Polygonatum cyrtonema Hua fructan ameliorates ulcerative colitis via gut microbiota modulation and follistatin targeting

Abstract

Ulcerative colitis (UC) is a chronic inflammatory bowel disease with persistent colonic inflammation and inadequate therapeutic options. The medicinal and edible plant Polygonatum cyrtonema Hua from Jiuhua Mountain contains polysaccharides with potent anti-inflammatory activities. In this study, a low-molecular-weight fructan (Mw = 2087 Da), designated PCP2, was isolated and purified from its rhizome. Biologically, PCP2 administration markedly alleviated disease severity in dextran sulfate sodium (DSS)-induced colitis mice, as shown by the improvement in multiple indicators of colon injury and inflammation. Fecal microbiota transplantation and antibiotic depletion experiments revealed that the protective effects of PCP2 are mediated through both modulation of the gut microbiota and additional microbiota-independent pathways. Importantly, through molecular dynamics simulations, microscale thermophoresis, and surface plasmon resonance assays, follistatin (Fst) is identified as a direct binding target of PCP2. Functional validation using siRNA-mediated Fst knockdown in Caco-2 cells, combined with adenovirus-mediated knockdown in the murine colon, confirmed that PCP2 exerts its therapeutic effect by directly interacting with Fst and suppressing the BMP4/Smad1/ID1 signaling axis. In summary, PCP2 ameliorates ulcerative colitis via dual mechanisms involving restoration of gut microbiota homeostasis and direct targeting of Fst. These findings establish a novel therapeutic strategy and support the clinical development of P. cyrtonema Hua from Jiuhua Mountain as a functional food for intestinal health.

Introduction

Ulcerative colitis (UC) is a chronic bowel disorder marked by persistent inflammation of the mucosal layer in the colon and rectum¹. Its clinical manifestations include recurrent bloody diarrhea, abdominal pain, and extraintestinal symptoms. The disease course alternates between periods of remission and activity². The pathogenesis of UC includes genetic susceptibility, immune dysregulation, intestinal flora imbalance, and environmental factors³. Currently, treatment primarily consists of 5-aminosalicylic acid, glucocorticoids, and biological agents⁴. However, these treatments are limited by insufficient efficacy, long-term side effects, and high recurrence rates. Surgical resection can cure the disease but significantly impacts quality of life⁵. Thus, developing more potent and individualized therapeutic modalities to improve patient outcomes is essential.

The gut microbiota (GM) functions as a pivotal modulator of intestinal health, orchestrating dietary fiber metabolism to generate immunomodulatory short-chain fatty acids (SCFAs) (e.g., butyrate), which maintain immune homeostasis and reinforce the mucosal barrier⁶. In UC, dysbiosis diminishes microbial diversity, depletes beneficial butyrate producers (e.g., Faecalibacterium prausnitzii), increases the abundance of pathobionts (e.g., Escherichia coli), and markedly decreases SCFAs⁷. These changes disrupt intestinal barrier integrity and anti-inflammatory responses, perpetuating a pro-inflammatory milieu⁸. Polysaccharides, as dietary fibers and functional substances, can improve inflammatory bowel diseases such as colitis by regulating the intestinal flora. Previous studies have shown that Abelmoschus manihot polysaccharides regulated the abundance of the intestinal probiotic Akkermansia muciniphila (A. muciniphila), promoting MUC2 secretion and enhancing the intestinal mucus barrier function⁹. The steamed Polygonatum cyrtonema polysaccharide exerts significant anti-inflammatory and protective effects on colitis by regulating the intestinal flora and restoring the mucosal barrier¹⁰. Consequently, targeting GM-driven mechanisms represents a potential avenue for UC therapeutics. P. cyrtonema is a prestigious plant with a long history of dual medicinal and edible use in East Asia, officially recognized as a “medicine-food homology” material. In particular, Polygonatum cyrtonema Hua grown on Jiuhua Mountain, a National Geographical Indication product in China, is renowned for the superior quality and high polysaccharide content of its rhizomes, which has been used for both medicinal and edible purposes¹⁰. Polysaccharides, as its main bioactive components, exhibit remarkable immunomodulatory, anti-inflammatory, and intestinal health-promoting properties¹¹, including potent anti-oxidative and intestinal barrier-protective activities. However, while crude extracts of P. cyrtonema have shown promise in regulating intestinal homeostasis, the specific therapeutic efficacy and underlying molecular targets of its purified, low-molecular-weight fructans in the context of UC remain to be fully elucidated.

Beyond microbiota-mediated effects, host-derived proteins are critical therapeutic targets. Follistatin (Fst), a multifunctional glycoprotein, is a potent antagonist of the TGF-β superfamily that primarily neutralizes activins and bone morphogenetic proteins (BMPs) to modulate inflammatory and regenerative pathways¹². Notably, Fst critically regulates the BMP/Smad signaling axis, balancing intestinal inflammation and mucosal repair¹³. Previous studies demonstrated that Fst improved 2,4,6-trinitrobenzenesulfonic acid and DSS-stimulated UC in mouse models, evidenced by increased survival rates, reduced pro-inflammatory cytokine levels, enhanced epithelial cell proliferation, and greater tissue repair capacity¹⁴. In addition, Fst has been shown to alleviate intestinal inflammation by attenuating the release of pro-inflammatory cytokines (e.g., TNF-α, and IL-6) and modulating anti-inflammatory cytokines such as IL-10, while simultaneously repairing the intestinal barrier through intestinal epithelial cell proliferation, thereby enhancing its anti-inflammatory properties¹⁴. Given the pivotal role of Fst, we hypothesized that Fst acts as a key modulator of the pathogenesis of colitis and therefore represents a potential therapeutic target.

Therefore, this study aimed to investigate the protective effects of Polygonatum cyrtonema polysaccharide (PCP2) against UC, and to elucidate its underlying mechanisms as a bioactive food-derived polysaccharide. A multi-faceted approach was employed, beginning with an in vivo assessment of PCP2’s therapeutic efficacy in a murine UC model. To comprehensively delineate the underlying mechanisms, 16S rRNA sequencing, metabolomics, and Olink proteomics were integrated, with causal inferences verified through fecal microbiota transplantation (FMT) and antibiotic perturbation experiments. Furthermore, the specific targeting of Fst by PCP2 was confirmed using molecular dynamics (MD) simulations, microscale thermophoresis (MST), and surface plasmon resonance (SPR). The functional necessity of Fst in mediating the protective effects of PCP2 was subsequently validated through both in vitro siRNA-mediated knockdown and in vivo adenovirus-mediated specific knockdown. Collectively, this research offers a solid foundation for the potential use of PCP2 in UC treatment and paves the way for further advancement of polysaccharide-based therapeutic agents from P. cyrtonema Hua sourced from Jiuhua Mountain.

Results

Extraction, purification, and structural analysis of PCP2

The extraction process of PCP2 is shown in Fig. 1A. The crude polysaccharides (PCPs) were fractionated on a DEAE-52 cellulose column eluted sequentially with deionized water and NaCl solutions of increasing concentration, and an elution profile was generated (Figure. S1A, Supporting Information). The fraction eluted with NaCl (0.1 M) was designated as PCP2, with a yield of 0.76% by weight. Subsequently, PCP2 was further purified using a Sephadex G-50 column. The high purity and homogeneous molecular weight profile of PCP2 were rigorously confirmed. HPGPC analysis revealed a single symmetrical peak (Fig. 1D), indicating high molecular homogeneity. Furthermore, UV-visible spectrum analysis (Fig. 1B) confirmed the absence of nucleic acid or protein contamination, as evidenced by the lack of absorption peaks at 260 and 280 nm, validating the high purity of PCP2. Consistent with these findings, the anthrone-sulfuric acid method determined a total sugar content of 92.3% (w/w), with no uronic acid detected, collectively affirming the high purity of PCP2.

Fig. 1. Separation, purification, and fundamental characterization of PCP2.

This figure was created with BioRender.com. A Extraction process of PCP2. B UV spectrum. C IR spectrum. D Molecular weight. E Monosaccharide composition. (Fig. 1A was created with BioRender.com).

The infrared (IR) spectrum of PCP2 showed seven characteristic peaks (Fig. 1C). The stretching peak at 3416.24 cm⁻¹ corresponded to the –OH stretching vibration, while that at 2933.56 cm⁻¹ was attributed to the C–H stretching vibration. The absence of a peak at 1740 cm⁻¹ indicated that uronic acids were not present in the polysaccharide structure¹⁵. The peak at 1632.45 cm⁻¹ corresponded to the C = O stretching vibration, and that at 1023.31 cm⁻¹ corresponded to the C–O–C stretching vibration, suggesting the presence of pyranose units in PCP2¹⁶. The stretching peak at 927.61 cm⁻¹ indicated the existence of furanose units in PCP2¹⁷. The weight-average molecular weight (Mw) of PCP2 was calculated as 2087 Da, with a number-average molecular weight (Mn) of 1599 Da, yielding a polydispersity index (PDI, Mw/Mn) of 1.305, which indicates a narrow molecular weight distribution and high homogeneity (Fig. 1D). Moreover, gas chromatography–mass spectrometry (GC-MS) analysis revealed the monosaccharide composition of PCP2. As shown in Fig. 1E, the carbohydrate portion of PCP2 consisted of 92.33% fructose and 7.67% glucose.

Structural analysis of PCP2

Glycosidic linkages between monosaccharide units in PCP2 were characterized by a methylation analysis, and the types of glycosidic bonds were further validated through a GC–MS analysis of fragment ions. As presented in Table 1, PCP2 consisted of the furanose form of fructose and the pyranose form of glucose, which aligned with the IR spectroscopy results. The monosaccharide composition of PCP2 included t-Fruf, t-Glcp, 1,2-Fruf, 2,6-Fruf, 1,6-Glcp, and 1,2,6-Fruf at molar ratios of 24.231:1.627:31.908:19.635:4.583:18.196. This finding was largely consistent with independently determined monosaccharide composition data.

Table. 1.

Partially methylated alditol acetates (PMAAs) and inferred glycosidic linkages of PCP2

Type of linkage	Methylated sugar	RT (min)	Molar ratio	Mass fragments (m/z)
2,5-di-O-acetyl-1,3,4,6-tetra-O-methyl mannitol/glucitol	t-Fruf	15.39 15.606	24.231	43, 59, 75, 87, 101, 113, 120, 130, 145, 162, 186, 205
1,5-di-O-acetyl-2,3,4,6-tetra-O-methyl glucitol	t-Glcp	16.749	1.627	43.1, 59, 71, 81, 87, 102, 113, 118, 129, 145, 157, 162, 174, 205
1,2,5-tri-O-acetyl-3,4,6-tri-O-methyl mannitol/glucitol	1,2-Fruf	19.469 19.579	31.908	43, 71, 81, 87, 100, 113, 129, 145, 161, 174, 190, 205, 234
2,5,6-tri-O-acetyl-1,3,4-tri-O-methyl mannitol/glucitol	2,6-Fruf	19.294 19.579	19.635	43, 59, 72, 87, 99, 113, 120, 129, 146, 162, 173, 189, 206, 233
1,5,6-tri-O-acetyl-2,3,4-tri-O-methyl glucitol	1,6-Glcp	20.475	4.583	43, 59, 71, 87, 102, 113, 118, 129, 143, 162, 173, 189, 233
1,2,5,6-tetra-O-acetyl-3,4-di-O-methyl-hexitol mannitol/glucitol	1,2,6-Fruf	23.393	18.196	43, 71, 87, 100, 113, 125, 129, 144, 159, 173, 190, 233

The structural characteristics of PCP2 were further elucidated by nuclear magnetic resonance (NMR) spectroscopy. ¹H NMR signals (Fig. 2A) mainly accumulated from 3.00 to 5.50 ppm, with the chemical shifts of the anomeric protons mostly observed in the 4.50–5.50 ppm range. Accordingly, the ¹³C NMR signals (Fig. 2B) were primarily found in the 60.0–100.0 ppm range, with the anomeric carbon chemical shifts located between 90 and 110 ppm. The α/β configuration of the residues was established based on the chemical shifts and coupling constants of their anomeric protons. By thoroughly examining the 1D and 2D NMR spectra of PCP2, along with comparisons to reported data, the ¹H and ¹³C NMR resonances of residues A, B, C, D, and E in PCP2 were assessed, as tabulated in Table 2.

Fig. 2. Structural analysis of PCP2.

A ¹H spectrum. B ¹³C spectrum. C HSQC spectrum. D ¹H–¹H COSY analysis. E HMBC analysis. F Structural prediction of PCP2.

Table. 2.

¹H and ¹³C NMR chemical shifts of PCP2 glycosidic bonds

Glycosyl residue		Chemical shift δ (ppm)
		1	2	3	4	5	6
A t-Fruf	H C	3.66 62.5	103.2	4.10 76.6	4.02 74.5	3.84 81.3	3.69 63.7
B 1,2-Fruf	H C	3.58 60.4	103.6	4.15 76.9	4.01 74.4	3.78 81.2	3.65 62.2
C 2,6-Fruf	H C	3.71 61.9	102.9	4.20 77.3	3.97 76.6	3.84 80.1	3.74 64.5
D 1,2,6-Fruf	H C	3.69 63.3	103.8	4.24 80.8	4.13 78.6	3.89 82.2	3.79 63.4
E 1,6-Glcp	H C	5.32 92.1	3.47 71.3	3.67 74.7	3.40 69.2	3.75 71.6	3.64 64.5

In the ¹H–¹H COSY spectrum (Fig. 2D), a clear correlation between δ_H 5.32 (H-1) and δ_H 3.47 enabled the assignment of δ_H 3.47 to the H-2 of residue E. Likewise, the resonances of H-3 through H-6 of residue E were attributed by the same correlation approach. Due to the low content of 1,6-Glcp, its anomeric carbon signal could not be detected in the ¹³C NMR spectrum. However, in the HSQC spectrum (Fig. 2C), the correlation between δ_H 5.32 (H-1) and δ_C 92.1 undoubtedly indicated that δ_C 92.1 should be assigned to C-1 of residue E. Then, according to the H-2 through H-6 signals identified in the ¹H–¹H COSY spectral data, the resonances of C-2 through C-6 of E were further determined based on the corresponding HSQC correlations. Finally, the anomeric configuration of residue E was assigned as α, as its anomeric carbon chemical shift was less than 100 ppm, which is consistent with the typical range for α-glycosidic bonds¹⁸. Thus, residue E was determined to be →1)-α-D-Glcp-(6 → . In the HSQC spectrum, no cross-peak signals of residues A, B, or C anomeric protons or anomeric carbons were observed. This was because fructose, as a ketose, lacks the anomeric protons necessary for heteronuclear correlation signals. Instead, through methylation analysis and comparisons with previous data, the carbon signals at δ_C 103.8, 103.6, 103.2, and 102.9 ppm were assigned to →1,6)-β-D-Fruf-(2 → , →1)-β-D-Fruf-(2 → , β-D-Fruf-(2 → , and →6)-β-D-Fruf-(2 → , respectively. By combining data from ¹H–¹H COSY and HSQC spectra with previously reported data, the resonances for H-1–H-6 and C-1–C-6 of residues A, B, C, and D were determined.

HMBC spectral data (Fig. 2E) indicated cross peaks between diverse residues, indicating glycosidic linkages and residue sequences. The key correlation from H-6 of residue D at δ_H 3.79 ppm to C-2 of residue C at δ_C 102.9 ppm demonstrated that C-2 of residue C was linked to C-6 of residue D. Similarly, the cross peaks in the HMBC spectrum between δ_H 3.74 ppm (H-6) of residue C and 103.2 ppm (C-2) of residue A, δ_H 3.69 ppm (H-1) of residue D and 103.6 ppm (C-2) of residue B, δ_H 3.64 ppm (H-6) of residue E and 103.6 ppm (C-2) of residue B, δ_H 3.58 ppm (H-1) of residue B and 103.2 ppm (C-2) of residue A, and δ_H 3.58 ppm (H-1) of residue B and 103.8 ppm (C-2) of residue D suggested that C-2 of residue A was linked to C-6 of residue C, C-1 of residue D was linked to C-2 of residue B, C-2 of residue B was linked to C-6 of residue E, C-2 of residue A was linked to C-1 of residue B, and C-2 of residue D was linked to C-1 of residue B.

Accordingly, the structure of PCP2 was determined to be a branched fructan. Its main chain consisted of →1)-β-D-Fruf-(2 → , →1,6)-β-D-Fruf-(2 → , β-D-Fruf-(2 → , and →6)-α-D-Glcp-(1→ residues, with the →1)-β-D-Fruf-(2→ linkage being the most prevalent. The branching units, identified as terminal β-D-Fruf-(2→ and →2)-β-D-Fruf-(6 → , were attached at the C-6 position of the →1,6)-β-D-Fruf-(2→ residues in the backbone. The inferred possible structure of PCP2 is shown in Fig. 2F.

Physicochemical properties of PCP2

The surface morphology of PCP2 was investigated using scanning electron microscopy (SEM) and atomic force microscopy (AFM). SEM imaging revealed that PCP2 possesses a distinct columnar stacking structure, composed of densely packed particles with prominent layering and vertical extensions (Fig. 3A). The surface appeared rough and was adorned with numerous small particulate attachments, a morphology potentially attributable to its highly branched architecture. Complementary nanoscale analysis by AFM further corroborated the granular and layered ultrastructure, showing a porous and spiked surface (Fig. 3B), which suggests strong intermolecular interactions and aggregation behavior consistent with a highly branched polysaccharide.

Fig. 3. PCP2 exists in a non-crystalline state and exhibits excellent thermal stability.

A SEM images of PCP2. B AFM images of PCP2. C XRD analysis of PCP2. D Thermogravimetric analysis of PCP2.

The conformational and structural properties of PCP2 were further examined. A Congo red assay confirmed the presence of a triple-helix conformation, as indicated by a characteristic redshift in the maximum absorption wavelength with increasing NaOH concentration (Fig. S1B, Supporting Information). X-ray diffraction (XRD) analysis showed a rapid initial rise in intensity followed by a gradual decline, with no sharp diffraction peaks detectable, confirming the amorphous nature of PCP2 (Fig. 3C).

Thermal stability was assessed by thermogravimetric analysis (TGA). PCP2 exhibited a three-stage decomposition pattern between 30–800 °C (Fig. 3D). The first stage (82–195 °C) involved gradual weight loss, attributed to the evaporation of free and bound water. A major decomposition phase occurred in the second stage (195–387 °C), marked by a sharp decline in weight and a distinct peak in the derivative thermogravimetric (DTG) curve, corresponding to the breakdown of polysaccharide chains and organic components. Above 400 °C, weight loss slowed significantly, indicating sample carbonization and the formation of thermally stable residues. Collectively, these results demonstrate that PCP2 maintains good thermal stability below 200 °C.

PCP2 alleviates intestinal damage caused by DSS in mice

To evaluate the therapeutic potential of PCP2 against UC, a mouse model of colitis was established using DSS, and the effects of different PCP2 doses were assessed (Fig. 4A). DSS challenge induced typical colitis symptoms, including progressive body weight loss (Fig. 4B) and elevated disease activity index (DAI) scores (Fig. 4C). PCP2 treatment dose-dependently ameliorated these changes, mitigating weight loss and reducing DAI scores. Consistent with these findings, colon length, a key indicator of colitis severity, was significantly shortened in DSS-treated mice but restored by PCP2 administration (Fig. 4D, E).

Fig. 4. PCP2 alleviates intestinal damage induced by DSS in mice.

A Experimental procedure and experimental groupings. This figure was created with BioRender.com. B Body weight changes (n = 10). C DAI score (n = 10). D Images of mouse colons. E Colonic length of mice. F Colonic tissue with H&E, PAS, and PSR staining (Arrows highlight the key pathological features, including inflammatory cell infiltration, crypt loss and mucosal damage. Contiguous sections were used for the different staining procedures). G Histological scoring of H&E-stained colon slices. H PAS staining area quantification. I Analysis of ZO-1, Occludin, and Claudin-1 protein levels in colonic tight junctions. J Protein levels of ZO-1. K Protein levels of Occludin. L Protein levels of Claudin-1. Data are presented as mean ± SEM. The symbol “^#” indicates a significant difference between the control and DSS model groups, and “*” indicates a significant difference between the DSS model and treatment groups. (Fig. 4A was created with BioRender.com).

Histopathological evaluation further confirmed the protective effect of PCP2. Hematoxylin and eosin (H&E) staining revealed extensive mucosal damage, crypt loss, and inflammatory infiltration in the DSS group, whereas PCP2 markedly alleviated these pathological alterations (Fig. 4F). Periodic acid–Schiff (PAS) staining showed a pronounced reduction in goblet cells and mucin secretion in DSS-treated mice, while PCP2 substantially restored both parameters. Additionally, Picrosirius Red (PSR) staining was used to assess early fibrotic changes, which revealed a significant increase in submucosal collagen deposition in the DSS group. This early-stage collagen deposition was markedly attenuated by PCP2 treatment. A quantitative assessment confirmed the reduction of histological injury (Fig. 4G) and a significant increase in the PAS-positive area in the PCP2-treated groups (Fig. 4H).

Given the critical role of epithelial barrier dysfunction in colitis pathogenesis¹⁹, we next examined the expression of key tight junction proteins. Western blot analysis demonstrated that DSS treatment markedly decreased the levels of ZO-1, Occludin, and Claudin-1, whereas PCP2 administration effectively restored their expression (Fig. 4I–L). Taken together, these results indicated that PCP2 exerted a protective effect against DSS-induced colitis by alleviating clinical symptoms, mitigating histological damage, restoring goblet cell function, preventing collagen deposition, and enhancing intestinal barrier integrity.

PCP2 exerts multifaceted protective effects against DSS-induced colitis

Impairment of the mucus barrier is a hallmark of ulcerative colitis, we therefore evaluated the expression of MUC2, the primary mucin secreted in the colon. Immunofluorescence staining showed that the fluorescence intensity of MUC2 was significantly lower in the DSS group than in the control group, while 5-ASA and PCP2 treatment restored MUC2 levels (Fig. S2A, C, Supporting Information). Consistently, immunohistochemical staining revealed that MUC2 levels were significantly reduced in the DSS group, and PCP2 treatment significantly increased the integrated optical density of MUC2 (Fig. S2B, D, Supporting Information). These findings indicated that PCP2 protected against DSS-induced colitis by enhancing MUC2 production, thereby preserving the integrity of the intestinal mucus barrier.

Inflammation and oxidative stress are the key factors contributing to colitis²⁰. To investigate the effects of PCP2 on inflammation and oxidative stress, we next evaluated cytokine and redox homeostasis in serum and colonic tissue. DSS challenge significantly elevated pro-inflammatory cytokines (IL-6 and TNF-α) and suppressed the anti-inflammatory cytokine IL-10 in both serum and colon tissues. PCP2 treatment reversed these alterations in a dose-dependent manner (Fig. S3A–C, S3H–J, Supporting Information). In addition, DSS exposure markedly decreased serum antioxidant indices, including levels of glutathione (GSH), glutathione peroxidase (GSH-Px), and superoxide dismutase (SOD), while increasing the lipid peroxidation product malondialdehyde (MDA). PCP2 administration significantly restored GSH, GSH-PX, and SOD levels and reduced MDA accumulation (Fig. S3D–G, Supporting Information). These results indicated that PCP2 alleviated intestinal inflammation by modulating pro- and anti-inflammatory cytokines, as well as mitigated oxidative stress, thereby contributing to its protective effects against colitis.

Uncontrolled tissue remodeling and the pathological accumulation of extracellular matrix proteins are key drivers of long-term intestinal damage in UC. To determine if PCP2 could mitigate the initial stages of this process, we examined key protein markers. Western blot analysis demonstrated that PCP2 significantly downregulated the expression of α-smooth muscle actin (α-SMA), a marker of myofibroblast activation, as well as Collagen-I and the proliferation-related protein Cyclin D1 (Fig. S3K–N). These results suggested that PCP2 suppressed early extracellular matrix remodeling and profibrotic signaling, thereby limiting the progression of tissue scarring. In summary, PCP2 alleviated experimental colitis by restoring mucin production, rebalancing inflammatory and oxidative responses, and attenuating early profibrotic activation in the colon.

PCP2 alleviates DSS-induced disruptions of the gut microbial composition

Studies have shown that polysaccharides are crucial for regulating the host physiological condition and reshaping the gut microbiota (GM), and an imbalance of the GM is related to UC²¹. To explore whether and how PCP2 affected the composition or abundance of the GM, 16S rRNA sequencing was performed to assess the gut microbiome of the colitis model mice. The findings revealed that compared to the DSS group, the ACE and Shannon indices of the PCP2 group were notably higher (Fig. 5A, B), indicating that PCP2 affected the diversity and richness of GM in colitis mice. Beta diversity analysis assesses the differences in species composition between different environments or groups and is an important indicator in microbial community profiling. Principal Co-ordinates Analysis (PCoA) or Non-metric Multidimensional Scaling (NMDS) was used to visualize the differences between samples. Based on the PCoA (Fig. 5C) and NMDS (Fig. 5D) results, the DSS and control groups showed distinct microbial communities, while the sample points of the PCP2 group were closer to those of the control group. This suggested that the composition of GM in the PCP2 group was relatively comparable to that of the control group.

Fig. 5. PCP2 improves the GM imbalance in DSS-induced UC mice.

A ACE index. B Shannon index. C PCoA analysis. D NMDS analysis. E Taxonomic distributions of the GM composition at the phylum level. F Heatmap of the GM composition at the genus level. G Relative abundances of A. muciniphila. H Relative abundance of Lachnospiraceae. I LEfSe cladogram from phylum to genus levels. Data are presented as mean ± SEM. The symbol “^#” indicates a significant difference between the control and DSS model groups, and “*” indicates a significant difference between the DSS model and treatment groups.

The taxonomic analysis revealed that PCP2 fundamentally restructured the GM, most notably by promoting the keystone species A. muciniphila. At the phylum level, this was reflected by a significant enrichment of its corresponding phylum, Verrucomicrobiota, in the PCP2 group compared to the DSS group (Fig. 5E). At the genus level, A. muciniphila was identified as one of the most dramatically enriched taxa following PCP2 intervention (Fig. 5F, G). This finding was particularly significant and provided a direct mechanistic link to our earlier observation that PCP2 restored MUC2 expression (Fig. S2, Supporting Information). A. muciniphila is a specialized mucin-degrading bacterium known to reside in the mucus layer, where it stimulates goblet cells to produce and secrete mucins, thereby reinforcing the integrity of the intestinal mucus barrier—a critical defense mechanism compromised in UC^9,22. In addition to fostering the growth of this key barrier-promoting symbiont, PCP2 also suppressed pathobionts and cultivated a community of beneficial SCFA-producing bacteria. The abundances of pro-inflammatory genera, including Desulfovibrio and Clostridium, which were markedly elevated by DSS, were significantly reduced by PCP2 treatment (Fig. 5F). Concurrently, PCP2 administration led to the enrichment of several well-known beneficial, butyrate-producing genera, including Lachnospiraceae, Eubacterium, and Alistipes (Fig. 5H). A Linear discriminant analysis Effect Size (LEfSe) analysis further confirmed these trends, identifying pathogenic Clostridia as a key biomarker of the DSS group, while the PCP2 group was characterized by an enrichment of SCFA-producing taxa such as Bacteroides (Fig. 5I).

Collectively, these findings suggested that PCP2 ameliorated colitis by broadly shifting the microbial composition, and by specifically fostering the bloom of the mucus-protective specialist A. muciniphila while simultaneously suppressing pathogens and promoting a beneficial, fermentative microbial community.

PCP2 ameliorates colitis via a transferable GM-dependent mechanism

To determine whether the protective effects of PCP2 depend on gut microbiota (GM) remodeling, fecal microbiota transplantation (FMT) was performed using donor feces from the DSS and PCP2-treated groups (Fig. 6A). Mice receiving PCP2-modulated microbiota (FMT-PCP2) showed significant mitigation of DSS-induced colitis, as reflected by attenuated weight loss, lower disease activity index (DAI) scores, and less colon shortening compared to the FMT-DSS group (Fig. 6B–E). Histological evaluation further confirmed that FMT-PCP2 alleviated inflammatory infiltration, restored mucin production (PAS staining), and reduced early collagen deposition (PSR staining) (Fig. 6F). Consistently, the FMT-PCP2 group exhibited lower levels of pro-inflammatory cytokines (IL-6, TNF-α), elevated IL-10, and enhanced expression of tight junction proteins (ZO-1, Claudin-1, Occludin) (Fig. 6G–J).

Fig. 6. PCP2 relies on the GM to alleviate intestinal damage.

A Experimental procedure and groupings. This figure was created with BioRender.com. B Body weight changes (n = 8). C DAI scores (n = 8). D Images of mouse colons. E Colon length of mice. F Colonic tissue H&E, PAS, and PSR staining (Arrows highlight the key pathological features, including inflammatory cell infiltration, crypt loss and mucosal damage. Contiguous sections were used for the different staining procedures). G Serum IL-6 levels (n = 8). H Serum TNF-α levels (n = 8). I Serum IL-10 levels (n = 8). J Fluorescent staining of mouse colonic tissue for Occludin, ZO-1, and Claudin-1. K Heatmap of the community composition at the phylum level. Data are presented as mean ± SEM. The symbol “*” indicates a significant difference between the DSS + FMT and PCP2 + FMT groups. (Fig. 6A was created with BioRender.com).

Microbiome analysis confirmed the successful transfer of a protective microbial community. The FMT-PCP2 group displayed restored microbial α- and β-diversity (Fig. S5A–C) and a normalized Firmicutes-to-Bacteroidota ratio (Fig. 6K). At the genus level, beneficial taxa such as Akkermansia, Lachnospiraceae, and Eubacterium were enriched, while harmful genera including Desulfovibrio and Clostridium were suppressed (Fig. S5D–H). LEfSe analysis identified the phylum Bacteroidota as a key biomarker of the FMT-PCP2 group (Fig. S5I). Importantly, the specific enrichment of Akkermansia and Bacteroides, along with the suppression of Desulfovibrio, was fully recapitulated in recipient mice, underscoring that a functionally active, PCP2-shaped microbiota is sufficient to convey anti-colitis effects.

To further link microbial changes to host physiology, we analyzed correlations between bacterial genera and host inflammatory/oxidative markers (Fig. S4A). Several PCP2-enriched genera were strongly associated with improved pathological parameters. In addition, consistent with the increase in SCFA-producing bacteria, high-dose PCP2 robustly restored colonic levels of butyric acid, acetic acid, and other SCFAs, which were significantly reduced in DSS-treated mice (Fig. S4B–E). These data collectively indicated that PCP2 remodeled the gut microbiota into an anti-inflammatory community with enhanced SCFA production, thereby playing a functional and transferable role in alleviating colitis.

PCP2 confers protection against colitis through GM-independent pathways

Having established that PCP2-modulated microbiota is sufficient to confer protection, we further investigated whether its efficacy depends entirely on the GM. To this end, a broad-spectrum antibiotic cocktail was administered to deplete the GM in C57BL/6 J mice (Fig. 7A). Quantitative fecal DNA analysis confirmed a pronounced reduction in bacterial load following antibiotic treatment (Fig. 7B). Under these GM-depleted conditions, PCP2 still significantly attenuated DSS-induced body weight loss, improved disease activity index (DAI) scores, and ameliorated colon shortening (Fig. 7C–F). Histological evaluation via H&E, PAS, and PSR staining further revealed that PCP2 alleviated inflammatory infiltration, restored mucin production, and reduced early collagen deposition even in the absence of an intact microbiota (Fig. 7G). Moreover, PCP2 treatment suppressed pro-inflammatory cytokines (IL-6, TNF-α) and elevated the anti-inflammatory cytokine IL-10 under these conditions (Fig. 7H–J). Collectively, although antibiotic-mediated GM depletion partially diminished the overall efficacy of PCP2, the polysaccharide retained a significant degree of protective activity. These results indicated that PCP2 ameliorated colitis through both GM-dependent and GM-independent mechanisms.

Fig. 7. The protective mechanism of PCP2 is not entirely dependent on the GM.

A Experimental procedure and groupings. This figure was created with BioRender.com. B The DNA content in feces (n = 8). C Body weight changes (n = 8). D DAI scores (n = 8). E Images of mouse colons. F Colon length of mice. G Colonic tissue H&E, PAS, and PSR staining (Arrows highlight the key pathological features, including inflammatory cell infiltration, crypt loss and mucosal damage. Contiguous sections were used for the different staining procedures). H Serum IL-6 levels (n = 8). I Serum TNF-α levels (n = 8). J Serum IL-10 levels (n = 8). Abx, antibiotics. The symbol “^#” indicates a significant difference between the DSS+Abx+PCP2 and DSS+Abx groups, and “*” indicates a significant difference between the DSS+Abx+PCP2 and DSS + PCP2 groups. (Fig. 7A was created with BioRender.com).

PCP2 directly targets follistatin (Fst) and inhibits the BMP4/Smad1/ID1 signaling pathway to ameliorate intestinal injury in DSS-induced colitis

To further investigate its underlying mechanism, we performed an Olink proteomics analysis. The results demonstrated that PCP2 alleviated intestinal injury by downregulating inflammatory factors, including TNF, IL-17, and chemokines, while upregulating repair-related factors, including Fst, Fas, Hgf, and Riox2 (Fig. 8A, B). Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis revealed that PCP2 primarily exerted its protective effects by inhibiting inflammatory signaling pathways and activating tissue repair and metabolism-associated pathways (Fig. 8C).

Fig. 8. PCP2 exerts protective effects in a DSS-stimulated intestinal damage model by regulating Fst levels.

A Heatmap of differentially expressed proteins. B Box plot of the levels of significantly differentially expressed proteins in each group. C KEGG analysis. D The binding region and binding diagram of PCP2 and Fst. E The 50-ns RMSD of the Fst and PCP2 complex. F The RMSF of amino acid residues in the PCP2–Fst complex. G The number of hydrogen bonds produced by the PCP2–Fst complex within 50 ns. H MST analysis of PCP2 binding to Fst. I SPR analysis of PCP2 binding to Fst. J Steady-state fitting of PCP2 and Fst. K Analysis of BMP4, p-Smad1, Smad1, and ID1 protein levels in colonic tissue. L Relative protein levels of BMP4. M Ratio of p-Smad1/Smad1 levels. N Relative protein levels of ID1. The symbol “^#” indicates a significant difference between the control and DSS model groups, and “*” indicates a significant difference between the DSS model and treatment groups.

To identify the most promising target from the upregulated repair-related factors, we performed molecular docking. The docking results indicated that the binding energy between PCP2 and Fst was −6.2 kcal/mol, lower (more favorable) than those for the other candidate proteins (Fas, Hgf, and Riox2). Therefore, we proposed that Fst is a potential target. (Fig. S6, Fig. S7A–D, Supporting Information). To further characterize this interaction, molecular dynamics (MD) simulations were performed. These simulations indicated that PCP2 forms stable hydrogen bonds with residues Asn124, Asp121, Leu146, Cys150, Gly103, and Pro102 in Fst, with bond lengths ranging from 3.1 to 4.0 Å. The overall binding energy was calculated to be −22.40 ± 5.58 kcal/mol (Fig. 8D). RMSD analysis showed that the PCP2–Fst complex reached a dynamic equilibrium after initial structural adjustments (Fig. 8E), while RMSF analysis confirmed that fluctuating residues were located outside the binding pocket and did not compromise binding stability (Fig. 8F). Hydrogen bond analysis further identified Gly103 as a key residue contributing to complex stability, maintaining interactions with a frequency of 19.58% during the simulation (Fig. 8G).

The PCP2–Fst interaction was further validated experimentally by microscale thermophoresis (MST) and surface plasmon resonance (SPR), which confirmed direct and dose-dependent binding (Fig. 8H–J). Given that Fst is a well-established endogenous antagonist of bone morphogenetic proteins (BMPs)²³, and considering prior evidence linking BMP4/Smad1/ID1 signaling to intestinal inflammation and impaired epithelial repair^13,24,25, we hypothesized that PCP2 alleviates colitis by binding Fst and consequently inhibiting BMP4 pathway activation. Western blot analysis confirmed that PCP2 treatment indeed suppressed BMP4/Smad1/ID1 signaling in DSS-challenged mice (Fig. 8K–N). In summary, these integrated findings demonstrated that PCP2 ameliorated intestinal inflammation and injury in colitis by directly targeting Fst and inhibiting the BMP4/Smad1/ID1 signaling pathway.

PCP2 exerts its therapeutic effects in an Fst-dependent manner

To establish whether the therapeutic efficacy of PCP2 functionally depends on Fst, we performed siRNA-mediated knockdown of Fst in Caco-2 cells. A cellular thermal shift assay (CETSA) confirmed the direct binding of PCP2 to Fst, as evidenced by a significant increase in Fst thermal stability upon PCP2 treatment (Fig. 9A, B). Following efficient Fst depletion (Fig. 9C, D), PCP2 failed to suppress the BMP4/Smad1/ID1 signaling pathway (Fig. 9E–H), indicating that Fst was required for PCP2-mediated inhibition of this pathway in vitro.

Fig. 9. Functional validation of Fst as a direct target of PCP2.

A Cellular thermal shift assay of Fst protein. B Intensity related to the cellular thermal shift assay of the Fst protein. C Western blot analysis of Fst protein levels. D Protein levels of Fst. E Analysis of BMP4, p-Smad1, Smad1, and ID1 protein levels in colonic tissue. F Relative protein levels of BMP4. G Ratio of p-Smad1/Smad1 levels. H Relative protein levels of ID1. I Schematic diagram of the in vivo adenovirus-mediated Fst knockdown experimental design. This figure was created with BioRender.com. J Images of mouse colons. K Colon length of mice. L Body weight changes (n = 10). M DAI scores (n = 10). N Serum TNF-α levels. O Serum IL-6 levels. P Serum IL-10 levels. Q Western blot analysis of Fst protein levels. R Protein levels of Fst. S Colonic tissue H&E, PAS, and PSR staining (Arrows highlight the key pathological features, including inflammatory cell infiltration, crypt loss and mucosal damage. Contiguous sections were used for the different staining procedures). *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001. (Fig. 9I was created with BioRender.com).

To verify this dependency in vivo, Fst was specifically knocked down in the mouse colon using an adenovirus-based approach (Fig. 9I). Western blot analysis confirmed successful reduction of Fst protein in colonic tissue (Fig. 9Q, R). Under these Fst-deficient conditions, the protective effects of PCP2 were markedly attenuated. Although partial mitigation of DSS-induced colon shortening was still observed (Fig. 9J, K), PCP2’s ability to ameliorate body weight loss (Fig. 9L), disease activity index (DAI) scores (Fig. 9M), pro-inflammatory cytokine dysregulation (Fig. 9N–P), and impaired mucosal repair (Fig. 9S) was substantially weakened compared to that in mice with normal Fst expression.

In summary, these genetic loss-of-function studies demonstrate that Fst was essential for PCP2 to exert its full therapeutic benefit. Fst targeting constitutes a key mechanistic pathway through which PCP2 acts, operating in concert with gut microbiota remodeling to alleviate intestinal inflammation and injury.

Discussion

The therapeutic landscape of UC is shifting towards interventions that can simultaneously restore the disrupted gut ecosystem and directly modulate host mucosal immunity. While plant-derived polysaccharides are widely recognized as potent prebiotics, their mechanisms are predominantly viewed through the lens of “microbiota-dependent” effects, often overlooking their potential to directly engage host signaling networks^9,26. This study expands upon the conventional microbiota-centric paradigm by elucidating a dual-pronged therapeutic mechanism for PCP2, a novel low-molecular-weight fructan derived from Polygonatum cyrtonema Hua. We provide evidence that PCP2 functions not only as an ecological modulator that promotes the proliferation of beneficial symbionts but also as a specific molecular ligand for the host protein Fst, thereby orchestrating a synergistic microbiota-host defense against colitis. In contrast to the prevalent view that polysaccharide actions are primarily mediated by microbial fermentation or non-specific interactions, our antibiotic depletion experiments revealed a critical divergence: PCP2 retained significant protective efficacy even under conditions of severe gut microbiota suppression, implying the existence of an additional, host-targeted mechanism. Indeed, we validated Fst as a direct binding partner of PCP2, demonstrating that this interaction antagonizes the pro-inflammatory BMP4/Smad1/ID1 signaling axis. The functional necessity of this axis was further confirmed by the attenuation of PCP2’s efficacy in Fst-deficient mice. Collectively, these findings suggest that specific polysaccharides such as PCP2 can act as macromolecular agents with distinct bioactivity, functioning in concert with their prebiotic effects.

Gut microbiota dysbiosis is a well-established trigger in UC pathogenesis²⁷. For instance, relevant studies demonstrated that patients with UC exhibit a significant reduction in GM diversity. Specifically, these patients experienced a marked decrease in beneficial bacteria, such as those belonging to the Firmicutes and Bacteroidetes phyla, while the abundance of potentially pathogenic bacteria from the Proteobacteria and Actinobacteria phyla notably increased. This dysbiosis of the GM compromised the integrity of the gut barrier, disrupting immune homeostasis and ultimately eliciting chronic inflammatory responses²⁸. In this context, as a potential prebiotic, PCP2 fundamentally reshapes the dysbiotic gut microbiota, most notably by promoting the bloom of Akkermansia muciniphila, which is a key mucin-degrading symbiont that stimulates goblet cells to secrete MUC2, establishing a positive feedback loop that thickens the mucus layer. This mechanism directly aligns seamlessly with our observed restoration of MUC2 expression and goblet cell counts. Moreover, the enrichment of SCFA producers such as Bacteroides and Lachnospiraceae, along with the subsequent elevation of butyrate levels, provides a sustained energy source for colonocytes. Crucially, FMT experiments demonstrated that PCP2 exerted its effects by regulating the intestinal microbiota. However, subsequent antibiotic exposure (Abx) experiments revealed that while PCP2 alleviated intestinal damage, its action did not entirely depend on the intestinal microbiota. Therefore, PCP2 acts both as an effective microecological regulator by improving the intestinal environment through microbiota optimization, and as a direct ligand that targets the host protein Fst independently of the microbiota (a pathway we later identified).

Fst, a glycoprotein significantly upregulated during the acute phase of colitis, is mainly produced by intestinal epithelial cells, macrophages, and endothelial cells¹². As a natural antagonist of the TGF-β superfamily, Fst binds and neutralizes hormone-like ligands. Research has demonstrated that pretreatment with Fst enhanced the survival rate of mice from 33% to 82%, while reducing plasma levels of IL-6 and amyloid proteins. Furthermore, it markedly alleviated tissue damage and myeloperoxidase (MPO) activity in 2,4,6-trinitrobenzenesulfonic acid and DSS-induced and IL-10 knockout models by promoting the proliferation of colonic epithelial cells^12,29. The role of bone morphogenetic protein 4 (BMP4) in UC is context-dependent. While it can exhibit homeostatic functions, its over-activation is pro-inflammatory and is antagonized by Fst, a well-established endogenous inhibitor of BMP signaling³⁰. In cases of tissue damage or disease, over-activated BMP4 becomes a powerful pro-inflammatory signal that attracts immune cells, intensifies the inflammatory response³⁰, and can promote fibrogenic responses including fibroblast activation, collagen deposition, and extracellular matrix synthesis²⁴. As an exogenous antagonist of BMP4, Fst was shown in this study to bind BMP4 and inhibit its activation of the downstream Smad signaling pathway, thus achieving a balance between promoting tissue repair and reducing inflammation¹³. While the protective role of the Fst/BMP4 axis in colitis is increasingly recognized, specific therapeutic agents that can directly engage and potentiate Fst activity are still lacking. Here, we identify the natural polysaccharide PCP2 as a novel direct binding partner of Fst. An Olink proteomics screen initially identified Fst as a key repair-related protein upregulated by PCP2. We subsequently validated the direct physical interaction between PCP2 and Fst, demonstrating that PCP2 forms stable hydrogen bonds with specific residues within the binding pocket, notably Glycine 103. The physiological requirement for this interaction was definitively established by loss-of-function experiments, where the protective effects of PCP2 were significantly compromised in Fst-deficient models, confirming Fst as an essential mediator of the observed anti-colitis effects.

This study provides compelling evidence for a “microbiota–target” dual mechanism through which PCP2 alleviates colitis. However, certain limitations should be acknowledged. Firstly, regarding mechanistic depth, while we established that PCP2 reprograms the gut microbiota (e.g., enriching Akkermansia muciniphila) and directly binds to Follistatin (Fst), key functional questions remain. It is unclear whether the protective effects of the modulated microbiota are mediated by specific metabolites beyond those identified, and how the PCP2-Fst interaction functionally alters Fst’s stability or activity. Secondly, concerning signal specificity, our focus was on the BMP4/Smad1/ID1 axis. Although Fst is a selective antagonist, we did not experimentally rule out potential effects on other TGF-β superfamily members (e.g., Activin A), leaving its selectivity within this family to be fully validated. Finally, regarding safety assessment, while PCP2 showed a favorable in vivo tolerability profile, as evidenced by its reversal of disease-related weight loss (Fig. 4) and the absence of macroscopic organ damage, a comprehensive toxicological evaluation is still lacking. Future studies must incorporate clinical chemistry, hematological analysis, and detailed histopathology to definitively establish its systemic safety window.

In conclusion, our integrated study, from structural elucidation to mechanistic dissection, establishes PCP2, a low-molecular-weight (Mw = 2087 Da) fructan from the medicinal-food plant Polygonatum cyrtonema Hua, as a potent agent for alleviating ulcerative colitis. Structurally, its main chain consists of →1)-β-D-Fruf-(2 → , →1,6)-β-D-Fruf-(2 → , β-D-Fruf-(2 → , and →6)-α-D-Glcp-(1→ residues, with the →1)-β-D-Fruf-(2→ linkage being the most prevalent. The branching units, identified as terminal β-D-Fruf-(2→ and →2)-β-D-Fruf-(6 → , are attached at the C-6 position of the →1,6)-β-D-Fruf-(2→ residues in the backbone. Mechanistically, a dual-pronged strategy was delineated wherein PCP2 not only restores intestinal homeostasis via a microbiota-dependent pathway through enrichment of beneficial bacteria such as Akkermansia muciniphila and suppression of pathobionts, but also directly targets the host protein Fst to inhibit the pro-inflammatory BMP4/Smad1/ID1 signaling axis. This conclusion is conclusively supported by proteomic, biophysical, and genetic evidence. The dual capacity to simultaneously target both microbial dysbiosis and host inflammatory pathways positions PCP2 as a promising therapeutic candidate for ulcerative colitis. Collectively, our work not only characterizes a promising anti-colitis candidate but also provides unprecedented evidence for a direct interaction between a plant-derived polysaccharide and a specific host protein, opening new avenues for understanding the mechanism of bioactive polysaccharides.

Methods

Extraction and purification

Fresh Polygonatum cyrtonema Hua rhizomes were dried and pulverized into a fine powder. The powder was extracted three times with deionized water at 90 °C for 2 h each. The combined extracts were concentrated using a rotary evaporator (N-1100, Eyela, Shanghai, China). Subsequently, four volumes of absolute ethanol were added to the concentrate, and the mixture was incubated at 4 °C for 12 h to precipitate polysaccharides. The precipitate was recovered by centrifugation at 8000 rpm for 10 min using a high-speed refrigerated centrifuge (3K30, Sigma Laborzentrifugen GmbH, Osterode am Harz, Germany). After redissolving the precipitate in deionized water, proteins and pigments were removed using papain and D101 macroporous resin (Solarbio, Beijing, China), respectively. The solution was then dialyzed against deionized water for 48 h using a 1000 Da molecular weight cutoff dialysis bag (Solarbio). Finally, the retentate was lyophilized using a vacuum freeze dryer (FreeZone, Labconco, Kansas City, MO, USA) to yield crude polysaccharides (PCPs). The crude PCPs were redissolved in deionized water and purified sequentially via a DEAE-52 cellulose column (2.5 × 40 cm) and a Sephadex G-50 column (1.6 × 60 cm; Solarbio). The polysaccharide content in the eluates was monitored using the anthrone-sulfuric acid method³¹. The major fractions were pooled, dialyzed, and lyophilized to obtain the purified polysaccharide, designated as PCP2. Homogeneity analysis was performed using an HPGPC system (LC-10avp, Shimadzu, Kyoto, Japan) equipped with a TSKgel G3000PWXL column (7.8 × 300 mm, Tosoh Corp., Tokyo, Japan) and a refractive index detector (RID-10A, Shimadzu). The sample (2 mg/mL) was eluted with deionized water at a flow rate of 0.7 mL/min³².

Determination of molecular weight

Briefly, 5 mg of the sample was accurately weighed and dissolved in 1 mL of 0.05 M NaCl solution to obtain a final concentration of 5 mg/mL. The solution was centrifuged at 8000 rpm for 10 min using a high-speed refrigerated centrifuge (3K30, Sigma Laborzentrifugen GmbH, Osterode am Harz, Germany). The supernatant was subsequently filtered through a 0.22-μm microporous membrane, transferred to a 2-mL injection vial, and analyzed using a high-performance gel permeation chromatography (HPGPC) system (LC-10avp, Shimadzu, Kyoto, Japan) equipped with a tandem column³³.

Monosaccharide composition analysis

Briefly, an appropriate volume of the supernatant was concentrated using a rotary evaporator (N-1100, Eyela, Shanghai, China) and dried under a nitrogen stream. Subsequently, 5 mg (±0.05 mg) of the polysaccharide sample was accurately weighed and hydrolyzed with 1 mL of 2 M trifluoroacetic acid (TFA) at 60 °C for 1 h. The hydrolysate was evaporated to dryness under nitrogen. To ensure the complete removal of residual TFA, the residue was co-evaporated with methanol three to four times. The final residue was reconstituted in sterile deionized water and transferred to a chromatography vial. The analysis was performed using a Dionex ICS-5000+ system (Thermo Fisher Scientific, Waltham, MA, USA) equipped with a Dionex™ CarboPac™ PA20 column (150 × 3.0 mm, 10 µm). The mobile phases consisted of H₂O (A), 0.1 M NaOH (B), and 0.1 M NaOH containing 0.2 M NaAc (C). The injection volume was 5 µL, and the column temperature was maintained at 30 °C. Monosaccharide detection was achieved using a pulsed amperometric detector (PAD)³⁴.

UV and FT-IR analyses

The UV-vis absorption spectrum of PCP2 was acquired by scanning a solution of the polysaccharide in deionized water from 200 to 400 nm using a NanoDrop One ultra-micro spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). For FT-IR analysis, 1–2 mg of the dried sample was mixed with 200 mg of KBr powder, ground homogeneously, and pressed into a pellet. The infrared spectrum was subsequently recorded in the wavenumber range of 4000 to 400 cm⁻¹ using a Fourier transform infrared spectrometer (e.g., Nicolet iS10, Thermo Fisher Scientific, Waltham, MA, USA)³⁵.

NMR analysis

The PCP2 sample was dissolved in deuterium oxide (D₂O) to a final concentration of 20 mg/mL. One-dimensional (1D) and two-dimensional (2D) NMR spectra, including ¹H NMR, ¹³C NMR, COSY, NOESY, HMBC, and HSQC, were recorded using a Bruker Avance III 600 MHz NMR spectrometer (Bruker, Rheinstetten, Germany) equipped with a dual probe³⁶.

Methylation analysis

The PCP2 sample was dissolved in anhydrous DMSO and alkalized with NaOH. Methylation was achieved by adding iodomethane in two stages. The reaction mixture was partitioned between water and dichloromethane, after which the organic phase was collected and evaporated to dryness. The residue was hydrolyzed with TFA at 90 °C, dried under a nitrogen stream, and reduced with NaBD₄. The reaction was quenched with acetic acid, and borate was removed by repeated co-evaporation with methanol. Acetylation was performed using acetic anhydride at 100 °C, and the resulting partially methylated alditol acetates (PMAAs) were extracted with dichloromethane. Analysis was conducted on a GC-MS system (7890 A GC coupled to a 5977B mass spectrometer, Agilent Technologies, Santa Clara, CA, USA) equipped with an HP-5MS capillary column (30 m × 0.25 mm × 0.25 μm, Agilent J&W Scientific), using high-purity helium (≥99.999%) as the carrier gas. The column temperature was programmed as follows: 50 °C for 1 min, increased to 130 °C at 50 °C/min, then to 230 °C at 3 °C/min, and finally held at 230 °C for 2 min.

Congo red analysis

The helical conformation of PCP2 was investigated according to the method of Ogawa et al.³⁷. Briefly, the PCP2 solution (1 mg/mL) was combined with a Congo red solution (100 mM). Subsequently, NaOH was added to adjust the final alkali concentration to a range of 0.00–0.50 M. After equilibration, the maximum absorption wavelength (λ_max) was determined by scanning the spectrum from 200 to 600 nm using a NanoDrop One spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). Pure Congo red solutions at the corresponding NaOH concentrations served as controls.

SEM analysis

The surface morphology of PCP2 was characterized using a scanning electron microscope (SEM; Zeiss Merlin Compact, Carl Zeiss, Oberkochen, Germany). The sample was mounted onto a conductive stub and sputter-coated with a thin gold layer to enhance conductivity. Images were acquired under high vacuum conditions at an accelerating voltage of 1.0 kV, with magnifications of 100× and 4000×. (The analysis was performed by Sanshu Biotech).

AFM analysis

The PCP2 sample was dissolved in distilled water to a final concentration of 10 μg/mL. A 5-μL aliquot of the solution was deposited onto the surface of a freshly cleaved mica sheet and air-dried at room temperature. The surface topography was subsequently characterized using an atomic force microscope (Dimension Icon, Bruker, Germany).

DSS-induced colitis mouse model

Specific pathogen-free (SPF) male C57BL/6 J mice (6–8 weeks old, 18–22 g) were purchased from Zhejiang Vital River Laboratory Animal Technology Co., Ltd. (Zhejiang, China). Mice were housed in a temperature-controlled environment (22 ± 2 °C, 50% ± 10% humidity) under a 12-h light/dark cycle with ad libitum access to standard chow and water. All animal experiments were approved by the Animal Ethics Committee of Wannan Medical College (Approval No. WNMC-AWE-2024398) and conducted in strict accordance with the Guide for the Care and Use of Laboratory Animals. Following a 7-day acclimation period, mice were randomly divided into five groups (n = 10 per group): Control, Model (DSS), 5-ASA (positive control), PCP2-L (low dose), and PCP2-H (high dose). To induce acute colitis, mice in the experimental groups were administered drinking water containing 3% (w/v) dextran sulfate sodium (DSS; Mw 36–50 kDa, MP Biomedicals, Solon, OH, USA) for 11 consecutive days, while the control group received standard drinking water. From day 3 to day 10, treatments were administered daily by oral gavage (0.2 mL per mouse) as follows: the 5-ASA group received mesalazine (100 mg/kg; dissolved in saline), and the PCP2-L and PCP2-H groups received PCP2 at doses of 200 mg/kg and 400 mg/kg, respectively. The Control and Model groups were gavaged with an equivalent volume of sterile saline. Body weight, stool consistency, and gross bleeding were monitored daily. On day 11, all mice were euthanized by cervical dislocation under isoflurane anesthesia, and colon tissues were harvested. Colon length was measured, and tissues were either fixed in 4% paraformaldehyde or stored at −80 °C for further analysis.

For fecal microbiota transplantation (FMT), FMT was performed to evaluate the contribution of the gut microbiota to the therapeutic effects of PCP2. Donor feces were collected from the Model and PCP2-H groups under sterile conditions on day 10. Fresh feces (100 mg) were suspended in 1 mL of sterile saline, homogenized, and centrifuged at 800 × g for 3 min at 4 °C to remove debris. The supernatant containing the microbial community was collected for transplantation. Recipient mice were pre-treated with a broad-spectrum antibiotic cocktail (1 g/L ampicillin, 1 g/L neomycin, 1 g/L metronidazole, and 0.5 g/L vancomycin) in their drinking water for 7 consecutive days to deplete the indigenous microbiota. Depletion was confirmed by 16S rRNA gene qPCR of fecal samples. Following a washout period, the microbiota-depleted mice were orally gavaged with 200 μL of the donor fecal suspension on alternate days. Colitis was induced by 3% DSS as described above, commencing 3 days after the initial FMT inoculation.

For Antibiotic Exposure (Abx), To assess the microbiota-dependency of PCP2’s effects, a pseudo-germ-free mouse model was established. C57BL/6 J mice were randomly assigned to three groups (n = 8): Abx+DSS, Abx+DSS + PCP2, and DSS + PCP2. Mice were administered the aforementioned broad-spectrum antibiotic cocktail in drinking water for 2 weeks to maximally deplete gut bacteria, followed by a maintenance dose (20% of the initial concentration) throughout the experiment. Under these microbiota-depleted conditions, colitis induction (3% DSS) and PCP2 treatment (400 mg/kg) were performed following the standard protocol described above.

To investigate the functional role of Follistatin (Fst), adenovirus vectors carrying a short hairpin RNA (shRNA) targeting murine Fst (Ad-Fst) and a scrambled negative control (Ad-NC) were constructed by Cyagen Biosciences (Suzhou, China). Three weeks prior to DSS induction, mice received a single tail vein injection of the adenovirus at a titer of 1 × 10¹¹ plaque-forming units (PFU)/mouse. Fst knockdown efficiency in colonic tissues was verified by Western blotting. After a 3-week incubation period to allow for stable gene silencing, the mice were subjected to the standard DSS-induced colitis and PCP2 treatment protocols.

DAI determination

The Disease Activity Index (DAI) was assessed daily by monitoring body weight, stool consistency, and the presence of fecal occult blood. Scores were assigned to each parameter according to the criteria outlined in Table 3. The DAI was calculated as the sum of the individual scores for weight loss, stool consistency, and rectal bleeding, using the formula: DAI = Weight Change Score + Stool Consistency Score + Bleeding Score³⁸.

Table 3.

DAI Scoring for Mice

Score	Bodyweight loss (%)	Loose stool	Bloody stool
0	<1	Solid/granular	Negative
1	1–5	Soft/granular	Weakly positive
2	5–10	Semi-formed	Positive
3	10–15	Unformed	Strongly positive
4	≥15	Liquid	Strongly positive with visible blood

Histopathological analysis

Colonic tissues were paraffin-embedded and sectioned. The sections were stained with hematoxylin and eosin (H&E), Picrosirius red (PSR), or periodic acid-Schiff (PAS) for histopathological evaluation. Images were acquired using an optical microscope (Olympus, Tokyo, Japan). All histological assessments were performed on longitudinal sections of the distal colon. Histological scoring was conducted on H&E-stained sections by two independent observers in a blinded manner. The total histological score (ranging from 0 to 10) was calculated as the sum of inflammation severity (0–3), inflammation depth (0–3), and crypt damage (0–4)³⁹. To quantify the mucus layer, the PAS-positive area was measured in five non-overlapping fields (200× magnification) per section using ImageJ software (NIH, Bethesda, MD, USA) and expressed as a percentage of the total mucosal area.

Evaluation of inflammatory cytokines

The concentrations of IL-10, IL-6, and TNF-α in serum and colonic tissue homogenates were quantified using commercial ELISA kits (Nanjing Jiancheng Bioengineering Institute, Nanjing, China), in strict accordance with the manufacturer’s protocols. Optical density (OD) was measured using a microplate reader (Model 550, Bio-Rad Laboratories, Hercules, CA, USA).

Determination of oxidative stress levels

The levels of superoxide dismutase (SOD), malondialdehyde (MDA), glutathione peroxidase (GSH-Px), and glutathione (GSH) in serum were quantified using commercial assay kits (Nanjing Jiancheng Bioengineering Institute, Nanjing, China), in strict accordance with the manufacturer’s protocols. The absorbance was measured using a microplate reader (Model 550, Bio-Rad Laboratories, Hercules, CA, USA).

Protein extraction and western blotting experiments

Total protein was extracted from colonic tissues (50 mg) by homogenization in RIPA lysis buffer containing 1% PMSF (Solarbio, Beijing, China). Protein concentration was quantified using a BCA assay kit (YaZyme Biology, Shanghai, China). Equal amounts of protein were resolved by SDS-PAGE, transferred onto PVDF membranes, and blocked with 5% non-fat milk for 2 h at room temperature. The membranes were incubated overnight at 4 °C with primary antibodies against Claudin-1 (1:3000), Occludin (1:1000), and ZO-1 (1:1000) (Abcam, Cambridge, UK). Subsequently, the membranes were incubated with HRP-conjugated secondary antibodies (1:1000) for 2 h at room temperature. Protein bands were visualized using ECL reagents (Millipore, Billerica, MA, USA) on a Tanon 5200 system (Tanon Science & Technology, Shanghai, China). Densitometric analysis was performed using ImageJ software (NIH, Bethesda, MD, USA). Target protein levels were normalized to β-actin, and phosphorylated proteins were normalized to their respective total protein levels.

Immunofluorescence assay

Paraffin-embedded colonic sections were deparaffinized, rehydrated, and subjected to antigen retrieval. After blocking with 5% BSA, the sections were incubated overnight at 4 °C with primary antibodies targeting Occludin, Claudin-1, and ZO-1 (Abcam, Cambridge, UK). Subsequently, the sections were washed with PBS and incubated with corresponding fluorophore-conjugated secondary antibodies for 1 h at room temperature. Nuclei were counterstained with DAPI. Images were captured using a Nikon Eclipse C1 immunofluorescence microscope (Nikon, Tokyo, Japan).

Immunohistochemistry

Paraffin-embedded colonic sections were deparaffinized in xylene and rehydrated through a graded ethanol series. Antigen retrieval was performed by heating in 0.01 M sodium citrate buffer (pH 6.0). After washing with PBS, endogenous peroxidase activity was blocked with 3% hydrogen peroxide for 30 min, followed by blocking with 5% goat serum for 30 min to prevent non-specific binding. The sections were incubated overnight at 4 °C with primary antibodies against Ki67 (1:200) and MUC2 (1:1500; Abcam, Cambridge, UK). Subsequently, the sections were incubated with biotin-conjugated goat anti-rabbit IgG (CWBIO, Beijing, China) for 2 h, followed by horseradish peroxidase (HRP)-streptavidin (CWBIO) for 2 h. Immunoreactivity was visualized using a DAB chromogenic reagent kit (ZSGB-BIO, Beijing, China), and nuclei were counterstained with hematoxylin. For negative controls, the primary antibody was replaced with PBS. Images were acquired using an optical microscope (Olympus, Tokyo, Japan).

16S rRNA analysis

Genomic DNA was extracted from fecal samples using the E.Z.N.A.® Stool DNA Kit (Omega Bio-tek, Norcross, GA, USA). The V3–V4 hypervariable regions of the 16S rRNA gene were amplified using universal primers (338 F/806 R) and sequenced on an Illumina NovaSeq 6000 platform (Illumina, San Diego, CA, USA) by Hangzhou Lianchuan Biotechnology Co., Ltd. (Hangzhou, China). Raw data were processed using QIIME2 and/or the LC-Bio Cloud platform. High-quality sequences were clustered into amplicon sequence variants (ASVs) and taxonomically classified against the SILVA database (v138). Alpha and beta diversity indices were calculated to assess microbial community structure.

SCFA determination

Fecal samples (20 mg) were accurately weighed into 2-mL grinding tubes and mixed with 800 μL of 0.5% phosphoric acid solution spiked with 2-ethylbutyric acid (10 μg/mL) as an internal standard. The samples were homogenized (50 Hz, 3 min), ultrasonicated for 10 min, and centrifuged at 13,000 × g for 15 min at 4 °C. Subsequently, a 200-μL aliquot of the supernatant was extracted with an equal volume of n-butanol. The mixture was vortexed for 10 s, sonicated in an ice bath for 10 min, and centrifuged again (13,000 × g, 5 min, 4 °C). The resulting supernatant was transferred to an autosampler vial and analyzed using an 8890B GC system coupled to a 7000D triple quadrupole mass spectrometer (Agilent Technologies, Santa Clara, CA, USA).

Olink proteomics

Protein quantification was performed using Olink® Proximity Extension Assay (PEA) technology. Briefly, pairs of oligonucleotide-labeled antibodies specifically bind to the target protein in the sample. Upon binding in close proximity, the oligonucleotides hybridize and are extended by a DNA polymerase to create a unique DNA reporter sequence. This sequence was subsequently amplified and quantified by real-time PCR using the Signature Q100 system (Olink Proteomics, Uppsala, Sweden). Assays were performed using 1 µL of sample. Data were output as Normalized Protein Expression (NPX) values on a log2 scale, where a higher NPX value indicates a higher protein abundance.

Molecular dynamics simulation

MD simulations were performed using AmberTools 20. The protein was parameterized with the AMBER ff19SB force field, while the ligand was described using the General AMBER Force Field (GAFF), with AM1-BCC partial charges assigned via Antechamber. The complex was solvated in an OPC water box with a 10-Å buffer distance and neutralized with Na⁺ counterions. Following sequential restrained and unrestrained energy minimization, the system was heated to 300 K in the NVT ensemble and equilibrated in the NPT ensemble. A 50-ns production run was conducted at 300 K using a Berendsen thermostat with a 2-fs time step. Binding free energies were calculated using the MM-PBSA method. Trajectory analysis and conformational clustering (DBSCAN) were performed using CPPTRAJ⁴⁰.

Microscale thermophoresis

The binding affinity between PCP2 and Fst was determined using a Monolith NT.115 instrument (NanoTemper Technologies, Munich, Germany). Briefly, the Fst protein was fluorescently labeled using the Protein Labeling Kit RED-NHS (NanoTemper) following the manufacturer’s instructions. Unreacted dye was eliminated using the provided dye-removal columns. Subsequently, the labeled Fst was mixed with a serial dilution of PCP2 to establish a concentration gradient. The samples were loaded into standard treated capillaries, and thermophoresis measurements were performed using optimized excitation and MST power settings. Data analysis and K_D calculation were conducted using MO. Affinity Analysis software (NanoTemper)⁴¹.

Surface plasmon resonance

SPR experiments were performed using a Biacore T200 system (GE Healthcare, Uppsala, Sweden) equipped with a CM5 sensor chip. The sensor surface was activated for 420 s by injecting a 1:1 mixture of 0.1 M N-hydroxysuccinimide (NHS) and 0.4 M 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC) at 10 μL/min. Fst (20 μg/mL) was immobilized on the active flow cell (Fc2) to a target density of approximately 12,600 resonance units (RU), using an unmodified flow cell (Fc1) as the reference. Remaining active esters were blocked with 1 M ethanolamine-HCl (pH 8.5). PCP2 analyte solutions were prepared in running buffer at eight concentrations (1–100 μM) and injected over both flow cells at 20 μL/min. The association and dissociation phases were monitored for 100 s and 180 s, respectively, followed by surface regeneration between cycles⁴².

Statistical analyses

Data analyses were performed using GraphPad Prism (v9, Windows, GraphPad). The data are presented as mean ± SEM. Statistical comparisons between groups of different parameters were performed using one-way ANOVA. The significance levels were set as follows: *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, ^#P < 0.05, ^##P < 0.01, ^###P < 0.001, ^####P < 0.0001.

Author contributions

Qiangbao Xu: writing—original draft, validation, formal analysis, data curation, conceptualization. Qiuyue Lv: investigation, formal analysis, data curation, writing—original draft. Zhu Yang: investigation, data curation. Yiping Yang: investigation, data curation. Zihan Li, Yingying Zhang, Sumiao Zhan, Lingzhi Chen, Guodong Wang: investigation. Hui Che: writing—review & editing. Jiangping Wu: writing—review & editing, validation, funding acquisition. Jun Han: visualization, funding acquisition, conceptualization, and writing—review & editing.

Data availability

The relevant data can be obtained by the corresponding authors upon reasonable request. The raw 16S rRNA sequencing data and Olink proteomics data are available in the NCBI Sequence Read Archive (SRA) under accession number SUB15811941.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at 10.1038/s41538-026-00729-3.