Application of decellularized tilapia skin in rabbit urethral reconstruction: an experimental study

Introduction

In the urethra tissue, urothelial epithelial cells, smooth muscle cells and their associated extracellular matrix (ECM) form a highly ordered elastic luminal structure which are vital for urine and semen transport (1,2). Trauma, inflammation, congenital anomalies and medically induced injuries can cause urethral strictures, urethral fistulas, and other conditions that require surgical reconstruction of the urethra in severe cases. However, there is no single urethral reconstruction procedure that is accepted by all clinicians, and various postoperative complications are common. Urethral reconstruction remains a clinical challenge today (3).

Currently, the gold standard in urethroplasty to treat urethral stricture are autologous grafts from buccal mucosa or genital skin (4,5). Among them, buccal mucosa has greatly improved the success rate of urethral reconstruction over the past two decades due to its excellent physicochemical properties and has become the preferred tissue for urethral reconstruction (1,6). However, patients undergoing replacement urethroplasty using buccal mucosal grafts or tissue flaps may have donor site lesions or insufficient donor material to meet reconstruction needs (7).

Tissue engineering (TE) combines strategies from biology, materials science, medicine, and engineering and provides a therapeutic approach to repair damaged tissues and organs using ECM or synthetic materials (8,9). The ideal material not only has reliable biocompatibility but also provides a biochemical and physical environment similar to that of native tissue, which promotes cell adhesion, proliferation, and production of ECM (10,11). In experimental and clinical urogenital TE, collagen-based materials such as small intestinal submucosa (SIS) and bladder acellularized matrix (BAM) have been successfully used to repair short-segment urogenital defects (4,7,12-14).

Tilapia is a tropical freshwater fish. During the processing of tilapia, a large number of by-products are discarded. These wastes account for 50–70% of the weight of the entire fish (15). The wide geographical distribution and consistent availability of tilapia make large-scale production of tilapia-derived biomaterials economically feasible. Effective utilization of these fish skins can reduce costs while also reducing environmental pollution (16).

Previously, Lv et al. (17) showed that tilapia skin decellularized matrix could promote the healing of large acute wounds in rats by directing cell infiltration, and angiogenesis, regulating the expression and secretion of inflammatory and skin repair-related factors as well. Lau et al. (18) used decellularized tilapia skin (DTS) in a rat cranial defect model to promote bone regeneration. The major component of DTS is collagen which is an important component of connective tissue, accounting for 25–30% of total mammalian protein, and is less immunogenic, making it widely used in TE (19,20) due to circumvent of religious constraints and the risk of zoonosis between mammalian species.

To explore an ideal alternative material for urethroplasty in this study, we decellularized fresh tilapia skins via combinations of physical, chemical and enzyme treatments, and then conducted a randomized controlled trial using DTS for urethroplasty in rabbits urethral defect model with SIS collagen-based scaffold as a positive control, to verify its effect on urethral defect reconstruction. We present this article in accordance with the ARRIVE reporting checklist (available at https://tau.amegroups.com/article/view/10.21037/tau-24-598/rc).

Methods

Chemical and biological reagents

Phosphate-buffered saline (PBS), dispase, Pierce universal nuclease, PicoGreen assay kit and type I collagenase were obtained from Thermofisher Scientific, Waltham, MA, USA. SIS was purchased from Cook Biotech, West Lafayette, IN, USA. Anti-α-smooth muscle actin (α-SMA) was purchased from Novus Biologicals, Englewood, CO, USA. Anti-tumor necrosis factor-α (TNF-α) was purchased from Abcam, Waltham, MA, USA. Secondary antibody was purchased from Cell Signaling Technology, Danvers, MA, USA. Other reagents were obtained from Sigma-Aldrich, Burlington, MA, USA, unless otherwise stated.

Preparation of DTS

The method for fish skin decellularization used in this study is followed our previous report with some modifications (18). Freshly harvested skins from Nile tilapia (Oreochromis niloticus) descaled were cleaned and then immersed into 0.5 mol/L NaOH solution and shaken at 100 rpm for 3 hours and then rinsed with deionized (DI) water until the pH is less than eight. Subsequently, they were then immersed into 1% sodium dodecyl sulfate (SDS) solution for 6 hours. Once completion, the fish skins were scraped from the surface with a blade to physically remove the epidermis. After washed with DI water to remove the black residue. Nuclease 25 U/mL in PBS solution was used to remove nucleic acids from the fish skin for 3 hours. After treatment, the fish skins were thoroughly washed with DI water and dried in a freeze-dryer for 24 hours after 3 hours freeze in −80 ℃ freezer. Finally, the decellularized fish skins was cut into 1 cm × 1 cm pieces (Figure 1) after ethylene oxide sterilization.

Figure 1 The process of DTS preparation. DTS, decellularized tilapia skin; PBS, phosphate‐buffered saline; SDS, sodium dodecyl sulfate.

Characterization of DTS

Residual DNA content measurement

Freeze-dried DTS were cut into 5 mm × 5 mm pieces (n=3). Each piece was weighed and placed into a 1.5 mL centrifuge tube. DNA content was quantified using the Favprep^TM Tissue Genomic DNA Extraction Mini Kit (Chiyoda Science, Tokyo, Japan) to extract the DNA from the DTS samples and then quantified using the PicoGreen^®dsDNA Assay Kit according to the instructions to verify the success of the decellularization process.

Morphology characterization

The morphologies of the DTS and SIS were observed by scanning electron microscopy (SEM). After freeze dry, the samples were fixed to the stages using conductive adhesive. Thereafter the samples were spattered with an ultrathin platinum layer for 90s. Finally, DTS and SIS samples were observed by ultra-high resolution SEM (SU8000, HITACHI, Tokyo, Japan).

Mechanical property characterization

Uniaxial tensile tests were used to determine the mechanical properties of DTS and SIS. Samples were cut into 30 mm × 5 mm strips and mounted on a Universal Testing Systems (Instron 5594, Instron, Norwood, MA, USA). The tensile testing software recorded the dimensions of each specimen. The load sensor was calibrated and the specimens were pulled at a rate of 10 mm/min until failure. The results were recorded by Universal Testing Systems, the maximum tensile stress and Young’s modulus were compiled. The test is repeated at least three times for each set of specimens.

Degradation

DTS samples with a square size of 1 cm × 1 cm were prepared, lyophilized, accurately weighed (Wo), and placed under sterile conditions in 10 mL simulated body fluid (SBF) containing 50 U/mL type I collagenase (18) to simulate in vivo enzymatic degradation. Tubes were capped and incubated at 37 ℃. Every one hour, the solution was removed with a syringe, and the samples were lyophilized and reweighed (Wr). The degradation curve was plotted based on percentage weight loss [(Wo − Wr)/Wo] * 100%.

Animal experiment

Surgical procedure

Experiments were performed under a project license (No. wydw-2022-0463) granted by the Experimental Animal Ethics Committee at Wenzhou Medical University, in compliance with the National Institutes of Health’s Guidelines for the care and use of animals.

During the animal experiments, we used 24 New Zealand rabbits. The weight of rabbits were between 2.5–3.0 kg, and rabbits were housed in the animal facilities of Wenzhou Medical University. All surgical procedures were performed under aseptic conditions. Before surgery, DTS and SIS were given a 15-minute rehydrating soak in sterile saline prior to surgery. Sodium pentobarbital (30–50 mg/kg) was administered intravenously via the ear margins to anesthetize rabbits. After an 8 F catheter was inserted, the ventral skin of the penis was incised with a 1.5 cm cut, with each layer being incised in turn and separated down to the urethra (Figure 2A,2B). Group A (n=8) was implanted with a DTS; group B (n=8) was implanted with a SIS; group C (control group, n=8) was not implanted, and the urethral defect was closed directly using 6-0 poly-p-dioxanone suture (PDS) (Figure 2C). After repair, the 6-0 PDS was used to close layer by layer, and 5-0 prolene sutures were left in place to mark the ends of the wound and the catheter was removed (Figure 2D).

Figure 2 Animal surgical procedures. (A) Suspension of the penis, exposing the ventral side of the penis. (B) Separating the ventral penis layer by layer to create a rabbit urethral defect model. (C) Covered the defect with repair material. (D) Sutured wounds.

Histological, immunohistochemical, and histomorphometric analyses

Overdoses of sodium pentobarbital were administered through rabbit alar vein at 2 and 4 weeks postoperatively. The penises were excised intact for gross examination and histological processing. All tissue samples were fixed in 4% paraformaldehyde for 24 hours and then paraffin embedded. Tissue paraffin blocks were cut into sections with a thickness of 5 µm. The histological analysis was done by hematoxylin and eosin (H&E) staining for pathological examination. The angiogenesis in samples were evaluated by numbers of blood vessel formation a blood vessel area measured using ImageJ software as previous reports (21,22). In addition, collagen distribution was evaluated using Masson’s staining. The detection of smooth muscle was accomplished through α-SMA staining in immunohistochemistry (IHC). Tissue inflammatory response detected by TNF-α staining in immunofluorescence (IF).

Statistical analysis

All experimental data were presented as mean ± standard deviation. The manual tracking and measurement software used for histomorphometry was NIS-Elements (version 5.11.01). Quantitative analysis of Masson staining, IHC, and IF results was done by ImageJ software (version 1.53t). Experimental data were tested for differences by applying analysis of variance (ANOVA) and t-test using SPSS (version 22.0). P value <0.05 was considered statistically significant. The experimental data were processed using a double-blind method.

Results

In Figure 3, the surfaces and cross sections of DTS and SIS were shown. Compared to fresh tilapia skins, melanin were removed from the DTS surface and the appearance of DTS was similar as that of SIS (Figure 3A,3D). We measured the residual DNA content of DTS. The results showed that DTS retained 38.78±5.84 ng/mg (dry tissue) after the decellularization, a value lower than the 50 ng/mg specified in the industry standard (23). This indicates that the DTS was successful in reducing the DNA content after treatment while maintaining the structural integrity of the fish skin as show in Figure 3. As shown in SEM images of DTS and SIS, DTS successfully preserved its three-dimensional microstructure during processing and appeared to be looser in its internal structure than SIS. In DTS, the collagen fibres showed a clear laminar arrangement, along with the presence of many pores, which render the high permeability properties.

Figure 3 Visual appearance and SEM images of DTS and SIS. (A,D) Visual appearance of the outer surface of DTS and SIS. (B,E) SEM images of the surface of DTS and SIS. (C,F) SEM images of the cross section of DTS and SIS. DTS, decellularized tilapia skin; SEM, scanning electron microscopy; SIS, small intestinal submucosa.

Through measument using a macrometer, it was found that the thickness of DTS and SIS were 0.39±0.04 and 0.22±0.01 mm, respectively. In order to compare the mechanical properties of DTS and SIS, uniaxial tensile tests were performed. From the results, it revealed excellent ductility of DTS with a maximum tensile stress of 20.97±4.46 MPa and Young’s modulus of 130.43±7.00 MPa, while the SIS presented less ductility with a maximum tensile stress of 37.32±3.97 MPa and Young’s modulus of 615.13±7.43 MPa which is in line with its dense structure (Figure 4A). In vitro degradation test, DTS experienced 49% and 86% weight lost after 2 and 4 hours in SBF with 50 U/mL collagenase, respectively. After 6 hours DTS was completely degraded, while 51.26%±5.42% of SIS still remained under the same treatment (Figure 4B) resulted from its dense structure.

Figure 4 The physical properties and degradation profile of DTS and SIS. (A) Representative stress-strain curves for DTS and SIS. (B) Degradation profile of DTS and SIS in 50 U/mL collagenase. DTS, decellularized tilapia skin; SIS, small intestinal submucosa.

All the experimental animals survived throughout the experimental phase. From morphological observation, the penile wounds in group A healed effectively without any significant penile body curvature and the absence of any fistula openings on the surface (Figure 5A). On the other hand, group B experienced a little curvature of the penis with a slight contracture visible in the wound, although the wound healed well overall (Figure 5B). Conversely, group C suffered from an inadequately healed penile wound featuring severe scarring, substantial contracture, and a ventral fistula opening that adversely impacted erectile function (Figure 5C).

Figure 5 Post-operative penile tissue specimens. (A) The wound healed well in group A. (B) Local contracture was seen on the ventral side of the penis in group B. (C) Severe contracture of the penis was seen in group C, with urethral fistula visible on the ventral side. Group A: implanted with a DTS. Group B: implanted with a SIS. Group C: not implanted. DTS, decellularized tilapia skin; SIS, small intestinal submucosa.

H&E staining at week 2 and week 4 in Figure 6 revealed an intact, stratified uroepithelium on the surface of the excised ventral urethra, with organized smooth muscle and blood vessels visible beneath the uroepithelium. At 2 weeks postoperatively, neither the DTS nor the SIS patch had completely disintegrated in the rabbit penis. In group A, the DTS in the penis showed flaky disintegration and resulted in local deposition of collagen fibers with fibroblast infiltration, but no significant inflammatory cells were detected around DTS fragments (Figure 6A). In group B, it showed significant delamination of the SIS patch under high magnification, with heavy infiltration of inflammatory cells around the SIS laminars (Figure 6B). Group C showed typical loose connective tissue with reticular fibrous structures and a few local inflammatory cells under microscopy (Figure 6C). At 4 weeks postoperatively, in both treatments (DTS and SIS implantations), there were no significant rejections were observed as shown in Figure 6D,6E. The collagen fibers between the tissues had deposited significantly to form a lamellar collagen structure, different from the sparse reticular structure in group C (Figure 6F). In group B, a few SIS patches were still visible in the microscopic field, but not completely fused to the tissue (Figure 6E). We examined the neovasculature in defect area in each group at 4 weeks postoperatively (as characterized by endothelialized lumens containing red blood cells) under a high-power field (HPF) microscope (Figure 7). In terms of the number of blood vessels formation, there was significant difference in group A (5.81±2.61/HPF) compared to group C (3.31±2.44/HPF) (Figure 7A) (P=0.006). There was no significant difference between group A and B (P=0.39). Also, there was significant difference in vascular area in group A [(3.20±1.96)×10⁴ pixel/HPF] compared to the group C [(1.47±1.59)×10⁴ pixel/HPF] (Figure 7B) (P=0.04). Similarly, there was no significant difference between group A and B in term of vascular area (P=0.88).

Figure 6 Images of H&E staining of penile tissue post-surgery. (A-C) H&E staining of penile cross-section in group A, B, C at 2 weeks post-surgery. (D-F) H&E staining of penile cross-section in group A, B, C at 4 weeks post-surgery. Group A: implanted with a DTS. Group B: implanted with a SIS. Group C: not implanted. H&E, hematoxylin and eosin; DTS, decellularized tilapia skin; SIS, small intestinal submucosa.

Figure 7 The results were quantified by pathological staining of postoperative penile tissues. (A,B) The quantification of microvessels’ numbers (A) and area (B). (C) Image analyses of inflammatory response at 2 and 4 weeks. (D) Changes in collagen content in the tissues of each group at 2 and 4 weeks post surgery. (E) After 4 weeks, the number of positive pixel points for each group was calculated by image analysis of α-SMA IHC staining. α-SMA, α-smooth muscle actin; DTS, decellularized tilapia skin; IHC, immunohistochemistry; SIS, small intestinal submucosa.

To clarify the severity of inflammation in each group, we detected TNF-α through IF (Figure 8). The results showed that the fluorescence intensity of TNF-α was weak in group A in where DTS was used for defect repair at both 2 and 4 weeks postoperatively (Figure 7C), and on the contrary, the fluorescence intensity of TNF-α in group B in where commercialized SIS was used for urethral defect repair was the strongest among those groups at both 2 and 4 weeks. At 4 weeks postoperatively, the fluorescence intensities increased in all groups compared with those at 2 weeks earlier, which may be related to the enhancement of background fluorescence due to the increases of collagen fibres in the tissues.

Figure 8 Images of TNF-α IF staining of penile tissue post-surgery. (A-C) TNF-α IF staining of penile cross-section in group A, B, C at 2 weeks post-surgery. (D-F) TNF-α IF staining of penile cross-section in group A, B, C at 4 weeks post-surgery. Group A: implanted with a DTS. Group B: implanted with a SIS. Group C: not implanted. IF, immunofluorescence; TNF-α, tumor necrosis factor-α; DTS, decellularized tilapia skin; SIS, small intestinal submucosa.

Masson’s staining was used to assess the deposition of collagen fibers during defect repair (Figure 9). At 2 and 4 weeks post surgery, collagen fibers significantly deposed over time in the penile interstitium of groups A and B. The area of collagen fibers in group A was (14.56±0.29)×10⁶ pixels/HPF, is the highest among those groups at each time points (2 and 4 weeks post surgery) (Figure 7D).

Figure 9 Images of Masson staining of penile tissue post-surgery. (A-C) Masson staining of penile cross-section in group A, B, C at 2 weeks post-surgery. (D-F) Masson staining of penile cross-section in group A, B, C at 4 weeks post-surgery. Group A: implanted with a DTS. Group B: implanted with a SIS. Group C: not implanted. DTS, decellularized tilapia skin; SIS, small intestinal submucosa.

At 4 weeks postoperatively, the hyperplastic collagen tissue was analyzed by IHC staining, and assess the proliferation of α-SMA positive cells as well (Figure 10). At ×400 magnification, the cells in the penile interstitium of group A were organized and dense, and the mean area of reached (16.13±1.72)×10⁵ pixels/HPF, which was significantly more hyperplastic compared to (13.42±1.27)×10⁵ pixels/HPF in group B and (13.70±3.51)×10⁵ pixels/HPF in group C (Figure 7E) (A to B, P=0.006; A to C, P=0.002).

Figure 10 Images of α-SMA IHC staining of penile tissue post-surgery. (A-C) α-SMA IHC staining of penile cross-section in group A, B, C at 4 weeks post-surgery. Group A: implanted with a DTS. Group B: implanted with a SIS. Group C: not implanted. α-SMA, α-smooth muscle actin; IHC, immunohistochemistry; DTS, decellularized tilapia skin; SIS, small intestinal submucosa.

Discussion

Urethral reconstruction is to restore urethral continuity by harvesting tissue from adjacent or other areas and shaping it for the urethra disorder treatment. It is used to repair or replace urethral strictures or defects caused by congenital defects, trauma, inflammation, or surgery to restore normal urethral structure and function, and therefore improve the patient’s quality of life (24,25). Compared to flaps and skin grafts, buccal mucosa has thicker epithelial cells, and grafting buccal mucosa for urethral reconstruction has been used successfully in clinical practice, showing higher success rates and patient satisfaction during phase I and multi-stage repairs (26-28). However, patients may experience complications such as pain, numbness, and postoperative scar contracture after a portion of the buccal mucosa is removed (29). Based on the available literature, multiple regimens using scaffolds and cells alone or in combination have achieved short-term success in animals or small human trials, but none have matured in the clinic (30-34). The development of urethral replacements with adequate strength, rapid functional recovery, and tissue regeneration rates is a long-term clinical goal (35).

Tilapia skin, as a natural biomaterial, has been gaining attention in recent years in the field of TE. It is rich in collagen and other bioactive components, as well as having excellent biocompatibility, and is therefore considered an ideal natural scaffold material (36,37). In TE, tilapia skin is often used as a three-dimensional scaffold for cell culture to provide space for cell growth and differentiation (18). Tilapia skin has now been successfully used for regeneration and repair of tissues such as skin, cartilage and bone (38). The residual nucleic acid in the tilapia skin is a potential antigen that can cause rejection, and there is still the possibility of sensitization or immune rejection, which poses a potential risk of clinical use and cannot meet the basic requirements for pre-market approval and regulation, and therefore it is necessary to remove it by decellularization technology. SIS has been considered as a promising biological scaffold due to its excellent biocompatibility, low immunogenicity, good mechanical property, and biodegradability. Therefore, SIS was approved by the U.S. Food and Drug Administration for urogenital surgery (39). It was initially reported as an almost-acellular material (40). However, further studies have shown porcine cells inside, which may cause inflammatory response after in vivo implantation which is in line with our results as shown in H&E image (Figure 6A). The DNA content in the DTS used in this experiment was 38.78±5.84 ng/mg which may effectively help avoid immune rejection during urethral reconstruction. We lyophilized and sterilized DTS to prolong the shelf-life and avoid soft tissue infections after implantation without changing the physicochemical properties of DTS as much as possible (18,41). By comparing the physical characterization of DTS with that of SIS, we found that DTS possessed a looser three-dimensional micropore structure (Figure 3) than that of SIS, and this is essential as a scaffold for cell migration and proliferation during the repair process. In addition, it also results in large deformation under stretching and exhibits good ductility (Figure 4). Under electron microscopy, its loose three-dimensional pore structure is a scaffold for the tissue to provide cell proliferation during the repair process. At 4 weeks postoperatively, DTS had fully fused with the penile tissue, while some obvious residues were still visible in the SIS patch. This result is agreed with the dense structure of SIS which may attenuates the water molecules infiltration. Angiogenesis is critical for the survival of implant as de novo blood vessel will provide nutrients and oxygen for cells proliferation and tissue growth. Our results in Figure 6 are in line with previous reports in which they concluded that the neovascularization induced by decellularized tissue is the key factor to promote de novo tissue growth and wound healing as well (21,22).

Nature of collagen renders DTS to inherent the advantages of low cytotoxicity, low immunogenicity, guiding the regeneration of cells, and good compatibility with the human body. After degradation, it can promote the deposition of collagen in the tissues. Collagen, as a major component of the ECM, and collagen deposition can be used to assess the effectiveness of wound healing (42). As shown in Figure 9, the area of collagen fiber in the DTS group were significantly higher than those in the SIS group and the control group. Compared with the sparse tissue structure in the control group, the deposited collagen made the tissue structure of the ventral side of the penis strengthened, which could effectively prevent the occurrence of complications such as urethral fistula. As shown in SEM, DTS has a regular and dense microporous and layered structure, which provides a scaffold for cells to anchor and grow in one way, while ensuring water vapor permeability and water absorption, and providing suitable conditions for postoperative wound wet healing in another way. In addition, while promoting urethral tissue repair, DTS has a thickness of 0.39±0.04 mm, which acts as a physical barrier to the external environment, blocking bacterial invasion and preventing postoperative wound infection (38).

TNF-α is a central regulator of inflammatory cascade and associated with inflammatory mechanisms related to implantation. Implants coated with TNF-α inhibitor could lead to significantly higher vessel densities, suggesting improved biocompatibility of implants (43). Overproduction of TNF-α is thought to affect wound healing through wound strength due to decreased collagen type I and type III expression (44). Collagen obtained by hydrolysis with different proteases usually possesses various biological activities such as antioxidant, bacteriostatic and immunomodulatory. This may be the reason why the degree of inflammatory response during DTS repair was significantly lower than that in the SIS and control groups. It can also be observed that a large number of inflammatory cells infiltration was seen around the postoperative SIS patch in the SIS group, which may be related to the fact that some porcine-derived cells still remained in the SIS thus leading to local inflammatory reactions after implantation in the animal and affecting the effect of urethral tissue repair (39). In addition, smooth muscle stain positive cell proliferation was more pronounced in urethral tissues after the application of DTS compared to SIS. It has been pointed out that collagen also retains, stores and transports growth factors, and plays an important role in cell proliferation and angiogenesis during tissue growth and repair (45-47).

To date, decellularized porcine and bovine skin have been developed into medical devices successfully used in the repair of clinically refractory wounds, and their function in promoting tissue regeneration and wound healing has been clinically proven (48). The use of decellularized matrix materials origin from porcine and bovine may be restricted due to the risk of zoonotic disease transmission such as mad cow disease and foot and mouth disease, coupled with religious factors (49). In the meantime, several studies have shown that marine animal collagen is a promising alternative due to its safety profile (17,41,50,51). Fish collagen has been shown to have several biological benefits, including immunomodulation, anti-inflammatory and antibacterial properties, and promoting wound healing (52-54). Today, decellularized matrix products are relatively expensive, and the skin of tilapia, one of the most extensively farmed fish in the world, is disposed of as a waste resource in fish processing, generating tens of thousands of tons of waste yearly. The structure of fish skin is similar to that of mammalian skin and religious ethical barriers can be effectively circumvented.

Therefore, the DTS matrix is expected to be an alternative for decellularized matrix from terrestrial animals for soft tissue repair. The limitation of this study is that we were not able to test the repaired urethra for erectile function, uroflowmetry, and excretory urethrography, but we did not find that the rabbits had severe urinary difficulties during the experiment. Regarding larger defects, a potential limitation of DTS could be its relatively slower neovascularization rate compared with that of smaller defects. In larger defects, the diffusion distance of nutrients and oxygen from the surrounding tissues is longer than in small defects. Another limitation might be related to the mechanical properties of the DTS. In larger defects, similar to other rigid or elastic biomaterials, DTS may not provide adequate long-term mechanical support. However, we are currently exploring methods such as the crosslinking of collagen fibers in DTS, to modify DTS and improve its mechanical properties for better performance in larger urethral defect repair.

Conclusions

In the present study, we successfully prepared a marine acellular dermal matrix material, DTS, and used it in a rabbit urethral defect model for repair. Compared with the repairing effect of SIS, the tissue collagen and smooth muscle content were significantly increased and the inflammatory response was milder during the urethral defect healing with DTS. As a TE material of marine animal origin with no religious restrictions, DTS is expected to be used in clinical urethral reconstruction due to its effective effect on defect healing and histocompatibility in future.

Acknowledgments

None.

Footnote

Reporting Checklist: The authors have completed the ARRIVE reporting checklist. Available at https://tau.amegroups.com/article/view/10.21037/tau-24-598/rc

Data Sharing Statement: Available at https://tau.amegroups.com/article/view/10.21037/tau-24-598/dss

Peer Review File: Available at https://tau.amegroups.com/article/view/10.21037/tau-24-598/prf

Funding: This work was supported by Key Laboratory of Structural Malformations in Children of Zhejiang Province (No. ZJET2303Z), Wenzhou Institute, University of Chinese Academy of Sciences (No. WIUCASQD2021025), and Zhejiang Provincial Department of Science and Technology (No. 2023C04043).

Conflicts of Interest: All authors have completed the ICMJE uniform disclosure form (available at https://tau.amegroups.com/article/view/10.21037/tau-24-598/coif). The authors have no conflicts of interest to declare.

Ethical Statement: The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. Experiments were performed under a project license (No. wydw-2022-0463) granted by the Experimental Animal Ethics Committee at Wenzhou Medical University, in compliance with the National Institutes of Health’s Guidelines for the care and use of animals.

Open Access Statement: This is an Open Access article distributed in accordance with the Creative Commons Attribution-NonCommercial-NoDerivs 4.0 International License (CC BY-NC-ND 4.0), which permits the non-commercial replication and distribution of the article with the strict proviso that no changes or edits are made and the original work is properly cited (including links to both the formal publication through the relevant DOI and the license). See: https://creativecommons.org/licenses/by-nc-nd/4.0/.

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