Primary BPH Stromal Cell Cultures Exhibit Heightened Androgen‐Responsiveness

2.1. Prostate Tissue Collection and Processing

Residual prostate tissue specimens used in this study included paraffin‐embedded tissue blocks and fresh specimens obtained from patients with low‐volume prostate cancer and BPH undergoing prostatectomy at the University of Pittsburgh Medical Center, under an approved Institutional Review Board protocol through the Pitt Biospecimen Core at the University of Pittsburgh. Fresh prostatic tissue specimens were dissected under sterile conditions and histologically classified by a board‐certified genitourinary pathologist (R. Dhir) immediately after surgery. Areas of BPH and NAP from the transition zone and/or central zone of the prostate were confirmed by examining frozen sections of tissues procured adjacent to the sample submitted for clinical histologic assessment. Normal adjacent tissues were without any histopathological changes. Thereafter, the tissue samples were transported to the cell culture laboratory in 50/50 Dulbecco's modified Eagles medium (DMEM)/F12 (10‐090‐CV, Corning Inc., Corning, NY, USA), supplemented with 1 µg/mL insulin‐transferrin‐selenium‐X (51500056, Invitrogen, Waltham, MA, USA), 0.4% bovine pituitary extract (13028014, Gibco, Waltham, Massachusetts, USA), 1% penicillin and streptomycin, and 3 ng/mL epidermal growth factor (S0155, Gibco).

2.2. Primary Stromal Cell Culture

BPH and NAP tissue specimens were minced and dissociated by incubation in 2.4 U/ml Dispase II (04942078001, Roche Applied Science, Penzberg, Germany) at 37°C for 1 h on a rocker to generate a suspension [17]. A volume of 1 ml cell suspension was diluted with 4 mL of 50/50 Dulbecco's modified Eagles medium (DMEM)/F12 included 1 µg/mL insulin‐transferrin‐selenium‐X, 0.4% bovine pituitary extract, 1% penicillin and streptomycin, and 3 ng/mL epidermal growth factor in 6 cm dishes. Plated cells were incubated at 37°C, 5% CO₂ for 5 days without disturbing. On day five, 2 mL of fresh medium was added to each dish to compensate for evaporation. Stromal cell proliferation was typically evident by day seven. When stromal cells reached 95% confluence, cells were serially passaged using a trypsin:EDTA (0.25%:0.53 mM) solution and neutralized with medium. Primary stromal cell identity was confirmed as previously described using established stromal markers [17]. Cultures were routinely monitored and consistently exhibited the characteristic spindle‐shaped morphology of prostate stromal cells, with no apparent epithelial‐like colonies observed during culture expansion. Primary cells with 5 or fewer passages were used in RNA sequencing, Western blotting, and co‐culture experiments.

2.3. Western Blotting

Primary cultures of BPH and NAP cells were collected for Western blot assay. RIPA lysate solution was used to prepare cell lysates, and the protein concentration was measured using a BCA Protein Assay Kit (23225, Thermo Fisher Scientific, Waltham, MA, USA). After Western transfer, the membranes were blocked and then incubated with primary antibodies AR (Cell Signaling Technology, ab‐46794, 1: 1000 dilution) and GAPDH (Santa Cruz Biotechnology, sc‐47724, 1:3000 dilution). Subsequently, the membranes were incubated with corresponding secondary antibodies and assessed with chemiluminescence. Quantification of AR protein levels was performed using ImageJ software and normalized to GAPDH.

2.4. RNA Sequencing

Primary stromal cells derived from BPH or NAP were seeded into 6 cm dishes and grown in medium with 1 nM R1881 or 1 nM R1881 plus 10 μM MDV3100 for 2 days to reach 100% confluence prior to collection. Total RNA was extracted from stromal cells using RNeasy Mini Kit (Qiagen, Cat No. 74104, Hilden, Germany). Three patient‐derived stromal cell cultures with robust growth, viability, and cellular morphology were selected for transcriptomic analysis. Sequencing was carried out by Novogene (Sacramento, CA, USA).

2.5. RNA‐Seq Data Processing and Differential Expression Analysis

The RNA seq data can be accessed in GEO with id GSE344492. Transcript abundance was quantified in terms of transcripts per million (TPM) using kallisto v0.46.0 [18], with an index constructed from the Ensembl GRCh38 reference transcriptome. The resulting TPM values for each sample were then compiled into a transcript expression matrix for subsequent analyses. tximport (1.30.0) was employed to convert the kallisto output into raw counts, which were subsequently analyzed for differential expression using edgeR (version 4.0.16). Differentially expressed genes were selected based on a log fold change (logFC) ≥ 1 and p‐value < 0.05 for upregulated genes, and logFC ≤ −1 and p‐value < 0.05 for downregulated genes.

2.6. Principal Component Analysis (PCA) and Functional Enrichment Analysis

PCA was performed using the FactoMineR package (version 2.11) in R, and the results were visualized with the ggord package (version 1.1.7). For functional enrichment analysis, KEGG pathway enrichment analysis was conducted using the clusterProfiler package (version 4.10.1) after mapping gene symbols to ENTREZ Gene IDs to identify significantly enriched biological pathways. Additionally, Gene Ontology (GO) enrichment was performed using g:Profiler, and gene sets with similar functions were clustered using EnrichmentMap.

2.7. Three‐Dimensional Culture of Prostatic Epithelial Cells in the Presence of Stromal Cell Conditioned Medium

Primary stromal cells derived from BPH or NAP were seeded into 6 cm dishes in complete medium. As soon as the stromal cells reached full confluency, the cells were cultured for an additional 2 days with fresh medium in the presence of 1 nM R1881 or vehicle prior to collection of conditioned medium (CM). CM was collected from 100% confluent plates by aspirating from the culture dish into a 15 mL falcon tube. The stromal CM was centrifuged at 180 x g for 5 min to remove cellular debris and the supernatant was collected and stored at 4°C. BPH‐1 or BHPrE1 (2500 cells) were mixed with 40 μL of Matrigel then added carefully at the center of each well of a 24‐well plate, and gels were allowed to solidify for 15 min at 37°C. After Matrigel solidification, 500 µL stromal CM plus 500 µL fresh complete medium were added to each well and cultured without disturbing for 2 days at 37°C. The media in each well was then replaced with a fresh mixture of 500 μL CM and 500 μL completed medium every 2 days; BPH‐1 or BHPrE1 cells were cultured in 3D culture for a total of 6 days. Epithelial spheroids (about 20 spheres/well) were imaged using an inverted phase–contrast microscope (Nikon TE2000‐U) equipped with Hamamatsu digital camera (Hamamatsu Inc) and NIS‐Elements Documentation v 4.6 software (Nikon Instruments Inc., Mellville, NY) with a 20x or 40x objective. On day 6, cells were fixed using 2% paraformaldehyde (PFA) at room temperature for 30 min. The diameter of the spheres was measured using ImageJ (Wayne Rasband, National Institutes of Health, USA).

2.8. Quantitative Real‐Time RT‐PCR (qPCR)

Validation of RNA‐seq results was performed with a separate experimental group of stromal cells treated as described for the RNA‐seq analysis experiment above. The RNA reverse transcription was carried out using the first‐strand cDNA synthesis kit (Promega, Madison, WI). Quantitative real‐time RT‐PCR (qPCR) was performed using SYBR green mix (Thermo Scientific, Rockford, IL). PCR amplification was carried out using Applied Biosystems StepOne™Plus™ Real‐Time PCR Systems (Applied Biosystems CA, USA). The expression of indicated genes was normalized with respect to the GAPDH mRNA level. Data were analyzed using a 2^‐ΔΔCt method. The primers for TNFRSF11B and IL32 are following: TNFRSF11B Forward: 5'‐ GTC TTT GGT CTC CTG CTA ACT C −3'; TNFRSF11B Reverse: 5' ‐ CCT CAC ACA GGG TAA CAT CTA TTC − 3'; IL‐32 Forward: 5' ‐ ATG TGC TTC CCG AAG GTC CTC TCT GA − 3'; and IL−32 Reverse: 5' ‐ TCA TTT TGA GGA TTG GGG TTC AGA GC − 3' Each experimental sample was assayed in triplicate.

2.9. Statistics and Calculations

Statistical analyses were performed with GraphPad Prism 7 (GraphPad Software, La Jolla, California, USA). Data were expressed as mean ± standard deviation. Differences between groups were analyzed by Multiple t‐tests‐one per row. Statistical significance was defined as p < 0.05.

3.1. Androgen Receptor Protein in Primary BPH Stromal Cells Is Elevated in a Subset of Patients

To explore the androgen responsiveness of BPH stromal cells, we first examined androgen receptor (AR) protein level in primary BPH stromal cells, along with paired NAP stromal cells derived from the same patient. As shown in Figure 1, AR protein levels varied in primary BPH stromal cells derived from 7 different patients but in 4 paired cultures, AR levels appeared similar in NAP versus BPH stromal cell cultures. In contrast, AR level in NAP stromal cells from 3 of the 7 patients was very low or not detected. This finding suggests that AR expression in BPH stromal cells is elevated as compared to the paired NAP stromal cells in the same prostate in a subset of patients.

3.2. Androgen‐Induced Gene Expression in BPH Is Heightened Compared to NAP Primary Stromal Cells

To evaluate androgen signaling in BPH and NAP stromal cells, we performed RNA‐Seq and identified androgen‐response genes in paired primary BPH and NAP stromal cells from 3 different patients with BPH. Primary stromal cells were cultured in the presence of 1 nM R1881 with or without 10 μM MDV3100 (enzalutamide), a highly potent and specific AR antagonist. The expression of androgen‐response genes should be inhibited by enzalutamide. Principal component analysis (PCA) of the RNA‐seq data demonstrated clear clustering among different treatment groups. Interestingly, when stimulated with androgen, BPH stromal cells show greater sensitivity compared to NAP, as evidenced by the larger distance between the stimulated (BR = BPH with 1 nM R1881) and control groups (BM = BPH with 1 nM R1881 plus 10 μM MDV3100) for BPH samples on the PCA plot (Figure 2A). To corroborate this result, we further analyzed the differentially expressed genes (DEGs) in BPH and NAP samples from different patients with androgen stimulation. In all three patient samples, BPH showed more DEGs compared to NAP samples from the same patient (Figure 2B). Venn diagram shows the intersection of the up‐ (red) and down‐regulated (blue) genes in BPH samples from different patients after androgen stimulation (Figure 2C). The results show that seven genes (ALCAM, ANO3, GAS1, IRAG1, JPH2, NEDD9, RHOJ) are consistently up‐regulated across all samples, while three genes (CYP2S1, GAL, INSIG1) are consistently down‐regulated. Due to significant individual differences among patients, we define genes that are simultaneously up‐regulated in two or more patients as AR induced up‐regulated genes, and genes that are simultaneously down‐regulated in two or more patients as AR induced down‐regulated genes. In total, we identified 82 up‐regulated genes (Table S1) and 58 down‐regulated genes (Table S2). Notably, 17 of the 82 up‐regulated genes (21%) are known or predicted secreted proteins, representing a significant enrichment over the 7 of 58 (12%) in down‐regulated genes [19]. This suggests that AR can promote stromal cells to release more secreted proteins, thereby regulating neighboring cells in the microenvironment through paracrine signaling.

GO analysis revealed that AR‐induced upregulated genes were mainly enriched in biological processes related to cell motility and migration, circulatory system and vascular development, and regulation of blood pressure and vasoconstriction, suggesting that androgen signaling may enhance stromal cell activity and promote paracrine‐mediated tissue remodeling. In contrast, the downregulated genes were significantly associated with ATP and nucleotide biosynthesis, mitotic regulation and cell cycle progression, and DNA repair and replication, indicating that androgen stimulation may influence stromal cell proliferation and metabolic activity (Figure 3A). To further explore the biological processes affected by androgen stimulation, we also performed KEGG pathway enrichment analysis using a broader set of DEGs, including genes differentially expressed in at least one patient, to capture more comprehensive androgen‐responsive pathways. The analysis revealed multiple pathways associated with cell proliferation, secretion, and intercellular communication, highlighting the diverse regulatory effects of AR signaling in stromal cells. Affected pathways included cell cycle, TGF‐beta signaling pathway, cytokine‐cytokine receptor interaction, Wnt signaling pathway, ECM‐receptor interaction, calcium signaling pathway, and vascular smooth muscle contraction, etc (Figure 3B). Many of these pathways are closely related to cell growth regulation and secretory activity. For instance, cell cycle and TGF‐beta signaling are central regulators of stromal cell proliferation, while cytokine‐cytokine receptor interaction and ECM‐receptor interaction are key mediators of paracrine communication between stromal and epithelial cells. Moreover, Wnt signaling and calcium signaling pathways are known to coordinate cell differentiation and contractile responses, both of which are critical for prostate tissue remodeling.

Collectively, both GO and KEGG analyses consistently indicate that androgen signaling in BPH stromal cells exerts multifaceted regulatory effects. AR activation appears to enhance cellular motility, vascular and extracellular matrix remodeling, and paracrine signaling, while simultaneously modulating cell cycle and metabolic processes. These coordinated changes suggest a shift in stromal cells toward a more active, secretory, and signaling‐oriented state, which may enhance androgen‐dependent stromal–epithelial interactions and contribute to prostate tissue growth and the hyperplastic phenotype observed in BPH.

3.3. Comparative Transcriptomic Analysis of BPH and NAP Stromal Cells

We subsequently conducted a comparative transcriptomic analysis between paired primary BPH and NAP stromal cells from three patients. Volcano plots showed genes that are up‐regulated and down‐regulated in primary BPH stromal cells compared to primary NAP stromal cells both under AR blockade (BM vs NM; Figure 4A) and under androgen stimulation (BR vs NR; Figure S2) in different patients. Venn diagram showed the intersection of differential genes originating from different patients, with up‐regulation on the left and down‐regulation on the right (Figure 4B). Similarly, we selected DEGs that were consistently up‐ or down‐regulated in at least two patient samples for subsequent GO enrichment analysis, while the union of DEGs detected in at least one sample was used for KEGG pathway analysis. In total, we obtained 72 up‐regulated genes and 154 down‐regulated genes and selected the known or predicted secreted proteins among them. Among the 72 up‐regulated genes, we identified 18 secreted proteins that may play a crucial role in regulating epithelial cell proliferation in the microenvironment through paracrine signaling in BPH stromal cells (Tables [Link], [Link]). One of these, placental growth factor (PGF), has been reported to be highly expressed at the mRNA level in BPH stromal cells [20]. Further study of PGF and other secreted proteins may provide crucial insights into BPH pathogenesis. The GO analysis of signature genes in BPH revealed that most of these genes are associated with biological processes such as development, differentiation, and proliferation. This is consistent with the phenotype of uncontrolled abnormal growth observed in BPH (Figure 4C). Consistently, KEGG pathway enrichment analysis further highlighted signaling pathways related to cell proliferation, adhesion, and intercellular communication, including TGF‐beta signaling, PI3K‐Akt signaling, TNF signaling, Hippo signaling, and focal adhesion. In addition, pathways such as cytokine‐cytokine receptor interaction, ECM‐receptor interaction, and calcium signaling pathway were also enriched, suggesting enhanced stromal‐epithelial crosstalk and extracellular matrix remodeling. Together, these results indicate that dysregulation of these pathways may contribute to the excessive proliferative and structural remodeling activities characteristic of BPH (Figure 4D). To propose potential therapeutic options for BPH based on BPH stromal signature genes, we employed the Connectivity Map analysis. This approach leverages transcriptional expression data to explore connections between small molecules, genes, and disease [21, 22]. Figure 4E displays the top 10 nominated compounds according to the CMAP score. These compounds include antiviral agents and AR (androgen receptor) targeting drugs, which align with the characteristics of BPH and current clinical AR‐targeted treatments. Additionally, the list includes ERK inhibitors and BCL inhibitors, among others. This information provides valuable insights for subsequent experimental studies on BPH.

To support the results of RNA Seq, we assayed several cytokines, such as IL32 and TNFRSF11B, that were differentially expressed in paired primary stromal cells from a different patient (TP17‐S573) through RT‐PCR. RNA‐Seq showed IL32 and TNFRSF11B were suppressed in BPH stromal cells in the presence of MDV3100. Quantitative PCR analysis revealed that IL32 and TNFRSF11B were elevated in BPH stromal cells compared to NAP stromal cells, which was consistent with RNA‐Seq results (Figure 5). Quantitative PCR also showed androgen induction of these genes in primary BPH stromal cells, but not in NAP stromal cells (Figure 5).

3.4. Stimulation of Epithelial Proliferation by Androgen‐Treated BPH Stromal Conditioned Medium

To explore whether the differences in R1881‐induced secretory factor gene expression in RNASeq analysis of BPH vs NAP cells is associated with a functional consequence, we first tested the effect of synthetic androgen R1881 on prostatic epithelial cell lines BPH‐1 and BHPrE1, as well as on primary stromal cells, in monolayer culture. R1881 did not significantly stimulate the proliferation of any of the cultured cells in monolayer (data not shown). Next, we assessed the impact of R1881 on the growth of BPH‐1 and BHPrE1 cells in a 3D culture system. Cells were cultured in Matrigel for 6 days in the presence of conditioned medium (CM) from stromal cells derived from either BPH tissues or paired normal adjacent tissues (NAP), with or without 1 nM R1881. BHPrE1 cells cultured alone in Matrigel and treated with R1881 showed no enhanced growth (Figure 6A). Similarly, no significant difference in spheroidal size was observed when BHPrE1 cells were treated with CM from NAP stromal cells and R1881 (Figure 6B). However, when BHPrE1 cells were treated with CM from BPH stromal cells in the presence of R1881, the spheroid size significantly increased (Figure 6C). The same results were observed in BPH‐1 cell lines (Figure 6D–F). These findings were reproduced using another pair of stromal cells (Supplemental Fig. S1). This suggests that androgens, through their effects on BPH‐derived stromal cells, can induce the growth of epithelial cells.