We utilized PubMed and Web of Science to investigate the clinical applications of the Chinese botanical drug extracts CEP, which named QianJinTengSu in Chinese. Additionally, the TCMIP database (http://www.tcmip.cn/TCMIP/index.php/) was employed to examine efficacy and treatment data related to CEP. Our objective was to obtain precise preliminary insights into the underlying mechanisms by which CEP may treat BPH.

The canonical SMILES notation and molecular structure of CEP were retrieved from the PubChem database (https://pubchem.ncbi.nlm.nih.gov/, accessed on 14 November 2024). Utilizing this structural information, we identified potential drug targets by employing various databases, including SwissTargetPrediction (http://www.swisstargetprediction.ch/), ChEMBL (https://www.ebi.ac.uk/chembl/), SEA Search Server (https://sea.bkslab.org/), and STITCH (http://stitch.embl.de/). To ensure data integrity and reliability, the search was restricted to targets specific to Homo sapiens. Following the aggregation of results, redundant targets were filtered out, resulting in a comprehensive and refined library of potential CEP drug targets (Shi S. et al., 2023).

The keywords “benign prostatic hyperplasia”, “BPH”, and “hyperplasia, prostatic” were employed to retrieve BPH-related targets from three disease-specific databases: GeneCards (https://www.genecards.org/), OMIM (https://omim.org/), and TTD (https://db.idrblab.net/ttd/). The retrieved BPH-related targets were consolidated, and duplicate entries were excluded. Using the “Relevance score” in the GeneCards database as the screening basis, only genes with scores higher than the median score of all targets are retained to ensure that the screened targets have a high correlation with the disease, thereby improving the reliability of subsequent analysis.

To explore the potential targets of CEP in treating BPH, we performed a cross-analysis of CEP and BPH-related Targets and generated a Venn diagram using Venny 2.1 tools (https://bioinfogp.cnb.csic.es/tools/venny/index.html) to identify overlaps. These intersecting targets were subsequently imported into the STRING database (http://string-db.org/) to construct a PPI network. The species was limited to Homo sapiens, and the minimum interaction score threshold was set to 0.4, with all other parameters maintained as default. The PPI network was visualized using Cytoscape 3.9.1 software, and the degree of each target was calculated using the CytoNCA plugin. Key targets were identified based on their degree values, and molecular interactions were further analyzed.

To elucidate the biological mechanisms underlying the potential targets of CEP in the treatment of BPH, we conducted Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analyses. Utilizing the DAVID database (DAVID Functional Annotation Tools), which provides comprehensive gene function annotations across biological processes (BP), cellular components (CC), and molecular functions (MF), we identified significant GO terms and KEGG pathways. Pathway enrichment was deemed statistically significant at a false discovery rate (FDR) of less than 0.05. The top 10 GO terms and top 20 KEGG pathways were thoroughly summarized.

Furthermore, KEGG pathway enrichment analyses for the central targets of CEP in BPH treatment were performed using multiple databases, including DAVID, FUMA (https://fuma.ctglab.nl/gene2func), and Metascape (https://metascape.org/gp/index.html#/). This comprehensive approach aimed to investigate the signaling pathways and biological processes mediated by core targets in BPH treatment, thereby elucidating key mechanisms. Finally, the GO and KEGG enrichment results were visualized and interpreted using the online tool WeiShengXin (https://www.bioinformatics.com.cn/), facilitating the exploration of drug-disease signaling pathways and biological processes, and highlighting the underlying mechanisms.

To elucidate the molecular interactions and binding modes between CEP and key target proteins, we conducted molecular docking simulations. This structure-based approach predicts receptor-ligand binding geometries and affinities. Ligand files of CEP components in SDF format were retrieved from the PubChem database (https://pubchem.ncbi.nlm.nih.gov/), and three-dimensional structural models of proteins RAC-alpha serine/threonine-protein kinase 1 (AKT1), epidermal growth factor receptor (EGFR), Proto-Oncogene Tyrosine-Protein Kinase Src (SRC), and fibronectin 1 (FN1) were sourced from AlphaFold2 (https://alphafold.com/), UniProt (https://www.uniprot.org/), and the Protein Data Bank (PDB) (https://www.rcsb.org/). Docking was performed using the CB-DOCK platform (https://cadd.labshare.cn/cb-dock2/index.php), and the resulting poses were analyzed and visualized with Discovery Studio 2019 software.

CEP was purchased from Selleck Chemicals (Houston, USA). The purity of CEP was ≥98% as determined by HPLC. Store at −20 °C in a dry place away from light. Before use, prepare a 10 mM stock solution in DMSO and aliquot to avoid repeated freeze-thaw cycles. All CEP working solutions used in experiments were freshly diluted from this stock solution. DMEM high glucose medium, fetal bovine serum, trypsin, and PBS solution was purchased from Thermo Fisher-Scientific (Massachusetts, USA). Penicillin-streptomycin solution was purchased from Merck (Darmstadt, Germany). Cell Counting Kit (CCK)-8 was purchased from Dojindo Molecular Technologies (Maryland, USA). EdU cell proliferation assay kit, Annexin V-APC/PI Apoptosis Kit and Annexin V-FITC/PI Apoptosis Detection kit were purchased from Elabscience (Wuhan, China). Antibodies against AKT (No. 75692), phosphorylated AKT (Ser473) (No. 4060), EGFR (No. 4267), phosphorylated EGFR (Tyr1068) (No. 3777), FN1 (No. A0056-3), and GAPDH (No. 2118) were purchased from Cell Signaling Technology, Inc. (Boston, USA). Anti-rabbit secondary antibody was purchased from Absin (Shanghai, China).

WPMY-1 (human prostate stromal cells) and BPH-1 (​benign prostatic hyperplasia-1) were purchased from American Type Culture Collection (ATCC) (Maryland, USA) and cultured in DMEM high-glucose medium supplemented with 10% fetal bovine serum and 1% penicillin-streptomycin double antibody solution at 37 °C in a humidified atmosphere of 5% CO₂. The culture medium was changed every 2–3 days according to the cell growth rate and state. When the cell confluence reached about 80%, the cells were passaged in proportion. All experiments were performed using cells between passage 3 and passage 20 to ensure phenotypic stability and minimize the effects of cellular senescence.​

10,000 WPMY-1 cells/well or 10,000 BPH-1 cells/well were seeded in a 96-well plate (100 μL culture medium per well) for 24 h (37 °C, 5% CO₂) to adhere to the wall. Then, DMSO and 2.5, 5, 10, and 20 μM CEP were added for 48 h. Three replicate wells were set for each concentration, and the edge wells were filled with PBS buffer to eliminate the evaporation effect. After the intervention, 10 μL CCK-8 reagent was added to each well, gently shaken and mixed, and incubated for 2 h. The absorbance at 450 nm was detected using an ELISA reader (Biotek, Vermont, USA), and blank wells and solvent control wells were set simultaneously to correct the background. The cell viability was calculated according to the formula: (OD₄₅₀ of the experimental group - OD₄₅₀ of the blank group)/(OD₄₅₀ of the DMSO control group - OD₄₅₀ of the blank group) × 100%. The four-parameter nonlinear regression dose-effect curve was fitted with CEP concentration (X-axis) and survival rate (Y-axis), and the drug concentration at which the inhibition rate reached 50% was calculated as IC₅₀. Subsequently, 10,000 WPMY-1 (or BPH-1) cells/well were inoculated and pre-cultured for 24 h using the same method, and then divided into a DMSO control group and a 5 μM CEP (for BPH-1, the CEP concentration was 2.5 μM) (CEP concentration determined according to IC₅₀) treatment group (3 replicates per group). At five time points of 0, 24, 48, 72, and 96 h, 10 μL of CCK-8 reagent was directly added to each well, and the absorbance at 450 nm was detected after incubation for 2 h (the blank well background was deducted). The proliferation rate was calculated according to the formula: (treatment group OD₄₅₀–0 h OD₄₅₀)/(control group OD₄₅₀–0 h OD₄₅₀) × 100%, and the time-proliferation rate curve was drawn.

A 96-well plate (final volume of 100 μL per well) inoculated with 10,000 WPMY-1 (or BPH-1) cells was divided into a DMSO control group and a 5 μM CEP (for BPH-1, the CEP concentration was 2.5 μM) (CEP concentration determined according to IC₅₀) treatment group. The cells were pre-cultured for 24 h before administration to ensure stable cell adhesion. Non-labeled live cell counts were performed using a Celigo full-view cell scanning analyzer (Nexcelom Bioscience, Boston, USA) at five time points: 0, 24, 48, 72, and 96 h. The instrument was set to bright field full-well scanning mode (resolution 1 μm/pixel), and the Direct Cell Counting Application was used to perform high-speed imaging of the entire plate. The software automatically segmented the cell image and counted the number of live cells in each well. To reduce batch errors, the same culture plate was used for dynamic tracking at multiple time points throughout the process, and the culture plate was gently shaken before each scan to redistribute the suspended cells.

The culture medium of WPMY-1 (or BPH-1) cells (96-well plate, 10,000 cells/well, 100 μL culture medium/well) treated with DMSO or 5 μM CEP (for BPH-1, the CEP concentration was 2.5 μM) (CEP concentration determined according to IC₅₀) for 48 h was discarded, 100 μL culture medium containing 10 μM EdU was added to each well, and incubated at 37 °C for 2 h to allow EdU to be incorporated into DNA; the EdU culture medium was discarded, the cells were washed twice with PBS, and 50 μL 4% paraformaldehyde was added to each well for fixation at room temperature for 30 min; the fixative was discarded, and 100 μL PBS containing 0.5% Triton X-100 was added for permeabilization for 10 min; the permeabilization solution was discarded, and 50 μL of Click reaction solution was prepared according to the proportion of the kit, and it was added to each well for reaction at room temperature for 30 min in the dark; the reaction solution was discarded, PBS was washed three times, and 100 μL DAPI (1 μg/mL) was added to each well for nucleus staining in the dark for 10 min; after washing with PBS, the cells were observed under a fluorescence microscope (Olympus, Tokyo, Japan), and the EdU⁺ (green) and DAPI⁺ (blue) cells were counted to calculate the proliferation rate.

The culture medium of WPMY-1 cells (96-well plate, 10,000 cells/well) treated with DMSO or 5 μM CEP for 48 h was discarded, and the cells were gently washed once with PBS. The adherent cells were digested with EDTA-free trypsin, and the floating cells (including apoptotic cells) were combined and collected by centrifugation at 1,000×g for 5 min. The supernatant was discarded, and the cells were washed twice with pre-cooled PBS and resuspended with 195 μL 1× Binding Buffer. 100 μL of the cell suspension was taken, 5 μL Annexin V-APC and 10 μL propidium iodide (PI) staining solution were added, and the mixture was gently vortexed and incubated at room temperature in the dark for 15 min 300 μL Binding Buffer was added to terminate the staining, and the cells were detected by flow cytometry within 1 h (Beckman, California, USA). Apoptosis rate analysis: Annexin V-APC⁺/PI⁻ represents early apoptotic cells, Annexin V-APC⁺/PI⁺ represents late apoptotic/necrotic cells, and the increase in apoptosis rate was calculated using the DMSO group as the control. BPH-1 cells were seeded in 6-well plates at a density of 2 × 10⁵ cells per well and treated with CEP at concentrations of 0 and 2.5 μM for 48 h. Cells were then collected for apoptosis analysis using an Annexin V-FITC/PI Apoptosis Detection kit (Elabscience, Wuhan, China) according to the manufacturer’s protocol. Briefly, cells were resuspended in 500 μL of 1× binding buffer and stained with 5 μL Annexin V-FITC and 5 μL PI in the dark at room temperature for 15–20 min. Apoptosis was assessed using a flow cytometer.

WPMY-1 cells or BPH-1 cells (6-well plate, 2 × 10⁵ cells/well) treated with DMSO, 2.5 μM and 5 μM CEP for 48 h were discarded from the culture medium, washed twice with pre-cooled PBS, and 100 μL RIPA lysis buffer was added to each well for lysis on ice for 30 min. The cells were scraped and transferred to a centrifuge tube, centrifuged at 12,000×g and 4 °C for 15 min, and the supernatant was taken. The protein concentration was determined by BCA method and adjusted to equal concentration. 20 μg protein sample was mixed with 5× loading buffer and denatured by boiling at 100 °C for 10 min. The cells were separated by 10% SDS-PAGE gel electrophoresis and transferred to PVDF membrane by wet transfer method. The cells were blocked with 5% skim milk at room temperature for 1 h, and primary antibodies were added: AKT, p-AKT, EGFR, p-EGFR, FN1, GAPDH, and incubated at 4 °C overnight. The cells were washed with TBST 3 times, and HRP-labeled secondary antibodies were added for incubation at room temperature for 1 h. After washing with TBST, ECL chemiluminescence was developed. ImageJ (NIH, USA) software was used to analyze the grayscale values ​​of the bands. The ratio of the target protein to GAPDH was used to correct the loading error. The phosphorylation level was calculated according to the ratio of p-AKT/AKT and p-EGFR/EGFR.

All cell experiments (including CCK-8, EdU, flow cytometry, and Western blot) were repeated three times independently. The data are expressed as mean ± standard deviation (mean ± SD). Multiple groups were compared using one-way analysis of variance (ANOVA) followed by Tukey’s post hoc test, and statistical significance was set at P < 0.05.

This study adhered to the four pillars of ethnopharmacology best practices: (1) Traditional use: CEP has a history of use in traditional Chinese medicine; (2) Quality assurance: HPLC-verified high-purity standards were used; (3) Pharmacological evaluation: Multiple in vitro assays were used to verify biological activity; and (4) Ethical: All cell lines were commercially available, and no human or animal experiments were performed.

The standardized name for CEP in the TCMIP database is Epistephanine, with a chemical formula of C₃₇H₃₈N₂O₆ and a drug-likeness score of 0.248. CEP is a natural alkaloid known for its inhibitory effects on TNF-α-mediated NF-κB activation, plasma membrane lipid peroxidation, platelet aggregation, and cytokine production. These properties contribute to its anti-inflammatory, antioxidant, and anti-tumor activities. Notably, in 2022, traditional Chinese medicine, including compounds like CEP, demonstrated efficacy in treating COVID-19.

Benign prostatic hyperplasia has been associated with various mechanisms, including sex hormone regulation, peptide growth factor signaling, inflammatory pathways, apoptosis, oxidative stress, and smooth muscle activity. Collectively, these findings lay a critical groundwork for subsequent systematic and comprehensive studies assessing the therapeutic potential of CEP in BPH.

This study initially identified 142 therapeutic targets of CEP from the SwissTargetPrediction, ChEMBL, SEA Search Server, and STITCH databases, and 5,081 targets highly associated with BPH through analysis of the GeneCards, OMIM, and TTD databases. By integrating and deduplicating these target sets, 96 overlapping targets were obtained as potential targets for CEP in the treatment of BPH (Figure 1A) (Supplementary Appendix 2).

We constructed a PPI network using the STRING database, comprising 96 nodes and 666 edges. The network exhibited an average node degree of 13.9, an average local clustering coefficient of 0.545, and a PPI enrichment p-value of <1.0e-16, indicating significant interactions among the targets.

The topological properties of the network nodes, including degree and betweenness centrality, were analyzed using Cytoscape software, resulting in an optimized PPI network visualization (Figure 1B). Through this analysis, 20 core targets of CEP for treating BPH were identified (Table 1). A focused PPI network was then generated (Figure 1C) to illustrate interactions among these core targets. The top 4 targets based on degree values were AKT1, EGFR, SRC, and FN1. These proteins regulate cell proliferation, cell cycle control, apoptosis, and signal transduction.

To further elucidate the interactions and potential mechanisms between CEP and four key targets (AKT1, EGFR, SRC, and FN1) in the treatment of BPH, we performed comprehensive molecular docking simulations. Using the CB-Dock online tool, we generated docking models for each target. Notably, all models exhibited binding energies lower than −5.0 kcal/mol, indicating a strong binding affinity between CEP and these targets. This suggests that CEP can spontaneously bind to these core targets, playing a significant role in its molecular mechanism for treating BPH. Detailed VINA scores and docking energies are provided in Table 2.

We utilized Discovery Studio software to visualize the lowest energy binding conformations between CEP and each key target, generating both 2D and 3D representations (Figure 3). These visualizations provided insights into the molecular interactions and binding modes between CEP and the target proteins, highlighting their potential roles in mediating the drug’s therapeutic effects.

CCK-8 assay showed that CEP exhibited a dose-dependent inhibition on the viability of WPMY-1 cells in the concentration range of 0–20 μM. As the concentration of CEP increased, the cell viability decreased in a step-like manner. The dose-effect curve fitted by four-parameter nonlinear regression showed that the drug concentration corresponding to a 50% inhibition rate was IC₅₀ = 6.396 μM (95%CI 5.915–6.912 μM) (Figure 4A). Based on this, 5 μM was selected for treatment in subsequent experiments: this concentration maintained the viability at 65%–70% (corresponding to a 35%–40% inhibition rate), which can effectively observe the drug effect and avoid the cytotoxic interference caused by high concentrations, thus ensuring the reliability of mechanism research. In addition, CCK-8 assay showed that CEP exhibited a dose-dependent inhibition on the viability of BPH-1 cells in the concentration range of 0–20 μM. As the concentration of CEP increased, the cell viability decreased in a step-like manner. The dose-effect curve fitted by four-parameter nonlinear regression showed that the drug concentration corresponding to a 50% inhibition rate was IC₅₀ = 2.355 μM (95%CI 2.103–2.632 μM) (Supplementary Appendix 3A). Based on this, 2.5 μM was selected for treatment in subsequent experiments.

WPMY-1 cells were divided into a DMSO control group and a 5 μM CEP treatment group. CCK-8 was used to detect cell proliferation activity at five time points: 0, 24, 48, 72, and 96 h. The results showed that 5 μM CEP inhibited WPMY-1 cell proliferation in a time-dependent manner. The cell activity of the DMSO control group continued to rise, reaching a peak at 72 h, and slightly declined at 96 h due to contact inhibition; the activity of the CEP treatment group was always significantly lower than that of the control group, indicating that CEP can effectively block the cell proliferation process of WPMY-1 (Figure 4B). In addition, non-labeled live cell counting was performed using the Celigo full-view cell scanning analyzer. The results also confirmed that 5 μM CEP showed a significant time-dependent inhibition of WPMY-1 cell proliferation: the number of cells in the DMSO control group continued to rise, reaching a peak at 72 h, and slightly declined at 96 h due to contact inhibition; the proliferation of the CEP treatment group was not significantly affected in the early stage (24 h), growth stagnation began at 48 h, and severe cell disintegration occurred at 72–96 h. Morphological analysis showed that the cell density in the CEP group dropped sharply after 48 h, and the widespread signal disappeared after 96 h, confirming that CEP may inhibit the cell activity of WPMY-1 through a two-stage mechanism of first blocking proliferation and then inducing cell death (Figures 4C,D).

Furthermore, BPH-1 cells were divided into a DMSO control group and a 2.5 μM CEP-treated group. Cell proliferation activity was assessed using the CCK-8 assay at five time points: 0, 24, 48, 72, and 96 h. Results showed that 2.5 μM CEP exhibited a time-dependent inhibitory effect on BPH-1 cell proliferation. Cell viability in the DMSO control group continued to increase, reaching a peak at 72 h, and then decreased significantly at 96 h due to contact inhibition. Cell viability in the CEP-treated group remained significantly lower than that in the control group, demonstrating that CEP effectively blocked BPH-1 cell proliferation (Supplementary Appendix 3B). Label-free viable cell counts were performed using a Celigo full-field cell analyzer. Results also confirmed that 2.5 μM CEP exhibited a significant time-dependent inhibitory effect on BPH-1 cell proliferation: Cell number in the DMSO control group continued to increase, reaching a peak at 72 h, and then decreased slightly at 96 h due to contact inhibition. In the CEP-treated group, cell proliferation showed no significant effect early in the 24-h period, but began to arrest at 48 h, and severe cell disintegration occurred between 72 and 96 h (Supplementary Appendix 3C).

EdU staining assay visually showed that CEP significantly inhibited the proliferation of WPMY-1 cells. In the DMSO control group, a large amount of EdU⁺ green signals were distributed in the DAPI⁺ blue nuclei, indicating that the cells were actively proliferating; the proportion of EdU⁺ cells in the CEP treatment group decreased sharply, and the green signals were sparsely distributed in a dot-like manner. Merge further confirmed that the cell density in the CEP group decreased and nuclear fragmentation increased, verifying the inhibitory effect of CEP on WPMY-1 cell proliferation from the perspective of DNA synthesis (Figures 4E,F).

Similarly, EdU staining experiments visually demonstrated that CEP also significantly inhibited BPH-1 cell proliferation. In the DMSO control group, abundant EdU⁺ green signals were distributed within DAPI⁺ blue nuclei, indicating active cell proliferation. In the CEP-treated group, the proportion of EdU cells decreased dramatically, and the green signals were sparsely distributed in a punctate pattern. Merge experiments further confirmed the decreased cell density and increased nuclear fragmentation in the CEP group, confirming CEP’s inhibitory effect on BPH-1 cell proliferation from the perspective of DNA synthesis (Supplementary Appendix 3E).

Annexin V-APC/PI double staining flow cytometry showed that 5 μM CEP treatment for 48 h significantly induced apoptosis of WPMY-1 cells. The proportion of late apoptotic cells (Annexin V⁺/PI⁺, upper right quadrant) in the CEP group increased significantly, while the proportion of early apoptotic cells (Annexin V⁺/PI⁻, lower right quadrant) also increased (Figures 4G,H). The characteristic distribution of apoptotic cells clearly migrated toward the Annexin V⁺ quadrant, suggesting that CEP inhibits the activity of prostate stromal cells by dually promoting early and late apoptosis processes.

Annexin V-FITC/PI double staining combined with flow cytometry was used to examine the inhibitory effect of CEP on apoptosis in BPH-1 cells. As shown in Supplementary Appendix 3D, after 48 h of treatment with 2.5 μM CEP, the proportion of apoptotic cells was significantly altered compared to the DMSO control group. The proportion of early apoptotic cells (Annexin V⁺/PI⁻) in the CEP-treated group was significantly increased, while the proportion of late apoptotic and necrotic cells (Annexin V⁺/PI⁺) also increased to a certain extent. These results indicate that 2.5 μM CEP treatment effectively induces apoptosis in BPH-1 cells, primarily by significantly increasing the proportion of early apoptotic cells.

For WPMY-1, Western blot analysis showed that CEP inhibited the activation of the EGFR/PI3K/AKT signaling pathway and the expression of FN1 in a concentration-dependent manner. Compared with the DMSO group, 2.5 μM CEP treatment for 48 h significantly reduced the expression of p-AKT and p-EGFR, and 5 μM CEP further aggravated the inhibitory effect; phosphorylation level analysis simultaneously showed that the ratios of p-AKT/AKT and p-EGFR/EGFR in the CEP group were significantly decreased. The expression of the downstream effector protein FN1 decreased in a step-by-step manner with the increase in CEP concentration, confirming that CEP inhibited the synthesis of FN1, a key component of the extracellular matrix, by blocking the EGFR/PI3K/AKT signaling cascade, thereby inhibiting the proliferation of WPMY-1 cells (Figure 5).

Results from the BPH-1 cell line showed that, compared with the DMSO control group, CEP treatment for 48 h at 2.5 μM and 5 μM decreased the expression of the extracellular matrix protein FN1 in a concentration-dependent manner. Importantly, while the total protein levels of key signaling pathway proteins EGFR and AKT did not change significantly, their phosphorylation levels (p-EGFR and p-AKT) decreased significantly with increasing CEP concentration, indicating that CEP can effectively inhibit activation of the EGFR/AKT signaling pathway. These results suggest that CEP may exert its inhibitory effect on cell viability by inhibiting the EGFR/AKT signaling pathway, thereby affecting the expression of downstream functional proteins such as FN1 (Supplementary Appendix 4).