Promoting Diabetic Wound Healing through a Hydrogel‐Based Cascade Regulation Strategy of Fibroblast‐Macrophage

Abstract

The management of diabetic wounds (DWs) continues to pose a significant challenge in the field of medicine. DWs are primarily prevented from healing due to damage to macrophage efferocytosis and fibroblast dysfunction. Consequently, a treatment strategy that involves both immunoregulation and the promotion of extracellular matrix (ECM) formation holds promise for healing DWs. Nevertheless, existing treatment methods necessitate complex interventions and are associated with increased costs, for example, the use of cytokines and cell therapy, both of which have limited effectiveness. In this study, a new type of ruthenium (IV) oxide nanoparticles (RNPs)‐laden hybrid hydrogel dressing with a double network of Pluronic F127 and F68 has been developed. Notably, the hybrid hydrogel demonstrates remarkable thermosensitivity, injectability, immunoregulatory characteristics, and healing capability. RNPs in hydrogel effectively regulate both fibroblasts and macrophages in a cascade manner, stimulating fibroblast differentiation while synergistically enhancing the efferocytosis of macrophage. The immunoregulatory character of the hydrogel aids in restoring the intrinsic stability of the immune microenvironment in the wound and facilitates essential remodeling of the ECM. This hydrogel therefore offers a novel approach for treating DWs through intercellular communication.

By regulating the immune system to create a suitable microenvironment for healing and enhancing the function of cells involved in wound healing, it becomes an ideal strategy for treating DWs. The hydrogel system promotes fibroblast differentiation to synergistically enhance the efferocytosis of macrophages. This effectively promotes the resolution of chronic inflammation in DWs, accelerating the healing process.

1. Introduction

≈1.3 billion people worldwide suffer from chronic wound problems related to diabetes, and the cost of wound repair accounts for ≈5% of total medical care expenses.^{[ 1} , ^{2 ]} With the continuous and rapid aging of the population, the incidence of DWs is expected to continue to rise.^{[ 3} , ^{4 ]} The main challenge in treating DWs is managing chronic inflammation.^{[ 4 ]} The persistence of apoptotic cells (ACs), particularly apoptotic neutrophils, at the injury site is the primary cause of chronic inflammation.^{[ 5} , ⁶ , ^{7 ]} However, existing wound protective dressings have limited therapeutic effects, often leading to frequent wound recurrence and a lack of response to changes in the wound microenvironment.^{[ 8} , ^{9 ]} There is still a clinical need for more effective methods to address chronic wounds in diabetes patients.

Among the various immune cells recruited to DWs, macrophages are particularly significant due to their abundance and primary role in chronic wound development.^{[ 10} , ¹¹ , ^{12 ]} The Tyro3‐Axl‐MerTK (TAM) receptor tyrosine kinase family in macrophages plays a critical role in maintaining homeostasis.^{[ 13} , ^{14 ]} Normally, macrophages can recognize phosphatidylserine expressed by ACs through TAM and remove it through efferocytosis, a phagocytic process proposed by Henson in 2003.^{[ 15 ]} Efferocytosis refers to the process through which engulfing cells (primarily macrophages) phagocytose and clear ACs undergoing programmed cell death.^{[ 16} , ¹⁷ , ^{18 ]} Additionally, macrophages involved in efferocytosis secrete more cytokines, such as TGF‐β and IL‐10, which promote inflammation resolution.^{[ 19} , ^{20 ]} Therefore, efferocytosis is crucial for preventing autoimmune diseases and inflammation. Nevertheless, studies have shown that the efferocytosis of macrophages in DWs is impaired, leading to abnormal accumulation and subsequent necrosis of apoptotic cells, which further promotes chronic and progressive inflammation.^{[ 21} , ^{22 ]} Macrophages exhibit a proinflammatory phenotype, aggravating inflammation maintenance, impairing wound healing, and increasing infection and amputation risks.

The crosstalk between fibroblast and macrophage presents a potential avenue for reactivating the efferocytosis of macrophages. Cancer‐associated fibroblasts play a regulatory role in promoting the repair phenotype of tumor‐associated macrophages.^{[ 23 ]} Fibroblasts can aid macrophages in the wound niche by releasing cytokines, such as monocyte colony‐stimulating factor (M‐CSF), during their differentiation, thereby promoting macrophage survival and proliferation.^{[ 24 ]} Moreover, fibroblasts play a central role in the wound healing process by reshaping the ECM and filling the wound through proliferation and differentiation, directly contributing to the repair process.^{[ 25 ]} However, in chronic wounds affecting diabetic patients, factors such as persistent inflammation and high oxidative stress in the pathological microenvironment severely impair the ability of fibroblasts to proliferate and differentiate.^{[ 26 ]} As a result, the formation of the ECM is insufficient, rendering it unable to provide the necessary initial barriers relied upon by immune cells, nerve cells, fibroblasts, endothelial cells, and keratinocytes.^{[ 27 ]} Therefore, it is imperative to explore therapeutic strategies that drive fibroblast differentiation to synergistically enhance macrophage efferocytosis.

Exposed wounds that are highly susceptible to infection and have irregular wound margins pose challenges in terms of wound protection and drug delivery.^{[ 28 ]} A hydrogel‐based sustained release system is an ideal choice for wound dressings.^{[ 29 ]} Additionally, the presence of scabbed tissue increases the need for efficient drug penetration, and drug stability becomes crucial in the microenvironment characterized by high oxidative stress and low pH in DWs.^{[ 30 ]} Previously, precious metal nanoparticles, such as Au and Ag nanoparticles, which possess unique physical and chemical properties, such as high permeability and stability, have garnered considerable interest in the biomedical field.^{[ 31 ]} However, researchers have given little attention to the biological potential of RNPs, one of the most stable precious metal oxides.^{[ 32} , ^{33 ]}

To address the aforementioned issues, we developed a dual‐network thermosensitive hydrogel release system containing RNPs, inspired by the strategy of crosstalk between fibroblast and macrophage (Scheme 1 ). Our results demonstrate that this system can enhance fibroblast differentiation, facilitate ECM reconstruction, accelerate inflammation regression, and promote DWs healing. In vitro studies have indicated that RNPs significantly stimulate fibroblasts proliferation and differentiation. Furthermore, the conditioned medium obtained from the coculture of RNPs with fibroblasts upregulated the expression of macrophage efferocytosis receptors, MerTK and TIM‐4, thereby enhancing macrophage efferocytosis. Intriguingly, direct treatment with RNPs alone did not yield similar results in terms of enhancing macrophage efferocytosis. Subsequently, RNA‐seq analysis revealed differential gene expression, and ELISA confirmed an increase in M‐CSF secretion after RNPs treatment, providing evidence for the crucial role of fibroblasts in ECM reconstruction and the activation of macrophage efferocytosis. This study proposes a cascade repair concept of immune remodeling‐ECM reconstruction that occurs through the healing initiation stage of DWs and provides new therapeutic ideas for the treatment of wounds.

Scheme 1.

Schematic diagram illustrating the preparation and regulatory mechanism of RHG. The prepared RNPs solution is mixed with the HG to form the RHG. Importantly, the RHG promotes the secretion of M‐CSF by fibroblasts through the release of RNPs, thus enhancing efferocytosis of macrophages and creating an immune microenvironment conducive to healing. Additionally, differentiated fibroblasts contribute to the reconstruction of the ECM, establishing a vital biological barrier essential for the regeneration process.

2. Results

2.1. Synthesis and Characterization of RNPs

RNPs were prepared using hydrothermal treatment as shown in Figure 1a. To clarify their microstructure, we conducted a transmission electron microscopy (TEM) analysis. TEM images and size distribution results depict the particle morphology of RNPs nanocrystals, with a particle size of 14.9 ± 2.3 nm (Figure 1b,c). A crystalline structure of RNPs with a lattice distance of 0.12 nm was shown in a high‐resolution TEM image (Figure 1b). Moreover, the dynamic light scattering result demonstrated a narrow peak distribution of the RNPs size (Figure S1, Supporting Information). The presence of functional groups on the surface of the RNPs was confirmed by Fourier transform infrared spectroscopy (FT‐IR) (Figure 1d). Furthermore, water‐dispersible RNPs exhibit significant absorption in aqueous solutions, ≈775 nm in the visible and near‐infrared regions (Figure S2, Supporting Information).

Figure 1.

Synthesis and characterizations of RNPs. a) Schematic diagram of the preparation process of RNPs. SC: sodium citrate; RNPs: RuO₂ nanoparticles. b) TEM of RNPs. Scale bar, 20 nm. Inset: HRTEM image. c) Diameter of RNPs. d) FT‐IR of RNPs. e) XPS survey spectrum of RNPs. f) XPS of Ru 3d spectrum of RNPs. g) XPS of O 1s spectrum of RNPs. h) SEM image and corresponding elemental mapping images of RNPs. Scale bar, 100 nm.

The crystal phase of the RNPs was further confirmed by X‐ray diffraction, and the pattern of the RNPs exhibited a number of well‐indexed peaks (JCPDS 43‐1027) (Figure S3, Supporting Information). Next, X‐ray photoelectron spectroscopy (XPS) measurements will be conducted to determine the surface chemical state of the RNPs. The measurement spectrum of the RNPs indicated the presence of Ru and O (Figure 1e). The Ru 3d and O 1s curves were fitted to the XPS data. A pair of peaks, located at 282.3 and 284.8 eV, was attributed to the corresponding 3d_5/2 and 3d_3/2 spin orbitals of Ru (IV) in the RNPs. The other two peaks, located at 283.8 and 286.3 eV, were attributed to Ru (III) in the hydrated Ru (III)‐OH species (Figure 1f). Additionally, the O1s energy level can be divided into three regions, including the regions with bond energies of 530.7, 532.3, and 535.4 eV (Figure 1g). These energies can be assigned to three different oxygen‐containing components in their chemical environment, specifically, O₂‐, OH‐, and H₂O. It is possible that these components are generated by lattice oxygen, five hydroxyl groups, and adsorbed water in the sample. The EDS elemental spectrum analysis shows a uniform distribution of these two elements (Figure 1h).

2.2. Preparation and Characterization of the RNPs‐Hydrogel (RHG)

As shown in Figure 2a, RHG was prepared and stored for further use by dispersing the previously prepared RNPs into the preprepared Pluronic F127 and F68 hydrogel (HG). At room temperature (25 °C), the HG and RHG maintain a high flow dynamic with irregular wound coverage. However, under the body temperature (37 °C), the HG and RHG undergo a phase transition to a condensed state, which is completed within 2 min (Figure 2b). The words “CMU” written with the RHG ink displayed exceptional injectability and fluidity, indicating that RGH has the capability to effectively cover irregular or deep wounds (Figure 2c). Next, the chemical structure of the RNPs before and after loading was detected by FT‐IR. The spectrum of the RHG shows that the peak at 1581 cm⁻¹ belongs to the C═O bond of the RNPs, confirming the grafting of the RNPs (Figure S4, Supporting Information). RHG exhibited a low swelling ratio consistent with that of HG, indicating good internal crosslinking density of the hydrogels (Figure S5, Supporting Information). However, the dehydration rate experiment showed that the dehydration rate of the RGH at a constant temperature of 37 °C was slightly greater than that of the HG (Figure S6, Supporting Information). The results of the drug sustained‐release experiment showed sustained release of the RNPs to ensure long‐term pharmacological effects (Figure S7, Supporting Information). It is worth noting that the sustained release of RHG occurs under the conditions of a PBS buffer solution, indicating that the sustained release effect of RHG can be achieved in the bodily fluid environment.

Figure 2.

Fabrication and characterizations of RHG. a) Schematic illustration of fabricating RHG. b) Images of HG/RHG transition triggered by temperature change. c) Images of RHG injectability. d) The curve of storage modulus and loss modulus as functions of the temperature. e) Angular frequency (γ = 1.0%) of hydrogels versus shear rate. f) The time‐dependent step strain rheological experiments of hydrogels. g) The G′ and G″ of the hydrogels from strain amplitude sweep (γ = 1.0–1000.0%) at a fixed angular frequency (ε = 10.0 rad s⁻¹). h) Viscosity of the hydrogels versus temperature.

To verify the mechanical properties of the hydrogels, we used a parallel plate rheometer to conduct rheological analysis at different frequencies and strains. We performed dynamic rheological characterization on hydrogels with and without RNPs. First, we tested the elastic modulus and viscous modulus of two hydrogels precursor solutions using temperature scanning. For the HG system, at temperatures below 23 °C, G“ is lower than G', indicating that the sample is in a liquid state. The gel temperature of the RHG system is ≈31 °C, meaning that it remains in a liquid state suitable for injection at 25 °C and transitions to a gel‐like state at 37 °C (Figure 2d). The results of dynamic strain scanning show that the hydrogel has a wide linear viscoelastic region at low strains. With increasing strain force, the G' and G” of the HG remained unchanged, while the RHG slightly decreased, indicating that the RHG exhibited shear thinning properties (Figure 2e). Based on the strain amplitude scanning test, we conducted continuous step strain scanning to evaluate the self‐healing performance of the HG and RHG. Under alternating high and low strains for three cycles, the modulus of the hydrogels recovered within 60 s without any loss (Figure 2f). This can be attributed to the dynamic network of the hydrogels, which exhibits good injectability and self‐healing performance. Furthermore, the viscosity of the hydrogels decreased with increasing shear rate (Figure 2g). The shear thinning characteristic is desirable for injectable systems and aligns with the previously mentioned modulus results. Additionally, the viscosity of both types of hydrogels significantly decreased with increasing temperature while maintaining relatively stable adhesion at body temperature (Figure 2h). These results indicate that the RHG can rapidly gel via a temperature response, forming viscoelastic hydrogels with strong shape adaptability. This property can provide more effective protection and integration for irregular wounds.

2.3. Biocompatibility and Cell Proliferation‐Promoting Capability of the RNPs

Good biocompatibility is a fundamental requirement for the clinical use of hydrogel dressings. Since wound dressings inevitably contact blood, we first conducted a hemolysis test to further evaluate the blood compatibility of the hydrogels. The HG and RHG were separately incubated with blood, and the hemolysis rate for both hydrogels was less than 5% (Figure S8a,b, Supporting Information), indicating that the possibility of hemolysis for both hydrogels was low. The cell counting kit (CCK‐8) and live/dead staining assays are considered the gold standards for evaluating biocompatibility in vitro. After coculturing fibroblasts with different concentrations of RNPs for 24 h, we performed a CCK‐8 assay. As shown in Figure S9, Supporting Information, compared with that in the control group, the proliferation of the cells significantly increased in a concentration‐dependent manner, indicating that fibroblast proliferation increased during RNPs incubation. Next, after the same treatment as the CCK‐8 assay, the staining results of live/dead cells showed that fibroblasts exhibited more intense green fluorescence (live cells) after treatment with different concentrations of RNPs than did the control group. Additionally, the presence of red fluorescence (dead cells) decreased sequentially with increasing RNPs concentration (Figure S10, Supporting Information). To further investigate the intrinsic genetic regulation of cell proliferation, we conducted corresponding 5‐ethynyl‐2′‐deoxyuridine (EdU) staining. The results demonstrated that different concentrations of RNPs increased the proportion of positive cells compared to that in the control group, indicating enhanced DNA replication activity and active proliferation of fibroblasts (Figure S11, Supporting Information). Given the potential challenges in clinical applications due to the systemic toxicity caused by the in vivo retention effect of nanomaterials, major organs were collected from diabetic mice at the end of treatment for hematoxylin and eosin (H&E) staining. Figure S12, Supporting Information demonstrates that there was no evident tissue damage in the organs of the RHG group in comparison to those of the untreated group. Thus, the RNPs display remarkable biocompatibility with enhanced fibroblast proliferation, making them highly promising for use as a wound repair system for individuals with diabetes.

2.4. RNPs Promote the Formation of ECM In Vivo

The ECM plays a crucial role in wound healing by regulating cell growth and phenotype, providing both structural and mechanical support in addition to providing biochemical signals for tissue development.^{[ 34 ]} As demonstrated in Figure 3k, the correlation between the TGF‐β1 pathway and ECM formation was the strongest. Consequently, the western blotting results revealed significant increases in the levels of TGF‐β, α‐SMA, COL I, and COL III as the concentration of RNPs increased (Figure 3f). Based on the protein expression results above, the corresponding quantitative analysis confirms the findings (Figure 3g–j). Similarly, the immunofluorescence results demonstrated that the fluorescence intensity of TGF‐β, α‐SMA, COL I, and COL III increased with increasing RNPs concentration (Figure 3a). The semiquantitative analysis of fluorescence intensity further supported these experimental findings (Figure 3b–e). These results suggest that RNPs activate the TGF‐β1 signaling pathway and promote the formation of the ECM in vitro. Taken together, these findings indicate that RNPs contribute to the formation of the ECM and facilitate wound closure.

Figure 3.

RNPs promote ECM formation in vitro. a) Representative fluorescence images of the fibroblasts after different RNPs treatments (green: COL I, TGF‐β, α‐SMA; red: COL III; blue: nucleus. Scale bar, 20 µm. b–e) Immunofluorescence quantitative analysis of TGF‐β, α‐SMA, COL I, and COL III in fibroblasts after different RNPs treatments (n = 5). f) Western blotting analysis showing the levels of TGF‐β, α‐SMA, COL I, and COL III after different RNPs treatments. g–j) The corresponding quantitative analysis of TGF‐β, α‐SMA, COL I, and COL III proteins expression (n = 3). k) Schematic diagram illustrating RNPs activate the TGF‐β1 pathway in fibroblasts to enhance the formation of the ECM. Data are expressed as mean ± SD. The significant difference was detected by one‐way ANOVA. Compared to control, *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

2.5. RHG Imparts Tissue Regeneration Properties

In vitro studies have indicated that RNPs show promise as potential drugs for treating DWs. Consequently, a diabetic mouse model was used to assess the ability of the RHG to regenerate tissue. C57BL/6J mice were injected with streptozotocin (STZ), which led to consistently elevated blood glucose levels exceeding 16.7 mmol L⁻¹. A full‐thickness wound with an 8 mm diameter was created on the back. The progress of wound healing was assessed by observation and measurement on days 0, 3, 6, and 12 (Figure 4a). Overall, visual examination revealed that HG and two concentrations of RHG (5 and 10 mg mL⁻¹) could expedite wound healing, with over 50% wound closure achieved by the sixth day. Notably, RHG (10 mg mL⁻¹) exhibited the highest efficacy in promoting healing (Figure 4b). By day 12, RHG (5 mg mL⁻¹ and 10 mg mL⁻¹) had achieved greater than 90% wound closure, while 17% and 26% of wounds remained unhealed in the HG group and control group, respectively (Figure 4c,d). Histological analysis of granulation tissue formation and tissue maturation 12 days postinjury was conducted by assessing the thickness of granulation tissue and collagen deposition. The results of the histological analysis indicated that the RHG group exhibited a significantly greater thickness of granulation tissue following treatment than did the other groups (p < 0.001) (Figure 4e,f). Analysis of Massion staining at the wound site revealed that the RHG group had the highest level of collagen deposition (p < 0.001) (Figure 4g,h), suggesting that the recovery of damaged tissue in the wound after treatment was most advanced in the RHG group. To further clarify the difference in efficacy between RHG and existing commercial dressings in the treatment of DWs, we selected Alginate Tegaderm dressing (Alg) as the positive control. After 12 days of treatment, the wounds in both the Alg group and RHG group were nearly completely closed, in comparison to the control group (Figure S13a, Supporting Information). By analyzing the wound healing dynamic demonstration chart and wound area change curve, we observed that the wounds in the RHG group healed faster following treatment, compared to the Alg group (Figure S13b,c, Supporting Information). Additionally, further examination using H&E and Masson staining showed that the granulation tissue thickness in the RHG group was greater and collagen deposition was significantly improved compared to the Alg group (Figure S13d–g, Supporting Information). These results suggest that RHG has great potential for clinical translation and promising market prospects.

Figure 4.

RHG promoted diabetic wound repair and regeneration in vivo. a) Schematic diagram of experimental design of C57BL/6J diabetic mouse wound model. b) Representative digital images of wound areas in response to different treatments were taken on days 0, 3, 6, and 12 (n = 5). Wound diameter = 8 mm. c) The contour map of the wound healing process. d) Quantitative analysis of wound area for each group (n = 5). e) The H&E staining of the wound area reveals the granulation tissue on day 12. f) The thickness of the granulation tissue is indicated by the blue arrow (n = 5). Scale bar, 400 µm. g) The Masson staining of the wound area indicates the deposition of collagen on day 12. Scale bar, 400 µm. h) Quantitative analysis of collagen deposition in the wound area (n = 5). Data are expressed as mean ± SD. The significant difference was detected by one‐way ANOVA. Compared to control, *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

Considering the variations in collagen deposition among the different groups, we further investigated the relationship between collagen deposition and TGF‐β1 signal activation. The results of immunofluorescence staining of tissue slices after treatment confirmed that TGF‐β1, along with its downstream proteins TGF‐β, COL I, and COL III, was significantly increased in the RHG group compared to the other groups (Figure 5a,b,d–f). The immunohistochemical results for α‐SMA were consistent with the trend observed by immunofluorescence (Figure S14a,b, Supporting Information), which was also in line with the in vitro cytological results. Additionally, the COL III/COL I ratio was higher in the RHG (10 mg kg⁻¹) group compared to the other groups (Figure S15, Supporting Information). This suggests that COL III plays a role in facilitating efficient wound healing, while RHG (10 mg kg⁻¹) has the most favorable effect in promoting the formation of COL III. The Ki67 histochemical staining results demonstrated that compared with the control group and the HG group, the RHG treatment group had greater proliferative activity in the wound tissue, and the RHG (10 mg kg⁻¹) group exhibited greater proliferative activity than the RHG (5 mg kg⁻¹) group (Figure 5c–g). The aforementioned results indicate that RHG effectively enhances local ECM formation in wounds by stimulating fibroblast proliferation and differentiation in vivo (Figure 5h).

Figure 5.

RHG promoted ECM formation in vivo. a,b) Representative images of TGF‐β, COL I, and COL III immunofluorescence staining on day 12 (green: TGF‐β, COL III; red: COL I; blue: nucleus). Scale bar, 100 µm. c) Ki67 immunohistochemical staining results of wound tissue on day 12. Scale bar, 50 µm. d–f) Immunofluorescence quantitative analysis of TGF‐β, COL I, and COL III in the wound tissue (n = 5). g) Quantitative analysis of Ki67⁺ cells in tissue slices obtained from different treatment groups (n = 5). h) Schematic diagram of RHG promoting ECM formation in vivo. Data are expressed as mean ± SD. The significant difference was detected by one‐way ANOVA. *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

2.6. RHG Promotes the Resolution of Inflammation by Enhancing Macrophage Efferocytosis

The dynamic transition of the immune microenvironment is necessary for wound repair and regeneration after injury, as it shifts from a pro‐inflammatory microenvironment (which neutralizes damage and clears ACs or injured tissue) to an anti‐inflammatory microenvironment.^{[ 19 ]} Efferocytosis plays a crucial role in resolving inflammation and restoring tissue homeostasis.^{[ 21 ]} Therefore, the initial step should be determining whether the RHG mediates the process of inflammation resolution and efferocytosis, considering its effective ability to promote rapid wound healing. Subsequently, we conducted immunofluorescence staining on macrophage phenotypes in wound slices during the proliferation phase (day 6). As shown in Figure 6a, compared to the other groups, the RHG group had significantly lower levels of iNOS, a positive marker of pro‐inflammatory macrophages, while the proportion and distribution of CD206, a positive marker of anti‐inflammatory macrophages, increased. The quantitative analysis of iNOS and CD206 further demonstrated that RHG promoted inflammation resolution (Figure 6b,c). We aimed to determine whether the regression of inflammation within DWs is linked to the efferocytosis of macrophages. We further examined the effect of the RNPs on the efferocytosis of macrophages in vitro. Surprisingly, treatment with RNPs did not influence the efferocytosis index of macrophages toward apoptotic cells or the expression of the efferocytosis receptors MerTK and TIM‐4 (Figure 6f,g).

Figure 6.

RHG promotes macrophage transition to anti‐inflammatory phenotype by enhancing efferocytosis. a) Representative images of iNOS and CD206 immunofluorescence staining on day 6 are shown (red: iNOS; green: CD206; blue: nucleus). Scale bar, 100 µm. b,c) Statistical data of the percentage of iNOS⁺ and CD206⁺ macrophages in tissue (n = 5). d) Illustration of the enhanced macrophage efferocytosis in vitro by conditional culture medium treated with RNPs. e) Representative images of macrophages performing efferocytosis. CM: conditional medium collected from L‐929; RNPs‐CM: conditional medium collected from L‐929 treated with RNPs for 24 h. Green: macrophages; red: ACs.The yellow arrowheads indicate the efferocytosis effect of macrophages on ACs. Scale bar, 25 µm. f) The macrophage efferocytosis index was assessed post‐treatment with various groups (n = 5). g) Western blotting analysis showing the levels of TIM‐4, MerTK, iNOS, and CD206. h,i) The flow cytometer is used to verify the results and analyze the efficiency of macrophage efferocytosis after various treatment groups (n = 3). Note: ACs:Mø = 10:1; RNPs, CM, and RNPs‐CM were used to pretreat macrophages for 2 h. Data are expressed as mean ± SD. The significant difference was detected by one‐way ANOVA. *p < 0.05, **p < 0.01, ***p < 0.001, and ****p < 0.0001.

Although the role of fibroblasts in maintaining macrophage's function is crucial, their specific contribution to macrophage efferocytosis remains uncertain. According to our previous findings, we hypothesize that RNPs may have immunomodulatory effects through an indirect pathway, enhancing the communication between fibroblast and macrophage. Thus, we aimed to clarify the impact of crosstalk between fibroblast and macrophage on the latter's efferocytosis. We collected fibroblasts (L‐929) culture medium (CM) and treated fibroblasts cultured with RNPs (RNPs‐CM) to cultivate macrophages (RAW264.7) for efferocytosis detection (Figure 6d). The confocal laser scanning microscope (CLSM) images visually demonstrated that, compared to the control group, the CM group and RNPs‐CM group exhibited greater efficiency in clearing ACs (Figure 6e). According to the statistical analysis, the RNPs‐CM group displayed the greatest efferocytosis efficiency among the groups (Figure 6f). Furthermore, flow cytometry validation revealed that compared to the control group (0.54%), RNPs group (0.37%), and CM group (2.0%), the RNPs‐CM group exhibited the highest efferocytosis rate (11.1%) (Figure 6h,i). Western blotting results showed that the upregulation of the efferocytosis receptors MerTK and TIM‐4 in the RNPs‐CM group was more significant than that in the other groups (Figure 6g). After treatment with RNPs‐CM, increased expression of efferocytosis receptors further supports the enhancement of macrophage efferocytosis (Figure S16, Supporting Information). Additionally, the western blotting results revealed that under the influence of RNPs‐CM, the pro‐inflammatory macrophages stimulated with LPS downregulated the expression of iNOS but significantly upregulated the expression of CD206 (Figure S17, Supporting Information). This indicates a phenotypic shift of macrophages toward an anti‐inflammatory phenotype, which signifies the resolution of inflammation. The above results indicate that the communication between fibroblast and macrophage plays a role in enhancing the efferocytosis of macrophages. In particular, RNPs further enhance cell‒cell communication, thereby improving the function of macrophage efferocytosis and effectively mediating the resolution of inflammation in DWs.

2.7. M‐CSF Is the Intrinsic Mechanism through Which the RNPs‐Fibroblast Pathway Enhances Macrophage Efferocytosis

The cell behavior of fibroblast promoting macrophage efferocytosis encourages us to explore the mechanism of the RNPs enhancing the communication between the fibroblast and macrophage. We then conducted a transcriptional analysis to reveal the mRNA variations in fibroblasts treated with the RNPs, to gain in‐depth understanding of the molecular mechanism underlying the enhancement of fibroblast‐macrophage crosstalk by the RNPs. The heat map reveals a notable disparity in transcriptome between the treatment group and the control group, indicating that the fibroblast population exposed to RNPs possesses distinctive expression traits (Figure S18, Supporting Information). We performed gene ontology (GO) analysis, including molecular function, biological process, and cellular component, to determine the biological functions of the RNPs treatment. The information obtained from the above results indicated that differentially expressed genes induced by the RNPs are primarily enriched in the ECM‐related pathways (Figure S19, Supporting Information). The treatment with the RNPs induced a total of 638 differentially expressed genes, including 326 upregulated mRNAs and 312 downregulated mRNAs (Figure 7a). Compared to the control group, the RNPs resulting in differential expression of 2524 genes (8.95%), 22 142 genes (78.51%) exhibit similar expression dynamics (Figure 7b).

Figure 7.

a) The volcano plot reveals the downregulation or upregulation of genes caused by RNPs compared to the control group. b) Heat map of upregulated and downregulated genes caused by RNPs (top 50, p < 0.05). c) Venn diagram for transcriptome analysis. d) ELISA analysis of the content of M‐CSF in the cell supernatant after fibroblasts treated with RNPs. e) Western blotting analysis showing the levels of MerTK and TIM‐4 under different concentrations of M‐CSF. f) Representative images of macrophages performing efferocytosis. Green: macrophages; red: ACs. The yellow arrowheads indicate the efferocytosis effect of macrophages on ACs. Scale bar, 25 µm. g) Diagram depicting the role of RNPs in promoting fibroblast differentiation and facilitating the release of M‐CSF to enhance macrophage efferocytosis function. Data are expressed as mean ± SD. The significant difference was detected by one‐way ANOVA. *p < 0.05.

To investigate the key pathways mediated by the RNPs in fibroblast‐macrophage interaction, we focused on the top 50 upregulated genes in RNPs group (Figure 7c). Among them, the gene that caught our attention is CSF1, which ranks third. It is well‐known that cytokines serve as messengers for intercellular communication, and CSF1 is the encoding gene of the important cytokine M‐CSF that maintains the macrophage niche.^{[ 35 ]} Therefore, we further verified the production of M‐CSF through ELISA, as shown in Figure 7d, which demonstrated a significant increase in the levels of M‐CSF in the cell supernatant of fibroblasts treated with the RNPs. Previous reports have indicated that M‐CSF has clear implications for macrophage proliferation and polarization, but it remains unclear if it plays a role in regulating the function of efferocytosis. Therefore, we experimentally added M‐CSF to macrophages and detected the expression of MerTK and TIM‐4 through western blotting. The results showed that M‐CSF can promote the expression of MerTK and TIM‐4 (Figure 7e and Figure S20, Supporting Information). Simultaneously, the efficiency of macrophages in clearing ACs was evaluated through the efferocytosis assay. The results demonstrated that exposure to M‐CSF significantly improved the macrophages' ability to clear ACs, suggesting that M‐CSF enhances the efferocytosis function of macrophages (Figure 7f). As shown in Figure 7g, the aforementioned results indicate that the RNPs have the following biological functions: promoting fibroblast differentiation, increasing M‐CSF secretion, and enhancing macrophage efferocytosis. Meanwhile, the M‐CSF is an intrinsic factor for the RNPs to regulate the local immunity in a cascading manner through the fibroblast‐macrophage pathway.

3. Discussion

The presence of chronic inflammation is a critical issue that results in non‐healing wounds.^{[ 36 ]} Macrophages have received significant attention from researchers due to their crucial role in both pathological condition and tissue homeostasis, particularly in alleviating persistent inflammation and facilitating the transition of non‐healing wounds.^{[ 37} , ^{38 ]} Among the various strategies employed, macrophage phenotype reprogramming is considered one of the important approaches. By directly altering the macrophages' phenotype and promoting the conversion from pro‐inflammatory to anti‐inflammatory, inflammation can be alleviated to some extent.

Nevertheless, we must not overlook the high plasticity exhibited by macrophages. If stimulus factors that induce the pro‐inflammatory phenotype persist, for instance, damage‐associated molecular patterns are produced through the secondary necrosis of dying cells in the wound, macrophage will revert from anti‐inflammatory to pro‐inflammatory state.^{[ 36} , ^{39 ]} To completely eliminate the inflammatory stimuli factors, we have designed and prepared a hydrogel system containing RNPs with dual network of Pluronic F127 and F68. This hydrogel system acts likely as a chemical catalyst to enhance cell‐cell communication between fibroblast and macrophage, promoting the efferocytosis function of macrophage, which plays a crucial role in tissue homeostasis recovery, tissue repair, and reconstruction. This process helps prevent inflammation by identifying and clearing ACs. Furthermore, it also secretes various factors such as PGE2 and TGF‐β, which effectively alleviate tissue damage. Therefore, the hydrogel enhances macrophage efferocytosis through the fibroblast‐macrophage pathway. This, in turn, helps remove ACs from the wound, reshapes immune homeostasis in damaged areas, and establishes a foundation for accelerating the processes of wound healing and repair.

Fibroblasts are the most important pioneer cell in the process of wound repair, responsible for the reconstruction of ECM and recruitment of various functional cells.^{[ 40 ]} However, the functionality of fibroblasts in DWs is severely impaired, leading to insufficient formation of ECM and the lack of innate barriers necessary for wound healing.^{[ 41 ]} In this study, we confirmed that the RNPs‐laden hydrogel could promote fibroblast differentiation and ECM formation, and the excellent biological function of RNPs in promoting ECM remodeling has been clarified. Moreover, we assessed the tissue regeneration capability of the RHG system through the establishment of a mouse DWs model. Encouragingly, the RHG (10 mg kg⁻¹) group exhibited the most prominent tissue regeneration capability, compared to the control, HG, and RHG (5 mg kg⁻¹) groups. Further histopathological analyses have clearly demonstrated that the RHG (10 mg kg⁻¹) performed outstanding ECM remodeling roles in the DWs. Although neither concentration 5 mg kg⁻¹ nor 10 mg mL⁻¹ of RHG treatment showed any toxic effects on the vital organs of the mice in the H&E staining, the higher concentration of RNPs in RHG was more effective in promoting wound healing. Therefore, RHG (10 mg kg⁻¹) also has greater application value and translational potential.

In the research field of DWs healing, previous studies have focused more on the direct repair effect of fibroblasts through the formation of ECM or the direct regulation of tissue repair and regeneration in wounds through growth factors derived from the fibroblasts (such as TGF‐β).^{[ 42} , ^{43 ]} Noteworthy, in the treatment of diabetic foot or non‐healing wounds, more attention should be paid to the potential immunomodulatory effects of the fibroblasts. In this study, the RNPs activated the TGF‐β1 pathway in the fibroblasts, promoting the expression of ECM‐related proteins, which is crucial for the process of wound contraction until complete closure. Simultaneously, the fibroblasts released M‐CSF during the differentiation, which acts on macrophages in the niche, causing the significant upregulation of MerTK and TIM‐4, the efferocytosis receptors on macrophages. The high expression of TIM‐4 on macrophages can bind to PtdSer receptors on the surface of ACs, anchoring them to the macrophages and then signaling to MerTK to enhance efferocytosis, thereby promoting the clearance of accumulated ACs in the wound area and avoiding prolonged inflammation. However, the macrophages that were directly treated with RNPs did not exhibit the aforementioned effects. Thus, the RHG achieved cascade regulation in the fibroblasts and macrophages, promoting ECM formation, while reshaping immune homeostasis, synergistically overcoming the two key issues that hinder DWs healing. Therefore, the RHG provides a novel and comprehensive solution for the treatment of DWs.

4. Conclusions

A dual‐network thermosensitive hydrogel was prepared that is capable of achieving cascade regulation of fibroblast and macrophage. It demonstrates excellent capabilities in promoting ECM regeneration and reshaping the immune microenvironment for DWs repair. The dual‐network thermosensitive hydrogel, composed of Pluronic F127, F68, and RNPs, undergoes gelation within 2 min when exposed to body temperature. Additionally, the hydrogel exhibits remarkable compatibility and promotes cell differentiation by activating the TGF‐β1 pathway. Most importantly, the hydrogel promotes fibroblast differentiation and synergistically enhances the efferocytosis of macrophages. The hydrogel restores the efferocytosis of macrophages by promoting the secretion of M‐CSF from fibroblasts, upregulating the expression of MerTK and TIM‐4 receptors related to efferocytosis, and enhancing the capability of macrophages to eliminate ACs. This results in the ultimate restoration of immune homeostasis in the DWs. Therefore, the RNPs‐laden Pluronic F127 and F68 hydrogel is a very good candidate for use as a wound dressing in DWs treatments, with great commercial potential.

5. Experimental Section

Chemicals and Reagents

Pluronic F‐127, F‐68, and RuCl₃ • xH₂O were purchased from Aladdin (Shanghai, China). The Alg was purchased from Minnesota Mining Manufacturing (3 m) (St. Paul, Minnesota, USA). The STZ was purchased from Biosharp (Shanghai, China). The calcein‐AM/propidium iodide double stain kit was purchased from Yeasen (Shanghai, China). The BCA protein assay kit was obtained from TransGen (Beijing, China). The polyvinylidene difluoride (PVDF) membrane was purchased from Merck (Darmstadt, Germany). The recombinant protein M‐CSF was obtained from MCE (MCEMerced, CA, USA). The EdU assay kit was obtained from Meilunbio (Dalian, Liaoning, China). The RIPA lysis buffer and H&E staining kit were purchased from Beyotime Biotech (Nantong, Jiangsu, China). Antibodies against TGF‐β, COL I, COL III, α‐SMA, iNOS, CD206, and Actin were obtained from Affinity (Affinity Bioscience, OH, USA); MerTK, and TIM‐4 were obtained from Cell Signaling Technology (Beverly, MA, USA). Superenhanced chemiluminescence (ECL) detection reagents were purchased from Yeasen (Shanghai, China).

RNPs Preparation

RNPs (hydrous RuO₂ nanoparticles) were prepared based on previous reports. 105.0 mg of RuCl₃ • xH₂O was added to a PTFE‐lined high‐pressure vessel (capacity 60.0 mL), with 25.0 mL of water at a concentration of 4.2 mg mL⁻¹ added and stirred for 30.0 min. Then, 160.0 mg of sodium citrate was added to the solution of ruthenium chloride and stirred until fully dissolved. Then the high‐pressure vessel was sealed with a stainless‐steel shell and heated in an oven to 210.0 °C for 12.0 h. After cooling to room temperature, RuO₂ • xH₂O deposited at the bottom. The product was purified by centrifugation (14 000 rpm, 20.0 min) and then washed with deionized water until the pH of the solvent reached 7.4. The repeat procedure was performed to purify the product. The final nanoparticles were dispersed in 20.0 mL of water under ultrasonic treatment. The collected nanoparticles were dried at 60.0 °C for 12.0 h to obtain the final RNPs.

RHG Preparation

To prepare RHG, F127 and F68 were mixed in a mass ratio of 18:5, and fully dissolved in water at room temperature. The mixture was then swollen in water at 4.0 °C for 12.0 h, resulting in HG. RNPs were added to HG according to the predetermined ratio and thoroughly stirred to evenly disperse the RNPs, resulting in RHG. The hydrogel transformation was completed by raising the temperature to 37.0 °C.

Characterization of RNPs

The Talos F200S G2 S/TEM electron microscopy was used to obtain scanning electron microscopy (SEM)/TEM images. XPS measurements were conducted using the Thermo Scientific K‐Alpha surface analysis system. X‐ray diffraction was performed with a Rigaku ULTIMA IV X‐ray diffractometer. FT‐IR was carried out on the Nexus 470 spectrometer.

Rheological Studies

The rheological properties of the hydrogel were analyzed using the DHR‐3 rheometer (TA Instruments, DE, USA). The experiments were conducted with 40.0 mm parallel plates, with a 1.0 mm gap between the plates. The viscoelasticity of HG/RHG was tested immediately after gelation. Frequency scanning was performed at an oscillation frequency ranging from 0.1 to 100.0 rad s⁻¹, with a strain level of 1.0%. For strain scanning, the oscillation frequency was fixed at 10.0 rad s⁻¹, while the applied strain varied from 1.0 to 1000.0%. To observe the damage healing characteristics of the hydrogel, the storage modulus (G′) and loss modulus (G″) were measured at a frequency of 10.0 rad s⁻¹, by continuously varying the strain between high strain (200.0%, 500.0%) and 1.0%.

Cell Culture

Cell line culture and in vitro studies were conducted using Raw 264.7, L‐929, and Jurkat Clone E6‐1 cells obtained from ATCC. The cells were cultured in RPMI 1640/DMEM (Hyclone, USA) supplemented with 10.0% FBS (Hyclone, USA) and 1.0% penicillin/streptomycin (Sigma, USA).

Live/dead Assay

The cytotoxicity of RNPs was evaluated using a live/dead assay. 1.0 × 10⁴ fibroblasts per well were placed into a 24‐well plate. The cells were allowed to fully adhere for 12.0 h. Different concentrations of RNPs (100, 200, 300, and 400 µg mL⁻¹) were mixed in the corresponding wells and incubated with the cells for 24.0 h. The control group consisted of untreated cells. The culture medium was removed and gently washed with PBS. The fibroblasts were stained with the live/dead detection kit following the protocol. The cells were observed and imaged under a laser confocal microscope (DIM8, Leica, Germany).

EdU Assay

According to the manufacturer's instructions, cell proliferation was evaluated using the EdU assay kit. Different concentrations of RNPs were co‐cultured with fibroblasts for 24.0 h, while the blank group consisted of untreated cells. The cells were then treated with 50.0 µm EdU and incubated for 2.0 h at 37 °C, after which they were fixed in 4.0% paraformaldehyde. After treating the cells with 0.5% Triton X‐100 for 10.0 min, they were mixed with 1× Apollo reaction cocktail and incubated for 30.0 min. Subsequently, the cell nuclei were stained with 100.0 µL of DAPI in the dark for 20.0 min. The cell proliferation was observed under a CLSM, and the proportion of EdU‐labeled cells to unlabeled cells was counted and calculated. All experiments were performed with three samples to obtain the average value.

Efferocytosis Assays

Jurkat cells were suspended in RPMI containing 5.0% fetal bovine serum and treated with ultraviolet C irradiation at a dose of 150.0 mJ cm⁻². The cells were then incubated for 4.0 h to induce apoptosis. To conduct the efferocytosis assays, mouse RAW264.7 cells were stained with PKH67, while apoptotic cells were stained with PI. Both staining was performed under dark conditions. The co‐culturing was done at a ratio of 1:10 (macrophages: ACs) for a duration of 24.0 h. Efferocytosis was evaluated using CLSM and flow cytometry. To assess the efferocytosis efficiency of macrophages toward ACs, flow cytometry analysis was utilized with the application of PI to label the ACs. Subsequently, these labeled ACs were co‐cultured with macrophages for a 24.0 h period. Following the co‐culture, any remaining ACs that were not engulfed by the macrophages were thoroughly eliminated by washing with PBS three times. The analysis of these washed ACs was carried out on a flow cytometer. The resulting data was then analyzed using FlowJo (Ashland, Oregon, USA).

Western Blotting Analysis

With a concentration of 2 × 10⁵ per well to be inoculated in the 6‐well plate. After treatment, total protein extracts were obtained from the cells using the Pierce, the BCA protein assay kit, and the concentration of isolated proteins was measured. The sample with an equal amount of total protein underwent thermal denaturation and was then subjected to electrophoresis on a 4–20% SDS‐PAGE gel. The separated protein was transferred to a PVDF membrane. The membrane was incubated at room temperature for 2.0 h in tris‐buffered saline with 5.0% skimmed milk and Tween 20. The expression of TGF‐β, COL I, COL III, α‐SMA, iNOS, CD206, MerTK, TIM‐4, and Actin was detected using a second antibody coupled with horseradish peroxidase and an ECL detection system.

DWs Healing Evaluation

The DWs healing was evaluated in C57BL/6J mice weighting 20–25 g. This plan has been approved by the Experimental Animal Ethics Committee of Zhejiang Cancer Hospital (IACUC) (plan number: 2023‐12‐009). Male mice were administered an intraperitoneal injection of 50.0 mg kg⁻¹ STZ, once a day for 5.0 consecutive days. Mice whose blood sugar levels exceeded 16.7 mmol were deemed to have successfully developed diabetes. The diabetic mice were then anesthetized, their back hair shaved and sterilized, and a round full‐thickness skin wound with a diameter of 8.0 mm was created on their backs using a biopsy punch.

Hemolysis Test

To conduct the hemolysis test, mouse blood was centrifuged at 116.0 × g for 10.0 min to obtain red blood cells. The red blood cells were then washed three times with PBS. The purified red blood cell concentration was diluted to 5.0% (v/v). After allowing HG/RHG to gel at 37 °C, 500.0 µL of the red blood cell suspension was added to a 1.5 mL test tube and gently mixed by pipetting. The sample was then incubated at 37.0 °C for 1.0 h and subsequently released. The clear supernatant was measured using a microplate reader at 540.0 nm. Triton X‐100 (0.1%) and PBS were used as positive and negative controls, respectively. Each sample was copied three times during the test. The percentage of hemolysis was calculated using the following equation.

$$\mathrm{Hemolysis}\mathrm{ratio}\left(\%\right)=\left(A_{\mathrm{h}}-A_{\mathrm{p}}\right)/\left(A_{\mathrm{t}}-A_{\mathrm{p}}\right)\times 100\%$$ (1)

A _h, A _p, and A _t represent the fractions of absorbance values for the supernatant samples, negative control (PBS), and positive control (Triton X‐100), respectively.

Histological Analysis

Wound samples were collected and frozen sections were utilized for H&E staining. The formation of granulation tissue and epithelialization in the control group, HG group, and RHG group was calculated. Additionally, Masson staining was performed simultaneously to examine collagen deposition. All images were obtained using a pathological scanning microscope (Leica).

Statistical Analysis

All experimental data in this study were subjected to statistical analysis using GraphPad Prism 9.0 software. The data were represented as the mean ± standard deviation (SD) of each replicate. Student's t‐test was employed to compare the means of two sample groups that exhibited a normal distribution and homogeneity of variance, Two‐way ANOVA analysis was used for comparing three or more groups. p < 0.05 was considered statistically significant.

Conflict of Interest

The authors declare no conflict of interest.

Supporting information

Supporting Information

Jin N., Wang Z., Tang X., Jin N., Wang X., Promoting Diabetic Wound Healing through a Hydrogel‐Based Cascade Regulation Strategy of Fibroblast‐Macrophage. Adv. Healthcare Mater. 2024, 13, 2400526. 10.1002/adhm.202400526

Data Availability Statement

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