Mechanically heterogeneous hydrogel with cell-programmed network restructuring promotes tissue regeneration by mechano-epigenetic modulation

Domain: neuroscience
Published: 2026-01-30
Editorial outcome: major_revision

Abstract
Stem cell differentiation dynamically remodels and stiffens the extracellular matrix (ECM), generating stage-specific biomechanical cues that guide tissue development. However, conventional biomaterials, designed to mimic mature ECM stiffness, neglect its spatiotemporal heterogeneity due to their static, non-evolvable nature. Herein, we develop a cell-programmed adaptative contraction (CPAC) hydrogel that enables mesenchymal stem cells (MSCs) to actively remodel their microenvironment through alkaline phosphatase (an early osteogenic marker)-mediated hydrophilic-to-hydrophobic transition and contraction of microgels. This cell-programmed remodeling establishes local mechanical heterogeneity and promotes osteogenesis through a positive feedback loop. Mechanistically, the evolving matrix enhances mechanotransduction-related microRNA expression, suppresses EZH2, and reduces H3K27 trimethylation to active osteogenic transcription. In vivo, MSC-laden CPAC hydrogels significantly enhance the repair of rat cranial defects. These findings introduce a paradigm of cell-instructed, dynamically evolvable biomaterials that recapitulate the adaptive nature of native ECM to orchestrate stem cell fate and tissue morphogenesis.

Introduction
During early bone development, mesenchymal stem cells (MSCs) play a critical role in the continuous remodeling of the extracellular matrix (ECM), a process essential for guiding their differentiation into osteoblasts and driving bone formation1–4. The non-mineralized early developing bone ECM is loosely crosslinked and mechanically homogeneous5–9. As development progresses (Fig. 1a), the dynamic interactions between cells and the ECM lead to progressive restructuring and maturation of ECM due to increasing level of molecular aggregation and cross-linking, thereby transforming the ECM into a complex and heterogeneous structure comprising both the stiff components (e.g., collagen fibrils) and relatively soft elements (e.g., polysaccharide ground substances)10–14. The biochemical modifications of ECM by cell-secreted markers such as enzymes drive this dynamic remodeling and generate the spatially heterogeneous mechanical signals of ECM that are critical for regulating stem cell fate in a timeline that matches the cellular development timescale15–21.

Previous studies have revealed that the biomaterials with prefabricated stiffness promote the commitment of stem cells towards osteogenesis through the activation of mechanotransduction signaling, such as the YAP/TAZ-dependent pathways22,23. However, few studies have demonstrated the ECM-emulating biomaterials that can dynamically remodel into mechanically heterogeneous structures in response to stem-cell-secreted biomarkers to timely provide the complex mechanical cues necessary for guiding the development of complex tissues such as bone based on stem cell differentiations. Previous approaches, such as sequential cross-linking of hydrogels using enzymatic or photoinitiated crosslinkings, have sought to replicate the dynamic stiffening process inherent to natural ECM24,25. However, the temporal and spatial resolution of these artificial stiffening mechanisms remains limited in the absence of cellular participation, leading to potential misalignment with the spatiotemporal requirements of stem cell differentiations. Therefore, the development of a biomimetic hydrogel that can dynamically respond to key differentiation markers of stem cells to undergo the cell-programmed heterogeneous remodeling in both space and time can potentially enhance the differentiation of stem cells and regeneration of tissue defects.

In this study, we develop a microgel-integrated hydrogel system capable of evolving into the ECM-mimetic heterogeneous structure in response to alkaline phosphatase (ALP, a key early osteogenic marker of stem cells), termed cell-programmed adaptive contraction (CPAC) hydrogel. Unlike conventional static microgels, the phosphatase-responsive microgel is designed to be enzymatically modified by ALP secreted by the encapsulated MSCs to undergo a hydrophilic-to-hydrophobic transition and volume contraction, leading to biomimetic structural heterogenization of the CPAC hydrogel with local network stiffening (Fig. 1b). This cell-driven remodeling closely recapitulates native bone ECM development and overcomes the spatiotemporal limitations of previous artificial stiffening strategies. This strategy promotes the mechanosensing and osteogenesis of encapsulated MSCs and, therefore, further elevates the ALP expression, thereby establishing a positive feedback loop that reinforces the osteogenic differentiation through mechanotransduction (Fig. 1c). Notably, the cell-triggered hydrogel local stiffening alters the expression of mechanotransduction-related microRNAs (miRNAs) by the encapsulated MSCs. These mechanosensitive miRNAs target and suppress the expression of EZH2, a key histone methyltransferase, thereby reducing H3K27 trimethylation (H3K27me3) and enhancing the transcriptional activation of osteogenic genes (Fig. 1d). This mechanotransduction–epigenetic coupling enabled by the CPAC hydrogel significantly enhances the osteogenic commitment of MSCs and in situ healing of critical-sized cranial defects in rats. Our findings highlight the critical importance of cell-driven biomaterial adaptation in establishing a spatiotemporally coordinated inductive biomechanical microenvironment for guiding choreographed stem cell differentiation and tissue regeneration. The CPAC hydrogel thus demonstrates a cell-instructed biomaterial system capable of undergoing dynamic, ALP-mediated remodeling to form mechanically heterogeneous structures that promote MSC osteogenesis through mechanotransduction–epigenetic coupling. This work advances the design of biomaterials that respond adaptively to cellular cues, offering a step toward more physiologically relevant ECM mimetics.

Results
Synthesis of phosphatase-responsive contractive microgels as the building blocks of CPAC hydrogels
We first designed and synthesized the phosphorylated hydroxypropyl methacrylate (Phos-HPMA) microgels capable of phosphatase-responsive hydrophilic-to-hydrophobic transition mediated by enzymatic activity (Fig. 2a). Specifically, the Phos-HPMA monomer was synthesized via phosphorylation of HPMA using phosphorus oxychloride26, preserving the integrity of the carbon-carbon double bond, as evidenced by the characteristic vinyl proton resonances at δ 5.7 and 6.1 ppm in the 1H NMR spectrum (Supplementary Fig. 1). Successful phosphorylation was further confirmed by the presence of a distinct peak in the ³¹P NMR spectroscopy (Supplementary Fig. 2). This phosphorylated monomer was subsequently copolymerized with N, N’-methylenebisacrylamide (MBA) and acrylic acid via reverse emulsion polymerization, forming a crosslinked microgel network with controlled composition and uniform morphology. As a control, phosphorylated hydroxyethyl acrylate (Phos-HEA) microgels were synthesized following a similar protocol, with a comparable phosphorylation degree validated by NMR (Supplementary Figs. 3 and 4). HPMA possesses two additional methyl groups and one extra methylene group compared to HEA, which confers enhanced hydrophobicity to HPMA, particularly in its polymerized form. Upon phosphorylation and subsequent microgel fabrication, both Phos-HPMA and Phos-HEA microgels initially demonstrate comparable hydrophilic characteristics. However, under sustained ALP activity, enzymatic cleavage of phosphate groups occurs, leading to a progressive increase in hydrophobicity of Phos-HPMA microgels accompanied by volume contraction. To evaluate the ALP-triggered responsiveness, an equal mass of Phos-HPMA and Phos-HEA microgels was incubated in phosphate-buffered solution (PBS) or 1 U/mL ALP solution (37°C, 72 h). Inverted microscopy revealed a striking size reduction in Phos-HPMA microgels upon ALP treatment, with the average diameter shrinking from 170 μm (as observed in PBS) to 100 μm (in ALP) (Fig. 2b, c). In contrast, the Phos-HEA microgels retained their size (168–165 μm) under both conditions, confirming their enzymatic inertness. Post-lyophilization scanning electron microscopy (SEM) observation further corroborated these findings, revealing substantial deswelling of Phos-HPMA microgels post-ALP exposure, consistent with phosphate ester hydrolysis-induced contraction (Fig. 2d). To ensure seamless integration of the microgels with the network of the proposed cell-laden CPAC hydrogels, all microgels were functionalized with polymerizable vinyl groups, facilitating covalent linkage during network formation (Supplementary Figs. 5 and 6).

ALP-mediated microgel contraction and local stiffening generate the mechanically heterogeneous structures of CPAC hydrogel
We next fabricated the CPAC hydrogel by photopolymerizing a precursor solution containing p-tert-butylphenyl-modified hyaluronic acid (HA-TP), gelatin, mono-acryloyl cyclodextrin (Ac-β-CD), and Phos-HPMA microgels. The successful synthesis of HA-TP and AC-β-CD was demonstrated by 1H NMR spectroscopy (Supplementary Figs. 7 and 8). The obtained CPAC hydrogel comprised covalently bonded microgels embedded within the supramolecular inter-microgel network, stabilized by the dynamic host–guest complexation between TP and β-CD (Fig. 2e). This design enables enzyme-responsive structural reorganization, where the contraction of the Phos-HPMA microgels triggered by cell-secreted ALP can induce inter-microgel network reorganization and structural heterogenization in the CPAC hydrogel. The non-contractile control hydrogel (NC hydrogel) was fabricated by replacing the Phos-HPMA microgels with the Phos-HEA microgels, which lack enzymatic responsiveness. Both hydrogels were formulated with identical gelatin concentrations to ensure consistent cell-adhesive ligand density. Rheological characterization demonstrated that CPAC and NC hydrogels exhibited comparable initial viscoelastic profiles, with a storage modulus (G’) of ~1 kPa, a loss modulus (G”) of ~0.2 kPa (Supplementary Fig. 9a), and Young’s modulus (E) of ~2.5 kPa (Supplementary Fig. 9b). Stress relaxation kinetics showed near-identical characteristic for half of the peak stress relaxation times (τ1/2 = 135 ± 6.85 s) for CPAC and NC hydrogels, confirming that microgel incorporation did not perturb the network dynamics (Supplementary Fig. 9c). Upon ALP treatment, SEM images demonstrated the substantial thinning of the inter-microgel network in the CPAC hydrogel compared to the NC hydrogel (Fig. 2f). Confocal microscopy images of the FITC-labeled hydrogel matrices (green fluorescence) further supported this observation, showing increased fluorescence intensity surrounding the non-fluorescent microgels (black voids) post-ALP treatment, revealing the enhanced local matrix density and structural heterogeneity (Fig. 2g). Rheological characterization documents progressive increase in bulk storage modulus for CPAC hydrogels, achieving 3 kPa after 72 h of ALP exposure, while NC specimens maintained their original bulk stiffness (~1 kPa) (Fig. 2h). Notably, nanoindentation mapping of the CPAC hydrogels revealed a striking increase in local stiffening at the microgel sites (2-fold increase, a 10 kPa stiffness surge), while the surrounding inter-microgel network (red zones) softened due to the microgel contraction-induced thinning of the local network. In contrast, no such changes in local stiffness were observed in the NC hydrogels (Fig. 2(i, ii); Supplementary Fig. 10). Moreover, the theoretical analysis of CPAC hydrogel deformation also confirmed the formation of the heterogeneous structure within the hydrogel, with stiffness increasing radially near the Phos-HPMA microgel surface (Supplementary Fig. 11). To validate the specific response of CPAC hydrogel to ALP, identical sets of CPAC hydrogel were treated with ALP/ALP inhibitor and acid phosphatase, respectively. It was observed that compared to the ALP-treated group, the internal structures of these two groups did not exhibit significant changes (Supplementary Fig. 12a–c), indicating that ALP is the primary factor responsible for the formation of the heterogeneous structure in CPAC hydrogel. We further examined the responsiveness of the hydrogels to cell-secreted ALP by encapsulating osteogenically-induced MSCs in the CPAC and NC hydrogels. As shown in Supplementary Fig. 12d, ALP levels in both groups gradually increased during culture, but CPAC hydrogels exhibited a markedly higher and sustained ALP release. This elevated ALP secretion strongly correlates with the heterogeneous stiffening pattern observed in CPAC hydrogels. After 3 days of culture, nanoindentation mapping (Fig. 2(iii)) showed a similar pattern of heterogeneous stiffening and softening within the CPAC hydrogel treated with exogenous ALP (local effective moduli ranged from ~0.5 kPa in softer regions to ~14 kPa in stiffer regions). This stiffening is driven by cell-mediated microgel contraction, confirming the autonomous remodeling capacity of the material. In contrast, the MSC-laden NC hydrogels showed no significant changes in local stiffness (Supplementary Fig. 10(iii)). Notably, CPAC hydrogels exhibited comparable behavior in vivo, as ALP secreted by MSCs after implantation reached sufficient levels to drive microgel contraction and the formation of heterogeneous structures within the hydrogel (Supplementary Fig. 13). Moreover, the CPAC hydrogel demonstrated rapid structural recovery within 60 s following 60% strain-induced fracture (Supplementary Fig. 9d), confirming robust self-healing behavior driven by reversible TP/β-CD interactions. The hydrogel also retained structural integrity after extrusion through a 21-gauge needle (Supplementary Fig. 14), supporting potential for minimally invasive delivery. This reversible nature of the host–guest interactions in the bulk network enables the adaptive remodeling by facilitating stress redistribution and network reorganization in response to microgel contraction. Additionally, we evaluated the mechanical stiffness and heterogeneous structure of the CPAC hydrogel in both parallel and perpendicular directions. As shown in the Supplementary Fig. 15, the CPAC hydrogel exhibits the same Young’s modulus (~11 kPa), consistent compressive stress–strain curves, and the local effective moduli ranged (from ~0.5 kPa in softer regions to ~14 kPa in stiffer regions), indicating that CPAC hydrogels are mechanically isotropic at the bulk level both in vitro and in vivo. Furthermore, to validate the stability of CPAC hydrogel under complex in vivo conditions, we incorporated variations in pH and temperature into the evaluation. As shown in Supplementary Fig. 16, CPAC hydrogels were able to maintain stable formation of heterogeneous structures within physiological ranges of pH (7.2–7.8) and temperature (33–39°C) for 3 days, with no significant impact on the hydrogel structure. These findings collectively confirm that both the exogenous and cell-secreted ALP can trigger the contraction of Phos-HPMA microgels, thereby inducing the heterogeneous restructuring and associated local stiffness changes in the CPAC hydrogel.

The cell-programmed local stiffening in CPAC hydrogel promotes osteogenic differentiation of MSCs
Before examining osteogenic differentiation, we first evaluated whether CPAC hydrogel supports the proliferation and survival of encapsulated MSCs. Compared to the NC group, CPAC hydrogels exhibited a higher number of EdU-positive cells, indicating an increase in newly synthesized DNA (Supplementary Fig. 17a). Consistently, live/dead staining confirmed robust cell viability (Supplementary Fig. 17b), and CCK-8 assays further verified significantly higher proliferation rates in the CPAC group relative to NC (Supplementary Fig. 17c). These results suggest that the heterogeneous structure formed during microgel contraction not only provides space for cell expansion but also creates a favorable microenvironment that sustains MSC proliferation. Furthermore, the results of Actin staining and the statistical analysis of cell spreading area for MSCs (Supplementary Fig. 18) indicated that MSCs spread over a larger area in the CPAC hydrogel. Subsequently, we evaluated the impact of cellular ALP-induced local stiffening in CPAC hydrogel on the osteogenesis of encapsulated MSCs by analyzing the expression of key osteogenic markers in encapsulated MSCs. After 3 days of culture, PCR results revealed significantly upregulated expression of crucial osteogenic genes, including runt-related transcription factor 2 (Runx2), ALP, Col1, and osteopontin (OPN) in MSCs cultured in CPAC hydrogels compared to those in NC hydrogels (Fig. 3a). The 7-day PCR data also demonstrated that MSCs exhibited a higher degree of osteogenic differentiation within the CPAC hydrogel over longer culture periods (Supplementary Fig. 19). Immunofluorescence staining, Masson’s trichrome staining and ALP color development kit confirmed enhanced protein expression of Col1 and ALP in the CPAC hydrogel group, with greater fluorescence intensity than in the NC hydrogel group (Fig. 3b–d, Supplementary Figs. 20 and 21). Consistently, WB analysis further verified the increased synthesis of Col1 and ALP proteins in the CPAC hydrogel group (Fig. 3e). The above results indicate that the ALP-triggered matrix remodeling and heterogeneous stiffness distribution in CPAC hydrogel can better facilitate stellate spreading and osteogenic differentiation of encapsulated MSCs.
Mechanistically, ALP secreted by differentiating MSCs catalyzes the degradation of phosphorylated Phos-HPMA microgels, leading to structural contraction and localized stiffening within the CPAC hydrogel matrix (Fig. 3f). The resulting mechanical heterogeneity creates spatially confined high-stiffness microdomains that promote cytoskeletal reorganization and focal adhesion formation—key drivers of mechanotransduction. This enhanced mechanical signaling upregulates osteogenic transcription factors (e.g., Runx2 and Col1), reinforcing ALP secretion and accelerating matrix heterogenization through a positive feedback loop. This self-amplifying mechanochemical cascade distinguishes the CPAC hydrogel as a responsive biomimetic matrix capable of undergoing cell-programmed restructuring to promote cellular functions.

The cell-triggered local stiffening of CPAC hydrogel promotes osteogenesis by activating mechanotransduction
To further isolate the pro-osteogenic effects of cell-programmed active matrix remodeling from passive modulus variations, we engineered another control hydrogel system incorporating non-degradable, high-modulus N,N-dimethylacrylamide (DMAA) microgels into the dynamic hydrogel network (Fig. 4a). This non-contractile hydrogel with localized stiffening (NCLS) platform enabled differentiation of the contribution of MSC-driven matrix remodeling from intrinsic stiffness gradients. As shown in Supplementary Fig. 10b (i), the initial effective Young’s modulus of NCLS hydrogels matched the peak modulus of CPAC hydrogels, and this modulus remained unchanged upon exposure to ALP or after 3 days of MSC culture (Supplementary Fig. 10b (ii, iii)). We subsequently evaluated the expression of Col1 and ALP across hydrogel types (Fig. 4b, c). CPAC hydrogels exhibited the highest Col1 and ALP expression compared to all groups, while NCLS hydrogels showed no significant difference in osteogenic marker expression relative to NC hydrogels.
To explore the underlying mechanotransduction pathways, we assessed the phosphorylation of focal adhesion kinase (FAK) at tyrosine 397 (pFAK) and myosin IIa at serine 1943 (pMyosin IIa). Immunofluorescence analysis demonstrated markedly higher pFAK, paxillin, and pMyosin IIa levels in CPAC hydrogels compared to control groups (Fig. 4d–f), with quantitative intensity profiling confirming these observations (Fig. 4g). Additionally, morphometric analysis revealed a significant increase in MSC spreading within CPAC hydrogels (Fig. 4h), suggesting increased cell—matrix interactions. Collectively, these findings confirm that the dynamic, heterogeneous matrix remodeling within CPAC hydrogels promotes osteogenic differentiation through enhanced mechanotransduction. The increased expression of pFAK, paxillin, and pMyosin IIa underscores the activation of mechanosensitive signaling cascades. This supports the established mechanobiology paradigm, where matrix stiffening and dynamic remodeling trigger cytoskeletal tension, reinforcing focal adhesion formation and osteogenic commitment27. These findings highlight that the enhanced mechanotransduction observed in CPAC hydrogels stems from cell-programmed active matrix remodeling, demonstrating the critical role of dynamic matrix heterogeneity guiding osteogenic outcomes.

Mechano-epigenetic regulation governs MSC osteogenesis in CPAC hydrogels
MiRNAs such as miR-124, miR-214, and miR-26a are established regulators of cellular mechanotransduction, influencing chromatin remodeling through the EZH2/H3K27me3 axis28. To investigate their role in osteogenic differentiation within CPAC hydrogels, we compared miRNA expression between hydrogel groups (Fig. 5a). Cells encapsulated in CPAC hydrogels exhibited significantly elevated levels of miR-124-3p, miR-214, and miR-26a compared to NC controls. This upregulation correlated with a marked reduction in the expression of EZH2 and its downstream target, H3K27me3, in CPAC hydrogels (Fig. 5b). Immunofluorescence analysis further confirmed these findings, showing diminished EZH2 and H3K27me3 fluorescence intensity in CPAC hydrogels compared to NC hydrogels (Fig. 5c–f). ChIP-qPCR then demonstrated that H3K27me3 was enriched at the RUNX2 promoter (Fig. 5i), and this reduction was accompanied by increased RUNX2 transcription (Fig. 3a) and enhanced protein expression of Col1 and ALP (Fig. 5g). In addition, to further validate the mechanistic link, we incorporated siRNA-based inhibitors of miR-124, miR-214, and miR-26a into the CPAC hydrogel cultures. Western blot analysis demonstrated that the inhibition of these miRNAs markedly increased the protein levels of EZH2 and H3K27me3, while simultaneously reducing the expression of osteogenic markers ALP and Col1 (Supplementary Fig. 22). These results demonstrate that the cell-programmed matrix heterogenization in CPAC hydrogel establishes an epigenetic landscape conducive to osteogenic differentiation through miRNA-mediated suppression of the EZH2/H3K27me3 axis.
To further validate the role of EZH2 in MSC osteogenic differentiation, we pharmacologically inhibited EZH2 activity in NC hydrogels using a selective EZH2 methyltransferase inhibitor. After 3 days of cell culture, immunofluorescence staining revealed a substantial decrease in EZH2 and H3K27me3 expression, accompanied by a significant increase in Col1 and ALP (Fig. 5g). The fluorescence intensity of Col1 and ALP was notably higher in the inhibitor-treated group compared to the untreated controls (Fig. 5h), reinforcing the role of EZH2 suppression in promoting osteogenic differentiation. Collectively, our findings support a mechano-epigenetic regulatory model in which MSC-driven formation of locally stiffened microdomains within CPAC hydrogels activates cellular mechanotransduction pathways. This mechanical stimulation upregulates the expression of miR-124-3p, miR-214, and miR-26a, which in turn suppress EZH2 synthesis28–31, reducing H3K27 trimethylation in the nucleus and enhancing the expression of osteogenic genes32 (Fig. 5j). The resultant increase in ALP secretion further promotes localized matrix heterogenization through enzymatic microgel contraction, establishing a positive feedback loop that reinforces the osteogenic differentiation of MSCs encapsulated in the heterogeneous hydrogel.

MSC-laden CPAC hydrogels promote bone regeneration in an animal critical defect model
Before evaluating the therapeutic efficacy of dynamic heterogeneous CPAC hydrogel in facilitating bone regeneration, we first assessed the long-term stability of the hydrogel both in vitro and in vivo. As shown in Supplementary Fig. 23, the hydrogel underwent gradual degradation over 4 weeks, consistent with the expected degradation trend of implantable materials. However, the heterogeneous structure in the remaining non-degraded regions remained stable (Supplementary Fig. 24). This indicates that the CPAC hydrogel can stably provide the heterogeneous structural extracellular microenvironment required by MSCs during bone defect repair. Moreover, to evaluate the therapeutic efficacy of dynamic heterogeneous CPAC hydrogel in facilitating bone regeneration, we employed a rat calvarial defect model. Both CPAC and NC hydrogels encapsulated with rat mesenchymal stem cells (rMSCs) were cultured in osteogenic medium for 3 days before being transplanted into the rat cranial defects (Fig. 6a). As shown by micro-CT imaging and quantitative analysis, the CPAC group promoted the regeneration process in the early (4 weeks) and late (8 weeks) stages of bone repair (Fig. 6b, Supplementary Fig. 25), revealing denser new bone formation than that in the NC group and untreated (“Blank”) group. Specifically, at 8 weeks postoperatively, the BV (Bone volume) of the CPAC hydrogel group measured at 11.42 ± 0.87 mm3 (Fig. 6c, Supplementary Fig. 26a, b), which were 4.7-, 3.1-, and 2.4-fold greater than those of the Blank, Control, and NC hydrogel groups, respectively. The BV/TV (bone volume/total volume) and BS/BV (bone surface/bone volume) values exhibited similar trends, further demonstrating markedly higher new bone volume in the CPAC hydrogel group compared to the other groups (Fig. 6d, e, Supplementary Fig. 26c, d). Histological staining provided additional evidence of enhanced bone matrix formation. Hematoxylin and eosin (H&E) staining and Masson’s trichrome staining revealed that the CPAC hydrogel group presented significantly greater new bone and fibrous tissue matrix formation than other groups (Fig. 6f, g, Supplementary Fig. 26e, f). Moreover, immunohistochemical (IHC) staining of OCN, a key osteogenic marker, showed markedly higher expression in the CPAC hydrogel group than in other groups, confirming enhanced osteoblastic activity (Fig. 6h, Supplementary Fig. 26g). Collectively, these results establish that CPAC hydrogels promote robust bone regeneration, highlighting the potential of CPAC hydrogels for improving critical bone defect repair.
While the CPAC hydrogel demonstrates robust osteoinductive performance, some limitations remain. The mechanical heterogeneity of the system arises from cell-secreted ALP activity rather than pre-defined gradients, and the precise spatial control of stiffness may vary with cell density or metabolic activity. In addition, long-term mechanical stability and remodeling behavior under complex in vivo conditions warrant further investigation.
In this work, we developed a cell-programmed heterogeneous dynamic hydrogel system, termed CPAC hydrogel, which replicates the mechano-microenvironment essential for MSC osteogenic commitment. This construct leverages host–guest interaction-mediated dynamic networks to mimic native ECM remodeling capabilities faithfully. Moreover, MSC-directed dephosphorylation of microgel components enables spatiotemporal modulation of network heterogeneity through precisely controlled hydrophilic-hydrophobic phase transitions. The resultant CPAC hydrogel establishes a locally stiffened matrix that promotes the osteogenic differentiation of encapsulated MSCs by activating the mechanotransduction signaling pathways. Notably, the structural dynamics and heterogeneity within the CPAC hydrogel upregulated mechanosensitive miRNA expression in MSCs, thereby suppressing EZH2-mediated histone H3K27 trimethylation for enhanced osteogenesis. Functional validation through a critical-sized rat calvarial defect model confirmed the therapeutic efficacy of MSC-laden CPAC hydrogels in facilitating cranial bone regeneration. These findings establish the critical role of biomimetic physical cues in next-generation regenerative matrix design, establishing CPAC hydrogel as a promising platform for bone tissue engineering.

Methods
Protocol approval
The experimental procedures in this study were conducted in strict compliance with relevant ethical guidelines. The use of animals was reviewed and approved by the Animal Care and Use Committee of South China University of Technology (Permit Number 2023088).

Materials
Sodium hyaluronate (HA-Na, Mw=4 kDa) was purchased from Bloomage Freda Biopharm (China). Dowex was purchased from sigma, tetrabutylammonium hydroxide di-tert-butyldicarbonate (Boc Anhydride), sodium chloride (NaCl), β-cyclodextrin (β-CD), 4-(t-Butyl)-phenylacetic acid, 4-dimethylaminopyridine (DMAP), 2-hydroxyethyl methacrylate, 2-hydroxypropyl methacrylate, N,N-dimethylacrylamide, phosphorus oxychloride, span 80, cyclohexane, N,N’-methylene diacrylamide, acrylic acid, and 3-Buten-1-amine hydrochloride were purchased from J&K Chemicals (China). (USA). PBS, Dulbecco’s modified Eagle medium, fetal bovine serum (FBS), penicillin/streptomycin, alpha-minimum essential medium (α-MEM), and L-glutamine were purchased from Gibco (USA). The Calcein-AM, propidium iodide, Cell Counting Kit-8 (CCK-8), DAPI, TRIzol, Alkaline Phosphatase (ALP) Assay Kit, H&E staining kit, were purchased from Beyotime.

Synthesis of phosphorylated hydroxyethyl acrylate (Phos-HEA) and phosphorylated hydroxypropyl methacrylate (Phos-HPMA)
First, phosphorus oxychloride (7.46 g, 1.1 eq) was mixed with tetrahydrofuran (33 mL) and transferred into a 250 mL round-bottom flask under a nitrogen atmosphere. The flask was then placed in an ice bath for cooling. 2-hydroxyethyl acrylate (5.14 g, 1 eq) and triethylamine (4.93 g, 1.1 eq) were mixed with tetrahydrofuran (7 mL) and added into a dropping funnel under nitrogen. The solution was slowly dripped into the round-bottom flask. After 3 h of thorough reaction, 7.5 mL of water was added to the dropping funnel and slowly dripped into the flask to complete the hydrolysis. The reaction mixture was heated to room temperature and washed three times with saturated sodium chloride solution. The tetrahydrofuran (THF) was removed by rotary evaporation, yielding a light brown oily product (Phos-HEA). HPMA (6.38 g, 1 eq) was used to synthesize the Phos-HPMA following the same procedure.

Preparation of Phos-HEA microgel, DMAA microgel, and Phos-HPMA microgel
Phos-HEA (280 mg, 1 eq) was weighed and dissolved in 10 mL of H2O. Acrylic acid (10 μL, 0.1 eq) was added to the solution. N,N-dimethyl-bisacrylamide (MBA, 22 mg, 0.1 eq) was then dissolved in the solution, followed by the addition of a KPS solution (KPS (7 mg, 0.018 eq) dissolved in 100 μL of DI water). The reaction mixture was maintained under a nitrogen atmosphere at a stirring speed of 1200 rpm, and then mixed with 40 mL of cyclohexane and 900 μL of Span 80 to form a homogeneous emulsion. The emulsion was heated to 70 °C and stirred for 2 h. Afterward, the mixture was centrifuged, washed five times with anhydrous ethanol/DI water, and freeze-dried to obtain white Phos-HEA microgel. DMAA (142 mg, 1 eq) and Phos-HPMA (320 mg, 1 eq) were used to prepare the DMAA microgel and Phos-HPMA microgel, respectively, following the same procedure.

Preparation of Ac-Phos-HEA microgel, Ac-DMAA microgel, and Ac-Phos-HPMA microgel
Phos-HEA (1 g) was weighed and dispersed in 50 mL of H2O. Nitrogen gas was bubbled through the solution for 30 min to remove oxygen. EDC (3.68 g, 19.2 mmol) and NHS (552 mg, 4.8 mmol) were weighed and dissolved in the above solution to activate the carboxyl groups on the microgel surface. The solution was stirred for 2 h, after which 3-buteneamine hydrochloride (258 mg, 2.4 mmol) was added. The pH of the solution was adjusted to 8 using triethylamine. The reaction was carried out under a nitrogen atmosphere at room temperature for 12 h. The reaction mixture was placed in a 3500 Da dialysis bag and dialyzed for 5 days. Finally, the product was freeze-dried, yielding 75%. The DMAA microgel (1 g) and Phos-HPMA microgel (1 g) were used to prepare the Ac-DMAA microgel and Ac-Phos-HPMA microgel following the same procedure.

Synthesis of 4-(t-Butyl)-phenylacetic acid modified hyaluronic acid (HA-TP)
A solution of sodium hyaluronate (HA‑Na, 2.0 g) in deionized water was treated with Dowex resin (6.0 g) for ion exchange, then neutralized with tetra‑n‑butylammonium hydroxide (TBAOH) to obtain an aqueous HA‑TBA solution at pH 7.0. After freeze‑drying, the product was redissolved in dimethyl sulfoxide (DMSO). HA‑TBA (1.0 g, 1.4 mmol in terms of disaccharide repeating units, 1 eq.), 4‑(tert‑butyl) phenylacetic acid (0.8 g, 4.2 mmol, 3 eq.), and 4‑dimethylaminopyridine (DMAP, 0.13 g, 1.05 mmol, 0.75 eq.) were dissolved in anhydrous DMSO (100 mL) at 25 °C. Under nitrogen atmosphere and at 25 °C, di‑tert‑butyldicarbonate (BOC₂O) was added dropwise. The mixture was warmed to 45 °C and stirred for 24 h. The resulting reaction mixture was dialyzed successively against DMSO, NaCl solution, and deionized water to remove low‑molecular‑weight impurities. Finally, the purified solution was freeze‑dried to afford the product as a white solid.

Synthesis of Ac-β-CD
Ac-β-CD was synthesized following established procedures from the literature33. In a typical preparation, β-cyclodextrin (β-CD, 10 g) was dissolved in dimethylformamide (DMF, 150 mL) and triethylamine (TEA, 7 mL). The solution was cooled to 0 °C with stirring. Acryloyl chloride (5 mL) was then introduced, and the reaction was allowed to proceed for 6 h. Following the reaction, the mixture was filtered to eliminate the precipitated trimethylamine hydrochloride salt. The filtrate was concentrated to roughly 20 mL via rotary evaporation. This concentrated solution was subsequently added dropwise into acetone (600 mL), which induced the formation of a white solid. The precipitate was collected, washed thoroughly with acetone, and finally subjected to freeze-drying to obtain the product.

Preparation of NC, NCLS, and CPAC hydrogel
For the NC hydrogel, HA-TP (1%), gelatin (3%), Ac-β-CD (5%), and Ac-Phos-HEA microgel (5%) were dissolved in PBS (900 μl). After complete dissolution, the solution was allowed to stand overnight. Subsequently, 100 μl of 0.5 wt% lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP) was added, and polymerization was initiated under blue light (λ=405 nm, 350 mW/cm²) for 5 min to obtain the NC hydrogel. For the NCLS and CPAC hydrogels, the Ac-Phos-HEA microgels were replaced with an equal mass of Ac-DMAA microgels and Ac-Phos-HPMA microgels, respectively, while keeping all other conditions identical.

3D cell culture within hydrogels
The human mesenchymal stem cells (hMSCs) were purchased from REGEN-αGEEK Biotechnology Co. Ltd (a 65-year-old male donor). Primary hMSCs were cultured in αMEM supplemented with 16.7% FBS (Excel), 1% penicillin-streptomycin (Gibco), and 1% L-glutamine (Gibco), and passaged to passage 4 (P4) under conditions of 37 °C and 5% CO2. After 5 min of digestion with 0.05% trypsin (Gibco), the cells were collected. Following cell counting, the cells were placed into individual 1.5 mL Eppendorf tubes and centrifuged at 290 × g for 3 min, with the supernatant discarded. For each type of hydrogel, the encapsulation density of hMSCs was 1 × 10⁷ cells per mL of hydrogel precursor solution (NC, NCLS, or CPAC). After mixing, 50 µL of precursor solution containing hMSCs was crosslinked under blue light (λ=405 nm) for 5 min and transferred into a 24-well plate. Constructs were continuously cultured in either growth or osteogenic medium (10 mM β-glycerophosphate, 50 µg/mL ascorbic acid 2-phosphate, and 100 nM dexamethasone). For the EZH2 inhibitor group, the inhibitor (Cat: HY-148458, MCE) was added to osteogenic medium at a final concentration of 10 μM. Constructs were cultured in the inhibitor-containing osteogenic medium for 12 h, after which the medium was replaced with fresh osteogenic medium for continued culture.

Microgel shrinkage quantification
Microgel shrinkage was quantified by capturing images of Phos-HEA and Phos-HPMA microgels before and after ALP treatment using an inverted microscope. Fifteen microgels were randomly selected from each group, and their diameters were measured using NanoMeasure software (Version 1.2). The shrinkage of the microgels was quantified using NanoMeasure (Version 1.2) with quantitative analysis of fluorescence intensity conducted using ImageJ software (NIH).

Fabrication and rheological characterization of the NC and CPAC hydrogels
Rheological characterizations were performed using a TA Discovery HR-30 hybrid rheometer. Samples were formed in situ by pipetting a 50 µL monomer solution between the bottom HR-30 hybrid rheometer plate and an 8 mm flat plate, and the experiment was initiated after photopolymerization (λ=365 nm, 396 mW/cm2). Evolution experiments were performed at 37 °C. Time sweep measurements were conducted at 37 °C with a fixed strain of 1% and a frequency of 10 rad/s for 120 s. For the alternating strain steps hear test, the temperature was maintained at 37 °C, with a frequency of 10 rad/s. The strain was alternated between 60% (45 s) and 1% (45 s) for three full cycles to obtain the step shear profile. The stress relaxation properties of the samples were measured from compression tests of the gel discs (8 mm in diameter, 2 mm thick). The gel discs were compressed to 15% strain with a deformation rate of 0.3 mm s−1 and held for 300 s to obtain the stress relaxation curve. The Young’s modulus was then calculated as the slope of the linear region of the resulting stress‑strain curve. Each sample group was tested independently three times to ensure reproducibility.

Phos-HPMA microgel mediated the heterogeneous distribution of matrix stiffness in the CPAC Hydrogel
Nanoindentation technology was applied to characterize the Young’s modulus distribution of both hydrogels under underwater conditions via a Piuma nanoindenter (Optics 11, Netherlands). Specifically, the hydrogels were maintained in PBS at 37 °C for 1 h before being loaded onto the plate. We randomly selected a 350 × 350 μm square matrix area on the surface of the samples and performed nanoindentation measurements at intervals of 50 μm. A total of 8 × 8 valid Young’s modulus data points were obtained, forming a heatmap. The selected areas were random, and the measurements were repeated three times.

Real-time quantitative PCR
Total RNA was isolated from cultured samples using TRIZOL reagent (Vazyme, R401-01) following the manufacturer’s protocol. RNA concentration and purity were assessed by spectrophotometric measurement (NanoDrop 300, Thermo Fisher Scientific). First-strand cDNA synthesis was performed using PrimeScript RT Master Mix (Takara, RR014A) according to the manufacturer’s specifications. Quantitative PCR was carried out using TB Green Premix Ex Taq (Takara, RR820A) on a LightCycler 480 instrument (Roche Diagnostics). The thermal cycling protocol comprised: initial denaturation at 95 °C for 30 s, followed by 40 cycles of amplification (95 °C for 5 s, 60 °C for 30 s), and a final dissociation stage (95 °C for 15 s, 60 °C for 1 min, and 95 °C for 15 s). GAPDH served as the endogenous control for normalization. Relative gene expression levels were determined using the comparative 2−ΔΔCT method. Primer sequences are provided in Supplementary Table 1.

Immunofluorescence staining
Following a brief PBS rinse, specimens were fixed in 4% paraformaldehyde (Aname, #153799) for 30 min at ambient temperature. After two additional PBS washes, samples were permeabilized and blocked for 2 h at room temperature using PBS containing 3% bovine serum albumin (Millipore, #S30) and 0.25% Triton X-100 (Sigma, #T8787). Primary antibody incubation was carried out overnight at 4 °C in blocking buffer. Unbound antibodies were removed through three washes with dilution buffer, followed by 1-h incubation with Alexa Fluor 488-conjugated secondary antibody (1:200 dilution). For cytoskeletal and nuclear visualization, samples were subsequently treated with phalloidin (1:400, 1 h) and DAPI (1:100, 30 min). Fluorescence imaging was performed using a Zeiss LSM780 confocal system, with quantitative analysis of fluorescence intensity conducted using ImageJ software (NIH).
The primary antibodies used were as follows: anti-Col 1 (Cat. #14695-1-AP, ProteinTech, 1:500), anti-ALP (Cat. #DF6225, Affinity, 1:500), anti-FAK (phospho Y397) (Cat. # 66258-1-lg, ProteinTech, 1:500), anti-paxillin (Cat. # 10029-1-lg, ProteinTech, 1:500), anti-phospho-MYH9 (Ser1943) (Cat. # AF4425, Affinity, 1:500), anti- EZH2 (Cat. #21800-1-AP, ProteinTech, 1:500), anti-H3K27me3 (Cat. #A2363, Abclonal, 1:500).
The secondary antibodies used were as follows: CoraLite488-conjugated goat anti-mouse IgG (H+L) (Cat. #SA0013-1, ProteinTech, 1:500), CoraLite488-conjugated goat anti-rabbit IgG (H+L) (Cat. #SA0013-2, ProteinTech, 1:500), and goat anti-rabbit Alexa Fluor 594 (Cat. #S0006, Affinity, 1:500).

Western blot
Cellular proteins were lysed using ice-cold RIPA buffer containing 1 mM phenylmethylsulfonyl fluoride. Nuclear and cytoplasmic fractions were isolated following the supplier’s protocol (Beyotime, Cat. #P0013B). Protein concentrations were determined using a bicinchoninic acid assay, followed by heat denaturation at 98 °C for 15 min. Protein samples (20 µg) were resolved on 10% SDS‒PAGE gels, initially at 80 V for 30 min and subsequently at 120 V for 80 min. Separated proteins were transferred onto PVDF membranes at 300 mA for 120 min. Membranes were blocked with 5% nonfat milk for 1 h at room temperature, followed by incubation with primary antibodies at 4 °C overnight. HRP-conjugated secondary antibodies were applied for 1 h at room temperature. Protein bands were detected using an enhanced chemiluminescence substrate (Epizyme, Cat. #SQ201) and imaged with a Bio-Rad system (Tanon, Shanghai, China). Band intensities were quantified using Image Lab software (Bio-Rad). A fixed rectangular selection tool was applied to each target band, and local background signals were subtracted to obtain net intensities. To normalize for loading differences, target protein signals were adjusted relative to internal controls (e.g., GAPDH) within the same lane. Triplicate experimental data were processed using GraphPad Prism 9.0 and expressed as mean ± SD. Statistical comparisons were performed via unpaired two-tailed Student’s t-test, with significance set at p<0.05.

Chromatin immunoprecipitation (ChIP) and ChIP-qPCR
Cells were fixed with 1% formaldehyde and subsequently quenched with glycine. After washing with PBS, the fixed samples were suspended in cell lysis buffer (5 mM PIPES-KCl, pH 8.0, 85 mM KCl, 0.5% NP40), and the collected nuclei were stored at −80 °C as nuclei pellets or nuclear lysates dissolved in nuclei lysis buffer (50 mM Tris-HCl pH 8.0, 10 mM EDTA, 1% SDS). For H3K27me3 ChIP, lysates were thawed and sonicated with Sonifier (BRANSON) to obtain chromatin fragments of 300–1000 bp, and 10-fold-diluted with ChIP dilution buffer (16.7 mM Tris-HCl pH 8.0, 1.2 mM EDTA, 0.01% SDS, 1.1% Triron 100×, 167 mM NaCl). After double-dilution with a sonication buffer (90 mM Hepes pH 7.9, 220 mM NaCl, 10 mM EDTA, 1% NP40, 0.2% sodium deoxycholate, 0.2% SDS), nuclei were homogenized with a Bioruptor (Toshodenki). After washing, a complex of nuclear proteins/DNAs and antibodies against H3K27me3 (Proteintech; 39157; 5 ug used) was retrieved with Protein A- and Protein G-Dynabeads (Life Technologies). After the cross-linking was reversed, chromatin fragments were treated with RNase A and proteinase K. DNA was purified with phenol-chloroform extraction or Ampure XP (Beckman Coulter). In the ChIP-qPCR analyses, the values from the immunoprecipitated samples were normalized to input DNA. Primer sequences are listed in Supplementary Table 2.

Ratskull defect repair model
All experimental procedures involving animals were conducted in accordance with protocols approved by the Animal Experimentation Ethics Committee of the South China University of Technology (AEC No. 2023088). Nine 8-week-old male Sprague-Dawley (SD) rats were acclimatized for 7 days under controlled conditions (24–26 °C, 70% humidity). The rats were randomly allocated into four groups (n=3 per group): (1) NC hydrogel, (2) CPAC hydrogel, (3) CPAC hydrogel without MSCs (Control), and (4) non-treated control (Blank). Following institutional guidelines, anesthesia was induced via intramuscular injection of a ketamine-xylazine mixture (0.2% ketamine, 0.1% xylazine in saline). Aseptic surgical exposure of the calvarium was performed, and two critical-size defects were created in the calvarium with a trephine drill. In each animal, the contralateral defect served as an internal control. Primary rMSCs were expanded to passage 3 (P3) prior to encapsulation in hydrogels. Both NC and CPAC hydrogels, each containing 5×106 cells, were preconditioned in osteogenic medium for 3 days before implantation into the calvarial defects. The surgical site was subsequently closed via suturing. Following an 8-week recovery period, the rats were euthanized for analysis. Skull specimens were initially subjected to micro-computed tomography (micro-CT) scanning, followed by decalcification in 10% ethylenediaminetetraacetic acid (EDTA, pH 7.4) for 2 months in preparation for immunohistochemical evaluation.

Quantification of micro-CT
Micro-CT Scanning and Analysis: Upon euthanasia, rat calvarial bones were isolated and fixed in 4% paraformaldehyde. Fixed samples were scanned using a Micro-CT system (Quantum GX2, Perkinelmer AG) under the following conditions: 70 kV for 140 ms with a resolution setting of 25 μm and 1024×1024 pixels. Three-dimensional reconstruction and subsequent data analysis of the defect region were performed to quantify bone regeneration. Key outcomes included bone volume (BV), bone volume/tissue volume (BV/TV), and bone surface/tissue volume (BS/TV).

Immunohistochemical staining
The skull specimens were first fixed in 4% paraformaldehyde for 48 h to ensure proper preservation of tissue morphology and antigens. Following fixation, the samples underwent mild decalcification with 10% EDTA solution (pH 7.4, 25 °C) for 6 weeks. Tissue dehydration was performed through a graded ethanol series (30–100%), with subsequent xylene clearing and paraffin embedding. Serial sections (4 μm thickness) were prepared for histological evaluation.
Tissue sections were subjected to H&E and Masson’s trichrome staining according to standard protocols, followed by coverslipping with mounting medium. For immunohistochemical detection, sections were incubated overnight at 4 °C with primary antibodies targeting osteocalcin (OCN; 1:100 dilution, DF12303, Affinity). Following primary antibody incubation, sections were treated with HRP-conjugated secondary antibody (room temperature, 30 min) and developed with diaminobenzidine substrate (10 min, room temperature). All stained sections were permanently mounted for microscopic evaluation.

Theoretical analysis of CPAC hydrogel deformation
Using elasticity theory, we analyzed how the hydrogel deforms in response to changes in the radius of the medium ball. Let the ball initially have radius R0, which then changes to R. Due to strong interactions at the interface, the surrounding hydrogel is perturbed accordingly. As a continuous elastic material, the hydrogel’s deformation can be described in spherical coordinates by the displacement components ur, uθ, uφ. Because of the spherical symmetry, only the radial displacement is nonzero, giving uθ=0 and uφ=0.
According to elasticity theory, the equilibrium is expressed as:
1 ∂ (r2Eu) = const (1)
r2 ∂r r
where E is the Young’s modulus. This system is subject to two boundary conditions: (1) At infinity (r→∞), the deformation vanishes, i.e., u→0; (2) At the ball’s surface (r=R0), the radial displacement equals the radius change, i.e., u=R-R0. Solving Eq. (1) under these conditions, we can obtain the expression for the radial displacement.
Generally, the Young’s modulus is assumed to be a fixed value (E=E0), then the solution of Eq. (1) yields:
u = (R-R0) R0^2 / r^2 (2)
The above equation shows that the hydrogel deforms radially around the ball, with greater deformation arising from larger radius changes.
For cases like the experiments conducted here, E is not fixed. When the hydrogel deforms, its local properties, such as polymer structures and interactions, change, which in turn alters E. The relationship between E and hydrogel deformation may be complex, but in the small deformation limit, we assume E is proportional to the deformation. Therefore, E can be approximated as a linear function, E=E0+kur, where E0 is the Young’s modulus of the undeformed hydrogel, and k is a constant that characterizes the dependence of E on ur. Solving Eq. (1) with this assumption yields:
E = 1/2 E0 + 1/2 sqrt(E0^2 + 4kbr^-2) (3)
where b=R0^2 k (R-R0)^2 + E0 (R-R0). Note that r≥R0 in the above equation. This expression shows that the Young’s modulus varies radially around the medium ball. As r decreases, E increases, meaning the Young’s modulus is greater closer to the ball’s surface. Equation (3) matches well with our experimental data, Supplementary Fig. 11a. By fitting the experimental data, we determine E0=270.26 Pa, and k= -21.26 Pa/μm or -2.13×10^7 N/m^3.
Solving Eq. (1) also provides the analytical solution for the radial displacement:
ur = -1/2k (E0 + sqrt(E0^2 + 4kbr^-2)) (4)
Equation (4) shows that the hydrogel deforms radially around the ball, with greater deformation arising from larger radius changes (Supplementary Fig. 11b).

Methods for ALP inhibitor use
ALP inhibitor (Beyotime, Cat# ST019) was prepared as a 10 mM stock solution using sterile 1× PBS. For experiments, the inhibitor was added to the culture medium at a final concentration of 0.1 mM and applied for 72 h. For the control group, an equal volume of PBS was added.

Methods for ALP activity quantification and the CCK-8 assay
The ALP activity in the culture medium was quantified using a commercial colorimetric assay kit (Beyotime, Cat# P0321S) based on the hydrolysis of p-nitrophenyl phosphate (pNPP). Specifically, after the designated treatments, the culture medium was collected and centrifuged to remove debris. Then, 50 µL of the supernatant was mixed with 50 µL of the pNPP substrate working solution in a 96-well plate and incubated in the dark at 37 °C for 30 min. The enzymatic reaction was terminated by adding the stop solution provided in the kit. The absorbance of the yellow reaction product, p-nitrophenol, was immediately measured at 405 nm using a microplate reader. A standard curve was established using p-nitrophenol standards, and the ALP activity was calculated for each sample based on the absorbance readings. All assays were performed in triplicate.
Cell viability was assessed using the Cell Counting Kit-8 (CCK-8; Beyotime, Cat# C0038). Following the designated treatments, 100 µL of the CCK-8 reagent was directly added to each well (900 µL culture medium with one hydrogel). The plate was then incubated at 37 °C in a humidified incubator with 5% CO2 for 2 h. After incubation, the absorbance of each well was measured at a wavelength of 450 nm using a microplate reader. Wells containing culture medium and CCK-8 reagent without cells were used as blanks for background subtraction. Cell viability was calculated as a percentage relative to the untreated cells (set as 100%), using the following formula:
Viability (%) = (As - Ab) / (Ac - Ab) × 100% (5)
where As is the absorbance of the sample (treated cells), Ac is the absorbance of the control (untreated cells), and Ab is the absorbance of the blank. All experiments were performed with three independent replicates. Day 0 indicates the viability of MSCs before encapsulation, while day 3 represents the viability after MSCs were loaded in NC and CPAC hydrogels for 3 days.

Methods for ALP activity quantification
The alkaline phosphatase (ALP) activity in the culture medium was quantified using a commercial colorimetric assay kit (Beyotime, cat# P0321S) based on the hydrolysis of p-nitrophenyl phosphate (pNPP). The specific procedure was as follows: After the designated treatments, the culture medium was collected and centrifuged. Then, 50 µL of the supernatant was mixed with 50 µL of the pNPP substrate working solution in a 96-well plate and incubated in the dark at 37 °C for 30 min. The enzymatic reaction was terminated by adding the stop solution provided in the kit. The absorbance of the yellow reaction product, p-nitrophenol, was immediately measured at 405 nm using a microplate reader. A standard curve was generated using p-nitrophenol standards, and the ALP activity was calculated for each sample based on the absorbance readings. All assays were performed in triplicate.

Methods for EdU incorporation assays and live/dead staining
The alkaline phosphatase (ALP) activity in the culture medium was quantified using a commercial colorimetric assay kit (Beyotime, cat# P0321S) based on the hydrolysis of p-nitrophenyl phosphate (pNPP). The specific procedure was as follows: After the designated treatments, the culture medium was collected and centrifuged. Then, 50 µL of the supernatant was mixed with 50 µL of the pNPP substrate working solution in a 96-well plate and incubated in the dark at 37 °C for 30 min. The enzymatic reaction was terminated by adding the stop solution provided in the kit. The absorbance of the yellow reaction product, p-nitrophenol, was immediately measured at 405 nm using a microplate reader. A standard curve was generated using p-nitrophenol standards, and the ALP activity was calculated for each sample based on the absorbance readings. All assays were performed in triplicate.
For live/dead staining, a staining solution was prepared in 1× PBS containing Calcein-AM and propidium iodide (PI), each at a final concentration of 1 µM. The hydrogels cultured for 3 days in a 24‑well plate were carefully removed and transferred to a new plate, and 1 mL of the prepared staining solution was added to each well. The plate was then incubated at 37 °C in the dark for 30 min. Following incubation, the hydrogels were washed three times with 1× PBS. Finally, imaging was performed using a confocal laser scanning microscope with detection channels set to 488 nm and 594 nm.

Methods for in vivo implantation model and sample processing for ALP, nanoindentation and mass quantification
According to the previously described method for creating critical-size bone defects in rat calvaria, hydrogels loaded with rMSCs were implanted into the calvarial defect areas. The implanted hydrogels were retrieved at different time points: before implantation (0 h), and at 24 h and 72 h after implantation. The retrieved hydrogels were added to 1 mL of 1× PBS, thoroughly homogenized using a tissue grinder, and then centrifuged. The alkaline phosphatase (ALP) activity in the hydrogel was quantified using a commercial colorimetric assay kit (Beyotime, cat# P0321S) based on the hydrolysis of p-nitrophenyl phosphate (pNPP). The specific procedure was as follows: 50 µL of the supernatant was mixed with 50 µL of the pNPP substrate working solution in a 96-well plate and incubated in the dark at 37 °C for 30 min. The enzymatic reaction was terminated by adding the stop solution provided in the kit. The absorbance of the yellow reaction product, p-nitrophenol, was immediately measured at 405 nm using a microplate reader. A standard curve was generated using p-nitrophenol standards, and the ALP activity was calculated for each sample based on the absorbance readings. All assays were performed in triplicate.
For nanoindentation testing, the extracted hydrogels were fixed onto glass slides. An 8×8 matrix of points, with each point spaced 50 μm apart, was randomly selected on the hydrogel surface. The effective Young’s modulus at each point in the matrix was measured using a nanoindenter.
For the degradation experiment, the implanted hydrogels were retrieved at different time points, freeze-dried, and weighed (recorded as Wx). Taking the mass of the freeze-dried hydrogel sample before implantation as W0, the degradation rate of the hydrogel was calculated as follows:
D = Wx / W0 × 100% (6)
All samples in each group were tested in three independent experiments to ensure reproducibility.

Method for alkaline phosphatase color development kit
After fixation with paraformaldehyde, the hydrogel was cryosectioned into 8 μm thin slices. The resulting slices were washed three times with PBS. A working solution was prepared according to the manufacturer’s instructions of the BCIP/NBT Alkaline Phosphatase Color Development Kit (Beyotime, cat# C3206). The sections were fully covered with the working solution and incubated at room temperature in the dark for 30 min. The color development reaction was terminated by washing twice, and images were captured using an inverted microscope.

Statistics and reproducibility
Experiments included triplicate samples (n=3), and results are shown as mean ± standard deviation. Statistical significance was determined using GraphPad Prism (version 9.0.0) and Origin (version 2018). Comparisons between two groups were analyzed by an independent t-test, while comparisons across more than two groups were analyzed by one-way ANOVA with Tukey’s multiple comparison test. In all analyses, statistical significance was defined as p<0.05.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "This study presents a novel hydrogel material—cell-programmed adaptive contraction (CPAC) hydrogel—that can respond to alkaline phosphatase (ALP) signals secreted by cells, leading to contraction of the embedded microgels within the hydrogel and resulting in local stiffness enhancement, ultimately forming a heterogeneous hydrogel structure. While the CPAC hydrogel demonstrates innovative bio-responsiveness and potential for bone repair, there are several shortcomings and potential gaps in the study. Therefore, we do not recommend the article for publication in Nat. Commun. at this stage.\n1.The design of the CPAC hydrogel relies solely on ALP as the key enzymatic marker for regulation. However, it remains unclear whether other enzymes within the phosphatase family could also influence this process, potentially leading to non-specific reactions. Additionally, the dynamics of ALP release during MSC osteogenic differentiation are not well defined.\n2.The CPAC hydrogel mimics the mechanical heterogeneity of natural ECM through the contraction of microgels and local stiffness enhancement. However, whether the hydrogel can achieve precise and controllable gradient variations in local stiffness remains a concern. In the complex in vivo biological environment, the hydrogel may be influenced by external factors such as temperature and pH, leading to unpredictable fluctuations in its mechanical properties. This could affect the cells' response to the material, particularly if the local stiffness of the hydrogel becomes uneven, potentially causing misalignment in stem cell differentiation.\n3.In this study, the CPAC hydrogel responds to ALP and alters its structure within a short period of time, but its stability in vivo or in vivo or in vivo or in vivo or in long-term experiments has not been sufficiently validated.\n4.Although the hydrogel promotes MSC osteogenic differentiation, the effects on cell proliferation and survival have not been adequately explored. The contraction of the microgels may lead to changes in the intercellular space, further affecting cell expansion and proliferation. If the hydrogel does not provide sufficient support or space during differentiation, it may limit the functional repair of the cells.\n5.The authors indicate that the expression levels of miR-124, miR-214, and miR-26a are elevated in the CPAC hydrogel group, and this is significantly correlated with a decrease in the expression of EZH2 and its downstream target H3K27me3. However, the lack of validation through anti-miR treatment raises concerns about the reliability of this correlation.\n6.In Figure 5g, the immunofluorescence images for the control group are not shown.\n7.In Figures 3b, 3c, and 5c, g, a significant proportion of suspected overexposed areas can be observed. This raises concerns about the accuracy of the semi-quantitative analysis based on fluorescence intensity.\n8.The regulatory effect of H3K27me3 on osteogenic-related genes has not been validated through relevant experiments.\n9.In the in vivo experiments, only data from 8 weeks after hydrogel implantation are provided. This duration does not adequately reflect the effects of local network stiffening in promoting MSC osteogenic differentiation.\n10.The size changes of CPAC hydrogel in a 1 U/mL ALP solution have been studied; however, the ALP concentration during in vivo or bone injury healing is unknown, and it remains unclear whether it would be sufficient to trigger the hydrogel's adaptive remodeling. Additionally, the hydrogel's contraction under in vivo conditions has not been characterized.", "reviewer_2": "The authors present a study focused on the development of a hydrogel system that mimics the dynamically evolving and heterogeneous extracellular matrix (ECM) environment during tissue development. Specifically, they engineer a cell-programmed adaptive contraction (CPAC) hydrogel containing ALP-responsive microgels that contract in response to alkaline phosphatase (ALP) activity. This induces overall hydrogel network contraction and local matrix stiffening, thereby promoting osteogenic differentiation of mesenchymal stromal cells (MSCs). The authors provide and test a hypothesis for the underlying mechanotransduction mechanism including upregulation of microRNA and demonstrate the translational potential of their system in a rat critical-size bone defect model, where they observed enhanced bone formation.\nThe study presents a dynamic hydrogel platform that responds to cell-released enzymes and modulates stem cell fate through dynamically changing the mechanical signals. The concept is interesting, and the authors have sought to perform both mechanistic investigation and in vivo validation. While the results are promising, additional experiments are required to support their main claims such as ‘‘ALP-induced gel contraction’’, ‘‘local stiffening’’ and ‘‘mechanical heterogeneity’’. Moreover, several aspects of the study require clarification, statistical information or further detail to strengthen its overall scientific rigor, reproducibility and impact.\nMajor comments:\n• Novelty and significance: It would be helpful if the authors can explain more why this work is novel and their findings are significant for a broad impact.\n• Study design: While this study probed an interesting concept, their main claims seem to have several gaps, across materials, in vitro- and in vivo models.\no From the materials aspect, the reliability relies on a thorough characterization of the mechanical properties and heterogeneity of their biomaterials.\no The range of mechanical stiffness such as magnitude, anisotropy seems unclear.\no From the cell biology perspective, whether the readouts are solely due to ALP, and cell-secreted ALP, or not, may require more rigorous investigation by setting up controls and perturbation experiments. It is known that cells also secrete other factors that block ALP acitivity.\no From the in vivo aspect, the complexity further increases in terms of proteins and cells in involved. The authors are recommended to add control experiments to demonstrate the enhanced bone formation is not by the gels along, nor other matrix factors, but exactly by their claimed mechanism.\n• Abstract clarity: The abstract does not clearly convey how the hydrogel system works. Key elements, such as the main components of the hydrogel as well as the control groups used in the study, are not mentioned but are critical for interpreting the results.\n• Introduction:\no The introduction would benefit from a comparison to naturally derived hydrogels such as collagen. How does the contractile behaviour of the presented system compare, and what advantages does it offer over existing ECM-mimetic materials?\no The exact composition of the hydrogel remains unclear in the introduction and Figure 1. Including a schematic that clearly explains the composition and assembly of the hydrogel from microgels and other components for all experimental hydrogel groups would greatly improve clarity. Also pay attention to complete & matching legends for the schematics.\n• Figure 2g: It would be helpful to include confocal images of the hydrogels prior to ALP treatment to demonstrate the changes induced by contraction.\n• Figure 2h: Please indicate which statistical test was used and annotate significance levels directly on the graph.\n• You claim that “CPAC hydrogel can better facilitate stellate spreading”. Please explain where this can be seen or provide more morphological data to support it.\n• Figures 3b/c: The CPAC hydrogels appear to have a higher cell density compared to NC controls. Is this a localized effect due to contraction, or does the material promote proliferation? Higher local cell density itself may influence osteogenic differentiation.\nRelated to this, can you comment on the macroscopic effect of ALP-mediated contraction on the entire hydrogel, does the hydrogel size change?\n• Can you specify how the “high-modulus N,N-dimethylacrylamide (DMAA) microgels” compare to the previous control microgels (Phos-HEA) in terms of mechanical properties?\n• The claim “NCLS hydrogels showed elevated osteogenic marker expression relative to NC hydrogels” is not evident from the data shown, please justify or explain more.\n• Figure 4: Please provide rationale for the selection of mechanotransduction markers (pFAK, pMyosin IIa, Paxillin).\n• Figure 4: Why are the microgels not visible in the confocal images like in Figures 3 & 5? The figure caption makes it a bit unclear, if cells were seeded on top of hydrogels rather than embedded.\n• Figure 4b: Collagen I appears to be predominantly intracellular. Is there an explanation for this observation?\n• The hydrogel is described as responsive to ALP secreted by MSCs, yet the immunofluorescence images show primarily intracellular ALP. Please clarify how this observation aligns with the proposed mechanism of hydrogel contraction, which relies on extracellular ALP activity.\n• A 3-day culture period is relatively short for observing osteogenic differentiation. Have longer culture periods been considered?\n• Materials and Methods in SI: Several important methodological aspects require clarification:\no How was the microgel shrinkage quantified? Was it done manually, and how many replicates were analysed?\no Please provide more details about your hydrogel preparation from your individual components including precise concentrations and volume ratios. Please also specify the light source, wavelenght and intensity (also for photorheology).\no What were the measurement parameters used for rheological analysis?\no In your nanoindentation measurements, was stiffness assessed in a single 50 µm × 50 µm region or across multiple areas? Are the results shown representative?\no What was the cell seeding density, and what was the source of the hMSCs?\no In the in vivo experiments, was there a contralateral defect used as an internal control in each animal as indicated in Figure 6a?\no Please describe the analysis of micro-CT data in more detail.\no You write that your skull specimens were decalcified for 2 months before fixation, whereas fixation is generally performed before decalcification to preserve tissue, cell and protein integrity. Could you please explain your choice of procedure and whether that could affect the staining results?\no The section “Immunohistochemical staining” describes the use of an anti-ALP antibody, whereas the data in Figure 6 shows OCN. Please correct.\no The section “3D cell culture within hydrogels” seems incomplete, especially regarding the use of the EZH2 inhibitor (starting from line 163).\nMinor comments:\n• The sequence of figure citations in the text is partially incorrect, for example, Supplementary Figure S11 is cited before S10. Also, the reference to Figure S11 is unclear, is it pertaining to stiffness (text) or ALP concentration (figure)?\n• Figure S12: Correct graph labellings “i, ii, iii, iv, v, vi” and their citations in the text.\n• The manuscript contains several typos that should be corrected, for example line 68 “promote the of commitment of stem cells”, line 102 “CAPC” instead of “CPAC” or line 272 “this modulus remained larged unchanged”.\n• Figure S9: The panel c described in the figure caption does not exist. Additionally, please clarify the meaning of the red and blue vertical lines in the existing panel c. Also the mention of “half of the peak stress relaxation times (τ1/2 = 150 ± 10 s)” doesn’t match the data shown in Figure S9c.\n• Figure 2f: Please label the images.", "reviewer_3": "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.", "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": "Thank you for this insightful and constructive comments. Regarding the specificity toward ALP, we do agree that other phosphatases (e.g., acid phosphatases) could potentially lead to the cleave of phosphate groups in our systems. Our design focused on ALP because of its well-established role as an early and strongly upregulated marker of osteogenic differentiation. Therefore, to test whether other phosphatases could also influence this system, we performed two control experiments: (i) CPAC hydrogels treated with ALP and an ALP inhibitor did not develop heterogeneous structures, and their storage modulus remained unchanged (Figure S12a, c), highlighting the dominant role of ALP in this heterogenization process. (ii) CPAC hydrogels treated with acid phosphatase (ACP) also failed to induce localized stiffening (Figure S12b), likely due to its limited activity under the neutral buffered conditions used. These results confirm that ALP is the primary enzyme driving the contraction-induced stiffening process in CPAC hydrogels.\nMoreover, to validate the dynamics of ALP release during MSC differentiation, we quantified ALP levels in CPAC and NC hydrogels over 0–168 hours (Figure S12d). Both groups showed a gradual increase in ALP during culture, but the levels in CPAC hydrogels rose more steeply, reaching ~10 U/mL at 168 h, compared to ~5 U/mL in NC hydrogels. This sustained elevation indicates that CPAC hydrogels not only permit ALP secretion but also amplify it through the feedback between microgel contraction, local stiffening, and osteogenic activation.\nThe updated paragraph in the section of “ALP-mediated microgel contraction and local stiffening generates the mechanically heterogenous structures of CPAC hydrogel ” now reads:\n…In contrast, no such changes in local stiffness were observed in the NC hydrogels (Figure 2i, ii; S10). Moreover, the theoretical analysis of CPAC hydrogel deformation also confirmed the formation of the heterogeneous structure within the hydrogel, with stiffness increasing radially near the Phos-HPMA microgel surface (Figure S11). To validate the specific response of CPAC hydrogel to ALP, identical sets of CPAC hydrogel were treated with ALP/ALP inhibitor and acid phosphatase, respectively. It was observed that compared to the ALP-treated group, the internal structures of these two groups did not exhibit significant changes (Figure S12a-c), indicating that ALP is the primary factor responsible for the formation of the heterogeneous structure in CPAC hydrogel. We further examined the responsiveness of the hydrogels to cell-secreted ALP by encapsulating osteogenically-induced MSCs in both hydrogels. As shown in Figure S12d, ALP levels in both groups gradually increased during culture, but CPAC hydrogels exhibited a markedly higher and sustained ALP release. This elevated ALP secretion strongly correlates with the heterogeneous stiffening pattern observed in CPAC hydrogels…\nFigure S12. Heatmaps displaying the distribution of local effective Young's modulus in the CPAC hydrogels after 72 hours of treatment with ALP inhibitor (a), ACP (b), and the storage modulus of CPAC hydrogel with different treat treatments for 3days (c), (d), The ALP concentration corresponding to two hydrogels loaded with MSCs at various culture time points. Data are presented as mean values ± SD, n = 3 independent hydrogels, ns, not significant, **p < 0.01, ***p < 0.001, ****p < 0.0001 (one-way ANOVA).\n2. The CPAC hydrogel mimics the mechanical heterogeneity of natural ECM through the contraction of microgels and local stiffness enhancement. However, whether the hydrogel can achieve precise and controllable gradient variations in local stiffness remains a concern. In the complex in vivo biological environment, the hydrogel may be influenced by external factors such as temperature and pH, leading to unpredictable fluctuations in its mechanical properties. This could affect the cells' response to the material, particularly if the local stiffness of the hydrogel becomes uneven, potentially causing misalignment in stem cell differentiation.\nAuthor Response:\nWe thank the reviewer for raising these important concerns regarding the precise control over local stiffness gradients and the influence of external factors like temperature and pH in the complex in vivo environment. Below, we address both of these aspects based on additional experiments:\n1) Precise and controllable gradient variations in local stiffness:\nWe agree that achieving precise and controllable gradients in local stiffness is a key consideration for biomimetic materials. In our current design, the mechanical heterogeneity is dynamically evolved through a cell-driven feedback mechanism, rather than being pre-programmed with static, absolute precision. The contraction of microgels and local stiffness enhancement are primarily triggered by ALP secreted by MSCs themselves during early osteogenic differentiation, leading to self-regulated structural heterogeneity. This design more accurately mimics the dynamic and reciprocal cell-matrix interactions found in natural ECM, where the matrix evolves in response to cellular activity. To characterize this dynamic process and assess its controllability, we performed the following analyses:\ni) Microscopic mechanical mapping: Nanoindentation mapping (Figure S16a) indicates that, while a degree of heterogeneity is present, as intended to mimic natural ECM, the stiffness variation occurs relatively uniformly without large, unpredictable fluctuations that would lead to detrimental misalignment in stem cell differentiation. This suggests that the cell-instructed heterogeneity may be more physiologically relevant than a perfectly engineered but static gradient.\nii) Macroscopic rheological analysis: Bulk storage modulus (G’) of the hydrogel was monitored over time during osteogenic differentiation. The data (Figure S16b) shows a consistent increase in overall stiffness, confirming that the CPAC hydrogel undergoes predictable and controlled temporal stiffening in response to cellular activity.\niii) Theoretical analysis of CPAC hydrogel deformation: To further understand the controllable gradient variations in local stiffness, we applied elasticity theory [1] to model how the CPAC hydrogel deforms in response to the contraction of Phos-HPMA microgels. This model showed that local stiffness varies radially around the microgel, with stiffer regions closer to the microgel surface, and this relationship closely matches our experimental data. This confirms that the CPAC hydrogel can precisely control stiffness gradients locally while maintaining uniform bulk mechanical properties.\nSpecifically, let the Phos-HPMA microgel initially have radius 𝑅 , which then changes to 0 𝑅 . Due to strong interactions at the interface, the surrounding hydrogel is perturbed accordingly. As a continuous elastic material, the hydrogel’s deformation can be described in spherical coordinates by the displacement components (𝑢 ,𝑢 ,𝑢 ) . Because of the 𝑟 𝜃 𝜑 spherical symmetry, only the radial displacement is nonzero, giving 𝑢 = 0 and 𝑢 = 0. 𝜃 𝜑 According to elasticity theory, the equilibrium is expressed as:\n1 𝜕 (𝑟2𝐸𝑢 ) = 𝑐𝑜𝑛𝑠𝑡. (1)\n𝑟2𝜕𝑟 𝑟\nwhere 𝐸 is the Young’s modulus. This system is subject to two boundary conditions: (1) At the infinity (𝑟 → ∞ ), the deformation vanishes, i.e., 𝑢 = 0 ; (2) At the Phos-HPMA ∞ microgel’s surface (𝑟 = 𝑅 ), the radial displacement equals the radius change, i.e., 𝑢 = 0 0 𝑅 −𝑅 . Solving Eq. (1) under these conditions, we can obtain the expression of the radial 0 displacement.\nGenerally, the Young’s modulus is assumed to be a fixed value (𝐸 = 𝐸 ), then the solution 0 of Eq. (1) yields:\n𝑢 = (𝑅−𝑅0 )𝑅 0 2 . (2)\n𝑟 𝑟2\nThe above equation shows that the hydrogel deforms radially around the Phos-HPMA microgel, with greater deformation arising from larger radius changes.\nFor cases like the experiments conducted here, 𝐸 is not fixed. When the hydrogel deforms, its local properties, such as polymer structures and interactions, change, which in turn alters 𝐸. The relationship between 𝐸 and hydrogel deformation may be complex, but in the small deformation limit, we assume 𝐸 is proportional to the deformation. Therefore, 𝐸 can be approximated as a linear function,𝐸 = 𝐸 +𝑘𝑢 , where 𝐸 is the Young’s modulus of the 0 𝑟 0 undeformed hydrogel, and 𝑘 is a constant that characterizes the dependence of 𝐸 on 𝑢 . 𝑟\nSolving Eq. (1) with this assumption yields:\n𝐸 = 1 𝐸 + 1 √𝐸2 +4𝑘𝑏𝑟−2, (3)\n0 0 2 2\nwhere 𝑏 = 𝑅2[𝑘(𝑅 −𝑅 )2 +𝐸 (𝑅 −𝑅 )]. Note that 𝑟 ≥ 𝑅 in the above equation. This 0 0 0 0 0 expression shows the Young’s modulus varies radially around the medium Phos-HPMA microgel. As 𝑟 decreases, 𝐸 increases, meaning the Young’s modulus is greater closer to the Phos-HPMA microgel’s surface. Eq. (3) matches well with our experimental data, Figure S11a. By fitting the experimental data, we determine 𝐸 = 270.26 𝑃𝑎 , and 𝑘 = 0 −21.26 𝑃𝑎/𝜇𝑚 or −2.13×107 𝑁/𝑚3.\nSolving Eq. (1) also provides the analytical solution for the radial displacement:\n𝑢 = 1 (−𝐸 +√𝐸2 +4𝑘𝑏𝑟−2), (4)\n𝑟 0 0 2𝑘\nEquation (4) shows that the hydrogel deforms radially around the Phos-HPMA microgel, with greater deformation arising from larger radius changes (Figure S11b).\n2) Potential influence of in vivo factors and hydrogel stability:\nRegarding the potential effects of temperature and pH fluctuations in the in vivo environment, we conducted stability tests to assess the hydrogel’s response to changes in these parameters. The CPAC hydrogels were incubated in buffers with pH ranging from 7.2 to 7.8 and at different temperatures from 33 to 39 °C. The hydrogels maintained their structural integrity within the physiological range of pH (7.2-7.8) and temperature (33- 39 °C), showing no significant degradation in mechanical properties or unpredictable fluctuations in stiffness (Figure S16c-f). While the enzymatic activity of ALP can be modulated by environmental pH and temperature, our results indicate that slight variations within physiological ranges do not induce measurable changes in the hydrogel’s structure or its ability to remodel over a relatively long culture period. These findings indicate that the CPAC hydrogel network, once formed, remains largely stable against transient fluctuations in pH and temperature within the physiological range without compromising its mechanical integrity or functionality.\nThe corresponding paragraphs in the section of “ALP-mediated microgel contraction and local stiffening generates the mechanically heterogenous structures of CPAC hydrogel” now reads:\n…In contrast, no such changes in local stiffness were observed in the NC hydrogels (Figure 2i, ii; S10). Moreover, the theoretical analysis of CPAC hydrogel deformation also confirmed the formation of the heterogeneous structure within the hydrogel, with stiffness increasing radially near the Phos-HPMA microgel surface (Figure S11)…\n…Furthermore, to validate the stability of CPAC hydrogel under complex in vivo conditions, we incorporated variations in pH and temperature into the evaluation. As shown in Figure S16, CPAC hydrogels were able to maintain stable formation of heterogeneous structures within physiological ranges of pH (7.2–7.8) and temperature (33–39 °C) for 3 days, with no significant impact on the hydrogel structure.\nFigure S11. (a) Young’s modulus varies with the distance to the center of the Phos-HPMA microgel. The curve is fitted using Eq. (3) with an initial radius of 𝑅 = 170 𝜇𝑚 and a 0 shrunk radius of 𝑅 = 70 𝜇𝑚 aata fittin iillds thl aaraeltlr aaluls 𝐸 = 270.26 𝑃𝑎 0 and 𝑘 = −21.26 𝑃𝑎/𝜇𝑚 (b) Radial deformation of the hydrogel induced by the shrinking microgel. As the microgel contracts from its initial radius of 170 μm to a reduced radius of 70 μm, the surrounding hydrogel network undergoes stronger deformation, quantified by the radial displacement 𝑢 . The displacement is calculated according to Eq. (4).\n𝑟\nFigure S16. (a) Heatmaps displaying the distribution of local effective Young's modulus in the CPAC hydrogels after 0, 1, 3 days of culture with MSC encapsulation. (b) The change in storage modulus over time. (c) The effect of different temperatures on the formation of heterogeneous structures in CPAC hydrogels under pH = 7.4, and the in vitro. (d) The effect of different pH on the formation of heterogeneous structures in CPAC hydrogels under temperature = 37 °C in vitro. (e-f) The change in storage modulus of CPAC hydrogel with different treatment. Data are presented as mean values ± SD, n = 3 independent hydrogels, ns, not significant, ****p < 0.0001 (one-way ANOVA).\n3. In this study, the CPAC hydrogel responds to ALP and alters its structure within a short period of time, but its stability in vivo or in long-term experiments has not been sufficiently validated.\nAuthor Response:\nWe are grateful to the reviewer for this excellent suggestion. In response, we have conducted additional experiments to evaluate the stability of the CPAC hydrogel both in vitro and in vivo over a period of 28 days. The results show that the CPAC hydrogel retained over 30-40 % of its original mass (Figure S23), while preserving its heterogeneous structure without noticeable deterioration (Figure S24). These findings indicate that CPAC hydrogels are sufficiently stable to match the typical timescale of the initial phase during bone repair, while still allowing gradual remodeling by cells.\nThe corresponding paragraphs in the section of “MSC-laden CPAC hydrogels promote bone regeneration in an animal critical defect model” now reads:\nBefore evaluating the therapeutic efficacy of dynamic heterogeneous CPAC hydrogel in facilitating bone regeneration, we first assessed the long-term stability of the hydrogel both in vitro and in vivo. As shown in Figure S23, the hydrogel underwent gradual degradation over four weeks, consistent with the expected degradation trend of implantable materials. However, the heterogeneous structure in the remaining non-degraded regions remained stable (Figure S24). This indicates that the CPAC hydrogel can stably provide the heterogeneous structural extracellular microenvironment required by MSCs during bone defect repair. Moreover, to evaluate the therapeutic efficacy of dynamic heterogeneous CPAC hydrogel in facilitating bone regeneration, we employed a rat calvarial defect model…\nFigure S23. Degradation profiles of CPAC hydrogel in vitro and in vivo over 28 days.\nFigure S24. Heatmaps displaying the distribution of local effective Young's modulus in CPAC hydrogels after in vitro culture (a) and in vivo (b).\n4. Although the hydrogel promotes MSC osteogenic differentiation, the effects on cell proliferation and survival have not been adequately explored. The contraction of the microgels may lead to changes in the intercellular space, further affecting cell expansion and proliferation. If the hydrogel does not provide sufficient support or space during differentiation, it may limit the functional repair of the cells.\nAuthor Response:\nWe thank the reviewer for raising this important point. To assess whether microgel contraction affects MSC proliferation and survival, we performed EdU incorporation assays, live/dead staining, and CCK-8 assays in both NC and CPAC hydrogels. As shown in the Figure S17, MSCs in CPAC hydrogels exhibited significantly higher proliferation efficiency compared to those in NC hydrogels, while cell viability remained consistently high in both groups. These findings suggest that the heterogeneous structure generated by microgel contraction does not hinder proliferation; instead, it creates larger network gaps (low-modulus regions) that provide additional space and a more permissive environment for MSC expansion. Thus, CPAC hydrogels support both osteogenic differentiation and sustained MSC growth, which is essential for effective tissue repair.\nThe corresponding paragraphs in the section of “The cell-programmed local stiffening in CPAC hydrogel promotes osteogenic differentiation of MSCs” now reads:\nBefore examining osteogenic differentiation, we first evaluated whether CPAC hydrogel supports the proliferation and survival of encapsulated MSCs. Compared to the NC group, CPAC hydrogels exhibited a higher number of EdU-positive cells, indicating an increase in newly synthesized DNA (Figure S17a). Consistently, live/dead staining confirmed robust cell viability (Figure S17b), and CCK8 assays further verified significantly higher proliferation rates in the CPAC group relative to NC (Figure S17c). These results suggest that the heterogeneous structure formed during microgel contraction not only provides space for cell expansion but also creates a favorable microenvironment that sustains MSC proliferation. Furthermore, the results of Actin staining and the statistical analysis of cell spreading area for MSCs…\nFigure S17. Proliferation and survival of MSCs in NC and CPAC hydrogels. (a) EdU assay showing newly synthesized DNA, (b) live/dead staining, and (c) CCK-8 assay confirming enhanced proliferation in CPAC hydrogels.\n5. The authors indicate that the expression levels of miR-124, miR-214, and miR-26a are elevated in the CPAC hydrogel group, and this is significantly correlated with a decrease in the expression of EZH2 and its downstream target H3K27me3. However, the lack of validation through anti-miR treatment raises concerns about the reliability of this correlation.\nAuthor Response:\nWe thank the reviewer for raising this important concern regarding the mechanistic role of the identified miRNAs. We performed additional validation experiments using siRNA-based inhibitors of miR-124, miR-214, and miR-26a. Western blot analysis demonstrated that inhibition of these miRNAs led to a markedly increase in EZH2 and H3K27me3 pritein levels, while significantly reducing the expression of the osteogenic markers ALP and Col 1. These findings provide direct functional evidence that the miR–EZH2–H3K27me3 axis mediates the enhanced osteogenesis observed in CPAC hydrogels.\nThe corresponding paragraphs in the section of “Mechano-epigenetic regulation governs MSC osteogenesis in CPAC hydrogels” now reads:\n…to further validate the mechanistic link, we incorporated siRNA-based inhibitors of miR- 124, miR-214, and miR-26a into the CPAC hydrogel cultures. Western blot analysis demonstrated that the inhibition of these miRNAs markedly increased the protein levels of EZH2 and H3K27me3, while simultaneously reducing the expression of osteogenic markers ALP and Col 1 (Figure S22). These results demonstrate that the cell-programmed matrix heterogenization in CPAC hydrogel establishes an epigenetic landscape conducive to osteogenic differentiation through miRNA-mediated suppression of the EZH2— H3K27me3 axis.\nFigure S22. Western blot analysis showing that inhibition of miR-124, miR-214, and miR- 26a in CPAC hydrogels restores EZH2 and H3K27me3 expression while reducing ALP and Col 1, confirming the involvement of the miR–EZH2–H3K27me3 axis in MSC osteogenesis.\n6. In Figure 5g, the immunofluorescence images for the control group are not shown.\nAuthor Response:\nWe thank the reviewer for this insightful comment. The immunofluorescence images of the control group have been added into Figure 5g.\nRevised Figure 5g now reads:\nFigure 5. (g) Immunofluorescent staining of EZH2, H3K27me3, Col 1, and ALP in NC hydrogels, with and without EZH2 inhibitor treatment (scale bar = 75 μm).\n7. In Figures 3b, 3c, and 5c, g, a significant proportion of suspected overexposed areas can be observed. This raises concerns about the accuracy of the semi-quantitative analysis based on fluorescence intensity.\nAuthor Response:\nWe thank the reviewer for pointing out this critical issue. The suspected overexposure in the original images was a technical artifact caused by suboptimal imaging settings during confocal acquisition, which were unintentionally set too high. This led to signal saturation in some regions, despite the underlying biological differences being genuine. To address this concern, we have now carefully re-optimized our imaging workflow by lowering both the antibody concentration (from 1:500 to 1:1000) and the laser excitation intensity to ensure that signals remain within the linear detection range. We then re-collected all immunofluorescence images and repeated the semi-quantitative fluorescence intensity analysis. The updated images and corrected quantifications are now included in the revised manuscript. Importantly, the new data fully support our original conclusions while avoiding the technical issue of signal saturation.\nRevised Figures 3 and 5 now read:\nFigure 3. Immunofluorescent staining of (b) Col 1 and (c) ALP in NC and CPAC hydrogels, respectively (scale bar = 75 μm). (d) Quantification of Col 1 and ALP immunofluorescence intensity in the two hydrogels (n = 10).\nFigure 5. (c-d) Immunofluorescent staining and statistical analysis of EZH2 in different hydrogel groups (scale bar = 75 μm; n = 10). (g) Immunofluorescent staining of EZH2, H3K27me3, Col 1, and ALP in NC hydrogels before and after EZH2 inhibitor treatment (scale bar = 75 μm). (h) Quantification of immunofluorescence for Col 1 and ALP in NC hydrogels before and after EZH2 inhibitor treatment (n = 10).\n8. The regulatory effect of H3K27me3 on osteogenic-related genes has not been validated through relevant experiments.\nAuthor Response:\nWe thank the reviewer for this constructive comment. Methylation of H3K27 serves as a critical epigenetic switch determining the lineage commitment of MSCs, with elevated H3K27me3 levels on osteogenic gene promoters reported to suppress osteogenic differentiation by silencing key transcription factors such as RUNX2 [2-7]. To directly validate this regulatory mechanism in our system, we performed a ChIP-qPCR assay targeting the RUNX2 promoter. The results showed that H3K27me3 was enriched specifically at the upstream promoter region of RUNX2 rather than at unrelated genomic regions (Figure R1a). In MSCs cultured within CPAC hydrogels, both immunofluorescence and Western blot analyses confirmed a significant decrease in global H3K27me3 levels (Figures R1b and R1c), while RT-qPCR demonstrated a marked upregulation of RUNX2 transcription (Figure R1d). Consistently, immunofluorescence staining further indicated increased expression of the osteogenic markers Col1 and ALP in MSCs of the CPAC group (Figure R1e). In summary, these findings demonstrate that H3K27me3 directly represses RUNX2 transcription, thereby limiting osteogenic differentiation, and that the CPAC hydrogel microenvironment alleviates this repression to promote osteogenesis.\nFigure. R1 (a) ChIP-qPCR analysis showing the occupancy of H3K27me3 at the promoter site of RUNX2, (b) Representative H3K27me3 staining image of two hydrogels (scale bar = 75 μm). (c) Western blot analysis of H3K27me3 in two hydrogel groups, (d) RT-qPCR analysis of relative RUNX2 expression in two hydrogel groups, (e) Representative Col 1 I and ALP staining image of two hydrogel groups (scale bar = 75 μm).\nThe corresponding paragraphs in the section of “Mechano-epigenetic regulation governs MSC osteogenesis in CPAC hydrogels” now reads:\n…This upregulation correlated with a marked reduction in the expression of EZH2 and its downstream target, H3K27me3, in CPAC hydrogels (Figure 5b). Immunofluorescence analysis further confirmed these findings, showing diminished EZH2 and H3K27me3 fluorescence intensity in CPAC hydrogels compared to NC hydrogels (Figure 5c-f). ChIP- qPCR then demonstrated that H3K27me3 was enriched at the RUNX2 promoter (Figure 5i), and This reduction was accompanied by increased RUNX2 transcription (Figure 3a) and enhanced protein expression of Col 1 and ALP (Figure 5g).\nSI materials and methods:\nChromatin immunoprecipitation (ChIP) and ChIP-qPCR\nCells were fixed with 1% formaldehyde and subsequently quenched with glycine. After washing with PBS, the fixed samples were suspended in cell lysis buffer (5 mM PIPES-KCl pH 8.0, 85 mM KCl, 0.5% NP40), and collected nuclei were stored at −80 °C as nuclear pellets or nuclear lysates dissolved in nuclei lysis buffer (50 mM Tris-HCl pH8.0, 10 mM EDTA, 1% SDS). For H3K27me3 ChIP, lysates were thawed and sonicated with Sonifier (BRANSON) to obtain chromatin fragments of 300–1,000 bp, and 10-fold-diluted with ChIP dilution buffer (16.7 mM Tris-HCl pH8.0, 1.2 mM EDTA, 0.01% SDS, 1.1% Triron100×, 167 mM NaCl). After double-dilution with a sonication buffer (90 mM Hepes pH 7.9, 220 mM NaCl, 10 mM EDTA, 1% NP 40, 0.2% sodium deoxycholate, 0.2% SDS), nuclei were homogenized with a Bioruptor (Tosho denki). After washing, a complex of nuclear proteins/DNAs and antibodies against H3K27me3 (Proteintech; 39157; 5ug used) was retrieved with Protein A- and Protein G-Dynabeads (Life Technologies). After the cross-linking was reversed, chromatin fragments were treated with RNase A and proteinase K. DNA was purified with phenol-chloroform extraction or Ampure XP (Beckman Coulter).\nIn the ChIP-qPCR analyses, the values from the immunoprecipitated samples were normalized to input DNA. Primer sequences are listed in Table S2, Supporting Information.\nTable S2: Primers sequence for quantitative polymerase chain reaction (miRNA and ChIP)\nHuean-eiR-124-3-1-F CACCGCGCGGTGAATGCCAAGAATG\nHuean-eiR-124-3-1-R AAACCATTCTTGGCATTCACCGCGC\nHuean-eiR-214-F CTGGCTGGACAGAGTTGTCAT\nHuean-eiR-214-R GCTGTACAGGTGAGCGGATG\nHuean-eiR26a-F GTGGCCTCGTTCAAGTAATCCA\nHuean-eiR26a-R CCCCGTGCAAGTAACCAAGAA\nChIP-RUNX2-F AGGCCTTACCACAAGCCTTT\nChIP-RUNX2-R AGAAAGTTTGCACCGCACTT\nCont-F AAAGCTCCATAAGGATACCCTGTATGAT\nCont-R CATGTTGGAGATCATAAAGAGAATCTTGTAAGT\nInhibitor-miR-124 ATCAAGGTCCGCTGTGAACACG\nInhibitor-miR-214 GCACAGCAAGTGTAGACAGGCA\nInhibitor-miR-26a AGCCTATCCTGGATTACTTGAA\nNo sequence due to the commercial secret\nInhibitor-NC\n(miR2N0000001-1-5, RIBO Biotechnology Co., LTD)\n9. In the in vivo experiments, only data from 8 weeks after hydrogel implantation are provided. This duration does not adequately reflect the effects of local network stiffening in promoting MSC osteogenic differentiation.\nAuthor Response:\nWe thank the reviewer for this valuable comment. We have supplemented the data from a 4-week in vivo study in a rat critical defect model with implanted hydrogels. As shown in the new results (Figure S25), the CPAC group exhibited a significant increase in bone volume compared to the Blank and the NC group at 4 weeks. This demonstrates that the heterogeneous extracellular matrix structure, autonomously remodeled by MSCs within the hydrogel, can markedly accelerate osteogenic differentiation and mineralization in vivo.\nThe corresponding paragraphs in “MSC-laden CPAC hydrogels promote bone regeneration in an animal critical defect model” now reads:\n…As shown by micro-CT imaging and quantitative analysis, the CPAC group promoted the regeneration process in the early (4 weeks) and late (8 weeks) stages of bone repair (Figure 6b, S25), revealing denser new bone formation than that in the NC group and untreated (“Blank”) groups. Specifically, at 8 weeks postoperatively, the BV (Bone volume) of the CPAC hydrogel group measured at 11.42 ± 0.87 mm3 (Figure 6c), which were 4.7-, 2.4-fold greater than those of the Blank and NC hydrogel groups, respectively. The BV/TV (bone volume/total volume) and BS/BV (bone surface/bone volume) values exhibited similar trends, further demonstrated markedly higher new bone volume in the CPAC hydrogel group compared to the other groups (Figure 6d-e)…\nFigure S25. CPAC hydrogel promotes in vivo bone regeneration in a rat cranial defect model. (a) Micro-CT images showing bone defects at 4 weeks post-implantation for the Blank, NC, and CPAC hydrogel groups. Quantification of (b) bone volume (BV), (c) bone volume/total volume (BV/TV), and (d) bone surface area/total volume (BS/TV) in the bone defect sites after 4 weeks, respectively (n = 3). (e) H&E staining, (f) Masson’s trichrome staining of the bone defects in the Blank, NC, and CPAC hydrogel groups, respectively (Scale bar = 200 μm). (g) OCN immunohistochemical staining in the Blank, NC, and CPAC hydrogel groups (Scale bar = 100 μm). Data are presented as mean ± SD; **p < 0.01, ****p < 0.0001.\n10. The size changes of CPAC hydrogel in a 1 U/mL ALP solution have been studied; however, the ALP concentration during in vivo or bone injury healing is unknown, and it remains unclear whether it would be sufficient to trigger the hydrogel's adaptive remodeling. Additionally, the hydrogel's contraction under in vivo conditions has not been characterized.\nAuthor Response:\nWe thank the reviewer for this valuable comment. To address this concern, we performed additional in vivo experiments. CPAC hydrogels encapsulating MSCs were implanted into a rat critical-sized cranial bone defect model and extracted after 3 days. Measurement of ALP activity within the explanted hydrogels revealed an average concentration of ~5 U/mL, confirming that the in vivo enzymatic environment provides sufficient ALP levels to trigger the adaptive remodeling of CPAC hydrogels. Consistently, nanoindentation mapping of the explanted hydrogels demonstrated the formation of a distinct heterogeneous structure, indicating significant microgel contraction driven by cell-secreted ALP. These data verify that the CPAC hydrogel undergoes phase transformation in vivo, ensuring its functional responsiveness during bone healing.\nThe corresponding paragraphs in the section of “ALP-mediated microgel contraction and local stiffening generates the mechanically heterogenous structures of CPAC hydrogel” now reads:\n…After 3 days of culture, nanoindentation mapping (Figure 2iii) showed a similar pattern of heterogenous stiffening and softening within the CPAC hydrogel treated with exogeneous ALP (local effective moduli ranged from ~0.5 kPa in softer regions to ~14 kPa in stiffer regions). This stiffening is driven by cell-mediated microgel contraction, confirming the autonomous remodeling capacity of the material. In contrast, the MSC- laden NC hydrogels showed no significant changes in local stiffness (Figure S10iii).\nNotably, CPAC hydrogels exhibited comparable behavior in vivo, as ALP secreted by MSCs after implantation reached sufficient levels to drive microgel contraction and the formation of heterogeneous structures within the hydrogel (Figure S13)..…\nFigure S13. (a) ALP concentration over time in CPAC hydrogels after in vivo implantation. (b-c) Heatmaps displaying the distribution of local effective Young's modulus in MSCs-laden CPAC hydrogels at 0 and 72 h post-implantation, respectively.\nReviewer #2:\nThe authors present a study focused on the development of a hydrogel system that mimics the dynamically evolving and heterogeneous extracellular matrix (ECM) environment during tissue development. Specifically, they engineer a cell-programmed adaptive contraction (CPAC) hydrogel containing ALP-responsive microgels that contract in response to alkaline phosphatase (ALP) activity. This induces overall hydrogel network contraction and local matrix stiffening, thereby promoting osteogenic differentiation of mesenchymal stromal cells (MSCs). The authors provide and test a hypothesis for the underlying mechanotransduction mechanism including upregulation of microRNA and demonstrate the translational potential of their system in a rat critical-size bone defect model, where they observed enhanced bone formation.\nThe study presents a dynamic hydrogel platform that responds to cell-released enzymes and modulates stem cell fate through dynamically changing the mechanical signals. The concept is interesting, and the authors have sought to perform both mechanistic investigation and in vivo validation. While the results are promising, additional experiments are required to support their main claims such as ‘‘ALP-induced gel contraction’’, ‘‘local stiffening’’ and ‘‘mechanical heterogeneity’’. Moreover, several aspects of the study require clarification, statistical information or further detail to strengthen its overall scientific rigor, reproducibility and impact.\nMajor comments:\n1. Novelty and significance: It would be helpful if the authors can explain more why this work is novel and their findings are significant for a broad impact.\nAuthor Response:\nWe thank the reviewer for this insightful comment. Previous biomaterials designed to mimic the ECM were predominantly based on preset static mechanical properties (e.g., uniform stiffness). While they could promote stem cell differentiation through mechanotransduction pathways (e.g., YAP/TAZ), they failed to replicate the spatiotemporally dynamic mechanical heterogeneity of natural ECM during development. The innovations and broad applicability of the CPAC hydrogel system proposed in this study are demonstrated in the following aspects:\n1) Cell-driven dynamic remodeling: Our CPAC hydrogel leverages stem cell-secreted ALP to induce a hydrophilic-to-hydrophobic transition and local microgel contraction. This results in progressive, cell-programmed heterogenization of the network, closely resembling native bone ECM remodeling during development. This dynamic remodeling overcomes the spatiotemporal resolution limitations of existing photo- or enzyme-crosslinking methods by synchronizing material stiffening with stem cell differentiation.\n2) Positive feedback regulation: The ALP-triggered local stiffening further amplifies ALP secretion, forming a closed-loop cycle of “mechanical signal-ALP release-hydrogel stiffening- amplified mechanical signal”. This reciprocal interaction better emulates the bidirectional crosstalk between developing ECM and cells, in contrast to the unidirectional mechanical cues of previous systems.\n3) Spatiotemporally coordinated microenvironment: By enabling MSCs to construct a heterogeneous and adaptive microenvironment, the CPAC hydrogel supports bone-like ECM evolution. Its successful application in repairing critical-sized cranial defects in rats, with performance superior to conventional homogeneous hydrogels, highlighting its potential in clinical translation and the regeneration of other complex tissues.\n4) Potential for personalized therapy: Since the remodeling process is driven by endogenous ALP secreted by the patient-derived stem cells, the CPAC system may reduce the need for exogenous growth factors, minimize immunological risks, and adapt to individual variations.\n5) Paradigm shift in biomaterial design: This work shifts the focus from “static ECM mimicry” toward “dynamic cell-material interactions”, establishing a new design principle where cells instruct materials, and materials feedback to cells.\nThe corresponding paragraphs now reads:\nPrevious studies have revealed that the biomaterials with prefabricated stiffness promote the commitment of stem cells towards osteogenesis through the activation of mechanotransduction signaling such as the YAP/TAZ-dependent pathways.22,23 However, few studies have demonstrated the ECM-emulating biomaterials that can dynamically remodel into mechanically heterogenous structures in response to stem-cell-secreted biomarkers to timely provide the complex mechanical cues necessary for guiding the development of complex tissues such as bone based on stem cell differentiations…\n…However, the temporal and spatial resolution of these artificial stiffening mechanisms remains limited in the absence of cellular participation, leading to potential misalignment with the spatiotemporal requirements of stem cell differentiations. Therefore, the development of a biomimetic hydrogel that can dynamically respond to key differentiation markers of stem cells to undergo the cell-programmed heterogenous remodeling in both space and time can potentially enhance the differentiation of stem cells and regeneration of tissue defects…\n2. Study design: While this study probed an interesting concept, their main claims seem to have several gaps, across materials, in vitro- and in vivo models. From the materials aspect, the reliability relies on a thorough characterization of the mechanical properties and heterogeneity of their biomaterials. The range of mechanical stiffness such as magnitude, anisotropy seems unclear.\nAuthor Response:\nWe thank the reviewer for raising this important point. To address these concerns, we performed additional characterizations of the CPAC hydrogel:\n(i) Stiffness range: The Young’s modulus of CPAC hydrogels increased from ~3 kPa at day 0 to ~11 kPa after 3 days of MSC culture both in vitro and in vivo. This stiffening is driven by cell-mediated microgel contraction, confirming the autonomous remodeling capacity of the material.\n(ii) Anisotropy: Compression tests were carried out in both parallel and perpendicular directions. The resulting stress-strain curves and moduli were consistent, indicating that CPAC hydrogels are mechanically isotropic at the bulk level.\n(iii) Heterogeneity: Nanoindentation mapping revealed marked internal heterogeneity in both parallel and perpendicular directions. Local effective moduli ranged from ~0.5 kPa in softer regions to ~14 kPa in stiffer regions, demonstrating the dynamic formation of a heterogeneous microenvironment both in vitro and in vivo.\nTogether, these results provide a more complete mechanical characterization of CPAC hydrogels, confirming their ability to dynamically span a physiologically relevant stiffness range while maintaining isotropic bulk mechanics and pronounced local heterogeneity.\nThe corresponding paragraphs in the section of “ALP-mediated microgel contraction and local stiffening generates the mechanically heterogenous structures of CPAC hydrogel” now reads:\n…Additionally, we evaluated the mechanical stiffness and heterogeneous structure of the CPAC hydrogel in both parallel and perpendicular directions. As shown in the Figure S15, the CPAC hydrogel exhibits the same Young's modulus (~11 kPa), the consistent compressive stress-strain curves, and the local effective moduli ranged (from ~0.5 kPa in softer regions to ~14 kPa in stiffer regions), indicating that CPAC hydrogels are mechanically isotropic at the bulk level both in vitro and in vivo. These findings collectively confirm that both the exogenous and cell-secreted ALP can trigger the contraction of Phos-HPMA microgels…\nFigure S15. Mechanical characterization of CPAC hydrogels. (a) Compression curves and (b) Young’s modulus of CPAC hydrogels at day 0. (c-d) Heatmaps showing local effective Young’s modulus distribution in parallel and vertical directions at day 0. (e-f) Compression curve and Young’s modulus after 3 days in vitro, with corresponding heatmaps displaying the distribution of local effective Young’s modulus in CPAC hydrogels in both parallel and perpendicular directions (g-h). (i-j) Compression curve and Young’s modulus after 3 days in vivo, with corresponding heatmaps displaying the distribution of local effective Young’s modulus in CPAC hydrogels in both parallel and perpendicular directions (k-l).\n3. From the cell biology perspective, whether the readouts are solely due to ALP, and cell-secreted ALP, or not, may require more rigorous investigation by setting up controls and perturbation experiments. It is known that cells also secrete other factors that block ALP acitivity.\nAuthor Response:\nWe thank the reviewer for this thoughtful comment. We agree that it is essential to rigorously evaluate whether ALP, particularly cell-secreted ALP, is the principal driver of CPAC hydrogel remodeling.\nTo this end, CPAC hydrogels were first cultured in PBS supplemented with ALP/ALP inhibitor. As shown in Figure S12a and S12c, inhibition of ALP abolished the formation of heterogeneous structures and prevented changes in storage modulus, confirming that ALP activity is required for CPAC remodeling.\nTo probe potential contributions from other phosphatases, CPAC hydrogels were treated with acid phosphatase. No appreciable heterogeneous structures were observed (Figure S12b), likely because acid phosphatase exhibits limited activity in the neutral PBS environment. Together with the fact that such phosphatases are secreted at low levels and primarily localized intracellularly, this suggests minimal impact on CPAC remodeling.\nTo address the possibility that MSCs secrete molecules capable of suppressing ALP activity, we measured ALP activity in the culture medium over time. The results showed a steady increase in ALP activity from 0 to 168 hours, indicating that any inhibitory factors are insufficient to block ALP function in this context.\nCollectively, these results demonstrate that MSC-secreted ALP is the dominant factor driving microgel contraction and heterogeneous structure formation in CPAC hydrogels.\nThe corresponding paragraphs in the section of “ALP-mediated microgel contraction and local stiffening generates the mechanically heterogenous structures of CPAC hydrogel” now reads:\n…In contrast, no such changes in local stiffness were observed in the NC hydrogels (Figure 2i, ii; S10). To validate the specific response of CPAC hydrogel to ALP, identical sets of CPAC hydrogel were treated with ALP/ALP inhibitor and acid phosphatase, respectively. It was observed that compared to the ALP-treated group, the internal structures of these two groups did not exhibit significant changes (Figure S12a-c), indicating that ALP is the primary factor responsible for the formation of the heterogeneous structure in CPAC hydrogel. We further examined the responsiveness of the hydrogels to cell-secreted ALP by encapsulating osteogenically-induced MSCs in both hydrogels. As shown in Figure S12d, ALP levels in both groups gradually increased during culture, but CPAC hydrogels exhibited a markedly higher and sustained ALP release. This elevated ALP secretion strongly correlates with the heterogeneous stiffening pattern observed in CPAC hydrogels…\nFigure S12. Heatmaps displaying the distribution of local effective Young's modulus in the CPAC hydrogels after 72 hours of treatment with ALP inhibitor (a), ACP (b), and the storage modulus of CPAC hydrogel with different treat treatments for 3days (c), (d), The ALP concentration corresponding to two hydrogels loaded with MSCs at various culture time points. Data are presented as mean ± SD; **p < 0.01, ****p < 0.0001.\n4. From the in vivo aspect, the complexity further increases in terms of proteins and cells in involved. The authors are recommended to add control experiments to demonstrate the enhanced bone formation is not by the gels along, nor other matrix factors, but exactly by their claimed mechanism.\nAuthor Response:\nWe thank the reviewer for highlighting this important point. To rule out potential confounding effects that the observed bone regeneration was due to the hydrogel material itself or other nonspecific matrix effects, we included an additional CPAC only control group (without MSCs). Micro-CT, H&E, Masson, and IHC analyses (Figure R2) revealed that the BV, BV/TV, and BS/TV values in the control group were significantly lower than those in the CPAC group. Importantly, bone formation in the control group was negligible compared to the robust regeneration observed in the CPAC group. This indicates that the enhanced bone regeneration is not attributable to the hydrogel alone but is instead mediated by the cell-secreted ALP-triggered heterogeneous remodeling of CPAC and the resulting mechano-epigenetic regulation of MSC osteogenesis. Our supplementary experiments also confirmed this: after implanting control group and CPAC hydrogels into rats for 3 days, nanoindentation mapping showed that the control group failed to form a distinct heterogeneous structure due to the lack of ALP activity (Figure R3).\nFigure R2. (a) Micro-CT images showing bone defects at 8 weeks post-implantation for the Control, and CPAC hydrogel groups. Quantification of (b) bone volume (BV), (c) bone volume/total volume (BV/TV), and (d) bone surface area/total volume (BS/TV) in the bone defect sites after 8 weeks, respectively (n = 3). (e) H&E staining, (f) Masson’s trichrome staining of the bone defects in the Blank, Hydrogel, NC, and CPAC hydrogel groups, respectively (Scale bar = 200 μm). (g) OCN immunohistochemical staining in the Blank, Hydrogel, NC, and CPAC hydrogel groups (Scale bar = 100 μm). Data are presented as mean ± SD; **p < 0.01, ***p < 0.001.\nFigure. R3 After 3 days in vivo, heatmaps displaying the distribution of local effective Young's modulus in the Control and CPAC hydrogels\n5. Abstract clarity: The abstract does not clearly convey how the hydrogel system works. Key elements, such as the main components of the hydrogel as well as the control groups used in the study, are not mentioned but are critical for interpreting the results.\nAuthor Response:\nWe thank the reviewer for highlighting this important point. We have revised the Abstract to explicitly describe the main components of the hydrogel, the control groups, and the principle underlying the formation of heterogeneous structures within the hydrogel.\nThe corresponding paragraphs in “ABSTRACT” now reads:\n…In this study, we developed a cell-programmed adaptative contraction (CPAC) hydrogel, synthesized by photo-crosslinking p-tert-butylphenyl-modified hyaluronic acid (HA-TP), acryloyl cyclodextrin (Ac-β-CD), and phosphorylated hydroxypropyl methacrylate (Phos-HPMA) microgels. The CPAC hydrogel enables mesenchymal stem cells (MSCs) to actively remodel the hydrogel matrix through alkaline phosphatase (ALP, an early osteogenic marker)-mediated hydrophilic-to-hydrophobic transition and contraction of integrated Phos-HPMA microgels. In contrast, control hydrogels containing phosphorylated hydroxyethyl acrylate microgels lack this remodeling ability. This cell-programmed remodeling drives local network stiffening and the emergence of heterogeneous structures, promoting MSC osteogenic differentiation through a positive feedback loop…\n6. The introduction would benefit from a comparison to naturally derived hydrogels such as collagen. How does the contractile behaviour of the presented system compare, and what advantages does it offer over existing ECM-mimetic materials?\nAuthor Response:\nWe thank the reviewer for this valuable suggestion. Traditional collagen hydrogels typically exhibit a relatively homogeneous network structure with limited dynamic responsiveness, where remodeling relies on slow enzymatic degradation to facilitate cell migration and mineralization. Their static modulus and sluggish remodeling prevent them from rapidly adapting to the evolving mechanical needs of differentiating cells. In contrast, the CPAC hydrogel features a host-guest crosslinked network that provides significantly greater dynamic properties, better supporting such cellular processes. More importantly, its remodeling is actively programmed by the cells themselves, i.e., ALP secreted by cells triggers a hydrophilic-to-hydrophobic transition in the integrated microgels, driving localized contraction and heterogeneous stiffening that mimic the spatiotemporal heterogeneity of bone ECM during development. Furthermore, this cell-triggered contraction further amplifies mechanical signals, creating a self-reinforcing positive feedback loop. Therefore, CPAC hydrogels overcome the passive, slow-responding nature of collagen and other naturally derived ECM mimetics, offering a more precise and timely regulation of stem cell differentiation.\nThe corresponding paragraphs in “INTRODUCTION” now reads:\nPrevious studies have revealed that the biomaterials with prefabricated stiffness promote the commitment of stem cells towards osteogenesis through the activation of mechanotransduction signaling such as the YAP/TAZ-dependent pathways.22,23 However, few studies have demonstrated the ECM-emulating biomaterials that can dynamically remodel into mechanically heterogenous structures in response to stem-cell-secreted biomarkers to timely provide the complex mechanical cues necessary for guiding the development of complex tissues such as bone based on stem cell differentiations...\n7. The exact composition of the hydrogel remains unclear in the introduction and Figure 1. Including a schematic that clearly explains the composition and assembly of the hydrogel from microgels and other components for all experimental hydrogel groups would greatly improve clarity. Also pay attention to complete & matching legends for the schematics.\nAuthor Response:\nWe thank the reviewer for the thoughtful suggestion. The updated schematic now explicitly illustrates the composition and assembly of the hydrogel, including both the microgel components and other structural elements, across all experimental groups. We have also revised the figure legend to ensure that it fully matches and explains each component and step depicted in the schematic.\nThe Figure 1 now reads:\n8. Figure 2g: It would be helpful to include confocal images of the hydrogels prior to ALP treatment to demonstrate the changes induced by contraction.\nAuthor Response:\nWe thank the reviewer for this constructive suggestion. We have now supplemented confocal images of NC and CPAC hydrogels prior to the ALP treatment to directly demonstrate the structural changes induced by ALP-mediated contraction.\nThe Figure 2g now shows:\n9. Figure 2h: Please indicate which statistical test was used and annotate significance levels directly on the graph.\nAuthor Response:\nWe thank the reviewer for this helpful suggestion. The two-tailed Student’s t test was used for comparisons between two groups. We have now annotated the corresponding significance levels directly in Figure 2h.\nThe Figure 2h now shows:\nFigure 2. (h) Time-dependent variation of storage modulus in NC and CPAC hydrogels after ALP treatment. Data are presented as mean values ± SD, n = 3 independent hydrogels, ns, not significant, *p < 0.05, ****p < 0.0001 (two-tailed Student’s t-test).\n10. You claim that “CPAC hydrogel can better facilitate stellate spreading”. Please explain where this can be seen or provide more morphological data to support it.\nAuthor Response:\nWe thank the reviewer for the helpful comment to better address our manuscript. We have carefully considered your comments and made the following revisions to our manuscript.\nWe designed the following experiments to demonstrate CPAC hydrogel can better facilitate stellate spreading. As shown in the Figures S18, Actin staining was performed on the MSC-loaded hydrogel (NC and CPAC), and statistical analysis of the MSC spread area revealed that MSCs spread over a larger area in CPAC hydrogel, exhibiting a stellate spreading morphology.\nThe corresponding paragraphs in the section of “The cell-programmed local stiffening in CPAC hydrogel promotes osteogenic differentiation of MSCs” now reads:\n…These results suggest that the heterogeneous structure formed during microgel contraction not only provides space for cell expansion but also creates a favorable microenvironment that sustains MSC proliferation. Furthermore, the results of Actin staining and the statistical analysis of cell spreading area for MSCs (Figure S18) indicated that MSCs spread over a larger area in the CPAC hydrogel. To evaluate the impact of cellular ALP-induced local stiffening in CPAC hydrogel on the osteogenesis of encapsulated MSCs, we analyzed the expression of key osteogenic markers in encapsulated MSCs…\nFigure S18. (a) Representative actin staining image of MSCs in NC and CPAC hydrogels. (b) Spread area of MSCs in NC and CPAC hydrogel, n = 5 independent hydrogels. Data are presented as mean ± SD, ****p < 0.0001 (two-tailed Student’s t-test).\n11. Figures 3b/c: The CPAC hydrogels appear to have a higher cell density compared to NC controls. Is this a localized effect due to contraction, or does the material promote proliferation? Higher local cell density itself may influence osteogenic differentiation.\nRelated to this, can you comment on the macroscopic effect of ALP-mediated contraction on the entire hydrogel, does the hydrogel size change?\nAuthor Response:\nWe thank the reviewer for this thoughtful comment. We performed EdU incorporation assays, live/dead staining, and CCK-8 assays in both NC and CPAC hydrogels. As shown in the Figure S17, MSCs in CPAC hydrogels exhibited significantly higher proliferation efficiency compared to those in NC hydrogels, while cell viability remained consistently high in both groups. This enhanced proliferation is likely facilitated by the heterogeneous architecture generated through ALP-mediated microgel contraction, which increases local network porosity and provides additional space for cell expansion. Thus, the higher apparent cell density in CPAC hydrogels arises from a combination of both increased proliferation and local spatial reorganization.\nSimultaneously, the ALP-triggered contraction of microgels does lead to macroscopic shrinkage of the entire hydrogel. Specifically, the average hydrogel diameter decreased from ~4.5 mm to ~3 mm after 72 hours of culture (Figure R4). This demonstrates that the CPAC hydrogel undergoes measurable bulk shrinkage while still maintaining sufficient internal space to support cell survival and proliferation. Together, these results indicate that the observed increase in local cell density is not an artifact of hydrogel contraction alone but reflects a genuine proliferation-promoting effect of the CPAC microenvironment.\nFigure S17. Proliferation and survival of MSCs in NC and CPAC hydrogels. (a) EdU assay showing newly synthesized DNA, (b) live/dead staining, and (c) CCK-8 assay confirming enhanced proliferation in CPAC hydrogels.\nFigure R4. The macroscopic changes of the two hydrogels after 3 days of culture.\n12. Can you specify how the “high-modulus N,N-dimethylacrylamide (DMAA) microgels” compare to the previous control microgels (Phos-HEA) in terms of mechanical properties?\nAuthor Response:\nWe thank the reviewer for highlighting this important point. As shown in Figure S10, the effective Young's modulus of DMAA microgel exceeds 10,000 Pa, which is significantly higher than that of Phos-HEA (~7,500 Pa) and is comparable to the modulus of Phos-HPMA microgels after MSC-mediated contraction (Figure 2(iii)). Meanwhile, neither DMAA nor HEA microgels exhibited significant changes in modulus after exposure to ALP and MSCs, confirming that they are non-reponsive to ALP. Therefore, Phos-HEA microgels serve as a low-modulus control for Phos-HPMA microgels in their initial state, while DMAA microgels serve a high-modulus control for Phos-HPMA microgels after ALP-induced stiffening.\n13. The claim “NCLS hydrogels showed elevated osteogenic marker expression relative to NC hydrogels” is not evident from the data shown, please justify or explain more.\nAuthor Response:\nWe thank the reviewer for this helpful reminder. Upon careful re-examination of our data, we agree that the NCLS and NC hydrogel groups did not show a significant difference in MSC osteogenic marker expression after 3 days of culture. We have therefore corrected the relevant statement in the manuscript to avoid overstating the effect.\nThe corresponding paragraphs in the section of “The cell-triggered local stiffening of CPAC hydrogel promotes osteogenesis by activating mechanotransduction” now reads:\n…CPAC hydrogels exhibited the highest Col 1 and ALP expression compared to all groups, while NCLS hydrogels showed no significant difference in osteogenic marker expression relative to NC hydrogels.\n14. Figure 4: Please provide rationale for the selection of mechanotransduction markers (pFAK, pMyosin IIa, Paxillin).\nAuthor Response:\nWe thank the reviewer for the comments. The selection of pFAK, pMyosin IIa, and Paxillin was based on their central and complementary roles in the focal adhesion-mediated mechanotransduction cascade. Specifically: 1) pFAK acts as the primary mecanosensor and signal amplifier, with its phosphorylation serving as an early indicator of force detection and conversion into biochemical signals; 2) pMyosin IIa functions as the intracellular force generator, reflecting the level of cell contractility and tension in response to the matrix; 3) Paxillin functions as a critical scaffolding protein, with its phosphorylation status reflecting the maturation, size, and stability of the focal adhesions necessary to effective force transmission. Monitoring these three markers together therefore provides an integrated readout of mechanotransduction, spanning force sensing, contractility, and adhesion reinforcement.\n15. Figure 4: Why are the microgels not visible in the confocal images like in Figures 3 & 5? The figure caption makes it a bit unclear, if cells were seeded on top of hydrogels rather than embedded.\nAuthor Response:\nWe appreciate the reviewer’s comments. To accurately quantify cell spreading areas, MSC-containing hydrogel precursor solutions were dropped on glass slides and photocrosslinked. Immunofluorescence imaging was then focused on the near-surface layers of the hydrogel rather than the interior, which allowed precise assessment of cell morphology and the expression of various mechanotransduction markers. The microgels were thus outside the field of view, preventing interference with cell area measurements.\n16. Figure 4b: Collagen I appears to be predominantly intracellular. Is there an explanation for this observation?\nAuthor Response:\nWe greatly appreciate your insightful question. As a key marker of osteogenesis, type I collagen (Col1) is initially synthesized intracellularly as procollagen before being secreted and deposited into the ECM. At this early time point (3 days), MSCs have not yet deposited substantial extracellular collagen, which explains the predominantly intracellular signal. In addition, the multiple washing and permeabilization steps required for immunofluorescence staining may partially remove deposited extracellular Col1, and any remaining matrix-associated collagen may display fluorescence intensity too weak to be distinguished from the stronger intracellular signal.\nTo address this concern, we complemented our IF analysis with Masson’s trichrome staining on hydrogel cryosections. The darker blue areas observed in the CPAC group confirmed enhanced extracellular Col 1 deposition, supporting the conclusion that CPAC hydrogels promote osteogenic differentiation of MSCs.\nThe corresponding paragraphs in the section of “The cell-programmed local stiffening in CPAC hydrogel promotes osteogenic differentiation of MSCs” now reads:\n…Immunofluorescence staining, Masson’s trichrome staining and alkaline phosphatase color development kit further confirmed enhanced protein expression of Col 1 and ALP in the CPAC hydrogel group, with greater fluorescence intensity than in the NC hydrogel group (Figures 3b-d, S20, S21). Consistently, WB analysis further verified the increased synthesis of Col 1 and ALP proteins in the CPAC hydrogel group (Figure 3e)…\nFigure S20. The Masson’s trichrome staining in the two hydrogel groups (scale bar = 50 um).\n17. The hydrogel is described as responsive to ALP secreted by MSCs, yet the immunofluorescence images show primarily intracellular ALP. Please clarify how this observation aligns with the proposed mechanism of hydrogel contraction, which relies on extracellular ALP activity.\nAuthor Response:\nWe thank the reviewer for this insightful question. ALP is predominantly localized on the plasma membrane (via GPI anchors) as well as in intracellular compartments such as endoplasmic reticulum, and Golgi apparatus. Importantly, membrane-associated ALP can be released into the extracellular space by cleavage of its GPI anchor through specific enzyme, allowing it to diffuse into the hydrogel and trigger microgel contraction.\nDuring the immunofluorescence staining process, the hydrogel undergoes sequential fixation, permeabilization, and multiple washing steps likely removed soluble or weakly bound ALP within the hydrogel matrix. Any residual extracellular ALP may also have been too faint to distinguish from the much stronger intracellular signal, explaining why the IF images appear to show primarily intracellular ALP.\nTo overcome this limitation, we additionally performed staining of hydrogel cryosections using a BCIP/NBT alkaline phosphatase color development kit. This assay confirmed the presence of extracellular ALP within the CPAC hydrogel matrix, with more intense staining compared to controls, supporting the conclusion that secreted ALP actively contributes to hydrogel remodeling.\nThe corresponding paragraphs in the section of “The cell-programmed local stiffening in CPAC hydrogel promotes osteogenic differentiation of MSCs” now reads:\n…Immunofluorescence staining, Masson’s trichrome staining and alkaline phosphatase color development kit further confirmed enhanced protein expression of Col 1 and ALP in the CPAC hydrogel group, with greater fluorescence intensity than in the NC hydrogel group (Figure 3b-d, S20, S21). Consistently, WB analysis further verified the increased synthesis of Col 1 and ALP proteins in the CPAC hydrogel group (Figure 3e)…\nFigure S21. The distribution of alkaline phosphatase in the two hydrogel groups. (scale bar = 50 um).\n18. A 3-day culture period is relatively short for observing osteogenic differentiation. Have longer culture periods been considered?\nAuthor Response:\nWe appreciate the reviewer’s valuable comment. We agree that a 3-day culture period may appear relatively short for assessing osteogenic differentiation. To address this, we also conducted longer-term culture experiments. At day 7, qPCR analysis demonstrated that the expression levels of osteogenic genes (RUNX2, ALP, Col 1, and OPN) were significantly higher in the CPAC hydrogel group compared to the NC group. This further confirms that the cell-programmed heterogeneous structure in CPAC hydrogels not only promotes early osteogenic signaling but also sustains and enhances osteogenic differentiation over longer culture periods.\nThe corresponding paragraphs in the section of “The cell-programmed local stiffening in CPAC hydrogel promotes osteogenic differentiation of MSCs” now reads:\nTo evaluate the impact of cellular ALP-induced local stiffening in CPAC hydrogel on the osteogenesis of encapsulated MSCs, we analyzed the expression of key osteogenic markers in encapsulated MSCs. After 3 days of culture, PCR results revealed significantly upregulated expression of crucial osteogenic genes, including runt-related transcription factor 2 (Runx2), ALP, Col 1, and osteopontin (OPN) in MSCs cultured in CPAC hydrogels compared to those in NC hydrogels (Figure 3a). The 7-day PCR data also demonstrated that MSCs exhibited a higher degree of osteogenic differentiation within the CPAC hydrogel over longer culture periods (Figure S19) .…\nFigure S19. Relative gene expression of selected osteogenic markers (Runx2, ALP, Col 1, OPN) after 7 days of osteogenic induction, Data are presented as mean values ± SD, n = 3 independent hydrogels, ****p < 0.0001 (two-tailed Student’s t-test).\n19. Materials and Methods in SI: Several important methodological aspects require clarification: How was the microgel shrinkage quantified? Was it done manually, and how many replicates were analysed?\nAuthor Response:\nWe thank the reviewer for this important comment. Microgel shrinkage was quantified by capturing images of Phos-HEA and Phos-HPMA microgels before and after ALP treatment using an inverted microscope. Fifteen microgels were randomly selected from each group, and their diameters were measured using Nano Measure software (Version 1.2). The replicates information has been added into the caption of Figure 2c.\nThe corresponding section in SI now reads:\nMicrogel shrinkage quantification\nMicrogel shrinkage was quantified by capturing images of Phos-HEA and Phos-HPMA microgels before and after ALP treatment using an inverted microscope. Fifteen microgels were randomly selected from each group, and their diameters were measured using Nano Measure software (Version 1.2). The shrinkage of the microgels was quantified using Nano Measure (Version 1.2)\nFigure 2. (c) diameter changes of Phos-HEA and Phos-HPMA microgels after 72 hours in PBS and ALP solution (scale bar = 100 μm). Data are presented as mean values ± SD, n = 15 independent microgels, ns, not significant, ****p < 0.0001 (two-tailed Student’s t-test).\n20. Please provide more details about your hydrogel preparation from your individual components including precise concentrations and volume ratios. Please also specify the light source, wavelenght and intensity (also for photorheology).\nAuthor Response:\nWe thank the reviewer for the suggestion. We have now added detailed descriptions of NC, NCLS, and CPAC hydrogel preparation, as well as specifications for the light source, wavelength, and intensity for both polymerization and photorheology in the SI.\nThe corresponding sections in SI now reads:\nSynthesis of NC，NCLS, and CPAC hydrogel\nFor the NC hydrogel, HA-TP (1%), gelatin (3%), Ac-β-CD (5%), and Ac-Phos-HEA microgel (5%) were dissolved in PBS (900 μl). After complete dissolution, the solution was allowed to stand overnight. Subsequently, 100 μl of 0.5 wt% lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP) was added, and polymerization was initiated under blue light (λ = 405 nm, 350 mW/cm²) for 5 minutes to obtain the NC hydrogel. For the NCLS and CPAC hydrogels, the Ac-Phos-HEA microgels were replaced with an equal mass of Ac-DMAA microgels and Ac-Phos-HPMA microgels, respectively, while keeping all other conditions identical.\nFabrication and rheological characterization of the NC and CPAC hydrogels\n…Samples were formed in situ by pipetting a 50 µL monomer solution between the bottom HR-30 hybrid rheometer plate and an 8 mm flat plate, and the experiment was initiated after photopolymerization (λ = 365 nm, 396 mW/cm2)…\n21. What were the measurement parameters used for rheological analysis?\nAuthor Response:\nWe appreciate the reviewer’s comment. We have now added detailed measurement parameters used for rheological analysis in the SI.\nThe corresponding section in SI now reads:\n…Time sweep measurements were conducted at 37 °C with a fixed strain of 1% and an frequency of 10 rad/s for 120 seconds. For the alternating strain step shear test, the temperature was maintained at 37 °C, with a frequency of 10 rad/s. The strain was alternated between 60% (45 seconds) and 1% (45 seconds) for three full cycles to obtain the step shear profile…\n22. In your nanoindentation measurements, was stiffness assessed in a single 50 µm × 50 µm region or across multiple areas? Are the results shown representative?\nAuthor Response:\nWe thank the reviewer for highlighting this point and apologize for the lack of clarity in our initial description. In our nanoidentation experiments, we randomly selected a 350 μm × 350 μm square area on the sample surface. Nanoindentation measurements were performed at intervals of 50 μm, yielding an 8 × 8 grid of valid Young's modulus data points, which were then compiled into heatmaps. To ensure representativeness, different regions were selected randomly, and the measurements were repeated three times independently.\nThe corresponding section in SI now reads:\nPhos-HPMA microgel mediated the heterogeneous distribution of matrix stiffness in the CPAC Hydrogel\nNanoindentation technology was applied to characterize the Young’s modulus distribution of both hydrogels under underwater conditions via a Piuma nanoindenter (Optics 11, Netherlands). Specifically, the hydrogels were maintained in PBS at 37 °C for 1 h before being loaded onto the plate. We randomly selected a 350 μm × 350 μm square matrix area on the surface of the samples and performed nanoindentation measurements at intervals of 50 μm. A total of 8 × 8 valid Young's modulus data points were obtained, forming heatmap. The selected areas were random, and the measurements were repeated three time.\n23. What was the cell seeding density, and what was the source of the hMSCs?\nAuthor Response:\nWe thank the reviewer’s comments. The human mesenchymal stem cells (hMSCs) were purchased from REGEN-αGEEK Biotechnology Co. Ltd (a 65-year-old male donor). For all experiments, hMSCs were encapsulated in the hydrogel at a density of 1×107 cells per milliliter. This information has now been added into the SI for clarity.\nThe corresponding paragraphs in SI now reads:\nThe human mesenchymal stem cells (hMSCs) were purchased from REGEN-αGEEK Biotechnology Co. Ltd (a 65-year-old male donor)…\n…For each type of hydrogel, the encapsulation density of hMSCs was 10 million cells per milliliter….\n24. In the in vivo experiments, was there a contralateral defect used as an internal control in each animal as indicated in Figure 6a?\nAuthor Response:\nYes, a contralateral defect was used as an internal control in each animal, as indicated in Figure 6a. This has now been clarified in the SI.\nThe corresponding sentences in SI now read:\n…Aseptic surgical exposure of the calvarium was performed, and two 5 mm diameter critical-sized defects were created in each rat skull. In each animal, one defect was filled with the hydrogel sample, while the contralateral defect served as an internal control…\n25. Please describe the analysis of micro-CT data in more detail.\nAuthor Response:\nWe thank the reviewer’s comment. We have now described the analysis of micro-CT data in detail in the revised manuscript.\nThe corresponding paragraphs in “MSC-laden CPAC hydrogels promote bone regeneration in an animal critical defect model” now reads:\n…As shown by micro-CT imaging and quantitative analysis, the CPAC group promoted the regeneration process in the early (4 weeks) and late (8 weeks) stages of bone repair (Figure 6b, S25), revealing denser new bone formation than that in the NC group and untreated (“Blank”) groups. Specifically, at 8 weeks postoperatively, the BV (Bone volume) of the CPAC hydrogel group measured at 11.42 ± 0.87 mm3 (Figure 6c), which were 4.7-, 2.4-fold greater than those of the Blank and NC hydrogel groups, respectively. The BV/TV (bone volume/total volume) and BS/BV (bone surface/bone volume) values exhibited similar trends, further demonstrated markedly higher new bone volume in the CPAC hydrogel group compared to the other groups (Figure 6d-e)…\n26. You write that your skull specimens were decalcified for 2 months before fixation, whereas fixation is generally performed before decalcification to preserve tissue, cell and protein integrity. Could you please explain your choice of procedure and whether that could affect the staining results?\nAuthor Response:\nWe thank the reviewer for this important comment. We apologize for the ambiguity in our original description. In our experiments, the skull specimens were first fixed in 4% paraformaldehyde for 48 hours to ensure proper preservation of tissue morphology and antigens. Following fixation, the samples underwent mild decalcification with 10% EDTA solution (pH 7.4, 25 °C) for 6 weeks. Compared with rapid decalcification agents, this method maximally preserves antigen activity and is widely regarded as the gold standard for immunohistochemistry (IHC). Finally, standard antigen retrieval using citrate buffer was performed on paraffin-embedded tissue sections to unmask epitopes prior to IHC staining. Our approach is consistent with established protocols reported in classical literature [8,9]. Therefore, fixation was indeed performed before decalcification, and the chosen method ensures reliable antigen preservation without compromising the IHC results.\nThe corresponding paragraphs in SI now reads:\nImmunohistochemical staining\nFollowing decalcification, specimens were fixed in 4% formaldehyde for 24 hours, followed by extensive washing with deionized water to remove residual fixative. Tissue dehydration was performed through a graded ethanol series (30-100%), with subsequent xylene clearing and paraffin embedding. Serial sections (4 μm thickness) were prepared for histological evaluation.\nTissue sections were subjected to hematoxylin and eosin (H&E) and Masson's trichrome staining according to standard protocols, followed by coverslipping with mounting medium. For immunohistochemical detection, sections were incubated overnight at 4°C with primary antibodies targeting osteocalcin (OCN; 1:100 dilution, DF12303, Affinity). Following primary antibody incubation, sections were treated with HRP-conjugated secondary antibody (room temperature, 30 minutes) and developed with diaminobenzidine substrate (10 minutes, room temperature). All stained sections were permanently mounted for microscopic evaluation.\n27. The section “Immunohistochemical staining” describes the use of an anti-ALP antibody, whereas the data in Figure 6 shows OCN. Please correct.\nAuthor Response:\nWe thank the reviewer for reminding this point. Now we have corrected it.\nThe corresponding paragraphs in SI now reads:\n…For immunohistochemical detection, sections were incubated overnight at 4°C with primary antibodies targeting osteocalcin (OCN; 1:100 dilution, DF12303, Affinity)…\n28. The section “3D cell culture within hydrogels” seems incomplete, especially regarding the use of the EZH2 inhibitor (starting from line 163).\nAuthor Response:\nWe thank the reviewer for this helpful suggestion. We have now completed this section by adding a full description of the culture procedure and the application of the EZH2 inhibitor.\nThe revised section in the SI “3D cell culture within hydrogels” now reads:\nThe Human mesenchymal stem cells (hMSCs) were purchased from REGEN-αGEEK Biotechnology Co. Ltd (a 65-year-old male donor). Primary hMSCs were cultured in αMEM supplemented with 16.7% fetal bovine serum (Excel), 1% penicillin-streptomycin (Gibco), and 1% L-glutamine (Gibco), and passaged to passage 4 (P4) under conditions of 37°C and 5% CO . After 5 minutes of digestion with 0.05% trypsin (Gibco), the cells were 2 collected. Following cell counting, the cells were placed into individual 1.5 mL Eppendorf tubes and centrifuged at 1200 rpm for 3 minutes, with the supernatant discarded. For each type of hydrogel, the encapsulation density of hMSCs was 1 × 10⁷ cells per mL of hydrogel precursor solution (NC, NCLS, or CPAC). After mixing, 50 µL of precursor solution containing hMSCs was crosslinked under blue light (λ = 405 nm) for 5 minutes and transferred into a 24-well plate. Constructs were continuously cultured in either growth or osteogenic medium (10 mM β-glycerophosphate, 50 µg/mL ascorbic acid 2-phosphate, and 100 nM dexamethasone). For the EZH2 inhibitor group, the inhibitor (Cat: HY-148458, MCE) was added to osteogenic medium at a final concentration of 10 μM. Constructs were cultured in the inhibitor-containing osteogenic medium for 12 hours, after which the medium was replaced with fresh osteogenic medium for continued culture.\nMinor comments:\n29. The sequence of figure citations in the text is partially incorrect, for example, Supplementary Figure S11 is cited before S10. Also, the reference to Figure S11 is unclear, is it pertaining to stiffness (text) or ALP concentration (figure)?\nAuthor Response:\nWe thank the reviewer for pointing out this issue. We have carefully corrected the sequence of figure citations in the manuscript. After revising the manuscript and adjusting the relevant figures, the original Figure S10 has been changed to Figure S12d, and the original Figure S11 has been moved to Figure S15. In addition, we clarified that the reference in the text pertains to ALP concentration, which is now correctly shown as Figure S12d.\nThe Figure S12d and Figure S15 now read:\nFigure S12d. (d) The ALP concentration corresponding to two hydrogels loaded with MSCs at various culture time points. Data are presented as mean ± SD, **p < 0.01, ***p < 0.001, ****p < 0.0001.\nFigure S14. Visualization of the injectability of CPAC hydrogel.\n30. Figure S12: Correct graph labellings “i, ii, iii, iv, v, vi” and their citations in the text.\nAuthor Response:\nWe thank the reviewer for this helpful comment. The original Figure S12 has been changed to Figure S10, and the labeling of Figure S10 has now been corrected to “i, ii, iii, iv, v, vi”, and the corresponding citations in the main text and figure legend have been revised accordingly to ensure consistency.\nThe Figure S12 now reads:\nFigure S10. Heatmaps displaying the distribution of local effective Young's modulus in the NC Hydrogel (a) and NCLS Hydrogel (b) after 72 hours of treatment with PBS (i), ALP (ii), and MSC-loaded (iii) respectively.\n31. The manuscript contains several typos that should be corrected, for example line 68 “promote the of commitment of stem cells”, line 102 “CAPC” instead of “CPAC” or line 272 “this modulus remained larged unchanged”\nAuthor Response:\nWe thank the reviewer for carefully pointing out these errors. All the mentioned typos have been corrected in the revised manuscript.\nLine 68 now reads:\n…Previous studies have revealed that the biomaterials with prefabricated stiffness promote the commitment of stem cells towards osteogenesis through the activation of mechanotransduction signaling…\nLine 102 now reads:\n…This mechanotransduction-epigenetic coupling enabled by the CPAC hydrogel significantly enhanced the osteogenic commitment of MSCs…\nLine 272 now reads:\n…the initial effective Young's modulus of NCLS hydrogels matched the peak modulus of CPAC hydrogels, and this modulus remained unchanged upon exposure to ALP or after 3 days of MSC culture…\n32. Figure S9: The panel c described in the figure caption does not exist. Additionally, please clarify the meaning of the red and blue vertical lines in the existing panel c. Also the mention of “half of the peak stress relaxation times (τ1/2 = 150 ± 10 s)” doesn’t match the data shown in Figure S9c.\nAuthor Response:\nWe thank the reviewer for the kind reminder. We have now corrected Figure S9 and its caption to ensure accuracy and consistency.\nThe Figure S9 now reads:\nFigure S9. (a) Time sweep of rheological tests revealing that both hydrogels exhibited similar macroscopic mechanical properties; (b) Young's modulus of the two hydrogels; (c) Stress relaxation curves with insert showing the relaxation times of the two hydrogels, (the red and blue vertical lines indicate the half-relaxation times, τ ); (d) Rheological shear ₁/₂ strain curves of CPAC hydrogel at high strain (60%) and low strain (1%).\n33. Figure 2f: Please label the images.\nAuthor Response:\nWe thank the reviewer for the kind reminder.\nThe Figure 2f now shows:\nReviewer #3:\nI 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.\nReviewer #4:\nI 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.\nReferences\n[1] Landau, L.D. et al. Theory of elasticity: volume 7. (Elsevier Press. 2012).\n[2] Li, Y. et al. Dysregulation of histone modifications in bone marrow mesenchymal stem cells during skeletal ageing: roles and therapeutic prospects. Stem Cell Res. Ther. 14, 166 (2023).\n[3] Li, C. et al. Programmed cell senescence in skeleton during late puberty. Nat Commun. 8, 1312 (2017).\n[4] Yang, R. et al. 1,25-Dihydroxyvitamin D protects against age-related osteoporosis by a novel VDR-Ezh2-p16 signal axis. Aging Cell. 19, e13095 (2020).\n[5] Su, X. et al. Epigenetic therapy attenuates oxidative stress in BMSCs during ageing. J Cell Mol Med. 26, 375–84 (2022).\n[6] Jing, H. et al. Suppression of EZH2 prevents the shift of osteoporotic MSC fate to adipocyte and enhances bone formation during osteoporosis. Mol. Ther. 24, 217–229 (2016).\n[7] Hemming, S. et al. EZH2 and KDM6A act as an epigenetic switch to regulate mesenchymal stem cell lineage specification. Stem Cells. 32, 802–815 (2014).\nHemming, S. et al. EZH2 deletion in early mesenchyme compromises postnatal bone microarchitecture and structural integrity and accelerates remodeling. Faseb J. 31, 1011–1027 (2017).\n[8] Wang, J. et al. Engineering Large-Scale Self-Mineralizing Bone Organoids with Bone Matrix-Inspired Hydroxyapatite Hybrid Bioinks. Adv. Mater. 36, 2309875 (2024).\n[9] Yao, J. et al. A Moldable Hydrogel based on Sericin and Zn2+/F- Dual-doped Hydroxyapatite Promotes Skull Defect Repair through the Synergistic Effects of Immunoregulation, Enhanced Angiogenesis and Osteogenesis. Chem. Eng. J. 491, 151789 (2024)."}, {"round": 2, "reviewer_comments": {"reviewer_1": "Overall, the authors have provided substantial additional experiments that address the principal concerns: they demonstrate that ALP is the dominant driver of material remodeling; add in vitro and in vivo evidence for mechanical heterogeneity; strengthen osteogenic outcomes and pathway validation; and improve methods and figure descriptions. The responses are generally satisfactory, and the manuscript has been significantly improved. We recommend publication in this journal after minor revision.\nRemaining issues to be addressed before acceptance:\n1. The ALP inhibitor condition lacks essential experimental details in the manuscript, including the inhibitor identity/composition, concentration, exposure regimen, and appropriate controls.", "reviewer_2": "The authors have improved the manuscript with revision and additional experiments. However, methods are not updated and the revised version still lacks sufficient scientific rigor and accuracy to be considered for publishing in NC at this stage. My major concerns are the authors just provided more results, but did not explain how they obtained the students in a rigorous manner. Several important method descriptions such as protein expression and in vivo control experiments still miss important details necessary for the reliability and reproducibility in the paper.\nTherefore, another revision is needed. Please address these comments and questions:\n1. Introduction: The authors stated ''The CPAC hydrogel thus represents a shift from “static ECM mimicry” toward “dynamic, cell instructed biomaterials, integrating structural remodeling and biomechanical evolution to regulate the mechanotransduction–epigenetic axis of interacting cells.'' This seems over-shooting as they should be grounded, focus on what exactly they have achieved and critically discuss the limitations.\n2. The abstract and Figure 1 contain unnecessary information and should be more compact to ensure good readability and clarity.\n3. The authors started with skull development (Figure 1) but later shifted to in vivo studies on bone regeneration. Note that development is not same as regeneration. Please correct to make it crystall-clear.\n4. The English language can be greatly improved.\nOther comments:\n1. New methods were only included for Fig S11 & Fig 5i/R1, please ensure all new data shown (esp. Figs S12-S24) are accompanied by the corresponding methods description, in particular:\na. Method for ALP quantification, Fig S12, S13\nb. Method for in vivo implantation model & sample processing for ALP, nanoindentation & mass quantification, Fig S13, S23. Also clarify what 0h in vivo means.\nc. Method for “Remaining mass (%)” quantification, specifically if it was the dry mass, or if it was the wet mass (mostly water): how the results should be interpreted taking into account the bulk volume shrinkage as well.\nd. Method for bulk Young’s modulus quantification, e.g. Fig. S9b, S15\ne. Methods for EdU incorporation assays, live/dead staining, and CCK-8 assays (incl. time points).\nf. Method for alkaline phosphatase color development kit.\n2. Follow-up to previous round Question 12: It is unclear how to deduce effective Young’s moduli values from all the nanoindentation heat map graphs. The values in the graphs don’t match the values of the color scale legend (e.g. Fig 2i(iii): graph at X(0), Y(300) shows ~1000Pa but color indicates ~4000Pa). Please double-check or clarify the meaning of the color scale values (they are linear instead of logarithmic as in graph).\n3. The new Figure 1b is much clearer now, but the text might be too small for publication. Also, the figure captions needs updating.\n4. Figure S17c: Recommended to labels as “Relative metabolic activity (%)“ or similar (not viability) when used as a proliferation assay.\n5. Regarding previous round Question 26: It seems like the method section has not been adapted to reflect the experimental procedure described in your response. The SI still reads:\n“Skull specimens were initially subjected to micro-computed tomography (micro-CT) scanning, followed by decalcification in 10% ethylenediaminetetraacetic acid (EDTA, pH 7.4) for two months in preparation for immunohistochemical evaluation. […] Following decalcification, specimens were fixed in 4% formaldehyde for 24 hours”\nwhich indicates the sequence 1) decalcification, 2) fixation. Maybe you mean: „Before decalcification, specimens were fixed in 4% formaldehyde for 24 hours“?\n6. Fig S18: the results should be normalized to cell number.\n7. Methods for quantification of micro-CT data should be updated to ensure reproducibility.\n8. Figure R2 (response to Q4) should be added to at least SI, given its importance to support the in vivo findings.\n9. Figure 3 & 5: WesternBlott: please explain why proteins of interests and reference protein are analyzed separately, instead of 1 combined image.\n10. Figure 4: it is unclear why the authors showed only single cells instead of showing tens of cells per image. How many cells in total have been analyzed?", "reviewer_3": "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.", "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": "Author Response:\nWe thank the reviewer for this insightful and constructive suggestion. We agree that the statement in the introduction was overly broad and could give the impression that our system completely resolves all challenges in dynamic ECM mimicry. We have refined this section to more accurately reflect the specific achievements of our study. Namely, the demonstration of an ALP-responsive, cell-instructed hydrogel capable of forming mechanically heterogeneous structures that promote osteogenic differentiation through mechanotransduction–epigenetic coupling.\nThe Introduction now reads:\nThe CPAC hydrogel thus demonstrates a cell-instructed biomaterial system capable of undergoing dynamic, ALP-mediated remodeling to form mechanically heterogeneous structures that promote MSC osteogenesis through mechanotransduction–epigenetic coupling. This work advances the design of biomaterials that respond adaptively to cellular cues, offering a step toward more physiologically relevant ECM mimetics.\nIn addition, we now explicitly acknowledge the current limitations of our work, including that the stiffness gradients are cell-regulated rather than pre-programmed with absolute precision, and that long-term in vivo stability and remodeling kinetics warrant further study.\nA new paragraph discussing the limitations now reads:\nWhile the CPAC hydrogel demonstrates robust osteoinductive performance, some limitations remain. The mechanical heterogeneity of the system arises from cell-secreted ALP activity rather than predefined gradients, and the precise spatial control of stiffness may vary with cell density or metabolic activity. In addition, long-term mechanical stability and remodeling behavior under complex in vivo conditions warrant further investigation.\n2. The abstract and Figure 1 contain unnecessary information and should be more compact to ensure good readability and clarity.\nAuthor Response:\nWe thank the reviewer for raising these important points. Now we have revised the abstract and Figure 1.\nThe Abstract now reads:\nStem cell differentiation dynamically remodels and stiffens the extracellular matrix (ECM), generating stage-specific biomechanical cues that guide tissue development. However, conventional biomaterials, designed to mimic mature ECM stiffness, neglect its spatiotemporal heterogeneity due to their static, non-evolvable nature. Herein, we develop a cell-programmed adaptative contraction (CPAC) hydrogel that enables mesenchymal stem cells (MSCs) to actively remodel their microenvironment through alkaline phosphatase (an early osteogenic marker)-mediated hydrophilic-to-hydrophobic transition and contraction of microgels. This cell-programmed remodeling establishes local mechanical heterogeneity and promotes osteogenesis through a positive feedback loop. Mechanistically, the evolving matrix enhances mechanotransduction-related microRNA expression, suppresses EZH2, and reduces H3K27 trimethylation to active osteogenic transcription. In vivo, MSC-laden CPAC hydrogels significantly enhance the repair of rat cranial defects. These findings introduce a paradigm of cell-instructed, dynamically evolvable biomaterials that recapitulate the adaptive nature of native ECM to orchestrate stem cell fate and tissue morphogenesis.\nThe Figure 1 now shows:\n3. The authors started with skull development (Figure 1) but later shifted to in vivo studies on bone regeneration. Note that development is not same as regeneration. Please correct to make it crystall-clear.\nAuthor Response:\nWe thank the reviewer for this thoughtful comment. We have corrected it now.\nThe Figure 1 now shows:\n4. The English language can be greatly improved.\nAuthor Response:\nWe thank the reviewer for the helpful comment to better address our manuscript. Now the English language of the revised manuscript has been significantly improved.\n5. New methods were only included for Fig S11 & Fig 5i/R1, please ensure all new data shown (esp. Figs S12-S24) are accompanied by the corresponding methods description, in particular:\na. Method for ALP quantification, Fig S12, S13\nAuthor Response:\nWe thank the reviewer for highlighting this important point. The ALP activity in the culture medium was quantified using a commercial colorimetric assay kit (Beyotime, Cat# P0321S) based on the hydrolysis of p-nitrophenyl phosphate (pNPP). Specifically, after the designated treatments, the culture medium was collected and centrifuged to remove debris. Then, 50 µL of the supernatant was mixed with 50 µL of the pNPP substrate working solution in a 96-well plate and incubated in the dark at 37 °C for 30 minutes. The enzymatic reaction was terminated by adding the stop solution provided in the kit. The absorbance of the yellow reaction product, p-nitrophenol, was immediately measured at 405 nm using a microplate reader. A standard curve was established using p-nitrophenol standards, and the ALP activity was calculated for each sample based on the absorbance readings. All assays were performed in triplicate.\nThe section of “The procedures for ALP activity quantification and the CCK-8 assay” in SI now reads:\nThe ALP activity in the culture medium was quantified using a commercial colorimetric assay kit (Beyotime, Cat# P0321S) based on the hydrolysis of p-nitrophenyl phosphate (pNPP). Specifically, after the designated treatments, the culture medium was collected and centrifuged to remove debris. Then, 50 µL of the supernatant was mixed with 50 µL of the pNPP substrate working solution in a 96-well plate and incubated in the dark at 37 °C for 30 minutes. The enzymatic reaction was terminated by adding the stop solution provided in the kit. The absorbance of the yellow reaction product, p-nitrophenol, was immediately measured at 405 nm using a microplate reader. A standard curve was established using p-nitrophenol standards, and the ALP activity was calculated for each sample based on the absorbance readings. All assays were performed in triplicate.\nb. Method for in vivo implantation model & sample processing for ALP, nanoindentation & mass quantification, Fig S13, S23. Also clarify what 0h in vivo means.\nAuthor Response:\nWe thank the reviewer for raising these important concerns. Method for in vivo implantation model & sample processing for ALP, nanoindentation & mass quantification:\nAccording to the previously described method for creating critical-size bone defects in rat calvaria, hydrogels loaded with rMSCs were implanted into the calvarial defect areas. The implanted hydrogels were retrieved at different time points: before implantation (0 h), and at 24 h and 72 h after implantation. The retrieved hydrogels were added to 1 mL of 1× PBS, thoroughly homogenized using a tissue grinder, and then centrifuged. The alkaline phosphatase (ALP) activity in the hydrogel was quantified using a commercial colorimetric assay kit (Beyotime, cat# P0321S) based on the hydrolysis of p-nitrophenyl phosphate (pNPP). The specific procedure was as follows: 50 µL of the supernatant was mixed with 50 µL of the pNPP substrate working solution in a 96-well plate and incubated in the dark at 37°C for 30 minutes. The enzymatic reaction was terminated by adding the stop solution provided in the kit. The absorbance of the yellow reaction product, p-nitrophenol, was immediately measured at 405 nm using a microplate reader. A standard curve was generated using p-nitrophenol standards, and the ALP activity was calculated for each sample based on the absorbance readings. All assays were performed in triplicate.\nFor nanoindentation testing, the extracted hydrogels were fixed onto glass slides. An 8×8 matrix of points, with each point spaced 50 μm apart, was randomly selected on the hydrogel surface. The effective Young's modulus at each point in the matrix was measured using a nanoindenter.\nFor the degradation experiment, the implanted hydrogels were retrieved at different time points, freeze-dried, and weighed (recorded as W ). Taking the mass of the freeze-dried hydrogel sample before implantation as W , the degradation rate of the hydrogel was calculated as follows:\n𝑊\n𝑋\n𝐷 = ×100%\n𝑊\nThe section of “Methods for in vivo implantation model & sample processing for ALP, nanoindentation & mass quantification” in SI now reads:\nAccording to the previously described method for creating critical-size bone defects in rat calvaria, hydrogels loaded with rMSCs were implanted into the calvarial defect areas. The implanted hydrogels were retrieved at different time points: before implantation (0 h), and at 24 h and 72 h after implantation. The retrieved hydrogels were added to 1 mL of 1× PBS, thoroughly homogenized using a tissue grinder, and then centrifuged. The alkaline phosphatase (ALP) activity in the hydrogel was quantified using a commercial colorimetric assay kit (Beyotime, cat# P0321S) based on the hydrolysis of p-nitrophenyl phosphate (pNPP). The specific procedure was as follows: 50 µL of the supernatant was mixed with 50 µL of the pNPP substrate working solution in a 96-well plate and incubated in the dark at 37 °C for 30 minutes. The enzymatic reaction was terminated by adding the stop solution provided in the kit. The absorbance of the yellow reaction product, p-nitrophenol, was immediately measured at 405 nm using a microplate reader. A standard curve was generated using p-nitrophenol standards, and the ALP activity was calculated for each sample based on the absorbance readings. All assays were performed in triplicate.\nFor nanoindentation testing, the extracted hydrogels were fixed onto glass slides. An 8×8 matrix of points, with each point spaced 50 μm apart, was randomly selected on the hydrogel surface. The effective Young's modulus at each point in the matrix was measured using a nanoindenter.\nFor the degradation experiment, the implanted hydrogels were retrieved at different time points, freeze-dried, and weighed (recorded as W ). Taking the mass of the freeze-dried hydrogel sample before implantation as W , the degradation rate of the hydrogel was calculated as follows:\n𝑊\n𝑋\n𝐷 = ×100%\n𝑊\nc. Method for “Remaining mass (%)” quantification, specifically if it was the dry mass, or if it was the wet mass (mostly water): how the results should be interpreted taking into account the bulk volume shrinkage as well.\nAuthor Response:\nWe thank the reviewer for the thoughtful suggestion. The remaining mass refers to the absolute dry mass of the hydrogel. Specifically, the absolute dry mass of the hydrogel before the degradation experiment is recorded as W0. After the hydrogel sample is taken out at specified time intervals and freeze-dried, it is weighed (recorded as W1). The remaining mass ratio (also called degradation rate (D)) is calculated as:\n𝑊\n𝑋\n𝐷 = ×100%\n𝑊\n0\nd. Method for bulk Young’s modulus quantification, e.g. Fig. S9b, S15\nAuthor Response:\nWe thank the reviewer for this helpful suggestion. The bulk Young's modulus of the hydrogel was measured using a rotational rheometer. Specifically, the hydrogel was prepared into cylindrical samples with a diameter of 8 mm and a thickness of 2 mm. The hydrogel was compressed from its initial state at a rate of 2 mm/min until a deformation of 15% was achieved. The Young's modulus was then calculated as the slope of the linear region of the resulting stress‑strain curve. Each sample group was tested independently three times to ensure reproducibility. Now we have added the relevant description in SI.\nThe section of “Fabrication and rheological characterization of the NC and CPAC hydrogels” in SI now reads:\nThe stress relaxation properties of the samples were measured from compression tests of the gel discs (8 mm in diameter, 2 mm thick). The gel discs were compressed to 15% strain with a deformation rate of 0.3 mm s-1 and held for 300 seconds to obtain the stress relaxation curve. The Young's modulus was then calculated as the slope of the linear region of the resulting stress‑strain curve. Each sample group was tested independently three times to ensure reproducibility.\ne. Methods for EdU incorporation assays, live/dead staining, and CCK-8 assays (incl. time points).\nAuthor Response:\nWe thank the reviewer for this valuable suggestion. The EdU-594 Cell Proliferation Detection Kit (Cat# C0078S) was purchased from Beyotime. A 2× EdU working solution (20 μM) was prepared by diluting EdU (10 mM) in culture medium. The working solution was mixed with an equal volume of fresh medium and added separately to well plates containing MSC-loaded NC and CPAC hydrogels. Following a 12‑h incubation, the medium was replaced with 1 mL of fixative for 15 min. After three PBS washes, 1 mL of permeabilization solution was applied for 10 min. Following another three PBS washes, 0.5 mL of Click reaction solution was added and incubated in the dark for 30 min. After three additional PBS washes, 1 mL of 1× Hoechst was added for 10 min in the dark. Finally, the samples were washed and subjected to fluorescence detection.\nFor live/dead staining, a staining solution was prepared in 1X PBS containing Calcein-AM and propidium iodide (PI), each at a final concentration of 1 µM. The hydrogels cultured for three days in a 24‑well plate were carefully removed and transferred to a new plate, and 1 mL of the prepared staining solution was added to each well. The plate was then incubated at 37 °C in the dark for 30 minutes. Following incubation, the hydrogels were washed three times with 1X PBS. Finally, imaging was performed using a confocal laser scanning microscope with detection channels set to 488 nm and 594 nm.\nCell viability was assessed using the Cell Counting Kit-8 (CCK-8; Beyotime, Cat# C0038). Following the designated treatments, 100 µL of the CCK-8 reagent was directly added to each well (900 µL culture medium with one hydrogel). The plate was then incubated at 37 °C in a humidified incubator with 5% CO₂ for 2 hours. After incubation, the absorbance of each well was measured at a wavelength of 450 nm using a microplate reader. Wells containing culture medium and CCK-8 reagent without cells were used as blanks for background subtraction. Cell viability was calculated as a percentage relative to the untreated cells (set as 100%), using the following formula:\n𝐴 − 𝐴\n𝑠 𝑏\n𝑉𝑖𝑎𝑏𝑖𝑙𝑖𝑡𝑦 (%) = ×100%.\n𝐴 − 𝐴\n𝑐 𝑏\nwhere A is the absorbance of the sample (treated cells), A is the absorbance of the control (untreated cells), and A is the absorbance of the blank. All experiments were performed with three independent replicates. Day 0 indicates the viability of MSCs before encapsulation, while day 3 represents the viability after MSCs loaded in NC and CPAC hydrogels for 3 days. Now we have added the relevant description in SI.\nThe section of “Methods for EdU incorporation assays and live/dead staining” in SI now reads:\nThe alkaline phosphatase (ALP) activity in the culture medium was quantified using a commercial colorimetric assay kit (Beyotime, cat# P0321S) based on the hydrolysis of p-nitrophenyl phosphate (pNPP). The specific procedure was as follows: After the designated treatments, the culture medium was collected and centrifuged. Then, 50 µL of the supernatant was mixed with 50 µL of the pNPP substrate working solution in a 96-well plate and incubated in the dark at 37 °C for 30 minutes. The enzymatic reaction was terminated by adding the stop solution provided in the kit. The absorbance of the yellow reaction product, p-nitrophenol, was immediately measured at 405 nm using a microplate reader. A standard curve was generated using p-nitrophenol standards, and the ALP activity was calculated for each sample based on the absorbance readings. All assays were performed in triplicate.\nFor live/dead staining, a staining solution was prepared in 1× PBS containing Calcein-AM and propidium iodide (PI), each at a final concentration of 1 µM. The hydrogels cultured for three days in a 24‑well plate were carefully removed and transferred to a new plate, and 1 mL of the prepared staining solution was added to each well. The plate was then incubated at 37 °C in the dark for 30 minutes. Following incubation, the hydrogels were washed three times with 1× PBS. Finally, imaging was performed using a confocal laser scanning microscope with detection channels set to 488 nm and 594 nm.\nThe section of “Procedures for ALP activity quantification and the CCK-8 assay” in SI now reads:\nCell viability was assessed using the Cell Counting Kit-8 (CCK-8; Beyotime, Cat# C0038). Following the designated treatments, 100 µL of the CCK-8 reagent was directly added to each well (900 µL culture medium with one hydrogel). The plate was then incubated at 37°C in a humidified incubator with 5% CO₂ for 2 hours. After incubation, the absorbance of each well was measured at a wavelength of 450 nm using a microplate reader. Wells containing culture medium and CCK-8 reagent without cells were used as blanks for background subtraction. Cell viability was calculated as a percentage relative to the untreated cells (set as 100%), using the following formula:\n𝑉𝑖𝑎𝑏𝑖𝑙𝑖𝑡𝑦 (%) = 𝐴𝑠 − 𝐴𝑏 ×100%. (5)\n𝐴𝑐 − 𝐴𝑏\nwhere A is the absorbance of the sample (treated cells), A is the absorbance of the control (untreated cells), and A is the absorbance of the blank. All experiments were performed with three independent replicates. Day 0 indicates the viability of MSCs before encapsulation, while day 3 represents the viability after MSCs loaded in NC and CPAC hydrogels for 3 days.\nf. Method for alkaline phosphatase color development kit.\nAuthor Response:\nWe thank the reviewer for this helpful reminder. After fixation with paraformaldehyde, the hydrogel was cryosectioned into 8 μm thin slices. The resulting slices were washed three times with PBS. A working solution was prepared according to the manufacturer's instructions of the BCIP/NBT Alkaline Phosphatase Color Development Kit (Beyotime, cat# C3206). The sections were fully covered with the working solution and incubated at room temperature in the dark for 30 minutes. The color development reaction was terminated by washing twice, and images were captured using an inverted microscope. Now we have added the relevant description in SI.\nThe section of “Method for alkaline phosphatase color development kit” in SI now reads:\nAfter fixation with paraformaldehyde, the hydrogel was cryosectioned into 8 μm thin slices. The resulting slices were washed three times with PBS. A working solution was prepared according to the manufacturer's instructions of the BCIP/NBT Alkaline Phosphatase Color Development Kit (Beyotime, cat# C3206). The sections were fully covered with the working solution and incubated at room temperature in the dark for 30 minutes. The color development reaction was terminated by washing twice, and images were captured using an inverted microscope.\n6. Follow-up to previous round Question 12: It is unclear how to deduce effective Young’s moduli values from all the nanoindentation heat map graphs. The values in the graphs don’t match the values of the color scale legend (e.g. Fig 2i(iii): graph at X(0), Y(300) shows ~1000Pa but color indicates ~4000Pa). Please double-check or clarify the meaning of the color scale values (they are linear instead of logarithmic as in graph).\nAuthor Response:\nWe thank the reviewer for the important comments. When creating the graph, we set the Z-axis to a logarithmic scale, but the color bar was not adjusted accordingly and remained linear. We apologize for this oversight. The color bar has now been updated entirely to a logarithmic scale. Thank you for bringing this to our attention.\nThe Figure 2i(iii) now shows:\n7. The new Figure 1b is much clearer now, but the text might be too small for publication. Also, the figure captions needs updating.\nAuthor Response:\nWe appreciate the reviewer’s comments. Now we have updated the figure caption.\nThe Figure 1b caption now reads:\nFigure 1. (b) Schematic showing the network heterogenization of MSCs-laden NC hydrogel (left), NCLS hydrogel (middle) and CPAC hydrogel (right) after ALP exposure.\n8. Figure S17c: Recommended to labels as “Relative metabolic activity (%) “or similar (not viability) when used as a proliferation assay.\nAuthor Response:\nWe greatly appreciate your insightful question. Now we have corrected it.\nThe Figure S17c now shows:\n9. Regarding previous round Question 26: It seems like the method section has not been adapted to reflect the experimental procedure described in your response. The SI still reads:\n“Skull specimens were initially subjected to micro-computed tomography (micro-CT) scanning, followed by decalcification in 10% ethylenediaminetetraacetic acid (EDTA, pH 7.4) for two months in preparation for immunohistochemical evaluation. […] Following decalcification, specimens were fixed in 4% formaldehyde for 24 hours”\nwhich indicates the sequence 1) decalcification, 2) fixation. Maybe you mean: „Before decalcification, specimens were fixed in 4% formaldehyde for 24 hours“?.\nAuthor Response:\nWe appreciate the reviewer’s valuable comment. We apologize for the ambiguity in our original description. In our experiments, the skull specimens were first fixed in 4% paraformaldehyde for 48 hours to ensure proper preservation of tissue morphology and antigens. Following fixation, the samples underwent mild decalcification with 10% EDTA solution (pH 7.4, 25 °C) for 6 weeks.\nThe corresponding paragraphs in SI now reads:\nImmunohistochemical staining\nThe skull specimens were first fixed in 4% paraformaldehyde for 48 hours to ensure proper preservation of tissue morphology and antigens. Following fixation, the samples underwent mild decalcification with 10% EDTA solution (pH 7.4, 25 °C) for 6 weeks. Tissue dehydration was performed through a graded ethanol series (30-100%), with subsequent xylene clearing and paraffin embedding. Serial sections (4 μm thickness) were prepared for histological evaluation.\n10. Fig S18: the results should be normalized to cell number.\nAuthor Response:\nWe appreciate the reviewer’s helpful suggestion. Now the results have been normalized to cell number.\nThe Figure S18b now shows:\n11. Methods for quantification of micro-CT data should be updated to ensure reproducibility.\nAuthor Response:\nWe thank the reviewer for the suggestion. Now the methods for quantification of micro-CT data have been updated to the supporting information.\nThe corresponding paragraphs in SI now reads:\nQuantification of micro-CT\nMicro-CT Scanning and Analysis: Upon euthanasia, rat calvarial bones were isolated and fixed in 4% paraformaldehyde. Fixed samples were scanned using a Micro-CT system (Quantum GX2, Perkinelmer AG) under the following conditions: 70 kV for 140 ms with a resolution setting of 25 lm and 1024×1024 pixels. Three-dimensional reconstruction and subsequent data analysis of the defect region were performed to quantify bone regeneration. Key outcomes included bone volume (BV), bone volume/tissue volume (BV/TV) and bone surface/tissue volume (BS/TV).\n12. Figure R2 (response to Q4) should be added to at least SI, given its importance to support the in vivo findings.\nAuthor Response:\nWe appreciate the reviewer’s comment. Now we have added it in SI as Figure S26.\nThe Figure S26 now reads:\nFigure S26. (a) Micro-CT images showing bone defects at 8 weeks post-implantation for the Control and CPAC hydrogel groups. Quantification of (b) bone volume (BV), (c) bone volume/total volume (BV/TV), and (d) bone surface area/total volume (BS/TV) in the bone defect sites after 8 weeks, respectively (n = 3). (e) H&E staining, (f) Masson’s trichrome staining of the bone defects in the Control and CPAC hydrogel groups, respectively (Scale bar = 200 μm). (g) OCN immunohistochemical staining in the Control and CPAC hydrogel groups (Scale bar = 100 μm). Data are presented as mean ± SD; **p < 0.01, ***p < 0.001.\n13. Figure 3 & 5: WesternBlott: please explain why proteins of interests and reference protein are analyzed separately, instead of 1 combined image.\nAuthor Response:\nWe thank the reviewer for raising these important concerns. The Western blot (WB) experiment was performed on the same gel. However, after electrophoresis, the gel was carefully cut into different sections corresponding to the molecular weight ranges of the target proteins and the reference protein (β-actin or GAPDH) for separate antibody incubation and exposure. This approach was adopted because different proteins often require distinct incubation conditions and exposure times for optimal signal detection. Running and exposing them separately avoids overexposure or underexposure of certain bands and prevents antigen loss that can occur during repeated stripping and reprobing. Therefore, the separate presentation of the blots more accurately reflects the true protein expression levels under each condition.\n14. Figure 4: it is unclear why the authors showed only single cells instead of showing tens of cells per image. How many cells in total have been analyzed?\nAuthor Response:\nWe thank the reviewer for this valuable comment. High-resolution single-cell images were presented to clearly display the detailed morphology and cytoskeletal organization of individual MSCs, which would otherwise be obscured in low-magnification images containing dense cell clusters. For quantitative analysis, 15 cells per experimental group were randomly selected and analyzed to ensure statistical rigor and reproducibility. Representative single-cell images were shown in the figures for clarity, while the quantified results are based on the full dataset."}]

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