All animal experimental procedures were performed on six-weeks-old male (150 g) RccHan:WIST rats (Envigo Laboratories, Indianapolis, IN, United States). Rats were housed in three animals per cage under a 12/12 h light/dark cycle at a constant temperature (23 °C–25 °C) with free access to food and water and were allowed to acclimate for at least 1 week after arrival before starting all experiments, which were conducted between 9:00 a.m. and 3:00 p.m. This study was executed according to the European Communities Council directive and Italian regulation (EEC Council 2010/63/EU and Italian D.Lgs. No. 26/2014) to replace, reduce, and refine the use of laboratory animals. All procedures were approved by the ethical committee of the University of Catania (OPBA) and by the Italian Ministry of Health (authorization no. 231/2023-PR). Animals were divided into three experimental groups: Control group (sham operation group), BPH group and BPH orally administered with pomegranate extract (PWE). Rats of the BPH group were injected IP (Intraperitoneal inoculation) with testosterone propionate (10 mg/kg) in corn oil, every 2 days for 2 weeks, in line with literature reports (Paterniti et al., 2018). Rats of the BPH model group were orally administrated with or without PWE as aqueous solution at 0.6%, added with 1% sucrose, which was replenished every 2 days. The commercial dry powdered pomegranate extract (PWE; Medgranate™, Medinutrex, Catania, Italy) used in this study was derived from “Wonderful” variety pomegranate by-products (exhausted peels, membranes, and arils) via industrial drying to 5% moisture, hydroalcoholic extraction (ethanol/water 60:40 v/v, 1:10 w/v, 24 h stirring), alcohol recovery, and spray-drying (∼30% yield), and has been previously characterized and validated for its polyphenolic profile and bioactivity in in vitro models of prostate cell lines demonstrating antioxidant and anti-proliferative effects (Consoli et al., 2023). At the end of the experiment, animals were euthanized by CO₂ inhalation with a flow rate of about 50% (5 L/min) and prostate, liver and periprostatic adipose tissue (PPAT) were excised. Prostate tissue was used for histological, immunohistochemical, and biochemical analyses, liver and PPAT tissues were processed for proteomic and biochemical analyses.

Rat prostate tissues were immediately fixed in 10% formalin for 24 h and were then paraffin-embedded and sectioned at a thickness of 5-μm. The sections were processed for hematoxylin-eosin (H&E) staying as previously described (Ginja et al., 2019). Briefly, selected sections were deparaffinized in xylene and rehydrated in alcohol concentrations. Subsequently, sections were stained with hematoxylin and eosin (H&E; Sigma-Aldrich, Egypt) for histological examination.

The expression and distribution of Platelet endothelial cell adhesion molecule (PECAM-1), Hemeoxygenase-1 (HO-1), and Nuclear factor erythroid 2-related factor 2 (Nfr2) in rat prostate tissue was evaluated through immunohistochemical analysis as previously described (D’Amico et al., 2013; D’Amico et al., 2023). Briefly, the sections were incubated overnight at 4 °C with the specific primary antibodies, and then incubated with 3,3′-diaminobenzidine solution (DAB substrate Kit; SK-4100, Vector Laboratories, Burlingame, CA, United States). The immunoreaction was visualized with an Axioplan Zeiss light microscope (Carl Zeiss), and the micropictures were captured using a digital camera (AxioCam MRc5, Carl Zeiss) via AxioVision Release 4.8.2—SP2 Software (Carl Zeiss Microscopy GmbH, Jena, Germany) (scale bar 50 μm).

For proteomic analysis, Filter-Aided Sample Preparation (FASP) of liver and prostate homogenates was performed by using an adapted protocol of Sielaff et al.(2017). Briefly, 40 μg of proteins were transferred onto a Nanosep 10-kDa-cutoff filter (Pall Corporation, Portage, MI, United States) previously rinsed with 100 μL of 1% (v/v) formic acid (FA), the filter was washed two times with 200 μL urea buffer (8 M urea and 100 mM Tris, pH 8.5 in Milli-Q water) to remove the detergents present in the extraction buffer. 100 μL of dithiothreitol (DTT) solution (8 mM DTT in urea buffer) was added to the filter unit, covered with tin foil, and incubated for 15 min at 56 °C in a dry block heating system to reduce proteins. After two washes with 100 μL of urea buffer, the proteins were alkylated by adding 100 μL of 2-iodoacetamide (IAA) solution (50 mM IAA in urea buffer) and incubating in the dark for 20 min at room temperature. 100 μL of ammonium bicarbonate (AB) buffer (50 mM in milliQ water) was added to the filter for the buffer exchange. The digestion was performed with trypsin: a trypsin stock solution (1 μg/μL) was prepared, dissolving the enzyme in 50 mM acetic acid; 50 μL of trypsin working solution (0.01 μg/μL in AB buffer) was added onto the filter and incubated overnight at 37 °C in a wet chamber. The peptide mixture was collected by centrifugation and acidified with 10% (v/v) trifluoroacetic acid (TFA) to a final TFA concentration of 0.2% (v/v). The peptides obtained with the FASP protein digestion were analyzed on a nanoflow liquid chromatography (nLC) system (Ultimate 3000 UHPLC) coupled to an Orbitrap Fusion mass spectrometer (Thermo Fisher Scientific, Waltham, MA, United States), operating in positive ionization mode with spray voltage set at 1.7 kV and source temperature at 275 °C, equipped with a nanoEASY-Spray ion source. The solvents used for the chromatographic system were: solvent A, consisting of 100% water with 0.1% formic acid, and solvent B, consisting of 80% acetonitrile with 0.1% formic acid. Samples were loaded on a PepMap100 C18 pre-column cartridge (5 µm particle size, 100 Å pore size, 300 µm i.d. × 5 mm length, Thermo Fisher Scientific, Waltham, MA, United States) and separated on an EASY-Spray PepMap RSLC C18 column (75 μm i.d., 3 μm particle size, 100 Å pore size, Thermo Fisher Scientific, Waltham, MA, United States) at a flow rate of 300 nL/min and a temperature of 40 °C. The gradient was as follows: from 5% to 15% B in 20 min, from 15% to 35% B in 20 min, from 35% to 50% B in 5 min, and from 50% to 90% B in 2 min, with total LC runtime of 60 min. Data were acquired with Xcalibur software v4.4 (Thermo Fisher Scientific, Waltham, MA, United States). Full mass scans (MS1) were recorded in the Orbitrap analyzer at a resolving power of 120,000 (at m/z 200), with a mass range of m/z 350–1,500, automatic gain control (AGC) target of 4.0 × 10⁵, and a maximum injection time of 50 ms. Data-dependent MS/MS (MS2) analysis was conducted in top speed mode with a 1-second cycle time, where the most abundant multiple-charged (²⁺-⁵⁺) precursor ions were selected for activation in order of abundance and detected in linear ion trap at rapid scan rate. Quadrupole isolation with a 1.6 m/z isolation window was used, and dynamic exclusion was enabled for 30 s after a single scan. For MS2, automatic gain control was 1.0 × 10³, and the maximum injection time was 35 ms. The fragmentation mode was the Higher-energy collisional dissociation (HCD) with 30% normalized collision energy. The raw data were processed using Proteome Discoverer version 2.5 (Thermo Fisher Scientific, Waltham, MA, United States) with the MS1 Precursor option. The post-translational modification (PTM) profile was set as follows: fixed cysteine carbamidomethylation (ΔMass: 57.02), variable methionine oxidation (ΔMass: 15.99). Non-specific cleavage was allowed to one end of the peptides, with a maximum of 2 missed cleavages and Trypsin enzyme specificity. The highest error mass tolerances for precursors and fragments were set at 10 ppm.

For quantification, Minora Feature Detector node was employed to detect chromatographic peaks, while Precursor Ion Quantifier node was used for precursor quantification, applying Unique peptides only. Normalization was performed based on the Total Peptide Amount. Protein abundance was calculated using the Top N Average method (N = 3), protein ratios were determined using a Pairwise ratio–based approach, and statistical significance was assessed using a Background-based t-test. Spectra were matched against the Rattus norvegicus (TaxID = 10,116) database downloaded from the Uniprot website (8,205 entries). A false discovery rate (FDR) threshold of 1% was applied at the protein level.

Western blot analysis was conducted to evaluate PECAM-1, NRF-2, HO-1, GPX4 and IL1R1 expression levels on prostatic tissue and PPAT. Samples were processed using RIPA buffer (Thermo Fisher Scientific, Paisley, United Kindom) in the presence of phosphatase and protease inhibitors (Thermo Fisher Scientific, Paisley, United Kindom) for protein extraction. Subsequently, each sample was homogenized and sonicated twice for 20 s using an ultrasonic probe (Vibra cell, Sonics, Newtown, United States). The protein samples (70 μg) were diluted in 2× Laemmli sample buffer (Bio Rad, Berkeley, CA, United States) and heated at 85 °C for 5 min. Proteins were separated via electrophoresis and then transferred as previously reported by Ciaffaglione et al. (2022). Membranes were incubated overnight with HO-1 (GTX101147, diluted 1:1000, GeneTex, Irvine, CA, United States), NRF-2 (ab62352, diluted 1:1000, Abcam), PECAM-1 (SAB5700639, Sigma Aldrich, St. Louis, MO, United States), GPX4 (ab231174, diluted 1:1000, Abcam) and β-actin (GTX109639, diluted 1:7000, GeneTex) primary antibodies. Appropriate secondary antibodies were used to detect blots (diluted 1:10000). Blots were scanned, and densitometric analysis was performed with the Odyssey Infrared Imaging System (LI-COR, Milan, Italy). Values were normalized to β-actin.

Concentration of non-protein thiol groups (RSH), reflecting about 90% of GSH cellular content, was measured in prostate tissue homogenates. RSH levels were evaluated by a spectrophotometric assay based on the reaction of thiol groups with 2,2-dithio-bis-nitrobenzoic acid (DTNB). DTNB solution and samples were mixed and incubated at room temperature for 20 min in the dark until a noticeable appearance of a yellow color was observed. After incubation, samples were centrifugated at 3,000 rpm for 10 min. The supernatant was collected and set in a black 96-well plate for measurement of absorbance in a microplate reader (Biotek Synergy-HT, Winooski, VT, United States) at λ = 412 nm. Experiments were conducted in quadruplicate. Results are expressed as pmoles/µL.

Levels of LOOH were evaluated through the oxidation of Fe²⁺ to Fe³⁺ in the presence of xylenol orange. Assay mixture consisted of 200 µg of sample (tissues homogenates), 100 µM xylenol orange, 250 µM ammonium ferrous sulphate, 90% ethanol, 4 mM butylated hydroxytoluene and 25 mM H₂SO₄. Samples were incubated at room temperature for 30 min and finally the absorbance was measured at λ = 560 nm using a microplate reader (Biotek Synergy-HT, Winooski, VT, United States). Calibration was obtained using hydrogen peroxide (0.2–20 µM). Results were expressed as fold change.

IL-10 levels were assessed on prostate tissues homogenates using enzyme-linked immunosorbent assays (ELISA) (RBELR-IL10-CL-1, RayBiotech, Norcross, GA, United States) according to the manufacturer’s instructions. The absorbance was measured using a microplate reader (Biotek Synergy-HT, Winooski, VT, United States). The same technique was used for Adiponectin and Leptin levels quantification on PPAT homogenates (JXE0758Ra-96, JXE0561Ra-96 Shanghai, Cina). The results are expressed as pg/mL for IL-10, ng/mg prot for Adiponectin, and pg/mg prot for Leptin.

Protein abundance ratios between groups were calculated from label-free quantification values and log₂-transformed prior to visualization. Heatmaps were generated to display relative abundance patterns across conditions, including proteins that did not reach statistical significance in differential analysis. These non-significant changes are presented for descriptive and hypothesis-generating purposes only and were not interpreted as confirmed effects.

The statistical analyses were performed using the software Prism v. 8.0.1 (GraphPad Software, San Diego, CA, United States) and Microsoft Excel v. 16.54. At least three independent experiments were performed for each analysis. Statistical significance (p < 0.05) of the differences between the experimental groups was determined by Fisher’s method for analysis of multiple comparisons. For comparison between treatment groups, the null hypothesis was tested by either a single-factor analysis of variance (ANOVA) for multiple groups or an unpaired t-test for two groups, and the data are presented as mean ± SEM.

Body weight was monitored throughout the entire treatment period, and no significant differences were observed among the experimental groups (Figure 2). This finding suggests that the treatments did not exert adverse systemic effects, thereby supporting the reliability of the subsequent analyses focused on prostate-specific outcomes. Indeed, the prostate did not exhibit a significant increase in volume, most likely reflecting an early stage of benign prostatic hyperplasia (BPH), in which only mild clinical symptoms are typically observed and substantial prostatic enlargement has not yet occurred.

The morphological examination based on H&E staining showed a prostate histoarchitecture in the control group (CTRL) or following testosterone (T) and PWE treatment (T + PWE). Figures 3, 4 show sections of the ventral prostate (A and B, ×10 and ×20 magnification, respectively). In the control group, the acini showed regular sizes and shapes with epithelium consisting of a single layer of cuboidal or columnar cells or a pseudostratified columnar epithelium resting on the basal lamina and with few villous projections. The cells’ nuclei are basally located. Testosterone (T) and PWE groups are characterized by columnar cells with basal nuclei and perinuclear cytoplasmic clear areas, villous projections (thick arrows) more in PWE than in testosterone, and pseudostratified columnar epithelium lining (dashed arrows).

To verify whether the histological alterations observed were associated with oxidative stress and inflammatory processes, the immunoreactivity and distribution of key proteins involved in these pathways, specifically PECAM-1, NRF2, and HO-1, were evaluated.

Immunodetection of PECAM-1, NRF2, and HO-1 in ventral prostate sections showed significant molecular alterations following testosterone administration and a modulatory effect induced by co-treatment with PWE (Figure 5). Testosterone exposure led to a pronounced reduction in PECAM-1 expression, consistent with endothelial impairment and early vascular remodeling events associated with early prostatic hyperplasia. In contrast, NRF2 and HO-1 levels were markedly upregulated in T-treated prostates, indicating activation of antioxidant and cytoprotective pathways in response to androgen-induced oxidative stress. Remarkably, co-treatment with PWE restored PECAM-1, NRF2, and HO-1 expression toward control levels, reflecting the extract’s capacity to mitigate testosterone-driven oxidative imbalance and preserve redox homeostasis within prostatic tissue (Figure 5). Western blot analysis further supported the immunohistochemical observations, confirming the modulatory effects of PWE co-treatment on testosterone-induced alterations. Quantitative protein evaluation revealed a significant decrease in PECAM-1 expression following T administration, consistent with endothelial dysfunction and vascular alteration within the prostatic microenvironment. Conversely, NRF2 and HO-1 protein levels were increased in T-treated groups, reflecting the activation of compensatory antioxidant defenses in response to oxidative stress. Importantly, co-treatment with PWE effectively normalized the expression of all three markers, restoring PECAM-1, NRF2, and HO-1 protein levels to values comparable to controls (Figures 6A,B). Analysis of redox biomarkers revealed that intracellular levels of reduced glutathione (GSH), measured as total thiol groups (RSH), remained unaltered across all experimental groups, indicating that neither testosterone administration nor PWE treatment significantly affected the cellular thiol antioxidant pool (Figure 6C). Conversely, lipid hydroperoxide (LOOH) levels exhibited a slight decrease in the PWE co-treatment group, reaching values marginally below those observed in control group (Figure 6D). This mild reduction suggests that PWE may exert a modest protective effect against lipid peroxidation, contributing to the maintenance of membrane integrity and redox homeostasis under testosterone-induced oxidative conditions. Moreover, IL1R1 expression was also increased upon T treatment, suggesting enhanced inflammatory signaling within the prostatic tissue, while PWE co-treatment restored protein levels similarly to untreated group (Figure 6F). In addition, IL-10 levels measured by ELISA revealed a slight but significant reduction after testosterone administration, consistent with a shift toward a pro-inflammatory microenvironment (Figure 6G). Notably, no alterations in IL-10 secretion were observed in the T + PWE group compared to controls, indicating that PWE co-treatment prevented the testosterone-induced decline of this anti-inflammatory cytokine.

Results from volcano plot analysis showed that comparing the protein expression profiles in the testosterone group with the control group, and testosterone group + PWE with the testosterone group, both down-expressed and overexpressed proteins were highlighted (Figures 6H,I).

In particular, several proteins such as Cytochrome c, Ubiquitin carboxyl-terminal hydrolase isozyme L1 (UCH-L1), Thioredoxin domain-containing protein 12, were down-expressed in testosterone group respect to control group. Such proteins may act as tumor suppressors counteracting oxidative stress (Figure 6H).

Evaluating the effect induced by PWE in the testosterone group we have highlighted that PWE treatment is able to counteract the effect of modulating the expression of Cytochrome c, Ubiquitin carboxyl-terminal hydrolase isozyme L1 (UCH-L1), Thioredoxin domain-containing protein 12 proteins, bringing the values back to those of the control. Moreover, PWE treatment in testosterone group, induced over-expression of Superoxide dismutase (SOD), Histone H4 and ATP synthase Subunit d, proteins that assisting in DNA repair during DNA replication and acting as superoxide anion scavenger in the prostate can reduce oxidative stress caused by testosterone (Figure 6I).

Data obtained in our experimental conditions highlighted also that Glutathione S-transferases (GSTs), and Tumor-associated calcium signal transducer 2 (TROP-2) were down-expressed in testosterone group + PWE respect to testosterone group. These results suggest that in testosterone group the expression of GSTs can represent an adaptive response of the prostate to testosterone, while TROP-2 expression could serve as a potential prognostic biomarker acting as intracellular calcium signal transducer linked to processes like self-renewal, proliferation and invasion. The treatment with PWE was able to reduce the expression of these proteins highlighting its ability to counteract the effect of testosterone and to decrease prostate cells proliferation (Figure 6I).

The heatmap summarizes how testosterone alone and T + PWE modulate the abundance of multiple proteins involved in cytoskeletal organization, metabolism, redox balance, and stress responses relative to control (Figure 6J). Many proteins are clearly upregulated by testosterone, as indicated by the intense red color in the T column, whereas the corresponding T + PWE values are often lighter, suggesting that the extract blunts several testosterone-driven changes. The heatmap also includes proteins below significance threshold to illustrate global patterns.

Periprostatic adipose tissue (PPAT), the fat surrounding the prostate, is increasingly recognized as an active endocrine and paracrine organ that influences BPH. This fat can release factors like hormones and cytokines that affect prostate reactivity and growth, potentially promoting BPH development and worsening urinary symptoms.

PPAT influences BPH releasing inflammatory mediators, including cytokines and hormones, that can increase prostate tissue’s growth and reactivity, contributing to BPH development. PPAT produce adipokines, such as leptin and adiponectin, and other signaling proteins that can affect various physiological processes in the prostate and that play a role in inflammation and tissue growth (Fibbi et al., 2010; Sacca and Calvo, 2022).

In the PPAT, a distinct redox-related response was observed (Figures 7A–D) following hormonal and phytochemical treatments. Specifically, testosterone administration led to a significant upregulation of HO-1, NRF2, and GPX4 expression, indicating the activation of adaptive antioxidant and cytoprotective mechanisms within the adipose microenvironment. Conversely, co-treatment with PWE markedly reduced the expression levels of all three markers compared to testosterone alone, restoring them toward basal or control-like values. These findings suggest that PWE counteracts testosterone-induced redox imbalance in PPAT.

Measurement of adipokines as adiponectin and leptin in PPAT samples showed that PWE co-treatment was able to increase adiponectin levels in PPAT compared to testosterone alone, indicating a shift toward a more anti-inflammatory and metabolically favorable adipose profile. In the context of androgen exposure, where PPAT typically acquires a pro-inflammatory secretory pattern, the upregulation of adiponectin suggests that PWE can partially restore protective adipokine signaling within PPAT tissue samples (Figure 7E). This effect is particularly relevant given adiponectin’s known ability to counteract cytokine-driven inflammation, improve insulin sensitivity, and modulate tissue remodeling, implying that higher adiponectin in PPAT may attenuate paracrine cues that promote prostatic growth and reactivity (Choi et al., 2020; Karnati et al., 2017). On the other hand, no significant alterations were observed in leptin levels analyzed in PPAT samples (Figure 7F).

To investigate whether testosterone, although not eliciting an increase in prostate volume, induced metabolic alterations at the hepatic level, a proteomic analysis was conducted. Results from volcano plot analysis showed that comparing the protein expression profiles in the testosterone group with the control group, down-expressed and overexpressed proteins were highlighted (Figures 7G,H).

In particular, Glutathione S-transferases (GSTs), Mitochondrial ATP synthase and Cytochrome c oxidase were down-expressed in testosterone group respect to control group. These results highlight that dysregulation of GSTs can lead to the accumulation of toxic metabolites, DNA damage and increased inflammation, as well as of Mitochondrial ATP synthase and Cytochrome c oxidase, are a consequence of oxidative stress (Figure 7G).

Several proteins such as, Cytochrome P450 A14 (CYP4A14), Cytochrome P450 A2, or CYP2A, 17-β-hydroxysteroid dehydrogenase type 6 (HSD17B6), Retinol dehydrogenases (RoDHs), Carnitine palmitoyltransferase (CPT), Elongation factor 1 alpha (EF-1α), Alkylglycerol monooxygenase (AGMO), claudin-3, Stearoyl-CoA desaturase (SCD), Caspase-6 were overexpressed in testosterone group respect to control group. These results highlight that CYP4A14 overexpression may be related to liver lipid accumulation and inflammation, while CYP2A enzymes in rats can catalyze testosterone 7α-hydroxylation (Figure 7G). It has been reported that enzymes such as liver HSD17B6, RoDHs, claudin-3 and EF-1α are induced by testosterone because they are involved in the regulation of androgen metabolism while Caspase-6 plays a critical role in liver injury promoting inflammation and cell death (Schiffer et al., 2018).

Results from volcano plot analysis showed that comparing the protein expression profiles in the T + PWE group with the testosterone group, overexpressed proteins are highlighted (Figure 7H). In particular, Mitochondrial ATP synthase, Thioredoxin, glutathione S-transferase (GST) and glutathione peroxidase (GPx) were overexpressed highlighting that PWE treatment was able to induce synthesis of proteins involved in maintaining mitochondrial membrane potential, in detoxification of harmful substances, in protection of cells from oxidative damage. These enzymes are part of the liver’s antioxidant defense system and reducing peroxides they promote regeneration of glutathione (GSH) to maintain cellular health (Asantewaa et al., 2024).

In liver, testosterone also induced a coordinated proteomic response, characterized by increased expression of cytoskeletal and polarization markers, lipid and xenobiotic-metabolizing enzymes, and proteins involved in translation, protein turnover, immune function, and apoptosis (Figure 7I). PWE co-treatment attenuated many of these testosterone-dependent changes, although several metabolic and signaling proteins remained elevated or were further increased, indicating that PWE potentially modulates rather than uniformly suppresses the hepatic response to androgen stimulation. The heatmap also includes proteins below significance threshold to illustrate global patterns.