Sciences version 22 software (SPSS Inc., IBM®, Armonk, NY, USA). Tukey’s multiple comparisons test, one-way analysis of variance, and the independent-samples t-test were used to assess significant differences between groups. A significance level of p < 0.05 was considered statistically significant. Microsoft Excel was used for the IC50 and enzymatic calculations.

The effects of mung bean and soybean-based medium on the growth of seven probiotic strains were evaluated after 72 h at 37°C. After a 24–72 h incubation period, all probiotic strains exhibited high growth in this medium. As shown in Table 1, the viable cell counts increased substantially compared to the initial time at 0 h. Based on these results, the mung bean–soybean medium provided a favorable environment for probiotic growth, as observed in another study on legume-based fermentation [31]. Notably, the lactic acid bacteria strains (L. salivarius, L. paracasei, L. acidophilus, L. rhamnosus, and L. reuteri) exhibited peak growth at 24 h, after which the viable cell count slightly declined at 72 h. This trend aligned with the findings of Ruiz de la Bastida et al. [32], where L. paracasei INIA P272, L. rhamnosus INIA P344, and L. rhamnosus GG reached their highest counts after 24 h of fermentation in soy beverages. Among the tested strains, Bacillus coagulans KPS-TF02 had the highest overall increase in viable count and maintained stable growth until 72 h, suggesting that B. coagulans is more resistant to environmental changes and can thrive during extended fermentation [33]. In addition, Saccharomyces boulardii KUKPS6005 exhibited steady growth but at a lower rate than the other bacterial strains. These findings confirm that mung bean and soybean are suitable substrates for probiotic fermentation, particularly for lactic acid bacteria and spore-forming Bacillus species. Future research should explore how different fermentation conditions (such as pH and temperature variations) influence probiotic survival and bioactive compound production [34]. Thus, the mung bean–soybean medium was used for the production of probiotic inoculum.

The antimicrobial activity of the fermented aqueous herbal extracts (turmeric, galingale, and gotu kola) using probiotic strains (L. salivarius, L. paracasei, L. acidophilus, L. rhamnosus, L. reuteri, B. coagulans, S. boulardii, and multi-strain) was compared against aqueous herbal extracts (negative control) and 0.01% streptomycin (positive control), as shown in Table 2 and Supplementary Figure S1 and S2. The antimicrobial activity against two major pathogens (A. hydrophila and B. cereus) was evaluated using the agar diffusion method. Based on the results, all the probiotics produced clear zones of inhibition, indicating their ability to suppress pathogen growth. The inhibition diameters of A. hydrophila (7.67–8.67 mm) and B. cereus (7.33–9.00 mm) suggested that the probiotic-fermented herbal extracts had moderate antimicrobial activity, likely due to the production of organic acids, hydrogen peroxide, and bacteriocins during fermentation [35]. With the other pathogens, the cells of Vibrio parahaemolyticus were destroyed, although no clear zone was formed (Supplementary Figure S1), indicating a different antimicrobial mechanism. The probiotic strains did not affect E. coli, Proteus vulgaris, S. aureus, S. epidermidis, Serratia marcescens, and S. Typhimurium (Supplementary data Figure S2). This may be due to the low concentration of active antimicrobial compounds or the high resistance of these bacteria. Due to the moderate inhibition activity, the minimum inhibitory concentration and minimum bactericidal concentration were not determined. However, other antibacterial activity tests, such as the co-inoculation test and adhesion activity, should be evaluated. Chumphon et al. [21] reported that several probiotic strains (L. paracasei, L. acidophilus, L. reuteri, L. rhamnosus, L. salivarius, B. coagulans, and S. boulardii) inhibited the growth of B. cereus, E. coli, S. aureus, S. marcescens, S. Typhimurium, P. vulgaris, V. harveyi, and V. parahaemolyticus. In addition, they reported that L. rhamnosus had strong antimicrobial activity against eight pathogens (B. cereus, Pseudomonas aeruginosa, V. parahaemolyticus, S. marcescens, S. Typhimurium, A. hydrophila, Candida albicans, and E. coli). In contrast, as reported elsewhere [36], B. coagulans inhibited any pathogens tested. The mechanism of probiotic antimicrobial activity was likely due to bacteriocins, which disrupt the bacterial cell wall, causing leakage of amino acids and inorganic salts and ultimately leading to pathogen inhibition [35]. Ensuring animal health and feed safety is a critical component of sustainable livestock and aquaculture production. Among the wide range of microbial hazards associated with animal production systems, A. hydrophila and B. cereus have gained increasing attention due to their ability to cause disease in animals and contaminate feed and feed ingredients. A. hydrophila exemplifies pathogens linked to aquatic environments and live animal disease, whereas B. cereus is a feed-borne hazard capable of surviving processing and producing harmful toxins. Together, they illustrate the interconnectedness of feed quality, environmental hygiene, and animal health. Safer feed formulations and alternative control strategies, such as probiotics, organic acids, and enhanced processing technologies. This aligns with global efforts to reduce AMR and promote sustainable animal production.

Figure 1 shows the viable cell counts of L. salivarius, L. paracasei, L. acidophilus, L. rhamnosus, L. reuteri, B. coagulans, and S. boulardii during fermentation. The prebiotic potential of the black sesame extract was confirmed by the enhanced viable cell counts of all the probiotic strains tested. Some strains, especially B. coagulans and L. salivarius, had higher growth rates, indicating that they effectively digested the prebiotic ingredients. Black sesame extract had a substantial impact on probiotic development compared with aqueous herbal extracts alone (t-test; p < 0.05). Several mechanisms alone or in combination could be responsible for the observed increase in probiotic viability, with the potential growth enhancement pathways depending on nutrient availability. For example, the oligosaccharides and vital minerals found in black sesame extract provide the fermentable energy source for probiotic metabolic activity [37]. According to Wei et al. [16], black sesame fiber and polyphenols act as prebiotic substrates and specifically promote the growth of beneficial gut bacteria. Black sesame contains many antioxidants, including sesamin and sesamolin, which may shield probiotic cells from oxidative stress and increase their survival during fermentation [38]. The results of this study were corroborated by Wei et al. [16], who showed that prebiotics increased probiotic viability and metabolic activity. According to Fukuda et al. [38], black sesame components altered the gut microbiota composition and boosted probiotic survival in fermented foods. Our results were similar to those of Ruiz-Ojeda et al. [37], who reported that polyphenol-rich prebiotics greatly enhanced the growth of Lactobacillus and Bifidobacterium species. Overall, the results supported the notion that black sesame extract was a useful prebiotic that improved probiotic viability in aqueous herbal extracts. This prebiotic-probiotic combination may be beneficial in the creation of nutraceuticals and functional fermented foods that enhance gut health.

Notably, fundamental quality control parameters, such as total curcumin content and selected marker compounds, should be systematically evaluated to ensure reproducibility and improved characterization of raw materials. Bioactive compounds should be characterized using Fourier Transform Infrared Spectroscopy or GC-MS to enhance the development of commercial probiotic products.

Fermentation with multi-strain probiotics significantly enhanced the antioxidant activities of most strains, as measured by 2,2-diphenyl-1-picrylhydrazyl (DPPH) radical scavenging activity and FRAP assay. The antioxidant potential varied across strains, with B. coagulans showing the most substantial improvement in both assays (Table 5). The DPPH radical scavenging activity improved in most probiotic strains after fermentation. However, B. coagulans (p < 0.05) and L. reuteri (p < 0.05) showed the greatest increases in antioxidant activity, indicating strong antioxidant potential. Zhao et al. [48] reported that probiotics enhanced radical scavenging through the metabolism of polyphenols. In contrast, L. paracasei and L. acidophilus produced only moderate increases in DPPH activity, suggesting that some strains were less efficient in generating antioxidant metabolites during fermentation. In addition, S. boulardii and the multi-strain probiotics maintained similar values before and after fermentation, indicating a less pronounced effect on radical scavenging.

The FRAP assay measures the ability of antioxidants to reduce Fe³⁺ to Fe²⁺ reflecting their overall redox stability and potential metal-chelating effects. Based on the current results, fermentation improved the FRAP values in some strains but reduced them in others. For example, B. coagulans (p < 0.05) demonstrated strong redox balance maintenance, whereas S. boulardii (p < 0.05) reduced oxidative stress. However, L. rhamnosus (p < 0.05) and L. reuteri (p < 0.05) were reduced, possibly due to metabolic shifts away from antioxidant compound production. B. coagulans and S. boulardii are ideal candidates for antioxidant-focused functional foods because of their superior antioxidant enhancement. L. reuteri and B. coagulans are known for their strong polyphenol metabolism, which may explain their high antioxidant enhancement [49]. S. boulardii produce extracellular metabolites that enhance antioxidant defense, contributing to high FRAP values. This variation could be attributed to differences in the metabolism of polyphenols and bioactive compounds in probiotic strains. Some probiotics produce extracellular metabolites that enhance antioxidant defense [50], while others degrade antioxidant compounds over time, leading to reduced FRAP activity.

The ability of probiotics to metabolize polyphenols varies by strain, influencing their antioxidant properties. Some probiotics break down complex antioxidant compounds into more bioavailable forms, thereby increasing antioxidant activity, whereas other strains may consume or degrade these bioactive compounds over extended fermentation periods, thereby reducing FRAP values. Probiotic strain selection and fermentation time optimization are crucial for maximizing antioxidant potential. The current results were consistent with other research in this respect, with Wang et al. [49] reporting that fermentation enhanced polyphenol bioavailability and antioxidant potential, particularly in B. coagulans and S. boulardii. Similarly, Hao et al. [50] noted that fermentation released bioactive peptides with antioxidant properties, consistent with the improved DPPH scavenging observed in the current study. However, some differences were observed compared with the results reported by Zhao et al. [51], in which FRAP values improved across all strains. In contrast, the current study identified certain strains with reduced FRAP values, suggesting that probiotic metabolism may shift during fermentation, affecting the stability of antioxidant metabolites. These findings highlight the strain-specific metabolic pathways influencing probiotic efficacy in enhancing antioxidant activity.

Antioxidant activity is fundamentally based on the suppression of free radical–mediated oxidation. Free radicals are highly reactive species containing unpaired electrons that are generated when covalent bonds are disrupted by external factors. They can induce cellular damage through oxidative stress. Phytochemicals mitigate these effects by interrupting free radical chain reactions, either through hydrogen atom donation or electron transfer to chelated metal ions.

In this study, fermentation of the aqueous herbal extract with probiotic strains significantly increased the levels of antioxidant compounds. Both the herbal extract and the probiotics enhanced the antioxidant capacity, likely through synergistic interactions that amplified the overall antioxidant activity. Further characterization of the bioactive compounds in the fermented products is required to fully elucidate the functional benefits of these probiotic formulations.