Phytosynbiotic Containing Double-Layer Microencapsulated Pediococcus acidilactici V202 and Tiliacora triandra Leaf Extract Improve Growth Performance and Gut Health in Broiler Chickens

TL;DR

A new plant-based feed supplement improves chicken growth and gut health by combining probiotics and plant extract.

Contribution

A novel phytosynbiotic with microencapsulated probiotics and plant extract is shown to enhance poultry productivity sustainably.

Findings

DMP improved growth rate, feed efficiency, and economic returns in broiler chickens.

DMP increased lactic acid and short-chain fatty acids in gut fermentation.

DMP enhanced intestinal morphology and enriched beneficial gut bacteria.

Abstract

The novel phytosynbiotic formulation of double-layer microencapsulated Pediococcus acidilactici V202 and Tiliacora triandra leaf extract (DMP) represents a green and sustainable nutritional approach to modern poultry production. This natural feed innovation integrates plant-based bioactive compounds with beneficial probiotics to support gut health, improve growth performance, and reduce the reliance on antibiotic growth promoters. Dietary supplementation with the DMP improved the growth rate, feed efficiency, productive index, and economic returns in broiler chickens. These benefits were associated with improved nutrient digestibility and favorable changes in gut fermentation, including increased lactic acid and short-chain fatty acid (SCFA) production. The DMP also promoted healthier intestinal morphology and selectively enriched beneficial gut taxa, such as Lachnospirales, Oscillospirales, and Lactobacillales, which are linked to intestinal function. Overall, the DMP demonstrates strong potential as an environmentally friendly and sustainable feed supplement that supports nutrient utilization and productive efficiency while aligning with responsible and safe broiler-farming practices.

This study investigated the effects of a novel phytosynbiotic feed supplement derived from double-layer microencapsulated Pediococcus acidilactici V202 and Tiliacora triandra leaf extract (DMP) on the growth performance, nutrient utilization, gut fermentation, intestinal morphology, and cecal microbiota of broiler chickens. A total of 250 one-day-old male Ross 308 broilers were randomly assigned to five dietary treatments: basal control diet, antibiotic growth promoter (AGP) with chlortetracycline at 0.07%, and DMP supplementation at 0.25, 0.50, or 1.00% (w/w) for 42 days. Compared with the control diet, feeding the DMP led to linear or quadratic responses (p < 0.05) on average daily gain, feed efficiency, productive index, and economic returns. Apparent digestibility of dry matter, crude protein, and apparent metabolizable energy was enhanced in DMP-fed broilers, indicating improved nutrient utilization efficiency. These performance responses were accompanied by pronounced alterations in cecal fermentation, characterized by increased lactic acid, total volatile fatty acids, and particularly acetic and butyric acid levels (p < 0.01). Microbiome analysis revealed that the DMP selectively enriched fermentative SCFA-producing bacterial orders, including Lachnospirales, Oscillospirales, and Lactobacillales. It also reduced the relative abundance of less desirable taxa. As evidenced by an increased villus height and surface area in the duodenum and jejunum, along with a higher villus height-to-crypt depth ratio in the ileum, feeding the DMP also enhanced small intestinal morphology. These coordinated morphological adaptations are indicative of enhanced epithelial maturation and reduced crypt hyperplasia, likely mediated by elevated microbial SCFA production in the gut. In conclusion, the DMP improved broiler growth performance by coordinating the modulation of the gut microbiota, SCFA levels, and intestinal morphology, resulting in enhanced nutrient digestibility and productivity. This phytosynbiotic strategy represents a sustainable plant-based alternative to antibiotic growth promoters for environmentally responsible poultry production.

Figures

Full text

1. Introduction

Poultry production under green nutrition strategies has become a central focus amid accelerating environmental, economic, and social transitions. These shifts have driven the industry toward sustainable systems that maximize resource efficiency and minimize ecological burden. Broiler production, one of the fastest-growing livestock sectors, supplies affordable and high-quality animal protein essential for global food security [1]. Despite rapid progress, intensive broiler systems face persistent challenges, including fluctuating feed costs, climate-induced resource competition, and inefficient nutrient utilization [2]. Life cycle assessments indicate that poor feed conversion increases nutrient excretion and aggravates environmental impacts [3]. Additionally, the global shift to antibiotic-free poultry production has intensified due to the widespread public health threat of antimicrobial resistance driven by excessive antibiotic use in livestock. Antibiotic growth promoters (AGPs) historically enhanced growth and, despite approximately 75% of AGPs being consumed by food animals, their use facilitated the emergence of multidrug-resistant bacteria [4,5]. Poultry production faced challenges in maintaining performance and gut health following the European Union’s 2006 ban on AGPs. This catalyzed research into natural alternatives such as phytogenics, probiotics, and prebiotics, which exhibit antimicrobial, antioxidant, and immunomodulatory effects. These feed supplements improve gut microbiota balance and immunity, supporting sustainable, antibiotic-free poultry production [6,7].

Pediococcus acidilactici is recognized as a promising probiotic lactic acid bacterium for broiler production due to its beneficial effects on growth and health. This bacterium exerts antibacterial effects against pathogens such as Salmonella enteritidis and Escherichia coli, likely through bacteriocin production and lactic acid secretion, which inhibit pathogenic populations in the gut [8]. There are other strains of P. acidilactici isolated from broiler chickens with strong antimicrobial activities against common pathogens, highlighting its probiotic potential in poultry feed [9]. The modulation of broiler gut microbiota by P. acidilactici contributes to enhanced intestinal innate immunity and antioxidant capacity, supporting better health, disease resistance, improved growth performance, and reduced abdominal fat [10]. The anti-inflammatory effects of P. acidilactici have been demonstrated through the regulation of intracellular signaling pathways that reduce inflammatory mediators, thereby promoting gastrointestinal health and nutrient absorption [11]. Although P. acidilactici generally tolerates acidic conditions and bile action similar to those in the gastrointestinal tract, strain-specific variability exists; for example, strain P10 retains about 55% viability in bile salts, indicating partial susceptibility under harsh gut conditions [12]. Acid tolerance also varies among strains, with many enduring low gastric pH but prolonged or extreme acidity, potentially impairing survival [13].

Encapsulation is a widely recognized technique to maintain the viability of probiotics during processing, storage, and gastrointestinal transit. This protection allows the simultaneous delivery of viable probiotics and stable herbal phytochemicals in functional food products [14]. However, selecting appropriate encapsulation materials is crucial, as they must exhibit safety and non-toxicity while offering adequate protection for probiotic microorganisms and enabling controlled release [15]. Polysaccharides, lipids, proteins, gums, and mucilages derived from diverse sources are commonly utilized for probiotic encapsulation [16,17]. Due to its rich composition of dietary fibers, proteins, and bioactive compounds that enhance probiotic stability and functionality, wheat bran has emerged as a promising biomaterial for probiotic encapsulation. Recently, wheat bran has been considered a wall material for the encapsulation of probiotic bacteria [18] because its fiber matrix serves as a natural prebiotic substrate that supports probiotic colonization and survival. It provides fermentable carbohydrates that stimulate beneficial bacterial growth and enhance gut microbial balance [19,20]. Thus, this material appears suitable as a wall matrix for probiotic encapsulation.

Among the various phytogenic ingredients, Tiliacora triandra, a traditional medicinal plant commonly consumed in Southeast Asia, has been extensively studied for its phytochemical composition and bioactive properties with promising pharmaceutical applications. Phytochemical profiling revealed that Tiliacora triandra contains diverse bioactive compounds such as alkaloids, flavonoids, phenolics, and other secondary metabolites that contribute to its antioxidant, antimicrobial, anti-inflammatory, and anticancer properties [21]. In fact, it has been incorporated into multi-herbal formulations exhibiting potent antioxidant, anti-inflammatory, anticancer, and antimigration effects against various cancer cell lines, highlighting its role in integrative medicinal applications [22]. While serving as a natural encapsulation matrix [18], its mucilaginous leaf extract also provides polysaccharides acting as prebiotic-like substrates that promote beneficial bacteria, including Lactobacillus and Bifidobacterium [23].

Recently, phytosynbiotics have been introduced as a novel category of next-generation feed supplements that strategically integrate phytobiotics with synbiotics into a unified multifunctional formulation [18]. They integrate bioactive compounds from Tiliacora triandra leaf extract with probiotics and wheat bran-derived prebiotics. Unlike conventional synbiotics that primarily focus on microbial modulation, phytosynbiotics employ a proprietary double-layer microencapsulation system using lyophilization to achieve both co-encapsulation and functional synergy. The inner layer, made of fermented Tiliacora triandra mucilage, acts as a natural hydrogel matrix protecting probiotics during processing and gastrointestinal passage. The outer porous cereal bran layer functions simultaneously as a prebiotic carrier and a structural scaffold, ensuring targeted release in the distal gut. However, despite rising interest in phytosynbiotics, limited information using in vivo models exists on the synergistic effects of double-microencapsulated probiotics combined with phytogenic extracts. Thus, our laboratory developed a phytosynbiotic formulation consisting of Tiliacora triandra leaf extract combined with double-layer microencapsulated P. acidilactici V202 (DMP). We hypothesized that DMP would optimize intestinal health, improve metabolic efficiency, modulate host immunity, and enhance broiler growth performance. Accordingly, this study comprehensively evaluated its effects on nutrient utilization, gut microbiota composition, intestinal morphology, immune responses, and overall performance, providing mechanistic insights into the potential of advanced phytosynbiotic feed strategies for sustainable and resilient poultry production in the post-antibiotic era.

2. Materials and Methods

2.1. Animal Ethics

The present study was carried out at the Animal Nutrition Laboratory, Faculty of Agriculture, Natural Resources and Environment, Naresuan University, Phitsanulok, Thailand. All procedures involving animals were reviewed and approved by the Naresuan University Animal Care and Use Committee (Approval ID: 68 01 008) and were conducted in accordance with institutional guidelines for the ethical use of animals in research.

2.2. DMP Preparation

The phytosynbiotic was prepared by extracting and pasteurizing Tiliacora triandra leaves, then fermenting the extract with Pediococcus acidilactici V202 (1.5% v/v) at 37 °C for 24 h under anaerobic conditions. The fermented extract was mixed with wall material, absorbed into pretreated wheat bran under mild vacuum, frozen at −18 to −20 °C for 24 h, and lyophilized at 0.5 mbar for 48 h to obtain a stable microencapsulated powder stored at 4 °C. The encapsulated Pediococcus acidilactici V202 retained over 97% viability, indicating excellent probiotic stability. The final product had a bulk density exceeding 35 g/100 mL, with phytochemical analysis showing total phenolics above 16 mg GAE/g and tannic acid over 12 mg/g, both key contributors to DMP bioactivity. The morphology of the DMP was examined by scanning electron microscopy to verify the efficient entrapment of probiotic cells within the porous matrix (Figure 1). The observed microstructure demonstrated the structural integrity and potential functional efficacy of the double-layer encapsulation system, supporting its suitability as a phytosynbiotic supplement.

2.3. Experimental Design, Birds, Housing, and Diets

A total of 250 one-day-old male Ross 308 broilers were obtained from a commercial hatchery (Charoen Pokphand Foods PCL, Lamphun, Thailand) and randomly assigned to five dietary treatments: a basal diet without supplement (CON) or CON supplemented with 0.07% chlortetracycline (AGP) or with DMP at 0.25, 0.50, or 1.00% (w/w). This study was arranged as a completely randomized design (CRD) comprising five dietary groups, each with five replicate pens (10 birds/pen). All birds were reared in an environmentally controlled house equipped with an evaporative cooling system. Pens were bedded with rice hulls to a depth of approximately 3–4 inches. During the first week, the brooding temperature in the pen area was maintained at ~32 °C using a 100 W incandescent lamp. Thereafter, the temperature was gradually reduced by approximately 2.8–3.0 °C per week until reaching a stable range of 25–28 °C. Birds had ad libitum access to feed and water throughout the experimental period (42 days), and standard management and biosecurity practices were followed. Vaccination of birds followed standard commercial protocols. On day 1, chicks received Marek’s disease vaccine and Newcastle disease + infectious bronchitis vaccines as applied by the hatchery. On day 7, ND + IB vaccines were administered again, followed by infectious bursal disease vaccination on day 14. Mortality and clinical signs were monitored daily, and necropsies were performed when necessary to ascertain cause of death. Experimental diets comprised corn–soybean meal and followed a two-phase program (Table 1): starter (days 0–21) and finisher (days 22–42), aligned with NRC nutrient specifications for broilers [24]. All basal diets were formulated to be isocaloric and isonitrogenous, with variations only due to the test feed supplement. Diets were mixed in single-phase batches and analyzed for proximate composition and gross energy using standard AOAC methods [25].

2.4. Evaluation of Growth Performance and Economic Benefit Return

Data on feed consumption, body weight, and mortality were recorded for each pen throughout the experimental period. Measurements were taken at three growth phases: starter (7–21 days), grower (22–35 days), finisher (36–42 days), and overall (7–42 days). Average daily feed intake (ADFI) was calculated as total pen feed consumption divided by bird count and period length. Average daily gain (ADG) was determined as body weight gain divided by period length. Feed efficiency ratio (FER) was calculated as the ratio of body weight gain to feed intake for each period. Additionally, economic efficiency was evaluated on a per-bird basis using feed cost and marketable output. Bird viability (%) was computed as surviving birds divided by initial bird number, multiplied by 100. Productive index (PI) or European production efficiency factor was calculated according to the following formula described by Peric et al. [26]. In addition, feed cost per unit gain (FCG) was calculated as total feed cost divided by total body weight gain per bird. Salable bird return (SBR) was based on the market value of live birds at the end of the trial. Net profit per bird (NPR) was calculated as the difference between SBR and production cost per bird. Return on investment (ROI) was expressed as the ratio of net profit to total production cost multiplied by 100, following the calculated method described by Nopparatmaitree et al. [27].

2.5. Apparent Nutrient Digestibility

Apparent nutrient digestibility of broiler chickens was determined using the indicator method as described by Fenton & Fenton [28]. To achieve this, broilers were offered the experimental diets supplemented with 0.3% chromium oxide (Cr_2_O_3_) as an indigestible marker. Chromium oxide was thoroughly mixed into diets to ensure uniform distribution. Representative feed samples were collected prior to feeding, while excreta samples were collected from each pen during the last 5 days of the trial (days 38–42) to allow adequate adaptation of birds to the marker. Feed samples were stored in moisture-proof polyethylene bags, whereas excreta samples were collected into bags containing 3% sulfuric acid (H_2_SO_4_) to stabilize nitrogen and prevent microbial degradation. All samples were stored at −20 °C until laboratory analysis. Before analysis, feed and excreta samples were oven-dried at 60 °C to constant weight, ground through a 1 mm mesh to obtain homogenous powders, and then subjected to proximate analysis. Laboratory analyses included the determination of dry matter (DM), crude protein (CP), crude fiber (CF), ether extract (EE), organic matter (OM), and gross energy (GE) according to AOAC procedures [25]. Chromium oxide concentrations in feed and excreta were determined using UV–visible spectrophotometry after acid digestion to enable marker-based calculations of digestibility. Apparent nutrient digestibility was calculated using the marker concentrations in the excreta and the feed following the equations by Smeets et al. [29]. Meanwhile, apparent metabolizable energy (AME) was calculated using the equations AME (kcal/kg) = GE_diet_ − GE_excreta_ × (Cd/Cf), where Nf represents the nutrient concentration in the excreta (% DM); Nd represents the nutrient concentration in the diet (% DM); Cf represents the Cr_2_O_3_ concentration in the excreta (% DM); and Cd represents the Cr_2_O_3_ concentration in the diet (% DM) according to Zhang et al. [30].

2.6. Intestinal Histological Alterations

At 42 days of age, broiler chickens (one bird per pen) were randomly selected and humanely euthanized. Prior to neck cutting, birds were rendered unconscious using electrical stunning (50 V, 50 Hz, for 3 s) to ensure immediate loss of consciousness in accordance with animal welfare guidelines. During necropsy, the duodenum (from the gizzard outlet to the pancreatic loop), jejunum (from the pancreatic loop to Meckel’s diverticulum), and ileum (from Meckel’s diverticulum to the ileocecal junction) were excised. Approximately 2–3 cm of each intestinal segment was collected and fixed in 4% neutral-buffered formalin to preserve tissue morphology. Tissue processing included sequential dehydration through graded ethanol concentrations (70–100%), clearing in xylene, and paraffin infiltration. Samples were embedded in paraffin blocks with consistent orientation to maintain villus and crypt integrity. Sections of 5 µm thickness were cut using a rotary microtome (Leica Biosystems, Wetzlar, Germany) mounted on glass slides and dried overnight at 40 °C. Slides were stained with hematoxylin and eosin using a Leica Autostainer XL (Leica Biosystems) to allow clear visualization of villi and crypt structures. Histomorphometric analyses were conducted under a light microscope equipped with a digital imaging system. For each intestinal segment, 10 well-oriented villi per sample were randomly selected for measurement. The evaluated parameters included villus height (VH), measured from the villus apex to the villus–crypt junction; villus width (VW), measured at the widest point of the villus; and crypt depth (CD), measured from the villus–crypt junction to the base of the crypt. Villus surface area (VSA) was calculated using the formula VSA = 2π × (VW/2) × VH and the villus height-to-crypt depth ratio (VH:CD) was calculated. All intestinal sample preparations and measurements were carried out according to the method of Incharoen et al. [31].

2.7. Analysis of Volatile Fatty Acid in Cecal Contents

At 42 days of age, cecal contents were collected from broiler chickens to evaluate the concentration of volatile fatty acids (VFAs), which serve as key indicators of microbial fermentation and gut health. Cecal samples were first homogenized thoroughly to ensure uniform distribution of the microbial metabolites and then centrifuged to separate the supernatant. Approximately 1 mL of the supernatant was transferred into an ampulla and immediately mixed with 0.2% meta-phosphoric acid to precipitate proteins and stabilize the VFAs. The mixture was kept on ice for 30 min to further enhance stabilization and prevent degradation of the target compounds. The stabilized supernatant was subsequently centrifuged at 10,844× g for 10 min to remove residual particulates [32]. The clarified supernatant was collected for VFA quantification using gas chromatography (HP 5890 Series II GC; Agilent J & W 30 m × 0.535 mm × 1.00 µm HP-FFAP column) equipped with a flame ionization detector. For accurate quantification, samples were co-injected with 4-methylvaleric acid (Alfa Aesar, Heysham, Lancashire, UK) as an internal standard, following the method described by Ribeiro et al. [33]. Individual VFAs, including acetic, propionic, and butyric acids, were quantified by comparing the peak areas of the analytes to those of standard solutions prepared at known concentrations. Calibration curves were constructed using serial dilutions of the standard VFAs to ensure linearity and precision [34].

2.8. Cecal Microbiome Analysis

Cecal microbiota composition of broiler chickens was assessed using NGS of the bacterial 16S rRNA gene. Cecal contents were aseptically collected, transferred into sterile 1.5 mL microcentrifuge tubes, and stored at −20 °C until DNA extraction. Approximately 200 mg of cecal content per sample was used to isolate genomic DNA using the QIAamp DNA Stool Mini Kit (Qiagen, Valencia, CA, USA) following the manufacturer’s protocol with modifications to enhance the DNA yield and reduce potential inhibitors. The DNA concentration and purity were measured using a spectrophotometer (NanoDrop 2000, Thermo Scientific, Wilmington, DE, USA), and DNA integrity was verified by agarose gel electrophoresis. The DNA samples were subsequently diluted to 50 ng/µL and stored at −20 °C.

The diluted DNA served as a template for PCR amplification of the V3-V4 hypervariable region of the 16S rRNA gene, using the forward primer ACTCCTACGGGAGGCAGCA and reverse primer GGACTACHVGGGTWTCTAAT [35]. Sequencing was conducted on the Illumina MiSeq platform to generate 250 bp paired-end reads. Paired-end reads were processed by trimming barcode and primer sequences with FLASH software (V1.2.8; http://ccb.jhu.edu/software/FLASH/ (accessed on 27 May 2025)). Reads were merged when they exhibited sufficient overlap from opposite ends of the same DNA fragment, yielding raw tags. High-quality clean tags were obtained through quality filtering of raw tags based on specific criteria, following the QIIME (V1.9.1; http://qiime.org/scripts/split_libraries_fastq.html (accessed on 30 May 2025)) quality control pipeline. Tags were then aligned against the Gold database using the UCHIME algorithm (http://www.drive5.com/usearch/manual/uchime_algo.html (accessed on 1 June 2025)) to identify and remove chimeric sequences, resulting in effective tags (clean reads).

Nucleotide sequences were analyzed with Uparse software (version 7.0.1001; http://drive5.com/uparse/ (accessed on 5 June 2025)), clustering effective tags with ≥97% similarity into operational taxonomic units (OTUs). Representative sequences for each OTU were selected for annotation. Using Mothur software (versions 1.47.0) against the SILVA database SSUrRNA (http://www.arb-silva.de/ (accessed on 5 June 2025)), representative sequences were annotated at the phylum level. Phylogenetic relationships among OTU representative sequences were determined using MUSCLE software (Version 3.8.31) for rapid multiple sequence alignment. OTU abundance was normalized to the sample with the lowest sequence count.

2.9. Statistical Analysis

All experimental data were analyzed using one-way analysis of variance (ANOVA) to evaluate the effects of dietary treatments in CRD. Prior to ANOVA, assumptions of normality and homogeneity of variance were assessed using standard diagnostic tests. If these assumptions were not met, non-parametric analysis using the Kruskal–Wallis test was used. The general linear model used for ANOVA was Yij = μ + Ti +εij, where Yij = observed value for the jth replicate of the ith treatment (j = 1–6, i = 1–5), μ = overall mean, Ti = effect of the ith treatment, and εij = random error associated with each observation [36]. Significant differences among treatment means were determined using Tukey’s honestly significant difference test at a significance level of α = 0.05 (p < 0.05). Additionally, orthogonal contrasts were performed to specifically compare (i) control vs. all DMP treatments and (ii) AGP vs. all DMP treatments. To evaluate the response to increasing levels of phytosynbiotic inclusion, trend analysis was conducted using polynomial contrasts to determine the linear, quadratic, and cubic effects of the feed supplement on the measured variables. This allowed the identification of dose–response relationships and determination of optimal supplementation levels. All statistical analyses were conducted using R software version 4.3.3 with the ‘Agricolae’ package (version 1.3-7), as described by R Core Team [37].

3. Results

3.1. Production Performance and Economic Benefit Return Assessment

The dietary inclusion of the DMP improved the growth performance of broiler chickens (Table 2). Compared with the control group, broilers receiving the DMP had a higher (p < 0.001) ADG during both the starter (7–21 days) and grower (22–35 days) phases, with clear linear or quadratic dose–response trends. The FER was also markedly enhanced in these periods, reflecting better nutrient utilization, while the ADFI remained unaffected.

Across the overall production period (7–42 days), compared with birds fed the CON diet, broilers receiving AGP or diets supplemented with the DMP had a greater ADG and FER. The ADG increased linearly in response to increasing levels of DMP inclusion (p < 0.001), whereas the FER followed a quadratic response (p < 0.001). Dietary treatments did not affect the viability rate, which remained consistently high across all experimental groups (98–100%). A high value of PI was observed in broilers receiving the AGP diet and all levels of DMP supplementation (p < 0.001).

Economic analysis indicated that the FCG was not influenced by the dietary treatments (p > 0.05). However, compared with those fed the CON diet, the SBR, NPR, and ROI were greater in broilers fed the AGP and phytosynbiotic-supplemented diets (p < 0.05). Both linear and quadratic trends were observed for the SBR, NPR, and ROI with increasing DMP levels (p < 0.01), indicating improved economic efficiency associated with DMP supplementation.

3.2. Nutrient Intake and Digestibility

Compared with the CON and AGP groups, dietary supplementation with the DMP did not affect daily nutrient intake, including DM, OM, CP, and EE (p > 0.05; Table 3). In contrast, apparent nutrient digestibility was improved by DMP supplementation. Compared with those fed the CON diet, broilers fed the DMP had a higher apparent digestibility of DM, OM, CP, and AME (p < 0.05). Apparent digestibility of DM increased significantly at 0.25% supplementation (p < 0.001), while OM digestibility and AME had significant quadratic responses to increasing DMP levels (p < 0.01). CP digestibility was also enhanced, with the highest value observed in broilers receiving 1.00% of the DMP (p < 0.05).

Regarding digestible nutrient intake, compared with the CON treatments, digestible CP intake was greater in broilers fed the DMP and AGP diets (p < 0.01), whereas digestible intakes of DM, OM, and EE were not affected by dietary treatments (p > 0.05). The greater digestible CP intake was attributable to improved CP digestibility rather than differences in CP intake, indicating the enhanced efficiency of nutrient utilization in DMP-supplemented broilers.

3.3. Morphological Change in the Small Intestine

Dietary supplementation with the DMP exerted significant modulatory effects on small intestinal morphology (Table 4). In the duodenum, VH was greater in response to DMP inclusion, with the 0.50% DMP group exhibiting the greatest VH, comparable to or exceeding that observed in the AGP group (p = 0.001). In the duodenum, VH was enhanced by DMP supplementation, with the 0.50% DMP group leading to the highest VH, which was comparable to the AGP group (p = 0.001). Orthogonal contrasts indicated significant differences between the control and DMP-supplemented groups (p = 0.003), along with a pronounced dose-dependent response (p < 0.001). Although VW was not affected (p > 0.05), VSA was enhanced by DMP supplementation (p = 0.036), displaying both linear and contrast effects relative to the CON and AGP groups. The CD increased with rising DMP levels (p < 0.001), exhibiting a quadratic response, while the VH:CD ratio remained unaffected. In the jejunum, DMP supplementation did not alter VH and VW (p > 0.05). However, compared with the control group, VSA was markedly increased in DMP-fed broilers (p = 0.019), particularly at the 0.25% and 0.50% inclusion levels. Orthogonal contrasts confirmed a significant improvement in VSA in the DMP groups relative to both the CON and AGP groups (p = 0.017). The CD increased at the 0.25% DMP level (p = 0.003), following a quadratic pattern, whereas the VH:CD ratio had only a numerical trend without statistical significance. In the ileum, DMP supplementation did not influence VH, VW, or VSA (p > 0.05). Despite this, CD had a quadratic response to increasing DMP inclusion levels (p = 0.004). Notably, the VH:CD ratio differed among treatments (p = 0.006), with the 1.00% DMP group resulting in the highest ratio, indicating improved epithelial maturation and reduced crypt hyperplasia at higher supplementation levels.

3.4. Volatile Fatty Acid Analysis in Cecal Contents

Supplementation of the DMP had a pronounced influence on the lactic acid and VFA concentrations in the cecum (Table 5). Broilers receiving the DMP had greater lactic acid concentrations compared with the CON group (p < 0.01), with the values increasing quadratically as the inclusion levels rose from 0.25 to 1.00%. Total VFAs followed a similar pattern, wherein the highest concentration was observed at 1.00% DMP supplementation (p < 0.01). Orthogonal contrasts revealed that both the CON vs. DMP and AGP vs. DMP comparisons were significant (p < 0.01), confirming the strong fermentative response driven by the supplemented product. Among individual VFA components, acetic acid (C2) increased markedly with the phytosynbiotic (p < 0.001), exhibiting a clear quadratic trend. Propionic acid (C3) was not affected by the treatments (p > 0.05). Concentrations of n-butyric acid and total butyric acid (C4) were significantly elevated in the phytosynbiotic groups (p < 0.01), with the highest responses noted at 0.25% and 1.00%, respectively. Iso-butyric acid and iso-valeric acid were not affected (p > 0.05). In contrast, valeric acid (C5) and n-valeric acid increased significantly with supplementation, with n-valeric acid having a linear trend (p < 0.01). Overall, as reflected by greater lactic acid and VFA production, the phytosynbiotic product enhanced cecal fermentation and performed comparably or better relative to the AGP treatment across most parameters.

3.5. Microbiome Analysis

Microbiome profiling using NGS demonstrated that the DMP modulated the cecal bacterial community structure (Figure 2). At the order level, the DMP produced clear shifts in microbial composition. CON birds exhibited greater proportions of Bacteroidales and Veillonellales–Selenomonadales, representing the typical microbiota of unsupplemented broilers. Supplementation with 0.25% and 0.50% DMP increased the fermentative bacterial orders, particularly Lachnospirales and Oscillospirales, both of which are key contributors to SCFA production and gut homeostasis. At higher DMP inclusion levels (0.50–1.00%), Lactobacillales became increasingly dominant, accompanied by a reduction in Bacteroidales, indicating a shift toward a probiotic-enriched and metabolically favorable microbial ecosystem. This microbial pattern reflects the selective enrichment of lactic acid-producing bacteria commonly associated with enhanced intestinal health. Minor orders, including Desulfovibrionales, Campylobacterales, and Synergistales, remained consistently low across treatments, confirming that the DMP did not promote the expansion of potential pathogens. Collectively, these results show that DMP supplementation effectively reshaped the cecal microbiota by increasing beneficial SCFA-producing taxa while limiting less desirable bacterial groups, thereby supporting improved gut function and overall intestinal health in broiler chickens.

4. Discussion

This study demonstrates that dietary supplementation with the DMP, combining P. acidilactici V202, wheat bran-derived prebiotics, and Tiliacora triandra phytogenics, enhances broiler health and performance. This effect is mediated through coordinated modulation of gut microbiota composition, microbial metabolism, intestinal morphology, and nutrient utilization. These findings align with previous research demonstrating that synergistic combinations of probiotics, prebiotics, and phytogenics more effectively enhance the microbial community structure, metabolic function, probiotic viability, nutrient efficiency, and immune modulation than single components, ultimately improving growth and feed efficiency in livestock [38,39,40].

Dietary supplementation with the DMP markedly modulated the cecal microbial composition and fermentative activity in broilers. At 0.25% and 0.50%, the DMP increased the fermentative bacterial orders Lachnospirales and Oscillospirales, both of which play important roles in the cecal ecology of poultry, contributing significantly to gut health and microbiota maturation. Lachnospirales, particularly members of the family Lachnospiraceae, are abundant in early dynamic cecal microbiota and are involved in fermentative metabolism that produces SCFAs. These SCFAs, including butyrate, are crucial for maintaining intestinal epithelial integrity, modulating immune responses, and providing energy to colonocytes, thereby supporting gut health and barrier function in poultry [41,42]. By balancing this group with others to reach an adult-like microbiome earlier in development, probiotic treatments can modulate the abundance of Lachnospiraceae, accelerating the maturation of cecal microbiota [42]. Oscillospirales, primarily from the family Ruminococcaceae, are also key butyrate producers and are associated with microbial functions related to fiber degradation and energy metabolism in the cecum. Their presence correlates with increased SCFA concentrations and overall metabolic activity, highlighting their role in sustaining microbial homeostasis and nutrient utilization in poultry [43]. Both Lachnospirales and Oscillospirales contribute to maintaining a balanced anaerobic environment in the ceca, inhibit pathogenic bacterial colonization through competitive exclusion, and promote a stable microbial ecosystem vital for optimal poultry health and performance [41,42].

The DMP formulation contains bioactive phytochemicals from Tiliacora triandra, which synergistically suppress pathogenic bacteria. Extracts from their leaves and stem bark exhibit broad antimicrobial activity against poultry and human pathogens, an effect attributable to polyphenols, flavonoids, alkaloids, and mucilage polysaccharides [44,45]. Through biochemical mechanisms involving the suppression of lipid peroxidation, attenuation of pro-inflammatory cytokine production, and restoration of endogenous antioxidant enzyme activities, its antioxidant phytochemicals mitigate neurobehavioral deficits, alongside the normalization of AChE function [46]. In poultry nutrition, Tiliacora triandra compounds act as effective phytobiotics that mitigate enteric pathogens such as E. coli and Salmonella, reduce the dependence on antibiotic growth promoters, and support probiotic viability with prebiotic-like gut effects [18]. Synergistic probiotic–phytobiotic interactions have been demonstrated to reduce ESBL-producing E. coli survival in broiler gut contents, promoting the favorable modulation of microbiota composition and metabolic activity in young broilers [47]. This ecological restructuring enhances microbial fermentation, sustains SCFA production, and contributes to long-term intestinal stability and health.

At higher doses of the DMP (0.50–1.00%), there was a progressive dominance of Lactobacillales and a reduction in Bacteroidales, signifying a shift toward a probiotic-enriched, metabolically favorable cecal ecosystem. This restructuring enriched health-associated taxa including Lactobacillus, Pediococcus, and Enterococcus alongside saccharolytic SCFA producers [48,49]. These microflora play essential roles in carbohydrate fermentation, SCFA productions, and intestinal metabolic homeostasis [50]. The microbial shifts likely stem from the DMP’s modulation of the gastrointestinal environment favoring beneficial microbes. Notably, P. acidilactici V202 rapidly produces lactic acid and bacteriocins, acidifying the lumen (pH ~5.5–6.5) to suppress pH-sensitive pathogens while promoting acid-tolerant, healthful bacteria [51,52]. Increased lactic acid-producing Lactobacillales contribute to pathogen exclusion via nutrient competition, bacteriocin-mediated inhibition, and maintaining low intestinal pH [53].

Dietary DMP supplementation significantly enhanced cecal fermentation in broilers, increasing lactic acid and total VFAs in a dose-dependent manner, with peak effects at 1.00% inclusion (p < 0.01). Notably, the acetic, butyric, and valeric acid levels rose, reflecting enhanced saccharolytic microbial activity, while propionic and branched-chain VFAs remained unchanged. These shifts stem from the DMP’s modulation of the gut microbiota, particularly fermentative activity by P. acidilactici V202, which elevates lactic acid and lowers intestinal pH, suppressing pathogens and favoring beneficial microbes [54,55]. This acidification supports a bifidogenic effect, promoting Bifidobacterium spp. critical for cecal microbial succession and fermentation efficiency [56]. Wheat bran-derived non-starch polysaccharides (NSPs), chiefly arabinoxylans and β-glucans, escape upper gut digestion to serve as fermentable substrates for saccharolytic bacteria such as Lactobacillales, Lachnospirales, Oscillospirales, and Bifidobacterium spp., which enzymatically degrade NSPs into fermentable sugars [57,58,59,60,61]. These sugars enter anaerobic glycolysis yielding VFAs, predominantly acetate, via the pyruvate–acetyl-CoA pathway, which regulates microbial interactions [62,63,64]. Lactate and acetate produced by LAB and bifidobacteria are substrates for butyrate-producing bacteria within Lachnospirales and Oscillospirales, enhancing butyrate pools via cross-feeding syntrophy [65,66]. Correspondingly, the DMP reshaped cecal microbiota by enriching saccharolytic SCFA producers while reducing Bacteroidales linked to proteolytic fermentation under carbohydrate scarcity, paralleling microbial shifts observed with probiotic and phytogenic supplements that improve fermentative efficiency and gut health.

The DMP’s multifaceted benefits extend to enhancing intestinal morphology and nutrient digestibility through integrated mechanisms involving microbial shifts, increased SCFAs, improved epithelial energy metabolism, and optimized nitrogen utilization. Elevated lactic acid levels in the DMP group likely stem from lactic acid bacteria, which play a pivotal role in modulating the gut microenvironment through carbohydrate fermentation. This process generates lactic acid that establishes and sustains an acidic milieu along intestinal villi, particularly adjacent to absorptive epithelial cells. Such localized pH gradients are essential for optimizing digestive and absorptive functions in the gut [67]. In addition, the increased butyric acid in the DMP groups plays a critical role in supporting mitochondrial ATP production, which may be essential for the development of intestinal villi and their absorptive functions. Butyrate serves as the primary energy substrate for colonocytes and intestinal epithelial cells, fueling mitochondrial oxidative phosphorylation to generate ATP vital for cellular maintenance and proliferation [68]. In the intestinal epithelium, efficient ATP production through butyrate oxidation underpins the maintenance of the complex structure and function of villi, which increase the surface area for nutrient absorption. This energy supports the epithelial barrier integrity, renewal, and absorptive capacity necessary for optimal digestive health [69].

Butyrate also modulates mitochondrial dynamics and counteracts dysfunction induced by inflammatory cytokines, enhancing epithelial cell bioenergetics and contributing to tissue homeostasis [70]. By promoting mitochondrial function and ATP generation, butyrate sustains the structural development of villi and their critical role in nutrient uptake, ultimately improving gut health, growth performance, and disease resistance in the host. Furthermore, by favoring the growth of beneficial acetate-producing Bifidobacteria, the high acetic acid levels across experimental treatments, except the CON group, likely promote intestinal microbial balance [71]. Acetic acid also has systemic impacts through the modulation of the gut–brain and gut–liver axes by influencing the microbial metabolism and immune responses. By modulating both the intestinal and central nervous system pathways, acetic acid can reduce intestinal fibrosis and has been implicated in the attenuation of neuroinflammation [72,73]. Increased concentrations of valeric acid were detected in the supplemented groups, which may positively influence gut health by modulating microbial composition and strengthening intestinal barrier function. In Helicobacter pylori-infected mice, Weizmannia coagulans BC99 supplementation elevated valeric acid, correlating with reduced gastric inflammation and oxidative stress [74]. These data suggest a role for valeric acid in promoting microbial homeostasis and mitigating inflammatory and oxidative damage in the gastrointestinal tract. Together, our findings highlight the essential roles of these VFAs in shaping a balanced gut ecology and maintaining intestinal structural and functional integrity.

An important outcome in the present study was that the DMP improves nutrient utilization without altering feed intake, increasing the apparent digestibility of DM, OM, CP, and AME, thereby elevating digestible protein intake independent of total protein consumption, suggesting enhanced protein utilization efficiency [75]. However, the direct factor of these changes to the animal remains uncertain and may involve indirect effects through the modulation of microbial communities or nutrient absorption. Notably, both the DMP and AGP treatments elicited comparable improvements in nutrient digestibility and small intestinal morphology relative to the controls, despite these parameters being indirectly linked to antimicrobial action. These convergent effects likely stem from shared non-antimicrobial mechanisms, including enhanced cecal fermentation and selective microbiota enrichment, which promote epithelial proliferation and nutrient transporter expression independently of pathogen reduction. While AGP may exert indirect modulation via altered microbial ecology, the DMP’s targeted synbiotic delivery achieves equivalent structural and functional outcomes through co-encapsulated prebiotic substrates and stable probiotic colonization, underscoring phytosynbiotics’ capacity to recapitulate multifaceted AGP benefits sans resistance risks.

Collectively, our developed phytosynbiotic demonstrates substantial feasibility as an AGP alternative in commercial broiler production. DMP supplementation at practical levels yielded superior or comparable growth performance, nutrient utilization, gut health, and economic returns relative to AGP supplementation, without increasing production costs. Its scalability is underpinned by cost-effective, locally sourced materials suitable for large-scale industrial processing while maintaining probiotic viability under harsh GI conditions. This positions the DMP as a sustainable, regulatory-compliant solution aligned with global AGP reduction initiatives, effectively addressing antimicrobial resistance concerns while preserving productivity and profitability in antibiotic-free poultry systems.

5. Conclusions

The DMP can be incorporated into commercial broiler diets to enhance growth performance, nutrient utilization, and gut health without increasing feed intake. It can do this by improving the ADG, feed efficiency, and economic returns, concomitant with improved nutrient digestibility and coordinated modulation of the gut ecology. These responses were characterized by selective enrichment of fermentative and SCFA-producing microbiota along with increased concentrations of lactic acid, acetic acid, n-butyric acid, butyric acid, and valeric acid. As a result of these responses, the intestinal morphology was enhanced and contributed to improved growth performance. Taken together, these results indicate that the DMP can be effectively incorporated at practical inclusion levels (0.5–1.0%) as a viable alternative to AGP. From an industrial perspective, the DMP represents a scalable and sustainable nutritional strategy aligned with global antibiotic reduction policies and sustainable development goals, particularly responsible production and animal health-oriented poultry systems.

6. Patents

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 phytosynbiotic products from Tiliacora triandra leaf extract and P. 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 Naresuan University.