Inflammation-free foreskin samples were collected from subjects who underwent circumcision, with their written consent. The foreskin was thoroughly disinfected with iodine, rinsed repeatedly with saline, and treated with a penicillin and streptomycin solution (4 × 10⁶ IU/L) for 10 min, followed by treatment with chloramphenicol (25 mg/L) for 5 min. The samples were then washed with PBS and placed in 1 mol/L NaCl, shaken at 200 rpm at 37°C for 24 h. Afterward, the epidermis was removed, and the dermis was treated with 0.5% sodium dodecyl sulfate (SDS) at room temperature for 2 h to remove cells. The decellularized dermal matrix was rinsed several times with PBS and then placed in PBS at 4°C for 48 h to eliminate residual SDS. Finally, the ADM was rinsed with PBS and carefully trimmed into a rectangular shape using scissors. Non-decellularized foreskin dermis was used as a control for subsequent histological analyses.

Both the non-decellularized foreskin dermis and ADM were fixed in 4% PFA overnight at 4°C. The samples were then paraffin-embedded, sectioned, de-waxed, and rehydrated. Hematoxylin and eosin (H&E) staining and Masson's trichrome staining were performed using kits from Pinuofei Biological (S191003 for H&E, S191006 for Masson's trichrome), following the manufacturer's instructions. The results were captured using a Nikon ECLIPSE-Ci microscope equipped with a digital camera.

With parental consent, urethral mucosa tissues were collected from children undergoing repair surgery for perineal hypospadias. The dissected tissues (∼0.5 cm × 0.5 cm) were immersed in DPBS containing penicillin/streptomycin (100,000 U/L) and amphotericin B (1 mg/L) and transported on ice to the laboratory. The tissues were then incubated with 2.24 U/ml Dispase II (Roche) overnight at 4°C. The following day, the mucosal layer was separated from the submucosal layer with forceps, cut into small pieces, and digested with 0.25% trypsin-0.02% EDTA at 37°C for 5 min. The tissues were then placed in DPBS, gently pipetted to form a cell suspension, and passed through a 100-mesh cell filter. The cell suspension was centrifuged at 1,000 rpm for 5 min, and the pellet was resuspended in DMEM/F-12 (1:1, Hyclone). All experiments were approved by the Ethics Committee of Jiangmen Maternity and Child Health Care Hospital ([2022]056).

For cell culture, the cells were seeded into laminin-coated 24-well plates at a density of 5 × 10⁴ cells/ml and cultured in DMEM + 10% FBS at 37°C with 5% CO₂. After 24 h, the culture medium was refreshed, and the medium was changed every 3 days. Once the cells reached 80%–90% confluence, they were passaged using 0.25% trypsin-0.02% EDTA at the same density.

MTT assay was performed to assess cell viability. Cells were seeded into 96-well plates at a density of 1 × 10⁴ cells per well (200 μl) and cultured for 48 h. Afterward, 10 μl MTT working solution (5 mg/ml, Yeasen) was added to each well, and the plates were incubated at 37°C for 4 h. Then, 150 μl DMSO was added to each well, followed by 10 min of agitation at 37°C. Finally, OD₄₉₀ values were measured using a microplate reader. The assays were performed in triplicate and repeated independently three times.

2 × 10⁴ cells were seeded onto each coverslip, which was placed in a 24-well plate and cultured overnight. The cells were then rinsed with PBS and fixed with 4% PFA for 30 min at 4°C, followed by another PBS wash. Cells were permeabilized with 0.5% Triton X-100/PBS for 10 min, washed with PBS three times, and blocked with 2% BSA for 1 h at room temperature. After blocking, the cells were incubated with primary antibody solutions overnight at 4°C. The following day, the cells were washed with PBS, incubated with secondary antibodies for 2 h at room temperature, washed with PBS, counterstained with DAPI, mounted, and imaged using a Nikon DS-Ri2 microscope. The antibodies used were Rabbit anti-CD31 (11265-1-AP, Proteintech, 1:200), Rabbit anti-Cytokeratin 14 (CK14, 10143-1-AP, Proteintech, 1:200), and Goat anti-Rabbit IgG (H + L) Alexa Fluor 488 (SA00006-2, Proteintech, 1:500).

2 × 10⁴ cells were seeded onto each coverslip, placed into a 24-well plate, and cultured overnight. The cells were then incubated with 1:1000 diluted CFSE/Hoechst solution for 4 h at 37°C, followed by PBS washing and imaging using a Leica confocal microscope.

5,000 cells were seeded into each well of a 96-well plate containing trimmed ADM and cultured until the indicated time points. At the time of assay, 10 μl CCK-8 solution was added to each well and incubated for 3 h at 37°C. The medium was transferred to another 96-well plate, and OD₄₅₀ values were measured using a microplate reader. The assays were performed in triplicate.

To assess the mechanical properties of foreskin ADM, urothelial-cell-seeded ADM, and urothelial cell sheets, samples were trimmed into strips (20 × 50 mm) and clamped onto the sample holders of an STB-1400 Cell Stretching System (Strex Cell) at a speed of 10 mm/min until tearing occurred. Maximum tensile strength and pull forces were recorded and quantified. The experiment was repeated three times independently.

To evaluate the host response to implanted ADM constructs, Male nude mice weighing 22–25 g were obtained from the Animal Experiment Center of Xiamen University and housed under standard conditions with controlled temperature (20–24°C), humidity (40%–60%), and a 12-h light/dark cycle, with free access to food and water. For the implantation experiments, sterile 8 × 8 mm ADM samples were surgically implanted into dorsal subcutaneous pockets of C57BL/6 mice, with three mice per experimental group. Following implantation, tissue samples were collected weekly and subjected to histological processing for H&E staining to assess inflammatory responses, ELISA for quantification of IL-6 and TNF-α cytokine levels, and CD31 immunohistochemistry to evaluate vascular ingrowth, while macroscopic observations were performed in parallel to document hair regrowth patterns throughout the study duration.

Statistical analysis and graph generation were conducted using GraphPad Prism (v8.0). One-way ANOVA followed by post hoc Tukey's HSD test was used for statistical analysis. Significance was determined at *P < 0.05, **P < 0.01, with ***P < 0.001 and ****P < 0.0001.

To assess the efficacy of decellularization, we conducted histological analyses on both untreated foreskin dermis and prepared foreskin ADM. H&E staining revealed that untreated foreskin dermis contained hematoxylin-positive nuclei (indicated by black arrowheads), while foreskin ADM lacked hematoxylin signals, indicating the absence of cellular material (Figures 1A,B). Additionally, to evaluate the collagen content, we performed Masson's trichrome staining. The results demonstrated that although the collagen content (blue staining) in foreskin ADM was slightly reduced, the majority of dermal collagen was retained compared to untreated foreskin dermis (Figures 1C,D). These results confirm the successful preparation of foreskin ADM. Meanwhile, the detection results of hydroxyproline (HYP) content for collagen and glycosaminoglycan (GAG) showed no significant differences between foreskin ADM and control foreskin dermis (Supplementary Figures S1F–G).

To characterize the isolated primary urothelium-derived cells, we first monitored their growth under a bright-field microscope over 96 h. The cells exhibited typical urothelial morphology and reached 100% confluence within 48 h after initial inoculation. The growth potential was maintained even after being passaged for five generations (Figure 2A). MTT assays were performed to quantify the viability of these cells, showing that their viability was slightly reduced after five passages (Figure 2B). Furthermore, IF staining revealed that these cells were negative for the endothelial marker CD31 but expressed the epithelial marker CK14 (Figure 2C), confirming their identity as urothelium-derived cells.

To assess any potential adverse effects of foreskin ADM on primary urothelium-derived cells, we soaked the ADM in culture medium to create a conditioned medium, which was then added to the urothelial cell culture. After 24 or 48 h, MTT assays were conducted to measure cell viability. The results indicated that the ADM-conditioned medium had minimal impact on the viability of primary urothelium-derived cells (Figure 3), demonstrating that foreskin ADM is biocompatible and safe for use with urothelium-derived cells.

To directly evaluate the growth of primary urothelium-derived cells on foreskin ADM, we labeled the urothelium-derived cells with CFSE (Figures 4A,B). These fluorescence-labeled cells were inoculated onto the foreskin ADM for co-culture. After 96 h, we observed that the majority of labeled cells remained viable (Figures 4C,D), with continued cell division throughout the 4-day culture period, as shown by the CCK-8 assay results (Figure 4E). Furthermore, we examined the biomechanical properties of the ADM, ADM with growing cells, and the urothelial cell sheet. The findings indicated that the maximum tensile strength and pull force of the cell-growing ADM were significantly higher than that of native ADM and urothelial cell sheets (Figures 4F,G). These results suggest that primary urothelium-derived cells can grow robustly on foreskin ADM.

To systematically assess the host response to ADM implantation, we established three experimental groups: Group 1 (ADM subcutaneous implantation), Group 2 (ADM dorsal subcutaneous implantation with cells), and Control (Sham operation). The ADM scaffolds were surgically implanted into the dorsal subcutaneous space of mice, with tissue samples harvested at 2 weeks post-operation (n = 3 per group). Histological evaluation through H&E staining and cytokine analysis via ELISA demonstrated comparable levels of IL-6 and TNF-α expression between experimental and control groups, indicating minimal inflammatory response (Supplementary Figures S1A,D–E). CD31 immunohistochemical analysis demonstrated higher expression levels in experimental groups compared to controls in the study period (Supplementary Figure S1C). Macroscopic examination confirmed the hairless nature of ADM implantation sites, in contrast to the normal hair growth observed in control areas (Supplementary Figure S1B). These collective findings suggest that foreskin-derived ADM exhibits excellent biocompatibility and promotes vascularization while maintaining its critical hairless property for urethral tissue engineering applications.