In vitro simulated digestion and fermentation of Lithocarpus litseifolius [Hance] Chun green tea polysaccharides and their modulation of the gut microbiota

Abstract

Lithocarpus litseifolius [Hance] Chun (L. litseifolius), a herbal tea rich in bioactive polysaccharides, has garnered attention for its health-promoting potential. Polysaccharides represent its principal bioactive component and play a significant role in regulating the gut microbiota. This study investigated the physicochemical characteristics and in vitro digestion bioactivities of special-grade polysaccharides (STPs) and first-grade polysaccharides (FTPs) from L. litseifolius green tea, alongside their modulatory effects on the gut microbiota following in vitro fermentation. The results demonstrated that STPs exhibited higher total carbohydrate and uronic acid content, higher molecular weight, and greater in vitro bioactivity compared to FTPs, despite structural similarities. During in vitro simulated digestion, polysaccharides underwent moderate physicochemical modifications accompanied by partial loss of bioactivity. Notably, STPs exhibited a greater extent of degradation compared to FTPs. Despite differential digestion-induced degradation, most STPs and FTPs remained largely intact upon reaching the colon and were thus accessible to the gut microbiota for fermentation. Fecal fermentation demonstrated efficient microbial utilization of STPs and FTPs, each modulating the architecture of the human gut microbiota, characterized by carbohydrate consumption, a decrease in pH, and an elevated relative abundance of beneficial bacterial phyla, including Firmicutes and Bacteroidota. Concurrently, a marked increase in short-chain fatty acid (SCFA) output—particularly acetate and propionate—was observed. STPs primarily enriched Actinobacteria, whereas FTPs favored Bacteroidota, both contributing to elevated acetate and propionate levels while suppressing potential pathogens such as Proteobacteria and Escherichia–Shigella. These findings underscore the potential of L. litseifolius green tea polysaccharides to serve as effective prebiotics for gut microbiota modulation.

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1. Introduction

Lithocarpus litseifolius [Hance] Chun (L. litseifolius), colloquially termed sweet tea, is a pharmaco-nutritional plant endemic to southern China.¹ It was designated as a novel food resource by Chinese regulatory authorities in 2017.² This species has gained scientific interest due to its abundance of bioactive constituents, including polysaccharides, flavonoids, triterpenoids, phenolic compounds, essential micronutrients, and amino acids.³ Polysaccharides represent a class of essential biomacromolecules composed of numerous monosaccharide units interconnected through glycosidic linkages. Characterized by their favorable safety profiles and minimal adverse effects, these polymeric carbohydrates have attracted substantial scientific attention over recent decades. Polysaccharides have emerged as pharmaceutically significant biopolymers, demonstrating multifaceted therapeutic potential through reactive oxygen species (ROS) neutralization, glycemic homeostasis modulation, antineoplastic activity, and immunoregulatory functions, coupled with their capacity to beneficially modulate intestinal microbial ecosystems.^4,5 Their unique physicochemical properties and biological compatibility have facilitated extensive investigation and practical utilization across multiple industrial domains, particularly in biochemical engineering, pharmaceutical development, and food technology applications.^6,7

The biological efficacy of polysaccharides depends on their absorption in the gastrointestinal system before engaging in metabolic processes. Recent advances in gastrointestinal simulation research have highlighted the transformation of polysaccharides in in vitro simulated digestion and colonic fermentation models, a topic of increasing significance in nutritional and pharmacological investigations.⁸ Digestion constitutes a fundamental biological process governing the catabolic transformation of macromolecular nutrients into assimilable metabolites.⁹ Modern in vitro digestion systems simulate human gastrointestinal conditions by precisely controlling enzyme activity, ionic concentrations, and pH levels.¹⁰ Besides, owing to its operational simplicity, rapid processing, and experimental reproducibility, in vitro simulated digestion has emerged as a robust experimental paradigm, serving as a viable alternative to in vivo digestive models in contemporary biomedical research.¹¹ Research has indicated that changes in pH levels and digestive enzymes within the gastrointestinal tract can alter the properties of polysaccharides, including the monosaccharide composition, chemical structure, and molecular weight. Studies have shown that Hordeum vulgare L. polysaccharides exhibit reduced chemical composition content (total carbohydrate, protein, and uronic acid) and altered molecular weight (M_w) following simulated in vitro digestion.¹² Similar alterations have been observed in Artemisia sphaerocephala Krasch polysaccharide during gastrointestinal digestion.¹³ However, although polysaccharides undergo partial degradation mediated by digestive enzymes during gastrointestinal digestion, a substantial proportion can still reach the colon. The colon harbors a dense microbial community, certain members of which produce specialized enzymes to catabolize and utilize these polysaccharides.¹⁴ Growing evidence suggests that polysaccharides can function as a prebiotic, modulating the gut microbiota by reducing pathogens, producing SCFAs, and increasing beneficial bacteria.¹⁵ Furthermore, polysaccharides play a crucial role in shaping the gut microbial community and promoting species diversity. Specifically, they tend to increase the richness of beneficial microbial species while simultaneously decreasing the abundance of pathogenic microbes. In prior studies, fresh tea leaf polysaccharides elevated the levels of Bifidobacterium, Bacteroides, and Lactobacillus while suppressing the populations of Escherichia, Shigella, and Enterococcus.¹⁶ Polysaccharides derived from dark tea enhance the proliferation of beneficial microorganisms, including Bifidobacterium, Prevotella, and Saccharimonadiales, consequently restructuring the gut microbial community and promoting SCFA biosynthesis.¹⁷L. litseifolius green tea polysaccharides have a positive impact on human health. However, their digestive and absorptive processes, along with their influence on the human intestinal flora and SCFA generation, are not yet fully comprehended. Therefore, systematic investigation of the gastrointestinal processing dynamics and colonic fermentation characteristics of L. litseifolius green tea polysaccharides represents an essential research priority.

Tea quality is governed by leaf grade, as the grade dictates the quality of the raw material. Consequently, leaf maturity becomes a critical factor, influencing final quality by regulating the spectrum and concentration of metabolites in the fresh leaves.^18,19 Specifically, for L. litseifolius green tea, this leads to a classification into special and first grades based on maturity and morphology.²⁰ The special grade originates from tender, reddish buds and leaves (length <10 cm and width <5 cm), and the first grade from mature, largely green leaves (length: 10–15 cm and width: 5–10 cm) with limited red pigmentation. These maturity-driven differences are key determinants of the tea's ultimate flavor characteristics, biochemical makeup, and bioactivity. Notably, biochemicals such as polysaccharides exhibit dynamic patterns during growth, creating distinct compositional signatures across the different grades. Current scientific evidence confirms that the polysaccharide molecular architecture and pharmacological efficacy exhibit significant dependency on multiple determinants, including source material characteristics (botanical origin and developmental stage), coupled with extraction protocols.²¹ In recent years, most current research on polysaccharides from L. litseifolius has primarily concentrated on chemical compositions and extraction techniques, while studies on the polysaccharides derived from L. litseifolius green tea are almost non-existent.⁴ However, there is currently an absence of studies on both the structural and chemical compositions of polysaccharides in different grades of L. litseifolius green tea, and the potential compositional and structural variations in these polysaccharides have not yet been rigorously characterized. Therefore, this study investigates the chemical structures, physicochemical properties, and bioactivities of special-grade L. litseifolius green tea polysaccharides (STPs) and first-grade L. litseifolius green tea polysaccharides (FTPs) during in vitro digestion and human fecal fermentation processes, while systematically analyzing their interactions with the intestinal microbiome and underlying mechanisms. This study proposes to elucidate the potential prebiotic effects of STPs and FTPs on human health through a comprehensive analysis of structure–activity relationships, thereby laying a scientific foundation for their functional exploitation and application.

2. Materials and methods

2.1. Materials and reagents

The first-grade green tea and the special-grade green tea of L. litseifolius were bought from Tree Wormwood Tea Farmers’ Specialized Cooperative (Lushan County, Ya'an City, Sichuan Province). Standard SCFAs (GC >98%) were obtained from Aladdin Biochemistry Science and Technology Co., Ltd (Shanghai, China). Bile salt, α-amylase (14 U mg⁻¹), pepsin (1 : 3000), pancreatic enzyme (8X USP), inulin (INL), and monosaccharide standards (glucose (Glu), xylose (Xyl), rhamnose (Rha), galactose (Gal), mannose (Man), and arabinose (Ara)) were purchased from Yuanye Biotechnology Co., Ltd (Shanghai, China). The other chemicals were procured from Sinopharm Chemical Reagent Co., Ltd (China).

2.2. Preparation of polysaccharides from L. litseifolius green tea

STPs and FTPs were respectively isolated from the special-grade and the first-grade L. litseifolius green tea through a standardized extraction protocol. Both tea grades were processed identically: sequential mechanical grinding followed by particle size fractionation using a 60-mesh sieve, generating precisely categorized special-grade and first-grade tea powders. The extraction of STPs and FTPs was carried out following the method previously established by our research team;^17,22 the extraction procedure commenced with 12-hour defatting in 95% ethanol, followed by centrifugation (5000g, 15 min) to collect the pelleted material. The ethanol-insoluble fractions underwent aqueous extraction (1 : 40 w/v) at 95 °C for 2 h, with subsequent centrifugation to recover soluble fractions. Concentrated supernatants (40 °C, rotary evaporation) underwent further analysis. Proteins were removed using Sevag reagent (chloroform : n-butanol, 4 : 1 v/v) through repeated cycles (8–10 times) to ensure complete removal. The sample was dialyzed using a 3000 Da molecular weight cut-off (MWCO) dialysis membrane. Final polysaccharide isolation was achieved through 95% ethanol precipitation (1 : 4 v/v, 48 h), yielding STPs and FTPs from respective tea grades after centrifugation. The yield of STPs from L. litseifolius green tea of special grade was 7.43%, while that of FTPs from first-grade tea was 6.77%.

2.3. Characterization of the physicochemical properties of STPs and FTPs

2.3.1. Chemical composition analysis. The constituent profile was quantified according to the established analytical protocol.²³ Total carbohydrate content was determined via the phenol–sulfuric acid assay, while reducing sugar levels were assessed through the DNS colorimetric method. Glucuronic acid levels were determined via the meta-hydroxyphenyl assay. Simultaneously, protein content analysis was conducted using the Coomassie Brilliant Blue colorimetric assay.

2.3.2. M _w analysis. The analysis was performed using high-performance size exclusion chromatography (HPSEC) coupled with multi-angle laser light scattering and refractive index detection (HPSEC-MALLS-RID, Wyatt Technology, Santa Barbara, CA, USA). The chromatographic system was operated under optimized parameters: the column temperature was maintained at 35 °C, the refractive index increment (dn/dc) value was 0.318 mL g⁻¹, the sample injection volume was 100 μL, and the mobile phase flow rate was 0.5 mL min⁻¹. Samples were prepared in a mobile phase consisting of 0.02% (w/v) NaN₃ containing 0.9% (w/v) NaCl, homogenized by vortex mixing, and filtered through 0.22 μm hydrophilic membranes.

2.3.3. Fourier transform infrared spectroscopy (FT-IR) analysis. FT-IR spectroscopy was conducted using a Nicolet iS50 spectrometer (Thermo Fisher Scientific) under ambient conditions (25 °C). Lyophilized polysaccharide samples (STPs/FTPs) were homogeneously blended with spectroscopic-grade potassium bromide (KBr) and pelletized under hydraulic pressure. Spectral acquisition was conducted across 4000–400 cm⁻¹ with a 4 cm⁻¹ resolution, accumulating 32 scans per measurement to optimize the signal-to-noise ratio.

2.3.4. Study of monosaccharide composition. Monosaccharide composition analysis was conducted following the prior method with some slight adjustments. The samples underwent acid hydrolysis in 2 M trifluoroacetic acid (TFA) at 120 °C for 2 h under a nitrogen atmosphere. The hydrolysates were sequentially treated with anhydrous methanol through three cycles of rotary evaporation to ensure complete TFA removal, followed by lyophilization. Derivatization was performed via the alditol acetate protocol: neutralized hydrolysates were reduced with sodium borohydride, acetylated with acetic anhydride-pyridine (1 : 1 v/v), and extracted with dichloromethane. Gas chromatography was used to analyze the monosaccharide composition (Agilent 7890B GC system). Quantitative determination utilized external calibration with authenticated monosaccharide standards, mannose (Man), rhamnose (Rha), xylose (Xyl), arabinose (Ara), glucose (Glc), and galactose (Gal).

2.4. Evaluation of biological activities

The biological activities comprise antioxidant and hypoglycaemic functions in vitro. Antioxidant activities and hypoglycaemic properties were measured in our previous study.²³ The antioxidant activity encompasses the DPPH radical, metal chelating capacities, scavenging, and reducing power. Hypoglycaemic activity involves inhibiting α-glucosidase and α-amylase actions.

2.5. In vitro simulated digestion of STPs and FTPs

In vitro gastrointestinal digestion was simulated following the INFOGEST model.¹⁰ We prepared reserve electrolyte fluids containing simulated saliva fluid (SSF), simulated gastric fluid (SGF), and simulated intestinal fluid (SIF). The digestion steps are as follows.

2.5.1. Saliva digestion process. SSF was heated to 37 °C. Then, 10 mg mL⁻¹ of STPs or FTPs, salivary amylase at 3700 U mg⁻¹, and calcium chloride dihydrate solution were introduced into 5 mL of SSF solution. Ultrapure water was added to the SSF solution in a 1 : 1 proportion to ensure that the concentration of calcium chloride dihydrate reached 1.5 mM and the enzyme activity of salivary amylase was 75 U mL⁻¹ (pH = 7). The oral simulants were put into an oscillating incubator and cultivated at 37 °C for 2 minutes. Once digestion was finished, the activity of salivary amylase was inactivated via high-temperature treatment.

2.5.2. Gastric digestion. Following oral processing, the chyme was combined with preheated simulated gastric fluid (SGF, 37.0 °C) at a 4 : 5 (v/v) ratio. Sequential introduction of porcine pepsin (≥250 U mg⁻¹) and calcium chloride dihydrate (CaCl₂·2H₂O) preceded system normalization through ultrapure water addition (final SGF : H₂O = 1 : 1). The reaction system was standardized to 2000 U mL⁻¹ pepsin activity and 1.5 mM Ca²⁺ concentration, with the pH maintained at 3 via 0.1 M HCl titration. Gastric digestion proceeded under physiomimetic conditions for 120 min, followed by enzyme deactivation through high-temperature treatment.

2.5.3. Intestinal digestion. The SIF was mixed with the gastrically digested polysaccharide solution mixture in a volume ratio of 4 : 5 at 37 °C, following the simulated gastric phase. Bile salts (10 mM), pancreatic enzymes (8× USP), and CaCl₂(H₂O)₂ were then added. Ultrapure water was incorporated to achieve a final SIF (the ratio was 1 : 1). The pancreatic enzyme activity was adjusted to 100 U mL⁻¹, and the CaCl₂(H₂O)₂ concentration was set to 10 mM, with the pH adjusted to 7. The gastric simulated digest was then maintained in an oscillating incubator at 37 °C for 2 h. After the digestion process, the enzyme activity was inactivated through high-temperature treatment.

Following the digestion cascade, the post-digestion mixture was subjected to centrifugation to partition the supernatant. This bioactive fraction was obtained through dialysis (500 Da, 24 h against ultrapure water) followed by lyophilization to obtain stable polysaccharide fractions.

2.6. In vitro fermentation of STPs and FTPs

In vitro fermentation experiments were based on adaptations of methods established by previous researchers.²⁴ The fecal material was collected from six human volunteers (3 females and 3 males, aged between 20 and 26 years). To ensure that the experiment was conducted accurately, they had abstained from antibiotics or probiotics for three months and experienced no gastrointestinal discomfort in the two weeks before the experiment. The fecal suspension (32%, w/v) was made by mixing feces and diluting with phosphate buffer solution and subsequently centrifuged at low speed. The basal medium was formulated with (per liter): yeast extract (2.0 g), peptone (2.0 g), NaCl (0.1 g), KH₂PO₄ (0.04 g), K₂HPO₄ (0.04 g), MgSO₄·7H₂O (0.01 g), CaCl₂ (0.01 g), NaHCO₃ (2 g), hemin (0.02 g), cysteine-HCl (0.5 g), bile salts (0.5 g), Tween 80 (2 mL), and 1% resazurin solution (1 mL), constituting the standardized fermentation matrix. (All volunteers provided stool samples voluntarily, having understood and signed the written informed consent form. The collection of stool samples was approved by the Academic Ethics and Welfare Committee of Sichuan Agricultural University and was conducted in accordance with the approved protocols.)

The experiment was subsequently divided into five groups: original group (OR), basal medium as a blank group (BLK), inulin as a positive control (INL), and STPs and FTPs as experimental groups without a carbon source. Samples from fermentation at 0, 6, 12, and 24 h were collected for further study.

2.7. Measurement of pH and SCFAs during fermentation

The pH was measured at 0, 6, 12, and 24 h of fermentation using a pH meter (FE28, Mettler Toledo). The SCFAs in the fermentation samples at these time points were analyzed by gas chromatography (GC) using Agilent Technologies equipment (Palo Alto, CA, USA).

2.8. Determination of the gut microbiota after in vitro fermentation

After 24 h of fermentation, the genomic DNA of OR, BLK, INL, STP, and FTP groups was extracted using the E.Z.N.A.® Soil DNA Kit. Subsequently, the 16S rRNA gene sequencing was performed by Majorbio Biotechnology Co., Ltd (Shanghai, China). Analyses of species taxonomy, community diversity, composition, and differences were based on Amplicon Sequence Variants (ASV).

2.9. Statistical analysis

All data are expressed as mean ± SD. Graphs were made using OriginPro 2021 (OriginLab Corporation) and GraphPad Prism 8.0.1. Statistical significance between groups (p < 0.05) was determined using SPSS 16.0 and Duncan's multiple range test.

3. Results and discussion

3.1. Changes in the physicochemical properties of STPs and FTPs during digestion

3.1.1. Changes in chemical compositions. As shown in Table 1, STPs had total carbohydrate, protein, and uronic acid contents of 74.166%, 8.204%, and 3.317%, respectively, while FTPs had contents of 60.173%, 8.009%, and 2.825%, respectively. STPs had higher total carbohydrate and uronic acid contents than FTPs, with similar protein contents (STPs: 8.204% and FTPs: 8.009%). The high uronic acid content in both STPs and FTPs suggests their potential classification as acidic polysaccharides.²⁵ Overall, the results demonstrate that STPs exhibit greater chemical diversity than FTPs. Nevertheless, the precise structural basis for this difference requires further in-depth investigation. Additionally, the chemical components of STPs and FTPs changed before and after digestion. Following oral digestion, the total carbohydrate and uronic acid content slightly declined. After gastrointestinal digestion, a reduction was observed in total carbohydrate and uronic acid levels, likely attributable to the acidic pH and intestinal enzymes within the gastrointestinal environment influencing the degradation of uronic acid.²⁶ Notably, the protein content of STPs and FTPs increased during all three digestion stages, likely because the glycoproteins within the polysaccharides were not easily digested and formed larger protein complexes with digestive enzymes, which led to the increase in protein content.²⁷

Table 1 The chemical compositions of STPs and FTPs during in vitro stimulated digestion

Samples	Stage	Total carbohydrate (%, w/w)	Protein (%, w/w)	Uronic acid (%, w/w)	Reducing sugars (mg mL^−1)
Each value is expressed as the mean ± SD (n = 3) of triplicate determinations. Different lowercase letters (a–d) indicate statistically significant differences (p < 0.05) for the same polysaccharide across different digestion time points. Undigested, STPs-S, STPs-G and STPs-I in STPs represent undigested STPs and saliva-digested, gastric-digested, and intestinal-digested STPs, respectively. Undigested, FTPs-S, FTPs-G and FTPs-I in FTPs represent undigested FTPs and saliva-digested, gastric-digested, and intestinal-digested FTPs, respectively.
STPs	Undigested	74.166 ± 0.250^a	8.204 ± 0.240^c	3.317 ± 0.030^a	0.073 ± 0.002^c
	STPs-S	72.662 ± 0.265^a	8.439 ± 1.128^b	3.006 ± 0.026^b	0.071 ± 0.001^c
	STPs-G	65.892 ± 0.178^b	9.790 ± 0.141^a	2.494 ± 0.005^c	0.096 ± 0.001^b
	STPs-I	60.142 ± 0.700^c	9.219 ± 0.737^a	1.678 ± 0.174^d	0.112 ± 0.001^a
FTPs	Undigested	60.173 ± 0.245^a	8.009 ± 0.247^c	2.825 ± 0.045^a	0.077 ± 0.000^c
	FTPs-S	58.697 ± 0.371^a	12.776 ± 0.039^b	2.654 ± 0.027^b	0.079 ± 0.000^c
	FTPs-G	54.303 ± 0.292^b	18.886 ± 0.058^a	2.414 ± 0.004^c	0.085 ± 0.002^b
	FTPs-I	40.577 ± 0.509^c	18.952 ± 0.171^a	1.598 ± 0.008^d	0.108 ± 0.000^a

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The change in reducing sugar levels can mirror the breakdown of polysaccharides and the breaking of glycosidic bonds during digestion.²⁸ After oral digestion, the reducing sugar content in STPs and FTPs showed no significant change, indicating their resistance to salivary amylase in the mouth. However, after gastrointestinal digestion, the reducing sugar content in STPs and FTPs increased. This may be due to the high-acid gastric condition breaking glycosidic bonds and exposing reducing ends.²⁹ Besides, pancreatic amylase and protease in the small intestine can partially degrade the polysaccharides, releasing small oligosaccharide monomers.³⁰ Following in vitro simulated digestion, STPs exhibited a more pronounced increase in reducing sugar content than FTPs (53.425% vs. 40.260%), which suggested that STPs are more susceptible to digestive effects in the gastrointestinal tract than FTPs, potentially attributable to structural differences that lead to more apparent hydrolysis. Whether this indeed occurs specifically during the digestive phase, however, requires further investigation.

3.1.2. Changes in M_w. As presented in Table 2, the polysaccharides from L. litseifolius green tea are characterized by high molecular weights. Comparative analysis revealed that STPs have a marginally lower M_w than FTPs, a difference that can likely be attributed to their distinct chemical compositions. Moreover, digestive degradation of polysaccharides can be indicated by alterations in their M_w. The M_w of STPs and FTPs decreased during digestion. Specifically, after oral digestion, the M_w of STPs increased from 1.111 × 10⁶ Da to 1.484 × 10⁶ Da, and that of FTPs increased from 1.723 × 10⁶ Da to 2.189 × 10⁶ Da. This may be due to the transglycosylation activity of salivary α-amylase under certain conditions, which can temporarily increase M_w by forming new glycosidic bonds, but it is uncertain whether STPs and FTPs are degraded. After further gastrointestinal digestion, the M_w of STPs was reduced to 6.977 × 10⁵ Da, and that of FTPs decreased to 9.620 × 10⁵ Da. The breakdown of polysaccharides during digestion is mainly affected by pH, with lower pH values enhancing their decomposition.³¹ STPs and FTPs are particularly sensitive to the acidic gastric environment. This sensitivity likely contributes to their significant reduction in M_w following gastric digestion. Besides, the rise in reducing sugar levels in both STPs and FTPs after gastric digestion further indicates glycosidic bond splitting and structural breakdown within the polysaccharides. This observation corresponds to findings from other studies.³²

Table 2 Effects of in vitro digestion on the molecular weight (M_w) and polydispersity (M_w/M_n) of STPs and FTPs

Samples	Stage	M w (Da)	M w/Mn
Each value is expressed as the mean ± SD (n = 3) of triplicate determinations. Different lowercase letters (a–d) indicate statistically significant differences (p < 0.05) for the same polysaccharide across different digestion time points. Undigested, STPs-S, STPs-G and STPs-I in STPs represent undigested STPs and saliva-digested, gastric-digested, and intestinal-digested STPs, respectively. Undigested, FTPs-S, FTPs-G and FTPs-I in FTPs represent undigested FTPs and saliva-digested, gastric-digested, and intestinal-digested FTPs, respectively.
STPs	Undigested	1.111 × 10^6 ± 0.052 × 10^6 ab	6.148^b
	STPs-S	1.484 × 10^6 ± 0.327 × 10^6 a	10.815^a
	STPs-G	1.389 × 10^6 ± 0.399 × 10^6 a	7.065^ab
	STPs-I	6.977 × 10^5 ± 0.33 × 10^5 b	10.757^a
FTPs	Undigested	1.723 × 10^6 ± 0.103 × 10^6 ab	7.165^b
	FTPs-S	2.189 × 10^6 ± 0.117 × 10^6 a	9.516^b
	FTPs-G	1.517 × 10^6 ± 0.492 × 10^6 b	7.903^b
	FTPs-I	9.620 × 10^5 ± 0.457 × 10^5 c	14.771^a

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3.1.3. Variation of FT-IR spectra. As shown in Fig. 1A and B, the absence of significant spectral shifts in the FTIR analysis between STPs and FTPs suggests a high degree of structural similarity, with both exhibiting the common functional groups diagnostic of polysaccharides. The digestion products of STPs and FTPs were similar in structure to the original polysaccharides, indicating that their structures remained stable during digestion. Specifically, a strong and broad absorption peak appeared between 3500 and 3200 cm⁻¹ in the spectra of STPs and FTPs, mainly due to the stretching oscillation of –OH in sugar molecules. Absorption peaks in the range of 2800–3000 cm⁻¹ were attributed to the C–H stretching vibrations in –CH₂.³³ Additionally, both polysaccharides exhibited strong absorption bands between 1600 and 1400 cm⁻¹, primarily caused by the asymmetric stretching vibrations of –COOH.³⁴ STPs had an absorption peak at 1647 cm⁻¹, while FTPs had one at 1646 cm⁻¹. These peaks are associated with C–H and C=O stretching vibrations, suggesting the potential presence of uronic acids in both STPs and FTPs. Furthermore, as detailed in section 3.1.1, quantitative analysis has already confirmed substantial uronic acid content in both polysaccharides. The FT-IR results thus provide additional confirmation of acidic functional groups, collectively indicating that STPs and FTPs are acidic polysaccharides. In the range of 1000–1200 cm⁻¹, there were three absorption peaks detected for the polysaccharides (STPs: 1017 cm⁻¹, 1072 cm⁻¹, 1101 cm⁻¹; FTPs: 1023 cm⁻¹, 1046 cm⁻¹, 1100 cm⁻¹). These peaks correspond to the stretching vibrations of C–O–H and C–O–C, signifying the presence of pyranose rings within the polysaccharides. Moreover, STPs displayed absorption bands at 851 cm⁻¹ and 935 cm⁻¹, whereas FTPs showed them at 862 cm⁻¹ and 938 cm⁻¹, implying that the predominant glycosidic bonds in the polysaccharides are α-glucosidic and β-glycosidic bonds.³⁵ Moreover, absorption peaks were detected at 1500 cm⁻¹ for STPs and 1488 cm⁻¹ for FTPs, which are characteristic of the C=O group in proteins.³¹ This suggests that the polysaccharides contain protein, which corroborates the prior chemical composition analysis.

Fig. 1 FT-IR spectra of STPs and FTPs during in vitro simulated digestion. (A) FT-IR spectra of STPs and (B) FT-IR spectra of FTPs. STPs-S, STPs-G, and STPs-I represent saliva-digested, gastric-digested, and intestinal-digested STPs and FTPs-S, FTPs-G, and FTPs-I represent saliva-digested, gastric-digested, and intestinal-digested FTPs respectively.

3.1.4. Changes of monosaccharide composition. STPs and FTPs differ in their monosaccharide composition. As shown in Table 3 and Fig. S1, STPs were composed mainly of rhamnose (17.229%), xylose (38.768%), mannose (3.051%), glucose (7.517%), and galactose (33.435%). In contrast, FTPs consisted primarily of rhamnose (2.467%), xylose (19.883%), arabinose (9.568%), mannose (12.949%), glucose (27.457%), and galactose (27.675%). The monosaccharide composition of STPs and FTPs changed during digestion. Galactose and rhamnose are major monosaccharides in STPs, while glucose and galactose are predominant in FTPs. These differences in monosaccharide contents and proportions are likely due to the varying maturity of the two grades of L. litseifolius [Hance] Chun green tea, which affects the types and amounts of bio-components accumulated during growth. After digestion, the types of monosaccharides in STPs and FTPs remained unchanged, but their proportions varied with the digestion stage. Furthermore, STPs showed a significant increase in glucose, xylose, and mannose, while FTPs saw a rise in galactose, xylose, and arabinose after digestion, and the content of other monosaccharides also changed dynamically during digestion. This may be due to the introduction of digestive enzymes and varying pH levels across the three digestive stages, which can cause the cleavage and reorganization of the glycosidic bonds linking monosaccharides, leading to dynamic changes in their proportions within the polysaccharides.³⁶ This is similar to other research study results.^5,29 In summary, these results demonstrate that STPs and FTPs diverged markedly in both the types and molar ratios of their monosaccharide constituents. Furthermore, while simulated digestion induced a quantitative redistribution of these monosaccharides, their qualitative profile remained unchanged.

Table 3 The monosaccharide composition of STPs and FTPs during in vitro stimulated digestion (%)

Samples	Stage	Rha	Xyl	Ara	Man	Glu	Gal
Each value is expressed as the mean ± SD (n = 3) of triplicate determinations. Different lowercase letters (a–d) indicate statistically significant differences (p < 0.05) for the same polysaccharide across different digestion time points. Undigested, STPs-S, STPs-G and STPs-I in STPs represent undigested STPs and saliva-digested, gastric-digested, and intestinal-digested STPs, respectively. Undigested, FTPs-S, FTPs-G and FTPs-I in FTPs represent undigested FTPs and saliva-digested, gastric-digested, and intestinal-digested FTPs, respectively. Rha, rhamnose; Xyl, xylose; Ara, arabinose; Man, mannose; Glu, glucose; and Gal, galactose.
STPs	Undigested	17.229 ± 1.081^a	38.768 ± 0.846^a	0.000 ± 0.000^a	3.051 ± 0.596^b	7.517 ± 0.454^d	33.435 ± 0.790^a
	STPs-S	6.182 ± 0.053^b	29.344 ± 0.200^b	0.000 ± 0.000^a	5.811 ± 0.189^b	25.310 ± 0.257^c	33.354 ± 0.594^a
	STPs-G	7.727 ± 0.442^c	21.466 ± 2.290^d	0.000 ± 0.000^a	6.852 ± 4.064^b	34.691 ± 2.049^a	29.264 ± 0.708^b
	STPs-I	3.862 ± 0.202^d	27.346 ± 2.040^c	0.000 ± 0.000^a	19.335 ± 4.327^a	29.049 ± 1.325^b	20.407 ± 3.410^c
FTPs	Undigested	2.467 ± 0.186^b	19.883 ± 4.670^b	9.568 ± 5.23^b	12.949 ± 2.886^b	27.457 ± 4.996^a	27.675 ± 2.002^a
	FTPs-S	2.753 ± 0.418^b	19.626 ± 2.211^b	8.103 ± 0.547^b	22.754 ± 0.910^a	22.346 ± 1.428^a	24.418 ± 0.636^a
	FTPs-G	2.451 ± 0.070^b	18.766 ± 2.232^b	9.533 ± 5.467^b	12.451 ± 3.074^b	28.170 ± 5.135^a	28.629 ± 0.861^a
	FTPs-I	5.808 ± 1.137^a	34.886 ± 6.329^a	21.428 ± 6.667^a	0.790 ± 0.159^c	8.117 ± 1.608^b	28.971 ± 4.470^a

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3.2. Variation of biological activities

3.2.1. Changes in antioxidant activity. As depicted in Fig. 2A–C and Table S1, the antioxidant activities were evaluated using DPPH, reducing power, and metal-chelating ability assays. STPs demonstrated significantly stronger antioxidant activity than FTPs across all evaluated assays. Specifically, STPs exhibited a higher DPPH radical scavenging capacity (26.899 ± 1.318 μg Vc eq. per mg) compared to FTPs (25.171 ± 0.812 μg Vc eq. per mg), a markedly greater reducing power (77.284 ± 3.030 μg BHT eq. per mg vs. 71.249 ± 1.807 μg BHT eq. per mg), and a superior metal chelating ability (64.101 ± 0.374 μg EDTA eq. per mg vs. 47.866 ± 0.242 μg EDTA eq. per mg). After simulated digestion, the residual antioxidant activities of STPs, measured using DPPH, reducing power, and metal-chelating ability, decreased from 97.920% to 55.492%, 79.088% to 35.621%, and 81.604% to 50.888%, respectively, while the residual activities of FTPs declined from 86.415% to 43.199%, 85.118% to 38.449%, and 93.699% to 68.109%, respectively. The reduction in antioxidant capacity during digestion is closely related to the chemical composition of the polysaccharides.³⁷ The antioxidant properties of polysaccharides are influenced by their content of reducing sugars, uronic acids, and proteins.³² Uronic acids, containing electrophilic groups like ketones and aldehydes, are potent antioxidants. As digestion continues, the glycosidic bonds in polysaccharides break, creating reducing ends on sugar chains and boosting reducing sugar content.³⁸ This rise in reducing sugars diminishes antioxidant capacity. Furthermore, the M_w of polysaccharides also affects their antioxidant activity.³⁹ Research has shown that polysaccharides with a lower M_w have weaker antioxidant activity compared to higher molecular weight ones.³¹ It is plausible that the molecular weight disparity between STPs and FTPs, as detailed above, accounts for the stronger antioxidant activity observed in STPs. The M_w of both polysaccharides gradually decreases after digestion, which may be another reason for the reduced antioxidant activity. However, although the antioxidant activities of STPs and FTPs decreased during digestion, the residual activities after digestion were still relatively high. This indicates that after intestinal digestion, the antioxidant activity of the polysaccharides decreased slightly but remained at a better level.

Fig. 2 Residual biological activities of STPs and FTPs during digestion. (A) Residual activity of DPPH; (B) residual activity of reducing power; (C) residual activity of metal chelation; (D) residual activity of α-amylase inhibitory activity; and (E) residual activity of α-glucosidase inhibitory activity. Different lowercase letters (a–d) indicate statistically significant differences (p < 0.05) for the same polysaccharide across different digestion time points.

3.2.2. Changes in hypoglycemic activity. α-Amylase and α-glucosidase, as key carbohydrate-hydrolyzing enzymes, demonstrate therapeutic potential in mitigating hyperglycemia and postprandial glycemic spikes. Specifically, α-amylase cleaves starch into oligosaccharides, whereas α-glucosidase mediates the enzymatic conversion of disaccharides to absorbable monosaccharides.⁴⁰ The inhibitory efficacy against these enzymatic targets serves as a critical metric for assessing glucose-lowering potential in vitro.⁴¹ As shown in Fig. 2D and E and Table S1, STPs and FTPs exhibited similar inhibitory effects on α-amylase, but FTPs showed better inhibition of α-glucosidase than STPs. We also investigated the variation in the hypoglycemic activities of STPs and FTPs during in vitro simulated digestion; the inhibitory activities of STPs and FTPs on α-amylase decreased from 89.447% to 76.144% and from 91.047% to 79.002%, respectively, while those on α-glucosidase dropped from 88.174% to 55.156% and from 78.871% to 57.915%, respectively. This indicates that digestion affects the hypoglycemic activities of STPs and FTPs. This variance could stem from stage-dependent compositional variances in polysaccharides and their interactions with the digestive enzymatic milieu. Mechanistically, empirical evidence demonstrates that α-amylase and α-glucosidase inhibition correlates with unbound carboxyl/hydroxyl structural motifs within polysaccharide matrices. These functional groups engage in stereospecific interactions with the catalytic residues of digestive hydrolases, inducing conformational changes that impair enzymatic activity.⁴² STPs and FTPs possess elevated uronic acid and free carboxyl group contents. These components can react with the amino acid residues of enzymes, resulting in significant enzyme inhibitory activity. Furthermore, changes in uronic acid content can serve as an indicator of variations in hypoglycemic activity. Following digestion, the inhibition of α-amylase and α-glucosidase by STPs decreased more markedly than that by FTPs, a phenomenon potentially attributable to differences in their chemical structures and compositions. Specifically, STPs exhibited a more pronounced reduction in uronic acid content after digestion compared to FTPs (Table 1), which may help explain the distinct decline in their respective hypoglycemic activities. Throughout digestion, the quantity of uronic acid components in both STPs and FTPs exhibited a downward trajectory, consistent with the reduction in α-amylase and α-glucosidase inhibitory activities. Similarly, although the hypoglycemic activities of STPs and FTPs are affected by the digestive tract, they still retain strong hypoglycemic effects after digestion, indicating their potential for effective blood sugar reduction.

3.3. Variation of characteristics in STPs and FTPs during in vitro fermentation

3.3.1. Variations in total carbohydrate and reducing sugar. Studies have confirmed that polysaccharides can reach the colon and are subsequently degraded through utilization by the colonic gut microbiota.³⁰Fig. 3A illustrates the changes in total carbohydrate content during fermentation at 0, 6, 12, and 24 h. We found that the total carbohydrate in the INL, STPs, and FTPs groups showed a decreasing trend as the fermentation time increased, indicating that carbohydrates are continuously utilized and degraded by the intestine. Meanwhile, the composition of reducing sugars also decreased with fermentation time (Fig. 3B). The experimental groups exhibited progressive reductions in residual polysaccharide concentrations during fermentation: INL decreased from 3.160 ± 0.032 mg mL⁻¹ to 0.485 ± 0.009 mg mL⁻¹, STPs decreased from 2.973 ± 0.062 mg mL⁻¹ to 1.006 ± 0.009 mg mL⁻¹, and FTPs decreased from 1.956 ± 0.006 mg mL⁻¹ to 0.473 ± 0.011 mg mL⁻¹ over 24 h. These findings are identical to those of fresh passion fruit peel polysaccharides and oolong tea polysaccharides during in vitro simulated digestion.^43,44 Following in vitro fermentation, STPs and FTPs were degraded by 66.2% and 75.8%, respectively. The degradation rates exceeding 50% indicate that both polysaccharides could be effectively utilized by the gut microbiota, with FTPs exhibiting a higher degree of microbial utilization compared to STPs. These results confirm their value as fermentable substrates for the colonic microbiota and demonstrate favorable bioavailability.

Fig. 3 Changes in reducing sugar contents, total carbohydrates, pH values, and SCFA content of STPs and FTPs during different fermentation times. (A) Total carbohydrate contents; (B) reducing sugar contents; (C) pH values; (D) total SCFA acid; (E) acetic acid; (F) propionic acid; (G) i-butyric acid; (H) n-butyric acid; (I) i-valeric acid; and (J) n-valeric acid. Different letters (a–d) denote significant differences (p < 0.05) for the same sample over the fermentation period (0, 6, 12, and 24 h). OR, BLK, and INL represent the original fecal group, blank group, and inulin group.

3.3.2. Changes of pH values. Generally, the dynamic variations in pH during in vitro fermentation of fecal samples serve as an indirect indicator of colonic microbiota-mediated degradation and utilization of natural polysaccharides.⁴⁵ The variations in the pH of the fermentation cultures for BLK, INL, STPs, and FTPs are presented in Fig. 3C. After 6 h of fermentation, all groups showed significant pH reductions, with the INL group exhibiting the most notable decrease and the lowest pH values. This phenomenon may be due to the rapid microbial utilization and degradation of inulin (a high-quality prebiotic) during the initial fermentation phase.⁴⁶ After 24 h of fermentation, pH stabilization across groups indicated near-complete microbial catabolism of polysaccharides and inulin. Besides, STP and FTP groups maintained pH levels intermediate between the BLK and INL groups, a discrepancy potentially associated with SCFA accumulation by the gut microbiota. Collectively, the observed pH dynamics suggest that microbial degradation of polysaccharides generates acidic metabolites capable of modulating the colonic pH.

3.3.3. Effects of STPs and FTPs on the gut microbiota composition. The gut microbiota is vital to the host, supplying nutrients, combating pathogens, and preserving intestinal equilibrium. It can metabolize polysaccharides to boost beneficial substances and adjust the gut microbiota's structure and abundance. To explore the impact of STPs and FTPs on the gut microbiota, we used 16S rRNA high-throughput sequencing to investigate bacterial community diversity. In total, 940 700 raw reads were retrieved from the 15 samples. Rarefaction curves assessed whether the sample size was sufficient for exploring the variety and abundance of the gut microbiota. The α-diversity index, indicating microbial community richness, was assessed using the Sobs, Ace, Chao, Shannon, Coverage, and Simpson indices, as shown in Table S3. Furthermore, as shown in Fig. 4A and B, the Chao curve became smoother with increasing sequencing depth, indicating high sequencing data coverage and good representation of species diversity. The Shannon curve, assessing microbial diversity in samples, also gradually smoothed with increased sequencing depth. These data suggest that sufficient sample data were obtained for subsequent analyses.

Fig. 4 Intestinal flora composition after 24 h of fermentation. (A) Chao curves; (B) Shannon curves; (C) PCoA analysis of the gut microbiota; and (D) hierarchical clustering tree. OR, BLK, and INL represent the original fecal group, blank group, and inulin group.

The connection between the microbial community beta-diversity and gut microbial community structure is frequently illustrated through principal coordinate analysis (PCoA) and cluster analysis. As depicted in Fig. 4C, PCoA was conducted at the ASV level, and the two axes—PC1 (56.14%) and PC2 (29.16%)—accounted for 85.30% of the total variance among samples. Comparing different samples revealed that the INL, STP, and FTP groups were distinct from the OR group, indicating significant differences. Moreover, the STP, FTP, and INL groups were distinctly separated from each other. STPs and FTPs also showed significant separation, suggesting that despite structural similarities, their compositional differences affect polysaccharide structure and modulate gut microbiota differently. Cluster analysis in Fig. 4D confirmed significant differences in community clustering among the four groups (OR, INL, STPs, and FTPs), validating the reliability of the PCoA results. In summary, these results indicate that STPs and FTPs significantly impact the gut microbial community structure, with differences in their regulatory effects.

At the phylum level (Fig. 5A), the dominant bacterial phyla in the samples were Firmicutes, Actinobacteria, Proteobacteria, and Bacteroidetes. Compared with the OR group, the microbial composition of the samples underwent alterations following in vitro fermentation. Desulfobacterota, a harmful bacterial group that thrives under anaerobic conditions, increased in the BLK group. In contrast, it was absent in the INL, STPs, and FTPs groups, indicating that polysaccharides can suppress the growth of harmful bacteria. In the STPs and FTPs groups, Firmicutes and Actinobacteria increased, but Bacteroidetes decreased. The lower pH values (STPs: 5.51 ± 0.03 and FTPs: 5.14 ± 0.02) after 24-hour fermentation might explain the increase in Firmicutes and Actinobacteria, as comparable findings have been reported in another polysaccharide research.⁴⁷ Furthermore, the STPs group was characterized by a distinct microbial profile compared to the FTPs group, featuring a lower relative abundance of Bacteroidetes (52.643% vs. 64.033%) alongside higher relative abundances of Proteobacteria (39.078% vs. 26.737%) and Actinobacteriota (0.07% vs. 0.04%). Bacteroidetes, capable of carbohydrate utilization and SCFA production in the human gut, exhibited a declining trend in these groups.⁴⁸ The STP and FTP groups also had significant differences in Firmicutes and Proteobacteria, while other bacterial groups had similar abundances. Actinobacteria, being Gram-positive, are beneficial to the gut microbiota, helping to regulate serum cholesterol and prevent intestinal diseases. An increase in Proteobacteria may lead to gut dysbiosis, mild inflammation, or chronic colitis, though they are commonly found in healthy human fecal microbiota.⁴⁹ Many members of Firmicutes are known for producing butyrate.²⁷Bacteroidetes can secrete more lytic enzymes and glycosidic bond hydrolases, breaking down polysaccharides into SCFAs. They also decompose the galactan side chains and rhamnogalacturonan I (RGI) main chains of polysaccharides, thus facilitating the degradation of intestinal polysaccharides.²³ These results show that fermentation can change microbial community composition, likely due to oxygen consumption. During fermentation, STPs and FTPs can modulate the gut microbiota structure, increasing community evenness by promoting the proliferation of anaerobes and welfare-promoting bacteria.

Fig. 5 The influence of STPs and FTPs on the intestinal flora composition after 24 h of fermentation. (A) Phylum level; (B) community heatmap analysis on the genus level; (C) histogram of difference analysis using the LDA score; (D) correlation heatmap between the genus-level gut microbiota and SCFAs; (E) LEfSe from the phylum to genus level. OR, BLK, and INL represent the original fecal group, blank group, and inulin group.

To further determine the specific changes in the gut microbiota caused by STPs and FTPs during fermentation, we assessed the genus level of microbial composition. As shown in Fig. 5B, Parabacteroides, Echeichia-Shigella, Prevotella, Phascolarctobacterium, and Bifidobacterium are the main genera in these samples. In contrast to the OR group, Shigella shows the largest increase in the BLK group, while its abundance decreases in the STPs and FTPs groups. As the most common pathogen of bacterial dysentery in humans, Shigella grows better under anaerobic conditions.³⁶ So, the anaerobic environment in the BLK group promotes Shigella growth. However, the addition of prebiotics (INL, STPs, and FTPs) effectively inhibits this growth, indicating that polysaccharides can suppress harmful bacteria in the gut. Compared with the BLK group, the STP and FTP groups harbor more beneficial bacteria. Following polysaccharide regulation, the abundance of dominant species like Bifidobacterium and Lachnospira increases. Bifidobacterium, a probiotic in the human gut, can modulate the immune system, exhibit anticancer activity, and notably alleviate metabolic disorders in obese individuals.⁵⁰Lachnospira is present in the intestines of most healthy individuals and may be a beneficial bacterium involved in various carbohydrate metabolic processes.⁵¹ The findings indicate that polysaccharides from L. litseifolius green tea can support the host gut microbiota health by boosting beneficial bacteria and curbing harmful ones.

Significantly different microbial groups after STPs and FTPs intervention were analyzed by Linear Discriminant Analysis (LDA). Fig. 5C and E mainly show differences in the microbial community at the phylum and genus levels, with an LDA score between 3.5 and 5, indicating statistically significant biomarkers. In the OR, BLK, INL, STPs, and FTPs groups, there are 10, 10, 10, 4, and 6 species, respectively. Besides, in the STP group, they were Megamonas, Negativicutes, Selenomonadaceae, Veillonellales-Selenomonadales, and Firmicutes. In the FTPs group, the dominant genera were Acidaminococcales, Phaseolactobacterium, Acidaminococcaceae, and Lachnospiraceae_UCG-004. Relative to other groups, the distinctive genera in the STPs and FTPs groups were within Firmicutes, many being beneficial SCFA-producing genera. LEfSe analysis validated that those polysaccharides boosted beneficial gut bacteria abundance, suggesting their prebiotic capacity to regulate the gut microbiota.

3.3.4. Variation of SCFAs during fermentation. SCFAs maintain normal gut function by influencing microbial enzyme activity and are crucial for inhibiting colorectal cancer and modulating immunity.⁵² Studies show that SCFAs are crucial metabolic outcomes from the fermentation of dietary polysaccharides by colonic microbes.³⁴ Therefore, we analyzed the SCFA levels during fermentation. Fig. 3D–J show the changes in total SCFA and six SCFA levels at 0, 6, 12, and 24 h. The results revealed that the content of SCFAs varied across groups at different fermentation times. After 24 h, the total acid levels in the BLK, INL, STP, and FTP groups increased from 0.655 ± 0.035 mM, 1.995 ± 0.117 mM, 1.679 ± 0.153 mM, and 1.332 ± 0.145 mM to 6.722 ± 0.415 mM, 25.506 ± 4.559 mM, 11.471 ± 0.608 mM, and 21.732 ± 0.967 mM, respectively. The STP and FTP groups had higher total SCFA levels than the BLK group, but lower than the INL group. Notably, the FTP group exhibited a higher content than the STP group after 24 h, indicating better fermentation performance of FTPs than that of STPs. Interestingly, acetic acid reached the highest concentration, followed by propionic acid, with valeric acid at the lowest after 24 h. In the INL group, acetic acid rose from 0.314 ± 0.077 mM to 5.864 ± 1.011 mM, and propionic acid from 0.411 ± 0.028 mM to 6.659 ± 0.759 mM. In the STP and FTP groups, acetic acid increased from 0.440 ± 0.141 mM and 0.236 ± 0.074 mM to 2.492 ± 0.161 mM and 6.257 ± 0.892 mM, respectively, and propionic acid rose from 0.183 ± 0.025 mM and 0.156 ± 0.013 mM to 2.079 ± 0.146 mM and 5.872 ± 0.318 mM, respectively. Furthermore, the level of n-butyric acid rose from 0.493 ± 0.080 mM to 1.679 ± 0.036 mM and from 0.389 ± 0.016 mM to 2.643 ± 0.124 mM in the STP and FTP groups, respectively.

Research has demonstrated that acetic acid acts as a major energy source for the heart, brain, and muscles, and plays a significant role in gluconeogenesis, lipogenesis, and cholesterol synthesis.⁵³ In the gut microbiota, Bifidobacterium breaks down polysaccharides to yield acetic acid. Besides, propionic acid cuts liver and blood fatty acids, curbs appetite, and aids in liver cholesterol metabolism. Bacteroides and Phascolarctobacterium are key producers of these beneficial short-chain fatty acids.³⁴ Butyrate is a primary energy source for the gut microbiota and significantly benefits human health. Multiple genera within the Lachnospiraceae family possess the ability to produce butyrate.²⁷ In the STPs and FTPs groups, the buildup of acetic and propionic acids is linked to the richness of Bifidobacterium, which is consistent with the previously stated microbial community composition. Interestingly, FTPs demonstrated a higher acid-producing capacity than STPs, suggesting more efficient microbial utilization of FTPs. The structural complexity and higher uronic acid content of STPs may hinder their degradation by common gut microorganisms, resulting in comparatively lower microbial utilization.⁵⁴ Previous studies have indicated that Ara can be metabolized by beneficial bacteria such as Bifidobacterium, Lactobacillus, and Megamonas to produce acetate and propionate. Furthermore, Ara-containing polysaccharides have been shown to indirectly suppress the growth of potential pathogens, thereby helping maintain gut microecological balance.⁵⁵ The absence of Ara in STPs may therefore contribute to their reduced acid-producing capacity and weaker regulatory effects on the gut microbiota compared to FTPs. In summary, differences in monosaccharide composition and chemical composition between STPs and FTPs lead to distinct responses in the gut microbial community, ultimately resulting in significant differences in their acid-producing capabilities.

3.3.5. Correlation between SCFAs and the gut microbiota. Spearman's rank correlation analysis was employed to assess the association between SCFAs and the gut microbiota during in vitro fermentation, as shown in Fig. 5D. Megamonas, Bifidobacterium, and Prevotella showed positive correlations with the majority of SCFAs, among which Bifidobacterium exhibited significant correlations with butyrate and propionate (P < 0.05). In contrast, genera such as Faecalibacterium, Lachnospiraceae and Phascolarctobacterium were significantly negatively correlated with SCFA levels (P < 0.05). After fermentation, the concentrations of all SCFAs increased, with acetate and propionate being the most abundant, a pattern consistent with the relative abundance of Bifidobacterium. These findings suggest that STPs and FTPs are likely degraded and utilized primarily by Bifidobacterium, resulting in acetate and propionate as their major metabolic end products.

4. Conclusion

Our research showed that polysaccharides from special-grade and first-grade polysaccharides from L. litseifolius green tea differ in their chemical composition, monosaccharide composition, M_w, and in vitro bioactivity, but not in the chemical structure. STPs have better in vitro bio-activities than FTPs. Besides, in vitro simulated digestion caused partial degradation of STPs and FTPs. Total carbohydrate, reducing sugar, uronic acid, M_w, and monosaccharide contents showed dynamic changes, but the chemical structure was not affected. At the same time, in vitro antioxidant and hypoglycemic activities were partially lost. However, most STPs and FTPs can still enter the colon. In vitro fermentation results confirmed that the STPs and FTPs could be utilized and had the ability to regulate the gut microbes. The total carbohydrate and reducing sugar levels in STPs and FTPs fell markedly. Meanwhile, both STPs and FTPs demonstrated the ability to remodel the gut microbiota structure by inhibiting pathogenic bacteria and enriching beneficial populations. These changes were accompanied by a reduction in intestinal pH and enhanced production of short-chain fatty acids (SCFAs), particularly acetate and propionate. FTPs demonstrated a superior ability to support gut microbial communities compared to STPs, as evidenced by more efficient utilization, enhanced SCFA production, and stronger modulation of the microbial structure with differences rooted in their distinct monosaccharide profiles and chemical characteristics. Importantly, both STPs and FTPs consistently promoted beneficial structural and functional changes in the gut microbiota. In summary, we suggest that STPs and FTPs could be potential health-promoting prebiotics that improve gut health. Our study reveals the composition and structure of L. litseifolius green tea polysaccharides, explains their changes during digestion and impact on the gut microbes using in vitro models, and offers new insights and a scientific basis for their use as functional ingredients and prebiotics. However, our study has certain limitations. in vitro simulated digestion and fermentation models serve primarily as preliminary investigative tools, and their outcomes are susceptible to factors such as digestion parameter settings and inter-individual variations in fecal samples. Therefore, further in vivo experiments are necessary to rigorously validate the metabolic trajectories within a complete biological system. Future research could also deeply analyze the structure–activity relationship between the polysaccharide molecular conformation and its observed bioactivity and probiotic effects.

Author contributions

Qingying Luo: validation, funding acquisition, and writing – review & editing. Xuewei Liao: methodology, formal analysis, visualization, and writing – original draft. Lijia Zhang: formal analysis, visualization, and writing – review & editing. Zhengfeng Fang: supervision and visualization. Hong Chen: supervision. Bin Hu: supervision. Yuntao Liu: methodology, conceptualization, supervision, validation, and funding acquisition. Zhen Zeng: conceptualization, supervision, validation, and funding acquisition.

Conflicts of interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Ethics statement

This statement is to certify that all evaluations performed in the study “In vitro simulated digestion and fermentation of Lithocarpus litseifolius [Hance] Chun green tea polysaccharides and their modulation of the gut microbiota” were in accordance with Sichuan Agricultural University Academic Ethical and Welfare Committee (Approval Number: 20250531).

Declaration of consent for the use of fecal samples

As the Principal Investigator of this research project, I hereby confirm that we have strictly adhered to research ethics and legal regulations, obtaining explicit, voluntary, and informed consent from all stool sample donors. The six participating donors voluntarily contributed their stool samples after achieving full comprehension of the research objectives, methodologies, potential risks, and personal privacy protection measures. They explicitly authorized the use of their samples for this specific study. We commit to strictly complying with relevant research ethics guidelines to ensure donor privacy and information security. The use of all stool samples will be strictly limited to the research purposes and scope specified in this project. Furthermore, we will ensure that sample storage, handling, and analysis conform to applicable biosafety standards.

Data availability

The data that support the findings of this study are available from the corresponding author upon reasonable request. Supplementary information (SI) is available. See DOI: https://doi.org/10.1039/d5fo04379d.

Acknowledgements

This work was financially supported by Sichuan Tea Innovation Team (SCCXTD-2024-10), the Project of Sichuan Province's Leading County for Agricultural Science and Technology Modernization Application, National College Students’ Innovation and Entrepreneurship Training Program (202310626013), Sichuan Provincial Joint Fund for Science and Technology Education General Project (2024NSFSC2059).

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Footnote

† These authors contributed equally to this article.

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