Cyclophosphamide (CTX) was purchased from Sigma-Aldrich (USA). Enzyme-linked immunosorbent assay (ELISA) kits for cytokines, including interleukin 6 (IL-6) and tumor necrosis factor α (TNF-α), were obtained from Beijing 4A Biotech Co., Ltd. (Beijing, China). The following assay kits were purchased from Nanjing Jiancheng Bioengineering Institute (Nanjing, China), including lactate dehydrogenase (LDH), acid phosphatase (ACP), superoxide dismutase (SOD), malondialdehyde (MDA), and glutathione peroxidase (GSH-Px). Primary antibodies against anti-phospho-c-Jun N-terminal kinase (p-JNK, mouse monoclonal, 1:500), JNK (rabbit monoclonal, 1:1000), anti-phospho-p38 (p-p38, rabbit monoclonal, 1:1000), p38 (rabbit monoclonal, 1:1000), anti-phospho-ERK (p-ERK, rabbit monoclonal, 1:1000), ERK (mouse monoclonal, 1:500), anti-phospho-NF-kappa-B p65 (p-p65, rabbit monoclonal, 1:500), p-65 (rabbit monoclonal, 1:500), β-actin (anti-rabbit antibody, 1:3000)), Zonula occludens-1 (rabbit monoclonal, 1:1000), Claudin-1 (rabbit monoclonal, 1:500) and Occludin (rabbit monoclonal, 1:500) were obtained from Wuhan Servicebio Biotechnology Co., Ltd. (Wuhan, China). Horseradish peroxidase (HRP)-conjugated secondary antibodies were used, including sheep anti-rabbit (1:3000) and sheep anti-mouse (1:1000). All other conventional chemicals were of the analytical grade.

According to the previous extraction methods of our research group (16, 18), polysaccharides from P. corylifolia were extracted using hot water extraction followed by ethanol precipitation. Proteins were removed by Sevage method, and the resulting extract was freeze-dried to obtain purified polysaccharides PPs, which was used in this study. The main active fraction of PPs was PCp-I, with molecular weight of 2.721×10⁴. PCp-I mainly contained galactose, arabinose, mannose, xylose, rhamnose and glucose. Its main backbone consisted of →5)-α-Araf-(1→, →2, 4)-α-Rhap-(→, →4)-α-Galp-(1→ and β-Glcp-(1→ linkages. Detailed structural information was provided in our previous work (16).

Male Balb/c mice (20 ± 2 g, 6–8 weeks old) from Zhengzhou Huaxing Animal Center (Henan) were housed under standard conditions (23-25 °C, 60–70% humidity, 12 h light/dark cycle) with free access to food and water at Huanghe Science and Technology College. All experiments complied with the ethical guidelines approved by the institution’s Academic Committee. Mice were randomly divided into five groups (n=12 per group) according to body weight. The experimental design was illustrated in Figure 1A. Blank control group (BC) and model control group (MC) were gavaged with 0.1 mL/10 g of normal saline, while positive control group (PC) received levamisole hydrochloride (LH, 10 mg/kg, 0.1 mL/10 g). The high-dose (HD) and low-dose (LD) groups were gavaged daily with PPs at 300 and 75 mg/kg, respectively, for 18 days. On days 16-18, mice in BC group were intraperitoneally injected with normal saline, whereas all other groups were injected with CTX (80 mg/kg/d) to induce immunosuppression. After fasting for 12 h, blood, spleen, liver, small intestine, and cecum contents were collected and stored at -80°C for subsequent analysis.

Body weights were recorded every two days from the first day of gavage for a total of nine measurements. The spleen and liver were excised, rinsed with cold saline, blotted, and weighed. Organ indices were calculated according to the following Formula.

Blood samples were collected from the retro-orbital plexus, allowed to stand for 2 h at room temperature, centrifuged at 3500 r/min for 10 min to obtain serum. Serum levels of IL-6 and TNF-α were measured using ELISA kits. The activities of LDH and ACP were assayed using corresponding commercial kits according to the manufacturer’s instructions.

Liver tissues were accurately weighed and homogenized with 9 volumes of cold normal saline (weight-to-volume ratio = 1: 9) to prepare a 10% homogenate. After centrifugation at 3500 r/min for 10 min, the supernatant was collected. The activities of SOD and GSH-Px, as well as the MDA content were determined according to the instructions of the corresponding assay kits.

Spleen and small intestine samples were fixed in 4% paraformaldehyde for at least 24 h, dehydrated, and embedded in paraffin. Paraffin blocks were sectioned into 3–4 μm slices using a microtome, deparaffinized, and rehydrated. Sections were stained with hematoxylin and eosin (H&E), dehydrated, cleared, and mounted with coverslips. Histological structures were observed using a bright-field microscope at 200× magnification.

The distribution of CD4⁺ and CD8⁺ T cells in small intestinal tissue were assessed using immunohistochemical (IHC) staining. Paraffin-embedded tissue sections were deparaffinized, rehydrated, and subjected to antigen retrieval. Endogenous peroxidase activity was quenched with 3% hydrogen peroxide. Sections were encircled with an IHC pen and blocked with 3% bovine serum albumin (BSA). After washing with PBS, primary antibodies were applied and incubated overnight at 4°C. After washing three times with PBS, the sections were incubated with species-matched secondary antibodies, developed with DAB chromogen, and counterstained with hematoxylin. Finally, sections were dehydrated, mounted, and examined under a light microscope.

Spleen or small intestine tissues were rinsed with pre-cooled PBS, minced, and transferred to homogenization tubes containing 10 volumes of lysis buffer and two 4 mm homogenization beads. After homogenization, lysates were kept on ice for 30 min and centrifuged at 12,000 rpm for 10 min at 4°C. The supernatant was collected as the total protein extract. Protein concentration was measured using a BCA assay kit. Samples were mixed with 5×loading buffer (v/v= 4:1), boiled for 15 min, and stored at -20°C. Proteins were separated by SDS-PAGE (5% stacking gel, 10% separating gel) and transferred to PVDF membranes activated in methanol. Membranes were blocked with 5% skim milk for 30 min, incubated overnight at 4 °C with primary antibodies, washed three times with TBST, and incubated with HRP-conjugated secondary antibodies for 30 min. Protein bands were visualized using an enhanced chemiluminescence (ECL) reagent and imaged with an SCG-W3000 PLUS imaging system. Band intensities were quantified using ImageJ software.

Cecum contents were collected for genomic DNA extraction and quality assessment. Amplification of 16S rDNA variable region, library construction, high-throughput sequencing, and subsequent bioinformatics analysis were performed by Jinwei Zhi Biotechnology Co., Ltd. (Suzhou, China).

Serum metabolites were analyzed using UPLC-MS/MS (Thermo Scientific Q Exactive HF). Raw data processed using Compound Discoverer 3.3 software (Thermo Fisher Scientific, USA) in combination with BMDB (BGI Metabolome Database), mzCloud database and ChemSpider online databases. Principal component analysis (PCA) and partial least squares discriminant analysis (PLS-DA) were used to evaluate metabolite variations among groups. Differential metabolites were screened based on variable importance in projection (VIP) ≥1, Fold Change ≥1.2 or ≤ 0.83, and P < 0.05. Metabolic pathways enrichment was conducted using the KEGG (http://www.genome.jp/kegg/) and HMDB (https://hmdb.ca/) databases. Spearman’s correlation analysis was performed to analyze associations between intestinal flora (genus level) and metabolites.

All statistical analyses were conducted using GraphPad Prism 8.0. Data were mean ± standard deviation (SD). Statistical differences among groups were determined using one-way analysis of variance (ANOVA). Differences were considered statistically significant at P < 0.05.

PPs exerted positive effects on both body weight and immune organs in mice. The results are shown in Figures 1B–D. In the MC group, a significant decrease in body weight was observed three days after CTX injection (days 16-18). The body weight then remained consistently lower than that in the BC group. In contrast, mice in the PPsH and PPsL groups showed less weight reduction, suggesting that PPs alleviated CTX-induced weight loss. As shown in Figures 1C, D, CTX administration significantly reduced the spleen index compared with the BC group (P < 0.001), whereas both PPsH and PPsL treatment increased the spleen index in CTX-treated mice (P < 0.05). The effect on liver weight was minimal. These results indicated that PPs could mitigate CTX-induced immune organ damage.

Histological analysis of the spleen was performed (Figure 1E). In the MC group, CTX administration caused clear morphological damage. This included blurred boundaries between the red and white pulp, disorganized cellular structures, and a disruption of the normal tissue architecture. In contrast, PPs treatment restored clear medullary boundaries and dense cellular arrangements, suggesting effective protection of splenic structure from CTX-induced damage.

IL-6 and TNF-α are key inflammatory markers in immune regulation. As shown in Figures 2A, B, CTX markedly reduced serum levels of both IL-6 and TNF-α compared with the BC group (P < 0.001), while both PPsH and PPsL significantly restored the concentrations of these cytokines (P < 0.001).

As shown in Figures 2C, D, the activities of ACP and LDH were significantly decreased in the MC group (P < 0.01). PPsH significantly increased the activities of both enzymes (P < 0.01 and P < 0.05). PPsL also produced a significant effect, notably elevating LDH activity (P < 0.05). These results indicate that PPs counteracted CTX-induced suppression of macrophage enzyme activity, thereby enhancing immune function.

The metabolism of CTX triggers excessive production of reactive oxygen species (ROS), leading to oxidative stress. This state was evidenced in our model by reduced activities of the key antioxidant enzymes SOD and GSH-Px, alongside elevated levels of the lipid peroxidation marker MDA in the liver. Both PPsH and PPsL significantly enhanced SOD and GSH-Px activities, reduced MDA content, and improved hepatic antioxidant capacity (Figures 2E–G). Collectively, these findings demonstrated the multifaceted protective effects of PPs. They alleviated CTX-induced suppression of cytokine secretion, enhanced hepatic antioxidant defenses, and thereby exerted overall anti-inflammatory effects.

The NF-κB/MAPK signaling pathways are signaling cascade mediating immune responses. Western blot analysis of spleen tissue (Figure 3) found that CTX inhibited phosphorylation of p38 (P < 0.001), p-ERK was lower than that of BC group. Unexpectedly, the phosphorylation levels of JNK showed no significant decrease in macrophages, which may be attributed to the specific time point of sample collection. while PPs treatment increased phosphorylation of ERK, JNK and p65 (P < 0.001 or P < 0.05). These results suggested that the immunoenhancing effect of PPs might be related to activation of the NF-κB/MAPK signaling pathways.

PPs could enhance the function of the intestinal barrier. Histopathological observation of the small intestine (Figure 4A) showed that the BC group exhibited intact mucosal architecture with well-defined villi and crypts. CTX treatment caused clear structural damage, disrupting epithelial integrity and inducing crypt deformation. These morphological changes compromised intestinal function and increased susceptibility to pathogens (19). In contrast, PPs treatment effectively restored the mucosal structure. This restoration indicated an improved capacity for intestinal barrier repair. CD4⁺ and CD8⁺ T lymphocytes play essential roles in adaptive immunity. Immunohistochemical analysis (Figures 4B–D) showed that CTX significantly reduced the counts of both CD4⁺ and CD8⁺ T cells (P < 0.001). Both PPsH and PPsL restored these cell populations, with the most potent effect observed for PPsL (P < 0.001). This suggests that PPs enhance T cell proliferation and activation in the small intestine, thereby supporting local immune responses and cytokine secretion. Western blot analysis of tight junction proteins (Figure 4E) revealed that CTX markedly decreased the expressions of Claudin-1, Occludin, and ZO-1. Conversely, PPs treatment significantly upregulated these proteins (P < 0.001). The increased expression of these key proteins indicated improved epithelial tightness and reduced intestinal permeability.

16S rDNA sequencing revealed a total of 2,818,562 valid reads from 30 cecal content samples across five groups. α-Diversity indices (ACE, Chao1, Simpson, and Shannon) showed no significant differences between CTX and PPs groups (P > 0.05, Figure 5A), indicating that PPs did not alter overall gut microbiota richness. However, β-diversity analysis (Figure 5B) revealed distinct clustering among groups, with PPsH and PPsL showing clear separation from the CTX group, suggesting compositional shifts in the gut microbiota.

At the phylum level, the microbial community structure was shown in Figure 5C. Firmicutes and Bacteroidetes dominated across all groups (> 93% relative abundance), PPs reduced the Firmicutes/Bacteroidetes (F/B) ratio. At the genus level (Figure 5D), the relative abundances of Lachnospiraceae_NK4A136_group and Bacteroides were higher in the BC group compared to other groups. CTX increased the relative abundances of f:Lachnospiraceae_Unclassified and Prevotellaceae_UCG-001. Compared to the MC group, the PPsH and PPsL groups showed higher relative abundances of Muribaculaceae and lower abundances of Prevotellaceae_UCG-001.

Linear Discriminant Analysis Effect Size (LEfSe) analysis identified group-specific biomarker taxa (Supplementary Figure 1). Among them, g_Lactobacillus was enriched in BC group with a high LDA score, g_Ruminococcus in MC group, while g_Helicobacter and g_Oscillibacter were characteristic PPsH and PPsL groups, respectively. It is noteworthy that a total of 30 bacterial biomarkers were differentially enriched in the PPs-treated groups. These results indicated that under immune-challenged conditions, these gut microbes responded to PPs intervention and collectively participated in modulating the host’s immune response.

Dietary polysaccharides serve as a major nutrient source for gut bacteria and play a key role in immune regulation, while metabolites act as critical mediators between the host and the microbiota. In previous studies, we investigated changes in the gut microbiome and metabolome. It has been demonstrated that supplementation with PPs altered metabolite profiles. To further investigate the systemic effects, we analyzed serum metabolites after PPs administration. This analysis was performed using a UPLC-Q-Exactive Orbitrap MS system. A total of 3,204 ion features were identified in both positive and negative ion modes. PCA and PLS-DA analyses revealed distinct separations among groups (Figures 6A, B). This indicated that CTX markedly altered the serum metabolic profile. Notably, PPs treatment partially reversed these CTX-induced metabolic changes.

Differential metabolites were screened according to predefined criteria, and the results are presented in Figures 6C, D. KEGG enrichment analysis was performed to identify affected metabolic pathways (Figures 6E, F). Based on the criteria of P < 0.05 and Rich Factor > 0.01, CTX was found to significantly affect 25 pathways (the top 20 are displayed). These included pathways such as D-amino acid metabolism, pantothenate and CoA biosynthesis, carbon metabolism, amino acid biosynthesis, glyoxylate and dicarboxylate metabolism, histidine metabolism, and protein digestion and absorption. PPs treatment, in contrast, significantly modulated six key metabolic pathways. These primarily involved protein digestion and absorption, cofactor biosynthesis, nicotinate and nicotinamide metabolism, lipoic acid metabolism, antifolate resistance, and branched-chain amino acid biosynthesis. Notably, when comparing the PPsH group to the MC group, only the protein digestion and absorption pathway showed a significant reversal, while the other pathways altered by CTX remained unchanged. Similarly, aside from protein digestion and absorption, none of these pathways were significantly altered between the MC and BC groups. These results suggested that PPs ameliorated CTX-induced metabolic disturbances via modulating these key pathways. Among them, the protein digestion and absorption pathway appeared to hold particular potential for alleviating the metabolic disorders caused by CTX.

To elucidate the mechanism of PPs-mediated immune modulation, we analyzed the correlations among gut microbiota, serum metabolites, and immunological parameters. As shown in Figure 7A, the relative abundances of f:Lachnospiraceae_Unclassified, Prevotellaceae_UCG-001, and Allopretrotella were negatively correlated with immune and antioxidant markers, including IL-6, TNF-α, ACP, LDH, GSH-Px, SOD and spleen index. Conversely, their abundances were positively correlated with MDA. On the contrary, Lachnospiraceae_NK4A136_group and Lactobacillus were positively correlated with IL-6, TNF-α, ACP, LDH, SOD, spleen index and liver index, whereas negatively correlated with MDA. Similarly, correlation analysis of the main differential metabolites and immunosuppression markers (Figure 7B) showed that the contents of arachidic acid, 8-iso prostaglandin A1, D-(+)-tryptophan, mesaconic acid, 3-furoic acid and D-ornithine were positively correlated with IL-6, TNF-α, ACP, LDH, GSH-Px, SOD and spleen index, but negatively correlated with MDA. In contrast, metabolites such as NP-020078, pentadecanoic acid and valine showed opposite trends. These findings suggested that PPs modulated gut microbial structure and serum metabolites, jointly contributing to the alleviation of immunosuppression.