Coordination‐Driven Cu²⁺‐Peptide Supramolecular Hydrogel‐PCL Scaffold for Synergistic Antibacterial Activity and Osteogenic Regeneration in Infectious Bone Defects

ABSTRACT

Infectious bone defects, characterized by persistent bacterial infection and impaired osteogenesis, remain a major clinical challenge. Here, a copper ion‐peptide supramolecular hydrogel integrated with a 3D‐printed polycaprolactone scaffold (CPSH@PCL) was developed for synergistic treatment of infectious bone defects. Cu²⁺‐mediated peptide coordination formed a stable nanofibrous network, enabling robust scaffold coating, enhanced mechanical support, and sustained Cu²⁺ release under both physiological and infection‐associated mildly acidic conditions. CPSH@PCL exhibited potent broad‐spectrum antibacterial and antibiofilm activity (>99% against methicillin‐resistant Staphylococcus aureus and Pseudomonas aeruginosa) and disrupted mature biofilms. It also scavenged reactive oxygen species and catalyzed oxygen generation, alleviating infection‐induced oxidative stress and hypoxia. At the cellular level, CPSH@PCL promoted proliferation and migration of bone marrow mesenchymal stem cells and osteoblasts, while markedly enhancing osteogenic differentiation through activation of HIF‐1α‐mediated aerobic glycolysis. In a rat cranial defect model, CPSH@PCL suppressed infection, significantly increased bone mineral density and bone volume fraction, and accelerated new bone formation with upregulated osteocalcin. This work demonstrated a metal ion‐driven peptide self‐assembly strategy that integrated antibacterial, antioxidant, and osteogenic functions in a single scaffold, offering a multifunctional biomaterial platform for treating complex infectious bone defects.

A copper–peptide supramolecular hydrogel (CPSH) integrated with a 3D‐printed PCL scaffold (CPSH@PCL) was developed for infectious bone defect repair. The composite scaffold exhibited antibacterial, antibiofilm, and antioxidant activities, while promoting bone regeneration by reprogramming glycolytic metabolism in bone mesenchymal stem cells (BMSCs), offering a multifunctional strategy for infection‐compromised bone healing.

1. Introduction

Bone defects complicated by bacterial infection are a major challenge in clinical orthopedics [1]. They typically arise from severe trauma, open fractures, or surgical intervention and are characterized by the coexistence of persistent infection and extensive bone loss [2]. The pathological microenvironment features continuous bacterial biofilm formation, severe inflammation, excessive reactive oxygen species (ROS) production, and impaired host osteogenic capacity, all of which synergistically hinder tissue repair [3, 4]. Current clinical management mainly relies on systemic antibiotics combined with surgical debridement [5]. However, the emergence of methicillin‐resistant Staphylococcus aureus (MRSA) and other multidrug‐resistant strains [6], the poor penetration of antibiotics into mature biofilms, systemic drug toxicity, and the lack of inherent osteogenic activity in most implant materials lead to unsatisfactory outcomes [7]. Recent progress in bone tissue engineering has underscored the importance of multifunctional scaffolds that can simultaneously regulate infection, inflammation, oxidative stress, angiogenesis, and osteogenic remodeling within the defect microenvironment [8]. However, for infectious bone defects, it remains highly challenging to achieve sustained bacterial eradication while maintaining a microenvironment conducive to tissue regeneration. At present, no universally recognized optimal strategy exists for infectious bone defects, and achieving simultaneous infection control and bone regeneration is still an unmet clinical need.

Antimicrobial peptides (AMPs), which constitute an important component of the innate immune defense system, have attracted increasing attention as potential alternatives to conventional antibiotics [9]. Unlike conventional antibiotics, which typically act on specific metabolic or enzymatic targets, many antimicrobial peptides rapidly kill bacteria by interacting with negatively charged bacterial membranes and inducing membrane permeabilization, pore formation, or membrane destabilization [10]. This membrane‐targeting, multi‐site mechanism confers broad‐spectrum antibacterial activity against both Gram‐positive and Gram‐negative bacteria and lowers the probability of resistance development [11]. Moreover, AMPs and their derivatives have been reported to interfere with bacterial adhesion, inhibit biofilm formation, and disrupt established biofilms, which is particularly relevant to implant‐associated infections and infectious bone defects [12]. Nevertheless, despite these advantages, the direct use of free AMPs is still constrained by several inherent limitations, including poor proteolytic stability, short in vivo half‐life, potential cytotoxicity arising from insufficient membrane selectivity, reduced activity under complex physiological conditions such as high‐salt or serum‐containing environments, rapid diffusion away from infected sites, and relatively high production costs [13, 14]. These limitations hinder free AMPs from providing sustained antibacterial protection during the prolonged and complex repair process of infectious bone defects.

Therapeutic metal ions have attracted significant attention due to their broad‐spectrum antibacterial activity with immunoregulatory and osteogenic functions [15]. Recent perspectives on metal‐ion‐based therapeutics have further emphasized that bioactive metal ions can serve as multifunctional regulators in regenerative medicine by modulating antibacterial activity, redox homeostasis, angiogenesis, immune responses, and osteogenesis [16]. Unlike conventional antibiotics, metal ions inhibit pathogens through multiple mechanisms, including disruption of bacterial membranes and interference with metabolic pathways, thereby reducing the risk of antibiotic resistance [17]. Among them, copper ions, an essential trace element, exhibit particularly strong activity against Gram‐positive and Gram‐negative bacteria, and fungi and can effectively disrupt biofilms, reducing infection recurrence and resistance development [18]. Nearly 300 copper‐based antibacterial materials are registered with the U.S. Environmental Protection Agency and are reported to kill up to 99.9% of pathogens, underscoring the robustness of copper‐based antimicrobial strategies [19]. Furthermore, Cu²⁺ can promote collagen maturation, enhance osteogenic differentiation and mineralization, and ultimately accelerate bone regeneration [20].

Despite these advantages, most existing metal‐ion‐encapsulated systems still suffer from burst release, limited structural stability, and poor coupling between the carrier microstructure and metal‐mediated functionality [21, 22]. Metal ions are typically introduced through surface adsorption or simple chelation (e.g., with carboxyl or histidine side chains), which provides poor control over release kinetics and compromises long‐term performance [23]. Therefore, there is an urgent need for functionalization strategies that can achieve stable loading, controlled release, and intimate integration of metal ions with the carrier architecture. In this context, metal‐ion‐coordinated hydrogels have recently emerged as an attractive strategy because coordination interactions can simultaneously act as dynamic crosslinking motifs, ion reservoirs, and bioactive regulatory units [24].

Hydrogels are attractive candidates for this purpose owing to their high‐water content, extracellular matrix (ECM)‐like porous microstructure, and excellent mass‐transport properties, which collectively support sustained ion release [25]. Peptide‐based self‐assembled hydrogels have outstanding advantages, including sequence designability, excellent biocompatibility, and the ability to present bioactive motifs as well as well‐defined coordination sites, thereby mitigating burst release [26]. Recent studies have further demonstrated that self‐assembling peptide hydrogels can serve as multifunctional supramolecular platforms for tissue regeneration, antibacterial therapy, immunomodulation, and the sustained delivery of bioactive agents [27, 28, 29]. However, the limited mechanical strength of hydrogels alone is often insufficient for load‐bearing bone repair [30]. In contrast, polycaprolactone (PCL) scaffolds possess excellent mechanical robustness and tunable biodegradability but lack antibacterial and osteoinductive functions [31]. In addition, recent discussions on implants have emphasized that long‐term biosafety, degradation‐associated ion release, batch‐to‐batch reproducibility, sterilization, scale‐up manufacturing, and regulatory evaluation should be carefully considered before clinical translation [32]. Combining peptide hydrogels with PCL can therefore integrate structural support with biological activity. Incorporating Cu²⁺ into the peptide hydrogel further endows the composite with potent antibacterial and osteogenic capabilities tailored to the complex microenvironment of infectious bone defects.

In this study, we constructed a coordination‐driven supramolecular hydrogel‐scaffold system for infectious bone defects. Consistent with emerging design principles of smart biomaterials for bone regeneration, this system aimed to integrate infection control, oxidative stress regulation, and osteogenic stimulation within a single scaffold platform [33]. A customized peptide Ac‐FEFEFE‐NH₂ (FE3), was coordinated with Cu²⁺ to form a mechanically reinforced Cu²⁺‐peptide supramolecular hydrogel (CPSH), which was subsequently infiltrated into and coated onto a 3D‐printed PCL scaffold, yielding a stable scaffold CPSH@PCL (Figure 1). CPSH@PCL achieved sustained Cu²⁺ release under both physiological and infection‐associated mildly acidic conditions, enabling biofilm disruption and inhibition of drug‐resistant bacteria, while its antioxidant activity scavenged local ROS and improved the metabolic microenvironment through regulation of glycolysis, thereby promoting osteogenic differentiation and maturation of bone marrow‐derived mesenchymal stem cells (BMSCs). This coordination‐driven CPSH@PCL provided a rational design framework for multifunctional scaffolds targeting infectious bone defects and offered a promising candidate for clinical applications requiring both robust antibacterial and osteogenic performance.

FIGURE 1.

Schematic illustration of the construction of copper ion‐peptide (FE3) supramolecular hydrogel (CPSH) integrated with a 3D‐printed polycaprolactone scaffold (CPSH@PCL) for the treatment of infectious cranial bone defects, and its underlying mechanisms enabling robust antibacterial activity and enhanced osteogenic regeneration. Figure created with BioRender.com.

2. Results and Discussion

2.1. Preparation of CPSH and CPSH@PCL

Coordination‐mediated supramolecular self‐assembly was employed as the central design strategy to construct structurally stable, bioactive hydrogels. The FE3 peptide was first synthesized (Figure S1), and coordination systems with different peptide: Cu²⁺ molar ratios (1:0.5, 1:1, 1:1.5; designated CPSH‐0.5, CPSH‐1, CPSH‐1.5, respectively) were prepared (Figure S2). Among these formulations, CPSH‐1.5 exhibited the most stable self‐supporting gel state after standing at room temperature for 30 min, whereas the lower‐Cu²⁺ groups showed weaker gelation or residual fluidity. Therefore, CPSH‐1.5 was selected as the standard hydrogel formulation (hereafter referred to as CPSH). Macroscopically, CPSH maintained its integrity when lifted with tweezers (Figure 2A), indicating sufficient mechanical rigidity. CPSH was then incorporated into a 3D‐printed PCL scaffold using an impregnation method (Figure 2B), resulting in a CPSH@PCL composite with a uniform blue‐green hydrogel coating on the scaffold surface and within the pores (Figure 2D). In contrast, the FE3 peptide solution without Cu²⁺ failed to form a stable coating on PCL (Figure 2C), underscoring the essential role of Cu²⁺‐mediated self‐assembly in gel formation and scaffold integration.

FIGURE 2.

Morphological characterization and molecular interactions of copper ion‐peptide supramolecular hydrogel (CPSH) and CPSH‐integrated polycaprolactone (PCL) scaffold (CPSH@PCL). (A) Photograph of CPSH. Optical microscopy images of scaffolds: (B) pristine PCL; (C) CPSH@PCL; (D) peptide‐loaded PCL (P@PCL). (E) Transmission electron microscope (TEM) image of CPSH (scale bar: 500 nm). (F) Scanning electron microscope (SEM) image of CPSH (scale bar: 20 µm). (G) SEM image of CPSH@PCL (scale bar: 200 µm); (H) SEM image of the pristine PCL scaffold (scale bar: 100 µm). (I) Schematic illustration of the molecular interactions: CPSH is stabilized by Cu²⁺‐peptide coordination and interchain π‐π stacking, while CPSH@PCL is further formed through hydrophobic interactions between the hydrogel and the PCL matrix.

2.2. Structural and Physicochemical Characterization of CPSH and CPSH@PCL

Transmission electron microscopy (TEM) revealed a regularly distributed fibrous network within CPSH, with continuous, uniformly distributed nanofibers interweaving into a three‐dimensional framework (Figure 2E). Scanning electron microscopy (SEM) further confirmed a highly porous ECM‐mimetic architecture (Figure 2F). Such nanoscale structural characterization is important for hydrogel‐based biomaterials because nanofiber morphology, mesh‐like architecture, and interfacial organization can strongly influence ion release, protein adsorption, cell‐material interactions, and subsequent biological responses [34]. CPSH@PCL exhibited a rough, adherent hydrogel coating that fully wetted and conformed to the scaffold struts (Figure 2H). In contrast, pristine PCL displayed a smooth surface (Figure 2G), indicating intimate and uniform integration of CPSH with the PCL scaffold (Figure 2I).

CPSH coating markedly enhanced the surface hydrophilicity of PCL (Figure 3A), a property favorable for cell adhesion, spreading, and penetration. Rheological measurements showed that, over the frequency range of 0.1–10 rad s⁻¹, the storage modulus (G′) consistently exceeded the loss modulus (G″), both above 1000 Pa, indicating typical solid‐like gel characteristics. The complex viscosity demonstrated efficient energy dissipation and a stable cross‐linked network (Figure 3B). It should be noted that these rheological data characterize the intrinsic viscoelastic properties of CPSH rather than the compressive mechanical performance of the CPSH@PCL composite scaffold. In the present design [35], the 3D‐printed PCL framework served as the primary structural‐supporting component, whereas the CPSH coating mainly provided antibacterial, antioxidant, and osteogenic bioactivity. This design rationale was consistent with our previous studies on self‐assembling peptide hydrogel/PCL composite scaffolds. In a SAPH‐PCL composite scaffold developed for rabbit segmental bone defect repair, the composite scaffolds exhibited stress points at approximately 220 N, corresponding to a modulus of approximately 11.2 MPa, and SAPH infilling did not noticeably alter the mechanical characteristics of the PCL scaffold. In addition, SAPH‐coated PCL scaffolds were previously shown to provide a stable biomimetic ECM‐like microenvironment and to support osteochondral regeneration while retaining the structural‐supporting role of the PCL framework.

FIGURE 3.

Performance and internal structure characterization of the copper‐peptide supramolecular hydrogel (CPSH). (A) Water contact angle measurements of pristine polycaprolactone (PCL) and CPSH@PCL. (B) Rheological properties of CPSH: left, storage modulus (G') and loss modulus (G'') as a function of frequency; right, complex viscosity with frequency. (C) Zeta potential of FE3 peptide, Cu²⁺, and CPSH. (D) Circular dichroism (CD) spectra of CPSH and peptide. (E) Fourier‐transform infrared (FTIR) spectra of CPSH and peptide. (F) Schematic illustration of the CPSH self‐assembly process obtained from molecular dynamics simulation. (G–I) Structural parameters of CPSH during molecular dynamics simulation: (G) root mean square deviation (RMSD) versus simulation time; (H) radius of gyration (Rg) versus simulation time; (I) solvent accessible surface area (SASA) versus simulation time.

Zeta potential analysis revealed that FE3 carried a high negative charge (−50 mV), while Cu²⁺ was positively charged (∼+10 mV). After coordination self‐assembly, the zeta potential of CPSH increased to −14.55 mV (Figure 3C), reflecting partial neutralization of carboxylate groups by Cu²⁺ and a transition from a highly negative to a more moderately charged, stable state. This charge modulation is consistent with Cu²⁺ binding to side‐chain carboxyl groups.

Circular dichroism (CD) spectroscopy was used to probe the assembly‐state changes of FE3 after Cu²⁺ coordination (Figure 3D). The FE3 peptide showed a negative band near 195 nm and positive CD signals above 200 nm, indicating that the peptide adopted a defined chiral assembly state in solution. After coordination with Cu²⁺, the CD signal in the 215–235 nm region decreased markedly, together with attenuation of the negative band near 195 nm. These changes suggest that Cu²⁺ coordination altered the supramolecular packing mode and reorganized the chiral assembly of FE3 during hydrogel formation. Fourier transform infrared (FT‐IR) spectroscopy further confirmed coordination between Cu²⁺ and peptide carboxyl groups (Figure 3E). The asymmetric stretching vibration of the carboxylate group [ν_{a s}(COO⁻)] shifted from 1531 to 1546 cm⁻¹, accompanied by the disappearance of the band at 1444 cm⁻¹. The corresponding Δν value (ν_{a s}—ν_s), calculated as 139 cm⁻¹ relative to the symmetric stretching vibration at 1407 cm⁻¹, indicated a modified coordination environment dominated by bridging or bidentate carboxylate interactions rather than monodentate binding (Figure S3). Taking potential peak overlap into consideration, this Δν value was indicative of a coordination mode dominated by bridging or bidentate carboxylate interactions rather than a purely monodentate configuration. The C─OH stretching band at 1238 cm⁻¹, presented in FE3, vanished in CPSH, indicating carboxyl deprotonation and a bidentate coordination. In addition, the amide I band shifted from 1633 to 1641 cm⁻¹, consistent with tighter peptide chain stacking after gelation. Together, these data supported a Cu²⁺‐driven coordination‐crosslinked network formed through carboxylate binding.

Cu²⁺ release from CPSH@PCL was evaluated under physiological (pH 7.4) and simulated infection‐associated acidic (pH 6.2) conditions (Figure S4). A typical biphasic profile was observed, characterized by an initial burst release within 24 h followed by sustained release over 216 h. Comparable release kinetics were obtained under both pH conditions, with only a modest increase in Cu^{2 +} release under acidic conditions after 72 h, statistical analysis showed no significant difference in cumulative Cu²⁺ release between pH 6.2 and pH 7.4 at all tested time points, indicating that Cu^{2 +} release from CPSH@PCL is only weakly influenced by pH within this range.

Matrix‐assisted laser desorption/ionization time‐of‐flight mass spectrometry (MALDI‐TOF MS) further confirmed the formation of FE3‐Cu²⁺ coordination complexes. In the negative ion mode, the deprotonated FE3 peptide peak was observed at m/z 886.2062 and assigned to [M‐H]⁻. A sodium‐adducted peptide peak appeared at m/z 908.1886, corresponding to [M‐2H+Na⁺]⁻. Importantly, two Cu²⁺‐containing peaks were detected at m/z 948.1239 and 970.1019, which were assigned to [M‐2H+Cu²⁺]⁻ and [M‐3H+Cu²⁺+Na⁺]⁻, respectively. The calculated and detected m/z values showed small deviations of approximately 0.16 Da (Table S1), supporting the reliability of the peak assignments. These results confirmed the formation of FE3‐Cu²⁺ coordination complexes (Figure S5). To further elucidate its self‐assembly mechanism, molecular dynamics (MD) simulations were performed in GROMACS with 20 FE3 and 30 Cu²⁺ ions (in a 2:3 ratio). FE3 and Cu²⁺ aggregated within 30 ns, reached a relatively stable configuration by 60 ns, and remained stable to 90 ns (Figure 3F). Cu²⁺ preferentially coordinated with peptide side chain carboxylates, consistent with experimental data.

Root mean square deviation (RMSD), radius of gyration (Rg), and solvent‐accessible surface area (SASA) all plateaued after ∼50 ns (Figures 3G–I). After stabilization, RMSD and Rg were 4.168 ± 0.02 and 1.808 ± 0.05 nm, respectively, while SASA decreased from ∼240 to 108.6 ± 3.19 nm², suggesting compact cluster formation. The binding energy between Cu²⁺ and FE3 was calculated using the MMPBSA method [36], and the results are presented in Table S2. Binding energy calculations showed that electrostatic interactions are dominant. The binding energy between Cu²⁺ and FE3 in the system is determined to be −5613.12 kJ mol⁻¹. Overall, the simulations corroborated a stable, Cu²⁺‐driven coordination assembly consistent with the experimental observations.

2.3. In Vitro Antibacterial Properties of CPSH@PCL

Antimicrobial resistance, especially in biofilm‐forming pathogens such as MRSA and P. aeruginosa, severely compromises current therapies for infectious bone defects. We therefore selected MRSA (Gram‐positive) and P. aeruginosa (Gram‐negative) as representative strains to evaluate the antibacterial performance of CPSH@PCL.

Plate colony counting revealed that, after co‐culture with the scaffolds, almost no colonies were detected in eluates from CPSH@PCL, whereas abundant bacterial growth persisted in the PCL, P@PCL, and blank control group (Figure 4A,B). Consistently, OD600 measurements showed a >99% reduction in bacterial growth for CPSH@PCL against MRSA and P. aeruginosa after 12 h compared with the control groups (p < 0.0001) (Figure 4C,D). MTT assays further confirmed that CPSH@PCL reduced bacterial metabolic activity by over 82%, while PCL and P@PCL showed negligible inhibition (Figure 4E,F). These data indicated that the potent bactericidal effect arose from the sustained release of Cu²⁺ within the CPSH network rather than from PCL or peptide alone.

FIGURE 4.

Antibacterial and anti‐biofilm performance of the copper‐peptide supramolecular hydrogel (CPSH) against methicillin‐resistant Staphylococcus aureus (MRSA) and Pseudomonas aeruginosa. (A,B) Standard plate counts of MRSA (A) and P. aeruginosa (B) after 12 h of co‐culture with each group (BLK, PCL, P@PCL, CPSH@PCL). (C,D) Bacterial growth (OD600) of MRSA (C) and P. aeruginosa (D) after 12 h cultivation with each group. (E,F) Metabolic activity (MTT assay) of MRSA (E) and P. aeruginosa (F) at different time points. (G–J) Crystal violet staining and quantitative analysis of biofilm inhibition for MRSA (G, I) and P. aeruginosa (H, J). (K–N) Crystal violet staining assay and quantitative analysis of mature biofilm removal for MRSA (K, M) and P. aeruginosa (L,N). Data are presented as mean ± SD, n = 4. **** p < 0.0001, ns, no significance.

Given the central role of biofilms in refractory infections, we next examined the antibiofilm activity of CPSH@PCL. In a 72‐h biofilm formation assay, crystal violet staining showed that CPSH@PCL almost completely prevented biofilm attachment by both strains (Figure 4G,H), which was corroborated by significantly reduced OD570 values after dye dissolution (Figure 4I,J). In a 72‐h biofilm eradication assay using preformed mature biofilms, CPSH@PCL also demonstrated excellent disruption capability: staining indicated extensive matrix removal (Figure 4K,L) and quantitative OD570 analysis (Figure 4M,N) showed a marked decrease in residual biofilm biomass. These results demonstrated that sustained, pH‐responsive Cu²⁺ release effectively inhibited initial bacterial adhesion and penetrated the extracellular polymeric substance matrix of mature biofilms, leading to efficient biofilm inhibition and removal.

Collectively, P@PCL did not show obvious antibacterial activity compared with pristine PCL, whereas CPSH@PCL markedly inhibited bacterial proliferation and biofilm formation. This comparison suggests that the antibacterial activity of CPSH@PCL is mainly derived from Cu²⁺ coordination rather than the FE3 peptide hydrogel matrix alone. CPSH@PCL exhibited robust broad‐spectrum antibacterial and antibiofilm activity, primarily through sustained membrane‐disruptive action of released Cu²⁺ ions, while the superhydrophilic CPSH coating further suppressed initial bacterial adhesion. This dual mode of antibacterial mechanism positioned CPSH@PCL as a promising multifunctional platform for the management of infectious bone defects and other challenging orthopedic infections.

2.4. In Vitro Antioxidant Properties of CPSH@PCL

Infectious bone defects are typically accompanied by excessive reactive oxygen species (ROS), which exacerbate inflammation, interfere with tissue regeneration, and induce cell damage [37]. To determine whether CPSH@PCL could simultaneously provide antibacterial and antioxidant functions, we evaluated both its catalytic ROS‐scavenging activity and its cytoprotective effects.

To clarify the origin of the enzyme‐mimetic activity, P@PCL and Cu@PCL were introduced as additional controls. In the qualitative H₂O₂‐triggered bubble‐generation assay, pristine PCL and P@PCL showed no obvious bubble generation after exposure to H₂O₂ (Figure S6A,B), whereas visible bubbles were observed in both Cu@PCL and CPSH@PCL groups (Figure S6C,D). These results indicated that the H₂O₂ decomposition‐related oxygen‐generation behavior was mainly associated with the presence of Cu²⁺, and that Cu²⁺ coordinated within the CPSH network retained catalytic activity. Consistently, in the TMB colorimetric assay, CPSH@PCL induced a significant increase in OD450 only in the presence of both H₂O₂ and TMB (Figure S7A). P@PCL showed negligible catalytic response under the same H₂O₂/TMB conditions (Figure S7B), whereas Cu@PCL markedly enhanced TMB oxidation (Figure S7C), confirming that Cu²⁺ served as the major catalytic active center. These results suggest that the enzyme‐mimetic activity of CPSH@PCL was mainly derived from Cu²⁺, while the FE3 peptide supramolecular network functioned as a coordination matrix to stabilize Cu²⁺ and regulate its catalytic presentation.

We first evaluated the cytotoxicity of free Cu²⁺. To maintain consistency with the scaffold extract culture system, free Cu²⁺ was prepared under the same extraction conditions as CPSH@PCL. The MTT results showed that free Cu²⁺ exhibited evident cytotoxicity, indicating that direct exposure to free Cu²⁺ was not suitable for subsequent cell‐based functional assays (Figure S8). In contrast, CPSH@PCL maintained good cytocompatibility through sustained Cu²⁺ release, suggesting that the peptide coordination network and scaffold‐based delivery effectively mitigated Cu²⁺‐associated cytotoxicity.

To assess cellular protection, BMSCs were pretreated with H₂O₂ to induce oxidative stress and then cultured with extracts from different scaffolds. Intracellular ROS levels measured by DCFH‐DA staining were markedly lower in the CPSH@PCL group than in the positive control and other groups (Figure 5A), indicating effective scavenging of intracellular ROS and improved tolerance of BMSCs to oxidative damage.

FIGURE 5.

Antioxidant performance and underlying mechanism of copper‐peptide supramolecular hydrogel‐loaded PCL composite scaffold (CPSH@PCL). (A) Intracellular reactive oxygen species (ROS) levels in BMSCs by DCFH‐DA staining. (B–D) Relative mRNA expression of antioxidant‐related genes SIRT1 (B), SOD2 (C), CAT (D) was determined by RT‐qPCR. Data are presented as mean ± SD, n = 4. *p<0.05, ** p < 0.01, *** p < 0.001, **** p <0.0001.

RT‐qPCR analysis further clarified the underlying mechanisms. Relative mRNA expression was normalized to Actin and calculated using the 2^−ΔΔCt method. BMSCs cultured with CPSH@PCL extract showed significant upregulation of key antioxidant genes compared with the control groups: SIRT1 increased approximately fivefold, SOD2 increased approximately sevenfold, and CAT also increased approximately sevenfold (p < 0.0001) (Figure 5B–D). These results indicated that CPSH@PCL not only exhibited direct catalase‐like activity but also activated the SIRT1‐SOD2‐CAT axis in BMSCs, thereby enhancing endogenous antioxidant defense.

Thus, CPSH@PCL exerted antioxidant protection through a dual‐pathway mechanism: on the one hand, its intrinsic enzyme‐mimicking activity directly catalyzed the decomposition of reactive oxygen species (ROS); on the other hand, it activated the intracellular SIRT1‐SOD2‐CAT signaling axis, thereby enhancing endogenous cellular antioxidant defenses and alleviating infection‐induced oxidative damage. This synergistic regulation was expected to improve cell survival and supported bone regeneration within the inflammatory microenvironment of infectious bone defects.

2.5. Biocompatibility of CPSH@PCL

For clinical translation, CPSH@PCL must couple strong antibacterial activity with excellent biocompatibility. We therefore evaluated hemocompatibility, cell viability and migration, and proliferation.

Hemolysis assays were performed to evaluate the hemocompatibility of the scaffold extracts. The 1, 3, and 5 h time points were selected to assess the early‐stage hemolytic response of red blood cells to leachable components from the scaffolds, while an additional 24 h time point was included to evaluate hemocompatibility under a longer exposure period. During the early incubation period of 1, 3, and 5 h, the hemolysis ratios of all scaffold extract groups, including PCL, P@PCL, and CPSH@PCL, remained below 2%, indicating negligible acute hemolytic activity (Figure S9A). After prolonged incubation for 24 h (Figure S9B), a slight increase in hemolysis ratio was observed in the scaffold extract groups, however, all values remained below the generally accepted 5% threshold referenced in ISO 10993–4. Notably, CPSH@PCL did not induce excessive hemolysis compared with the control scaffold groups. These results indicated that the leachable components from the Cu²⁺‐coordinated peptide hydrogel coating did not cause obvious erythrocyte damage and that CPSH@PCL exhibited acceptable hemocompatibility under the tested conditions. The cytocompatibility of CPSH@PCL was evaluated using BMSCs and osteoblasts (OBs). Live/dead staining showed abundant viable cells (green) and very few dead cells (red) in all groups, with the CPSH@PCL group comparable to the negative control (Figure 6A,B). These findings indicated that the Cu²⁺ released from CPSH@PCL remained within a safe range and did not produce obvious toxicity.

FIGURE 6.

Biocompatibility and cellular response of copper‐peptide supramolecular hydrogel‐loaded PCL composite scaffold (CPSH@PCL). (A, B) Live/Dead staining assay of BMSCs (A) and Osteoblasts (OBs, B) cultured with each group of materials. (C, D) Scratch wound healing assay evaluating the migration of BMSCs (C) and OBs (D) on different scaffolds. (E, F) Quantitative analysis of the migration rates of BMSCs (E) and OBs (F) from the scratch assay. (G, H) Cell proliferation of BMSCs (G) and OBs (H) on different scaffolds assessed by CCK‐8. All data were obtained from a minimum of n = 3 independent experiments and are expressed as mean ± SD. *p<0.05, ** p < 0.01, *** p < 0.001, **** p <0.0001.

Cell migration, a key process during early bone repair, was assessed by scratch assays. CPSH@PCL significantly promoted wound closure for both BMSCs and OBs at 12 and 24 h compared with control scaffolds (Figure 6C,D), as confirmed by quantitative analysis (Figure 6E,F). Notably, OB migration at 12 h on CPSH@PCL exceeded that of BMSCs, suggesting that this composite may preferentially and rapidly stimulate mature osteoblasts at the bone‐implant interface.

CCK‐8 assays demonstrated that CPSH@PCL supported continuous proliferation of BMSCs and OBs over 5 days, with proliferation rates comparable to those on PCL and P@PCL (Figure 6G,H), indicating that the composite provided a suitable microenvironment for cell adhesion, metabolism, and growth.

Overall, CPSH@PCL exhibited low hemolysis, low cytotoxicity, and favorable effects on cell migration and proliferation. These properties were attributable to the sustained, controllable release of Cu²⁺ from the CPSH network, which avoided toxic transient peaks while supplying bioactive Cu^{2 +} levels that modulate cell behavior and support.

2.6. In Vitro Osteogenic Performance of CPSH@PCL

We next investigated whether CPSH@PCL could promote osteogenic differentiation and meet the elevated energy and biosynthetic demands of bone formation. Osteogenic phenotypes, gene expression, and metabolic regulation were evaluated in vitro.

Alkaline phosphatase (ALP) and Alizarin Red S (ARS) staining were used to assess early osteogenic activity and matrix mineralization, respectively. After 7 and 14 days of culture, CPSH@PCL group induced markedly stronger ALP and ARS staining than control scaffolds (Figure 7A). Intense ALP staining indicated enhanced early osteogenesis, while numerous dense red mineralized nodules indicated significantly improved extracellular matrix mineralization, confirming that CPSH@PCL effectively promoted the entire osteogenic differentiation process.

FIGURE 7.

Validation of in vitro Osteogenic Mineralization Function and Metabolic Reprogramming Analysis of CPSH@PCL. (A) Representative images of alkaline phosphatase (ALP) staining after 7 days and Alizarin Red S (ARS) staining after 14 days of osteogenic induction with extracts from different scaffolds. Scale bar = 100 µm; (B) Analysis of gene expression of osteogenic markers (ALP, OPN, RUNX2, OCN) after 7 days of co‐culture via real‐time quantitative polymerase chain reaction (RT‐qPCR); (C) Analysis of gene expression of glycolysis markers (GLUT1, HK2, PDK1, LDHA, HIF‐1α) via real‐time quantitative polymerase chain reaction (RT‐qPCR). Data are presented as mean ± SD, n = 4. *p<0.05, ** p < 0.01, *** p < 0.001, **** p <0.0001; ns, no significance.

RT‐qPCR analysis of key osteogenic genes showed that BMSCs cultured with CPSH@PCL extract exhibited significantly elevated expression of early osteogenic marker ALP, osteopontin (OPN), and the master osteogenic transcription regulator RUNX2, as well as osteocalcin (OCN), a late marker of mature mineralizing osteoblasts (Figure 7B). The coordinated upregulation of these genes confirmed that CPSH@PCL drove BMSCs through early commitment, matrix secretion, and late‐stage mineralization.

Because osteogenesis is tightly coupled to cellular metabolism, we further examined the expression of five glycolysis‐related genes (Figure 7C). CPSH@PCL significantly upregulated glucose transporter 1 (GLUT1), hexokinase 2 (HK2), pyruvate dehydrogenase kinase 1 (PDK1), and lactate dehydrogenase A (LDHA). Notably, expression of hypoxia‐inducible factor‐1α (HIF‐1α), a key transcriptional regulator of glycolysis, was also significantly enhanced. HIF‐1α activation promoted the shift from oxidative phosphorylation to aerobic glycolysis by coordinately regulating GLUT1, HK2, PDK1, and LDHA, thereby enhancing ATP production and providing biosynthetic precursors (nucleotides, amino acids, lipids) necessary for rapid extracellular matrix deposition and osteogenic differentiation.

To further evaluate the potential effect of CPSH@PCL on glycolytic metabolism, lactate production in the cell culture supernatant was measured after treatment with different scaffold extracts. As shown in Figure S10, lactate production was significantly increased in the CPSH@PCL group compared with both the PCL group (p < 0.01) and the P@PCL group (p < 0.05). Since lactate was the major end product of glycolysis, this result indicated that CPSH@PCL could enhance glycolytic activity, thereby providing metabolic support for osteogenic differentiation.

Together, these results suggested that CPSH@PCL promoted osteogenesis through a dual regulatory mechanism: direct activation of osteogenic gene programs and HIF‐1α‐mediated metabolic reprogramming toward glycolysis to support the high energy and anabolic demands of bone formation.

2.7. In Vivo Antibacterial and Osteogenic Properties of CPSH@PCL

To evaluate the therapeutic efficacy of CPSH@PCL in a clinically relevant setting, we established a rat model of infectious critical‐size calvarial defects, a classical model for studying bone regeneration under infectious conditions (Figure S11). A 5 mm full‐thickness skull defect was prepared, MRSA was locally inoculated into the defect, and scaffolds (PCL, P@PCL, and CPSH@PCL) were implanted immediately. In vivo antibacterial and osteogenic outcomes were evaluated over 4 weeks.

Effective control of local infection was a prerequisite for bone regeneration. One week after implantation, Tissues from the defect area were homogenized and plated for bacteriological analysis. Few bacterial colonies were detected in the CPSH@PCL group, while dense colonies were observed in the PCL and P@PCL groups (Figure S12). To further support this observation, quantitative analysis of colony‐covered area was performed using ImageJ. Because some colonies were densely distributed and partially merged, colony‐covered area rather than exact colony number was used as a semi‐quantitative indicator of bacterial burden. The CPSH@PCL group exhibited a significantly lower colony‐covered area than the PCL and P@PCL groups (Figure S13), suggesting that CPSH@PCL reduced the local bacterial burden at the infected defect site. These results demonstrated that the sustained Cu²⁺ release from the CPSH hydrogel network maintained strong antibacterial activity in vivo, effectively clearing MRSA infection and providing a sterile microenvironment conducive to subsequent bone repair.

At 4 weeks, micro‐computed tomography (micro‐CT) was used to evaluate new bone formation. Three‐dimensional reconstruction revealed the greatest bone ingrowth and most continuous defect bridging in the CPSH@PCL group, with new bone extending from the defect margins toward the center (Figure 8A). Quantitative analysis indicated a bone regeneration rate of 40% in the CPSH@PCL group, significantly higher than that in the PCL and P@PCL groups (Figure 8B). Morphometric parameters further demonstrated superior osteogenic quality: CPSH@PCL significantly increased bone mineral density (BMD), bone volume fraction (BV/TV), and trabecular number (Tb.N) compared with controls, while trabecular thickness (Tb.Th) showed no significant intergroup differences (Figure 8C–F). These results indicated that CPSH@PCL enhances both the quantity and microarchitectural quality of regenerated bone.

FIGURE 8.

In vivo evaluation of infected bone defect regeneration. (A) 3D reconstructed microcomputed tomography (micro‐CT) images of rat cranial bones after 4 weeks; (B) quantitative analysis of new bone formation after 4 weeks; (C–F) quantitative analysis of bone morphometric parameters: (C) bone mineral density (BMD), (D) bone volume/total volume ratio (BV/TV), (E) trabecular number (Tb.N), (F) trabecular thickness (Tb.Th); (G) hematoxylin and eosin (H&E) histological staining; (H) Masson histological staining; (I) immunohistochemical (IHC) staining for Osteocalcin (OCN). Data are presented as mean ± SD, n = 4. *p<0.05, ** p < 0.01, *** p < 0.001, **** p <0.0001, ns, no significance.

Histological staining corroborated the imaging findings. Hematoxylin‐eosin (H&E) staining (Figure 8G) showed significantly reduced inflammatory cell infiltration in the CPSH@PCL group and abundant, orderly arranged osteoblasts on the surfaces of newly formed bone. In contrast, control defects displayed a persistent inflammatory response and fibrous tissue encapsulation. Blinded ImageJ‐based quantitative analysis further confirmed that the relative inflammatory infiltration area, normalized to the PCL group, was significantly lower in the CPSH@PCL group than in the PCL and P@PCL groups (Figure S14A), indicating effective attenuation of local inflammatory responses. Masson's trichrome staining (Figure 8H) showed extensive, and well‐organized collagen matrix deposition (blue) and substantial new mineralized bone formation (red) in the CPSH@PCL group. At high magnification, the gradual transition from osteoid into mature mineralized bone with clear lacunar structures was clearly observed, indicating ongoing osteogenic activity and tissue maturation. Quantitative analysis demonstrated that the collagen‐positive area was significantly increased in the CPSH@PCL group compared with the PCL and P@PCL groups (Figure S14B), suggesting enhanced extracellular matrix deposition during bone repair.

Immunohistochemical (IHC) staining for OCN, a late osteogenic marker, further validated the enhanced osteogenesis (Figure 8I). The CPSH@PCL group showed strong, widespread OCN expression in the newly formed bone matrix, whereas control groups exhibited weak and scattered staining. Blinded ImageJ‐based quantitative analysis showed that the OCN‐positive area was significantly higher in the CPSH@PCL group than in the control groups (Figure S14C). These data demonstrated that CPSH@PCL not only supported robust new bone formation but also promoted maturation into functional bone tissue in vivo.

Overall, CPSH@PCL demonstrated sustained in vivo antibacterial efficacy and markedly enhanced bone regeneration in an infected calvarial defect model. By continuously suppressing infection and local inflammation while improving the regenerative microenvironment, CPSH@PCL integrated antibacterial, antibiofilm, antioxidant, and osteogenic functions within a single scaffold, underscoring its potential as an intelligent bioactive platform for infected bone defect repair.

Importantly, the osteogenic enhancement was not driven by a single inductive cue but arose from coordinated metabolic regulation. The sustained release of Cu²⁺ activated HIF‐1α‐associated metabolic pathways, promoting energy production and biosynthetic capacity required for bone matrix formation. Consequently, metabolic reprogramming emerged as a central nexus linking infection control, oxidative stress alleviation, and osteogenic regeneration. These in vivo findings confirmed that the multifunctional synergy of CPSH@PCL was preserved under infectious conditions, enabling robust and high‐quality bone regeneration.

3. Conclusion

In summary, we developed a coordination‐driven Cu²⁺‐peptide supramolecular hydrogel‐PCL scaffold (CPSH@PCL) for the treatment of infected bone defects. By integrating Cu²⁺ as a structural coordination unit within a peptide self‐assembly network integrated into a mechanically robust PCL scaffold, CPSH@PCL achieved synchronized control of Cu²⁺ release, durable antibacterial and antibiofilm activity, modulation of oxidative stress, and promotion of osteogenic differentiation. Mechanistic studies indicated that CPSH@PCL coupled infection control with bone regeneration in part through HIF‐1α‐mediated metabolic reprogramming toward glycolysis. This integrated structure‐function strategy provided a versatile blueprint for designing infection‐tolerant bone repair materials and broadened the application potential of multifunctional biomaterials in regenerative medicine.

4. Experimental Section/Methods

4.1. Preparation of Copper‐Peptide Supramolecular Hydrogel (CPSH)

FE3 peptide (99.95%, DongHeng, China) was dissolved in ultrapure water, and the pH was adjusted to 7.0 ± 0.2 with 1 M NaOH. CuCl₂ solution was added at peptide‐to‐Cu²⁺ molar ratios of 1:0.5, 1:1, and 1:1.5. After vortexing for 30 s, the mixtures were incubated at 25 °C for 30 min to form CPSH.

4.2. Fabrication of 3D‐Printed PCL Scaffolds

PCL scaffolds were fabricated by a melt‐extrusion 3D printing (CREALITY, China) at 60 °C. The printing parameters were set to a layer height of 0.2 mm, a printing speed of 1000 mm min⁻¹, and an infill density of 27 %. Cylindrical scaffolds (Ø5 × 1 mm) were sterilized in 75% ethanol and UV irradiation.

4.3. Preparation of Copper‐Peptide Supramolecular Hydrogel Integrated Polycaprolactone Scaffolds (CPSH@PCL)

PCL scaffolds were completely immersed in CPSH, followed by ultrasonic treatment for 5 min to remove trapped air. The scaffolds were then incubated under ambient pressure for 12 h to allow hydrogel infiltration and surface coating, yielding CPSH@PCL. P@PCL were prepared using the same procedure without Cu²⁺.

4.4. Transmission Electron Microscopy (TEM)

Hydrogel samples were diluted 50‐fold with deionized water, deposited onto carbon‐coated copper grids, negative‐stained with 2 wt% phosphotungstic acid (pH 6.8), and imaged at 100 kV using a TEM (Thermo Fisher Scientific, USA).

4.5. Scanning Electron Microscopy (SEM)

Freeze‐dried CPSH, PCL, and CPSH@PCL samples were sputter‐coated with platinum (20 mA, 45 s) and observed by SEM (JEOL, Japan).

4.6. Water Contact Angle Measurement

Surface wettability was evaluated using a sessile drop method (SINDIN, China). A 3 µL droplet of deionized water was placed on scaffold surfaces, and contact angles were recorded within 10 s.

4.7. Cu²⁺ Release

Scaffolds (0.1 g) were immersed in 1 mL PBS (pH 7.4 or 6.2) at 37 °C. Released Cu²⁺ was quantified using a TMB‐based colorimetric assay (Macklin, China) with a microplate reader (Tecan, Switzerland) and calculated based on a CuCl₂ standard curve.

4.8. Zeta Potential Measurement

Zeta potential measurements were performed using a Zetasizer Nano ZS (Malvern Panalytical, UK) at 25°C. Samples were diluted in ultrapure water and measured in triplicate.

4.9. Circular Dichroism Spectroscopy (CD)

CD spectra were recorded in the range of 185–250 nm using a CD Spectrometer (Applied Photophysics, UK) with a 0.5 cm quartz cuvette.

4.10. Fourier Transform Infrared Spectroscopy (FTIR)

FTIR spectra were acquired using an FTIR spectrometer (Thermo Fisher Scientific, USA). Freeze‐dried CPSH samples were ground with dry KBr and pressed into transparent pellets prior to spectral analysis.

4.11. Matrix‐Assisted Laser Desorption/Ionization Time‐of‐Flight Mass Spectrometry (MALDI‐TOF MS)

CPSH samples were mixed with α‐cyano‐4‐hydroxycinnamic acid (CHCA) matrix (15 mg mL⁻¹ in 70% ACN/0.1% TFA). Spectra were collected in negative ion reflection mode using a Bruker Ultraflextreme MALDI‐TOF/TOF mass spectrometer.

4.12. Molecular Dynamics (MD) Simulation

All‐atom molecular dynamics simulations were conducted using GROMACS 2023.3 with the AMBER99SB‐ILDN force field. The simulation system consisted of 20 FE3 peptides and 30 Cu²⁺ ions placed randomly in an 8 × 8 × 8 nm cubic box solvated with TIP3P water molecules. After energy minimization using the steepest descent algorithm, the system was equilibrated under NPT conditions for 100 ps. Production simulations were performed for 90 ns at 298 K and 1 bar. Trajectory visualization and analyses, including RMSD, Rg, and SASA, were carried out using built‐in GROMACS tools and VMD software.

4.13. Preparation of Scaffold Extracts

Sterile CPSH@PCL were prepared under aseptic conditions. Scaffold extracts were obtained in accordance with ISO 10993‐12:2021, scaffolds were immersed in sterile Luria‐Bertani (LB) broth for bacterial assays or low‐glucose DMEM (L‐DMEM) for cell‐based assays at a ratio of 0.1 g mL⁻¹ and incubated at 37 °C for 72 h. The extracts were collected and stored at 4 °C prior to use. Extracts from blank PCL scaffolds and P@PCL were prepared using the same protocol.

4.14. Bacterial Culture

MRSA and P. aeruginosa were kindly provided by Prof. Wenjun Miao (Nanjing Tech University, China). These strains, representing Gram‐positive and Gram‐negative pathogens, respectively, were used as model organisms to evaluate the antibacterial performance of CPSH@PCL. Bacteria were cultured in 5 mL LB broth at 37°C with shaking at 180 rpm for 12 h. Exponentially growing bacteria were harvested the following day and resuspended in PBS (pH 7).

4.15. In Vitro Antibacterial Activity

Bacterial suspensions (1 × 10⁶ CFU mL⁻¹ in PBS) were incubated with scaffold extracts (100 µL per well) in 96‐well plates. Optical density at 600 nm (OD600) was measured at 0 and 12 h. For colony counting, suspensions were serially diluted (10⁶ fold), plated on LB agar, and incubated at 37 °C for 12 h. Bacterial viability was further assessed using an MTT assay (0.5 mg mL⁻¹, 30 min), followed by DMSO solubilization and OD570 measurement.

4.16. Biofilm Inhibition and Eradication Assays

For biofilm inhibition, bacterial suspensions (1 × 10⁶ CFU mL⁻¹) were cultured with scaffold extracts for 72 h. For eradication assays, mature biofilms were first established by culturing bacteria in LB broth for 72 h, followed by treatment with extracts for 12 h. Biofilms were fixed with methanol, stained with 0.5% crystal violet, and quantified after dissolution with 33% acetic acid by measuring OD570.

4.17. ROS Scavenging Capacity

For qualitative observation of oxygen generation, scaffold samples were immersed in 100 µL H₂O₂ solution (100 mM) and immediately observed under an optical microscope. Bubble formation on the scaffold surface was recorded as an indicator of catalase‐like decomposition of H₂O₂. Cu@PCL was prepared as an additional copper‐only control. Briefly, sterile PCL scaffolds were immersed in an aqueous CuCl₂ solution containing an equivalent amount of Cu²⁺ to that used for CPSH preparation, followed by incubation under the same conditions as CPSH@PCL. For extract‐based evaluation, 40 µL extract was mixed with 40 µL H₂O₂ (100 mM) and 10 µL TMB solution. After 30 min incubation, reactions were terminated with 10 µL 1 M HCl, and absorbance at 450 nm was recorded.

4.18. Intracellular ROS Detection

BMSCs were seeded in 24‐well plates at 5× 10³ cells per well. Upon reaching 80–90% confluence, cells were exposed to ROS‐inducing medium containing 400 µM H₂O₂ solution for 12 h. The medium was then replaced with scaffold extract medium and incubated for another 12 h. Intracellular ROS levels were detected using DCFH‐DA staining, with nuclei counterstained by DAPI. Fluorescence images were captured using an inverted fluorescence microscope.

4.19. The Cytotoxicity of Free Cu²⁺ and Scaffold Extracts Was Assessed by MTT Assay

Extracts from PCL, P@PCL, and CPSH@PCL were prepared according to Section 4.13. Free Cu²⁺ solution was prepared from CuCl₂ under the same extraction conditions as CPSH@PCL, with the CuCl₂ dosage equivalent to that used in CPSH fabrication (at a Cu²⁺ concentration of 30 mM). BMSCs were seeded in 96‐well plates at a density of 5 × 10³ cells per well and cultured overnight. The medium was then replaced with the extracts. After 24 h of incubation, MTT solution was added and incubated at 37 °C for 4 h. The formed formazan crystals were dissolved in DMSO, and the absorbance was measured at 570 nm using a microplate reader. Cell viability was expressed as a percentage relative to the control group.

4.20. RT‐qPCR Analysis of Antioxidant and Bone Repair‐Related Gene Expressions

BMSCs were seeded in six‐well plates at 4×10⁴ cells per well with scaffold extracts. Total RNA was extracted using TRIzol reagent (Vazyme, China), and quantified by NanoDrop (Thermo Fisher Scientific, USA). Reverse transcription was performed using the HiScript III First‐Strand cDNA Synthesis Kit (Vazyme, China). Quantitative PCR was performed using SYBR qPCR Master Mix (Vazyme, China) in a 20 µL reaction volume for 40 cycles. Relative gene expression was calculated using 2^−ΔΔCt method. Primer sequences were listed in Table S3.

4.21. Hemolytic Assay

Hemocompatibility was evaluated by an in vitro hemolysis assay. Whole blood was collected from Sprague‐Dawley rats and centrifuged to isolate red blood cells (RBCs), which were washed repeatedly with saline. The RBC suspension was then mixed with scaffold extracts and incubated at 37°C for 1, 3, 5, and 24 h. Saline and distilled water served as negative (0% hemolysis) and positive (100% hemolysis) controls, respectively. After centrifugation, absorbance at 540 nm was measured, and hemolysis percentages were calculated.

4.22. In Vitro Cytocompatibility

Cytocompatibility was evaluated using Live/Dead staining and CCK‐8 assays. BMSCs and osteoblasts were cultured in 96‐well plates at 5 × 10³ cells per well and cultured with scaffold extracts. Live/Dead staining was performed after 3 days using Calcein‐AM and propidium iodide. For CCK‐8 assays, absorbance at 450 nm was measured after 1, 3, and 5 days of culture.

4.23. Cell Migration Assay

BMSCs and osteoblasts were seeded in six‐well plates (4 × 10⁴ cells per well). After reaching 90% confluence, a linear scratch was introduced using a pipette tip. Cells were cultured in serum‐free scaffold extract medium, and wound closure was monitored at 0, 12, and 24 h. Migration rates were quantified using ImageJ.

4.24. In Vitro Osteogenic Differentiation

BMSCs were cultured in osteogenic induction medium prepared with scaffold extracts. Cells were seeded at 4 × 10⁴ cells per well and cultured until 80–95% confluence, followed by replacement with fresh osteogenic induction extract every 48 h.

4.25. ALP Staining

After 7 days of osteogenic induction, BMSCs were fixed with 4% paraformaldehyde and stained using an ALP staining kit. Images were acquired by optical microscopy.

4.26. Alizarin Red S Staining

After 14 days of osteogenic induction, BMSCs were fixed and stained with Alizarin Red S solution to evaluate mineralized nodule formation.

4.27. Lactate Assay

BMSCs were seeded in six‐well plates at 4 × 10⁴ cells per well with scaffold extracts. After 48 h, collect the cell culture supernatant and determine lactate production using an L‐lactic acid assay kit (Beyotime, Shanghai) in accordance with the manufacturer's instructions.

4.28. Animal Study

Twenty‐four male Sprague‐Dawley rats (≈300 g) were used. All animal procedures were approved by the Institutional Animal Care and Use Committee of the Jiangsu Center for Safety Evaluation of Drugs (IACUC‐20240710‐01).

4.29. Infected Calvarial Defect Model

Rats were randomly assigned to PCL, P@PCL, and CPSH@PCL groups (n = 8). Under general anesthesia, a 5‐mm full‐thickness calvarial defect was created using a trephine bur. MRSA suspension (1.5 µL, 1 × 10⁸ CFU mL⁻¹) was inoculated into the defect, followed by implantation of the assigned scaffold.

4.30. In Vivo Antibacterial Assessment

One week post‐implantation, four rats per group were euthanized. Harvested skulls were incubated in PBS at 37°C for 12 h, and aliquots were plated on LB agar to qualitatively assess antibacterial activity. The obtained images were analyzed for bacterial colony coverage area using ImageJ software, which served as a semi‐quantitative indicator of bacterial load. The plate images were then imported into ImageJ for quantitative analysis of colony coverage percentage.

4.31. Micro‐CT Analysis

Bone regeneration was evaluated by micro‐CT (70 kV, 329 µA, 6 µm resolution). A cylindrical VOI (5.0 mm diameter, 1.0 mm depth) was analyzed for BMD, BV/TV, Tb.N, and Tb.Th.

4.32. Histological Analysis

Samples were fixed in 4% paraformaldehyde, embedded in paraffin, and sectioned. H&E and Masson's trichrome staining were performed to evaluate bone regeneration.

4.33. Immunofluorescence Staining

Sections were subjected to antigen retrieval, permeabilization, and blocking, followed by incubation with anti‐OCN primary antibodies and fluorescent secondary antibodies. Nuclei were counterstained with DAPI, and images were acquired by fluorescence microscopy (BX51, Olympus, Japan).

4.34. ImageJ‐Based Quantitative Histological Analysis

Quantitative histological analysis was performed using ImageJ. All images were anonymized and randomly coded before analysis. Two independent investigators blinded to the treatment groups performed the quantification. The entire defect region in each image was defined as the region of interest, and the mean value from the two investigators was used for final statistical analysis.

Hematoxylin‐positive inflammatory cell‐rich areas in H&E‐stained sections, collagen‐positive areas in Masson's trichrome‐stained sections, and OCN‐positive areas in OCN immunostained sections were segmented using the color threshold function in ImageJ. Positive areas were calculated as percentages of the total region of interest. The inflammatory infiltration area was further normalized to the mean value of the PCL group. Identical thresholding parameters were applied to all images within the same staining batch.

4.35. Statistical Analysis

All data are presented as mean ± SD. Statistical analyses were conducted using one‐way ANOVA with Tukey's post hoc test using GraphPad Prism 8.4 (GraphPad Software, USA). A p‐value < 0.05 was considered statistically significant.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: adhm71342‐sup‐0001‐SuppMat.docx.

Data Availability Statement

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