{"paper_id":"0979bace-953c-4e09-95c3-00be686a93cd","body_text":"A Core–Shell Microneedle Platform for the Spatiotemporal Codelivery of Dual-Agent Therapeutics Precisely Orchestrates Diabetic Wound Healing | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Research Article A Core–Shell Microneedle Platform for the Spatiotemporal Codelivery of Dual-Agent Therapeutics Precisely Orchestrates Diabetic Wound Healing Yuhang Zhan, Zhihan Zhang, Zhiyue Zhang, Hui Liu, Mengru Lin, and 6 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-7689935/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 17 Mar, 2026 Read the published version in Journal of Nanobiotechnology → Version 1 posted 10 You are reading this latest preprint version Abstract Chronic non-healing of diabetic wound (DW) remains a critical clinical challenge worldwide. Sustained oxidative stress and prolonged inflammatory responses disrupt the wound microenvironment, while bacterial colonization and biofilm formation on the wound bed compromise drug penetration, consequently leading to suboptimal outcomes with conventional approaches. Here, we developed a core-shell structured microneedle (MN) patch system, designated as MN@Ple/Exo Q10 , to precisely regulate the DW microenvironment through sequential drug release. The MN@Ple/Exo Q10 features a biphasic agent release profile, comprising a natural antimicrobial peptide (Pleurocidin) in the outer shell layer, that exerts anti-infective effects during the initial phase of wound healing, while engineered exosomes (Exos Q10 ) in the core layer alleviate oxidative stress and modulate immune responses subsequently. This study details that the sustained release of Exos Q10 effectively inhibits high glucose (HG)-induced ferroptosis in vitro, demonstrating potent antioxidant activity and anti-inflammatory capacity. Furthermore, in a S. aureus -infected diabetic mouse wound model, MN@Ple/Exo Q10 demonstrates potent antibacterial activity while mitigating oxidative stress, suppressing inflammation and promoting angiogenesis, thereby accelerating wound healing. Collectively, the developed spatiotemporally controlled MN system overcomes bacterial barriers and stabilizes exosomal delivery, enabling comprehensive regulation of the microenvironment in DWs. This breakthrough approach presents a novel and translational strategy for DW therapy. Core-shell Microneedle Diabetic wound healing Exosomes Ferroptosis Antibacterial activity Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Figure 8 Figure 9 1. Introduction Diabetes, a metabolic disorder, has exhibited a steady increase in global prevalence over the past few decades, with projections indicating that the number of affected individuals will reach 693 million by 2045 [1] . Over 30% of individuals with diabetes are likely to develop DWs [2] , which are characterized as chronic, non-healing lesions primarily resulting from diabetic neuropathy and vascular complications [3] . Notably, the management of DW is a prolonged and complex process, and severe diabetic foot ulcers frequently culminate in amputation [4] , imposing substantial clinical, caregiving, and economic burdens on society [5] . DWs are typified by hyperglycemia, ischemia, hypoxia, excessive inflammation, and persistent infection [6] . Single interventions such as pharmacological agents, wound dressings or surgical procedures are insufficient to modulate the pathological and physiological disturbances within the wound microenvironment [7] . Therefore, there is an urgent need to develop an ideal therapeutic strategy capable of comprehensively addressing these multifaceted pathophysiological challenges in order to accelerate DW healing. Exosomes (Exos), double-layer lipid membrane-bound extracellular vesicles, mediate intercellular communication through the delivery of functional proteins [8] , and Exos extracted from human umbilical cord mesenchymal stem cells (HucMSCs) have garnered significant attention in the field of regenerative medicine due to the capacity for tissue repair and immune modulation [9–10] .Nevertheless, the HucMSC-derived Exos exhibited minimal efficacy in mitigating HG-triggered ferroptosis of endothelial cells within the wound bed—a programmed cell death process marked by pathological reactive oxygen species (ROS) and Fe 2+ overload, exacerbating oxidative damage and immune imbalance in DWs [11] . Therefore, preparing engineered HucMSC-derived Exos by pretreating HucMSCs to enhance their capacity to inhibit HG-induced ferroptosis is crucial for the repair of DWs. Previous studies have demonstrated that pretreatment of mesenchymal stem cells (MSCs) with pharmacological agents [12] , cytokines [13] and physical stimuli [14] represents an effective strategy to improve the biological activity and therapeutic functionality of MSCs. Coenzyme Q10 (CoQ10), an endogenously synthesized lipophilic antioxidant, potently suppresses ferroptosis [15] and may serve as a potential enhancer for therapeutic Exos. Here, we hypothesize that the engineered Exos prepared by pre-stimulating HucMSCs with CoQ10 could inhibit ferroptosis associated with DWs and accelerate wound healing. Notably, simple Exo therapy cannot effectively target DWs infected by bacteria [16] , as bacterial pathogens and their biofilms can significantly compromise Exos functionality [17] . Thus, targeted strategies to enhance antimicrobial capacity are critically needed. Antimicrobial peptides (AMPs) are unique molecules with molecular weights below 10 kDa that have attracted considerable attention as potential alternatives to conventional antibiotics due to their broad-spectrum antimicrobial activity and high specificity [18] . Pleurocidin (Ple), a natural antimicrobial peptide isolated from various species of Atlantic flounder, is composed of 25 amino acid residues [19] . Previous studies have demonstrated that Ple exhibited broad-spectrum antimicrobial activity with low cytotoxicity, disrupted bacterial membranes, interfered with intracellular processes, and inhibited biofilm formation [20]−[21] . However, as a linear peptide, Ple is susceptible to inactivation and denaturation in the complex wound microenvironment, while its molecular weight (2.7 kDa) hinders effective transdermal delivery through the stratum corneum via conventional formulations. Consequently, conjugating Ple with functional polymers to develop multifunctional biomedical materials that preserve antimicrobial efficacy while enabling localized delivery is a promising approach [22] . Microneedle (MN) patches represent a promising delivery platform capable of preserving the bioactivity of therapeutic agents while enabling multifaceted and integrated treatment approaches [23] , whereas conventional MN-based therapies are limited by either single-agent loading or simple blending of multiple agents, which often leads to constrained agent-loading capacity, physicochemical incompatibility between agents, and conflicting release kinetics [24]−[25] . Therefore, to address the complex pathophysiological microenvironment of DW, the development of temporally differentiated release systems is imperative for achieving synergistic multi-component combination therapy. Here, considering the distinct dynamic functions of different components in the DW microenvironment during each healing phase, we developed a near-infrared (NIR)-responsive core-shell structured MN system with biphasic agent release capability, designated as MN@Ple/Exo Q10 . Specifically, Ple was encapsulated into Gelatin-methacryloyl (GelMA) utilizing freeze-drying and spraying technology to form the shell layer of MN@Ple/Exo Q10 , while the core layer was constructed by incorporating freeze-dried engineered Exos Q10 into methacryloyl hyaluronic acid (HAMA). Furthermore, dopamine (DA) was self-polymerized and incorporated into both the shell and core layers of the MN patch as polydopamine nanoparticles (PDA NPs), endowing the MN patch with superior tissue adhesion and photothermal conversion capabilities. The process in which NIR irradiation is converted into localized hyperthermia is defined as photothermal therapy (PTT) [26] , effectively killing bacteria by disrupting their cell membranes and denaturing intracellular proteins and enzymes while also promoting wound healing [27] . Significantly, this system aims to sequentially deliver therapeutic agents to modulate the pathological microenvironment of DW. On one hand, penetration of MN@Ple/Exo Q10 through the wound barrier causes rapid GelMA shell swelling and Ple release, which synergizes with PTT to concurrently eradicate bacteria and disrupt biofilms. On the other hand, the HAMA core layer undergoes slow degradation following the breakdown of the shell, ensuring sustained and stable release of Exos Q10 within the wound bed. Accordingly, the released Exos Q10 suppress ferroptosis associated with DWs, mitigate oxidative stress, and modulate immune responses, thereby accelerating wound healing in diabetic mice. Overall, this innovative sequential microneedle therapeutic strategy demonstrates considerable potential for the effective management of chronic non-healing diabetic wounds. 2. Results and discussion 2.1. Preparation and Characterization of MN patch As illustrated in Fig. 1 A, MN@Ple/Exo Q10 consisted of a Gelatin methacryloyl (GelMA)-based MN shell layer loaded with Ple via freeze-drying and spray techniques and methacryloyl hyaluronic acid (HAMA)-based MN core layer encapsulating Exos Q10 , and the detailed preparation protocol for MN@Ple/Exo Q10 is provided in the Methods section. The rational design of distinct structural differences between the shell and core layers is critical for achieving stage-specific agent release in diabetic wound treatment. GelMA, a gelatin derivative modified with methacryloyl (MA) groups, exhibits excellent drug loading/storage capacity and biocompatibility [28] .Moreover, GelMA with a higher degree of methacryloyl substitution for constructing the MN shell layer possesses greater mechanical strength, thereby facilitating more effective skin penetration [29]−[30] . Hyaluronic acid (HA), a non-sulfated glycosaminoglycan (GAG), is a major component of the extracellular matrix (ECM) in skin tissue [31] , chemically modified into HAMA through esterification with MA. HAMA retains the intrinsic properties of HA, including high hydrophilicity, moisturizing capacity and wound healing functionality, while showing improved stability and resistance to enzymatic degradation, thus ensuring the structural integrity and sustained release of Exos [32]−[33] , which is crucial for the core layer of the MN patch. Figure 1 B presented a representative image of the MN patch, consisting of 100 needle tips arranged in a 10 × 10 configuration. Optical microscopy revealed that the needle tips displayed conical morphology, and the incorporation of PDA imparted a black coloration to the MN surface, as shown in Fig. 1 C. The MN patch demonstrated a tip height of 650 µm, a base diameter of 500 µm, and a tip-to-tip spacing of 550 µm. Further morphological characterization of MN@Ple/Exo Q10 was performed through a scanning electron microscope (Fig. 1 D). Subsequently, the fluorescent dyes 5(6)-Carboxyfluorescein and Rhodamine B were employed to label the GelMA shell layer and the HAMA core layer, respectively. The core-shell structure of the MN patch was confirmed using LSCM (Fig. 1 E). Fluorescence imaging showed a uniform distribution of green and red signals on the outer layer and core of the needle tips, respectively, thereby confirming the successful fabrication of the core-shell MN patch. Furthermore, Energy-dispersive X-ray Spectroscopy (EDS) elemental mapping demonstrated that carbon (C), nitrogen (N), and oxygen (O) were evenly distributed across the MN patch, whereas sodium (Na) and silicon (Si) exhibited low concentrations and showed no signs of aggregation (Fig. S1 ). The mechanical strength of the prepared MN patches was evaluated through compression testing on a universal testing machine. Force-displacement curves were illustrated in Fig. 1 F. The blank MN with a core-shell structure exhibited a continuous upward deformation trend and a higher stress load compared to the blank MN (Core) composed solely of the HAMA core layer. The compressive force at full-tip-height deformation for the blank MN was approximately 1.25 N/Needle, significantly higher than that of the blank MN (Core) (0.94 N/Needle), which indicated that the GelMA shell layer contributed substantially to the mechanical strength of the MN patch, endowing it with excellent mechanical properties that facilitate effective penetration of human skin [34] . The compression testing results revealed continuous force-displacement profiles, indicating robust structural stability of the MNs post-skin penetration without fracture risk. Additionally, the MN@Ple/Exo Q10 maintained a force-displacement behavior comparable to that of the blank MN, achieving a mechanical performance of 1.36 N/Needle—superior to that of the blank MN. This observation demonstrated that agent loading did not compromise the mechanical integrity of the MN patch, while the freeze-drying cycles involved in the agent-loading process could enhance the hardness of MN tips. As displayed in Fig. 1 G, MN@Ple/Exo Q10 was effectively inserted into the skin of mice, forming uniformly distributed microchannels. The structural integrity of the needle tips was further confirmed by optical microscopy. To evaluate adhesion performance, peel adhesion forces were measured among different MNs. The MN@Ple/Exo Q10 and blank MN exhibited maximum peel adhesion forces of 0.27 N and 0.28 N, respectively, indicating excellent and comparable adhesive properties. In contrast, the blank MN (Core) showed a significantly lower maximum peel adhesion force of only 0.02 N (Fig. S5). This difference might be attributed to the presence of collagen-derived RGD sequences on the GelMA molecular chains within the MN patch shell layer, which can interact with functional groups on the tissue surface, thereby enabling strong and stable adhesion to biological tissues [35] . As demonstrated in Fig. 1 H, the GelMA shell layer coated with Ple and the HAMA core layer loaded with Exos Q10 exhibited distinct in vitro release profiles in PBS solution. The GelMA shell layer showed a rapid release profile of Ple, with a cumulative release rate of 81.69 ± 3.01% achieved within 8 hours and a plateau reached by 24 hours (Fig. 1 I). This rapid release behavior was attributed to the surface loading of Ple through the freeze-drying and spray method, which facilitated efficient sustained release within a short timeframe and provided a clear advantage over conventional drug-loading strategies [36] . By contrast, Exos Q10 from the HAMA core layer showed a sustained release profile, with cumulative release rates of 18.27 ± 2.81%, 38.80 ± 3.77%, and 60.83 ± 2.41% observed on days 4, 10, 18, respectively (Fig. 1 J). This slow yet consistent and prolonged release pattern ensured the continuous delivery of Exos Q10 . According to the above findings, Ple underwent a burst release during the initial phase upon MN application to the wound, rapidly attaining a plateau, whereas Exos Q10 in the core layer revealed a sustained release throughout the entire duration of wound healing. The integration of these two release systems with distinct kinetic profiles conferred MN@Ple/Exo Q10 with significant potential for spatiotemporal therapeutic applications. The photothermal functionality was engineered through incorporation of PDA within the MN patch [37] . As depicted in Fig. 1 K, infrared thermographs of MN@Ple/Exo Q10 were obtained at various time points under two irradiation power densities (1 W/cm 2 and 0.5 W/cm 2 ) to monitor temperature fluctuations. Following separate laser irradiations for 5 minutes at both power densities, the temperature of MN@Ple/Exo Q10 increased from the baseline (25.0°C) to 46.8°C and 36.8°C, respectively, both of which fall within the therapeutic temperature range suitable for mild photothermal therapy in vivo [38] . Furthermore, photothermal curve analysis revealed that the 1 W/cm 2 group exhibited a faster temperature rise rate compared with the 0.5 W/cm 2 group (Fig. 1 L), indicating enhanced photothermal response efficiency and temperature tunability of MN@Ple/Exo Q10 . Additionally, thermal cycling experiments demonstrated that following five cycles of laser activation and deactivation at 1 W/cm 2 using an 808 nm laser, the maximum temperature achieved by the MN patch did not show significant degradation, with a consistently stable heating rate observed in each cycle (Fig. 1 M). This robust photothermal stability supported the potential for repeated applications in photothermal therapy [39] . 2.2. Characterization and internalization determination of Exo Q10 As detailed in the Methods section, Exos derived from CoQ10-preconditioned HucMSCs were prepared and subsequently characterized in terms of morphology, size distribution, and marker protein expression utilizing transmission electron microscopy (TEM), nanoparticle tracking analysis (NTA), and Western blotting, respectively. As shown in Fig. 1 N, HucMSC-Exos Q10 exhibited a spherical ultrastructural morphology and demonstrated an average particle size of approximately 124.8 nm (Fig. 1 O). Western blot analysis was performed to evaluate the expression of exosomal-specific markers, including the transmembrane protein CD63, the cytoplasmic protein TSG101, and the negative marker Calnexin. The results indicated that both CD63 and TSG101 were positively expressed in HucMSC-Exos and HucMSC-Exos Q10 with no significant difference between the two groups, whereas the negative marker Calnexin was detected exclusively in the HucMSCs group (Fig. 1 P), confirming the successful isolation of HucMSC-Exos Q10 and their acceptable purity. Subsequently, the cellular internalization of HucMSC-Exos Q10 was evaluated employing human umbilical vein endothelial cells (HUVECs) and L929 cells as in vitro models. Following a 6-hour co-incubation period with Dil-labeled HucMSC-Exos Q10 , LSCM revealed a uniform distribution of Dil-Exos around the nuclei of both cell types (Fig. 1 Q and R), thereby confirming the efficient uptake of HucMSC-Exos Q10 by these cells. 2.3. In vitro antibacterial performance of MN patches Bacterial infections significantly impede the healing process of diabetic wounds. Compared to traditional antibiotics, the antimicrobial peptide, Ple, exhibits broad-spectrum antibacterial activity and remarkable inhibitory effects against both Gram-positive and Gram-negative bacteria. Ple targets the bacterial cell membrane through a distinct mechanism, leading to disturbances in membrane structure and disruption of bacterial physiological functions, which substantially reduces the likelihood of developing bacterial resistance [40] . Moreover, the MN patch designed in this study exhibited photothermal conversion performance due to the incorporation of PDA, thereby enhancing the antibacterial function and promoting difficult diabetic wound healing. In this study, Staphylococcus aureus ( S. aureus ) and Escherichia coli ( E. coli ) were chosen as representative strains of Gram-positive and Gram-negative bacteria, respectively, to assess the in vitro anti-infective efficacy of MN@Ple/Exo Q10 . The plate counting method was employed to assess the survival rates of two bacterial species following exposure to various treatment conditions. Numerous colonies of S. aureus and E. coli were evident on the agar plates in both the control and MN groups. By contrast, the MN@Ple and MN@Ple/Exo Q10 groups demonstrated a significant reduction in colony counts relative to the control group, with the lowest bacterial density observed in the MN@Ple/Exo Q10 +NIR group (Fig. 2 A). The bacterial survival rates of S. aureus in the control, MN, MN@Ple, MN@Ple/Exo Q10 and MN@Ple/Exo Q10 +NIR groups were 100.70 ± 1.34%, 98.25 ± 2.52%, 16.71 ± 0.59%, 15.76 ± 0.36% and 1.71 ± 0.36%, respectively. Comparatively, the bacterial survival rates of E. coli in the same groups were 100.10 ± 1.10%, 100.40 ± 1.26%, 23.21 ± 0.73%, 18.90 ± 0.21% and 1.86 ± 0.17%, respectively (Fig. 2 B). As presented in Fig. 2 D and E, bacterial absorbance at a wavelength of 600 nm was measured hourly over a 9-hour period, and the resulting data were used to construct growth curves for S. aureus and E. coli . The absorbance growth of both bacterial strains in the control and MN groups was markedly higher than that in the other groups. The maximum absorbance values for S. aureus and E. coli in the control group reached approximately 1.63 and 1.52, respectively. In contrast, bacterial growth in the MN@Ple and MN@Ple/Exo Q10 groups was significantly inhibited, with absorbance increases plateauing around 6 hours post-treatment. Notably, bacterial activity in the group subjected to 10 minutes of NIR irradiation remained consistently low, with absorbance values for S. aureus and E. coli maintained at approximately 0.11 and 0.12. Subsequently, live/dead fluorescence staining was conducted on the two bacterial species across different experimental groups, followed by observation and imaging via LSCM, with quantitative analysis conducted through heat map visualization. The results demonstrated that live bacteria labeled with SYTO 9, which emitted green fluorescence, were mainly present in the control and MN groups, whereas minimal red fluorescence, indicative of dead bacteria labeled with propidium iodide (PI), was detected in these groups. In contrast, green fluorescence was markedly reduced in the MN@Ple and MN@Ple/Exo Q10 groups, while red fluorescence showed a moderate increase. Additionally, in the NIR group, the vast majority of bacteria exhibited intense red fluorescence, indicating the most potent antibacterial effect (Fig. 2 C). Figure 2 F showed the SEM images of S. aureus and E. coli after treatment in different groups. The two bacterial species in the control group possessed smooth surfaces and intact cell membranes, suggesting intact cellular morphology. MN treatment did not induce significant morphological alterations in the two bacterial species, while in the MN@Ple and MN@Ple/Exo Q10 groups, some S. aureus and E. coli cells displayed mild surface wrinkles and structural deformations. However, both bacterial species in the NIR group demonstrated irregular morphological shrinkage and severe membrane disruption, collectively indicating substantial impairment of bacterial integrity. The biofilm formed during a wound infection protects bacteria from conventional antibiotics and contributes to impaired wound healing. Therefore, the capability to disrupt bacterial biofilms is an essential characteristic of an effective antibacterial material applied to wounds [41] . In this study, the formation of bacterial biofilms in S. aureus and E. coli was examined using LSCM and crystal violet staining. As shown in Fig. 2 G, live/dead staining of the biofilms formed by S. aureus and E. coli revealed that both the control and MN groups exhibited continuous biofilm layers with high structural integrity, as evidenced by the strong green fluorescence. In contrast, the biofilms in the MN@Ple and MN@Ple/Exo Q10 groups displayed decreased green fluorescence and reduced continuity. Remarkably, treatment with MN@Ple/Exo Q10 +NIR resulted in severe disruption and dispersion of the biofilms in both bacterial strains, accompanied by intense red fluorescence, indicating extensive bacterial cell death. For crystal violet staining, the control group exhibited complete purple biofilms of S. aureus and E. coli , with absorbance values of approximately 2.61 and 2.49, respectively, as determined by quantitative analysis. Conversely, biofilm formation in the NIR group was nearly undetectable, showing significantly reduced absorbance values of approximately 0.47 and 0.49 for S. aureus and E. coli , respectively, which were markedly lower than those observed in the MN@Ple/Exo Q10 group (1.34 and 1.22) (Fig. 2 H and I). Overall, MN@Ple/Exo Q10 demonstrated significant antimicrobial activity against both Gram-negative and Gram-positive bacteria, which could be attributed to the synergistic effects of Ple and PTT. On the one hand, the amphiphilic α-helical structure of the antimicrobial peptide Ple, derived from flounder, induces membrane perturbation and permeabilization, enabling Ple to enter bacterial cells and bind to DNA, thus interfering with normal cellular functions [42] . On the other hand, the PTT of the MN patch induced by NIR irradiation can rapidly elevate the local temperature of the MN patches, triggering abnormal expression of bacterial heat shock proteins (HSPs) and depleting the bacteria’s repair capacity [43] . Furthermore, the combination of PTT and Ple effectively disrupted the extracellular polymeric substance (EPS) in bacterial biofilms, and the thermal effect of periodic NIR irradiation then enabled Ple to rapidly penetrate into the damaged interior of the biofilm, allowing for a thorough attack on the bacteria [44] . 2.4. The in vitro biocompatibility of MN patches A co-culture model based on HUVECs and L929 cells was established in the transwell chamber under different grouping conditions (Fig. 3 A). The in vitro compatibility of the MN patches in different groups was verified employing live/dead cell staining, a cell counting kit-8 (CCK-8) assay and hemolysis test. As illustrated in Fig. 3 B, live/dead cell staining was conducted on L929 cells pre-treated for 24 and 72 hours with different groups. The results indicated that none of the MN groups interfered with the normal proliferation of L929 cells, moreover, no significantly increased number of red fluorescent-labelled dead cells were observed in any of the experimental groups compared with the control group. The CCK-8 assay demonstrated a modest reduction in cell proliferation activity over 24 hours and 72 hours within the MN@Ple, MN@Ple/Exo Q10 and MN@Ple/Exo Q10 +NIR groups, presumably due to the addition of Ple. Notably, the cell viability across all MN groups remained above 95% at each time point (Fig. 3 C). For the hemolysis test, the positive control group treated with ddH 2 O exhibited evident red blood cell lysis, whereas the negative control group treated with PBS showed no hemolytic activity. No significant hemolysis was observed in any of the remaining MN groups (hemolysis ratio < 5%), and NIR irradiation did not elicit any notable hemolysis either (Fig. 3 D). The above biocompatibility test results indicated that the primary materials utilized in the fabrication of MNs demonstrated favorable in vitro biocompatibility. As a member of the antimicrobial peptide family, Ple has demonstrated excellent biocompatibility in previous research [45] . When incorporated at an appropriate concentration (20 µM) into the prepared MN patch, it demonstrated minimal cytotoxicity and hemolytic activity, thereby satisfying established human safety standards [46] . In addition, the safety features of MNs remained unaffected by the administration of Exos and appropriate NIR hyperthermia. Overall, the MN@Ple/Exo Q10 developed in this study demonstrated adequate suitability and safety, meeting the fundamental criteria necessary for clinical applications in wound treatment. 2.5. MN patches promoted wound healing and angiogenesis in vitro As demonstrated in Figure (Fig. 3 E), the impact of various groups of MNs on cell migration and angiogenesis in vitro was evaluated through scratch assays, migration assays and tube formation assays. The scratch wound healing experiment revealed that the migration rate of L929 cells after 24 hours in the HG-treated control group (39.85 ± 2.78%) was significantly lower than that observed in the negative control group (74.59 ± 1.74%) treated with normal culture medium. This discrepancy was attributed to the oxidative stress state and ferroptosis of cells induced by the HG environment in vitro, which subsequently resulted in a reduced cell migration speed [47] . The cell migration rates corresponding to the MN, MN@Ple, MN@Ple/Exo Q10 and MN@Ple/Exo Q10 +NIR groups were determined quantitatively as follows: 48.70 ± 3.79%, 50.56 ± 2.92%, 61.59 ± 3.42% and 75.91 ± 2.07%, respectively. The enhancement of cell migration observed in the MN and MN@Ple groups was relatively modest. Conversely, a marked improvement in cell migration status was attained through combined treatment with MN@Ple/Exo Q10 patch and photothermal therapy (Fig. 3 F and I). For the Transwell assay, the MN@Ple/Exo Q10 +NIR group exhibited the maximum number of cells that migrated across the Transwell chamber. Additionally, the quantity of migrating cells in the MN@Ple/Exo Q10 group without NIR irradiation was greater than that observed in other HG-treated groups and was even slightly higher than that in the negative control group (Fig. 3 G and J). Tube formation experiments based on HUVECs revealed that MN patches supplemented with Exos Q10 effectively mitigated the impairment of capillary-like tube formation induced by HG conditions (Fig. 3 H). This intervention resulted in enhanced tube quality, longer total tube lengths and a greater number of Nb nodes. Furthermore, the NIR group further augmented the promotion of angiogenesis through MN@Ple/Exo Q10 (Fig. 3 K and L). In conclusion, both blank MN and MN@Ple without Exo Q10 showed a relatively weak effect on cell migration and angiogenesis. In contrast, the release of Exos Q10 from MNs significantly enhanced cell migration and promoted microtubule formation. This finding corroborates previous research indicating that HucMSC-derived Exos accelerated angiogenesis and facilitated wound healing in vitro [48] . Concurrently, the elevated temperature of the MN@Ple/Exo Q10 patch, induced by periodic NIR irradiation, exerted a substantial influence on HUVECs and L929 cells, thereby promoting cellular migration and angiogenesis. Moreover, increased molecular thermal motion, induced by elevated temperatures, results in greater Exos Q10 release [49]−[50] . 2.6. MN treatment alleviated HG-induced ferroptosis in endothelial cells As illustrated in Fig. 4 A, the pathological mechanism by which a HG environment induces oxidative stress in cells and subsequently triggers ferroptosis through the accumulation of lipid peroxidation serves as the cellular basis for diabetic wounds [51] . Specifically, HG induces lipid damage in the inner mitochondrial membrane, leading to an increased release of ROS. This event subsequently triggers the transcriptional downregulation of SLC7A11, which functions as a cystine antiporter and directly contributes to a substantial decrease in the GSH/GSSG ratio. Additionally, the inhibition of glutathione peroxidase 4 (GPX4) activity leads to impaired clearance of lipid peroxidation [52] . Alternatively, excessive glucose upregulates long-chain acyl-CoA synthase 4 (ACSL4) and lysophosphatidylcholine acyltransferase 3 (LPCAT3), resulting in the accumulation of polyunsaturated fatty acids (PUFAs), thereby aggregating phospholipid hydroperoxide (PLOOH), which can impair membrane structures and facilitate the release of ROS [53]−[54] . To further elucidate the mechanism by which MN@Ple/Exo Q10 , in conjunction with NIR, mitigates HG-induced ferroptosis in vitro, an investigation was conducted encompassing four key aspects: cellular ROS levels, accumulation of lipid peroxidation and Fe 2+ , mitochondrial morphology and structure, as well as the expression of ferroptosis-related genes. As demonstrated in Fig. 4 D, the Dichlorofluorescein-Diacetate (DCFH-DA) fluorescent probe was employed as an indicator for ROS to assess intracellular ROS levels. Fluorescence imaging and quantitative analysis across various treatment groups revealed that the green fluorescence of HUVECs subjected to HG conditions was significantly elevated compared to that of the negative control group. Moreover, both MN and MN@Ple treatments did not exhibit a significant reduction in fluorescence intensity relative to the control group. Nevertheless, the MN@Ple/Exo Q10 group demonstrated a green fluorescence intensity that was only 0.33 times that of the control group, while only a minimal number of cells exhibited this signal in the NIR group (Fig. 4 G). JC-1, serving as a probe for assessing mitochondrial membrane potential, forms red aggregates within normal mitochondria displaying high membrane potential. Conversely, when mitochondria are compromised and exhibit low membrane potential, JC-1 exists as green monomers and fails to aggregate in the mitochondrial matrix [55] . After labeling with the JC-1 fluorescent probe, HUVECs subjected to different treatments were quantitatively analyzed for JC-1 fluorescence images, and the red/green ratio was subsequently calculated. The results indicated a significant increase in JC-1 monomers alongside a notable decrease in JC-1 aggregates within the HG-treated control group, whereas the negative control group exhibited opposite trends. Furthermore, the red/green ratio observed in both the MN and MN@Ple groups closely mirrored that of the control group. In the MN@Ple/Exo Q10 and NIR groups, there was a substantial increase in JC-1 aggregates emitting red fluorescence compared to other HG-treated groups, while levels of JC-1 monomers emitting green fluorescence were significantly diminished (Fig. 4 E and G). GSH and MDA serve as crucial biomarkers for assessing LOOH in endothelial cells undergoing ferroptosis [56] . HUVECs from various treatment groups were lysed and collected for the determination of GSH and MDA levels. The results suggested that the production of MDA in the HG-treated control group was significantly higher compared to the negative control group, while GSH levels were markedly lower. In contrast, the MDA level in the MN@Ple/Exo Q10 group was approximately 0.29 times that of the HG-treated control group, while GSH levels increased by about 2.8 times. This finding showed that Exo Q10 significantly reduced lipid peroxidation levels in cells, importantly, NIR treatment did not affect the efficacy of Exo Q10 (Fig. 4 B and C). The FerroOrange fluorescent probe was utilized to evaluate free Fe 2+ accumulation in the cytoplasm of HUVECs (Fig. 4 F). Cells within the HG control group exhibited pronounced orange-red fluorescence signals, indicating a significant aggregation of Fe 2+ . However, both the MN@Ple/Exo Q10 and NIR groups demonstrated reduced fluorescence signals (Fig. 4 G). As displayed in Fig. 4 J, scanning electron microscopy (SEM) was employed to visualize the morphological alterations of mitochondria in HUVECs subjected to various treatment groups. In comparison to the negative control group, mitochondria in the HG control group showed atrophy, increased membrane density, and reduced or absent mitochondrial cristae (indicated by red arrows). Furthermore, mitochondrial damage was also observed in the MN and MN@Ple groups. Remarkably, structural damage to mitochondria was significantly reduced in both the MN@Ple/Exo Q10 and NIR groups, showing no significant difference from the negative control. Above results indicated that the MN@Ple/Exo Q10 patch effectively alleviated HG-induced mitochondrial damage by releasing Exo Q10 alongside NIR therapy. The expression levels of ferroptosis-related markers in HUVECs were detected through Western blot analysis. The result revealed that HG significantly reduced the expression levels of GPX4 and SLC7A11 in HUVECs, while increasing ACSL4 levels. However, there were no significant differences in the expression levels between the HG control, MN and MN@Ple groups. Notably, GPX4 expression was upregulated to approximately 0.78 and 0.72 in cells co-cultured with MN@Ple/Exo Q10 and MN@Ple/Exo Q10 +NIR, respectively. Meanwhile, SLC7A11 levels increased to 0.67 and 0.77 in these groups and ACSL4 expression was downregulated to 0.40 and 0.359, respectively (Fig. 4 H and I). For the RT-qPCR (Fig. 4 K), MN@Ple/Exo Q10 significantly improved the mRNA expression levels of GPX4, SLC7A11, ACSL4 and 4 LPCAT3. Specifically, the expressions of GPX4 and SLC7A11 in the HG control group were approximately 0.15 and 0.37 times those in the negative control group, while ACSL4 and LPCAT3 showed expression levels of about 3.2 and 2.1 times, respectively. Conversely, the expression levels in the MN@Ple/Exo Q10 group were found to be 0.90 for GPX4, 0.78 for SLC7A11, as well as 1.7 and 1.3 for ACSL4 and LPCAT3, respectively. Alternatively, the NIR group did not impair the ability of MN@Ple/Exo Q10 to regulate ferroptosis-related genes; rather, a modest enhancement was observed (Fig. 4 L-O). Additionally, although HucMSC-derived Exos regulated diabetic wound healing across various dimensions in previous studies, the pathological microenvironment of intracellular oxidative stress–lipid peroxidation burst–ferroptosis could not be improved [57]−[58] . In this study, the MN@Ple/Exo Q10 patch served as a carrier to release Exos Q10 to modulate the SLC7A11-GSH-GPX4 antioxidant pathway and the PUFA-ACSL4-PLOOH axis, both closely linked to ferroptosis, while this regulatory process was unaffected by intermittent temperature increases. In summary, MN@Ple/Exo Q10 exhibited the capability to alleviate cellular ferroptosis in vitro under NIR irradiation. 2.7. In vitro anti-inflammatory capacity of MN patches Immune regulation disorder is a critical factor in the management of DW. The elevated oxidative stress caused by HG results in mitochondrial damage, which further inhibits the metabolic transition from glycolysis to oxidative phosphorylation in macrophages, disrupting the normal immune equilibrium [59] . At the cellular level, classically activated M1 macrophages are unable to transition smoothly into alternatively activated M2 macrophages [60] . The impaired transition impedes the shift in the DW microenvironment from a persistent inflammatory response to tissue repair, ultimately resulting in an inflammatory cytokine storm and increased cell death leading to ROS accumulation, thus establishing a detrimental cycle of inflammation-oxidative stress [61] . In this study, Raw264.7 cells were employed as an in vitro model to investigate the regulatory effects of MN@Ple/Exo Q10 on macrophage polarization, through the application of immunofluorescence, Western blot, qRT-PCR, and flow cytometry. Raw264.7 cells were polarized into M1 macrophages by lipopolysaccharide (LPS), displaying irregular pseudopodia and a strong ability to produce inflammatory mediators. In contrast, M2 macrophages, induced by an appropriate concentration of IL-4 stimulation, were characterized by slender spindle morphology and the secretion of anti-inflammatory cytokines (Fig. 5 A and B). In immunofluorescence staining, Raw264.7 cells pre-stimulated with LPS (20 ng/mL) were respectively labeled as markers of M1 and M2 macrophages: CD86 and CD206, while the immunofluorescence images and their corresponding heat maps are shown in Fig. 5 C. Compared with the negative control group, the LPS control group exhibited significantly enhanced red fluorescence representing CD86 and nearly undetectable green fluorescence representing CD206, indicating successful polarization of M1 macrophages. Cells co-cultured with MN group revealed high red fluorescence comparable to that of the LPS group, and the MN@Ple group displayed a slight reduction in red fluorescence, whereas both the MN and MN@Ple groups showed low levels of green fluorescence. By contrast, the MN@Ple/Exo Q10 and NIR groups exhibited markedly reduced red fluorescence and significantly enhanced green fluorescence, suggesting a considerable enhancement in M2 activation. Flow cytometry further confirmed the capacity of MN@Ple/Exo Q10 to promote the reprogramming of macrophages to the M2 phenotype. The percentages of CD206⁻CD86⁺ (M1) cells in the MN@Ple/Exo Q10 and MN@Ple/Exo Q10 +NIR groups were 37.3% and 30.8%, respectively, significantly lower than the 67.2% observed in the LPS control group. Conversely, the percentages of CD206 + CD86 − (M2) in these two groups were approximately 18.1- and 23.7-fold higher than those in the LPS group. These results demonstrate that MN@Ple/Exo Q10 effectively facilitates the transition of macrophages from the pro-inflammatory M1 phenotype to the anti-inflammatory M2 phenotype (Fig. 5 D). Western blot and qRT-PCR were employed to quantitatively assess the differential expression of M1 and M2 macrophage-related markers at the gene level among various experimental groups. For the Western Blot, LPS pretreatment elevated the protein expression of inducible nitric oxide synthase (iNOS), while the protein level of CD206 remained low. Neither the MN group nor the MN@Ple group displayed a significant reduction in iNOS expression or an increase in CD206 expression. However, the MN@Ple/Exo Q10 and NIR groups exhibited a marked decrease in iNOS protein levels and a significant upregulation of CD206 expression (Fig. 5 E and F). The expression levels of pro-inflammatory cytokines (TNF-α and IL-6) associated with M1 macrophages, and anti-inflammatory factors (Arg-1 and IL-4) linked to M2 macrophages across different experimental groups were analyzed by qRT-PCR (Fig. 5 G). The relative mRNA expression levels of pro-inflammatory cytokines TNF-α and IL-6 were markedly higher in the LPS-treated control group. Following MN treatment among various experimental groups, these cytokines demonstrated differing levels of downregulation, with the most pronounced reduction observed in the NIR group. Instead, the mRNA levels of anti-inflammatory markers, including Arg-1 and IL-4, were significantly upregulated in the MN@Ple/ Exo Q10 +NIR group compared to the other groups (Fig. 5 H-K). In conclusion, MN@Ple/Exo Q10 demonstrated anti-inflammatory potential in vitro. Specifically, it inhibited ferroptosis by delivering Exo Q10 , accelerating the clearance of intracellular lipid peroxidation and reducing its accumulation. Consequently, the intervention effectively disrupted the oxidative stress-ferroptosis-inflammation cycle in Raw264.7 cells, thus facilitating M2 macrophage reprogramming, enhancing the secretion of anti-inflammatory cytokines, and expediting the repair and tissue reconstruction phase [62]−[63] . Meanwhile, Ple exhibited a notable anti-inflammatory capacity in the present study, consistent with findings from previous investigations [64] . Furthermore, safe and periodic NIR irradiation could enhance the immunomodulatory function of MN patches [65] . 2.8. Transcriptomic analyses revealed the molecular mechanisms underlying the antioxidant and anti-inflammatory effects mediated by MN@Ple/Exo Q10 To investigate the molecular mechanisms by which MN@Ple/Exo Q10 regulates macrophage polarization toward the M2 phenotype, this study conducted whole-transcriptome sequencing analysis on Raw264.7 cells treated with MN@Ple/Exo Q10 . The results revealed that volcano plot and heatmap analyses identified a total of 1,459 differentially expressed genes (DEGs), among which 893 genes were significantly downregulated and 566 genes were significantly upregulated (Fig. 6 A and B). Functional enrichment analysis showed that in Gene Ontology (GO) analysis, these DEGs were primarily enriched in biological processes such as regulation of inflammatory response, oxidoreductase activity, and protein kinase binding (Fig. 6 D), suggesting a close association between these processes and the therapeutic effects of MN@Ple/Exo Q10 . Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis further demonstrated that MN@Ple/Exo Q10 significantly activated inflammatory regulatory pathways such as the NF-κB signaling pathway, TNF signaling pathway, and C-type lectin receptor signaling pathway, while also modulating oxidative stress-related pathways including the p53 signaling pathway, apoptosis, and AMPK signaling pathway (Fig. 6 E). ROS exhibit cytotoxicity by inducing oxidative stress through reactions with proteins, lipids, and nucleic acids. Consequently, precise cellular responses to ROS production are critical for preventing further oxidative damage and maintaining cell viability. In this study, Gene Set Enrichment Analysis (GSEA) confirmed that MN@Ple/Exo Q10 treatment significantly activated NF-κB, MAPK, and PI3K/AKT signaling pathways (Fig. 6 F, G, and H). Notably, enhanced NF-κB activity induced the expression of antioxidant proteins such as NAD(P)H quinone oxidoreductase 1 (NQO1), heme oxygenase-1 (HO-1), and glutathione peroxidase 1 (GPX1), thereby counteracting oxidative stress injury. Concurrently, activation of the PI3K/AKT pathway not only suppressed oxidative stress-induced apoptosis but also promoted M2 macrophage polarization to exert anti-inflammatory effects. Ferroptosis is closely linked to the progression of numerous diseases and involves multiple disease-associated signaling pathways. The MAPK pathway inhibits ferroptosis through phosphorylation and activation of nuclear factor erythroid 2-related factor 2 (Nrf2). As a cellular energy sensor, AMPK activation modulates redox homeostasis and iron metabolism to inhibit ferroptosis. Additionally, GSEA results of the ferroptosis suppressor gene set indicated that MN@Ple/Exo Q10 treatment upregulated the expression of ferroptosis-inhibiting genes, including GPX4 and Stat3 (Fig. 6 I). Further CIBERSORT immune infiltration analysis demonstrated that MN@Ple/Exo Q10 significantly enhanced macrophage polarization toward the M2 phenotype (Fig. 6 C, Fig. S6). Collectively, these results indicate that MN@Ple/Exo Q10 remodels the diabetic wound immune microenvironment by synergistically attenuating inflammatory responses, enhancing antioxidant capacity, and suppressing ferroptosis. 2.9. MN patches promoted infected diabetic wound healing in vivo This study utilized a full-thickness infected diabetic wound (IDW) model in BALB/c mice to evaluate the in vivo efficacy of the MN@Ple/Exo Q10 patch in promoting wound healing. As shown in Fig. 7 A, BALB/c mice were induced to develop type I diabetes via streptozotocin (STZ) administration, and subsequently, a circular full-thickness skin wound measuring 1 cm in diameter was created on the dorsum of each mouse. Thereafter, a suspension of S. aureus was injected into the diabetic wounds to establish IDW. Wound images of diabetic mice were recorded macroscopically on days 0, 3, 7 and 14 following treatment with different biomaterial groups, and the residual wound areas were then calculated (Fig. 7 B). On day 3, the relative residual wound area (RRWA) did not show significant differences among the control, MN, MN@Ple, and MN@Ple/Exo Q10 groups, whereas the MN@Ple/Exo Q10 +NIR group exhibited a slightly reduced RRWA (53.49 ± 3.62%). Following 7 days of treatment, the RRWA of the control group was approximately 51.88 ± 3.80%, while that of the MN and MN@Ple groups was 45.58 ± 3.98% and 35.23 ± 1.98%, respectively, showing a modest decrease compared to the control group. However, the relative residual wound area of the MN@Ple/Exo Q10 group (22.85 ± 3.36%) and the NIR group (12.53 ± 1.56%) decreased significantly, with the NIR group showing the most notable reduction. On day 14, the wound area in the NIR group was nearly completely healed (4.34 ± 1.34%), while the MN@Ple/Exo Q10 group also exhibited a significantly lower residual wound area (7.01 ± 1.83%), which was notably smaller than that observed in the MN group (16.39 ± 2.68%) and the MN@Ple group (13.97 ± 1.22%). In contrast, a substantial proportion of wound areas in the control group remained unhealed (26.20 ± 2.41%) (Fig. 7 C). It is noteworthy that the MN and MN@Ple groups exhibited limited improvement in wound healing within the first 7 days, whereas the MN@Ple/Exo Q10 group began to demonstrate a discernible therapeutic effect on wound healing starting from day 3. Furthermore, the NIR group consistently displayed a significantly higher wound healing rate compared to all other groups at every observed time point. In addition, bacteria were isolated from the collected wound tissues treated with various groups and cultured on agar plates to further quantify the in vivo anti-infective capacity. As illustrated in Fig. S8, the control and MN groups exhibited a high bacterial load in the wound area, whereas the bacterial counts were markedly reduced in MN@Ple and MN@Ple/Exo Q10 groups. Notably, the MN@Ple/Exo Q10 +NIR group, which demonstrated superior antibacterial efficacy, displayed a significantly lower colony count compared to the other treatment groups. In summary, during the early stage of IDW treatment, the shell layer of the MN patch in the putamen structure was fully degraded and completely released the loaded Ple, thereby exerting an antibacterial effect. As the core layer became exposed, it gradually released the loaded Exos Q10 . Simultaneously, PTT applied via NIR irradiation synergized with the sustained release of Exo Q10 to further accelerate wound healing, enabling sequential and effective treatment of IDW. 2.10. MN patch for transdermal and sustained delivery of Exos Q10 in vivo The capacity of the MN patch to achieve transdermal and sustained release within the wound bed is crucial for nano-scale drug delivery platforms [66] . Exos Q10 were pre-labeled with Dil dye, and the wound treated with the MN patch was subsequently compared to two distinct local Exos delivery approaches, followed by imaging through LSCM and in vivo imaging system (IVIS) (Fig. 7 D). As illustrated in Fig. 7 E, both the topical administration (TA) group and the MN group exhibited intense red fluorescence localized in the epidermis on the day of treatment. By contrast, the MN group demonstrated the capability to penetrate into the dermal layer, forming a wedge-shaped opening and producing a significantly enhanced fluorescence signal in that region. With the progression of the treatment period, by day 7, the skin tissue in the MN group still exhibited noticeable red fluorescence from Dil-Exo Q10 , which gradually diminished by day 14. Conversely, the TA group showed only faint residual red fluorescence on the skin surface. Particularly, the enhanced transdermal delivery capability of the MN patch ensured a sustained presence of Exos Q10 within the dermal tissue throughout all stages of wound healing, a significant advantage over conventional topical treatments. To assess the efficacy of the MN patch in achieving sustained delivery of Exos Q10 to the wound bed, circular full-thickness skin defect mice were treated with intradermal injection (ID) of Exos Q10 and the MN patch for 14 days. Then, the distribution and retention of Exos Q10 were monitored and quantified via IVIS on days 3, 7, 10, and 14 (Fig. 7 F and G). The results demonstrated that the red fluorescence emitted from the wound defects on the backs of mice in the MN group gradually decreased as the wound healed, yet remained detectable by day 14. However, the red fluorescence in the ID group began to diminish from day 7 and was no longer observable on days 10 and 14. Overall, the MN treatment enabled the sustained release of Exos Q10 in vivo for at least 14 days, in contrast to the ID treatment, while the MN system developed in this study functions as an effective Exo delivery platform, capable of achieving complete transdermal penetration and prolonged, stable release of Exos Q10 , thereby ensuring its continuous therapeutic efficacy in diabetic wound healing. 2.11. Histological evaluation of MN treatment in infected diabetic wound healing Hematoxylin-eosin (H&E) and Masson staining were conducted to assess wound diameter, granulation tissue formation, epithelialization, and collagen deposition in IDW. According to the H&E staining (Fig. 8 A), quantitative analysis of wound length on day 7 demonstrated that the MN@Ple/Exo Q10 +NIR group exhibited the shortest wound length (3500.00 µm), followed by the MN@Ple/Exo Q10 group (5384.62 µm). The MN and MN@Ple groups showed wound lengths of approximately 7307.69 µm and 6769.23 µm, respectively, representing a significant reduction compared to the control group (7692.31 µm). Additionally, the wound lengths of the control, MN, MN@Ple, MN@Ple/Exo Q10 and NIR groups after 14 days were 6076.92 µm, 4653.85 µm, 4230.77 µm, 3384.62 µm and 2423.08 µm, respectively. Significantly, except for the control group, the wound lengths of all groups were shorter than those at the previous time point, with the NIR group exhibiting the shortest length. This result corresponded to the wound photographs of IDW across different treatment groups observed previously (Fig. 8 C). The thickness of granulation tissue and the degree of epithelial reconstruction are key indicators for assessing wound healing. Histological analysis via HE staining on day 7 revealed that both the MN@Ple/Exo Q10 group and the NIR group displayed thicker granulation tissue and ongoing epithelial regeneration, with the NIR group demonstrating the most pronounced effects. However, the remaining three groups showed minimal epithelial formation and significantly reduced granulation tissue thickness, particularly in the control group. In contrast, the control group still lacked newly formed epithelium on day 14, and no significant changes in granulation tissue were observed. Nevertheless, the MN@Ple/Exo Q10 and NIR groups revealed a marked reduction in granulation tissue thickness compared to day 7, with the NIR group displaying the thinnest granulation tissue along with a fully matured thin epithelial layer. Furthermore, the MN and MN@Ple groups showed relatively thicker granulation tissue and incompletely formed epithelium that appeared immature and uneven (Fig. 8 D). This accelerated healing process might be attributed to Exo Q10 facilitating the transition from the inflammatory to the proliferative phase of wound repair. Additionally, PTT promoted cell migration and the release of growth factors, which, in combination with Exo Q10 , synergistically enhanced the healing of IDW [67] . Moreover, H&E staining of the heart, liver, spleen, lung and kidney of mice in different groups presented that MN patches induced no evident organ toxicity (Fig. S9). As illustrated in Fig. 8 B, Masson staining revealed the collagen deposition patterns in IDW across different treatment groups. On day 7, the control group exhibited minimal blue-stained collagen fibers, whereas the MN and MN@Ple groups displayed only a limited amount of loosely arranged collagen fibers. Conversely, the MN@Ple/Exo Q10 +NIR group demonstrated markedly increased collagen deposition compared to the other groups. Particularly, the NIR group showed a larger area of collagen deposition characterized by a denser and more compact arrangement on day 14, indicating a transition from fragile, disorganized type III collagen to robust and well-organized type I collagen fibers [68] . However, in the control, MN and MN@Ple groups, the collagen deposition was still irregular and disordered on day 14, and the relative collagen deposition was less compared with that of the MN@Ple/Exo Q10 group (Fig. 8 E). Considering the healing mechanism of MN@Ple/Exo Q10 , the diffusion of Ple from the MN shell layer created a favorable microenvironment for IDW healing through its antibacterial effects. Nonetheless, the wound healing capacity of the MN@Ple group remained limited, as evidenced by wounds that persisted in the proliferative phase on day 14, characterized by the presence of thick granulation tissue, incomplete epithelialization, and disorganized type III collagen. Moreover, the HA fragments composed of 6–20 disaccharides within the MN core layer promoted the migration and proliferation of dermal fibroblasts, then inducing the deposition of type III collagen and contributing to ECM formation [69] . Notably, Exos Q10 released from MN@Ple/Exo Q10 suppressed oxidative stress in IDW and modulated immune responses, thereby accelerating the transition from the inflammatory phase to the proliferative phase, and subsequently facilitating a faster progression into the remodeling phase, which was characterized by the gradual maturation and thinning of granulation tissue, the development of a mature, thin epithelial layer, and a more organized and dense deposition of collagen. In addition, NIR further synergized with this process to facilitate wound healing. 2.12. In vivo evaluation of antioxidant and anti-Inflammatory for MN patches The in vitro antioxidant and immunomodulatory effects of MN@Ple/Exo Q10 were confirmed through prior cellular experiments. Diabetic mice infected with S. aureus were treated with different groups of MN patch for seven days. Then, wound tissues were collected and analyzed by utilizing immunofluorescence staining, immunohistochemical staining, and enzyme-linked immunosorbent assay (ELISA) to assess the in vivo antioxidant and anti-inflammatory capacities of MNs. As shown in Fig. 9 A, the ROS levels in IDW were examined using the dihydroethidium (DHE) fluorescence probe, which is oxidized by ROS present in the wound bed, producing a red fluorescence. The intense red fluorescence observed in the wound bed of the control group indicated a pronounced oxidative stress state in the untreated IDW. The MN and MN@Ple groups exhibited similarly strong red fluorescence, with no statistically significant reduction compared to the control group. However, the MN@Ple/Exo Q10 group showed a marked decrease in red fluorescence intensity. Additionally, quantitative analysis revealed that the relative fluorescence intensity in the MN@Ple/Exo Q10 group was only 0.13 times that of the control group, while the MN@Ple/Exo Q10 +NIR group exhibited a relative fluorescence intensity of nearly 0.09 times that of the control group, with almost no detectable red ROS signal (Fig. 9 G). GPX4, a selenium-dependent antioxidant enzyme responsible for reducing peroxides [70] , was evaluated for its expression level in the wound tissue through immunohistochemical staining (Fig. 9 D and K). The results demonstrated that GPX4 expression levels in the MN@Ple/Exo Q10 and NIR groups were considerably elevated in comparison with those in the other groups, and these findings were in agreement with the ELISA data. Moreover, ELISA analysis revealed low expression levels of 4-HNE in the control, MN, and MN@Ple groups, whereas significantly reduced levels were observed in the MN@Ple/Exo Q10 group and NIR group (Fig. 9 N). Essentially, MN@Ple/Exo Q10 displayed robust antioxidant capacity in diabetic mice, and the sustained-release Exos Q10 from the MN core layer effectively mitigated ferroptosis-associated lipid peroxide accumulation in the wound, thereby alleviating oxidative stress induced by excessive ROS. IDW tissues were harvested after seven days of treatment across different groups to investigate the immunomodulatory efficacy of MN treatment during the inflammatory phase. As illustrated in Fig. 9 B, CD86/CD206 double immunofluorescence staining on day 7 was employed to assess the expression levels of M1 and M2 macrophages. The results demonstrated that the control group exhibited intense red fluorescence associated with CD86-positive cells and minimal green fluorescence associated with CD206-positive cells, suggesting a predominance of M1 macrophages in untreated IDWs. In contrast, the MN and MN@Ple groups showed a moderate reduction in relative fluorescence intensity for CD86 compared to the control group, while no significant change was observed in the relative fluorescence intensity for CD206, indicating that although the proportion of M1 macrophages responsible for inflammatory responses decreased, the wound remained in the inflammatory phase and had not transitioned to the repair-oriented proliferative phase. Fluorescence quantitative analysis revealed that the MN@Ple/Exo Q10 group demonstrated a reduction in relative fluorescence intensity (CD86) to 0.21-fold of the control group through sustained release of Exos Q10 , accompanied by a 3.09-fold increase in relative fluorescence intensity (CD206). Furthermore, the NIR group displayed significantly higher green fluorescence intensity compared to other groups, with a relative fluorescence intensity (CD206) reaching 4.5-fold that of the control group (Fig. 9 H and I). The inhibition of inflammatory response in IDWs by MN@Ple/Exo Q10 was further verified by immunohistochemical staining of myeloperoxidase (MPO), which is a critical biomarker that reflects both the intensity of inflammatory responses and the extent of immune cell activation. Meanwhile, the persistent hyperglycemic environment in diabetic wounds promotes neutrophil activation, resulting in their excessive accumulation at the wound site [71] . As demonstrated in Fig. 9 E and L, the relative expression level of MPO in the control group was evidently elevated compared to the other experimental groups. Following treatment with MN and MN@Ple, MPO expression in IDWs was reduced, yet remained higher than that observed in the MN@Ple/Exo Q10 group. Notably, MPO expression was nearly undetectable in the NIR group. Furthermore, ELISA analysis of wound tissue on day 7 indicated that the expression of the M1 cytokine (IL-6) in the MN and MN@Ple groups was reduced compared to the control group, whereas the expression of the M2 cytokine (IL-10) remained at low levels across the control, MN, and MN@Ple groups. Significantly, in both the MN@Ple/Exo Q10 and NIR groups, IL-6 levels were markedly suppressed, while IL-10 expression was significantly upregulated (Fig. 9 N). In summary, following application of the MN@Ple/Exo Q10 patch to IDWs, the linear anionic acidic polysaccharide HA in the core layer regulated inflammatory cell activity through binding to cell surface receptors, thereby partially controlling the wound inflammatory response [72] . However, the efficacy of HA was constrained by pathological conditions such as persistent oxidative stress and HG in diabetic wounds. Moreover, the release of Exos Q10 in combination with mild photothermal therapy effectively inhibited cellular ferroptosis, reduced intracellular lipid peroxide levels, and alleviated oxidative stress in diabetic wounds, thereby modulating the wound immune response. 2.13. MN treatment accelerated angiogenesis in vivo Adequate angiogenesis is essential for IDW healing. Following the inflammatory phase of wound healing, newly formed blood vessels progressively infiltrate the wound during the proliferative phase, delivering oxygen and nutrients to the wound site while removing ROS and metabolic waste products, thereby enhancing the supply of critical components required for tissue regeneration and accelerating wound repair [73] . The expression levels of CD31 and α-SMA were analyzed using double immunofluorescence staining to investigate the distribution patterns of vascular endothelial cells and pericytes in skin wounds among different treatment groups after 14 days of intervention (Fig. 9 C). Specifically, only minimal red and green fluorescence corresponding to CD31 and α-SMA was observed in the control group, and the MN group exhibited a modest increase in CD31 and α-SMA fluorescence intensity compared to the control group, whereas the MN@Ple group demonstrated a more pronounced enhancement. This improvement might be attributed to the role of HA in the core MN layer in regulating fibroblast and vascular endothelial cell differentiation, thus promoting angiogenesis and new tissue formation [74] . Additionally, Ple-mediated suppression of S. aureus in IDWs reduced bacterial oxygen consumption, alleviating the local hypoxic environment, which increased the oxygen partial pressure and directly stimulated endothelial cell proliferation, thereby accelerating neovascularization [75] . Notably, significant angiogenesis was observed in the MN@Ple/Exo Q10 group, while the NIR group displayed the highest proportion of CD31-positive endothelial cells and α-SMA-positive pericytes (Fig. 9 J). The immunohistochemical staining results illustrated that the expression level of the vascular marker VEGF in the control group was significantly lower than that in the other groups, with the NIR group exhibiting the highest relative VEGF expression. As a ligand, VEGF binds to and activates VEGFR-2, thereby triggering downstream signaling pathways involved in the angiogenic signal transduction cascade [76] , ultimately promoting wound angiogenesis (Fig. 9 F). In addition, the ELISA results further confirmed that the VEGF expression level in the NIR group was the highest, reaching approximately 384.9 pg/mgprot. In comparison, the MN@Ple/Exo Q10 group exhibited the second-highest VEGF expression, with a value of approximately 288.13 pg/mgprot (Fig. 9 N). The above results indicated that Exos Q10 , derived from HucMSCs, delivered various bioactive molecules such as ANG-2 and Wnt4, which promoted both angiogenesis and vascular maturation [77] . Furthermore, when combined with photothermal therapy, the system induced local vasodilation, enhanced blood flow velocity and oxygen concentration, thereby delivering increased nutrients, immune cells, and oxygen to the non-healing wound. 3. Materials and methods 3.1. Materials Gelatin methacryloyl (GelMA, EFL-GM-60,150kDa), Methacryloyl hyaluronic acid (HAMA, EFL-HAMA-400, 400kDa) and Photoinitiator (lithium phenyl-2,4,6-trimethylbenzoylphosphinate, LAP) were bought from Suzhou Intelligent Manufacturing Research Institute (Suzhou, China). Pleurocidin (HY-P5641), Trehalose (T9531), Coenzyme Q10 (HY-N0111) and Acetone (179124) were bought from MedChemExpress (USA). Dopamine hydrochloride was purchased from Macklin Biochemical Technology (Shanghai, China). Tris hydrochloride solution (PM12630) and glutaraldehyde solution (PM20835) were acquired from PERFEMIKER (Shanghai, China).Polydimethylsiloxane (PDMS) was purchased from Dow Corning (Midland, USA). Anhydrous ethanol, Rhodamine B, 5(6)-Carboxyfluorescein were acquired from Aladdin Bio-Chem Technology Co., Ltd. (Shanghai, China). Modified Eagle Medium (DMEM), Fetal bovine serum (FBS), Penicillin-Streptomycin and Phosphate Buffered Saline (PBS) were bought from Gibco (USA). LB nutrient agar, LB broth, Crystal Violet Staining solution, 4% paraformaldehyde Fix Solution, lipopolysaccharide (LPS), DAPI Staining Solution and Triton X-100 were bought from Beyotime Biotechnology (Shanghai, China). H&E kit, and Masson kit were purchased from Solarbio (Beijing, China). 3.2. Antibodies CD63 (1:500, goat polyclonal antibody, SC-15363, Santa Cruz); TSG101 (1:8000, rabbit polyclonal antibody, 28283-1-AP, Proteintech); Calnexin (1:1000, rabbit polyclonal antibody, ab22595, Abcam); GPX4 (1:1000, rabbit polyclonal antibody, 14432-1-AP, Proteintech); SLC7A11 (1:2000, rabbit polyclonal antibody, AF7992, Beyotime); ACSL4 (1:5000, rabbit polyclonal antibody, 22401-1-AP, Proteintech); iNOS (1:2000, rabbit polyclonal antibody, ab3523, Abcam); CD206 (1:1000, rabbit polyclonal antibody, 18704-1-AP, Proteintech); CD86 (1:500, rabbit polyclonal antibody, 13395-1-AP, Proteintech); CD31 (1:1000, rabbit polyclonal antibody, 11265-1-AP, Proteintech); α-SMA (1:1000, rabbit polyclonal antibody, 14395-1-AP, Proteintech); MPO (1:100, rabbit polyclonal antibody, AF7494, Beyotime); VEGF (1:500, rabbit monoclonal antibody, YA013, MCE); GAPDH (1:5000, rabbit polyclonal antibody, 10494-1-AP, Proteintech); β-actin (1:10000, mouse monoclonal antibody, 60008-1-Ig, Proteintech); Goat Anti-Rabbit IgG H&L (HRP) (1:2000, ab7090, Abcam); Alexa Fluor® 488 Goat Anti-Rabbit IgG H&L (1:400, ab150077, Abcam); Alexa Fluor® 488 Goat Anti-Mouse IgG H&L (1:400, ab150113, Abcam). 3.3. Isolation, identification and internalization assay of Exo Q10 Before Exo extraction, 1 mM Coenzyme Q10 (CoQ10) was added to culture dishes containing HucMSCs with a confluence of 90%, followed by a 24-hour pretreatment. Next, the culture medium was replaced with Exo-depleted medium, and the cells were cultured for an additional 48 hours. Finally, the supernatant was collected by centrifugation at 2000 × g for 10 minutes. Then, the supernatant was centrifuged at 10,000 g for 30 minutes, followed by ultracentrifugation at 100,000 g for 70 minutes. After washing with PBS, ultracentrifugation was performed again at 100,000 g for another 70 minutes. The Exos were washed with PBS and then carefully resuspended in sterile PBS. Meanwhile, Exos derived from HucMSCs that were not co-cultured with CoQ10 were prepared utilizing the same method for identification experiments. The morphology and structure of Exos Q10 were detected by transmission electron microscopy (TEM, Thermo, USA), and the size of Exos Q10 was determined by Nanoparticle Tracking Analysis (NTA, PARTICLE METRIX GmbH, Germany). The expression of exosomal marker proteins (CD63, TSG101 and Calnexin) was detected by Western blot to confirm the successful separation of Exo Q10 . Exos Q10 were labeled by the Dil staining kit (Beyotime, C1991S) according to the manufacturer's instructions. L929 cells and HUVECs were respectively co-cultured with the culture medium containing Dil-labeled Exos Q10 for 6 hours. The cultured cells were fixed with 4% paraformaldehyde for 15 minutes, permeabilized with 0.1% Triton X-100 for 10 minutes, and then blocked with 5% BSA for 20–30 minutes. Following incubation with FITC at room temperature for 30 minutes, the cell nuclei were then stained with DAPI. The internalization of Exos Q10 was subsequently visualized using a laser scanning confocal microscope (LSCM, Leica, Germany). 3.4. Preparation method of MN patches First, DA hydrochloride (50 mg) was dissolved in 0.9% (wt/v) Tris solution (pH 8.5) to prepare a mixture, which was magnetically stirred for 24 hours to yield a black solution. Next, the mixture was washed three times with acetone and anhydrous ethanol to remove unreacted monomers, and the resulting Polydopamine nanoparticles (PDA NPs) were collected via centrifugation. Gelatin methacryloyl (GelMA) was mixed with a 0.25% (w/v) LAP initiator solution to achieve a final 15% (w/v) GelMA solution, and 0.02 wt% PDA NPs were incorporated as the pre-working solution for constructing the shell layer. The resulting mixture was poured into the molding holes of a PDMS negative mold, and bubbles were removed under negative pressure via a freeze dryer for 2 minutes. Thereafter, the PDMS positive mold was placed on top, and the composite was cross-linked under 405 nm UV light. After demolding, the MN shell layer was successfully fabricated. Then, Ple was dissolved in ddH₂O to prepare a 20 µM aqueous antimicrobial peptide solution, and 5% trehalose was added to preserve protein activity. Then, the preformed shell MN was rapidly immersed in liquid nitrogen and frozen to -196°C, and the Ple solution was precisely sprayed onto the tips of the MN in a vertical manner. Following rapid cooling in liquid nitrogen, the MN shell layer was transferred to a freeze dryer and subjected to two drying cycles to remove free water. Through repeated freeze-drying cycles, a sufficient quantity of Ple was effectively deposited at the tips of the MN. To prepare the Exo Q10 -loaded MN core layer, Exos Q10 (3 × 10¹⁰ particles mL-1) were mixed with 5% (w/v) trehalose, and the mixture was slowly frozen at -80°C at a cooling rate of 0.5°C/min over a period of 2 hours. The sample underwent two freeze-drying cycles to obtain an Exos lyophilized powder. This lyophilized powder was then added to a pre-polymer solution composed of 5% (w/v) Hyaluronic Acid Methacrylate (HAMA) and 0.25% (w/v) LAP, with the addition of 0.02 wt% PDA NPs. The mixture was reconstituted by incubation in a 37°C water bath with shaking for 10 minutes before being poured into the remaining PDMS cavities of the shell layer. Air bubbles were removed under negative pressure using a vacuum pump, and the core layer was cross-linked under 405 nm UV light. Finally, the core-shell MN patch was gently demolded and designated as MN@Ple/Exo Q10 , while it was subjected to secondary freeze-drying, sealed, and stored in the dark at 2–8°C until further use. In addition, blank MN, MN@Ple and MN@Ple/Exo Q10 patches with different loading compositions were prepared utilizing the aforementioned method for subsequent experiments, while a core-only blank MN patch (Core) was also fabricated. 3.5. Morphological characterization of MN patch The surface of the obtained sample was gold-coated prior to observation of the morphological characteristics of MN with a Scanning Electron Microscope (SEM, Zeiss, Germany). Elemental distribution across the MN array was analyzed via Energy-Dispersive X-Ray Spectroscopy (EDS, Zeiss, Germany). Moreover, the overall morphology of MN was examined using an optical microscope (BX53M, Olympus, Japan). To visualize the core-shell structure, the fluorescent dyes 5(6)-Carboxyfluorescein and Rhodamine B were incorporated into the shell and core solutions, respectively, during the MN fabrication process. The core-shell architecture was subsequently observed and imaged by LSCM. 3.6. Determination of mechanical properties of MN patches Compression tests were performed on MN patches from different experimental groups utilizing a universal testing machine (AllroundLine, ZwickRoell, Germany). The MN patches were positioned vertically on a rigid stainless steel platform with the needle tips oriented upward. The detector probe then compressed the needle tips at a constant rate of 5 mm/min in the vertical direction. The resulting stress-strain curves were recorded and analyzed to evaluate the mechanical properties of the MN patches. To evaluate the skin penetration capability of the MN@Ple/Exo Q10 , the MN patch was applied to the dorsal skin of Balb/C mice by thumb pressure for 3 minutes, followed by visual observation and photography. The tissue from the pressed area was then excised and examined under an optical microscope to assess the penetration depth of the needle tips into the mouse skin. To evaluate the tissue adhesion performance of the MNs, porcine skin was mounted on the baseplate of a stainless steel platform, and various MN samples were attached to a sensor probe with the needles oriented downward. The needle tips were inserted into the pig skin tissue to establish the initial contact position. Thereafter, a universal testing machine (AllroundLine, ZwickRoell, Germany) was employed to lift the probe vertically at a constant speed of 0.5 mm/min until complete separation of the MN patches from the tissue occurred. The resulting force-time curves were generated based on the recorded data. 3.7. In-vitro release assay of MN patches In vitro release determination of Ple: Serial dilutions of Ple solutions were prepared in 96-well plates, and their optical density values were measured utilizing a UV-Vis Spectrophotometer (Thermo, USA). A standard curve was subsequently generated utilizing GraphPad Prism software. For the release study, 0.5 mg of shell MN, fabricated via the previously described method, was immersed in 5 mL of PBS. At predetermined time points (1, 2, 4, 6, 8, 16, 24, 32, 40, and 48 hours), 1 mL of the release medium was withdrawn and replaced with 1 mL of fresh PBS. The collected samples were analyzed using the UV-Vis Spectrophotometer to measure the absorbance at 280 nm. The cumulative release amount of Ple was then calculated based on the standard curve. In vitro release determination of Exos Q10 : To evaluate the release of Exos Q10 from the core layer of MNs, 0.5 mg of previously prepared core layer MNs were immersed in 5 mL of PBS preheated to 37°C. At predetermined time points (1, 2, 4, 6, 8, 10, 12, 14, 16, and 18 days), 1 mL of the release medium was withdrawn and replaced with 1 mL of fresh PBS. The total protein concentration of Exos Q10 from different groups was quantified by the BCA protein assay kit (Thermo Scientific) in accordance with the manufacturer's instructions. The amount of released Exos Q10 was subsequently analyzed by multi-mode microplate reader (Thermo, USA), and the cumulative release percentage of Exos Q10 was calculated as follows: Cumulative release rate(%) = Rt / R0 × 100% (Rt: cumulative release amount at each time point; R0: Total release amount) 3.8. Photothermal property of MN patch The dried MN@Ple/Exo Q10 was placed on the sample platform and irradiated with an 808 nm near-infrared (NIR) laser at 0.5 W/cm² and 1 W/cm² for 5 minutes. Thermal images were captured every minute using an infrared thermal imager to monitor the temperature changes over time, and a temperature-time curve was plotted accordingly. To evaluate the thermal stability of the MN patch, five heating/cooling cycles were conducted by alternately switching the NIR laser on and off, during which temperature fluctuations were recorded. 3.9. The antibacterial performance of MN patches Staphylococcus aureus (S. aureus) and Escherichia coli (E. coli) liquid cultures were incubated in a shaking incubator at 37°C for 24 hours until the bacterial populations reached the logarithmic growth phase. MN patches were pre-sterilized via ultraviolet light for 2 hours and stored under sterile conditions prior to use. Subsequently, the bacterial suspensions were co-cultured with MN patches from different experimental groups for 24 hours. In the NIR group, samples were exposed to an 808 nm NIR laser for 10 minutes every 6 hours to induce photothermal effects. The colony count was performed according to the plate count method. Briefly, bacterial suspensions with varying concentrations obtained after co-culturing were gradient diluted 105 times, then uniformly spread onto agar plates, and incubated at 37°C in a constant temperature incubator for 24 hours. Colony growth of the two bacterial strains was recorded by photography, and the bacterial survival rate was calculated based on the colony counts. To assess the growth dynamics of bacteria across different experimental groups, bacterial cultures in the logarithmic growth phase were co-cultured with various MN patches for a duration of 9 hours. The optical density at 600nm (OD₆₀₀) was measured hourly for each group, and the resulting bacterial growth curves were generated using GraphPad Prism software. The morphological changes of S. aureus and E. coli were analyzed employing SEM (Zeiss, Germany). Treated bacterial samples were collected via centrifugation at 5000 rpm, fixed with 2.5% glutaraldehyde solution, and the supernatant was subsequently removed by centrifugation. The bacterial pellets were then washed three times with PBS, dehydrated through a graded ethanol series (25%, 50%, 75%, and 100%), and finally dropped onto silicon wafers for SEM observation. For the live/dead bacterial staining. Bacterial suspensions from different experimental groups were incubated in confocal dishes under dark conditions for 20 minutes using a SYTO9/PI live and Dead Bacteria Stain Kit (Thermo, L7007), following the manufacturer's protocol. Then, the samples were examined and imaged through LSCM. In vitro anti-biofilm activity assay: In vitro anti-biofilm activity assay: 1 mL of S. aureus and E. coli bacterial suspensions in the logarithmic growth phase were seeded into 24-well plates and incubated at 37°C for 48 h to allow biofilm formation. Next, the original culture medium was replaced with medium pre-treated with different experimental samples. For the NIR group, samples were irradiated with an 808 nm NIR laser for 10 min every 6 h. After a total incubation time of 24 h, the biofilms were gently rinsed twice with PBS and stained with 0.1% crystal violet solution for 20 min. Following image capture, the stained biofilms were solubilized in 95% ethanol, and the absorbance was measured at 590 nm using a UV-Vis spectrophotometer (Thermo, USA) to quantitatively assess the anti-biofilm efficacy of the MNs. To investigate the three-dimensional architecture of biofilms, biofilms of the two bacterial strains were cultivated in confocal dishes and subjected to the same treatment procedures as previously described. After rinsing of the established biofilms, the S. aureus and E. coli biofilms were stained with STYO9/PI reagent. Then, three-dimensional structural images of the biofilms were captured and reconstructed by LSCM. 3.10. Cells isolation and culture system Human umbilical cord mesenchymal stem cells (HucMSC, GPC0191, CCTCC), mouse fibroblasts (L929, GDC106, CCTCC), human umbilical vein endothelial cells (HUVEC, GDC166, CCTCC), and mouse mononuclear macrophage leukemia cells (Raw264.7, GDC0143, CCTCC) were purchased from the China Center for Type Culture Collection (CCTCC, Wuhan, China), and the cells were cultured in accordance with the manufacturer's instructions. Specifically, cells were cultured in DMEM medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin (100×) under standard conditions (37°C, 5% CO2). To simulate the in vitro high glucose (HG) environment of diabetic wounds, cells were cultured in the DMEM medium containing 50 mM glucose. 15 mm × 15 mm MN samples from different experimental groups were immersed in 10 mL of culture medium for 1 week to prepare the sample extracts, then the original cell culture medium was replaced with the corresponding extract to establish a co-culture system between MNs and cells. 3.11. Biocompatibility of MN patches in vitro Hemocompatibility: MN extracts were prepared by incubating MN samples from different experimental groups with PBS prior to testing. A volume of 200 µL of freshly prepared mouse red blood cell suspension was added to each of the following groups: double-distilled water (ddH2O), PBS, and MN extracts, followed by incubation in a 37°C water bath for 2 hours. Following incubation, all samples were centrifuged at 1000 rpm for 5 minutes, and the resulting supernatants were collected. The absorbance of each supernatant was then measured at 550 nm using a multi-mode microplate reader (Thermo, USA). The hemolysis rate was calculated using the following formula: Hemolysis ratio(%) = (AM-Ab) / (Ap-Ab) × 100% (AM: the absorbance of MN group; Ap: the absorbance of positive group; Ab: the absorbance of blank group) Cell Counting Kit-8 assay (CCK-8) assay: L929 cells were seeded into 96-well plates (2000 cells/well) and cultured for 24 hours. Next, the culture medium was replaced with MN-conditioned media from different experimental groups, and the cells were further incubated for 24 and 72 hours, respectively. Cell viability was assessed by the CCK-8 assay kit (Solarbio, CA1210), which was measured at three time points based on the following formula: Cell viability(%) = [(AM - Ab) / (Ac - Ab)] × 100% (AM: the absorbance of the MN group; Ac: the absorbance of the control group; Ab: the absorbance of blank hole) Live-dead cell staining: L929 cells were seeded into 24-well plates at a density of 1 × 104 cells/well and allowed to adhere prior to treatment. Following co-cultivation with MNs from different experimental groups for 24 and 72 hours, respectively, the cells were then stained with Calcein-AM/PI (Beyotime, C2015L) according to the manufacturer's instructions. Fluorescent images were captured using an inverted fluorescence microscope (IX73, Olympus, Japan), and the fluorescence intensity of Calcein-AM was quantitatively analyzed through ImageJ software. 3.12. Scratch test L929 fibroblasts were seeded into 6-well plates at a density of 1 × 106 cells/well and incubated for 24 hours until the cells reached more than 90% confluence. Following 24 hours of serum starvation, wounds were created using a 10 µL pipette tip. Then, the cells were co-cultured with DMEM complete medium, HG medium, or extract solutions from MNs of different experimental groups for 24 hours. Fluorescent staining was performed with reagent at 0 h and 24 h post-wounding, and images were captured with an inverted fluorescence microscope (IX73, Olympus, Japan). The cell migration rate was calculated according to the following formula: Migration rate(%) = (1-At / A0) × 100% (A0: the scratched area at 0 h, At: the scratched area at 24 h) 3.13. Transwell assay L929 cells were seeded at a density of 1 × 104 cells per well in the upper chamber of Transwell-24 plates, and 250 µL of serum-free medium was subsequently added. Meanwhile, 600 µL of DMEM, HG medium and extract solutions from MNs of different experimental groups were placed into the lower chambers for 24-hours incubation. Following two washes with PBS, the cells were fixed with 4% paraformaldehyde for 20 minutes and stained with 0.1% crystal violet dye. Non-migrated cells on the upper surface of the membrane were carefully removed with a cotton swab. Finally, migrated cells were visualized via an inverted microscope (BX53M, Olympus, Japan), and the number of migrated cells was quantified by ImageJ software. 3.14. Tube formation assay The pre-cooled Matrigel (Corning, 356234) was evenly spread onto the bottom of 24-well plates and subsequently incubated in a humidified chamber at 37°C with 5% CO₂ for approximately 30 minutes to allow gel polymerization. Starved HUVECs were then seeded at a density of 2 × 104 cells per well on the solidified Matrigel and co-cultured under various treatment conditions for 6 hours. Tube formation was observed and captured using an inverted microscope (Nikon, Eclipse TS100, Japan), and the total tube length as well as the number of nodes was quantified by ImageJ software. 3.15. MDA determination The content of malondialdehyde (MDA) in HUVECs was measured by the Lipid Peroxidation MDA Assay Kit (Beyotime, S0131S). HUVECs were seeded in 6-well plates and treated in different groups. Then, HUVECs were lysed with cell lysis buffer and the supernatant was obtained by centrifugation. 100 µL of the lysate was mixed with 200 µL of MDA detection working buffer. The samples were heated in a 100°C water bath for 15 minutes, then cooled to room temperature and centrifuged at 1000 ×g for 10 minutes. 200 µL of the supernatant was added to a 96-well plate, and the absorbance was measured at 532 nm using a microplate reader. The MDA concentration was calculated relative to the total protein concentration. 3.16. GSH determination The intracellular total glutathione level was determined using the Total Glutathione Assay Kit (Beyotime, S0052). The GSH working solution and standard curve were prepared in accordance with the manufacturer's instructions. Following various treatments, well-cultured HUVECs were supplemented with three times the volume of protein removal reagent relative to the cell pellet. The cell pellet was incubated at 4°C for 5 minutes and subsequently centrifuged at 10,000 ×g for 10 minutes to collect the supernatant. This supernatant was used for total glutathione measurement. In a 96-well plate, 10 µL of sample, 150 µL of total glutathione detection working solution, and 50 µL of NADPH solution were sequentially added and thoroughly mixed. Finally, the absorbance of A412 was measured and the corresponding GSH concentration was found on the standard curve. 3.17. Intracellular ROS scavenging determination HUVECs were seeded in confocal dishes and exposed to designated treatments. Following treatment, the cells were rinsed with PBS and incubated with the DCFH-DA fluorescent probe (Solarbio, D6470) at 37°C for 30 minutes to assess intracellular ROS levels. After incubation, excess dye was removed by washing the cells three times with serum-free DMEM, and representative images were acquired by LSCM. 3.18. JC-1 staining To evaluate the mitochondrial membrane potential (MMP) in HUVECs, the JC-1 detection kit (Thermo, M34152) was employed. Following the designated treatments, live cells were incubated with freshly prepared JC-1 staining solution in accordance with the manufacturer's instructions, and then maintained in the dark at 37°C in a cell culture incubator for 30 minutes. After incubation, the cells were rinsed twice with JC-1 staining buffer, followed by the addition of DMEM, and representative fluorescence images were observed and acquired by LSCM. 3.19. Determination of intracellular free Fe 2+ The intracellular Fe²⁺ concentration was assessed using the FerroOrange probe (Dojindo, F374). HUVECs were seeded in 24-well plates and incubated in a constant-temperature incubator for 24 hours, then treated with different groups. Following 48 hours of treatment, the cells were rinsed twice with serum-free medium and subsequently incubated with a 10 µM FerroOrange working solution diluted in culture medium at 37°C for 30 minutes. After staining, nuclei were counterstained with Hoechst (Beyotime, C1026), and representative fluorescence images were captured using the LSCM. 3.20. TEM analysis of mitochondrial ultrastructure The morphological alterations of mitochondria in HUVECs were examined by TEM. HUVECs were pre-cultured with different media prior to analysis. Following treatment, the cells were fixed with 2.5% glutaraldehyde at room temperature for 1 hour. Next, cells were scraped from the plates and centrifuged at 800 rpm for 5 minutes. The supernatant was removed, and the cell pellet was resuspended in 1 mL of pre-cooled 2.5% glutaraldehyde and thoroughly mixed. The samples were then stored at 4°C overnight. After fixation, the cells underwent dehydration, embedding, ultrathin sectioning and staining. Mitochondrial morphology was ultimately visualized and analyzed by TEM. Finally, TEM was utilized to observe. 3.21. Western blotting RIPA lysis buffer (Solarbio,R0010) was added to cells, followed by rinsing the samples with TBS buffer (Solarbio, T1081) two to three times. The cells were then centrifuged at 12,000 ×g for 5 minutes at 4°C until complete lysis was achieved. The total cellular protein was quantified using the BCA protein concentration determination kit (Servicebio, G2026-1000T). Protein extracts were loaded onto sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels, separated, and subsequently transferred to PVDF membranes (Solarbio, R0010), which were blocked with 5% fat-free milk for 1 hour. The membranes were then incubated with primary antibodies overnight at 4°C, washed three times with TBST buffer, and incubated with the corresponding secondary antibodies for 1 hour at room temperature. Finally, the target proteins were visualized by the ECL chemiluminescence detection kit (Beyotime, P0018AS), and the optical density of the resulting bands was analyzed utilizing the IPWIN60 software system. ImageJ software was employed to quantify the gray values, that reflect the levels of protein expression. The Western blot antibodies used in this study are detailed in the supporting information. 3.22. Analysis of qRT-PCR The pre-treated cells from different groups were collected by centrifugation and stored for subsequent use. Total RNA was extracted from the cell pellets via Trizol Reagent (Servicebio, G3013). RNA concentration was measured using a Nanodrop 2000 spectrophotometer (Thermo, ND-2000). cDNA was synthesized from the extracted RNA following the manufacturer's instructions for the cDNA synthesis kit (Thermo, K1651). Next, amplification was carried out with HieffTM qPCR SYBR® Green Master Mix (Shanghai Yeasen BioTechnologies, 11201ES03). In accordance with the manufacturer's instructions, the cDNA, primers, nuclease-free water, and gDNA elimination mixture were combined in a 96-well PCR plate. Then, real-time quantitative PCR analysis was conducted by Quantagene q225 real-time PCR system (Kubo Technology, China). Finally, the data of target gene expression levels were calculated utilizing the 2-ΔΔCT method and the forward and reverse primer sequences were demonstrated in supplementary information. 3.23. Immunofluorescence staining of Raw264.7 cells Raw 264.7 cells from different groups were cultured for 48 hours, then fixed with 4% paraformaldehyde and processed for immunofluorescence staining. Following fixation, the cells were washed twice with PBS buffer and blocked with goat serum at room temperature for 1 hour. After fixation, the cells were incubated with the primary antibody overnight at 4°C. Afterward, the samples were incubated with the secondary antibody at 37°C for 1 hour, washed twice with PBS, and counterstained with DAPI working solution for 10 minutes. Immunofluorescence images were acquired by LSCM, and the relative fluorescence intensity was quantified using ImageJ software. 3.24. Flow Cytometry Raw 264.7 cells treated with different groups were harvested and washed three times with PBS. Subsequently, the cells were separately co-incubated on ice for 30 minutes with a phycoerythrin (PE)-conjugated CD206 antibody and an allophycocyanin (APC)-conjugated CD86 antibody. After that, the cells were washed twice with PBS. The fluorescence intensity of the target markers was measured using a flow cytometer (Beckman Coulter, USA), and the acquired data were analyzed with FlowJo software. 3.25. Transcriptomic Profiling and Differential Expression Analysis Total RNA was extracted from Raw264.7 macrophages treated with either DMEM (control group) or MN@Ple/Exo Q10 (MN group) for RNA sequencing. After quality control using FastQC, raw reads were preprocessed, aligned to a reference genome, and normalized via RPKM. Differentially expressed genes (DEGs) were identified using thresholds of |log₂FC|>1 and false discovery rate (FDR)-adjusted P < 0.05. 3.26. Functional Annotation and Pathway Enrichment Gene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analyses were performed utilizing the clusterProfiler R package (v4.10.0), with KEGG-specific analysis executed via the enrichKEGG function. Immune cell subsets, particularly M1/M2 macrophage polarization states, were quantified from TPM-normalized expression matrices using the CIBERSORT algorithm integrated with a murine-specific signature matrix. 3.27. Genome-Scale Pathway Characterization Gene Set Enrichment Analysis (GSEA) characterized key pathways at the whole-genome level, including ferroptosis suppressor gene sets[78] and the top 20 significantly enriched KEGG pathways. Processed data were visualized through the Bioinformation Cloud Platform ( http://www.bioinformatics.com.cn ). 3.28. MN patch capable of transdermal and sustained delivery of Exos Q10 To evaluate the transdermal and sustained delivery performance of Exos Q10 via MN patches, isolated Exos Q10 were pre-incubated with the fluorescent dye Dil for 20 minutes to prepare the labeling solution. For the topical administration (TA) group, the Dil-Exo Q10 solution was sprayed directly onto the wound surface, whereas for the MN group, Dil-Exos Q10 were loaded into HAMA MN and applied directly to the wound site. Wound tissues from both groups were fixed with 4% formalin solution and subsequently incubated with DAPI working solution at room temperature. LSCM was employed to visualize and capture images of the stained tissues. To validate the sustained and stable delivery of Exos Q10 by MN, two delivery methods were compared: sustained release via MN versus direct intradermal injection of Exos Q10 . Specifically, Dil-Exos Q10 were administered via intradermal injection (ID) into the wound area of diabetic mice, while in the MN group, Exos Q10 were delivered through MN@Ple/Exo Q10 . Whole-body fluorescence images were acquired employing an in vivo imaging system (IVIS, Lumina III, PerkinElmer, Waltham, MA, USA), and fluorescence intensity within the wound area was quantitatively analyzed by Living Image software. 3.29. Establishment of infected diabetic wound model 100 male BALB/c mice (6–8 weeks old, weighing 25–27 g) were purchased from Wuhan Wanqian Jiaying Biotechnology Co., Ltd., and all animal experiments were implemented according to the guidelines for laboratory animals established by the Wuhan University Center for Animal Center Experiment/A3-Lab. This research was approved by the Animal Welfare and Ethics Committee of Zhongnan Hospital of Wuhan University (Approval number: ZN2025067). The mice were randomly assigned to five experimental groups (n = 20 per group): Control group, MN group, MN@Ple group, MN@Ple/Exo Q10 group, and MN@Ple/Exo Q10 +NIR group. Mice in different groups were housed separately under standardized conditions. Type I diabetes was induced via intraperitoneal injection of streptozotocin (STZ, 50 mg/kg). A diabetic model was considered successfully established when the blood glucose level exceeded 16.7 mmol/L. Prior to surgery, the mice were anesthetized by isoflurane inhalation. A full-thickness skin wound with a diameter of 10 mm was created on the dorsal surface of each mouse using a skin puncher, followed by the injection of 100 µL of Staphylococcus aureus suspension (1 × 108 CFU/mL) into the wound to establish an infected diabetic wound (IDW) model. 3.30. Infected diabetic wound healing determination After establishment of IDW, the control group received sterile gauze coverage, while the remaining groups were treated with corresponding 15 mm × 15 mm micro-needle patches fully covering the wound area. In the NIR group, the MN patches were irradiated with an 808 nm NIR laser at a power density of 1 W/cm − 2 vertically above the wound area twice daily for 10 minutes per session. Afterwards, on days 3, 7, and 14 following treatment in the respective groups, wound areas were observed and documented by a digital camera, and the residual wound area ratio was quantified using ImageJ software. Subsequently, bacterial specimens were collected from the wound injury sites of each group, and the number of bacteria was assessed via agar plate technique. 3.31. Histological analysis of wounds Tissue samples from each wound group were collected on days 7 and 14 post-injury and fixed in 4% paraformaldehyde. Following paraffin embedding, the tissues were sectioned into 5 µm slices and stained with hematoxylin and eosin (H&E) as well as Masson's trichrome for histopathological evaluation of IDW healing. On day 7 after MN patches treatment, heart, liver, spleen, lung, and kidney tissues from mice were subjected to H&E staining, and images were captured through an optical microscope (BX53M, Olympus, Japan). For immunohistochemical analysis, sections were incubated overnight at 4°C with primary antibodies against GPX4, MPO and VEGF. Subsequently, samples were incubated with secondary antibodies at room temperature for 1 hour. Fluorescent images were acquired using a fluorescence microscope and quantitatively analyzed with ImageJ software. 3.32. Immunofluorescence staining of wounds The ROS level in wound tissues on post-injury day 7 was assessed by the DHE ROS detection kit (Beyotime, S0064). Double immunofluorescence staining for CD86/CD206 was performed on skin wound sections collected on post-injury day 7, while double immunofluorescence staining for CD31/α-SMA was conducted on day 14. Briefly, skin wound tissues were sectioned into 4 µm-thick slices. The harvested tissue sections were blocked with 1% bovine serum albumin for 1 hour and permeabilized with 2% Triton X-100 for 30 minutes. Then, the sections were incubated with primary antibodies at 4°C overnight, washed with PBS buffer, and then incubated with secondary antibodies in the dark for 1 hour. Finally, nuclei were counterstained with DAPI working solution at room temperature. Fluorescent signals were visualized through LSCM and quantitatively analyzed with ImageJ software. 3.33. Enzyme-linked immunosorbent assay (ELISA) Wound tissues from different groups of mice were harvested on post-injury days 7 and 14 and immediately frozen in liquid nitrogen. Following homogenization, the samples were centrifuged to obtain clear tissue extracts. Protein concentration in the supernatant was determined by BCA Protein Assay Kit (Abcam, ab102536). Subsequently, ELISA kits of GPX4 (Reddot Biotech, RD-GPX4-Mu), 4-HNE (SPBIO, SP14767), IL-6 (Abcam, ab222503), IL-10 (Proteintech, KE10103), and VEGF (Beyotime, PV957) were utilized to determine total protein concentrations. 3.34. Statistical analysis Statistical analysis was conducted using GraphPad Prism 9.0 software (GraphPad Software Inc., USA). All measurements were repeated three times by the same operator. Student's t-test was performed to determine statistical differences between two groups, and one-way ANOVA was used to calculate the differences across multiple groups. Data are shown as mean ± SD. ∗P < 0.05, ∗∗P < 0.01, ∗∗∗P < 0.001 and ∗∗∗∗P < 0.0001, respectively. The * indicated the statistical difference between various groups. 4. Conclusion In this study, we developed a differentially released multifunctional core-shell structured MN patch, termed MN@Ple/Exo Q10 , designed to sequentially release the shell-loaded antimicrobial peptide (Ple) and the core-encapsulated therapeutic exosomes (Exos Q10 ) for dynamic modulation of the diabetic wound microenvironment. The core-shell architecture and transdermal mechanical properties of the MN@Ple/Exo Q10 patch were thoroughly characterized. In vitro antibacterial assays demonstrated that Ple released from the MN shell, combined with the photothermally generated heat derived from the excellent light-to-thermal conversion capability of the MN patch, potently eradicated bacteria and disrupted their biofilms. In vitro cellular experiments confirmed that the engineered Exos Q10 effectively suppressed HG-induced endothelial ferroptosis, enhanced the antioxidant capacity of the patch via regulating oxidative stress-related pathways, and modulated macrophage polarization. Additionally, in vivo studies validated that Exo Q10 was efficiently and sustainably delivered into the wound bed by the MN patch, and comprehensively accelerated the healing of infected diabetic wounds in mice through synergistic antibacterial, antioxidant, anti-inflammatory, and pro-angiogenic effects. In conclusion, this MN-based therapeutic platform provides a practical and effective strategy for the synergistic management of diabetic wounds via spatiotemporal regulation of the wound microenvironment. Abbreviations DW, diabetic wound; MN, microneedle; Exo, exosome; CoQ10, Coenzyme Q10; Exos Q10 , Engineered exosomes pretreated with CoQ10 ; HG, high glucose; HucMSCs, human umbilical cord mesenchymal stem cells; MSCs, mesenchymal stem cells; ROS, reactive oxygen species; AMPs, Antimicrobial peptides; Ple, Pleurocidin; NIR, near-infrared; MA, methacryloyl; GelMA, Gelatin-methacryloyl; HAMA, methacryloyl hyaluronic acid; GAG, glycosaminoglycan; DA, dopamine; PDA NPs, polydopamine nanoparticles; PTT, photothermal therapy; ECM, extracellular matrix; EDS, Energy-dispersive X-ray Spectroscopy; TEM, transmission electron microscopy; SEM, scanning electron microscopy; NTA, nanoparticle tracking analysis; HUVECs, human umbilical vein endothelial cells; S. aureus , Staphylococcus aureus; E. coli , Escherichia coli; PI, propidium iodide; HSPs, heat shock proteins; EPS, extracellular polymeric substance; CCK-8, cell counting kit-8; GPX4, glutathione peroxidase 4; ACSL4, long-chain acyl-CoA synthase 4; LPCAT3, lysophosphatidylcholine acyltransferase 3; PUFAs, polyunsaturated fatty acids; PLOOH, phospholipid hydroperoxide; DCFH-DA, Dichlorofluorescein-Diacetate; DEGs, differentially expressed genes; GO, Gene Ontology; KEGG, Kyoto Encyclopedia of Genes and Genomes; GSEA, Gene Set Enrichment Analysis; NQO1, NAD(P)H quinone oxidoreductase 1; HO-1, heme oxygenase-1; GPX1, glutathione peroxidase 1; Nrf2, nuclear factor erythroid 2-related factor 2; IDW, infected diabetic wound; STZ, streptozotocin; RRWA, relative residual wound area; IVIS, in vivo imaging system; TA, topical administration; ID, intradermal injection; H&E, Hematoxylin-eosin; ELISA, enzyme-linked immunosorbent assay; DHE, dihydroethidium; MPO, myeloperoxidase; Declarations Data availability No datasets were generated or analysed during the current study. Acknowledgements The authors would like to thank the use of BioRender.com for creating schemas. The authors would like to thank the support from Zhongnan Hospital of Wuhan University Science and Education Project (KYXM2022015), the National Natural Science Foundation of China (82102626), the Joint Foundation for Translational Medicine and Interdisciplinary Research, Zhongnan Hospital of Wuhan University (ZNJC202310) and the Science and Technology Achievements Transformation Fund, Zhongnan Hospital of Wuhan University (2023CGZH-ZD005). Funding This research was supported by Zhongnan Hospital of Wuhan University Science and Education Project (KYXM2022015), the National Natural Science Foundation of China (82102626), the Joint Foundation for Translational Medicine and Interdisciplinary Research, Zhongnan Hospital of Wuhan University (ZNJC202310) and the Science and Technology Achievements Transformation Fund, Zhongnan Hospital of Wuhan University (2023CGZH-ZD005). Author Contributions The manuscript was written through contributions of all authors. Yuhang Zhan 1 , Zhihan Zhang 1 and Zhiyue Zhang 1 contributed equally to this work. Yuhang Zhan : Data curation; Methodology; Writing – review & editing; Writing – original draft. Zhihan Zhang : Formal analysis; Investigation; Visualization; Writing – review & editing. Zhiyue Zhang : Writing – original draft, Data curation, Methodology, Conceptualization. Hui Liu : Software; Funding acquisition; Data curation. Mengru Lin : Investigation, Software, Methodology. Dilihumaer·Abulimiti : Data curation, Software. Shami·Aihemaiti : Conceptualization, Methodology. Yun Kan : Conceptualization; Investigation. Jinhai Tan : Funding acquisition; Methodology; Supervision. Xi Chen : Conceptualization, Methodology, Resources. Shengxiang Tao : Funding acquisition, Project administration, Supervision, Writing – review & editing. Ethics declarations Ethics approval and consent to participate All animal experiments were implemented according to the guidelines for laboratory animals established by the Wuhan University Center for Animal Center Experiment/A3-Lab. This research was approved by the Animal Welfare and Ethics Committee of Zhongnan Hospital of Wuhan University (Approval number: ZN2025067). Consent for publication All authors have approved the manuscript and agree for the submission. Competing interests The authors declare no competing interests. References Cho N H , Shaw J E , Karuranga S , et al. \"IDF Diabetes Atlas: Global estimates of diabetes prevalence for 2017 and projections for 2045.\" Diabetes Research & Clinical Practice, (2018):271. Noor S , Zubair M , Ahmad J .Diabetic foot ulcer—A review on pathophysiology, classification and microbial etiology[J].Diabetes & metabolic syndrome, 2015. Patel S, Srivastava S, Singh MR, Singh D. 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Supplementary Files supplementoryinformation.docx scheme1.png Scheme 1: The core-shell structured microneedle patch loaded with antimicrobial peptide and engineered exosomes enables chronological management of diabetic wound through by addressing multiple factors. (A) Synthesis Process of Engineered Exosomes: Exo Q10 . (B) Fabrication Protocol for core-shell microneedle patch: MN@Ple/Exo Q10 . (C) Exo Q10 inhibits oxidative stress by modulating intracellular ferroptosis. (D) NIR irradiation of MN@Ple/Exo Q10 patch temporally regulates the wound microenvironment of diabetic mice to promote healing. 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13:31:47\",\"extension\":\"xml\",\"order_by\":42,\"title\":\"\",\"display\":\"\",\"copyAsset\":false,\"role\":\"acdc-reference\",\"size\":263689,\"visible\":true,\"origin\":\"\",\"legend\":\"\",\"description\":\"\",\"filename\":\"eef04c90db404bd6a5474276ac7c32931structuring.xml\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/dc169fa02430aa9b2f98e909.xml\"},{\"id\":93144347,\"identity\":\"597d2548-6cdc-469d-9696-d3edea82378c\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:31:45\",\"extension\":\"html\",\"order_by\":43,\"title\":\"\",\"display\":\"\",\"copyAsset\":false,\"role\":\"acdc-reference\",\"size\":287572,\"visible\":true,\"origin\":\"\",\"legend\":\"\",\"description\":\"\",\"filename\":\"earlyproof.html\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/0a6d5c5585b6b2a09acf9b5b.html\"},{\"id\":93144328,\"identity\":\"54ac9ad3-9ac8-4c59-8f58-2bccedafa991\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:31:44\",\"extension\":\"png\",\"order_by\":1,\"title\":\"Figure 1\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":5264765,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003eThe core-shell structured microneedle patch loaded with antimicrobial peptide and engineered exosomes enables chronological management of diabetic wound through by addressing multiple factors. (A) Synthesis Process of Engineered Exosomes: Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e. (B) Fabrication Protocol for core-shell microneedle patch: MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e. (C) Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e inhibits oxidative stress by modulating intracellular ferroptosis. (D) NIR irradiation of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch temporally regulates the wound microenvironment of diabetic mice to promote healing.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"floatimage2.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/b3c665a7aa0b4dbe1f88989b.png\"},{\"id\":93145252,\"identity\":\"91517998-0b7e-4d3a-bfdb-e59a654b1848\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:39:45\",\"extension\":\"png\",\"order_by\":2,\"title\":\"Figure 2\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":6513192,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003eIn vitro antibacterial performance of MN patches. (A) Photographs of \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003ecolonies on agar plates after various treatments. (B) Quantitative analysis of bacterial survival rate was performed by counting the colony numbers of \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e (n = 3). (C) Representative live/dead fluorescence images and heatmap analysis of bacteria (Scale bar: 10 μm). Growth curves of (D) \\u003cem\\u003eS. aureus\\u003c/em\\u003e and (E) \\u003cem\\u003eE. coli\\u003c/em\\u003e with different treatments for 9 h (n = 3). (F) SEM images of \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003eafter various treatments (Scale bar: 400 nm and 500 nm). (G) Representative 3D LSCM images of biofilms constructed by \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e (Scale bar: 20 μm). (H) Representative images of biofilms stained with crystal violet and (I) quantitative analysis of corresponding biomass absorbance of biofilms at 595 nm (n = 3). The results are shown as the mean ± standard deviation (SD). *P \\u0026lt; 0.05, **P \\u0026lt; 0.01, ***P \\u0026lt; 0.001.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"floatimage3.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/29802b10bd20028264623420.png\"},{\"id\":93143891,\"identity\":\"fc1f01d7-3093-487d-9748-c74686effb49\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:23:43\",\"extension\":\"png\",\"order_by\":3,\"title\":\"Figure 3\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":9875893,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003eEvaluation of in vitro wound healing and angiogenesis capacity of MN patches. (A) Co-culture model of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch and L929 cells and HUVECs. (B) Live/dead cell staining of L929 cells after incubation with different groups for 24 and 72 h (Scale bar: 200 μm). (C) CCK-8 results after co-incubation with MN patches for 24 and 72 hours (n = 3). (D) Hemolysis tests following MN patches exposure (n = 3). (E) Schematic illustration of the promotion of cell migration and angiogenesis under MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch co-culture. (F) Scratch wound healing test on L929 cell migration from 0 to 24 h (Scale bar: 200 μm). (G) Transwell migration images of L929 cells after co-incubation for 48 h (Scale bar: 200 μm). (H) Tube formation assay of HUVECs after co-incubation for 6 h (Scale bar: 200 μm). Quantitative analysis for (I) migration rate, (J) number of migratory cells, (K) total tube length and (L) nb nodes (n = 3). The results are shown as the mean ± standard deviation (SD). *P \\u0026lt; 0.05, **P \\u0026lt; 0.01, ***P \\u0026lt; 0.001.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"floatimage4.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/3ad756de8078737dfd5b4e4b.png\"},{\"id\":93144000,\"identity\":\"c3becdfa-fe2f-485a-b6a8-02c24e4da5fd\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:23:47\",\"extension\":\"png\",\"order_by\":4,\"title\":\"Figure 4\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":6135743,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003eMN patches modulated HG-induced ferroptosis and oxidative stress in vitro. (A) The potential mechanism of Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e in inhibiting HG-induced intracellular ferroptosis. Intracellular (B) MDA and (C) GSH levels of HUVECs (n = 3). (C) Representative DCFH-DA fluorescence images of HUVECs after various treatments (Scale bar: 10 μm). (D) Representative confocal images of JC-1 staining (Scale bar: 50 μm). (E) Representing fluorescent images indicating intracellular Fe\\u003csup\\u003e2+\\u003c/sup\\u003e concentrations in HUVECs treated with different groups (Scale bar: 10 μm). (F) Quantitative analysis of DCFH-DA, FerroOrange and JC-1 staining (n = 5). (H-I) Western blot images and quantitative analysis of GPX4, SLC7A11 and ACSL4 expression in HUVECs (n = 3). (J) TEM images of mitochondrial morphology in HUVECs after different interventions (The yellow arrows and red arrows respectively indicated mitochondria with different structures) (Scale bar: 1 μm). (K) The heatmap of the relative mRNA levels of ferroptosis-related markers in HUVECs. (L-O) Relative mRNA expression of GPX4, SLC7A11, ACSL4 and LPCAT3 among different groups (n = 3). The results are shown as the mean ± standard deviation (SD). *P \\u0026lt; 0.05, **P \\u0026lt; 0.01, ***P \\u0026lt; 0.001.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"floatimage5.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/c02daedc3bc516c0cea61d36.png\"},{\"id\":93143947,\"identity\":\"2f5b0f1c-20f1-4b57-9725-25fea5747133\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:23:44\",\"extension\":\"png\",\"order_by\":5,\"title\":\"Figure 5\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":5507462,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003eImmunomodulatory effects of MN patches on Raw264.7 cells in vitro. (A) Schematic diagram of MN@Ple/Exo\\u003csup\\u003eQ10 \\u003c/sup\\u003einhibiting M1 macrophage polarization while promoting M2 polarization in LPS-induced inflammatory environment. (B) Morphological changes of Raw264.7 cells under different polarization conditions observed by optical microscope (Scale bar: 20 μm). (C) Immunofluorescence images and corresponding heatmaps of M1 (CD86) and M2 (CD206) macrophage markers (Scale bar: 50 μm). (D) Flow cytometry analysis of macrophage polarization markers: CD86 for M1 macrophages and CD206 for M2 macrophages. (E-F) Protein expression of iNOS and CD206 in macrophages analyzed by western blot (n = 3). (G) The heatmap of the relative mRNA levels of inflammation-related markers in Raw264.7 cells. (H-K) Relative mRNA expression of TNF-α, IL-6 and Arg-1 and IL-4 among different groups (n = 3). The results are shown as the mean ± standard deviation (SD). *P \\u0026lt; 0.05, **P \\u0026lt; 0.01, ***P \\u0026lt; 0.001.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"floatimage6.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/3f760269b46ab4270c032d5d.png\"},{\"id\":93144009,\"identity\":\"a594699a-f1bf-4355-83a1-fa369bbf5c2b\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:23:49\",\"extension\":\"png\",\"order_by\":6,\"title\":\"Figure 6\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":5374237,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003eMechanistic profiling of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e in modulating macrophage polarization and oxidative stress. (A) Hierarchical clustering heatmap of DEGs between MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e-treated and control cohorts (n = 4). (B) Volcano plot illustrating transcriptomic alterations. (C) Immune cell infiltration landscape quantified by CIBERSORT algorithm. (D) GO enrichment analysis of DEGs within biological processes. (E) KEGG pathway enrichment profiling. (F) Integrated GSEA of PI3K/AKT signaling pathway and associated DEG expression dynamics. (G) GSEA of MAPK signaling cascade coupled with core DEG expression patterns. (H) GSEA of NF-κB signaling pathway and correlated DEG regulatory networks. (I) Enrichment trajectory of ferroptosis suppressor gene set via GSEA.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"floatimage7.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/6453633e15ad922366bf00ba.png\"},{\"id\":93143897,\"identity\":\"a8eec17d-c2bd-4eda-a599-0b5b0c62987d\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:23:43\",\"extension\":\"png\",\"order_by\":7,\"title\":\"Figure 7\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":5286584,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003eIn vivo performance of MN patches on infected diabetic wound (IDW) healing, and the MN patch for transdermal and sustained delivery of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e in vivo. (A) Schematic illustration of constructing and treating IDWs in diabetic mice at various stages. (B) Representative wound images after various treatments on days 3, 7, and 14 (Scale bar: 2 mm). (C) Quantitative analysis of relative residual wound area (n = 5). (D) Schematic illustration for evaluating the performance of the MN patch in transdermally and continuously delivering Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e by comparing it with topical administration (TA) and intradermal injection (ID) methods, respectively. (E) Fluorescent images of Dil-labeled Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e distribution in wounds after treatment with the MN patch and TA (Scale bar: 50 μm). (F-G) IVIS images and the corresponding quantitative analysis of wounds treated with the MN patch and ID (n = 3). The results are shown as the mean ± standard deviation (SD). *P \\u0026lt; 0.05, **P \\u0026lt; 0.01, ***P \\u0026lt; 0.001.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"floatimage8.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/7ce56119959f26722dd3de2d.png\"},{\"id\":93144006,\"identity\":\"a7b09fbb-b85e-4220-9b6d-e26b13b8a4f3\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:23:48\",\"extension\":\"png\",\"order_by\":8,\"title\":\"Figure 8\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":7641359,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003eHistological analysis of IDW following various treatments. (A) H\\u0026amp;E staining images of wound areas in different groups at days 7 and 14 (Wound area length and thickness of granulation were represented by black arrows and blue vertical segments, respectively. ) (Scale bar: 2 mm and 200 μm). (B) Masson's trichrome staining images of wound areas in different groups at days 7 and 14 (Scale bar: 2 mm and 100 μm). Quantitative analysis of (C)Wound area length and (D) thickness of granulation tissue in H\\u0026amp;E-stained tissue sections (n = 5). (E) Quantitative analysis of relative collagen deposition in Masson's trichrome-stained tissue sections (n = 5). The results are shown as the mean ± standard deviation (SD). *P \\u0026lt; 0.05, **P \\u0026lt; 0.01, ***P \\u0026lt; 0.001.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"floatimage9.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/cacad05b0c195d8e5183785b.png\"},{\"id\":93143991,\"identity\":\"fbd3f535-1a91-44d3-88ed-5d17c77c345d\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:23:46\",\"extension\":\"png\",\"order_by\":9,\"title\":\"Figure 9\",\"display\":\"\",\"copyAsset\":false,\"role\":\"figure\",\"size\":7549562,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003eIn vivo regulation of anti-inflammatory, antioxidant, and angiogenic effects of MN patches. (A) Immunofluorescent staining of ROS content in wounds after different treatments detected utilizing DHE probe (Scale bar: 100 μm). (B) Double immunofluorescence staining for CD86 (red) and CD206 (green) in wound tissues of different groups on day 7 (Scale bar: 50 μm). (C) Double immunofluorescence staining for CD31 (red) and α-SMA (green) in wound tissues of different groups on day 14 (Scale bar: 100 μm). Representative immunohistochemical images of (D) GPX4, (E) MPO and (F) VEGF in wound tissues of different groups (Scale bar: 100 μm). Quantitative analysis of relative fluorescence intensity for (G) DHE, (H) CD86, (I) CD206, and (J) CD31and α-SMA (n = 5). Quantitative analysis of (K) GPX4, (L) MPO and (M) VEGF immunohistochemistry (n = 5). (N) Protein concentration of GPX4, 4-HNE, IL-6, IL-10 and VEGF of wound tissues by ELISA test (n = 5). The results are shown as the mean ± standard deviation (SD). *P \\u0026lt; 0.05, **P \\u0026lt; 0.01, ***P \\u0026lt; 0.001.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"floatimage10.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/5892ee8a39b0d4ddb1e8b82f.png\"},{\"id\":105224830,\"identity\":\"0259410e-e220-4b05-b3a6-bd0afa484dc2\",\"added_by\":\"auto\",\"created_at\":\"2026-03-23 16:16:37\",\"extension\":\"pdf\",\"order_by\":0,\"title\":\"\",\"display\":\"\",\"copyAsset\":false,\"role\":\"manuscript-pdf\",\"size\":61114997,\"visible\":true,\"origin\":\"\",\"legend\":\"\",\"description\":\"\",\"filename\":\"manuscript.pdf\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/91f780f2-77b1-4ac1-925d-ecdea72d11e8.pdf\"},{\"id\":93143899,\"identity\":\"493af10a-f71f-4362-8c5f-250114f71e0f\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:23:43\",\"extension\":\"docx\",\"order_by\":1,\"title\":\"\",\"display\":\"\",\"copyAsset\":false,\"role\":\"supplement\",\"size\":6902406,\"visible\":true,\"origin\":\"\",\"legend\":\"\",\"description\":\"\",\"filename\":\"supplementoryinformation.docx\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/09399988e207e96255ad8da1.docx\"},{\"id\":93143957,\"identity\":\"3c41d6e8-6984-4f43-9f63-c83c621a2b56\",\"added_by\":\"auto\",\"created_at\":\"2025-10-09 13:23:44\",\"extension\":\"png\",\"order_by\":2,\"title\":\"\",\"display\":\"\",\"copyAsset\":false,\"role\":\"supplement\",\"size\":948590,\"visible\":true,\"origin\":\"\",\"legend\":\"\\u003cp\\u003e\\u003cstrong\\u003eScheme 1: \\u003c/strong\\u003eThe core-shell structured microneedle patch loaded with antimicrobial peptide and engineered exosomes enables chronological management of diabetic wound through by addressing multiple factors. (A) Synthesis Process of Engineered Exosomes: Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e. (B) Fabrication Protocol for core-shell microneedle patch: MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e. (C) Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e inhibits oxidative stress by modulating intracellular ferroptosis. (D) NIR irradiation of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch temporally regulates the wound microenvironment of diabetic mice to promote healing.\\u003c/p\\u003e\",\"description\":\"\",\"filename\":\"scheme1.png\",\"url\":\"https://assets-eu.researchsquare.com/files/rs-7689935/v1/a9c28b41c0ac874a018a3860.png\"}],\"financialInterests\":\"No competing interests reported.\",\"formattedTitle\":\"A Core–Shell Microneedle Platform for the Spatiotemporal Codelivery of Dual-Agent Therapeutics Precisely Orchestrates Diabetic Wound Healing\",\"fulltext\":[{\"header\":\"1. Introduction\",\"content\":\"\\u003cp\\u003eDiabetes, a metabolic disorder, has exhibited a steady increase in global prevalence over the past few decades, with projections indicating that the number of affected individuals will reach 693\\u0026nbsp;million by 2045\\u003csup\\u003e[1]\\u003c/sup\\u003e. Over 30% of individuals with diabetes are likely to develop DWs\\u003csup\\u003e[2]\\u003c/sup\\u003e, which are characterized as chronic, non-healing lesions primarily resulting from diabetic neuropathy and vascular complications\\u003csup\\u003e[3]\\u003c/sup\\u003e. Notably, the management of DW is a prolonged and complex process, and severe diabetic foot ulcers frequently culminate in amputation\\u003csup\\u003e[4]\\u003c/sup\\u003e, imposing substantial clinical, caregiving, and economic burdens on society \\u003csup\\u003e[5]\\u003c/sup\\u003e. DWs are typified by hyperglycemia, ischemia, hypoxia, excessive inflammation, and persistent infection \\u003csup\\u003e[6]\\u003c/sup\\u003e. Single interventions such as pharmacological agents, wound dressings or surgical procedures are insufficient to modulate the pathological and physiological disturbances within the wound microenvironment\\u003csup\\u003e[7]\\u003c/sup\\u003e. Therefore, there is an urgent need to develop an ideal therapeutic strategy capable of comprehensively addressing these multifaceted pathophysiological challenges in order to accelerate DW healing.\\u003c/p\\u003e\\u003cp\\u003eExosomes (Exos), double-layer lipid membrane-bound extracellular vesicles, mediate intercellular communication through the delivery of functional proteins\\u003csup\\u003e[8]\\u003c/sup\\u003e, and Exos extracted from human umbilical cord mesenchymal stem cells (HucMSCs) have garnered significant attention in the field of regenerative medicine due to the capacity for tissue repair and immune modulation\\u003csup\\u003e[9\\u0026ndash;10]\\u003c/sup\\u003e.Nevertheless, the HucMSC-derived Exos exhibited minimal efficacy in mitigating HG-triggered ferroptosis of endothelial cells within the wound bed\\u0026mdash;a programmed cell death process marked by pathological reactive oxygen species (ROS) and Fe\\u003csup\\u003e2+\\u003c/sup\\u003e overload, exacerbating oxidative damage and immune imbalance in DWs\\u003csup\\u003e[11]\\u003c/sup\\u003e. Therefore, preparing engineered HucMSC-derived Exos by pretreating HucMSCs to enhance their capacity to inhibit HG-induced ferroptosis is crucial for the repair of DWs. Previous studies have demonstrated that pretreatment of mesenchymal stem cells (MSCs) with pharmacological agents \\u003csup\\u003e[12]\\u003c/sup\\u003e, cytokines\\u003csup\\u003e[13]\\u003c/sup\\u003e and physical stimuli\\u003csup\\u003e[14]\\u003c/sup\\u003e represents an effective strategy to improve the biological activity and therapeutic functionality of MSCs. Coenzyme Q10 (CoQ10), an endogenously synthesized lipophilic antioxidant, potently suppresses ferroptosis\\u003csup\\u003e[15]\\u003c/sup\\u003e and may serve as a potential enhancer for therapeutic Exos. Here, we hypothesize that the engineered Exos prepared by pre-stimulating HucMSCs with CoQ10 could inhibit ferroptosis associated with DWs and accelerate wound healing. Notably, simple Exo therapy cannot effectively target DWs infected by bacteria \\u003csup\\u003e[16]\\u003c/sup\\u003e, as bacterial pathogens and their biofilms can significantly compromise Exos functionality \\u003csup\\u003e[17]\\u003c/sup\\u003e. Thus, targeted strategies to enhance antimicrobial capacity are critically needed.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003cp\\u003eAntimicrobial peptides (AMPs) are unique molecules with molecular weights below 10 kDa that have attracted considerable attention as potential alternatives to conventional antibiotics due to their broad-spectrum antimicrobial activity and high specificity\\u003csup\\u003e[18]\\u003c/sup\\u003e. Pleurocidin (Ple), a natural antimicrobial peptide isolated from various species of Atlantic flounder, is composed of 25 amino acid residues \\u003csup\\u003e[19]\\u003c/sup\\u003e. Previous studies have demonstrated that Ple exhibited broad-spectrum antimicrobial activity with low cytotoxicity, disrupted bacterial membranes, interfered with intracellular processes, and inhibited biofilm formation \\u003csup\\u003e[20]\\u0026minus;[21]\\u003c/sup\\u003e. However, as a linear peptide, Ple is susceptible to inactivation and denaturation in the complex wound microenvironment, while its molecular weight (2.7 kDa) hinders effective transdermal delivery through the stratum corneum via conventional formulations. Consequently, conjugating Ple with functional polymers to develop multifunctional biomedical materials that preserve antimicrobial efficacy while enabling localized delivery is a promising approach\\u003csup\\u003e[22]\\u003c/sup\\u003e. Microneedle (MN) patches represent a promising delivery platform capable of preserving the bioactivity of therapeutic agents while enabling multifaceted and integrated treatment approaches\\u003csup\\u003e[23]\\u003c/sup\\u003e, whereas conventional MN-based therapies are limited by either single-agent loading or simple blending of multiple agents, which often leads to constrained agent-loading capacity, physicochemical incompatibility between agents, and conflicting release kinetics\\u003csup\\u003e[24]\\u0026minus;[25]\\u003c/sup\\u003e. Therefore, to address the complex pathophysiological microenvironment of DW, the development of temporally differentiated release systems is imperative for achieving synergistic multi-component combination therapy.\\u003c/p\\u003e\\u003cp\\u003eHere, considering the distinct dynamic functions of different components in the DW microenvironment during each healing phase, we developed a near-infrared (NIR)-responsive core-shell structured MN system with biphasic agent release capability, designated as MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e. Specifically, Ple was encapsulated into Gelatin-methacryloyl (GelMA) utilizing freeze-drying and spraying technology to form the shell layer of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, while the core layer was constructed by incorporating freeze-dried engineered Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e into methacryloyl hyaluronic acid (HAMA). Furthermore, dopamine (DA) was self-polymerized and incorporated into both the shell and core layers of the MN patch as polydopamine nanoparticles (PDA NPs), endowing the MN patch with superior tissue adhesion and photothermal conversion capabilities. The process in which NIR irradiation is converted into localized hyperthermia is defined as photothermal therapy (PTT) \\u003csup\\u003e[26]\\u003c/sup\\u003e, effectively killing bacteria by disrupting their cell membranes and denaturing intracellular proteins and enzymes while also promoting wound healing \\u003csup\\u003e[27]\\u003c/sup\\u003e. Significantly, this system aims to sequentially deliver therapeutic agents to modulate the pathological microenvironment of DW. On one hand, penetration of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e through the wound barrier causes rapid GelMA shell swelling and Ple release, which synergizes with PTT to concurrently eradicate bacteria and disrupt biofilms. On the other hand, the HAMA core layer undergoes slow degradation following the breakdown of the shell, ensuring sustained and stable release of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e within the wound bed. Accordingly, the released Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e suppress ferroptosis associated with DWs, mitigate oxidative stress, and modulate immune responses, thereby accelerating wound healing in diabetic mice. Overall, this innovative sequential microneedle therapeutic strategy demonstrates considerable potential for the effective management of chronic non-healing diabetic wounds.\\u003c/p\\u003e\"},{\"header\":\"2. Results and discussion\",\"content\":\"\\u003cdiv id=\\\"Sec3\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.1. Preparation and Characterization of MN patch\\u003c/h2\\u003e\\u003cp\\u003eAs illustrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eA, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e consisted of a Gelatin methacryloyl (GelMA)-based MN shell layer loaded with Ple via freeze-drying and spray techniques and methacryloyl hyaluronic acid (HAMA)-based MN core layer encapsulating Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e, and the detailed preparation protocol for MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e is provided in the Methods section. The rational design of distinct structural differences between the shell and core layers is critical for achieving stage-specific agent release in diabetic wound treatment. GelMA, a gelatin derivative modified with methacryloyl (MA) groups, exhibits excellent drug loading/storage capacity and biocompatibility \\u003csup\\u003e[28]\\u003c/sup\\u003e.Moreover, GelMA with a higher degree of methacryloyl substitution for constructing the MN shell layer possesses greater mechanical strength, thereby facilitating more effective skin penetration \\u003csup\\u003e[29]\\u0026minus;[30]\\u003c/sup\\u003e. Hyaluronic acid (HA), a non-sulfated glycosaminoglycan (GAG), is a major component of the extracellular matrix (ECM) in skin tissue\\u003csup\\u003e[31]\\u003c/sup\\u003e, chemically modified into HAMA through esterification with MA. HAMA retains the intrinsic properties of HA, including high hydrophilicity, moisturizing capacity and wound healing functionality, while showing improved stability and resistance to enzymatic degradation, thus ensuring the structural integrity and sustained release of Exos\\u003csup\\u003e[32]\\u0026minus;[33]\\u003c/sup\\u003e, which is crucial for the core layer of the MN patch.\\u003c/p\\u003e\\u003cp\\u003eFigure\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eB presented a representative image of the MN patch, consisting of 100 needle tips arranged in a 10 \\u0026times; 10 configuration. Optical microscopy revealed that the needle tips displayed conical morphology, and the incorporation of PDA imparted a black coloration to the MN surface, as shown in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eC. The MN patch demonstrated a tip height of 650 \\u0026micro;m, a base diameter of 500 \\u0026micro;m, and a tip-to-tip spacing of 550 \\u0026micro;m. Further morphological characterization of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e was performed through a scanning electron microscope (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eD). Subsequently, the fluorescent dyes 5(6)-Carboxyfluorescein and Rhodamine B were employed to label the GelMA shell layer and the HAMA core layer, respectively. The core-shell structure of the MN patch was confirmed using LSCM (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eE). Fluorescence imaging showed a uniform distribution of green and red signals on the outer layer and core of the needle tips, respectively, thereby confirming the successful fabrication of the core-shell MN patch. Furthermore, Energy-dispersive X-ray Spectroscopy (EDS) elemental mapping demonstrated that carbon (C), nitrogen (N), and oxygen (O) were evenly distributed across the MN patch, whereas sodium (Na) and silicon (Si) exhibited low concentrations and showed no signs of aggregation (Fig. \\u003cspan refid=\\\"MOESM1\\\" class=\\\"InternalRef\\\"\\u003eS1\\u003c/span\\u003e).\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003cp\\u003eThe mechanical strength of the prepared MN patches was evaluated through compression testing on a universal testing machine. Force-displacement curves were illustrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eF. The blank MN with a core-shell structure exhibited a continuous upward deformation trend and a higher stress load compared to the blank MN (Core) composed solely of the HAMA core layer. The compressive force at full-tip-height deformation for the blank MN was approximately 1.25 N/Needle, significantly higher than that of the blank MN (Core) (0.94 N/Needle), which indicated that the GelMA shell layer contributed substantially to the mechanical strength of the MN patch, endowing it with excellent mechanical properties that facilitate effective penetration of human skin \\u003csup\\u003e[34]\\u003c/sup\\u003e. The compression testing results revealed continuous force-displacement profiles, indicating robust structural stability of the MNs post-skin penetration without fracture risk. Additionally, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e maintained a force-displacement behavior comparable to that of the blank MN, achieving a mechanical performance of 1.36 N/Needle\\u0026mdash;superior to that of the blank MN. This observation demonstrated that agent loading did not compromise the mechanical integrity of the MN patch, while the freeze-drying cycles involved in the agent-loading process could enhance the hardness of MN tips. As displayed in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eG, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e was effectively inserted into the skin of mice, forming uniformly distributed microchannels. The structural integrity of the needle tips was further confirmed by optical microscopy. To evaluate adhesion performance, peel adhesion forces were measured among different MNs. The MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and blank MN exhibited maximum peel adhesion forces of 0.27 N and 0.28 N, respectively, indicating excellent and comparable adhesive properties. In contrast, the blank MN (Core) showed a significantly lower maximum peel adhesion force of only 0.02 N (Fig. S5). This difference might be attributed to the presence of collagen-derived RGD sequences on the GelMA molecular chains within the MN patch shell layer, which can interact with functional groups on the tissue surface, thereby enabling strong and stable adhesion to biological tissues\\u003csup\\u003e[35]\\u003c/sup\\u003e.\\u003c/p\\u003e\\u003cp\\u003eAs demonstrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eH, the GelMA shell layer coated with Ple and the HAMA core layer loaded with Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e exhibited distinct in vitro release profiles in PBS solution. The GelMA shell layer showed a rapid release profile of Ple, with a cumulative release rate of 81.69\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;3.01% achieved within 8 hours and a plateau reached by 24 hours (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eI). This rapid release behavior was attributed to the surface loading of Ple through the freeze-drying and spray method, which facilitated efficient sustained release within a short timeframe and provided a clear advantage over conventional drug-loading strategies\\u003csup\\u003e[36]\\u003c/sup\\u003e. By contrast, Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e from the HAMA core layer showed a sustained release profile, with cumulative release rates of 18.27\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;2.81%, 38.80\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;3.77%, and 60.83\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;2.41% observed on days 4, 10, 18, respectively (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eJ). This slow yet consistent and prolonged release pattern ensured the continuous delivery of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e. According to the above findings, Ple underwent a burst release during the initial phase upon MN application to the wound, rapidly attaining a plateau, whereas Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e in the core layer revealed a sustained release throughout the entire duration of wound healing. The integration of these two release systems with distinct kinetic profiles conferred MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e with significant potential for spatiotemporal therapeutic applications.\\u003c/p\\u003e\\u003cp\\u003eThe photothermal functionality was engineered through incorporation of PDA within the MN patch\\u003csup\\u003e[37]\\u003c/sup\\u003e. As depicted in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eK, infrared thermographs of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e were obtained at various time points under two irradiation power densities (1 W/cm\\u003csup\\u003e2\\u003c/sup\\u003e and 0.5 W/cm\\u003csup\\u003e2\\u003c/sup\\u003e) to monitor temperature fluctuations. Following separate laser irradiations for 5 minutes at both power densities, the temperature of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e increased from the baseline (25.0\\u0026deg;C) to 46.8\\u0026deg;C and 36.8\\u0026deg;C, respectively, both of which fall within the therapeutic temperature range suitable for mild photothermal therapy in vivo\\u003csup\\u003e[38]\\u003c/sup\\u003e. Furthermore, photothermal curve analysis revealed that the 1 W/cm\\u003csup\\u003e2\\u003c/sup\\u003e group exhibited a faster temperature rise rate compared with the 0.5 W/cm\\u003csup\\u003e2\\u003c/sup\\u003e group (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eL), indicating enhanced photothermal response efficiency and temperature tunability of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e. Additionally, thermal cycling experiments demonstrated that following five cycles of laser activation and deactivation at 1 W/cm\\u003csup\\u003e2\\u003c/sup\\u003e using an 808 nm laser, the maximum temperature achieved by the MN patch did not show significant degradation, with a consistently stable heating rate observed in each cycle (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eM). This robust photothermal stability supported the potential for repeated applications in photothermal therapy\\u003csup\\u003e[39]\\u003c/sup\\u003e.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec4\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.2. Characterization and internalization determination of Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e\\u003c/h2\\u003e\\u003cp\\u003eAs detailed in the Methods section, Exos derived from CoQ10-preconditioned HucMSCs were prepared and subsequently characterized in terms of morphology, size distribution, and marker protein expression utilizing transmission electron microscopy (TEM), nanoparticle tracking analysis (NTA), and Western blotting, respectively. As shown in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eN, HucMSC-Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e exhibited a spherical ultrastructural morphology and demonstrated an average particle size of approximately 124.8 nm (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eO). Western blot analysis was performed to evaluate the expression of exosomal-specific markers, including the transmembrane protein CD63, the cytoplasmic protein TSG101, and the negative marker Calnexin. The results indicated that both CD63 and TSG101 were positively expressed in HucMSC-Exos and HucMSC-Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e with no significant difference between the two groups, whereas the negative marker Calnexin was detected exclusively in the HucMSCs group (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eP), confirming the successful isolation of HucMSC-Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e and their acceptable purity. Subsequently, the cellular internalization of HucMSC-Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e was evaluated employing human umbilical vein endothelial cells (HUVECs) and L929 cells as in vitro models. Following a 6-hour co-incubation period with Dil-labeled HucMSC-Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e, LSCM revealed a uniform distribution of Dil-Exos around the nuclei of both cell types (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig1\\\" class=\\\"InternalRef\\\"\\u003e1\\u003c/span\\u003eQ and R), thereby confirming the efficient uptake of HucMSC-Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e by these cells.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec5\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.3. In vitro antibacterial performance of MN patches\\u003c/h2\\u003e\\u003cp\\u003eBacterial infections significantly impede the healing process of diabetic wounds. Compared to traditional antibiotics, the antimicrobial peptide, Ple, exhibits broad-spectrum antibacterial activity and remarkable inhibitory effects against both Gram-positive and Gram-negative bacteria. Ple targets the bacterial cell membrane through a distinct mechanism, leading to disturbances in membrane structure and disruption of bacterial physiological functions, which substantially reduces the likelihood of developing bacterial resistance \\u003csup\\u003e[40]\\u003c/sup\\u003e. Moreover, the MN patch designed in this study exhibited photothermal conversion performance due to the incorporation of PDA, thereby enhancing the antibacterial function and promoting difficult diabetic wound healing. In this study, Staphylococcus aureus (\\u003cem\\u003eS. aureus\\u003c/em\\u003e) and Escherichia coli (\\u003cem\\u003eE. coli\\u003c/em\\u003e) were chosen as representative strains of Gram-positive and Gram-negative bacteria, respectively, to assess the in vitro anti-infective efficacy of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e.\\u003c/p\\u003e\\u003cp\\u003eThe plate counting method was employed to assess the survival rates of two bacterial species following exposure to various treatment conditions. Numerous colonies of \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e were evident on the agar plates in both the control and MN groups. By contrast, the MN@Ple and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e groups demonstrated a significant reduction in colony counts relative to the control group, with the lowest bacterial density observed in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR group (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eA). The bacterial survival rates of \\u003cem\\u003eS. aureus\\u003c/em\\u003e in the control, MN, MN@Ple, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR groups were 100.70\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;1.34%, 98.25\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;2.52%, 16.71\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;0.59%, 15.76\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;0.36% and 1.71\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;0.36%, respectively. Comparatively, the bacterial survival rates of \\u003cem\\u003eE. coli\\u003c/em\\u003e in the same groups were 100.10\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;1.10%, 100.40\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;1.26%, 23.21\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;0.73%, 18.90\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;0.21% and 1.86\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;0.17%, respectively (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eB). As presented in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eD and E, bacterial absorbance at a wavelength of 600 nm was measured hourly over a 9-hour period, and the resulting data were used to construct growth curves for \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e. The absorbance growth of both bacterial strains in the control and MN groups was markedly higher than that in the other groups. The maximum absorbance values for \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e in the control group reached approximately 1.63 and 1.52, respectively. In contrast, bacterial growth in the MN@Ple and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e groups was significantly inhibited, with absorbance increases plateauing around 6 hours post-treatment. Notably, bacterial activity in the group subjected to 10 minutes of NIR irradiation remained consistently low, with absorbance values for \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e maintained at approximately 0.11 and 0.12.\\u003c/p\\u003e\\u003cp\\u003eSubsequently, live/dead fluorescence staining was conducted on the two bacterial species across different experimental groups, followed by observation and imaging via LSCM, with quantitative analysis conducted through heat map visualization. The results demonstrated that live bacteria labeled with SYTO 9, which emitted green fluorescence, were mainly present in the control and MN groups, whereas minimal red fluorescence, indicative of dead bacteria labeled with propidium iodide (PI), was detected in these groups. In contrast, green fluorescence was markedly reduced in the MN@Ple and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e groups, while red fluorescence showed a moderate increase. Additionally, in the NIR group, the vast majority of bacteria exhibited intense red fluorescence, indicating the most potent antibacterial effect (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eC). Figure\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eF showed the SEM images of \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e after treatment in different groups. The two bacterial species in the control group possessed smooth surfaces and intact cell membranes, suggesting intact cellular morphology. MN treatment did not induce significant morphological alterations in the two bacterial species, while in the MN@Ple and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e groups, some \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e cells displayed mild surface wrinkles and structural deformations. However, both bacterial species in the NIR group demonstrated irregular morphological shrinkage and severe membrane disruption, collectively indicating substantial impairment of bacterial integrity.\\u003c/p\\u003e\\u003cp\\u003eThe biofilm formed during a wound infection protects bacteria from conventional antibiotics and contributes to impaired wound healing. Therefore, the capability to disrupt bacterial biofilms is an essential characteristic of an effective antibacterial material applied to wounds\\u003csup\\u003e[41]\\u003c/sup\\u003e. In this study, the formation of bacterial biofilms in \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e was examined using LSCM and crystal violet staining. As shown in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eG, live/dead staining of the biofilms formed by \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e revealed that both the control and MN groups exhibited continuous biofilm layers with high structural integrity, as evidenced by the strong green fluorescence. In contrast, the biofilms in the MN@Ple and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e groups displayed decreased green fluorescence and reduced continuity. Remarkably, treatment with MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR resulted in severe disruption and dispersion of the biofilms in both bacterial strains, accompanied by intense red fluorescence, indicating extensive bacterial cell death. For crystal violet staining, the control group exhibited complete purple biofilms of \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e, with absorbance values of approximately 2.61 and 2.49, respectively, as determined by quantitative analysis. Conversely, biofilm formation in the NIR group was nearly undetectable, showing significantly reduced absorbance values of approximately 0.47 and 0.49 for \\u003cem\\u003eS. aureus\\u003c/em\\u003e and \\u003cem\\u003eE. coli\\u003c/em\\u003e, respectively, which were markedly lower than those observed in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group (1.34 and 1.22) (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig3\\\" class=\\\"InternalRef\\\"\\u003e2\\u003c/span\\u003eH and I). Overall, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e demonstrated significant antimicrobial activity against both Gram-negative and Gram-positive bacteria, which could be attributed to the synergistic effects of Ple and PTT. On the one hand, the amphiphilic α-helical structure of the antimicrobial peptide Ple, derived from flounder, induces membrane perturbation and permeabilization, enabling Ple to enter bacterial cells and bind to DNA, thus interfering with normal cellular functions\\u003csup\\u003e[42]\\u003c/sup\\u003e. On the other hand, the PTT of the MN patch induced by NIR irradiation can rapidly elevate the local temperature of the MN patches, triggering abnormal expression of bacterial heat shock proteins (HSPs) and depleting the bacteria\\u0026rsquo;s repair capacity\\u003csup\\u003e[43]\\u003c/sup\\u003e. Furthermore, the combination of PTT and Ple effectively disrupted the extracellular polymeric substance (EPS) in bacterial biofilms, and the thermal effect of periodic NIR irradiation then enabled Ple to rapidly penetrate into the damaged interior of the biofilm, allowing for a thorough attack on the bacteria\\u003csup\\u003e[44]\\u003c/sup\\u003e.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec6\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.4. The in vitro biocompatibility of MN patches\\u003c/h2\\u003e\\u003cp\\u003eA co-culture model based on HUVECs and L929 cells was established in the transwell chamber under different grouping conditions (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eA). The in vitro compatibility of the MN patches in different groups was verified employing live/dead cell staining, a cell counting kit-8 (CCK-8) assay and hemolysis test. As illustrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eB, live/dead cell staining was conducted on L929 cells pre-treated for 24 and 72 hours with different groups. The results indicated that none of the MN groups interfered with the normal proliferation of L929 cells, moreover, no significantly increased number of red fluorescent-labelled dead cells were observed in any of the experimental groups compared with the control group. The CCK-8 assay demonstrated a modest reduction in cell proliferation activity over 24 hours and 72 hours within the MN@Ple, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR groups, presumably due to the addition of Ple. Notably, the cell viability across all MN groups remained above 95% at each time point (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eC). For the hemolysis test, the positive control group treated with ddH\\u003csub\\u003e2\\u003c/sub\\u003eO exhibited evident red blood cell lysis, whereas the negative control group treated with PBS showed no hemolytic activity. No significant hemolysis was observed in any of the remaining MN groups (hemolysis ratio\\u0026thinsp;\\u0026lt;\\u0026thinsp;5%), and NIR irradiation did not elicit any notable hemolysis either (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eD). The above biocompatibility test results indicated that the primary materials utilized in the fabrication of MNs demonstrated favorable in vitro biocompatibility. As a member of the antimicrobial peptide family, Ple has demonstrated excellent biocompatibility in previous research \\u003csup\\u003e[45]\\u003c/sup\\u003e. When incorporated at an appropriate concentration (20 \\u0026micro;M) into the prepared MN patch, it demonstrated minimal cytotoxicity and hemolytic activity, thereby satisfying established human safety standards \\u003csup\\u003e[46]\\u003c/sup\\u003e. In addition, the safety features of MNs remained unaffected by the administration of Exos and appropriate NIR hyperthermia. Overall, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e developed in this study demonstrated adequate suitability and safety, meeting the fundamental criteria necessary for clinical applications in wound treatment.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec7\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.5. MN patches promoted wound healing and angiogenesis in vitro\\u003c/h2\\u003e\\u003cp\\u003eAs demonstrated in Figure (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eE), the impact of various groups of MNs on cell migration and angiogenesis in vitro was evaluated through scratch assays, migration assays and tube formation assays. The scratch wound healing experiment revealed that the migration rate of L929 cells after 24 hours in the HG-treated control group (39.85\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;2.78%) was significantly lower than that observed in the negative control group (74.59\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;1.74%) treated with normal culture medium. This discrepancy was attributed to the oxidative stress state and ferroptosis of cells induced by the HG environment in vitro, which subsequently resulted in a reduced cell migration speed \\u003csup\\u003e[47]\\u003c/sup\\u003e. The cell migration rates corresponding to the MN, MN@Ple, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR groups were determined quantitatively as follows: 48.70\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;3.79%, 50.56\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;2.92%, 61.59\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;3.42% and 75.91\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;2.07%, respectively. The enhancement of cell migration observed in the MN and MN@Ple groups was relatively modest. Conversely, a marked improvement in cell migration status was attained through combined treatment with MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch and photothermal therapy (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eF and I). For the Transwell assay, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR group exhibited the maximum number of cells that migrated across the Transwell chamber. Additionally, the quantity of migrating cells in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group without NIR irradiation was greater than that observed in other HG-treated groups and was even slightly higher than that in the negative control group (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eG and J). Tube formation experiments based on HUVECs revealed that MN patches supplemented with Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e effectively mitigated the impairment of capillary-like tube formation induced by HG conditions (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eH). This intervention resulted in enhanced tube quality, longer total tube lengths and a greater number of Nb nodes. Furthermore, the NIR group further augmented the promotion of angiogenesis through MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig8\\\" class=\\\"InternalRef\\\"\\u003e3\\u003c/span\\u003eK and L). In conclusion, both blank MN and MN@Ple without Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e showed a relatively weak effect on cell migration and angiogenesis. In contrast, the release of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e from MNs significantly enhanced cell migration and promoted microtubule formation. This finding corroborates previous research indicating that HucMSC-derived Exos accelerated angiogenesis and facilitated wound healing in vitro\\u003csup\\u003e[48]\\u003c/sup\\u003e. Concurrently, the elevated temperature of the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch, induced by periodic NIR irradiation, exerted a substantial influence on HUVECs and L929 cells, thereby promoting cellular migration and angiogenesis. Moreover, increased molecular thermal motion, induced by elevated temperatures, results in greater Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e release\\u003csup\\u003e[49]\\u0026minus;[50]\\u003c/sup\\u003e.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec8\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.6. MN treatment alleviated HG-induced ferroptosis in endothelial cells\\u003c/h2\\u003e\\u003cp\\u003eAs illustrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eA, the pathological mechanism by which a HG environment induces oxidative stress in cells and subsequently triggers ferroptosis through the accumulation of lipid peroxidation serves as the cellular basis for diabetic wounds\\u003csup\\u003e[51]\\u003c/sup\\u003e. Specifically, HG induces lipid damage in the inner mitochondrial membrane, leading to an increased release of ROS. This event subsequently triggers the transcriptional downregulation of SLC7A11, which functions as a cystine antiporter and directly contributes to a substantial decrease in the GSH/GSSG ratio. Additionally, the inhibition of glutathione peroxidase 4 (GPX4) activity leads to impaired clearance of lipid peroxidation \\u003csup\\u003e[52]\\u003c/sup\\u003e. Alternatively, excessive glucose upregulates long-chain acyl-CoA synthase 4 (ACSL4) and lysophosphatidylcholine acyltransferase 3 (LPCAT3), resulting in the accumulation of polyunsaturated fatty acids (PUFAs), thereby aggregating phospholipid hydroperoxide (PLOOH), which can impair membrane structures and facilitate the release of ROS\\u003csup\\u003e[53]\\u0026minus;[54]\\u003c/sup\\u003e. To further elucidate the mechanism by which MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, in conjunction with NIR, mitigates HG-induced ferroptosis in vitro, an investigation was conducted encompassing four key aspects: cellular ROS levels, accumulation of lipid peroxidation and Fe\\u003csup\\u003e2+\\u003c/sup\\u003e, mitochondrial morphology and structure, as well as the expression of ferroptosis-related genes.\\u003c/p\\u003e\\u003cp\\u003eAs demonstrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eD, the Dichlorofluorescein-Diacetate (DCFH-DA) fluorescent probe was employed as an indicator for ROS to assess intracellular ROS levels. Fluorescence imaging and quantitative analysis across various treatment groups revealed that the green fluorescence of HUVECs subjected to HG conditions was significantly elevated compared to that of the negative control group. Moreover, both MN and MN@Ple treatments did not exhibit a significant reduction in fluorescence intensity relative to the control group. Nevertheless, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group demonstrated a green fluorescence intensity that was only 0.33 times that of the control group, while only a minimal number of cells exhibited this signal in the NIR group (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eG). JC-1, serving as a probe for assessing mitochondrial membrane potential, forms red aggregates within normal mitochondria displaying high membrane potential. Conversely, when mitochondria are compromised and exhibit low membrane potential, JC-1 exists as green monomers and fails to aggregate in the mitochondrial matrix \\u003csup\\u003e[55]\\u003c/sup\\u003e. After labeling with the JC-1 fluorescent probe, HUVECs subjected to different treatments were quantitatively analyzed for JC-1 fluorescence images, and the red/green ratio was subsequently calculated. The results indicated a significant increase in JC-1 monomers alongside a notable decrease in JC-1 aggregates within the HG-treated control group, whereas the negative control group exhibited opposite trends. Furthermore, the red/green ratio observed in both the MN and MN@Ple groups closely mirrored that of the control group. In the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and NIR groups, there was a substantial increase in JC-1 aggregates emitting red fluorescence compared to other HG-treated groups, while levels of JC-1 monomers emitting green fluorescence were significantly diminished (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eE and G).\\u003c/p\\u003e\\u003cp\\u003eGSH and MDA serve as crucial biomarkers for assessing LOOH in endothelial cells undergoing ferroptosis\\u003csup\\u003e[56]\\u003c/sup\\u003e. HUVECs from various treatment groups were lysed and collected for the determination of GSH and MDA levels. The results suggested that the production of MDA in the HG-treated control group was significantly higher compared to the negative control group, while GSH levels were markedly lower. In contrast, the MDA level in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group was approximately 0.29 times that of the HG-treated control group, while GSH levels increased by about 2.8 times. This finding showed that Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e significantly reduced lipid peroxidation levels in cells, importantly, NIR treatment did not affect the efficacy of Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eB and C). The FerroOrange fluorescent probe was utilized to evaluate free Fe\\u003csup\\u003e2+\\u003c/sup\\u003e accumulation in the cytoplasm of HUVECs (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eF). Cells within the HG control group exhibited pronounced orange-red fluorescence signals, indicating a significant aggregation of Fe\\u003csup\\u003e2+\\u003c/sup\\u003e. However, both the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and NIR groups demonstrated reduced fluorescence signals (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eG). As displayed in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eJ, scanning electron microscopy (SEM) was employed to visualize the morphological alterations of mitochondria in HUVECs subjected to various treatment groups. In comparison to the negative control group, mitochondria in the HG control group showed atrophy, increased membrane density, and reduced or absent mitochondrial cristae (indicated by red arrows). Furthermore, mitochondrial damage was also observed in the MN and MN@Ple groups. Remarkably, structural damage to mitochondria was significantly reduced in both the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and NIR groups, showing no significant difference from the negative control. Above results indicated that the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch effectively alleviated HG-induced mitochondrial damage by releasing Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e alongside NIR therapy.\\u003c/p\\u003e\\u003cp\\u003eThe expression levels of ferroptosis-related markers in HUVECs were detected through Western blot analysis. The result revealed that HG significantly reduced the expression levels of GPX4 and SLC7A11 in HUVECs, while increasing ACSL4 levels. However, there were no significant differences in the expression levels between the HG control, MN and MN@Ple groups. Notably, GPX4 expression was upregulated to approximately 0.78 and 0.72 in cells co-cultured with MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR, respectively. Meanwhile, SLC7A11 levels increased to 0.67 and 0.77 in these groups and ACSL4 expression was downregulated to 0.40 and 0.359, respectively (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eH and I). For the RT-qPCR (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eK), MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e significantly improved the mRNA expression levels of GPX4, SLC7A11, ACSL4 and 4 LPCAT3. Specifically, the expressions of GPX4 and SLC7A11 in the HG control group were approximately 0.15 and 0.37 times those in the negative control group, while ACSL4 and LPCAT3 showed expression levels of about 3.2 and 2.1 times, respectively. Conversely, the expression levels in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group were found to be 0.90 for GPX4, 0.78 for SLC7A11, as well as 1.7 and 1.3 for ACSL4 and LPCAT3, respectively. Alternatively, the NIR group did not impair the ability of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e to regulate ferroptosis-related genes; rather, a modest enhancement was observed (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig9\\\" class=\\\"InternalRef\\\"\\u003e4\\u003c/span\\u003eL-O). Additionally, although HucMSC-derived Exos regulated diabetic wound healing across various dimensions in previous studies, the pathological microenvironment of intracellular oxidative stress\\u0026ndash;lipid peroxidation burst\\u0026ndash;ferroptosis could not be improved\\u003csup\\u003e[57]\\u0026minus;[58]\\u003c/sup\\u003e. In this study, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch served as a carrier to release Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e to modulate the SLC7A11-GSH-GPX4 antioxidant pathway and the PUFA-ACSL4-PLOOH axis, both closely linked to ferroptosis, while this regulatory process was unaffected by intermittent temperature increases. In summary, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e exhibited the capability to alleviate cellular ferroptosis in vitro under NIR irradiation.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec9\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.7. In vitro anti-inflammatory capacity of MN patches\\u003c/h2\\u003e\\u003cp\\u003eImmune regulation disorder is a critical factor in the management of DW. The elevated oxidative stress caused by HG results in mitochondrial damage, which further inhibits the metabolic transition from glycolysis to oxidative phosphorylation in macrophages, disrupting the normal immune equilibrium\\u003csup\\u003e[59]\\u003c/sup\\u003e. At the cellular level, classically activated M1 macrophages are unable to transition smoothly into alternatively activated M2 macrophages\\u003csup\\u003e[60]\\u003c/sup\\u003e. The impaired transition impedes the shift in the DW microenvironment from a persistent inflammatory response to tissue repair, ultimately resulting in an inflammatory cytokine storm and increased cell death leading to ROS accumulation, thus establishing a detrimental cycle of inflammation-oxidative stress\\u003csup\\u003e[61]\\u003c/sup\\u003e. In this study, Raw264.7 cells were employed as an in vitro model to investigate the regulatory effects of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e on macrophage polarization, through the application of immunofluorescence, Western blot, qRT-PCR, and flow cytometry.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003cp\\u003eRaw264.7 cells were polarized into M1 macrophages by lipopolysaccharide (LPS), displaying irregular pseudopodia and a strong ability to produce inflammatory mediators. In contrast, M2 macrophages, induced by an appropriate concentration of IL-4 stimulation, were characterized by slender spindle morphology and the secretion of anti-inflammatory cytokines (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig10\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eA and B). In immunofluorescence staining, Raw264.7 cells pre-stimulated with LPS (20 ng/mL) were respectively labeled as markers of M1 and M2 macrophages: CD86 and CD206, while the immunofluorescence images and their corresponding heat maps are shown in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig10\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eC. Compared with the negative control group, the LPS control group exhibited significantly enhanced red fluorescence representing CD86 and nearly undetectable green fluorescence representing CD206, indicating successful polarization of M1 macrophages. Cells co-cultured with MN group revealed high red fluorescence comparable to that of the LPS group, and the MN@Ple group displayed a slight reduction in red fluorescence, whereas both the MN and MN@Ple groups showed low levels of green fluorescence. By contrast, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and NIR groups exhibited markedly reduced red fluorescence and significantly enhanced green fluorescence, suggesting a considerable enhancement in M2 activation. Flow cytometry further confirmed the capacity of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e to promote the reprogramming of macrophages to the M2 phenotype. The percentages of CD206⁻CD86⁺ (M1) cells in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR groups were 37.3% and 30.8%, respectively, significantly lower than the 67.2% observed in the LPS control group. Conversely, the percentages of CD206\\u003csup\\u003e+\\u003c/sup\\u003eCD86\\u003csup\\u003e\\u0026minus;\\u003c/sup\\u003e (M2) in these two groups were approximately 18.1- and 23.7-fold higher than those in the LPS group. These results demonstrate that MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e effectively facilitates the transition of macrophages from the pro-inflammatory M1 phenotype to the anti-inflammatory M2 phenotype (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig10\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eD).\\u003c/p\\u003e\\u003cp\\u003eWestern blot and qRT-PCR were employed to quantitatively assess the differential expression of M1 and M2 macrophage-related markers at the gene level among various experimental groups. For the Western Blot, LPS pretreatment elevated the protein expression of inducible nitric oxide synthase (iNOS), while the protein level of CD206 remained low. Neither the MN group nor the MN@Ple group displayed a significant reduction in iNOS expression or an increase in CD206 expression. However, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and NIR groups exhibited a marked decrease in iNOS protein levels and a significant upregulation of CD206 expression (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig10\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eE and F). The expression levels of pro-inflammatory cytokines (TNF-α and IL-6) associated with M1 macrophages, and anti-inflammatory factors (Arg-1 and IL-4) linked to M2 macrophages across different experimental groups were analyzed by qRT-PCR (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig10\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eG). The relative mRNA expression levels of pro-inflammatory cytokines TNF-α and IL-6 were markedly higher in the LPS-treated control group. Following MN treatment among various experimental groups, these cytokines demonstrated differing levels of downregulation, with the most pronounced reduction observed in the NIR group. Instead, the mRNA levels of anti-inflammatory markers, including Arg-1 and IL-4, were significantly upregulated in the MN@Ple/ Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR group compared to the other groups (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig10\\\" class=\\\"InternalRef\\\"\\u003e5\\u003c/span\\u003eH-K). In conclusion, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e demonstrated anti-inflammatory potential in vitro. Specifically, it inhibited ferroptosis by delivering Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, accelerating the clearance of intracellular lipid peroxidation and reducing its accumulation. Consequently, the intervention effectively disrupted the oxidative stress-ferroptosis-inflammation cycle in Raw264.7 cells, thus facilitating M2 macrophage reprogramming, enhancing the secretion of anti-inflammatory cytokines, and expediting the repair and tissue reconstruction phase\\u003csup\\u003e[62]\\u0026minus;[63]\\u003c/sup\\u003e. Meanwhile, Ple exhibited a notable anti-inflammatory capacity in the present study, consistent with findings from previous investigations\\u003csup\\u003e[64]\\u003c/sup\\u003e. Furthermore, safe and periodic NIR irradiation could enhance the immunomodulatory function of MN patches \\u003csup\\u003e[65]\\u003c/sup\\u003e.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec10\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.8. Transcriptomic analyses revealed the molecular mechanisms underlying the antioxidant and anti-inflammatory effects mediated by MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e\\u003c/h2\\u003e\\u003cp\\u003eTo investigate the molecular mechanisms by which MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e regulates macrophage polarization toward the M2 phenotype, this study conducted whole-transcriptome sequencing analysis on Raw264.7 cells treated with MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e. The results revealed that volcano plot and heatmap analyses identified a total of 1,459 differentially expressed genes (DEGs), among which 893 genes were significantly downregulated and 566 genes were significantly upregulated (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig11\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eA and B). Functional enrichment analysis showed that in Gene Ontology (GO) analysis, these DEGs were primarily enriched in biological processes such as regulation of inflammatory response, oxidoreductase activity, and protein kinase binding (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig11\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eD), suggesting a close association between these processes and the therapeutic effects of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e. Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway analysis further demonstrated that MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e significantly activated inflammatory regulatory pathways such as the NF-κB signaling pathway, TNF signaling pathway, and C-type lectin receptor signaling pathway, while also modulating oxidative stress-related pathways including the p53 signaling pathway, apoptosis, and AMPK signaling pathway (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig11\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eE).\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003cp\\u003eROS exhibit cytotoxicity by inducing oxidative stress through reactions with proteins, lipids, and nucleic acids. Consequently, precise cellular responses to ROS production are critical for preventing further oxidative damage and maintaining cell viability. In this study, Gene Set Enrichment Analysis (GSEA) confirmed that MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e treatment significantly activated NF-κB, MAPK, and PI3K/AKT signaling pathways (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig11\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eF, G, and H). Notably, enhanced NF-κB activity induced the expression of antioxidant proteins such as NAD(P)H quinone oxidoreductase 1 (NQO1), heme oxygenase-1 (HO-1), and glutathione peroxidase 1 (GPX1), thereby counteracting oxidative stress injury. Concurrently, activation of the PI3K/AKT pathway not only suppressed oxidative stress-induced apoptosis but also promoted M2 macrophage polarization to exert anti-inflammatory effects. Ferroptosis is closely linked to the progression of numerous diseases and involves multiple disease-associated signaling pathways. The MAPK pathway inhibits ferroptosis through phosphorylation and activation of nuclear factor erythroid 2-related factor 2 (Nrf2). As a cellular energy sensor, AMPK activation modulates redox homeostasis and iron metabolism to inhibit ferroptosis. Additionally, GSEA results of the ferroptosis suppressor gene set indicated that MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e treatment upregulated the expression of ferroptosis-inhibiting genes, including GPX4 and Stat3 (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig11\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eI). Further CIBERSORT immune infiltration analysis demonstrated that MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e significantly enhanced macrophage polarization toward the M2 phenotype (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig11\\\" class=\\\"InternalRef\\\"\\u003e6\\u003c/span\\u003eC, Fig. S6). Collectively, these results indicate that MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e remodels the diabetic wound immune microenvironment by synergistically attenuating inflammatory responses, enhancing antioxidant capacity, and suppressing ferroptosis.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec11\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.9. MN patches promoted infected diabetic wound healing in vivo\\u003c/h2\\u003e\\u003cp\\u003eThis study utilized a full-thickness infected diabetic wound (IDW) model in BALB/c mice to evaluate the in vivo efficacy of the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch in promoting wound healing. As shown in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig12\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eA, BALB/c mice were induced to develop type I diabetes via streptozotocin (STZ) administration, and subsequently, a circular full-thickness skin wound measuring 1 cm in diameter was created on the dorsum of each mouse. Thereafter, a suspension of \\u003cem\\u003eS. aureus\\u003c/em\\u003e was injected into the diabetic wounds to establish IDW. Wound images of diabetic mice were recorded macroscopically on days 0, 3, 7 and 14 following treatment with different biomaterial groups, and the residual wound areas were then calculated (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig12\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eB). On day 3, the relative residual wound area (RRWA) did not show significant differences among the control, MN, MN@Ple, and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e groups, whereas the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR group exhibited a slightly reduced RRWA (53.49\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;3.62%). Following 7 days of treatment, the RRWA of the control group was approximately 51.88\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;3.80%, while that of the MN and MN@Ple groups was 45.58\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;3.98% and 35.23\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;1.98%, respectively, showing a modest decrease compared to the control group. However, the relative residual wound area of the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group (22.85\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;3.36%) and the NIR group (12.53\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;1.56%) decreased significantly, with the NIR group showing the most notable reduction. On day 14, the wound area in the NIR group was nearly completely healed (4.34\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;1.34%), while the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group also exhibited a significantly lower residual wound area (7.01\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;1.83%), which was notably smaller than that observed in the MN group (16.39\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;2.68%) and the MN@Ple group (13.97\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;1.22%). In contrast, a substantial proportion of wound areas in the control group remained unhealed (26.20\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;2.41%) (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig12\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eC). It is noteworthy that the MN and MN@Ple groups exhibited limited improvement in wound healing within the first 7 days, whereas the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group began to demonstrate a discernible therapeutic effect on wound healing starting from day 3. Furthermore, the NIR group consistently displayed a significantly higher wound healing rate compared to all other groups at every observed time point. In addition, bacteria were isolated from the collected wound tissues treated with various groups and cultured on agar plates to further quantify the in vivo anti-infective capacity. As illustrated in Fig. S8, the control and MN groups exhibited a high bacterial load in the wound area, whereas the bacterial counts were markedly reduced in MN@Ple and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e groups. Notably, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR group, which demonstrated superior antibacterial efficacy, displayed a significantly lower colony count compared to the other treatment groups. In summary, during the early stage of IDW treatment, the shell layer of the MN patch in the putamen structure was fully degraded and completely released the loaded Ple, thereby exerting an antibacterial effect. As the core layer became exposed, it gradually released the loaded Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e. Simultaneously, PTT applied via NIR irradiation synergized with the sustained release of Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e to further accelerate wound healing, enabling sequential and effective treatment of IDW.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec12\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.10. MN patch for transdermal and sustained delivery of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e in vivo\\u003c/h2\\u003e\\u003cp\\u003eThe capacity of the MN patch to achieve transdermal and sustained release within the wound bed is crucial for nano-scale drug delivery platforms\\u003csup\\u003e[66]\\u003c/sup\\u003e. Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e were pre-labeled with Dil dye, and the wound treated with the MN patch was subsequently compared to two distinct local Exos delivery approaches, followed by imaging through LSCM and in vivo imaging system (IVIS) (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig12\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eD). As illustrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig12\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eE, both the topical administration (TA) group and the MN group exhibited intense red fluorescence localized in the epidermis on the day of treatment. By contrast, the MN group demonstrated the capability to penetrate into the dermal layer, forming a wedge-shaped opening and producing a significantly enhanced fluorescence signal in that region. With the progression of the treatment period, by day 7, the skin tissue in the MN group still exhibited noticeable red fluorescence from Dil-Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, which gradually diminished by day 14. Conversely, the TA group showed only faint residual red fluorescence on the skin surface. Particularly, the enhanced transdermal delivery capability of the MN patch ensured a sustained presence of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e within the dermal tissue throughout all stages of wound healing, a significant advantage over conventional topical treatments. To assess the efficacy of the MN patch in achieving sustained delivery of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e to the wound bed, circular full-thickness skin defect mice were treated with intradermal injection (ID) of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e and the MN patch for 14 days. Then, the distribution and retention of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e were monitored and quantified via IVIS on days 3, 7, 10, and 14 (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig12\\\" class=\\\"InternalRef\\\"\\u003e7\\u003c/span\\u003eF and G). The results demonstrated that the red fluorescence emitted from the wound defects on the backs of mice in the MN group gradually decreased as the wound healed, yet remained detectable by day 14. However, the red fluorescence in the ID group began to diminish from day 7 and was no longer observable on days 10 and 14. Overall, the MN treatment enabled the sustained release of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e in vivo for at least 14 days, in contrast to the ID treatment, while the MN system developed in this study functions as an effective Exo delivery platform, capable of achieving complete transdermal penetration and prolonged, stable release of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e, thereby ensuring its continuous therapeutic efficacy in diabetic wound healing.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec13\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.11. Histological evaluation of MN treatment in infected diabetic wound healing\\u003c/h2\\u003e\\u003cp\\u003eHematoxylin-eosin (H\\u0026amp;E) and Masson staining were conducted to assess wound diameter, granulation tissue formation, epithelialization, and collagen deposition in IDW. According to the H\\u0026amp;E staining (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig13\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eA), quantitative analysis of wound length on day 7 demonstrated that the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR group exhibited the shortest wound length (3500.00 \\u0026micro;m), followed by the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group (5384.62 \\u0026micro;m). The MN and MN@Ple groups showed wound lengths of approximately 7307.69 \\u0026micro;m and 6769.23 \\u0026micro;m, respectively, representing a significant reduction compared to the control group (7692.31 \\u0026micro;m). Additionally, the wound lengths of the control, MN, MN@Ple, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and NIR groups after 14 days were 6076.92 \\u0026micro;m, 4653.85 \\u0026micro;m, 4230.77 \\u0026micro;m, 3384.62 \\u0026micro;m and 2423.08 \\u0026micro;m, respectively. Significantly, except for the control group, the wound lengths of all groups were shorter than those at the previous time point, with the NIR group exhibiting the shortest length. This result corresponded to the wound photographs of IDW across different treatment groups observed previously (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig13\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eC). The thickness of granulation tissue and the degree of epithelial reconstruction are key indicators for assessing wound healing. Histological analysis via HE staining on day 7 revealed that both the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group and the NIR group displayed thicker granulation tissue and ongoing epithelial regeneration, with the NIR group demonstrating the most pronounced effects. However, the remaining three groups showed minimal epithelial formation and significantly reduced granulation tissue thickness, particularly in the control group. In contrast, the control group still lacked newly formed epithelium on day 14, and no significant changes in granulation tissue were observed. Nevertheless, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and NIR groups revealed a marked reduction in granulation tissue thickness compared to day 7, with the NIR group displaying the thinnest granulation tissue along with a fully matured thin epithelial layer. Furthermore, the MN and MN@Ple groups showed relatively thicker granulation tissue and incompletely formed epithelium that appeared immature and uneven (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig13\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eD). This accelerated healing process might be attributed to Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e facilitating the transition from the inflammatory to the proliferative phase of wound repair. Additionally, PTT promoted cell migration and the release of growth factors, which, in combination with Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, synergistically enhanced the healing of IDW\\u003csup\\u003e[67]\\u003c/sup\\u003e. Moreover, H\\u0026amp;E staining of the heart, liver, spleen, lung and kidney of mice in different groups presented that MN patches induced no evident organ toxicity (Fig. S9).\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003cp\\u003eAs illustrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig13\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eB, Masson staining revealed the collagen deposition patterns in IDW across different treatment groups. On day 7, the control group exhibited minimal blue-stained collagen fibers, whereas the MN and MN@Ple groups displayed only a limited amount of loosely arranged collagen fibers. Conversely, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR group demonstrated markedly increased collagen deposition compared to the other groups. Particularly, the NIR group showed a larger area of collagen deposition characterized by a denser and more compact arrangement on day 14, indicating a transition from fragile, disorganized type III collagen to robust and well-organized type I collagen fibers\\u003csup\\u003e[68]\\u003c/sup\\u003e. However, in the control, MN and MN@Ple groups, the collagen deposition was still irregular and disordered on day 14, and the relative collagen deposition was less compared with that of the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig13\\\" class=\\\"InternalRef\\\"\\u003e8\\u003c/span\\u003eE). Considering the healing mechanism of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, the diffusion of Ple from the MN shell layer created a favorable microenvironment for IDW healing through its antibacterial effects. Nonetheless, the wound healing capacity of the MN@Ple group remained limited, as evidenced by wounds that persisted in the proliferative phase on day 14, characterized by the presence of thick granulation tissue, incomplete epithelialization, and disorganized type III collagen. Moreover, the HA fragments composed of 6\\u0026ndash;20 disaccharides within the MN core layer promoted the migration and proliferation of dermal fibroblasts, then inducing the deposition of type III collagen and contributing to ECM formation\\u003csup\\u003e[69]\\u003c/sup\\u003e. Notably, Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e released from MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e suppressed oxidative stress in IDW and modulated immune responses, thereby accelerating the transition from the inflammatory phase to the proliferative phase, and subsequently facilitating a faster progression into the remodeling phase, which was characterized by the gradual maturation and thinning of granulation tissue, the development of a mature, thin epithelial layer, and a more organized and dense deposition of collagen. In addition, NIR further synergized with this process to facilitate wound healing.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec14\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.12. In vivo evaluation of antioxidant and anti-Inflammatory for MN patches\\u003c/h2\\u003e\\u003cp\\u003eThe in vitro antioxidant and immunomodulatory effects of MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e were confirmed through prior cellular experiments. Diabetic mice infected with \\u003cem\\u003eS. aureus\\u003c/em\\u003e were treated with different groups of MN patch for seven days. Then, wound tissues were collected and analyzed by utilizing immunofluorescence staining, immunohistochemical staining, and enzyme-linked immunosorbent assay (ELISA) to assess the in vivo antioxidant and anti-inflammatory capacities of MNs. As shown in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eA, the ROS levels in IDW were examined using the dihydroethidium (DHE) fluorescence probe, which is oxidized by ROS present in the wound bed, producing a red fluorescence. The intense red fluorescence observed in the wound bed of the control group indicated a pronounced oxidative stress state in the untreated IDW. The MN and MN@Ple groups exhibited similarly strong red fluorescence, with no statistically significant reduction compared to the control group. However, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group showed a marked decrease in red fluorescence intensity. Additionally, quantitative analysis revealed that the relative fluorescence intensity in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group was only 0.13 times that of the control group, while the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR group exhibited a relative fluorescence intensity of nearly 0.09 times that of the control group, with almost no detectable red ROS signal (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eG). GPX4, a selenium-dependent antioxidant enzyme responsible for reducing peroxides\\u003csup\\u003e[70]\\u003c/sup\\u003e, was evaluated for its expression level in the wound tissue through immunohistochemical staining (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eD and K). The results demonstrated that GPX4 expression levels in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and NIR groups were considerably elevated in comparison with those in the other groups, and these findings were in agreement with the ELISA data. Moreover, ELISA analysis revealed low expression levels of 4-HNE in the control, MN, and MN@Ple groups, whereas significantly reduced levels were observed in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group and NIR group (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eN). Essentially, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e displayed robust antioxidant capacity in diabetic mice, and the sustained-release Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e from the MN core layer effectively mitigated ferroptosis-associated lipid peroxide accumulation in the wound, thereby alleviating oxidative stress induced by excessive ROS.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003cp\\u003eIDW tissues were harvested after seven days of treatment across different groups to investigate the immunomodulatory efficacy of MN treatment during the inflammatory phase. As illustrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eB, CD86/CD206 double immunofluorescence staining on day 7 was employed to assess the expression levels of M1 and M2 macrophages. The results demonstrated that the control group exhibited intense red fluorescence associated with CD86-positive cells and minimal green fluorescence associated with CD206-positive cells, suggesting a predominance of M1 macrophages in untreated IDWs. In contrast, the MN and MN@Ple groups showed a moderate reduction in relative fluorescence intensity for CD86 compared to the control group, while no significant change was observed in the relative fluorescence intensity for CD206, indicating that although the proportion of M1 macrophages responsible for inflammatory responses decreased, the wound remained in the inflammatory phase and had not transitioned to the repair-oriented proliferative phase. Fluorescence quantitative analysis revealed that the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group demonstrated a reduction in relative fluorescence intensity (CD86) to 0.21-fold of the control group through sustained release of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e, accompanied by a 3.09-fold increase in relative fluorescence intensity (CD206). Furthermore, the NIR group displayed significantly higher green fluorescence intensity compared to other groups, with a relative fluorescence intensity (CD206) reaching 4.5-fold that of the control group (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eH and I). The inhibition of inflammatory response in IDWs by MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e was further verified by immunohistochemical staining of myeloperoxidase (MPO), which is a critical biomarker that reflects both the intensity of inflammatory responses and the extent of immune cell activation. Meanwhile, the persistent hyperglycemic environment in diabetic wounds promotes neutrophil activation, resulting in their excessive accumulation at the wound site\\u003csup\\u003e[71]\\u003c/sup\\u003e. As demonstrated in Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eE and L, the relative expression level of MPO in the control group was evidently elevated compared to the other experimental groups. Following treatment with MN and MN@Ple, MPO expression in IDWs was reduced, yet remained higher than that observed in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group. Notably, MPO expression was nearly undetectable in the NIR group. Furthermore, ELISA analysis of wound tissue on day 7 indicated that the expression of the M1 cytokine (IL-6) in the MN and MN@Ple groups was reduced compared to the control group, whereas the expression of the M2 cytokine (IL-10) remained at low levels across the control, MN, and MN@Ple groups. Significantly, in both the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e and NIR groups, IL-6 levels were markedly suppressed, while IL-10 expression was significantly upregulated (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eN). In summary, following application of the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch to IDWs, the linear anionic acidic polysaccharide HA in the core layer regulated inflammatory cell activity through binding to cell surface receptors, thereby partially controlling the wound inflammatory response\\u003csup\\u003e[72]\\u003c/sup\\u003e. However, the efficacy of HA was constrained by pathological conditions such as persistent oxidative stress and HG in diabetic wounds. Moreover, the release of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e in combination with mild photothermal therapy effectively inhibited cellular ferroptosis, reduced intracellular lipid peroxide levels, and alleviated oxidative stress in diabetic wounds, thereby modulating the wound immune response.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec15\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e2.13. MN treatment accelerated angiogenesis in vivo\\u003c/h2\\u003e\\u003cp\\u003eAdequate angiogenesis is essential for IDW healing. Following the inflammatory phase of wound healing, newly formed blood vessels progressively infiltrate the wound during the proliferative phase, delivering oxygen and nutrients to the wound site while removing ROS and metabolic waste products, thereby enhancing the supply of critical components required for tissue regeneration and accelerating wound repair\\u003csup\\u003e[73]\\u003c/sup\\u003e. The expression levels of CD31 and α-SMA were analyzed using double immunofluorescence staining to investigate the distribution patterns of vascular endothelial cells and pericytes in skin wounds among different treatment groups after 14 days of intervention (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eC). Specifically, only minimal red and green fluorescence corresponding to CD31 and α-SMA was observed in the control group, and the MN group exhibited a modest increase in CD31 and α-SMA fluorescence intensity compared to the control group, whereas the MN@Ple group demonstrated a more pronounced enhancement. This improvement might be attributed to the role of HA in the core MN layer in regulating fibroblast and vascular endothelial cell differentiation, thus promoting angiogenesis and new tissue formation\\u003csup\\u003e[74]\\u003c/sup\\u003e. Additionally, Ple-mediated suppression of \\u003cem\\u003eS. aureus\\u003c/em\\u003e in IDWs reduced bacterial oxygen consumption, alleviating the local hypoxic environment, which increased the oxygen partial pressure and directly stimulated endothelial cell proliferation, thereby accelerating neovascularization\\u003csup\\u003e[75]\\u003c/sup\\u003e. Notably, significant angiogenesis was observed in the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group, while the NIR group displayed the highest proportion of CD31-positive endothelial cells and α-SMA-positive pericytes (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eJ). The immunohistochemical staining results illustrated that the expression level of the vascular marker VEGF in the control group was significantly lower than that in the other groups, with the NIR group exhibiting the highest relative VEGF expression. As a ligand, VEGF binds to and activates VEGFR-2, thereby triggering downstream signaling pathways involved in the angiogenic signal transduction cascade \\u003csup\\u003e[76]\\u003c/sup\\u003e, ultimately promoting wound angiogenesis (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eF). In addition, the ELISA results further confirmed that the VEGF expression level in the NIR group was the highest, reaching approximately 384.9 pg/mgprot. In comparison, the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group exhibited the second-highest VEGF expression, with a value of approximately 288.13 pg/mgprot (Fig.\\u0026nbsp;\\u003cspan refid=\\\"Fig18\\\" class=\\\"InternalRef\\\"\\u003e9\\u003c/span\\u003eN). The above results indicated that Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e, derived from HucMSCs, delivered various bioactive molecules such as ANG-2 and Wnt4, which promoted both angiogenesis and vascular maturation\\u003csup\\u003e[77]\\u003c/sup\\u003e. Furthermore, when combined with photothermal therapy, the system induced local vasodilation, enhanced blood flow velocity and oxygen concentration, thereby delivering increased nutrients, immune cells, and oxygen to the non-healing wound.\\u003c/p\\u003e\\u003c/div\\u003e\"},{\"header\":\"3. Materials and methods\",\"content\":\"\\u003cdiv id=\\\"Sec17\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.1. Materials\\u003c/h2\\u003e\\u003cp\\u003eGelatin methacryloyl (GelMA, EFL-GM-60,150kDa), Methacryloyl hyaluronic acid (HAMA, EFL-HAMA-400, 400kDa) and Photoinitiator (lithium phenyl-2,4,6-trimethylbenzoylphosphinate, LAP) were bought from Suzhou Intelligent Manufacturing Research Institute (Suzhou, China). Pleurocidin (HY-P5641), Trehalose (T9531), Coenzyme Q10 (HY-N0111) and Acetone (179124) were bought from MedChemExpress (USA). Dopamine hydrochloride was purchased from Macklin Biochemical Technology (Shanghai, China). Tris hydrochloride solution (PM12630) and glutaraldehyde solution (PM20835) were acquired from PERFEMIKER (Shanghai, China).Polydimethylsiloxane (PDMS) was purchased from Dow Corning (Midland, USA). Anhydrous ethanol, Rhodamine B, 5(6)-Carboxyfluorescein were acquired from Aladdin Bio-Chem Technology Co., Ltd. (Shanghai, China). Modified Eagle Medium (DMEM), Fetal bovine serum (FBS), Penicillin-Streptomycin and Phosphate Buffered Saline (PBS) were bought from Gibco (USA). LB nutrient agar, LB broth, Crystal Violet Staining solution, 4% paraformaldehyde Fix Solution, lipopolysaccharide (LPS), DAPI Staining Solution and Triton X-100 were bought from Beyotime Biotechnology (Shanghai, China). H\\u0026amp;E kit, and Masson kit were purchased from Solarbio (Beijing, China).\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec18\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.2. Antibodies\\u003c/h2\\u003e\\u003cp\\u003eCD63 (1:500, goat polyclonal antibody, SC-15363, Santa Cruz); TSG101 (1:8000, rabbit polyclonal antibody, 28283-1-AP, Proteintech); Calnexin (1:1000, rabbit polyclonal antibody, ab22595, Abcam); GPX4 (1:1000, rabbit polyclonal antibody, 14432-1-AP, Proteintech); SLC7A11 (1:2000, rabbit polyclonal antibody, AF7992, Beyotime); ACSL4 (1:5000, rabbit polyclonal antibody, 22401-1-AP, Proteintech); iNOS (1:2000, rabbit polyclonal antibody, ab3523, Abcam); CD206 (1:1000, rabbit polyclonal antibody, 18704-1-AP, Proteintech); CD86 (1:500, rabbit polyclonal antibody, 13395-1-AP, Proteintech); CD31 (1:1000, rabbit polyclonal antibody, 11265-1-AP, Proteintech); α-SMA (1:1000, rabbit polyclonal antibody, 14395-1-AP, Proteintech); MPO (1:100, rabbit polyclonal antibody, AF7494, Beyotime); VEGF (1:500, rabbit monoclonal antibody, YA013, MCE); GAPDH (1:5000, rabbit polyclonal antibody, 10494-1-AP, Proteintech); β-actin (1:10000, mouse monoclonal antibody, 60008-1-Ig, Proteintech); Goat Anti-Rabbit IgG H\\u0026amp;L (HRP) (1:2000, ab7090, Abcam); Alexa Fluor\\u0026reg; 488 Goat Anti-Rabbit IgG H\\u0026amp;L (1:400, ab150077, Abcam); Alexa Fluor\\u0026reg; 488 Goat Anti-Mouse IgG H\\u0026amp;L (1:400, ab150113, Abcam).\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec19\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.3. Isolation, identification and internalization assay of Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e\\u003c/h2\\u003e\\u003cp\\u003eBefore Exo extraction, 1 mM Coenzyme Q10 (CoQ10) was added to culture dishes containing HucMSCs with a confluence of 90%, followed by a 24-hour pretreatment. Next, the culture medium was replaced with Exo-depleted medium, and the cells were cultured for an additional 48 hours. Finally, the supernatant was collected by centrifugation at 2000 \\u0026times; g for 10 minutes. Then, the supernatant was centrifuged at 10,000 g for 30 minutes, followed by ultracentrifugation at 100,000 g for 70 minutes. After washing with PBS, ultracentrifugation was performed again at 100,000 g for another 70 minutes. The Exos were washed with PBS and then carefully resuspended in sterile PBS. Meanwhile, Exos derived from HucMSCs that were not co-cultured with CoQ10 were prepared utilizing the same method for identification experiments. The morphology and structure of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e were detected by transmission electron microscopy (TEM, Thermo, USA), and the size of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e was determined by Nanoparticle Tracking Analysis (NTA, PARTICLE METRIX GmbH, Germany). The expression of exosomal marker proteins (CD63, TSG101 and Calnexin) was detected by Western blot to confirm the successful separation of Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e.\\u003c/p\\u003e\\u003cp\\u003eExos\\u003csup\\u003eQ10\\u003c/sup\\u003e were labeled by the Dil staining kit (Beyotime, C1991S) according to the manufacturer's instructions. L929 cells and HUVECs were respectively co-cultured with the culture medium containing Dil-labeled Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e for 6 hours. The cultured cells were fixed with 4% paraformaldehyde for 15 minutes, permeabilized with 0.1% Triton X-100 for 10 minutes, and then blocked with 5% BSA for 20\\u0026ndash;30 minutes. Following incubation with FITC at room temperature for 30 minutes, the cell nuclei were then stained with DAPI. The internalization of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e was subsequently visualized using a laser scanning confocal microscope (LSCM, Leica, Germany).\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec20\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.4. Preparation method of MN patches\\u003c/h2\\u003e\\u003cp\\u003eFirst, DA hydrochloride (50 mg) was dissolved in 0.9% (wt/v) Tris solution (pH 8.5) to prepare a mixture, which was magnetically stirred for 24 hours to yield a black solution. Next, the mixture was washed three times with acetone and anhydrous ethanol to remove unreacted monomers, and the resulting Polydopamine nanoparticles (PDA NPs) were collected via centrifugation.\\u003c/p\\u003e\\u003cp\\u003eGelatin methacryloyl (GelMA) was mixed with a 0.25% (w/v) LAP initiator solution to achieve a final 15% (w/v) GelMA solution, and 0.02 wt% PDA NPs were incorporated as the pre-working solution for constructing the shell layer. The resulting mixture was poured into the molding holes of a PDMS negative mold, and bubbles were removed under negative pressure via a freeze dryer for 2 minutes. Thereafter, the PDMS positive mold was placed on top, and the composite was cross-linked under 405 nm UV light. After demolding, the MN shell layer was successfully fabricated. Then, Ple was dissolved in ddH₂O to prepare a 20 \\u0026micro;M aqueous antimicrobial peptide solution, and 5% trehalose was added to preserve protein activity. Then, the preformed shell MN was rapidly immersed in liquid nitrogen and frozen to -196\\u0026deg;C, and the Ple solution was precisely sprayed onto the tips of the MN in a vertical manner. Following rapid cooling in liquid nitrogen, the MN shell layer was transferred to a freeze dryer and subjected to two drying cycles to remove free water. Through repeated freeze-drying cycles, a sufficient quantity of Ple was effectively deposited at the tips of the MN.\\u003c/p\\u003e\\u003cp\\u003eTo prepare the Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e-loaded MN core layer, Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e (3 \\u0026times; 10\\u0026sup1;⁰ particles mL-1) were mixed with 5% (w/v) trehalose, and the mixture was slowly frozen at -80\\u0026deg;C at a cooling rate of 0.5\\u0026deg;C/min over a period of 2 hours. The sample underwent two freeze-drying cycles to obtain an Exos lyophilized powder. This lyophilized powder was then added to a pre-polymer solution composed of 5% (w/v) Hyaluronic Acid Methacrylate (HAMA) and 0.25% (w/v) LAP, with the addition of 0.02 wt% PDA NPs. The mixture was reconstituted by incubation in a 37\\u0026deg;C water bath with shaking for 10 minutes before being poured into the remaining PDMS cavities of the shell layer. Air bubbles were removed under negative pressure using a vacuum pump, and the core layer was cross-linked under 405 nm UV light. Finally, the core-shell MN patch was gently demolded and designated as MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, while it was subjected to secondary freeze-drying, sealed, and stored in the dark at 2\\u0026ndash;8\\u0026deg;C until further use. In addition, blank MN, MN@Ple and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patches with different loading compositions were prepared utilizing the aforementioned method for subsequent experiments, while a core-only blank MN patch (Core) was also fabricated.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec21\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.5. Morphological characterization of MN patch\\u003c/h2\\u003e\\u003cp\\u003eThe surface of the obtained sample was gold-coated prior to observation of the morphological characteristics of MN with a Scanning Electron Microscope (SEM, Zeiss, Germany). Elemental distribution across the MN array was analyzed via Energy-Dispersive X-Ray Spectroscopy (EDS, Zeiss, Germany). Moreover, the overall morphology of MN was examined using an optical microscope (BX53M, Olympus, Japan). To visualize the core-shell structure, the fluorescent dyes 5(6)-Carboxyfluorescein and Rhodamine B were incorporated into the shell and core solutions, respectively, during the MN fabrication process. The core-shell architecture was subsequently observed and imaged by LSCM.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec22\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.6. Determination of mechanical properties of MN patches\\u003c/h2\\u003e\\u003cp\\u003eCompression tests were performed on MN patches from different experimental groups utilizing a universal testing machine (AllroundLine, ZwickRoell, Germany). The MN patches were positioned vertically on a rigid stainless steel platform with the needle tips oriented upward. The detector probe then compressed the needle tips at a constant rate of 5 mm/min in the vertical direction. The resulting stress-strain curves were recorded and analyzed to evaluate the mechanical properties of the MN patches. To evaluate the skin penetration capability of the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, the MN patch was applied to the dorsal skin of Balb/C mice by thumb pressure for 3 minutes, followed by visual observation and photography. The tissue from the pressed area was then excised and examined under an optical microscope to assess the penetration depth of the needle tips into the mouse skin.\\u003c/p\\u003e\\u003cp\\u003eTo evaluate the tissue adhesion performance of the MNs, porcine skin was mounted on the baseplate of a stainless steel platform, and various MN samples were attached to a sensor probe with the needles oriented downward. The needle tips were inserted into the pig skin tissue to establish the initial contact position. Thereafter, a universal testing machine (AllroundLine, ZwickRoell, Germany) was employed to lift the probe vertically at a constant speed of 0.5 mm/min until complete separation of the MN patches from the tissue occurred. The resulting force-time curves were generated based on the recorded data.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec23\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.7. In-vitro release assay of MN patches\\u003c/h2\\u003e\\u003cp\\u003eIn vitro release determination of Ple: Serial dilutions of Ple solutions were prepared in 96-well plates, and their optical density values were measured utilizing a UV-Vis Spectrophotometer (Thermo, USA). A standard curve was subsequently generated utilizing GraphPad Prism software. For the release study, 0.5 mg of shell MN, fabricated via the previously described method, was immersed in 5 mL of PBS. At predetermined time points (1, 2, 4, 6, 8, 16, 24, 32, 40, and 48 hours), 1 mL of the release medium was withdrawn and replaced with 1 mL of fresh PBS. The collected samples were analyzed using the UV-Vis Spectrophotometer to measure the absorbance at 280 nm. The cumulative release amount of Ple was then calculated based on the standard curve.\\u003c/p\\u003e\\u003cp\\u003eIn vitro release determination of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e: To evaluate the release of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e from the core layer of MNs, 0.5 mg of previously prepared core layer MNs were immersed in 5 mL of PBS preheated to 37\\u0026deg;C. At predetermined time points (1, 2, 4, 6, 8, 10, 12, 14, 16, and 18 days), 1 mL of the release medium was withdrawn and replaced with 1 mL of fresh PBS. The total protein concentration of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e from different groups was quantified by the BCA protein assay kit (Thermo Scientific) in accordance with the manufacturer's instructions. The amount of released Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e was subsequently analyzed by multi-mode microplate reader (Thermo, USA), and the cumulative release percentage of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e was calculated as follows:\\u003c/p\\u003e\\u003cp\\u003eCumulative release rate(%)\\u0026thinsp;=\\u0026thinsp;Rt / R0 \\u0026times; 100% (Rt: cumulative release amount at each time point; R0: Total release amount)\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec24\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.8. Photothermal property of MN patch\\u003c/h2\\u003e\\u003cp\\u003eThe dried MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e was placed on the sample platform and irradiated with an 808 nm near-infrared (NIR) laser at 0.5 W/cm\\u0026sup2; and 1 W/cm\\u0026sup2; for 5 minutes. Thermal images were captured every minute using an infrared thermal imager to monitor the temperature changes over time, and a temperature-time curve was plotted accordingly. To evaluate the thermal stability of the MN patch, five heating/cooling cycles were conducted by alternately switching the NIR laser on and off, during which temperature fluctuations were recorded.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec25\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.9. The antibacterial performance of MN patches\\u003c/h2\\u003e\\u003cp\\u003eStaphylococcus aureus (S. aureus) and Escherichia coli (E. coli) liquid cultures were incubated in a shaking incubator at 37\\u0026deg;C for 24 hours until the bacterial populations reached the logarithmic growth phase. MN patches were pre-sterilized via ultraviolet light for 2 hours and stored under sterile conditions prior to use. Subsequently, the bacterial suspensions were co-cultured with MN patches from different experimental groups for 24 hours. In the NIR group, samples were exposed to an 808 nm NIR laser for 10 minutes every 6 hours to induce photothermal effects.\\u003c/p\\u003e\\u003cp\\u003eThe colony count was performed according to the plate count method. Briefly, bacterial suspensions with varying concentrations obtained after co-culturing were gradient diluted 105 times, then uniformly spread onto agar plates, and incubated at 37\\u0026deg;C in a constant temperature incubator for 24 hours. Colony growth of the two bacterial strains was recorded by photography, and the bacterial survival rate was calculated based on the colony counts.\\u003c/p\\u003e\\u003cp\\u003eTo assess the growth dynamics of bacteria across different experimental groups, bacterial cultures in the logarithmic growth phase were co-cultured with various MN patches for a duration of 9 hours. The optical density at 600nm (OD₆₀₀) was measured hourly for each group, and the resulting bacterial growth curves were generated using GraphPad Prism software.\\u003c/p\\u003e\\u003cp\\u003eThe morphological changes of S. aureus and E. coli were analyzed employing SEM (Zeiss, Germany). Treated bacterial samples were collected via centrifugation at 5000 rpm, fixed with 2.5% glutaraldehyde solution, and the supernatant was subsequently removed by centrifugation. The bacterial pellets were then washed three times with PBS, dehydrated through a graded ethanol series (25%, 50%, 75%, and 100%), and finally dropped onto silicon wafers for SEM observation.\\u003c/p\\u003e\\u003cp\\u003eFor the live/dead bacterial staining. Bacterial suspensions from different experimental groups were incubated in confocal dishes under dark conditions for 20 minutes using a SYTO9/PI live and Dead Bacteria Stain Kit (Thermo, L7007), following the manufacturer's protocol. Then, the samples were examined and imaged through LSCM.\\u003c/p\\u003e\\u003cp\\u003eIn vitro anti-biofilm activity assay: In vitro anti-biofilm activity assay: 1 mL of S. aureus and E. coli bacterial suspensions in the logarithmic growth phase were seeded into 24-well plates and incubated at 37\\u0026deg;C for 48 h to allow biofilm formation. Next, the original culture medium was replaced with medium pre-treated with different experimental samples. For the NIR group, samples were irradiated with an 808 nm NIR laser for 10 min every 6 h. After a total incubation time of 24 h, the biofilms were gently rinsed twice with PBS and stained with 0.1% crystal violet solution for 20 min. Following image capture, the stained biofilms were solubilized in 95% ethanol, and the absorbance was measured at 590 nm using a UV-Vis spectrophotometer (Thermo, USA) to quantitatively assess the anti-biofilm efficacy of the MNs. To investigate the three-dimensional architecture of biofilms, biofilms of the two bacterial strains were cultivated in confocal dishes and subjected to the same treatment procedures as previously described. After rinsing of the established biofilms, the S. aureus and E. coli biofilms were stained with STYO9/PI reagent. Then, three-dimensional structural images of the biofilms were captured and reconstructed by LSCM.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec26\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.10. Cells isolation and culture system\\u003c/h2\\u003e\\u003cp\\u003eHuman umbilical cord mesenchymal stem cells (HucMSC, GPC0191, CCTCC), mouse fibroblasts (L929, GDC106, CCTCC), human umbilical vein endothelial cells (HUVEC, GDC166, CCTCC), and mouse mononuclear macrophage leukemia cells (Raw264.7, GDC0143, CCTCC) were purchased from the China Center for Type Culture Collection (CCTCC, Wuhan, China), and the cells were cultured in accordance with the manufacturer's instructions. Specifically, cells were cultured in DMEM medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin (100\\u0026times;) under standard conditions (37\\u0026deg;C, 5% CO2). To simulate the in vitro high glucose (HG) environment of diabetic wounds, cells were cultured in the DMEM medium containing 50 mM glucose. 15 mm \\u0026times; 15 mm MN samples from different experimental groups were immersed in 10 mL of culture medium for 1 week to prepare the sample extracts, then the original cell culture medium was replaced with the corresponding extract to establish a co-culture system between MNs and cells.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec27\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.11. Biocompatibility of MN patches in vitro\\u003c/h2\\u003e\\u003cp\\u003eHemocompatibility: MN extracts were prepared by incubating MN samples from different experimental groups with PBS prior to testing. A volume of 200 \\u0026micro;L of freshly prepared mouse red blood cell suspension was added to each of the following groups: double-distilled water (ddH2O), PBS, and MN extracts, followed by incubation in a 37\\u0026deg;C water bath for 2 hours. Following incubation, all samples were centrifuged at 1000 rpm for 5 minutes, and the resulting supernatants were collected. The absorbance of each supernatant was then measured at 550 nm using a multi-mode microplate reader (Thermo, USA). The hemolysis rate was calculated using the following formula:\\u003c/p\\u003e\\u003cp\\u003eHemolysis ratio(%) = (AM-Ab) / (Ap-Ab) \\u0026times; 100% (AM: the absorbance of MN group; Ap: the absorbance of positive group; Ab: the absorbance of blank group)\\u003c/p\\u003e\\u003cp\\u003eCell Counting Kit-8 assay (CCK-8) assay: L929 cells were seeded into 96-well plates (2000 cells/well) and cultured for 24 hours. Next, the culture medium was replaced with MN-conditioned media from different experimental groups, and the cells were further incubated for 24 and 72 hours, respectively. Cell viability was assessed by the CCK-8 assay kit (Solarbio, CA1210), which was measured at three time points based on the following formula:\\u003c/p\\u003e\\u003cp\\u003eCell viability(%) = [(AM - Ab) / (Ac - Ab)] \\u0026times; 100% (AM: the absorbance of the MN group; Ac: the absorbance of the control group; Ab: the absorbance of blank hole)\\u003c/p\\u003e\\u003cp\\u003eLive-dead cell staining: L929 cells were seeded into 24-well plates at a density of 1 \\u0026times; 104 cells/well and allowed to adhere prior to treatment. Following co-cultivation with MNs from different experimental groups for 24 and 72 hours, respectively, the cells were then stained with Calcein-AM/PI (Beyotime, C2015L) according to the manufacturer's instructions. Fluorescent images were captured using an inverted fluorescence microscope (IX73, Olympus, Japan), and the fluorescence intensity of Calcein-AM was quantitatively analyzed through ImageJ software.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec28\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.12. Scratch test\\u003c/h2\\u003e\\u003cp\\u003eL929 fibroblasts were seeded into 6-well plates at a density of 1 \\u0026times; 106 cells/well and incubated for 24 hours until the cells reached more than 90% confluence. Following 24 hours of serum starvation, wounds were created using a 10 \\u0026micro;L pipette tip. Then, the cells were co-cultured with DMEM complete medium, HG medium, or extract solutions from MNs of different experimental groups for 24 hours. Fluorescent staining was performed with reagent at 0 h and 24 h post-wounding, and images were captured with an inverted fluorescence microscope (IX73, Olympus, Japan). The cell migration rate was calculated according to the following formula:\\u003c/p\\u003e\\u003cp\\u003eMigration rate(%) = (1-At / A0) \\u0026times; 100% (A0: the scratched area at 0 h, At: the scratched area at 24 h)\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec29\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.13. Transwell assay\\u003c/h2\\u003e\\u003cp\\u003eL929 cells were seeded at a density of 1 \\u0026times; 104 cells per well in the upper chamber of Transwell-24 plates, and 250 \\u0026micro;L of serum-free medium was subsequently added. Meanwhile, 600 \\u0026micro;L of DMEM, HG medium and extract solutions from MNs of different experimental groups were placed into the lower chambers for 24-hours incubation. Following two washes with PBS, the cells were fixed with 4% paraformaldehyde for 20 minutes and stained with 0.1% crystal violet dye. Non-migrated cells on the upper surface of the membrane were carefully removed with a cotton swab. Finally, migrated cells were visualized via an inverted microscope (BX53M, Olympus, Japan), and the number of migrated cells was quantified by ImageJ software.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec30\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.14. Tube formation assay\\u003c/h2\\u003e\\u003cp\\u003eThe pre-cooled Matrigel (Corning, 356234) was evenly spread onto the bottom of 24-well plates and subsequently incubated in a humidified chamber at 37\\u0026deg;C with 5% CO₂ for approximately 30 minutes to allow gel polymerization. Starved HUVECs were then seeded at a density of 2 \\u0026times; 104 cells per well on the solidified Matrigel and co-cultured under various treatment conditions for 6 hours. Tube formation was observed and captured using an inverted microscope (Nikon, Eclipse TS100, Japan), and the total tube length as well as the number of nodes was quantified by ImageJ software.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec31\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.15. MDA determination\\u003c/h2\\u003e\\u003cp\\u003eThe content of malondialdehyde (MDA) in HUVECs was measured by the Lipid Peroxidation MDA Assay Kit (Beyotime, S0131S). HUVECs were seeded in 6-well plates and treated in different groups. Then, HUVECs were lysed with cell lysis buffer and the supernatant was obtained by centrifugation. 100 \\u0026micro;L of the lysate was mixed with 200 \\u0026micro;L of MDA detection working buffer. The samples were heated in a 100\\u0026deg;C water bath for 15 minutes, then cooled to room temperature and centrifuged at 1000 \\u0026times;g for 10 minutes. 200 \\u0026micro;L of the supernatant was added to a 96-well plate, and the absorbance was measured at 532 nm using a microplate reader. The MDA concentration was calculated relative to the total protein concentration.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec32\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.16. GSH determination\\u003c/h2\\u003e\\u003cp\\u003eThe intracellular total glutathione level was determined using the Total Glutathione Assay Kit (Beyotime, S0052). The GSH working solution and standard curve were prepared in accordance with the manufacturer's instructions. Following various treatments, well-cultured HUVECs were supplemented with three times the volume of protein removal reagent relative to the cell pellet. The cell pellet was incubated at 4\\u0026deg;C for 5 minutes and subsequently centrifuged at 10,000 \\u0026times;g for 10 minutes to collect the supernatant. This supernatant was used for total glutathione measurement. In a 96-well plate, 10 \\u0026micro;L of sample, 150 \\u0026micro;L of total glutathione detection working solution, and 50 \\u0026micro;L of NADPH solution were sequentially added and thoroughly mixed. Finally, the absorbance of A412 was measured and the corresponding GSH concentration was found on the standard curve.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec33\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.17. Intracellular ROS scavenging determination\\u003c/h2\\u003e\\u003cp\\u003eHUVECs were seeded in confocal dishes and exposed to designated treatments. Following treatment, the cells were rinsed with PBS and incubated with the DCFH-DA fluorescent probe (Solarbio, D6470) at 37\\u0026deg;C for 30 minutes to assess intracellular ROS levels. After incubation, excess dye was removed by washing the cells three times with serum-free DMEM, and representative images were acquired by LSCM.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec34\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.18. JC-1 staining\\u003c/h2\\u003e\\u003cp\\u003eTo evaluate the mitochondrial membrane potential (MMP) in HUVECs, the JC-1 detection kit (Thermo, M34152) was employed. Following the designated treatments, live cells were incubated with freshly prepared JC-1 staining solution in accordance with the manufacturer's instructions, and then maintained in the dark at 37\\u0026deg;C in a cell culture incubator for 30 minutes. After incubation, the cells were rinsed twice with JC-1 staining buffer, followed by the addition of DMEM, and representative fluorescence images were observed and acquired by LSCM.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec35\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.19. Determination of intracellular free Fe\\u003csup\\u003e2+\\u003c/sup\\u003e\\u003c/h2\\u003e\\u003cp\\u003eThe intracellular Fe\\u0026sup2;⁺ concentration was assessed using the FerroOrange probe (Dojindo, F374). HUVECs were seeded in 24-well plates and incubated in a constant-temperature incubator for 24 hours, then treated with different groups. Following 48 hours of treatment, the cells were rinsed twice with serum-free medium and subsequently incubated with a 10 \\u0026micro;M FerroOrange working solution diluted in culture medium at 37\\u0026deg;C for 30 minutes. After staining, nuclei were counterstained with Hoechst (Beyotime, C1026), and representative fluorescence images were captured using the LSCM.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec36\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.20. TEM analysis of mitochondrial ultrastructure\\u003c/h2\\u003e\\u003cp\\u003eThe morphological alterations of mitochondria in HUVECs were examined by TEM. HUVECs were pre-cultured with different media prior to analysis. Following treatment, the cells were fixed with 2.5% glutaraldehyde at room temperature for 1 hour. Next, cells were scraped from the plates and centrifuged at 800 rpm for 5 minutes. The supernatant was removed, and the cell pellet was resuspended in 1 mL of pre-cooled 2.5% glutaraldehyde and thoroughly mixed. The samples were then stored at 4\\u0026deg;C overnight. After fixation, the cells underwent dehydration, embedding, ultrathin sectioning and staining. Mitochondrial morphology was ultimately visualized and analyzed by TEM. Finally, TEM was utilized to observe.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec37\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.21. Western blotting\\u003c/h2\\u003e\\u003cp\\u003eRIPA lysis buffer (Solarbio,R0010) was added to cells, followed by rinsing the samples with TBS buffer (Solarbio, T1081) two to three times. The cells were then centrifuged at 12,000 \\u0026times;g for 5 minutes at 4\\u0026deg;C until complete lysis was achieved. The total cellular protein was quantified using the BCA protein concentration determination kit (Servicebio, G2026-1000T). Protein extracts were loaded onto sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels, separated, and subsequently transferred to PVDF membranes (Solarbio, R0010), which were blocked with 5% fat-free milk for 1 hour. The membranes were then incubated with primary antibodies overnight at 4\\u0026deg;C, washed three times with TBST buffer, and incubated with the corresponding secondary antibodies for 1 hour at room temperature. Finally, the target proteins were visualized by the ECL chemiluminescence detection kit (Beyotime, P0018AS), and the optical density of the resulting bands was analyzed utilizing the IPWIN60 software system. ImageJ software was employed to quantify the gray values, that reflect the levels of protein expression. The Western blot antibodies used in this study are detailed in the supporting information.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec38\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.22. Analysis of qRT-PCR\\u003c/h2\\u003e\\u003cp\\u003eThe pre-treated cells from different groups were collected by centrifugation and stored for subsequent use. Total RNA was extracted from the cell pellets via Trizol Reagent (Servicebio, G3013). RNA concentration was measured using a Nanodrop 2000 spectrophotometer (Thermo, ND-2000). cDNA was synthesized from the extracted RNA following the manufacturer's instructions for the cDNA synthesis kit (Thermo, K1651). Next, amplification was carried out with HieffTM qPCR SYBR\\u0026reg; Green Master Mix (Shanghai Yeasen BioTechnologies, 11201ES03). In accordance with the manufacturer's instructions, the cDNA, primers, nuclease-free water, and gDNA elimination mixture were combined in a 96-well PCR plate. Then, real-time quantitative PCR analysis was conducted by Quantagene q225 real-time PCR system (Kubo Technology, China). Finally, the data of target gene expression levels were calculated utilizing the 2-ΔΔCT method and the forward and reverse primer sequences were demonstrated in supplementary information.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec39\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.23. Immunofluorescence staining of Raw264.7 cells\\u003c/h2\\u003e\\u003cp\\u003eRaw 264.7 cells from different groups were cultured for 48 hours, then fixed with 4% paraformaldehyde and processed for immunofluorescence staining. Following fixation, the cells were washed twice with PBS buffer and blocked with goat serum at room temperature for 1 hour. After fixation, the cells were incubated with the primary antibody overnight at 4\\u0026deg;C. Afterward, the samples were incubated with the secondary antibody at 37\\u0026deg;C for 1 hour, washed twice with PBS, and counterstained with DAPI working solution for 10 minutes. Immunofluorescence images were acquired by LSCM, and the relative fluorescence intensity was quantified using ImageJ software.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec40\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.24. Flow Cytometry\\u003c/h2\\u003e\\u003cp\\u003eRaw 264.7 cells treated with different groups were harvested and washed three times with PBS. Subsequently, the cells were separately co-incubated on ice for 30 minutes with a phycoerythrin (PE)-conjugated CD206 antibody and an allophycocyanin (APC)-conjugated CD86 antibody. After that, the cells were washed twice with PBS. The fluorescence intensity of the target markers was measured using a flow cytometer (Beckman Coulter, USA), and the acquired data were analyzed with FlowJo software.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec41\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.25. Transcriptomic Profiling and Differential Expression Analysis\\u003c/h2\\u003e\\u003cp\\u003eTotal RNA was extracted from Raw264.7 macrophages treated with either DMEM (control group) or MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e (MN group) for RNA sequencing. After quality control using FastQC, raw reads were preprocessed, aligned to a reference genome, and normalized via RPKM. Differentially expressed genes (DEGs) were identified using thresholds of |log₂FC|\\u0026gt;1 and false discovery rate (FDR)-adjusted P\\u0026thinsp;\\u0026lt;\\u0026thinsp;0.05.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec42\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.26. Functional Annotation and Pathway Enrichment\\u003c/h2\\u003e\\u003cp\\u003eGene Ontology (GO) and Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analyses were performed utilizing the clusterProfiler R package (v4.10.0), with KEGG-specific analysis executed via the enrichKEGG function. Immune cell subsets, particularly M1/M2 macrophage polarization states, were quantified from TPM-normalized expression matrices using the CIBERSORT algorithm integrated with a murine-specific signature matrix.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec43\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.27. Genome-Scale Pathway Characterization\\u003c/h2\\u003e\\u003cp\\u003eGene Set Enrichment Analysis (GSEA) characterized key pathways at the whole-genome level, including ferroptosis suppressor gene sets[78] and the top 20 significantly enriched KEGG pathways. Processed data were visualized through the Bioinformation Cloud Platform (\\u003cspan class=\\\"ExternalRef\\\"\\u003e\\u003cspan class=\\\"RefSource\\\"\\u003ehttp://www.bioinformatics.com.cn\\u003c/span\\u003e\\u003cspan address=\\\"http://www.bioinformatics.com.cn\\\" targettype=\\\"URL\\\" class=\\\"RefTarget\\\"\\u003e\\u003c/span\\u003e\\u003c/span\\u003e).\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec44\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.28. MN patch capable of transdermal and sustained delivery of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e\\u003c/h2\\u003e\\u003cp\\u003eTo evaluate the transdermal and sustained delivery performance of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e via MN patches, isolated Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e were pre-incubated with the fluorescent dye Dil for 20 minutes to prepare the labeling solution. For the topical administration (TA) group, the Dil-Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e solution was sprayed directly onto the wound surface, whereas for the MN group, Dil-Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e were loaded into HAMA MN and applied directly to the wound site. Wound tissues from both groups were fixed with 4% formalin solution and subsequently incubated with DAPI working solution at room temperature. LSCM was employed to visualize and capture images of the stained tissues. To validate the sustained and stable delivery of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e by MN, two delivery methods were compared: sustained release via MN versus direct intradermal injection of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e. Specifically, Dil-Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e were administered via intradermal injection (ID) into the wound area of diabetic mice, while in the MN group, Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e were delivered through MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e. Whole-body fluorescence images were acquired employing an in vivo imaging system (IVIS, Lumina III, PerkinElmer, Waltham, MA, USA), and fluorescence intensity within the wound area was quantitatively analyzed by Living Image software.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec45\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.29. Establishment of infected diabetic wound model\\u003c/h2\\u003e\\u003cp\\u003e 100 male BALB/c mice (6\\u0026ndash;8 weeks old, weighing 25\\u0026ndash;27 g) were purchased from Wuhan Wanqian Jiaying Biotechnology Co., Ltd., and all animal experiments were implemented according to the guidelines for laboratory animals established by the Wuhan University Center for Animal Center Experiment/A3-Lab. This research was approved by the Animal Welfare and Ethics Committee of Zhongnan Hospital of Wuhan University (Approval number: ZN2025067). The mice were randomly assigned to five experimental groups (n\\u0026thinsp;=\\u0026thinsp;20 per group): Control group, MN group, MN@Ple group, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e group, and MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e+NIR group. Mice in different groups were housed separately under standardized conditions. Type I diabetes was induced via intraperitoneal injection of streptozotocin (STZ, 50 mg/kg). A diabetic model was considered successfully established when the blood glucose level exceeded 16.7 mmol/L. Prior to surgery, the mice were anesthetized by isoflurane inhalation. A full-thickness skin wound with a diameter of 10 mm was created on the dorsal surface of each mouse using a skin puncher, followed by the injection of 100 \\u0026micro;L of Staphylococcus aureus suspension (1 \\u0026times; 108 CFU/mL) into the wound to establish an infected diabetic wound (IDW) model.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec46\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.30. Infected diabetic wound healing determination\\u003c/h2\\u003e\\u003cp\\u003eAfter establishment of IDW, the control group received sterile gauze coverage, while the remaining groups were treated with corresponding 15 mm \\u0026times; 15 mm micro-needle patches fully covering the wound area. In the NIR group, the MN patches were irradiated with an 808 nm NIR laser at a power density of 1 W/cm\\u0026thinsp;\\u0026minus;\\u0026thinsp;2 vertically above the wound area twice daily for 10 minutes per session. Afterwards, on days 3, 7, and 14 following treatment in the respective groups, wound areas were observed and documented by a digital camera, and the residual wound area ratio was quantified using ImageJ software. Subsequently, bacterial specimens were collected from the wound injury sites of each group, and the number of bacteria was assessed via agar plate technique.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec47\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.31. Histological analysis of wounds\\u003c/h2\\u003e\\u003cp\\u003eTissue samples from each wound group were collected on days 7 and 14 post-injury and fixed in 4% paraformaldehyde. Following paraffin embedding, the tissues were sectioned into 5 \\u0026micro;m slices and stained with hematoxylin and eosin (H\\u0026amp;E) as well as Masson's trichrome for histopathological evaluation of IDW healing. On day 7 after MN patches treatment, heart, liver, spleen, lung, and kidney tissues from mice were subjected to H\\u0026amp;E staining, and images were captured through an optical microscope (BX53M, Olympus, Japan). For immunohistochemical analysis, sections were incubated overnight at 4\\u0026deg;C with primary antibodies against GPX4, MPO and VEGF. Subsequently, samples were incubated with secondary antibodies at room temperature for 1 hour. Fluorescent images were acquired using a fluorescence microscope and quantitatively analyzed with ImageJ software.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec48\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.32. Immunofluorescence staining of wounds\\u003c/h2\\u003e\\u003cp\\u003eThe ROS level in wound tissues on post-injury day 7 was assessed by the DHE ROS detection kit (Beyotime, S0064). Double immunofluorescence staining for CD86/CD206 was performed on skin wound sections collected on post-injury day 7, while double immunofluorescence staining for CD31/α-SMA was conducted on day 14. Briefly, skin wound tissues were sectioned into 4 \\u0026micro;m-thick slices. The harvested tissue sections were blocked with 1% bovine serum albumin for 1 hour and permeabilized with 2% Triton X-100 for 30 minutes. Then, the sections were incubated with primary antibodies at 4\\u0026deg;C overnight, washed with PBS buffer, and then incubated with secondary antibodies in the dark for 1 hour. Finally, nuclei were counterstained with DAPI working solution at room temperature. Fluorescent signals were visualized through LSCM and quantitatively analyzed with ImageJ software.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec49\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.33. Enzyme-linked immunosorbent assay (ELISA)\\u003c/h2\\u003e\\u003cp\\u003eWound tissues from different groups of mice were harvested on post-injury days 7 and 14 and immediately frozen in liquid nitrogen. Following homogenization, the samples were centrifuged to obtain clear tissue extracts. Protein concentration in the supernatant was determined by BCA Protein Assay Kit (Abcam, ab102536). Subsequently, ELISA kits of GPX4 (Reddot Biotech, RD-GPX4-Mu), 4-HNE (SPBIO, SP14767), IL-6 (Abcam, ab222503), IL-10 (Proteintech, KE10103), and VEGF (Beyotime, PV957) were utilized to determine total protein concentrations.\\u003c/p\\u003e\\u003c/div\\u003e\\u003cdiv id=\\\"Sec50\\\" class=\\\"Section2\\\"\\u003e\\u003ch2\\u003e3.34. Statistical analysis\\u003c/h2\\u003e\\u003cp\\u003eStatistical analysis was conducted using GraphPad Prism 9.0 software (GraphPad Software Inc., USA). All measurements were repeated three times by the same operator. Student's t-test was performed to determine statistical differences between two groups, and one-way ANOVA was used to calculate the differences across multiple groups. Data are shown as mean\\u0026thinsp;\\u0026plusmn;\\u0026thinsp;SD. \\u0026lowast;P\\u0026thinsp;\\u0026lt;\\u0026thinsp;0.05, \\u0026lowast;\\u0026lowast;P\\u0026thinsp;\\u0026lt;\\u0026thinsp;0.01, \\u0026lowast;\\u0026lowast;\\u0026lowast;P\\u0026thinsp;\\u0026lt;\\u0026thinsp;0.001 and \\u0026lowast;\\u0026lowast;\\u0026lowast;\\u0026lowast;P\\u0026thinsp;\\u0026lt;\\u0026thinsp;0.0001, respectively. The * indicated the statistical difference between various groups.\\u003c/p\\u003e\\u003c/div\\u003e\"},{\"header\":\"4. Conclusion\",\"content\":\"\\u003cp\\u003eIn this study, we developed a differentially released multifunctional core-shell structured MN patch, termed MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, designed to sequentially release the shell-loaded antimicrobial peptide (Ple) and the core-encapsulated therapeutic exosomes (Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e) for dynamic modulation of the diabetic wound microenvironment. The core-shell architecture and transdermal mechanical properties of the MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e patch were thoroughly characterized. In vitro antibacterial assays demonstrated that Ple released from the MN shell, combined with the photothermally generated heat derived from the excellent light-to-thermal conversion capability of the MN patch, potently eradicated bacteria and disrupted their biofilms. In vitro cellular experiments confirmed that the engineered Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e effectively suppressed HG-induced endothelial ferroptosis, enhanced the antioxidant capacity of the patch via regulating oxidative stress-related pathways, and modulated macrophage polarization. Additionally, in vivo studies validated that Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e was efficiently and sustainably delivered into the wound bed by the MN patch, and comprehensively accelerated the healing of infected diabetic wounds in mice through synergistic antibacterial, antioxidant, anti-inflammatory, and pro-angiogenic effects. In conclusion, this MN-based therapeutic platform provides a practical and effective strategy for the synergistic management of diabetic wounds via spatiotemporal regulation of the wound microenvironment.\\u003c/p\\u003e\"},{\"header\":\"Abbreviations\",\"content\":\"\\u003cp\\u003eDW,\\u0026nbsp;diabetic wound; MN, microneedle; Exo, exosome; CoQ10, Coenzyme Q10; Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e, Engineered exosomes pretreated with CoQ10 ; HG, high glucose; HucMSCs, human umbilical cord mesenchymal stem cells; MSCs, mesenchymal stem cells; ROS, reactive oxygen species; AMPs, Antimicrobial peptides; Ple, Pleurocidin; NIR, near-infrared; MA, methacryloyl; GelMA, Gelatin-methacryloyl; HAMA, methacryloyl hyaluronic acid; GAG, glycosaminoglycan; DA, dopamine; PDA NPs, polydopamine nanoparticles; PTT, photothermal therapy; ECM, extracellular matrix; EDS, Energy-dispersive X-ray Spectroscopy;\\u0026nbsp;TEM, transmission electron microscopy; SEM, scanning electron microscopy; NTA, nanoparticle tracking analysis; HUVECs, human umbilical vein endothelial cells; \\u003cem\\u003eS. aureus\\u003c/em\\u003e, Staphylococcus aureus; \\u003cem\\u003eE. coli\\u003c/em\\u003e, Escherichia coli; PI, propidium iodide; HSPs, heat shock proteins; EPS, extracellular polymeric substance; CCK-8, cell counting kit-8; GPX4, glutathione peroxidase 4; ACSL4, long-chain acyl-CoA synthase 4; LPCAT3, lysophosphatidylcholine acyltransferase 3; PUFAs, polyunsaturated fatty acids; PLOOH, phospholipid hydroperoxide; DCFH-DA, Dichlorofluorescein-Diacetate; DEGs, differentially expressed genes; GO, Gene Ontology; KEGG, Kyoto Encyclopedia of Genes and Genomes; GSEA, Gene Set Enrichment Analysis; NQO1, NAD(P)H quinone oxidoreductase 1; HO-1, heme oxygenase-1; GPX1, glutathione peroxidase 1; Nrf2, nuclear factor erythroid 2-related factor 2; IDW, infected diabetic wound; STZ, streptozotocin; RRWA, relative residual wound area; IVIS, in vivo imaging system; TA, topical administration; ID, intradermal injection; H\\u0026amp;E, Hematoxylin-eosin; ELISA, enzyme-linked immunosorbent assay; DHE, dihydroethidium; MPO, myeloperoxidase;\\u0026nbsp;\\u003c/p\\u003e\"},{\"header\":\"Declarations\",\"content\":\"\\u003cp\\u003e\\u003cstrong\\u003eData availability\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eNo datasets were generated or analysed during the current study.\\u003c/p\\u003e\\n\\u003cp\\u003eAcknowledgements\\u003c/p\\u003e\\n\\u003cp\\u003eThe authors would like to thank the use of BioRender.com for creating schemas. The authors would like to thank the support from Zhongnan Hospital of Wuhan University Science and Education Project (KYXM2022015), the National Natural Science Foundation of China (82102626), the Joint Foundation for Translational Medicine and Interdisciplinary Research, Zhongnan Hospital of Wuhan University (ZNJC202310) and the Science and Technology Achievements Transformation Fund, Zhongnan Hospital of Wuhan University (2023CGZH-ZD005).\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eFunding\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eThis research was supported by Zhongnan Hospital of Wuhan University Science and Education Project (KYXM2022015), the National Natural Science Foundation of China (82102626), the Joint Foundation for Translational Medicine and Interdisciplinary Research, Zhongnan Hospital of Wuhan University (ZNJC202310) and the Science and Technology Achievements Transformation Fund, Zhongnan Hospital of Wuhan University (2023CGZH-ZD005).\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eAuthor Contributions\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eThe manuscript was written through contributions of all authors.\\u0026nbsp;Yuhang Zhan\\u003csup\\u003e1\\u003c/sup\\u003e, Zhihan Zhang\\u003csup\\u003e1\\u003c/sup\\u003e and Zhiyue Zhang\\u003csup\\u003e1\\u003c/sup\\u003e contributed equally to this work.\\u0026nbsp;\\u003cstrong\\u003eYuhang Zhan\\u003c/strong\\u003e: Data curation; Methodology; Writing – review \\u0026amp; editing; Writing – original draft. \\u003cstrong\\u003eZhihan Zhang\\u003c/strong\\u003e: Formal analysis; Investigation; Visualization; Writing – review \\u0026amp; editing.\\u003cstrong\\u003e\\u0026nbsp;Zhiyue Zhang\\u003c/strong\\u003e: Writing – original draft, Data curation, Methodology, Conceptualization. \\u003cstrong\\u003eHui Liu\\u003c/strong\\u003e: Software; Funding acquisition; Data curation. \\u003cstrong\\u003eMengru Lin\\u003c/strong\\u003e: Investigation, Software, Methodology. \\u003cstrong\\u003eDilihumaer·Abulimiti\\u003c/strong\\u003e: Data curation, Software. \\u003cstrong\\u003eShami·Aihemaiti\\u003c/strong\\u003e: Conceptualization, Methodology. \\u003cstrong\\u003eYun Kan\\u003c/strong\\u003e: Conceptualization; Investigation. \\u003cstrong\\u003eJinhai Tan\\u003c/strong\\u003e: Funding acquisition; Methodology; Supervision. \\u003cstrong\\u003eXi Chen\\u003c/strong\\u003e: Conceptualization, Methodology, Resources. \\u003cstrong\\u003eShengxiang Tao\\u003c/strong\\u003e: Funding acquisition, Project administration, Supervision, Writing – review \\u0026amp; editing.\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eEthics declarations\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eEthics approval and consent to participate\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eAll animal experiments were implemented according to the guidelines for laboratory animals established by the Wuhan University Center for Animal Center Experiment/A3-Lab. This research was approved by the Animal Welfare and Ethics Committee of Zhongnan Hospital of Wuhan University (Approval number: ZN2025067).\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eConsent for publication\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eAll authors have approved the manuscript and agree for the submission.\\u003c/p\\u003e\\n\\u003cp\\u003e\\u003cstrong\\u003eCompeting interests\\u003c/strong\\u003e\\u003c/p\\u003e\\n\\u003cp\\u003eThe authors declare no competing interests.\\u003c/p\\u003e\"},{\"header\":\"References\",\"content\":\"\\u003col\\u003e\\n\\u003cli\\u003eCho N H , Shaw J E , Karuranga S , et al. \\u0026quot;IDF Diabetes Atlas: Global estimates of diabetes prevalence for 2017 and projections for 2045.\\u0026quot; Diabetes Research \\u0026amp; Clinical Practice, (2018):271.\\u003c/li\\u003e\\n\\u003cli\\u003eNoor S , Zubair M , Ahmad J .Diabetic foot ulcer\\u0026mdash;A review on pathophysiology, classification and microbial etiology[J].Diabetes \\u0026amp; metabolic syndrome, 2015.\\u003c/li\\u003e\\n\\u003cli\\u003ePatel S, Srivastava S, Singh MR, Singh D. 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Histone lysine crotonylation accelerates ACSL4-mediated ferroptosis of keratinocytes via modulating autophagy in diabetic wound healing. Pharmacological Research, 2025(213): 107632.\\u003c/li\\u003e\\n\\u003cli\\u003eLiming Wang, Jun Chen, Jia Song, et al. Activation of the Wnt/\\u0026beta;-catenin signalling pathway enhances exosome production by hucMSCs and improves their capability to promote diabetic wound healing. Journal of Nanobiotechnology, 2024(373).\\u003c/li\\u003e\\n\\u003cli\\u003eJinlang Fu, Ding Wang, Zinan Tang, et al. NIR-responsive electrospun nanofiber dressing promotes diabetic-infected wound healing with programmed combined temperature-coordinated photothermal therapy. Journal of Nanobiotechnology, 2024(384).\\u003c/li\\u003e\\n\\u003cli\\u003eJunjie Zeng, Minhui Lu, Yu Wang,et al. Photothermal Fish Gelatin-Graphene Microneedle Patches for Chronic Wound Treatment. Small, 2024(20): 2405847.\\u003c/li\\u003e\\n\\u003cli\\u003eCui S, Liu X, Liu Y, Hu W, Ma K, Huang Q, Chu Z, Tian L, Meng S, Su J, Zhang W, Li H, Fu X, Zhang C. Autophagosomes Defeat Ferroptosis by Decreasing Generation and Increasing Discharge of Free Fe2+ in Skin Repair Cells to Accelerate Diabetic Wound Healing. Adv Sci (Weinh). 2023 Sep;10(25):e2300414.\\u003c/li\\u003e\\n\\u003cli\\u003eHadian K, Stockwell BR. SnapShot: Ferroptosis. Cell. 2020 May 28;181(5):1188-1188.e1.\\u003c/li\\u003e\\n\\u003cli\\u003eShuangwen Li, Yuan Li, Ziyu Wu, et al. Diabetic ferroptosis plays an important role in triggering on inflammation in diabetic wound. American Journal of Physiology-Endocrinology and Metabolism, 2021(321): E509-E520.\\u003c/li\\u003e\\n\\u003cli\\u003eZhang W, Feng J, Ni Y, Li G, Wang Y, Cao Y, Zhou M, Zhao C. The role of SLC7A11 in diabetic wound healing: novel insights and new therapeutic strategies. Front Immunol. 2024 Sep 3;15:1467531.\\u003c/li\\u003e\\n\\u003cli\\u003ePerelman A, Wachtel C, Cohen M, Haupt S, Shapiro H, Tzur A. JC-1: alternative excitation wavelengths facilitate mitochondrial membrane potential cytometry. Cell Death Dis. 2012 Nov 22;3(11):e430. \\u003c/li\\u003e\\n\\u003cli\\u003eZhang S, Hu R, Geng Y, Chen K, Wang L, Imam MU. The Regulatory Effects and the Signaling Pathways of Natural Bioactive Compounds on Ferroptosis. Foods. 2021 Dec 1;10(12):2952.\\u003c/li\\u003e\\n\\u003cli\\u003eWu Q, Guo Y, Li H, Zhang D, Wang S, Hou J, Cheng N, Huang M, Luo L, Li Y, Zhao Y, Tan H, Jin C. Recombinant human collagen I/carboxymethyl chitosan hydrogel loaded with long-term released hUCMSCs derived exosomes promotes skin wound repair. Int J Biol Macromol. 2024 Apr;265(Pt 1):130843.\\u003c/li\\u003e\\n\\u003cli\\u003eYang J, Chen Z, Pan D, Li H, Shen J. 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Exosomes Derived from Human Umbilical Cord Mesenchymal Stem Cells Accelerate Cutaneous Wound Healing by Enhancing Angiogenesis through Delivering Angiopoietin-2. Stem Cell Rev and Rep 17, 305\\u0026ndash;317 (2021).\\u003c/li\\u003e\\n\\u003c/ol\\u003e\"},{\"header\":\"Scheme 1\",\"content\":\"\\u003cp\\u003eScheme 1 is available in the Supplementary Files section.\\u003c/p\\u003e\"}],\"fulltextSource\":\"\",\"fullText\":\"\",\"funders\":[],\"hasAdminPriorityOnWorkflow\":false,\"hasManuscriptDocX\":true,\"hasOptedInToPreprint\":true,\"hasPassedJournalQc\":\"\",\"hasAnyPriority\":false,\"hideJournal\":false,\"highlight\":\"\",\"institution\":\"\",\"isAcceptedByJournal\":true,\"isAuthorSuppliedPdf\":false,\"isDeskRejected\":\"\",\"isHiddenFromSearch\":false,\"isInQc\":false,\"isInWorkflow\":false,\"isPdf\":false,\"isPdfUpToDate\":true,\"isWithdrawnOrRetracted\":false,\"journal\":{\"display\":true,\"email\":\"info@researchsquare.com\",\"identity\":\"journal-of-nanobiotechnology\",\"isNatureJournal\":false,\"hasQc\":true,\"allowDirectSubmit\":false,\"externalIdentity\":\"jnan\",\"sideBox\":\"Learn more about [Journal of Nanobiotechnology](http://jnanobiotechnology.biomedcentral.com)\",\"snPcode\":\"12951\",\"submissionUrl\":\"https://submission.nature.com/new-submission/12951/3\",\"title\":\"Journal of Nanobiotechnology\",\"twitterHandle\":\"@BioMedCentral\",\"acdcEnabled\":true,\"dfaEnabled\":true,\"editorialSystem\":\"em\",\"reportingPortfolio\":\"BMC/SO AJ\",\"inReviewEnabled\":true,\"inReviewRevisionsEnabled\":true},\"keywords\":\"Core-shell Microneedle, Diabetic wound healing, Exosomes, Ferroptosis, Antibacterial activity\",\"lastPublishedDoi\":\"10.21203/rs.3.rs-7689935/v1\",\"lastPublishedDoiUrl\":\"https://doi.org/10.21203/rs.3.rs-7689935/v1\",\"license\":{\"name\":\"CC BY 4.0\",\"url\":\"https://creativecommons.org/licenses/by/4.0/\"},\"manuscriptAbstract\":\"\\u003cp\\u003eChronic non-healing of diabetic wound (DW) remains a critical clinical challenge worldwide. Sustained oxidative stress and prolonged inflammatory responses disrupt the wound microenvironment, while bacterial colonization and biofilm formation on the wound bed compromise drug penetration, consequently leading to suboptimal outcomes with conventional approaches. Here, we developed a core-shell structured microneedle (MN) patch system, designated as MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e, to precisely regulate the DW microenvironment through sequential drug release. The MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e features a biphasic agent release profile, comprising a natural antimicrobial peptide (Pleurocidin) in the outer shell layer, that exerts anti-infective effects during the initial phase of wound healing, while engineered exosomes (Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e) in the core layer alleviate oxidative stress and modulate immune responses subsequently. This study details that the sustained release of Exos\\u003csup\\u003eQ10\\u003c/sup\\u003e effectively inhibits high glucose (HG)-induced ferroptosis in vitro, demonstrating potent antioxidant activity and anti-inflammatory capacity. Furthermore, in a \\u003cem\\u003eS. aureus\\u003c/em\\u003e-infected diabetic mouse wound model, MN@Ple/Exo\\u003csup\\u003eQ10\\u003c/sup\\u003e demonstrates potent antibacterial activity while mitigating oxidative stress, suppressing inflammation and promoting angiogenesis, thereby accelerating wound healing. Collectively, the developed spatiotemporally controlled MN system overcomes bacterial barriers and stabilizes exosomal delivery, enabling comprehensive regulation of the microenvironment in DWs. This breakthrough approach presents a novel and translational strategy for DW therapy.\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\\u003cp\\u003e\\u003c/p\\u003e\",\"manuscriptTitle\":\"A Core–Shell Microneedle Platform for the Spatiotemporal Codelivery of Dual-Agent Therapeutics Precisely Orchestrates Diabetic Wound Healing\",\"msid\":\"\",\"msnumber\":\"\",\"nonDraftVersions\":[{\"code\":1,\"date\":\"2025-10-09 13:23:19\",\"doi\":\"10.21203/rs.3.rs-7689935/v1\",\"editorialEvents\":[{\"type\":\"communityComments\",\"content\":0},{\"type\":\"decision\",\"content\":\"Revision requested\",\"date\":\"2025-10-27T14:35:05+00:00\",\"index\":\"\",\"fulltext\":\"\"},{\"type\":\"editorInvitedReview\",\"content\":\"\",\"date\":\"2025-10-27T08:06:21+00:00\",\"index\":\"hide\",\"fulltext\":\"\"},{\"type\":\"editorInvitedReview\",\"content\":\"\",\"date\":\"2025-10-22T14:10:50+00:00\",\"index\":\"hide\",\"fulltext\":\"\"},{\"type\":\"reviewerAgreed\",\"content\":\"96580160434057629566416112550177845668\",\"date\":\"2025-10-01T07:58:39+00:00\",\"index\":\"hide\",\"fulltext\":\"\"},{\"type\":\"reviewerAgreed\",\"content\":\"153829947757688043995268964290304566351\",\"date\":\"2025-09-29T00:43:57+00:00\",\"index\":\"hide\",\"fulltext\":\"\"},{\"type\":\"reviewerAgreed\",\"content\":\"95488578525072123282190897495416686565\",\"date\":\"2025-09-29T00:26:35+00:00\",\"index\":\"hide\",\"fulltext\":\"\"},{\"type\":\"reviewersInvited\",\"content\":\"\",\"date\":\"2025-09-28T15:40:40+00:00\",\"index\":\"\",\"fulltext\":\"\"},{\"type\":\"editorAssigned\",\"content\":\"\",\"date\":\"2025-09-24T04:06:08+00:00\",\"index\":\"\",\"fulltext\":\"\"},{\"type\":\"checksComplete\",\"content\":\"\",\"date\":\"2025-09-24T04:01:05+00:00\",\"index\":\"\",\"fulltext\":\"\"},{\"type\":\"submitted\",\"content\":\"Journal of Nanobiotechnology\",\"date\":\"2025-09-23T05:30:20+00:00\",\"index\":\"\",\"fulltext\":\"\"}],\"status\":\"published\",\"journal\":{\"display\":true,\"email\":\"info@researchsquare.com\",\"identity\":\"journal-of-nanobiotechnology\",\"isNatureJournal\":false,\"hasQc\":true,\"allowDirectSubmit\":false,\"externalIdentity\":\"jnan\",\"sideBox\":\"Learn more about [Journal of Nanobiotechnology](http://jnanobiotechnology.biomedcentral.com)\",\"snPcode\":\"12951\",\"submissionUrl\":\"https://submission.nature.com/new-submission/12951/3\",\"title\":\"Journal of Nanobiotechnology\",\"twitterHandle\":\"@BioMedCentral\",\"acdcEnabled\":true,\"dfaEnabled\":true,\"editorialSystem\":\"em\",\"reportingPortfolio\":\"BMC/SO AJ\",\"inReviewEnabled\":true,\"inReviewRevisionsEnabled\":true}}],\"origin\":\"\",\"ownerIdentity\":\"0d3a5b2d-f187-4120-a6d5-21b6a992971f\",\"owner\":[],\"postedDate\":\"October 9th, 2025\",\"published\":true,\"recentEditorialEvents\":[],\"rejectedJournal\":[],\"revision\":\"\",\"amendment\":\"\",\"status\":\"published-in-journal\",\"subjectAreas\":[],\"tags\":[],\"updatedAt\":\"2026-03-23T16:14:19+00:00\",\"versionOfRecord\":{\"articleIdentity\":\"rs-7689935\",\"link\":\"https://doi.org/10.1186/s12951-026-04155-1\",\"journal\":{\"identity\":\"journal-of-nanobiotechnology\",\"isVorOnly\":false,\"title\":\"Journal of Nanobiotechnology\"},\"publishedOn\":\"2026-03-17 15:58:33\",\"publishedOnDateReadable\":\"March 17th, 2026\"},\"versionCreatedAt\":\"2025-10-09 13:23:19\",\"video\":\"\",\"vorDoi\":\"10.1186/s12951-026-04155-1\",\"vorDoiUrl\":\"https://doi.org/10.1186/s12951-026-04155-1\",\"workflowStages\":[]},\"version\":\"v1\",\"identity\":\"rs-7689935\",\"journalConfig\":\"researchsquare\"},\"__N_SSP\":true},\"page\":\"/article/[identity]/[[...version]]\",\"query\":{\"redirect\":\"/article/rs-7689935\",\"identity\":\"rs-7689935\",\"version\":[\"v1\"]},\"buildId\":\"XKTyCvWXoU3ODBz1xrDgd\",\"isFallback\":false,\"isExperimentalCompile\":false,\"dynamicIds\":[84888],\"gssp\":true,\"scriptLoader\":[]}","source_license":"CC-BY-4.0","license_restricted":false}