Ginsenoside F4 inhibits colorectal cancer progression by boosting dendritic cell maturation and remodeling the tumor microenvironment

Abstract

BACKGROUND

Immunotherapy that employs dendritic cells (DCs) to activate the patient’s immune system has emerged as a promising therapeutic strategy to combat cancer; however, effective targeting agents are still limited. Ginsenoside F4, as a rare ginsenoside found in Panax ginseng, exhibits stronger antitumor and immunomodulatory activities than primary ginsenosides. However, its therapeutic effects on various diseases remain limited.

AIM

To investigate the antitumor effect of Ginsenoside F4 and mechanism on the maturation of DCs in colorectal cancer (CRC).

METHODS

The changes in mature DC markers and cytokines generated after DCs were exposed to F4 were assessed using flow cytometry and enzyme-linked immunosorbent assay, respectively. The viability of CRC CT26 cells co-cultured with T lymphocytes was monitored by cell counting kit-8 assay. Furthermore, the histopathological characteristics and immune cell infiltration in tumor tissues of CT26-bearing mice were analyzed by hematoxylin-eosin and immunofluorescent staining. The expressions of apoptosis-relative proteins were detected by western blot assay.

RESULTS

Treatment with F4 promoted the maturation of DCs, elevated the expressions of cluster of differentiation (CD) 83 and CD86, increased the secretion of interleukin (IL)-2, IL-10, and IL-12 p70, and upregulated the expressions of phosphorylated phosphoinositide 3-kinase, phosphorylated protein kinase B, and nuclear factor kappa-B (NF-κB) phosphorylated p65 in DCs, which enhanced antigen-specific CD8+ T-cell responses. However, these benefits could be reversed by the sphingosine-1-phosphate 1 (S1PR1) inhibitor fingolimod hydrochloride. Furthermore, oral administration with F4 inhibited tumor growth and increased DC and CD8+ T-cell infiltration in the tumor tissues of CT26-bearing mice.

CONCLUSION

The results demonstrated that F4 inhibited the growth of CRC by maturing DCs through activating S1PR1-mediated phosphoinositide 3-kinase/protein kinase B and NF-κB pathways, which triggered the antitumor effects of CD8+ T cells. Therefore, F4 could serve as an antitumor immunomodulator for CRC treatment.

Key Words: Colorectal cancer; Cytotoxic T lymphocyte; Dendritic cells; Ginsenosides; Sphingosine-1-phosphate 1; Tumor microenvironment

Core Tip: This study provides novel insights into the antitumor effects of natural ginsenoside F4 on the apoptosis of colorectal cancer (CRC) cells by activating the sphingosine-1-phosphate 1-mediated phosphoinositide 3-kinase/protein kinase B and nuclear factor kappa-B pathways in dendritic cells, which subsequently triggers the antitumor effects of cluster of differentiation (CD)8+ T cells. Further investigation revealed that F4 inhibited tumor growth in CRC-bearing mice by recruiting dendritic cells and CD8+ T cells into the tumor immune microenvironment. These findings underscore the promising role of ginsenoside F4 in the management of CRC, highlighting the potential of natural ginsenosides as an adjuvant treatment for this disease.

INTRODUCTION

Colorectal cancer (CRC) is one of the most prevalent malignant tumors of the gastrointestinal tract and the leading cause of cancer-related deaths globally[1]. Surgery combined with radiotherapy or chemotherapy is the primary treatment for CRC. However, the high heterogeneity and invasiveness of CRC often lead to unsatisfactory postoperative outcomes. The inevitable recurrence and metastasis remain significant challenges of mortality among patients. In recent years, immunotherapy has emerged as an effective strategy to treat malignant tumors and metastases in clinical practice, gradually transforming the approach to CRC treatment[2].

As potent antigen-presenting cells, dendritic cells (DCs) play a crucial role in antitumor response by recognizing tumor-associated antigens and triggering cytotoxic T lymphocyte (CTL) responses against tumor cells. Recently, the combination of DCs with other tumor treatments, such as radiotherapy, chemotherapy, and immune checkpoint inhibitors, has become a feasible strategy to enhance antitumor efficiency[3,4]. However, there are several limitations to tumor immunotherapy. Clinical studies have demonstrated that the impaired function of DCs in cancer patients is strongly associated with the onset and progression of tumors[5,6]. The number of mature DCs (mDCs) in the tumor tissues, lymph nodes, and peripheral blood of patients is significantly reduced, and their ability to present antigens and activate CTL responses is diminished[7]. Moreover, immature DCs (iDCs), which are the predominant subtype in the peritumoral area of CRC, play a significant role in promoting immune tolerance and immunosuppression within the tumor microenvironment (TME)[8]. Consequently, the promotion of DC maturation and CTL activation has become a prominent research focus.

Natural products exhibit a range of immunoregulatory activities and can remodel the tumor-immunosuppressive microenvironment, which can prevent tumors from evading recognition and attack by the immune system, thereby enhancing immunotherapy efficiency. Panax ginseng, a traditional herbal medicine, has antioxidant, anti-inflammatory, and anticancer potential for the prevention of various aging-related diseases, including cardiovascular diseases, metabolic diseases, neurodegenerative diseases, and cancers[9-12]. Its aqueous extracts and the main active components of ginsenosides inhibit tumor growth and metastasis by suppressing angiogenesis and enhancing the host immune response[13-15]. Moreover, it can mitigate chemotherapy-induced myelosuppression, cardiotoxicity, nephrotoxicity, and cancer-related fatigue[16-19]. Rare ginsenosides [e.g., red ginseng (Rg) 3, Rh2, F4, and compound K (CK)], which are low in abundance in Panax ginseng, are intestinal metabolites of primary ginsenosides (e.g., Rb1, Rb2, Re, and Rg1) and exhibit stronger anticancer activity than primary ginsenosides[20]. These rare ginsenosides remarkably inhibit CRC growth in vivo by inactivating the Wnt/β-catenin, CCAAT enhancer binding protein β/nuclear factor kappa-B (NF-κB), and Axl pathways. However, their IC₅₀ values against the proliferation of CRC cell lines in vitro exceed 100 μM, which is five times higher than the threshold for screening anticancer candidates[21-23]. Ginseng extract and rare ginsenosides exhibit potent anticancer effects by suppressing the programmed cell death 1 (PD-1)/programmed cell death ligand 1 (PD-L1) pathway in CRC cells and enhancing CD8+ T-cell infiltration into tumors[24,25]. Ginsenoside metabolites, such as CK, Rh1, and Rg3, exhibit dual roles in DC maturation; in other words, they can significantly inhibit DC activation in inflammatory diseases, but promote it in autoimmune diseases and cancers at low concentrations[26-29].

Therefore, we hypothesized that rare ginsenosides might enhance antitumor CTL responses through their immunoregulatory effects on DC activation. However, the exact underlying mechanisms remain unclear. To validate this hypothesis, we aimed to investigate the anticancer effects of ginsenoside F4, a primary active metabolite derived from the primary ginsenosides in Panax ginseng, on the phenotype and function of DCs and CD8+ T cells in the CRC microenvironment.

MATERIALS AND METHODS

Reagents

Lipopolysaccharides (LPS) were purchased from Sigma-Aldrich (St. Louis, MO, United States). F4 was provided by professor Gao RL, Zhejiang Provincial Traditional Chinese Medicine Hospital (Hangzhou, Zhejiang Province, China), and kept in the specimen chamber (No. 2023Y00165) of the Zhejiang Provincial Laboratory Animal Centre. Capecitabine (CAP) was purchased from Aladdin (Shanghai, China). Fluorescein isothiocyanate (FITC)-cluster of differentiation (CD) 11c, PE-CD83, FITC-CD86, and Brilliant Ultra Violet™ 737 (BUV737)-major histocompatibility complex (MHC) II anti-mouse antibodies were purchased from BD Biosciences (NJ, United States). The cell-staining buffer was purchased from BioLegend (CA, United States). All mouse cytokine enzyme-linked immunosorbent assay (ELISA) kits were purchased from Multisciences (Hangzhou, Zhejiang Province, China). The antibodies, including phosphoinositide 3-kinase (PI3K), phosphorylated PI3K (p-PI3K), protein kinase B (AKT), phosphorylated AKT (p-AKT), NF-κB p65, phosphorylated NF-κB p65 (NF-κB p-p65), and sphingosine-1-phosphate 1 (S1PR1), used for western blot analysis were purchased from Proteintech (Wuhan, Hubei Province, China).

Isolation and maturation of DCs

Murine bone marrow-derived DCs (BMDCs) were isolated as previously reported[30]. Briefly, the bone marrow obtained from the femurs and tibiae of C57BL/6J mice was washed with phosphate-buffered saline (PBS) to prepare the cell suspension. After centrifugation at 2000 rpm for 5 minutes, the cell precipitate was resuspended in Roswell Park Memorial Institute 1640 medium (Gibco, MA, United States) containing 1000 U/mL granulocyte-macrophage colony-stimulating factor (GM-CSF), 800 U/mL interleukin (IL)-4, and 10% fetal bovine serum. CD11c-positive BMDCs were purified using immunomagnetic beads and their purity was determined using a flow cytometer (BD Biosciences). Isolated BMDCs were stimulated with GM-CSF (1000 U/mL) and IL-4 (800 U/mL) for 48 hours and then incubated with LPS (10 μg/mL) as a positive control or F4 (10, 25, and 50 μg/mL) for 24 hours. Cell viability was detected using the cell counting kit-8 (CCK8) assay and the expression of the DC biomarkers CD83 and CD86 was detected using flow cytometry (BD, United States). The shapes of DCs were imaged using a light microscope (Olympus, Japan).

Co-culture of CT26 cells and DCs

CD8+ T lymphocytes derived from the spleen of C57BL/6J mice were prepared by passing the dispersed cells over nylon wool columns (Xiangbo Biotech Co., Guangzhou, Guangdong Province, China) and purified using magnetic beads as previously reported[31,32]. Mouse CRC CT26 cells were obtained from Zhejiang Provincial Laboratory Animal Centre, Linan, China. These cells were frozen and lysed to obtain tumor antigens, and then mixed with BMDCs that had been pretreated with F4 for 72 hours. The ratio of F4-pretreated DCs to tumor antigens was maintained at 1:10. The sensitized DCs were then mixed with CD8+ T lymphocytes at ratios of 1:5, 1:10, 1:20, and 1:40 in 96-well plates for 24 hours. Mixtures of DCs and CD8+ T cells were used as controls. Cell viability in the co-culture system was detected using the CCK8 assay to calculate the proliferation rate of T cells. In addition, to investigate the specific killing of DC-induced CTL response, sensitized CD8+ T lymphocytes were mixed with CT26 cells at ratios of 10:1, 20:1, and 50:1, respectively, in 96-well plates for 24 hours. Mixtures of CT26 and CD8+ T cells were used as controls. Cell viability in the co-culture system was detected using the CCK8 assay. After 48 hours of incubation, 10⁵ CTL and 10⁴ CT26 cells in each well were mixed and incubated at 37 °C for 24 hours. The apoptotic rate of CT26 cells was determined by flow cytometry (Agilent, United States).

Animals and animal treatment

Balb/c male mice (8 weeks old, body weight 22-25 g) were purchased from Shanghai Slaughter Laboratory Animals Co. Ltd., China. All mice were kept under a temperature of 23 ± 2 °C, humidity of 40%-60%, and 12-hour light/dark cycle conditions in a specific pathogen-free laboratory at the Zhejiang Provincial Laboratory Animal Centre. The experiments (No. 2023R001099) were approved by the Animal Ethics Committee of Zhejiang Provincial Laboratory Animal Center.

CT26 cells in the logarithmic growth phase were harvested and diluted in serum-free Dulbecco’s modified eagle medium (DMEM) to a concentration of 1 × 10⁷/mL. Cell suspensions were inoculated subcutaneously into the axilla of the right forelimb of the mice (0.2 mL/mouse). After 7 days, mice (n = 25) with a tumor volume of 100 mm³ were selected and randomly divided into five groups: Control group (saline), CAP-treated group (500 mg/kg), F4-L group (treated with 25 mg/kg of F4), F4-M group (treated with 50 mg/kg of F4), and F4-H group (treated with 100 mg/kg of F4). The dose of F4 treatment in the in vivo experiment was chosen as per previous studies[22,24,33]. The chemical compound F4 was dissolved in saline and orally administered daily for 7 days. The tumor volume and body weight were recorded daily. All mice were sacrificed by carbon dioxide asphyxia 7 days after drug administration.

Flow cytometry assay

BMDCs obtained from the co-culture system were resuspended in cell-staining buffer and the cell concentration was adjusted to 1 × 10⁶ cells/mL. The cell suspension (300 μL) was then incubated with PE-CD83, FITC-CD86, or FITC-CD11c antibodies at 4 °C for 30 minutes in the dark, and centrifugated at 2000 rpm for 20 minutes to discard the supernatant. The cell suspension was washed twice with PBS and resuspended in 500 μL of cell-staining buffer. Similarly, the anticoagulated blood obtained from each tumor-bearing mouse was treated with erythrocyte lysate at room temperature for 15 minutes in the dark, centrifuged at 2000 rpm for 5 minutes to discard the supernatant, and incubated with PE-CD83, FITC-CD86, or FITC-CD11c antibodies.

Half of the tumor tissue was sheared in 2 mL of digestion solution (DMEM containing 1 mg/mL collagenase I, 20 μg/mL DNAzyme I, and 5% fetal bovine serum) and digested for 30 minutes at 37 °C. Digestion was terminated with DMEM containing 5% fetal bovine serum. The cell suspension was filtered using a 40-μm cell strainer and centrifuged at 2500 rpm for 5 minutes. The cell precipitates were mixed with 1 mL of cell-staining buffer and used to resuspend the cells. Next, 100 μL of the suspension was incubated with 1 μL of FITC-CD11c and BUV737-MHC II at 4 °C for 30 minutes in the dark and centrifuged at 2000 rpm for 5 minutes to discard the supernatant. The levels of mDC surface biomarkers in all cell samples were detected using a FACSCalibur flow cytometer (BD Biosciences, United States).

ELISA assay

Culture supernatants were collected from each group of BMDC after treatment with appropriate drugs. The levels of IL-2, IL-10, and IL-12 p70 in the culture supernatants were detected using ELISA kits according to the manufacturer’s instructions.

Quantitative real-time-polymerase chain reaction assay

A quantitative real-time-polymerase chain reaction (qRT-PCR) was performed according to the manufacturer’s instructions. Briefly, total RNA was extracted using an RNA extraction kit (Qiagen, Hilden, Germany). Total RNA was reverse-transcribed into complementary DNA using HiScript II Q RT SuperMix for qPCR (Vazyme, Nanjing, Jiangsu province, China). Complementary DNA was extracted and amplified using Taq Pro Universal SYBR qPCR Master Mix (Vazyme, Nanjing, Jiangsu province, China), and evaluated using the CFX Connect™ Fluorescent Quantitative PCR System (BIO-RAD, United States). β-actin was used as an internal reference for message RNA (mRNA) expression analysis. Primers were designed and synthesized by Sangon Biotech Co. Ltd. (Shanghai, China) as shown in Table 1.

Table 1 Sequences of primers used for the real-time-polymerase chain reaction assay.

Gene	Primer sequences (5’ to 3’)
β-actin	F: GGCTGTATTCCCCTCCATCG
	R: CCAGTTGGTAACAATGCCATGT
Bcl-2	F: CCATGTGTCCATCTGAC
	R: GCCATATAGTTCCACAA
Bax	F: TGAAGACAGGGGCCTTTTTG
	R: AATTCGCCGGAGACACTCG
Caspase-3	F: TGGTGATGAAGGGGTCATTTATG
	R: TTCGGCTTTCCAGTCAGACTC
IL-12 p70	F: ATGTGGAATGGCGTCTC
	R: GTCTCCTCGGCAGTTGG
IL-12R	F: CACCCCAGTGAACTACCTT
	R: GGAGTCATAGCCCATCTTT
STAT4	F: AAACCTGAGCCCAACGA
	R: CAAAGCCGAAGAAATAA

IL-12R: Interleukin-12 receptor.

Western blot assay

Protein lysates from tumor tissues were prepared using radio immunoprecipitation assay lysis buffer. The total protein concentration was determined using a bicinchoninic acid assay quantification kit. Samples were denatured in 5 × loading buffer for 5 minutes at 100 °C. Ten micrograms of protein samples were separated by 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis. Subsequently, the proteins on the gel were transferred to polyvinylidene fluoride membranes. The membranes were blocked with 5% skim milk powder at room temperature for 1.5 hours. The cells were incubated with the primary antibody at 4 °C overnight. The membrane was then incubated with secondary antibody for 1 hour at room temperature. Finally, the target proteins were detected using an ImageQuant 800 system (GE, United States), with β-actin serving as an internal reference.

Immunohistochemistry

Paraffin-embedded sections of tumor tissue were dewaxed and rehydrated. The tissue sections were then placed in a repair cassette containing citric acid antigen repair buffer [potential of hydrogen (pH) = 6.0] and heated in a microwave oven for antigen retrieval. Endogenous peroxidase activity was blocked using a 3% hydrogen peroxide solution. The samples were subsequently blocked with 3% bovine serum albumin at 37 °C for 30 minutes. The samples were then incubated with the primary antibody at 4 °C overnight. The samples were then treated with a reaction enhancement solution at room temperature for 30 minutes, followed by incubation with the secondary antibody at room temperature for an additional 30 minutes. The samples were stained with diaminobenzidine and the nuclei were re-strained. Finally, the sections were dehydrated, sealed, and subjected to microscopic examination and image collection using the SlideView VS200 system (Olympus, Japan).

Immunofluorescence

Frozen sections of tumor tissue were dewaxed and rehydrated. The tissue sections were then placed in a repair cassette containing citric acid antigen repair buffer (pH = 6.0) and placed in a microwave oven for antigen repair. Endogenous peroxidase was blocked with 3% hydrogen peroxide solution. The samples were blocked with 3% bovine serum albumin at 37 °C for 30 minutes. The samples were incubated with the primary antibody at 4 °C overnight. The samples were incubated with the reaction enhancement solution at room temperature for 30 minutes and then with the secondary antibody at room temperature in the dark for 1 hour. The nuclei were re-strained with 4’,6-diamidino-2-phenylindole. An anti-fluorescence burst sealer was added dropwise to the tissues and the sections were sealed and imaged using a SlideView VS200 system (Olympus, Japan).

mRNA sequencing and molecular docking analysis

Total RNA was extracted using TRIzol reagent (Invitrogen, United States), fragmented into 100-200 nt pieces, and prepared into an mRNA-Seq library, followed by 2 × 150 bp paired-end sequencing on an Illumina NovaSeq 2000. The profiles of differentially expressed mRNAs (fold change > 1.5, P < 0.05) were analyzed using Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) enrichment to investigate the regulatory mechanism of F4 in DC maturation.

The potential targets of F4 were predicted using the SuperPred and SwissTarget prediction systems according to our previously reported method[34-36]. The genes involved in cytokine–cytokine receptor interactions, viral protein interactions with cytokines and cytokine receptors, hematopoietic cell lineage, chemokine signaling pathways, and pathways in cancer (which were primarily displayed in KEGG enrichment) were prioritized. Reverse docking identified an interaction between the target and saponin with a binding energy of less than -7.0 kcal/mol. Molecular docking was conducted using the AutoDock tool and the results were visualized using PyMOL software.

F4 solutions (0.625-10 μM) were prepared in PBS and analyzed using a Biacore 100 T biosensor detector (GE Healthcare, United States) with a binding time of 120 seconds and flow rate of 20 μL/minute. The affinity constants (K_D) were determined by computerized fitting analysis.

Statistical analysis

All data are presented as the mean ± SD. GraphPad Prism 10 software was used to analyze the results. One-way analysis of variance analysis and least significant difference test were used for comparison between groups, and t-test was used for comparison between the two groups. P values of less than 0.05 were considered significant for all data sets.

RESULTS

F4 promoted the proliferation and maturation of BMDCs

As shown in Figure 1A, the IC₅₀ value of F4 for the proliferation of CT26 cells was 97.1 μg/mL, indicating weak anticancer activity. Then, DC was identified by flow cytometry, whose purity was 96% and could be employed for further investigation (Figure 1B). Treatment with F4 at the concentrations of 10-150 μg/mL facilitated DC proliferation in vitro (Figure 1C). Unlike iDCs, which exhibited a round shape, F4-treated DCs extended numerous pseudopodia as the concentration increased (Figure 1D). Thus, treatment with F4 at concentrations of 10-50 μg/mL, which did not affect the viability of CT26 cells, but promoted the proliferation and maturation of DCs in vitro, was employed for further investigation of DC maturation. Furthermore, treatment with F4 upregulated the expression of mDC biomarkers CD83 and CD86 and enhanced the release of IL-2, IL-10, and IL-12 p70 in mDCs (Figure 1E and F). Treatment with 10 μg/mL LPS and 50 μg/mL F4 exhibited similar effects on BMDC maturation. These results indicate that F4 promoted the proliferation and maturation of BMDCs in vitro.

Figure 1 F4 promoted the maturation of bone marrow-derived dendritic cells in vitro. A: Cytotoxicity of F4 on the proliferation of mouse colon CT26 cells; B: The purity (representative data) of bone marrow-derived dendritic cells (BMDCs) identified by flow cytometry; C: Cytotoxicity of F4 on the proliferation of BMDCs; D: Effects of F4 on the morphological change of BMDCs; E: Effects of F4 on the expressions of DC surface biomarkers cluster of differentiation (CD) 83 and CD86; F: Effects of F4 on the release of cytokines in the culture mediums. All data were shown as mean ± SD (n = 5). ^aP < 0.05. ¹P vs control group. ²P vs lipopolysaccharides group. Ctrl: Control; 5-FU: 5-fluorouracil; CD: Cluster of differentiation; SSC-H: Side scatter high; BMDCs: Bone marrow-derived dendritic cells; LPS: Lipopolysaccharides; IL: Interleukin.

Cytotoxicity of CD8+ T cells against the proliferation of CT26 cells

Compared to the control group, the number of CD8+ T cells cultured with DCs at ratios of 1:5, 1:10, 1:20, and 1:40 sharply increased (Figure 2A). Among the four co-culture groups, the highest proliferative ratios were observed in the CD8+ T/DC cell ratios of 1:5 and 1:10, without a significant difference. As a higher proliferation rate correlated with an increased number of CD8+ T cells, our results indicate that mDCs stimulated by F4 could promote the proliferation of CD8+ T cells in vitro. To manage the costs of the in vitro experiments, CD8+ T cells were mixed with DCs at a 1:5 ratio for further investigation. Additionally, these sensitized CD8+ T cells promoted CT26 cell death in a concentration-dependent manner. As the ratio of CD8+ T cells to CT26 cells increased from 10:1 to 50:1, the apoptosis of CT26 cells, along with the mRNA expression of Bax and caspase-3, significantly decreased, while the mRNA expression of Bcl-2 increased (Figure 2B-E). These results suggested that DCs cultured with F4 enhanced the cytotoxicity of CD8+ T cells against the proliferation of CT26 cells in vitro.

Figure 2 Dendritic cells cultured with F4 enhanced the cytotoxicity of cluster of differentiation 8 + T cells against CT26 cells. A: Effects of F4-stimulated dendritic cells (DCs) on the proliferation of T cells; B: Cytotoxicity of tumor-specific cytotoxic T lymphocyte (CTL) responses against CT26 cells by F4-treated DCs; C and D: DC-stimulated cluster of differentiation 8 + T cells induced the apoptosis of CT26 cells; E: The relative levels of Bcl-2, Bax and caspase-3 message RNA in CTL-stimulated CT26 cells. All data were shown as mean ± SD (n = 5). ^aP < 0.05. ¹P vs control group. ²P vs lipopolysaccharides group. Ctrl: Control; LPS: Lipopolysaccharides; PI: Propidium iodide.

F4 promoted the maturation of DCs via targeting S1PR1

To investigate the immunoregulation of F4, we analyzed the mRNA profiles of DCs treated with or without F4 using transcriptome sequencing. A total of 97 mRNAs were significantly upregulated, whereas 103 were significantly downregulated (Figure 3A). These 200 differentially expressed mRNAs were enriched in biological functions associated with GO terms such as inflammatory and immune responses. They were also mainly involved in the regulation of cytokine-cytokine receptor interactions, viral protein interactions with cytokines and cytokine receptors, hematopoietic cell lineage, chemokine signaling pathways, and pathways in cancer, as indicated by KEGG enrichment analysis. Five common targets of F4, predicted by the SuperPred and SwissTarget systems (Figure 3B), all ranked within the top 10 signaling pathways identified in the KEGG enrichment analysis. Among these five targets, the affinity between S1PR1 and F4 was lowest at -7.60 kcal/mol. F4 primarily interacted with the TRP117, LYS34, and PHE291 sites of S1PR1 through conventional hydrogen bonds as well as with various other sites via alkyl and π-alkyl interactions (Figure 3C). The results of the surface plasmon resonance (SPR) assay also showed that the K_D value for the S1PR1-F4 interaction was 3.81 μM (Figure 3D). Furthermore, the expression of S1PR1 in the F4-treated group at different temperatures was higher than that in the control group, suggesting that F4 binds S1PR1 to form a tight complex, thereby enhancing the thermostability of S1PR1 (Figure 3E). Collectively, these findings indicated that S1PR1 is the primary target of F4.

Figure 3 The regulation mechanism of F4 on the maturation of dendritic cells. A: The volcano plot of differently expressed genes in dendritic cells treated with F4 or saline. The pathways of the involved differently expressed genes were enriched by Gene Ontology and Kyoto Encyclopedia of Genes and Genomes analysis; B: The targets of F4 were predicted by SuperPred and SwissTargetPrediction web-tools. The binding energies among targets and F4 were detected by using AutoDock tool; C: The interaction between target and F4 was evaluated by using AutoDock tool; D: The schematic diagram of interaction between sphingosine-1-phosphate 1 (S1PR1) and F4; E: The interaction between S1PR1 and F4 was detected by surface plasmon resonance assay and verified by thermostability assay. FC: Fold change; TNF: Tumor necrosis factor; GO: Gene Ontology; LPS: Lipopolysaccharides; ERK: Extracellular regulated protein kinases; JAK: Janus kinase; STAT: Signal transducer and activator of transcription; PI3K: Phosphatidylinositol 3-kinase; AKT: Protein kinase B; S1PR1: Sphingosine-1-phosphate 1.

F4 promoted DC maturation via activation of PI3K/AKT and NF-κB pathways

As mentioned, F4 treatment accelerated the proliferation of DCs, increased the expression levels of CD83 and CD86, and enhanced the production of IL-2, IL-10, and IL-12 p70. However, the effects of F4 on DC maturation were reversed by the S1PR1 inhibitor fingolimod hydrochloride (FTY720) in a concentration-dependent manner (Figure 4A-D). Exposure to FTY720 did not affect the proliferation and maturation of DCs, indicating a specific effect on DC maturation. Furthermore, treatment with F4 upregulated the expression of p-PI3K, p-AKT, and phosphorylated NF-κB p65 (NF-κB p-p65) in DCs, which was reversed by FTY720 in a concentration-dependent manner (Figure 4E).

Figure 4 F4 promoted the maturation of dendritic cells via activation of phosphoinositide 3-kinase/protein kinase B and nuclear factor kappa-B pathways. A and B: F4 promoted the maturation of dendritic cells (DCs) and the production of inflammatory cytokines, which could be reversed by sphingosine-1-phosphate 1 (S1PR1) inhibitor fingolimod hydrochloride (FTY720); C and D: F4-stimulated DCs enhanced cytotoxic T lymphocyte response against the proliferation of CT26 cells, which could be depleted by S1PR1 inhibitor FTY720; E: F4 upregulated the expressions of phosphorylated phosphatidylinositol 3-kinase, phosphorylated protein kinase B and nuclear factor kappa-B phosphorylated-p65 in DCs, which could be reversed by S1PR1 inhibitor FTY720. All data were shown as mean ± SD (n = 5). ^aP < 0.05. ¹P vs control group (0 μg/mL F4 + 0 μM fingolimod hydrochloride). ²P vs 50 μg/mL F4-treated group. CD: Cluster of differentiation; IL: Interleukin; FTY720: Fingolimod hydrochloride; mRNA: Message RNA; PI3K: Phosphatidylinositol 3-kinase; p-PI3K: Phosphorylated phosphatidylinositol 3-kinase; AKT: Protein kinase B; p-AKT: Phosphorylated protein kinase B; NF-κB: Nuclear factor kappa-B.

F4 inhibited tumor growth and increased DC and CD8+ T-cell infiltration in CT26-bearing mice

Oral administration of F4 significantly inhibited tumor growth in CT26-bearing mice (Figure 5A). Notably, oral administration of F4 at a dose of 100 mg/kg and CAP at 500 mg/kg resulted in comparable inhibition of tumor size, highlighting the potent anticancer activity of F4. The in vitro results showed that treatment with F4 not only dose-dependently increased the counts of DCs and effective CD8+ (eCD8+) T cells in the peripheral blood, but also elevated the expression of CD83 and CD86 in DCs, as well as the serum levels of IL-2, interferon (IFN)-γ, and IL-12 p70 (Figure 5B and C). However, it could not impact the peripheral proportions of native CD8+ (nCD8+) T cells and memory CD8+ (mCD8+) T cells. These findings suggested that F4 promotes DC maturation, thereby facilitating the transformation of CD8+ T cells into e CD8+ T cells.

Figure 5 Effects of F4 on the tumor growth of CT26-bearing mice. A: Inhibition of F4 on the tumor size and tumor/body weight ratio in CT26-bearing mice; B: The landscapes of peripheral immune cells detected by using 15-color spectral flow cytometry; C: The levels of interferon-γ, interleukin-2, and interleukin-12 p70 in the serum detected by enzyme-linked immunosorbent assay. All data were shown as mean ± SD (n = 5). ^aP < 0.05. P vs model group. Ctrl: Control; CAP: Capecitabine; L: Low; M: Middle; H: High; CD: Cluster of differentiation; eCD: Effective cluster of differentiation; mCD: Memory cluster of differentiation; nCD: Native cluster of differentiation; FSC-H: Forward scatter height; FSC-A: Forward scatter area; APC: Allophycocyanin; BV: Brilliant violet; UV: Ultraviolet; PE: Phycoerythrin; SSC-A: Side scatter area; PE-A: Phycoerythrin area; DC: Dendritic cells; IFN: Interferon; IL: Interleukin.

Tumor cells in the model group were tightly aligned, exhibited larger nuclei, and showed high Ki67 protein expression. In contrast, the tumor cells in the F4-treated groups were loosely packed, had smaller nuclei and thinner cytoplasm, and displayed lower levels of Ki67 expression (Figure 6A and B). Additionally, few DCs and CD8+ T cells were observed in the tumor tissues of the model group, whereas the number of immune cells in the tumor tissues of the F4-treated groups was remarkably elevated in a dose-dependent manner, indicating a high level of immune cell infiltration (Figure 6C-E). Similarly, the populations of DCs and CD8+ T cells in the tumor tissues of the F4-treated groups were 5-8 times higher than those in the model group (Figure 7A and B). Compared with the model group, the expression of IFN in CD8+ T cells were markedly increased in a dose-dependent manner, suggesting the activation of CD8+ T cells. Owing to the limited number of CD8+ T-cell subtypes in mouse tumor tissues and the large error produced by flow cytometry, the proportion changes in eCD8+, nCD8+, and mCD8+ T cells were not obtained, and the subtype transformation of CD8+ T cells in tumor tissues after F4 intervention should be further verified. Furthermore, Bax and cleaved caspase-3 expression were barely detectable in the model group, whereas these proteins were upregulated in F4-treated groups; the expression of Bcl-2 exhibited the opposite trend (Figure 7C). These findings demonstrated that F4 effectively stimulated DCs in tumors to promote CD8+ T-cell-mediated antitumor immunity (Figure 8).

Figure 6 Effects of F4 on the immune cell infiltration in the tumor tissues of colon cancer CT26-bearing mice. A: Histological change of tumor tissues; B: Representative immunohistochemical images of Ki67 protein expressions in the tumor tissue; C: Representative immunofluorescent images indicated the infiltration of dendritic cells (DCs); D: Cluster of differentiation (CD) 8+ T cells in the tumor tissues. DCs were stained with Texas red-labeled CD11c antibody (red), CD8+ T cells were stained with Texas red-labeled CD8 antibody (red), and nuclei was stained with 4’,6-diamidino-2-phenylindole (blue); E: Quantification of DC or CD8+ T-cell infiltration was visualized by using three-dimensional Gaussian density analysis. All data were shown as mean ± SD (n = 5). ^aP < 0.05. P vs model group. Ctrl: Control; CAP: Capecitabine; L: Low; M: Middle; H: High; DC: Dendritic cells; CD: Cluster of differentiation; MFI: Mean fluorescence intensity.

Figure 7 F4-enhanced cytotoxic T lymphocyte induced the apoptosis of CT26 cells. A: The proportions of dendritic cells in the tumor tissues, which were detected by flow cytometry; B: The proportions of cluster of differentiation (CD) 8+ T cells in the tumor tissues, and the interferon-γ expression in the CD8+ T cells, which were detected by flow cytometry; C: The expressions of apoptosis-related proteins in the tumor tissues of CT26-bearing mice. All data were shown as mean ± SD (n = 5). ^aP < 0.05. P vs model group. MHC: Major histocompatibility complex; Ctrl: Control; CD: Cluster of differentiation; CAP: Capecitabine; L: Low; M: Middle; H: High; LPS: Lipopolysaccharides; SSC: Side scatter; IFN: Interferon.

Figure 8 Schematic diagram of antitumor effects of ginsenoside F4 on colorectal cancer progression by boosting dendritic cell maturation and remodeling the tumor microenvironment. S1PR1: Sphingosine-1-phosphate 1; CTL: Cytotoxic T lymphocyte; CD: Cluster of differentiation; IL: Interleukin.

DISCUSSION

Ginsenosides are the main active components of plants of the Panax genus and can be isolated and purified relatively easily. These saponins primarily include protopanaxadiol saponins (Rb1, Rb2, Rb3, and Rc) and protopanaxatriol saponins (Re, Rg1, and Rg2)[37]. Low-polarity minor ginsenosides, which are derived from partial glycosylation or structural modifications of these native saponins by the gut microbiota, including diol ginsenosides (CK, Rg3, Rg5, Rk1, and Rh2) and triol ginsenosides (Rg2, Rg6, F4, Rh1, Rh4, and Rk3), can be found in wild ginseng, red ginseng, and ginseng fruit[38]. However, their concentrations in ginseng plants are generally low and there are limited research reports on low-polarity ginsenosides. Recent studies have highlighted the more pronounced pharmacological effects of these low-polarity minor ginsenosides, including antitumor, immunoregulatory, and antiviral properties, compared to the primary ginsenosides[9-12,39]. In this study, we investigated for the first time the novel anticancer mechanism of F4, the primary active compound in Painengda capsules, on the enhancement of DC-mediated antitumor T-cell response via activation of S1PR1-mediated PI3K/AKT and NF-κB pathways.

Ginsenoside F4 is a rare saponin found in steamed notoginseng and Rg. It possesses potent antidiabetic, anti-inflammatory, and antileukemic properties[40-43]. However, the anticancer effects of F4 in CRC remain unclear. In this study, although F4 inhibited CT26 cell proliferation, the IC₅₀ value of F4 against the viability of CT26 cells was nearly 100 μg/mL, which is consistent with previous studies on other triol ginsenosides such as Rg2 and Rh4[44,45]. Given that the China Food and Drug Administration officially recognizes an IC₅₀ value of less than 20 μg/mL as significant[46], these results suggest that F4 does not exhibit substantial cytotoxicity against CRC cell proliferation. Notably, F4 treatment inhibited tumor growth and size in CT26-bearing mice. Since dammarane ginsenosides are a class of natural products with the hormone-like activity of triterpenoid saponins[47], we hypothesized that F4 exerts therapeutic effects by stimulating the immune system rather than directly inhibiting tumor cell proliferation.

As the “soil” for tumor growth, the TME is the ecological niche that provides the stromal environment for tumor development[48]. Although a variety of immune cells are present in this environment, functional abnormalities not only diminish the antitumor response, but also promote the rapid growth, invasion, and metastasis of tumor cells. The abnormal or defective function of DCs within the TME represents a significant barrier to effective antitumor efficacy. DCs are the most potent antigen-presenting cells identified to date and are capable of activating naive T cells. DC maturation is a specialized process triggered by pathogen-associated or damage-associated molecular patterns, as well as inflammatory cytokines (e.g., IFN-γ, tumor necrosis factor-α). During maturation, DCs upregulate costimulatory molecules (CD80/CD86, CD40) and MHC class II, switch chemokine receptors (CCR5/CCR7), and extend dendrites to enhance T-cell interaction. This process enables DCs to migrate to lymph nodes, present antigens, and prime naive T cells (Th1/Th2/Th17) or regulate immune tolerance key steps in initiating adaptive immunity[49]. Inflammation is a nonspecific response to infection or injury, driven by cytokines (e.g., IL-1β, IL-6) and characterized by vascular changes (edema, vasodilation) and immune cell recruitment (neutrophils, macrophages). Unlike DC maturation, inflammation focuses on rapid pathogen clearance via phagocytosis, oxidative bursts, and acute-phase proteins, often resolving without engaging adaptive immunity[50]. While inflammation can support DC maturation (e.g., via tumor necrosis factor-α), its primary role is localized tissue defense rather than antigen-specific T-cell activation[51]. During the progression of CRC, the impaired function of DCs results in their inability to effectively present tumor antigens and leads to low expression of costimulatory molecules, such as CD83 and CD86, as well as adhesion molecules. This dysfunction ultimately hinders the effective activation of T cells and facilitates tumor immune escape[52]. Our results demonstrated that F4 treatment promoted the proliferation of DCs in a concentration-dependent manner, increased the expression of the mDC surface biomarkers CD83 and CD86, and elevated the secretion of IL-2, IL-10, and IL-12 p70, indicating the great potential of F4 in DC maturation. Similar findings were observed in animal experiments, where few DCs and CD8+ T cells were detected in the peripheral blood and tumor tissues of CT26-bearing mice. However, the number of activated DCs and IFN-γ + CD8+ T cells increased dramatically as F4 dose increased. Compared with the model group, the expression levels of Bax and cleaved caspase-3 in the F4-treated groups also sharply declined in a dose-dependent manner. These results demonstrated that F4-treated DC enhanced CD8+ T-cell-mediated antitumor responses, subsequently inducing apoptosis in CT26 cells.

S1PR1, also known as endothelial differentiation gene 1, is a G-protein-coupled receptor for sphingosine-1-phosphate. It is primarily expressed in vascular endothelial cells and various immune cells, including DCs, macrophages, and T cells, and plays a pivotal role in tumorigenesis and immune surveillance[53]. S1PR1 signaling activates several downstream effectors including PI3K/AKT, extracellular regulated protein kinase 1/2, NF-κB, and signal transducer and activator of transcription 3[54]. Oral administration of the S1PR1 agonist SEW2871 significantly increased the number of circulating DCs, whereas the S1PR1 inhibitor FYT720 diminished endothelial migration of iDCs[55]. Therefore, S1PR1 has emerged as a potential target for DC-centric immunotherapies. In this study, we identified S1PR1 as the primary F4 target that enhances DC maturation. F4 interacts with the active sites of S1PR1 via various conventional hydrogen bonds and π-alkyl interactions. The interaction between F4 and S1PR1 was further confirmed by SPR and thermostability assays. Additionally, treatment with F4 not only increased the expression levels of CD83 and CD86, as well as the production of inflammatory cytokines, but also upregulated the expression of p-PI3K, p-AKT, and NF-κB p-p65 in DCs, all of which could be reversed by FYT720. Collectively, these findings suggest that F4 promotes the maturation of DCs through the activation of S1PR1-mediated PI3K/AKT and NF-κB signaling pathways. Despite extensive previous research indicating that certain saponins exhibit glucocorticoid-like effects[47], the oral administration of F4 in this study did not elicit significant adverse effects, demonstrating its favorable safety profile. However, its potential side effects require further comprehensive investigation.

In this study, the CT26 colon cancer cell-bearing C57 mouse model was utilized, which can simulate an intact immune system in the body. However, the murine immune system significantly differs from humans in lymphocyte subsets (e.g., regulatory T cells ratios), cytokine networks, and checkpoint expression (e.g., PD-1/PD-L1 interactions); furthermore, the simplified TME lacks the genetic heterogeneity and immune complexity of human CRC, potentially overestimating drug efficacy[56-58]. These limitations highlight the need for cautious interpretation of CT26 data when translating findings to human trials. Recently, humanized immune models, neoantigen-rich systems, and patient-derived tumor organoid models are employed to predict immune-related adverse events and ultimately improve patient outcomes. Therefore, integrating these humanized models with multi-omics approaches will allow more accurate assessment of immunotherapy outcomes and reveal new therapeutic mechanisms.

CONCLUSION

In this study, our results revealed for the first time that F4 exhibited antitumor effects by enhancing the DC-mediated antitumor T-cell response, leading to apoptosis of CRC cells, rather than by directly inhibiting CRC cell proliferation. Mechanistically, F4 promoted the maturation of iDCs by activating S1PR1-mediated PI3K/AKT and NF-κB signaling pathways, thereby inducing robust responses from sensitized CD8+ T cells to exert antitumor effects against CRC progression. Therefore, F4 has the potential to be developed as a potent antitumor immunomodulator or adjuvant for DC vaccines that target CRC.

ACKNOWLEDGEMENTS

We acknowledge the technical supports of the Shared Instrumentation Core Facility of the Hangzhou Institute of Medicine, Chinese Academy of Sciences.