Rosa roxburghii Tratt fruit polysaccharides inhibit proliferation and induce apoptosis in human cervical cancer cell line

Introduction

Cervical cancer is one of the most common malignant tumors among women globally, with its incidence and mortality rates continuously rising, posing a severe challenge to public health (1). The development of cancer is a multi-stage and multi-factorial process, typically progressing through carcinogenesis, promotion, and evolution stages, characterized by abnormal cell proliferation, uncontrolled differentiation, and enhanced invasiveness and metastatic potential (2). Current clinical treatments mainly rely on surgery, chemotherapy, and radiotherapy. Although these methods have certain therapeutic effects, they are often accompanied by issues such as drug resistance, side effects, and insufficient targeting (3). Therefore, the development of highly effective, low-toxicity novel anti-cancer drugs has become a research hotspot.

Natural polysaccharides, with their multi-target, low toxicity, high safety, and synergistic potential with chemotherapy drugs, have attracted significant attention in anti-cancer drug development (4). The mechanisms of their action can be broadly categorized into direct and indirect effects: direct effects include the inhibition of tumor cell proliferation, migration, and invasion, the induction of apoptosis, regulation of autophagy, and cell cycle arrest; indirect effects involve the modulation of the tumor microenvironment, such as activating macrophages, enhancing immune responses, and inhibiting tumor angiogenesis (2,5). Several studies have confirmed the anti-cancer activity of polysaccharides from different sources. For example, Angelica sinensis polysaccharides (ASP) inhibit the Wnt/β-catenin pathway by regulating the miR-3187-3p/PCDH10 axis, suppressing the proliferation, migration, and invasion of breast cancer cells (6); Rosa roxburghii polysaccharides inhibit liver cancer HepG2 and lung adenocarcinoma A549 cells through immune regulation and anti-angiogenesis effects in a zebrafish model (7); edible mushroom polysaccharides inhibit tumor-associated macrophages (TAMs), enhancing immune surveillance and stimulating lymphocyte activity (8); additionally, Astragalus polysaccharides, Ganoderma lucidum polysaccharides, and Lentinus edodes polysaccharide have shown anti-tumor activity in the cervical cancer HeLa cell line (9-11). These findings highlight the significant value of polysaccharides as low-toxicity, multi-target anti-cancer candidates.

Among various natural polysaccharides, RTFPs have garnered attention due to their rich bioactivities. Rosa roxburghii Tratt, a perennial deciduous shrub primarily distributed in the northwest of Yunnan and Guizhou provinces in China, produces fruits known as “Wenxian fruit”, which are rich in polyphenols, polysaccharides, flavonoids, and other bioactive components, with significant medicinal and nutritional value (12-14). As one of its main active components, RTFP exhibits various bioactivities, including antioxidant, hypoglycemic, anti-inflammatory, gut microbiota regulation, and anti-tumor effects (14). A study has shown that RTFP has strong antioxidant activity comparable to that of standard antioxidants, as well as dose-dependent immunomodulatory effects, including enhancing phagocytic activity and regulating key immune mediators (15). In addition, RTFP has shown good potential in lowering blood glucose, acting as a prebiotic, and alleviating colitis induced by a high-fat diet (16-18), and has exhibited significant inhibitory activity against breast cancer MCF-7 cells and prostate cancer DU145 cells (19,20). Although RTFP has been confirmed to have various bioactivities and demonstrated anti-tumor effects in certain tumor models, its potential role in cervical cancer and related molecular mechanisms have not been systematically reported. Therefore, in this study, human cervical cancer HeLa cells were used as a model to explore the effects of RTFP on proliferation, migration, invasion, and apoptosis, and to preliminarily analyze its relationship with the expression of cell cycle and apoptosis-related proteins, aiming to provide in vitro experimental evidence for the development of RTFP-based strategies for cervical cancer prevention and treatment. We present this article in accordance with the MDAR reporting checklist (available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-aw-2431/rc).

Methods

Extraction of RTFP

The fruits of Rosa roxburghii Tratt were purchased from a traditional Chinese medicine wholesale market in Dali Bai Autonomous Prefecture, Yunnan Province, China, and were authenticated botanically by Prof. Chen (Dali University). The extraction procedure was adapted from a previously reported method (21). Briefly, the dried fruits were ground into particles of 10–20 mesh, defatted with petroleum ether (3× volume, refluxed twice for 3 h each), and the residue was filtered and dried at 55 ℃ to constant weight. Polysaccharides were extracted by hot-water stirring (material-to-liquid ratio 1:30 g/mL) at 70 ℃ for 3 h, and the extraction was repeated three times. The combined extracts were concentrated under reduced pressure, followed by deproteinization using the Sevage method (chloroform: n-butanol =4:1) twice (Fuchen Chemical Reagent Factory, Tianjin, China). Polysaccharides were precipitated by adding four volumes of ethanol and standing overnight at 4 ℃. The precipitate was collected, washed sequentially with anhydrous ethanol, acetone, and diethyl ether, and then decolorized with macroporous resin (Macklin Biochemical Co., Ltd., Shanghai, China). After freeze-drying, the refined polysaccharide fraction (RTFP) was obtained (Figure 1A). The polysaccharide yield was determined using the anthrone-sulfuric acid method (22). To evaluate purity, RTFP was dissolved in water (1 mg/mL) and scanned by ultraviolet-visible (UV-Vis) (Jinghua Scientific Instrument Co., Ltd., Shanghai, China) spectrophotometry (260–280 nm) to detect potential protein or nucleic acid contamination.

Figure 1 Extraction procedure and preliminary purity evaluation of RTFP. (A) Schematic diagram of RTFP extraction. (B) UV-Vis absorption spectrum of RTFP (1 mg/mL in water) in the 260–280 nm wavelength range, used to detect protein and nucleic acid impurities. RTFP, Rosa roxburghii Tratt fruit polysaccharides; UV-Vis, ultraviolet-visible.

Cell culture and treatment

HeLa cells (catalog number: XB0421-1785) were obtained from Wuhan Pricella Biotechnology Co., Ltd. (Wuhan, China). HeLa cells were cultured in minimum essential medium (MEM) supplemented with 10% fetal bovine serum (FBS) (GIBCO, Grand Island, NE, USA) and 1% penicillin/streptomycin, and maintained in a humidified incubator (Shenli Scientific Instrument Co., Ltd., Shanghai, China) at 37 ℃ with 5% CO₂. Cells in the logarithmic growth phase were harvested and subcultured. RTFP was dissolved in MEM (GIBCO) to prepare stock solutions at the experimental concentrations (0–10 mg/mL). Prior to each experiment, the solution was sterilized by filtration through a disposable 0.22 µm filter (Millex^®-GP, Merck Millipore, Darmstadt, Germany) twice, and then immediately applied to HeLa cells at the predetermined concentrations for subsequent experiments. Throughout the entire study, no signs of nonspecific immune activation were observed.

Cell proliferation assay

HeLa cells (1×10³ cells/well) were seeded in 96-well plates (NEST, Wuxi, China), and divided into three groups: blank (100 µL MEM medium), control (100 µL cell suspension in MEM medium), and RTFP (100 µL cell suspension with RTFP at 2, 4, 6, 8, and 10 mg/mL). After treatment for 24, 48, and 72 h, cell morphology was directly observed under an inverted microscope to perform preliminary sensitivity screening experiments. Cell viability was assessed using the Cell Counting Kit-8 (CCK-8) assay (Med Chem Express, Shanghai, China). Briefly, under the same culture conditions, the original medium was discarded, and each well was replaced with a mixture of 90 µL fresh MEM medium and 10 µL CCK-8 reagent (final volume 100 µL). After incubation at 37 ℃ for 1 h, the absorbance was measured at 450 nm using a microplate reader (Huisong Technology Development Co., Ltd., Shenzhen, China). Wells containing medium and CCK-8 reagent without cells were set as blank controls. All experiments were performed in triplicate. The half-maximal inhibitory concentration (IC₅₀) value was calculated using SPSS software (version 20.0; SPSS Inc., Chicago, IL, USA). Cell viability (%) was calculated as:

Cellviability=ARTFP−ABlankAControl−ABlank×100%[1]

where A_RTFP indicates absorbance of the experimental group (HeLa cells + RTFP + MEM + CCK-8); A_Control indicates absorbance of the control group (HeLa cells + MEM + CCK-8); and A_Blank indicates absorbance of the blank group (MEM + CCK-8).

Scratch assay

HeLa cells (1×10⁵ cells/well) were seeded in six-well plates (NEST) and cultured in complete MEM until >80% confluence. A sterile 200 µL pipette tip was used to create a uniform vertical scratch in the cell monolayer. After gently washing twice with phosphate-buffered saline (PBS) (Servicebio, Wuhan, China) to remove detached cells, the control group received 2.5 mL FBS-free MEM medium, while the RTFP groups received 2.5 mL FBS-free MEM medium containing RTFP at 3 and 6 mg/mL. All groups were performed in triplicate. Scratch images were captured at 0, 24, and 48 h post-scratching using an inverted microscope (Olympus CKX41, Tokyo, Japan) equipped with a digital camera at 100× magnification. For each well, images were taken at the same marked positions to ensure consistency. The scratch width was measured at three random sites per image using ImageJ software (National Institutes of Health, Bethesda, MD, USA).

Transwell assay

HeLa cells (1×10⁵ cells/mL, 200 µL) were seeded into the upper chambers of Trans well inserts (8.0-µm pore size; Corning Inc., Beijing, China) with 800 µL of complete MEM added to the lower chambers. Prior to seeding, the upper surface of the Transwell membrane was coated with Matrigel (Corning Inc.). After 8 h attachment period, the medium in the upper chambers was replaced with test media: serum-free MEM (control) or serum-free MEM supplemented with RTFP (3 and 6 mg/mL). All groups were performed in triplicate. After 24 and 48 h of incubation, the chambers were washed with PBS (Servicebio), fixed with 4% paraformaldehyde for 15 min, and stained with Giemsa for 15 min. Invaded cells on the lower membrane surface were counted in five randomly selected non-overlapping fields of view per membrane using an inverted microscope (Olympus) and semi-automatically quantified with ImageJ software.

Cell cycle assay

HeLa cells (1×10⁶ cells) were seeded into six-well plates (NEST) and cultured for 12 h. The medium was then replaced with FBS-free MEM medium, and the cells were incubated for an additional 12 h. Subsequently, the Control group was supplemented with 2.5 mL of complete MEM medium, while the RTFP group was treated with 2.5 mL of complete MEM medium containing RTFP at 3 and 6 mg/mL, with three replicates per group. After 48 h, the cells were harvested via trypsinization (Vazyme, Nanjing, China), centrifuged at 900 rpm for 4 min, and fixed in 75% pre-chilled ethanol for 24 h. Prior to analysis, the cells were centrifuged again at 900 rpm for 4 min, stained with 500 µL of propidium iodide (PI) (Beyotime, Shanghai, China) staining solution, and incubated at 37 ℃ in the dark for 30 min. Cell cycle distribution was subsequently analyzed using a BD FACSCanto™ II flow cytometer (BD Biosciences, San Jose, CA, USA). At least 20,000 events were collected for analysis for each sample.

Apoptosis assay

HeLa cells (1×10⁶ cells) were seeded in a six-well plate (NEST) and treated according to the experimental groups for 48 h. Cells were harvested by centrifugation at 900 rpm for 4 min, stained with 210 µL Annexin V-fluorescein isothiocyanate (FITC)/PI reagent (Beyotime), and incubated in the dark on ice for 30 min. Apoptosis was analyzed using a BD FACSCanto™ II flow cytometer (BD Biosciences). At least 20,000 events were collected for analysis for each sample.

Real-time polymerase chain reaction (PCR) for gene expression

HeLa cells were treated according to the experimental groups for 48 h. Total RNA was extracted using Trizol reagent (Vazyme), and RNA integrity was verified by electrophoresis. Total RNA was reverse-transcribed into complementary DNA (cDNA) using the HiScript III 1^st Strand cDNA Synthesis Kit (Vazyme). All groups were performed in triplicate. Gene expression was analyzed using real-time PCR with cDNA as the template. Primer sequences for human cyclin B1, CDK-1, caspase-3, caspase-8, caspase-9, and β-actin were retrieved from the NCBI Gene Bank database and designed using Primer 5.0 software (Table 1). PCR conditions were as follows: 95 ℃ for 30 s, followed by 40 cycles of 95 ℃ for 10 s, 60 ℃ for 30 s, and 60 ℃ for 60 s. Relative gene expression was calculated using the 2^−ΔΔCT method. The reactions were performed using an ABI PRISM 7500 Sequence Detection System (Applied Biosystems, Foster City, CA, USA).

Western blot analysis for protein expression

HeLa cells were lysed in radioimmunoprecipitation assay (RIPA) buffer (Beyotime) supplemented with 1 mM phenylmethylsulfonyl fluoride (PMSF) to inhibit protease degradation. Protein concentration was determined using a bicinchoninic acid (BCA) reagent kit (Solarbio, Beijing, China). Proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) (Solarbio) and transferred to a polyvinylidene difluoride (PVDF) membrane (Millipore, Burlington, MA, USA). The membrane was blocked with 5% skim milk powder for 2 h at room temperature and then washed three times with Tris-buffered saline with Tween^® 20 (TBST; Servicebio). The membranes were incubated overnight at 4 ℃ with the following rabbit primary antibodies from Cell Signaling Technology (CST, Danvers, MA, USA): CDK-1 (1:1,000, Cat. No. AF2515), cyclin B1 (1:1,000, Cat. No. 4135S), Bax (1:1,000, Cat. No. 15071T), Bcl-2 (1:500, Cat. No. 5023T), caspase-3 (1:1,000, Cat. No. 14220S), caspase-8 (1:1,000, Cat. No. 4790S), caspase-9 (1:1,000, Cat. No. 9508S), and β-actin (1:1,000, Cat. No. 4967S) as the loading control. After washing with TBST, the membranes were incubated for 2 h at room temperature with horseradish peroxidase (HRP)-conjugated goat anti-rabbit secondary antibody (1:2,000, Beyotime, Cat. No. A0208). Bands were visualized using enhanced chemiluminescence (ECL) plus solution (Meilunbio, Dalian, China). The experiment was repeated three times, and band intensities were quantified using ImageJ software (version 2023).

Statistical analysis

Statistical analysis was conducted using GraphPad Prism software (version 7.0) and SPSS software (version 20.0; SPSS Inc.); Image J 2023, Adobe Photoshop 2021. All experiments were performed independently at least three times (biological replicates, n≥3), with each biological replicate containing three technical replicates. Data are presented as mean ± standard deviation (SD) calculated from the biological replicates (technical replicates were averaged within each biological replicate prior to analysis). Differences were considered statistically significant at *, P<0.05; **, P<0.01 [two-tailed Student’s t-test or one-way analysis of variance (ANOVA) followed by post-hoc tests, as appropriate].

Results

Extraction of RTFP

The polysaccharide content, as determined by the anthrone-sulfuric acid method, was 25.15%±0.78% [weight by weight (w/w)]. Although the monosaccharide composition and molecular weight of RTFP were not characterized in detail in this study, the UV-Vis spectrophotometry results of RTFP (Figure 1B) showed no characteristic absorption peaks at 260 or 280 nm, indicating that RTFP contains negligible protein or nucleic acid contamination. This confirms that the preparation method effectively removed major non-polysaccharide impurities, ensuring the stability and quality of RTFP for subsequent in vitro activity screening.

RTFP inhibits proliferation in HeLa cells

Following treatment of HeLa cells with RTFP for 24, 48, and 72 h, the most pronounced effects on both cell viability and morphology were observed at 48 h (Figure 2). Microscopic examination (100× magnification) revealed that, compared to the control, increasing concentrations of RTFP led to a significant reduction in cell density and notable morphological alterations, including cell shrinkage and diminished adhesion, indicating a dose-dependent inhibitory effect on HeLa cell proliferation. CCK-8 assay results (Table 2) further demonstrated that at 24 h, 2 mg/mL RTFP did not significantly affect cell viability (P>0.05). In contrast, at 48 h, cell viability decreased progressively with increasing RTFP concentration, exhibiting statistically significant, time- and dose-dependent inhibition (P<0.01). Based on the 48-h dose-response curve, the IC₅₀ of RTFP against HeLa cells was calculated to be 3.323 mg/mL. Based on this IC₅₀ value, two treatment groups were established for subsequent experiments: a low-dose group (3 mg/mL, approximately the IC₅₀) and a high-dose group (6 mg/mL, approximately 2 × IC₅₀ value).

Figure 2 Morphological changes in HeLa cells after 48 h treatment with RTFP (×100 magnification; n=3). (A) 0 mg/mL (control): cells exhibited normal polygonal morphology with clear cell borders. (B) 2 mg/mL: partial cell rounding was observed. (C) 4 mg/mL: obvious cell rounding with decreased cell density. (D) 6 mg/mL: cytoplasmic condensation and rounding, accompanied by reduced confluence. (E) 8 mg/mL: pronounced cell shrinkage and detachment, with markedly reduced adherent cells. (F) 10 mg/mL: widespread cell fragmentation and formation of apoptotic bodies. RTFP, Rosa roxburghii Tratt fruit polysaccharides.

RTFP inhibits migration in HeLa cells

RTFP inhibited HeLa cell migration in a dose- and time-dependent manner (Figure 3). At 24 h, the residual wound areas significantly increased to 184,915.05±11,684.59 µm² (3 mg/mL, P<0.05) and 203,433.11±8,236.04 µm² (6 mg/mL, P<0.01), compared with 173,362.38±7,580.23 µm² in the control group. The inhibitory effect was more pronounced at 48 h, with migration inhibition rates of approximately 35% (3 mg/mL) and 74% (6 mg/mL), respectively, relative to the control. Correspondingly, the residual wound areas were 137,543.83±4,721.27 µm² (3 mg/mL, P<0.05) and 177,361.02±8,136.19 µm² (6 mg/mL, P<0.01), vs. 102,013.92±1,073.68 µm² in the control group.

Figure 3 Migration inhibition of RTFP on HeLa cells (n=3). (A) Representative phase-contrast images of scratch assay showing RTFP effect on HeLa cell migration at 0 (control), 3, and 6 mg/mL after 24 and 48 h (×40 magnification). (B) Quantitative analysis of migration changes. **, P<0.01 vs. control group. Data are presented as mean ± SD. RTFP, Rosa roxburghii Tratt fruit polysaccharides; SD, standard deviation.

RTFP inhibits invasion in HeLa cells

Increasing concentrations of RTFP led to a progressive reduction in the number of cells migrating through the Transwell membrane (Figure 4). Quantitative results showed that after 24 h, invasive cell numbers were 140±6 (control), 101±2 (3 mg/mL, P<0.05), and 49±5 (6 mg/mL, P<0.01). After 48 h of treatment, the number of invading cells was 106±5 in the control group, 69±4 in the 3 mg/mL RTFP group (P<0.05), and 21±4 in the 6 mg/mL RTFP group (P<0.01). This corresponded to approximate reductions of 35% and 80%, respectively, relative to the control. These findings suggest that RTFP inhibits HeLa cell invasion in a time- and dose-dependent manner.

Figure 4 Inhibitory effect of RTFP on invasion of HeLa cells (n=3). (A) Representative images of Transwell assay demonstrating the effect of RTFP on HeLa cell invasion after 24 and 48 h treatment at 0 (control), 3, and 6 mg/mL (Giemsa staining, ×200 magnification). (B) Quantitative analysis of RTFP-induced changes in HeLa cell invasion after 24 and 48 h exposure using the Transwell assay. **, P<0.01 vs. control group. Data are presented as mean ± SD. RTFP, Rosa roxburghii Tratt fruit polysaccharides; SD, standard deviation.

RTFP arrests HeLa cell cycle at G2/M phase

RTFP arrested the cell cycle at the G2/M phase in HeLa cells in a dose-dependent manner (Figure 5). After 48 h, the percentage of G2/M phase cells increased significantly from 6.66%±0.65% in the control group to 11.57%±0.38% (3 mg/mL, P<0.05) and 23.37%±1.68% (6 mg/mL, P<0.01), while the proportions of cells in the G0/G1 and S phases decreased accordingly. These data indicate that RTFP effectively induces G2/M phase arrest in HeLa cells.

Figure 5 Effect of RTFP on HeLa cell cycle (n=3). (A) Representative flow cytometry plots showing RTFP effect on HeLa cell cycle at 0 (control), 3, and 6 mg/mL after 48 h (PI staining), cells synchronized in FBS-free MEM medium, fixed in 75% ethanol. (B) Quantitative analysis of G2/M phase changes. ns, not significant (P>0.05); **, P<0.01 vs. control group. Data are presented as mean ± SD. FBS, fetal bovine serum; MEM, minimum essential medium; PI, propidium iodide; RTFP, Rosa roxburghii Tratt fruit polysaccharides; SD, standard deviation.

RTFP downregulates CDK-1 and cyclin B1 gene expression in HeLa cells

RTFP treatment for 48 h significantly downregulated the messenger RNA (mRNA) expression levels of CDK-1 and cyclin B1 in HeLa cells in a dose-dependent manner (P<0.05). Compared to the control group, the relative expression of CDK-1 decreased to 0.55±0.13 and 0.26±0.11 at RTFP concentrations of 3 and 6 mg/mL, respectively. Similarly, cyclin B1 expression was reduced to 0.83±0.06 and 0.38±0.09 at the corresponding doses. A more pronounced suppression of both genes was observed at the higher dose (6 mg/mL, P<0.01), indicating a clear dose-response relationship. These findings suggest that RTFP may inhibit HeLa cell proliferation by modulating the expression of key genes involved in the cell cycle.

RTFP downregulates CDK-1 and cyclin B1 protein expression in HeLa cells

Compared with the Control group, RTFP significantly downregulated the expression of CDK-1 and cyclin B1 proteins (P<0.01) (Figure 6). These findings suggest that RTFP may arrest the HeLa cell cycle at the G2/M phase by downregulating CDK-1 and cyclin B1 protein expression.

Figure 6 Effect of RTFP on the expression of CDK-1 and cyclin B1 proteins (n=3). (A) Representative Western blot images showing RTFP effect on CDK-1 and cyclin B1 expression in HeLa cells at 0 (control), 3, and 6 mg/mL after 48 h (β-actin normalized) (B) Quantitative analysis of protein expression changes. **, P<0.01 vs. control group. Data are presented as mean ± SD. RTFP, Rosa roxburghii Tratt fruit polysaccharides; SD, standard deviation.

RTFP promotes HeLa cell apoptosis

Apoptosis was assessed by flow cytometry after 48 h of RTFP treatment (Figure 7). The total apoptosis rates were 2.96%±0.15% (control), 6.78%±0.38% (3 mg/mL RTFP), and 17.68%±0.68% (6 mg/mL RTFP), with the treated groups showing significant increases compared to the control (P<0.01). Detailed quadrant analysis revealed early apoptosis rates of 1.13%, 2.96%, and 9.16%, and late apoptosis rates of 1.83%, 3.82%, and 8.52% for the control, 3 mg/mL, and 6 mg/mL groups, respectively. These findings indicate that RTFP induces HeLa cell apoptosis in a dose-dependent manner.

Figure 7 RTFP promotes HeLa cell apoptosis (n=3). (A) Representative flow cytometry plots showing RTFP effect on HeLa cell apoptosis at 0 (control), 3, and 6 mg/mL after 48 h (Annexin V-FITC/PI staining), cells harvested and stained. (B) Quantitative analysis of apoptosis changes. **, P<0.01 vs. control group. Data are presented as mean ± SD. FITC, fluorescein isothiocyanate; PI, propidium iodide; RTFP, Rosa roxburghii Tratt fruit polysaccharides; SD, standard deviation.

RTFP upregulates caspase-3, caspase-8, and caspase-9 gene expression in HeLa cells

Compared with the control group, RTFP treatment significantly upregulated the mRNA expression of pro-apoptotic genes caspase-3, caspase-8, and caspase-9 in HeLa cells in a dose-dependent manner. At a concentration of 3 mg/mL, the relative expression levels of caspase-3, caspase-8, and caspase-9 increased to 1.46±0.11 (P<0.01), 1.23±0.11 (P<0.05), and 1.21±0.11 (P<0.05), respectively. When the concentration was increased to 6 mg/mL, the expression further increased to 2.44±0.17 (P<0.01), 1.90±0.03 (P<0.01), and 2.11±0.27 (P<0.01). These results demonstrate that RTFP effectively activates the apoptosis-related genes caspase-3, caspase-8, and caspase-9, suggesting its potential role in promoting apoptosis in HeLa cells.

RTFP regulates apoptosis-related protein expression

Compared with the control group, RTFP significantly upregulated the expression of pro-apoptotic proteins Bax, caspase-3, caspase-8, and caspase-9 (P <0.05) and downregulated the expression of anti-apoptotic protein Bcl-2 (P<0.01) in a dose-dependent manner (Figures 8,9).

Figure 8 Effect of RTFP on the expression of apoptosis related proteins Bax and Bcl-2 (n=3). (A) Representative Western blot images showing RTFP effect on Bax and Bcl-2 expression in HeLa cells at 0 (control), 3, and 6 mg/mL after 48 h. (B) Quantitative analysis of protein expression changes using ImageJ software. **, P<0.01 vs. control group. Data are presented as mean ± SD. RTFP, Rosa roxburghii Tratt fruit polysaccharides; SD, standard deviation.

Figure 9 Effect of RTFP on the expression of apoptosis related proteins caspase-3, caspase-8, and caspase-9 (n=3). (A) Representative Western blot images showing RTFP effect on caspase-3, caspase-8, and caspase-9 expression in HeLa cells at 0 (control), 3, and 6 mg/mL after 48 h. (B) Quantitative analysis of protein expression changes using ImageJ software. *, P<0.05; **, P<0.01 vs. control group. Data are presented as mean ± SD. RTFP, Rosa roxburghii Tratt fruit polysaccharides; SD, standard deviation.

Discussion

RTFP significantly inhibited the proliferation, migration, and invasion of HeLa cells in a time- and dose-dependent manner. Cell cycle analysis revealed that RTFP induced G2/M phase arrest, accompanied by downregulation of the key regulatory proteins CDK-1 and cyclin B1. The CDK-1/cyclin B1 complex plays a central role in G2/M transition; its inhibition thus blocks cell cycle progression and suppresses uncontrolled tumor cell proliferation (23,24). This mechanism aligns with the antitumor effects of other plant-derived polysaccharides, which interfere with cell cycle progression (2,5). Regarding apoptosis, RTFP upregulated pro-apoptotic proteins (Bax, caspase-3, caspase-8, and caspase-9) while downregulating the anti-apoptotic protein Bcl-2. Caspases are core executors of apoptosis: caspase-8 initiates the extrinsic death receptor pathway, caspase-9 drives the intrinsic mitochondrial pathway, and caspase-3 serves as the key effector that cleaves substrates to execute cell death (25-28). These changes suggest that RTFP activates both intrinsic and extrinsic pathways, increasing the Bax/Bcl-2 ratio to enhance mitochondrial outer membrane permeabilization, trigger caspase-9 activation, and initiate the caspase cascade.

These findings align with the antitumor mechanisms of other polysaccharides. For instance, lentinan edodes polysaccharide induces HeLa cell apoptosis via a mitochondria-dependent pathway, with elevated Bax/Bcl-2 ratio and activated caspase-9 and caspase-3 (9). Similarly, Rosa roxburghii polysaccharide also induces apoptosis in DU145 prostate cancer cells by upregulating caspase-3, caspase-8, caspase-9, and Bax (20), while polysaccharides from Gynura divaricata and Hemerocallis citrina Baroni modulate Bax/Bcl-2 and caspase-3 or induce G2/M arrest via Wnt/β-catenin inhibition (29,30). Collectively, these studies confirm that polysaccharides can inhibit tumor progression by regulating the cell cycle and apoptotic pathways. The present study is among the first to reveal the similar activity of RTFP in a cervical cancer model.

Although this study provides preliminary in vitro evidence of RTFP’s anti-cervical cancer effects and underlying mechanisms, several limitations should be acknowledged. First, a total polysaccharide extract was used. Systematic purification, detailed structural characterization (e.g., molecular weight distribution, monosaccharide composition, and branching structure), batch-to-batch consistency assessment, and comprehensive toxicological evaluation have not yet been performed. Therefore, the possibility of nonspecific effects cannot be ruled out. Second, the findings are confined to in vitro models, lacking validation from in vivo animal studies or clinical data. Thus, the in vivo efficacy, safety profile, pharmacokinetics, and bioavailability of RTFP remain to be evaluated. Finally, the upstream signaling pathways involved and the potential interactions between RTFP and the tumor microenvironment warrant further investigation.

Future studies should: (I) isolating and purifying the active components of RTFP, followed by detailed structural analysis to elucidate structure-activity relationships; (II) verifying the broad applicability and reproducibility of its effects in multiple cervical cancer cell lines, primary cells, and tumor-bearing animal models; (III) evaluating the combination potential of RTFP with existing chemotherapeutic agents to explore synergistic and toxicity-reducing effects; (IV) employing multi-omics approaches (e.g., transcriptomics, proteomics) and network pharmacology to systematically reveal upstream mechanisms and potential targets; and (V) conducting comprehensive​ preclinical toxicological and pharmacodynamic evaluations to support its translational potential.

Conclusions

This study demonstrates that polysaccharides from RTFP effectively inhibit the proliferation, migration, and invasion of human cervical cancer HeLa cells. The underlying mechanisms involve the induction of G2/M phase cell cycle arrest and the activation of related apoptosis pathways. These findings provide preliminary in vitro evidence for the anti-cervical cancer activity of RTFP and lay a foundation for further research. Future studies should focus on purified fractions, in vivo validation, and deeper mechanistic exploration to assess its potential for future development.

Acknowledgments

We would like to thank the laboratory colleagues for their valuable support and assistance in the preparation of this article.

Footnote

Reporting Checklist: The authors have completed the MDAR reporting checklist. Available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-aw-2431/rc

Data Sharing Statement: Available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-aw-2431/dss

Peer Review File: Available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-aw-2431/prf

Funding: This work was supported by the National Natural Science Foundation of China (No. 31860252), the Joint Special Project for General Research of Natural Science Foundation of Local Universities in Yunnan Province (Nos. 202301BA070001-043 and 202401BA070001-089), the Scientific Research Fund of Yunnan Provincial Department of Education (No. 2025J0767), the Open Project of Yunnan Provincial Key Laboratory of Entomological Biopharmaceutical R&D (No. AG2022006), and the Key R&D Program of the Science and Technology Plan in Biomedicine of Dali Bai Autonomous Prefecture (No. 20242903B030014).

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://tcr.amegroups.com/article/view/10.21037/tcr-2025-aw-2431/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved.

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