A Peritoneum‐Inspired Biomimetic Asymmetric Patch for Functional Repair and Anti‐Adhesion in Abdominal Wall Defects

ABSTRACT

Poor repair outcomes of abdominal wall soft tissue defects often lead to hernia recurrence and tissue adhesion, thereby increasing the medical burden. An ideal repair material should simultaneously fulfill the requirements of anti‐adhesion properties, wet tissue adhesion capability, biodegradability, and the ability to promote functional tissue regeneration. Herein, inspired by the natural abdominal wall architecture, we designed a biomimetic Janus‐structured repair patch, denoted as PS‐PDH. This patch integrates a biodegradable poly(4‐hydroxybutyrate) (P4HB) electrospun fibrous membrane with a glutathione‐responsive degradable poly(sulfobetaine methacrylate) (PSBMA) hydrogel via polydopamine (PDA). It exhibits excellent asymmetric adhesive properties: the fibrous side facilitates cell adhesion and proliferation, while the hydrogel side effectively resists cell and protein adhesion. Furthermore, PS‐PDH possesses robust mechanical properties, enabling it to adapt to dynamic tissue deformation. Notably, the patch effectively scavenges free radicals and intracellular reactive oxygen species (ROS), while promoting macrophage polarization from the pro‐inflammatory M1 to the pro‐regenerative M2 phenotype. In a mouse full‐thickness abdominal wall defect model, PS‐PDH effectively prevented postoperative adhesion and fibrosis, modulated the inflammatory microenvironment, and promoted functional muscle and vascular regeneration. Therefore, this biomimetic Janus‐structured patch offers a reliable and highly clinically translatable strategy for the functional repair of abdominal wall soft tissue defects.

A biomimetic Janus patch (PS‐PDH) for repairing abdominal wall defects is fabricated through in situ polydopamine deposition on a P4HB electrospun fibrous membrane, followed by integration with a PSBMA zwitterionic hydrogel. The resulting Janus patch, which mimics the abdominal wall structure, demonstrates effective anti‐adhesion, mechanical support, and promotion of functional regeneration at abdominal wall defect sites.

1. Introduction

Abdominal wall defects represent a significant global clinical challenge, requiring over 20 million surgical interventions annually [1, 2]. These defects arise from diverse causes, including traffic accidents, tumor resection, and war‐related trauma, posing serious threats to human health and quality of life [3, 4]. Minor abdominal wall injuries can lead to pathological adhesion formation between visceral organs and the abdominal wall, restricting organ mobility and causing long‐term complications such as chronic pain and bowel obstruction. In contrast, full‐thickness abdominal wall defects not only pose adhesion risks but also readily progress to hernias due to insufficient resistance against intra‐abdominal pressure, further precipitating life‐threatening consequences, including intestinal necrosis and enterocutaneous fistulas [5, 6].

Currently, the most widely adopted clinical repair materials are polypropylene (PP) meshes. Despite their high mechanical strength and deformation resistance, direct contact with abdominal organs frequently triggers severe adhesion formation. Moreover, the non‐degradable nature of PP meshes results in chronic foreign body reactions, persistent pain, and infection‐related complications, limiting their utility in full‐thickness abdominal wall reconstruction [7, 8]. Although absorbable meshes such as Dexon and Vicryl can partially alleviate foreign body responses, their design remains predominantly focused on mechanical support and fails to effectively prevent postoperative adhesions [9, 10]. At the site of abdominal wall defects, oxidative stress is significantly elevated, and excessive accumulation of reactive oxygen species (ROS) persistently activates pro‐inflammatory signaling pathways, driving macrophage polarization toward the M1 pro‐inflammatory phenotype and triggering excessive collagen deposition with fibrotic responses. This sustained inflammation‐fibrosis cascade not only exacerbates tissue adhesion formation but also suppresses satellite cell‐mediated myogenic differentiation and angiogenesis, ultimately impeding functional muscle regeneration [11, 12, 13, 14]. Therefore, an ideal abdominal wall repair material should possess not only appropriate mechanical strength, stable anti‐adhesion properties, and biodegradability, but also the capability to scavenge excessive ROS at the injury site, modulate local inflammatory responses, and promote functional muscle regeneration.

From a structural perspective, the native abdominal wall itself represents a highly compartmentalized bifunctional architecture. The deep muscular and fascial layers provide mechanical support, while the overlying peritoneal mesothelial layer secretes mucus to minimize friction and prevent adhesion. Inspired by this natural structure, a growing number of Janus‐structured patches have recently been reported for abdominal wall defect repair. However, critical limitations remain at both the structural and functional levels. At the structural level, composite strategies relying predominantly on physical lamination result in weak interfacial bonding between layers [15, 16, 17], rendering the constructs unable to withstand the long‐term dynamic shear and tensile stresses within the abdominal cavity. Although infiltration of hydrogel precursors into the underlying substrate may improve interfacial integration, it often compromises the morphological integrity of the substrate layer [18, 19]. At the functional level, most Janus patches have focused solely on validating anti‐adhesion performance and promoting collagen deposition. These patches lack active regulation of the injury‐site microenvironment and fail to efficiently facilitate muscle regeneration and neovascularization [20, 21]. Furthermore, many patches still rely on suture fixation, which increases surgical complexity and carries the risk of secondary tissue injury [22, 23]. Therefore, simultaneously achieving robust bonding at heterogeneous interfaces, self‐adhesive fixation of the patch to wet tissue, and synergistic multi‐functional modulation of the injury microenvironment represents the three core challenges that must be addressed to construct high‐performance Janus‐structured repair patches for abdominal wall defects. Polydopamine (PDA), with its abundant catechol and amine functional groups, can form robust adhesion with diverse substrates in complex wet environments [24] and has been widely employed for surface functionalization of biomaterials and development of tissue adhesives [25, 26]. Nevertheless, the strategy of utilizing PDA to hetero‐integrate zwitterionic hydrogels with electrospun membranes for constructing Janus structures remains unexplored.

Therefore, this study presents a strategy utilizing PDA as both an interfacial bridge and tissue adhesive to robustly integrate FDA‐approved biodegradable poly(4‐hydroxybutyrate) (P4HB) electrospun nanofibrous membranes with poly(sulfobetaine methacrylate) (PSBMA) zwitterionic hydrogels to construct a heterogeneous biomimetic abdominal wall repair patch (PS‐PDH). This patch synergistically combines mechanical support and anti‐adhesion functionalities. The heterogeneous interfacial stability, wet tissue adhesion of the fibrous side, and the dual‐sided differential cell and protein adhesion properties were systematically evaluated. The free‐radical scavenging ability of PDA and its role in modulating macrophage polarization were further validated. Finally, the therapeutic efficacy of PS‐PDH, including mechanical reinforcement, immunomodulation, anti‐adhesion performance, and promotion of muscle and vascular regeneration, was comprehensively assessed using a murine full‐thickness abdominal wall defect model (Scheme 1). This work provides a novel material design strategy for achieving functional regeneration of abdominal wall defects.

SCHEME 1.

(A) Schematic diagram of the preparation of PS‐PDH patch. (B) The application of PS‐PDH patch in abdominal wall defect. PS‐PDH exhibits excellent wet tissue adhesion properties, which can effectively prevent adhesion, alleviate inflammation, induce macrophage polarization toward the M2 anti‐inflammatory phenotype, and promote muscle regeneration (differentiation of satellite cells into myoblasts followed by fusion into multinucleated myotubes), as well as angiogenesis at the defect site.

2. Results and Discussion

2.1. Preparation and Characterization of PS‐PDH

Inspired by the structural characteristics of natural abdominal walls, a biomimetic Janus patch was fabricated. Initially, PDA nanoparticles were in situ deposited onto electrospun P4HB fibers (PH), yielding a PDA‐decorated P4HB (PDH) membrane. This membrane was then integrated with an ultrathin PSBMA hydrogel (PS) layer by leveraging the wet adhesion of PDA. As illustrated in Figure 1A, the PH exhibited a randomly oriented morphology, and the attachment of PDA nanoparticles on the fibrous membrane was clearly observable after in situ deposition. Consistent with this observation, Fourier Transform Infrared (FTIR) spectroscopy results (Figure 1B) show the characteristic absorption peaks of PDA, including broad absorption peaks corresponding to the stretching vibrations of hydroxyl (─OH) and amino (─NH₂) groups (∼3400 cm^{− 1}) as well as the skeletal vibrations of benzene rings (1400–1600 cm^{− 1}) [27, 28]. The PDH membrane displayed an average fiber diameter of 160.0 ± 31.3 nm (Figure 1C), closely resembling the architecture of the ECM. The surface of the PS hydrogel featured a porous architecture with an average pore size of 2.12 ± 0.56 µm (Figure 1D), which is significantly smaller than the dimensions required for cellular infiltration. Cross‐sectional characterization of the PS‐PDH patch demonstrated a distinct layered structure, and the thicknesses of the fiber layer and the hydrogel layer were approximately 123 and 33 µm, respectively. Robust interfacial bonding between the two layers was achieved primarily through mechanical interlocking, as the PS hydrogel infiltrated the porous fiber network during fabrication (indicated by blue arrows in Figure 1A). Additionally, non‐covalent interactions between PSBMA and PDA also play an important role. A schematic illustration of the intermolecular hydrogen bonding and cation‐π interactions is presented in Figure 1E. Specifically, hydrogen bonds form between the sulfonate groups of PSBMA and the phenolic hydroxyl groups of PDA, with the lone‐pair electrons on the sulfonate oxygen atoms acting as hydrogen‐bond acceptors for the hydrogen atoms of the phenolic ─OH. The cation‐π interactions originate from the strong electrostatic attraction between the electron‐rich π‐electron cloud of the aromatic rings in PDA and the quaternary ammonium cations on the side chains of PSBMA [29, 30, 31].

FIGURE 1.

Characterization of PS‐PDH patch. (A) Surface SEM images of PH, PDH, PS hydrogel, and the cross‐section of PS‐PDH (red arrows indicate PDA particles deposited on the fibers; blue arrows indicate mechanical interlocks formed by the penetration of PS hydrogel into the fiber pores). (B) ATR‐FTIR spectra of PH and PDH. (C) Fiber diameter distribution of PDH membrane. (D) Pore diameter distribution on the surface of PS hydrogel. (E) Schematic diagram of PDA‐mediated interfacial adhesion mechanism between PS hydrogel and PDH membrane. (F) Tensile strength and elongation at break of PH, PDH, and PS‐PDH (n = 3). (G) Young's modulus of PH, PDH, and PS‐PDH (n = 3). (H) Weight loss of PS hydrogels incubated in GSH solutions of various concentrations (0, 5, 10, 20, and 30 µm) at 37°C (n = 3). Data were represented as mean ± SD.

To simulate the mechanical behavior of the patch under in vivo implantation conditions, the tensile strength and elongation at break of PH, PDH, and PS‐PDH were evaluated in the wet state (Figure 1F and Figure S1). All samples exhibited excellent ductility, ensuring damage resistance during surgical manipulation. Notably, the PS‐PDH Janus patch possessed a Young's modulus of 0.037 ± 0.002 MPa (Figure 1G), which was comparable to that of human peritoneum (0.01–0.1 MPa) [32, 33], indicating favorable biomechanical compatibility. Nevertheless, during clinical translation, further targeted optimization of patch parameters, such as thickness, is required to accommodate different types of abdominal wall hernias and varying defect sizes, so as to meet specific clinical application requirements. To ensure the overall degradability of the material, the PS hydrogel was crosslinked using a disulfide‐containing crosslinker that can be reductively cleaved by glutathione (GSH). Given that GSH concentrations vary significantly across different pathological and physiological conditions, the in vitro degradation behavior of the PS hydrogel was investigated under varying GSH concentrations. As shown in Figure 1H, the PS hydrogel remained stable in pure PBS, with a weight loss of only 4.7% at day 14. As the GSH concentration increased, the degradation rate progressively accelerated. The concentration of GSH in human blood and body fluids is about 20 µm [34]. At GSH concentrations of 20 µm or below, the PS hydrogel maintained a minimal weight loss over 10 days, owing to the overall stability of the crosslinking network. However, after 12 days, the cumulative reductive cleavage of disulfide bonds led to a substantial reduction in crosslinking density, triggering rapid weight loss. This degradation profile allows the PS hydrogel to maintain its structural integrity throughout the high‐risk period for adhesion formation (within 7 days post‐injury), providing effective protection to the wound site. In practical applications, the degradation behavior of the PS hydrogel can be precisely tuned by adjusting the crosslinker content, enabling flexible adaptation to different clinical application scenarios.

2.2. Tissue Adhesion, Swelling Behavior, and Surface Lubricity Evaluation

To visually evaluate the tissue adhesion capability of the fibrous side of the PS‐PDH patch, it was directly attached to the heart, liver, spleen, lung, kidney, and muscle tissues of rats. The PS‐PDH patch demonstrated robust adhesion, successfully lifting all tested tissues, including the heaviest heart weighing approximately 600 mg (Figure 2A). Furthermore, when adhered to porcine skin, the patch maintained robust adhesion without detachment even under stretching, bending, and twisting manipulations (Figure 2B). Quantitative adhesion strength tests on porcine skin confirmed that both the PDH membrane and the fibrous side of the PS‐PDH patch exhibited significantly higher adhesive strength compared to the PH membrane (Figure 2C). This enhanced adhesion is attributed to the electrospun structure's ability to rapidly absorb interfacial moisture through capillary action [35, 36], which facilitates effective contact between the patch's catechol groups, quinone groups, and the functional groups (─NH₂, ─OH, ─SH, ─CONH─, etc.) present in tissue proteins and polysaccharides. Strong bonds are subsequently formed through hydrogen bonding, Michael addition, and Schiff base reactions [37, 38] (Figure S2). Such excellent tissue adhesion prevents displacement and eliminates the need for suturing, thereby shortening operative time and avoiding secondary tissue damage.

FIGURE 2.

Evaluation of tissue adhesion, swelling behavior, and surface lubricity of PS‐PDH patch. (A) Adhesion of the fibrous side of the PS‐PDH patch to major rat organs. (B) Adhesion performance of PDH and PS‐PDH on porcine skin under different mechanical deformations. (C) Adhesion strength of PH, PDH, and PS‐PDH to porcine skin (n = 3). (D) Swelling behavior of the PS hydrogel in deionized water and PBS. (E) Swelling rate of PS hydrogel in deionized water and PBS (n = 3). (F) Directional deformation of PS‐PDH induced by swelling. (G) COF of the different surfaces of PS‐PDH (n = 3). (H) COF‐time curve of different surfaces of PS‐PDH. Data were represented as mean ± SD. ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, ^**** p < 0.0001, ns means no significance.

Subsequently, the swelling behavior of the PS hydrogel was evaluated in different media. As shown in Figure 2D,E, the PS hydrogel exhibited minimal swelling in deionized water, but rapidly swelled to 335% of its original mass within 2 h in PBS, reaching equilibrium at 396% after 6–12 h. This phenomenon is attributed to the strong intra‐ and intermolecular electrostatic interactions of PSBMA [39, 40], leading to a low swelling ratio in aqueous solutions. In PBS, ions such as Na⁺, K⁺, Cl⁻, and PO₄ ²⁻ disrupt these intermolecular electrostatic interactions, leading to rapid swelling of the PS hydrogel (Figure S3). Interestingly, when the PS‐PDH patch was immersed in PBS, the swelling of the PS hydrogel layer caused the patch to deform, bending toward the PDH side (Figure 2F). This deformation can provide additional tension to resist abdominal pressure when the patch is applied to an abdominal wall defect (Figure S4).

Finally, the surface lubricity of both sides of the PS‐PDH patch was characterized by measuring the coefficient of friction (COF). As illustrated in Figure 2G,H, the fibrous surface and hydrogel surface exhibited COF values of 0.223 and 0.084, respectively. Notably, the COF of the PS hydrogel surface was significantly lower than that of the fibrous surface. Furthermore, during a prolonged 1200 s cyclic friction test, the PS hydrogel side maintained an exceptionally smooth and stable friction profile, confirming its durability and high lubricity. This low‐friction characteristic is functionally analogous to the natural serosal surface of the abdominal wall.

2.3. Biocompatibility and Asymmetric Adhesion

The biocompatibility of the PS‐PDH patch was evaluated by incubating it with L929 cells. Cell proliferation and viability were assessed using live/dead staining and the CCK‐8 assay. As shown in Figure 3A, few dead cells (red fluorescence) were observed in both the control and PS‐PDH groups on days 1, 3, and 5, with cell density increasing over time. CCK‐8 assay results (Figure 3B and Figure S5) revealed no significant differences in cell viability or OD values between the two groups on days 1 and 3. Notably, on day 5, the PS‐PDH group exhibited significantly higher cell viability and OD values compared to the control group. These results confirmed the excellent cytocompatibility of PS‐PDH and further suggest that it could actively promote cell survival and proliferation, particularly at higher cell densities. Hemocompatibility, a critical biosafety parameter for implantable materials, was evaluated via a hemolysis assay. As shown in Figure 3C, incubation of PDH, PS hydrogel, or PS‐PDH with mouse red blood cells (RBCs) resulted in supernatants that were nearly colorless and transparent, comparable to the negative control. Quantitative analysis further revealed an extremely low hemolysis rate (0.13%) for PS‐PDH, demonstrating its excellent blood compatibility.

FIGURE 3.

Biocompatibility and asymmetric adhesion properties of the PS‐PDH patch. (A) Live/dead staining of L929 cells co‐cultured with PS‐PDH for 1, 3, and 5 days. (B) Viability of L929 cells co‐cultured with PS‐PDH patch for 1, 3, and 5 days (n = 4). (C) Hemolysis rate of PS‐PDH patch and corresponding digital image (n = 3). (D) Adhesion images of L929 cells (Calcein/PI staining) on the different sides of PS‐PDH patch. (E) Quantification of FITC‐BSA protein adsorption on the different sides of PS‐PDH patch. (n = 3). (F) Fluorescence images of FITC‐BSA adsorption on the different sides of PS‐PDH patch. (G) Schematic illustration of cell adhesion on different surfaces of the PS‐PDH patch. Data were represented as mean ± SD. ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, ^**** p < 0.0001, ns means no significance.

To further elucidate cell‐patch interactions, L929 cells were seeded on both the fibrous and hydrogel sides of the patch, and cell adhesion was observed after 24 h of culture. As shown in Figure 3D, cells adhered well to the fibrous side of PS‐PDH, exhibiting a spread morphology with a typical spindle‐like or polygonal shape. In contrast, only a small number of cells were observed on the hydrogel side, and they remained round, which indicates that cells find it difficult to adhere to the surface of the PS hydrogel. Additionally, the protein adsorption behavior was evaluated using BSA‐FITC as a model protein. As shown in Figure 3E,F, the electrospun nanofiber layer adsorbed a large amount of BSA, while almost no adsorption was detected on the PS hydrogel layer, indicating that the latter can effectively resist protein adsorption. The above phenomena can be attributed to the heterogeneous surface design of the material: the electrospun fiber structure mimics the ECM, and the surface‐deposited PDA further promotes cell recognition, adhesion, and proliferation. Conversely, the PS hydrogel forms a stable hydration layer, which effectively prevents the adsorption of biomolecules such as fibronectin and vitronectin [41], thereby blocking integrin‐mediated cellular recognition and adhesion at the interface (Figure 3G). In summary, the PS‐PDH patch exhibits excellent biocompatibility and distinct Janus characteristics, enabling simultaneous fulfillment of the dual requirements for promoting tissue regeneration and preventing postoperative adhesion in abdominal wall repair.

The L929 fibroblast cell line was specifically chosen for biocompatibility evaluation because it represents a well‐established in vitro cytotoxicity model with well‐characterized and sensitive responses to potentially toxic substances [42]. Moreover, it is the recommended cell line for cytotoxicity tests in the biological evaluation of medical devices as specified by ISO 10993–5:2009. It was chosen for the cell adhesion assay since fibroblasts serve as the key effector cells in pathological adhesion formation [43]. However, the L929 cell line used in this study is an immortalized murine fibroblast line, and its phenotypic characteristics and responses to biomaterials may not fully recapitulate those of primary human fibroblasts in vivo. Furthermore, in vitro experiments are inherently limited in mimicking the complex in vivo microenvironment. Therefore, future studies using primary human cells will be necessary to further validate these findings.

2.4. Antioxidant and In Vitro Immunomodulatory Properties

Oxidative stress is intimately linked to macrophage phenotype regulation and tissue regeneration outcomes. The free radical scavenging efficiency of PS‐PDH was evaluated using two widely adopted models (DPPH• and ABTS•⁺). As shown in Figure 4A,B, both PDH and PS‐PDH exhibited time‐dependent free radical scavenging capabilities. Specifically, more than 90% of DPPH• radicals were eliminated within 6 h, while nearly complete scavenging of ABTS•⁺ radicals was achieved within 12 h. In contrast, the PH membrane and PS hydrogel showed negligible scavenging effects on both radicals.

FIGURE 4.

Antioxidant and in vitro immunomodulatory properties of PS‐PDH. (A) DPPH• and (B) ABTS•⁺ radical scavenging kinetics of PH, PDH, PS, and PS‐PDH (n = 3). (C) Schematic illustration of the dopamine‐mediated radical scavenging mechanism and intracellular ROS elimination. (D) Intracellular ROS scavenging by PS‐PDH in LPS‐stimulated RAW264.7 macrophages. (E) Representative flow cytometry analysis of CD80 and CD206 expression in RAW264.7 cells under different treatments. (F) Quantitative analysis of intracellular ROS scavenging efficiency (n = 3). (G) Percentage of CD80⁺ (M1‐like) macrophages across treatment groups (n = 3). (H) Percentage of CD206⁺ (M2‐like) macrophages across treatment groups (n = 3). Data were represented as mean ± SD. ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, ^**** p < 0.0001, ns means no significance.

To further assess intracellular antioxidant performance, the ROS‐scavenging ability of PS‐PDH was examined in RAW264.7 macrophages. As illustrated in Figure 4D,F, compared with the positive control group, the PDH and PS‐PDH groups scavenged 99.7% and 99.5% of intracellular ROS, respectively, demonstrating high scavenging efficiency. The PS hydrogel alone exhibited a modest ROS scavenging effect (≈19%), which may be attributed to the dynamic disulfide bonds (─S─S─) within its crosslinked network that can participate in redox reactions with H₂O₂. The superior antioxidant activity of PDH and PS‐PDH is primarily associated with the catechol/quinone redox couples and abundant phenolic hydroxyl groups within the polydopamine (PDA) coating, which facilitate continuous electron transfer reactions with free radicals [44] (Figure 4C).

Given the intimate link between oxidative stress and macrophage phenotypic regulation, the effect of PS‐PDH on macrophage polarization was subsequently investigated. LPS‐stimulated RAW264.7 macrophages were co‐cultured with PDH, PS hydrogel, or PS‐PDH, and their phenotypes were analyzed by flow cytometry. As shown in Figure 4E,G,H, and LPS stimulation induced a pronounced M1 polarization, with M1 macrophages accounting for 72.1% of the population. In contrast, co‐culture with PDH, PS hydrogel, and PS‐PDH significantly reduced the M1 proportion to 40.9%, 63.0%, and 36.2%, respectively. Notably, PS‐PDH treatment markedly promoted macrophage polarization toward the M2 phenotype, increasing its proportion from 5.9% in the LPS group to 20.7%. These results demonstrate that PS‐PDH effectively suppresses LPS‐induced M1 polarization while concurrently promoting M2 polarization. This immunomodulatory effect is likely associated with the material's efficient ROS scavenging capacity, which suppresses the sustained oxidative stress‐mediated activation of pro‐inflammatory pathways such as NF‐κB, thereby disrupting the self‐amplifying feedback loop that sustains M1 polarization and favoring a shift toward an anti‐inflammatory, pro‐regenerative macrophage phenotype [45, 46]. Collectively, these findings confirm the successful integration of PDA's intrinsic antioxidant properties into the P4HB electrospun membrane and highlight its potential to modulate the inflammatory microenvironment through redox‐mediated immune regulation.

RAW264.7 cells were selected for ROS scavenging and macrophage polarization experiments, as this classical murine macrophage cell line retains the core biological functions of primary macrophages. It can produce ROS under inflammatory stimulation and possesses the capacity for phenotypic polarization [47], which well simulates the dynamic changes of macrophages in the in vivo inflammatory microenvironment. However, RAW264.7 exhibits inherent limitations similar to those of L929 cells. Future studies are required to better recapitulate the complex in vivo microenvironment and to adopt human‐derived cell models in subsequent translational research.

2.5. In Vivo Evaluation of Abdominal Wall Defect Repair and Anti‐Adhesion

A full‐thickness abdominal wall defect model in mice was established to evaluate the anti‐adhesion efficacy and tissue repair performance of PS‐PDH. The abdominal cavities were reopened to observe the adhesion conditions at 14 and 28 days postoperatively (Figure 5A). At day 14, all four mice in both the Control group and the PP group developed severe adhesions, with adhesions in the PP group extending to the liver and requiring sharp dissection for separation. The average adhesion scores were 8.75 for the Control group and 9.5 for the PP group. In contrast, the PDH group showed significantly reduced adhesion formation, with an average adhesion score of 3.25. Notably, no adhesions were observed in any of the four mice treated with PS‐PDH, yielding an average adhesion score of 0, which was significantly lower than that of the Control, PP and PDH groups (Figure 5B). At day 28, the control group and PP group still exhibited severe adhesions, with average scores of 7.5 and 9.75, respectively. The PDH group developed localized adhesions between the membrane and the cecum, with the average score decreasing to 2.5, whereas the PS‐PDH group remained completely free of adhesions (Figure 5C). The mild and localized adhesions in the PDH group may be partially attributed to the intrinsic adhesive properties of PDA. Nevertheless, PDA modulated the wound microenvironment and synergized with the extracellular matrix‐mimetic structure of the electrospun membrane, thereby alleviating adhesion severity. The superior anti‐adhesion performance of the PS‐PDH is attributed to the PS hydrogel layer, which mimics the natural peritoneal serosa. Its strong hydration capacity prevents fibrin adsorption as well as cell recognition and adhesion, while its aqueous lubricity reduces friction between injured tissues.

FIGURE 5.

In vivo evaluation of PS‐PDH for abdominal wall defect repair and anti‐adhesion efficiency in a mouse model. (A) Gross observations of intra‐abdominal adhesion in Control, PP, PDH, and PS‐PDH groups at 14 and 28 days post‐surgery. (B,C) Adhesion severity scores based on macroscopic evaluation at (B) 14 days and (C) 28 days post‐surgery (n = 4). (D) Representative H&E and Masson's trichrome staining of different groups at 14 days post‐surgery (red triangles: PDH membrane; red pentagrams: PS‐PDH patch; red arrows: collagen deposition; AW: Abdominal wall; CE: cecum). (E) Quantification of collagen deposition based on Masson's trichrome staining (n = 4). (F) Monitoring of postoperative body weight changes of mice (n = 4). Data were represented as mean ± SD. ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, ^**** p < 0.0001, ns means no significance.

To further evaluate the anti‐adhesion efficacy and abdominal wall repair performance of each group, histological analyses of the injured tissues were conducted using H&E and Masson's trichrome staining. As shown in Figure 5D, in the Control and PP groups, the injured cecum and the defective abdominal wall were connected by a thick adhesive layer, rendering the two tissues difficult to distinguish. In the PP group, obvious staining voids corresponding to polypropylene fibers were observed. Given the nondegradable nature of PP, its long‐term retention at the injury site may contribute to chronic inflammation and persistent postoperative pain. Although local adhesion was observed in the PDH group, the connection between the membrane and the cecum was loose, and certain regeneration of the cecal serosa could be seen. In contrast, no adhesions were detected between the abdominal wall and the cecum in the PS‐PDH group; the cecal serosa was completely repaired, and the abdominal wall defect area was significantly reduced. H&E staining results at 28 days postoperatively (FigureS6) showed that the size of the abdominal wall muscle defect in the PS‐PDH group was further diminished, indicating its potential to promote functional abdominal wall repair. In addition, histological evaluation of the local inflammatory response at both 14 and 28 days post‐surgery revealed massive infiltration of neutrophils and macrophages around the polypropylene fibers in the PP group. In contrast, no excessive accumulation of inflammatory cells was observed at the interface between the PS‐PDH patch and the host tissue, indicating favorable tissue integration and excellent biocompatibility. As illustrated in Figure 5D,E, Masson staining and quantitative analysis of collagen deposition revealed extensive collagen fiber accumulation within the adhesive regions of the Control and PP groups, indicative of pathological fibrosis. The PDH group showed reduced collagen deposition, whereas the PS‐PDH group exhibited the lowest collagen content among all groups. These results suggest that the PS‐PDH patch preferentially directs tissue regeneration toward functional repair rather than pathological scar formation.

Body weight changes were monitored over 14 days postoperatively (Figure 5F). Mice in the PDH and PS‐PDH groups exhibited the most rapid recovery of body weight, with increases exceeding 10% of initial weights. In contrast, weight recovery was slower in the Control and PP groups, likely due to pain and impaired intestinal motility caused by severe adhesions, which subsequently affected feeding and digestion. These data visually reflect the favorable therapeutic effect of PS‐PDH in abdominal wall defect repair. To evaluate the in vivo biocompatibility of PS‐PDH, major organs (heart, liver, spleen, lung, and kidney) were harvested at 28 days post‐implantation for histological analysis. H&E staining revealed no apparent pathological abnormalities or inflammatory lesions in any of the examined organs across all groups, comparable to those observed in the Control group (Figure S7). This favorable systemic biosafety can be largely attributed to the safe degradation profiles of the patch components. The degradation product of P4HB is 4‐hydroxybutyrate monomer, a natural metabolite in the human body with low acidity and rapid metabolism [48]. The crosslinked PSBMA hydrogel degrades into linear fragments. These fragments possess excellent biocompatibility and can be eliminated from the body through renal pathways [49, 50, 51]. The degradation products of PDA are similar to those of natural melanin and can also be excreted via urine [52, 53, 54]. However, systematic monitoring of the complete in vivo degradation cycle is still required in future studies.

2.6. Effects of PS‐PDH on the Inflammatory Microenvironment and Macrophage Polarization

The level of inflammation and the polarization of macrophage phenotypes (M1/M2) play crucial roles in tissue adhesion and defect repair. To systematically evaluate the in vivo immunomodulatory effects of PS‐PDH, the immune microenvironment at the defect site was analyzed 14 days postoperatively. As shown in Figure 6A, the expression levels of the pro‐inflammatory cytokines TNF‐α and IL‐1β were markedly reduced in the PDH and PS‐PDH groups compared to the Control and PP groups, indicating effective suppression of the inflammatory response. Further immunofluorescence analysis of macrophage phenotypes revealed pronounced infiltration of M1 pro‐inflammatory macrophages in the injured regions of the Control and PP groups, which was consistent with their elevated levels of pro‐inflammatory cytokines. In contrast, the PDH and PS‐PDH groups exhibited a substantial decrease in M1 macrophage infiltration, accompanied by a significant increase in M2 macrophages (Figure 6B). Quantitative analyses of inflammatory cytokines and macrophage polarization further demonstrated that the PDH and PS‐PDH groups exhibited the lowest expression of TNF‐α and IL‐1β (Figure 6C,D), with no significant difference between the two groups (p > 0.05). Notably, the PS‐PDH group showed the fewest M1 macrophages and the highest number of M2 macrophages (Figure 6E,F). These findings indicate that PS‐PDH can effectively modulate the local inflammatory response and promote a pro‐regenerative immune microenvironment favorable for tissue repair and regeneration.

FIGURE 6.

Effects of PS‐PDH on the inflammatory microenvironment and macrophage polarization at the injured site. (A) Representative immunofluorescence staining of TNF‐α (red) and IL‐1β (red) at injured site on day 14. (B) Representative immunofluorescence staining of CD86 (red) and CD206 (green) at injured site on day 14. (C,D) Quantitative analysis of the positive area coverage of TNF‐α (C) and IL‐1β (D) (n = 4). (E,F) Quantification of the positive area of CD86⁺ (E) and CD206⁺ (F) macrophages (n = 4). Data were represented as mean ± SD. ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, ^**** p < 0.0001, ns means no significance.

2.7. Effects of PS‐PDH on Muscle Healing and Angiogenesis

The abdominal wall muscle belongs to skeletal muscle tissue. Although it possesses an intrinsic regenerative capacity, effective regeneration is frequently compromised in complex abdominal wall defects. This limitation stems from the lack of appropriate topographical cues within the defect region, as well as the intense inflammatory microenvironment and excessive collagen deposition following injury, both of which can inhibit satellite cell‐mediated myogenic differentiation [55]. Satellite cells are primary initiators of skeletal muscle regeneration, with PAX7 serving as their specific transcriptional marker. Activated satellite cells begin to express MYOD1, and the resulting differentiated myoblasts subsequently fuse to form multinucleated myotubes expressing MYH3 [56, 57]. As shown in Figure 7A,B, only limited PAX7⁺ satellite cells were observed in the Control and PP groups at 14 days postoperatively. In contrast, satellite cell migration into the defect was markedly increased in the PDH group, and the PS‐PDH group exhibited the highest abundance of PAX7⁺ cells. Furthermore, the PS‐PDH group also displayed a significantly greater number of MYOD1⁺ activated satellite cells than the other groups (Figure 7A,C), indicating a more active myogenic differentiation process. Concurrently, MYH3 expression was most pronounced in the PS‐PDH group, confirming enhanced formation of nascent myotubes (Figure 7A,D). Given the high metabolic demand of skeletal muscle, a rich capillary network is essential. CD31 immunofluorescence staining (Figure 7A,E) revealed that the Control and PP groups had very few newly formed vessels, whereas the PS‐PDH group possessed the highest density of neovascularization. Collectively, these results demonstrate that PS‐PDH synergistically promotes satellite cell migration and activation, enhances myogenic differentiation, accelerates myotube formation, and effectively induces angiogenesis, thereby facilitating functional muscle regeneration at the defect site.

FIGURE 7.

Effects of PS‐PDH on muscle healing and angiogenesis at injured site. (A) Immunofluorescence staining of PAX7 (green), MYOD1 (red), MYH3 (red), and CD31 (green) at injured site on day 14. (B–D) Quantitative analysis of the positive area coverage of myogenic markers PAX7 (B), MYOD1 (C), and MYH3 (D) (n = 4). (E) Quantification of the CD31‐positive area coverage (n = 4). Data were represented as mean ± SD. ^* p < 0.05, ^** p < 0.01, ^*** p < 0.001, ^**** p < 0.0001, ns means no significance.

Importantly, this regenerative outcome fundamentally differs from that achieved by conventional mesh materials, which primarily depend on collagen deposition to provide mechanical reinforcement. The superior regenerative performance of PS‐PDH could be attributed to its bilayer synergistic design. Specifically, the electrospun fibrous layer offers a biomimetic microarchitecture, scavenges ROS, and modulates the immune microenvironment, while the hydrogel layer effectively prevents excessive fibroblast activation and inflammation amplification associated with tissue adhesions, both of which are critical factors that hinder myogenic regeneration. Consequently, effective anti‐adhesion is an indispensable prerequisite for achieving high‐quality muscle regeneration in abdominal wall repair.

3. Conclusion

In summary, a biomimetic Janus‐structured PS‐PDH patch for abdominal wall defect repair was successfully developed. Through in situ deposition of PDA on P4HB fibers, robust interfacial integration between the electrospun membrane and the PS hydrogel was achieved. The nanofibrous layer, with a porous structure and PDA nanoparticles, effectively dissipated interfacial water, enabling strong adhesion to diverse tissue surfaces. Meanwhile, the PS hydrogel layer exhibited a persistent and ultra‐low coefficient of friction, providing sustained lubrication between the patch and the abdominal organs. In vitro studies demonstrated that the nanofibrous side promoted cell adhesion and proliferation, whereas the PS hydrogel side effectively resisted cell and protein adhesion. Furthermore, PS‐PDH exhibited efficient free‐radical scavenging capability and could polarize macrophages toward the M2 phenotype. In a mouse full‐thickness abdominal wall defect model, the material exhibited excellent anti‐adhesion performance, immunomodulatory function, and the ability to promote muscle regeneration. Additionally, the PS‐PDH patch is fully biodegradable, easy for large‐scale production, and possesses excellent biocompatibility. These collective findings demonstrate that the PS‐PDH patch addresses key limitations of existing repair materials and holds great potential for clinical translation.

4. Experimental Section

4.1. Materials

P4HB was synthesized according to the previously reported method [58]. The number‐average molecular weight (M _n) and polydispersity index (M _w/M _n) of P4HB are 33200 g/mol and 2.42, respectively. Methacryloyloxyethyl sulfobetaine (SBMA) was purchased from Shanghai Yuanye Bio‐Technology Co., Ltd. Hexafluoroisopropanol (HFIP), (N, N'‐(dithiodi‐2,1‐ethanediyl)bis(acrylamide)) (BACA), ammonium persulfate (APS), and (N, N, N', N'‐tetramethylethylenediamine) (TEMED) were purchased from Shanghai Aladdin Biochemical Technology Co., Ltd. Dopamine hydrochloride was purchased from Shanghai Macklin Biochemical Technology Co., Ltd. The L929 mouse fibroblasts (RRID: CVCL_0462; catalog number: CL0339) and the RAW264.7 mouse macrophages (RRID: CVCL_0493; catalog number: CL0266) were obtained from Hunan Fenghui Biotechnology Co., Ltd. Dulbecco's modified Eagle's medium (DMEM), fetal bovine serum (FBS), phosphate buffered saline (PBS, pH 7.4), penicillin‐streptomycin and trypsin were purchased from Wisent Biotechnology (Nanjing) Co., Ltd. Cell Counting Kit‐8 (CCK‐8) and Calcein/PI Cell Viability/Cytotoxicity Assay Kit were purchased from Beyotime. PP mesh was purchased from Pride Medical Instruments Technology Jiangsu Co., Ltd.

4.2. Electrospinning

P4HB was dissolved in HFIP at a concentration of 10% (w/v) and loaded into a syringe equipped with a 21‐gauge needle. Electrospinning was performed using a flat collector under the following conditions: applied voltage, 15 kV; solution flow rate, 0.9 mL/h; tip‐to‐collector distance, 15 cm; ambient temperature, 25°C. After collection, the samples were placed in a vacuum oven at 25°C for 3 days to remove residual solvent. The prepared P4HB membranes were designated as PH.

4.3. Preparation of PSBMA Hydrogel

BACA (5 mg) was dissolved in deionized water (2.5 mL), followed by the addition of SBMA (1 g) and APS (10 mg). After complete dissolution, TEMED (5 µL) was added to initiate polymerization. The mixture was drop‐cast into a punched polyimide film mold, covered with a glass slide, and polymerized at 37°C overnight. The resulting hydrogel was then immersed in deionized water, with the water changed every 12 h for 3 cycles to remove unreacted monomers.

4.4. Preparation of PS‐PDH

A Tris‐HCl buffer (pH 8.5) containing dopamine hydrochloride at a concentration of 2 mg/mL was prepared. The PH membrane was immersed in the aforementioned buffer solution and incubated at 37°C for 24 h. Subsequently, the membrane was thoroughly rinsed with deionized water to remove unbound PDA particles. The prepared PDA‐deposited P4HB membranes were designated as PDH. Thereafter, the PDH was placed onto the surface of the pre‐prepared PS hydrogel and fixed with glass slides for 12 h. The final composite patch was designated as PS‐PDH.

4.5. Characterization

The surface morphologies of the samples were observed using a JSM‐7500 scanning electron microscope (SEM, JEOL, Tokyo, Japan). Prior to imaging, the hydrated samples were freeze‐dried, and all specimens were sputter‐coated with a thin layer of gold via plasma deposition.

Mechanical properties were tested using a CMT‐4204 electric universal testing machine (Shenzhen, China). The membranes were cut into 20 × 5 mm long strips with a thickness of 150–220 µm. The tensile test was conducted at a speed of 10 mm/min under a controlled temperature of 25°C, with three parallel specimens prepared for each group.

Fourier transform infrared (FT‐IR) spectra of the samples were recorded using a PerkinElmer Spectrum 100 instrument (Waltham, USA) equipped with an attenuated total reflectance (ATR) accessory. The measurements were performed over the wavenumber range of 4000–400 cm⁻¹ at a resolution of 4 cm⁻¹ with 32 cumulative scans.

4.6. Degradation of PS Hydrogel

The degradation test of the PS hydrogel was conducted in PBS solutions containing glutathione (GSH) at concentrations of 30, 20, 10, 5, and 0 µm, at 37°C under constant shaking (100 rpm). At predetermined time points of 1, 2, 4, 6, 8, 10, 12, and 14 days, the hydrogel samples were washed, lyophilized, and weighed. The weight loss of the samples was calculated using the following formula: Weight loss (%)  =  [(W0–Wt)/W0]  ×  100%, where W₀ represents the initial lyophilized dry mass of the hydrogel, and W_t represents the lyophilized dry mass of the hydrogel collected at each time point.

4.7. Adhesiveness Tests

The adhesive strength of the samples to tissue was evaluated using an HP‐20 pull‐and‐push dynamometer (Yueqing, China). Specimens of PH, PDH, and PS‐PDH (each with the fibrous side as the adhesive surface) were cut into 10 mm × 10 mm squares and firmly attached to fresh porcine skin. A vertical tensile test was then performed at a speed of 2 mm/min until complete separation occurred between the sample and the skin. The maximum force recorded during detachment was normalized to the initial contact area to calculate the adhesive strength. Triplicate specimens were tested for each group, and the results were averaged.

4.8. Swelling Assay

The swelling behavior of the PS hydrogel was evaluated by recording its initial weight before immersion in PBS or deionized water. At predetermined time points (10 min, 30 min, 1 h, 2 h, 3 h, 6 h, 12 h, 24 h, and 48 h), the hydrogel was removed from the solution, blotted dry with filter paper, and immediately weighed. The swelling ratio was calculated according to the following formula: Swelling ratio (%)  =  [(Wt ‐ W0)/W0]  ×  100%, where W₀ represents the initial weight of the hydrogel, and W_t represents the weight at each time point. All tests were conducted at 37°C, and triplicate specimens were used for each group.

4.9. Tribological Tests

The tribological test was conducted in rotational mode using a tribometer (UMT‐2, Bruker Nano Inc., Germany). The samples served as the lower specimen and were securely affixed to glass slides. A SiN ball with a diameter of 4 mm was used as the upper counterpart, which slid against the patch surface under the following conditions: rotational speed of 50 mm/min, normal load of 0.5 N, rotation radius of 3 mm, and deionized water as the lubricating medium. All tests were conducted at room temperature (25°C).

4.10. Biocompatibility Assay

In vitro cytocompatibility assay: The CCK‐8 assay and live/dead cell staining kit assay were used to evaluate cytocompatibility. L929 cells were cultured in DMEM supplemented with 10% FBS and 1% penicillin‐streptomycin, under a humidified atmosphere containing 5% CO₂ at 37°C. Cell viability was assessed using the CCK‐8 assay. Cells were seeded in a 96‐well plate at a density of 5 × 10³ cells per well (suspended in 200 µL of culture medium). After 12 h of incubation to allow cell attachment, sterilized samples (4 mm‐diameter circular discs) were co‐incubated with the cells. At 1, 3, and 5 days of co‐incubation, the samples and original medium were removed, and 150 µL of CCK‐8 working solution was added to each well, and the plate was incubated at 37°C for 1 h in the dark. Subsequently, 100 µL of the solution from each well was transferred to a new blank 96‐well plate, and the optical density (OD) was measured at a wavelength of 450 nm using a Multiskan MK3 microplate reader (Thermo Fisher Scientific, Waltham, USA). Cell viability was calculated by the following formula: Cell viability  =  (ODs/ODc)  ×  100%, where OD_s represents the absorbance intensity of the sample group, and OD_c represents the absorbance intensity of the cells incubated with culture medium only. For live/dead staining, the cells were seeded in 35 mm‐diameter confocal dishes at a density of 5 × 10⁴ cells per dish (suspended in 1 mL of culture medium). After 12 h of incubation to allow cell attachment, sterilized samples (10 mm‐diameter circular discs) were co‐incubated with the cells. At 1, 3, and 5 days of co‐incubation, the samples and original medium were removed, and the cells were rinsed with PBS. Subsequently, 1 mL of working concentration staining solution containing calcein acetoxymethyl ester (calcein‐AM) and propidium iodide (PI) was added to each dish, followed by incubation at 37°C for 30 min in the dark. Finally, the cells were observed and imaged using a confocal laser scanning microscope (CLSM, Leica TCS SP8, Germany).

Hemolysis assessment: Mouse whole blood was collected into EDTA‐coated tubes and centrifuged at 1500 rpm (208 × g) for 15 min. The collected red blood cells (RBCs) were washed three times with saline and diluted to a 20% (v/v) suspension. For the experimental groups, 60 µL of the RBC suspension was mixed with 240 µL of saline, and the mixture was then incubated with sterilized samples (PDH, PS hydrogel, or PS‐PDH) at 37°C for 4 h. The negative control consisted of 60 µL of RBC suspension mixed with 240 µL of saline in the absence of any material (0% hemolysis), while the positive control consisted of 60 µL of RBC suspension mixed with 240 µL of deionized water (100% hemolysis). After incubation, the samples were centrifuged at 3000 rpm (835 × g) for 5 min, images were captured, and 100 µL of the supernatant was transferred to a 96‐well plate and measured at a wavelength of 540 nm. The hemolysis rate was calculated using the following formula: Hemolysis rate (%)  =  (ODS ‐ ODN)/(ODP  −  ODN)  ×  100%, where OD_S, OD_N, and OD_P were the absorbance of the experimental group, negative control group, and positive control group, respectively.

4.11. Cell and Protein Adhesion Assays

Cell adhesion assay: To evaluate cell adhesion on both surfaces of PS‐PDH patches, the patches were placed in glass‐bottom confocal dishes with either the nanofibrous surface or the hydrogel surface facing upward. L929 cells were suspended in culture medium at a density of 1 × 10⁵ cells/mL, and 1 mL of the cell suspension was seeded onto the designated surface. After 24 h of incubation, the medium was removed, and the surfaces were gently rinsed three times with PBS. Subsequently, the cells were stained following the aforementioned live/dead assay protocol. The patches were then transferred to glass slides, and cell adhesion on the patch surfaces was observed and imaged using CLSM.

Protein adhesion assay: PS‐PDH patches (10 mm × 10 mm) were immersed in 50 µg/mL fluorescein isothiocyanate‐labeled bovine serum albumin (FITC‐BSA) solution (dissolved in PBS, pH 7.4) and incubated at 37°C for 2 h. After incubation, the patches were rinsed three times with PBS. Protein adhesion on both surfaces of the PS‐PDH patches was visualized by CLSM.

4.12. Antioxidant Activity Tests

The antioxidant activities of materials were evaluated using 1,1‐diphenyl‐2‐picrylhydrazyl (DPPH•) and 2,2′‐azino‐bis(3‐ethylbenzothiazoline‐6‐sulfonic acid) radical (ABTS•⁺) scavenging assays. For DPPH• scavenging assay, DPPH was dissolved in an ethanol/water mixture (1:1, v/v) to prepare a 0.1 mm solution. The samples (10 mm × 10 mm) were immersed in the solution and incubated at 37°C under constant shaking (100 rpm). At predetermined time intervals, 80 uL of the solution was taken out, and the absorbance at 517 nm was measured. For ABTS•⁺ scavenging assay, stock solutions of 0.1 mm ABTS and 0.03 mm potassium persulfate (KPS, K₂S₂O₈) were prepared in deionized water, mixed at a 1:1 (v/v) ratio, and incubated in the dark at room temperature for 12 h to generate ABTS•⁺. Prior to use, the resulting ABTS•⁺ solution was diluted with deionized water to obtain the working solution. The same co‐incubation procedure as described for the DPPH• assay was applied. The DPPH• and ABTS•⁺ scavenging rates were calculated using the following equation: Scavenging rate  =  (ODc ‐ ODs)/ODc  ×  100%, where OD_c represents the absorbance of the control group, and OD_s represents the absorbance of the sample group. Each assay was performed in triplicate.

4.13. Intracellular ROS Scavenging Assay

RAW264.7 macrophages were cultured in DMEM supplemented with 10% FBS and 1% penicillin‐streptomycin, and maintained at 37°C in a humidified atmosphere containing 5% CO₂. The cells were seeded into 35 mm‐diameter confocal dishes at a density of 2.5 × 10⁵ cells/dish (1 mL cell suspension per dish) and cultured for 12 h to allow adherence. Subsequently, the control group was added with serum‐free medium, while the remaining groups were added with serum‐free medium containing lipopolysaccharide (LPS, 100 ng/mL) and cultured for another 24 h. Then, the PDH, PS hydrogel, and PS‐PDH were added to each group separately, and the cells were further cultured for 24 h. Thereafter, the cells were stained using a DCFH‐DA Reactive Oxygen Species Detection Kit, incubated at 37°C in the dark for 20 min. After being washed three times with PBS, the cells were observed and photographed under a CLSM with an excitation wavelength (λ_ex) of 488 nm and an emission wavelength (λ_em) of 525 nm. The fluorescence intensity was quantitatively analyzed using ImageJ software, and each group was tested in triplicate.

4.14. Macrophage Polarization

RAW264.7 cells were seeded into 6‐well plates at a density of 4 × 10⁵ cells/well (2 mL cell suspension per well) and cultured for 12 h. Subsequently, the control group was cultured in serum‐free DMEM. The remaining groups were cultured in serum‐free DMEM containing LPS (100 ng/mL) and further cultured for 24 h. Subsequently, PDH, PS hydrogel, and PS‐PDH patches (10 mm × 10 mm) were separately added to each group. After 24 h of incubation, the materials were removed. Subsequently, the cells were stained respectively with FITC‐CD80 antibody (1 µg/test, Cat. No. 65076, Proteintech) and PE‐CD206 antibody (0.5 µg/test, Cat. No. 98031, Proteintech) at 4°C for 30 min. After staining, the cells were washed twice with cold PBS and immediately subjected to flow cytometry analysis (CytoFLEX, Beckman Coulter, USA). The percentages of CD80⁺ (M1) and CD206⁺ (M2) macrophages were quantified using FlowJo software.

4.15. Repair of Abdominal Wall Defects

All animal experiments were performed according to a protocol approved by the Institutional Animal Care and Use Committee at the China‐Japan Friendship Hospital (No. zryhyy21‐22‐08‐09). BALB/c mice (female, 6 weeks old) were randomly divided into four groups, including Control, PP (a commercialized mesh), PDH, and PS‐PDH. Sample size was first estimated by a priori power analysis using G*Power 3.1, with the abdominal adhesion score defined as the primary evaluation endpoint (effect size f = 0.7, α = 0.05, power = 0.8), yielding a theoretical sample size of n = 7 per group. However, in accordance with the 3R principle and considering the strong anti‐adhesion performance observed in pilot experiments, the final sample size was set to n = 4 per group. Post hoc sensitivity analysis demonstrated that the final sample size achieved adequate statistical power (> 0.80), thereby validating the reliability of the experimental design.

Following abdominal hair removal, mice were anesthetized with isoflurane in an induction chamber. Under aseptic conditions, a midline laparotomy was performed to expose the abdominal cavity. A full‐thickness abdominal wall defect (5 mm diameter) was created on the right lateral wall using a biopsy punch, excising the peritoneum, muscle, and fascia while preserving the overlying skin. The cecum was abraded with sterile surgical gauze until a petechial hemorrhage formed on its surface. In the treatment groups, PP, PDH, or PS‐PDH patches (with the nanofibrous surface oriented toward the abdominal wall defect) were positioned to fully cover the defect, while no implant was placed in the Control group. Finally, the midline incision was closed with 4‐0 absorbable sutures.

4.16. Morphological Observation and Histological Evaluation

Mice were euthanized at 14 and 28 days post‐surgery, and the severity of adhesions was observed and scored. Adhesion scoring was performed according to the criteria shown in Table S1. The abdominal wall and cecal tissues at the modeling sites of mice in each group were dissected and immediately fixed in 4% paraformaldehyde fixative for 24 h. After fixation, the tissues were dehydrated through a graded ethanol series, embedded in paraffin, and then sectioned. Hematoxylin and eosin (H&E) staining and Masson's trichrome staining were performed to evaluate tissue morphological changes and collagen fiber deposition, respectively.

Before immunofluorescence staining, the sections were subjected to heat‐mediated antigen retrieval in sodium citrate buffer (pH 6.0), and non‐specific binding sites were blocked with 5% goat serum. The sections were incubated overnight at 4°C with the following primary antibodies: TNF‐α, IL‐1β, CD86, CD206, and CD31 (Abcam; diluted 1:200), as well as PAX7, MYH3, and MYOD1 (Proteintech; diluted 1:200). Then, fluorescently labeled secondary antibodies (Jackson, diluted 1:400) were added and incubated for 1 h in the dark. The cell nuclei were counterstained with DAPI. Images were acquired using a digital slide scanner, and the fluorescence intensity was quantitatively analyzed using ImageJ software.

4.17. Statistical Analysis

All quantitative data were expressed as the mean ± standard deviation (SD). Student's t‐test or one‐way analysis of variance (ANOVA) was used to analyze the statistical differences. Statistical significance was set at ^* p < 0.05, and high statistical significance was set at ^** p < 0.01, ^*** p < 0.001, ^**** p < 0.0001.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: adhm71346‐sup‐0001‐SuppMat.docx.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.