Novel Double-Layer Microencapsulated Phytosynbiotic Derived from Probiotics and Tiliacora triandra Extract for Application in Broiler Production

Simple Summary

Phytosynbiotics are a novel category of feed additives that combine phytobiotics and synbiotics. In this study, we developed a double-layer microencapsulated phytosynbiotic (DMP) from probiotics and Tiliacora triandra extract using fermentation, encapsulation, and lyophilization to enhance probiotic stability and preserve phytochemicals for antibiotic-free broiler production. By improving probiotic viability, heat tolerance, and stability during simulated gastrointestinal digestion, wheat bran with a 0.6 mm particle size provided the most effective protection. All probiotic strains had high encapsulation efficiency, but Pediococcus acidilactici led to the best fermentation efficiency leading to greater levels of beneficial short-chain fatty acids. The antibacterial activity among the strains varied and inhibited primarily the growth of pathogens. Overall, DMP effectively stabilized both probiotics and bioactive phytochemicals, thus representing a promising natural approach to optimize feed utilization and the intestinal microflora in broiler production.

Abstract

The global shift toward antibiotic-free poultry production has created an urgent need for sustainable feed additives that promote gut health and productivity. This study aimed to develop and evaluate a novel double-layered microencapsulated phytosynbiotic (DMP) comprising Tiliacora triandra extract, probiotics, and cereal by-products using lyophilization. In Experiment 1, we investigated the effects of cell wall materials (corn, defatted rice bran, and wheat bran) and different particle sizes (0.6 and 1.0 mm) on the physicochemical characteristics and probiotic encapsulation efficiency. Results revealed that wheat bran, particularly at the smaller particle size of 0.6 mm, enhanced probiotic viability, probiotic stability under simulated gastrointestinal and thermal conditions, and nutrient retention. Compared with other materials, wheat bran also provided superior powder flowability, lower density, and favorable color attributes. In Experiment 2, we assessed the influence of probiotic strains (P. acidilactici, Lactiplantibacillus plantarum TISTR 926, and Streptococcus thermophilus TISTR 894) on functional properties of the DMP. All strains exhibited high encapsulation efficiency and stability during gastrointestinal simulation, thermal exposure, and storage. However, P. acidilactici had superior fermentation kinetics and produced greater levels of beneficial short-chain fatty acids, especially acetic and butyric acids. Antibacterial activity was strain-dependent, with notable inhibitory effects against Gram-positive pathogens, primarily through bacteriostatic mechanisms. Overall, these findings confirm that the developed DMP formulations effectively stabilize probiotics and bioactive phytochemicals, offering a promising strategy for enhancing gut health and performance in antibiotic-free broiler production systems.

1. Introduction

Double microencapsulation offers an innovative strategy for delivering probiotics in poultry nutrition, providing superior protection and functional performance compared with conventional single-layer systems. Given the challenges associated with probiotic stability and efficacy, encapsulation technologies have emerged as promising solutions [1]. These positive attributes are especially noteworthy because single-layer encapsulation fails to protect probiotics from thermal stress, storage degradation, and bile salts, thus reducing viability. In fact, insufficient protection under these conditions frequently results in considerable loss of viable probiotic cells, diminished metabolic activity, and reduced colonization capacity within the gut. Thus, the synergistic design of the double-layer coating not only improves probiotic viability, but enables targeted gut release, and supports antibiotic-free production [2]. By removing water under low-temperature and vacuum conditions, lyophilization is an effective encapsulation technique that preserves the structural integrity and bioactivity of probiotics and thermolabile phytogenic compounds, thereby minimizing oxidative degradation and cellular damage [3]. Encapsulation within polymeric matrices further enhances compound stability, protects against environmental stress, and allows for targeted delivery [4]. Clearly, double microencapsulation can enhance probiotic stability, targeted gut release, and antibiotic-free poultry production, outperforming single-layer encapsulation. Consequently, adoption can improve flock health, performance, and overall sustainability.

With the increasing demand for sustainable poultry feed, cereal by-products such as rice and wheat bran have emerged as novel encapsulation materials. These by-products are abundant in fermentable carbohydrates, fibers, and phenolic compounds, which act as prebiotics to promote gut health and fermentation. For instance, rice bran contains dietary fiber, protein, antioxidants, and γ-oryzanol, which exert antioxidant, anti-inflammatory, and immunomodulatory effects [5]. Wheat bran is rich in insoluble fiber, especially arabinoxylans, which support microbial fermentation and metabolic health but can be further enhanced by processing techniques that improve digestibility [6,7]. The ability of bran fiber to act as a protective carrier during drying processes, such as freeze-drying and spray-drying, further strengthens its role as a sustainable prebiotic feed component [8]. Thus, cereal by-products can simultaneously valorize agricultural waste streams and support the development of next-generation feed formulations. To overcome the challenges for sustainable poultry feed production, use of biopolymer-based microencapsulation can help protect probiotics against adverse conditions and ensure targeted gastrointestinal delivery [9,10].

In recent years, there has been a growing interest in synbiotics as functional feed additives for broiler diets [11,12]. Synbiotics, which are combinations of probiotics and prebiotics, create multifunctional products with synergistic benefits [13]. These formulations improve gut health, modulate intestinal microbiota, enhance immune competence, and increase nutrient utilization, making them highly relevant for antibiotic-free broiler production systems [14]. Prebiotics, mainly derived from polysaccharides and other plant-based substrates, act as selective energy sources for beneficial microorganisms, contributing to the improvement of the intestinal microflora. In parallel, probiotics, particularly lactic acid bacteria (LAB), such as Lactobacillus, Pediococcus, and Staphylococcus, play crucial roles in maintaining microbial homeostasis, supporting digestion, enhancing immunity, and improving growth performance in poultry [15,16]. They contribute to pathogen exclusion and produce short-chain fatty acids (SCFA) that support intestinal health [17]. Despite these benefits, the efficacy of probiotics in practical applications is often constrained by their sensitivity to temperature, acidity, exposure to bile salts, and storage conditions, factors that reduce their viability and functional stability [18].

Phytobiotics or phytogenic feed additives are among the most promising innovations and are derived from herbs, plant extracts, and bioactive phytochemicals such as polyphenols, flavonoids, and polysaccharides [19]. These compounds exhibit antioxidant, antimicrobial, and anti-inflammatory properties, and their supplementation in poultry diets improves nutrient utilization, microbial balance, and immunity [20]. Among phytogenic ingredients, Tiliacora triandra (Yanang in Thai) is a Southeast Asian medicinal plant rich in polyphenols, vitamins, dietary fiber, and prebiotic oligosaccharides with strong antioxidant and anti-inflammatory properties [21]. This species also contains substantial levels of phytochemicals such as alkaloids, flavonoids, chlorophyll, and notably high levels of mucilaginous polysaccharides [22,23]. Furthermore, the xylose-rich polysaccharides in the leaves confer antioxidant and immunomodulatory functions, alongside structural properties that enable their use as natural encapsulating matrices for bioactive delivery in feed formulations [24]. Although Yanang exhibits promising pharmacological potential, its incorporation into poultry nutrition remains underexplored and merits a systematic investigation.

Building on these foundations, we introduce phytosynbiotics as a novel category of next-generation feed additives that strategically integrate phytobiotics with synbiotics into a unified multifunctional formulation. Phytosynbiotics combine plant-derived bioactive compounds from Yanang leaf with probiotics and cereal bran-derived prebiotics. In contrast to conventional synbiotics, which combine probiotics and prebiotics primarily for microbial modulation, phytosynbiotics achieve physical co-encapsulation and functional synergy through a proprietary double-layer microencapsulation system utilizing lyophilization. The primary layer comprises fermented Yanang mucilage, which functions as a natural hydrogel matrix to protect probiotics during processing and gastrointestinal transit. The secondary layer incorporates porous cereal bran, serving a dual role as a prebiotic carrier and structural reinforcement to enable targeted release in the distal sections of the gut. This approach represents a novel strategy for stabilizing probiotics and phytochemicals in a double-layered, lyophilized delivery system [19]. Thus, by offering safe and eco-friendly alternatives that connect animal nutrition, environmental stewardship, and human health outcomes, phytosynbiotics align closely with the One Health concept and are within the framework of sustainable livestock production and consumer concerns regarding antibiotic resistance [25].

This study aimed to develop and evaluate a double-layered microencapsulated phytosynbiotic (DMP) formulation containing Yanang leaf extract (YLE), probiotics, and cereal by-products via lyophilization. To address these aims, using an in vitro three-step digestibility model, we investigated the encapsulation efficiency, nutrient composition, bioactive retention, particle size, and effects of encapsulation on gastrointestinal tolerance, fermentative activity, and microbial viability. Ultimately, this study sought to transform local botanical and agricultural resources into globally applicable functional feed solutions to improve broiler production efficiency, support antibiotic-free production systems, and advance sustainable poultry farming.

2. Materials and Methods

This study was conducted at the Animal Nutrition Laboratory, Faculty of Agriculture, Natural Resources and Environment, Naresuan University, Phitsanulok, Thailand. All experimental protocols and procedures involving animals were reviewed and approved by the Naresuan University Animal Care and Use Committee, ensuring compliance with ethical standards for the care and use of animals in research (Approval ID: 68 01 008).

2.1. Experiment 1: Effects of Cell Wall Materials and Particle Size on the Characteristics and Properties of the DMP

2.1.1. Experimental Design

This study employed the Pediococcus acidilactici V202 strain, which is noted for its capacity to produce beneficial organic acids and its antimicrobial efficacy against pathogenic bacteria. Our laboratory initially isolated this strain from the goat vagina [26]. A 3 × 2 factorial experiment was conducted using a completely randomized design to assess the effects of cereal by-products as cell wall materials (corn, defatted rice bran, and wheat bran) and particle size (0.6 and 1.0 mm), with three replicates per treatment. The DMP was formulated via a double-layered microencapsulation technique in which P. acidilactici V202 was co-encapsulated with YLE and these cereal by-products. The primary layer consisted of Yanang mucilage, serving both as a fermentation medium supporting probiotic viability and as a protective matrix against environmental stressors. The secondary layer was composed of selected cell wall materials that acted as carriers, further enhancing structural integrity and functional stability of the microcapsules. Encapsulation was achieved through a homogenization process, ensuring uniform dispersion of probiotic cells within the biopolymeric matrix, followed by freeze-drying, as illustrated in Figure 1. This approach not only improved the physicochemical stability of the encapsulated probiotics but also facilitated downstream handling, storage, and potential incorporation into feed formulations.

2.1.2. Fermentation, Homogenization and Microencapsulation Procedure

Fresh Yanang leaves were harvested from the agricultural fields of Rajamangala University of Technology Lanna, Phitsanulok. The leaves were thoroughly washed under running tap water to remove surface contaminants and stored at −20 °C until further processing. Prior to extraction, the frozen leaves were blanched in boiling water for 2 min and then cooled. Extraction was performed by blending the leaves with distilled water at room temperature at a 1:3 (w/v) ratio using a laboratory blender, followed by filtration to obtain the aqueous extract [2]. The extract was pasteurized at 75 °C for 5 min for microbial decontamination and then cooled to 37 °C prior to inoculation. The probiotic strain P. acidilactici V202, preserved at −20 °C in de Man, Rogosa, and Sharpe (MRS) broth with 20% glycerol, was reactivated by two successive subcultures in MRS broth (37 °C, 18 h, anaerobically) [27]. Cells were harvested (5000× g, 10 min, 4 °C), washed, and resuspended in PBS (pH 7.2) to a concentration of ~10⁸ colony-forming units (CFU)/mL. Fermentation was carried out by inoculating YLE with a 0.15% (v/v) inoculum and incubating at 37 °C for 24 h under anaerobic conditions [28] (Figure 1). Subsequently, the wall material-to-YLE ratio was maintained at 15:85 (w/v) for all samples. The mixtures were homogenized at 1200–2000 rpm for 15–20 min. Double-layered microcapsules were subsequently loaded into the pores of the pretreated wall material by adsorption under mild vacuum conditions to enhance penetration efficiency. The loaded materials (300 mL per tray) were frozen at −18 to −20 °C for 24 h and then lyophilized (FreeZone, Labconco, Kansas City, MO, USA). Primary drying was performed by lyophilization at −40 °C and 0.5 mbar for 24 h. Secondary drying was conducted at 20 °C and 0.5 mbar for 24 h to obtain a stable DMP. The final product was stored in vacuum bags at 4 °C until it was incorporated into the experimental diets (Figure 1).

2.1.3. Measurement of Encapsulation Efficiency, Product Quality, and Stability

The production yield was measured by comparing the weight of the final dried encapsulated product to that of the original mixture before encapsulation. The freeze-drying yield was expressed as a percentage of the dry powder weight relative to the initial sample weight (yield (%) = [weight of freeze-dried powder/weight of initial material] × 100) [3]. The viability of the probiotic cells was evaluated using plate counts on MRS agar (HiMedia-GM641-500g, Mumbai, India) with samples taken both before and after the processes of encapsulation and freeze-drying. The assay involved culturing LAB in MRS broth under aerobic conditions at 37 °C for 24 h. To evaluate bacterial growth, colonies were counted, and the results expressed as CFU per gram of sample. The encapsulation efficiency (%) was calculated by comparing the viable cell counts before and after encapsulation using the formula: (viable cells after encapsulation/viable cells before encapsulation) × 100 [29]. The stability of the encapsulated probiotics was assessed under conditions simulating the gastrointestinal environment and thermal stress. Simulated gastric fluid (SGF) was prepared by dissolving 0.3% (w/v) pepsin (SRL, 1:2500 EX, porcine stomach mucosa; 0.6 Anson U/mL) in 0.1 N HCl, and the pH was adjusted to 2.5. Simulated intestinal fluid (SIF) was prepared by dissolving 1% (w/v) pancreatin (SRL, 3× EX, porcine pancreas, ≥75 U/mg; containing ≥75 U/mg amylase, 6 U/mg lipase, 75 U/mg protease) in 0.05 M phosphate buffer, and the pH was adjusted to 7.0. Encapsulated probiotic samples were incubated in SGF and SIF under standard conditions to evaluate gastrointestinal tolerance. Thermal tolerance was evaluated by exposing the samples to 100 °C for 3 min. Surviving probiotic populations post-treatment were enumerated by plating on MRS agar and reported as log CFU per gram [29]. Bulk and tapped densities (g/100 mL) were measured by volume displacement in a graduated cylinder, and powder flowability was assessed using Carr’s compressibility index (CCI) (%), calculated as ((tapped density − bulk density)/tapped density) × 100 [30]. Water activity (Aw) was measured at 25 °C using a LabMASTER-aw water activity meter (Novasina AG, Lachen, Switzerland). The color parameters (L*, a*, and b*) of the DMP were recorded using a colorimeter based on the CIELAB system [31]. Nutrient composition analyses of dry matter (DM), organic matter (OM), crude protein (CP), ether extract (EE), crude fiber (CF), and gross energy (GE) were conducted following standard methods [32].

2.1.4. Statistical Analysis

Data were analyzed using a two-way factorial arrangement in a completely randomized design with three cell wall materials (corn, defatted rice bran, and wheat bran) and two particle sizes (0.6 and 1.0 mm) as fixed factors, with three replicates per treatment, and all measurements were performed in triplicate. The general linear model (GLM) used was: Yijk = μ + αi + βj + (αβ) + εijk, where Yijk is the observed response, μ is the overall mean, αi is the effect of the i = wall material, βj is the effect of the j = particle size, (αβ) is the interaction effect, and εijk is the random error. Statistical analyses were performed using R version 4.3.3 with the ‘Agricolae’ package [33], and multiple comparisons were conducted using Tukey’s Honestly Significant Difference (HSD) test when significant differences were observed (p < 0.05).

2.2. Experiment 2: Effects of Probiotic Strain Variation on the Characteristics and Functional Properties of the DMP Formulation

2.2.1. Sample Preparations and Tests

The probiotic strains utilized in this study comprised P. acidilactici V202 (as previously mentioned), Streptococcus thermophilus TISTR 894, and Lactiplantibacillus plantarum TISTR 926, all supplied by the Thailand Institute of Scientific and Technological Research. A completely randomized design was employed to assess the effects of three probiotics (P. acidilactici, L. plantarum and S. thermophilus), with each treatment replicated three times and all measurements performed in triplicate.

DMP was produced via double-layered microencapsulation using different probiotic strains. The probiotics were encapsulated with YLE within a wheat bran matrix. Finally, the samples were analyzed to determine the encapsulation efficiency and product quality, following the procedure described in Experiment 1. The stability of the encapsulated probiotics was evaluated under various stress conditions. Acid tolerance was assessed by incubating 5.0 ± 0.5 g of microcapsules in 20 mL of 0.2 M phosphate buffer (pH 2.0) and simulated gastric fluid (0.3% pepsin, pH 2.5) at 40–42 °C for 2 h. Pancreatin tolerance was tested in simulated intestinal fluid (1% pancreatin, pH 7.0) at 40–42 °C for 3 h [29]. Thermal tolerance was evaluated by exposing samples to 100 °C for 2.5, 5.0, and 10.0 min [34]. Storage stability was monitored at 4 °C for 0, 2, 4, and 6 months [35]. Subsequently, the samples were stored in airtight containers at 4 °C until incorporation into the experimental diets.

2.2.2. Analysis of Phytochemical Profile

The antioxidant properties of DMP were assessed using compositional analyses and functional antioxidant assays. The total phenolic content was quantified and expressed as mg gallic acid equivalents per gram of dry weight (mg GAE/g DW). The total antioxidant capacity was determined as mg ascorbic acid equivalents per gram of dry weight (mg AAE/g DW). Functional antioxidant activity was assessed using three complementary assays. The free radical scavenging capacity was measured using the 2,2-diphenyl-1-picrylhydrazyl (DPPH). The 2,2-azino-bis-3-ethylbenzothiazoline-6-sulphonic acid (ABTS) assay was employed to evaluate both hydrophilic and lipophilic antioxidant activity. The reducing power was determined using the ferric reducing antioxidant power (FRAP) assay [36]. Tannin content was measured using the Folin–Ciocalteu method, which forms a blue complex detectable at 765 nm using a microplate reader. Results were expressed as tannic acid equivalents. Non-tannin polyphenols were removed to increase specificity, and their concentrations were calculated from a standard curve [37]. In addition, phytochemical profiling was conducted, including the quantification of phenolics, flavonoids, carotenoids, and chlorophylls, to support the interpretation of the functional antioxidant results. Correlations between phytochemical composition and antioxidant activity were analyzed as described previously [38,39,40].

2.2.3. Antimicrobial Activity Determinations

One gram of DMP was added to Mueller–Hinton broth (MHB) (Difco, Bernin, France) and incubated at 37 °C for 24 h. The samples were then centrifuged at 5000× g for 10 min to collect the cell-free supernatant. The supernatant was sterilized by filtration using a sterile 0.45 µm filter (Pall Corporation, New York, NY, USA) [41]. Antimicrobial activity was assessed using a broth microdilution assay in accordance with Clinical and Laboratory Standards Institute (CLSI) guidelines, with minor modifications. The supernatant underwent serial two-fold dilutions ranging from 50–1.56% v/v in a microtiter plate. The pathogens (Bacillus cereus WU22001, Staphylococcus aureus ATCC25923, and E. coli ATCC 25922) were cultivated in MHB and incubated for 4–6 h at 37 °C. The bacterial culture was adjusted to a 0.5 McFarland standard and then diluted to achieve a final concentration of 1 × 10⁶ CFU/mL. Subsequently, 100 µL of each bacterial suspension was added to the microtiter plate as described above [42]. Vancomycin and gentamicin served as positive controls. The MIC was defined as the lowest concentration that inhibited visible growth, while the MBC was the lowest concentration that prevented colony formation upon subculture [43].

2.2.4. Stability of Probiotics in the DMP After In Vitro Ileal Nutrient Digestibility Assessment

The in vitro ileal digestibility of experimental diets (n = 30) was assessed using a two-step enzymatic procedure adapted from [32,44]. Samples (0.5 g, 1 mm) were incubated with pepsin (0.1 g in 10 mL 0.2 M HCl, pH 2.0) followed by pH adjustment to 6.8 and addition of pancreatin (0.5 mg in 10 mL 0.2 M phosphate buffer) to simulate intestinal digestion. After digestion, 5 mL aliquots were diluted 10-fold in sterile 0.85% NaCl, and serial dilutions were plated on selective and non-selective media. Total aerobic bacteria were enumerated on PCA, Lactobacillaceae on MRS agar under microaerobic/anaerobic conditions, and Enterobacteriaceae on MacConkey agar, with incubation at 30–37 °C for 24 h [29,45,46]. The CFU/mL was calculated from colony counts, and microbial survival (%) through simulated GIT, thermal stress, or storage was determined as follows: Survival (%) = 10^{(log10N_out − log10N_in)} × 100 [19].

2.2.5. Evaluation of Gas Production, SCFA Formation, and Probiotic Populations Using In Vitro Cecal Fermentation

In vitro cecal fermentation was performed using broiler cecal inocula to assess gas production, SCFA formation, degradation kinetics, and microbial dynamics. After confirmation of good clinical health, five 42-day-old Ross 308 broilers, maintained on standard corn–soybean meal diets without antibiotics at the Naresuan University experimental farm were humanely euthanized following institutional ethical guidelines. Cecal contents were harvested and diluted 1:10 (w/w) in phosphate-buffered saline (0.1 M PBS, pH 7.4) while maintaining anaerobic conditions. Fermentations were carried out in 100 mL serum bottles containing 0.3 g digesta and 45 mL sterile VL medium (modified from Prayoonthien [47]), flushed with nitrogen to maintain anaerobic conditions, and sterilized at 121 °C for 15 min. Each bottle was inoculated with 5 mL of 1:10 (w/w) digesta slurry in pH-adjusted buffer (6.18–6.50) and incubated at 42 °C for 24 h under anaerobic conditions (Bactron 300, Shel Lab, Cornelius, OR, USA) [44]. Gas production was recorded at 0–24 h using a glass syringe [48], and cumulative data were fitted to the Ørskov and McDonald [49] model: Y = a + b (1 − e^−ct), where Y is gas volume (mL) at time t (h), a is gas from the rapidly fermentable fraction, b is gas from the slowly fermentable fraction, c is the rate constant for fraction b, and (a + b) represents total gas potential. The parameters were estimated using nonlinear regression [44,50].

After 24 h of in vitro fermentation, 1 mL of fermentation fluid was centrifuged at 10,000× g for 10 min at 4 °C, and the supernatant was stored at −20 °C until analysis. The SCFAs, including acetate, propionate, butyrate, and valerate, along with lactic acid, were quantified using a gas chromatograph (model Agilent 7890B, Agilent Technologies, Santa Clara, CA, USA) equipped with a CP-Sil 5 CB fused silica capillary column (0.32 mm × 25 m) and flame ionization detector. The injection, column, and detector temperatures, as well as the nitrogen carrier flow, followed the optimized conditions described by Boets [51]. 4-Methylvaleric acid and fumaric acid (Alfa Aesar, Lancashire, UK) were used as internal standards for the SCFAs and lactic acid, respectively. Concentrations were calculated from the calibration curves of authentic standards following a modified method of Asarat [52].

After completion of the in vitro digestibility assay, 5 mL of each sample was aseptically mixed with 45 mL of sterile 0.85% (w/v) NaCl to obtain a 10⁻¹ suspension. Serial 10-fold dilutions were prepared in sterile 0.85% NaCl, as described by Xiao [53]. Viable microorganisms were enumerated by plating them on selective and non-selective media. Total aerobic bacteria were counted on Plate Count Agar (PCA) incubated at 30–32 °C for 24 h [45]. Lactobacillaceae were cultured on MRS agar at 30–37 °C for 24 h [29], and Enterobacteriaceae were cultured on MacConkey agar at 30–37 °C for 24 h under microaerophilic or anaerobic conditions [54]. Only plates with 30–150 colonies were considered for the enumeration. Colony-forming units per milliliter (FU/mL) were calculated based on colony counts and dilution factors, and the Lactobacillaceae-to-Enterobacteriaceae ratios were determined accordingly.

2.2.6. Statistical Analysis

Data were subjected to one-way analysis of variance (ANOVA) using the model Yij = μ + τi + εij where Yij is the observed response, μ is the overall mean, τi is the effect of the i = bacterial treatment, and εij is the random error. Statistical analyses were conducted in R version 4.3.3 with the ‘Agricolae’ package [33], and multiple comparisons were performed using Tukey’s HSD test when significant differences were detected (p < 0.05).

3. Results

3.1. Experiment 1: Effects of Cell Wall Materials and Particle Size on the Characteristics of the DMP Formulation

3.1.1. Encapsulation Efficiency and Stability

The encapsulation yield and probiotic viability were significantly influenced (p < 0.001) by both the wall material and particle size (

Table 1. Effects of wall material and particle size on encapsulation efficiency and stability of DMP formulations.

Item	Encapsulation Efficiency		Stability Under Different Conditions
	Yield (%)	Viability (%)	Pepsin Tolerance at pH 2.5	Pancreatin Tolerance at pH 7.0	Thermal Tolerance at 100 °C for 3 min
Interaction A × B
Corn
0.6 mm	20.07	97.56 AB	95.71 AB	94.35	95.10
1.0 mm	18.77	93.88 D	91.75 C	92.59	92.78
Defatted rice bran
0.6 mm	20.31	97.30 AB	96.27 AB	96.87	97.72
1.0 mm	18.59	95.18 C	94.75 B	94.79	94.28
Wheat bran
0.6 mm	20.25	98.05 A	96.70 A	96.63	98.03
1.0 mm	18.48	96.74 B	94.98 AB	93.61	94.61
Factor A (Wall material)
Corn	19.41	95.72 B	93.73 B	93.47 B	93.94 B
Defatted rice bran	19.54	96.24 B	95.51 A	95.83 A	96.00 A
Wheat bran	19.78	97.40 A	95.84 A	95.12 A	96.32 A
Factor B (Particle size)
0.6 mm	20.21 A	97.64 A	96.23 A	95.95 A	96.95 A
1.0 mm	18.75 B	95.27 B	93.83 B	93.66 B	93.89 B
SEM	0.234	0.362	0.413	0.417	0.481
p-value
Factor A	0.945	<0.001	<0.001	0.003	<0.001
Factor B	0.001	<0.001	<0.001	<0.001	<0.001
Factor A × B	0.907	<0.001	0.014	0.506	0.401

SEM, standard error of the mean. ^A,B,C,D Different superscripts in the same column indicate significant differences (p < 0.01).

). Wheat bran exhibited the highest viability (97.40%), followed by defatted rice bran (96.24%) and corn (95.72%) (p < 0.001). Smaller particle size (0.6 mm) resulted in greater yield (121.24%) and viability (97.64%) than the larger size (1.0 mm; 112.22% and 95.27%, respectively; p < 0.001). No interaction between wall material and particle size was detected for the yield (p = 0.907).

Stability under simulated gastrointestinal and thermal conditions also varied significantly (p = 0.003). Wheat and defatted rice bran conferred greater acid, pancreatin, and thermal tolerance than corn. Compared to 1.0 mm particles, encapsulation with 0.6 mm particles consistently improved acid tolerance (96.23%), pancreatin tolerance (95.95%), and thermal resistance (96.95%) (p < 0.001). A significant interaction between wall material and particle size was observed only for acid tolerance (p = 0.014). The greatest acid tolerance was observed in wheat bran at 0.6 mm (96.70%), whereas corn at 1.0 mm had the lowest (91.75%). No interaction effects were detected for pancreatin or thermal tolerance (p ≥ 0.401).

3.1.2. Physical Properties

The physical properties of DMP were influenced by both the wall material and particle size (

Table 2. Effects of wall material and particle size on physical properties of DMP formulation.

Item	Density (g/100 mL)		Carr’s Index (%)	Water Activity (Aw)	Product Color
	Bulk	Tapped			L*	a*	b*
Interaction A × B
Corn
0.6 mm	50.91	75.77	33.28 A	0.17	50.47 B	2.53 E	22.93 A
1.0 mm	47.05	69.61	31.36 AB	0.15	50.17 B	2.46 E	22.58 A
Defatted rice bran
0.6 mm	45.49	60.37	24.49 C	0.19	54.53 A	3.05 C	20.59 C
1.0 mm	41.49	55.70	27.97 B	0.19	54.04 A	2.73 D	20.52 C
Wheat bran
0.6 mm	36.98	50.85	28.06 B	0.19	50.31 B	3.88 A	21.58 B
1.0 mm	32.63	46.57	29.88 AB	0.19	50.73 B	3.40 B	19.84 C
Factor A (Wall material)
Corn	48.98 A	72.69 A	32.32 A	0.16 B	50.32 B	2.50 C	25.25 A
Defatted rice bran	43.49 B	58.03 B	26.23 C	0.19 A	53.29 A	2.89 B	20.55 B
Wheat bran	34.81 C	48.71 C	28.97 B	0.19 A	50.52 B	3.64 A	20.71 B
Factor B (Particle size)
0.6 mm	44.46 A	62.33 A	28.61	0.19	51.77 A	3.15 A	21.70 A
1.0 mm	40.39 B	57.29 B	29.73	0.18	49.65 B	286 B	19.98 B
SEM	1.504	2.483	0.721	0.004	0.821	0.106	0.246
p-value
Factor A	<0.001	<0.001	<0.001	<0.001	<0.001	<0.001	0.005
Factor B	<0.001	<0.001	0.080	0.472	<0.001	<0.001	<0.001
Factor A × B	0.801	0.358	0.008	0.537	<0.001	<0.001	<0.001

SEM, standard error of the mean. ^A,B,C,D,E Different superscripts in the same column indicate significant differences (p < 0.01).

). The bulk and tapped densities were the highest in corn-based formulations (48.98 and 72.69 g/100 mL, respectively), intermediate in defatted rice bran, and lowest in wheat bran (p < 0.001). The DMP produced with 0.6 mm particles had greater bulk and tapped densities than that produced with 1.0 mm particles (p < 0.001). Significant differences in flowability quantified by Carr’s index were observed among different cell wall materials (p < 0.001), and corn showed the greatest value of 32.32%. A significant interaction between wall material and particle size was found in Carr’s index (p = 0.008), with the greatest value (33.28%) produced by corn at a particle size of 0.6 mm. Water activity varied with wall material (p < 0.001), with defatted rice bran and wheat bran powders exhibiting greater values (0.19 and 0.19, respectively) than corn (0.16).

The particle size did not significantly affect the water activity (p = 0.472). Product color parameters (L*, a*, and b*) were significantly affected by wall material and particle size (p < 0.001). Defatted rice bran produced the brightest powder (L* = 53.29), wheat bran exhibited the highest redness (a* = 3.64), and corn had the highest yellowness (b* = 25.25). Significant interaction effects were observed for all color attributes (p < 0.001), with defatted rice bran at 0.6 mm exhibiting the greatest lightness (L* = 54.53) and wheat bran at the same particle size displaying the most intense red hue (a* = 3.88).

3.1.3. Nutrient Composition

DM and OM were unaffected (p > 0.05) by the wall material, particle size, or their interaction (

Table 3. Effects of wall material and particle size on nutrient composition of DMP formulations.

Item	Nutrient Composition (%)
	DM	OM	CP	EE	CF	GE (kcal/kg)
Interaction A × B
Corn
0.6 mm	99.97	99.24	9.62	3.24	1.31	4332.14 c
1.0 mm	99.97	99.30	9.64	3.27	1.31	4334.60 c
Defatted rice bran
0.6 mm	99.98	99.23	18.49	0.85	6.35	4453.83 b
1.0 mm	99.97	99.18	18.45	0.85	6.31	4451.05 b
Wheat bran
0.6 mm	99.97	99.25	18.59	2.92	7.60	4543.53 a
1.0 mm	99.95	99.19	18.61	2.92	7.60	4540.15 a
Factor A (Wall material)
Corn	99.97	99.27	9.63 B	3.26 A	1.31 C	4333.37 C
Defatted rice bran	99.98	99.21	18.47 A	0.85 C	6.33 B	4452.44 B
Wheat bran	99.96	99.22	18.60 A	2.92 B	7.60 A	4541.84 A
Factor B (Particle size)
0.6 mm	99.97	99.24	15.56	2.35	5.08	4443.17
1.0 mm	99.96	99.22	15.56	2.34	5.08	4441.94
SEM	0.005	0.013	1.018	0.260	0.658	20.718
p-value
Factor A	0.315	0.084	<0.001	<0.001	<0.001	<0.001
Factor B	0.458	0.433	0.977	0.649	0.662	0.216
Factor A × B	0.888	0.076	0.934	0.855	0.594	0.050

SEM, standard error of the mean; DM, dry matter; OM, organic matter; CP, crude protein; EE, ether extract; CF, crude fiber; GE, gross energy. ^a,b,c Different superscripts in the same column indicate significant differences (p ≤ 0.05). ^A,B,C Different superscripts in the same column indicate significant differences (p < 0.01).

). The CP content cell wall materials differed significantly among (p < 0.001). Corn-based DMP had the lowest CP content (9.63%), whereas defatted rice bran (18.47%) and wheat bran (18.60%) had significantly greater values. The opposite trend (p < 0.001) was observed for EE, with the highest level in corn (3.26%), intermediate in wheat bran (2.92%), and the lowest in defatted rice bran (0.85%). The CF also varied with the wall material (p < 0.001). Wheat bran exhibited the greatest CF (7.60%), followed by defatted rice bran (6.33%), whereas corn contained the lowest CF (1.31%). The particle size did not influence CP, EE, or CF (p > 0.05). The GE was strongly affected by wall material (p < 0.001), being the highest in wheat bran (4541.84 kcal/kg), moderate in defatted rice bran (4452.44 kcal/kg), and lowest in corn (4333.37 kcal/kg). Particle size did not have a significant effect (p = 0.216), but a marginal interaction was observed (p = 0.050), with 0.6 mm wheat bran exhibiting the greatest GE of 4543.53 kcal/kg.

3.1.4. Morphological Assessments

The structure and surface characteristics of the DMP formulation were analyzed using scanning electron microscopy (Figure 2). Scanning electron micrographs clearly depict the effective incorporation of P. acidilactici V202 and YLE within the porous framework of the wheat bran matrix. Microencapsulation via lyophilization (freeze-drying) technology resulted in a well-defined, stable double-layered structure that potentially protects the bioactive components during gastrointestinal enzyme action and heat. Porous wheat bran served as an effective carrier matrix, providing physical protection and controlled release properties for the encapsulated materials.

3.2. Experiment 2: Effect of Probiotic Strain Variation on the Characteristics and Functional Properties of the DMP Formulations

3.2.1. Encapsulation Efficiency, Product Quality, and Nutrient Composition

The encapsulation efficiency of DMP remained high across all treatments, with yields ranging from 20.08% for S. thermophilus to 20.38% for L. plantarum (

Table 4. Effect of probiotic strain variation on encapsulation efficiency, product quality, and nutrient composition of DMP formulations.

Item	DMP Formulation			SEM	p-Value
	P. acidilactici	L. plantarum	S. thermophilus
Encapsulation efficiency
Yield (%)	20.21	20.38	20.08	0.115	0.644
Viability (%)	97.65	97.66	97.38	0.256	0.905
Water activity (aw)	1.95	1.96	1.93	0.010	0.623
Color
Lightness (L*)	52.47	52.04	52.32	0.819	0.982
Redness (a*)	3.61	3.68	3.68	0.050	0.830
Yellowness (b*)	21.38	22.12	21.47	0.626	0.898
Bulk density (g/100 mL)	35.80	35.87	36.30	0.331	0.845
Tapped density (g/100 mL)	50.57	50.62	50.61	0.544	0.999
Compressibility of powder ratio (%)	29.13	29.08	28.25	0.875	0.922
Nutrient composition (%)
Dry matter	99.95	99.94	99.96	0.012	0.853
Organic matter	99.18	99.18	99.14	0.037	0.911
Crude protein	18.39	18.40	18.34	0.090	0.970
Ether extract	2.89	2.91	2.84	0.020	0.445
Crude fiber	7.58	7.59	7.43	0.041	0.194
Gross energy (kcal/kg)	4521.46	4524.30	4505.73	10.924	0.805

SEM, standard error of the mean; DMP, double-layered microencapsulated phytosynbiotic.

). No significant differences were detected among the treatments (p > 0.05). Probiotic viability after lyophilization remained consistently high (97.38–97.66%; p > 0.05). The water activity was low and uniform across all formulations (1.93–1.96; p > 0.05). The DMP exhibited similar color profiles, with lightness (L*) ranging from 52.04 to 52.47, redness (a*) from 3.61 to 3.68, and yellowness (b*) from 21.38 to 22.12 (p > 0.05). Bulk density (35.80–36.30 g/100 mL) and tapped density (50.57–50.62 g/100 mL) were comparable, resulting in similar compressibility ratios (28.25–29.13%; p > 0.05). Nutrient composition was consistent among treatments: DM exceeded 99.94%; OM ranged from 99.14 to 99.18%; CP from 18.34 to18.40%; EE from 2.84 to 2.91%; and CF from 7.43 to7.59%. The GE content varied minimally between 4505.73 and 4524.30 kcal/kg, with no significant differences among the probiotic strains (p > 0.05).

3.2.2. Stability of Probiotics Within the DMP Formulation

DMP demonstrated high stability under simulated gastrointestinal conditions, thermal stress, and refrigerated storage (

Table 5. Effect of probiotic strain variation on their stability against gastrointestinal enzymes, thermal processing, and long-term storage of DMP formulations.

Item	DMP Formulation			SEM	p-Value
	P. acidilactici	L. plantarum	S. thermophilus
Pepsin tolerance at pH 2.5 (%)	95.67	94.83	94.38	0.373	0.409
Pancreatin tolerance at pH 7.0 (%)	94.78	94.00	94.29	0.347	0.711
Thermal treatment at 100 °C (%)
2.5 min	98.90	98.18	99.45	0.354	0.234
5.0 min	95.39	96.13	95.43	0.333	0.173
10.0 min	0.00	0.00	0.00	0.000	-
Storage time at 4 °C (%)
0 months	100.00	100.00	100.00	0.000	-
2 months	99.94	99.61	99.45	0.155	0.472
4 months	100.00	99.61	99.81	0.128	0.529
6 months	99.82	99.28	99.57	0.257	0.746

SEM, standard error of the mean; DMP, double-layered microencapsulated phytosynbiotic.

). Survival rates under pepsin (pH 2.5) ranged from 94.38% (S. thermophilus) to 95.67% (P. acidilactici), while survival under pancreatin (pH 7.0) varied from 94.00% to 94.78%, with no significant differences among strains (p > 0.05). Thermal tolerance at 100 °C was maintained during short exposure times, with 2.5 min and 5 min treatments yielding survival rates between 98.18% and 99.45% and between 95.39% and 96.13%, respectively. Prolonged heating for 10 min led to complete enzyme inactivation. During storage at 4 °C, viability remained consistently above 99% across all strains for up to six months, with no significant differences (p > 0.05). These findings indicate that DMPs possess excellent stability under conditions simulating gastrointestinal passage, short-term heat exposure, and long-term cold storage.

3.2.3. Phytochemical Composition and Biological Activities

The DMP formulations, developed by combining YLE with double-layered microencapsulated probiotics within a wheat bran matrix, exhibited consistent phytochemical profiles and bioactive across all evaluated probiotic strains (

Table 6. Phytochemical composition and biological activities of DMP formulations.

Item	DMP Formulation			SEM	p-Value
	P. acidilactici	L. plantarum	S. thermophilus
Phytochemical composition
Tannic acid (mg/g)	12.69	12.45	12.61	0.220	0.312
Total phenolic (mg GAE/g)	16.85	16.60	16.79	0.250	0.278
Total flavonoids (mg catechin/g)	10.56	10.42	10.52	0.075	0.419
Total carotenoid (mg/g)	4.26	4.22	4.27	0.045	0.435
Total chlorophyll (mg/g)	0.49	0.47	0.48	0.030	0.396
Total antioxidant (mg AEAC/g)	20.41	20.30	20.39	0.150	0.372
Biological activities
DPPH (IC50, mg/g)	1.99	0.20	0.20	0.008	0.421
ABTS (IC50, mg/g)	0.08	0.08	0.01	0.005	0.458
FRAP (mmol Fe2+/g)	0.06	0.05	0.05	0.002	0.389

SEM, standard error of the mean; DMP, double-layered microencapsulated phytosynbiotic; DPPH, 2,2-diphenyl-1-picrylhydrazyl; ABTS, 2,2-azino-bis-3-ethylbenzothiazoline-6-sulphonic acid; FRAP, ferric reducing antioxidant power.

). Quantitative analysis revealed similar concentrations of tannic acid, total phenolics, flavonoids, carotenoids, and chlorophyll among P. acidilactici, L. plantarum and S. thermophilus (p > 0.05). Furthermore, the total antioxidant capacity, assessed via DPPH, ABTS, and FRAP assays, did not differ significantly among the treatments (p > 0.05). These findings indicate that probiotic strain variation does not influence phytochemical retention or preservation of bioactivity. The double-layered microencapsulation process, which utilizes Yanang mucilage as the primary protective matrix, wheat bran as the secondary carrier, and lyophilization for stabilization, serves as the primary determinant of bioactive compound stability. Consequently, the observed antioxidant capacities in DMP formulations are attributable to the intrinsic phytochemical constituents of the YLE rather than strain-specific microbial metabolism.

3.2.4. Ratio of the Minimal Inhibitory Concentration to the Minimal Bactericidal Concentration

The antibacterial potential of the DMP culture supernatants was assessed against B. cereus, S. aureus, and E. coli (

Table 7. Effect of probiotic strain variation of DMP formulations on ratio of minimal inhibitory concentration (MIC) to minimal bactericidal concentration (MBC).

Item	MIC/MBC (%v/v)
	B. cereus	S. aureus	E. coli
P. acidilactici	50/>50	25/>50	25/>50
L. plantarum	50/>50	50/>50	25/>50
S. thermophilus	50/>50	50/>50	25/>50
Vancomycin	1/2	0.5/1	NA
Gentamicin	NA	NA	0.5/1

MIC, minimal inhibitory concentration; MBC, minimal bactericidal concentration; NA, not applicable.

). All tested formulations inhibited the growth of B. cereus at 50% (v/v), although no bactericidal effect was observed at this concentration (>50% v/v). Against S. aureus, and P. acidilactici exhibited the lowest minimum inhibitory concentration (MIC) at 25% (v/v), whereas L. plantarum and S. thermophilus required 50% (v/v). In all cases, the minimum bactericidal concentrations (MBCs) exceeded 50% (v/v), indicating a predominant bacteriostatic activity. The growth of E. coli was inhibited at 25% (v/v) for all strains, with no detectable bactericidal effect. These findings indicate that DMP formulations exert moderate, strain-dependent antibacterial activity, which is more pronounced against Gram-positive bacteria and primarily mediated through bacteriostatic mechanisms rather than bacterial cell death.

3.2.5. Probiotic Counts and Viability of the DMP Formulations Following In Vitro Ileal Digestibility

The total viable counts (TVC) of the encapsulated probiotics remained consistently high across all formulations (

Table 8. Viability and population dynamics of probiotics of DMP formulations during post in vitro ileal digestion.

Item (Log CFU/mL)	DMP Formulation			SEM	p-Value
	P. acidilactici	L. plantarum	S. thermophilus
Total viable count
Initial	8.64	8.68	8.60	0.025	0.465
Final	8.46	8.51	8.45	0.024	0.603
Viability (%)	97.89	97.99	98.20	0.286	0.920
Lactobacillaceae
Initial	8.55	8.51	8.55	0.017	0.739
Final	8.21	8.29	8.27	0.066	0.368
Viability (%)	96.73	97.18	96.25	0.608	0.251

SEM, standard error of the mean; DMP, double-layered microencapsulated phytosynbiotic.

). The initial TVC ranged from 8.60 log CFU/mL in the S. thermophilus group to 8.68 log CFU/mL in the L. plantarum group. Following in vitro ileal digestion, final TVC values decreased slightly but remained robust, ranging from 8.45 to 8.51 log CFU/mL, corresponding to overall survival rates of 97.89–98.20%, with no statistically significant differences among the probiotic strains (p > 0.05). These results indicate that the encapsulation matrix effectively protected probiotics from digestive stress during the ileal phase.

3.2.6. Gas Production and Degradation Kinetics of the DMP Formulations During In Vitro Cecal Fermentation

During the early incubation period (4–16 h), gas production differed significantly among the probiotic strains (p < 0.05;

Table 9. Influence of probiotic strain variation on gas production and degradation kinetics of DMP formulations during in vitro cecum fermentation.

Item	DMP Formulation			SEM	p-Value
	P. acidilactici	L. plantarum	S. thermophilus
Gas production in different incubation times (mL)
4 h	9.36 a	7.40 b	6.65 b	0.415	0.005
8 h	17.76 a	15.75 b	14.53 b	0.534	0.018
12 h	24.62 a	21.65 b	21.45 b	0.583	0.022
16 h	31.26 a	28.57 ab	27.54 b	0.590	0.040
20 h	34.78	33.84	32.90	0.583	0.465
24 h	38.50	38.41	37.62	0.586	0.825
Kinetics of degradation *
P (mL)	30.38	31.19	30.82	0.459	0.7941
a (mL)	−0.95 a	−1.94 b	−2.32 b	0.229	0.022
b (mL)	56.01 b	71.09 a	75.08 a	2.946	0.004
c (% hour)	0.05 a	0.04 b	0.03 b	0.003	0.009
d (mL)	56.97 b	73.03 a	77.40 a	3.131	0.004

* Degradation kinetic were measured as: P, the volume of gas produced (mL) at incubation time t (h); a, represents gas production from the rapidly fermentable (upper gut digestible) fraction; b, denotes gas production from the slowly fermentable (cecal fermentation) fraction; c, is the rate constant of gas production for fraction b; d = (a + b) potential extent of gas production. SEM, standard error of the mean; DMP, double-layered microencapsulated phytosynbiotics. ^a,b Different superscripts in the same row indicate significant differences (p < 0.05).

and Figure 3). Compared with L. plantarum and S. thermophilus, P. acidilactici generated the greatest gas volumes at 4 h (9.36 mL), 8 h (17.76 mL), 12 h (24.62 mL), and 16 h (31.26 mL). No significant differences were observed among the treatments at 20 h and 24 h (p > 0.05), indicating that gas production had plateaued at these later time points. Furthermore, the degradation kinetics varied significantly depending on the probiotic strain. The rapidly fermentable fraction (a) was highest in P. acidilactici (−0.95 mL) and lower in L. plantarum (−1.94 mL) and S. thermophilus (−2.32 mL; p = 0.022), suggesting faster initial substrate utilization by P. acidilactici. The slowly fermentable fraction (b) was greater in L. plantarum (71.09 mL) and S. thermophilus (75.08 mL) than in P. acidilactici (56.01 mL; p = 0.004), indicating extended cecal fermentation for these strains. The total degradable fraction (d) mirrored this trend (L. plantarum, 73.03 mL; S. thermophilus, 77.40 mL; P. acidilactici, 56.97 mL; p = 0.004). Additionally, the fermentation rate constant (c) of fraction b was highest for P. acidilactici (0.05%/h; p = 0.009), reflecting a faster initial fermentation rate relative to the other strains.

3.2.7. Lactic Acid, Volatile Fatty Acids, and Cecal Microbiology of the DMP Formulation After In Vitro Cecal Fermentation

The lactic acid concentration was overall greatest with the P. acidilactici treatment (15.16 mM), which was significantly greater (p = 0.004) than that observed with L. plantarum (14.26 mM) and S. thermophilus (14.55 mM) (

Table 10. Evaluation of lactic acid, short-chain fatty acids, and cecal microbiology of DMP formulations subjected to in vitro cecal fermentation.

Item	DMP Formulation			SEM	p-Value
	P. acidilactici	L. plantarum	S. thermophilus
Lactic acid content (mMol)	15.16 A	14.26 B	14.55 B	0.145	0.004
Volatile fatty acids content (mMol)
Acetic acid (C2)	22.99 A	21.04 B	21.06 B	0.326	<0.001
Propionic acid (C3)	5.17 a	4.93 b	5.07 ab	0.044	0.052
Butyric acid (C4)	0.84 A	0.78 B	0.82 A	0.011	0.007
Valeric acid (C5)	0.79 A	0.46 B	0.39 B	0.062	<0.001
Total VFAs	30.12 A	27.50 B	27.66 B	0.450	0.001
Cecal microbiology content (Log CFU/mL)
Total viable count	9.17	9.13	9.16	0.017	0.697
Lactobacillaceae	8.59	8.62	8.87	0.046	0.101
Enterobacteriaceae	6.94	7.05	6.84	0.045	0.188
L:E ratio	1.23	1.26	1.24	0.027	0.632

SEM, standard error of the mean; DMP, double-layered microencapsulated phytosynbiotics; L, Lactobacillaceae; E, Enterobacteriaceae. ^a,b Different superscripts in the same row indicate significant differences (p < 0.05). ^A,B Different superscripts in the same row indicate significant differences (p < 0.01).

). Total volatile fatty acid (VFA) production followed a similar pattern, with P. acidilactici producing 30.12 mM compared with 27.50–27.66 mM for the other formulations (p = 0.001). Analysis of individual VFAs revealed that acetic acid (C2) and butyric acid (C4) were significantly greater in P. acidilactici (22.99 and 0.84 mM, respectively; p < 0.01). In addition, valeric acid (C5) exhibited strain-dependent differences (p < 0.001), while there was a tendency for propionic acid (C3) to increase (p = 0.052). The cecal microbial populations remained largely unaffected by the different formulations. The total viable counts ranged from 9.13 to 9.17 log CFU/mL (p = 0.697), and Lactobacillaceae abundance varied between 8.59 and 8.87 log CFU/mL (p = 0.101). Enterobacteriaceae counts were similar across treatments (6.84–7.05 log CFU/mL; p = 0.188), resulting in L:E ratios of 1.23–1.26 without significant differences (p = 0.632). These findings indicate that P. acidilactici–based DMP enhances lactic acid and VFA production during in vitro cecal fermentation while maintaining stable cecal microbial populations.

4. Discussion

The present study demonstrated that both wall material and particle size significantly influenced the encapsulation efficiency, stability, physical characteristics, and nutritional composition of DMP formulations composed of P. acidilactici V202 and YLE. Wheat bran emerged as the most effective wall material, yielding the highest probiotic viability (97.40%) and superior tolerance to gastrointestinal and thermal stress compared with defatted rice bran and corn. The enhanced performance of wheat bran can be attributed to its highly porous, fibrous structure, high β-glucan content, and abundant dietary fiber [55], which collectively provide a protective matrix for probiotic cells during lyophilization and subsequent storage [10]. A smaller particle size (0.6 mm) further improved encapsulation efficiency, probiotic viability, and stress tolerance by increasing the surface area and reducing interparticle voids, thereby promoting more uniform coverage and immobilization of probiotics within the wall matrix [56]. The observed interaction between wall material and particle size for acid tolerance underscores the importance of optimizing both factors to maximize probiotic survival under harsh conditions.

The DMP formulation, composed of P. acidilactici V202 combined with YLE and double-layer microencapsulation within wheat bran, exhibited a well-defined morphology, as confirmed by scanning electron microscopy (Figure 2), demonstrating the successful incorporation of both probiotics and YLE within the porous wheat bran matrix. The lyophilization process produced a stable, double-layered structure capable of protecting bioactive components from gastrointestinal enzymes and thermal stress while maintaining the structural integrity of encapsulated probiotics. During fermentation, the viscous YLE likely formed a protective hydrogel-like coating, enhancing bacterial stability and tolerance to environmental stressors, particularly at low temperatures [57]. This observation aligns with previous studies [58], which reported that biopolymer-based coatings in probiotic microencapsulation improved cell viability by providing physical protection and mitigating stress during processing and storage. This protective effect is likely due to the high polysaccharide content and unique rheological properties of YLE. Additionally, extracellular polysaccharides produced by lactic acid bacteria during fermentation [59] contribute to both physical and biochemical barriers, creating a microenvironment with antioxidant activity that mitigates oxidative stress and preserves bacterial viability during fermentation and storage [60].

The superior performance of wheat bran at 0.6 mm particle size can be mechanistically attributed to its unique physicochemical properties. Wheat bran’s high arabinoxylan content (approximately 70% of the total dietary fiber) [6], characterized by feruloylated arabinoxylan side chains, endows it with an exceptional water-holding capacity, which is instrumental in maintaining microcapsule hydration during lyophilization, thereby preventing desiccation of probiotic cells. In addition, the water-holding and gel-forming properties of the extract maintain cellular hydration and reduce osmotic and freeze–thaw stress, further supporting the viability of probiotics [61]. Our data suggest that the 0.6 mm particle size optimizes the surface area-to-volume ratio (increased by 45% vs. 1.0 mm), thus enhancing adsorption of Yanang mucilage into the bran pores. The porous structure and internal matrix of wheat bran serve as an effective physical carrier, providing an extensive surface area and a network of insoluble and fermentable dietary fibers, including arabinoxylans and cellulose [10,62], which facilitate microbial attachment and protection against environmental stressors, such as acidity, heat, and oxygen exposure.

The diffusion of viscous, polysaccharide-rich YLE into wheat bran pores enables deeper immobilization of probiotics [63], forming a double-layer encapsulation system that enhances shielding from gastric acid and bile while supporting gradual release in the intestine [64]. This composite strategy not only improves encapsulation efficiency and storage stability under varying conditions but also enables the controlled release of both probiotics and Yanang-derived bioactive compounds, maximizing their bioavailability and functional effects. Furthermore, the small particle size, high porosity, and β-glucan content of wheat bran contributed to superior probiotic viability (97.40%), outperforming that of defatted rice bran (96.24%) and corn (95.72%). Its porous matrix provides physical protection, retains bioactive compounds, and combined with its inherent prebiotic properties, supports microbial colonization and gut health. The polysaccharides and dietary fibers in wheat bran synergistically reinforce the encapsulation matrix, further promoting survival during storage and gastrointestinal transit [65]. Collectively, these findings demonstrate that the combination of YLE and wheat bran constitutes a robust and efficient strategy for protection during processing, storage, and delivery, highlighting its potential application in functional feed and food systems, where stability, bioactivity, and controlled release of probiotics are critical for efficacy.

The physical and nutritional characteristics of DMP are largely determined by the type of wall material and particle size, which collectively influence the density, flowability, water activity, color, and macronutrient composition, ultimately affecting the stability and functionality of the encapsulated probiotics. Corn-based powders exhibit the highest bulk and tapped densities [66] owing to their dense and starch-rich structure, which facilitates tighter particle packing. Wheat bran powders have the lowest densities, reflecting their highly porous and fibrous architecture [67]. Smaller particle sizes (0.6 mm) increased packing efficiency and reduced interparticle voids [68], particularly in corn powders, demonstrating a significant wall material × particle size interaction in powder handling and flowability. Flowability, evaluated via the compressibility index, was lowest in corn powders, while wheat bran and defatted rice bran powders exhibited higher flowability due to their fibrous and porous matrices that reduce interparticle cohesion [69]. The higher water activity observed in wheat bran and defatted rice bran powders was attributed to the hygroscopic nature of dietary fibers, which provide water-binding sites that can influence probiotic survival and shelf life [70]. Color attributes are also dependent on wall material and particle size [71]. Defatted rice bran powders were lighter (higher L*), wheat bran powders exhibited higher redness (a*), and corn powders displayed greater yellowness (b*). Smaller particles generally enhance lightness and uniformity by promoting an even microcapsule distribution. From a nutritional perspective, wheat bran powders exhibit the highest levels of CP, CF, and GE [72], providing a supportive matrix for both probiotic proliferation and prebiotic activity. In contrast, corn powder is richer in EE [73] but contains lower protein and fiber levels, whereas defatted rice bran has intermediate nutritional values. These differences in macronutrient composition, together with the structural characteristics of the cell wall materials, play a key role in influencing the probiotic encapsulation efficiency and stability [74]. Overall, the nutrient content was predominantly determined by the type of wall material, with particle size exerting only a minor effect, highlighting that the intrinsic properties of the carrier largely dictate the nutritional profile of the DMP.

The current findings demonstrate that DMP, prepared using YLE and wheat bran for double microencapsulation of probiotics, successfully preserves its phytochemical profile and bioactive functions, regardless of the probiotic strains involved. Tannins, total phenolics, flavonoids, carotenoids, and chlorophyll contents were comparable among P. acidilactici, L. plantarum, and S. thermophilus, demonstrating that the microencapsulation process preserves the integrity of Yanang-derived bioactive compounds from degradation caused by oxygen, moisture, and adverse conditions [75]. Antioxidant capacity, as evaluated by DPPH, ABTS, and FRAP assays, remained consistent across probiotic strains, indicating that the choice of probiotic does not compromise the free radical scavenging potential or reducing power of the powders [76]. These results align with previous reports showing that polysaccharide-rich plant extracts and hydrogel-forming carriers [77], such as wheat bran, provide a protective environment that stabilizes phytochemicals during processing and storage, preventing degradation under thermal or oxidative stress. In addition, the antibacterial activity of DMP was moderate and primarily bacteriostatic [78], with greater efficacy against Gram-positive (B. cereus and S. aureus) than Gram-negative bacteria (E. coli). The observed differential inhibition likely reflects the structural differences in the bacterial cell wall. Gram-positive bacteria possess a thick peptidoglycan layer that is more susceptible to tannins and polyphenols [79], which can bind to cell wall proteins, disrupt membrane integrity, and interfere with enzymatic activities [80]. In contrast, the outer membrane of Gram-negative bacteria confers partial resistance, limiting the penetration of phytochemicals. Among the probiotic strains, P. acidilactici supernatant exhibited significant inhibitory activity against S. aureus, with a minimum inhibitory concentration of approximately 10–20% v/v, primarily exerting a bacteriostatic effect by delaying bacterial growth rather than directly killing the cells [81]. This inhibition is likely driven by the synergistic action of organic acids, such as lactic acid, and bacteriocin-like compounds, including pediocin, which are known to target Gram-positive pathogens [82,83]. The susceptibility of S. aureus can be attributed to its thick peptidoglycan cell wall [84], which is more vulnerable to tannins and polyphenols [85]. These plant-derived bioactives can bind to cell wall proteins, compromise membrane integrity [86], and interfere with enzymatic functions, thereby enhancing bacteriostatic effects [87]. Although bactericidal activity was not observed, the combined action of P. acidilactici metabolites and YLE bioactives effectively limited pathogen proliferation while maintaining the overall microbial balance. This suggests that P. acidilactici inhibits the growth of S. aureus primarily via bacteriostatic mechanisms, with its activity enhanced by tannins and polyphenols, offering an effective approach for pathogen control in the developed DMP formulations.

In the present study, phytosynbiotic formulations derived from YLE and containing double-microencapsulated probiotics exhibited distinct strain-dependent influences on in vitro cecal fermentation dynamics. The DMP formulation with P. acidilactici resulted in higher gas production during the early incubation period (4–16 h) and had the greatest rapidly fermentable fraction and fermentation rate, whereas L. plantarum and S. thermophilus exhibited slower and more prolonged fermentation. P. acidilactici also enhanced lactic acid and total SCFA production, particularly acetic and butyric acids, which are critical for maintaining gut barrier integrity [62], supporting energy metabolism, and facilitating microbial cross-feeding. However, the enhanced production of SCFA by P. acidilactici occurs through its heterofermentative phosphoketolase pathway rather than the homofermentative metabolism route used by S. Thermophilus [88]. These effects occurred without altering the total cecal microbial populations, indicating that the observed changes were primarily driven by the metabolic activity of the encapsulated probiotics [89]. Combined with the high survival rate during simulated ileal digestion (97.88–98.20%) and stable retention of phytochemicals, these results confirm the functional efficacy of the formulations.

The superior nutrient utilization and early fermentation of P. acidilactici can be attributed to its strong fermentative capacity, including rapid acid production and efficient exopolysaccharide (EPS) synthesis [90]. EPS contributes to substrate degradation, feed digestibility, and antioxidant protection against reactive oxygen species and lipid peroxidation, thereby supporting gut health [90]. The associated increase in SCFA production, particularly acetic and butyric acids, reinforces intestinal barrier function, supplies energy to colonocytes, modulates immune responses, and preserves microbial homeostasis [91]. Butyric acid, in particular, serves as a key energy source for colonocytes and promotes anti-inflammatory processes [92]. Enhanced SCFA production by P. acidilactici supports microbial cross-feeding in the gut, amplifying its beneficial effects on gut health. Specifically, P. acidilactici mainly produces acetic acid, which serves as a substrate for other gut microbes to convert into butyric acid, a critical SCFA for colonocyte energy and intestinal health [62]. This cross-feeding between microbes boosts butyrate levels, helping to maintain intestinal barrier integrity, modulate immune responses, reduce oxidative stress, and promote anti-inflammatory processes [93].

Increased SCFA production also correlates with a higher abundance of beneficial bacteria, such as Actinobacteria, Faecalibacterium, Lachnospiraceae, and Ruminococcaceae [94,95], which further contribute to gut homeostasis by continuously fermenting dietary fibers and supporting gut barrier function [96]. Thus, the interaction between P. acidilactici and other commensal microbes via SCFA cross-feeding enhances the overall gut microbial ecosystem and its health benefits. Furthermore, supplementation with P. acidilactici has been linked to increased antioxidant enzyme activity, reduced oxidative stress, and a higher abundance of beneficial gut bacteria, contributing to a balanced intestinal microbiome. Collectively, these attributes highlight P. acidilactici as a promising functional feed additive for improving gut fermentation efficiency, SCFA production, microbial balance, and overall gastrointestinal health in chickens.

While this study has demonstrated excellent in vitro performance of the DMP formulations, the findings constitute preliminary evidence that necessitates validation through in vivo trials with broilers. Assertions regarding the promotion of gut health, enhancement of productivity, optimal inclusion rates, cost-effectiveness, carcass quality, and improvements in immune parameters remain to be substantiated in practical settings. As such, future research should aim to establish practical dosage levels, assess long-term efficacy under commercial production conditions, evaluate economic viability in comparison to antibiotic alternatives, and investigate the impacts on broiler performance metrics and immune biomarkers. These critical next steps will determine the commercial applicability of DMP in antibiotic-free poultry systems.

5. Conclusions

We developed a double-layer microencapsulated phytosynbiotic by combining YLE, probiotics, and cereal by-products via lyophilization. Wheat bran (0.6 mm) provided superior encapsulation efficiency (20.2%), viability (97.4%), and stability under gastrointestinal conditions and thermal stress, particularly when combined with P. acidilactici, which exhibited superior fermentation activity and higher lactic acid and SCFA production. Together, these responses indicate enhanced probiotic gut health, antioxidant capacity, and moderate antibacterial activity against Gram-positive pathogens. These findings demonstrate the potential of scalable bioactive feed additives to support antibiotic-free poultry production. As such, compared with single-layer systems, the DMP advances One Health by valorizing Thai botanicals through lyophilization technology, achieving superior probiotic delivery. Despite all these positive attributes, limitations to the use of DMP include in vitro-only validation and a single-strain focus. To confirm the efficacy and economic viability of antibiotic-free systems, future research should be conducted in vivo with broilers to optimize multi-strain formulations, evaluate pelleted feed integration, and to assess commercial scalability.

6. Patents

A Thailand petty patent application (No. 2503004945) was filed on 15 December 2025, for the invention related to this research. The title of the invention is “The production process of phytosinbiotic products from Tiliacora triandra (Yanang) leaf extract and Pediococcus acidilactici V202 probiotics in double-layer microcapsule form using wheat bran as a carrier material.” The inventors are Tossaporn Incharoen, Manatsanun Nopparatmaitree and Noraphat Hwanhlem. All rights to this invention are owned by the Naresuan University.