Human prostate cancer (PC) cell lines PC3 and DU145 were obtained from the Shanghai Cell Bank, Type Culture Collection Center, Chinese Academy of Sciences. Cells were cultured in RPMI-1640 medium (HyClone, USA) supplemented with 10% fetal bovine serum (HyClone, USA) and 100 units/ml penicillin-streptomycin, and maintained in an incubator at 37°C with 5% CO2. Ginsenoside Rh2 (GRh2), mitochondrial division inhibitor 1 (Mdivi-1), and Ferrostatin-1 (Fer-1, 2 μM, purity >98%) were purchased from MedChemExpress (MCE, USA).

PC cells were cultured in 96-well plates, and cell viability was assessed using the CCK-8 kit (Solarbio, China). After incubation with CCK-8 reagent for 2 hours, absorbance at 450 nm was measured using a microplate reader (Infinite F50, Switzerland).

The colony formation assay reflects cell proliferation capacity and population dependency. PC3 and DU145 cells in the logarithmic growth phase were trypsinized, counted, and seeded at 1,000 cells per plate in normal medium, GRh2-containing medium, or GRh2 + Mdivi-1-containing medium. The cells were cultured in a 37°C incubator with 5% CO2 for 2–3 weeks until visible colonies formed. After incubation, cells were washed with PBS three times and fixed with 5 mL of 4% paraformaldehyde for 15 minutes. Following fixation, the paraformaldehyde was removed, and the cells were stained with Giemsa stain for 10–30 minutes. After gently washing with PBS and air drying, the plates were inverted, and images were taken to count the number of colonies. The colony formation rate was calculated as:

Matrigel (Thermo, USA) and RPMI-1640 (Gibco, USA) were mixed in a 1:4 ratio, and 60 μL of the mixture was added to the upper chamber of a Transwell chamber (8 μm pore size, Millipore, USA). The chamber was incubated at 37°C for 1 hour to allow Matrigel to solidify. Cells were then treated with GRh2 and GRh2 + Mdivi-1 for 48 hours. A total of 1.0 × 10⁴ cells were seeded in 100 μL of medium in the lower chamber, subjected to serum chemotaxis. After the treatment, 4% paraformaldehyde was removed and 600 μL of it was added to a new well of a 24-well plate. The Transwell chamber was then placed into this new well for 20 minutes, followed by 5 minutes in a fresh well. Cells were fixed and stained with 0.1% crystal violet. The stained membranes were examined under an inverted microscope.

A wound healing assay was conducted to assess the migratory ability of PC3 and DU145 cells. Cells were cultured in 6-well plates and allowed to adhere until they reached full confluence. Afterward, they were treated with normal, GRh2-containing, and GRh2 + Mdivi-1-containing complete media. A scratch was made in the cell monolayer using a 10 μL pipette tip. The cells were rinsed three times with PBS and cultured in RPMI-1640. Images were taken at pre-labeled locations immediately after the scratch and again at 24 and 48 hours. The wound area was quantified using the Image-Pro image analysis system.

Mitochondrial membrane potential changes were assessed using JC-1 (Beyotime, China) according to the manufacturer’s instructions. PC3 and DU145 cells were seeded in a six-well plate. After aspirating the culture medium, the cells were washed once with PBS and then replenished with fresh culture medium. The JC-1 staining working solution was added and thoroughly mixed. The cells were incubated at 37°C for 20 minutes, after which the supernatant was aspirated, and the cells were washed twice with JC-1 staining buffer. Fresh culture medium was added, and the cells were observed under a fluorescence microscope (Nikon Eclipse 80i, Japan).

BBcellProbe (BestBio, China) was used to measure the mitochondrial reactive oxygen species (MtROS) content in PC3 and DU145 cells. Following the product instructions, the staining working solution was prepared by diluting the BBcellProbe Fluorescent Dye 10-fold with Dye Diluent, then further diluting it 10-fold with Buffer. The cells were cultured in appropriate dishes, and the preheated probe-containing working solution was added to the cells. After incubating for 10 minutes in the dark at 37°C, fluorescence intensity was measured using a fluorescence microscope (Nikon Eclipse 80i, Japan) with a maximum excitation/emission wavelength of 510/580 nm.

The adenosine triphosphate (ATP) content in the experimental samples was measured using the ATP detection kit (Beyotime, China), following the manufacturer’s instructions. The ATP content was then normalized to the cell number.

The content of adenosine diphosphate (ADP) and adenosine triphosphate (ATP) in the experimental samples was determined using the ADP/ATP Ratio Assay Kit (Abnova, Wuhan, China).

A. ATP measurement: Cells were lysed using the working solution provided in the kit. ATP reacts with the substrate D-luciferase to produce fluorescence, which is then measured to determine the ATP concentration.

B. ADP measurement: ADP is converted to ATP through an enzymatic reaction, after which the ATP concentration is measured as described in step A.

Cells from each group were collected and washed with PBS. The samples were fixed with 2.5% glutaraldehyde for 1.5 hours, followed by rinsing. Afterward, the samples were further fixed with 2% osmium tetroxide for 1.5 hours and rinsed again. The fixed cell samples were then dehydrated through a graded series of acetone. Next, the samples were infiltrated with pure acetone and embedding solution at room temperature for 1 to 1.5 hours, followed by overnight infiltration with pure embedding solution at room temperature. Ultrathin sections (70 nm) were prepared using an ultramicrotome. Double staining was performed with 2% uranyl acetate and lead citrate. Finally, the samples were observed under a transmission electron microscope.

Firstly, total RNA was extracted from the samples according to the Trizol extraction protocol, and the RNA concentration and purity were assessed using a spectrophotometer. Next, a sequencing library was constructed. DNA was then amplified into clusters, and high-throughput sequencing was performed to obtain fastq data. All data were analyzed using the Dr. Tom Multi-Organomics Data Mining System, with a cutoff of Log2FC absolute value > 1 and Q value < 0.05 to identify differentially expressed genes. Subsequently, KEGG pathway enrichment and gene ontology (GO) enrichment analyses were performed.

Firstly, proteins were extracted from PC3 cells, and protein concentration was determined using the BCA method. Equal amounts of protein were loaded onto a 10% polyacrylamide gel for SDS-PAGE, followed by electrophoresis, membrane transfer, blocking, and incubation with primary and secondary antibodies. Finally, protein bands were detected using ECL chemiluminescence, and images were captured. The antibodies used in this study, including PINK1, Parkin, VDAC1, TOM20, SLC7A11, GPX4, and GAPDH, were purchased from Proteintech Wuhan.

Six-week-old immunodeficient BALB/C nude mice which were purchased from Beijing Laboratory Animal Research Centre were housed under sterile, aseptic conditions. To assess the effect of GRh2 on tumor growth, the mice were randomly assigned to three groups: the blank group, GRh2 group, and GRh2 + Mdivi-1 group. PC3 cells (1 × 10^7 cells/mL) were subcutaneously inoculated bilaterally on the medial aspect of the right upper limb-trunk junction. The cells were mixed with Matrigel (BD Biosciences, San Jose, CA, USA) and diluted 1:3 in RPMI-1640. Mice were treated according to the designated group for 21 days. Tumor size was monitored regularly with a Vernier caliper and calculated using the formula (1/2 × W × H). When the tumor volume reached 1.5 cm³, mice were euthanized by cervical dislocation. Tumor specimens were then fixed with formaldehyde and stored at -80°C for future analysis. All animal experiments were approved by the Ethics Committee of Tianjin Medical University. At the same time, we confirmed that all methods were performed in accordance with the relevant guidelines and regulations from the Ethics Committee of Tianjin Medical University.

Fe²⁺ content in the cell samples was measured using a Ferrous Iron Colorimetric Assay Kit (Elabscience), following the manufacturer’s instructions. Absorbance was read at 593 nm using a microplate reader.

Relative lipid ROS levels in cells were assessed using C11-BODIPY dye (Thermo Fisher Scientific, D3861). Cells were incubated with 5 μM C11-BODIPY for 30 minutes, then washed three times with PBS and resuspended in 500 μL PBS. Fluorescence intensity was measured using a fluorescence microscope (Nikon Eclipse 80i, Japan) with excitation/emission wavelengths of 581/591 nm (Texas Red filter) for vat dyes and 488/510 nm (FITC filter) for oxidation dyes.

PC cells were seeded in cell culture dishes and exposed to the designated treatment groups. After incubation, cells were counted, and an equal number of cells were collected for analysis. Malondialdehyde (MDA) levels were measured using a lipid peroxide malondialdehyde assay kit (Beyotime). Glutathione (GSH) levels were determined according to the manufacturer’s instructions (kktb1600, Abbkine).

Statistical analysis was conducted using GraphPad Prism 8 software (GraphPad, USA). The specific statistical tests used were chosen based on data type and experimental design. Comparisons between two groups were analyzed using a two-tailed Student’s t-test. Comparisons across multiple groups were analyzed using one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test for pairwise comparisons. Cell proliferation curves were analyzed using two-way ANOVA followed by Sidak’s post hoc test for comparisons at specific time points. All data are presented as the mean ± standard error of the mean (SEM). A p-value of < 0.05 was considered statistically significant.

To investigate the inhibitory effect of GRh2 ( Figure 1a ) on the growth of prostate cancer cells, PC3 cells were exposed to eight different concentrations of GRh2 (0 μg/ml, 5 μg/ml, 10 μg/ml, 20 μg/ml, 40 μg/ml, 60 μg/ml, 80 μg/ml, and 100 μg/ml) for 48 hours, and cellular viability was assessed by CCK-8 assay. As shown in Figure 1b , the viability of PC3 cells gradually decreased with increasing GRh2 concentrations, with the half-maximal inhibitory concentration (IC50) calculated to be 19.3 μg/ml. To further verify the effect of GRh2 at the IC50 concentration, we evaluated its impact on cell proliferation at three time points: 0h, 24h, and 48h. The results revealed a significant reduction in cell proliferation following GRh2 exposure ( Figure 1c ). Plate colony formation assays also indicated a diminished cloning capability of PC3 cells upon GRh2 treatment ( Figure 1d ).

Subsequently, we assessed the effects of GRh2 on the invasion and migration capacities of PC3 cells. Transwell assays showed a marked reduction in the number of invaded PC3 cells after GRh2 treatment ( Figure 1e ). Additionally, wound healing assays demonstrated a significant decrease in migration in the GRh2-treated group compared to the control group ( Figure 1f ). Similar results were observed in DU145 cells ( Supplementary Figure S1 ).

These findings collectively suggest that GRh2 effectively inhibits the viability, proliferation, migration, and invasion of prostate cancer cells.

Mitochondria are the powerhouses of cells, and mitochondrial damage plays a critical role in regulating cell death. To explore whether GRh2 induces mitochondrial damage leading to cell death, we first examined the mitochondrial membrane potential in PC3 cells. As shown in Figure 2a , JC-1 staining revealed a significant reduction in mitochondrial membrane potential following GRh2 treatment. However, the mitochondrial membrane potential was restored by the addition of Mdivi-1, a mitochondrial division inhibitor.

Mitochondrial ROS is a key indicator of mitochondrial dysfunction, so we assessed the production of mitochondrial ROS using the BBcell probe. Fluorescence microscopy results showed that GRh2 markedly increased mitochondrial ROS levels in PC3 cells. This increase was reversed by the addition of Mdivi-1, confirming the role of mitochondrial damage in GRh2-induced effects ( Figure 2b ).

We also measured ATP levels and the ADP/ATP ratio, both of which were significantly altered in GRh2-treated cells compared to controls ( Figures 2c , 2d ). To further validate these findings, we performed transmission electron microscopy (TEM). As depicted in Figure 2e , GRh2 treatment induced mitochondrial swelling, an increased density of bilayer membranes, and a reduction or complete disappearance of mitochondrial cristae in PC3 cells. These observations collectively indicate that GRh2 induces mitochondrial damage, leading to cellular dysfunction and death.

Similar results were obtained in DU145 cells ( Supplementary Figure S2 ), supporting the consistency of GRh2’s effect on mitochondrial impairment across different prostate cancer cell lines.

To investigate the molecular mechanisms underlying GRh2’s repressive effects on prostate cancer (PC), RNA sequencing was conducted on PC3 cells from two groups: control and GRh2-treated. The differentially expressed genes (DEGs) between the two groups are shown in Figures 3a , 3b . Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis revealed that the mitophagy pathway was the top enriched pathway among the DEGs in the GRh2-treated group compared to the control group ( Figure 3c ).

Further analysis of the DEGs associated with mitophagy ( Figure 3d ) confirmed the involvement of this pathway in GRh2’s effects on PC3 cells. Based on our previous observations of mitochondrial damage following GRh2 treatment, we sought to validate these findings by assessing mitophagy in PC cells. Transmission electron microscopy (TEM) revealed that GRh2 treatment induced mitochondrial damage, with damaged mitochondria surrounded by membranous structures forming autophagosomes ( Figure 3e ).

Additionally, the expression levels of key mitophagy markers, including PINK1, Parkin, VDAC1, and TOM20, were analyzed by western blotting. Our results showed that PINK1 and Parkin were significantly upregulated, whereas VDAC1 and TOM20 were downregulated in the GRh2-treated group compared to the control group ( Figure 3f ). These changes were reversed by the addition of Mdivi-1, further supporting the role of mitochondrial damage in GRh2-induced mitophagy.

Taken together, these findings suggest that GRh2 induces mitophagy in PC cells, contributing to its anti-cancer effects.

To further investigate the effect of GRh2 on the progression of prostate cancer cells via mitophagy, we performed a series of experimental verifications. Firstly, we treated PC3 cells with four different groups (control, GRh2, Mdivi-1, and GRh2 + Mdivi-1) for 48 hours and assessed cell viability using the CCK8 assay. As shown in Figure 4a , GRh2 exposure led to a decrease in cell viability, which was restored by the addition of Mdivi-1. Additionally, colony formation assays revealed an increase in the clonogenic capacity of PC3 cells when treated with GRh2 + Mdivi-1 compared to the GRh2-only group ( Figure 4b ).

Next, we examined the effect of GRh2 with and without Mdivi-1 on the invasion and migration capacity of PC3 cells. Wound healing assays ( Figure 4c ) showed that cell migration was significantly increased in the GRh2 + Mdivi-1 group compared to the GRh2-only group. Similarly, Transwell assays indicated that the number of invaded PC3 cells was higher in the GRh2 + Mdivi-1 group than in the GRh2 group ( Figure 4d ).

To further explore whether GRh2 affects prostate cancer progression in vivo, we established a subcutaneous PC model in BALB/c nude mice using PC3 cells. The mice were randomly assigned to the control group, GRh2 group, and GRh2 + Mdivi-1 group. The results showed that tumor growth was significantly suppressed in the GRh2 group compared to the control group, with tumor volume in the GRh2 group being approximately five times smaller than that in the control group. However, after the addition of Mdivi-1, both tumor growth rate and volume increased compared to the GRh2-only group ( Figure 4e ).

In summary, our results indicate that GRh2 inhibits the progression of prostate cancer cells through the induction of mitophagy.

In our previous study, we observed that GRh2 exposure to prostate cancer cells resulted in mitochondrial damage, as well as changes in mitochondrial morphology and structure. Based on these findings, we hypothesized a potential link between GRh2 exposure and ferroptosis induction in PC cells, and conducted the following studies to explore this connection.

To investigate whether GRh2 exposure promotes ferroptosis, we treated PC3 cells with GRh2, GRh2 + Fer-1 (ferroptosis inhibitors), and control groups. Lipid ROS, a key marker of ferroptosis, was measured, and our results showed that the green fluorescence intensity in the GRh2-exposed group was significantly stronger than in the other groups, indicating a marked increase in intracellular lipid ROS levels ( Figure 5a ).

We further assessed several ferroptosis-related markers, including Fe2+, GSH, and MDA levels. The GRh2-exposed group showed significantly higher Fe2+ and MDA levels, while GSH levels were significantly reduced compared to the Fer-1 and control groups ( Figures 5b–d ). To further validate these findings, we performed a CCK8 assay on PC3 cells treated with GRh2, Fer-1, or GRh2 + Fer-1 for 48 hours. The results ( Figure 5e ) indicated that GRh2 exposure decreased cell viability, but this effect was reversed by the addition of Fer-1.

GPX4, a key GSH peroxidase, plays an essential role in ferroptosis by inhibiting lipid peroxidation (16). We observed that GRh2 exposure decreased intracellular GSH levels, suggesting a potential link to GPX4 inhibition. Additionally, SLC7A11, an upstream regulator of GPX4, is known to promote ferroptosis when downregulated (17). We examined the expression levels of GPX4 and SLC7A11 by Western blotting and found that their expression was significantly reduced in the GRh2-exposed group compared to the control ( Figure 5f ).

In summary, our findings suggest that GRh2 exposure induces ferroptosis in prostate cancer cells by repressing the SLC7A11/GPX4 pathway, leading to decreased cell viability.