A smart drug delivery microgel system with phased intervention capabilities and dual physical state of use promotes healing of diabetic infected wounds

Abstract

Effectively managing infected diabetic wounds involves the elimination of bacteria, neutralization of reactive oxygen species (ROS), suppression of inflammation, and induction of angiogenesis. This study describes the development of a multifunctional hyaluronic acid (HA)-based microgel system capable of serving as either an injectable wet microgel or dry microspheres (MSs). After initially engineering Fe²⁺/tea polyphenol (TP) metal–polyphenol network (MPN)-functionalized HAMA MS, these particles were found to suppress inflammation and facilitate ROS scavenging. A deferoxamine (DFO)-loaded zinc-based metal–organic framework (ZIF-8@DFO) was then coated using phenylboronic acid (PBA)-functionalized ε-polylysine (PPL) to produce PPZD nanoparticles with antibacterial and pro-angiogenic properties. The dynamic loading of PPZD into MPN-functionalized MS (MMS) via boron ester bonds then yielded a pH/ROS-responsive microgel system (MMS@PPZD). PPL coating endowed the prepared materials with antimicrobial properties while mitigating cytotoxic effects resulting from the rapid release of Zn²⁺ and DFO in acidic micro-environments. This microgel system showed superior biocompatibility and phased intervention activities aligned with the various stages of the wound healing process in vitro and in vivo. Specifically, under acidic conditions, the system sequentially released TP, PL, Zn²⁺, and DFO, enabling effective ROS scavenging, suppressing inflammation, exhibiting antibacterial activity, and inducing angiogenesis. Overall, this environmentally-responsive, multifunctional, versatile microgel system offers significant promise for infected diabetic wound management.

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1. Introduction

Diabetes affected 536.6 million people throughout the world in 2021, and this figure is expected to rise to 783.2 million by 2045.¹ Chronically infected wounds, including diabetic foot ulcers (DFUs), commonly develop in diabetic individuals,² resulting in high therapeutic costs and a high risk of gangrene, amputation, and mortality.³ Normally, wound healing progresses through hemostatic, inflammatory, proliferative, and remodeling phases.⁴ Infected diabetic wounds, however, often stall in the inflammatory phase, associated with persistent elevation of inflammatory cytokines, growth factor, and ROS levels, along with the acidification of the wound microenvironment and the impairment of angiogenesis.⁵ Wounds in patients with diabetes that remain persistently infected with bacteria, due to the hyperglycemic wound environment, contribute to further deterioration of the healing process.⁶ Roughly half of all diabetes patients have peripheral artery disease,⁷ leading to further deterioration of the hypoxic wound environment and a greater consequent risk of amputation.⁸ Effective therapeutic strategies for infected diabetic wound management thus center on eliminating bacteria, neutralizing ROS, suppressing inflammation, and promoting angiogenesis.

Conventional treatments for infected diabetic wounds are often ineffective and associated with side effects,^9,10 underscoring the need for new strategies. A large number of flexible biomaterials have shown promise in treating these wounds,¹¹ including hydrogel microspheres (MSs),^12,13 hydrogels,^14,15 films,¹⁶ foams/sponges,¹⁷ nanoparticles,¹⁸ and extracellular matrix (ECM)-derived scaffolds.¹⁹ Compared to other biomaterials like scaffolds and films, hydrogel MSs offer several advantages. Specifically, when applied to infected wounds that are extensive, irregularly shaped, exudate-rich, or challenging to inject, MS dry powder provides significant benefits. Additionally, wet MSs can also be used in a manner similar to traditional hydrogels. The tunable properties and flexible states of these MS-based preparations can enable their use in treating a wide range of wound types.¹²

Hyaluronic acids (HAs) are attractive candidates for use in wound healing applications given the ease with which they can be acquired and synthesized, their excellent biocompatibility and biodegradability, and their ability to retain moisture while absorbing wound exudates.^20,21 HA-based hydrogel MSs have been a focus of particular interest as a platform for drug delivery as they are modifiable.²² The coordination of polyphenols with metal ions (Mn, Cu, Gd, Ti, and Fe) enables the preparation of films consisting of a supramolecular self-assembled metal–polyphenol network (MPN).^23,24 These adaptable MPN films have universal surface binding affinity and pH responsivity such that they can be effectively applied as responsible, multifunctional coating materials.^25,26 MPN-based surface functionalization processes are straightforward, rapid, and stable such that they have been applied in the preparation of nanoparticles,²⁶ stem cell microspheres,²⁷ titanium alloys,²⁸ and other scaffold materials.²⁹ Tea polyphenols (TPs) are natural tea-derived polyphenolic compounds with significant anti-inflammatory and antioxidant properties.³⁰ Here, a TP and Fe²⁺ based MPN was used as a coating for hyaluronan methylacrylate (HAMA) MSs, yielding MSs with enhanced anti-inflammatory and antioxidant activity. To adequately address the chronically infected diabetic wounds, biomaterials that exhibit antibacterial and angiogenic activity are also needed.

Iron chelator deferoxamine (DFO) functions by activating HIF-1α signaling, leading to the transactivation of secreted factors including VEGF that promote vasculogenesis.³¹ Even though there are several studies documenting the ability of DFO to improve angiogenesis when applied in the context of wound healing, it has a short half-life, is rapidly cleared, and exhibits poor biocompatibility, thus limiting its clinical use.^32,33 There is thus a pressing need to design new approaches for delivering DFO. ZIF-8, the zinc-based metal–organic framework (MOF), is a biodegradable platform with good capacity for drug loading and excellent drug delivery activity such that it is a promising tool for efficient drug delivery mediated through pH-sensitive drug release.³⁴ ZIF-8 also exhibits antimicrobial properties associated with the release of Zn²⁺.^35,36 DFO-loaded ZIF-8 (ZIF-8@DFO) is thus an ideal platform for the treatment of infected wounds in patients with diabetes. ZIF-8 has been linked to negative effects including neurobehavioral disorders in zebrafish,³⁷ neutrophil aggregation and inflammation,³⁸ and increased ROS biogenesis.³⁹ In addition, infected wounds with a low pH undergo more rapid ZIF-8 degradation, resulting in higher levels of local DFO and Zn²⁺ release that may result in toxic side effects. Efforts to mitigate these effects have focused on the surface modification of ZIF-8 using coating materials, including HA, cell membranes, exosomes, and polydopamine (PDA).⁴⁰ These modifications, however, tend to result in the overly slow release of Zn²⁺, hampering the antibacterial activity. Interestingly, ε-polylysine (PL) is a natural cationic peptide that has emerged as an attractive research target given its high levels of antibacterial activity, superior tissue and cellular adhesion, and cost-effectiveness.⁴¹ PL is thus an ideal material for use in the coating of ZIF-8, providing a means of enhancing the underlying antibacterial efficacy, reducing associated side effects, and improving the bio-adhesion and tissue compatibility of the resultant preparations.

Drug delivery systems with spatiotemporally sequential release functionality have been demonstrated to be effective in the repair of infected wounds.⁴² In this study, 4-carboxyl phenylboronic acid (PBA) grafted ε-polylysine (PPL) was used to coat the ZIF-8@DFO (PPZD), after which it was dynamically loaded onto TP/Fe²⁺ MPN-coated HAMA MSs with boronate ester bonds, producing a ROS/pH-responsive microgel system (MMS@PPZD) for infected diabetic wound treatment (Fig. 1). The MPN and boronate ester bond in the resultant preparation were responsive to acidic conditions, initially releasing TP to suppress inflammation and neutralize ROS. PPZD can then detach from the MSs to exert antibacterial effects, while DFO is released from the ZIF-8 system, promoting angiogenesis. Here, a multifunctional microgel system with intelligent delivery and sequential release functionality was developed, capable of neutralizing ROS, suppressing inflammation, and inducing angiogenesis, ultimately leading to enhanced diabetic wound healing. This system can also be leveraged in the form of both an injectable wet microgel and as a dry powder, providing further versatility.

Fig. 1 Schematic overview of the microgel system developed herein and its mechanistic effects when deployed for the treatment of infected diabetic wounds. HA: hyaluronic acid; MA: methacrylic anhydride; HAMA: hyaluronan methylacrylate; MS: microsphere; TP: tea polyphenol; MMS: TP/Fe²⁺ metal–polyphenol network (MPN)-coated HAMA MS; PL: ε-polylysine; PBA: 4-carboxyphenylboronic acid; DFO: deferoxamine; 2-MI: 2-methylimidazole.

2. Materials and methods

HA (M_w 2 500 000), ε-polylysine (PL) (M_w 4100), methacrylic anhydride (MA), N-hydroxysuccinimide (NHS), 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC), 4-carboxyphenylboronic acid (PBA), zinc acetate dihydrate, 2-methylimidazole, anhydrous ferric chloride (FeCl₃), anhydrous ferrous chloride (FeCl₂), tea polyphenols (TPs), and deferoxamine (DFO) were from Macklin (Shanghai, China). Deferoxamine – Rhodamine B (DFO-RhB) was from QIYUE BIOLOGY (Xi’an, China). Corning (Shanghai, China) was the source of all culture media, PBS, trypsin, enzyme-free water, and culture dishes and plates. Abcam (Shanghai, China) and Affinity (Jiangsu, China) were the sources of all antibodies. Solarbio (Beijing, China) was the source of the DAPI and live/dead staining kit used for this study.

2.1. PPL, ZIF-8@DFO and PPZD synthesis and characterization

PPL (4-carboxyphenylboronic acid-grafted ε-polylysine) synthesis was achieved with the previously described EDC/NHS method.^43,44 Briefly, 1.5 mL of deionized (DI) water was used to dissolve 150 mg of PL (solution A), while 20 mL of DMSO was used to dissolve 146 mg of PBA, 172 mg of EDC, and 106 mg of NHS (solution B), and this latter solution was stirred for 1 h at 250 rpm at 25 °C. Solution B was then added to Solution A, and the mixture was stirred for 48 h at 250 rpm at 25 °C, thereby yielding PPL. PPL was then purified through dialysis (MWCO: 1 kDa), after which it was lyophilized. The success of PPL synthesis was confirmed through ¹H NMR spectroscopy.

A one-pot synthesis method was used to incorporate DFO into ZIF-8 as in a prior report.⁴⁵ First, 10 mL of methanol was used to dissolve 325 mg of 2-methylimidazole, followed by stirring for 30 min at room temperature to produce a homogenous mixture (solution A). In addition, 5 mL of methanol was combined with DFO (7.5 mg) and zinc acetate dihydrate (109 mg) followed by sonication for 5 min at room temperature to ensure uniform mixing, producing Solution B. Solution A was then added in a dropwise manner to Solution B, and the resultant mixture was stirred at 1500 rpm for 30 min at 37 °C to produce ZIF-8@DFO. Methanol was used to wash the prepared ZIF-8@DFO solution followed by centrifugation (15 min, 12 000 rpm), repeating this process three times and collecting the supernatant to assess the DFO loading capacity. Precipitated ZIF-8@DFO samples were dried for 24 h under vacuum conditions. A similar approach was also used for pure ZIF-8 synthesis. As prior studies noted the highest drug encapsulation efficiency and loading efficiency values when using 7.5 mg of DFO,³¹ this same amount of DFO was used in this study.

PPL was next used to coat ZIF-8@DFO preparations. Briefly, 5 mL of DI water was used to dissolve 10 mg of PPL (Solution A), while 2 mL of DI water was used to dissolve 10 mg of ZIF-8@DFO, with sonication for 5 min (Solution B). Solution B was then gradually added to Solution A under conditions of continuous stirring at 600 rpm over 5 min, followed by incubation for 6 h at room temperature. This mixture was then rinsed with DI water, centrifuged (15 min, 12 000 rpm), and this washing step was performed three times. Supernatants were also collected to assess residual DFO in the context of the PPL coating of ZIF-8@DFO. The resultant PPZD precipitate was harvested and dried for 24 h under vacuum conditions.

The crystal morphology of ZIF-8, ZIF-8@DFO, and PPZD was analyzed through scanning electron microscopy (SEM), transmission electron microscopy (TEM), and X-ray diffraction (XRD) approaches. TEM-based energy-dispersive X-ray spectroscopy (EDX) analyses were used to assess the presence of sulfur (S) and boron (B), which are respectively only present in DFO and PBA. Particle size and ζ-potential values were also assessed through Zetasizer spectroscopy.

DFO encapsulation efficiency (EE) and loading efficiency (LE) in PPZD were assessed by using UV-vis spectrophotometry to quantify the residual levels of DFO in the supernatant fraction. DFO can function by chelating Fe³⁺ to generate a complex that is detectable with a UV-vis spectrophotometer at 430 nm.³¹ To assess the sustained and delayed release properties of the preparations established herein under acidic conditions following PPL coating, PBS (pH 5.0) was used to disperse ZIF-8@DFO and PPZD. DFO release into the supernatant was then assessed via UV-vis spectrophotometry at defined intervals. The Zn²⁺ concentrations in the samples were measured by ICP-OES. To test the ability of PPL coating to reduce ZIF-8@DFO toxicity, L929 cells were treated with ZIF-8@DFO and PPZD extracts. Briefly, after disinfecting ZIF-8@DFO (120 μg mL⁻¹) and PPZD (200 μg mL⁻¹), they were submerged for 48 h in complete media (pH 5.0). After culturing L929 cells for 24 h, their media were exchanged for these extract preparations. CCK-8 assays and Calcein-AM/PI staining were used to assess the viability and proliferation of these cells after 24 and 48 h.

2.2. MMS@PPZD preparation and characterization

HAMA synthesis was performed as described previously.⁴⁶ Briefly, MA (2 mL) and HA (5 g) were dissolved in 250 mL of carbonate buffer and allowed to react for 12 h. The reactants were then dialyzed and lyophilized to obtain HAMA. HAMA MS preparation was performed using a microfluidic system. The aqueous phase consisted of 8 wt% HAMA and 0.5 wt% photoinitiator in deionized (DI) water, while the oil phase was composed of paraffin oil containing 5 wt% Span 80. Syringes were used to inject these solutions into the inlets of the microfluidic device, and a syringe pump was used to set the flow rates for both phases to 10 μL min⁻¹ using a syringe pump. To initiate polymerization, samples were exposed to UV light, yielding cured HAMA MSs that were then washed thrice with DI water.

HAMAS MSs were then coated with an MPN prepared from TP and Fe²⁺. For this study, a TP to FeCl₂ mass ratio of 2 : 1 was used, as in prior reports.^27,47 The MPN coating of MS samples (to prepare MMS) was achieved by combining 5 mL of MS with 5 mL of a 10 mg mL⁻¹ solution of TP, followed by constant shaking for 10 min. Next, 1 mL of 25 mg mL⁻¹ FeCl₂ was added, followed by gentle shaking for a further 2 h. MMS samples were then rinsed thrice with DI water to remove the remaining TP and FeCl₂, after which lyophilization was used to obtain the MMS.

The prepared PPZD was loaded into MMS samples as follows (producing MMS@PPZD): lyophilized PPZD (10 mg) was suspended in 10 mL of DI water with sonication, followed by mixing with MMS (10 mL). An alkaline buffer was used to adjust the solution to a pH of 8.5, and it was shaken overnight at room temperature at low speeds. MMS@PPZD was then rinsed thrice with DI water to remove the remaining PPZD, after which lyophilization was used to obtain the prepared MMS@PPZD. The supernatants were collected, centrifuged (12 000 rpm, 15 min), and lyophilized to assess the residual PPZD.

¹H NMR spectroscopy was used to assess HAMA structural features. MS, MMS, and MMS@PPZD water retention testing was performed as described in the ESI.† SEM and mapping strategies were used to assess the surface structures and morphology of different MS preparations and the impact of MMS and PPZD loading on MMS@PPZD. The PPZD loading of MMS samples with RhB-DFO was additionally visualized via fluorescence microscopy (FM, Ex: 540 nm; Em: 570 nm).

The environmentally responsive release of Zn²⁺ and DFO from MMS@PPZD was evaluated by incubating the prepared microgel in PBS samples designed to simulate various microenvironments (pH 7.4, pH 5.0, and pH 5.0 with 100 μM H₂O₂) at 37 °C. DFO-RhB was utilized to assess the DFO release dynamics. Supernatants were collected at appropriate time intervals and the associated fluorescent signal was assessed with a Bio Synergy H1 microplate reader (Ex: 540 nm; Em: 570 nm). The release of Zn²⁺ was also measured by ICP-OES and 1,1-diphenyl-2-picrylhydrazyl (DPPH) was used to assess ROS-scavenging activity, as described in the ESI.†

2.3. In vitro antibacterial assays

The minimum inhibitory concentration (MIC) was evaluated using the broth microdilution method, as described in the ESI.† Additionally, the antibacterial ratio at different concentrations of PPZD and the optimal antibacterial concentration of PPZD were determined, as detailed in the ESI.†

After irradiation for disinfection purposes, PPZD (200 μg mL⁻¹), freeze-dried MS (100 μL mL⁻¹), MMS (100 μL mL⁻¹), and MMS@PPZD (100 μL mL⁻¹) were submerged in Luria-Bertani (LB) broth (pH 5) for 48 hours, respectively. The ability of the different materials to exert antimicrobial effects was evaluated by culturing S. aureus or E. coli (10 μL, 1 × 10⁸ CFU mL⁻¹) for 12 h in 24-well plates in the presence of 1 mL of extracts of the different materials (PPZD, MS, MMS, MMS@PPZD), using 1 mL LB broth (pH = 5.0) and 50 μg Cefepime in 1 mL LB broth (pH = 5.0) as the respective blank and positive control. One-hundred microliter suspensions of bacteria from these wells were then transferred to solid LB agar plates and cultured for 12 h at 37 °C.³ The OD 600 values for these bacterial samples were quantified. The plates were imaged and the colonies were counted. Bacterial live–dead staining was conducted by treating the cells with SYTO9 and propidium iodide (PI) for 15 min in the dark and then imaging the bacteria under a fluorescence microscope (FM, excitation: 488 nm; emission: 536 nm).³

2.4. Assessment of microgel biocompatibility

Microgel biocompatibility was evaluated by using the prepared extracts to treat L929 cells and HUVECs. Briefly, after irradiating PPZD (200 μg mL⁻¹), freeze-dried MS (100 μL mL⁻¹), MMS (100 μL mL⁻¹), and MMS@PPZD (100 μL mL⁻¹) for the purposes of disinfection, they were submerged for 48 h in complete media (pH = 5.0). After culturing L929 cells and HUVECs for 24 h, their media was exchanged for these prepared extracts. Shifts in cell viability were then evaluated after 24 and 48 h through CCK-8 and Calcein-AM/PI staining approaches. Microgels were also used to conduct a hemolysis assay as reported previously.⁴⁸

2.5. Wound healing experiment

After culturing L929 cells to confluence in 6-well plates, they were treated for 2 h with H₂O₂ (100 μM), and the monolayer was subsequently scratched with a 10 μL pipette tip. After rinsing the cells three times with PBS, the cells were treated with appropriate material extracts (the same as those used in the assessment of microgel biocompatibility), and a fluorescence microscope was used to image the healing of these simulated wounds over time.

2.6. In vitro antioxidant, anti-inflammatory, and angiogenic assays

The ability of microgels to scavenge ROS was assessed with H₂O₂ (100 μM)-pretreated L929 and RAW 264.7 cells as reported previously.^3,35 Intracellular ROS detection was achieved with 2,7-dichlorofuorescein diacetate (DCFH-DA; APExBIO, USA). After incubating RAW 264.7 macrophages in DMEM with LPS (5 μg mL⁻¹) for 2 h, they were incubated for a further 24 h in the presence of 2 mL of the prepared material extracts (the same as those used in the assessment of microgel biocompatibility). Altered cytokine expression was then evaluated through qPCR, Western immunoblotting, and immunofluorescent staining approaches. HUVECs were additionally treated with these prepared extracts for 48 h to analyze angiogenic activity, and cytokine expression was assessed using the same approaches as above. For further details regarding the HUVEC angiogenesis assays, see the ESI.†

2.7. In vivo analyses

A previously reported approach was used to establish a rat model of diabetes.³ The ability of the prepared microgel to promote healing was assessed as in prior reports.^3,48 Briefly, male Sprague-Dawley rats (280–320 g) were randomized into Control, PPZD, MS, MMS, and MMS@PPZD groups. To model diabetes, rats in the appropriate groups were intravenously injected with streptozotocin once per day until their blood glucose levels were above 16.7 mM. These rats were then anesthetized and a wound 15 mm in diameter was generated on each animal. A 200 μL volume of a suspension containing both S. aureus and E. coli was then applied to the wound site. After a 24-h rest interval, the wound sites in appropriate groups were treated with 100 μL of PBS, PPZD (200 μg mL⁻¹), or wet microgel (MS, MMS, or MMS@PPZD), as appropriate (n = 6). The relative performance of 100 μL of wet MMS@PPZD was also compared to that of the corresponding lyophilized powder as a tool for wound treatment. The process of wound healing was assessed on days 0, 3, 5, 7, and 14 after wound generation. On days 7 and 14, samples of wound tissue were harvested for hematoxylin and eosin (H&E) staining, Masson's trichrome staining, and/or immunofluorescent staining. Histological analyses of major organs including the heart, lungs, kidneys, liver, and spleen were conducted on day 14. For further details regarding these histological staining approaches, see the ESI.† The animal protocols received ethical approval (number 20240417006) from the Sichuan University Animal Care and Use Committee and abided by the Principles of Laboratory Animal Care regulated by the National Society for Medical Research.

2.8. Statistical analysis

Data are means ± SD from a minimum of three replicate analyses. One-way ANOVAs and Tukey's test were used to compare the results in GraphPad Prism. P < 0.05 was deemed significant (NS: not significant; *P < 0.05, **P < 0.01, ***P < 0.001).

3. Results

3.1. PPL synthesis

To prepare microgels responsive to pH/ROS levels, PBA grafting of the PL side chains was performed. Amidation reactions between the PBA –COOH group and the PL –NH₂ group were facilitated using EDC and NHS, and ¹H NMR analyses confirmed that PPL had been successfully synthesized (Fig. 2(a)). Relative to PL samples, samples of PPL presented with additional k–m peaks in the 7.00–7.50 (ppm) range attributable to the hydrogens present in BPA aromatic rings. A decrease in the area values for peak a for PPL samples was also observed as compared to PL samples, and a new peak f was observed in the 4.15–4.40 (ppm) range in these PPL samples attributable to the conversion of some amino groups into amide bonds following the amidation reactions. The calculated grafting ratio was 53%.

Fig. 2 PPZD fabrication and characterization. (a) ¹H NMR spectra for PL and PL-PBA. (b) TEM and (c) SEM images of ZIF-8, ZIF-8@DFO, and PPZD. (d) EDS images of PPZD (scale bar: 200 μm). (e) ZIF-8, ZIF-8@DFO, and PPZD sizes and ζ-potentials. (f) XRD and (g) UV-vis absorption analyses of PPZD, ZIF-8@DFO, and ZIF-8. (h) 0–72 h DFO release curves for PPZD and ZIF-8@DFO in PBS (pH 5.0). Data are means ± SD, n = 3, *P < 0.05.

3.2. PPZD fabrication and characterization

A one-pot synthesis approach was used to successfully prepare ZIF-8 and ZIF-8@DFO nanoparticles from 2-MIM, Zn(CH₃COO)₂·2H₂O, and DFO (Fig. 1). PPL was then used to coat ZIF-8@DFO surfaces, thereby fabricating PPZD (Fig. 2(b) and (c)). The grafting of PBA endows PPL with the capability to stably coat ZIF-8@DFO through π–π stacking interactions due to the presence of benzene rings,⁴⁹ as well as the formation of B–N coordination bonds between the boron in PBA and the nitrogen atoms of ZIF-8.⁵⁰ Sample morphology was assessed via SEM and TEM (Fig. 2(b) and (c), and Fig. S1, ESI†), revealing the regular dodecahedral structures of ZIF-8 and ZIF-8@DFO crystals consistent with those of synthetic ZIF-8. Elemental mapping analyses revealed that sulfur (S) characteristic of DFO was present in ZIF-8@DFO and PPZD preparations, whereas it was absent from pure ZIF-8 (Fig. 2(d) and Fig. S1, ESI†). Similarly, PBA-associated elemental boron (B) was present in PPZD samples but absent from ZIF-8 and ZIF-8@DFO (Fig. 2(d) and Fig. S1, ESI†). The S peak concentrations in the ZIF-8@DFO and PPZD samples were 0.10% and 0.06%, respectively. The respective diameters of ZIF-8, ZIF-8@DFO, and PPZD were measured as 174.1 nm, 190.6 nm, and 384.6 nm (Fig. 2(e)), and the corresponding ζ-potential values measured for these samples were 31.4 mV, 26.1 mV, and 17.8 mV (Fig. 2(e)). The lower ζ-potential of ZIF-8@DFO relative to ZIF-8 was potentially attributable to DFO molecules with partially obscured Zn²⁺ binding sites.³¹ Coating with PPL further decreased the ζ-potential of ZIF-8@DFO, which was attributed to several factors: first, PPL was grafted with PBA through NH₂ groups; second, the PPL chains likely increased the surface steric hindrance, thus reducing the density of the positive charge; lastly, as the coating thickness increased, the contact area available for cations on the ZIF-8@DFO surface decreased.

XRD analyses of these nanoparticle preparations revealed good consistency in the observed patterns as compared to characteristic peaks for simulated ZIF-8⁵¹ (Fig. 2(f)), suggesting that ZIF-8@DFO nanoparticle synthesis and PPL coating did not disrupt the standard ZIF-8 crystal structures.

DFO encapsulation and loading efficiency (EE and LE) in PPZD preparations were assessed based on DFO-mediated absorption of Fe³⁺, taking advantage of its ability to chelate Fe³⁺ and form the DFO–Fe³⁺ complex with a characteristic ∼430 nm absorption peak⁵² (Fig. 2(g)). The characteristic change in the color of the prepared solutions resulting from DFO–Fe³⁺ complex formation was used to prepare a standard curve combining 3 mM FeCl₃ and various DFO concentrations (0–300 μM) (Fig. S2, ESI†). Using this standard curve, the EE and LE for DFO in PPZD samples were respectively measured at 37.49 ± 1.79 wt% and 8.36 ± 0.48 wt%.

SEM, TEM, EDS mapping, ζ potential values, and specific UV-vis absorption peaks for DFO@ZIF-8 and PPZD confirmed the successful encapsulation of DFO within ZIF-8@DFO as well as the successful coating of ZIF-8@DFO with PPL.

The functionalization of ZIF-8@DFO nanoparticles aims to increase their efficacy while mitigating any associated adverse effects. The acidic microenvironment associated with infected chronic wounds can trigger rapid ZIF-8@DFO degradation, leading to the release of high levels of Zn²⁺ and DFO that can cause toxic side effects. Zn²⁺ and DFO release curves for ZIF-8@DFO and PPZD in PBS at pH 5.0 revealed that PPL coating was able to protect against this explosive release of Zn²⁺ and DFO under acidic conditions (Fig. 2(h) and Fig. S3, ESI†). Following incubation with ZIF-8@DFO and PPZD extracts for 24 and 48 h, viability assays and live/dead staining confirmed that the PPZD group exhibited fewer dead cells and greater viability, consistent with the ability of PPL coating to mitigate any toxicity stemming from rapid ZIF-8@DFO degradation (Fig. S4, ESI†).

3.3. MMS@PPZD construction

Through ¹H NMR analyses, successful HA used to synthesize HAMA was confirmed (Fig. 3(a)), with the resultant HAMA MS particles exhibiting sizes in the 190–230 μm range (Fig. 3(b)–(d)). Dynamic PPZD loading was achieved by using TP/Fe²⁺ MPN for the functionalization of HAMA MS through surface coating. Following this coating process, the previously colorless MS exhibited a bluish-purple coloration but no significant changes in their morphology or size on microscopic examination (Fig. 3(d)). Dynamic PPZD loading is achieved through reversible boronic ester formation between MMS phenolic hydroxyl groups and PPZD phenylboronic acid groups.⁵³ Successful RhB-DFO loading into HAMA MS particles was confirmed through fluorescence microscopy (Fig. 3(e)). SEM analyses indicated that MMS samples exhibited porous MS (Fig. S5, ESI†) that were encapsulated within an MPN network structure (white arrow) (Fig. 3(f)). EDS analyses confirmed the presence of iron derived from the TP/Fe²⁺ MPN on the MS surfaces (Fig. 3(h)). SEM analyses indicated that PPZD (red arrow) was homogenously distributed on the MMS surface (Fig. 3(g)), and the successful combination of MMS and PPZD and was confirmed based on visible Zn and B in the resultant EDS spectroscopic scans (Fig. 3(i)). One milliliter of freeze-dried MMS@PPZD was confirmed to contain 2.35 mg of PPZD.

Fig. 3 MMS@PPZD construction. (a) HA and HAMA ¹H NMR spectra. (b) Light microscopy images of MS. (c) HAMA MS size distribution. (d) Light microscopy images of MMS. (e) Fluorescence images of RhB-DFO and MMS@PPZD. SEM images of MMS (f) and MMS@PPZD (g). EDS images of MMS (h) and MMS@PPZD (i) (scale bar: 50 μm). (j) Images of MMS@PPZD adherence to different tissues. (k) DFO release rates from PPZD under the indicated conditions. Data are means ± SD, n = 3, *P < 0.05.

While a slight reduction in HAMA MS water retention capacity was observed following MPN coating (Fig. S6, ESI†), in a water loss test both the MMS and MMS@PPZD samples nonetheless presented with significant water retention activity attributable to the dense MS surfaces and the high capacity of HA for water retention. These results align well with prior studies of the capacity of similar materials for water retention.⁴⁶

Both wet and dry MMS@PPZD samples were able to adhere to tested organs including the lungs, kidneys, heart, liver, spleen, and skin (Fig. 3(j)), consistent with excellent adhesive properties.

3.4. Evaluation of Zn²⁺ and DFO release from the prepared microgels

Precisely regulated DFO release is vital for the ability of a prepared microgel to effectively promote angiogenic activity within infected diabetic wounds. The goal of this study was to generate a smart PPZD nanoplatform capable of dynamically and sequentially releasing different compounds to facilitate appropriate local drug delivery. Infected diabetic wounds often stall in the inflammatory phase of the healing process, establishing a microenvironment characterized by high ROS levels and acidic conditions.³ To achieve the in vitro simulation of these conditions, an H₂O₂-containing solution with a pH of 5.0 was prepared. Relative to incubation in PBS, significantly faster Zn²⁺ and DFO release was observed under conditions of low pH and H₂O₂ exposure (Fig. 3(k) and Fig. S3, ESI†). The release of Zn²⁺ was faster than that of DFO. ZIF-8 was also responsive to acidic conditions, and this dual pH- and ROS-responsive mechanism enabled the precise release of DFO, providing an opportunity to optimize therapeutic efficacy while minimizing the risk of adverse effects.⁵⁴ The incorporation of a PL coating also provided the microgel with antibacterial properties while delaying Zn²⁺ and DFO release (Fig. 2(h) and Fig. S3, ESI†), thereby preventing any cytotoxicity stemming from excessive Zn²⁺ and DFO release (Fig. S4, ESI†). The results of the DPPH assay of MMS@PPZD at different time points demonstrated that the ROS-scavenging activity increased rapidly within 24 hours, indicating the early and rapid release of TP (Fig. S7 and S8, ESI†). This strategy was thus able to achieve sustained, sequential release of TP, Zn²⁺, and DFO, thereby promoting angiogenesis after achieving ROS-scavenging and antibacterial effects.

3.5. In vitro analyses of the microgel's antibacterial activity

The MIC of PPZD was found to be 80 μg mL⁻¹ for S. aureus and 40 μg mL⁻¹ for E. coli. The OD 600 measurements showed that at PPZD concentrations of 160 μg mL⁻¹ or above, the antibacterial ratio exceeded 90% against both S. aureus and E. coli (Fig. S9, ESI†). According to the CCK-8 assays, significantly reduced L929 cell viability was observed at a PPZD concentration of 400 μg mL⁻¹ (Fig. S10, ESI†). Tube formation assays using HUVECs demonstrated optimal tube formation activity at PPZD concentrations of 200 and 300 μg mL⁻¹ (Fig. S11, ESI†). Ultimately, after weighing the considerations of cytotoxicity, antibacterial efficacy, and pro-angiogenic effects, we determined that the optimal concentration of PPZD for subsequent experiments was 200 μg mL⁻¹.

The prepared PPZD and MMS@PPZD samples exhibited strong antibacterial activity against S. aureus (>95.0%) and E. coli (>93.5%) attributable to PL and Zn²⁺ release (Fig. 4(a) and (b)). The antibacterial activity of these materials was additionally assessed by spreading the bacterial suspensions on plates and assessing S. aureus and E. coli colonies after incubation in the presence of these preparations for 12 h (Fig. 4(c) and Fig. S12, ESI†). These results were confirmed through the live/dead staining of these bacteria. The majority of the bacteria included in this assay were dead following incubation in the presence of PPZD and MMS@PPZD preparations (Fig. 4(d)), while significantly weaker antibacterial activity was evident in the absence of PPZD, confirming that the antibacterial activity of these preparations was primarily attributable to PL and Zn²⁺ incorporation.

Fig. 4 In vitro analyses of the antibacterial activity of the prepared materials. (a) and (b) Analyses of the antibacterial properties of different preparations when combined with S. aureus and E. coli. Data are means ± SD, n = 3, ***P < 0.001. (c) Images of agar plates from assays testing the antibacterial activity of the indicated preparations against S. aureus and E. coli. (d) Live–dead staining of S. aureus and E. coli in the indicated treatment groups (scale bar: 50 μm).

3.6. Analyses of the microgel's biocompatibility and antioxidant performance

L929 cells and HUVECs were selected as models to assess the biocompatibility of the synthesized microgels, given the key roles that these cell types play in angiogenesis, collagen production, formation of the ECM, and wound contraction.^3,55 Following incubation with various material extracts for 24 and 48 h, all of these materials were confirmed to exhibit excellent cytocompatibility through live–dead staining, as few dead cells were visible (Fig. 5(a), (b) and Fig. S13, ESI†). In hemolysis assays, the preparations also exhibited strong hematological biocompatibility, as evidenced by <5% hemolysis in the treatment groups (Fig. 5(c)), falling well within the ISO/TR 740654 safety guidelines.⁵⁶

Fig. 5 Evaluation of microgel biocompatibility and antioxidant properties. (a) and (b) CCK-8 and live–dead staining results for L929 cells under the indicated treatment conditions (scale bar: 100 μm). (c) Hemolysis assay results for the indicated treatment conditions. (d) and (e) L929 cell wound healing assay results in the indicated treatment groups (scale bar: 200 μm). (f) and (g) DPPH scavenging analyses for the indicated groups. (h) Representative DFCH-DA fluorescence in the indicated groups (scale bar: 50 μm). Data are means ± SD, n = 3 or 6, ***P < 0.001.

The process of wound healing is strongly dependent on cellular migration. To assess how the synthesized materials impact cellular migration, wound healing assays were thus performed, using H₂O₂ to mimic the oxidative stress and ROS accumulation attributable to localized inflammatory activity during the wound healing process, which can lead to apoptosis and senescence.³ After stimulation with H₂O₂ (100 μM), a scratch wound was generated in the L929 cell monolayer and the cells were observed (Fig. 5(d) and (e)). Significantly enhanced migration was observed for cells treated with MMS and MMS@PPZD preparations. This improved migratory activity is likely a result of the strong capacity of TP for ROS scavenging given the abundance of natural catechol/pyrogallol groups in TP. Compared with the control group, the PPZD and MS groups were also able to enhance cell migration. This was likely attributable to a certain degree of antioxidant activity in ZIF-8,⁵⁷ PL, and HA. Additionally, PL⁵⁸ and HA⁵⁹ have been confirmed to modulate biological processes that promote the migration of fibroblasts.

To confirm that the prepared MPN-coated materials (MMS and MMS@PPZD) exhibit antioxidant activity, a DPPH scavenging assay was performed in which both preparations were confirmed to exhibit significant capacity for DPPH scavenging (Fig. 5(f), (g) and Fig. S7, ESI†). These results are consistent with the migration assay findings, providing further support for the ability of TP to facilitate ROS scavenging. In patients with diabetes, hyperglycemia contributes to excessive ROS biogenesis, impairing the wound healing process.⁶⁰ The ability of the prepared TP-containing materials to protect L929 cells from ROS-induced damage was also evaluated after subjecting these cells to H₂O₂ treatment using DCFH-DA to detect ROS.⁶¹ Following incubation with these MPN-coated materials, L929 cells exhibited a reduction in DCFH-DA fluorescence relative to control H₂O₂-treated cells consistent with reduced levels of intracellular oxidative stress (Fig. 5(h)). TP can thus help attenuate oxidative stress in these cells.

Together, these data attest to the biocompatibility of these prepared TP-loaded microgels and their antioxidant activity when exposed to inflammatory microenvironmental conditions, enabling them to mitigate oxidative stress and promote fibroblast proliferation.

3.7. Assessment of the microgel's anti-inflammatory properties

Inflammatory activity is closely linked to enhanced ROS biogenesis, which can further perpetuate the cascade of inflammatory signaling.⁶² Macrophages are key mediators of local and systemic inflammation, in addition to serving as essential regulators of wound healing.⁶³ The ROS-scavenging antioxidant activity of TP was regarded as the primary source of anti-inflammatory activity for the prepared microgel system, inhibiting M1 macrophage polarization in favor of M2 polarization such that anti-inflammatory cytokine production is enhanced and pro-inflammatory cytokine production is suppressed.⁶⁴ Intracellular ROS levels were reduced following the TP-loaded microgel treatment of RAW 264.7 macrophages that were pre-activated with 100 μM H₂O₂ (Fig. S14, ESI†).

After pre-activating RAW 264.7 macrophages using LPS (5 μg mL⁻¹), these cells were incubated for 48 h in the presence of various material extracts. Then, qPCR (Fig. 6(f)–(g)) and immunofluorescent staining (Fig. 6(g)–(j)) were used to evaluate the M1 and M2 polarization of these cells based on iNOS and CD206 expression, respectively, revealing strongly enhanced M2 polarization in response to TP-loaded microgel treatment. While IL-10 is an anti-inflammatory cytokine capable of suppressing effector T cell responses,⁶⁵ TNF-α is a pro-inflammatory cytokine capable of stimulating ongoing inflammation.⁶⁶ These changes in TNF-α and IL-10 expression were confirmed at the mRNA level in the MMS and MMS@PPZD groups by qPCR (Fig. 6(a) and (b)), and the same shifts were observed through immunofluorescent staining (Fig. 6(c)–(e)). These results highlight the ability of TP loading to imbue the prepared microgels with robust anti-inflammatory activity.

Fig. 6 Evaluation of microgel anti-inflammatory activity. (a) and (b) IL-10 and TNF-α were detected by qPCR in the indicated groups. (c)–(e) Representative immunofluorescent staining for IL-10 and TNF-α with corresponding quantification (scale bar: 50 μm). (f) and (g) CD206 and iNOS levels were assessed by qPCR in the indicated groups. (h)–(j) Representative immunofluorescent staining for CD206 and iNOS with corresponding quantification (scale bar: 50 μm). Data are means ± SD, n = 3, ***P < 0.001.

3.8. Assessment of the microgel's angiogenic activity

The angiogenic process entails several important stages, including endothelial matrix degradation, endothelial cell (EC) migration and proliferation, the formation of lumen-containing ducts, and the onset of blood flow.⁶⁷ HUVEC wound healing and tube formation assays are widely used to model in vivo angiogenic activity. DFO has additionally been used to induce the proliferation of ECs and the growth of blood vessels during wound healing, as it is capable of stabilizing HIF-1α and upregulating VEGF.²⁹

In wound healing assays, the highest migration levels were evident in the DFO-loaded treatment groups (PPZD and MMS@PPZD) (Fig. 7(a) and (b)), and PPZD or MMS@PPZD treatment similarly enhanced HUVEC tube lengths in a tube formation, with DFO-loaded microgel treatment thus having significantly enhanced the overall degree of angiogenesis (Fig. 7(c) and (d)). The effects of hyaluronic acid (HA) and its derivative, HAMA, as well as TP, on the biological activity of HUVECs include their enhancement of cell proliferation, motility, and the promotion of VEGF secretion.^68,69 Consequently, both cell migration and tube formation in the MS and MMS groups showed some improvement compared to the control group. In line with these results, PPZD and MMS@PPZD extract-treated HUVECs exhibited significantly increased HIF-1α and VEGF expression after 48 h, as detected via qPCR (Fig. 7(e) and (f)) and immunofluorescent staining (Fig. 7(g)–(i)). MMS@PPZD was thus able to drive the effective upregulation of growth factors in ECs, providing a foundation for in vivo research aimed at validating these findings.

Fig. 7 Evaluation of the pro-angiogenic properties of prepared microgels. (a) and (b) HUVEC wound healing rate and scratch images in the indicated treatment groups (scale bar: 200 μm). (c) and (d) Tube formation assay images and corresponding quantification (scale bar: 200 μm). (e)–(i) HIF-1α and VEGF expression in the indicated groups was assessed by qPCR (e) and (f), and immunofluorescence (g)–(i) (scale bar: 50 μm). Data are means ± SD, n = 3 or 6, ***P < 0.001.

3.9. Evaluation of the in vivo wound healing performance of the prepared microgels

To test the suitability of the prepared microgel system as a tool for the management of infected diabetic wounds in vivo, a chronically infected diabetic wound model was established in rats (Fig. 8(a)), using S. aureus and E. coli as model pathogens. More rapid wound closure was evident in the MMS@PPZD treatment group relative to other groups (Fig. 8(b)), with 80% closure on day 7 and almost complete wound closure on day 14. The superiority of MMS@PPZD treatment was confirmed through quantitative analyses (Fig. 8(d)). Similarly improved wound closure rates were observed for both the dry MMS@PPZD and wet MMS@PPZD treatment groups (Fig. S15, ESI†), with these benefits being attributable to the hydrating, antibacterial, anti-inflammatory, angiogenic, and antioxidant activity of these microgel preparations due to their incorporation of PL, Zn²⁺, TP, DFO, and HAMA.

Fig. 8 Evaluation of the in vivo wound healing activity of the prepared microgels. (a) Wound images at different time points and corresponding wound bacterial cultures on day 7. (b) Images of wounds and corresponding traces of the healing process on days 0, 3, 5, 7, and 14 in the indicated groups. (c) Quantification of the antibacterial efficiency of different biomaterials relative to the blank control. (d) Wound healing rates in the indicated treatment groups over time. Data are means ± SD, n = 3, **P < 0.01, ***P < 0.001.

To evaluate the in vivo antibacterial activity of the prepared biomaterials, quantification of the bacterial load in the wound site was performed on day 7. Significantly fewer bacteria were present in PPZD and MMS@PPZD-treated wounds, confirming the robust antibacterial activity of the PL and Zn²⁺ in these microgel preparations (Fig. 8(c)). A limited amount of antibacterial activity was also evident for MPN in the MMS, as reported previously.⁷⁰

Histological analyses of regenerated skin quality were performed on days 7 and 14. Through H&E and Masson's trichrome staining, the wound sites from animals in the MMS@PPZD treatment group were found to harbor fewer inflammatory cells, more fibroblasts, and greater vascularity as compared to other experimental groups (Fig. 9(a) and Fig. S16, ESI†). These improvements are likely related to the antibacterial activity achieved through rapid PL and Zn²⁺ release, the suppression of inflammation mediated by MPN, and the induction of angiogenesis through the sustained release of DFO. As of day 14, wounds in the control group were surrounded by substantial granulation tissue, whereas a largely regular epithelial layer with near-total dermal regeneration was evident in the MMS@PPZD treatment group. Masson's trichrome staining revealed significantly enhanced collagen deposition in the MMS@PPZD group relative to the control group on day 14 (Fig. 9(a)). Collagen fibers in the MMS@PPZD group were denser, thicker, and more regularly arranged, and the samples from this group also presented with a thicker epidermal layer, more hair follicles, and significantly greater vascular density (Fig. 9(b)–(e)).

Fig. 9 Assessment of the in vivo wound healing and antibacterial properties of different biomaterials. Yellow arrows: newly regenerated blood vessels; blue arrows: newly regenerated hair follicles. (a) Representative H&E and Masson's trichrome staining results from day 14 (scale bar: 200 μm). Quantification of epithelial thickness (b), hair follicle growth rates (c), collagen deposition (d), and vascular density (e) in the indicated groups. Data are means ± SD, n = 3, ***P < 0.001.

Immunofluorescence staining provided further confirmation of the higher levels of collagen I deposition in the MMS@PPZD group (Fig. 10(a) and (e)), emphasizing the synergetic effects of different microgel components. This synergy is likely attributable to the anti-inflammatory activity of MPN in combination with the robust angiogenic and antibacterial effects of PPZD throughout the process of wound healing.

Fig. 10 Evaluation of the ability of the prepared microgels to promote in vivo wound healing. (a)–(g) Representative iNOS, TNF α, IL-10, Col-I, VEGF, and CD31 staining images on day 14 (a) (scale bar: 50 μm) and the corresponding quantification (b)–(g). Data are means ± SD, n = 3, ***P < 0.001.

Macrophage M1 and M2 polarizations are respectively linked to pro- and anti-inflammatory activity within a given microenvironment, with iNOS serving as an M1 macrophage biomarker. Markedly reduced iNOS expression was evident in the wound site in the MMS and MMS@PPZD treatment groups relative to the control group (Fig. 10(a) and (b)). Moreover, higher TNF-α levels were observed in wounds in the control group, whereas they were reduced in the MMS@PPZD group, while the opposite was true for IL-10 (Fig. 10(a), (c) and (d)).

CD31 and VEGF were used as biomarkers of angiogenesis in the indicated groups, both of which were upregulated in the MMS@PPZD group relative to the control group on day 14 (Fig. 10(a), (f) and (g)). This upregulation provided further confirmation of the ability of the MMS@PPZD system to induce angiogenesis via the sustained release of DFO following the elimination of pathogenic bacteria in the wound site and the abrogation of inflammatory activity.

The established MMS@PPZD system presented with strong drug release activity that was spatiotemporally controlled, inducing the M2 polarization of macrophages while inhibiting their M1 polarization. This resulted in higher levels of IL-10 production and reduced TNF-α release, ultimately suppressing inflammation. This, in turn, led to the eventual upregulation of CD31 and VEGF expression and associated angiogenic activity, leading to the enhanced healing of infected chronic diabetic wounds.

3.10. Assessment of in vivo microgel biocompatibility

Microgels must exhibit a satisfactory safety profile in order to be feasibly used for the treatment of infected wounds. To test for any potential side effects associated with the use of these microgels, histomorphological analyses of major organs (lungs, kidneys, liver, heart, spleen) were performed on day 14 after treatment. H&E staining of these tissues revealed no evidence of any histological anomalies or other adverse effects in any of the treatment groups (Fig. 11). The prepared microgels thus exhibit a high degree of in vivo biosafety.

Fig. 11 Evaluation of in vivo microgel toxicity. (a) Histologic analyses of the heart, liver, spleen, lungs, and kidneys following the indicated treatments for 14 days (scale bar: 200 μm).

4. Conclusion

In this study, a versatile microgel-based system was developed that can be deployed to treat infected diabetic wounds in the form of either an injectable wet microgel or dry powder MS preparation. This system was established by engineering Fe²⁺/TP MPN-functionalized HAMA MS, yielding the innovative MMS@PPZD system with robust anti-inflammatory and ROS scavenging properties. The PPZD component of this system consists of a PBA-functionalized PL coating on ZIF-8@DFO, with boron ester bonds used to load into onto MPN-functionalized MS. The resultant microgel system can thereby respond to microenvironmental conditions, releasing PPZD in a dynamic fashion. The PPL coating process also simultaneously imbues the microgel with stronger antibacterial activity while preventing the excessive and cytotoxic release of Zn²⁺and DFO under acidic conditions, instead promoting gradual, sustained Zn²⁺and DFO release.

In summary, this novel microgel-based drug delivery platform exhibits advantageous environmental responsivity and phased interventional activity that aligns well with the different stages of infected wound healing. When used to treat infected diabetic wounds, these microgel preparations were able to obviate local inflammation and oxidative stress, eliminate bacteria, induce angiogenesis, and promote collagen deposition to accelerate the overall process of wound healing. Importantly, the biocompatibility of this microgel was excellent in vitro and in vivo, and it was able to readily abrogate inflammatory activity by releasing TPs, kill bacteria through the release of PL and Zn²⁺, and promote angiogenesis through the sustained, regulated release of DFO. These results highlight the promise of MMS@PPZD as a multifunctional tool for the care of diabetic wounds, while also emphasizing the feasibility of exploring the development of other environmentally-responsive, sequentially-activated biomaterials.

Author contributions

Ma Fei and Yuheng Liu: conceptualization, data curation, investigation, methodology, resources, software, validation, writing – original draft. Wang Yu: data curation, formal analysis, investigation, validation. Chirume, Walter Munesu: data curation, formal analysis, investigation, validation. Dengbo Yao: investigation, methodology, validation. Weiqiang lan and Zhen Zhao: data curation, formal analysis, investigation. Xueyuan Xu and Weifei Zhang: data curation, formal analysis. Chuan Guo: conceptualization, funding acquisition, investigation, project administration, resources, supervision, writing – review & editing. Qingquan Kong: conceptualization, funding acquisition, investigation, project administration, resources, supervision, writing – review & editing.

Data availability

Data will be made available on request.

Conflicts of interest

The authors declare no competing financial interest.

Acknowledgements

This work was partially supported by the Sichuan Science and Technology Program (2024NSFSC1814), the China Postdoctoral Science Foundation (GZC20231805), the National Natural Science Foundation of China (82202747, 823724478), Post-Doctor Research Project, West China Hospital, Sichuan University (2023HXBH077, 2023HXBH082), the Science and Technology Major Project of Tibetan Autonomous Region of China (XZ202201ZD0001G), and the Science and Technology Project of Tibet Autonomous Region (XZ202301YD0027C). The authors would like to thank and express their heartfelt gratitude to Xuanhe You, Diwei Wu, Mingjie Xu from the Orthopedic Research Institute (Core Facilities of West China Hospital, Sichuan University), and Li Chai, Yi Li, and Xing Xu from the Core Facilities of West China Hospital, Sichuan University for their assistance and guidance.

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Footnotes

† Electronic supplementary information (ESI) available. See DOI: https://doi.org/10.1039/d4tb02474e

‡ These authors contributed equally to this work and shared the first authorship.

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