Decidualization-empowered ECM hydrogel integrating sustained Tβ4 release drives endometrial regeneration in intrauterine adhesions

Domain: biology
Published: 2026-01-21
Editorial outcome: major_revision

Abstract
Intrauterine adhesions (IUA), a leading cause of female infertility, result from a pathological switch in the uterine injury response from regeneration to fibrotic scarring. Current treatments are often inadequate as they fail to address this fundamental shift. Here, we report a “decidualization-empowered” hydrogel that reverses this pathology by synergistically combining a bioactive extracellular matrix from decidualized endometrium (DEndo-UdECM) with the sustained release of the anti-fibrotic peptide Thymosin β4 (Tβ4). In a murine IUA model, a single administration of the hydrogel restores endometrial architecture, resolves fibrosis, and, most critically, leads to a near-complete recovery of fertility. Mechanistically, the hydrogel orchestrates a pro-regenerative niche by reprogramming macrophages to an M2 phenotype while dually inhibiting pyroptosis-driven inflammation and the canonical TGF-β/Smad3 fibrotic cascade. Together, these findings demonstrate that mimicking the biological intelligence of a physiological niche can resolve complex fibrotic diseases and restore organ function.

Introduction
Intrauterine adhesions (IUA), characterized by fibrotic scarring of the endometrium, represent a major cause of female infertility and a formidable clinical challenge1. With incidence rates exceeding 30% in high-risk populations, it poses a significant global health burden, culminating in debilitating outcomes including menstrual disorders, recurrent pregnancy loss, and infertility2,3. Current treatments, primarily surgical resection, even when supplemented with physical barriers or hormonal therapies, are plagued by high rates of recurrence (often >60%) and fail to restore fertility in most cases2,4. This therapeutic impasse highlights a fundamental flaw in existing approaches: they do not address the underlying pathology—a pathological switch in the uterine microenvironment from a pro-regenerative to a pro-fibrotic state.

The persistence of IUA is driven by a self-sustaining vicious cycle of inflammation and fibrosis5,6. Following the initial injury, potent inflammatory cascades are initiated, involving NLRP3 inflammasome activation, M1 macrophage polarization, and excessive cytokine secretion7. This inflammatory milieu propagates detrimental cell death pathways, such as pyroptosis8–10, and fuels the aberrant activation of the canonical TGF-β1/Smad3 pathway, which orchestrates excessive collagen deposition and matrix stiffening11,12. Ultimately, this process culminates in the formation of a non-permissive scar tissue that physically and biochemically impedes neo-angiogenesis, hinders the function of endometrial stem/progenitor cells, and obstructs functional regeneration3,13,14.

Addressing the dual challenge of promoting regeneration while suppressing fibrosis demands an integrated biomaterial strategy3,14. However, current approaches are often too simplistic. Many function merely as passive physical barriers or deliver single growth factors, failing to address the multifaceted pathology15–17, while simple hydrogels are limited by their inherent bio-inertness6,18. Decellularized extracellular matrix (dECM) hydrogels represent an advance by retaining intrinsic bioactivity from the source tissue19,20. While even tissue-specific uterine dECM (UdECM) has shown promise21–25, such scaffolds are derived from a quiescent state and thus may lack the potent, specialized cues of the dynamically remodeling, pro-regenerative microenvironment required for true functional restoration.

Nature provides a compelling blueprint for scarless endometrial repair in physiological decidualization, a process that establishes a unique pro-regenerative and immunomodulatory milieu12,26–28. While prior studies inspired by this phenomenon have explored the therapeutic potential of decidual cells or their soluble derivatives29–31, translating cell-based therapies faces clinical hurdles, and these approaches may not fully leverage the persistent cues embedded within the insoluble matrix framework32,33. We therefore hypothesized that a decellularized ECM derived directly from in vitro decidualized endometrium (DEndo-UdECM) could capture this unique, pro-regenerative matrix. This would offer a superior, cell-free, “decidualization-empowered” biomaterial that leverages the persistent physical and biochemical cues of the insoluble scaffold to drive functional repair.

However, to overcome the potent fibrotic drive in severe IUA, we reasoned that this pro-regenerative scaffold required a synergistic partner: a sustained anti-fibrotic agent. For this, we selected Thymosin β4 (Tβ4), a pleiotropic peptide known not only as a potent inhibitor of TGF-β signaling but also for its pro-angiogenic and immunomodulatory activities34,35. To overcome its short biological half-life, we engineered its prolonged delivery via PLGA microspheres to provide a sustained therapeutic pressure36. This creates a dual-action system designed for a synergistic “division of labor”: the DEndo-UdECM scaffold provides the immediate pro-regenerative blueprint, while Tβ4@PLGA acts as a sustained brake on fibrosis.

Here, we develop and validate a composite hydrogel that synergistically integrates the bioactive DEndo-UdECM scaffold with sustained Tβ4 release. We show that in a stringent murine IUA model, a single administration of this hydrogel not only restores endometrial architecture and resolves fibrosis but, most critically, leads to a near-complete recovery of fertility as summarized in Fig. 1. We further elucidate the underlying mechanisms, demonstrating that the hydrogel orchestrates a pro-regenerative microenvironment by reprogramming macrophages, dually inhibiting pyroptosis and TGF-β signaling, and restoring the molecular signature of endometrial receptivity.

Results
Establishment and characterization of a mouse uterine decidualization model
To obtain an ECM reflecting a dynamic uterine microenvironment, we first established a physiologically relevant mouse model of decidualization37. Pseudopregnant mice received intrauterine sesame oil injection on day 4 post-coitus, and uteri were harvested on day 8 (Supplementary Fig. 1a). Decidualization induced marked structural changes, including pronounced bilateral uterine horn distension (Supplementary Fig. 1b). Quantitative analysis revealed significant increases in uterine length (2.516 vs 1.280 cm, P < 0.0001), area (2.296 vs 0.801 cm², P < 0.0001), and weight (814.91 vs 169.68 mg, P < 0.0001) in the decidualized (Dec) group compared to controls (Supplementary Fig. 1c–f). Molecular assessment employed markers for angiogenesis (CD31)38, hormone response (Progesterone Receptor, PR)39, stromal differentiation (HAND2)40, and proliferation (PCNA). Immunofluorescence (IF) staining revealed distinct changes: CD31 expression showed an elevated and continuous pattern in decidualized uterine vessels (75.14% positive area fraction vs controls, P < 0.0001); PR displayed increased nuclear intensity in decidualized stromal cells (P < 0.0001); the PCNA-positive area fraction was significantly higher in the Dec group (12.56% vs 1.71%, P < 0.0001); and HAND2 expression was also significantly upregulated (P < 0.0001) (Supplementary Fig. 1g–j). These morphological and molecular changes confirmed the successful establishment of the mouse uterine decidualization model.

Characterization of DEndo-UdECM scaffolds
Following model establishment, decellularized ECM scaffolds derived from decidualized endometrium (DEndo-UdECM) were prepared and systematically characterized. Macroscopically, DEndo-UdECM scaffolds appeared translucent white, contrasting with the opaque native tissue, while retaining the gross 3D architecture (Fig. 2a). Histological analysis using Hematoxylin and Eosin (H&E) staining demonstrated the absence of nuclear remnants in DEndo-UdECM (Fig. 2b). While collagen fiber density appeared slightly reduced visually compared to native tissue, the overall fibrillar architecture was maintained (Fig. 2b). Masson’s trichrome staining confirmed the preservation of collagen fiber spatial organization (Fig. 2c), and quantitative analysis showed no significant difference in the collagen area fraction between native decidualized tissue and DEndo-UdECM (P > 0.05) (Fig. 2d). Effective decellularization was confirmed by a >98% reduction in DNA content in DEndo-UdECM scaffolds (17.22 ± 4.940 ng/mg) compared to native tissue (1259.07 ± 216.582 ng/mg; P < 0.0001) (Fig. 2e). Key ECM components (Collagen I, Collagen IV, Fibronectin) were assessed by IF21. Compared to native non-decidualized tissue, decidualization significantly increased the area fraction of Collagen I (by ~20%; P < 0.05; Fig. 2f) and Collagen IV (by ~30%; P < 0.01; Fig. 2g). Fibronectin showed a non-significant upward trend in decidualized tissue (P > 0.05; Fig. 2h). The decellularization process resulted in a slight, non-significant decrease in the fluorescence intensity of these three ECM proteins (P > 0.05). The absence of nuclei in DEndo-UdECM was further confirmed by negative DAPI staining (Fig. 2f–h).

To elucidate compositional distinctions, quantitative proteomic analysis compared uterine decellularized ECM (UdECM) from control (CEndo-UdECM) and decidualized (DEndo-UdECM) endometrium. Proteomic profiling identified 198 proteins unique to CEndo-UdECM and 300 unique to DEndo-UdECM, with 8513 proteins common to both (Fig. 3a). Alignment with the Matrisome database identified 20 CEndo-specific, 24 DEndo-specific, and 348 shared matrisome proteins (Fig. 3b). Differential abundance analysis revealed significant matrisome remodeling during decidualization, with 99 proteins significantly upregulated and 126 significantly downregulated in DEndo-UdECM relative to CEndo-UdECM (Fig. 3c; adjusted p-value < 0.05, Fold Change > 1.5). Classification into core matrisome (Collagens, Glycoproteins, Proteoglycans) and matrisome-associated categories highlighted compositional shifts (Fig. 3d). DEndo-UdECM showed enrichment in the relative abundance of ECM regulators (13.4% vs 9.9%), Secreted factors (11.6% vs 3.4%), and Glycoproteins (20.9% vs 14.3%), while CEndo-UdECM maintained higher relative proportions of Collagens (42.3% vs 31.4%), Proteoglycans (11.5% vs 5.7%), and ECM-affiliated proteins (18.8% vs 17.1%). Analysis of the top 10 most abundant ECM proteins revealed shared structural elements and unique signatures (Fig. 3e). While components like Collagen type I (Col1a1, Col1a2), Collagen type VI (Col6a1, Col6a2), Biglycan (Bgn), and Annexins (Anxa2, Anxa6) were prominent in both, CEndo-UdECM was distinguished by high levels of Tgm2, Col6a4, and Dcn. In contrast, DEndo-UdECM featured S100a10, Anxa1, and S100a6. Category-specific protein abundance analysis further detailed these differences (Fig. 3f). DEndo-UdECM was enriched in Glycoproteins like Fibrinogen chains (Fga, Fgb), Fbn1, specific Laminin subunits (Lamb2, Lamc1, Lama5), and Postn, whereas Fn1 and TGFB1 characterized CEndo-UdECM. Collagen VI subunits predominated in CEndo-UdECM, while Collagen I levels were relatively higher in DEndo-UdECM. Among ECM regulators, while Serpinh1 and Tgm2 were shared, DEndo-UdECM uniquely featured Pzp/A2M and Itih3. Shared proteoglycans included Bgn, Dcn, and Hspg2, but Prelp, Lum, and Ogn were enriched in CEndo-UdECM. Annexins (Anxa1, 2, 3, 5, 6) were abundant ECM-affiliated proteins in both. Notably, DEndo-UdECM showed specific enrichment in secreted S100 family members (S100a4, S100a6, S100a10, S100a11).

Gene Ontology (GO) enrichment analysis of the 99 significantly upregulated matrisome proteins in DEndo-UdECM (Fig. 3g) revealed significant associations (adjusted p-value < 0.05) with biological processes including “wound healing”, “positive regulation of response to wounding”, “collagen-containing extracellular matrix”, and “cell-matrix adhesion”. Pathways related to matrix dynamics (“regulation of peptidase activity”, “regulation of collagen metabolic process”, “negative regulation of TGFβ receptor signaling pathway”), hemostasis/angiogenesis (“coagulation”, “fibrinolysis”, “angiogenesis involved in wound healing”, “positive regulation of VEGF signaling pathway”), and cellular processes (“regulation of PI3K/PKB signal transduction”, “regulation of epithelial cell proliferation”, “negative regulation of apoptotic signaling pathway”, “stem cell development”) were also prominent. These distinct proteomic profiles indicate that the decellularization process preserved the overall ECM architecture while reflecting the unique compositional features induced by decidualization.

Fabrication and characterization of DEndo-UdECM/Tβ4@PLGA composite hydrogels
The DEndo-UdECM/Tβ4@PLGA composite hydrogel system was fabricated by homogeneously suspending Tβ4@PLGA microspheres within the DEndo-UdECM precursor solution (Fig. 4a). Rheological measurements showed that the hydrogel precursor solution exhibited shear-thinning behavior, with viscosity decreasing as the shear rate increased (Fig. 4b), indicative of good injectability. This property was visually demonstrated by smoothly extruding the hydrogel precursor through a fine-gauge needle to form stable, high-resolution patterns (Supplementary Fig. 6a). To assess its gelation kinetics at physiological temperature, time-sweep rheology was performed. This revealed a rapid sol-gel transition, with the crossover of the storage modulus (G’) and loss modulus (G”) occurring at approximately 52 s (Fig. 4c), confirming its ability to form a stable scaffold quickly upon injection. The mechanical properties of the resulting gel were then characterized by frequency-sweep analysis, which showed that the storage modulus (G’, ≈ 1.0–1.3 kPa) was consistently higher than the loss modulus (G”, ≈ 0.6–0.7 kPa), indicative of a stable, elastic-dominant scaffold suitable for supporting soft tissue regeneration (Fig. 4d). SEM imaging of the pure DEndo-UdECM hydrogel revealed a porous, interconnected microstructure conducive to cell infiltration and nutrient exchange (Fig. 4i, left panel). Furthermore, its behavior in a physiological environment was assessed. The hydrogel demonstrated a controlled swelling profile, reaching a modest maximum swelling ratio of approximately 18.2% and indicating structural stability without the risk of uncontrolled expansion (Supplementary Fig. 6b). Its in vitro biodegradation profile showed near-complete degradation by day 14 (Supplementary Fig. 6c), a timeframe conducive to guiding endometrial repair.

To enable sustained therapeutic delivery, thymosin β4 (Tβ4) was encapsulated in PLGA microspheres. Particle size analysis revealed a highly consistent and narrow size distribution for both unloaded (mean diameter: 35.5 μm) and Tβ4-loaded (mean diameter: 35.6 μm) microspheres, confirming that drug loading did not significantly alter their physical dimensions (Fig. 4e, f). SEM imaging showed that both unloaded PLGA and Tβ4-loaded microspheres were spherical with smooth surfaces (Fig. 4g). To assess their degradation profile, the microspheres were incubated in vitro under physiological conditions and sampled at various time points. Bright-field microscopy showed a gradual loss of their sharp, spherical morphology over 10 days, while SEM imaging of parallel samples revealed the evolution from a smooth, intact surface to a more porous and eroded structure, indicating progressive degradation (Fig. 4h). Quantitative analysis determined the encapsulation efficiency (EE) to be 76.2 ± 1.6%, with the loading capacity (LC) reaching a plateau at initial Tβ4 concentrations above 300 ng/mL (Fig. 4i). When incorporated into the DEndo-UdECM hydrogel precursor, these microspheres were observed to be homogeneously dispersed throughout the resulting hydrogel matrix, as confirmed by cross-sectional SEM (Fig. 4j, right panel, Supplementary Fig. 7 and Supplementary Movie 1). Critically, the drug release profile was evaluated both for the microspheres alone and for the composite hydrogel system. While Tβ4-loaded microspheres alone exhibited a burst release with nearly 70% released within 3 days, their incorporation into the DEndo-UdECM hydrogel resulted in a significantly more sustained and controlled release profile. This dual-component system effectively mitigated the initial burst and prolonged delivery over the 10-day period, ensuring prolonged local availability of the therapeutic peptide (Fig. 4k). These experiments confirmed the successful construction and characterization of the composite therapeutic hydrogel system, providing the material basis for subsequent biological evaluation.

DEndo-UdECM hydrogel promotes key cellular functions for endometrial regeneration in vitro
Following physicochemical characterization, we systematically investigated the in vitro biological effects of the hydrogel matrices to validate our design rationale, using gelatin as a baseline control. First, we confirmed the fundamental cytocompatibility of the materials. Live/Dead (AM/PI) staining revealed exceptionally high cell viability (>98%) across all groups, with no significant difference observed between the Gelatin, CEndo-UdECM, and DEndo-UdECM hydrogels, establishing the biocompatibility of our ECM-derived materials (Fig. 5a, b; P > 0.05). Next, we assessed the core pro-regenerative functions. We began by evaluating cellular proliferation using two distinct but complementary assays. An EdU incorporation assay demonstrated that both CEndo-UdECM and DEndo-UdECM significantly promoted the proliferation of Ishikawa endometrial epithelial cells compared to the Gelatin control (Fig. 5c, d; P < 0.05). This finding was independently corroborated by Ki67 IF staining, which similarly showed a marked increase in the percentage of Ki67-positive cells in both ECM groups (Fig. 5e, f; P < 0.01). Critically, in both assays, the DEndo-UdECM group exhibited the most potent pro-proliferative effect, underscoring its superior regenerative potential.

Cell migration, a critical process for wound closure, was evaluated using a scratch assay with endometrial stromal cells. Both CEndo-UdECM and DEndo-UdECM significantly accelerated wound closure over 12 h compared to the Gelatin control, with DEndo-UdECM again demonstrating the most robust effect (Fig. 5g, h; 12 h, P < 0.0001). Recognizing that robust angiogenesis is a cornerstone of endometrial regeneration, we then assessed the pro-angiogenic potential using a gold-standard tube formation assay with HUVECs. Qualitative observation over 12 h showed a clear progression towards more complex and stable capillary-like networks in the DEndo-UdECM group (Fig. 5i). To rigorously quantify these striking differences, we performed a comprehensive, multi-parameter analysis at multiple timepoints (3 h, 6 h, 12 h). This analysis provided unambiguous, time-dependent evidence of DEndo-UdECM’s pro-angiogenic superiority. Compared to the Gelatin control, the DEndo-UdECM group induced a dramatic and statistically significant increase in all four key angiogenic metrics: Total tube length (Fig. 5j), Number of junctions (Fig. 5k), Number of meshes (Fig. 5l) Coverage area (Fig. 5m). Taken together, these robust, multi-faceted in vitro data provide compelling evidence that the DEndo-UdECM matrix itself creates a potent pro-regenerative microenvironment by powerfully stimulating epithelial cell proliferation, stromal cell migration and proliferation, and, most critically, the formation of mature vascular networks.

Separately, in an in vitro cellular fibrosis model induced by TGF-β1 in endometrial stromal cells, treatment with 50 nM Tβ4 significantly decreased the expression levels of fibrosis markers α-SMA (P < 0.05), Col1a1 (P < 0.0001), Fibronectin (P < 0.05), COL4 (P < 0.01), and LAMININ (P < 0.05) (Supplementary Fig. 2a–i). To provide quantitative biochemical validation for these phenotypic observations, we performed Western Blot (WB) analysis. The results confirmed that Tβ4 treatment significantly suppressed the TGF-β1-induced upregulation of both α-SMA and COL1A1 protein levels (Supplementary Fig. 2j–l; P < 0.0001), offering direct molecular evidence of its anti-fibrotic activity.

DEndo-UdECM/Tβ4@PLGA hydrogel promotes endometrial regeneration and reverses fibrosis in a murine IUA model
To assess the in vivo therapeutic efficacy of our composite hydrogel, we established a severe murine model of IUA via 95% ethanol-induced chemical injury9, followed by a single administration of the respective treatments (Fig. 6a). Fourteen days post-treatment, we evaluated endometrial regeneration and the resolution of fibrosis. Gross morphological assessment revealed that uteri from the Model group exhibited severe luminal stenosis and cavity dilation, pathologies that were only minimally improved by Tβ4@PLGA monotherapy. In stark contrast, the DEndo-UdECM/Tβ4@PLGA treatment group displayed a uterine morphology most comparable to the healthy Sham group, with no apparent stenosis or fluid retention (Fig. 6b). Histological analysis via H&E staining corroborated these macroscopic findings. The Model group displayed a severely thinned, damaged endometrium with extensive luminal adhesion and a notable absence of endometrial glands. While hydrogels or Tβ4@PLGA alone offered partial recovery, the DEndo-UdECM/Tβ4@PLGA composite hydrogel induced the most profound structural restoration, characterized by a well-organized, thick endometrial layer and numerous regenerated glands (Fig. 6c). Quantification confirmed these observations: the DEndo-UdECM/Tβ4@PLGA treatment led to a significant increase in both endometrial thickness (Fig. 6d) and the number of glands (Fig. 6e) compared to all other groups (P < 0.0001 vs. Model; P < 0.0001 vs. CEndo-UdECM/Tβ4@PLGA). To further validate glandular regeneration, we stained for FOXA2, a key marker for glandular epithelium. IF analysis showed a near-complete loss of FOXA2 signal in the Model group, which was robustly restored by the DEndo-UdECM/Tβ4@PLGA treatment to levels approaching the Sham group and significantly higher than all other treatments (Fig. 6f, g). Along side structural regeneration, a critical component of endometrial repair is the resolution of pathological fibrosis3. Masson’s trichrome staining revealed extensive collagen deposition (indicated by dense blue staining) in the Model group, signifying severe fibrosis. The DEndo-UdECM/Tβ4@PLGA hydrogel treatment resulted in the most substantial reduction in the fibrotic area, significantly outperforming all other groups (Fig. 6h, i; P < 0.0001 vs. Model; P < 0.01 vs. CEndo-UdECM/Tβ4@PLGA). To dissect the molecular underpinnings of this anti-fibrotic effect, we assessed the expression of key pro-fibrotic markers, alpha-smooth muscle actin (α-SMA) and Collagen I (COL1). IF staining showed a dramatic upregulation of both α-SMA and COL1 in the endometrial stroma of the Model group. Conversely, the DEndo-UdECM/Tβ4@PLGA treatment most effectively suppressed the expression of both markers (Fig. 6j), leading to a significant reduction in the mean fluorescence intensity of α-SMA (Fig. 6k) and COL1 (Fig. 6l) compared to the Model and other treatment groups.

Collectively, these results demonstrate that the DEndo-UdECM/Tβ4@PLGA hydrogel orchestrates a dual therapeutic action, concurrently promoting robust structural and glandular regeneration while potently suppressing the pathological fibrotic response that characterizes severe IUA.

DEndo-UdECM/Tβ4@PLGA treatment effects on fertility in IUA mice
To assess functional recovery, the “gold standard“ of fertility was evaluated by quantifying embryo implantation sites and live birth numbers after mating (Fig. 7a). The Sham group exhibited 8 ± 1.0 implantation sites per mouse (sum of both uterine horns), whereas the Model group showed only 2 ± 0.2 site. The Tβ4@PLGA group had 2 ± 1.0 sites (P > 0.05 vs. Model). CEndo-UdECM and DEndo-UdECM treatments increased the number of sites to 3 ± 0.5 and 5 ± 0.5 (P < 0.0001 vs. Model), respectively. The CEndo-UdECM/Tβ4@PLGA group had 5 ± 1.0 sites (P < 0.01 vs. Model). Notably, the DEndo-UdECM/Tβ4@PLGA group reached 7 ± 1.0 sites, significantly higher than the Model group (P < 0.0001) and approaching the levels of the Sham group (Fig. 7b, d). Consistent with implantation data, live birth rates were significantly decreased in the Model group compared to the Sham group (which had 12 ± 2.0 pups per litter) (P < 0.0001). The DEndo-UdECM/Tβ4@PLGA group achieved a live birth rate of 12 ± 0.8 pups per litter, comparable to the Sham group (P > 0.05 vs. Sham) and significantly higher than the Model group (P < 0.0001) and other treatment groups (Fig. 7c, e). To assess the long-term viability and health of the offspring, we monitored their postnatal development. Crucially, all live-born offspring, regardless of the treatment group, exhibited a 100% survival rate to weaning with no observable physical abnormalities (Table S2). This uniform health was further corroborated by comparative imaging of weaned pups, which revealed no discernible differences in body size or physical condition among the groups (Fig. 7f). Collectively, these fertility experiments confirm that the DEndo-UdECM/Tβ4@PLGA hydrogel not only restores implantation rates but, crucially, achieves a comprehensive functional recovery enabling the birth and healthy development of a normal-sized litter.

The composite hydrogel exhibits systemic safety and local biocompatibility in vivo
To evaluate the translational potential of our hydrogel systems, we conducted a comprehensive safety assessment focusing on both systemic toxicity and local tissue compatibility. Systemic safety was first confirmed through histological analysis of major organs (heart, liver, spleen, lungs, and kidneys) harvested from the IUA model animals 14 days post-treatment. H&E staining revealed no evidence of inflammation, necrosis, or other pathological abnormalities in any hydrogel-treated group, with tissue architecture indistinguishable from controls (Supplementary Fig. 3a). For local biocompatibility, we subcutaneously implanted the hydrogels and monitored the tissue response over 21 days. H&E staining provided compelling evidence of the material’s gentle integration; even at day 7, the fundamental skin structure remained intact and well-organized, showing no significant signs of cellular infiltration or structural disruption (Supplementary Fig. 3b). This seamless integration was followed by a complete, scar-free resolution by day 21. Furthermore, Masson’s trichrome staining critically showed no evidence of excessive collagen deposition or the formation of a dense fibrous capsule, confirming the absence of a significant fibrotic foreign body response (Supplementary Fig. 3c). To dissect the molecular underpinnings, time-course immunohistochemical analysis for the pro-inflammatory cytokines IL-6 and IL-1β revealed only a transient, low-level expression at day 7 that was comparable to the PBS control and resolved to baseline by day 21, providing direct molecular evidence that our hydrogels do not elicit a chronic inflammatory state (Supplementary Fig. 3d, e). Taken together, these systemic and multi-level local safety assessments—validated at the histological, architectural, and molecular levels—robustly demonstrate in vivo biocompatibility of the Tβ4@PLGA@CEndo-UdECM and Tβ4@PLGA@DEndo-UdECM hydrogels, supporting their strong potential for future clinical applications.

DEndo-UdECM/Tβ4@PLGA treatment rebalances the immune microenvironment towards a pro-regenerative M2 phenotype
Given that macrophage polarization is a critical determinant of the inflammatory-reparative balance during tissue repair3, we performed a comprehensive four-marker IF analysis to rigorously investigate the influence of our composite hydrogel on macrophage phenotype at the injury site. We assessed two canonical M1 markers (CD86 and iNOS) and two canonical M2 markers (Arginase-1 and CD206).

First, we examined the pro-inflammatory M1 phenotype. Expression of CD86 was significantly upregulated in the Model group compared to the Sham group (P < 0.001) (Fig. 8a, b). Tβ4@PLGA treatment significantly reduced CD86+ cell density (P < 0.001 vs. Model). Both CEndo-UdECM and DEndo-UdECM treatments further decreased CD86+ cell density (P < 0.0001 vs. Model). The DEndo-UdECM/Tβ4@PLGA group exhibited the lowest CD86+ cell density, comparable to the Sham group (P > 0.05 vs. Sham). Consistent with these findings, the expression of the M1 marker iNOS followed a nearly identical pattern, with the DEndo-UdECM/Tβ4@PLGA treatment most effectively suppressing its expression to levels approaching the Sham group (Fig. 8c, d).

Conversely, we assessed the pro-reparative M2 phenotype. Expression of Arginase-1 (Arg-1) was significantly diminished in the Model group compared to Sham (Fig. 8e, f). Tβ4@PLGA treatment alone did not significantly alter Arg-1+ cell density compared to the Model group (P > 0.05). Both CEndo-UdECM and DEndo-UdECM treatments significantly increased Arg-1+ cell density compared to the Model group. The DEndo-UdECM/Tβ4@PLGA group displayed the highest Arg-1+ cell density, similar to the Sham group. This pro-regenerative shift was powerfully corroborated by the analysis of CD206, another key M2 marker. The DEndo-UdECM/Tβ4@PLGA group demonstrated the most profound upregulation of CD206 expression, significantly higher than all other treatment and control groups (Fig. 8g, h).

To further corroborate these IF findings with a robust, single-cell quantitative method, we performed flow cytometric analysis on single-cell suspensions from the regenerated endometrial tissues. After gating on the total macrophage population (F4/80⁺), we quantified the relative proportions of M1-like (CD86⁺) and M2-like (CD206⁺) cells (Supplementary Fig. 8a). This analysis revealed a starkly elevated M1/M2 ratio in the Model group, confirming a dominant pro-inflammatory state. In stark contrast, treatment with the DEndo-UdECM/Tβ4@PLGA hydrogel induced a profound reduction in the M1/M2 ratio to a level even lower than that of the Sham group, significantly outperforming all other control treatments (Supplementary Fig. 8b). This flow cytometry data provides unequivocal quantitative validation of our immunohistochemical observations.

Taken together, this comprehensive four-marker analysis, powerfully corroborated by quantitative flow cytometry, unequivocally demonstrates that the DEndo-UdECM/Tβ4@PLGA treatment orchestrates a profound shift in the local immune microenvironment. It does so by concurrently suppressing the pro-inflammatory M1 phenotype (evidenced by reduced CD86 and iNOS) and promoting the pro-regenerative M2 phenotype (evidenced by elevated Arg-1 and CD206), thereby creating optimal conditions for endometrial repair.

DEndo-UdECM/Tβ4@PLGA treatment effects on markers of regeneration
Effective tissue repair requires active promotion of regeneration alongside inhibition of detrimental processes. Therefore, the pro-regenerative capacity of the hydrogel was assessed by examining markers for epithelialization, stem/progenitor cells, angiogenesis, and proliferation. Co-staining of Pan-Cytokeratin 7 (Pan-CK, epithelial marker) and Aldehyde Dehydrogenase 1 Family Member A1 (ALDH1A1, stem/progenitor cell-associated marker) was performed (Fig. 9a). In the Model group, both Pan-CK+ epithelial structures and ALDH1A1+ cells were largely absent (P < 0.0001 vs. Sham for both) (Fig. 9–c). CEndo-UdECM and Dendo-UdECM single treatments resulted in scattered Pan-CK+ signals (P < 0.001 vs. Model) and a small number of ALDH1A1+ cells (P < 0.001 vs. Model). Tβ4@PLGA treatment showed no significant improvement in Pan-CK+ and ALDH1A1+ signal (P > 0.05 vs. Model). The Dendo-UdECM/Tβ4@PLGA group showed the highest levels of both ALDH1A1+ cells (P < 0.0001 vs. Model, Cendo-UdECM/Tβ4@PLGA, and Dendo-UdECM) and Pan-CK+ epithelial coverage (P < 0.0001 vs. Model, Cendo-UdECM/Tβ4@PLGA, and Dendo-UdECM). Angiogenesis was assessed by CD34 (vascular endothelial marker) IF (Fig. 9d, e). CD34+ cells/vessels were largely absent in the Model group endometrium (P < 0.0001 vs. Sham). No significant improvement was observed in the Tβ4@PLGA (P > 0.05 vs. Model) or Cendo-UdECM (P > 0.05 vs Model) groups. Dendo-UdECM and Cendo-UdECM/Tβ4@PLGA treatments resulted in a small number of scattered CD34+ signals (P < 0.0001 vs. Model). Dendo-UdECM/Tβ4@PLGA treatment resulted in a significant increase in CD34+ cell density compared to the Model group (P < 0.0001), restoring levels comparable to the Sham group and significantly higher than other treatment strategies (P < 0.0001 vs. Cendo-UdECM/Tβ4@PLGA; P < 0.0001 vs. Dendo-UdECM). Cell proliferation was evaluated by Ki67 staining (Fig. 9f, g). Ki67+ proliferating cells were almost undetectable in the Model group (P < 0.0001 vs. Sham). No significant change was observed in the Tβ4@PLGA group (P > 0.05 vs. Model). Cendo-UdECM and Dendo-UdECM treatments showed a marked increase in Ki67+ cells (P < 0.0001 vs. Model). Cendo-UdECM/Tβ4@PLGA also increased Ki67+ cells (P < 0.0001 vs. Model). The Dendo-UdECM/Tβ4@PLGA treatment resulted in the highest Ki67+ cell density, comparable to the Sham group and significantly higher than other treatment strategies (P < 0.0001 vs. Cendo-UdECM/Tβ4@PLGA; P < 0.0001 vs. Dendo-UdECM). Assessment of markers related to epithelialization, stem/progenitor cells, angiogenesis, and proliferation indicated enhanced regenerative processes following Dendo-UdECM/Tβ4@PLGA treatment.

Dendo-UdECM/Tβ4@PLGA treatment effects on pyroptosis and TGF-β/Smad3 signaling
To elucidate the molecular mechanisms underlying the therapeutic efficacy of our composite hydrogel, we investigated its impact on two critical pathological pathways implicated in IUA: pyroptosis-driven inflammation and TGF-β-mediated fibrosis10 (Fig. 10a). WB analysis of uterine tissues revealed a dramatic activation of the pyroptosis cascade in the Model group. This was evidenced by a significant upregulation of Pro-caspase-1 and, critically, a marked increase in the cleaved, active N-terminal fragment of Gasdermin D (GSDMD-NT), the ultimate executor of pyroptotic cell death, while full-length GSDMD levels remained relatively stable (Fig. 10b–e). Consequently, the downstream pro-inflammatory cytokines IL-1β and IL-18 were substantially elevated in the Model group (Fig. 10f, g). While individual components of the treatment showed modest effects, the DEndo-UdECM/Tβ4@PLGA hydrogel potently suppressed this entire inflammatory axis, significantly reducing the levels of Pro-caspase-1, GSDMD-NT, IL-1β, and IL-18 to levels approaching those of the healthy Sham group.

Concurrently, we assessed the canonical pro-fibrotic TGF-β/Smad signaling pathway. IF staining and WB analysis both confirmed a significant upregulation of the master fibrogenic cytokine TGF-β1 in the injured endometrium of the Model group (Fig. 10k–m). This led to a corresponding increase in the phosphorylation of the downstream effector Smad3, as indicated by an elevated p-Smad3/Smad3 ratio (Fig. 10i). The DEndo-UdECM/Tβ4@PLGA treatment most effectively blocked this pathway, resulting in the strongest reduction of both TGF-β1 expression and Smad3 phosphorylation. This upstream inhibition translated directly to a significant downstream reduction in the key fibrotic proteins α-SMA and COL1A1 (Fig. 10h, j), quantitatively corroborating our histological anti-fibrosis findings.

In summary, these molecular analyses demonstrate that the DEndo-UdECM/Tβ4@PLGA hydrogel orchestrates endometrial repair through a powerful dual mechanism: it simultaneously quenches the pro-inflammatory fire of the pyroptosis pathway and dismantles the pro-fibrotic machinery of the TGF-β/Smad axis, thereby creating a microenvironment conducive to regeneration rather than scarring.

DEndo-UdECM/Tβ4@PLGA hydrogel restores the molecular signature of endometrial receptivity
To provide a molecular basis for the observed restoration of fertility, we performed a comprehensive assessment of key markers that define the endometrial receptive state41. The IUA Model group exhibited a molecular profile characteristic of a non-receptive endometrium. Critical pro-receptivity factors were significantly downregulated, including Progesterone Receptor (PR) in both epithelial and stromal compartments, Estrogen Receptor α (ERα), and the essential embryo attachment molecules Osteopontin (OPN) and Integrin β3 (Supplementary Fig. 9a–i). In striking contrast, treatment with the DEndo-UdECM/Tβ4@PLGA hydrogel robustly restored the expression of all these positive markers, elevating their levels to closely approximate those seen in the healthy Sham group.

Conversely, we examined markers associated with a non-receptive or hostile endometrial state. In the Model group, the luminal epithelium displayed aberrantly high levels of the proliferation marker PCNA, indicating a failure to enter the necessary quiescent state required for the implantation window (Supplementary Fig. 9j–l). Furthermore, the anti-adhesion mucin MUC1, which can physically block embryo attachment, was significantly upregulated (Supplementary Fig. 9m, n). The DEndo-UdECM/Tβ4@PLGA treatment effectively reversed these pathological changes, normalizing the expression of PCNA and significantly downregulating MUC1 to levels comparable with the Sham group.

Collectively, these data demonstrate that our composite hydrogel not only physically repairs the endometrium but also fundamentally re-establishes the complex molecular milieu required for successful embryo implantation. This provides a clear mechanistic underpinning for the functional recovery of fertility observed in our study.

Methods
Animal experiments
Sexually mature female ICR mice (6–8 weeks old, 25–30 g), vasectomized male ICR mice (8–10 weeks old), and fertile male ICR mice (8–10 weeks old) were used in this study. All animals were procured from the Experimental Animal Center of Shanxi Medical University (Animal Production License No: SCXK(Jin)2024-0004) and housed under specific pathogen-free conditions at Shanxi Medical University. The animal facility was maintained at 23 ± 2°C with a humidity of 40–70% and a 14 h light/10 h dark cycle. All mice had ad libitum access to standard chow and water.

Materials
Poly(lactic-co-glycolic acid) (PLGA, LA:GA=50:50, Mw 15 kDa, cat. no. P1066) was purchased from Jinan Daigang Biomaterial. Polyvinyl Alcohol (PVA, Mw 30–70 kDa, cat. no. 363138), sesame oil (cat. no. S3507), Triton X-100 (cat. no. T8787), Sodium Dodecyl Sulfate (SDS, cat. no. L3771), and 0.25% Trypsin-EDTA (cat. no. 25200056) were purchased from Gibco. Hank’s Balanced Salt Solution (HBSS, cat. no. H8264) was from Merck, and Penicillin-Streptomycin Solution (cat. no. SV30010) from HyClone. Matrigel Basement Membrane Matrix (cat. no. 356255) was obtained from Corning. For assays and staining, Paraformaldehyde (PFA, cat. no. P1110), H&E Staining Kit (cat. no. G1120), and Masson’s Trichrome Staining Kit (cat. no. G1340) were from Solarbio. Calcein-AM/PI Live/Dead Cell Staining Kit (cat. no. C2015S), BeyoClick™ EdU Cell Proliferation Kit (cat. no. C0071S), RIPA Lysis Buffer (cat. no. P0013B), BCA Protein Assay Kit (cat. no. P0012), DAPI Staining Solution (cat. no. C1005), and Antifade Mounting Medium (cat. no. P0126) were purchased from Beyotime. The Tβ4 ELISA Kit (cat. no. JL19928) was from JONLNBIO, and the DNeasy Blood & Tissue Kit (cat. no. 69504) from Qiagen. Zombie Aqua™ Fixable Viability Kit (cat. no. 423101) was obtained from BioLegend. Detailed information regarding antibodies used for IF, immunohistochemistry (IHC), and WB analysis is provided in Supplementary Table 1.

Preparation and characterization of decidualized uterine extracellular matrix (DEndo-UdECM)
Pseudopregnancy was induced in sexually mature female ICR mice via mating with vasectomized males (1:1 ratio), with the day of vaginal plug detection designated as day 1 (D1). On D4, artificial decidualization was triggered by intrauterine injection of 10 μL sesame oil into each uterine horn under isoflurane anesthesia. Fully decidualized uteri (DEndo-Uteri) were harvested on D8, while untreated, age-matched uteri served as controls (CEndo-Uteri). Successful decidualization was verified by histological assessment of endometrial thickening and vascularization, and IF staining for the decidual marker HAND2. For decellularization, fresh tissues were thoroughly rinsed with deionized water and subjected to a sequential protocol consisting of: (1) incubation in 1% Triton X-100 containing 0.1% EDTA for 1.5 h; (2) incubation in 1% SDS containing 0.1% EDTA for 48 h; and (3) treatment with 200 U/mL DNase I in PBS containing 10 mM MgCl2 at 37°C for 4 h. All steps except enzymatic digestion were performed at room temperature with continuous agitation, interspersed with extensive PBS washes. The resulting matrices were sterilized in 75% ethanol for 30 min, lyophilized, and ground into powder. Decellularization efficacy was evaluated by H&E and Masson’s trichrome staining, and residual DNA was quantified using a DNeasy Blood & Tissue Kit (Qiagen) and a Nanodrop 2000 spectrophotometer (Thermo Fisher Scientific). Retention of key ECM components (COL1A1, COL4, Fibronectin) was confirmed via IF. To compare matrisome profiles, proteins were extracted using SDT lysis buffer (4% SDS, 100 mM Tris-HCl, 0.1 M DTT) and digested via the FASP method using trypsin. Peptides were separated on a C18 analytical column (75 μm × 15 cm) using a 120-min gradient on an UltiMate 3000 RSLC nanosystem and analyzed on a timsTOF Pro 2 mass spectrometer (Bruker Daltonik) in diaPASEF mode. Data were processed using Spectronaut™ Pulsar X software, annotated against the MatrisomeDB database, and differentially expressed proteins (fold change > 2, adjusted P < 0.05) were analyzed via GO enrichment.

Fabrication of Tβ4@PLGA microspheres
Tβ4-loaded PLGA microspheres (Tβ4@PLGA) were fabricated using an optimized water-in-oil-in-water (W/O/W) double emulsion solvent evaporation technique. Specifically, Tβ4 was dissolved in PBS to serve as the inner aqueous phase (W1, 5% w/v), while PLGA was dissolved in DCM to form the oil phase (O, 5% w/v). Under ice-bath conditions, 0.5 mL of W1 was emulsified into 4 mL of O using a probe sonicator (35% power, pulse mode: 4 s on/4 s off, 5 min) to generate the primary W/O emulsion. This primary emulsion was immediately injected into 200 mL of deionized water containing 2% (w/v) PVA (outer aqueous phase, W2) under stirring at 800 rpm to form the secondary W/O/W emulsion. The system was continuously stirred at 600 rpm for > 12 h to allow for complete solvent evaporation and microsphere solidification. The resulting Tβ4@PLGA microspheres were collected via centrifugation, washed three times with deionized water to remove residual PVA and unencapsulated protein, and lyophilized for at least 10 h. The final powder was stored at −80°C. Blank PLGA microspheres were prepared using the same procedure without the addition of Tβ4.

Characterization of Tβ4@PLGA microspheres
The surface morphology of the microspheres was examined using scanning electron microscopy (SEM, ZEISS Gemini SEM 300, Germany) after sputter-coating with gold. Particle size distribution was determined using a dynamic light scattering instrument (DLS, Malvern Mastersizer 2000, UK) after resuspending the microspheres in water. To quantify LC and EE, a precise amount of lyophilized microspheres was dissolved in DCM, and Tβ4 was extracted into an aqueous buffer. The Tβ4 concentration was measured using a commercial ELISA kit according to the manufacturer’s instructions. LC and EE were calculated using the following formulas:

LC (%) = (weight of encapsulated Tβ4 / weight of microspheres) × 100% (1)

EE (%) = (weight of encapsulated Tβ4 / initial input Tβ4) × 100% (2)

For in vitro release kinetics, 10 mg of microspheres were incubated in 1 mL of PBS (pH 7.4) at 37°C with constant shaking (100 rpm). At predetermined time points (days 0, 1, 2, 3, 5, 7, and 10), the supernatant was collected for ELISA analysis and immediately replaced with fresh pre-warmed PBS. Degradation behavior was assessed by incubating 20 mg of microspheres in 1 mL of PBS at 37°C with gentle agitation (300 rpm). At defined intervals, aliquots were imaged using an inverted microscope (Nikon Ti2) to observe hydrated morphology. Parallel samples were washed with deionized water, lyophilized, and examined using SEM (Phenom Pro X) at an accelerating voltage of 10 kV to monitor morphological erosion.

Preparation of DEndo-UdECM/Tβ4@PLGA composite hydrogels
Lyophilized UdECM powder was solubilized in 0.01 M HCl at a concentration of 25 mg/mL containing pepsin (2.5 mg/mL; matrix-to-enzyme ratio of 10:1, w/w) and stirred magnetically at room temperature for 5 days to ensure limited digestion. Following digestion, the pH was neutralized to 7.2–7.4 using 10 N NaOH, and the ionic strength was adjusted with 10× PBS (Ca²⁺/Mg²⁺-free) to yield a final UdECM concentration of 10 mg/mL. The resulting pre-gel solution was maintained on ice to prevent premature gelation. Control 1% gelatin solutions were prepared in 1× PBS. To fabricate the composite hydrogels (DEndo-UdECM/Tβ4@PLGA and CEndo-UdECM/Tβ4@PLGA), pre-weighed Tβ4@PLGA microspheres were dispersed into the ice-cold UdECM pre-gel solution. The mixture was homogenized by gentle pipetting to ensure uniform distribution without introducing air bubbles.

Physicochemical and functional characterization of composite hydrogels
Rheological properties were analyzed using a rotational rheometer (Anton Paar MCR 302) with a 25 mm parallel plate geometry (1 mm gap). Gelation kinetics were monitored via time-sweep oscillatory tests (1 Hz, 1% strain) as the temperature was ramped from 4°C to 37°C. Upon reaching equilibrium, frequency sweep assays (0.1–10 Hz, 1% strain) were conducted to determine the viscoelastic properties of the formed hydrogels. For microstructural analysis, hydrogels formed at 37°C were fixed in 2.5% glutaraldehyde, dehydrated through a graded ethanol series (30–100%), and critical point dried or lyophilized. The internal morphology was visualized by SEM (ZEISS Gemini SEM 300) after gold sputter-coating. Injectability was assessed by extruding the pre-gel solution through a 27G needle.

In vitro degradation was evaluated by incubating weighed hydrogels (approx. 100 µL) in PBS containing 0.01% NaN₃ at 37°C with agitation. The remaining wet weight was recorded at defined intervals over 14 days. Swelling behavior was determined gravimetrically by immersing lyophilized hydrogels in PBS at 37°C and calculating the swelling ratio [(Ws − Wd)/Wd × 100%] over 48 h. To quantify Tβ4 release, composite hydrogels (100 µL) were incubated in PBS at 37°C. Supernatants were collected at predetermined time points, replaced with fresh buffer, and analyzed using a Tβ4 ELISA kit. Cumulative release was calculated relative to the total encapsulated protein. For 3D structural visualization, Rhodamine B-labeled Tβ4@PLGA microspheres were encapsulated within a FITC-labeled DEndo-UdECM hydrogel. Z-stack images were acquired using a spinning-disk confocal microscope (Evident SIM-ultimate) and reconstructed using cellSens software to visualize the spatial distribution of microspheres within the matrix.

Cell culture
Human endometrial stromal cells (hESC, CRL-4003) were obtained from ATCC. The human endometrial adenocarcinoma cell line (Ishikawa) was kindly provided by Prof. Zengming Yang (Guizhou University), and human umbilical vein endothelial cells (hUVEC) were provided by Dr. Jianbing Liu (Shanxi Medical University). hESCs were cultured in DMEM/F12 supplemented with 10% charcoal-stripped fetal bovine serum (CS-FBS) and 1% penicillin-streptomycin (P/S). Ishikawa cells were maintained in DMEM/F12 containing 10% FBS and 1% P/S, while hUVECs were cultured in DMEM/F12 supplemented with 10% FBS, 30 µg/mL ECGS, and 0.1 mg/mL heparin. All cells were incubated at 37°C in a humidified atmosphere containing 5% CO₂. The hESC line was authenticated by the supplier via short tandem repeat profiling. The Ishikawa and hUVEC lines were verified in-house based on cellular morphology and specific marker expression. All cell lines tested negative for mycoplasma contamination. Experiments were performed using hESCs at passages 3–8 and hUVECs at passages 3–6.

In vitro anti-fibrotic and cell migration assays
To assess anti-fibrotic efficacy, hESCs (5 × 10⁴ cells/well) were seeded in 24-well plates and subjected to one of three conditions: basal medium (Control), medium containing 10 ng/mL TGF-β1 (Fibrosis Model), or medium containing 10 ng/mL TGF-β1 supplemented with 100 nM Tβ4 (Treatment). After 48 h, cells were fixed and immunostained for fibrosis markers α-SMA and COL1A1 (antibody details in Supplementary Table 1). Fluorescence intensity was quantified from at least five random fields using ImageJ software (NIH). For migration analysis, a scratch assay was performed on confluent hESC monolayers (2 × 10⁵ cells/well). Following the creation of a cell-free gap using a pipette tip, cells were washed and incubated in low-serum medium (1% CS-FBS) containing extracts from different hydrogels (prepared by incubating hydrogels in serum-free medium for 24 h). Wound closure was monitored at 0, 6, and 12 h using an inverted microscope (Nikon Ti2), and migration distance was analyzed using ImageJ.

Cell proliferation and cytotoxicity analyses
A Transwell co-culture system (0.4 μm pore size, Corning) was established to evaluate the paracrine effects of the hydrogels. Pre-formed hydrogels (100 µL; CEndo-UdECM, DEndo-UdECM, or Gelatin) were placed in the upper chambers, while Ishikawa cells (2 × 10⁴ cells/well) were seeded in the lower chambers. For proliferation analysis, cells were co-cultured for 48 h and then incubated with 10 μM 5-ethynyl-2′-deoxyuridine (EdU) for 2 h using the BeyoClick™ EdU Cell Proliferation Kit with Alexa Fluor 488. Alternatively, cell proliferation was assessed via Ki67 IF staining. The percentage of proliferative cells (EdU⁺ or Ki67⁺) relative to total nuclei (DAPI⁺) was calculated. Cytotoxicity was evaluated using a Calcein-AM/PI Live/Dead Cell Staining Kit. After 48 h of co-culture, cells were stained with Calcein-AM (2 μM) and propidium iodide (PI, 4 μM). Live (green) and dead (red) cells were visualized using fluorescence microscopy.

In vitro angiogenesis assays
The angiogenic potential of the hydrogels was assessed using a tube formation assay. hUVECs (1 × 10⁴ cells/well) were seeded onto Matrigel-coated (30 min at 37°C) 24-well plates. Transwell inserts containing the respective hydrogels were added to the wells to initiate co-culture. Tubular network formation was imaged at 3, 6, and 12 h using a Nikon Ti2 inverted microscope. Quantitative analysis was performed on binary images using the Angiogenesis Analyzer plugin in ImageJ. Key parameters, including total tube length, coverage area, number of junctions, and number of meshes, were calculated from three to five random fields per well.

Animal models and study design
A murine model of intrauterine adhesion (IUA) was established using mechanical and chemical injury. Female mice were anesthetized with isoflurane, and a midline abdominal incision was made to expose the left uterine horn. The uterine segment was clamped, and 50 μL of 95% ethanol was infused into the lumen for 1 min, followed by thorough flushing with sterile saline. The contralateral horn served as an uninjured internal control. Mice were randomized into seven groups (n=24 per group): Sham, Model (PBS), Tβ4@PLGA (microspheres only), CEndo-UdECM, DEndo-UdECM, CEndo-UdECM/Tβ4@PLGA, and DEndo-UdECM/Tβ4@PLGA. Immediately post-injury, 30 μL of the respective therapeutic agents or controls were injected into the uterine cavity. To evaluate in vivo biocompatibility, healthy female ICR mice (n=9 per group) received subcutaneous dorsal injections (100 µL) of PBS, Tβ4@PLGA@CEndo-UdECM, or Tβ4@PLGA@DEndo-UdECM. Skin tissues were harvested at days 7, 14, and 21 for histological examination.

Histological and morphometric analysis
Uterine tissues (n=6 biologically independent animals per group) were harvested 14 days post-surgery, fixed in 4% PFA for 24 h, and embedded in paraffin following standard dehydration through a graded ethanol series. Serial transverse sections (5 µm) were stained with H&E to evaluate tissue architecture, glandular density, and inflammatory infiltration. Endometrial thickness was quantified using ImageJ software by averaging four equidistant perpendicular measurements taken from the myometrial-endometrial junction to the luminal surface in sections displaying a patent lumen. In cases of severe injury manifesting as luminal obliteration (observed in the Model and Tβ4@PLGA groups), endometrial thickness was biologically defined as zero to reflect regenerative failure. To assess fibrosis, sections were subjected to Masson’s trichrome staining. The endometrial stroma was manually delineated as the region of interest (ROI), strictly excluding the myometrium, and collagen deposition was visualized as blue staining. The area of fibrosis was isolated using the color deconvolution plugin in ImageJ, and the fibrosis percentage was calculated as the ratio of the blue-stained area to the total endometrial ROI area. Quantitative analyses were derived from at least three non-consecutive sections per animal, with 3–5 random non-overlapping fields evaluated per section.

Flow cytometry analysis of macrophage polarization
Uterine tissues (n=3 biologically independent animals per group) were harvested, minced into small fragments (~1–2 mm³), and dissociated using a sequential enzymatic digestion protocol. Tissues were incubated in HBSS containing 1% trypsin, 6 mg/mL dispase, and 1 mg/mL DNase I for 1 h at 4°C, 1 h at room temperature, and 10 min at 37°C, followed by secondary digestion with 0.15 mg/mL collagenase I for 35 min at 37°C with gentle agitation. The reaction was neutralized with FACS buffer (DPBS with 2% FBS and 1 mM EDTA), and suspensions were filtered through a 70-µm strainer. Single-cell suspensions (1–2 × 10⁶ cells) were stained with the Zombie Aqua™ Fixable Viability Kit to exclude dead cells and incubated with anti-mouse CD16/32 to block Fc receptors. Surface staining was performed on ice for 30 min using fluorochrome-conjugated antibodies against CD45, F4/80, and CD86. Cells were subsequently fixed with 4% PFA, permeabilized with 0.2% Triton X-100, and stained intracellularly with anti-CD206 for 35 min. Data were acquired on a BD FACSAria™ Fusion Flow Cytometer and analyzed using FlowJo software (v10.8).

The gating strategy was performed as follows: (1) exclusion of debris based on forward scatter (FSC) and side scatter; (2) exclusion of doublets using FSC-H versus FSC-A; (3) identification of live cells (Zombie Aqua-negative); (4) selection of leukocytes (CD45⁺); and (5) identification of macrophages (F4/80⁺). Within the F4/80⁺ population, M1-like (CD86⁺) and M2-like (CD206⁺) phenotypes were quantified based on fluorescence minus one controls.

Reproductive performance and endometrial receptivity
To assess functional fertility restoration, female mice (n=6 biologically independent animals per group) were co-housed with proven fertile male ICR mice (1:1 ratio) two weeks post-surgery. Vaginal plugs were monitored daily to confirm mating. On gestation day 19 (D19), pregnant dams were euthanized, and the number of viable pups was recorded. For implantation analysis, a separate cohort (n=6 per group) was mated two weeks post-treatment. The presence of a vaginal plug was designated as gestation day 1 (D1). On D8, the uteri were harvested, and the number of visible implantation sites, identified as distinct swellings along the uterine horns, was quantified.

To evaluate endometrial receptivity during the implantation window, uterine tissues were collected from a third cohort (n=6 per group) on day 4 post-coitus (D4). The expression of key receptivity markers—including progesterone receptor (PR), estrogen receptor alpha (ERα), integrin β3, OPN, PCNA, and mucin 1 (MUC1)—was analyzed via IHC or IF. The detection method for each marker was selected based on validation assays to maximize signal specificity and preservation of morphological context. For IHC, antigen-antibody complexes were visualized using DAB chromogen and counterstained with hematoxylin. For IF, sections were incubated with appropriate fluorochrome-conjugated secondary antibodies and counterstained with DAPI. Quantitative analysis was performed on representative images based on staining intensity or the percentage of positive cells.

Immunohistochemistry, immunofluorescence, and Western blotting
For histological analysis, uterine tissues were fixed, embedded in paraffin, and sectioned at 5 µm. Sections were deparaffinized in xylene, rehydrated through graded ethanol, and subjected to antigen retrieval using citrate buffer (pH 6.0) or EDTA buffer (pH 9.0) via heat induction. Non-specific binding was blocked using 5% normal serum for 1 h at room temperature.

For IF, sections were incubated with specific primary antibodies (detailed in Supplementary Table 1) overnight at 4°C. Following washing, sections were incubated with species-specific Alexa Fluor-conjugated secondary antibodies for 1 h at room temperature. Nuclei were counterstained with DAPI, and images were acquired using a Nikon Ti2 fluorescence microscope.

For IHC, endogenous peroxidase activity was quenched prior to blocking. Sections were incubated with primary antibodies overnight at 4°C. Detection was performed using a horseradish peroxidase (HRP)-conjugated secondary antibody system followed by color development with 3,3’-diaminobenzidine (DAB). Sections were counterstained with hematoxylin and imaged using light microscopy. Quantitative analysis of staining intensity, positive area, or cell counts was performed using ImageJ software.

For Western blotting, frozen uterine tissues were homogenized in RIPA lysis buffer supplemented with protease and phosphatase inhibitors. Lysates were clarified by centrifugation (12,000 rpm, 15 min, 4°C), and protein concentrations were quantified using a BCA Protein Assay Kit. Equal amounts of protein (20–40 µg) were separated by SDS-PAGE and transferred onto PVDF membranes. Membranes were blocked with 5% non-fat milk or BSA and probed with primary antibodies against GSDMD, Caspase-1, IL-1β, IL-18, Smad3, p-Smad3, and GAPDH overnight at 4°C (Supplementary Table 1). After incubation with HRP-conjugated secondary antibodies, protein bands were visualized using an enhanced chemiluminescence system (Tanon 4600). Band densities were quantified using ImageJ and normalized to GAPDH levels.

Statistical analysis
Statistical analyses were performed using GraphPad Prism 8.0 software. All quantitative data are presented as mean ± standard deviation (SD) derived from at least three independent biological replicates. Data distribution and variance homogeneity were assessed using Shapiro–Wilk and Levene’s tests, respectively. Comparisons between two groups were determined using two-tailed unpaired Student’s t-tests. For comparisons involving multiple groups, one-way analysis of variance (ANOVA) was employed, followed by Tukey’s or Dunnett’s post hoc tests. Non-parametric data were analyzed using the Mann–Whitney U test or Kruskal–Wallis test with Dunn’s multiple comparisons adjustment. A P value < 0.05 was considered statistically significant (*P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This study introduced a newly developed decellularized ECM derived from decidualized uteri and were the first to demonstrate its efficacy in the treatment of IUA. DEndo-UdECM was shown to promote endometrial regeneration and angiogenesis both in vivo and in vitro. Building on these findings, the authors proposed a novel synergistic therapeutic strategy combining DEndo-UdECM with Tβ4 peptides, which are known to enhance tissue repair, stimulate angiogenesis, and critically, inhibit inflammatory and fibrotic pathways, including the TGF-β1/Smad3 signalling axis. In a mouse model of IUA, Tβ4 was administered in combination with DEndo-UdECM as a mixed formulation, functioning as a dual-action system. This approach created a pro-regenerative microenvironment by modulating immune responses, specifically by promoting M2 macrophage polarization, while concurrently suppressing detrimental processes such as pyroptosis and canonical TGF-β/Smad3 signalling.\n\nMajor comment\n1. The rationale for selecting decellularized ECM derived from decidualized endometrium (DEndo-UdECM) remains unclear to this reviewer. The authors stated that “prior studies have explored the therapeutic potential of decidual stromal cells or their derivatives (e.g., exosomes) for IUA, underscoring the value of harnessing decidualization-associated factors” (lines 116–118), and further hypothesise that DEndo-UdECM may retain key bioactive components of the decidual niche (lines 120–122). However, it is well established that non-structural derivatives such as soluble cytokines and exosomes are typically removed during the decellularization process. This raise concerns that DEndo-UdECM may not confer the same regenerative efficacy as reported for decidual cell-derived soluble factors in previous studies. The authors are encouraged to clarify the justification for using decidualized ECM under these considerations and to discuss potential limitations regarding the retention of bioactive molecules following decellularization.\n2. The authors used uterus tissue subjected to sesame oil-induced decidualization as the source for DEndo-UdECM. Is there a specific reason why the authors chose to use a decidualization-induced model instead of utilizing naturally decidualized uterus from mated animals? While this model is commonly used to induce decidualization via artificial mechanical stimulation, it does not fully recapitulate the physiological context of natural decidualization process. I am not fully convinced that the ECM derived from artificially decidualized tissue can adequately mimic a true decidual niche. The authors should justify the choice of this model over physiologically decidualized uteri and discuss how the mode of decidualization might impact the ECM composition and its regenerative efficacy. In order to stick to this model system, the authors should provide a screening data that demonstrate the similarity between sesame oil-induced and naturally-induced decidualization.\n3. In Results 2–4, the authors present that the use of DEndo-UdECM enhances wound healing in endometrial stromal cells and promotes the proliferation of Ishikawa cells (endometrial epithelial cells). However, these findings appear to contradict the well-established concept of the decidualization niche. Decidualization of the human endometrium is commonly defined as a process involving the dramatic morphological and functional differentiation of endometrial stromal cells (ESCs), which is essential for the establishment of a successful pregnancy. This process is typically characterized by cellular differentiation rather than proliferation, migration, or regenerative behaviour. These results therefore raise questions as to whether the DEndo-UdECM truly mimics the characteristics of a decidualized niche or not, as claimed by the authors.\n4. Immunofluorescent images presented both in the main and supplementary figures are not clear at all. The zoomed-in panels appear blurry, as if they have been artificially smoothed or are out of focus. Please clarify whether this issue is due to image resolution degradation or suboptimal imaging during acquisition. All images should be revised and if magnified images are provided, it would be very much helpful.\n\nMinor comments\n1. In Figure 8B, the representative image for the model group does not appear to match the corresponding data points shown in the graph (Figure 8D). According to the experimental design, both uterine horns should have undergone modelling. Could the authors clarify whether each horn received a different treatment, or if there is an inconsistency between the image and the quantitative data?\n2. In Figure 2H-I: The images show fibronectin expression, whereas the main text refers to laminin. This inconsistency should be corrected.\n3. The manuscript currently includes 12 main figures, which makes the presentation somewhat fragmented and potentially overwhelming for the reader. I recommend consolidating the figures where possible to improve clarity and narrative flow. A more compact and focused presentation would enhance the overall readability and impact of the manuscript.\n4. In Figure 7A and 7C, Masson's trichrome staining results appear visually similar across the different groups, with limited red staining observed. It is unclear whether the staining was optimally performed. Additionally, the authors should clarify whether collagen quantification was based on the entire cross-sectional area of the uterus or selected regions of interest. A more detailed explanation of the quantification method and representative staining quality would be helpful to validate the conclusions drawn from this data.", "reviewer_2": "The manuscript presents a conceptually innovative and well-structured study that introduces a “decidualization-empowered” strategy for intrauterine adhesion (IUA) therapy. By integrating extracellular matrix derived from decidualized endometrium (DEndo-UdECM) with sustained release of thymosin β4 (Tβ4) via PLGA microspheres, the authors demonstrate enhanced endometrial regeneration, suppression of fibrosis, and improved fertility restoration in a murine model. While the biological outcomes are thoroughly validated, several critical concerns, particularly related to materials characterization, experimental design transparency, and figure presentation, must be addressed to ensure reproducibility and mechanistic clarity. Therefore, the manuscript requires major revision.\n1. Although the hydrogel system is described as injectable and biodegradable, the manuscript lacks comprehensive characterization of key material parameters that are essential for reproducibility and mechanistic interpretation. For instance,\n(a) There is no direct evidence that Tβ4 is successfully loaded onto or encapsulated within the PLGA microspheres. SEM images of PLGA and Tβ4@PLGA microspheres look indistinguishable.\n(b) The actual Tβ4 loading capacity per unit weight of PLGA microsphere is not reported. Was encapsulation efficiency confirmed via ELISA or other quantification methods?\n(c) There is no information on microsphere porosity: Are the particles solid or porous? If porous, what is the average pore size, and does it fall within micro-, meso-, or macroporous range?\n(d) The distribution of microspheres within the hydrogel is not evaluated. Is it uniform? Was any imaging used to confirm homogeneous dispersion?\n(e) The mechanical properties of the hydrogel (e.g., compressive modulus, tensile strength) are not reported. Given that the hydrogel is injected into the uterine cavity, its mechanical support capacity is critical for functional application.\n(f) The swelling behavior of the hydrogel is not reported. Given the secretory and fluid-rich environment of the uterine cavity, it is essential to assess the anti-swelling capacity of the hydrogel.\n2. The manuscript claim that “time-series SEM visualized the gradual degradation of the embedded microspheres over time” lacks methodological detail and raises interpretive concerns. SEM is inherently a dry-state imaging technique and cannot capture dynamic degradation in situ. The manuscript does not clarify whether the microspheres were extracted from the hydrogel, whether samples were freeze-dried and fractured, or how the time series was established. As such, it remains unclear whether the observed morphological changes truly reflect degradation kinetics within the hydrogel or simply post-processing artifacts.\n3. Figure 1, the schematic appears to depict a surgically opened uterine cavity, which is inconsistent with the experimental protocol. Please revise accordingly. The timeline in the bottom row proceeds right-to-left; for clarity and convention, consider reordering it to left-to-right, ending with “healthy growth.”\n4. Figures 2, 5–7, 9–12, and S1, DAPI staining across multiple figures is too faint to discern nuclear localization. Please adjust imaging brightness/contrast or use enhanced labeling to improve clarity.\n5. Figure 4E, the meaning of “Con” and “Dec” is not defined.\n6. Figure 5, the meaning of “Geltin” is unclear and lacks a corresponding legend. Please clarify. Is “Mre” meant to indicate “merge”? Ensure consistent and explicit labeling. EdU (A, C) and AM-PI staining (B, D) appear to use endometrial cancer cells. Given the focus on regeneration rather than malignancy, this may be inappropriate—please justify or replace. The figure layout does not match the order of data presentation. Reorganize panels accordingly (e.g., C ↔ B, F ↔ H, J ↔ G) to enhance logical flow. Include quantitative analysis of angiogenesis, such as average capillary length or number of branch points in tube formation assays.\n7. Figure 6, a schematic illustration showing how endometrial thickness was measured (e.g., anatomical landmarks, measurement direction, number of sections) would improve reproducibility.\n8. Line 609, the statement that “functional restoration of glands was evidenced by the near-complete recovery of the key transcription factor FOXA2” should be elaborated. Please explain the physiological relevance of FOXA2 and how it reflects glandular function.\n9. Figure 7, Panel order needs reorganization for narrative consistency. The blue channel indicating DNA (DAPI) staining is not labeled. A large discrepancy is observed between fibrotic marker expression in IHC and immunofluorescence—please explain potential methodological or interpretive reasons.\n10. Figure 8, only the postnatal development of pups from the DEndo-UdECM/Tβ4@PLGA group is shown. Were offspring from other treatment groups monitored? Comparative data should be provided to validate the functional benefit.\n11. Figure 9, consider supplementing M2 macrophage polarization data with flow cytometry to enhance rigor and quantification.\n12. Figure 10, the single-channel image for CD34 lacks visible fluorescence signal. Verify if signal acquisition or channel intensity was appropriate. If necessary, adjust exposure or contrast.\n13. Figure 11, a schematic depiction of the TGFβ1/Smad signaling pathway and its activation by the treatment would help contextualize the molecular findings.\n14. Figure 12, clarify the rationale for choosing IHC versus immunofluorescence for each marker. Is it based on antibody availability, signal specificity, or detection sensitivity?\n15. Figure S3, the toxicity assay employed is insufficient to conclude biocompatibility. Standard in vivo biocompatibility evaluation should be conducted—e.g., subcutaneous implantation followed by histology and IHC at defined time points.", "reviewer_3": "This manuscript presents the development of a decellularized extracellular matrix hydrogel incorporating Tβ4-loaded PLGA microspheres. Notably, the study introduces an innovative approach by utilizing decidualized mouse uterus for the preparation of dECM hydrogel, demonstrating favorable regenerative efficacy. However, the scientific evidence remains insufficiently robust and the experimental design requires refinement. Given the current scope and depth of the work, I conclude that this study does not meet the standards for publication in Nature Communications and therefore recommend rejection. Detailed comments are provided below.\nMajor remarks:\n1. Section 3.1 mentions that the dECM hydrogel exhibits thermosensitive gelation properties. Experimental validation of this property, including quantification of the sol-gel transition time, should be provided.\n2. The manuscript only presents the release curve of Tβ4 from microspheres. Please also show the release behavior of Tβ4 from the hydrogel.\n3. The in vitro and in vivo degradation properties of the composite hydrogel should be investigated.\n4. Figure 5, the authors only evaluated in vitro regeneration effects of CEndo-UdECM and DEndo-UdECM. Since the introduction highlights Tβ4's role in enhancing tissue repair and promoting angiogenesis, DEndo-UdECM/Tβ4@PLGA group should be added\n5. A large number of immunofluorescence and histological experiments were used in in vitro and in vivo experiments to observe the phenotype. Please add Western blotting experiments for in vitro experiments to increase credibility\n6. The in vitro anti-fibrosis experiments are insufficient. Additional studies are needed to evaluate the overall effect of the hydrogel.\n7. All immunofluorescence images suffer from poor clarity and overexposure. Please improve the image quality. Non-specific fluorescence signals are evident in multiple immunofluorescence images, particularly in Figures 9A, 10A, and 11H.\n8. As the manuscript described, the established animal model is in fact an endometrium injured-model, other than an IUA model which should present thin endometrium and severe fibrosis.\nMinor remarks:\n1. Caption of Figure 2, “Characterization of UdECM scaffolds before and after decidualization” , herein “decidualization” should be “decellularization”.\n2. Figure 11A, please add the number of samples in each group on the western blot bands.\n3. Figure 11A, please add western blot bands of Total Smad3, and Fibrosis related proteins (e.g. FN, COL1A1…)\n4. In figure. 5B, C, D, the PI/AM staining results of the gelatin hydrogel were not displayed.", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Response to Reviewers for Manuscript ID NCOMMS-25-36427A\nTitle: Decidualization-Empowered ECM Hydrogel integrating sustained Tβ4\nrelease Drives Endometrial Regeneration in Intrauterine Adhesions\nAuthors: Yuxiang Liang#12, Zhaowei Yu#1, Shaobo Du1, Yuqian Guo1, Jing Li1, Yujia Yan1,\nShanshan Jin1, Wenjing Liang1, Mengyuan Li1, Jiao Yang1, Zhiwei Peng1, Zhaoyang Chen2,\nZhizhen Liu*1, Qizhi Shuai*1, Liping Li*1, Jun Xie*1\nIn this response letter, all revisions made to the manuscript text are highlighted\nin blue for clarity.\nPoint-by-Point Responses to Reviewer #1: ... [Full response text omitted for brevity as per instructions to parse the provided text]"}, {"round": 2, "reviewer_comments": {"reviewer_1": "Overall, the revised manuscript presents a coherent and mechanistically substantiated narrative that advances the study from a phenomenological report to a conceptually unified demonstration of a “decidualization-empowered” bioactive matrix. The work now offers clear scientific novelty, methodological rigor, and tangible translational relevance in uterine regenerative medicine. Minor refinements—such as clarifying methodological details related to ECM batch variability—could further enhance reproducibility, but no substantive deficiencies remain.", "reviewer_2": "The authors have revised some figures and text in response to the previous review and have addressed a few of the questions raised. However, many critical issues remain unresolved, and several responses contain inaccuracies or appear to downplay key concerns. Therefore, I recommend major revision or rejection of the manuscript.\n1. Fig. 4i: The image does not clearly demonstrate a homogeneous distribution of Tβ4@PLGA microspheres; local aggregation appears evident. Fluorescence labeling combined with confocal microscopy is recommended to more accurately verify the spatial distribution of the microspheres within the hydrogel.\n2. The authors claim that the hydrogel is self-healing, yet no experimental evidence is provided to support this statement.\n3. Fig. S6c: The x-axis label should indicate days rather than hours.\n4. Time-series SEM images: If these images represent the degradation behavior of standalone microspheres rather than those embedded within the hydrogel, the data should be relocated to the earlier section describing microsphere characterization. In addition, the methodological description in lines 236–255 remains insufficient, which hinders proper evaluation of this experiment. Please provide detailed procedural information.\n5. Regarding the use of endometrial cancer cells (Ishikawa cells) in the experiments from the original Fig. 5 to validate the material’s ability to promote normal tissue regeneration, the authors provided no response. This is a critical issue. Furthermore, in this section of their response, the four references cited under \"Regarding the Rationale for Using HUVECs\" do not actually contain content related to tube formation assays.\n6. We recommended incorporating quantitative statistical analysis of the tube formation assay results. However, the authors failed to implement this. Instead, they cited four references unrelated to tube formation assays to assert that such assays are a gold standard. For reference, quantifiable parameters for tube formation assays include, but are not limited to, the following:\nTube length: Measurement of the total length of tubular structures, reflecting endothelial cell migration and assembly capability.\nCoverage area: Calculation of the area occupied by the tubular network, indicating the extent of angiogenesis.\nNumber of loops/meshes: Enumeration of loop or mesh structures within the tubular network, reflecting vascular network complexity.\nNumber of nodes/branch points: Counting of branching points in the tubular structures, indicating the richness of vascular branching.\n7. The authors partially addressed our concerns regarding endometrial thickness measurements. According to their response, they measured endometrial thickness using the myometrium as the boundary. However, would it be more appropriate to perform these measurements on Masson-stained sections?\n8. In the original Fig. 8, the \"Postnatal development of pups in the DEndo-UdECM/Tβ4@PLGA combination treatment group\" was presented. What about the offspring in other treatment groups? Were there any differences? This question was not directly answered. The authors stated, \"We have added a new column for 'Postnatal Survival Rate (%)' to our new Supplementary Table (Table S1), where every group with live births is listed with a value of '100%'.\" However, the current Table S1 contains antibody-related data, not the relevant survival data. Thus, our query remains unaddressed. Additionally, we still recommend including offspring data in the figure, similar to Fig. 7c, to enhance data reliability.\n9. We suggested that the authors use flow cytometry to detect M2-polarized macrophages in the original Fig. 9, as this would improve data credibility. While the authors provided additional immunohistochemical images for other markers, they did not directly address the use of flow cytometry. We maintain that employing multiple methodologies would enhance data reliability and still recommend using flow cytometry, a relatively more objective method, for statistical analysis.\n10. Please provide IHC analysis of tissue sections after subcutaneous implantation to evaluate whether the material induces an inflammatory response. This is crucial for assessing biocompatibility, for example, by examining inflammatory markers such as IL-1.", "reviewer_3": "The revised manuscript seems to be improved much after addressing the reviewers' comments and is ready to be considered for acceptance.", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": "Response to Reviewers for Manuscript ID NCOMMS-25-36427A\nTitle: Decidualization-Empowered ECM Hydrogel integrating sustained Tβ4\nrelease Drives Endometrial Regeneration in Intrauterine Adhesions\nAuthors: Yuxiang Liang#12, Zhaowei Yu#1, Shaobo Du1, Yuqian Guo1, Jing Li1, Yujia Yan1,\nShanshan Jin1, Wenjing Liang1, Mengyuan Li1, Jiao Yang1, Zhiwei Peng1, Zhaoyang Chen2,\nZhizhen Liu*1, Qizhi Shuai*1, Liping Li*1, Jun Xie*1\nIn this response letter, all revisions made to the manuscript text are highlighted\nin blue for clarity. ... [Full response text omitted for brevity]"}, {"round": 3, "reviewer_comments": {"reviewer_2": "The revised manuscript seems to be improved much after addressing the reviewers' comments and is ready to be considered for acceptance.", "reviewer_4": "I co-reviewed this manuscript with one of the reviewers who provided the listed reports. This is part of the Nature Communications initiative to facilitate training in peer review and to provide appropriate recognition for Early Career Researchers who co-review manuscripts."}, "author_response": ""}]

Decision letter